Perspective Cite This: J. Agric. Food Chem. 2018, 66, 8672−8678
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Management of Cyst and Root Knot Nematodes: A Chemical Ecology Perspective Baldwyn Torto,*,† Laura Cortada,‡ Lucy K. Murungi,§ Solveig Haukeland,† and Danny L. Coyne‡ †
International Centre of Insect Physiology and Ecology (icipe), P.O. Box 30772-00100, Nairobi, Kenya International Institute of Tropical Agriculture (IITA), P.O. Box 30772-00100, Nairobi, Kenya § Department of Horticulture, Jomo Kenyatta University of Agriculture and Technology (JKUAT), P.O. Box 62000-00200, Nairobi, Kenya J. Agric. Food Chem. 2018.66:8672-8678. Downloaded from pubs.acs.org by UNIV OF SOUTH DAKOTA on 08/29/18. For personal use only.
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ABSTRACT: Plant parasitic nematode infection of crops can be highly detrimental to agricultural production. Since the discovery that plant roots release chemicals that attract the infective stage of plant parasitic nematodes some 80 years ago, significant progress in identifying the signaling molecules has occurred only relatively recently. Here, we review the literature on chemical ecological studies of two major plant parasitic nematode groups: root knot nematodes in the genus Meloidogyne and cyst nematodes in the genus Globodera because of the negative impact their parasitism has on farming systems in Africa. We then highlight perspectives for future directions for their management. KEYWORDS: Globodera spp., Meloidogyne spp., semiochemicals, solanoeclepin A, volatile organic compounds
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INTRODUCTION Plant parasitic nematodes constitute some of the greatest threats to crop production. The two most important groups of plant parasitic nematodes are root knot nematodes (RKNs) (Meloidogyne spp.) and cyst nematodes (Globodera and Heterodera spp.). Virtually every cultivated crop is prone to RKN attack, with Meloidogyne incognita, being the most important crop pathogen globally.1 Cyst nematodes have a similar biology to RKNs with a distinct egg, juvenile stages (J1), (J2) the infective stage, (J3), (J4) and adult stages. However, cyst nematodes tend to be much more host specific and require host stimulus for egg hatching. The J2 of RKN and cyst nematodes react to stimuli from suitable host roots to attract and guide them to their food source. Previous work has shown that with increased human mobility, the accidental introduction of alien species to new environments will occur, as witnessed with the recent report of potato cyst nematode (PCN) in Kenya.2 Global warming will also likely facilitate the further distribution of tropical RKN and cyst species to regions where they are currently not present, as shown by the spread of the tropical RKN species M. incognita, M. javanica, and M. arenaria, which were observed among the most rapidly spreading pests globally.3 Given that many of the nematicides formerly relied upon to manage plant parasitic nematodes have been withdrawn from the market due to environmental concerns,4 there is even greater pressure to identify suitable and environmentally sensitive tools to effectively control RKN and cyst nematodes. In this regard, understanding and exploiting knowledge of the chemical ecology of RKN and cyst nematode and their interactions with crops for their management is critical.
species. The mediating chemicals are commonly referred to as “semiochemicals” (message-bearing chemicals). Semiochemicals are low to medium molecular weight volatile and nonvolatile organic compounds synthesized from various pathways and can be exploited for the management of pests including plant parasitic nematodes. During the previous half a century or so, we have witnessed the rapid development and sensitivity in chemical instrumentation, resulting in the discovery of a broad range of semiochemicals from a wide range of above- and below-ground (rhizosphere) interactions. Comparatively, most of the discoveries on semiochemicals have been made from above- as opposed to below-ground interactions because of the complexity of the interactions in the rhizosphere (Figure 1). To improve our knowledge on below-ground interactions, better techniques are needed to collect and identify the molecules, especially volatile organic compounds (VOCs). Chemo-ecological studies are typically driven by simple, reliable, and reproducible behavioral assays. For plant parasitic nematode−host plant interactions, two main types of behavioral assays are recognized: stylet thrusting and chemotaxis. In the stylet thrusting assay, measurements are made on the rate at which the nematode thrusts its stylet (microscopic spear-like structure) into a chemical stimulus obtained from the host root. Stylet thrusting has been successfully recorded in both water and Pluronic gel formulations (a synthetic block copolymer),5 suggesting that the nematode can move around freely in various liquid media and at the same time thrust its stylet to detect chemical stimuli. More recently, it was shown that electrophysiological recordings of the stylet can be made,6 demonstrating that some of the techniques that have provided
ADVANCES IN CHEMICAL ECOLOGY OF ROOT KNOT AND CYST NEMATODES Chemical ecology is the study of the role of natural chemical interactions between organisms of the same or different
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© 2018 American Chemical Society
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April 13, 2018 July 11, 2018 July 24, 2018 July 24, 2018 DOI: 10.1021/acs.jafc.8b01940 J. Agric. Food Chem. 2018, 66, 8672−8678
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Figure 1. Physico-chemical, physiological, and microbiological factors influencing populations’ dynamics of plant parasitic nematodes at the rhizosphere level. Plant roots produce semiochemical molecules (volatile and nonvolatile) as defense cues to repel nematodes and minimize root infestation (blue-dotted arrow-shaped icon). Conversely, some of these root allelochemicals are used by plant parasitic nematodes as indicators of the presence of a suitable host, and once these are detected they trigger a cascade of hatching mechanisms in the nematode that end up in the releasing of infective juveniles into the soil to infest the plant root system (red solid arrow-shaped icon); additionally, a few cyst nematodes species (i.e., PCN) can maximize their chances of persisting in the soil until a suitable host is planted by reducing spontaneous hatching through inner inhibiting hatching mechanisms (i.e., proteases and proteases inhibitors) hosted in the cuticle of the cysts (green- dotted arrow-shaped icon). Population dynamics of root-knot nematodes (RKN) and potato cyst nematodes (PCN) are further influenced by the presence of microbiological control agents in the soil that act either as obligate parasites or as antagonist, both with the ability of reducing the incidence of parasitic nematodes in the soil under adequate environmental conditions.
disadvantage of this method is that tissue material crushed at even low temperatures can lead to production of artifacts in the volatile emissions associated with mechanical damage. Using such methods would need to be backed by direct sampling of the root emitted volatiles into the sand. Our research shows that pulling air from the sand through SPME fibers enclosed in glass tubes collects predominantly sand-associated volatiles. To overcome the masking of root volatiles with sand-associated volatiles, other researchers have successfully developed and used a direct sampling method for roots in the field or for uprooted plants in the laboratory using a probe inserted directly into the sand. Air is then pulled through the probe and then an adsorbent attached at the end of the probe.12,14,15 The advantage of this method is that it allows for comparison of laboratory-collected volatiles from uprooted plants and field roots to eliminate artifacts associated with uprooted plants. It also allows for continuous sampling of root volatiles. Analysis of root secretions mainly consists of two approaches depending upon whether the target secretion is VOCs or nonvolatiles. The most common method used for the analysis of VOCs is coupled gas chromatography−mass spectrometry (GC−MS). The second approach is analysis of nonvolatile compounds by coupled high-performance liquid chromatography−time-of-flight-mass spectrometry (LC−QTOF-MS). In the analysis by GC−MS, some researchers have argued that solvent extracted volatiles from inert adsorbents do not reflect the complete natural volatile profile since low molecular weight compounds (C-2 to C-4 derivatives) such as acetaldehyde,
breakthroughs to identifying semiochemicals associated with insect olfaction are applicable to plant parasitic nematodes. However, current electrophysiological techniques developed for plant parasitic nematodes are yet to be optimized to isolate physiologically active compounds. Although stylet thrusting indicates detection of a chemical stimulus, using this technique does not allow for full classification of the chemical stimulus as an attractant (stimulant) or repellent (deterrent).7 On the other hand, using chemotaxis (directional responses of nematodes to chemical stimuli) assays, allows for a full description of the stimulus as an attractant or repellent. Previously chemotaxis assays were screened in 1% agar discs,8−10 but recent successes using sand-filled dual choice olfactometer11−15 have allowed for testing nematode directional responses to intact plant root secretions, extracts, and synthetic compounds identified from these root secretions. Another reason why studies on below-ground interactions lag above-ground interactions is the complexity involved in collecting and identifying the mediating molecules from the roots in the sand. To collect volatile organic compounds emitted by plant roots, inert polymer adsorbents including solid phase microextraction (SPME) fibers, Super Q, Porapak Q, HayeSepQ, which have been used previously to collect odors for various interactions (e.g., plant−plant, plant−insect, insect−insect, plant−microbe) are used in various sampling techniques. For example, some researchers have used SPME fibers to collect volatiles from excised plant roots crushed at very low temperatures such as in liquid nitrogen.11 The 8673
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Recently, it has been shown that small lipophilic molecules extracted in tomato (Solanum lycopersicum) and rice (Oryza sativa) root exudates had a nematotoxic or nematostatic effect on M. incognita and M. graminicola.21 Whether these small lipophilic molecules constitute downstream signaling components is yet to be established. More recent studies have shown that M. incognita is attracted to volatile organic compounds released by roots of different cultivars of the solanaceous plant pepper (Capsicum annum) (Table 1), such as methyl salicylate,
ethyl acetate, propanal and butanal may not be captured in the adsorbed volatiles. An alternative approach suggested is sampling on an adsorbent such as Tenax TA which can be thermally desorbed in the GC-MS system. Although thermal desorption is less complicated, it has the disadvantage of degrading sensitive samples.16 However, using specific behavioral assays combined with these analytical methods, have revealed that parasitic nematode chemical ecology is dependent on several factors including nematode species, root part (zone of elongation of growing roots), plant species, pH, concentration of chemical in the root exudate and interactions with other rhizosphere organisms especially microbes. In the following sections, we will provide highlights on chemical ecological studies of two of the most important plant parasitic nematode groups; RKN in the genus Meloidogyne and cyst nematodes in the genus Globodera because of the detrimental impact of their parasitism on agricultural productivity, and then highlight perspectives for future directions. Chemical Ecology of Meloidogyne spp.-Host Plant Interactions. In nematode-plant interactions, the discovery that chemicals associated with plant roots attracted infective stage juveniles of parasitic nematodes was first made in 1939.17 Subsequent experiments carried out after this discovery occurred almost two decades later, focusing on excised plant roots and interactions with infective juvenile stages of various parasitic nematode species from the genus Meloidogyne.18,19 Although these follow up studies improved our understanding of nematode−plant interactions, they failed to identify the signaling molecules mediating interactions. Since then, several signaling molecules have been identified to show that Meloidogyne−host plant interactions are complex, involving diverse molecules derived from different chemical classes. Some of these chemicals may serve as long- or short-range signaling molecules depending upon their volatility, interaction with sand particles (adsorption and desorption) polarity, and solubility in water. The simplest chemical identified is carbon dioxide, considered a long-range attractant,18 perhaps because previous work demonstrated that parasitic nematodes congregated around anaerobic surfaces associated with release of carbon dioxide. However, carbon dioxide is considered a nonspecific signal because of the wide range of sources (decomposing plant, metabolism of root mutualistic microbes, etc.) that emit it in the rhizosphere and which when in solution can lead to acidic gradients that attract or repel parasitic nematodes.5 Assays carried out in a 23% aqueous solution of pluronic gel confirmed the attraction of M. hapla to acidic gradients ranging between pH of 4.5 and 5.4 formed by acetic acids,5 suggesting that knowledge of the salinity of the medium in which the root is developing is critical to an understanding of Meloidogyne responses to root secretions. Evidently, the complexity of the interactions with host plant roots found by other researchers and in our group, is because of the responsiveness shown by different Meloidogyne species to a wide range of volatile and nonvolatile organic compounds, including phytohormones, terpenoids, esters and phenols.15 To illustrate this complexity, the signaling pathway of the phytohormone ethylene was shown to modulate the attractiveness of host roots to M. hapla.20 However, components of the ethylene-signaling pathway directly affecting root length did not appear to be responsible for modulating root attractiveness; instead other unidentified components of downstream signaling modulated the responses of M. hapla.
Table 1. Compounds Identified in the Volatiles Emitted by Host Plant Roots of the Root Knot Nematode Meloidogyne incognita cmpd
source
α-pinene camphene β-pinene δ-3-carene 6-methyl-5-hepten-2-one myrcene decane limonene (Z)-β-ocimene p-cymene undecane camphor sabinene 2-isopropyl-3-methoxypyrazine 2-methoxy-3-(1-methylpropyl)-pyrazine dodecane methyl salicylate thymol tridecane tetradecane geosmin α-cedrene β-cedrene γ-himachalene allo-aromadendrene α-muurolene 4,5-diepi-aristolochene γ-gurjunene
pepper, tomato, spinach tomato tomato, spinach tomato tomato tomato pepper pepper, tomato, spinach pepper, tomato pepper pepper pepper tomato tomato, spinach pepper, tomato, spinach pepper pepper, tomato pepper pepper, tomato pepper spinach tomato, spinach tomato, spinach pepper pepper pepper pepper pepper
α-pinene, limonene, and tridecane,15 with subtractive assays demonstrating that the aromatic ester methyl salicylate was the most potent attractant in the blend of compounds identified. This study also showed that the host finding process in M. incognita can be disrupted by thymol, released in the VOCs of the root of another cultivar of pepper.15 These findings mark a significant step in the process of developing novel eco-friendly strategies for control of M. incognita via plant breeding, whereby specific molecular pathways for suppressing root production of methyl salicylate or incorporating genes responsible for thymol production in the roots of pepper can be explored to protect pepper from RKN infection. Similar studies in our laboratory have demonstrated the importance of plant species in altering RKN responses to root VOCs using tomato and spinach, (Spinacea oleracea; Amaranthaceae) (Table 1).22 A recent study showed that tomato root-secreted nonvolatiles including phenol 2, 6-ditert-butyl-p-cresol, and esters L-ascorbyl 2, 6-dipalmitate, dibutyl phthalate, and dimethyl 8674
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Journal of Agricultural and Food Chemistry phthalate influenced the behavior of J2s differentially.23 Among these compounds, only dibutyl phthalate repelled J2s. These compounds may be artifacts because they are typical contaminants detected in samples analyzed by GC−MS. Chemical Ecology of Cyst Nematode−Host Plant Interactions. Potato cyst nematodes (PCN), comprising two species of Globodera (G. pallida and G. rostochiensis) represent quarantine pests in over 100 countries24 and are considered the primary potato pests globally. Cysts of PCN may contain ∼500−700 eggs and can persist in the soil for more than 20 years, withstanding extreme temperatures (−15 °C) and/or prolonged desiccation.25 The reactivation of eggs within the cysts is triggered by the presence of a suitable host crop, primarily potato (Solanum tuberosum) but also to a lesser extent other Solanaceae species such as tomato (Solanum lycopersicum), eggplant (Solanum melongena), or Capsicum spp. Root secretions from several plant species have been identified as the triggering factors for the hatching of diverse cyst nematode species. Some of these root effectors are responsible for prompting changes in the permeability of the PCN three-layer egg’s membrane. For instance, calcium ions which diffuse into the soil activate osmotic changes in the membrane to release trehalose. This allows water to be absorbed through the egg’s membrane to hydrate it, which in turn ceases the diapause in the cyst, leading to hatching and emergence of metabolically active J2s. The J2s then locate their hosts by detection of root secretions including volatile organic compounds.26 Interestingly, several root secreted metabolites have been identified as stimulating PCN hatching, but solanoeclepin A, a tetranortriterpene derived from gonanane27 present in the root exudates of potato and tomato roots is the most studied.28 It shows nanomolar activity as a PCN-hatching factor, indicating the sensitivity of the hatching process in response to the appropriate chemical stimulus. Related studies have also identified hatching factors including glycinoeclepin A in soybean (Glycine max) and kidney bean (Phaseolus vulgaris) as well as glycinoeclepin B and C in the latter species, for the soybean cyst nematode Heterodera glycines.29 These three compounds stimulating hatching in cyst nematodes are triterpenoids sharing a common biosynthetic origin30 like solanoeclepin A. Structurally, because they are highly oxygenated and will bind to water molecules, low concentrations of them can be absorbed through the egg’s membrane to trigger hatching. Because plant roots also release volatile organic compounds, their role in PCN J2 host location requires investigation. Additional molecules that play an important role in the plant-nematode recognition signaling are the hatching factor stimulants: their presence in the soil enhances and/or inhibits hatching through chemical interaction with the already existing root secretions available in the soil. The glycoalkaloids αchaconine and α-solanine are both able to stimulate J2 hatching of Globodera spp.;31 the variable hatching stimulus of these two molecules32 is linked to their concentration in the rhizosphere, which seems to vary with the age of the plant; for instance, α-solanine is present at higher concentrations in younger plants. Similarly, these two glycoalkaloids are also highly oxygenated, thereby potentially using a similar mechanism to that of solanoeclepin A, glycinoeclepin A, glycinoeclepin B and C to trigger hatching in PCNs. Hatching of cyst nematodes is triggered by highly specific root exudate cues, but it is additionally controlled by hatching inhibitors including proteases and protease inhibitors that have been
identified from the cyst wall.33 These cyst inner inhibitors are influenced by the rhizosphere composition and appear in higher concentrations during unfavorable periods for cysts to be persistent in the soil until a suitable host crop is planted.33,34 Such hatching inhibitors have been found in both the plant and cyst cuticles,7 and therefore these have been identified as an additional nematode-resistance mechanism that prevents cysts from hatching under suboptimal conditions. This points toward a strong adaptation by cyst nematodes to their hosts through the chemical-dialogue established at the rhizosphere level.35 While disruption of the diapause stage of PCN is highly dependent on the specific presence of host root secretions in the rhizosphere, some cyst nematode species, such as the soybean cyst nematode hatch more spontaneously. Therefore, use of its natural hatching effectors has been highlighted as a sustainable method to manage PCN in the absence of a suitable host. Over the years, these hatching factors (i.e., asparagusic acid) have been proposed to be developed into nematicides.33,36 However, the complexity in the structures of these molecules have precluded their large-scale synthetic production. Instead, less complex oxygenated synthetic hatching effectors have been developed, such as metavanadate and picrolonic acid, with the latter being effective only with G. rostochiensis.31 In the absence of a commercial product derived from the identified hatching factors, a strategy currently being explored to manage PCN is the disruption of the chemical signaling between the potato root secretions and the J2s. Recently, the use of dead-end trap crops to suppress PCN in soil has been attempted with varying degrees of success using Solanaceae plants, for example, the sticky nightshade (Solanum sysimbriifolium).37 Using trap crops, PCN are stimulated to hatch by nightshade root exudates but are then unable to develop on the resistant plant thus acting as a dead-end trap crop. In a previous study,38 90 accessions of Solanaceae (nontuber bearing) were screened for their hatching stimulatory effect on PCN as well as for resistance. All these Solanum species induced hatching of PCN and was highest for the S. nigrum complex. The hatching stimulatory effects of these solanaceous plants may be associated with a similar class of compounds previously identified in the root exudates of tomato and potato. Sticky nightshade and two cultivars of S. nigrum induced high levels of hatching as well as providing resistance against PCN and which were considered good candidates as trap crops. Interestingly, other researchers39 have demonstrated the effect and potential of some non-Solanaceae crops, such as lupine (Lupinus mutabilis (Fabaceae)) and Oxalis tuberosa (Oxalidaceae) as trap crops for PCN. As a result, lupine is used as a green manure crop by Bolivian potato farmers to mitigate against PCN attack. The underlying mechanisms leading to lupine to serve as a trap crop needs investigating. In Kenya, where PCN is a recent introduction,2 ongoing studies in our laboratory indicate that at least two species of the African indigenous vegetables (AIVs), African nightshade (Solanum scabrum and S. vilosum), act as potential trap crops. Our field studies show that after three seasons of planting these nightshade species, the PCN population density declined by 80%. Given these findings, elucidation of the chemicals involved and degree of hatching they cause (vital for success as a dead-end trap crop) and level of resistance are necessary. Furthermore, the sequence of cropping, in terms of restimulation for PCN hatching, should also be studied. 8675
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hatching and localization of the feeding site by J2s,50 although the identity and proportion in each of these fractions contributing to bioactivity is yet to be determined. Some studies have demonstrated the involvement of plant hormonal signaling in PCN host plant interaction. For example, salicylic acid (SA)-deficient mutants were found to be more susceptible to the beet cyst nematode, H. schachtii, than nonmutants, suggesting that an intact salicylic (SA) pathway is required for the defense mechanisms of sugar beet.51 Similar findings have been reported with tomato for jasmonic acid, which requires an intact signaling pathway to retain the Mi-1 mediated resistance response against RKN. These findings suggest that more studies are needed to elucidate the chemical ecology of the interactions between these plant parasitic nematodes and their host plants.
Thus, the importance of these nightshade species as nutritious food crops and for income (unlike S. sisymbriifolium, which is a weed) is an added value within such a cropping system for African smallholder farmers. Screening of other AIVs as trap crops for PCN management therefore provides potential value and may result in identifying better trap crops as well as new hatching factors. Regarding the effects of potential nonsolanaceous crops as trap crops or inhibitors of PCN hatching, much work remains on the chemical ecology and interactions with PCN. The management of plant parasitic nematodes in biofumigated and amended soils is attributed to a combination of different mechanisms, such as (i) the enhancement of the production of nematicide chemical molecules at the rhizosphere level; (ii) the increase/introduction of antagonist microorganisms in the soil; and/or (iii) boosting of plant defense mechanisms.40 Neem (Azadirachta indica) and several species from the Asteraceae family (Tagetes spp., Artemisia dracunculus, Chrysanthemum spp., Calendula spp., and Crotolaria spp.) have been successfully used to amend soils and decrease nematode populations due to their ability to release pre-existing nematicidal phytochemicals, mainly limonoids, sesquiterpene lactones, and polythienyls (especially αterthienyl) into the rhizosphere; a few of these species have been used commercially, although a high variability on the efficacy of the biofumigation process has been reported, depending on the plant species used or on the soil type.40−42 Botanical species of the Brassicaceae family are rich in glucosinolates, which transform into isothiocyanates upon degradation of plant tissues and are also used for their nematicidal effects in biofumigation programs.43 The incorporation of Brassica species as green manures has been reported as a potential management option for the control of PCN, RKN, and other plant parasitic nematodes.42,44 Studies show that the precursors of 2-propenyl isothiocyanate excreted by mustard (Brassica juncea), both in liquid and in gas phase, can successfully inhibit G. pallida hatching (50% decline) in vitro, although it was less successful under field conditions.45 Despite the mechanisms linked to PCN hatching inhibition being unclearly elucidated, it has been observed that 2-propenyl isothiocyanate triggers the expression of heat-shock proteins in the free living nematode Caenorhabditis elegans; the overexpression of such proteins has been linked to an acceleration of the metabolic activities of J1 within the egg and the subsequent depletion of their energetic reserves that leads to its death and/or prevents J2s from infecting its hosts. It has been proposed that soil micronutrient composition, specifically sulfur, could also have an impact on the efficiency of the metabolism of the glucosinolate production by Brassica species and should be further investigated. The degradation of soil organic acids into short-chain fatty acids under anoxic conditions has also been reported to have a suppressive effect on PCN.46,47 As shown in Figure 1, plant hormonal signaling pathway can also influence RKN and cyst nematode host finding. For instance, an active ethylene (ET) signaling pathway has been reported to repel M. hapla20 and the soybean cyst nematode H. glycines but attracts the sugar beet cyst nematode (Heterodera schachtii),48 while Arabidopsis mutants overexpressing ethylene were found to be more susceptible to the cyst nematodes.49 For G. pallida, both volatile and nonvolatile compounds have been reported to play a role in plant-nematode signaling: the water-soluble components combined with volatile cues elicit
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FUTURE PERSPECTIVES What we know from the review of the chemical ecology of the two genera Meloidogyne and Globodera is that most studies have exclusively focused on their interaction with roots of crops, which is a binary system. However, crop roots typically have a close association with mutualistic rhizosphere microorganisms. Therefore, what we have yet to establish is how these mutualistic associations influence the chemical ecology of plant parasitic nematodes and how such knowledge can be exploited for RKN and cyst nematode management. The rhizosphere is characterized by a complexity of interactions (Figure 1); for instance, microbial root mutualists may influence plant fitness by altering the composition and concentration of secondary metabolites and nutrients. Similarly, plant chemistry may also influence microbial root mutualist fitness. For instance, a trap crop or biotransformed crop may be effective against plant parasitic nematodes due to the secondary metabolites of the plant, or alternatively the composition of microbial root mutualists, or even both. As such, it would serve a useful exercise to identify beneficial microbial root mutualists for a specific crop or cropping system and investigate their influence on the chemical signaling in the rhizosphere, via studies such as microbe−microbe, microbe− plant, plant−plant, and microbe−plant−nematode interactions. Also, the interaction between microbial root mutualists and nematodes may influence, or even determine, the fitness of the nematode to locate host roots. Understanding the chemical communication of such microbe−nematode interactions may be critical to developing effective nematode management methods. Furthermore, studies on the effects that these interactions have on plant parasitic nematode natural enemies in the rhizosphere are also warranted. Understanding these interactions would provide opportunities for developing methods to enhance and exploit the use of natural enemies in nematode control. Since genetic pathways may drive these binary and tritrophic interactions, some researchers have investigated the expression of genes and demonstrated that certain genes are either up- or down-regulated due to nematode infection in certain crops. These studies have also exclusively focused on the binary system: nematode−plant interaction. Therefore, knowledge of the genes that are expressed due to the microbial root mutualist associations and understanding how they influence the chemical ecology of nematode−host interactions is required. Such knowledge can be exploited for use in the biotransformation of crops to mitigate against nematode attack. 8676
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thogenic nematodes by insect-damaged maize roots. Nature 2005, 434, 732−737. (12) Ali, J. G.; Alborn, H. T.; Stelinski, L. L. Subterranean herbivoreinduced volatiles released by citrus roots upon feeding by Diaprepes abbreviatus recruit entomopathogenic nematodes. J. Chem. Ecol. 2010, 36, 361−368. (13) Ali, J. G.; Alborn, H. T.; Stelinski, L. L. Constitutive and induced subterranean plant volatiles attract both entomopathogenic and plant parasitic nematodes. J. Ecol. 2011, 99, 26−35. (14) Ali, J. G.; Alborn, H. T.; Campos-Herrera, R.; Kaplan, F.; Duncan, L. W.; Rodríguez-Saona, C.; Koppenhofer, A. M.; Stelinski, L. L. Subterranean, herbivore-induced plant volatile increases biological control activity of multiple beneficial nematode species in distinct habitats. PLoS One 2012, 7 (6), e38146. (15) Kihika, R.; Murungi, L. K.; Coyne, D.; Ng’ang’a, M.; Hassanali, A.; Teal, P. E. A.; Torto, B. Parasitic nematode Meloidogyne incognita interactions with different Capsicum annum cultivars reveal the chemical constituents modulating root herbivory. Sci. Rep. 2017, 7, 2903. (16) Agelopoulos, N. G.; Pickett, J. A. Headspace analysis in chemical ecology: effects of different sampling methods on ratios of volatile compounds present in headspace samples. J. Chem. Ecol. 1998, 24, 1161−1172. (17) Linford, B. M. Attractiveness of roots and excised shoot tissues to certain nematodes. Proc. Helm. Soc. Wash. 1939, 6, 11−18. (18) Bird, A. F. The attractiveness of roots of plants to the plant parasitic nematodes Meloidogyne javanica and M. hapla. Nematologica 1959, 4, 322−335. (19) Bird, A. F. Additional notes on the attractiveness of roots to plant parasitic nematodes. Nematologica 1960, 5, 217. (20) Fudali, S. L.; Wang, C.; Williamson, V. M. Ethylene signaling pathway modulates attractiveness of host roots to the root-knot nematode Meloidogyne hapla. Mol. Plant-Microbe Interact. 2013, 26, 75−86. (21) Dutta, T. K.; Powers, S. J.; Hari, S. J.; Gaur, H. S.; Birkett, M.; Curtis, R. H. C. Effect of small lipophilic molecules in tomato and rice root exudates on the behavior of Meloidogyne incognita and M. graminicola. Nematology 2012, 14, 309−320. (22) Murungi, L. K.; Kirwa, H.; Coyne, D.; Teal, P. E. A.; Beck, J. J.; Torto, B. Identification of key root volatiles signaling preference of tomato over spinach by the root knot nematode Meloidogyne incognita. J. Agric. Food Chem. 2018, 66, 7328. (23) Yang, G.; Zhou, B.; Zhang, X.; Zhang, Z.; Wu, Y.; Zhang, Y.; Lü, S.; Zou, Q.; Gao, Y.; Teng, L. Effects of tomato root exudates on Meloidogyne incognita. PLoS One 2016, 11, e0154675. (24) PM 7/40 (3) Globodera rostochiensis and Globodera pallida. Bull. OEPP 2013, 43, 119−138, . (25) Hunt, D. J.; Luc, M.; Manzanilla-López, R. H. Identification, morphology and biology of plant parasitic nematodes. In Plant Parasitic Nematodes in Subtropical Agriculture; Luc, M., Sikora, R. A., Bridge, J. B., Eds.; CABI Publishing: Wallingford, Oxfordshire, U.K., 2005; pp 1−917. (26) Farnier, K.; Bengtsson, M.; Becher, P. G.; Witzell, J.; Witzgall, P.; Manduríc, S. Novel bioassay demonstrates attraction of the white potato cyst nematode Globodera pallida (Stone) to non-volatile and volatile host plant cues. J. Chem. Ecol. 2012, 38, 795−801. (27) Schenk, H.; Driessen, R. A. J.; de Gelder, R.; Goubitz, K.; Nieboer, H.; Brü ggemann-Rotgans, I. E.M.; Diepenhorst, P. Elucidation of the structure of solanoeclepin A, a natural hatching factor of potato and tomato cyst nematodes, by single-crystal X-ray diffraction. Croat. Chem. Acta 1999, 72, 593−606. (28) Kleinnijenhuis, R. A.; Timmer, B. J. J.; Lutteke, G.; Smits, J. M. M.; De Gelder, R.; Van Maarseveen, J. H.; Hiemstra, H. Formal synthesis of solanoeclepin A: enantioselective allene diboration and intramolecular [2 + 2] photocycloaddition for the construction of the tricyclic core. Chem. - Eur. J. 2016, 22, 1266−1269. (29) Masamune, T.; Fukuzawa, A.; Furusaki, A.; Ikura, M.; Matsue, H.; Kaneko, T.; Abiko, A.; Sakamoto, N.; Tanimoto, N.; Murai, A. Glycinoeclepins, natural hatching stimuli for the soybean cyst
To conclude, sedentary RKN and cyst nematodes pose a serious threat to crop productivity across the globe but especially in the tropics and SSA.52 Given the apparent importance of chemical signaling between nematode pest and crop host roots for host recognition, host finding, and location there appears a whole gamut of potential avenues to explore for exploiting this toward nematode pest management in small holder farming systems.
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AUTHOR INFORMATION
Corresponding Author
*Phone: +254-20-2000. Fax: +254-20-2001. E-mail: btorto@ icipe.org. ORCID
Baldwyn Torto: 0000-0002-5080-9903 Funding
We gratefully acknowledge the financial support for this research by the following organizations and agencies: Swedish International Development Cooperation Agency (Sida), U.K.’s Department for International Development (DFID), the Swiss Agency for Development and Cooperation (SDC), the Austrian Development Agency (ADA), USAID, and the Kenyan Government. The views expressed herein do not necessarily reflect the official opinion of the donors. Notes
The authors declare no competing financial interest.
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REFERENCES
(1) Trudgill, D. L.; Blok, V. C. Apomictic, polyphagous root-knot nematodes: exceptionally successful and damaging biotrophic root pathogens. Annu. Rev. Phytopathol. 2001, 39, 53−77. (2) Mwangi, J. M.; Kariuki, G. M.; Waceke, J. W.; Grundler, F. M. First report of Globodera rostochiensis infesting potatoes in Kenya. New Dis. Rep. 2015, 31, 18. (3) Bebber, D. P.; Holmes, T.; Gurr, S. J. The global spread of crop pests and pathogens. Global Ecol Biogeogr. 2014, 23, 1398−1407. (4) United Nations Environmental Program (UNEP). Montreal Protocol on Substances that Deplete the Ozone Layer, UNEP Technology and Economic Assessment Panel, April 2000 Report, http://www.teap.org (Accessed February 25, 2018). (5) Wang, C.; Bruening, G.; Williamson, G. M. Determination of preferred pH for Root-Knot Nematode Aggregation using Pluronic F127 Gel. J. Chem. Ecol. 2009, 35, 1242−1251. (6) Hu, C.; Kearn, J.; Urwin, P.; Lilley, C.; O’Connor, V.; HoldenDye, L.; Morgan, H. StyletChip: A microfluidic device for recording host invasion behavior and feeding of plant parasitic nematodes. Lab Chip 2014, 14, 2447−2455. (7) Reynolds, A. M.; Dutta, T. K.; Curtis, R. H. C.; Powers, S. J.; Gaur, H. S.; Kerry, B. R. Chemotaxis can take plant-parasitic nematodes to the source of a chemo-attractant via the shortest possible routes. J. R. Soc., Interface 2011, 8, 568−577. (8) Wieser, W. The attractiveness of plants to larvae of root-knot nematodes. 1. The effect of tomato seedlings and excised roots on Meloiodogyne hapla Chitwood. Proc. Helm. Soc. Wash. 1955, 22, 106− 112. (9) Wieser, W. The attractiveness of plants to larvae of root-knot nematodes. 2. The effect of excised bean, eggplant and soybean roots on Meloiodogyne hapla Chitwood. Proc. Helm. Soc. Wash. 1956, 23, 59−64. (10) Wallace, H. R. Observations on the emergence from cyst and the orientation of larvae of three species of the genus Heterodera in the presence of host plant roots. Nematologica 1958, 3, 236−243. (11) Rasmann, S.; Köllner, T. G.; Degenhardt, J.; Hiltpold, I.; Toepfer, S.; Kuhlmann, U.; Turlings, T. Recruitment of entomopa8677
DOI: 10.1021/acs.jafc.8b01940 J. Agric. Food Chem. 2018, 66, 8672−8678
Perspective
Journal of Agricultural and Food Chemistry nematode, Heterodera glycines. II Structural Elucidation. Bull. Chem. Soc. Jpn. 1987, 60, 1001−1014. (30) van Dam, N. M.; Bouwmeester, H. J. Metabolomics in the rhizosphere: tapping into belowground chemical communication. Trends Plant Sci. 2016, 21, 256−265. (31) Clarke, A. J.; Shepherd, A. M. Picrolonic acid as a hatching agent for the potato cyst nematode, Heterodera rostochiensis woll. Nature 1966, 211, 546. (32) Byrne, J.; Twomey, U.; Maher, N.; Devine, K. J.; Jones, P. W. Detection of hatching inhibitors and hatching factor stimulants for golden potato cyst nematode, Globodera rostochiensis, in potato root leachate. Ann. Appl. Biol. 1998, 132, 463−472. (33) Masler, E. P.; Chitwood, D. J. Evaluation of proteases and protease inhibitors in Heterodera glycines cysts obtained from laboratory and field populations. Nematology 2017, 19, 109−120. (34) Okada, T. Hatching inhibitor factors in the cysts contents of the soybean cyst nematode, Heterodera glycines ICHINOE (Tylenchida: Heteroderidae). Appl. Entomol. Zool. 1972, 7, 99−102. (35) Hirsch, A.; Bauer, W. D.; Bird, D. M.; Cullimore, J.; Tyler, B.; Yoder, J. Molecular signals and receptors: controlling rhizosphere interactions between plants and other organisms. Ecology 2003, 84, 858−868. (36) Bohlmann, S. Introductory Chapter on the basic biology of cyst nematodes. In Plant Nematode Interactions: A View on Compatible Interrelationships; Escobar, C., Fenoll, C., Eds.; Elsevier: San Diego, CA, 2015; pp 1−421. (37) Belair, G.; Dauphinais, N.; Mimee, B. Evaluation of cultural methods for the management of the golden nematode (Globodera rostochiensis) in Quebec, Canada. Can. J. Plant Pathol. 2016, 38, 209− 217. (38) Scholte, K. Screening of non-tuber bearing Solanaceae for resistance to and induction of juvenile hatch of potato cyst nematodes and their potential for trap cropping. Ann. Appl. Biol. 2000, 136, 239− 246. (39) Franco, J.; Main, G.; Oros, R. Trap crops as a component for the integrated management of Globodera spp. (potato cyst nematode) in Bolivia. Nematropica 1999, 29, 51−60. (40) Oka, Y. Mechanisms of nematode suppression by organic soil amendments-A review. Appl. Soil Ecol 2010, 44, 101−115. (41) Barros, A. F.; Campos, V. P.; da Silva, J. C. P.; Pedroso, M. P.; Medeiros, F. H. V.; Pozza, E. A.; Reale, A. L. Nematicidal activity of volatile organic compounds emitted by Brassica juncea, Azadirachta indica, Canavalia ensiformis, Mucuna pruriens and Cajanus cajan against Meloidogyne incognita. Appl. Soil Ecol 2014, 80, 34−43. (42) Fourie, H.; Ahuja, P.; Lammers, J.; Daneel, M. Brassicaceabased management strategies as an alternative to combat nematode pests: A synopsis. Crop Prot. 2016, 80, 21−41. (43) Wu, H.; Wang, C. J.; Bian, X. W.; Zeng, S. Y.; Lin, K. C.; Wu, B.; Zhang, G. A.; Zhang, X. Nematicidal efficacy of isothiocyanates against root-knot nematode Meloidogyne javanica in cucumber. Crop Prot. 2011, 30, 33−37. (44) Valdes, Y.; Viaene, N.; Moens, M. Effects of yellow mustard amendments on the soil nematode community in a potato field with focus on Globodera rostochiensis. Appl. Soil Ecol 2012, 59, 39−47. (45) Brolsma, K. M.; van der Salm, R. J.; de Ron, G. G. Hatching of Globodera pallida cysts is inhibited by allyl isothiocyanate but not by Brassica juncea. Biofumigation 2014, 84, 1−22. (46) Ebrahimi, N.; Viaene, N.; Aerts, J.; Debode, J.; Moens, M. Agricultural Waste Amendments Improve Inundation Treatment of Soil Contaminated with Potato Cyst Nematodes, Globodera rostochiensis and G. pallida. Eur. J. Plant Pathol. 2016, 145, 755−775. (47) Ebrahimi, N.; Viaene, N.; Vandecasteele, B.; D’Hose, T.; Debode, J.; Cremelie, P.; De Tender, C. A.; Moens, M. Traditional and new soil amendments reduce survival and reproduction of potato cyst nematodes, except for biochar. Appl. Soil Ecol 2016, 107, 191− 204. (48) Hu, Y.; You, J.; Li, C.; Williamson, V. M.; Wang, C. Ethylene response pathway modulates attractiveness of plant roots to soybean cyst nematode Heterodera glycines. Sci. Rep. 2017, 7, 41282.
(49) Cabrera, J.; Díaz-Manzano, F. E.; Fenoll, C.; Escobar, C. Developmental pathways mediated by hormones in nematode feeding sites. In Plant Nematode Interactions: A View on Compatible Interrelationships ; Escobar, C., Fenoll, C., Eds.; Elsevier: San Diego, CA, 2015; pp 1−421. (50) Farnier, K.; Bengtsson, M.; Becher, P. G.; Witzell, J.; Witzgall, P.; Manduríc, S. Novel bioassay demonstrates attraction of the white potato cyst nematode Globodera pallida (Stone) to non-volatile and volatile host plant cues. J. Chem. Ecol. 2012, 38, 795−801. (51) Bellafiore, S.; Briggs, S. P. Nematode effectors and plant responses to infection. Curr. Opin. Plant Biol. 2010, 13, 442−448. (52) Coyne, D. L.; Cortada, L.; Dalzell, J. J.; Claudius-Cole, A. O.; Haukeland, S.; Luambano, N.; Talwana, H. Plant Parasitic Nematodes and Food Security in Sub-Saharan Africa. Annu. Rev. Phytopathol. 2018, 56, 1 DOI: 10.1146/annurev-phyto-080417-045833.
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