Penetration of roots and formation of syncytium

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Chapter 1: Research Proposal

The secreted SPRY Domain-Containing proteins (SPRYSECs) are a large family of proteins secreted from the dorsal gland cell of cyst nematodes (Qin, et al., 2000). There are approximately 400 different SPRYSEC genes in the genome of Globodera pallida (unpublished data, J. Jones, SCRI). Different members of the SPRYSEC family of proteins from G. pallida show different subcellular localization patterns in plants, with some localized to the cytoplasm and others to the nucleus and nucleolus. Differences in subcellular localization may reflect diverse functional roles for each individual protein or, more likely, variety in the compartmentalization of plant proteins targeted by the nematode (Jones, et al., 2009). Two members of this gene family may be involved in interactions a plant resistance gene by now (Rehman et al., 2009; Sacco, et al., 2009). The currently going-on G. pallida genome sequencing project will reveal the full complement of potential effectors present in this species. Our aim here is to clone highly expressed genes in SPRYSEC family and see if they can suppress PAMP-triggered immunity (PTI) or effector-triggered immunity (ETI) with different functional assays.

Chapter 2: Literature Review


Nematodes are pseudocoelomate, unsegmented worm-like animals, commonly described as filiform or thread-like, a characteristic reflected by the taxon name nema (Greek, nema=thread) and its nominative plural nemata (Decraemer W & Hunt D J, 2006). The alimentary tract that extends from the head to tail and the reproductive organs are the two most prominent structures in the anatomy of nematodes. They are the most abundant multicelluar animals on earth, which are found in practically all different environments that sustain life.

Nematodes are generally grouped by different feeding types on the basis of food source and/or feeding habit, i.e. microbial feeders, predacious species, animal parasites and plant parasites. For instance, the world most famous nematode, Caenorhabditis elegans, is a microbial feeder. C. elegans has proven to be an excellent model for studies on animal genetics and development. It was the first animal whose genome was completely sequenced (Consortium, 1998). Enoploides Longispiculosus is an example of a predacious nematode species that lives in marine sediments (Moens T, 2000) while Onchocerca volvulus and Steinernema weiseri are species that infect other animals (Lindblade, et al., 2007 ; Unlu et al., 2007).

Plant parasitic nematodes consume nutrient from cytoplasm of plant cells and cause substantial losses in variable crops throughout the world. Annual estimated worldwide yield losses to major crops by nematodes action is about 12.3% and it is about 14% in developing countries (Sasser & Freckman, 1987). The cost of world agriculture loss by nematode was estimated recently to be US$ 125 billion annually (Chitwood, 2003). To reduce these losses to control nematode globally an estimated amount of US$ 500 million is spent every year (Keren-Zur et al., 2000). It's widely acknowledged that root knot (Meloidogyne spp) and cyst forming (Heterodera and Globodera spp) nematodes are considered economically most devastating plant parasitic nematodes. Root knot nematodes have a wide host range, while the host range of individual cyst nematodes species is restricted to only one or a few related families of plant species (Evans & Stones, 1977). There are kinds of measures used to control nematode populations in the field and in the green house, such as resistant cultivars, crop rotation, biological control and the application of chemical pesticides. Due to the harmful side effects of chemicals and low efficiency of biocontrol, the focus of nematode management shifts currently to nematode resistant host-plant. Insight in the nematode genes that are essential in the plant-nematode interaction is a prerequisite to make productive use of both natural and bio-engineered resistance in host plants.

Plant parasitism

Nematode adaptations to parasitism

Parasitism is the most harmful form of symbiosis, which is a general term describing situation where two dissimilar organisms live together in close physical association (OED2 1998).

The nematode body plan has proved to be a remarkably adaptable platform (Bird & Bird, 1991), upon which a wide range of modifications have evolved including morphological and developmental specializations (Bird & Alan, 2001). Morphological adaptations for plant parasitism are surprisingly similar among all plant-parasitic nematodes. Most notably all plant-parasitic nematodes are equipped with a stylet (hollow mouth spear) to pierce cell walls and allow solute exchange between plant and parasite (Baum, et al., 2007). The extensible stylet of plant parasitic nematodes is connected to a well-developed pharynx containing three or five gland cells. The most evolutionary advanced adaptations for plant parasitism by nematodes are the products of parasitism genes expressed in their esophageal gland cells and secreted through their stylet into host tissue to control the complex process of parasitism (Hussey, et al., 2002). Undoubtedly one of the key adaptations that have permitted nematodes to become such successful parasites is the ability to suspend development so as to couple their biology temporally with that of the host or other environmental cues. The canonical example of developmental arrest is provided by the dauer larva of C. elegans (Riddle & Albert, 1997) where it serves as an environmentally resistant, dispersal stage.

Distinct feeding strategies

The plant parasitic nematodes have specialized to be three groups with different kinds of feeding strategies. The first is called the ectoparasites (e. g. Trichodorus and Xiphinema) that mainly feed on epidermal cells and root hairs or on the outer cortex cells underneath the epidermal cell layer using their stylet. The migratory endoparasites form the second group of nematodes that have either specialized into herbaceous (e.g. Aphelenchoides) or woody plants (e.g. Bursaphelenchus). They are able to penetrate plant tissue, migrate intracellularly through several cell layers and use the ectoplasm of cells they come across as food source. Finally, there is the sedentary endoparasites (e.g. Meloidogyne and Nacobbus) which can establish a feeding site in the plant and feed for weeks. The potato cyst nematodes (Globodera rostochiensis and G. pallida) belong to the last group.

the Potato cyst nematode: Globodera pallida

host range, history and damage

Globodera pallida - potato cyst nematode (family Heteroderidae, order Tylenchida) is a sedentary endoparasite. Potato cyst nematodes have limited host range including Solanaceous plants, such as potato, tomato, and eggplant (Evans & Stones, 1977). Potato cyst nematode was introduced in Europe probably in the 19th century together with potato breeding material from Andes in South America. Since then it has become a major pest of potato, especially in countries with temperate climate. These nematodes cause damages valued at over?50 million to potato crops in the UK alone. A survey has shown that over 60% of potato growing land in the UK is infested with PCN and, of this land over 90% is infested with G. pallida (Jones J.T. & Perry R.N., 2004).

Life cycle

Potato cyst nematodes have a relatively narrow host range including Solanceae only. The life cycle of the cyst nematodes consists of four juvenile stages followed by adult stage (Lee, 2002).The first stage juvenile is contained inside the egg shell and this is where the first moulting will take place. Hatching process is generally stimulated by root diffuses of host plants, which may also help the infective juvenile in localizing the host plant (Sharma & Sharma, 1998). The pre-parasitic J2 migrates towards and subsequently invades the plant root in the cell elongation zone (von Mende et al., 1998). They move through the cortex towards the stele intracellularly where they select a single cell that becomes a starting point of a feeding site called syncytium formed by a group of cells in which cytoplasmic continuity is maintained. Syncytium is the only source of nutrients throughout the nematode's life. Its multiple nuclei are enlarged and amoeboid, resulting from endoreduplication, in which DNA synthesis occurs in the absence of nuclear or cellular division (Gheysen et al., 1997).Sooner after feeding cell induction, the nematode loses its ability to move and becomes sedentary, while feeding from the plant. During this feeding process the nematode undergoes three additional moults into J3, J4 and finally into the adult stage. Adult males regain their mobility and leave the root to mate with the female that remains sedentary. Females retain several hundreds of fertilized eggs inside their body wall, which eventually hardens to form a protective cyst. Inside the cyst eggs can remain viable for as long as 15 years (Dropkin, 1989).

Penetration of roots and formation of syncytium

Physical damage due to penetration of and migration through the root as well as induction and maintenance of syncytium are the two main events during the life cycle which contribute directly to the pathological effect on the infested crops. Freshly hatched J2 probes the selected area of the epidermal cells of the root, usually behind the growing tip, using stylet whose effect can be enhanced in the mean time by producing and releasing cell wall degrading enzymes (Smant et al.1998). After approximately one hour the slit created in the cell wall is sufficiently large for the juvenile to wedge it head through it. Once inside the root the juvenile migrates intracellularly. The nematode's way through the cortical cells toward the vascular cylinder is marked by necrosis of the cells aligning the track. Upon arrival in the inner cortex the nematode starts to search for an appropriate initial feeding cell while probing the neighbouring cells. However, the exact criteria involved in this selection are still largely unknown.

The nematode becomes immobile after the initial cell of syncytium has been selected. It inserts the stylet into the cell cytoplasm for several hours which is called the preparation period. The fully differentiated cortex cell changes into a metabolically highly active cell with dense cytoplasm that contains small secondary vacuoles, numerous organelles and enlarged nucleus with prominent nucleoli (Rice et al., 1985). After this preparation period, the nematode starts to feed while the initial syncytial cell expands towards the vascular bundle through the progressive local cell wall dissolution. Plant enzymes recruited by nematodes also can contribute to the breaking down of cell walls. Cells are fused to the initial feeding cell and become similar in their structure. During the course of a week the syncytium becomes increasingly hypertrophied and acts as metabolic sink from which nematode uptakes large amounts of nutrients.

Plant parasitic nematode sequencing projects

Globodera pallida sequencing project

The sequencing project of G. pallida is currently led by four institutes, i.e., Scottish Crop Research Institute (SCRI), Leeds University, Wellcome Trust Sanger Institute and Rothamsted Research from the UK. This project is to sequence and analyse the nuclear and mitochondrial genomes of G. pallida with the goal of producing a reference quality genome sequence. The sequencing is being done using a combination of capillary sequencing, Roche (454) and Illumina (Solexa) sequencing technology platforms ( ). Globodera pallida sequence data are being made available on the Sanger Institute Blast Sever as they become available ( ).

An unexpected finding in this sequencing project was the multipartite structure of mitochondrial DNA (mtDNA) genome (Armstrong, 2000). According to this paper, six circular DNAs ranging from ~ 6.3 to 9.5 kb have been amplified from a British population of G. pallida by PCR, and additional components of the G. pallida mtDNA remain uncharacterized. All of these potential G. pallida mtDNAs contain sequences similar to known mitochondrial genes, with most containing sequences that show highest sequence similarity to previously described nematode mitochondrial genes. The complete sequence of the largest putative mtDNA reveals that it contains seven full-length open reading frames similar in sequence to nematode mitochondrial genes. It suggests that these small, circular mitochondrial DNAs (scmtDNAs) are present together in population of G. pallida, although their relative frequencies vary considerably between populations.

There are three practical outputs brought by G. pallida sequencing project. First, the genome sequence will provide researches with details of all the genes present in PCN and provide us with a complete list of the potential nematode targets for novel control strategies. Second, a G. pallida genome sequence will provide a complete list of the potential nematode avirulence genes that trigger resistance in plants. Identification of avirulence genes will allow functional sceens for germplasm carrying resistance genes making plant breeding more rapid and will allow monitoring of PCN for resistance- breaking populations that contain changed versions of the avirulence genes. Third, the availability of the sequence of natural enemy (Pasteuria penetrans) and that of G. pallida will allow dissection of the interaction between these organisms. Of particular interest are the mechanisms underlying recognition and binding of Pasteuria to PCN. This information will improve the prospects for biological control of G. pallida.

Meloidogyne spp sequencing project

Root knot nematodes (Meloidogyne spp) also gained much attention in this area due to the highly damaging ability. They have an intimate interaction with their hosts. Within the host root, adult females induce the redifferentiation of root cells into specialized 'giant cells', upon which they feed continuously. Of most practical interest in RKN are highly damaging and polyphagous species, for instance. M. hapla and M. incognita. My intent here is to present some brief introduction to genome discoveries of both species. Assemlies of both genomes are available in the public databases, including GenBank and project Web sites ( and )

The northern root knot nematode, Meloidogyne hapla, is emerging as a model species for research on sedentary endoparasites (Opperman, 2009). Sequence and genetic map of Meloidogyne hapla were presented in 2008 as a compact nematode genome for plant parasitism (Opperman, et al., 2008). At 54 Mbp, M. hapla represents not only the smallest nematode genome yet completed, but also the smallest metazoan (Opperman, et al., 2008). As mentioned in this paper, many isolates of M. hapla reproduce by facultative meiotic parthenogenesis. The authors exploited this unique genetic system for construction of a linkage map. A striking finding of this project is that M. hapla encodes about 5,500 fewer protein-coding genes than does C. elegans. A hypothesis was further substantiated that horizontal gene transfer (HGT) played a role in evolution of parasitism. The acquisition of the M. hapla sequence and gene map represents a major step in the understanding of biological information on both the nature of nematode parasitism of plants and its evolution.

The Southern root-knot nematode Meloidogyne incognita, as an obligatory sedentary parasite that reproduces by mitotic parthenogenesis, is able to infect the roots of almost all cultivated plants and perhaps is the most damaging of all crop pathogens. M. incognita can infect Arabidopsis thaliana, making it a key model system for the understanding of metazoan adaptations to plant parasitism (Marie, et al., 2008). Genome sequence of M. incognita was reported last year (Abad, et al., 2008). According to this paper, the M. incognita was sequenced using whole-genome shotgun strategy and the assembled sequence reads gave a total size of 86 Mbps suggesting that it is a fixed heterozygous organism. Noncoding DNA repeats and transposable elements represent 36% of the M. incognita genome. They identified 19,212 protein-coding genes and 69% of protein sequences were < 95% identical to any other. An unprecedented set of 61 plant cell wall-degrading, carbohydrate-active enzymes (CAZymes), 20 candidate expansins and associated invertases was probably acquired by horizontal gene transfer, as the most similar proteins were bacterial homologs. M. incognita also has four secreted chorismate mutases that most closely resemble bacterial enzymes. Apart from genes restricted to M. incognita, they also identified gene families showing substantial expansion compared to C. elegans. However, M. incognita has fewer genes encoding peroxidases than C. elegans to produce cytotoxic oxygen radicals for protection against environmental stresses. The genome sequence of M. incognita provides insights into the adaptations required by metazoans to successfully parasitize and counter defenses of immunocompetent plants, and suggests new antipratasitic strategies (Abad, et al., 2008).

Plant parasitic nematodes are among the most damaging and difficult-to-control pests of world agriculture. Meeting current and future worldwide demands for food, fiber and bioenergy will necessitate minimizing these losses and will require development of new control paradigms. These genome sequences provide a new first step toward this goal (Opperman, et al., 2008).

Molecular aspects of Plant-nematode Interactions

Nematode secretions

Nematode secretions can originate from different sources such as epidermis (Jones, 2004), the amphids (Jones, 2000), the excretory/secretory system and rectal glands. However, the major secretory organs are pharyngeal glands. Each gland is a large single secretory cell with a wider basal region and a long narrow extension at the apical region, which terminates in an ampullae. The valve of a single dorsal gland is located near the base of the stylet while the two subventral glands empty into the esophagus just posterior to the metacorporal pump chamber (Hussey & Mims, 1991).

The secretions of pharyngeal gland cells through the hollow stylet play important roles in the host-parasite interaction. The products of the subventral gland cells are important in the early stages of parasitism, during invasion and perhaps during feeding site induction, while the products of the dorsal gland cell play a role later in the parasitic process, probably in feeding site development or maintenance (Gheysen, G. & Jones J.T., 2006).

Proteins secreted in the subventral pharyngeal glands

Most of the identified subventral pharyngeal gland proteins can be classified as cell wall degrading proteins that enable the migrating nematode to degrade all major types of polysaccharides in the cell wall of a host plant. The first parasitism genes cloned from plant parasitic nematodes were B-1, 4-endoglucanases (cellulases) developmentally expressed in the two subventral gland cells of Heterodera glycines and Globodera rostochiensis (Smant et al., 1998). Subsequent studies have shown that this is not the only plant cell wall degrading enzyme produced by plant parasitic nematodes. A family of pectate lyases is present in both root-knot and cyst nematodes, and root-knot nemaotdes also contain xylanase and polygalacturonase. It has also been known that plant-parasitic nematodes secrete expansins during migration which could greatly enhance the accessibility of the other polysaccharide chains to the activity of the cellulases and pectate lyases (Gheysen & Jones, 2006). Furthermore, cell wall binding proteins produced in the subventral pharyngeal glands of various plant parasitic nematodes (Ding et al., 1998; Gao, et al., 2003) may also be indirectly involved in the cell wall degradation.

Several extracellular proteins expressed in these subventral pharyngeal glands had strong similarity to secretory venom allergen AG5-like produced by hymenopteran insects (Ding, 2000). But their function in parasitism is still obscure.

Chorismate mutase gene, expressed both in subventral and dorsal glands, was identified in cyst nematodes (Johns et al., 2003) and root-knot nematodes (Lambert et al., 2009). Chorismate mutase is an enzyme in the shikimate pathway, an important metabolic route in plants.

The calcium-binding protein-calreticulin (Jaubert et al., 2002) is also an interesting protein expressed in subventral pharyngeal glands of root knot nematode. Calreticulin has multiple functions from regulation of cell cycle in animals to cell-to-cell trafficking and pressure support in plants.

Chitinase, acid phosphatase and sodium/calcium/potassium exchanger (Gao, et al., 2002; Huang et al., 2003)were novel proteins whose genes may create the biggest challenge for functional analysis and give us a glimpse how complex and unique the nematode plant relationship is. The third SXP/RAL-2 protein was reported specifically expressed in the subventral pharyngeal glands (Tytgat, 2005).Although they are also found in other nematodes which are not plant parasitic, it does not rule out that Mi-SXP-I could be involved in feeding cell induction.

Proteins secreted in the dorsal pharyngeal glands

The dorsal gland proteins are thought to be responsible for induction and maintenance of the syncytium because dorsal pharyngeal gland is activated at the onset of parasitism during feeding site initiation. They may directly or indirectly alter the gene expression in the recipient cells, either by binding to the plant cell receptors in order to elicit the required signal transduction pathways, or by entering the nucleus and direct modification of the gene expression.

The RanBPM like proteins may be responsible for plant microtubule stabilization in the syncytium and therefore for shunting of the M-phase of cell cycle (Qin, 2001). A domain search revealed that H. glycines is producing a protein similar to CLAVATA3 plant signal peptide, which is involved in differentiation of stem cells in shoot meristems (Olsen & Skriver, 2003).No functional evidence for the role of this peptide in host-parasite interaction is currently available. However, previous studies have shown that another cyst nematode, G. rostochiensis, secrets a small (but unidentified) peptide that can induce cell division in protoplasts. These findings taken together suggest that CLAVATA3-like peptides may have a role in feeding site formation in cyst nematodes (Gheysen & Jones, 2006). An ubiquitin extension protein was identified by comparing the gene expression profile between anterior and posterior part of H. glycines (Tytgat, et al., 2004). The author speculates that it plays a regulatory role in feeding cell formation and is possibly involved in protein turnover in highly active cells. S-phase kinase-associated proteins (skp-1) and RING-H2 proteins (Gao, et al., 2003)which are subunits of the complex that transfers UBI tags to target proteins, are expressed in gland cells of H. glycines and may be secreted into plants.

The secreted SPRY Domain-Containing proteins (SPRYSECs) are a large family of proteins secreted from the dorsal gland cell of cyst nematodes (Qin, et al., 2000). It has been shown that some of the proteins are localised to the nucleus of plant cells after they are secreted while others stay in the cytoplasm (Jones, et al., 2009). Another group has shown that one SPRYSEC interacts with a plant resistance gene (Sacco, et al., 2009).

Functional analysis

Sequence data is most useful when one can transform the obtained information into biological relevant knowledge (Grant & Viney, 2001). A considerable number of genes with known function enabling the parasitism are already identified. Undoubtedly, the sequencing projects of plant parasitic nematodes (mentioned in 2.4) and the use of expressed sequence tags (EST) (Popeijus, et al., 2000; Jones, 2009) will largely facilitate understanding parasitism mechanisms. However, a substantial part of candidate parasitism genes are novel genes without homologues in the database (Gao, et al., 2003; Huang, et al., 2003). This would simply mean that their homologs are not yet identified, or they could represent a very unique set of genes that has evolved only in plant parasitic nematodes. As a result of these facts, functional analysis becomes more and more attractive and necessary.

Several molecular techniques could be utilized to elucidate the role of such genes in plant nematode interaction. The RNA-mediated gene silencing (RNAi) is currently under scrutiny and some promising result indicated that indeed this method is applicable to some plant parasitic nematodes (Jones, et al., 2003; Urwin, et al., 2002; Chen, et al., 2005). Alternatively, overexpression of nematode genes in the whole plant (Goverse & Karczmarek, 1999) or transient expression in plant protoplast (Qin, 1999) may be used to asses the changes in host in response to nematode protein accumulation. Localisation of the secreted protein in plants (Wang, 1999) at a cellular or subcellular level might help us to better understand its specific task in the parasitic process with the help of green fluorescent protein (GFP).Regardless of the method, understanding how the parasitism factors interact with each other and with host molecules to lead to the successful parasitism is a major challenge in the future and may open novel perspectives on development of nematode resistant plants.

Plant defense

Instead of being passive to pathogens' attack, plants usually are able to react to parasites by employing a defensive system against pathogen's attack of which the first one to be encountered is the physical barrier provided by the waxy cuticle and the cell wall. Some chemicals produced by plants during the interaction, like phenolic compounds, also belong to this kind of pre-exsiting defences. Besides, plants have an immune system that is similar to animal innate immune system (Ausubel, 2005). Jones and Dangl presented a zigzag model in 2006 illustrating that the output of the plant immune system depends on the interplay between plant defense and pathogen effector molecules. The conserved pathogen-associated molecular patterns (PAMPs), like flagellin or chitin, are perceived by plant pattern recognition receptors (PRRs) and then PAMP-triggered immunity (PTI) will be induced (Jones & Dangl, 2006).PTI is also known as basal defense and occurs usually less than 10 minutes after exposure to a pathogen. Basal defense is characterized by a.o.callose deposition, production or accumulation of ethylene and reactive oxygen and nitrogen species and change in gene expression (Ingle et al., 2006). All those responses mainly aim to physically isolate the pathogen, prevent spread and attack the invader. Oxygen and nitrogen may be directly damaging to the pathogen, but they are also important molecules in plant defence signaling pathways.

The R-gene mediated effector triggered immunity (ETI) is a more specific form of defense which occurs 2-3 hours later in the host-pathogen interaction, upon delivery of pathogen effectors into the host cytoplasm or apoplast. Avirulence effectors are responsible to elicit ETI. ETI shares some similar features with PTI, and may even share some molecular meachanisms (Abramovitch et al., 2006). However, ETI is generally much stronger and featured by the localized programmed cell death known as hypersensitive response (HR). There are two different models which are normally used to describe recognition of pathogen effectors by R-gene products: the gene-for-gene model and the guard-model. In a gene-for -gene interaction, a resistance gene is only effective if a specific effector is produced by the pathogen. In the guard model, presence of the pathogen is sensed by R-gene products through a modified state of guarded host protein. These host proteins may be the virulence target of the effector protein in compatible interaction.

defense in the plant-cyst nematode interaction

The interaction between non-host Arabidopsis thaliana and the soybean cyst nematodes Heterodera glycines is characterized by a prolonged invasion and migration period, and browning of the tissue at the infection site. The juvenile in vascular cylinder confronted with a strong HR. Additionally, callose or callose-like material is deposited in vascular cylinder and around the nematodes' head (Grundler et al., 1997). Furthermore, hydrogen peroxide is produced by damaged cells but also by cells free of contact with nematodes or syncytium.

Host resistance against cyst nematode is often featured by local cell death in cells at the periphery of the initial feeding cell, therefore inhibiting the expansion of the syncytium. The avirulent nematode effectors are recognized early in feeding site formation according to the timing of response (Cabrera Poch, et al., 2006). Several nematode R-genes have been cloned, for instance Gpa2 (Bakker, et al., 2003), Hero (Ganal, et al., 1995), against G. pallida in potato and G.rostochiesis in tomato respectively. Interestingly, those genes belong to two different classes of NB-LRR-genes, either the class of the LZ-NB-LRR proteins, containing a leucine zipper (LZ) domain, a nucleotide binding domain (NB) and a leucine rich repeated region (LRR) or to the class of Tir-NB-LRR proteins containing a TIR domain (Ausubel, 2005).

The H1 gene from Solanum tuberosum ssp. andigena confers high levels of resistance to the potato cyst nematode Globodera rostochiensis and its unidentified elicitor comply with the gene-for-gene model, which explains the recognition specificity of major R genes. Recently, the Rbp-1 protein from Globodera pallida was shown to induce a HR on Nicotiana benthamiana leaves expressing Gpa2, a resistance gene conferring resistance to G. pallida (Sacco, et al, 2007). In 2009, they demonstrated that this secreted protein RBP-1 from Globodera pallida elicits defense responses including HR through the NB-LRR protein Gpa2. Recognition of Gp-RBP-1 by Gpa2 correlated to a single amino acid polymorphism at position 187 in the Gp-RBP-1 SPRY domain (Sacco, et al., 2009). A large gene family with sequence similarity to Rbp-1, the SPRYSEC family, has been shown to be specifically expressed in the dorsal esophageal gland of G. rostochiensis (Qin, et al., 2000). The secreted SPRY Domain-Containing protein (SPRYSEC) from G. rostochiensis can interact with a CC-NB-LRR protein from a susceptible tomato but does not trigger a resistance response. Alternatively, this SPRYSEC may bind to the immune receptor to downregulate its activity (Rehman, et al., 2009).

Suppression & avoidance of host defense

Plant parasitic nematodes have responded to host defences by evolving a series of physical and biochemical adaptations that help them either avoid eliciting a host response or to reduce the toxic effects of any plant defence response. (Gheysen & Jones, 2006)

The nematode cuticle surface is overlaid with a coat largely made up of proteins. This surface coat may mimic host tissues in an attempt to avoid eliciting a defense response (Jones & Robertson, 1997), which may ultimately result in effector triggered susceptibility (ETS). A peroxiredoxin that specifically break down hydrogen peroxide has been shown to be present in the surface coat of Globodera rostochiensis (Jones, et al., 2004).

Cyst nematodes may suppress plant defense by secreting effectors into the host cell apoplast or cytoplasm. Chorismate mutase (CM) is a secreted nematode effector produced in the esophageal gland of both root-knot and cyst nematodes (Lambert, et al., 1999). CM may be responsible to manipulate the plant's shikimate pathway which is involved in defense (Bekal, et al., 2003). It may suppress plant defense by breaking down chorismate-derived compounds, like salicylic acid or phenolic phytoalexins.

Other mechanisms of suppressing host defences exist and are well characterized in other pathosystems. Many gram-negative plant and animal pathogenic bacteria employ a type III secretion system (T3SS) to subvert and colonize their respective host organisms. The T3SS injects effector proteins directly into the cytosol of eukaryotic cells and thus allows the manipulation of host cellular activities to the benefit of the pathogen (Daniela, et al., 2009). A primary function of Pseudomonas syringae type III effectors appears to be the suppression of ETI and PTI. Seven type III effectors from P. syringae pv. tomato DC3000 were capable of suppressing an HR induced by P. fluorescens (PHIR 11) and the majority of the tested 35 DC3000 type III effectors can suppress the HR induced by HopA1 (Guo M, et al., 2009). The author also identified a new type III effector called HopS2 with particularly strong HR suppression activity. Another bacterial type III effector protein AvrPtoB can ubiquitinate the Arabidopsis LysM receptor kinase CERK1 in vitro and targets CERK1 for degradation in vivo (Selena, et al., 2009). The authors' results reveal a new pathway for plant immunity against bacteria and a role for AvrPtoB E3-ligase activity in suppressing PTI.

Several effectors that target plant defenses have now also been identified in pathogenic fungi and oomycetes. A well-studied case is the Cladosporium fulvum AVR2 protein. AVR2 is a cysteine proteinase inhibitor that binds to, and inhibits RCR3, an extracellular papainlike Cys protease (PLCP) that is induced and secreted by tomato as a defense response against pathogens (Kru¨ger, et al., 2002). Such secreted proteases can be involved in various ways in plant defenses: for example, by acting directly on the invading pathogen, or by being part of signaling cascades for the induction of the HR (Van der Hoorn & Jones, 2004). AVR2 is a pathogen effector with a suppressor activity: in the absence of the tomato resistance gene Cf2, AVR2 suppresses the action of a PLCP; but in the presence of Cf2 the AVR2 /RCR3 complex triggersETI-dependentHR(Rooney et al., 2005).A study on the oomycete pathogen Phytophthora sojae has led to the cloning of a glucanase-inhibiting protein (GIP) that was shown to form a complex in vitro and in vivo with a soybean endoglucanase (PR-2 protein) and to inhibit its activity. This interaction also inhibits the release of a glucan elicitor from Ph. sojae cell walls in vitro. Thus, GIP1 is an example of a suppressor of a plant defense response produced by a virulent pathogen of soybean (Rose, et al., 2002; Metraux, et al., 2009).

Charpter3: References

  • Abad P, Gouzy J, Aury JM, Castagnone-Sereno P, Danchin EGJ, et al. (2008). Genome sequence of the metazoan plant-parasitic nematode Meloidogyne incognita. Nat. Biotech. 8, 909-915.
  • Abramovitch, R.B., Anderson, J.C. & Martin, G.B. (2006). Bacterial elicitation and evasion of plant innate immunity. Nat Rev Mol Cell Biol, 7, 601-611.
  • Armstrong, M.R., Blok V.C. & Phillips M.S. A multipartite mitochondrial genome in the potato cyst nematode Globodera pallida. Genetics, 154: 181192, 2000. PMC1460896.
  • Ausubel, F.M. (2005). Are innate immune signaling pathways in plants and animals conserved? Nature Immunology, 6, 973-979.
  • Bakker, E., Butterbach, p., Rouppe Van Der Voort, J., Van Der Vossen, E., Van Vliet, J., Bakker, J., et al. (2003). Genetic and physical mapping of homologues of the virus resistance gene Rx1 and the cyst nematode resistance gene Gpa2 in potato. Theoretical and Applied Genetics, 106, 1524-1531.
  • Baum, T.J., Hussey, R.S. & Davis, E.L. (2007). Root-knot and cyst nematode parasitism genes: the molecular basis of plant parasitism. J.k. Setlow. (eds) Genetic Engineering, 28, 17-43.
  • Bekal, S., Niblack, T.L., & Lambert, K.N. (2003). A chorismate mutase from the soybean cyst nematode Heterodera glycines shows polymorphisms that correlate with virulence. Mol Plant Microbe Interact, 16,439-446.
  • Bird, A. F. & Bird, J. (1991). The structure of nematodes. 2nd edn. Academic Press, San Diego, pp316.
  • Cabrera Poch, H.L., Manzanilla Lopez, R.H. & kanyuka, K. (2006). Functionality of resistance gene Hero, which controls plant root-infecting potato cyst nematodes, in leaves of tomato. Plant, Cell and Evironment, 29, 1372-1378.
  • Chen, Q. Rehman, S., Smant, G. Jones J.T. (2005). Functional analysis of pathogenicity
  • Chitwood, D.J., (2003). Research on plant-parasitic nematode biology conducted by the United States Department of Agriculture-Agricultural Research Services. Pest Manage. Sci. 59, 748-753.
  • Consortium, T. C. e. S. (1998). Genome sequence of the nematode C.elegans: a platform for investigating biology. Science, 282, 2012-2018.
  • Daniela B. Sheng, Y.H. (2009). Type III Protein Secretion in Plant Pathogenic Bacteria. Plant Physiology, 150: 1656-1664.
  • David, M.B. & Alan. F., Bird. (2001). Plant-parasitic nematodes. In: M. W. Kennedy & W. Harnett, Parasitic Nematodes, CAB international 2001, pp139-166.
  • Decraemer, W. & Hunt, D.J. (2006). Structure and Classification. In: Perry, R. N. & Moens, M. (Eds) Plant Nematology. Wallingford, UK, CABI publishing, pp. 3-32.
  • Ding, X., J.Shields, R. Allen & R.S.Hussey. (1998). A secretory cellulose-binding protein cDNA cloned from the root-knot nematode (Meloidogyne incognita). Mol. Plant Microbe Interact. 11, 952-959.
  • Ding, X., J.Shields, R.Allen & R.S.Hussey. (2000). Molecular cloning and characterization of a venom allergen AG5-like cDNA from Meloidogyne incognita. Int J Parasitol 30, 77-81.
  • Dropkin, V.H.(1989). Introduction to plant nematology. New York: John Wiley and Sons Inc.
  • Evans, K. & Stones A. R. (1977) A Review of the Distribution and Biology of the potato Cyst Nematodes Globodera rostochiensis and G.pallida.PNAS,23,178-189.
  • GANALM.W, SIMONR.; BROMMONSCHENKELS., ARNDTM. ,PHILLIPSM. S. , TANKSLEYS. D & KUMARA.(1995). Genetic mapping of a wide spectrum nematode resistance gene (Hero) against Globodera rostochiensis in tomato. Molecular plant-microbe interactions,8, 886-891.
  • Gao, B., Allen, R., Maier, T., Davis, E.L., Baum, T.J., Hussey, R.S., 2003.The parasitome of the phytonematode Heterodera glycines. Mol. Plant-Microb. Interact. 14, 1247-1254.
  • Gao, B., R. Allen, T. Maier, J.P. McDermott, E.L.Davis, T.J.Baum & R.S.Hussey. (2002). Characterisation and developmental expression of a chitinase gene in Heterodera glycines. Int J Parasitol. 32, 1293-1300.
  • Gheysen, G. & Jones J.T. (2006) Molecular aspects of plant-nematode interaction. In: Perry, R. N. & Moens, M. (Eds) Plant Nematology. Wallingford, UK, CABI publishing, pp. 234-254.
  • Gheysen, G., Engler, J. D. A. & Montagu, M.V. (1997). Cell cycle regulation in nematode feeding sites. In: Fenoll, C., Grundler, F.M.W. & Ohl, S.A. (ed. ^ eds.) Cellular and molecular aspects of plant-nematode interactions. Kluwer Academic Publishers, pp.120-132.
  • Guo M., Tian, F., Wanboldt, Y. & Alfano, J.R. (2009). The majority of the Type III effector Inventory of Pseudomona syringae pv. Tomato DC3000 Can Suppress Plant Immunity. Molecular Plant-Microbe Interactions. 22, 1069-1080.
  • Huang, G., Gao, B., Maier, T., Allen, R., Davis, E.L., Baum, T.J., Hussey,R.(2003). A profile of putative parasitism genes expressed in the esophageal gland cells of the root-knot nematode Meloidogyne incognita. Mol. Plant-Microb. Interact. 16, 376-381.
  • Hussey, R.S. & C.W. Mims. (1991). Ultrastructure of feeding tubes formed in giant cells induced in plants by the root-knot nematode Meloidogyne incognita. Protoplasma, 162, 99-107.
  • Hussey, R.S., Davis, E.L. & Baum, T.J. (2002). Secrets in secretions: genes that control nemaotdes parasitism of plants. Braz. J. Plant Physiol., 14, 183-194.
  • Ingle, R.A., Carstens, M., & Denby, K.J. (2006). PAMP recognition and the plant pathogen arms race. BioEssays, 28, 880-889.
  • Jaubert, S., Laffaire, J.-B., Abad, P., Rosso, M.-N., (2002b). A polygalacturonase of animal origin isolated from the root-knot nematode Meloidogyne incognita. FEBS Lett. 522, 109-112.
  • Jaubert, S., Ledger, T.N., Laffaire, J.-B., Piotte, C., Abad, P., Rosso, M.-N., (2002a). Direct identification of stylet secreted proteins from root-knot 24 B. Vanholme et al. / Gene 332 (2004) 13-27 nematodes by a proteomic approach. Mol. Biochem. Parasitol. 121,112- 205.
  • Jones, J.D.G. & Dangl, J.L. (2006). The plant immune system. Nature, 444, 323-329.
  • Jones, J.T. & Robertson, W.M. (1997). Nematode secretions. In: (Fenoll, C., Grundler, F.M.W. & Ohl, S.A., ed.^ eds.).Cellular and molecular aspects of plant-nematode interactions. Kluwer Academic Publishers, pp. 98-106.
  • Jones, J.T. Reavy, B., Smant, G & Piror, A.E. (2004) Glutathione peroxidases of the potato cyst nematode Globodera rostochiensis. Gene, 324, 47-54.
  • Jones, J.T., Furlanetto, C., Bakker, E., Banks, B., Blok, V., Chen, Q., Phillips, M., Prior, A., ( 2003). Characterization of a chorismate mutase from the potato cyst nematode Globodera pallida. Mol. Plant Pathol. 4, 43-50.
  • Jones, J.T., Smant, G., Block, V.C. (2000). SXP/RAL-2 proteins of the potato cyst nematode Globodera rostochiensis: secreted proteins from the hypodermis and amphids. Nematology 2, 887- 893.
  • Jones, T.J. & Perry, R.N. (2004) Plant parasitic nematodes- small animals, big impact. Biologist, 51, 1-5.
  • JONES, J.T, A. KUMAR, LILIYA A. PYLYPENKO, AMARNATH THIRUGNANASAMBANDAM, L. CASTELLI, S. CHAPMAN,PETER J. A. COCK, E. GRENIER, CATHERINE J. LILLEY, MARK S. PHILLIPS & V. C. BLOK. (2009). Identification and functional characterization of effectors in expressed sequence tags from various life cycle stages of the potato cyst nematode Globodera pallida. Molcular Plant Pathology.10, 815-828.
  • Keren-Zur, M. Antonov, J. Bercovitz, A. Feldman, A. Keram, G. Morcov & N. Rebhum. (2000). Bacillus firmus formulations for the safe control fo root-knot nematodes. The BCPC conference. Pests and Disease, Bringhton, UK. 307-311.
  • Kru¨ger, J., Thomas, C. M., Golstein, C., Dixon, M. S., Smoker, M., Tang, S.,Mulder, L. and Jones, J. D. G. (2002). A tomato cysteine protease required for Cf-2-dependent disease resistance and suppression of autonecrosis. Science 296, 744-747.
  • Lambert, K.N., Allen, K.D., Sussex, I.M., (1999). Cloning and Characterization of an esophageal-gland-specific chorismate mutase from the phytoparasitic nematode Meloidogyne javanica. Mol. Plant-Microb. Interact.12, 328- 336.
  • Lee, D.L. (2002) Life cycles. In: The biology of Nemaotdes. (Lee, D.L., ed. ^ eds.) London: Taylor and Francis, pp.61-72.
  • Lindblade KA, Arana B, Zea-Flores G, Rizzo N, Porter CH, Dominguez A, Cruz-Ortiz N, Unnasch TR, Punkosdy GA, Richards J, Sauerbrey M, Castro J, Catú E, Oliva O, Richards FO Jr.(2007). Elimination of Onchocercia volvulus transmission in the Santa Rosa focus of Guatemala. Am. J. Trop. Med. Hyg., 77, 334-341.
  • Marie, C.C., Geraldine, D., Michael, Q., Laetitia P.B., Philippe, L., Janice de A.E., Pierre, A., Marie N.R., & Bruno. (2008). F. Root-knot nematode manipulate plant cell functions during a compatible interaction. Journal of Plant Physiology.165, 104-113.
  • Metraux, J.P., Jackson, R.W., Schnettler, E., & Goldbach, R.W. (2009) Plant pathogens as suppressors of host defense. In: L.C.Van Loon (eds), Plant Innate Immunity. 51, 39-89.
  • Moens T, Herman P, Verbeeck L, Steyaert M, Vincx M. (2000). Predation rates and prey selectivity in two predacious estuarine nematode species. Marine Ecology-Progress Series.205, 185-193.
  • Olsen, A.N. & K.Slriver. (2003). Ligand mimicry? Plant-parasitic nematode polypeptide with similarity to CLAVATA3. Trends Plant Sci 8, 55-57.
  • Opperman C.H., Bird D.M., Williamson, V.M., Rokhsar D.S., Burke M., Cohn, J., Cromer J., Diener S., Gajan J., Graham S., Houfek T.D., Liu Q.L., Mitros T., Schaff J., Schaffer R., Scholl E., Sosinsk B.R., Thomas V.P. & Windham E.(2008) Sequence and genetic map of Meloidogyne hapla: a compact nematode genome for plant parasitism. PNAS. 105: 14802-14807.
  • Opperman, C.H., Bird, D.M., & Schaff J.E. (2009) Genomic analysis of the root-knot nematode genome. In: Cell biology of plant nematode parasitism. Springer Berlin/ Heidelberg. pp: 221-237.
  • Oxford English Dictionary, (1998). Edited by J. Simpson & E. weiner. 2nd ed. 20 vols. Oxford: Clarendon Press.
  • Popeijus, H., Blok, V.C., Cardle, L., Bakker, E., Phillips, M.S., Helder, J., Smant, G. & Jones, J.T. (2000). Analysis of genes expressed in second stage juveniles of the potato cyst nematodes Globodera rostochiensis and G. pallida using the expressed sequence tag approach. Nematology 2: 567-574.
  • Poulin, R. & S. Morand. (2000). The diversity of parasites. Q Rev Biol 75, 277-293.
  • proteins of the potato cyst nematode Globodera rostochiensis using RNAi. MPMI 18, 621-625.
  • Qin, L. (2001). Identification of a RanBPM-like gene family specifically expressed in the dorsal glands of infective juveniles of the potato cyst nematode, Laboratory of Nematology. Plant Science, Wageningen University and Research Centrum, Wageningen.
  • Qin, L., Overmars, H., Helder, J., Popeijus, H., van der Voort, J.R.Groenink, W., et al. (2000). An efficient cDNA-AFLP based strategy for the identification of putative pathogenicity factors from the potato cyst nematode Globodera rostochiensis. Mol Plant Microbe Interact, 13, 830-836.
  • Rehman, S., Postma, W., Tytgat, T., Prins, P, Qin, L., Overmars H., et al. A secreted SPRY Domain-Containing Protein (SPRYSEC) from the Plant-Parasitic Nematode Globodera rostochiensis Interacts with a CC-NB-LRR protein from a Susceptible Tomato. MPMI, 22, 330-340.
  • Rice, S.L., B.S.C. Leadbeater & A.R. Stone. (1985). Changes in cell structure in roots of resistant potatoes parasitized by potato cyst nematodes.1. Potatoes with resistance gene H1 derived from Solanum tuberosum ssp. Andigena. Physiol. Plant Pathol. 27, 219-234.
  • Riddle, D. L., Blumenthal, T., Meyer, B.J. & Priess, J.R. (1997). C.elegans II. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, pp1222.
  • Rooney, H. C. E., Van't Klooster, J. W., Van der Hoorn, R. A. L.,Joosten, M. H. A. J., Jones, J. D. G. and De Wit, P. J. G. M. (2005).Cladosporium Avr2 inhibits tomato Rcr3 protease required for Cf-2-dependent disease resistance. Science 308, 1783-1786.
  • Rose, J. K. C.,Ham,K.-S., Darvill, A.G. and Albersheim, P. (2002).Molecular cloning and characterization of glucanase inhibitor proteins:Coevolution of a counterdefense mechanism by plant pathogens. The Plant Cell 14, 1329-1345.
  • Sacco, M.A., Koropacka, K., Grenier, E., Jaubert, M.J., Blanchard, A., Goverse, A., Smant, G & Moffett, P. (2009). The cyst nematode SPRYSEC protein RBP-1 elicits Gpa2 and RanGAP2 dependent plant cell death. Plos pathogens. 5, 1-14.
  • Sacco, M.A., Mansoor, S. & Moffett, P. (2007). A RanGAP protein physically interacts with the NB-LRR protein Rx, and is required for Rx-mediated viral resistance. Plant Journal, 52, 82-93.
  • Sasser, J.N., & D.W. Freckman. (1987). A world perspective on Nematology: the role of the society. In: J.A. Veech and D.W. Dickson. (Eds) Vistas on Nematology, Society of Nematology, Hyattsville, Maryland, pp.7-14.
  • Selena G.I., Dagmar, R.H., Vardis N., Elena, P., Volker, L. & J.P. Rathjen. (2009) AvrPtoB Targets the LysM Receptor Kinase CERK1 to Promote Bacterial Virulence on Plants. Current Biology, 19, 423-429.
  • Sharma, S., & Sharma, R. (1998) Hatch and emergence. In: Sharma, S., (ed. ^ eds) The cyst nematodes. Dordrecht: Kluwer Academic Publishers.
  • Smant, G., J.P. Stokkermans, Y.Yan, J.M. de Boer, T.J. Baum, X. Wang, R.S. Hussey, F.J.Gommers, B. Henrissat, E.L.Davis, J.Helder, A.Schots & J.Bakker. (1998). Endogenous cellulases in animals: isolation of beta-1,4-endoglucanase genes from two species of plant parasites cyst nematodes. Proc.Natl.Acad.Sci.U.S.A 95,4906-4911.
  • Tytgat T., Isabel, V., Bartel, V., Jan De M., Isabelle V., Greetje, G., G. Borgonie, A. Coomans, G.Gheysen. (2005). An SXP/RAL-2 protein produced by the subventral pharyngeal glands. Parasitol. Res, 95, 50-54.
  • Tytgat, T., B. Vanholme, J.De Meutter, M. Claeys, M. Couvreur, I.Vanhoutte, G.Gheysen, W.Van Criekinge, G.Borgonie & A.Coomans. (2004). A new class of ubiquitin extension proteins secreted by the dorsal pharyngeal gland in plant parasitic cyst nematodes. Mol Plant Microbe Interact, 17, 846-852.
  • UNLU, I.O., EHLERS, R.-U. & SUSURLUK, A. (2007). Additional data and first record of the entomopathogenic nematode Steinernema weiseri from Turkey. Nematology 9, 739-741.
  • Van der Hoorn, R. A. L. and Jones, J. D. G. (2004). The plant proteolytic machinery and its role in defence. Current Opinion in Plant Biology 7, 400-407.
  • Von Mende, N., Gravato-Nobre, M. J. & Perry, R. N. (1998). Host finding, invasion and f eeding. In: Sharma, S., (ed. ^ eds) The cyst nematodes. Dordrecht: Kluwer Academic Publishers.
  • Wang, X., Meyers, D., Yan, Y., Baum, T., Smant, G., Hussey, R., Davis, E. (1999). In planta localization of a B-1,4-endoglucanase secreted by Heterodera glycines. Mol. Plant-Microbe. Interact. 12, 64-67.
  • Zemin Ning, Anthony J. Cox, & James C. Mullikin. SSAHA: a fast search method for large DNA databases. Genome Research, 11: 17251729, October 2001. PMC311141.