Biotechnological application of functional genomics towards plant-parasitic nematode control
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Plant-parasitic nematodes are primary biotic factors limiting the crop production. Current nematode control strategies include nematicides, crop rotation and resistant cultivars, but each has serious limitations. RNA interference (RNAi) represents a major breakthrough in the application of functional genomics for plant-parasitic nematode control. RNAi-induced suppression of numerous genes essential for nematode development, reproduction or parasitism has been demonstrated, highlighting the considerable potential for using this strategy to control damaging pest populations. In an effort to find more suitable and effective gene targets for silencing, researchers are employing functional genomics methodologies, including genome sequencing and transcriptome profiling. Microarrays have been used for studying the interactions between nematodes and plant roots and to measure both plants and nematodes transcripts. Furthermore, laser capture microdissection has been applied for the precise dissection of nematode feeding sites (syncytia) to allow the study of gene expression specifically in syncytia. In the near future, small RNA sequencing techniques will provide more direct information for elucidating small RNA regulatory mechanisms in plants and specific gene silencing using artificial microRNAs should further improve the potential of targeted gene silencing as a strategy for nematode management.
Plant-parasitic nematodes represent a major biotic stress on world crops, causing over $100 billion in annual crop losses (Chitwood, 2003). Plant-parasitic nematodes exhibit a variety of feeding habits, ranging from migratory browsers to sedentary endoparasites with specialized host associations. The latter group, including the root-knot nematodes (RKNs; Meloidogyne spp.) and the cyst nematodes (CNs; Heterodera and Globodera spp.) are among the most important crop pests worldwide. Migratory endoparasitic nematodes, such as the root-lesion nematodes (RLNs; Pratylenchus spp.) and the burrowing nematodes (BN; Radopholus spp.), also are crop pests of widespread economic importance. Radopholus similis, for example, is found on more than 250 different plant species and is responsible for significant economic losses to the citrus industry (Sarah et al., 1996).
Sedentary endoparasites such as CNs and RKNs feed from modified host cells for prolonged periods of time. The infective second-stage juveniles (J2) of these nematodes establish multinucleate feeding sites in selected vascular tissues of host roots, using their stylets to pierce plant cell walls, inject salivary secretions and withdraw nutrients (Atkinson et al., 1988; Davis et al., 1992, 2000). Although the feeding cells of CNs and RKNs are formed by different processes, they uniformly serve as nutrient sinks for the growth and reproduction of sedentary stages (Sijmons et al., 1994; Hussey and Grundler, 1998; Gheysen and Fenoll, 2002). The feeding cell of CNs consists of a syncytium that contains around 200 merged root cells from the stele (Jones, 1981; Jung and Wyss, 1999), while RKNs induce multinucleate giant cells through acytokinetic mitosis (Sijmons et al., 1994; Gheysen and Fenoll, 2002). During feeding, a semi-permeable, blind-ended structure known as a feeding tube extends into the cytoplasm of the feeding cell from the stylet. In both groups, juveniles develop into either adult males or females, with adult males, when present, becoming motile, leaving the feeding site and, in the case of amphimictic species, mating with females. Reproduction typically occurs as amphimixis for CN species and through parthenogenesis for the RKN species (Siddiqi, 2000). Females produce hundreds of eggs that are laid in a gelatinous matrix (RKN) or mostly retained inside their body (CN).
Migratory endoparasitic nematodes remain mobile throughout their postembryonic life cycle and feed destructively from numerous plant cells. For example, RLNs and Radopholus similis puncture plant cell walls using their stylets, and with the aid of cell-wall degrading enzymes, they burrow through the root cortex causing extensive mechanical damage (Castillo and Vovlas, 2007; Haegeman et al., 2010). All life stages of females outside of the egg (J2-adult) are infective and capable of entering and exiting roots. Reproduction is either amphimictic or parthenogenetic depending on the species, and eggs are laid singly in root tissue or in the rhizosphere.
Limitations of current nematode control strategies
Management of sedentary plant-parasitic nematodes currently relies heavily on host-plant resistance, often derived from a relatively narrow genetic base through traditional breeding methods. In some cases, high levels of genetic diversity both within and among nematode populations confounds the use and limits the effectiveness of existing resistant cultivars, and selection pressure often results in resistance-breaking biotypes of the pathogens. Most populations of the soybean CN H. glycines, for instance, are now able to reproduce on soybean cultivars derived from PI88788, the most widely used source of H. glycines resistance (Mitchum et al., 2007; Hershman et al., 2008).
Nematicides have been widely used to control both migratory and sedentary plant-parasitic nematodes, but these compounds are often associated with detrimental environmental effects leading to substantially reduced availability in recent years. For example, methyl bromide, one of the most important chemical fumigants used to control nematodes and other pests, affects a wide range of organisms, including beneficial organisms, and was defined as a chemical that contributes to the depletion of the Earth’s ozone layer (Carpenter et al., 2001). As a result of the Montreal Protocol of 1991, methyl bromide has been phased out in 2005 in developed countries and will be phased out in developing countries by 2015. The neurotoxic organophosphate nematicide fenamiphos and carbamate nematicide carbofuran both were withdrawn from the U.S. market in 2007.
Crop rotation, a cultural practice effective for many biotic stresses, also is an important strategy for managing plant-parasitic nematodes, although its effectiveness is limited in many cases. Many nematodes, including RLN and RKN species, exhibit wide host ranges, encompassing both monocotyledonous and dicotyledonous plants, complicating management through crop rotation. Additionally, H. glycines cysts consist of many eggs that are able to remain dormant in the soil for up to 9 years (Inagaki and Tsutsumi, 1971), again limiting the effectiveness of crop rotation. Based on the limitations of current nematode control methods, the development of new strategies for plant-parasitic nematodes control should be a priority.
RNAi and its application in plant-parasitic nematode control
RNA interference (RNAi) refers to the suppression of gene expression through the use of sequence-specific, homologous RNA molecules. This phenomenon was first reported in Caenorhabditis elegans by Guo and Kemphues (1995) and subsequently demonstrated by Fire et al. (1998), who discovered that the presence of double-stranded RNA (dsRNA), formed from the annealing of sense and antisense strands present in the in vitro RNA preparations, is responsible for producing the interfering activity. Later, Bernstein et al. (2001) indicated that Dicer, a ribosome III-like enzyme, was responsible for processing dsRNA to ∼21 nucleotide (nt) sequences. The DCR-2/R2D2 complex binds to these small interfering RNA molecules (siRNAs) (Liu et al., 2003), then incorporated siRNAs into a multisubunit complex called the RNA induced silencing complex (RISC), which then dictates the degradation of any RNA molecules sharing sequence complementation (Hammond et al., 2000). This phenomenon of gene silencing occurs in several eukaryotic organisms, including nematodes and higher plants (Hammond et al., 2001). Researchers have been able to add in vitro-synthesized RNA directly to cells to obtain a gene knockdown. For example, Klink and Wolniak (2001) had been able to knockdown the centrin mRNA by dsRNA synthesized in vitro, and for the knockout effects, they demonstrated that dsRNA was at least ten times more effective than sense RNA or antisense RNA. To achieve plant-parasitic nematode control using an RNAi strategy, the RNAi mechanism is partially performed in planta and partially in the nematodes. siRNAs are generated by Dicer in plants expressing dsRNAs of nematode genes. When nematodes feed on plants, the plant-derived siRNAs or dsRNAs are taken up by nematodes through stylets, and RISC binds siRNAs to induce specific nematode genes’ degradation. siRNAs are then amplified in nematodes with the help of RNA-dependent RNA polymerase (RDRP) (Zamore and Haley, 2005; Chapman and Carrington, 2007).
This siRNA-mediated gene silencing is highly sequence-specific. For example, Tuschl and colleagues demonstrated that even a single base mismatch between a siRNA and its mRNA target prevented gene silencing (Elbashir et al., 2001). Even though it is gene specific, it is possible to silence gene families with a single sequence (Miki et al., 2005) or with a chimeric hairpin construction (Allen et al., 2004). The RNAi effect has been documented to spread not only from cell to cell (Fagard and Vaucheret, 2000; Kehr and Buhtz, 2008), but also throughout the plant (Yoo et al., 2004). Indirect evidence indicates that RNA silencing moves over long distances through the phloem and, upon unloading, spreads from cell to cell through plasmodesmata in recipient tissues (Voinnet et al., 1998; Jorgensen, 2002; Mlotshwa et al., 2002). Likewise, Himber et al. (2003) demonstrated that both localized and long-distance movement of the RNAi effect can occur. Limpens et al. (2004) also found that silencing signal was transported systemically from Arabidopsis roots to shoots, although extent of silencing was limited and greatly variable.
Although transformation of plant-parasitic nematodes represents one approach for deploying RNAi strategy on nematodes control, there are no successful reports of engineering these nematodes, partly because of their obligate nature of parasitism. In addition, this approach would have numerous regulatory hurdles for releasing these GMO nematodes into the environment. In C. elegans, RNAi has been used to transiently knockdown the expression of nearly all C. elegans genes and phenotypic effects including lethality have been observed for thousands of the C. elegans genes (Fraser et al., 2000; Gönczy et al., 2000; Maeda et al., 2001; Kamath et al., 2003). RNAi can be accomplished in C. elegans by ingestion of bacteria expressing dsRNA (Timmons and Fire, 1998), oral uptake of dsRNA from solution (Tabara et al., 1998) or microinjection (Mello and Conte, 2004). Of these three methods, soaking nematodes in dsRNA solutions and host-delivered RNAi are two strategies that have been widely explored. RNAi by feeding on bacteria expressing dsRNAs is not feasible for plant-parasitic nematodes because they are primarily obligate parasites.
Plant-parasitic nematodes are capable of ingesting large macromolecules from plants
Size exclusion of molecules moving from plants to nematodes has been a concern for delivery of nematicidal dsRNAs and proteins to parasites (Urwin et al., 1997, 1998). To better understand the molecular size cut-off for RKN ingestion, transgenic tomato roots expressing 54-kDa Bacillus thuringiensis (Bt) crystal (Cry) proteins Cry6A were challenged with M. incognita, and they observed RKNs successfully ingested this large molecule (Li et al., 2007), substantially increasing the known size of molecular uptake into RKNs. Therefore, based on molecule size, ingestion of dsRNAs is unlikely to be a challenge for RKNs. Similarly, CNs were also demonstrated to uptake large macromolecules. The feeding tube of Heterodera schachtii could uptake dextran molecules 20 kDa in size but not 40 kDa (Bockenhoff and Grundler, 1994). Urwin et al. (2002) proved using octopamine to stimulate dsRNAs uptake by preparasitic second-stage juveniles of H. glycines. dsRNA molecules ranging in size from 42 to 1300 bp have all proved effective in inducing RNAi in both CNs and RKNs (Chen et al., 2005; Huang et al., 2006; Kimber et al., 2007). For example, Huang et al. (2006) generated transgenic Arabidopsis plants expressing dsRNAs of full length (271-bp) and truncated (42-bp) 16D10 gene, and these transgenic lines all showed 63%–90% reduction for the RKN galls. In another case, dsRNA of 227 and 316-bp Globodera pallida flp gene were confirmed efficient to inhibit the worm growth (Kimber et al., 2007).
RNAi by soaking nematodes in dsRNA solutions
dsRNAs can be taken up by nematode intestinal cells after ingestion or through the cuticle during soaking. Delivery of dsRNAs by ingestion is difficult, as the sedentary nematodes typically only feed following the establishment of a feeding site inside the root and do not ingest substances prior to this stage. However, Urwin et al. (2002) demonstrated that with the use of octopamine, they could stimulate the uptake of dsRNAs by preparasitic second-stage juveniles of two CNs, H. glycines and G. pallida. Following this new technique for facilitating nematode uptake, several groups had successfully suppressed mRNAs of target nematode genes and reduced the number of established nematodes or suppressed the development of nematodes by soaking plant-parasitic nematodes in dsRNAs (Bakhetia et al., 2005a, 2007; Chen et al., 2005; Lilley et al., 2005; Huang et al., 2006; Alkharouf et al., 2007; Shingles et al., 2007). Other research groups have also efficiently suppressed the development of nematodes with or without the addition of different chemicals to induce uptake (Fanelli et al., 2005; Dubreuil et al., 2007; Kimber et al., 2007; Park et al., 2008). Although many of these reports did not mention the stability of gene silencing, it appears that soaking nematodes in dsRNA solutions is transitory in the reports that examined these phenomena. In H. glycines, cellulase transcript recovery was observed 6–10 days after soaking (Bakhetia et al., 2007); in M. incognita, recovery of calreticulin, polygalacturonase and GST transcripts was found 2–3 days after soaking (Rosso et al., 2005). The general conclusion from these reports is that the soaking method of preparasitic nematodes can be effective in evaluating gene functions, although in some cases, transitory nature enabled nematodes to recover in short time after soaking with dsRNA solutions.
Host-delivered RNAi to silence nematode genes
The pathogenic nematodes discussed in this review are obligate parasites of plant roots, making host-delivered dsRNAs an ideal strategy for silencing genes essential to the parasites, as well as for providing direct evidences of the function of the genes. Accumulated research has confirmed the feasibility and effectiveness of host-delivered RNAi for nematode control. Yadav et al. (2006) reported that RNAi was induced by using dsRNAs of two genes encoding an integrase and a splicing factor in the plant-parasitic nematode M. incognita, leading to protection against nematode infection in tobacco. The expression of RKN parasitism gene 16D10 dsRNA in transgenic Arabidopsis plants resulted in resistance against four major RKN species (Huang et al., 2006), while Sindhu et al. (2009) obtained reductions in H. schachtii females ranging from 23% to 64% in transgenic Arabidopsis lines expressing RNAi of four parasitism genes. RNA interference appears to be similarly effective against H. glycines in transformed soybean lines. Steeves et al. (2006) successfully produced transgenic soybean lines using this RNAi strategy targeted to a major sperm protein of H. glycines. Bioassay data indicated transgenic plants had up to a 68% reduction in eggs/g root tissue. The effects of plant-derived dsRNA molecules appeared to continue into the next generation. Klink et al. (2009b) effectively inhibited female H. glycines cyst formation by inverted repeats of several H. glycines homologs. Recently, Li et al. (2010a,b) successfully suppressed H. glycines reproduction in soybean plants by RNAi targeting three different H. glycines genes. Direct molecular evidences of host-derived RNAi down-regulating target nematode genes were observed from nematodes feeding on transgenic roots via real time RT-PCR analysis (Sindhu et al., 2009; Li et al., 2010a). Transgenic plants expressing dsRNAs of nematode genes have been more successful in RKN than in CNs. The likely reasons are the size exclusion limit of RKNs is considerably larger than that of CNs (Bockenhoff and Grundler, 1994; Lilley et al., 2007) and another possible reason is RNAi may be more sensitive in RKNs than in CNs.
The traditional transformation methodologies for many important plant species take at least several months to produce stable transgenic lines with substantial greenhouse space and labour. To rapidly assess target genes in planta, high-throughput composite or chimeric hairy root systems have been established in several systems including soybean (Cho et al., 2000; Klink et al., 2009b; Li et al., 2010b), sugar beet (Kifle et al., 1999; Cai et al., 2003) and tomato (Remeeus et al., 1998). For instance, using a composite hairy root method, Li et al. (2010b) have reduced the screening process by 8 months in comparison of traditional soybean transgenic approach. Klink et al. (2009b) and Li et al. (2010a,b) have identified several effective H. glycines genes using these hairy root systems. The effective nematode genes identified by the hairy root transgenic system can then be stably transformed into soybean plants to obtain transgenic seeds for further analysis. Another hurdle for large scale screening of target genes is that the traditional RNAi construction need to ligate together the sense, antisense and a linker into the destination vector, which is labour intensive and time-consuming. Therefore, Klink et al. (2009b) and Li et al. (2010a,b) established the Gateway cloning system for RNAi construction, which substantially reduced the time for cloning genes into RNAi constructs.
Characterization of target genes
To control plant-parasitic nematodes, several research groups tried to deploy RNAi strategy to block life cycle of nematodes, prevent the parasitism or block the transcription or translation process of nematode genes. In the past few years, a number of nematode genes have been targets for gene silencing. As a result of these investigations, Table 1 lists the target genes reported to be effective in suppressing nematode populations. The target genes can be divided into the following three categories: parasitism genes, developmental genes and mRNA metabolism-related genes.
Table 1. Plant-parasitic nematodes effectively controlled by RNAi targeting different genes
|hgctl, AF498244||C-type lectin||H. glycines||Soaking||41% decrease in no. of established nematodes||Urwin et al. (2002)|
|hgcp-I||Cysteine proteinase||H. glycines||Soaking||40% decrease in no. of established nematodes||Urwin et al. (2002)|
|pMiDuox1, DQ082753||Dual oxidase||M. incognita||Soaking||Up to 70% decrease in no. of established nematodes |
Decrease in fecundity
|Bakhetia et al. (2005a)|
|Gr-eng-1, AF004523||β-1,4-endoglucanase pharyngeal||G. rostochiensis||Soaking||Around 50% decrease in no. of established nematodes||Chen et al. (2005)|
|Gr-ams-1, AJ270995||Secreted amphid protein||G.rostochiensis||Soaking||Reduced ability to locate and invade roots||Chen et al. (2005)|
|AY013285||Chitin synthase||M. artiellia||Soaking||Delayed egg hatch||Fanelli et al. (2005)|
|Hg-amp-1, AY883023||Aminopeptidase||H. glycines||Soaking||61% decrease in number of female reproductive||Lilley et al. (2005)|
|16D10, DQ841121–DQ841123||Secreted peptide||M. incognita||Soaking||74%–81%` decrease in no. of established nematodes||Huang et al. (2006)|
|Hg-rps-23,BF014259||Ribosomal protein||H. glycines||Soaking||Decrease in J2 viability||Alkharouf et al. (2007)|
|hg-eng-1,AF006052||β-1,4-endoglucanase||H. glycines||Soaking||Decrease in no. of established nematodes||Bakhetia et al. (2007)|
|hg-syv46, AF273728||Secreted peptide SYV46||H. glycines||Soaking||Decrease in no. of established nematodes||Bakhetia et al. (2007)|
|Mi-gsts-1, EL784458||Glutathione-S transferase||M. incognita||Soaking||52%–71% decreased in fecundity||Dubreuil et al. (2007)|
|Flp||FMRFamide-like peptides||G. pallida||Soaking||Inhibition of motility||Kimber et al. (2007)|
|Mi-cpl-1||Cysteine proteinase||M. incognita||Soaking||60% decrease in no. of established nematodes||Shingles et al. (2007)|
|Bx-hsp-1, DQ785812||Heat shock protein 70, cytochrome C||B. xylophilus||Soaking||J2-J3 viability reduced at high temperature||Park et al. (2008)|
|Bx-myo-3||Myosin heavy chain, tropomyosin||B. xylophilus||Soaking||J2-J3 viability reduced at high temperature abnormal locomotion||Park et al. (2008)|
|AW871671||Integrase||M. incognita||In planta||>90% reduction in established nematodes||Yadav et al. (2006)|
|AW828516||Splicing factor||M. incognita||In planta||>90% reduction in established nematodes||Yadav et al. (2006)|
|16D10, DQ841121–DQ841123||Secreted peptide||M. arenaria, M. incognita, M. javanica, M. hapla, M. arenaria||In planta||63%–90% reduction no. of galls and gall size||Huang et al. (2006)|
|MSP||Major sperm protein||H. glycines||In planta||Up to 68% reduction in nematode eggs||Steeves et al. (2006)|
|Hg-rps-3a, CB379877||Ribosomal protein 3a||H. glycines||In planta||87% reduction in female cysts||Klink et al. (2009b)|
|Hg-rps-4, CB278739||Ribosomal protein 4||H. glycines||In planta||81% reduction in female cysts||Klink et al. (2009b)|
|Hg-spk-1, BI451523.1||Spliceosomal SR protein||H. glycines||In planta||88% reduction in female cysts||Klink et al. (2009b)|
|Hg-snb-1,BF014436||Synaptobrevin||H. glycines||In planta||93% reduction in female cysts||Klink et al. (2009b)|
|4G06, AF469060||Ubiquitin-like||H. schachtii||In planta||23%–64% reduction in developing females||Sindhu et al. (2009)|
|3B05, AF469058||Cellulose binding protein||H. schachtii||In planta||12%–47% reduction in developing females||Sindhu et al. (2009)|
|8H07, AF500024||SKP1-like||H. schachtii||In planta||>50% reduction in developing females||Sindhu et al. (2009)|
|10A06, AF502391||Zinc finger protein||H. schachtii||In planta||42% reduction in developing females||Sindhu et al. (2009)|
|Y25, CB824330||Beta subunit of the COPI complex||H. glycines||In planta||81% reduction in nematode eggs||Li et al. (2010a,b)|
|Prp-17, AF113915||Pre-mRNA splicing factor||H. glycines||In planta||79% reduction in nematode eggs||Li et al. (2010a)|
|Cpn-1, GU074018||Unknown protein||H. glycines||In planta||95% reduction in nematode eggs||Li et al. (2010a)|
During the feeding process, some proteins encoded by nematode parasitism genes are secreted through the nematode stylets into plant tissues. RNAi experiments on parasitism genes suggested that they may play a very important roles for nematodes invading plants. The importance of dual oxidases (peroxidase and NADPH oxidase) associated with extracellular matrix of nematodes was investigated in M. incognita by in vitro RNAi studies (Bakhetia et al., 2005a). Hg-syv46 protein product has similarity to the CLE family of A. thaliana, which is involved in controlling the balance between meristem cell proliferation and differentiation, therefore, it may produce a ligand that disrupts the normal pattern of plant cell differentiation at the feeding site (Bakhetia et al., 2007). Some nematode secreted proteasome members such as SKP-1 or Ring-H2 as well as ubiquitin-like proteins may regulate host cell protein degradation to their parasitic advantage (Sindhu et al., 2009). H. glycines and G. rostochiensisβ-1-4 endoglucanases are degradating plant tissues to facilitate invasion and migration (Chen et al., 2005; Bakhetia et al., 2007). Both cysteine proteinase genes from M. incognita and H. glycines displayed very important roles for nematode invading plants as RNAi of them resulted in significant decrease in established nematodes on plants, and in situ hybridization analysis suggested its product is a digestive enzyme (Urwin et al., 2002; Shingles et al., 2007). Parasitism gene 16D10 encodes a conserved RKN secretory peptide that stimulates root growth and functions as a ligand for a putative plant transcription factor, and this essential parasitism gene led to the availability of a resistance gene effective against four major RKN species (Huang et al., 2006). C-type lection has two lectin domains providing sequence homology with a cobra venom coagulation factor, macrophage mannose receptor and proteoglycan core proteins such as aggrecan (Urwin et al., 2002). One secreted amphid protein from G. rostochiensis is critical for host location as RNAi of the gene encoding the amphid protein resulted in reduced ability of nematode to locate and invade roots (Chen et al., 2005). The observed Mi-gsts-1 expression in the oesophageal secretory glands and the results of functional analyses based on RNA interference suggest that glutathione S transferases are secreted during parasitism and are required for completion of the nematode life cycle in its host (Dubreuil et al., 2007).
Genes involved in nematode development
Genes involved in nematode development, especially those in embryogenesis, juvenile development and reproduction are of particular interests because they have the potential to disrupt the life cycle of parasitic nematodes. M. artiellia egg hatching was delayed because a chitin synthase gene which is responsible for chitin production in the eggshells was suppressed by RNAi (Fanelli et al., 2005). Steeves et al. (2006) reported transgenic soybean plants expressing dsRNAs of a gene encoding a major sperm protein significantly reduced the reproductive potential of H. glycines, and progeny nematodes were also impaired in their ability to reproduce. Kimber et al. (2007) confirmed that flp gene disruption in potato CN G. pallida reveals motor dysfunction and unusual neuronal sensitivity to RNA interference.
Potential targets for RNAi suppression of plant-parasitic nematodes can be selected based on homologs of known embryonic or maternal lethal or sterile RNAi phenotypes/phenocopies in C. elegans. For instance, Alkharouf et al. (2007) used bioinformatics to yield 1508 candidate H. glycines genes whose homologous genes of C. elegans have lethal or sterile phenotypes/phenocopies when silenced in C. elegans, and through in vitro RNAi tests, they found one of the genes among them encoding ribosomal protein rps-23 is important for H. glycines development. Klink et al. (2009b) demonstrated that 32 of 150 conserved H. glycines homologs of C. elegans genes with RNAi lethal phenotypes/phenocopies were induced during feeding site establishment, and subsequently inhibited female development by engineering transgenic soybean plants with tandem inverted repeats of four selected homologs encoding small ribosomal protein 3a and 4, a spliceosomal SR protein and synaptobrevin. Using the same strategy, Li et al. (2010a,b) showed the RNAi of three genes coding for a beta subunit of the coatomer (COPI) complex, a pre-mRNA splicing factor and an unknown protein led to significant reduction for H. glycines cysts and eggs.
Genes associated with mRNA metabolism
RNAi of nematode genes involved in mRNA metabolism appears to be very effective for suppressing nematode development or reproduction. Yadav et al. (2006) reported that double-stranded RNA fragments of two genes encoding an integrase and a splicing factor induced RNAi in the plant-parasitic nematode M. incognita and protected tobacco against infection. Greater than 95% lethality of pi-J2 was observed for H. glycines soaked in dsRNA solution of a ribosomal gene Hg-rps-23 (Alkharouf et al., 2007). In addition, soybean roots expressing inverted repeat constructs of three genes Hg-rps-3a, Hg-rps-4 and Hg-spk-1 involved in different aspects of mRNA metabolism displayed 81%–88% reductions in numbers of H. glycines cysts (Klink et al., 2009b). RNAi of the H. glycines Prp-17 gene which encodes a mRNA splicing factor resulted in significant reductions in cysts/g root tissue (53% reduction) and eggs/g root tissue (79% reduction) in the transgenic soybean plants inoculated with H. glycines, which supported the important role of Prp-17 gene in C. elegans (Li et al., 2010a). All of these genes are involved in mRNA metabolism, suggesting that genes involved in mRNA metabolism may be particularly sensitive to RNAi and should, therefore, make good target genes for parasitic nematode control.
The benefits of RNAi strategy
RNAi as a resistance strategy has several benefits to offer compared to current nematode control methods. First, interactions with nontarget organisms can be minimized. RNAi is a highly specific method for gene silencing. For example, Tuschl and colleagues demonstrated that even a single base mismatch between a siRNA and its mRNA target prevented gene silencing (Elbashir et al., 2001). There is evidence, however, of off-target effects. In C. elgans, gene silencing can be induced if an ‘off-target’ gene has 30–50 nucleotide similarity with the sequence targeted for silencing (Rual et al., 2007). Proper selection of target sequence—that is selecting gene sequence targets that have limited homology with plants, higher animals and beneficial, free-living nematodes—is paramount to minimize off-target effects. Second, even though RNAi is gene specific, it is possible to silence gene families with a single sequence if RNAi is targeting a gene sequence conserved among gene families. Targeting several genes with a single construction should correlate with a greater RNAi effect. Third, current genetic resistance is based on a multigenic complex, making breeding for nematode resistance difficult. RNAi strategy could create a single resistant locus, which will facilitate the transfer of nematode resistance to other crop cultivars. Fourth, the long-range durability of nematode protection may also be achieved by deployment this type of resistance strategy. Depending on the target gene and sequences selected for the RNAi construction, durability can be maintained with the RNAi strategy, especially if a mutation in the targeted gene would be considered lethal. By selecting sequences that show high homology among nematode populations, it should be possible to maximize the effectiveness for durable resistance. However, long-term field and greenhouse studies will be needed to verify durability. In addition, it is possible to engineer an RNAi locus to target multiple genes, making an RNAi-tolerant nematode caused by a random mutation highly unlikely.
Other available functional genomics tools for facilitating target genes selection for gene silencing
Laser capture microdissection (LCM) and microarray
To facilitate plant resistance against parasitic nematodes, a few gene expression approaches such as microarray and LCM were deployed for studying the interaction between nematodes and plants. To specifically study gene expression at the feeding site, LCM was used to isolate syncytial cells of H. glycines (Klink et al., 2005, 2007). Distinct differences in gene expression were observed between syncytia undergoing resistant and susceptible responses by microarray expression of LCM isolated syncytia (Klink et al., 2007).
Moreover, using the Affymetrix GeneChip soybean genome array, 429 genes were identified showing significantly differential expression between nematode infected and noninfected root tissues for a susceptible reaction between soybean and H. glycine (Ithal et al., 2007). These genes included those encoding enzymes involved in primary metabolism; biosynthesis of phenolic compounds, lignin and flavonoids; genes related to stress and defence responses; cell-wall modification; cellular signalling; and transcriptional regulation. In the same year, another microarray analysis was conducted on the soybean cultivar ‘Peking’ infected with the incompatible H. glycines population NL1-RHg, HG-type 7. The results indicated that 71 genes were induced in the incompatible NL1-RHg population, compared with H. glycines compatible TN8 populations (baseline) (Klink et al., 2007). Furthermore, Klink et al. (2009a) performed gene expression analysis of PI 548402 (Peking) root syncytia undergoing a resistant reaction after soybean cyst nematod (SCN) infection and found lipoxygenase-9 and lipoxygenase-4 are most highly induced genes, and identified the components of the phenylpropanoid pathway was induced, suggesting that the jasmonic acid and phenylpropanoid signalling pathway at the site of the syncytium may play important roles for the resistant reaction. For another resistant reaction of G. max ([PI 88788]) to H. glycines ([NL1-RHg/HG-type 7]), Klink et al. (2010a) further strongly confirmed that the jasmonic acid defence pathway acts as a factor involved in the localized resistant reaction. Recently, a comparative microarray investigation was performed using detection call methodology to identify genes possibly cell-type specific and/or involved in important aspects of plant nematode interactions during the resistance response (Klink et al., 2010b).
Genomic and EST sequencing of essential genes from plant-parasitic nematodes
James McCarter’s group at Divergence Inc has led the public sequencing of over 250 000 ESTs from 30 nematode species including >100 000 plant parasite ESTs (Parkinson et al., 2004; McCarter et al., 2003; http://www.nematode.net). The M. hapla genome was sequenced and mapped (Opperman et al., 2008). In addition, the genome sequence from M. incognita has recently become available (Abad et al., 2008; McCarter, 2008). To increase the likelihood of identifying genes beyond those represented by public ESTs, Monsanto and Divergence generated ∼3X coverage of the H. glycines 93 megabase genome (Boukharov et al., 2007). In the year 2008, Divergence and Monsanto jointly released their draft H. glycines genome through NCBI. All the genomic and EST sequences of parasitic nematodes provided remarkably valuable sequence information that will enable researchers to find suitable target genes for continued RNAi experiments.
Enhance nematode resistance in transgenic plants by modifying current RNAi strategies
In general, the RNAi mechanism for gene silencing is based on a large amplification of siRNA molecules that bind to a specific gene transcript. The current methodology produces only siRNAs that correspond to the specific sequence fragments found in the RNAi constructs. The quantity of siRNA species does not increase exponentially because the nematode gene target is not found in the plants (Gheysen and Vanholme, 2007). Moreover, the absence of an endogenous, homologous gene expression in plant cells might induce off-target effects (Bakhetia et al., 2005b). To increase the amount of siRNAs in transgenic plants, over-expressing a nematode gene in plants to obtain RNA templates should be a good option. Stacking two or more effective genes’ RNAi cassettes in one RNAi vector should provide more durable or robust resistance against plant-parasitic nematodes, as engineering an RNAi locus to target multiple genes should make an RNAi-tolerant nematode caused by a random mutation highly unlikely.
Artificial microRNAs (amiRNAs)
MicroRNAs (miRNA) is an alternative method for gene silencing. miRNAs are generated by Dicer from short hairpin structures miRNA precursors (pre-miRNA) which are initially derived from longer primary miRNA transcripts (pri-miRNA) (Brodersen and Voinnet, 2006). The mature miRNAs are 20–24 nucleotides long endogenous single-stranded small RNA molecules, which are incorporated into RISC to guide mRNA degradation. Vaucheret et al. (2004) demonstrated that several nucleotides changes within an 21-nt miRNA sequence did not affect its biogenesis. Following this report, artificial miRNAs (amiRNAs) have successfully been shown to specifically regulate gene expression by RNA silencing in various plant organisms such as rice (Wang et al., 2010), Arabidopsis (Schwab et al., 2006; Liu et al., 2010) and Nicotiana benthamiana (Tang et al., 2010). The main advantage of amiRNA vectors compared to traditional RNAi vectors containing inverted repeats of target genes is that amiRNA vectors can lead to more specific gene silencing effect as amiRNA vectors contain a single 21-nt sequence complementary to target gene. The current or ‘traditional’ RNAi vectors usually contain inverted repeats of more than 100-bp sequences resulting in a population of at least several different species of siRNAs and the potential for ‘off-target’ gene silencing. These ‘off-target’ effects in the host plants can become a crucial consideration in the RNAi experiments (Gheysen and Vanholme, 2007; Sukno et al., 2007). Recently, Melito et al. (2010) deployed amiRNA technology to study the role of rhg1 locus LRR-kinase gene for the interaction between soybean and SCN and observed rhg1 locus LRR-kinase had no significant impact for SCN resistance. Taken together, there is little doubt that this technique will play a major role in nematode resistance in the future.
Small RNA sequencing
Small RNAs, especially microRNAs (miRNAs), have been implicated in a variety of physiological and morphological processes through computational and cloning approaches. Deep sequencing of small RNA populations in animals, plants, fungi and protozoa have helped to elucidate the regulatory mechanisms of small RNAs (Lu et al., 2005; Nobuta et al., 2008; Tuteja et al., 2009; Wu et al., 2010). For example, small RNA sequencing results supported hypothesis of how endogenous gene suppression functions in maize (Houmard et al., 2007) and petunia (De et al., 2009). Small RNA sequencing of transgenic plants expressing RNAi vectors for nematode resistance could provide insight to populations of siRNA species generated and may help to find the specific target regions to design artificial miRNA vectors. However, no small RNA sequencing results from transgenic plants expressing dsRNAs of nematode genes nor nematodes paratizing such plants were published to our knowledge. Nonetheless, small RNA sequencing is a field that is growing and should continue to provide more direct information for elucidating small RNA regulatory mechanisms in plants and other organisms.
Andrew Fire and Craig Mello were awarded 2006 Nobel Prize award in Physiology or Medicine for the discovery of ‘RNA interference—gene silencing by double-stranded RNA’. In addition to the numerous applications this technology has in other systems, the RNAi strategy has been explored by many researchers as an efficient tool for identifying genes’ functions in plant-parasitic nematodes. Results from RNAi experiments have been very encouraging in regard to sedentary plant-parasitic nematode controls, although additional research is needed. It also remains to be seen whether migratory nematode species are equally susceptible to plant-delivered RNAi. This novel approach to nematode control can be made environmentally friendly but selection of target genes is essential to minimize or prevent off-target interactions. It will also be necessary to perform proper ecological risk assessment with this new technology. Scientists are increasing the use of functional genomics tools such as genome sequencing or microarray to facilitate finding more suitable gene targets for silencing by RNAi. Small RNA sequencing approach will provide more direct information for understanding RNAi machinery in plants, and new approach artificial miRNA promises to bring more precision and predictability to the RNAi technology.
This research was supported by Kansas State University and the Kansas Soybean Commission. This article is contribution no. 11-141-J from the Kansas Agricultural Experimental Station, Kansas State University, Manhattan, Kansas.