Journal Search Engine
Search Advanced Search Adode Reader(link)
Download PDF Export Citaion korean bibliography PMC previewer
ISSN : 1225-8504(Print)
ISSN : 2287-8165(Online)
Journal of the Korean Society of International Agricultue Vol.24 No.4 pp.485-493

RNAi 기술을 이용한 뿌리혹선충 저항성 식물 개발 동향

헤마바티 아자팔라, 심준수, 한범수
농촌진흥청 국립농업과학원
RNAi 기법은 예쁜꼬마선충을 비롯하여 초파리, 생쥐, 인체 및 식물들을 포함한 다양한 생물종에서 작용기작이 잘 알려져있고, 특정 유전자들의 발현을 제어하기 위해 사용되어 왔다. 본 논문은 RNAi 기법을 통해 구축된 뿌리혹선충 저항성 형질전환식물체들의 개발현황, 발전 및 응용 가능성에 대해 고찰하고자 하였다.
지난 10여년에 걸쳐 진행된 연구들을 통해 다양한 분화단계의 뿌리혹선충으로부터 분석된 79,978개의 EST가 GenBank에 등록되었고, 고구마뿌리혹선충(M. incognita)과 당근뿌리혹선충(M. hapla)의 전체 게놈 염기서열이 해독됨으로써 뿌리혹선충의 기생에 관련된 단백질들 및 식물세포벽 분해 관련 효소들에 대한 정보를 이용할 수 있게 되었다. RNAi의 기본 기작은 모든 진핵생물종에서 잘 보존되어 있고, 최근 식물기생선충들에서 RNAi 효과에 대한 연구결과들이 많이 발표되었다. 서로 다른 뿌리혹선충들에서 현재까지 22종 이상의 RNAi를 위한 목표 유전자들이 보고되었다. 제2령 식물기생선충(second-stage juvenile of plant parasitic nematodes)에게 dsRNA의 섭취를 유도하는 octopamine, resorcinol, serotonin 등의 화합물들이 발견되었고, 이를 이용하여 선충 유전자의 발현제어를 쉽게 판별할 수 있는 새로운 기술의 활용이 가능하게 되었다.
최근에 RNAi 기술의 응용 및 발전을 통해 형질전환식물체에서 발현된 dsRNA에 의한 식물기생선충 유전자들의 발현제어가 증명되었고, 이를 위한 핵심 요소들로서 적절한 선충 표적 유전자의 선택, 식물체 내 높은 함량의 dsRNA의 발현 및 선충이 섭취할 수 있는 충분한 양의 dsRNA의 운반 등이 중요하다는 것이 밝혀졌다. 특히, 비표적 유전자서열(off-target gene sequence)의 발현제어를 피하기 위해 다음과 같은 사항들이 고려되어야 한다. 1)비표적 유전자서열을 확인할 수 있는 소프트웨어의 개발 및 이를 통한 비표적 유전자서열을 제거한 RNAi 벡터를 제작하여야 한다. 2)식물과 동물에서 상동성이 높은 표적 유전자의 발현을 피해야 한다. 3)전사해석틀(open reading frames)의 염기서열들 보다 상동성이 낮은 5' 혹은 3'-비해석부위(untranslated regions)로부터 표적 유전자를 설계하여야 한다.

RNA Interference Silencing in Root-knot Nematodes

Bum-Soo Hahn, Hemavathi Ajjappala, Joon-Soo Sim
National Academy of Agricultural Science, RDA
Received Oct 15, 2012 / Revised Nov. 20, 2012 / Accepted Dec. 7, 2012


The root-knot nematodes (RKN) belonging to the genus Meloidogyne are obligate andsedentary endoparasites that infect wide range of plant species. By parasitizing the root system, they dis-rupt the water and nutrient uptake thus affecting the whole plant causing significant reduction in thecrop yield. They pose huge losses of several million dollars worldwide. Use of chemical nematicide,including soil fumigants such as methyl bromide was the most reliable strategy of controlling the root-knot nematodes. Recently, molecular strategies such as RNA interference (RNAi) mediated nematodegene silencing, R gene transformation, expression of proteins detrimental to RKN are gaining impor-tance. RNAi is used for downregulating the expression of specific gene in several organisms, particularlyfree-living nematode Caenorhabditis elegans. RNAi strategy has been widely used for silencing the targetgenes in plant parasitic nematodes. Nearly 79,978 ESTs analysed from various developmental stages ofMeloidogyne species has gained entry into the GenBank. The complete genome sequencing of M. incog-nita and M. hapla provides information regarding the parasitism genes and their success in parasitizingthe plants. Recent RNAi approach has resulted in successful reports suggesting suppression of genesessential for nematode development, survival and parasitism in plants. These studies may further serveas a foundation for identification of novel genes and their role in parasitism and could be used as targetgenes for silencing thus conferring resistance to plants. This review focuses on the progress madetowards the development of transgenic plants resistant to RKN through RNAi approach.

Plant parasitic nematodes are obligate parasites which are found with almost all crops that are of economic impor-tance. According to the 1987 international opinion survey, an estimated annual crop yield loss due to nematodes was found to be US $ 125 billion (Sasser & Freckman, 1987). Commer-cially important crops such as rice, maize, wheat, tomato, grape, banana, tobacco, coffee, and cotton are among the crops that are severely affected by nematodes. Comparison of 1987 international loss estimates by crop to that of 2001 data for crops, yield loss in US dollars is represented in Fig. 1.

Fig. 1. Annual crop yield loss in US dollars (billion) by nematodes in 2001 based on 1987 international yield loss survey (Sasser & Freckman, 1987). Ten major crops include rice, maize, potato, wheat, tomato, grape, sugarcane, yam, soybean, banana. Figure adapted and modified from McCarter, 2009.

The root-knot nematodes (RKN),  Meloidogyne  species, are sedentary endoparasitic species which are economically the most important and wide spread throughout the world with a broad host range. Meloidogyne chitwoodi was the first species described from the Pacific Northwest of the USA in 1980 (Golden et al., 1980) which caused severe damage to potato tubers.  Meloidogyne species such as  Meloidogyne incognita  and  Meloidogyne javanica  have been found at destructive levels in almost all maize growing regions of the world.  Meloidogyne africana  and  Meloidogyne arenaria have been also found infecting maize in India and in Pakistan(McDonald & Nicol, 2005). The symptoms of root knot nematode infection ranged from stunting, leaf chlorosis and patchy growth with the formation of root galls which are small or large, terminal or sub-terminal.

 The life cycle of root-knot nematodes includes pre-para-sitic and parasitic stages. The pre-parasitic stage of Meloid-ogyne species begins with the eggs laid in the gelatinous matrix by female and partially embedded in the root of a host plant (Table 1). The development of the eggs begins with in few hours of deposition and formed into coiled first stage juvenile within egg membrane. The J2 is hatched from the egg under favorable conditions and released from egg mass(Fig. 2). Once released J2s move through the soil and finally directed towards the nearest root tip of host plants. The infective second-stage juveniles enter the root at the elonga-tion zone by the dissolution of the cell walls with the cellu-lases and pectinases released through the stylet. After root penetration, that migrate intercellularly within the root and stops near xylem differentiation region and secrete proteins that induce the vascular parenchyma cells to become the giant cells. Secretory proteins synthesized by the nematode in one dorsal and two subventral esophageal gland cells can either be secreted into the plant extracellular space (the apo-plast) or into the host plant cell (the symplast) (Huang et al., 2006). Proteins secreted into the cytoplasm cause changes in gene expression and cell development of the host plant cell. The cells then undergo successive nuclear divisions result-ing in the formation of large multinucleate cells called 'giant cells' (Bird & Koltai, 2000). After two sequential molts (J3 and J4), juvenile embedded within the gall enters the adult stage (Fig. 2). In the later stage the body size of the adult female increases and body become flask shaped. The stylet and esophageal bulb disappears as the female complete sec-ond and third molts. After the fourth molt, the stylet and median bulb are regenerated, uterus and vagina are formed. The reproduction in Meloidogyne species is usually parthe-nogenesis (mitotic). The cells present at the distal end of the reproductive system divide many times to form oogonia. Further, the advanced oogonia stops dividing and become oocytes passing through a long growth zone. One more mitotic division occurs and the eggs become oval and form flexible shell which finally passes through the vagina and are deposited in the egg mass. Female produces eggs laid in a gelat-inous matrix outside the root.

Table 1. Comparison of properties of Meloidogyne incognita and Caenorhabditis elegans.

Fig. 2. A. Schematic representation of the infection and prorogation of Meloidogyne incognita on the roots of tomato plants. Formation of root knots (white arrow heads) (a). Egg mass (white arrow) formed on roots knots after 5 weeks of infection (b) and single egg mass(white arrow) stained in acid fuchsin (c). Root knot (white arrow head) showing female (star) producing egg mass (white arrow) in the root (d). Egg produced in gonads (e). B. Exophytic and endophytic stages of Meloidogyne incognita. Egg; J1 (first-stage juvenile) nematode residing inside the egg case; Motile J2 (second-stage or infective juvenile); After infection J2 subsequently molts within the root to become J3 and J4; In J4 stage, the juvenile progress to either globose adult female or vermiform adult males. The males emerge as adults from the J4 cuticle and migrate out of roots without mating. Female develops in to flask shaped structure and lays eggs in the gelatinous matrix outside the root and a single female nematode produces 500-1000 eggs. matrix outside the root. Scale bar = 60 μm.

 Conventional strategies like crop rotation, chemical fumi-gant application, transfer of naturally occurring plant resis-tant genes, breeding resistant plant cultivars, and transfer of Bt toxin and protease inhibitors to the plants have been adopted for the control of parasitic nematodes. Since the dis-covery of RNAi mechanism in C. elegans, it has been used as an important research tool to elucidate the nematode gene function by transcript knock-down leading to the aberrant phenotype. Soaking second-stage juveniles (J2) with dou-ble-stranded RNA (dsRNA) coding for amphidial secretory protein (AMS-1) greatly reduced the transcript levels there by disrupting the ability of nematodes to locate the host plants (Chen et al., 2005). RNAi could be more effective in controlling RKN through transgenic plant-derived dsRNA and by targeting multiple parasitism genes. Here we aim to review the current progress in the development of plant-derived RNAi for resistance against plant parasitic root-knot nematode and the future application of the technology.


 Over the last decade, the entry in GenBank for Meloidog-yne  reveals 79,978 ESTs analysed from various develop-mental stages of Meloidogyne  species. Recently, complete genome sequencing of two Meloidogyne spp. (M. hapla and M. incognita) have simultaneously been carried out and reported by two groups (Abad et al., 2008; Opperman et al., 2008). Sequencing of M. incognita and M. hapla genomes revealed that the diversity of plant cell wall-degrading endo-glucanases, expansins and pectate lyases were observed to be derived from multiple horizontal gene transfers from bac-terial sources and are believed to be involved in the adapta-tions to parasitism. In both the cases horizontal gene transfer played a role in the origin and evolution of parasitism. The M. hapla genome encodes ~ 5,500 less protein-coding genes and significantly smaller gene families compared to C. ele-gans. Data mining with ESTs has revealed the identification of transcripts encoding parasitism related proteins in plant parasitic nematodes, e.g. pectate lyase (Popeijus  et al., 2000),  β-1,4-endoglucanase,  β-1,4-endoxylanase (Dautova et al., 2001), polygalacturonase (Jaubert et al., 2002), glu-tathione peroxidase (Jones  et al., 2004), and chorismate mutase (Jones et al., 2003) etc. Analysis of ESTs, not only gives insight in identifying the family specific genes (Kang et al., 2010), but also revealed novel horizontally transferred gene candidates in Meloidogyne species.

 The haploid genome size of  M. incognita is 86 MB, almost twice the previously estimated genome size (47- to 51-Mb) with 79,978 ESTs. Out of these 1,625 EST clusters were formed and classified by function using Gene ontology(GO) hierarchy and the Kyoto KEGG database (McCarter et al., 2003). Assuming 19,200 total genes in M.  incognita(Abad  et al., 2008), these clusters are likely to represent approximately 17% of all genes in  M.  incognita.  High throughput in situ hybridization with cDNA clones encod-ing signal peptides resulted in probes of 37 unique clones specifically hybridizing to transcripts (parasitism genes) accumulating within the subventral (13 clones) or dorsal (24 clones) esophageal gland cells of M. incognita and several proteins of unknown function (Huang et al., 2003).


 RNA interference is a cellular mechanism in which dou-ble-stranded RNA induces gene silencing in a sequence spe-cific manner by targeting complementary mRNA for degradation (Bosher  et al., 2003). Silencing of gene via RNAi was first demonstrated in  C. elegans where the dsRNA molecules efficiently trigger the sequence specific silencing of endogeneous target genes. First step in the dsRNA triggered RNAi silencing in C. elegans involves the uptake of dsRNA and is carried out by SID-2 (systemic RNA interference deficient-2), a single pass transmembrane protein which in turn activated by SID-1 (systemic RNA interference deficient-1), a protein with large amino terminal extracellular domain and 11 transmembrane domains. Fur-ther, the dsRNA binding protein RDE-4 (RNAi deficient-4) and the RNaseIII related enzyme Dicer bind to the dsRNA, triggers and cleaves them into 21 to 25 nucleotide primary small interfering RNAs (siRNAs). Argonaute protein RDE-1 (RNAi deficient-1) then associate with the primary siRNA to form the RISC (RNA induced silencing complex). The RISC then uses the guiding strand of the siRNA to find tar-get mRNAs and induces the degradation of the mRNA by Argonaute, the catalytic component of the RISC (Gregory et al., 2005) leading to the gene silencing effect (Fig. 3). An additional RNA amplification step in C. elegans results in efficient RNAi silencing of the target gene (Akoi et al., 2007). In this step, RNA dependent RNA polymerases(RdRP) are guided to the homologous mRNAs by pri-mary siRNAs further cleaving them in to secondary-type siRNAs in a Dicer-independent manner. Abundant sec-ondary-type siRNAs with 5'-triphosphate-termination drives post-transcriptional silencing more effectively than primary siRNAs.

Fig. 3. Mechanism of RNA interference in Meloidogyne incognita similar to exogeneous RNAi pathway of Caenorhabditis elegans. A. Female of Meloidogyne incognita feeding on dsRNA expressed inside the tomato root. B. RNAi trigger within the dsRNA fed Meloidogyne incognita. SID-1 and SID-2: systemic RNA interference deficient (trans membrane binding protein), Dicer: RNase III family of nucleases, DCR-1; dicer protein, siRNA: small interfering RNA, RISC: RNA-induced silencing complex, RDE-1: RNAi deficient-1 (dsRNA binding protein), AGO: Argonaute, RdRP: RNA-dependent RNA polymerase, RRF-1: secondary RNA-dependent RNA polymerase. Triangle indicates RDE-4: RNAi deficient-4 (dsRNA binding protein); hexagon indicates DCR-1; square indicates RDE-1. N: not found in Mi; dcr-1, rde-1, rrf-1 found in Mi. Figures modified and adapted from Hutvagner & Simard (2008) and Rosso et al. (2009).

 Argonaute proteins play an important role in the final step of RNAi pathway through the multicomponent protein com-plex RISC. Argonaute proteins, an integral to RISC with characteristic nuclease activity binds to the siRNAs and cleaves target mRNA (Hammond, 2005). The first Argo-naute protein associated with the RNAi response is RDE-1 and is mainly required during exogeneous RNAi pathway in C. elegans (Yigit et al., 2006). Functional study of this pro-tein led to the discovery of ALG-1 (Argonaute-1) and ALG-2 (Argonaute-2) which are essential for the endogeneous miRNA silencing pathway in C. elegans. Interestingly, sec-ondary Argonaute proteins and other Argonaute protein family members are found in both exogeneous and endoge-neous RNAi pathways bind secondary siRNAs probably produced by the RDE-1 protein in complex with primary siRNA. They further recognizes the target mRNA and induces the synthesis of an antisense strand by RNA-depen-dent RNA polymerases (RdRP). The specific binding of an Argonaute protein to either primary or secondary siRNAs depends on the number of phosphates found at the 5' ends of either primary or secondary siRNAs in  C. elegans. In microRNAi (miRNA) pathway Argonaute associated with miRNA binds to the 3'-untranslated region of the target mRNA and induces deadenylation of the polyadenylated 3'-end causing mRNA degradation and or heterchromatin for-mation leading to post-translational silencing.

 The basic mechanism of RNAi appears to be conserved in all eukaryotic organisms. However, they vary in their nature to take up the foreign dsRNA molecules and use it in the RNAi pathway. The effects of RNAi can be both systemic and heritable in dicot plant Arabidopsis thaliana and C. ele-gans, although not in fruit fly Drosophila melanogaster or mammals. Recently, many reports suggest the efficiency of RNAi in plant parasitic nematodes. Several genes targeted for RNAi in different root-knot nematode species, their phe-notypic characterization and the molecular effects have been listed in Table 2.

Table 2. Target genes of Meloidogyne incognita silenced by plant mediated method and soaking method.


 The dsRNAs ingested by the pathogenic nematodes are taken up by the intestinal cells or through the cuticle during soaking. Initially, the delivery of dsRNA in to the plant para-sitic nematodes was a challenge as the infective stages of plant parasitic nematodes are small in size which makes microinjection with dsRNA a difficult task. Further, the sed-entary nematodes feed only after establishing the feeding site inside the root and do not ingest substances prior to this stage. However, Urwin et al. (2002) for the first time dem-onstrated the use of octopamine to induce the uptake of dsR-NAs by preparasitic second-stage juveniles of two cyst nematodes, H. glycines and G. pallida. Following this new technique, several groups had successfully silenced the nematode genes which resulted in reduced number of estab-lished nematodes or suppressed the development of nema-todes by soaking plant-parasitic nematodes in dsRNAs(Bakhetia et al., 2005, 2007; Chen et al., 2005; Huang et al., 2006; Matsunaga et al., 2012; Shingles et al., 2007). Other research groups have also efficiently suppressed the devel-opment of nematodes with or without the addition of differ-ent chemicals to induce uptake (Dubreuil  et al., 2007; Fanelli et al., 2005; Park et al., 2008). Resorcinol and sero-tonin are also found to induce dsRNA uptake by second stage juvenile of M. incognita  and may be more effective than octopamine (Rosso et al., 2005). Plant parasitic nema-todes are found to ingest varying sizes of dsRNA molecules ranging in size from 42 to 1300 bp and have proved to be effective in inducing RNAi (Chen et al., 2005; Huang et al., 2006). Recently, transgenic tomato roots expressing 54-kDa Bacillus thuringiensis (Bt) crystal (Cry) proteins Cry6A challenged with M. incognita, and RKNs could successfully ingested this large molecule (Li et al., 2007), substantially increasing the known size of molecular uptake into RKNs.


 Recent progress in the application of RNAi technology demonstrates the silencing of plant parasitic nematodes genes by  in planta expression of dsRNA homologous to nematode and there by displaying resistance to infection(Gheysen & Vanholme et al., 2007). Previous research find-ings has confirmed the feasibility of silencing nematode genes using dsRNA produced in the host plant (Huang et al., 2006; Yadav et al., 2006). Yadav et al. (2006) reported that silencing the genes encoding an integrase and a splicing fac-tor lead to the decrease in root knot and nematode number thereby conferring resistance to tobacco plants against the plant-parasitic nematode M. incognita. Targeting the esoph-ageal gland expressed parasitism gene 16D10 in transgenic Arabidopsis thaliana resulted in 63-90% reduction in root knot formation and egg production in M. incognita, M. jav-anica,  M. hapla and  M. arenaria (Huang  et al., 2006). Recently, high level resistance was achieved in soybean roots by targeting four M. incognita genes (Ibrahim et al., 2011). The result showed up to 92.0-94.7% reduction in root knot when tyrosine phosphate and mitochondrial stress-70protein precursor genes were targeted in soybean plants. The size of the M. incognita was 5.4 and 6.5 times less in the transformed roots compared to the non transformed roots. Silencing of two Mi candidate genes such as a dual oxidase gene (Miduox), involved in tyrosine cross-linking of the developing cuticle and a subunit of signal peptidase(Mispc3), required for the processing of secreted proteins resulted in conferring partial resistance in Arabidopsis roots. Only 50% reduction in the number of nematodes and retar-dation in the female development was observed. Enhanced reduction in the nematode numbers and nematode develop-ment ability was resulted by combined expression of the two dsRNAs by crossing the Arabidopsis lines (Charlton, 2010). Expression of putative transcription factor, MjTis11 contain-ing hairpin structures in Nicotiana tabacum plants did not result in significant decrease in nematode number or egg hatching rate indicating that MjTis11gene is not an ideal can-didate or the levels of down-regulation were not sufficient for efficient silencing (Fairbairn et al., 2007).


 The feasibility of using RNAi in the crop protection against root knot nematodes has been demonstrated well. RNAi technology is believed to be one of the most promis-ing techniques for the study of functional genetics in nema-todes in efficiently down regulating the expression of a candidate gene. Recently, several papers have described the transgenic plants producing dsRNAs directed against nema-tode genes. The key factors for this approach are the identifi-cation of suitable nematode target which when silenced should induce lethal effect in nematodes. The other factor is the high amount expression of dsRNA through in planta and delivering sufficient amounts of dsRNA for uptake by the nematodes. The regulation of gene expression by RNAi appears to be common among the eukaryotic organisms. The complete silencing of the target gene occurs by the large amplification of the siRNA molecules which bind to a spe-cific gene transcript. siRNAs production is usually less as the plants lack nematode gene targets (Gheysen & Van-holme, 2007). Therefore, it is necessary to increase the level of the nematode target gene in plants to obtain elevated amounts of siRNAs. Although, RNAi is found to be highly gene specific, cross hybridization with the gene sequences with partial homology with that of dsRNA sequence might induce the silencing of the non-target genes, resulting in an unexpected mutant phenotype (Ma et al., 2006). Improve-ments have been made in order to avoid off-target effects such as; (i) development of software to identify the off-tar-get sequences and excluding them from the RNAi constructs(Koberle et al., 2006), (ii) avoiding the expression of a target gene that is highly conserved across plants and animals, (iii) designing sequences from the 5' or 3' untranslated regions(UTRs), as these are less conserved than those sequences coding for open reading frames.

 An important advantage of RNAi mediated resistance is the lack of ability by dsRNA to produce a functional protein indicating minimal non-target effect in engineered plants. RNAi-based nematode control can be adopted as a biosafe approach with the careful selection of target genes to prevent off target interactions and with proper ecological risk assess-ment of this technology, a favorable outcome is expected. The biosafety level could be upgraded by the use of promot-ers that can efficiently deliver the nematode targeted dsRNA into the active feeding cells. Improvement in inducing sys-temic RNAi response in the nematodes enables to target multiple genes. Finally, combination of existing RNAi tech-nology together with novel technologies could provide most effective, durable and biosafe control of nematodes than the chemical or other biotechnological approaches of the RKN in future. Further, progress on developing plant resistance against plant parasitic nematodes have restricted to either engineering model plants (eg. Arabidopsis thaliana) or con-ducting the experiments to assess the resistance at the green house level. Therefore, it is important to focus the future research for the evaluation of the nematode resistance in economically important crop plants and at the field level.


 This project (PJ008541) was supported by grant (to B.S.H.) from the National Academy of Agricultural Sci-ence, RDA, Korea. This study was supported by 2012 Post-doctoral Fellowship Program (to H.A.) of National Academy of Agricultural Science, RDA, Korea.


1.Abad, P., J. Gouzy, J. M. Aury, and P. Castagnone-Sereno. 2008. Genome sequence of the metazoan plant-parasitic nema-tode Meloidogyne incognita. Nat. Biotechnol. 26: 909-915.
2.Aoki, K., H. Moriguchi, T. Yoshioka, K. Okawa, and H. Tabara. 2007. In vitro analyses of the production and activity of secondary small interfering RNAs in C. elegans. EMBO J. 26: 5007-5019.
3.Bakhetia, M., W. Charlton, H. J. Atkinson, and M. J. McPher-son. 2005. RNA interference of dual oxidase in the plant nema-tode Meloidogyne incognita. Mol. Plant-Mic. Inter. 18: 1099-1106.
4.Bakhetia, M., P. E. Urwin, and H. J. Atkinson 2007. qPCR anal-ysis and RNAi define pharyngeal gland cell-expressed genes of Heterodera glycines required for initial interactions with the host. Mol. Plant-Mic. Inter. 20: 306-312.
5.Bird, D. M., and H. Koltai. 2000. Plant Parasitic Nematodes: Habitats, Hormones, and Horizontally-Acquired Genes. J. Plant Grow. Regul. 19: 183-194.
6.Bosher, J. M., B. S. Hahn, R. Legouis, S. Sookhareea, R. M. Weimer, A. Gansmuller, A. D. Chisholm, A. M. Rose, J. L. Bessereau, and M. Labouesse. 2003. The Caenorhabditis ele-gans vab-10 spectraplakin isoforms protect the epidermis against internal and external forces. J Cell Biol. 161(4): 757-768.
7.Charlton, W. C., H. Y. M. Harel, M. Bakhetia, J. K. Hibbard, H. J. Atkinson, and M. J. McPherson. 2010. Additive effects of plant expressed double-stranded RNAs on root-knot nema-tode development. Inter. J. Parasitol. 40: 855-864.
8.Chen, Q., S. Rehman, G. Smant, and J. T. Jones. 2005. Func-tional analysis of pathogenicity proteins of the potato cyst nem-atode Globodera rostochiensis using RNAi. Mol. Plant-Mic. Inter. 18: 621-625.
9.Dalzell, J. J., N. D. Warnock, M. A. Stevenson, A. Mousley, C. C. Fleming, and A. G. Maule. 2010. Short interfering RNA-mediated knockdown of drosha and pasha in undifferentiated Meloidogyne incognita eggs leads to irregular growth and embryonic lethality. Int. J. Parasitol. 40: 1303-1310.
10.Dautova, M., M. N. Rosso, P. Abad, F. J. Gommers, J. Bakker, and G. Smant. 2001. Single pass cDNA sequencing-a power-ful tool to analyse gene expression in preparasitic juveniles of the southern root-knot nematode Meloidogyne incognita. Nem-atology 3: 129-139.
11.Dubreuil, G., M. Magliano, E. Deleury, P. Abad, and M. N. Rosso. 2007. Transcriptome analysis of root-knot nematode functions induced in the early stages of parasitism. New Phytol. 176: 426-436.
12.Fairbairn, D., A. Cavallaro, M. Bernard, J. Mahalinga-Iyer, M. Graham, and J. Botella. 2007. Host-delivered RNAi: an effective strategy to silence genes in plant parasitic nematodes. Planta 226: 1525-1533.
13.Fanelli, E., M. Di Vito, J. T. Jones, and C. De Giorgi. 2005. Analysis of chitin synthase function in a plant parasitic nema-tode, Meloidogyne artiellia, using RNAi. Gene 349: 87-95.
14.Gheysen, G., and B. Vanholme. 2007. RNAi from plants to nem-atodes. Tren. Biotechnol. 25: 89-92.
15.Golden, A. M., J. H. O'Bannon, G. S. Santo, and A. M. Finley. 1980. Description and SEM observations of Meloidogyne chit-woodi n. sp. (Meloidogynidae), a root-knot nematode on potato in the Pacific Northwest. J. Nematol. 12: 319-327.
16.Gregory, R. I., T. P. Chendrimada, N. Cooch, and R. Shiekhat-tar. 2005. Human RISC Couples MicroRNA biogenesis and posttranscriptional gene silencing. Cell. 123: 631-640.
17.Hammond, S. M. 2005. Dicing and slicing: The core machinery of the RNA interference pathway. FEBS Lett. 579: 5822-5829.
18.Huang, G., B. Gao, T. Maier, R. Allen, E. L. Davis, T. J. Baum, and R. S. Hussey. 2003. A profile of putative parasitism genes expressed in the esophageal gland cells of the root-knot nema-tode Meloidogyne incognita. Mol. Plant-Mic. Inter. 16: 376-381.
19.Huang, G. Z., R. Allen, E. L. Davis, T. J. Baum, and R. S. Hus-sey. 2006. Engineering broad root-knot resistance in transgenic plants by RNAi silencing of a conserved and essential root-knot nematode parasitism gene. Proc. Nat. Aca. Sci. USA. 103:14302-14306.
20.Huang, G., R. Dong, R. Allen, E. L. Davis, T. J. Baum, and R. S. Hussey. 2006. A root-knot nematode secretory peptide func-tions as a ligand for a plant transcription factor. Mol. Plant-Mic. Inter.19: 463-470.
21.Hutvagner, G., and M. J. Simard. 2008. Argonaute proteins: key players in RNA silencing. Nat. Rev. Mol. Cell Biol. 9: 22-32.
22.Ibrahim, H. M. M., N. W. Alkharouf, S. L. F. Meyer, M. A. M. Aly, A. Y. Gamal El-Din, E. H. A. Hussein, and B. F. Mat-thews. 2011. Post-transcriptional gene silencing of root-knot nematode in transformed soybean roots. Expt. Parasitol. 127:90-99.
23.Jaubert, S., J. B. Laffaire, P. Abad, and M. N. Rosso. 2002. A polygalacturonase of animal origin isolated from the root-knot nematode Meloidogyne incognita. FEBS Lett. 522: 109-112.
24.Jones, J. T., C. Furlanetto, E. Bakker, B. Banks, V. Blok, Q. Chen, M. Phillips, and A. Prior. 2003. Characterization of a chorismate mutase from the potato cyst nematode Globodera pallida. Mol. Plant Pathol. 4: 43-50.
25.Jones, J. T., B. Reavy, G. Smant, and A. E. Prior. 2004. Glu-tathione peroxidases of the potato cyst nematode Globodera rostochiensis. Gene 324: 47-54.
26.Kang, M. J., Y. H. Kim, and B. S. Hahn. 2010. Expressed sequence tag analysis generated from a normalized full-length cDNA library of the root-knot nematode (Meloidogyne incog-nita). Genes Genom. 32: 553-562.
27.Koberle, C., S. H. Kaufmann, and V. Patzel. 2006. Selecting effective siRNAs based on guide RNA structure. Nat. Prot. 1:1832-1839.
28.Li, X. Q., J. Z. Wei, A. Tan, and R. V. Aroian. 2007. Resistance to root-knot nematode in tomato roots expressing a nematicidal Bacillus thuringiensis crystal protein. Plant Biotech. J. 5: 455-464.
29.Ma, Y., A. Creanga, L. Lum, and P. A. Beachy. 2006. Preva-lence of off-target effects in Drosophila RNA interference screens. Nature 443: 359-363.
30.Matsunaga, Y., K. Kawano, T. Iwasaki, and T. Kawano. 2012. RNA interference-mediated growth control of the southern root-knot nematode Meloidogyne incognita. Biosci. Biotec. Biochem. 76: 378-380.
31.McCarter, J. P., M. D. Mitreva, J. Martin, M. Dante, T. Wylie, U. Rao, D. Pape, Y. Bowers, B. Theising, C. V. Murphy, A. P. Kloek, B. J. Chiapelli, S. W. Clifton, D. M. Bird, and R. H. Waterston. 2003. Analysis and functional classification of transcripts from the nematode Meloidogyne incognita. Gen. Biol. 4: R26.
32.McCarter, J. P. 2009. Molecular approaches toward resistance to plant-parasitic nematodes. Cell biology of plant nematode par-asitism. In R. H. Berg, and C. G. Taylor. (ed.) Series: Plant Cell Monographs. Springer-Verlag Press, Berlin, DE.
33.McDonald, A. H., and J. M. Nicol. 2005. Nematode parasites of cereals. p 131-191. In M. Luc et al. (ed.) Plant-parasitic nema-todes in subtropical and tropical agriculture. CAB Interna-tional, Wallingford, US.
34.Opperman, C. H., D. M. Bird, V. M. Williamson, D. S. Rokh-sar, M. Burke, J. Cohn, J. Cromer, S. Diener, J. Gajan, S. Graham, T. D. Houfek, Q. Liu, T. Mitros, J. Schaff, R. Schaffer, E. Scholl, B. R. Sosinski, V. P. Thomas, and E. Windham. 2008. Sequence and genetic map of Meloidogyne hapla: A compact nematode genome for plant parasitism. Proc. Nat. Aca. Sci. USA. 105: 14802-14807.
35.Park, J. E., K. Y. Lee, S. J. Lee, W. S. Oh, P. Y. Jeong, T. Woo, C. B. Kim, Y. K. Paik, and H. S. Koo. 2008. The efficiency of RNA interference in Bursaphelenchus xylophilus. Mol. Cells 26: 81-86.
36.Popeijus, H., H. Overmars, J. Jones, V. Blok, A. Goverse, J. Helder, A. Schots, J. Bakker, and G. Smant. 2000. Degrada-tion of plant cell walls by a nematode. Nature 406: 36-37.
37.Rosso, M. N., M. P. Dubrana, N. Cimbolini, S. Jaubert, and P. Abad. 2005. Application of RNA interference to root-knot nematode genes encoding esophageal gland proteins. Mol. Plant-Mic. Inter. 18: 615-620.
38.Rosso, M. N., J. T. Jones, and P. Abad. 2009. RNAi and func-tional genomics in plant parasitic nematodes. Ann. Rev. Phyto-pathol. 47: 207-232.
39.Risher, J. F., F. Mink, and J. F. Stara. 1987. The toxicologic effects of the carbamate insecticide aldicarb in mammals: a review. Environ. Heal. Perspec. 72: 267-281.
40.Sasser, J. N., and D. W. Freckman. 1987. A world perspective on nematology: the role of the society. p 7-14. In J. A. Veech and D. W. Dickson. (ed.) Vistas on nematology, Society of Nema-tology, Hyattsville, Maryland, US.
41.Shingles, J., C. J. Lilley, H. J. Atkinson, and P. E. Urwin. 2007. Meloidogyne incognita: Molecular and biochemical characteri-sation of a cathepsin L cysteine proteinase and the effect on parasitism following RNAi. Exper. Parasitol. 115: 114-120.
42.Steeves, R. M., T. C. Todd, J. S. Essig, and H. N. Trick. 2006. Transgenic soybeans expressing siRNAs specific to a major sperm protein gene suppress Heterodera glycines reproduction. Funct. Plant Biol. 33: 991-999.
43.Urwin, P. E., C. J. Lilley, and H. J. Atkinson. 2002. Ingestion of double-stranded RNA by preparasitic juvenile cyst nematodes leads to RNA interference. Mol. Plant-Mic. Inter. 15: 747-752.
44.Yadav, B. C., K. Veluthambi, and K. Subramaniam. 2006. Host-generated double stranded RNA induces RNAi in plant-parasitic nematodes and protects the host from infection. Mol. Biochem. Parasitol. 148: 219-222.
45.Yigit, E., P. Batista, Y. Bei, K. M. Pang, C. C. G. Chen, N. H. Tolia, L. Joshua-Tor, S. Mitani, M. J. Simard, and C. C. Mello. 2006. Analysis of the C. elegans Argonaute family reveals that distinct Argonautes act sequentially during RNAi. Cell 127: 747-757.