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Rice is regarded as one of the most utilized staple food for a major part of the world's population, especially in Asia, Latin America, the Middle East, and the West Indies. Around 90 percent of total rice is produced in Asia alone (Mohanty, 2009). Other then Asia, Rice is also becoming an important cereal food and supports lives of many people in other continents like Central America, Europe and Africa. About 50 million tons of rough rice needs to be increased per year by 2015 to meet the world's demand, and the projected demand for Asia is an additional 38 million tons (Pandey, 2008; Mohanty, 2009). Rice is the most important staple cereal crop for more than two billion people in Asia (Mew et al., 2004) and accounts for more than 40 percent of the calorie consumption of most Asians (Pandey, 2008) Interestingly, India is the world's second largest producer of white rice accounting for about 80% of all world rice production. Rice in India is the most dominant crop, and a staple food for the people of the eastern and southern parts of India. The earliest remains of cultivated rice was found in the north and west parts of India and date from around 2000 BC. In India, Rice production is an important part of the national economy (Library of congress, 1995).
With the increasing population, demands for food have been increasing. Increasing the productivity of food by decreasing the severity of biotic and abiotic factors will be a challenging task. Plant-parasitic nematodes are the hidden enemy of crops and are one of the many groups of harmful organisms which depend on plants for their survival and reproduction (Khan, 2008). Plant-parasitic nematodes can cause significant damage to almost all kinds of crops but due to their subterranean habit and microscopic size they remain invisible to the naked eye. The estimated annual yield losses due to plant-parasitic nematodes in the world's major crops are recorded at about 12.3% and 14% in the developing countries (Sasser and Freckman, 1987). In India, recent studies show that plant parasitic nematodes are responsible for both the quantitatively and qualitative yield losses amounting around Rs.240 billion every year (Khan, 2008). Besides this direct damage, these nematodes also serve as predisposing agents in development of disease complexes in association with other microbial agents including fungi, bacteria and viruses. Root-knot nematodes, such as Meloidogyne graminicola are known to cause substantial decrease in rice production. M. graminicola has increasing importance on lowland rice farming systems where water conservation results in intermittent flooding of fields. Yield losses in rice caused by Meloidogyne graminicola range from 20-80% in upland and about 11-73% in intermittently flooded conditions (De Waele & Elsen, 2007). Tolerance to plant-parasitic nematodes is also dependent on water management practices. Rice varieties showing resistance to nematodes may represent the practical and effective means of managing these nematodes in small scale rice farming systems. The absence of classical plant breeding solutions and the limitations of chemical treatment also represent an excellent opportunity for biotechnological applications. Because of the lack in knowledge about the nematode and plant interaction, the nematode problems are not recognized properly. More knowledge on the interaction between rice and nematodes is important and necessary to get the nematode problem under control.
The most evolutionary advanced adaptations for plant parasitism by plant parasitic nematodes are the products called parasitism genes (Gao et al., 2002). The parasitism proteins are secreted by the nematode and play a direct role in plant parasitism. These parasitism proteins secretions mostly originate from the pharyngeal gland cells, but secretions from the chemosensory amphids might also be important (Davis et al., 2004). Many parasitism genes have already been reported from plant-parasitic nematodes including Globodera rostochiensis, Meloidogyne incognita, Heterodera schachtii etc. However, surprisingly no parasitism genes from Meloidogyne graminicola have been reported so far. With the above background, the aims and objectives of the present study are
To clone putative plant cell wall degrading enzymes from the rice root-knot nematode Meloidogyne graminicola
To analyze the cloned putative plant cell wall degrading enzymes from the rice root-knot nematode Meloidogyne graminicola by application of various molecular tools and techniques and to study its expression at different life stages of nematodes.
The experimental work will involve the following steps:
1) Finding out whether these genes are really present in the DNA/RNA of Meloidogyne graminicola by Polymerase Chain Reaction (PCR)
2) Functional analyses by in situ hybridization to check their expression in the gland cells of the nematode
3) To check the expression of the genes in different life stages of Meloidogyne graminicola.
The ability to infect or to parasitize host plants by the rice root-knot nematode Meloidogyne graminicola indicates that the nematodes release certain plant cell wall modifying proteins to disrupt the plant cell wall in addition to the mechanical shearing of plant cell wall by the stylet of the nematode. The parasitism proteins secreted by Meloidogyne graminicola might include plant cell wall modifying enzymes likely to be of prokaryotic origin besides proteins that are capable of localizing in the host plant cell nucleus, suppressors of host defense, proteins that can mimic plant proteins as reported in other cyst and root-knot nematodes.
2. LITERATURE REVIEW
2.1. THE HOST PLANT: RICE
Rice (Oryza sativa) is cultivated in about 150 million ha of land worldwide (Bouman et al., 2006) and the major staple food in south-east Asia and also commonly grown in West Africa and South America. Rice feeds about one half of the world's population. The average yield of rice is about 3-4 tons per ha (Padgham et al., 2004). Rice is the grain with the second-highest production in the world next to maize and the most important grain with respect to calorie intake and human nutrition thereby providing more than one fifth of the calories consumed worldwide (Smith, 1998). The cultivation of rice is well-suited to countries and regions having low labor costs accompanied with high rainfall. Although the parent species of rice are native to regions of South Asia and some parts of Africa, trade and exportation of rice over centuries have made it commonplace in many cultures worldwide.
2.2. RICE FARMING SYSTEMS
2.2.1. IRRIGATED RICE
Irrigated rice farming systems account for about 55% of the world area under rice cultivation and account for 75% of the world total production (Bridge et al., 2005) Generally, land for rice cultivation is prepared while wet and water kept held in the reservoirs. In this system, seeds are usually pre germinated and grown in the wet seed-beds for a period of about 2 weeks (UNCTAD, 2010). Whereas in direct seeding, seeds are frequently pre germinated and can be broadcasted by hand or spread over the water by using aero planes as practice in developed countries. In this farming system, production ranges from 5 tons/ ha in the case of rainy season while more than 10 tons/ ha in dry season, when adapting modern technologies (UNCTAD, 2010).
2.2.2. THE RAINFED LOWLAND RICE
The rainfed lowland rice accounts for 17% of the world production just next to the irrigated rice farming system (UNCTAD, 2010). Bridge et al. (2005) reported that the world rice area planted under rain fed lowland farming system was approximately 31%. It grows on compacted soil and in bunded fields that are able to retain water between 25-50 centimeters of medium level and 0-25 centimeters of low level. This non-irrigated rice is feed by the rainwater or by a local reception tank. The major concern in this type of production system is the risk of drought and unexpected floods (UNCTAD, 2010).
2.2.3. UPLAND OR DRYLAND RICE
The upland or dryland rice represents approximately around 13% of the total world area under rice cultivation (Bridge et al., 2005) and accounts for 4% percent of global rice production (UNCTAD, 2010). Prot and Rahman, (1994) observed that root parasitic nematodes in this ecosystems were highly diversified. In this production system, land is prepared for planting rice and the rice is dry-seeded. This type of ecosystem is common in Brazil, Madagascar, India and other parts of Southeast Asia. It was reported that Meloidogyne spp. (M. graminicola M. incognita, M. javanica and M. oryzae) and Pratylenchus zeae occurred in this environment. However, other researchers reported different species of Heterodera in upland rice farming system, such as H. oryzicola (Rao & Jayaprakash, 1978), H. elachista (Ohshima, 1974) and H. sacchari (Coyne & Plowright, 2000).
2.2.4. DEEPWATER OR FLOOD-PRONE RICE
Deepwater or flood-prone rice is common in regions of Southeast Asia including Bangladesh, Thailand, Cambodia, and in West Africa and South America also (UNCTAD, 2010). Bridge et al. (2005) reported that deepwater rice farming system occurred in the river deltas of South and South-east Asia and it occupied about 3% of the world rice area. In this production system, water level is 1 to 5 meters deep and supplied by rivers, lakes, tides etc. Rice production in this system is usually low due to natural calamities like drought, flood and also due to the low production potential of the cultivars which are grown with few inputs. Interestingly, this rice farming system supports more than 100 million people in the world and most of them living on small family farms and mainly in the rural areas (UNCTAD, 2010).
2.3. DEFENSE RESPONSE OF RICE
Plants are capable to protect themselves against this invasion by inducing complex defense mechanisms, such as changes in biochemical, morphological and molecular characteristics including production of antipathogenic compounds, expression of defense related genes and apoptosis (Van Loon et al., 2006). Dangl and Jones (2001) described that the evolution of mechanisms of virulence which help pathogens overcome basal defense responses in plants is responsible for the evolution of specific disease resistance. Plants are known to defend themselves from pathogen attack by activating a multi component defense mechanism. In host plant defense, the invasion of pathogens is recognized by proteins encoded by plant disease resistance genes (R-gene) that bind specifically to the pathogen-derived avirulence proteins (Avr-gene). Odjakova and Hadjiivanova (2001) described that R- and Avr-gene mediated recognition as gene for gene recognition. Hussey and Janssen (2002) mentioned resistance genes in wild potato species against several Meloidogyne spp. and many of them had been successfully incorporated into modern potato cultivars and other economically important crops. Bari and Jones (2009) and Jones and Dangl (2006) reported a 'zigzag' model of the plant immune system. In this system, pathogen associated molecular patterns (PAMPs) are recognized by the host encoded PRRs (pattern recognition receptors) which results in PTI (PAMP triggered immunity). Successful pathogens secrete effectors that suppress PTI. Thus, disease is induced by pathogen effector triggered susceptibility (ETS). Plants recognize a given effector and activate effector-triggered immunity (ETI) results in disease resistance. Plant disease resistance is enhanced and pathogen growth is restricted by the activation of PAMP triggered immunity (PTI) or effector-triggered immunity (ETI).
Fig.Â A zigzag model illustrates the quantitative output of the plant immune system. Source-. Jones and Dangl, 2006.
According to Odjakova & Hadjiivanova (2001), a hypersensitive response (HR) is induced and different defense genes are expressed by the action of those compounds as secondary messengers. Zinov'eva et al. (2004) mentioned that after the reaction against plant-parasitic nematodes, two types of cell death occurred of which the first one was cell necrosis and the second type was genetically programmed cell death or apoptosis. Systemic resistance against new infections to other plant parts could be acquired by the plant after this primary infection and this phenomenon was described by Durrant and Dong (2004) as SAR (systemic acquired resistance). Systemic acquired resistance induces defense that confers long-lasting immunity against a wide range of microorganisms. The defense response of rice to M. graminicola infection has not yet been observed and there is limited information of defense response of rice to other nematodes also.
2.4. NEMATODE PROBLEMS IN RICE
More than 35 genera and 200 species (Prot et al., 1994) of nematodes have been reported in rice. These nematodes can be a major constraint to the high yields of rice. In upland rice, Meloidogyne spp. and Pratylenchus spp. are known to cause more damage while in deep water rice, very few nematodes have been reported including M. graminicola, Ditylenchus dipsaci, causing the ufra disease. Aphelenchoides besseyi, causes white tip disease in rice and is found in most ecosystems (Bridge et al., 1990), while other nematodes are not distributed homogeneously across the ecosystems (Prot et al., 1994). The foliar nematode parasites, Aphelenchoides besseyi and Ditylenchus dipsaci, cause visible symptoms in the foliage and hence are detected easily. In addition to Aphelenchoides besseyi, Hirshmanniella spp., cause major problems in irrigated rice. However, there have been reports of M. graminicola or other Meloidogyne species infesting deep water rice apart from the observations that a species of Meloidogyne, referred to as M. exigua, occurs in the deep water rice farming region in Thailand (Hashioka, 1963 ; Kanjanasoon, 1962 ; Ou, 1972).
2.5. THE ROOT-KNOT NEMATODES: Meloidogyne GOLDI, 1892
The root-knot nematodes are known to be responsible for billions of dollars of economic loss worldwide each year on over 5,000 host species (Sasser and Freckman, 1987). Meloidogyne spp. is considered as one among the most economically damaging genera of plant-parasitic nematodes on horticultural and field crops. It is distributed worldwide, obligate parasites of the roots of thousands of plant species, including monocotyledon and dicotyledon herbaceous and woody plants. Root-knot nematode is distributed both in tropics and temperate regions (Dhandaydham et al., 2008). According to the description of Eisenback & Triantaphyllou (1991), root knot nematodes, Meloidogyne spp., are sexually dimorphic; females are globose and 0.3-0.7 mm in diameter with a slender neck embedded in root tissue; their vulva is present near the anus and is subterminal; body cuticle is thin, annulated and whitish. Male nematodes are vermiform, 1-2 mm long and free-living in soil; they have robust spicules but bursa is absent. As these nematodes are endoparasites, their stylet is short and moderately sclerotized. According to Hunt et al. (2005), the excretory pore of these nematodes is often located near the stylet base; their eggs are deposited in a gelatinous matrix which is present outside the body; the juveniles (J2) of these nematodes are vermiform, slender and about 450 Âµm long having a weakly sclerotized stylet. According to Shurtleff and Averre (2002), the average length of the stylet of the root-knot nematodes is 10-20 Âµm but may have a little variation among J2, males and females. The systematic position of root-knot nematodes as mentioned by De Ley & Blaxter (2002) is: Phylum: Nematoda; Class: Chromadorea; Order: Rhabditida; Suborder: Tylenchina; Infraorder: Tylenchomorpha; Superfamily: Tylenchoidae; Family: Meloidogynidae; Genus: Meloidogyne. De Waele & Elsen (2007) reviewed that 92 nominal Meloidogyne species had been described by 2006. Major species of Meloidogyne include M. graminicola, M. mayaguensis. M. chitwoodi, M. hapla, M. incognita, M. javanica, M. arenaria, M. exigua and M. falax. Among these, M. graminicola is considered as the most common and important nematode for rice root-knot nematodes according to Hunt et al. (2005); De Waele and Elsen (2007); Jepson (1987).
2.6. THE ROOT-KNOT NEMATODE: Meloidogyne graminicola
2.6.1. Meloidogyne graminicola
M. graminicola is a common species of the tropics and subtropics where it infects rice and it is a facultative, meiotic parthenogen, with a haploid chromosome number of 18. M. graminicola is a sedentary endoparasitic nematode and first reported by Golden and Birchfield in 1965 from grasses in the United States. The rice root-knot nematode M. graminicola is known to be established in India, China, Nepal, Bangladesh, Laos, Thailand, United States and Vietnam (Yik et al., 1977, 1979; Kihn, 1982; Poudyal, 2005). De Waele and Elsen (2007) described Meloidogyne graminicola as the most damaging Meloidogyne species on rice in shallow intermittently flooded land and upland conditions and also reported that it was not only distributed in main rice producing area, specially the South and Southeast Asia, but was also found to be distributed in Brazil, Colombia, USA and South Africa and is prevalent in rainfed (upland), irrigated (lowland) and deepwater rice ecosystem. Meloidogyne graminicola has been reported from rice growing regions of India (Isreal, Rao & Rao, 1963; Roy, 1973), Laos (Manser, 1968), Thailand (Buangsuwon et al., 1971), U.S.A. (Golden & Birchfield, 1968; Yik & Birchfield, 1979), and Bangladesh (Hoque & Talukdar, 1971 ; Page et al., 1979). It has been reported mainly from rice growing in upland conditions and nurseries (Buangsuwon et al., 1971 ; Israel, Rao & Rao, 1963 ; Manser, 1968 ; Rao & Israel, 1971, 1972), and also reported to be absent from the rice crop grown in flooded fields (Buangsuwon et al., 1971 ; Manser, 1968).
2.6.2. SYMPTOMS OF Meloidogyne graminicola ON RICE
The common symptoms due to Meloidogyne graminicola infection on rice include roots with hooked-like galls, young leaves with distorted and crinkled sign in margins, slow growth, young plants cholorosis and the heavily infected rice mature and flower earlier than the healthy rice. Deep water rice varieties can elongate to come above the water surface; however, upon severe infection with M. graminicola they are unable to grow and drown, leaving patches of open water in flooded fields (Bridge & Page, 1982). Small galls appear on the roots as beaded, clubbed or spindle-shaped, which coalesce upon heavy infection. Characteristic hooked and swollen root tips are apparent which prevent root elongation (De Waele & Elsen, 2007).
Fig. Meloidogyne graminicola root galls on rice seedlings. Source-http://www.warda.org/publications/Warda_Nemaotde.pdf. 6thDec2010.
Fig.Characterisics hooked tips caused by Meloidogyne graminicola. Source-http://www.warda.org/publications/Warda_Nemaotde.pdf. 6thDec2010.
2.6.3. LIFE STAGES OF Meloidogyne graminicola
The second stage juveniles or J2 stages of M. graminicola penetrate the root behind the root tip. This leads to giant cell formation due to feeding by juveniles and hypertrophy of cells which further leads to the development of galls. The gall then continues to swell, with females of J4 and males of J4 inside the galls. As the life cycle goes on, the male leaves the root while the female stays inside the root and starts laying eggs in a gelatinous matrix outside the root. J2 hatch from the eggs and are attracted towards the root and in this way, the life cycle carries on. The life cycle depending on the environmental conditions takes around one to two months. However, there are reports that the life cycle of the rice root-knot nematodes under favorable conditions at 25-30Â°C takes 19 days (Bridge & Page, 1982). M. graminicola normally feeds on the cells and interferes with nutrient uptake, water uptake, and translocation due to root damage (Caillaud et al., 2008). Gheysen and Jones (2006) described that juveniles of Meloidogyne entered the root tip and migrated intercellularly towards the root tip. After reaching the root apex, they turned back and migrated again intercellularly until they found a suitable place near vascular cylinder for feeding site induction. Karsen and Moens (2006) also reported that like other root-knot nematodes, M. graminicola started feeding on a group of giant cells established in the phloem or adjacent parenchyma. Rao and Israel (1973) reported that this nematode completes its life cycle in 26-51 days in India on the other hand; Bridge and Page (1982) reported that this nematode species completes its life cycle in less than 3 weeks at 22-29Â°C in Bangladesh, resulting in building up of huge population densities during a single crop cycle.
Fig. Life cycle of Root knot nematode. Source-
2.6.4. RICE YIELD IN RESPONSE TO M. graminicola INFECTION
The rice root-knot nematode can attack the rice plant during all the growth stages and considered as one of the limiting factors in rice production. There is an estimated reduction of 2.6 percent in the yield of rice for every 1000 nematodes present in young seedlings as recorded in upland rice farming system and in irrigated rice (Rice knowledge bank, 2010). Greatest yield decline is reported under non flooded conditions (Plowright & Bridge, 1990; Prot & Matias, 1995; Tandingan et al., 1996; Soriano et al., 2000; Soriano & Reversat, 2003) however, great yield losses can also occur in drought-prone rain fed lowland systems (Padgham et al., 2004). Soriano et al. (2000) recorded yield losses due to M. graminicola which ranged from 11% to 73% in simulations of intermittently flooded rice.
2.6.5. MANAGEMENT PRINCIPLES OF Meloidogyne graminicola
There are various control measures that are available for managing rice root-knot nematodes. This includes biological, physical, chemical, cultural, resistant cultivars and mechanical control. The common cultural controls that are put into practices include continuous flooding of the rice field, raising the rice seedlings in flooded soils and also crop rotation. M. graminicola control can be achieved by continuous flooding of rice plants in deep water (Bridge et al., 2005). Rao and Israel (1971) and Soriano et al. (2000) proposed that early-season flooding can reduce M. graminicola damage on rice as the Meloidogyne juveniles cannot invade the roots in flooded conditions. Crop rotation with marigold (Tagetes sp.) is also found to be effective in lowering the root-knot nematode populations due to its nematicidal properties. There are several nematicidal compounds which can be used as chemical control. Other control methods includes the application of chemicals like fensulfothion or oxamyl, carbofuran, phorate etc. Soriano and Reversat (2003) reported that carbofuran controlled the rice root-knot nematode and improved yield of the first rice crop but did not affect the second rice crop.
2.7. NEMATODE PARASITISM GENES
Root-knot nematodes which are highly successful parasites evolved a very specialized feeding relationship with the host plant or crops to cause the destructive root-knot disease. They initiate their parasitic relationship with the host by releasing their secretions into root cells which in turn stimulate the root cells of the host to become specialized feeding cells which are considered as the single source of nutrients essential for the nematode's survival (Baum et al., 2007). A deeper understanding of the basic principles and mechanisms of root-knot nematode parasitism is critical for discovering new targets in root-knot nematodes to develop novel crop resistance using biotechnology tools.
Fig. Model of potential interactions of some secreted products of nematode parasitism genes with host plant cells. Source- Davis et al., 2004
The parasitism genes expressed in the root-knot nematode's esophageal gland cells encode secretory proteins that are released through its stylet to direct the interactions of the nematode with its host plants (Huang et al., 2005). The products which are collectively called parasitome, of parasitism genes secreted into susceptible host tissue modulate the complex changes in function, morphology and gene expression in host root cells to form feeding cells. Plant-parasitic nematodes are well equipped with a stylet to tear the cell walls and allow exchange of solute between plant and parasite and also have well-developed secretory gland cells associated with the esophagus that produce secretions released through the stylet into host. In order to be parasitic, the nematode must be able to penetrate the roots of host plants and migrate through root tissues. Despite the small size of root-knot and cyst nematode infective J2 stage, plant cell walls serve as an obstacles so nematodes release a mixture of cell-wall-digesting enzymes to break structural plant cell-walls (Baum et al., 2007) Above this, the most interesting things appear to be the nematode-directed formation of the feeding cells by both the root-knot nematodes as giant-cells and cyst nematodes as syncytia. Gheysen and Jones (2006) also suggested that both induction and maintenance of giant cells were controlled by stylet secretions. These secretions were originated from dorsal and subventral pharyngeal glands of feeding nematodes. They also reported that the nematode genes expressed solely in subventral gland cells were most similar to the genes that produced cell wall degradation enzymes from bacteria and were not present in symbiotic bacteria. Zinov'eva et al. (2004) summarized several gene products isolated from subventral glands of nematodes which included lipoprotein, cellulose-binding protein, endoglucanase, chitinase, pectinase and proteinase.
2.8. PARASITISM GENE IDENTIFICATION
A number of approaches to identify the nematode parasitism genes and proteins have been devised and tried. Many of these approaches mainly focus on esophageal-glands due to their active involvement in parasitism. The identification of parasitism genes has proven to be difficult due to the microscopic size of the plant-parasitic nematodes, which makes it hard to collect enough material for analysis (Vanholme et al., 2004). The peptide sequence from an antigen purified with an esophageal-gland-specific monoclonal antibody was used to isolate the first parasitism gene from a plant-parasitic nematode (Smant et al., 1998), encoding a Î²-1,4-endoglucanase enzyme (cellulase). Immunoaffinity purification was used to enrich secreted proteins, which resulted in the finding of a secreted protein (endo-1, 4- Î²-glucanase) from the subventral glands of the cyst nematode Globodera rostochiensis (Smant et al., 1998). Another method to be mentioned is the analysis of collected nematode secretions by 2D gel electrophoresis and microsequencing. This has also proved successful for the beet cyst nematode Heterodera schachtii (De Meutter et al., 2001) and the root-knot nematode Meloidogyne incognita (Jaubert et al., 2002a). Recently, mass spectometry has also been used for direct identification of proteins secreted by M. incognita, revealing proteins with host cell reprogramming potential (Bellafiore et al., 2008). Further studies also reveal the expression of cyst nematode endoglucanase genes and their associated products specifically from subventral gland cells of the nematode, cellulolytic activity of the enzymes, and secretion of cellulases from the stylet in the plant during migration of infective J2 stage inside the host roots (Wang et al., 1999). These studies also confirmed and refined the previous reports of secreted cell-wall-modifying enzymes from plant parasitic nematodes (Deubert et al. 1971). An important discovery of the nematode endoglucanases was their strong resemblance to prokaryote (glycosyl hydrolase family 5) endoglucanases and little similarity to endoglucanases of eukaryotes and no similarity to any gene of Caenorhabditis elegans (Smant et al., 1998)
2.9. ORIGINS OF NEMATODE PARASITISM GENES
Plant-parasitism is believed to have evolved at least three times independently (Blaxter et al., 1998). The genes that were evolved from nematode ancestors of contemporary species are one likely possible mechanism for the origin of nematode parasitism genes and the other mechanism may be horizontal gene transfer (HGT). It was reported that those genes expressed in the esophageal gland cells of plant-parasitic nematodes show strongest similarities to the bacterial genes which strengthened the existing hypothesis that parasitism genes in plant-nematodes may have been acquired, at least in part, by horizontal gene transfer from bacteria and other microorganisms that inhabit the same parasitic environment. The genes Mj-cm-1and Mi-cbp-1 shows strongest similarities to the genes of bacteria (Ding et al., 1998: Lambert et al., 1999). The complementation of a bacterial mutant with Mj-cm-1 was also used to provide functional analysis of the gene (Lambert et al., 1999). Most of the parasitism genes are found to be highly similar to bacterial sequences thereby suggesting that these parasitism genes could have been acquired from bacteria through horizontal gene transfer (Danchin et al., 2010). For example, the nematode endo-1,4-Î²-glucanases from the Tylenchomorpha, which belong to Glycosyl Hydrolase Family (GHF5), show less similarity to plant endoglucanases but show resemblance to the bacteria. The genes encoding the cellulases enzymes of both nematode and bacteria may have evolved from an ancient cellulase of a common ancestor of both the bacteria and nematodes. The endoglucanases from nematodes show the highest similarity with the bacterial ones, which also points to a HGT from bacteria to an ancestor of the cyst nematode. However, it is not possible and advisable to provide the conclusive evidence for a horizontal gene transfer (HGT) from one organism to another organism germ line. There are examples of putative cases of horizontal gene transfer from eukaryote to prokaryote, prokaryote to prokaryote and from prokaryote to eukaryote (Smith et al., 1992: Syvanen, 1994). On the other hand, the presence of bacterial symbionts in nematode ancestors, such as the bacterium Wolbachia symbiont found in filarial nematodes (Blaxter et al., 1999), may also represent a source for transfer of genetic material from bacteria to nematodes.
2.10. PLANT CELL WALL STRUCTURE AND COMPOSITION
Plant cell walls are complex mixture of carbohydrates, proteins, lignin, water and other substances including cutin, suberin, with inorganic compounds that differ from plant to plant species and even cell types. This composition and structural variation is further increased due to various developmental events and exposure to various abiotic and biotic stresses (Showalter, 1993). The major carbohydrates in the growing plant cell wall consist of cellulose, hemicellulose and pectin. The cellulose microfibrils are linked to hemicellulose forming the network of cellulose-hemicellulose which is embedded in the pectin matrix. Plants cell walls are composed of three types of layers- the middle lamella, the primary cell wall and in some case the secondary cell wall. The middle lamella is found to be rich in pectin and deposited after mitosis and connects the two adjacent plant cells (Cosgrove, 2005). The major polysaccharides in the primary cell wall are cellulose, pectin and hemicellulose whereas secondary cell walls are composed of cellulose, lignin, and hemicellulose like xylan, glucuronoxylan, arabinoxylan and glucomannan. Structural proteins are also found in most plant cell walls and are classified as Hydroxyproline rich glycoproteins (HRGP), Glycine-rich proteins (GRPs), Arabinogalactan proteins (AGP) and Proline-rich proteins (PRPs). These proteins are found to be concentrated in specialized cells and in cell corners also. Plant cell walls of the epidermis and endodermis may also contain suberin or cutin, two polyester-like polymers that protect the cell from herbivores (Moire et al., 1999). Plant cells walls also contain numerous enzymes, such as hydrolases, peroxidases, Î±-mannosidases, Î²-mannosidases, Î²-1,3-glucanases, Î²-1,4-glucanases, polygalacturonase, invertases, malate dehydrogenase, arabinosidases, pxylosidases, proteases and ascorbic acid oxidase (Varner and Lin, 1989).
2.11. NEMATODE PLANT CELL WALL DEGRADING ENZYMES
The most widely studied cell wall degrading enzyme in nematodes till date is cellulase or endo-1, 4-Î²-glucanase. These enzymes degrade the cellulose which is the structural component of the plant cell wall. The cellulases were identified in the sedentary nematode genera Heterodera, Globodera and Meloidogyne (Smant et al., 1998). It belongs to the glycosyl hydrolase family 5 (GHF5) and are believed to be adopted from bacteria through horizontal gene transfer (HGT) (Jones et al., 2005; Danchin et al., 2010). The plant cell wall digesting enzymes cellulase and pectinase have been described for root-knot nematodes (Huang et al., 2004; Huang et al., 2003; Huang et al., 2005; Rosso et al., 1999) and cyst nematode species (Smant et al., 1998: Gao et al., 2003; Wang et al., 2001; Yan et al., 2001). The first major achievement in parasitism gene discovery was the discovery of cellulase genes from the soybean and potato cyst nematodes. The discovery of cellulase genes was very important since no cellulase genes had been reported from animals at that time (Smant et al., 1998). The enzymes beta-1, 4-endoglucanase genes from Pratylenchus penetrans (Uehara et al., 2001), a migratory parasite that also requires enzymes to enter the plant cell-walls was reported later. An enzyme of the beta-1, 3-endoglucanase type was also recently reported from Bursaphelenchus xenophilus, the pinewood nematode where it is hypothesized of being involved in nematode feeding from the fungal mycelium (Kikuchi et al., 2005).
2.11.2. PECTATE LYASE
The enzyme pectate lyase can cleave the internal (1, 4)-Î±-linkages of pectate and was identified in Meloidogyne spp., but has also been reported from other genera such as Heterodera, Globodera and Bursaphelenchus (Popeijus et al., 2000; Doyle & Lambert, 2002; Huang et al., 2005; Kikuchi et al., 2006; Vanholme et al., 2007; Abad et al., 2008). The pectinase proteins obtained from nematode were of the type pectate lyase which are found in fungi and bacteria, cyst and root-knot nematodes; (Popeijus et al., 2000; Huang et al., 2003; Huang et al., 2005; De Boer et al., 2002; Doyle & Lambert, 2002) or to the polygalacturonase type of bacteria (Jaubert et al., 2002). The involvement of these enzymes in penetration and migration is well supported by the evidence that these enzymes are produced and released during nematode penetration and migration and to a smaller extent, or not at all during the sedentary stages of the nematodes (Huang et al., 2005; Rosso et al., 1999; De Boer et al., 1999; Goellner et al., 2000). These types of cell wall digesting enzymes are also reported from outside the community of sedentary nematodes.
The Mi-pg-1 gene encoding a functional polygalacturonase (PG) from M. incognita was the first known animal polygalacturonase (PG) cloned (Jaubert et al., 2002). These enzymes help in catalyzing the hydrolysis of pectic polygalacturonic acid and in turn release oligogalacturonides. PGs are classified into two classes namely exo-PGs and endo-PGs depending on their mode of action. The gene Mi-pg-1 encodes a 633 amino acid protein. Phylogenetic analysis reveals that Mi-pg-1 is closer to PGs from prokaryotes than to eukaryotic enzymes. The close similarity between bacterial PGs and Mi-pg-1 provides strong evidence supporting the hypothesis that the parasitism genes in nematodes may have been acquired through gene transfer from microorganisms. More interestingly, M. incognita PG could play an important role in weakening the plant cell walls of root tissue during nematode penetration and intercellular migration by the parasite like other nematode parasitism genes.
The most abundant polysaccharide in nature next to cellulose is xylan and is composed of (1, 4)-Î²-linked xylopyranose units (Collins et al., 2005). The enzymes endo-1,4-Î²-xylanases depolymerise the nonhydrolysed xylan polymer by cleaving the xylan backbone (Subramaniyan & Prema, 2002). The characterization of the first functional animal endo-1, 4-Î²-xylanase gene was reported from Meloidogyne incognita, the southern root-knot nematode. The nematode endoxylanase, Mi-XYL1, has similarity to bacterial endoxylanases (Mitreva- Dautova et al., 2006). Most of these enzymes have been designated to the family GHF5, although there is some confusion about this classification since the proteins have similarity to GHF30 enzymes as well. Expressed sequence tag (EST) study on Radopholus similis, the migratory nematode revealed some interesting genes, including an EST with homology to an endo-1,4-Î²-xylanase that was reported recently by Jacob et al. (2008) and analyzed by Haegeman et al. (2009).
There is evidence that the potato cyst nematode also secretes a protein that has the ability to break the non-covalent bonds in plant cell walls in addition to the ability of breaking down the covalent bonds found in plant cell-walls through the enzyme cellulase and pectinase (Baum et al, 2007). This type of activity is accomplished by an expansin-like protein found in the potato cyst nematodes (Qin et al., 2004), which is also the first confirmed report of such protein from outside the plant kingdom. Expansins are involved in softening the plant cell-walls by breaking the non-covalent bonds between cell-wall-fibrils. The resultant cell-wall softening could also be demonstrated for the potato cyst nematode expansin parasitism protein (Qin et al., 2004).
2.12. OTHER PARASITISM GENES
2.12.1. CHORISMATE MUTASE
Chorismate is the precursor for a number of compounds like cellular aromatic amino acids and the plant hormone indole-3-acetic acid, related to salicyclic acid and other secondary metabolites (Dewick, 1998). This chorismate-derived compounds plays an important role in plant growth and development, in plant defense, and also in interactions with other organisms (Schmid and Amrhein, 1995; Weaver and Hermann, 1997). The enzyme chorismate mutase catalysed the pericyclic claisen-like rearrangement of chorismate to prephenate in the shikimate pathway, which is a primary metabolic pathway found in plants and other micro-organisms (Romero et al, 1995). This enzyme is well characterized from microbes and plants, and not described from any other animals outside the plant-parasitic nematodes (Roberts et al., 1998; Romero et al., 1995; Schmid and Amrhein, 1995). The first animal chorismate mutase gene (Mj-cm-1) was cloned from Meloidogyne javanica and found to be expressed in the oesophageal gland cells of the nematode (Lambert et al., 1999). The enzyme chorismate mutase is known to be involved in early development of the feeding sites induced by the plant parasitic nematodes, but how this enzyme alters the development of plant cells is not properly known (Doyle and Lambert, 2003). Chorismate mutase was also identified from soybean and potato cyst nematodes (Bekal et al., 2003; Gao et al., 2003; Jones et al., 2003).
Chitinase is a putative parasitism protein, identified from the subventral glands of the soybean cyst nematode (Gao et al., 2002). This parasitism protein has a clearly defined function but no clear role for this function during the production of protein. The occurrence of chitin in nematode has been found only in the egg shell (Bird et al., 1991) and the presence of this parasitism protein chitinases has been discussed as having a role in the hatching of nematode (Baum et al., 2007). In situ expression (Gao et al., 2002) and microarray expression studies demonstrate that chitinase is not found to be expressed in the eggs of nematodes but has a strong expression peak in the early phases of parasitism after penetration inside the plants (Baum et al., 2007).
Annexin genes represent a family that codes for calcium dependent phospholipid binding proteins and has a broad range of reported functions. The mRNA for a secretory isoform of an annexin-like protein was also reported to be expressed in the dorsal gland of the soybean cyst nematode (Gao et al., 2002). However, no clear confirmation about its role in parasitism can be drawn at this time. This gene is also reported from Globodera pallida, the potato cyst nematode (Baum et al., 2007).
Calreticulin-like proteins are also reported to be secreted from other plant parasitic nematodes and are regarded as good candidates for a role in parasite-host interactions (Nakhasi et al., 1998, Pritchard et al., 1999). A calreticulin-like protein preceded by a signal peptide was reported to be secreted from the subventral glands of a root knot nematode (Jaubert et al., 2002). The confusing array of putative or demonstrated calreticulin functions reported (Nakhasi et al., 1998) make it difficult to confirm its role in host parasitism by the root-knot nematodes.
2.12.5. SMALL BIOACTIVE PEPTIDES
The most commonly expressed parasitism gene in Heterodera glycines, the soybean cyst nematode was first identified as clone HG-SYV46 (Wang et al., 2001). The computational analyses found out that the C-terminal domain of HG-SYV46 is related to the members of the CLAVATA3-ESR-like (CLE) family of signaling proteins in Arabidopsis (Olsen and Skriver, 2003). The CLAVATA3 in Arabidopsis has been identified as a key factor determining shoot meristem differentiation (Fletcher et al., 1999). The expression of the cDNA of Heterodera glycines CLAVATA3-like peptide in the clavata3 (clv3) Arabidopsis mutant was found to restore the wild-type phenotype being the first report of ligand mimicry in plant nematode interactions (Wang et al., 2005). It will be interesting to find out the role of the small C-terminal extension of the cyst nematode ubiquitin extension proteins when considering the importance of small peptides in signaling roles of plant development and plant nematode interactions (Tytgat et al., 2004; Gao et al., 2003). The role of small peptides in nematode-plant interactions is also confirmed by an unknown peptide fraction smaller than 3 kilo Dalton isolated from potato cyst nematode secretions that affects plant cell division (Goverse et al., 1999).
2.13. GENOMES OF M. incognita AND M. hapla
The complete genome sequencing of two root-knot nematodes reveals that M. hapla encodes approximately 14,200 genes in a compact 54 Mbp genome whereas the 86 Mbp of M incognita genome encodes approximately 19,200 genes (Abad et al., 2008; Bird et al., 2009). It was reported that M. incognita has a set of 61 plant cell wall degrading carbohydrate enzymes but a few of these enzymes were identified previously in some plant-parasitic nematodes and in insect (Caillaud et al., 2008; Davis et al., 2004; Wei et al., 2006). All together, 21 cellulases and 6 xylanases enzymes from family GHF5, 2 polygalacturonases enzymes from family GHF28 and 30 pectate lyases enzymes from family PL3 were identified from M. incognita (Abad et al., 2008). Two additional plant cell wall degrading enzymes arabinases (family GH43) and two invertases (family GH32) were also identified. Invertase is an enzyme that catalyses the conversion of sucrose into glucose and fructose, which the M. incognita uses as a carbon source. The enzymes expansins and invertases were probably acquired through horizontal gene transfer as most proteins were similar to bacteria. M. incognita also secreted four metabolic enzymes chorismate mutases (Lambert et al., 1999), which are highly similar to bacterial enzymes. Almost half of the genes in Meloidogyne hapla show the highest similarities to Caenorhabditis elegans genes as based on database similarity and the second largest group showing similarity is the animal-parasitic nematodes (Opperman et al., 2008) M. hapla encodes the enzyme cellulases at six loci. Only four of this has EST support which suggests that two of them are pseudogenes. The most attracting example of a root knot nematode gene acquired through horizontal gene transfer is pectate lyase. It was originally discovered as a secreted protein (Davis and Mitchum, 2005; Vanholme et al., 2004). The genome sequence of M. hapla sequence reveals that M. hapla encodes a family of 22 pectate lyases which is common in plant pathogenic fungi (Opperman et al., 2008).