- freely available
Insects 2014, 5(1), 139-166; doi:10.3390/insects5010139
Published: 6 January 2014
Abstract: This review focuses on common insecticidal virulence factors from entomopathogenic bacteria with special emphasis on two insect pathogenic bacteria Photorhabdus (Proteobacteria: Enterobacteriaceae) and Bacillus (Firmicutes: Bacillaceae). Insect pathogenic bacteria of diverse taxonomic groups and phylogenetic origin have been shown to have striking similarities in the virulence factors they produce. It has been suggested that the detection of phage elements surrounding toxin genes, horizontal and lateral gene transfer events, and plasmid shuffling occurrences may be some of the reasons that virulence factor genes have so many analogs throughout the bacterial kingdom. Comparison of virulence factors of Photorhabdus, and Bacillus, two bacteria with dissimilar life styles opens the possibility of re-examining newly discovered toxins for novel tissue targets. For example, nematodes residing in the hemolymph may release bacteria with virulence factors targeting neurons or neuromuscular junctions. The first section of this review focuses on toxins and their context in agriculture. The second describes the mode of action of toxins from common entomopathogens and the third draws comparisons between Gram positive and Gram negative bacteria. The fourth section reviews the implications of the nervous system in biocontrol.
For many decades, naturally occurring microbial insect pathogens, including bacteria, fungi, microsporidia, protists, viruses, and nematodes have been considered as non-chemical alternatives for suppression of insect pests [1,2,3]. Positive attributes such as their high specificity and less damage to non-target fauna and flora has certainly contributed to the consideration in management practices of agricultural and forest insect pests .
Of all entomopathogens, bacteria have been the most extensively used organisms to date. Indeed, Bacillus thuringiensis (Bt) (Firmicutes: Bacillaceae), originally discovered in 1901 by Ishiwata  and later rediscovered by Berliner , has been the most studied and broadly used in microbial control. In particular, the discovery that Bt spore-associated toxins are extremely virulent and can persist in the environment with high potency, prompted the development of bacterial spray formulations  and transgenic plants expressing bacterial toxins [8,9]. Advancements in the characterization of bacterial pathogens including purification and culturing methods, molecular identification of virulence factors, and whole genome characterization and comparisons have prompted the discovery of novel pest management tools. In this vein, insecticidal molecules expressed and secreted by various entomopathogenic bacteria have been targeted for genetic manipulation to enhance toxicity [10,11].
Recently, other insect pathogenic bacteria with modes of action similar to Bt have been hailed as agriculturally relevant . Certainly, insect pathogenic bacteria of diverse taxonomic groups and phylogenetic origin have been shown to have striking similarities in the virulence factors they produce. . Bacterial virulence factors are often encoded on mobile genetic elements, such as plasmids and bacteriophages, and can easily be spread through horizontal gene transfer. The discovery that both Gram-negative and Gram-positive bacteria produce analogous insect-specific toxins infers history of gene transfer between these organisms . For example, Photorhabdus luminescens (Proteobacteria: Enterobacteriaceae), the bacterial symbiont of the entomopathogenic nematode Heterorhabditis bacteriophora (Rhabditida: Heterorhabditidae), has been shown to have similar virulence factors to that of Bt .
Herein, we review the commonalities between virulence factors produced by entomopathogenic bacteria. Special emphasis is placed on those with potential for control of lepidopteran pests. Additionally, we discuss and propose novel tissue targets of virulence factors for their potential application in pest management.
2. Toxins Produced by Entomopathogenic Bacteria
Insecticidal toxins used in agriculture are predominantly from Gram-positive bacteria and derived mostly from Bacillus thuringiensis. Foliar sprays containing B. thuringiensis represent an organic alternative to synthetic foliar sprays. Transgenic plants expressing Cry toxins are now viewed as an overwhelming ecological success .
Entomopathogenic Gram-negative bacteria also produce toxins. Based on their targeted tissues, they can be categorized into three types (a) neurotoxins; (b) digestive toxins; and (c) cytotoxins. Members in the Enterobacteriaceae such as Photorhabdus, Xenorhabdus, Serratia, and Yersinia spp. produce insecticidal toxins with oral toxicity similar to that of Bt toxins, but have not yet been fully utilized.
This section describes the virulence factors associated with both Gram-negative and Gram-positive bacteria as well as their mode of action.
2.1. Insecticidal Toxin Genes Produced by Gram Positive Bacteria
Bacillus spp.: Within this genus three species, B. thuringiensis, Bacillus cereus, Bacillus anthracis are the most widely studied taxa in terms of insecticidal toxins. Recent phylogenetic studies based on 16S and 23S rRNA sequence data consider B. thuringiensis, together with B. cereus, B. anthracis and Bacillus mycoides (Firmicutes: Bacillaceae) nearly identical. However, their unique pathogenicity properties and the diverse modes of actions of their insecticidal toxins (Table 1) support their distinctiveness, giving each separate species names .
Bacillus thuringiensis is a phylloplane and soil-dwelling bacterium that is aerobic and endospore-forming. It has been isolated from many insect carcasses and a few plants, where it has been reported as an endophyte. Upon sporulation, B. thuringiensis forms crystals of proteinaceous insecticidal δ-endotoxins, called crystal proteins or Cry proteins, which are encoded by cry genes . The Cry toxins, are insecticidal to many insect orders including Lepidoptera, Diptera [18,19], Coleoptera , and Hymenoptera , and also have activity against nematodes [22,23]. Upon ingestion by Lepidoptera larvae, Cry protoxins are first solubilized by the alkaline pH of the gut, then proteolytically activated by proteases. The insecticidal form of these toxins has been shown to bind to specific receptors on the columnar cells of the midgut epithelium causing pore formation and midgut cell death. Cry toxins are commonly classified as gut poisons, as they compromise the epithelial-hemocoel layer and ultimately lead to starvation and/or septicemia and larval death . Cry toxins bind, form pores, and damage midgut epithelial goblet cells in mixed midgut cell cultures  and in vivo  and have also been shown to bind to the insect’s peritrophic matrix . The Cry toxins from B. thuringiensis are almost exclusively considered digestive toxins; however, they have homology to neurotoxins and effect diverse tissues of lepidopteran larvae. They have been shown to kill larval neurons of the cerebral ganglia in vitro , invade liposomes causing morphological changes to lipid bilayers in vitro , initiate apoptosis-like pathway in ovary-derived cells expressing a Bt receptor , and bind ATP-binding cassette (ABC) transporters . In this respect, mutated ABC transporters are correlated with resistance to Bt toxins in domesticated silkworm, Bombyx mori (Lepidoptera: Bombycidae) , tobacco budworm, Heliothis virescens (Lepidoptera: Noctuidae) , diamond back moth, Plutella xylostella (Lepidoptera: Plutellidae), and cabbage looper, Trichoplusia ni (Lepidoptera: Noctuidae) .
|Table 1. Mode of action of entomopathogenic bacterial toxins with toxicity to lepidopteran larvae.|
|Bacillaceae||B. thuringiensis||Crystal toxins produced during sporulation, pore forming toxins that act on gut columnar cells||Cry toxins|
|Pore forming toxin that acts on gut columnar cells, causes death to insect brains cells||Cry1C toxin|
|Vegetative insecticidal protein produced during the vegetative growth cycle, binds to midgut receptors, gut paralysis, lysis of midgut columnar cells||Vip3 toxin|
|Associated with the presence of certain cry and vip genes, toxic to mammalian cells, growth inhibition, secreted during vegetative growth cycle||Β-exotoxin I|
|B. thuringiensis israelensis||TccC and TcaCBA toxin homologs to Photorhabdus and Yersinia||Strain bthur0013|
|B. subtilis||Biosurfactant, destructive effect on gut epithelial cells by vesicle formation in the apical region and cellular vacuolization||SPB1|
|B. cereus||Secreted non-proteinous exotoxins, pathogenic to humans and insects||Exotoxin|
|L. sphaeriscus||Cholesterol binding cytolysin, lyses neurons and hemocytes||Sphaericolysin|
|Clostridiaceae||C. botulinum||Binary actin binding-ADP-ribosylating exotoxin||C2|
|C. difficile||Exotoxin which reorganizes cytoskeleton, hydrophobic central domain is shared between the Mcf1 and Mcf2 toxins, cytotoxic motif||Cytotoxin B|
|Enterobacteriaceae||S. entomophila||Shows high similarity to P. luminescens TcbA, TcdA, TcaB, and TccB, tripeptide cell-binding motif Arg-Gly-Asp implicates the ability to bind to eukaryotic cells||SepA|
|Shows similarity to P. luminescens TcaC and SpvB||SepB|
|Shows similarity to P. luminescens TccC, Shows similarity to Bacillus subtilis WapA, E. coli Rhs, Streptomyces coelicolor A3, Coxiella burnetii||SepC|
|Antifeeding prophage causes cessation of feeding||Afp|
|Y. frederiksenii||Orally toxic, not subcutaneously toxic, high homology to sep genes in Serratia||TcYF1 and TcTF2|
|P. luminescens||Assists TcC to translocate into the cell, transmembrane pore formation, contains an integrin binding motif||TcA (TcdA1)|
|Potentiates expressed toxin genes tcdA1, tcaA, and tcaB||TcdB1, TccC1, TcaC, TcdB2, TccC3|
|Assists TcC (TccC3) to translocate in the cell||TcB (TcdB2)|
|Actin-clustering, defects in phagocytosis and cell death||TcC (TccC3)|
|Formation of actin aggregates, ADP-ribosylates actin at the threonine-148 with TccC3 sub-complex||Tc Complex|
|Makes caterpillars floppy phenotype, induce apoptosis, rearrange actin||Mcf|
|Degrades peritrophic matrix, causes midgut damage, midgut cell sloughing, fat body nuclear degradation||Txp40|
|Photorhabdus virulence cassettes, phage-like, kills and condenses actin of hemocytes||PVC|
|Primarily hemoplymph-based insecticidal activity||pirA2B2, locus plu4437-plu4436|
|Similar to Serratia-type hemolysins||phlBA operon|
|Binds actin ADP-ribosylating, inhibits actin polymerization||Photox|
|X. nematophila||Midgut, intestinal sloughing||A24tox|
|Fed to neonates and caused inhibitory growth||Xin|
|From the cosmid CHRIM1, analogs to sep Serratia genes, oral toxicity||XptA1, XptA2, XptB1, XptC1|
|Alphaxenorhabdolysin triggers apoptosis in hemocyte cells, cytotoxic and hemolysin effects, has analogs in Photorhabdus, P. entomophila, P. syringae, Y. enterocolitica, and Proteus mirabilis||xaxAB|
|High similarity to GroEL, injectable toxicity to G. mellonella, innate immune response of increased phenoloxidase activity stimulated by injection||HIP57|
|One gene in the xenocin operon with RNAse and cytotoxic activity||xciA|
|Pseudomonadaceae||P. fluorescens||Related to the Mcf from P. luminescens, hemolymph-based insecticidal activity||Fit|
|Regulators of Fit insect toxin expression for biocontrol||FitG|
|P. entomophila||TccC-type toxins||PSEEN2485, PSEEN2697, PSEEN2788|
|Tcc-C type toxins||PSEEN701, PSEEN702|
|Exotoxins with hemolytic activity||PSEEN3925, PSEEN0968, PSEEN3843|
|Lipases||PSEEN709, PSEEN1065, PSEEN2195, PSEEN3432|
|P. syringae||Tcc-C type toxins||PSEEN701, PSEEN702|
Bacillus cereus has been mostly studied for its role in food contamination and human digestive food poisoning; however, this bacterium associates with soil and plants much like Bt . As mentioned above, B. cereus and B. thuringiensis are two very closely related taxa, however they produce different types of toxins. Unlike Bt, the endospore of B. cereus is not insecticidal. Both B. thuringiensis and B. cereus produce non-proteinous insecticidal exotoxins during their vegetative growth cycle. β-Exotoxin I is produced by B. thuringiensis and a small, proteinous exotoxin is produced by B. cereus . Although, B. cereus has been shown to grow and proliferate in the insect gut, this bacterium is mostly regarded as an opportunistic pathogen with the production of virulence factors that are most effective when titers are high. Entomopathogenic toxin genes have also been found in the genome of strain B. thuringiensis israelensis bthur0013, a member of the B. cereus sensu lato .
Other Bacillus species known to produce insecticidal components are Bacillus circulans (Firmicutes: Bacillaceae) and sphaeriscus (Firmicutes: Bacillaceae) (=Lysinibacillus sphaeriscus). Virulence factors produced by the former sp. have been shown to affect insects (mostly dipterans) and other invertebrates such as nematodes, and mollusks . Sphaericolysin is a toxin produced during the vegetative growth cycle of L. sphaeriscus . This toxin has heamoceolic toxicity toward Blattela germanica (Blattodea: Blattellidae) and Spodoptera litura (Lepidoptera: Noctuidae) .
Clostridium spp.: Clostridium are anaerobic spore-forming bacteria and, similar to Bacillus, comprise over one hundred species. Similar to Bacillus, Clostridium spp. produce binary proteinous toxins that are proteolytically activated by serine proteases  (Table 1). For example, Clostridium bifermentans serovar malaysia produces larvacidal toxin proteins that are active against mosquitoes . Clostridium perfringens (Firmicutes: Clostridiaceae) has an iota toxin which binds actin by ADP-ribosylation and has a C-domain structure similar to that of the vegetative insecticidal protein of Bt, VIP2, although this protein ultimately targets mammals . The Clostridium difficile cytotoxin B is an exotoxin which causes the reorganization of cytoskeletons, similar to Mcf (Makes Caterpillars Floppy) toxins produced by Gram-negative P. luminescens (See Section 2.2). Moreover, they have a three domain structure consisting of receptor-binding, translocation, and one catalytic domain .
The actin binding C2 toxin of another Clostridium sp., C. botulinum, is a binary toxin with four domains of activation, pore formation, and receptor recognition. The C2 toxin is similar to the toxin complex, TccC3, produced by P. luminescens because they both have ADP-ribosylating activity and bind actin. One difference between these two toxins is that TccC3 forms a complex with other Tc toxins, TcdA1 and TccC3, and is therefore not binary. Although ADP-ribosylation occurs in both cases, TccC3 ribosylates at a threonine while C2 toxins ribosylate at arginine, dysfunctioning actin by inducing actin polymerization in the former and inhibition in the latter . A toxin found in P. luminescens has an identical mode of action to that of C2 toxin. The translated product of the plu0822 gene, termed Photox toxin, stops actin polymerization by targeting an arginine with ADP-ribotransferase activity .
2.2. Insecticidal Toxin Genes Produced by Gram Negative Bacteria
Photorhabdus spp.: Until now three species have been identified within this genus: P. luminescens, Photorhabdus temperata and Photorhabdus asymbiotica . All Photorhabdus species are mutualistically associated with insect parasitic Heterorhabditis nematodes. However, P. asymbiotica has also been found associated with skin injuries in human patients . This bacterium is currently considered an emerging human pathogen model system . Unlike B. thuringiensis, Photorhabdus spp. are facultative anaerobes and cannot live freely in the soil environment. These bacteria are vectored by the nematodes and together they form an insecticidal complex that kills the insect and use the carcass for reproduction and nutrition. Once delivered by the nematodes in the insect hemocoel, the bacteria first evade the insect immune system and then produce toxins which kill the insect and break down insect epithelial tissues. The Photorhabdus genome contains a multitude of pathogenicity islands with an abundance of toxin genes . The major virulence factors characterized so far consist of the mcf1 and mcf2 (makes caterpillars floppy) genes, the tc (toxin complex) genes, Pir (Photorhabdus insect related) operon, and a multitude of virulence factors associated with the Photorhabdus virulence cassettes (PVC) (Table 1) .
The Mcf toxins are known to both rearrange actin cytoskeletons and induce apoptosis in both hemocytes and epithelial tissue, resulting in an abundance of tissue damage. The intensity of the tissue damage is so great that there is a complete loss of turgor pressure throughout the infected insect .
The Tc proteins, similar to Bt Cry toxins, are orally toxic compounds that have been discovered to be insecticidal to a wide range of insect taxa including Coleoptera, Lepidoptera, and Hemiptera. It has been shown that P. luminescens is highly pathogenic when injected into lepidopteran hosts such as the African cotton leafworm, Spodoptera littoralis (Lepidoptera: Noctuidae), and P. xylostella. The oral toxicity of the Tc proteins is peculiar because Photorhabdus are delivered directly into the hemocoel of the target insect. Furthermore, the Tc proteins are active in the lumen side of the insect’s midgut epithelium and not in the basal side of this tissue, which would be the expectation for a hemocoel-initiating pathogen .
Pir proteins are other Photorhabdus toxins that have been identified to have hemolymph  and oral  toxicity. The Pir proteins have homology to a neurotoxin commonly called leptinotarsin [55,56,57] and are binary . The binding and destructive effects on neural tissue could be a major factor in toxicity when Photorhabdus are injected into susceptible insects.
It can be speculated that there are many more toxins yet to be characterized from the Photorhabdus genome which are responsible for hemolymph-based toxicity. The Txp40 protein has been identified in 59 different strains of both Photorhabdus and Xenorhabdus. It has injectable toxicity to many lepidopteran insects including the greater wax moth, Galleria mellonella (Lepidoptera: Pyralidae), Indian meal moth, Plodia interpunctella (Lepidoptera: Pyralidae), corn earworm, Helicoverpa armigera (Lepidoptera: Noctuidae), and the Dipterans such as the Australian sheep blowfly, Lucilia cuprina (Diptera: Calliphoridae) . A txp40 gene, when expressed by E. coli, has also been shown to be insecticidal to P. xylostella . The Txp40 protein is involved in causing damage to both the insect midgut and the fat body and has also been shown to exhibit cytotoxicity in vitro in dipteral and lepidopteran cell lines .
Xenorhabdus spp.: Similar to Photorhabdus, bacteria in this genus are also non-free-living, though they are symbiotically associated with nematodes in the genus Steinernema . Similar to Heterorhabditis, Steinernema nematodes also play a key role in vectoring these bacteria from one insect host to another. Xenorhabdus also produce a large number of insecticidal toxins to help them garner nutrients from the insect hosts (Table 1). One example is a toxin from Xenorhabdus nematophila (Proteobacteria: Enterobacteriaceae) called A24tox. A24tox is a 42 kD protein that has been shown to kill G. mellonella and H. armigera. This toxin has a hypothetical homolog in Photorhabdus but has no significant matches outside of this group . Another example is the xenocin operon. The xenocin operon consists of two genes, xciA and ximB. When expressed, these molecules are secreted by the flagellar type II secretion pathway. Xenocin, the xciA gene, has RNAse activity and cytotoxicity. The immunity protein co-expressed in the xenocin operon has antimicrobial effects killing competing microbes in insect larvae .
The xpt gene products from X. nematophila have oral insecticidal activity to different lepidopteran larvae based upon how the differentially expressed xpt genes interact with each other. In this respect, the translated products of Tc genes are similar to the Tc proteins from Photorhabdus. The Tc gene products can be categorized into toxins and potentiators. Potentiators synergize with their Tc toxin counterpart for full insecticidal activity . When xptA1, xptA2, xptB1, and xptC1 genes are expressed in Escherichia coli and are fed to larvae, they confer oral insecticidal toxicity to various lepidopteran spp. including: cabbage butterfly, Pieris brassicae (Lepidoptera: Pieridae), white butterfly, Pieris (Lepidoptera: Pieridae), and H. virescens. Interestingly, when the interactions of all four genes are separated from each other their insecticidal toxicity is different. For example, E. coli expressed xptA1, xpbtB1, and xptC1 are needed to be insecticidal toward P. rapae and P. brassicae. The addition of xptA2 and the removal of xptA1 is all that is needed for insecticidal toxicity toward H. virescens. Thus, different combinations of the xpt E. coli lysates show different insecticidal specificities to different lepidopteran larvae . The intracellular Xin toxic protein from X. nematophila has also been demonstrated to have an inhibitory growth effect on cotton bollworm, Helicoverpa (Lepidoptera: Noctuidae), after ingestion with artificial diet incorporating the Xin protein .
The xaxAB genes from X. nematophila expressed in E. coli induce apoptosis when incubated with hemocytes derived from S. littoralis. Additionally, the order in which Xax toxins (XaxA and XaxB) are added to cells in vitro affects their toxicity . This is similar to xpt and tc genes that are expressed in E. coli.
Xenorhabdus bacteria also produce an insecticidal protein called HIP57 that have high similarity to chaperonins like GroEL produced by E. coli. GroEL chaperonins have been shown to help proteins fold from non-native to native structures . Moreover, they help combat problems such as aggregation when nascent proteins have hydrophobic residues exposed before reaching a fully folded native state . HIP57 has been shown to have injectable toxicity to G. mellonella. The insecticidal property of HIP57 is a novel function for the GroEL superfamily of proteins. Moreover, an innate immune response of increased prophenoloxidase activity is stimulated by HIP57 injection .
Serratia spp.: These bacteria often exist as endophytes possessing fungicidal properties, but can also associate with insects  and nematodes [70,71] in a facultative manner. Genome studies have found several insecticidal genes in the Serratia genome (Table 1). Other Serratia sp. are responsiblefor causing amber disease in grass grubs, Costelytra zealandica (Coleoptera: Scarabaeidae) . Contrastingly, Serratia marcescens (Proteobacteria: Enterobacteriaceae) infects other host such as poorly reared H. virescens . The pADAP plasmid from Serratia entomophila contains the genes sepA, sepB, and sepC. These genes are similar to the Tc genes described in P. luminescens and the xpt genes identified from X. nematophila. There is no need for the entire pADAP plasmid to be associated with the sep genes for amber disease to occur and cause death. However, when the sep genes are expressed in E. coli without the entire plasmid, the scarab beetles do not cease feeding  which is one symptom of amber disease. The virulence factor associated with pADAP that causes cessation of feeding in amber disease is the anti-feeding prophage (Afp) . Therefore both the sep genes and Afp are needed for full virulence of Serratia in grass grubs.
Yersinia spp.: Members of this genus are facultative anaerobes. In particular, Yersinia pestis the causative agent of bubonic disease (aka plague) is associated with fleas, humans and rodent intermediates. Two other species which are pathogenic to humans are Yersinia enterocolitica and Yersinia pseudotuberculosis often causing diarrheal disease and fever with inflammation, respectively. Yersinia entomophaga (Proteobacteria: Enterobacteriaceae), Yersinia frederiksenii , which can cause disease in grass grubs, Y. enterocolitica , Y. pseudotuberculosis, Yersinia mollaretii (Proteobacteria: Enterobacteriaceae)  and Y. pestis  have insecticidal genes of the Tc super family.
The pathogenicity island YAPIYe of Y. enterocolitica strain W22703 contains the homologues tcaA, tcaB, tcaC, and tccC and is insecticidal to the tobacco hornworm, Manduca sexta (Lepidoptera: Sphingidae), when fed orally; a tcaA mutant of the same Y. enterocolitica strain lost insecticidal lethality to M. sexta . Two Y. enterocolitica strains, W22703 and WA314, were shown to have nematocidal activity against C. elegans when the strains contained TcaA, though the E. coli strain DH5α expressing TcaA clones did not cause pathogenicity in C. elegans, implicating additional virulence factors are necessary for full nematicidal activity . The pathogenicity island of Y. entomophaga, termed PAIYe96, is composed of insecticidal chitinases and Tc components that cause virulence to C. zealandica. Liquid culture supernatants of Y. entomophaga were also shown to be insecticidal to the redheaded cockchafer, Adoryphorus couloni (Coleoptera: Scarabaeoidae), blackheaded pasture cockchafer, Acrossidius tasmaniae (Coleoptera: Scarabaeidae), and P. xylostella. In the case of P. xylostella, initial apical swelling of gut columnar cells occurred after ingestion of purified Tc from Y. entomophaga, followed by complete dissolution of the gut lining .
Pseudomonas spp.: The renowned anti-fungal properties of root-associated bacteria of the genus Pseudomonas have been widely used for crop protection and further investigated for viable natural products in both agriculture and medicine. The insecticidal properties (Table 1) of Pseudomonas entomophila, Pseudomonas syringae and Pseudomonas fluorescens were discovered subsequently.
Pseudomonas fluorescens insecticidal toxin, or Fit, is an insecticidal proteinous toxin that causes complete loss of turgor pressure and melanization. This toxin has hemocoel-based toxicity in M. sexta, and G. mellonella . Based on comparisons with Pseudomonas strains not expressing the Fit toxin, Fit has been shown to be orally toxic to S. littoralis, H. virescens, and P. xylostella. Pseudomonas chlororaphis (Proteobacteria: Pseudomonadaceae) also expresses a Fit toxin and has oral insecticidal activity . A factor of P. fluorescens which correlates to insecticidal activity is the global regulator gene gacA. The expression of this gene influences components of the type VI secretion system  and gacA mutants do not have full insecticidal action .
Pseudomonas entomophila destroys gut cells of the common fruit fly, Drosophila melanogaster (Diptera: Drosophilidae). Based on GC content analysis and the presence of transposases, three prophages, a pyocin-like phage, and a lambdoid phage in the genome, some insecticidal genes of P. entomophila were likely acquired through lateral gene exhange. This amount of gene acquisition is less than detected in the genomes of Pseudomonas putida (Proteobacteria: Pseudomonadaceae) and Pseudomonas syringae (Proteobacteria: Pseudomonadaceae). Although the genome of P. syringae does contain tc homologs, Tcc and TcdB, similar to those seen in the genome of P. fluorescens . Other insecticidal components of P. fluorescens, besides the Fit toxin, consist of three TccC toxins, lipases, and exotoxins with hemolytic activity.
4. Neurobiology of Hemolymph-Based Toxicity
This section summarizes the current knowledge of hemolymph-based toxicity caused by various entomopathogenic bacteria in relation to neurobiology of insect hosts. We also discuss potential application of this knowledge for use in the management of lepidopteran pests.
4.1. Neurotoxins and Their Application in Pest Management
Synthetic pesticides are the most widely accepted strategies for insect pest management. Most of the synthetic pesticides, such as organophosphates, pyrethroids, and phenylpyrazoles, target the nervous system of insects. When synthetic pesticides are applied as foliar sprays or used in urban settings they have the potential to persist in the environment with deleterious effects [127,128]. Furthermore, these insecticides lack pest specificity and promote development of pest resistance [129,130,131]. Synthetic pesticide resistance leads to a problem often referred to as the pesticide treadmill, meaning that the more often a pesticide is used, the more potent and highly concentrated a pesticide must be for efficacious future use .
The lack of specificity of synthetic pesticides results in similar toxic effects between insects, humans  and on other non-target organisms . One example is the synthetic pesticide ingredient amitraz. This insecticide is an acaracide and an octopamine receptor agonist of the nervous system of invertebrates . In the United States, it has been most commonly used to control ticks and fleas on dogs, mites on cotton, and insect pests on pear trees . Amitraz also stimulates the α2-adrenoceptors of the human nervous system . Some natural products used in organic agriculture have modes of action similar to synthetic pesticides despite their natural origins. For example, pyrethrum from chrysanthemums, Tanacetum cinerariifolium (Asterales: Asteraceae), have pyrethrins, chrysanthemates and pyrethrates with neurotoxic action similar to that of synthetic pyrethroids . Another example is spinosyns from soil actinomycetes which disrupt acetyocholine neurotransmission . One organic agricultural product with insect specificity is the azadirachtins class of organic chemicals, which is derived from the Indian neem tree, Azadirachta indica (Sapindales: Meliaceae), which block insect molting hormones thereby disrupting insect development .
One way to circumvent the damaging non-target effects of synthetic pesticides, which share common modes of action between humans and insects, is the consideration of naturally occurring entomopathogens and/or the toxins they produce . For example, Bt foliar sprays are organic, have high specificity, and have negligible environmental impact compared to their synthetic counterparts . Another alternative to synthetic pesticides is the use of toxins produced by mites, spiders, and other venomous organisms which also have been shown to have neurotoxic effects on insects . Various studies have shown that some of the genes encoding these toxins can be expressed in transgenic plants and contribute to a decrease of the target insect population .
As mentioned, many neurotoxins have been discovered from invertebrates. For example, leptinotarsin, isolated and identified from the Colorado potato beetle, has been shown to disrupt the release of acetylcholine at the presynaptic nerve terminal of rat synaptosomes [109,110]. Although peptides of leptinotarsin have shown homology to both juvenile hormone esterase (JHE) of insects and Cry toxin of Bt, there is no evidence that leptinotarsin has JHE activity. Because of the way proteinous neurotoxins interact and disrupt neural tissue function, they can be used to study the physiological consequences of the nervous system dysfunction . Understanding their mode of action and interaction with the insect nervous system can make them a powerful tool with application in insect pest management.
4.1.1. Insect Paralysis Caused by Bt
Once Bt Cry toxins are ingested by lepidopteran larvae a number of effects concurrent to toxicity occur including paralysis. Paralysis is such a predominant characteristic that early studies reported paralytic effects caused by Bt as a preliminary way to discriminate among different insecticidal Bacillus species and strains . Paralysis induced by Bt can be categorized by insect type. Type I insects are described by the symptom of whole body paralysis. The larvae have been described as becoming inactive causing them to fall off their host plant (Table 2). This has been demonstrated to be caused by increased pH in the hemolymph. In type II insects, paralysis symptoms are limited to gut movement. Paralysis is thought to be caused by the breakdown of the epithelial integument, thereby inhibiting function and movement . Gut paralysis in Type II insects has been described as cessation of feeding, frass production, or microscopic evidence of muscles surrounding the gut relaxing (Table 2).
Very few studies have discussed the potential role of the insect nervous system, brain, or neuromuscular junction in insect gut or whole body paralysis. Peristalsis in Lepidoptera is controlled by the stomatogastric nervous system . The frontal ganglion controls the ability of the foregut to empty food into the midgut. Peristalsis is controlled by the frontal connectives so if digestion has ceased then this aspect of the nervous system may have been targeted. However, larva regurgitation is a common defense response during ingestion of plant defense molecules. Midgut paralysis, the cessation of muscles emptying food from the foregut to the midgut, causing regurgitation, is not usually distinguished from regurgitation as a defense response. The toxins of insect pathogens which have both oral and injectable toxicity could selectively silence specific insect nervous tissues surrounding digestion and midgut muscle movement. Understanding toxins of this nature would help determine the function of these tissues in insect behavior, especially regarding food consumption.
One Cry toxin of Bt, Cry1C, targets the nervous system  and the gut  of various Lepidoptera species. Although Cry toxins are orally toxic compounds acting as gut poisons, upon observing a range of gut paralytic effects the role of the nervous system has been discussed . Some of the neurotoxic effects of Cry toxins may be explained by their homology to both Pir toxin from P. luminescens and neurotoxin from L. decemlineata .
4.1.2. Neurotoxicity in Other Bt Strains
Neurotoxic symptoms have been observed when Bt var. israelensis (Bti) are directly injected into T. ni. Higher doses of Bti stops heart activity when injected into the hemolymph. Moreover, a loss of motor activity, paralysis and flaccidity was observed. Similar but more benign symptoms occurred when Bti was injected into mice. Oral feeding of Bt kurstaki strain to T. ni resulted in vomiting while injection had no adverse neuromuscular affects. The proteins conferring neurotoxicity were components of the crystal endotoxin from Bti .
The neurotoxic effects of Bti were further explored in the American cockroach, Periplaneta americana (Blattodea: Blattidae). The presynaptic nerve terminal function was suspected to be blocked whereas the postsynaptic membranes and axons in the ventral nerve cord remained unaffected. The sixth abdominal ganglion transmitter release, calcium uptake, and complete blockage of transmitters were observed. Rat muscle cultures treated with a purified crystal protein from Bti resulted in degeneration. The mode of action described was related to Na+/K+-ATPase damage upon incubation and K+ levels decrease while Na+ levels increase within muscles cells with increasing Ca2+ influx .
|Table 2. Paralytic effects of ingested Bt on various lepidopteran families and species.|
|Noctuidae||Spodoptera spp.||Not specified||No paralysis|
|H. virescens||Bt var. kurstaki||Midgut paralysis; Intermittent whole body paralysis with recovery to body paralysis|
|T. ni||Not specified||Type I paralysis|
|Saturniidae||Philosamia ricini||Bt var. sotto crystals||Whole body paralysis, Type I|
|Crambidae||Ostrinia nubilalis||Bt var. thuringiensis crystalline paraspores||Gut paralysis|
|Pyralidae||Not specified||Cry proteins||Paralysis|
|G. mellonella||Spores and crystals derived from Thuricide||No paralysis, Type III most susceptible to spores|
|Ephestia cautella||Not specified||Type II paralysis|
|Sphingidae||Phlegathontius quinquemaculatus||Thuricide (International Minerals and Chemical Corp., Libertyville, IL, USA)||Abnormally quiescent, cessation of feeding and slow death, no paralytic effect which was directly compared to synthetic insecticide induce paralysis|
|Erebidae||L. dispar||Not specified||Type II paralysis|
|Plutellidae||P. xylostella||Bt biological products||Decreased movement with subsequent paralysis|
|P. xylostella||Bt var. kurstaki-HD1, sprayed bacterial suspensions||Reduction of movements to stoppage, limp, loss of agility and movements slow, unresponsive to touch|
|Papilionidae||Papilio demoleus||Bti Berliner spore||Fairly rapid paralysis followed by an increase of blood alkalinity after ingestion of spores|
|Gelechiidae||Pectinophora gossypella||Bt δ-endotoxin||Evidence of gut paralysis by histological investigation where gut muscles surrounding the disorganized epithelium are relaxed|
|Bombycidae||B. mori||Bacillus sotto||Paralysis within four hours|
|B. mori||Bt var. sotto||Paralysis|
|B. mori||Not specified||Type I paralysis|
|Protoparce quinquemaculata||Bt||General paralysis|
|P. quinquemaculata||Not specified||Type I paralysis|
|Protoparce sexta||Bt crystals||General paralysis|
|Antheraea pernyi||Bt crystals||General paralysis|
|Pieridae||Colias eurytheme||Bt var. thuringiensis||No paralysis|
|Pieris rapae||Not specified||Type II paralysis|
|Hesperiidae||Urbanus acawoios||Bt var. kurstaki wettable powder||Decreased movement after 10 h, classified as likely Type II because no general paralysis|
|Tortricidae||Choristoneura fumiferana||Bt Dipel foliar spray||Interruption of feeding due to gut paralysis resulted in reducing rate of development|
5. Areas for Future Study
The recent discoveries of numerous insect toxin analogs have raised questions about the origin of virulence factor genes of entomopathogenic bacteria. This is because it is those genes which define the very nature of entomopathogens and their pathogenic relationships with their target hosts. There are also symptoms which occur in entomopathogenicity which have no defined virulence factor associated, raising the question of who is the culprit. This section addresses some of these questions and briefly summarizes the work that has been done to raise these questions.
5.1. Why Are There so Many Insecticidal Genes in Bacteria?
One potential answer to the question of why there are so many insecticidal genes lies in the nature of Lepidopteran larvae behavior. Many larval stages spend most of their time ingesting food and bacteria along with it. Bacteria exist in habitats where these insects reside such as soil, phylloplane, or in an endophytic stage. Therefore, it is no surprise that potent virulence factors of entomopathogenic bacteria have an oral route for toxicity, like the Tc superfamily and Cry toxins.
The gut of lepidopteran larvae is also a reservoir of diverse bacterial populations that interact with one another, competing for both space and nutrients. It is known that organisms in constant competition for resources are in an atmosphere which fosters adaptation . For example, P. luminescens has adapted to outcompete surrounding microbes by secreting antibiotics. As a result, after infecting an insect host, P. luminescens dominates the insect cadaver .
Previous to molecular genome comparisons, bacteria were categorized by their specific pathogenic effect. For example species within the B. cereus sensu lato are distinguished by the diseases they cause in humans and insects. Now we come to find through bioinformatic analysis that similar insecticidal toxins have been identified in their genomes. The high degree of similarity between genomes of different Bacillus spp. and the identification of common insecticidal genes is probably due to plasmid shuffling, horizontal gene transfer events, and phage elements surrounding the toxin genes. It can be hypothesized that these similarities could be explained by their shared environment, including a common host, which promoted the exchange of genetic material. For example, recent genome analysis of Yersinia and Serratia have shown evidence of similar plasmid shuffling events in Y. pseudotuberculosis  and transposable elements in S. entomophila .
As more bacterial genomes are sequenced, more virulence factors will be identified between different taxa. So far, analogs within the tc gene superfamily, originally identified in P. luminescens, have also been identified in Yersinia, Xenorabdus, Bacillus, Clostridium, Salmonella and Serratia  potentially with more to come. Photorhabdus Pir proteins have homology to Bt Cry toxins and Mcf toxins have analogs in both Clostridium and Pseudomonas .
5.2. Why Nematode-Vectored Entomopathogenic Bacteria that Are Directly Delivered to the Host’s Hemocoel Produce Oral Toxins?
Photorhabdus luminescens toxins have both oral and hemolymph-based toxic capability. Hemolymph-based toxicity from an insect pathogen which can be delivered directly into the insect makes sense but the orally toxic nature of specific and highly pathogenic toxins is curious. The tc genes are found in many organisms which are not directly delivered into the hemolymph. The origin of the tc genes could be better understood by investigating the ancestry of bacteria with Tc toxins and the origins of nematode association with bacteria.
5.3. Why Is the Mechanism of Gut Paralysis Limited to Bt?
As mentioned above, Bt toxins are known to cause gut and whole body paralysis. It has been demonstrated that Bt causes whole body paralysis by the release of K+ from the compromised midgut into the hemolymph or from intracellular sources such as K+ channels, in the absence of midgut K+ . Paralysis effects can be mimicked, by injecting alkaline buffers directly into the hemocoel . Most neurotoxins, in particular those derived from insect hunting arthropods (i.e., spiders), act on major ion channels to produce their paralyzing effect . Bt has been previously reported to disrupt ion channels in the midgut [158,159] and ion channel formation at gut neuromuscular junctions could be one way Bt causes paralysis. The binding of Bt to ATP binding cassettes also has the potential to disrupt ion transport across a membrane. ATP binding cassettes utilize ATP to transport various substrates across membranes and have recently been identified as Bt receptors. Insects harboring mutant ATP binding cassettes in their genome are resistant to Bt. This may be another mechanism to investigate regarding Bt gut paralysis.
Without any doubt, Bt, whether incorporated into a foliar spray or toxins expressed in transgenic plants, is regarded as the premier entomopathogen used in insect pest management. Recently, similar toxins to that of Bt have been identified throughout the bacterial kingdom. These finding may have two types of implications: (1) the development of novel tools for insect pest management and (2) the gain of further knowledge of the origin of entomopathogens and their associated virulence factors. For example, it would be interesting to investigate if the combination of virulence factors of two different entomopathogenic bacteria such as Photorhabdus and Bacillus delay resistance. Combining toxins with different modes of action may delay the onset of resistance by forcing insects to develop two separate mechanisms of resistance. Moreover, their combination could result in a more lethal insecticide that would result in a better control tactic for a problematic insect pest.
Furthermore, the delivery mechanism of Photorhabdus, i.e., directly released by nematodes into the insect’s hemocoel, suggests the existence of virulence factors with novel tissue targets in the Lepidoptera that can be further investigated. If proven successful the combination of newly identified hemolymph-based toxins with conventional orally-based toxins would expand their application in insect pest management.
Entomopathogens in general have a history of horizontal gene transfer events shuffling toxin-containing plasmids and pathogenicity islands between each other. The acquisition of insecticidal genes may be a strategy that developed to onset virulence once bacteria were ingested, thereby, broadening the availability of possible nutrient sources. Recognizing that insect virulence factor genes are present throughout the bacterial kingdom can broaden our knowledge of the nature of entomopathogenicity and the role of bacterial symbionts as entomopathogenic partners.
The authors would like to acknowledge the Postdoctoral Program for Excellence in Research and Teaching (PERT) at The University of Arizona Center for Insect Science, funding through the National Institute of Health training grant number 1K12 GM 00708, IRACDA Program, for supporting A. Castagnola during the time of the writing of this manuscript. We also acknowledge funding from NEMASYM (Nematode-Bacterium Symbioses) Research Coordination Network (NSF-IOS 0840932 to S. P. Stock).
The opinions shared in this manuscript are not necessarily a reflection of those of the National Institute of Health or the National Science Foundation.
Conflicts of Interest
The authors declare no conflict of interest.
- Steinhaus, E.A. Living insecticides. Sci. Am. 1956, 195, 96–106. [Google Scholar] [CrossRef]
- Lacey, L.A.; Frutos, R.; Kaya, H.K. Insect pathogens as biological control agents: Do they have a future? Biol. Contr. 2001, 21, 230–248. [Google Scholar] [CrossRef]
- Lacey, L.A.; Kaya, H.K. Field Manual of Techniques in Invertebrate Pathology, 2nd ed.; Springer: Dorcrecht, The Netherlands, 2007. [Google Scholar]
- Kaya, H.K.; Vega, F.E. Insect Pathology, 2nd ed.; Elsevier Academic Press: San Diego, CA, USA, 2012. [Google Scholar]
- Ishiwata, S.C. One kind of severe flacherie (sotto disease). Dainihon Sans Kaiho 1901, 114, 1–5. [Google Scholar]
- Berliner, E. Ueber die schlaffsucht der ephestia kuhniella und Bacillus thuringiensis n. sp. Z Angew. Entomology 1915, 2, 21–56.(in German). [Google Scholar]
- Wilcox, D.R.; Shivakumar, A.G.; Melin, B.E. Genetic Engineering of Bioinsecticides. In Protein Engineering Applications in Science, Medicine, and Industry, 1st ed.; Inouye, M., Sarma, R., Eds.; Academic Press: Orlando, FL, USA, 1986; pp. 395–412. [Google Scholar]
- Fischhoff, D.A.; Bowdish, K.S.; Perlak, F.J. Insect tolerant transgenic tomato plants. Nat. Biotechnol. 1987, 5, 807–813. [Google Scholar] [CrossRef]
- Hilder, V.A.; Gatehouse, A.M.; Sheerman, S.E. A novel mechanism of insect resistance engineered into tobacco. Nature 1987, 330, 160–163. [Google Scholar] [CrossRef]
- Watkins, P.R.; Huesing, J.E.; Margam, V. Insects, Nematodes, and Other Pests. In Plant Biotechnology and Agriculture Prospects for the 21st Century, 1st ed.; Altman, A., Hasegawa, P.M., Eds.; Elsevier: London, UK, 2012; pp. 353–370. [Google Scholar]
- Gatehouse, J.A. Biotechnological prospects for engineering insect-resistant plants. Plant Physiol. 2008, 146, 881–887. [Google Scholar] [CrossRef]
- Chattopadhyay, A.; Bhatnagar, N.B.; Bhatnagar, R. Bacterial insecticidal toxins. Crit. Rev. Microbiol. 2004, 30, 33–54. [Google Scholar] [CrossRef]
- Priest, F.G.; Barker, M.; Baillie, L.W. Population structure and evolution of the Bacillus cereus group. J. Bacteriol. 2004, 186, 7959–7970. [Google Scholar] [CrossRef]
- Ochman, H.; Lawrence, J.G.; Groisman, E.A. Lateral gene transfer and the nature of bacterial innovation. Nature 2000, 405, 299–304. [Google Scholar] [CrossRef]
- Nielsen-LeRoux, C.; Gaudriault, S.; Ramarao, N. How the insect pathogen bacteria Bacillus thuringiensis and Xenorhabdus/Photorhabdus occupy their hosts. Curr. Opin. Microbiol. 2012, 15, 220–231. [Google Scholar] [CrossRef]
- Shelton, A.M.; Zhao, J.Z.; Roush, R.T. Economic, ecological, food safety, and social consequences of the deployment of Bt transgenic plants. Annu. Rev. Entomol. 2002, 47, 845–881. [Google Scholar] [CrossRef]
- Höfte, H.; Whiteley, H.R. Insecticidal crystal proteins of Bacillus thuringiensis. Microbiol. Rev. 1989, 53, 242–255. [Google Scholar]
- Gough, J.M.; Akhurst, R.J.; Ellar, D.J. New isolates of Bacillus thuringiensis for control of livestock ectoparasites. Biol. Contr. 2002, 23, 179–189. [Google Scholar] [CrossRef]
- Chilcott, C.N.; Ellar, D.J. Comparative toxicity of Bacillus thuringiensis var. israelensis crystal proteins in vivo and in vitro. J. Gen. Microbiol. 1988, 134, 2551–2558. [Google Scholar]
- Kreig, A.; Huger, A.M.; Langenbruch, G.A. New results on Bacillus thuringiensis var tenebrionis with special regard to its effect on the colorado beetle (Leptinotarsa decemlineata). Anz Schadlingskunde Pflanzenschutz Umweltschutz 1984, 57, 145–150. [Google Scholar] [CrossRef]
- Rose, E.A.; Harris, R.J.; Glare, T.R. Possible pathogens of social wasps (Hymenoptera: Vespidae) and their potential as biological control agents. New Zeal. J. Zool. 1999, 26, 179–190. [Google Scholar] [CrossRef]
- Wei, J.Z.; Hale, K.; Carta, L.; Plazter, E.; Wong, C.; Fang, S.C.; Aroian, R.V. Bacillus thuringiensis crystal proteins that target nematodes. Proc. Natl. Acad. Sci. USA 2003, 100, 2760–2765. [Google Scholar]
- Hui, F.; Scheib, U.; Hu, Y. Structure and glycolipid binding properties of the nematicidal protein Cry5B. Biochemistry 2012, 51, 9911–9921. [Google Scholar] [CrossRef]
- Schnepf, E.; Crickmore, N.; van Rie, J. Bacillus thuringiensis and its pesticidal crystal proteins. Microbiol. Mol. Biol. Rev. 1998, 62, 775–806. [Google Scholar]
- Loeb, M.J.; Martin, P.A.; Hakim, R.S. Regeneration of cultured midgut cells after exposure to sublethal doses of toxin from two strains of Bacillus thuringiensis. J. Insect. Physiol. 2001, 47, 599–606. [Google Scholar] [CrossRef]
- Griego, V.M.; Fanchier, L.J.; Spence, K.D. Scanning electron microscopy of the disruption of tobacco hornworm, Manduca sexta, midgut by Bacillus thuringiensis, endotoxin. J. Invertebr. Pathol. 1980, 35, 186–189. [Google Scholar] [CrossRef]
- Bravo, A.; Jansens, S.; Peferoen, M. Immunocytochemical localization of Bacillus thuringiensis insecticidal crystal proteins in intoxicated insects. J. Invertebr. Pathol. 1992, 60, 237–246. [Google Scholar] [CrossRef]
- Cerstiaens, A.; Verleyen, P.; van Rie, J. Effect of Bacillus thuringiensis Cry1 toxins in insect hemolymph and their neurotoxicity in brain cells of Lymantria dispar. Appl. Environ. Microb. 2001, 67, 3923–3927. [Google Scholar] [CrossRef]
- Haider, M.Z.; Ellar, D.J. Mechanism of action of Bacillus thuringiensis insecticidal δ-endotoxin: Interaction with phospholipid vesicles. Biochim. Biophys. Acta 1989, 978, 216–222. [Google Scholar] [CrossRef]
- Zhang, X.; Candas, M.; Griko, N.B. A mechanism of cell death involving an adenylyl cyclase/PKA signaling pathway is induced by the Cry1Ab toxin of Bacillus thuringiensis. Proc. Natl. Acad. Sci. USA 2006, 103, 9897–9902. [Google Scholar] [CrossRef]
- Tanaka, S.; Miyamoto, K.; Noda, H. The ATP-binding cassette transporter subfamily C member 2 in Bombyx mori larvae is a functional receptor for cry toxins from Bacillus thuringiensis. FEBS J. 2013, 280, 1782–1794. [Google Scholar] [CrossRef]
- Atsumi, S.; Miyamoto, K.; Yamamoto, K. A single amino acid mutation in an ABC transporter causes resistance to Bt toxin Cry1Ab in the silkworm, Bombyx mori. Proc. Natl. Acad. Sci. USA Plus 2012, 109, E1591–E1598. [Google Scholar] [CrossRef]
- Gahan, L.J.; Pauchet, Y.; Vogel, H. An ABC transporter mutation is correlated with insect resistance to Bacillus thuringiensis Cry1Ac toxin. PLoS Genet. 2010, 6, e1001248. [Google Scholar]
- Baxter, S.W. Parallel evolution of Bacillus thuringiensis toxin resistance in lepidoptera. Genetics 2011, 189, 675–679. [Google Scholar] [CrossRef]
- Arneson, L.P.; Fagerlund, A.; Granum, P.E. From soil to gut: Bacillus cereus and its food poisoning toxins. FEMS Microbiol. Rev. 2008, 32, 579–606. [Google Scholar] [CrossRef]
- Perchat, S.; Buisson, C.; Chaufaux, J. Bacillus cereus produces several nonproteinaceous insecticidal exotoxins. J. Invertebr. Pathol. 2005, 90, 131–133. [Google Scholar] [CrossRef]
- Zwick, M.E.; Joseph, S.J.; Didelot, X. Genomic characterization of the Bacillus cereus sensu lato species: Backdrop to the evolution of Bacillus anthracis. Genome Res. 2012, 22, 1512–1524. [Google Scholar] [CrossRef]
- Singer, S. The Utility of Strains of Morphological Group II Bacillus. In Advances in Applied Microbiology; Academic Press: San Diego, CA, USA, 1996; Volume 42, pp. 219–259. [Google Scholar]
- Berry, C. The bacterium, Lysinibacillus sphaericus, as an insect pathogen. J. Invertebr. Pathol. 2012, 109, 1–10. [Google Scholar] [CrossRef]
- Nishiwaki, H.; Nakashima, K.; Ishida, C. Cloning, functional characterization and mode of action of a novel insecticidal pore-forming toxin, Sphaericolysin, produced by Bacillus sphaericus. Appl. Environ. Microb. 2007, 73, 3404–3411. [Google Scholar] [CrossRef]
- Barth, H.; Aktories, K.; Popoff, M. Binary bacterial toxins: Biochemistry, biology, and applications of common Clostridium and Bacillus proteins. Microbiol. Mol. Biol. Rev. 2004, 68, 373–402. [Google Scholar] [CrossRef]
- Nicolas, L.; Charles, J.F.; Debarjac, H. Clostridium bifermentans servovar malaysia; characterization of putative mosquito larvicidal proteins. FEMS Microbiol. Lett. 1993, 113, 23–29. [Google Scholar] [CrossRef]
- Tsuge, H.; Nagahama, M.; Nishimura, H. Crystal structure and site-directed mutagenesis of enzymatic components from Clostridium perfringens iota-toxin. J. Mol. Biol. 2003, 325, 471–483. [Google Scholar] [CrossRef]
- Just, I.; Gerhard, R. Large clostridial cytotoxins. Rev. Physiol. Biochem. Pharmacol. 2005, 152, 23–47. [Google Scholar] [CrossRef]
- Aktories, K.; Schwan, C.; Papatheodorou, P. Bidirectional attack on the actin cytoskeleton. bacterial protein toxins causing polymerization or depolymerization of actin. Toxicon 2012, 60, 572–581. [Google Scholar] [CrossRef]
- Visschedyk, D.D.; Perieteanu, A.A.; Turgeon, Z.J. Photox, a novel actin-targeting mono-ADP-ribosyltransferase from Photorhabdus luminescens. J. Biol. Chem. 2010, 285, 13525–13534. [Google Scholar] [CrossRef]
- Fischer-Le Saux, M.; Viallard, V. Spodoptera littoralis Photorhabdus and proposal of new taxa: P. luminescens subsp. luminescens subsp. nov., P. luminescens subsp. akhurstii subsp. nov., P. luminescens subsp. laumondii subdp. nov., P. temperata sp. nov. and P. asymbiotica sp. nov. Int. J. Syst. Bacteriol. 1999, 49, 1645–1656. [Google Scholar] [CrossRef]
- Gerrard, J.G.; Joyce, S.A.; Waterfield, N.R. Nematode symbiont for Photorhabdus asymbiotica. Emerg. Infect. Dis. 2006, 12, 1562–1564. [Google Scholar] [CrossRef]
- Gerrard, J.; Waterfield, N.; Vohra, R. Human infection with Phtotorhabdus asymbiotica: An emerging bacterial human pathogen. Microbes Infect. 2004, 6, 229–237. [Google Scholar] [CrossRef]
- Duchaud, E.; Rusniok, C.; Frangeul, L. The genome sequence of the entomopathogenic bacterium Photorhabdus luminescens. Nat. Biotechnol. 2003, 21, 1307–1313. [Google Scholar] [CrossRef]
- Rodou, A.; Ankrah, D.; Stathopoulos, C. Toxins and secretion systems of Photorhabdus luminescens. Toxins 2010, 2, 1250–1264. [Google Scholar] [CrossRef]
- Daborn, P.J.; Waterfield, N.; Silva, C.P. A single Photorhabdus gene, makes caterpillars floppy (mcf), allows Escheria coli to persist within and kill Insects. Proc. Natl. Acad. Sci. USA 2002, 99, 10742–10747. [Google Scholar]
- Waterfield, N.; Hares, M.; Yang, G. Potentiation and cellular phenotypes of the insecticidal toxin complexes of Photorhabdus bacteria. Cell. Microbiol. 2005, 7, 373–382. [Google Scholar] [CrossRef]
- Waterfield, N.; Kamita, S.G.; Hammock, B.D. The Photorhabdus Pir toxins are similar to a developmentally regulated insect protein but show no juvenile hormone esterase activity. FEMS Microbiol. Lett. 2005, 245, 47–52. [Google Scholar] [CrossRef]
- Blackburn, M.B.; Farrar, R.R.; Novak, N.G. Remarkable susceptibility to the diamondback moth (Plutella xylostella) to ingestion of Pir toxins from Photorhabdus luminescens. Entomol. Exp. Appl. 2006, 121, 31–37. [Google Scholar] [CrossRef]
- Ffrench-Constant, R.H.; Dowling, A.; Waterfield, N. Insecticidal toxins from Photorhabdus bacteria and their potential use in agriculture. Toxicon 2007, 49, 436–451. [Google Scholar] [CrossRef]
- Crosland, R.; Fitch, R.; Hines, H. Characterization of beta-leptinotarsin and the effects of calcium flux antagonists on its activity. Toxicon 2005, 45, 829–841. [Google Scholar] [CrossRef]
- Brown, S.E.; Cao, A.T.; Dobson, P. Txp40, a ubiquitous insecticidal toxin protein from Xenorhabdus and Photorhabdus bacteria. Appl. Environ. Microb. 2006, 72, 1653–1662. [Google Scholar] [CrossRef]
- Park, J.M.; Kim, M.; Min, J. Proteomic identification of a novel toxin protein (Txp40) from Xenorhabdus nematophila and its insecticidal activity against larvae of Plutella xylostella. J. Agric. Food Chem. 2012, 60, 4053–4059. [Google Scholar] [CrossRef]
- Brown, S.E.; Cao, A.T.; Hines, E.R.; Akhurst, R.J.; East, P.D. A novel secreted protein toxin from the insect pathogenic bacterium Xenorhabdus nematophila. J. Biol. Chem. 2004, 279, 14595–14601. [Google Scholar] [CrossRef]
- Sicard, M.; Le Brun, N.; Pages, S. Effect of native Xenorhabdus on the fitness of their Steinernema hosts: Contrasting types of interaction. Parasitol. Res. 2003, 91, 520–524. [Google Scholar] [CrossRef]
- Singh, P.; Park, D.; Forst, S. Xenocin export by the flagellar type III pathway in Xenorhabdus nematophila. J. Bacteriol. 2013, 195, 1400–1410. [Google Scholar] [CrossRef]
- Sergeant, M.; Jarrett, P.; Ousley, M. Interactions of insecticidal toxin gene products from Xenorhabdus nematophilus PMFI296. Appl. Environ. Microb. 2003, 69, 63344–63349. [Google Scholar]
- Pan, Y.; Jian, H.; Zhang, J. An intracellular toxic protein (Xin) isolated from Xenorhabdus nematophilus strain BJ. Prog. Nat. Sci. 2002, 12, 310–312. [Google Scholar]
- Vigneux, F.; Zumbihl, R.; Jubelin, G. The xaxAB genes encoding a new apoptotic toxin from the insect pathogen Xenorhabdus nematophila are present in plant and human pathogens. J. Biol. Chem. 2007, 282, 9571–9580. [Google Scholar] [CrossRef]
- Viitanen, P.V.; Gatenby, A.A.; Lorimer, G.H. Purified chaperonin 60 (groEL) interacts with the nonnative states of a multitude of Escherichia coli proteins. Protein Sci. 1992, 1, 363–369. [Google Scholar] [CrossRef]
- Ellis, J.R. Chaperomics: In vivo GroEL function defined. Curr. Biol. 2005, 15, R661–R663. [Google Scholar] [CrossRef]
- Yang, J.; Zeng, H.M.; Lin, H.F. An insecticidal protein from Xenorhabdus budapestensis that results in prophenoloxidase activation in the wax moth, Galleria mellonella. J. Invertebr. Pathol. 2012, 110, 60–67. [Google Scholar] [CrossRef]
- Lamelas, A.; Gosalbes, M.J.; Manzano-Marín, A. Serratia symbiotica from the aphid Cinara cedri: A missing link from facultative to obligate insect endosymbiont. PLoS Genet. 2011, 7, 1–11. [Google Scholar]
- Rae, R.; Riebesell, M.; Dinkelacker, I. Isolation of naturally associated bacteria of necromenic Pristionchus nematodes and fitness consequences. J. Exp. Biol. 2008, 211, 1927–1936. [Google Scholar] [CrossRef]
- Abebe, E.; Abebe-Akele, F.; Morrison, J. An insect pathogenic symbiosis between a caenorhabditis and serratia. Virulence 2011, 2, 158–161. [Google Scholar] [CrossRef]
- Jackson, T.A.; Boucias, D.G.; Thaler, J.O. Pathobiology of amber disease, caused by Serratia spp., in the New Zealand grass grub, Costelytra zealandica. J. Invertebr. Pathol. 2001, 78, 232–243. [Google Scholar] [CrossRef]
- Sikorowski, P.P.; Lawrence, A.M.; Inglis, G.D. Effects of Serratia marcescens on rearing of the tobacco budworm (Lepidoptera: Noctuidae). Am. Entomol. 2001, 47, 51–60. [Google Scholar]
- Hurst, M.R.H.; Glare, T.R.; Jackson, T.A.; Ronson, C.W. Plasmid-located pathogenicity determinants of Serratia entomophila, the causal agent of amber disease of grass grub, show similarity to the insecticidal toxins of Photorhabdus luminescens. J. Bacteriol. 2000, 182, 5127–5138. [Google Scholar] [CrossRef]
- Hurst, M.R.; Beard, S.S.; Jackson, T.A. Isolation and characterization of the Serratia entomophila antifeeding prophage. FEMS Microbiol. Lett. 2007, 270, 42–48. [Google Scholar] [CrossRef]
- Dodd, S.J.; Hurst, M.R.; Glare, T.R. Occurrence of sep insecticidal toxin complex genes in Serratia spp. and Yersinia frederiksenii. Appl. Environ. Microb. 2006, 72, 6584–6592. [Google Scholar] [CrossRef]
- Tennant, S.M.; Skinner, N.A.; Joe, A. Homologues of insecticidal toxin complex genes in Yersinia enterocolitica biotype 1A and their contribution to virulence. Infect. Immun. 2005, 73, 6860–6867. [Google Scholar] [CrossRef]
- Fuchs, T.M.; Bresolin, G.; Marcinowski, L. Insecticidal genes of Yersinia spp.: Taxonomical distribution, contribution to toxicity towards Manduca sexta and Galleria mellonella, and evolution. BMC Microbiol. 2008, 8, 1–11. [Google Scholar] [CrossRef]
- Parkhill, J.; Wren, B.W.; Thomson, N.R. Genome sequence of Yersinia pestis, the causitive agent of plague. Nature 2001, 413, 523–527. [Google Scholar] [CrossRef]
- Bresolin, G.; Morgan, J.A.; Illgen, D. Low temperature-induced insecticidal activity of Yersinia enterocolitica. Mol. Microbiol. 2006, 59, 503–512. [Google Scholar] [CrossRef]
- Spanier, B.; Starke, M.; Higel, F. Yersinia enterocolitica infection and tcaA-dependent killing of Ceaenorhabditis elegans. Appl. Environ. Microb. 2010, 76, 6277–6285. [Google Scholar] [CrossRef]
- Hurst, M.R.; Jones, S.A.; Binglin, T. The main virulence determinant of Yersinia entomophaga MH96 is a broad-host-range toxin complex active against insects. J. Bacteriol. 2011, 193, 1966–1980. [Google Scholar] [CrossRef]
- Péchy-Tarr, M.; Bruck, D.J.; Maurhofer, M. Molecular analysis of a novel gene cluster encoding an insect toxin in plant-associated strains of Pseudomonas fluorescens. Environ. Microbiol. 2008, 10, 2368–2386. [Google Scholar] [CrossRef]
- Ruffner, B.; Péchy-Tarr, M.; Ryffel, F. Oral insecticidal activity of plant-associated pseudomonads. Environ. Microbiol. 2013, 3, 751–763. [Google Scholar] [CrossRef]
- Hassan, K.A.; Johnson, A.; Shaffer, B.T. Inactivation of the GacA response regulator in Pseudomonas fluorescens Pf-5 has far-reaching transcriptomic consequences. Environ. Microbiol. 2010, 12, 899–915. [Google Scholar] [CrossRef]
- Vodovar, N.; Vallenet, D.; Cruveiller, S. Complete genome sequence of the entomopathogenic and metabolically versatile soil bacterium Pseudomonas entomophila. Nat. Biotechnol. 2006, 24, 673–679. [Google Scholar] [CrossRef]
- Margo, G.; Porcar, M. Ecological mysteries: Is Bacillus thuringiensis a real insect pathogen? Bt Res. 2012, 3, 1–2. [Google Scholar]
- Yuan, Y.M.; Hu, X.M.; Liu, H.Z. Kinetics of plasmid transfer among Bacillus cereus group strains within lepidopteran larvae. Arch. Microbiol. 2007, 187, 425–431. [Google Scholar] [CrossRef]
- Hu, X.M.; Mahillon, J. Life Cycle and Gene Exchange. In Endospore-forming Soil Bacteria, 1st ed.; Logan, N.A., DeVos, P., Eds.; Springer-Verlag Berlin: Berlin, Germany, 2011; Volume 27, pp. 89–113. [Google Scholar]
- Mohan, M.; Selvakumar, G.; Sushil, S.N. Entomopathogenicity of endophytic Serratia marcescens strain SRM against larvae of Helicoverpa armigera (Noctuidae: Lepidoptera). World J. Microb. Biotechnol. 2011, 27, 2545–2551. [Google Scholar] [CrossRef]
- Boucher, Y.; Douady, C.J.; Papke, R.T. Lateral gene transfer and the origins of prokaryotic groups. Annu. Rev. Genet. 2003, 37, 283–328. [Google Scholar] [CrossRef]
- Andersson, J.O. Lateral gene transfer in eukaryotes. Cell. Mol. Life Sci. 2005, 62, 1182–1197. [Google Scholar] [CrossRef]
- Acuna, R.; Padilla, B.E.; Florez-Ramos, C.P. Adaptive horizontal gene transfer of a bacterial gene to an invasive insect pest of coffee. Proc. Natl. Acad. Sci. USA 2012, 109, 4197–4202. [Google Scholar]
- Pauchet, Y.; Heckel, D.G. The genome of the mustard leaf beetle encodes two active xylanases originally acquired from bacteria through horizontal gene transfer. Proc. R. Soc. B Biol. Sci. 2013, 280, 1–7. [Google Scholar]
- Wheeler, D.; Redding, A.J.; Werren, J.H. Characterization of an ancient lepidopteran lateral gene transfer. PLoS One 2013, 8, 1–9. [Google Scholar]
- Wybouw, N.; Balabanidou, V.; Ballhorn, D.J. A horizontally transferred cyanase gene in the spider mite Tetranychus urticae is involved in cyanate metabolism and is differentially expressed upon host plant change. Insect Biochem. Mol. 2012, 42, 881–889. [Google Scholar] [CrossRef]
- Thomas, D.J.; Morgan, J.A.; Whipps, J.M. Plasmid transfer between Bacillus thuringiensis subsp. israelensis strains in laboratory culture, river water, and dipteran larvae. Appl. Environ. Microb. 2001, 67, 330–338. [Google Scholar] [CrossRef]
- Blackburn, M.B.; Martin, P.A.; Kuhar, D.; Farrar, R.R., Jr.; Gundersen-Rindal, D.E. The Occurrence of Photorhabdus-Like Toxin Complexes in Bacillus thuringiensis. PLoS One 2011, 6. [Google Scholar] [CrossRef]
- Chen, G.; Zhang, Y.; Li, J. Chinitase activity of Xenorhabdus and Photorhabdus species, bacterial associates of entomopathogenic nematodes. J. Invertebr. Pathol. 1996, 68, 101–108. [Google Scholar] [CrossRef]
- Cabral, C.M.; Cherqui, A.; Pereira, A. Purification and characterization of two distinct metalloproteases secreted by the entomopathogenic bacterium Photorhabdus sp strain Az29. Appl. Environ. Microb. 2004, 70, 3831–3838. [Google Scholar] [CrossRef]
- Sergeant, M.; Baxter, L.; Jarret, P. Identification, typing, and insecticidal activity of Xenorhabdus isolates from entomopathogenic nematodes in United Kingdom soil and characterization of the xpt toxin loci. Appl. Environ. Microb. 2006, 72, 5895–5907. [Google Scholar]
- Chaston, J.M.; Suen, G.; Tucker, S.L. The entomopathogenic bacterial endosymbionts Xenorhabdus and Photorhabdus: Convergent lifestyles from divergent genomes. PLoS One 2011, 6, e27909. [Google Scholar] [CrossRef]
- Goodrich-Blair, H.; Clarke, D.J. Mutualism and pathogenesis in Xenorhabdus and Photorhabdus: Two roads to the same destination. Mol. Microbiol. 2007, 64, 260–268. [Google Scholar] [CrossRef]
- Bowen, D.J.; Ensign, J.C. Purification and characterization of a high-molecular-weight insecticidal protein complex produced by the entomopathogenic bacterium Photorhabdus luminescens. Appl. Environ. Microb. 1998, 64, 3029–3035. [Google Scholar]
- Waterfield, N.; Dowling, A.; Sharma, S. Oral toxicity of Photorhabdus luminescens W14 toxin complexes in Escherichia coli. Appl. Environ. Microbiol. 2001, 67, 5017–5024. [Google Scholar] [CrossRef]
- Liu, D.; Burton, S.; Glancy, T. Insect resistance conferred by 283-kDa Photorhabdus luminescens protein TcdA in Arabidopsis thaliana. Nat. Biotechnol. 2003, 21, 1307–1313. [Google Scholar] [CrossRef]
- Pinheiro, V.B.; Ellar, D.J. Expression and insecticidal activity of Yersinia pseudotuberculosis and Photorhabdus luminescens toxin complex proteins. Cell. Microbiol. 2007, 9, 2372–2380. [Google Scholar]
- Ffrench-Constant, R.; Waterfield, N. An ABC guide to the bacterial toxin complexes. Adv. Appl. Microbiol. 2005, 58, 169–183. [Google Scholar] [CrossRef]
- Yoshino, E.; Baxter, D.E.; Hsiao, T.H. Release of acetylcholine from rat brain synaptosomes stimulated with leptinotarsin—New neurotoxin. J. Neurochem. 1980, 34, 635–642. [Google Scholar] [CrossRef]
- McClure, W.O.; Abbott, B.C.; Baxter, D.E. Leptinotarsin—A presynaptic neurotoxin that stimulates release of acetylcholine. Proc. Natl. Acad. Sci. USA 1980, 77, 1219–1223. [Google Scholar] [CrossRef]
- Shrestha, Y.K.; Lee, K. Oral toxicity of Photorhabdus culture media on gene expression of the adult sweetpotato whitefly, Bemisia tabaci. J. Invertebr. Pathol. 2012, 109, 91–96. [Google Scholar] [CrossRef]
- Hurst, M.R.; Glare, T.R.; Jackson, T.A. Cloning Serratia entomophila antifeeding genes—A putative defective prophage active against the grass grub Costelytra zealandica. J. Bacteriol. 2004, 186, 5116–5128. [Google Scholar] [CrossRef]
- Yang, G.; Dowling, A.J.; Gerike, U. Photorhabdus virulence cassettes confer injectable insecticidal activity against the wax moth. J. Bacteriol. 2006, 188, 2254–2261. [Google Scholar] [CrossRef]
- Chavez, C.V.; Jubelin, G.; Courties, G. The cyclomodulin Cif of Photorhabdus luminescens inhibits insect cell proliferation and triggers host cell death by apoptosis. Microbes Infect. 2010, 12, 1208–1218. [Google Scholar] [CrossRef]
- Wolters, M.; Boyle, E.C.; Lardong, K. Cytotoxic necrotizing factor-Y boosts Yersinia effector translocation by activating Rac. J. Biol. Chem. 2013, 288, 23543–23553. [Google Scholar] [CrossRef]
- Brugirard-Ricaud, K.; Duchaud, E.; Givaudan, A. Site-specific antiphagocytic function of the Photorhabdus luminescens type III secretion system during insect colonization. Cell. Microbiol. 2004, 7, 363–371. [Google Scholar] [CrossRef]
- Gendlina, I.; Held, K.G.; Bartra, S.S. Identification and type III-dependent secretion of the Yersinia pestis insecticidal-like proteins. Mol. Microbiol. 2007, 64, 1214–1227. [Google Scholar] [CrossRef]
- Waterfield, N.; Hares, M.; Hinchliffe, S. The insect toxin complex of Yersinia. Adv. Exp. Med. Biol. 2007, 603, 247–257. [Google Scholar] [CrossRef]
- Spinner, J.L.; Jarrett, C.O.; LaRock, D.L. Yersinia pestis insecticidal-like toxin complex (Tc) family proteins: Characterization of expression, subcellular localization, and potential role in infection of the flea vector. BMC Microbiol. 2012, 12, 1–14. [Google Scholar] [CrossRef]
- Odinokov, G.N.; Eroshenko, G.A.; Krasnov, J.M. Analysis of insect toxin complex gene variability of Yersinia pestis and Yersinia pseudotuberculosis strains. Russ. J. Genet. 2011, 47, 10–17. [Google Scholar]
- Hares, M.C.; Hinchliffe, S.J.; Strong, P.C. The Yersinia pseudotuberculosis and Yersinia pestis toxin complex is active against cultured mammalian cells. Microbiol. SGM 2008, 154, 3503–3517. [Google Scholar] [CrossRef]
- Jeong, H.U.; Mun, H.Y.; Oh, H.K. Evaluation of insecticidal activity of a bacterial strain, Serratia, sp EML-SE1 against diamondback moth. J. Microbiol. 2010, 48, 541–545. [Google Scholar] [CrossRef]
- Brurberg, M.B.; Nes, I.F.; Eijsink, V.G. Comparative studies of chitinases A and B from Serratia marcescens. Microbiol. UK 1996, 142, 1581–1589. [Google Scholar] [CrossRef]
- Asano, S.; Suzuki, K.; Hori, H. Synergistic effects of the supernatants from Serratia marcescens culture on larvicidal activity of Bacillus thuringiensis Cry1C toxin against common cutworm, Spodoptera litura. J. Pestic. Sci. 1999, 24, 44–48. [Google Scholar] [CrossRef]
- Abby, S.S.; Rocha, E.P. The non-flagellar type III secretion system evolved from the bacterial flagellum and diversified into host-cell adapeted sytems. PLoS Genet. 2012, 8, 1–15. [Google Scholar]
- Vodovar, N.; Vinals, M.; Liehl, P. Drosophila host defense after oral infection by an entomopathogenic Psuedomonas species. Proc. Natl. Acad. Sci. USA 2005, 102, 11414–11419. [Google Scholar] [CrossRef]
- Younglai, E.V.; Wu, Y.J.; Foster, W.G. Reproductive toxicology of environmental toxicants: Emerging issues and concerns. Curr. Pharm. Des. 2007, 13, 3005–3019. [Google Scholar]
- Werner, I.; Hitzfeld, B. 50 years of exotoxicology since silent spring—A review. Gaia 2012, 21, 217–224. [Google Scholar]
- Davies, T.G.; Field, L.M.; Williamson, M.S. The re-emergence of the bed bug as a nuisance pest: Implications of resistance to the pyrethroid insecticides. Med. Vet. Entomol. 2012, 26, 241–254. [Google Scholar] [CrossRef]
- Durand, R.; Bouvresse, S.; Berdjane, Z. Insecticide resistance in head lice: Clinical, parasitological, and genetic aspect. Clin. Microbiol. Infect. 2012, 18, 338–344. [Google Scholar] [CrossRef]
- Ranson, H.; N’Guessan, R.; Lines, J. Pyrethroid resistance in African anopheline mosquitoes: What are the implications for malaria control? Trends Parasitol. 2011, 27, 91–98. [Google Scholar] [CrossRef]
- Knight, A.L. Economics of agricultural pesticide resistance in arthropods. Annu. Rev. Entomol. 1989, 34, 293–313. [Google Scholar] [CrossRef]
- Mostafalou, S.; Abdollahi, M. Pesticides and human chronic diseases: Evidences, mechanisms, and perspectives. Toxicol. Appl. Pharm. 2013, 268, 157–177. [Google Scholar] [CrossRef]
- Grue, C.E.; Gibert, P.L.; Seeley, M.E. Neurophysiological and behavioral changes in non-target wildlife exposed to organophosphate and carbamate pesticides: Thermoregulation, food consumption, and reproduction. Am. Zool. 1997, 37, 369–388. [Google Scholar]
- Prullage, J.B.; Tran, H.V.; Timmons, P. The combined mode of action of fipronil and amitraz on the motility of Rhipicephalus sanguineus. Vet. Parasitol. 2011, 179, 302–310. [Google Scholar] [CrossRef]
- Jorens, P.G.; Zandijk, E.; Belmans, L. An unusual poisoning with the unusual pesticide amitraz. Hum. Exp. Toxicol. 1997, 16, 600–601. [Google Scholar] [CrossRef]
- Marrs, T.C. Toxicology of insecticides to mammals. Pest Manag. Sci. 2012, 68, 1552–1556. [Google Scholar] [CrossRef]
- Dayan, F.E.; Cantrell, C.L.; Duke, S.O. Natural products in crop protection. Bioorg. Med. Chem. 2009, 17, 4022–4034. [Google Scholar] [CrossRef]
- Nicholson, G.M. Fighting the global pest problem: Preface to the special toxicon issue on insecticidal toxins and their potential for insect pest control. Toxicon 2007, 49, 413–422. [Google Scholar] [CrossRef]
- Castagnola, A.; Jurat-Fuentes, J.L. Bt Crops: Past and Future. In Bacillus Thuringiensis Biotechnology, 1st ed.; Sansinenea, E., Ed.; Springer: New York, NY, USA, 2012; Volume 392, pp. 283–304. [Google Scholar]
- Windley, M.J.; Herzig, V.; Dziemborowicz, S.A. Spider-venom peptides as bioinsecticides. Toxins 2012, 4, 191–227. [Google Scholar] [CrossRef]
- Khan, S.A.; Zafar, R.W.; Briddon, K.A. Spider venom toxin protects plants from insect attack. Transgenic Res. 2006, 15, 349–357. [Google Scholar] [CrossRef]
- Schiavo, G.; Matteoli, M.; Montecucco, C. Neurotoxins affecting neuroexocytosis. Physiol. Rev. 2000, 80, 717–766. [Google Scholar]
- Heimpel, A.M.; Angus, T.A. The taxonomy of insect pathogens related to Bacillus cereus frankland and frankland. Can. J. Microbiol. 1958, 4, 531–541. [Google Scholar] [CrossRef]
- Heimpel, A.M.; Angus, T.A. The site of action of crystalliferous bacteria in lepidopteran larvae. J. Insect Pathol. 1959, 1, 152–170. [Google Scholar]
- Hartenstein, V. Development of the insect stomatogastric nervous system. Trends Neurosci. 1997, 20, 421–427. [Google Scholar] [CrossRef]
- Rodriguez-Cabrera, L.; Trujillo-Bacallao, D.; Borras-Hidalgo, O. Molecular characterization of Spodoptera frugiperda Bacillus thuringiensis Cry1Ca toxin interaction. Toxicon 2008, 51, 681–692. [Google Scholar] [CrossRef]
- Aronson, A.I.; Shai, Y. Why Bacillus thuringiensis insecticidal toxins are so effective: Unique features of their mode of action. FEMS Microbiol. Lett. 2001, 195, 1–8. [Google Scholar] [CrossRef]
- Cheung, P.Y.; Roe, R.M.; Hammock, B.D. The apparent in vivo neuromuscular effects of the δ-endotoxin of Bacillus thuringiensis var. israelensis in mice and insects of four orders. Pest. Biochem. Physiol. 1985, 23, 85–94. [Google Scholar] [CrossRef]
- Cahan, R.; Shainberg, A.; Pechatnikov, I. A 28,000 mol. Wt toxin from Bacillus thuringiensis israelensis induces cation transport in rat muscle cultures. Toxicon 1995, 33, 943–951. [Google Scholar] [CrossRef]
- Münch, A.; Stingl, L.; Jung, K. Photorhabdus luminescens genes induced upon insect infection. BMC Genomics 2008, 9, 1–17. [Google Scholar] [CrossRef]
- Katrukha, G.S.; Zarubina, A.P.; Iudina, T.P. Antibiotics produced by Photorhabdus luminescens ZMI, a symbiont of entomopathogenic nematodes. Antibiot Khimioter 2009, 54, 11–16. [Google Scholar]
- Lesic, B.; Zouine, M.; Ducos-Galand, M. A natural system of chromosome transfer in Yersinia pseudotuberculosis. PLoS Genet. 2012, 8, e1002529. [Google Scholar] [CrossRef]
- Hurst, M.R.; Becher, S.A.; O’Callaghan, M. Nucleotide sequence of the Serratia entomophila plasmid pADAP and the Serratia proteamaculans pU143 plasmid virulence associated region. Plasmid 2011, 65, 32–41. [Google Scholar] [CrossRef]
- Ffrench-Constant, R.H.; Bowen, D.J. Novel insecticidal toxins from nematode-symbiotic bacteria. Cell. Mol. Life Sci. 2000, 57, 828–833. [Google Scholar] [CrossRef]
- Pendleton, I.R. Sodium and potassium fluxes in Philosamia ricini during Bacillus thuringiensis protein crystal intoxication. J. Invertebr. Pathol. 1970, 16, 313–314. [Google Scholar] [CrossRef]
- Bloomquist, J.R. Ion channels as targets for insecticides. Annu. Rev. Entomol. 1996, 41, 163–190. [Google Scholar] [CrossRef]
- Fast, P.G.; Angus, T.A. Effects of parasporal inclusions of Bacillus thuringiensis var. sotto ishiwata on the permeability of the gut wall of Bombyx mori (Linnaeus) larvae. J. Invertebr. Pathol. 1965, 7, 29–32. [Google Scholar] [CrossRef]
- Griego, V.M.; Moffet, D.; Spence, K.D. Inhibition of active K+ transport in the tobacco hornworm (Manduca sexta) midgut after ingestion of Bacillus thuringiensis endotoxin. J. Insect Physiol. 1979, 25, 283–288. [Google Scholar] [CrossRef]
© 2014 by the authors; licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution license (http://creativecommons.org/licenses/by/3.0/).