Molecular Biotechnology: Principles and Applications - Glick B., Pasternak J. 2002

Molecular Biotechnology of Microbial Systems
Microbial Insecticides
Toxin synthesized by Bacillus thuringiensis

MECHANISM OF ACTION and application

The term “microbial insecticide” sometimes refers to a microorganism that either synthesizes a toxic substance that selectively acts on specific insects, or infects a target insect, leading to its death. The most studied, effective, and frequently used Microbial Insecticides are the Bacteria B. thuringiensis. They are represented by many strains and subspecies (subsp.), each synthesizing a toxin specific to certain insects (Table 15.1). For example, B. thuringiensis subsp. kurstaki is toxic to lepidopteran larvae (including moths and butterflies), skipper larvae, mermithids, and spruce budworm caterpillars. B. thuringiensis subsp. israelensis kills dipterans: mosquitoes and blackflies. B. thuringiensis subsp. tenebrionis (also known as san diego) is effective against coleopterans, including the Colorado potato beetle and the boll weevil. Other strains of B. thuringiensis have also been described, each toxic to specific insects. The insecticide (protein toxin) of B. thuringiensis subsp. kurstaki and other strains is present in The Cell as a so-called parasporal crystal—a Structure formed during bacterial sporulation. This structure does not serve any special biological function. It accounts for 20 to 30% of the dry weight of the sporulating culture and consists mainly of protein (~95%) and a small amount of CARBOHYDRATES (~5%). The crystal is actually a protein aggregate that dissociates into subunits in weak alkali. The subunits can be further dissociated in vitro by Treatment with ß-mercaptoethanol, which reduces Disulfide Bonds (Fig. 15.1).

Class="center">Table 15.1. Some properties of insecticidal toxins synthesized by various strains of B. thuringiensis 1)

¹⁾ From Lereclus et al., pp. 37–69, in Entwistle et al., (eds.), Bacillus thuringiensis, an Environmental Biopesticide: Theory and Practice, 1993.

All insecticidal toxins isolated from various strains of B. thuringiensis can be grouped into four major classes based on their toxicity: CryI, CryII, CryIII, and CryIV. CryI Proteins are toxic to lepidopterans, CryII to lepidopterans and dipterans, CryIII to coleopterans, and CryIV to dipterans. The classes can be further divided into subclasses (A, B, C, ...) and subgroups (a, b, c, ...) according to The nucleotide sequences of the corresponding toxin genes. For example, the cryI Gene class includes six subclasses (cryIA to F), and the cryIA subclass includes three subgroups [cryIA(a) to (c)]. Additionally, based on immunological properties, approximately 30 different serotypes of B. thuringiensis are distinguished (Table 15.1). Each serotype differs from the others by a specific set of antigenic determinants on the cell surface of a particular B. thuringiensis strain. In the parasporal crystal, the insecticide is usually in an inactive form: upon solubilization of the crystal, the protein is released as a protoxin, a precursor of the active toxin. The protoxin of the CryI toxin class has a molecular mass of approximately 130 kDa (Fig. 15.1). After the insect ingests the parasporal crystal, the protoxin is activated in the gut under alkaline pH conditions (7.5–8.0) and, by the action of specific digestive proteinases, is cleaved into an active toxin with a molecular mass of approximately 68 kDa (Fig. 15.1). In this form, it inserts into the membrane of the insect's gut epithelial Cells and forms an ion channel, through which a significant portion of cellular ATP is believed to leak (Fig. 15.2). Approximately 15 minutes after The formation of this ion channel, Cellular METABOLISM is blocked, the insect stops feeding, dehydration occurs, and death eventually ensues. Since The conversion of the protoxin to the active toxin occurs only under alkaline pH conditions and in the presence of specific proteinases, the likelihood of any harmful effect of the toxins on humans and farm animals is low.

Fig. 15.1. Schematic representation of a B. thuringiensis parasporal crystal consisting of the protein protoxin CryI. Each protein subunit has a molecular mass of 250 kDa and contains two Polypeptides with a molecular mass of 130 kDa each. Molecular masses were determined by Polyacrylamide gel Electrophoresis; rounded values are shown. The Conversion of the protoxin (130 kDa) to the active toxin (68 kDa) occurs only under weakly alkaline conditions (pH 7.5–8) in the presence of a specific proteinase (or proteinases). These are precisely the conditions found in the insect gut.

Fig. 15.2. Insertion of the B. thuringiensis toxin into the membrane of an insect gut epithelial cell and the formation of an ion channel.

The mode of action of B. thuringiensis toxins imposes certain limitations on their application. To kill the insect, B. thuringiensis must be ingested into its gut; otherwise, there will be no effect. B. thuringiensis is most commonly sprayed, and the bacteria are usually mixed with insect attractants to increase the likelihood that the pest will ingest the toxin. However, for insects living inside plant Tissues or on roots, B. thuringiensis toxin applied this way is unlikely to pose any threat. With this in mind, attempts have been made to develop alternative strategies for protecting plants against such pests. One approach is to generate Transgenic Plants that carry and express the B. thuringiensis toxin gene, thereby remaining protected from insect pests throughout the entire growing season.

The second limitation on The Use of the B. thuringiensis toxin is that it only affects the insect at a specific stage of development. It is at this precise moment that the treatment must be applied.

The strain B. thuringiensis subsp. kurstaki was isolated in 1901, but interest in it as a valuable commercial product arose only in 1951, and over the last ten years, this bacterium has become the primary tool for controlling spruce budworm caterpillar populations in Canada. In 1979, B. thuringiensis subsp. kurstaki was sprayed over only 1% of Canadian forests treated with insecticides to eradicate this insect (corresponding to approximately 2 million hectares); the remaining areas were treated with chemical insecticides. By 1986, the scale of B. thuringiensis subsp. kurstaki use had increased to 74%. In other countries, B. thuringiensis subsp. kurstaki is used to control tent caterpillars, gypsy moths, mermithids, cabbage moths, and hawkmoths. The main obstacle to even wider use of B. thuringiensis subsp. kurstaki is its high cost: the price of such a preparation is 1.5–3 times higher than that of chemical insecticides.

For the biocontrol of insect pest populations, 1,5 ∙ 10⁹—2,5 ∙ 10⁹ spores of B. thuringiensis subsp. kurstaki are sprayed per square meter of the treated area. The treatment is carried out when the number of larvae in the target insect population is at its maximum, as parasporal crystals are sensitive to sunlight and degrade rapidly. Under laboratory conditions in the light, more than 60% of Tryptophan residues in parasporal crystal proteins degrade within 24 h, whereas in the environment, depending on light levels, the crystals can persist from one day to one month. Such instability of the insecticidal protoxin implies that The Emergence of resistant insects is unlikely.

However, when B. thuringiensis subsp. kurstaki was used as an insecticide under low-light conditions (for example, in grain storage facilities), toxin-resistant insects emerged after several generations. One reason for this inherited resistance is a modification in a membrane protein of the gut cells, which normally Functions as the B. thuringiensis subsp. kurstaki toxin receptor. It probably arises because the protoxin does not degrade under these conditions and acts as a selective agent. Consequently, the emergence of insects resistant to B. thuringiensis subsp. kurstaki is most easily avoided by limiting the use of this microorganism to field conditions. However, given the scale of B. thuringiensis use, it cannot be ruled out that it may accumulate in the environment in quantities sufficient to trigger Selection mechanisms. And since B. thuringiensis is being used increasingly widely in various regions, the likelihood of resistant insect populations emerging will continue to grow.

Identification of toxin genes

To develop B. thuringiensis strains that synthesize insecticides more efficiently and have a broad host range, it was necessary to identify and characterize the protoxin gene(s), and first of all, to determine their localization: on a plasmid or in chromosomal DNA. To test the hypothesis of plasmid localization, a toxin-producing strain of B. thuringiensis can be conjugated with a strain lacking insecticidal activity. Chromosomal DNA transfer during conjugation is extremely rare, and if the 'defective' strain acquires The ability to synthesize the insecticide, it means the toxin genes are localized on a plasmid.

Standard Procedures are used to identify the gene encoding the protoxin.

B. thuringiensis is grown in culture and the cells are lysed. Total cellular DNA is isolated and centrifuged in a CsCl density gradient to separate plasmid and chromosomal DNA. If the protoxin genes are part of The Genome, a chromosomal DNA clone library is constructed. If they are contained in a plasmid, the plasmid DNA is fractionated by size using sucrose density gradient centrifugation. This enriches the plasmid fraction that will subsequently serve as the Starting Material for identifying the protoxin genes (Fig. 15.3). The protoxin gene of B. thuringiensis subsp. kurstaki is located on one of seven Plasmids with sizes of 2.0, 7.4, 7.8, 8.2, 14.4, 45, and 71 kb. To determine which one, plasmid DNA was centrifuged in a sucrose density gradient. Three fractions were isolated, concentrating small (2.0 kb), medium (7.4, 7.8, 8.2, and 14.4 kb), and large (45 and 71 kb) plasmids. Small plasmids were excluded from consideration because they could not encode a protein with a molecular mass of 130 kDa. The length of The nucleotide sequence encoding a protein of this size must exceed 4.0 kb. Medium and large plasmids were subjected to partial Digestion with the restriction endonuclease Sau3AI, and the fragments were inserted into the BamHI site of the plasmid pBR322. The resulting recombinant plasmids were used to transform E. coli cells, followed by immunological screening According to the following scheme:

1. Cell colonies were transferred from Agar to a nitrocellulose filter.

2. The transferred cells were partially lysed with organic Solvents.

3. All non-specific binding sites for Primary and secondary Antibodies on the filter were blocked using bovine serum albumin (BSA).

4. The BSA-treated filters were incubated with rabbit antibodies against the target insecticide.

5. The filters were washed to remove unbound antibodies and treated with 1²⁵¹-labeled Staphylococcus aureus protein A, which interacts with the Fc fragment of the bound antibodies.

6. Regions on the filter corresponding to colonies actively synthesizing the insecticide were detected using autoradiography.

Fig. 15.3. Isolation and Fractionation of plasmids, one of which carries the protoxin gene.

Using the identified protoxin gene as a Hybridization probe, it was established that the corresponding nucleotide sequence is contained in a 71-kb plasmid of B. thuringiensis subsp. kurstaki. A similar cloning and screening scheme was used to identify toxin genes localized on plasmids or, less frequently, in the chromosomal DNA of other B. thuringiensis strains.

Introduction/32.html">Genetic Engineering OF B. thuringiensis toxin genes

Following the identification of the B. thuringiensis toxin gene, the Primary Structure of its encoded protein was determined. Comparison of the Amino acid sequences of various protein toxins revealed that proteins from some strains share a common domain responsible for toxicity. In addition, a segment of the full coding sequence was subcloned to synthesize a truncated protein that fully retained its toxicity. Thus, subsequent genetic engineering manipulations can utilize the intact toxin gene, its fragment, or a chemically synthesized oligonucleotide.

Under natural conditions, most B. thuringiensis protoxins are synthesized only during sporulation, meaning that the parasporal crystal is formed only at a specific stage of the microorganism's development. If the toxin gene were expressed throughout the entire life cycle, it would be possible to significantly increase the yield of the toxin and reduce its synthesis time. Furthermore, this would allow toxin synthesis to be a continuous process, thereby substantially lowering the cost of the product, as continuous Fermentation utilizes smaller, and therefore less expensive, bioreactors and equipment compared to batch fermentation (for more details, see Chapter 16).

During B. thuringiensis sporulation, a specific Transcription initiation factor (sigma factor) binds to the promoters of genes that function only at this stage of the bacterial life cycle, resulting in the synthesis of sporulation-specific messenger RNAs (mRNAs). Consequently, to achieve continuous expression of the B. thuringiensis insecticidal gene(s), it (they) must be placed under the control of a promoter that functions throughout the entire life cycle.

To achieve this, a DNA fragment containing the toxin gene without its own promoter was inserted into a plasmid under the control of an active constitutive promoter of the tetracycline resistance gene, which had previously been excised from a Bacillus cereus plasmid and introduced into B. thuringiensis. A fully functional toxic protein was continuously synthesized at all Stages of the microorganism's development (Fig. 15.4). Furthermore, when a sporulation-deficient mutant strain of B. thuringiensis was transformed with this construct, the toxin was still synthesized. Moreover, the process was much more efficient than in the case of wild-type B. thuringiensis: the yield of the product was higher, while the substrate consumption and synthesis time were significantly lower. A further improvement of this system could involve integrating the toxin gene, expressed throughout The life cycle, into the chromosomal DNA of a sporulation-deficient B. thuringiensis strain. This would ensure gene stability during continuous fermentation, which is not always achieved with plasmid localization due to plasmid instability.

Fig. 15.4. Cloning of a B. thuringiensis subsp. kurstaki toxin gene fragment under the control of the tetracycline resistance gene promoter (pTet). The native promoter is removed from the isolated B. thuringiensis gene using restriction Enzymes RE1 and RE2. The resulting fragment is inserted into a plasmid vector adjacent to the pTet promoter, replacing the tetracycline resistance gene removed with restriction enzymes RE1 and RE2, and ligated using T4 DNA ligase.

Unlike most other B. thuringiensis toxin (cry) genes, the expression of the cryIIIA gene is normally controlled by a vegetative promoter rather than a sporulation-specific promoter. The cryIIIA gene encodes a toxin effective against coleopteran larvae. As a result of transforming a sporulation-deficient mutant strain of B. thuringiensis with a plasmid carrying the cloned cryIIIA gene, the toxin was synthesized more efficiently and was more stable than when synthesized in the wild-type strain. Based on this result, researchers hypothesized that overexpression of other cry genes, which are normally expressed only during sporulation, could be achieved by placing them under the control of the cryIIIA promoter and transforming a sporulation-deficient mutant strain of B. thuringiensis with the resulting construct.

Since many crops are damaged by several insect species simultaneously, developing microbial insecticides targeted against a broad spectrum of insect pests would be extremely beneficial. A broad-spectrum toxin can be obtained in two ways: 1) by transferring a given toxin gene (for example, a toxin effective against dipterans) into a B. thuringiensis strain that synthesizes another species-specific toxin (for example, one effective against coleopterans); 2) by fusing portions of two genes encoding different species-specific toxins to create a sequence that encodes a unique dual-action toxin (hybrid toxin).

To evaluate the possibility of obtaining a broad-spectrum toxin, the toxin genes of B. thuringiensis subsp. aizawai and tenebrionis were cloned into shuttle vectors capable of replicating in both B. thuringiensis and Escherichia coli. These genetic constructs were then introduced via electroporation into B. thuringiensis subsp. aizawai, kurstaki, israelensis, and tenebrionis. The toxicity of the resulting transformed strains was tested on larvae from three insect orders.

In all cases, the toxicity mediated by the host's endogenous toxin gene(s) was retained, and in most instances, the introduced gene conferred the toxic Specificity characteristic of the donor strain from which it was isolated (Table 15.2). Furthermore, one experiment yielded a highly surprising result: when the toxin gene from B. thuringiensis subsp. tenebrionis was introduced into B. thuringiensis subsp. israelensis, the resulting transformants exhibited some degree of toxicity toward Pieris brassicae (cabbage white butterfly), which was unaffected by any of the products of the parental genes.

Plasmid Vectors carrying cloned cry genes are often unstable in B. thuringiensis; even in the absence of selective pressure, some or all of them are lost. Interestingly, in nature, most cry genes are localized precisely on plasmids. This plasmid instability underscores the feasibility of integrating the cloned cry gene(s) into the chromosomal DNA. Let us consider one successful example of such integration.

Table 15.2. Toxicity of wild-type and recombinant subspecies of B. thuringiensis to the insects Pieris brassicae (cabbage butterfly), Aedes aegypti (mosquito), and Phaedon cochleariae (beetle)¹⁾

Toxicity to²⁾

¹⁾ From Crickmore et al., Biochem. J. 270: 133—136, 1990.

²⁾ Symbols: ++, 0 to 5% leaf damage (for Phaedon and Pieris) or 100% insect mortality within 1 h (Aedes); +, 5 to 50% leaf damage (Phaedon and Pieris) or 50 to 100% mortality within 24 h (Aedes); —, more than 50% leaf damage (Phaedon and Pieris) or no mortality within 24 h (Aedes). Experiments were conducted on cabbage leaves (for Pieris) or turnip leaves (for Phaedon).

The strain B. thuringiensis subsp. kurstaki typically contains five different toxin genes: cryIA(a), cryIA(b), cryIA(c), cryIIA, and cryIIB. Their products are toxic to various lepidopterans but are ineffective against Spodoptera spp. The cryIC gene, which is normally present only in B. thuringiensis subsp. aizawai and subsp. entomocidus, was integrated into the chromosomal DNA of the B. thuringiensis subsp. kurstaki strain. The transformed B. thuringiensis subsp. kurstaki strain was six times more effective against Spodoptera exigua larvae than the wild-type strain.

As previously mentioned, another approach to obtaining a broad-spectrum toxin involves fusing the coding regions of two different toxin genes. This possibility has been tested in the laboratory. Several hybrid toxins targeting only lepidopterans were developed; some of these toxins were more effective than the products of either parental gene, and in one case, the hybrid protein exhibited an entirely novel biological activity.

Three domains are responsible for the toxic activity of the Cry protein. Domain I, located in the N-terminal region of the protein molecule, mediates the specific binding of the toxin to a receptor On the surface of the insect's midgut epithelial cells. Domain III, located in the C-terminal region of the molecule, is presumably responsible for toxicity. Resistance to B. thuringiensis toxins is typically caused by mutational change(s) in the receptor protein(s) on The surface of the insect's gut cells, which prevent the receptor from recognizing the Cry protein. However, if the toxin gene is modified so that the toxin can bind to other surface proteins, the likelihood of resistance developing will decrease.

The CryIC and CryIE proteins are toxic to lepidopterans but exhibit different species specificities: CryIC is active against S. exigua, Mamestra brassicae, and Manduca sexta, whereas CryIE is active only against M. sexta. In one laboratory, a hybrid CryIC-CryIE protein was created, and its toxicity was tested on various insects. Its ability to bind to different receptors was also investigated (Fig. 15.5). The hybrid toxin G27, containing domain III of the CryIC protein, was toxic to S. exigua larvae, even though it bound only to the CryIE receptor and not to the CryIC receptor. Conversely, the hybrid toxin F26 had no effect on S. exigua larvae, despite binding to the CryIC receptor. Because the CryIC and G27 proteins, which are toxic to S. exigua, bind to different receptors on the surface of the insect's gut cells, simultaneous or sequential treatment of S. exigua with these two toxins could reduce the likelihood of a resistant strain emerging, as this would require mutational changes to occur simultaneously in two different proteins.

Fig. 15.5. Toxicity of CryIC, CryIE proteins and hybrid toxins G27 and F26, and their receptor-binding specificity. Binding specificity was determined in toxin-receptor interaction experiments. Unlabeled CryIC or CryIE was added to the complex of the protein receptor on the surface of S. exigua gut cells and radiolabeled toxin, and the binding of the labeled toxin was determined. (From Bosch et al., Bio/Technology 12:915-918, 1994.)

The insecticide produced by B. thuringiensis subsp. israelensis exerts its toxic effects when ingested by mosquito larvae. However, if this toxin, which exists as parasporal crystals, is sprayed over Water, the crystals rapidly sink, removing the toxin from the mosquito larvae's food chain. To solve this problem, the toxin gene can be introduced into an Organism that serves as food for mosquito larvae. These could be, for example, photosynthetic cyanobacteria Synechocystis and Synechococcus spp., which proliferate in the surface water layer where sunlight is abundant and where the larvae typically reside.

Another organism that can be used to express B. thuringiensis toxin genes is the bacterium Caulobacter crescentus, which is widespread in the aquatic environments where mosquito larvae live. The insecticide synthesized by transformed cyanobacteria or C. crescentus was toxic to mosquito larvae under laboratory conditions. However, in the field, the transformed cyanobacteria and C. crescentus died rapidly, and the expression levels of the cloned genes were very low.

The Gram-negative aerobic bacterium Asticcacaulis excentricus, which inhabits the surface of water bodies, can also be used as a host organism for foreign cry genes encoding proteins toxic to mosquitoes. Experiments were conducted in which A. excentricus was transformed with a broad-host-range plasmid vector carrying the protein toxin genes of Bacillus sphaericus (a bacterium similar to B. thuringiensis). The genes were under the control of the tacI promoter, a variant of the tac promoter. The resulting transformant synthesized protein toxins with molecular masses of 51 and 42 kDa and was almost as toxic to Anopheles and Culex mosquito larvae as the highly toxic wild-type strains of B. sphaericus. However, unlike B. sphaericus, A. excentricus did not present settling problems when sprayed over water bodies. Furthermore, culturing A. excentricus is much cheaper because this microorganism grows on a simpler medium than B. sphaericus and B. thuringiensis. It is characterized by low proteinase activity, so the toxin does not undergo immediate degradation. A. excentricus is well adapted to conditions such as relatively high UV light intensity. However, before recombinant A. excentricus bacteria can be used to control mosquito populations in nature, it must be ensured that the integrated toxin genes do not contain sequences determining Antibiotic Resistance.

Insecticides produced by B. thuringiensis, when applied to leaves and stems, do not affect insects that damage plant roots. To circumvent this difficulty, the B. thuringiensis toxin gene can be introduced into a strain of a bacterial species that inhabits the soil layer immediately adjacent to the roots (the rhizosphere). When introduced into the soil, such recombinant bacteria will secrete the insecticidal toxin directly into the rhizosphere, protecting the roots from insects for as long as they remain in the soil. This eliminates The Need for repeated treatment of plants with biological or chemical insecticides.

The toxin gene of B. thuringiensis subsp. kurstaki was integrated into the chromosomal DNA of a Pseudomonas fluorescens strain that colonizes corn roots as follows (Fig. 15.6).

1. Transposon Tn5 was inserted into a plasmid, its left and right flanking sequences were genetically modified, and the transposase gene was deleted. Such a modified transposon cannot excise from the plasmid even in the presence of exogenous transposase.

2. The B. thuringiensis subsp. kurstaki toxin gene was inserted into the middle of the modified Tn5 transposon so that it was under the control of a constitutive promoter.

3. Wild-type transposon Tn5 was integrated into the chromosomal DNA of the ROOT-colonizing P. fluorescens strain.

4. The plasmid carrying the modified Tn5 transposon with the integrated toxin gene was introduced into the bacterium that had the wild-type Tn5 transposon integrated into its chromosomal DNA.

5. Homologous Recombination via double crossover was performed between the non-transposable Tn5 element carrying the toxin gene and the wild-type Tn5 transposon integrated into the chromosome. As a result of this integration, the modified Tn5 transposon with the toxin gene became integrated into the chromosomal DNA, while the wild-type Tn5 transposon was eliminated.

Fig. 15.6. Generation of a recombinant P. fluorescens strain with the B. thuringiensis toxin gene integrated into its chromosomal DNA. The gene is inserted into a non-excisable Tn5 transposon located on a plasmid. This construct is introduced into a P. fluorescens strain containing a wild-type Tn5 transposon in its chromosomal DNA. As a result of homologous recombination, the non-excisable Tn5 transposon carrying the B. thuringiensis toxin gene becomes integrated into the P. fluorescens chromosome.

In this form, the toxin gene is unlikely to be lost during large-scale cultivation of the recombinant microorganisms under laboratory conditions or after their release into the environment. Furthermore, the likelihood of transferring this gene to other Microorganisms in the environment is extremely low. Preliminary studies have shown that the recombinant P. fluorescens strain is toxic to hornworm larvae. Future plans include testing the ability of this recombinant microorganism to minimize root damage by insect pests in greenhouse and field trials.

B. thuringiensis toxin genes have been introduced into the chromosomal DNA of A wide variety of microorganisms. For instance, cryIA(c) genes were introduced into the DNA of P. fluorescens, which protects sugarcane plants against Eldana saccharina. Transformation with this gene of the bacterium Clavibacter xyli subsp. cynodontis, which normally inhabits the xylem of Bermuda grass, enables the recombinant bacteria to protect corn plants against the European corn borer (Ostrinia nubilalis).