Rice (Oryza sativa L.) is considered as one of the most important food crops in the world and more than half of the global population are dependent on rice for their food (Matsumoto et al., 2005). Although rice is a carbohydrate-rich food, it also supplies a substantial proportion of protein, fat, vitamin, minerals and antioxidants (Kennedy & Burlingame, 2003; Yawadio et al., 2007; Itani & Ogawa, 2004; Suzuki et al., 2004; Lee et al., 2008; Chen et al., 2006). Rice has been established as a model monocot because of its small genome size (390Mb) compared to other major cereals (Sasaki et al., 2005), its synteny with other cereals (Gale & Devos, 1998) and transformation efficiency (Ashikari et al., 2005). The importance of rice is increasing rapidly in Africa and Latin America, although Asia is the main center of production and consumption of rice. Moreover, exports of rice to the USA and Australia are of major economic importance to the rice trade in the world .
Rice is grown in almost all environmental conditions such as in lowland, uplands, deep water, in tidal wetlands, irrigated land and also in rain fed conditions (Chopra & Prakash, 2002). Wetland and upland systems are the two major production systems of rice , with 85% area under wetland production of the total rice production system (Bouman et al., 2007). As a result, rice production is one of the largest water consumers in the world (Chapagain & Hoekstra, 2011). However, the productivity of Asia’s irrigated rice systems is being increasingly threatened by water scarcity (Bouman et al., 2005). An increasing scarcity of water necessitates the development of production systems that require less water than traditional flooded rice. Aerobic rice system is such a production system, in which rice is grown in non-flooded and non-saturated soil under supplemental irrigation as necessary (Bouman et al., 2005).
Rice suffer from various biotic stress factors, like fungal, bacterial, viral and nematode diseases (Webster & Gunnell, 1992). Moreover, aerobic rice suffers from various abiotic (increase of soil pH, and deficiencies of Mn and Fe) stress factors. Among biotic stress, pathogenic fungi (Fusarium sp., and a Rhizoctonia-like species), oomycete Pythium spp, and root-knot nematodes are isolated from root samples (Kreye et al., 2009). Moreover, it is also reported that Pythium spp. and nematodes caused most of the damages in the root system of rice (Mew et al., 2004; Luc et al., 2005).
The large scale introduction and continuous cultivation of rice in the aerobic production system is promoting the buildup of large populations of Meloidogyne graminicola (De Waele & Elsen, 2007). Yield losses on rice caused by M. graminicola range in between 20% and 70% in many countries (Pokharel et al., 2007; Soriano et al., 2000; Padgham et al., 2004). As soil-borne pathogens attack the same roots, there are many interactions going on between the pathogens. In many cases double infection by nematodes and fungal pathogens or oomycetes cause increased damage to the plants compared to single infections (Bhattarai et al., 2009). Interestingly, in some cases an increased yield has been recorded after double infection compared to single infection. For example, double infection of P. arrhenomanes and M. graminicola on two rice cultivars has been shown to give a higher yield compared to single infection by M. graminicola (Kreye et al., 2010b). Here, we want to study the underlying mechanism that causes the increased yield caused by an interaction between P. arrhenomanes, and M. graminicola in rice roots.
2. Pythium as Oomycete Pathogen: Yield loss and distribution
Based on scientific and economic importance Pythium has been ranked as one of the 10 most important oomycetes in the world Kamoun et.al.,2015). Pythium spp. are responsible for yield loss in rye (Scott, 1987), in maize (Sumner et al., 1990) and in wheat and barley (Waller, 1979). In rice growing areas, Pythium spp. are one of the most important groups of seedling disease pathogens. Different pathogenic oomycetes e.g. P. monospermum, P. echinocarpum, P. nagaii, and P. oryzae are isolated from diseased seeds from the various localities in Japan (Ito & Tokunaga, 1933). P. graminicola is the most frequently isolated species among 91 isolates from rice fields in Korea affecting the germination capacity of the rice seed (Sung et al., 1983). P. aristosportum inhibits the growth of rice seedlings in the flooded rice field in Japan (Furuya et al., 2003). In Calfornia (USA), P. torulosum causes damping-off of wild rice (Zizania palustris L.) (Marcum & Davis, 2006). P. arrhenomanes is first detected from maize roots in Wisconsin and Illinois state, USA (Drechsler, 1928). It is later found in 47% of rice soils sampled across the Riverine Plain in New South Wales, Australia (Cother & Gilbert, 1992). P. arrhenomanes and P. myriotylum reduce the root development of rice significantly (Chun & Schneider, 1998). P. arrhenomanes causes seedling blight of rice in Japan (Beneventi et al., 2013). P. arrhenomanes is responsible for pre- and post-emergence death of rice seedlings and cause the growth reduction of root and shoot of the surviving seedlings by 70% and 48 % respectively and is considered to be the most pathogenic to rice (Cother & Gilbert, 1993; Eberle et al., 2007). P. arrhenomanes, P. graminicola and P. inflatum are isolated from aerobic rice fields in the Philippines, among which all the isolates of P. arrhenomanes are pathogenic to rice and reduce the growth of rice seedlings (Van Buyten & Hofte, 2013).
2.1 Life cycle and behavior of Pythium
In absence of the host plant, Pythium spp. survive in the soil both as sporangia (asexual resting spores) and oospores (sexually produced, thick-walled resting spores). Under unfavorable environmental conditions, Pythium spp. undergo a sexual life cycle. Most of the Pythium spp. are of homothallic-type (compatible male and female gametes are produced within the same sporangium). However, there are some heterothallic-type (compatible male and female gametes are produced in different sporangium ) Pythium spp., where oogonial hyphae of the female mating type produce a hormone to stimulate the formation of antheridia of the male mating type. At first, the female oogonim is fertilized by the male antheridium, which results in the formation of typical thick-walled oospores (Martin, 2009). The oospores are dormant and resistant to desiccation and can survive in the soil for up to 12 years in absence of a suitable host or organic substrates (Martin & Loper, 1999; van West et al., 2003). Under favorable conditions, oospores germinate to germ tubes (infective propagule), which form sporangia (Martin & Loper, 1999; van West et al., 2003). This happens in response to suitable chemical stimuli for example amino acids, carbohydrates, volatile compounds (ethanol or aldehydes) present in root and seed exudates, plant debris, or organic matter (Stanghellini & Hancock, 1971; Lifshitz et al., 1986; Nelson & Craft, 1989; Nelson, 1991; Paulitz, 1991). However, sporangia of some Pythium spp. produce zoospores in response to certain nutrients and high moisture content present in the soil. Zoospores are attracted to newly germinating seeds or roots of young seedlings in presence of free water where they penetrate into the host. To infect susceptible plants these motile, biflagellate zoospores can survive in the field for a limited period of time. Under adverse conditions, the zoospores may encyst in the soil, and may remain viable until suitable conditions such as favorable soil moisture content and temperature return (Martin & Loper, 1999; van West et al., 2003). After entering the plant, Pythium spp. consume nutrients from the living host tissues. When nutrients are depleted, they secrete toxins or cell wall degrading to kill the host-cells and feed on the death tissues (Chérif et al., 1991; Mojdehi et al., 1991). However, the infectivity of the propagules (Sporangium, hypha and appressorium) is dependent on the age of the host plant, soil moisture, soil pH, soil organic matter contents and soil temperature (Martin & Loper, 1999). The infectivity of in vitro produced propagules is lower than the infectivity of naturally produced propagules.
Fig. 1. Life cycle of a typical Pythium species that infect root (van West et al., 2003)
3. Root- knot nematode on Rice: Distribution and Yield Loss
Around 300 nematode species of 35 genera have been reported to infect rice, among which Meloidogyne spp. are considered as the number one plant- parasitic nematodes based on scientific and economic importance (Jones et al., 2013). Several species of Meloidogyne have been found in rice for example M. javanica has been found on upland rice in Egypt, Nigeria, Ivory Coast, Comoro Islands, and in Brazil ; M. arenaria has been found in Nigeria, Egypt, and South Africa and M. salasi has been reported in Panama and Costa Rica ,whereas M. oryzae has been found in Surinam (Bridge et al., 1990). M. incognita is reported in Ivory Coast, Costa Rica, Cuba, Egypt, Nigeria, South Africa and Japan (Bridge et al., 1990). All species are responsible for reduction of tillering, delayed maturation, and yield loss (Fortuner & Merny, 1979). Among the root-knot nematodes Meloidogyne graminicola is the most important species on rice. This species causes 17% to 32% yield losses in rice and has been found in India, Bangladesh, Burma, Thailand, Vietnam, Laos, China, Philippines, Nepal, South Africa, Colombia, Brazil and USA (Chantanao, 1962; Roy, 1973). In field studies, yield losses due to M. graminicola and M. incognita are as high as 70% (Bridge et al., 1990).
3.1 Host Finding of nematodes
In order to infect the plant, first of all nematodes have to localize their host plant. Their ability of host finding is stimulated by a number of factors remaining in the rhizosphere zone of the host plant. For example, the infective second stage juveniles (J2) are attracted to the host roots upon perception of gradients of attractants released from the roots (Karssen et al., 2013). Carbon dioxide has been reported as long distance attractant for plant-parasitic nematodes (Klingler, 1965; Prot, 1980; Robinson, 1995), whereas allelochemicals released from various parts of the roots as well as sites of previous penetration or injured root surfaces, or lateral root emergence zone, in the rhizosphere might act as short distance attractants of second stage juveniles (Curtis et al., 2009). Gradients of pH also act as a local attractant (Perry, 2005; Wang et al. 2009). On the other hand, 1M NaCl solution has been reported as repellent of Meloidogyne spp.(Jonathon et al, 2011). The behavior of host finding of M. incognita is stimulated by potassium nitrate and can act as both attractant and repellent depending on the concentration gradient (Hida et al., 2015).
Inorganic salts of K+, NH 4 + , Cs+, NO 3 − , and Cl− strongly repel the infective second-stage juvenile of Meloidogyne incognita. Some of these salts are beneficial to plant growth suggesting a novel means of plant protection against root-not nematode (Castro et.al., 1990).Temperature play an important role in the attraction of nematodes- Ditylenchus dipsaci. At 100C, more no. of nematodes are attracted than at 15 and 20o C. Similarly distance between root and nematode, plays an important role in the host finding and subsequent infection by Ditylenchus dipsaci for example at 12.5 mm distance more no. of Ditylenchus dipsaci are attracted than 25mm and 50 mm distance. The juveniles of M. hapla are strongly attracted to the root apical meristem and the region of elongation but very few no. of juveniles are attracted to the upper hypocotyl and the stem apical meristem of both resistant and susceptible seedlings of alfalfa plant (Griffin and Waite , 1971).
Around 1 to 2-mm apical region ensheathed by border cells strongly attract the second stage juvenile of M. incognita than the region of elongation. The chemotactic response of M. incognita is dependent on the species of host plant as well as combination of host plants for example no response in response to snap bean and alfalfa cv.Lahonton; attraction in response to pea and alfalfa cv. Thor; repulsion in response to alfalfa cv. Moapa 69 (Zhao et al., 2000).
Root exudates of eto3 (ethylene-overproducing mutant) of Arabidopsis attract more J2 of Heterodera schachtii than root exudates of the wild-type (Wubben et al. 2001).
More no. of Meloidogyne hapla are attracted to Arabidopsis roots exposed to an ethylene (ET)-synthesis inhibitor whereas ET-overproducing mutants attract less no. of nematodes. Seedlings with ET-insensitive mutant of Arabidopsis attract more no. of nematodes whereas mutations resulting in constitutive signaling attract less no. of nematodes. Similarly, Roots of Never ripe (Nr), an ET-insensitive tomato mutant attract more no. of nematodes than wild type, indicating that ET signaling modulate the attraction of Meloidogyne hapla to tomato plants (Fudali et al., 2013).
Upon attack by corn rootworm larvae (Diabrotica virgifera virgifera) the roots of maize plants, produce (E)-β-caryophyllene, to attract entomopathogenic nematodes. These entomopathogenic nematodes attack and kill the larvae of Diabrotica virgifera virgifera (Kollner, et al. 2008; Rasmann, et al., 2005).
Bacteria present in the rhizosphere attract both free living and plant-parasitic nematodes. Among bacterial strains, Burkholderia has been reported as the most attractive strain for Meloidogyne incognita. This bacterium favors the maximum aggregations of root-knot nematode – Meloidogyne incognita (Tahseen et al., 2014).
Arbuscular mycorrhizal fungi (AMF), Glomus mosseae continuously suppresses the penetration and subsequent life stage development of root-knot nematode Meloidogyne incognita in tomato (Vos et al., 2012). But we do not know the role of Pythium arrhenomanes on the host finding, penetration, and migration of Meloidogyne graminicola.
IAA binds to the surface cuticle, amphids and phasmids of M. incognita and this IAA acts as a signal that orientates the nematode to localize the root surface of the host and/or inside the root tissue to promote the nematode infection (Curtis, 2008).One hundred (100) μM auxin in the pluronic F-127 gel favors the attraction and aggregation of Aphelenchoides besseyi. In addition, the rice plant containing higher level of endogenous auxin promote the migration of this nematode and favors reproduction therein. On the other hand, rice plant containing insufficient auxin level in the pollen does not promote the migration and reproduction of Aphelenchoides besseyi . In addition rice plants treated with 2,3,5-triiodobenzoic acid (auxin transport inhibitor) attract fewer nematodes in the seeds whereas young panicles of rice treated with 1-naphthaleneacetic acid produce more seeds and attract more nematodes than untreated control. These results indicating that auxin might play an important role in the migration and reproduction of Aphelenchoides besseyi (Feng et.al., 2014). However, we do not know the role of auxin in the host finding of Meloidogyne graminicola so far.
Several methods have been reported to study the host finding behavior of plant-parasitic nematodes. Among them, pluronic F-127 is an useful medium for studying the host finding behaviour of root-knot nematodes (Wang et.al., 2009).In addition, agar sensory assay and pipette bulb assay has been reported as novel bioassays for the host finding of plant-parasitic nematodes (Fig. 1).
Figure 2 Layout of agar sensory assay (left) and pipette bulb assay(right) (Dalzell et al., 2011).
More recently, a very sophisticated method has been reported to study the chemotaxis assay of nematodes (Figure 3).
Figure 3 Schematic illustration of in-gel chemotaxis assay (Hida et al., 2015).
3.2 Life cycle and behavior of root-knot nematodes
After finding a host the J2s have to penetrate into the roots. To do so, J2s release a cocktail of enzymes to degrade the cell wall of the host plant. They usually penetrate at the root elongation zone where they cause physical damage through repeated thrusting by the stylet. After entering the root, the J2s migrate intercellularly to the root tip where they make a U-turn around the meristematic tissue to enter the stele. Immediately after that, they move upwards in the vascular cylinder towards the area of cell differentiation. There they secrete effectors into selected host cells to stimulate the transformation of around 2 to 12 vascular cells into large multinucleate feeding cells (so-called giant cells) (Gheysen & Mitchum, 2011; Jones et al., 2013; Kyndt et al., 2013). Giant cells are formed after repeated nuclear division without cell division. At the same time, root tissue around the nematode undergoes hyperplasia (abnormal increase in the number of individual cells) and hypertrophy (abnormal increase in the size of individual cells) to form a characteristic root gall. Galls are usually developed within 1 or 2 days after J2s have entered the root. The size of the galls depends on the number of J2 that enter the root, nematode species and host plant. After establishing feeding sites J2s become sedentary and undergo three moultings within the gall to become either adult males or females (Karssen et al., 2013).The adult male become vermiform, comes out from the roots, and searches for amphimictic females for sexual reproduction. Because of absence any functional stylet J3 and J4 stages are non-feeding. The adult females keep the giant cells alive while feeding on them, and as a result they swell and become pear-shaped. M. graminicola normally reproduces through facultative meiotic parthenogenesis (Dutta et al., 2012; Karssen et al., 2013). Adult females lay eggs in a protective gelatinous egg mass inside the galls or on the root surface. Inside the root of the host J2s hatch from the eggs and are capable of establishing a new feeding site within the same root (Bridge & Page, 1982). This adaptive behavior enables M. graminicola to survive and multiply within host tissue even under flooded conditions (Bridge et al., 2005). However, the newly hatched J2 can also go outside to infect other roots. Under favorable conditions, the eggs of M. graminicola can survive for at least one year.
Depending on temperature and flooding conditions M. graminicola completes its life cycle within 2-3 weeks (Bridge & Page, 1982; Bridge et al., 2005; Dabur et al., 2004; Fernandez et al., 2014; Win et al., 2013). Normally, the populations of M. graminicola decline rapidly after 4 months, but some egg masses can survive up to 14 months in waterlogged soil (MacGowan & Langdon, 1989). The populations of M. graminicola are also capable to survive in flooded soil for at least 5 months. During flooding conditions the J2s do not invade rice, but after draining out of excess water quickly invade the host roots. Twenty to 30% soil moisture and soil dryness at tillering and panicle initiation stage of rice are the favorable conditions for M. graminicola.
Figure 4 Disease cycle of Meloidogyne (Karssen et al., 2013)
3.3 Effect of Root-knot nematodes on physiology and morphology of rice
Root-knot nematodes induce giant cell formation by reprogramming the major morphological and physiological processes e.g. changes in cellular metabolism, cell wall architecture, cell cycle, protein synthesis, transport, signal transduction, cytoskeleton, hormone homeostasis, and epigenetic mechanisms of the root cells (Kyndt et al., 2014). During the infection period, the root-knot nematodes suppress the cell death and keep the giant cells alive. At the same time M. graminicola induce the expression of photosynthesis-related genes to produce chloroplast-like structures inside dark-grown giant cells. In addition, M. graminicola induces the strong upregulation of genes involved in chromatin remodeling, DNA methylation, small RNA formation, and histone modifications (Ji et.al., 2013).
Root-knot nematode infection in rice strongly upregulate the genes involved in ‘photosynthesis’, ‘DNA metabolic process’, ‘cellular component organization’, ‘generation of precursor metabolites and energy’, and ‘cell cycle’ in the giant cells at 14 dai, whereas downregulation of genes involved in ‘secondary metabolic process’, ‘response to stimulus’, and ‘cell death’ is observed at 14 dai. Genes involved in ‘hydrolase’ and ‘nuclease’ activity are generally upregulated. In most of the cases, similar expression patterns are observed both at 7dai and 14 dai but less strong induction of genes involved in ‘metabolic processes’, ‘structural molecule activity’, ‘organelle part’, ‘membrane-enclosed lumen’, and ‘macromolecular complex’ is observed at 14 dai than at 7 dai. A distinct modification is observed in the pathways of starch and sucrose metabolism, trehalose metabolism, tetrapyrrole synthesis, and the phenyl- propanoid , light reactions, flavonoid production, and cell-wall-related pathways in the giant cells (Ji et.al., 2013).
Root-knot nematode infection downregulate the transcription of genes coding for nicotianamine synthase, phytosiderophore biosynthesis; a glucan endo-1,3-β-glucosidase precursor; α- DOX2, synthesis of oxylipins; transcription factor WRKY71; four thionin-like peptides; flavonol synthase, and phenylalanine ammonia lyase activity, whereas strongly upregulate the expression of genes coding for starch synthase; Cullin-1; AP2-like ethylene- responsive transcription factor AINTEGUMENTA (regulator of growth and cell numbers during organogenesis); roothairless-1; and cell-cycle control (genes encoding cyclin-T1-1, cyclin-dependent kinase A-1, and cyclin-dependent kinase C-2) (Ji et.al., 2013).
The plant immune system is largely controlled by phytohormones. To suppress the immune system, M. graminicola or M. incognita strongly suppress the transcription of key immune regulatory genes in early developing galls of rice (Kyndt et al., 2012;Nahar et al., 2011; Nguyen et al., 2014)
The giant cells are characterized by dense cytoplasm, multiple enlarged nuclei, small vacuoles. In addition, proliferation of smooth endoplasmic reticulum, ribosomes, mitochondria, and plastids are also found in the giant cells (Gheysen and Fenoll 2002).
After successful infection by M. graminicola, hooked like gall formation is the typical below-ground symptom on the rice root tips (Bridge et al., 2005). However, several above-ground symptoms i.e. yellowing of plant, delayed maturation, are also visible (Chantanao, 1962; Roy, 1973). The severity of symptoms varies based on the number of penetrated and established nematodes within the root system (Hussey, 1985), time of infection, age of the plants, and the proportion between the number of nematodes and food resources supplied by the plants (Bird, 1974). M. graminicola adversely affects the plant growth both in seedbeds and in fields conditions. Due to high initial population of M. graminicola, seedling wilt as well as severe reduction in growth parameters are observed (Plowright & Bridge, 1990). M. graminicola caused reduction of the plant height, shoot and root weight of rice, tiller number and leaf area index (Bimpong et al., 2010). Nevertheless, Meloidogyne also adversely affects several physiological processes such as absorption as well as upward translocation of water and nutrient by the root system (Bridge et al., 2005).
4. Interaction between host and pathogen
In order to infect/parasitize the host plants successfully, pathogens have to overcome pre-existing structural defenses, like wax layers, thick and dense cell wall. Pathogens do this by exerting physical forces in combination with secreting chemical metabolites. In the meantime the host induces some structural and biochemical defenses in response to the pathogen attack, which also need to be overcome by the pathogen for successful infection. More specifically, at the very beginning of infection, the pathogens sometimes secrete some conserved molecules (pathogen-associated molecular pattern-PAMPs) to attack the host, the host detect this PAMPs by cell surface pattern recognition receptors and subsequently activate pattern triggered immunity (PTI). To overcome the PTI, pathogens adapt themselves by secreting effector protein inside the host cells to manipulate host-cell structure and function, more specifically to facilitate infection (virulence factors and toxins) or trigger defense reaction (avirulence factors and elicitors) or both (Huitema et al., 2004; Kamoun, 2007). In turn, the plant evolved a second layer of immune receptors leading to effector triggered immunity (ETI), where pathogen effectors can be recognized by resistance genes, leading to localized cell death. Then an evolutionary battle continues between pathogen evolving new effectors and plants gaining new resistance genes (Jones and Dangle, 2006).
Figure 5 Zigzag model of host pathogen interaction (Jones and Dangle, 2006).
Interestingly, pathogen effectors might function either in the extracellular spaces within plant tissues to degrade the cell wall and to inhibit the plant enzymes invading pathogens or within the cytoplasm of plant cells to exploit defense pathways of the host (Hardham & Cahill, 2010).
To prevent the invading pathogen, the host plant produces toxic oxygen radicals and systemic signaling compounds as well as activate defense genes leading to the development of structural barriers and production of other toxins.
4.1 Interaction between Oryza sativa and Pythium
Pythium spp. overcome the pre-existing structural defense of rice by direct penetration of the root tissue with appressorium-like structures (Van Buyten, 2013).They also secrete effectors to overcome plant defense. To degrade the cell wall of the host Pythium spp secrete a polysaccharide lyase enzyme for the degradation of the host cell wall (Adhikari et al., 2013). The secretomes of P.vexans are enriched with the transmembrane transport as well as sugar binding and sugar modification activities whereas glycosyl bonds hydrolase activity is mostly enriched in the secretomes of P. irregular (Adhikari et al., 2013).
During interaction plants produce hydrolytic enzymes like chitinases, glucanases, and proteases to attack Pythium spp. To counteract with these hydrolytic enzymes, P. ultimum produces 43 proteins/enzyme inhibitors, for example Agrin-like protein, cysteine-type peptidase, and four domain protease inhibitor (Levesque et al., 2010). P. ultimum and P. oligandrum secrete several enzymes such as lipin- acyltransferase, chologlycine hydrolase, triacylglycerol lipase into host cells to take up and utilize the host nutrients (Horner et al., 2012).
To facilitate cell death of the host plant, Pythium spp. produce 26 types of Crinklers- effectors with an amino terminal YXSL motif, which target the host nucleus (Torto et al., 2003; Win et al., 2007; Schornack et al., 2010; Horner et al., 2012; van Damme et al., 2012). P. ultimum and P. aphanidermatum produce Necrosis and Ethylene inducing peptide (NEP1) like proteins (NLPs), which induce necrosis and immunity related responses in the host plant (Levesque et al., 2010; Ottmann et al., 2009). Several necrotrophic members of Pythium spp. secrete apoplastic effectors (toxins) to facilitate the development of the pathogen by triggering host cell death during the necrotrophic phase (Stassen & Van den Ackerveken, 2011).
P. ultimum produces (3R, 5 Z)-( -)- hydroxy-5-dodecenoic acid (a phytotoxic compound) which causes black root disease in sugar beet. The phytotoxicity of this compound on rice seedlings has been examined, root and shoot growth of rice seedlings are inhibited to 52.0% and 31.0% respectively at 500 ppm concentration of this toxin. Even at 25 ppm, the toxin shows an 14.0% inhibition to the growth of the shoot and an 8.8% inhibition to the growth of the root (Ichihara et al., 1985). Similarly, Pythium arrhenomanes produced heterogeneous toxic metabolites in the culture filtrates and caused the general browning of root tissues of wheat seedlings (Mojdehi et al., 1990). Notably, a number of effectors (i.e. elicitin, cutinase, glycoside hydrolases, pectate lyase and peptidase inhibitors) are found in the secretions of P. arrhenomanes and reported to play important roles in plant parasitism (Adhikari et al., 2013).
4.2 Interaction between Oryza sativa and root-knot nematode
Similar to Pythium spp, plant-parasitic nematodes also secrete effectors from the pharyngeal glands, cuticle as well as amphidsand injected into plant tissue through the syringe like- stylet of the nematode. These effectors are needed to overcome plant defense as well as to facilitate penetration, migration and feeding site induction (Gheysen & Mitchum, 2011; Huang et al., 2003; Davis et al., 2004). Cell walls primarily consists of cellulose and hemicellulose (polymer of xylose, mannose, galactose, rhamnose, arabinose and other sugars). These molecules act as a barrier to penetration for plant- parasitic nematodes. Nematodes overcome this plant barrier by combination of stylet activity and secretion of effectors like cellulases from their pharyngeal glands (Smant et al., 1998; Dautova et al., 2001; De Meutter et al., 2001). The nematodes also secrete xylanase to degrade the hemicellulose in the cell wall of the host (Hussey et al., 2002). A cell wall binding protein is secreted by nematodes at the very beginning of nematode infection. To facilitate nematode parasitism this protein directly interacts with pectin methylesterase protein 3 (PME3) thereby activating and targeting this enzyme (Hewezi et al., 2008). For loosening cell walls, plant-parasitic nematode also secrete expansin to weaken the non-covalent interactions between cell wall components (Qin et al., 2004). For the intercellular migration through the host tissue plant-parasitic nematodes need to degrade the pectic polysaccharides embedded in the middle lamella between plant cells. They do it by secreting pectatelyases and polygalacturonases (Popeijus et al., 2000a; Jaubert et al., 2002). Several proteases effectors such as cathepsin-like proteinases, serine protease, cysteine proteinases have been found in root- knot and cyst nematodes and might be important to facilitate the degradation of plant proteins (Jacob et al., 2008).
During parasitism to suppress the basal defense of plant Meloidogyne incognita secretes Mi-CRT, a calreticulin (CRT) into the apoplasm of the infected tissues (Jaouannet et al., 2013).In the secretions of Meloidogyne, Globodera, Heterodera and Hirschmanniella; chorismate mutase (CM) has been reported (Lambert et al., 1999; Popeijus et al., 2000b; Bekal et al., 2003; Jones et al., 2003; Bauters et al., 2014). During parasitism chorismate mutase might be secreted into the host cell to alter the auxin balance and might take away chorismate from SA synthesis pathway to suppress plant defense response (Doyle & Lambert, 2003; Gheysen & Mitchum,2011).Root-knot nematodes have been shown to produce a Nodulation L (NODL) like proteins which might facilitate giant cell formation ( Gheysen and Jones 2006).
In response to nematode attack the host plant produces reactive oxygen species. To detoxify the full range of host reactive oxygen species (ROS) plant-parasitic nematodes produce a range of antioxidant proteins. For example Globodera rostochiensis secretes glutathione peroxidase in its hypodermis to metabolize H2O2 produced by the host cell (Jones et al., 2004). In secretions of G. rostochiensis and Meloidogyne incognita, superoxide dismutase has been found, which acts as a detoxifying agent for ROS (Robertson et al., 1999; Bellafiore et al., 2008).M. graminicola has been reported to secrete peroxiredoxin, glutathione peroxidase, and glutathione-S-transferase for the detoxification of reactive oxygen species (ROS).In addition, M. graminicola also secretes fatty acid and retinol-binding protein, annexin, chitinase, transthyretin-like protein, mitogen-activated protein (MAP-1), venom allergen-like protein, galectin, C-type lectin and 14-3-3 (Haegeman et.al., 2013)
5. Hormonal defense signaling in plants
Various types of phytohormones such as auxins, gibberellins (GA), cytokinins (CK), abscisic acid (ABA), jasmonates (JA), salicylic acid (SA), ethylene (ET), brassinosteroids (BR), and strigolactones are produced in the plants to regulate growth and development of plants as well as to play an important role in plant defense to both abiotic and biotic stress (Robert- Seilaniantz et al., 2007; Gomez-Roldan et al., 2008; Navarro et al., 2008; Umehara et al., 2008; Bari & Jones, 2009). These hormones play either positive or negative roles against various necrotrophic and biotrophic pathogens depending on the type of host-pathogen interaction (Robert-Seilaniantz et al., 2007; Bari & Jones, 2009). There is also extensive cross- talk among different hormonal pathways.
5.1 Auxin in Pythium -Oryza sativa- Root-knot nematode interaction
Nearly every aspect of plant growth and development is controlled by auxin. Auxin is produced in the plant from the precursor molecule tryptophan. Genes encoding the proteins, TAA ( Tryptophan Amino transferase) and YUC ( YUCCA Flavin monooxygenase) regulate the biosynthesis of auxin in a tryptophan dependent pathway. The enzymes peroxidases promote the oxidation of indole ring leading to the decarboxylation of the side chain which result in the loss of auxin activity. Moreover, most of the auxins remain in conjugated form. Through the (de)-activation of a set of genes, auxin modulate the development of plant. However, under low auxin concentration, the Aux/AUX repressors inactivate the transcription factors, ARF. In the presence of auxin, an auxin co-receptor of Aux/IAA and TIR1/AFB is formed. then polyubiquitination of Aux/IAA is occurred by SCF TIR1/AFB complex leading to the proteosomal degradation of repressor. The repression of the ARF transcription factor by the degradation of Aux/IAA repressor allow the auxin responsive gene transcription.
Xanthomonas oryzae pv oryzae infection in rice induces auxin signaling and triggers the expression of expansins to promote the loosening of cell wall thereby facilitating pathogen entry and allowing increased nutrient leakage (Ding et al., 2008). On the other hand, overexpression of GH 3-8 (an auxin responsive gene), resulting in less free auxin, makes the plant resistant to bacterial infection, independent of SA and JA signaling (Ding et al., 2008). Both OsIAA9, an auxin responsive gene, and expansin OsEXPB3, are upregulated in the rice roots infected with P. graminicola. Upregulation of the expansin might lead to increased flexibility of plant cell walls to facilitate systemic infection by Pythium spp (Van Buyten, 2013). Interestingly, Pythium spp produced auxin under in-vitro conditions (Rey et.al., 2001; Van Buyten, 2013). Auxin is also required as a trigger for initiation of giant cells (Hutangura et.al., 1999, Karssen et.al., 2013). Meloidogyne spp. regulate auxin distribution in the root to trigger giant cell initiation (Karczmarek et al., 2004). Root-knot nematode produce auxin conjugates that may interfere with the existing hormonal balance of the plant cell (de Meutter et al., 2003 and de Meutter et al., 2005).
Root knot nematodes activate cell cycle for giant cell initiation and auxin is considered as a key factor in controlling cell cycle progression in plants (Goverse et al., 2000). Auxin induces the transcription of the cyclin-dependent kinases (CDK) and the mitotic cyclins correlated with the expression of cdcaAt and cdc2Pet (Hemerly et al., 1993; Trehin et al., 1998). Upon nematode infection, local auxin balance is changed and auxin signaling is essential for the nematotode feeding site formation (Richardson and Price, 1984; Goverse et al., 1998a; Helder et al., 1998). Indole compounds are accumulated in galls formed by M. incognita, M. hapla and M. javanica (Balasubramanian and Rangarwami, 1962; Yu and Viglierchio, 1964; Setty and Wheeler, 1968; Viglierchio and Yu, 1968). Auxin-like substances are present in adult females of M. javanica (Bird, 1962) and in exudates of J-2 of M. incognita (Setty and Wheeler, 1968) and M. hapla (Yu and Vigliergio, 1964). On the other hand, M. hapla (Viglierchio and Yu, 1965) and M. javanica (Bird, 1966) have been reported to release an auxin-inactivating compound.
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