An Essay On Plant Pathogenic Nematodes Life

Abstract

Plant-parasitic nematodes (PPN) are responsible for substantial damages within the agriculture industry every year, which is a challenge that has thus far gone largely unimpeded. Chemical nematicides have been employed with varying degrees of success, but their implementation can be cumbersome, and furthermore they could potentially be neutralizing an otherwise positive effect from the entomopathogenic nematodes (EPN) that coexist with PPNs in soil environments and provide protection for plants against insect pests. Recent research has explored the potential of employing EPNs to protect plants from PPNs, while providing their standard degree of protection against insects. The interactions involved are highly complex, due to both the three-organism system and the assortment of variables present in a soil environment, but a strong collection of evidence has accumulated regarding the suppressive capacity of certain EPNs and their mutualistic bacteria, in the context of limiting the infectivity of PPNs. Specific factors produced by certain EPN complexes during the process of insect infection appear to have a selectively nematicidal, or at least repellant, effect on PPNs. Using this information, an opportunity has formed to adapt this relationship to large-scale, field conditions and potentially relieve the agricultural industry of one of its most substantial burdens.

Keywords: Entomopathogenic nematodes, Plant parasitic nematodes, Infection, Immunity, Parasitism

Graphical Abstract

1. Introduction

Nematodes are a fairly vast phylum, and many of the species in the group also happen to be parasitic, opportunistically inhabiting a range of hosts that include plants, insects and animals (Dillman and Sternberg, 2012; L’Ollivier and Piarroux, 2013; Quist et al., 2015). Naturally, depending on the type of organism a nematode has infected and the context in which the infection is taking place, a nematode’s success in terms of survival and parasitism can either be in line with, or sharply opposed to, human health or economic interests. Research into the interactions between the host immune system and nematode virulence mechanisms have therefore garnered considerable interest and support in the hope that these interactions can be mediated beneficially (Castillo et al., 2011; Babu and Nutman, 2014; Goverse and Smant, 2014).

In particular, nematodes can have a large impact on agriculture through their effect on populations of insects and plants alike. Those nematodes that are entomopathogenic, or insect parasitic, can generally be thought of as advantageous, and the Heterorhabditis and Steinernema genera of entomopathogenic nematodes (EPNs) have been employed specifically and intentionally as biocontrol agents for insect pests (Ehlers, 2001; Ffrench-Constant et al., 2007). Plant-infectious nematodes, on the other hand, account for approximately 5% of crop yield loss by limiting root growth, plant size and photosynthetic rate (Gheysen and Mitchum, 2011; Kyndt et al., 2013). One of the confounding issues with this situation then is that because nematodes present in a plant’s environment may be having opposite effects, an unspecific nematicidal treatment is eliminated as a viable strategy for crop protection, especially if EPNs are maintaining a population of insect pests below a harmful threshold, which may not be immediately apparent. These two kinds of nematodes may also have little competitive effect on each other, as efforts to suppress plant-parasitic Meloidogyne partityla nematodes with Steinernema feltiae produced inconsistent and marginal results (Shapiro-Ilan et al., 2006), although this may be a species-dependent effect, as other studies using different pairings of EPNs and plant-parasitic nematodes (PPNs) have shown more affirmative findings (Molina et al., 2007).

With this dynamic in place, guiding nematode interactions to a desired result may require a significant amount of tact and subtlety that will rely on a thorough knowledge of plant and insect immunity as it relates to nematode infection. It is important to consider as well that these two forms of immune response will have strong and fundamental differences. In general, insect innate immunity consists of mechanisms that lead to the production of antimicrobial peptides (AMPs) and reactive oxygen species (ROS), and cellular functions involving phagocytosis, encapsulation and nodulation (Welchman et al., 2009; Viljakainen, 2015), but plants, lacking a cellular response, will be incapable of responding to nematodes with functions such as encapsulation, which is the most common insect immune response to a metazoan invader (Jones and Dang, 2006; Muthamilarasan and Prasad, 2013; Honti et al., 2014; Vlisidou and Wood, 2015). A further examination of these two immune systems and their respective responses to nematode parasites will outline the primary mechanisms of insect and plant resistance.

2. The insect immune response to EPNs

A nematode infection of an insect host begins when a nematode of the infectious juvenile (IJ) stage attaches to the cuticle of the insect, penetrates through the various natural openings, such as the spiracles or mouth, and establishes itself in the hemolymph after advancing into the body cavity (Griffin, 2012). Once established, the nematode will release its mutualistic bacteria into the host hemolymph, Photorhabdus bacteria in Heterorhabditis nematodes and Xenorhabdus bacteria in Steinernema nematodes, either through regurgitation or esophageal pumping of the bacteria down through the intestine and out of the anus (Goodrich-Blair, 2007; Waterfield et al., 2009). These bacteria then go on to release toxic and immunosuppressive compounds, eventually leading to the death of the host by septicemia (Ffrench-Constant et al., 2007; Herbert and Goodrich-Blair, 2007). The release of nematode mutualistic bacteria does not occur immediately, however, and is instead delayed, by 30 min in the case of Heterorhabditis, and 4 – 6 h for Steinernema (Li et al., 2007). This means that the host insect has a window, granted of a variable timeframe depending on the species, in which it may neutralize the parasite before being forced to compensate for the additional challenge of the bacterial infection. In a general way, nematode avirulence is primarily achieved by initiating hemolymph clotting, activating a melanization reaction, and encapsulating the nematode in layers of hemocytes. Clot formation is based on the activity of soluble factors in the hemolymph, including transglutaminase, which will bind foreign bodies, including Photorhabdus (in the case of transglutaminase), and form microclots that can be incorporated into networks of fibers produced by hemolectin and triggrin that will further isolate the pathogen (Hyrsl et al., 2011; Toubarro et al., 2013). The melanization reaction, which is technically part of the humoral response, although it functions in close association with the cellular response, is comprised of the conversion of the inactive precursor prophenoloxidase to active phenoloxidase (Eleftherianos and Revenis, 2011), which generates the indole groups used to form melanin that then binds the nematode and supports the destruction of the parasite with ROS (Castillo et al., 2011). The cellular response, although thus far underrepresented, then plays perhaps the most crucial role by rapidly encapsulating the pathogen in compacted layers of hemocytes, which is a process that can be initiated within minutes of exposure, and potentially prevents the release of bacteria into the hemolymph (Satyavathi et al., 2014).

The release of bacteria by nematodes is clearly a challenge to the host that, once initiated, is difficult to overcome, as each nematode can release 50 – 200 bacteria directly into the hemolymph (Goodrich-Blair, 2007; Wu et al., 2014). The products of these bacteria can undermine much of the immune system, both cellular and humoral, by releasing toxic components that are capable of damaging hemocytes, and enzymes such as the RTX-like metalloprotease of Photorhabdus that can cleave hemolymph proteins involved in regulating host immune effector genes (Bowen et al., 2003; Rodou et al., 2010; Vlisidou et al., 2012). The success of the insect immune system in overcoming an infection by nematode parasites is therefore based largely on its ability to prevent the release of these bacteria and their immunosuppressive products, and indeed, a correlation between survival and the degree of encapsulation has been demonstrated (Li et al., 2007; Eleftherianos et al., 2010). A number of variables in the immune response, as viewed between different species of insects, reflect this point through their ability to influence host survival (Castillo et al., 2011). One such variable is the starting point of encapsulation. As mentioned previously, nematodes eject their mutualistic bacteria from the mouth and anus (Snyder et al., 2007), and therefore it would follow that the most effective method of encapsulation would be to cover these openings first in order to prevent the release of bacteria. Concordantly, encapsulation of Heterorhabditis by Manduca sexta hemocytes is initiated at the head and tail of the nematode (Li et al., 2007). In the Colorado potato beetle, Leptinotarsa decemlineata, however, encapsulation is initiated in the middle of the nematode, near the esophageal region, possibly in response to secretory-excretory pore exudates (Ebrahimi et al., 2011). This mechanism could represent a disadvantage for the insect, as the nematode could potentially eject bacteria during the encapsulation process and well before the process is complete. The time frame of the encapsulation process measured against the timing of bacterial release is also important, as both are represented by a spectrum of a fairly wide range. Encapsulation may begin within minutes of exposure, but completion of the task involves multiple stages, each with their own timing (Stanley et al., 2009; Jiang et al., 2010). Depending on the species involved in the interaction, the formation of multiple layers of hemocytes can take 45 min – 2 h, the complete encapsulation of the nematode 2 – 4 h, and partial melanization 16 – 24 h, as was observed with combinations of Heterorhabditis bacteriophora, L. decemlineata, and Galleria mellonella (Ebrahimi et al., 2011). These differences could be crucial in determining the survival of the insect based on the delay of bacterial release by the nematode, which as mentioned previously can be as brief as 30 min or as long as 4 – 6 h.

The insect immune response has been described in some detail, but one factor that has not yet been discussed is that nematode parasites are also capable of evading detection by the immune system, in which case the mechanics of a cellular response would be largely irrelevant (Brivio et al., 2005; Castillo et al., 2011). Lytic surface coat proteins, hydrophobic exudates, and lipopolysaccharide-like binding proteins produced by nematodes can all facilitate the parasite’s evasion of encapsulation (Li et al., 2009; Brivio et al., 2010; Mastore et al., 2014). Overall, the interaction can then be characterized as a highly complex interplay between the genotypes of the insect, the nematode and its mutualistic bacteria, which although perhaps difficult to predict, does provide a number of potential targets for control that could be beneficial to agriculture if applied appropriately. In the context of eliminating insect pests, strains could be developed that produce the proteins necessary for evasion, or nematodes could be generated that have significantly decreased delays in the ejection of their bacterial endosymbionts. Future research may do well to investigate the factors that generate differences in ejection timing, as nematodes that can overwhelm the insect immune system before being encapsulated would likely serve as much more efficient biocontrol agents.

3. The plant response to root knot nematodes (RKNs)

PPNs, and in particular, Root Knot Nematodes (RKNs) of the genus Meloidogyne carry out a slightly different parasitic protocol than their entomopathogenic counterparts (Curtis, 2007). RKNs initiate an infection by penetrating host root tips as second stage juveniles (J2) after hatching from eggs in the soil (Caillaud et al., 2008). Pharyngeal secretions from the nematode then induce proximal cells to undergo mitosis repeatedly without cytokinesis, such that a cluster of hypertrophic, multinucleated “giant” cells, referred to as a “gall”, is formed and serves as a food source for the nematode, which is then sedentary (Melillo et al., 2014; Davies et al., 2015). In addressing this invasion, the plant is limited to a program more similar to the insect humoral system than cellular system, as plants lack the specialized cell types required for a cellular response. The specific programs that moderate resistance to RKNs are still poorly understood, but progress has been made toward clarifying how some plants are able to gain relative advantages when employing their immune mechanisms against a nematode parasite.

A number of fundamental mechanisms appear to play a role in RKN resistance, including R genes such as Mi 1.2 in tomato plants, which encodes a member of the nucleotide binding leucine rich repeat gene family (Villeth et al., 2015), and the hypersensitive response, which appears to be a commonly employed mechanism for limiting RKN feeding capacities. In particular, a resistance gene referred to as RMc1(blb) in potatoes is known to limit nematode virulence by influencing the concentration of Ca2+ in root cells near the nematode (Davies et al., 2015). In this system, RMc1(blb) apparently induces the upregulation of the calcium-dependent protein kinase CDPK4, which may play a role in the release of ROS in response to a Ca2+ influx, possibly then inducing cell death characteristic of the hypersensitive response and preventing the nematode from forming giant cells as a resource reservoir. It is important to note, however, that the hypersensitive response is not always required for resistance to RKNs. Alfalfa for instance, does not display a hypersensitive phenotype in response to infection, but instead is apparently capable of resistance by denying nematodes access to the developing vascular cylinder of the root (Postnikova et al., 2015).

The salicylic acid (SA) and jasmonic acid (JA) pathways have also been identified as potential proponents of immunity. JA biosynthesis and signaling component genes have been found to be upregulated after RKN infection, and exogenous application of JA has been found to reduce root egg masses in tomato plants (Zhou et al., 2015). Exogenous application of the SA analog benzothiadiazole (BTH) to tomato plants also led to a decrease in gall formation, which is believed to be the result of cell-wall stiffening due to increased H2O2 production, lignin accumulation, and peroxidase activity in response to the BTH exposure (Melillo et al., 2014). Furthermore, when exposed to BTH, parenchymal cells surrounding the feeding site underwent a change in Tap1 anionic peroxidase expression and subsequently appeared to entrap the nascent giant cells, potentially preventing them from expanding further, perhaps similar to a rudimentary, extemporaneous cellular response.

Enhancing resistance in crop plants to Meloidogyne would be remarkably beneficial to agriculture as an industry, but there are still significant voids in the available information about the plant immune mechanisms that are capable of conferring this resistance. Much of the information available is limited to a level of detail that refers only to functional groups such as oxidative stress or ubiquitination as means of resistance (Kyndt et al., 2012; Villeth et al., 2015), and in cotton, overexpression of Meloidogyne-induced cotton 3 (MIC3) was able to dramatically reduce egg production by Meloidogyne incognita, but the function of MIC3 remains entirely unknown, and it does not appear to have any identifiable functional domains (Wubben et al., 2015). The interaction between plant parasitic nematodes and their hosts therefore represents an exciting field, potentially rich with novel interactions that in the future could be used to develop more stable and resistant crop plants.

4. The suppression of PPNs with EPNs

There is a collection of research describing interactions between EPNs and PPNs with regard to the ability or tendency of EPNs to interfere with the normal activity of PPNs. Fortunately, the available research allows a number of stepwise conclusions to be made about this system, based on specific experimental characteristics, such as the species of EPN being used and the presence or absence of an insect host, which indicate strongly that the symbiotic bacteria of EPNs may be responsible for the observed adverse effects on PPNs due to the production of deterrent factors from these symbiotic bacteria together with the factors that are produced to weaken or kill an insect host. The fundamental concepts of the interactions are generally straightforward, but the array of details that can affect the system will prove to be remarkably complex.

Using EPNs to deter PPNs for the benefit of a plant is a task that becomes rapidly more and more complex once the environment is transitioned from a controlled laboratory setting to the endlessly variable conditions present in an agricultural soil environment. Control of the system requires a method that accounts for, or simplifies, interactions between both types of nematodes and the plants, the two groups of nematodes, and certain insect species that act as pests of the plants and hosts of the EPNs. Moreover, each of the four parties encompasses a myriad of species that are not all likely to interact in the same way. Certain EPNs may be more effective against certain species of PPNs, and some plant species may not be as attuned to the benefits of specific EPNs. A number of studies have, however, thoroughly examined specific instances of the four-part interaction, providing individual pieces of information that have begun to form a pattern. Superficially the results can appear to lack a robust quality, or otherwise look inconsistent, but the specific successes that have appeared demonstrate a clear potential for the biological control of PPNs with EPNs.

One notable characteristic that separates the studies done to date is the application method used to introduce the EPNs, which can involve direct application or the use of an insect host cadaver as a delivery vessel. The more common method examined so far has been the direct introduction of IJ EPNs in aqueous solution (Kepenekci et al., 2015), although this specific technique has produced fairly mixed results. In another study, EPNs were applied in greenhouse and laboratory assays in order to monitor their effect on the Meloidogyne javanica infection process in tomatoes and soybeans (Fallon et al., 2002). The specific EPNs used were Heterorhabditis indica, as well as S. feltiae and Steinernema riobrave, but the only effect observed was a reduction in M. javanica root penetration after the introduction of S. feltiae. All other treatments were similar to water control treatments with regard to root penetration, M. javanica egg production and plant biomass, which generally can be interpreted as an insufficient degree of effect to be considered suppression. Pérez and Lewis (2002) however, observed strong suppressive effects, albeit with a significantly altered approach in terms of experimental setup. In this study, the ability of the same Steinernema spp., S. feltiae and S. riobrave, together with H. bacteriophora, was monitored for protective capacity when co-incubated under laboratory conditions with tomato seedlings and M. incognita (PPN) eggs. The results showed that the seedlings treated with the two Steinernema spp. had fewer M. incognita juveniles infecting their roots, and that these nematodes also went on to produce fewer eggs. Furthermore, all EPN introduction time points from 2 weeks prior to M. incognita infection to 2 weeks after were effective, and increases in the concentration of Steinernema were correlated with an enhanced suppressive effect, although low and high concentrations both resulted in suppression. Grewal et al. (1999) also examined tomato root penetration by M. incognita in sterilized sand, and the ability of Steinernema spp. to suppress it, but found that the application of live Steinernema did not have a protective effect. The scattered positive results that have arisen from the application of IJs seem to indicate a potential capacity of EPNs for direct interference with PPNs, but arguments could be made that this is merely an effect of highly variable experimental designs. In the case of the Pérez and Lewis (2002) study for instance, a control was not in place to identify any differences between control and treatment trials in reference to the ability of M. incognita eggs to hatch properly. These assays may have simply appeared to provide evidence of protection due to the partial destruction or manipulation of the M. incognita eggs by the Steinernema IJs.

Another significant factor that undoubtedly guides the results of these studies is that they have been performed under greenhouse or laboratory conditions, which highlights the importance of another study that identified a suppressive capacity in EPNs. The authors performed a field study of golf course turf grass that included M. incognita and aqueous application of S. riobrave, and found that S. riobrave was as effective as chemical nematicide, if not more so (Grewal et al., 1997). One consideration that comes with this result is that the natural life cycle of EPNs, and namely their interactions with and the ability to infect insect hosts, may play a role in the suppression of PPNs, even if the nematodes alone do not possess the ability to interfere with the infection process. EPNs that have infected a host have been found to display greater infectivity, increased dispersal and increased survival compared with nematodes in aqueous suspension (Shapiro and Glazer, 1996; Shapiro and Lewis, 1999; Pérez et al., 2003). Consistent with this notion, a number of studies have performed similar tests to those completed with IJs in aqueous suspension, but instead co-incubated plants and PPNs with insect cadavers that had been infected with EPNs prior to the assay. Some of these studies also performed duplications of their assays with IJs in aqueous suspension, allowing for a direct comparison between the two methods within the same experimental design. One such study found that insect hosts infected with Steinernema carpocapsae, S. feltiae, and S. riobrave were all capable of repelling M. incognita from tomato roots during a greenhouse trial, while the application of IJs failed to promote a decrease in root penetration (Grewal et al., 1999). Other similar studies, however, remain mottled with inconsistency or limited results, although with a trend towards suppression. For example, a previous study found an 18% reduction in egg masses from Meloidogyne partityla during greenhouse assays in pecans after the application of S. riobrave-infected hosts as well as increased dry-root weight when plants were treated with S. feltiae-infected hosts (Shapiro-Ilan et al., 2006). Generally, however, the results were not strong enough to conclude that M. partityla could be suppressed with the entomopathogenic species tested. Interestingly, another study found evidence of suppression by an H. bacteriophora isolate (Rama Caida) when used as a treatment for M. incognita infection in pepper and summer squash (Del Valle et al., 2013). The number of M. incognita eggs was reduced in both plants 60 days p.i. Oddly, the Steinernema spp. tested did not produce suppressive results, although the species was Steinernema diaprepesi rather than the more commonly used S. feltiae or S. riobrave.

The next challenge then is to explain why a suppressive effect might generally be more likely to occur if the EPN being used as a treatment is first allowed to infect an insect host. Perhaps the most striking change between these two treatments is the fact that if the EPN has infected a host, its mutualistic bacteria will have been released into the host where they will have begun to secrete components that aggressively enhance the infection process. As mentioned, PPNs do not infect cooperatively with mutualistic bacteria, so a reasonable assumption is that they would be less tolerant of the products of an EPN’s mutualistic bacteria. Some evidence has been generated to support this notion and even demonstrate that the cell-free culture supernatants of these bacteria may be sufficient to repel PPNs. As early as 1997, investigators demonstrated this effect with cell-free culture filtrates of the mutualistic bacteria of S. carpocapsae, S. feltiae and S. riobrave, namely Xenorhabdus nematophilus, Xenorhabdus bovienii and Xenorhabdus R, respectively (Grewal et al., 1997). All three of these filtrates were nematicidal and demonstrated 98 – 100% mortality when tested against M. incognita IJs, thus building on previous research indicating that Xenorhabdus spp. produce ammonia, indole and stilbene derivatives, and that the stilbene derivatives and ammonia have selective nematicidal capacities (Paul et al., 1981; Hu et al., 1988; Richardson et al., 1988). In their 2002 study, Pérez and Lewis also suggested that their results may have been partially due to the activity of Xenorhabdus spp. (Pérez and Lewis 2002), and in a 2015 study, the authors were able to demonstrate that dipping tomato plants in X. bovienii spent medium supernatants is capable of suppressing M. incognita egg mass numbers and increasing plant height compared with infected, but untreated, controls (Kepenecki et al., 2015). This result was also compared with a less effective Photorhabdus luminescens treatment, which is consistent with the more stable suppressive results observed when S. feltiae is used for biocontrol rather than H. bacteriophora (Lewis and Grewal, 2005).

The suppressive effects of Xenorhabdus filtrates have therefore been somewhat well demonstrated in the context of PPNs, but moreover, the effects of Xenorhabdus have also been well examined in the context of repelling scavengers in general, likely as a method of preventing the nematode host of the bacteria from being destroyed while reproducing within an infected insect (Zhou et al., 2002). The authors characterized a component of X. nematophila supernatants that is capable of repelling the Argentine ant Linepithema humile. The repellent component was found to be filterable, heat stable, acid sensitive and small enough to pass through a 10 kDa pore-size membrane, as well as present in some form in P. luminescens, although optimally present in supernatants after 132 h compared with the 108 h of Xenorhabdus cultures, which may provide at least a partial explanation of the more consistent efficacy of S. feltiae. These observations were later expanded in a well-controlled study in which the feeding behaviors of ants, crickets and wasps on nematode, freeze and axenic nematode-killed insects were monitored (Gulcu et al., 2012). The results showed that in general, the tested species would not feed on an insect killed at least 2 days previously by a nematode with its mutualistic bacteria. Crickets were, however, found to feed on insects infected by axenic nematodes, insects infected with Serratia marcescens, and freeze-killed insects, strongly indicating that the mutualistic bacteria were responsible for the deterrent effect.

EPNs therefore have demonstrated a strong potential, through their mutualistic bacteria, as a possible biocontrol method for PPNs (Fig. 1). Applying this potential would represent a tremendous advantage to the agriculture industry, but some of the subtleties involved in effectively introducing and sustaining the suppressive effects will require some additional work and research. The current information available seems to indicate that S. feltiae, and specifically X. bovienii, is the strongest candidate. As mentioned previously, S. feltiae has had the most consistent suppressive results, and its mutualistic X. bovienii has been suggested to produce a higher or otherwise more effective concentration of the deterrent factor (Gulcu et al., 2012). The aforementioned studies seem to indicate that S. feltiae contributes to suppression primarily as a vessel, which means a number of methods could be used to deploy X. bovienii in the field, although each with relative strengths and weaknesses. For instance it was suggested that plants could be dipped in Xenorhabdus culture filtrates, but this may only be effective transiently, as the stability of the nematicidal substances and deterrent factor in soil remain unknown (Kepenecki et al., 2015). Employing this method commercially would also require that nematodes still be used to harbor the Xenorhabdus at least occasionally, as Xenorhabdus can revert to a secondary form spontaneously in long-standing laboratory cultures, and fail to produce the substances that are associated with its cooperative action in Steinernema spp. (Zhou et al., 2002). The application of IJs carries its own challenges as well, however. The common application rate of IJs that has been shown to be effective is 2.5 billion IJs per hectare (Lewis et al., 2001), but this concentration is naturally reduced rapidly after application (Smits, 1996) due to a lack of hosts, temperature fluctuations, UV radiation, dessication (Kaya, 1990), and antagonists present in the soil (Kaya, 2002). A treatment that effectively suppresses PPNs will therefore require some enhancement in the stability or maintenance of Steinernema, or their associated Xenorhabdus spp., in a soil environment. This is a broad statement, but the possible solutions are just as broad. Additional research may develop a stable Xenorhabdus strain that innocuously associates with a variety of plant species, or a method for efficiently co-seeding a soil environment with EPNs and an insect host could be designed. Regardless of the specific method eventually used to address the biocontrol of PPNs, the presently available information leaves the field poised for a solution to a long-standing and high priority obstacle to agriculture.

Fig. 1

A representative illustration of the context in which an entomopathogenic nematode (EPN) (Steinernema nematodes) can interfere with the infection process of a plant pathogenic nematode (PPN) (Meloidogyne nematodes). The EPNs are seen entering the insect...

Additional research would also need to be done in order to determine whether the deterrent factor(s) being produced by Xenorhabdus is specific to certain nematode species or if the nematodes are responding to a broad-spectrum factor. EPNs may benefit from specifically deterring PPNs in that PPNs damage plants, and that these plants are also a food source for the insect pests that the EPNs consume, but a concomitantly plausible theory would be that certain insect pests could also benefit from the activity of PPNs, which may exhaust the immune resources of the plant and reduce its capacity for defense against insect herbivory. The exact balance and thresholds of benefit and detriment are likely to be highly variable however, as the species involved could dramatically affect long term outcomes based on specific immune responses as well as relative consumption and reproduction rates, among other factors. Despite this difficulty, an attempt to isolate a factor specific to the repulsion or killing of PPNs would likely be a worthwhile pursuit because the field use of one specific factor could greatly increase efficiency and reduce the possibility of detrimental, non-specific effects.

5. Concluding remarks

Nematodes are ubiquitously involved in many human interests, in terms of health, as over 100 species of nematodes are human parasites, and with regard to the global economy as nematodes can function as plant, livestock or insect parasites. In particular, because nematodes play a dual role in agriculture as both biocontrol agents of harmful insect pests and pathogens of plants themselves, a more detailed and complete understanding of how nematodes interact with the immune systems of their respective hosts could provide a strong advantage in developing strategies to mitigate the damage caused by plant pathogens and enhance the efficacy of EPNs as a biocontrol mechanism. An immunological approach may be especially beneficial as it can target specific nematode groups rather than employing a broad nematicidal agent, which could equally promote deleterious effects. Some fields of research that may be of particular interest include the pathways that dictate the timing of mutualistic bacteria ejection in EPNs, the mechanisms by which plants are able prevent RKN species from entering their root systems, and the precise nature of the relationship between EPNs and PPNs, especially with regard to the effect of EPN bacterial factors on PPNs, as this field may provide methods for control that are both efficiently applicable and effective.

Acknowledgments

We thank members the Columbian College of Arts and Sciences at George Washington University, Washington DC, USA for funding. Research in the laboratory of Ioannis Eleftherianos is supported in part by National Institutes of Health, USA, grants 1R01AI110675-01A1, 1R56AI110675-01 and 1R21AI109517-01A1.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  • Babu S, Nutman TB. Immunology of lymphatic filariasis. Parasite Immunol. 2014;36:338–346.[PMC free article][PubMed]
  • Bowen DJ, Rocheleau TA, Grutzmacher CK, Meslet L, Valens M, Marble D, Dowling A, Ffrench-Constant R, Blight MA. Genetic and biochemical characterization of PrtA, an RTX-like metalloprotease from Photorhabdus. Microbiology. 2003;149:1581–1591.[PubMed]
  • Brivio MF, Mastore M, Nappi AJ. A pathogenic parasite interferes with phagocytosis of insect immunocompetent cells. Dev Comp Immunol. 2010;34:991–998.[PubMed]
  • Brivio MF, Mastore M, Pagani M. Parasite-host relationship: a lesson from a professional killer. Invertebr Surv J. 2005;2:41–53.
  • Caillaud MC, Dubreuil G, Quentin M, Perfus-Barbeoch L, Lecomte P, de Almeida Engler J, Abad P, Rosso MN, Favery B. Root-knot nematodes manipulate plant cell functions during a compatible interaction. J Plant Physiol. 2008;165:104–113.[PubMed]
  • Castillo JC, Reynolds SE, Eleftherianos I. Insect immune responses to nematode parasites. Trends Parasitol. 2011;27:537–547.[PubMed]
  • Curtis RH. Plant parasitic nematode proteins and the host parasite interaction. Brief Funct Genomic Proteomic. 2007;6:50–58.[PubMed]
  • Davies LJ, Brown CR, Elling AA. Calcium is involved in the R Mc1 (blb)-mediated hypersensitive response against Meloidogyne chitwoodi in potato. Plant Cell Rep. 2015;34:167–177.[PubMed]
  • Del Valle EE, Lax P, Duenas JR, Doucet ME. Effects of insect cadavers infected by Heterorhabditis bacteriophora and Steinernema diaprepesi on Meloidogyne incognita parasitism in pepper and summer squash plants. Ciencia e Investigacion Agraria. 2013;40:109–118.
  • Dillman AR, Sternberg PW. Entomopathogenic nematodes. Curr Biol. 2012;22:R430–R431.[PMC free article][PubMed]
  • Ebrahimi L, Niknam G, Dunphy GB. Hemocyte responses of the Colorado potato beetle, Leptinotarsa decemlineata, and the greater wax moth, Galleria mellonella, to the entomopathogenic nematodes, Steinernema feltiae and Heterorhabditis bacteriophora. J Insect Sci. 2011;11:75.[PMC free article][PubMed]
  • Ehlers RU. Mass production of entomopathogenic nematodes for plant protection. Appl Microbiol Biotechnol. 2001;56:623–633.[PubMed]
  • Eleftherianos I, Ffrench-Constant RH, Clarke DJ, Dowling AJ, Reynolds SE. Dissecting the immune response to the entomopathogen Photorhabdus. Trends Microbiol. 2010;18:552–560.[PubMed]
  • Eleftherianos I, Revenis C. Role and importance of phenoloxidase in insect hemostasis. J Innate Immun. 2011;3:28–33.[PubMed]
  • Fallon DJ, Kaya HK, Gaugler R, Sipes BS. Effects of Entomopathogenic Nematodes on Meloidogyne javanica on Tomatoes and Soybeans. J Nematol. 2002;34:239–245.[PMC free article][PubMed]
  • Ffrench-Constant RH, Dowling A, Waterfield NR. Insecticidal toxins from Photorhabdus bacteria and their potential use in agriculture. Toxicon. 2007;49:436–451.[PubMed]
  • Gheysen G, Mitchum MG. How nematodes manipulate plant development pathways for infection. Curr Opin Plant Biol. 2011;14:415–421.[PubMed]
  • Goodrich-Blair H. They’ve got a ticket to ride: Xenorhabdus nematophila-Steinernema carpocapsae symbiosis. Curr Opin Microbiol. 2007;10:225–230.[PubMed]
  • Goverse A, Smant G. The activation and suppression of plant innate immunity by parasitic nematodes. Annu Rev Phytopathol. 2014;52:243–265.[PubMed]
  • Grewal PS, Martin WR, Miller RW, Lewis EE. Suppression of plant-parasitic nematode populations in turfgrass by application of entomopathogenic nematodes. Bio Sci Technol. 1997;7:393–399.
  • Grewal PS, Lewis EE, Venkatachari S. Allelopathy: a possible mechanism of suppression of plant-parasitic nematodes by entomopathogenic nematodes. Nematology. 1999;1:735–743.
  • Griffin CT. Perspectives on the behavior of entomopathogenic nematodes from dispersal to reproduction: traits contributing to nematode fitness and biocontrol efficacy. J Nematol. 2012;44:177–184.[PMC free article][PubMed]
  • Gulcu B, Hazir S, Kaya HK. Scavenger deterrent factor (SDF) from symbiotic bacteria of entomopathogenic nematodes. J Invertebr Pathol. 2012;110:326–333.[PubMed]
  • Herbert EE, Goodrich-Blair H. Friend and foe: the two faces of Xenorhabdus nematophila. Nat Rev Microbiol. 2007;5:634–646.[PubMed]
  • Honti V, Csordás G, Kurucz É, Márkus R, Andó I. The cell-mediated immunity of Drosophila melanogaster: hemocyte lineages, immune compartments, microanatomy and regulation. Dev Comp Immunol. 2014;42:47–56.[PubMed]
  • Hu KJ, Li JX, Wang WJ, Wu HM, Lin H, Webster JM. Comparison of metabolites produced in vitro and in vivo by Photorhabdus luminescens, a bacterial symbiont of the entomopathogenic nematode Heterorhabditis megidis. Can J Microbiol. 1988;44:1072–1077.
  • Hyrsl P, Dobes P, Wang Z, Hauling T, Wilhelmsson C, Theopold U. Clotting factors and eicosanoids protect against nematode infections. J Innate Immun. 2011;3:65–70.[PubMed]
  • Jiang H, Vilcinskas A, Kanost MR. Immunity in lepidopteran insects. Adv Exp Med Biol. 2010;708:181–204.[PubMed]
  • Jones JD, Dang JL. The plant immune system. Nature. 2006;444:323–329.[PubMed]
  • Kaya HK. Soil ecology. In: Gaugler R, Kaya HK, editors. Entomopathogenic Nematodes in Biological Control. CRC Press; Boca Raton, FL, USA: 1990. pp. 93–116.
  • Kaya HK. Natural enemies and other antagonists. In: Gaugler R, editor. Entomopathogenic Nematology. CABI; New York, USA: 2002. pp. 189–204.
  • Kepenekci I, Hazir S, Lewis EE. Evaluation of entomopathogenic nematodes and the supernatants of the in vitro culture medium of their mutualistic bacteria for the control of the root-knot nematodes Meloidogyne incognita and M. arenaria. Pest Manag Sci. 2015 doi: 10.1002/ps.3998.[PubMed][Cross Ref]
  • Kyndt T, Denil S, Haegeman A, Trooskens G, Bauters L, Van Criekinge W, De Meyer T, Gheysen G. Transcriptional reprogramming by root knot and migratory nematode infection in rice. New Phytol. 2012;196:887–900.[PubMed]
  • Kyndt T, Vieira P, Gheysen G, de Almeida-Engler J. Nematode feeding sites: unique organs in plant roots. Planta. 2013;238:807–818.[PubMed]
  • L’Ollivier C, Piarroux R. Diagnosis of human nematode infections. Expert Rev Anti Infect Ther. 2013;11:1363–1376.[PubMed]
  • Lewis EE, Grewal PS. Interactions with plant-parasitic nematodes. In: Grewal PS, Ehlers R-U, Shapiro-Ilan DI, editors. Nematodes as Biocontrol Agents. CABI; New York, USA: 2005. pp. 349–362.
  • Lewis EE, Grewal PS, Sardanelli S. Interactions between the Steinernema feltiae-Xenorhabdus bovienii insect pathogen complex and the root-knot nematode Meloidogyne incognita. Biol Control. 2001;21:55–62.
  • Li XY, Cowles RS, Cowles EA, Gaugler R, Cox-Foster DL. Relationship between the successful infection by entomopathogenic nematodes and the host immune response. Int J Parasitol. 2007;37:365–374.[PubMed]
  • Li X, Cowles EA, Cowles RS, Gaugler R, Cox-Foster DL. Characterization of immunosuppressive surface coat proteins from Steinernema glaseri that selectively kill blood cells in susceptible hosts. Mol Biochem Parasitol. 2009;165:162–169.[PubMed]
  • Mastore M, Arizza V, Manachini B, Brivio MF. Modulation of immune responses of Rhynchophorus ferrugineus (Insecta: Coleoptera) induced by the entomopathogenic nematode Steinernema carpocapsae (Nematoda: Rhabditida) Insect Sci. 2014 doi: 10.1111/1744-7917.12141.[PubMed][Cross Ref]
  • Melillo MT, Leonetti P, Veronico P. Benzothiadiazole effect in the compatible tomato-Meloidogyne incognita interaction: changes in giant cell development and priming of two root anionic peroxidases. Planta. 2014;240:841–854.[PubMed]
  • Molina JP, Dolinski C, Souza RM, Lewis EE. Effect of Entomopathogenic Nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) on Meloidogyne mayaguensis Rammah and Hirschmann (Tylenchida: Meloidoginidae) Infection in Tomato Plants. J Nematol. 2007;39:338–342.[PMC free article][PubMed]
  • Muthamilarasan M, Prasad M. Plant innate immunity: an updated insight into defense mechanism. J Biosci. 2013;38:433–449.[PubMed]
  • Paul VJ, Frautschy S, Fenical W, Nealson KH. Antibiotics in microbial ecology. Isolation and structure assignment of several new antibacterial compounds from the insect-symbiotic bacteria Xenorhabdus spp. J Chem Ecol. 1981;7:589–597.[PubMed]
  • Pérez EE, Lewis EE. Use of entomopathogenic nematodes to suppress Meloidogyne incognita on greenhouse tomatoes. J Nematol. 2002;34:171–174.[PMC free article][PubMed]
  • Pérez EE, Lewis EE, Shapiro-Ilan DI. Impact of the host cadaver on survival and infectivity of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) under desiccating conditions. J Invertebr Pathol. 2003;82:111–118.[PubMed]
  • Postnikova OA, Hult M, Shao J, Skantar A, Nemchinov LG. Transcriptome analysis of resistant and susceptible alfalfa cultivars infected with root-knot nematode Meloidogyne incognita. PLoS One. 2015;10:1–17.[PMC free article][PubMed]
  • Quist CW, Smant G, Helder J. Evolution of Plant Parasitism in the Phylum Nematoda. Annu Rev Phytopathol. 2015;53:289–310.[PubMed]
  • Richardson WH, Schmidt TM, Nealson KH. Identification of an anthraquinone pigment and a hydroxystilbene antibiotic from Xenorhabdus luminescens. Appl Environ Microbiol. 1988;54:1602–1605.[PMC free article][PubMed]
  • Rodou A, Ankrah DO, Stathopoulos C. Toxins and secretion systems of Photorhabdus luminescens. Toxins. 2010;2:1250–1264.[PMC free article][PubMed]
  • Satyavathi VV, Minz A, Nagaraju J. Nodulation: an unexplored cellular defense mechanism in insects. Cell Signal. 2014;26:1753–1763.[PubMed]
  • Shapiro DI, Glazer I. Comparison of entomopathogenic nematode dispersal from infected hosts versus aqueous suspension. Biol Control. 1996;25:1455–1461.
  • Shapiro DI, Lewis EE. Comparison of entomopathogenic nematode infectivity from infected hosts versus aqueous suspension. Biol Control. 1999;28:907–911.
  • Shapiro-Ilan DI, Nyczepir AP, Lewis EE. Entomopathogenic Nematodes and Bacteria Applications for Control of the Pecan Root-Knot Nematode, Meloidogyne partityla, in the Greenhouse. J Nematol. 2006;38:449–454.[PMC free article][PubMed]
  • Smits P. Post-application persistence of entomopathogenic nematodes. Bio Sci Technol. 1996;6:379–387.
  • Snyder H, Stock SP, Kim SK, Flores-Lara Y, Forst S. New insights into the colonization and release processes of Xenorhabdus nematophila and the morphology and ultrastructure of the bacterial receptacle of its nematode host, Steinernema carpocapsae. Appl Environ Microbiol. 2007;73:5338–5346.[PMC free article][PubMed]
  • Stanley D, Miller J, Tunaz H. Eicosanoid actions in insect immunity. J Innate Immun. 2009;1:282–290.[PubMed]
  • Toubarro D, Avila MM, Montiel R, Simoes N. A pathogenic nematode targets recognition proteins to avoid insect defenses. PLoS One. 2013;8:1–13.[PMC free article][PubMed]
  • Villeth GR, Carmo LS, Silva LP, Fontes W, Grynberg P, Saraiva M, Brasileiro AC, Carneiro RM, Oliveira JT, Grossi-de-Sá MF, Mehta A. Cowpea-Meloidogyne incognita interaction: Root proteomic analysis during early stages of nematode infection. Proteomics. 2015;15:1746–1759.[PubMed]
  • Viljakainen L. Evolutionary genetics of insect innate immunity. Brief Funct Genomics. 2015 pii, elv002. [PMC free article][PubMed]
  • Vlisidou I, Waterfield N, Wood W. Elucidating the in vivo targets of Photorhabdus toxins in real-time using Drosophila embryos. Adv Exp Med Biol. 2012;710:49–57.[PubMed]
  • Vlisidou I, Wood W. Drosophila blood cells and their role in immune responses. FEBS J. 2015;282:1368–1382.[PubMed]
  • Waterfield NR, Ciche T, Clarke D. Photorhabdus and a host of hosts Annu. Rev Microbiol. 2009;63:557–574.[PubMed]
  • Welchman DP, Aksoy S, Jiggins F, Lemaitre B. Insect immunity: from pattern recognition to symbiont-mediated host defense. Cell Host Microbe. 2009;6:107–114.[PubMed]
  • Wu G, Zhao Z, Liu C, Qiu L. Priming Galleria mellonella (Lepidoptera: Pyralidae) larvae with heat-killed bacterial cells induced an enhanced immune protection against Photorhabdus luminescens TT01 and the role of innate immunity in the process. J Econ Entomol. 2014;107:559–569.[PubMed]
  • Wubben MJ, Callahan FE, Velten J, Burke JJ, Jenkins JN. Overexpression of MIC-3 indicates a direct role for the MIC gene family in mediating Upland cotton (Gossypium hirsutum) resistance to root-knot nematode (Meloidogyne incognita) Theor Appl Genet. 2015;128:199–209.[PubMed]
  • Zhou J, Jia F, Shao S, Zhang H, Li G, Xia X, Zhou Y, Yu J, Shi K. Involvement of nitric oxide in the jasmonate-dependent basal defense against root-knot nematode in tomato plants. Front Plant Sci. 2015;6:193.[PMC free article][PubMed]
  • Zhou X, Kaya HK, Heungens K, Goodrich-Blair H. Response of ants to a deterrent factor(s) produced by the symbiotic bacteria of entomopathogenic nematodes. Appl Environ Microbiol. 2002;68:6202–6209.[PMC free article][PubMed]

Highlights

  • Entomopathogenic nematodes coexist with plant pathogenic nematodes

  • Nematodes interact with the immune systems of their respective hosts

  • Plant pathogenic nematodes suppress entomopathogenic nematodes

  • Factors from entomopathogenic nematodes can repel plant pathogenic nematodes

  • Novel strategies are required to control plant pathogenic nematodes in the field

Root-knot nematodes are plant-parasiticnematodes from the genus Meloidogyne. They exist in soil in areas with hot climates or short winters. About 2000 plants worldwide are susceptible to infection by root-knot nematodes and they cause approximately 5% of global crop loss.[1] Root-knot nematode larvae infect plant roots, causing the development of root-knot galls that drain the plant's photosynthate and nutrients. Infection of young plants may be lethal, while infection of mature plants causes decreased yield.

Economic impact[edit]

Root-knot nematodes (Meloidogyne spp.) are one of the three most economically damaging genera of plant-parasitic nematodes on horticultural and field crops. Root-knot nematodes are distributed worldwide, and are obligate parasites of the roots of thousands of plant species, including monocotyledonous and dicotyledonous, herbaceous and woody plants. The genus includes more than 90 species,[2] with some species having several races. Four Meloidogyne species (M. javanica, M. arenaria, M. incognita, and M. hapla) are major pests worldwide, with another seven being important on a local basis.[3]Meloidogyne occurs in 23 of 43 crops listed as having plant-parasitic nematodes of major importance, ranging from field crops, through pasture and grasses, to horticultural, ornamental and vegetable crops.[4] If root-knot nematodes become established in deep-rooted, perennial crops, control is difficult and options are limited.

Meloidogyne spp. were first reported in cassava by Neal in 1889.[5] Damage on cassava is variable depending on cultivar planted, and can range from negligible to serious.[6] Early-season infection leads to worse damage.[7] In most crops, nematode damage reduces plant health and growth; in cassava, though, nematode damage sometimes leads to increased aerial growth as the plants try to compensate. This possibly enables the plant to maintain a reasonable level of production. Therefore, aerial correlations to nematode density can be positive, negative or not at all.[8]Vegetable crops grown in warm climates can experience severe losses from root-knot nematodes, and are often routinely treated with a chemical nematicide. Root-knot nematode damage results in poor growth, a decline in quality and yield of the crop and reduced resistance to other stresses (e.g. drought, other diseases). A high level of damage can lead to total crop loss. Nematode-damaged roots do not use water and fertilisers as effectively, leading to additional losses for the grower. In cassava, it has been suggested that levels of Meloidogyne spp. that are sufficient to cause injury rarely occur naturally.[8] However, with changing farming systems, in a disease complex or weakened by other factors, nematode damage is likely to be associated with other problems.[9]

Control[edit]

Root-knot nematodes can be controlled with biocontrol agents Paecilomyces lilacinus, Pasteuria penetrans[10] and Juglone.[11]

Life cycle[edit]

All nematodes pass through an embryonic stage, four juvenile stages (J1–J4) and an adult stage. Juvenile Meloidogynes parasites hatch from eggs as vermiform, second-stage juveniles (J2), the first moult having occurred within the egg. Newly hatched juveniles have a short free-living stage in the soil, in the rhizosphere of the host plants. They may reinvade the host plants of their parent or migrate through the soil to find a new host root. J2 larvae do not feed during the free-living stage, but use lipids stored in the gut.[3]

An excellent model system for the study of the parasitic behaviour of plant-parasitic nematodes has been developed using Arabidopsis thaliana as a model host.[12] The Arabidopsis roots are initially small and transparent, enabling every detail to be seen. Invasion and migration in the root was studied using M. incognita.[13] Briefly, second stage juveniles invade in the root elongation region and migrate in the root until they became sedentary. Signals from the J2 promote parenchyma cells near the head of the J2 to become multinucleate[14] to form feeding cells, generally known as giant cells, from which the J2 and later the adults feed.[15] Concomitant with giant cell formation, the surrounding root tissue gives rise to a gall in which the developing juvenile is embedded. Juveniles first feed from the giant cells about 24 hours after becoming sedentary.

After further feeding, the J2s undergo morphological changes and become saccate. Without further feeding, they moult three times and eventually become adults. In females, which are close to spherical, feeding resumes and the reproductive system develops.[3] The life span of an adult female may extend to three months, and many hundreds of eggs can be produced. Females can continue egg laying after harvest of aerial parts of the plant and the survival stage between crops is generally within the egg.

The length of the life cycle is temperature-dependent.[16][17] The relationship between rate of development and temperature is linear over much of the root-knot nematode life cycle, though it is possible the component stages of the life cycle, e.g. egg development, host root invasion or growth, have slightly different optima. Species within the Meloidogyne genus also have different temperature optima. In M. javanica, development occurs between 13 and 34 °C, with optimal development at about 29 °C.

Gelatinous matrix[edit]

Root-knot nematode females lay eggs into a gelatinous matrix produced by six rectal glands and secreted before and during egg laying.[18] The matrix initially forms a canal through the outer layers of root tissue and later surrounds the eggs, providing a barrier to water loss by maintaining a high moisture level around the eggs.[19] As the gelatinous matrix ages, it becomes tanned, turning from a sticky, colourless jelly to an orange-brown substance which appears layered.[20]

Egg formation and development[edit]

Egg formation in M. javanica has been studied in detail,[21] and is similar to egg formation in the well studied, free-living nematode Caenorhabditis elegans.[22] Embryogenesis has also been studied, and the stages of development are easily identifiable with a phase contrast microscope following preparation of an egg mass squash. The egg is formed as one cell, with two-cell, four-cell and eight-cell stages recognisable. Further cell division leads to the tadpole stage, with further elongation resulting in the first stage juvenile, which is roughly four times as long as the egg. The J1 stage of C. elegans has 558 cells, and the J1 of M. javanica likely has a similar number, since all nematodes are morphologically and anatomically similar.[22] The egg shell has three layers, with the vitelline layer outermost, then a chitinous layer and a lipid layer innermost.

Egg hatching[edit]

Preceded by induced changes in eggshell permeability, hatching may involve physical and/or enzymatic processes in plant-parasitic nematodes.[23]Cyst nematodes, such as Globodera rostochiensis, may require a specific signal from the root exudates of the host to trigger hatching. Root-knot nematodes are generally unaffected by the presence of a host, but hatch freely at the appropriate temperature when water is available. However, in an egg mass or cyst, not all eggs will hatch when the conditions are optimal for their particular species, leaving some eggs to hatch at a later date. Ammoniumions have been shown to inhibit hatching and to reduce the plant-penetration ability of M. incognita juveniles that do hatch.[24]

Reproduction[edit]

Root-knot nematodes exhibit a range of reproductive modes, including sexuality (amphimixis), facultative sexuality, meiotic parthenogenesis (automixis) and mitotic parthenogenesis (apomixis).

Species[edit]

References[edit]

  1. ^Sasser JN, Carter CC: Overview of the International Meloidogyne Project 1975–1984. In An Advanced Treatise on Meloidogyne. Edited by: Sasser JN, Carter CC. Raleigh: North Carolina State University Graphics; 1985:19-24.
  2. ^Moens, Maurice, Roland N Perry, and James L Starr. 2009. "Meloidogyne Species: a Diverse Group of Novel and Important Plant Parasites." In Root-knot Nematodes, ed. Roland N Perry, Maurice Moens, and James L Starr, 1–17. Wallingford, UK: CABI Publishing.
  3. ^ abcEisenback, J. D. & Triantaphyllou, H. H. 1991 Root-knot Nematodes: Meloidogyne species and races. In: Manual of Agricultural Nematology, W. R. Nickle. (Ed). Marcel Dekker, New York. pp 281 – 286.
  4. ^Stirling, G. R.; Stanton, J. M.; Marshall, J. W. (1992). "The importance of plant-parasitic nematodes to Australian and New Zealand agriculture". Australasian Plant Pathology. 21: 104–115. doi:10.1071/app9920104. 
  5. ^Neal, J. C. 1889. The root-knot disease of the peach, orange and other plants in Florida due to the work of Anguillula Bull. I.S. Bur. Ent.20.31pp.
  6. ^Jatala, P., bridge, J. 1990. Nematode parasites of root and tuber crops. In Plant parasitic nematodes in sub-tropical and tropical agriculture., pp 137-180. Luc, M., Sikora, R.A., Bridge, J., CABI Publishing, Wallingford, UK.
  7. ^Makumbi-kidza, N. N., Speijer and Sikora R. A. 2000. Effects of Meloidogyne incognita on Growth and Storage-Root Formation of Cassava (Manihot esculenta). J Nematol.; 32(4S): 475–477.
  8. ^ abGapasin, R.M. 1980. Reaction of golden yellow cassava to Meloidogyne spp. Inoculation. Annals of Tropical Research 2:49-53. .
  9. ^Theberge, R. L. (eds). 1985. Common African Pests and Diseases of cassava, Yam, Sweet Potato and Cocoyam. International Institute of Tropical Agriculture (IITA). Ibadan, Nigeria 107 p.
  10. ^Charles, Lauren; Carbone, Ignazio; Davies, Keith G.; Bird, David; Burke, Mark; Kerry, Brian R.; Opperman, Charles H. "Phylogenetic Analysis of Pasteuria penetrans by Use of Multiple Genetic Loci". Journal of Bacteriology. 187 (August 2005): 5700–5708. doi:10.1128/JB.187.16.5700-5708.2005. PMC 1196054. PMID 16077116. Retrieved 2014-09-13. 
  11. ^Dama, L.B.; Poul, B.N.; Jadhav, B.V.; Hafeez, M.D. (1999). "Effect of Herbal "Juglone" on Development of the plant parasitic nematode (Meloidogyne Spp.) on Arachis hypogaea". Journal of Ecotoxicology and Environmental Monitoring. 9: 73–76. 
  12. ^Sijmons, P. C.; Grundler, F. M. W.; von Mende, N.; Burrows, P. R.; Wyss, U. (1991). "Arabidopsis thalliana as a new model host for plant-parasitic nematodes". The Plant Journal. 1: 245–254. doi:10.1111/j.1365-313x.1991.00245.x. 
  13. ^Wyss, U., Grundler, F.M.W. & Munch, A. 1992 The parasitic behaviour of second stage juveniles of Meloidogyne incognita in roots of Arabidopsis thaliana. Nematologica, 38, 98 - 111.
  14. ^Hussey, R. S. & Grundler, F. M. W. 1998 Nematode parasitism of plants. In: The Physiology and Biochemistry of free-living and plant-parasitic nematodes. Perry, R. N. & Wright, D. J. (Eds), CABI Publishing, UK. pp 213 – 243.
  15. ^Sijmons, P. C.; Atkinson, H. J.; Wyss, U. (1994). "Parasitic strategies of root nematodes and associated host cell responses". Annual Review of Phytopathology. 32: 235–259. doi:10.1146/annurev.phyto.32.1.235. 
  16. ^Madulu, J. & Trudgill, D. L. 1994 Influence of temperature on Meloidogyne javanica. Nematologica, 40, 230 - 243.
  17. ^Trudgill, D. L. 1995 An assessment of the relevance of thermal time relationships to nematology. Fundamental and Applied Nematology, 18, 407 - 417.
  18. ^Maggenti, A. R. & Allen, M. W. 1960 The origin of the gelatinous matrix in Meloidogyne. Proceedings of the Helminthological Society of Washington, 27, 4 - 10.
  19. ^Wallace, H. R. 1968 The influence of soil moisture on survival and hatch of Meloidogyne javanica. Nematologica, 14, 231-242.
  20. ^Bird, A. F. 1958 The adult female cuticle and egg sac of the genus Meloidogyne Goeldi, 1887. Nematologica, 3, 205 - 212.
  21. ^McClure, M. A.; Bird, A. F. (1976). "The tylenchid (Nematoda) egg shell: formation of the egg shell in Meloidogyne javanica". Parasitology. 72: 29–39. doi:10.1017/s003118200004316x. 
  22. ^ abWood, W. B. 1988 Introduction to C.elegans. In::The Nematode Caenorhabditis elegans, W. B. Wood (Ed), Cold Spring Harbour Laboratory, New York. pp 1 – 16.
  23. ^Norton, D. C. & Niblack, T. L. 1991 Biology and ecology of nematodes. In: Manual of Agricultural Nematology, Nickle, W. R. (Ed), Marcel Dekker, New York. pp 47 – 68.
  24. ^Surdiman; Webster, J. M. (1995). "Effect of ammonium ions on egg hatching and second-stage juveniles of Meloidogyne incognita in axenic tomato root culture". Journal of Nematology. 27: 346–352. 

External links[edit]

0 thoughts on “An Essay On Plant Pathogenic Nematodes Life”

    -->

Leave a Comment

Your email address will not be published. Required fields are marked *