Bt toxins have been known as molecules that are active against insects and nematodes since the beginning of the previous century. They are synthesized by the soil-borne gram-positive bacterium Bacillus thuringiensis (Bt). About 400 Bt toxins are known so far produced by diverse B. thuringiensis strains (Crickmore et al. 2009). All of them have a crystal structure, therefore named Cry toxins. Because of the natural origin of the toxins, they occupy the position of the world's leading bio-pesticide.
Cry proteins bind to glycoprotein receptors that are located within the membrane of target insects' epithelium and afterwards inserted irreversibly into the membrane leading to the formation of a pore. Reasonably, alterations of these glycoprotein receptors can cause as a reason for toxin resistance of insects to a particular Bt-protein (Knight et al. 1994, 1995; Malik et al. 2001; Griffitts et al. 2005). B. thuringiensis strains produce different crystal proteins with specific activity against distinct species: Cry1A, Cry1B, Cry1C, Cry1H, Cry2A against lepidoptera, Cry3A, Cry6A, Cry 12A, Cry13A against nematodes, Cry3A, Cry6A against coleoptera and Cry10A, Cry11A against diptera. The toxins are effective tools for controlling lepidopteran and coleopteran insect pests, but application of Bt toxins as an insecticide by spraying is not efficient because the protein is unstable and has no systemic effect. In contrast, when synthesized by transgenic plants, Cry protoxins are taken up by sucking insects. Within the insect gut, protoxins are proteolytically cleaved to produce the active toxin, finally leading to affection on epithelial cells. So far, Bt toxins have been introduced into a wide range of crop plants like soybean, maize and cotton (see Chaps. 16, 19, 25). More than 20 transgenic crop varieties carry Cry genes (Bruderer and Leitner 2003). For instance, Cry1Ab is integrated into the genome of the transgenic maize varieties M0N810 and Bt176 (Bruderer and Leitner 2003), where it is particularly active against the european corn borer (Ostrinia nubilalis). In cotton the variety "Bollgard" expresses the Bt toxin Cry1Ac that is efficient for controlling the cotton bollworm (Helicoverpa armigera). To increase the expression levels of Bt toxins in transgenic plants, considerable changes to the Bt toxin genes are required such as change in codon-usage and the use of plant-specific processing signals in different events.
Even though immense advantages have been given by the use of Bt toxin in various transgenic crop plants (Romeis et al. 2006), the utilization of Bt toxin within transgenic plants is still controversially discussed, especially in Europe. Up to now, insect resistance against Bt toxins has not been observed under field conditions, only under laboratory conditions (Christou 2006), which is thought to be caused by a decreased fitness of resistant individuals (Christou et al. 2006; Soberon et al. 2007; Tabashnik et al. 2008). For instance, monitoring the pink bollworm (Pectinophora gossypiella) for eight years showed no increase of resistance to Bt (Tabashnik et al. 2005). The same result come from monitoring corn borers (Sesamia nonagrioides, Ostrinia nubilalis) in Spain over a period of five years (Farinos et al. 2004). Furthermore, an overview about environmental effects of Bt proteins was made by Clark et al. (2005). A negative effect on non-target organisms under true conditions was not observed (Romeis et al. 2006). By contrast, a meta-analysis showed an increased abundance of non-target invertebrates on Bt-transgenic cotton and maize fields, compared to non-transgenic fields managed with insecticides, as reported by Marvier et al. (2007). It is generally believed that the durability of resistance will be extended, e.g. by establishing refuges with areas of susceptible plants or by growing transgenic crops with a multi-gene, multi-mechanistic resistance (Boulter et al. 1993). The strategy of pyramiding effector genes within crops has two follow two major aims. One potential effect is to broaden insecticidal activity by combining genes with different specificity to control insect and nematode pests. The second effect is to enhance the durability of genetically engineered plant resistance because single mutation events do not break the insecticidal effect (Maqbool et al. 2001). Developing different strategies to protect the insecticidal effect of Bt toxins remains a great challenge (McGaughey et al. 1992; Frutos et al. 1999; Bates et al. 2005).
The potential for Bt toxin as a nematicide was reported by Marroquin et al. (2000). A preliminary study with transgenic tomato plants expressing the Bt endo-toxin CryIab after inoculation with Meloidogyne spp. resulted in a reduction in egg mass per gram of root of about 50% (Burrows and Waele 1997). The nematicidal effects were determined to result from a similar gut-damaging mechanism to that which occurs in insects: the activated toxin binds receptors in the intestine and forms a pore, causing lysis of the Gut (Wei et al. 2003; Li et al. 2007). Tomato hairy roots expressing the Bt crystal protein variant cry6A were challenged with M. incognita and supported significantly reduced amounts of nematode reproduction, although gall-forming ability was not affected (Li et al. 2007). The nematode feeding tube acts as a molecular sieve, permitting the uptake of certain molecules and excluding others. It is believed that root-knot nematodes are able to ingest larger molecules than cyst nematodes (Li et al. 2007). The size exclusion limit for H. schachtii has been determined to be approx. 23 kDa (Urwin et al. 1998). Therefore, the size exclusion limit (Bockenhoff and Grundler 1994; Urwin et al. 1997a, 1998; Li et al. 2007) severely restricts the agronomic application of trans-genic Bt as a broad-spectrum nematode control strategy (Fuller et al. 2008).
The expression of proteinase inhibitors (PIs) of digestive proteinases in plants is a promising strategy of engineering insect and nematode resistance. Compared to Bt toxin, the beneficial properties of proteinase inhibitors are their small size and stability for their expression in transgenic plants. A direct proof of activity against insects was shown in transgenic tobacco plants which were resistant against a bud worm mediated by the expression of a trypsin inhibitor (Hilder et al. 1987).
PIs represent a well studied class of plant defense proteins which are generated within storage organs. Proteinase inhibitors are an important element of natural plant defense strategies (Ryan 1990) and are anti-feedants known to reduce the capacity of certain parasites to use dietary protein, so delaying their development and reducing their fecundity (Hilder et al. 1987). In addition, it has been shown that PIs are induced as part of defense cascades, e.g. by insect attack, mechanical wounding, pathogen attack and UV exposure (Ryan 1999). Different kinds of proteinase inhibitors are known to reduce the digestibility of the nutrients through oral uptake by insects and nematodes. The inhibitor binds to the active site of the enzyme to form a complex with a very low dissociation constant, thus effectively blocking the active site.
There are ten groups of PI characterized from plants spanning all four classes of proteinases: cysteine, serine, metallo- and aspartyl. The majority of proteinase inhibitors studied in the plant kingdom originates from three main families, namely Leguminosae, Solanaceae and Gramineae (Rao et al. 1991). The cowpea trypsin inhibitor (CpTI) is a serine inhibitor used in the first transgenic approach to confer insect resistance. CpTI in an amount of 1% of the solouble protein in the transgenic plant has an effect on the lepidopteran insect Heliothis virens in tobacco (Hilder et al. 1987) and inhibits insect development up to 50%. The gene was also transferred into potato, rice and other plants, where it showed similar activity. Another effective gene is the sweet potato trypsin inhibitor (SpTI) that is active against Spodoptera litura when it is expressed in tobacco and Brassica spp. (Yeh et al. 1997b; Ding et al. 1998). Another group of PI, cysteine proteinases, is common in animals, eukaryotic microorganisms and bacteria, as well as in plants. Recent studies have shown that other classes of proteases are also found in insect guts, such as cysteine proteinase (Wolfson and Murdock 1990). Brioschi et al. (2007) reported that adaptation of the insects to proteinase inhibitors appears through upregulation of proteinases, trypsins and chymotrypsins by insects.
The potential of plant proteinase inhibitors (PIs) for engineering nematode resistance has been demonstrated in several laboratories (Vain et al. 1998; Urwin et al. 2000; Cai et al. 2003). Both serine and cysteine proteinases are present in plant-parasitic nematodes (Koritsas and Atkinson 1994; Lilley et al. 1997). Their activities have been detected in the nematode intestine where they are involved in digestion of dietary proteins (Lilley et al. 1996). Broad nematode resistance has been achieved in potato plants by expressing a cystatin from rice, even when the proteinase inhibitor was preferentially expressed in feeding sites of G. pallida and M. incognita (Lilley et al. 2004). A cysteine proteinase inhibitor based transgenic resistance to the cyst nematode Globodera pallida in potato plants proved to be effective, even under field conditions (Urwin et al. 2001), demonstrating its great potential.
We demonstrated that sporamin, a tuberous storage protein of sweet potato is a functionally trypsin proteinase inhibitor. The full-length sporamin gene encodes a 23-kDa mature protein (Yao et al. 2001). It can be taken up through the feeding tube and the stylet and delivered within the nematode, where it can exhibit effective inhibition. After its transfer into the sugar beet hairy roots, a significant reduction of developed females was observed in sporamin expressing roots but with variation in their inhibitory effects. Thereby the trypsin inhibitory activity was found to be a critical factor for nematode inhibition (Cai et al. 2003).
Nevertheless, there are no transgenic varieties carrying a proteinase inhibitor commercially available. It was discussed that parasites are able to modify there proteinase pattern and to bypass the inhibited protein digestion pathways (Broadway et al.1997; Giri et al. 1998). Thus, the source of the PI used in transgenic plants is critical to avoid development of insect insensibility (Ranjekar et al. 2003). Analogous to the case of Bt toxins, a combination of different PIs targeting a set of proteases would be a promising alternative to engineer a stable and broad resistance against insects and nematodes as well.
Lectins are a structurally heterogeneous group of carbohydrate-binding proteins which play biological roles in many cellular processes. More than 500 different plant lectins have been isolated and (partially) characterized. Application of lectins as insecticidal protein has mainly been focused on homopteran, e.g. planthoppers, leafhoppers and aphids (Habibi J et al. 1993; Hussain et al. 2008). Because of their low level of susceptibility to proteinase inhibitors, lectins were considered to be a suitable insecticidal agent.
The toxic effect of lectins to insects and nematodes is still poorly understood. The proteins seem to bind to cells of the insect/nematode midgut disrupting the cell function like digestive processes and nutrient assimilation. Insect-feeding studies with purified lectins and experiments with transgenic plants confirmed that at least some lectins enhance the plant's resistance against insects and nematodes. Several lectins from plants have been reported to confer broad insect resistance against Lepidoptera, Coleoptera, Diptera and Homoptera (Carlini and Crossi-de-Sa 2002). A gene encoding a sugar-binding protein derived from pea (Pisum sativum) was the first example of a lectin which was used to generate transgenic plants with an enhanced insect resistance (Boulter et al. 1990). Another famous example of a lectin used in transgenic plants is the Galanthus nivalis agglutinin (GNA), which confers resistance against insects in rice, e.g. planthoppers (Rao et al. 1998; Nagadhara et al. 2004). Moreover, expression of GNA in potato has been shown to confer enhanced resistance to lepidopterans like Lacanobia oleracea and homo-pteran insects like aphids (Down et al. 1996; Gatehouse et al. 1997). Rapeseed was successfully transformed with a pea lectin, which leads to a reduced weight of pollen beetle larvae that was correlated to lectin expression (Melander et al. 2003).
Also, a significant reduction of G. pallida females was reported after transfer of the gene encoding the snowdrop (Galanthus nivalis) lectin GNA into potato plants (Burrows et al. 1997). It is believed that, analogous to insects, these proteins could be targeted to interact with the nematode at different sites: within the intestine; on the surface coat; or with amphidial secretions.
A set of proteins from various organisms were tested for their activity against parasites. For instance, an insecticidal protein from scorpions enhances resistance to cotton bollworm (Heliothis armigera) larvae (Wu et al. 2008), toxins from endosymbionts of nematodes from the genus Photorhabdus and Xenorhabdus seems to have a broad insecticidal effect (Chattopadhyay et al. 2005).
Chitinases are known to be part of the plant defense system and are antifungal. A possible target is believed to be the nematode eggshell, which largely consists of chitin. We demonstrated recently that transgenic sugar beet roots and potato plants overexpressing a chitinase from the entomopathogenic fungus Paecilomyces javanicus confer broad-spectrum resistance to sedentary plant parasitic nematodes in transgenic sugar beet (B. vulgaris) and potato (Solanum tuberosum) plants (Thurau et al., unpublished data). The development of females was suppressed and the number of females was drastically reduced of both cyst nematodes Hetero-dera schachtii and Globodera pallida. In addition, the development of knots and egg sacks formed by root-knot nematode Meloidogyne incognita was also found to be severely affected. Although the mechanism underlying is not yet resolved and chitin has been reported to be present only in the egg shell of plant-parasitic nematodes so far, our results strongly suggest an active role of chitin also in the parasitic process of various nematodes, thus providing an effective target for genetic engineering of broad nematode-resistant crops.
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