Nematophagous Fungi

Egg-parasitic fungi P32 32 6.2 8.5

Ver112 32 - 10 Chitinases/chitosanases

P. rubescens CHI43 43 7.6

P. chlamydosporia CHI43 43 7.9

References

Tunlid, Rosen, Ek, and Rask (1995) Ähman et al (1996) Zhao, Mo, and Zhang (2004)

R.B. Wang, Yang, Lin, Y. Zhang, and K.Q. Zhang (2006)

Lopez-Llorca (1990)

Olivares-Bernabeu

Lopez-Llorca and Robertson (1992b) Segers, Butt, Kerry, and Peberdy (1994); Segers, Butt, Keen, Kerry, and Peberdy (1995) Bonants et al. (1995)

5.2-5.7 Tikhonov et al (2002) 5.2-5.7 Tikhonov et al (2002) 6 Chen, Cheng, Huang, and Li (2005)

Another serine protease from A. oligospora (Aoz1), with a molecular mass of 38 kDa showing 97% homology with PII was recently described (Zhao et al., 2004). Other serine proteases have been isolated and characterized from the nematode-trapping fungi Arthrobotrys (syn. Monacrosporium) microscaphoides designated Mlx (M. Wang et al., 2006) and Arthrobortys (syn. Dactylella) shizishanna (Ds1) (R. B. Wang et al., 2006) both showing high homology with the A. oligospora serine proteases (M. Wang et al., 2006).

Nematode eggshells mostly contain protein and chitin (Clarke, Cox, & Shepherd, 1967) organized in a microfibrillar and amorphous structure (Wharton, 1980). Therefore, a search for extracellular enzymes degrading those polymers was carried out. A 32 kDa serine protease (P32) was first purified and characterized from the egg parasite P. rubescens (Lopez-Llorca, 1990). Involvement of the enzyme in pathogenesis was suggested by quick in vitro degradation (Fig. 3i) of Globodera pallida egg shell proteins (Lopez-Llorca, 1990), but most of all by its immunolocalization (Fig. 3f, 3g) in appressoria of the fungus infecting Heterodera schachtii eggs (Lopez-Llorca & Robertson, 1992b).

Although pathogenesis is a complex process involving many factors, inhibition of P32 with chemicals and polyclonal antibodies reduced egg infection and penetration (Lopez-Llorca et al., 2002b). The similar species P. chlamydosporia also produces an extracellular protease (VcP1) (Segers et al., 1994) which is immunologically related to P32 and similar enzymes from entomopathogenic fungi (Segers et al., 1995). VcP1-treated eggs were more easily infected than untreated eggs, suggesting a role of the enzyme in eggshell penetration by egg-parasitic fungi.

Recently a serine protease (Ver112) was isolated and characterized from Lecanicillium psalliotae showing similarities with the Arthrobotrys proteases (PII and Aoz1) of ca 40%, and ca 60% homology with serine proteases of egg-parasitic fungi (Yang et al., 2005a, 2005b).

Other proteases from nematophagous fungi have been partly characterized, e.g. a chymotrypsin-like protease from conidia of the endoparasite D. coniospora (Jansson & Friman, 1999), and a collagenase produced by the nematode-trapping Arthrobotrys tortor (Tosi, Annovazzi, Tosi, Iadarola, & Caretta, 2001). Non-nematophagous fungi such as the mycoparasites Trichoderma harzianum and Clonostachys rosea (syn. Gliocladium roseum) are also sources of serine proteases with nematicidal activity (Suarez, Rey, Castillo, Monte, & Llobell, 2004; Li, Yang, Huang, & Zhang, 2006).

Several chitinolytic enzymes of Pochonia rubescens and P. chlamydosporia have been detected. One of those accounting for most of the activity was a 43 kDa endochitinase (CHI43) (Tikhonov et al., 2002). When G. pallida eggs were treated with both P32 and CHI43 damage to eggshell was more extensive than with each enzyme alone, suggesting a cooperative effect of both enzymes to degrade egg shells (Tikhonov et al., 2002). Recently a chitosanase was isolated and characterized from the egg-parasitic fungus P. lilacinus (Chen et al., 2005).

3.4. Fungal Pathogen Genomics andProteomics

In the era of genomics, fungal pathogens are suitable candidates for the analysis under this new paradigm in modern biology. In the dawn of fungal pathogen genomics under the Fungal Genome Initiative, important fungal pathogens have been or are being sequenced (Xu, Peng, Dickman, & Sharon, 2006). A direct bonus is the finding of unique fungal genes and characterization of genome structure and function. Available gene predictions in genomes of fungal plant pathogens indicate 30% of no homologues. This situation, which could be similar in nematophagous fungi, indicates that new fungal genes or gene products (e.g. proteins of unknown function) can soon be discovered.

The re-evaluation of the study of fungal pathogenicity-related genes with a genomic approach is underway. One example is appressorium development. This awaits to be applied in nematophagous fungi. Signalling/reception are other fields which will follow.

Proteomic approaches complement genomics. There are expression, localization and interactions, which are unique to this global strategy. Our preliminary results indicate that plant-host fungal invertebrate pathogen "crosstalk" can be approached this way

The assembly of the Fungal Tree of Life project (Spatafora, 2005; Kuramae, Robert, Snel, Weiss, & Boekhout, 2006) which is at a very advanced stage, could represent a useful tool for deciding on how to proceed to establish genomic approaches. EST approaches to understand the pathogenicity of nematophagous fungi are already being used (Ahren et al., 2005).

4. SOIL AND RHIZOSPHERE ENVIRONMENT 4.1. Activities in Soil

Nematophagous fungi are generally regarded as soil organisms (Dackman, Jansson, & Nordbring-Hertz, 1992), although there are reports on their frequent occurrence also in aquatic environments, especially in shallow, unpolluted water (Hao, Mo, Su, & Zhang, 2005). Most nematophagous fungi can live saprophytically in soil, but in presence of a host they change from a saprophytic to a parasitic stage. The exact mechanism behind this is not known. Nematophagous fungi inhabit soil pores where infection structures are formed and nematodes are captured (Fig. 5). The zoosporic fungi are obviously dependent of soil water films for their function.

When nematophagous species have to be applied to manage plant parasitic nematodes they have to be delivered to soil. Several approaches for introducing them have been used (see Stirling, 1991), but very little efforts have been paid to follow the fate of nematophagous fungi in the soil/rhizosphere environment, after their release.

Nematophagous fungi grow in almost all types of soil, but are generally regarded as being more frequent in soils with high organic matter (Duddington, 1962). Generally, they have few nutritional and vitamin requirements for growth, and hence are ubiquitous. Additions of glucose (Cooke, 1962) and chopped organic matter, e.g. grass (Duddington, 1962) increased activity of nematode-trapping species. This effect was probably due to an increase in the numbers of microbivorous nematodes. Arthrobotrys spp. have a teleomorph in Orbilia, which are weak wood decomposers (Pfister, 1997), and the wood decomposing Pleurotus spp. suggests that decomposition of wood may be an important supply of carbon and energy for the fungi. Capturing nematodes may hence support the fungi with nitrogen (Barron, 1992). In Petri dishes and sterilized microcosms there is a heavy reduction of nematodes due to nematophagous fungi (Jansson, 1982b), and a density dependence relationship exists between nematodes and endoparasites (Jaffee, Gaspard, & Ferris, 1989).

In field soil, there is no clear correlation between nematophagous fungi and nematodes (Persmark, Banck, & Jansson, 1996a) and nematode-trapping fungi are known to be sensitive to soil mycostasis (Cooke & Satchuthananthavale, 1968), as well as to feeding by soil enchytraeids (Jaffee, 1999).

Dactylella Ellipsospora Nematodes

Figure 5. Low temperature scanning electron micrographs (LTSEM) of nematophagous fungi in soil. (a) Conidiophores with conidia of the nematode-trapping fungus Arthrobortys superba (bar = 100 ¡im). (b) Constricting ring traps of Drechslerella dactyloides (bar = 50 ¡m). (c) Nematode captured in constricting ring of D. dactyloides (bar = 50 ¡m). From Jansson, Persson, and Odselius (2000), courtesy of Mycological Society of America.

Figure 5. Low temperature scanning electron micrographs (LTSEM) of nematophagous fungi in soil. (a) Conidiophores with conidia of the nematode-trapping fungus Arthrobortys superba (bar = 100 ¡im). (b) Constricting ring traps of Drechslerella dactyloides (bar = 50 ¡m). (c) Nematode captured in constricting ring of D. dactyloides (bar = 50 ¡m). From Jansson, Persson, and Odselius (2000), courtesy of Mycological Society of America.

Introduction of nematophagous fungi, and most microbial biocontrol agents, to soil has been problematic due to both biotic and abiotic factors. Biocontrol experiments using the egg-parasite P. chlamydosporia showed low control efficiency against root-knot nematodes, and furthermore, the fungus was detected at very low rates, mainly in the rhizosphere of the test plants (Verdejo-Lucas, Sorribas, Ornat, & Galeano, 2003). One of the reasons for this may be that the soil was not receptive to the fungus.

We have used an in vitro assay to be able to easily study soil receptivity for nematophagous fungi (Monfort, Lopez-Llorca, Jansson, & Salinas, 2006). Using a soil-membrane technique 0, 25, 50, 75 and 100% sterilized soil was inoculated with several isolates of the nematophagous fungi P. chlamydosporia and P. lilacinus. After 4 weeks, colony radius was measured (expressed as relative growth) as well as hyphal density on the membrane placed on top of the soils.

When comparing two sandy soils (Spanish and Australian) with similar physico-chemical properties, large differences between the receptivity to the fungi were found, both regarding isolates as well as between soils. For instance, an Australian isolate of P. chlamydosporia was most inhibited in the Spanish soil, but the least inhibited in the Australian soil. The result suggests that a soil can be more receptive to indigenous isolates than to non-indigenous ones.

4.2. Nematophagous Fungi as Root Endophytes

Since nearly all plant-parasitic nematodes attack plant roots, the rhizosphere biology of nematophagous fungi is important from the point of view of a biological control strategy. Nematode-trapping fungi (Peterson & Katznelson, 1965; Gaspard & Mankau, 1986; Persmark & Jansson, 1997) and egg-parasitic fungi (Bourne, Kerry, & De Leij, 1996; Kerry, 2000) have been found to be more frequent in the rhizosphere than in the bulk soil.

External root colonisation varies between plant species. The pea rhizosphere harboured by far the highest frequency and diversity of nematode-trapping fungi compared to other plant species tested (Persmark & Jansson, 1997). In an investigation on chemotropic growth of nematophagous fungi towards roots of several plants, only isolates of A. oligospora were attracted (Bordallo et al., 2002). In a 3-month pot experiment, Dactylellina ellipsospora (syn. Monacrosporium ellipsosporum) and D. dactyloides were especially competent in colonising tomato roots (Persson & Jansson, 1999).

Several nematode-trapping fungi are able to form so-called conidial traps in response to roots and root exudates (Persmark & Nordbring-Hertz, 1997). The external root colonisation by the egg-parasite Pochonia chlamydosporia also varied with plant species and was increased when plants were infected with the root-knot nematode Meloidogyne incognita (Bourne et al., 1996). This effect is possibly due to increased leakage of root exudates after damage to the root surface by the nematodes.

In recent investigations we studied the endophytic root colonization of the four groups of nematophagous species. The nematode-trapping species A. oligospora, D. dactyloides (Figs. 6a, b), and N. robustus (Figs. 6b, c) were all capable of endophytic colonization of barley roots. Similar root colonization was also detected for the egg-parasite P. chlamydosporia (Figs. 6e, f) and the toxin-producing P. djamor. The only fungi which did not show root colonization were the endoparasitic fungi H. rhossiliensis and N. pachysporus (Lopez-Llorca, Bordallo, Salinas, Monfort, & Lopez-Serna., 2002a; Bordallo et al., 2002; Lopez-Llorca, Jansson, Macia Vicente, & Salinas, 2006). The fungi grew inter- and intracellularly, formed appressoria when penetrating plant cell walls of epidermis and cortex cells, but never entered vascular tissues (Lopez-Llorca et al., 2002a; Bordallo et al., 2002). In contrast to Pochonia spp., appressoria had never been observed previously in A. oligospora.

Using histochemical stains it was possible to reveal the plant defence reactions, e.g. papillae and other cell wall appositions induced by nematophagous fungi, but these never prevented root colonization. Nematophagous fungi grew extensively especially in monocotyledon plants producing abundant mycelia, conidia and chlamydospores. Necrotic areas of the roots were observed at initial stages of colonization by the nematode-trapping and toxin-producing fungi tested, but were never seen at later stages, even when the fungi proliferated in epidermal and cortical cells.

Nematophagous Fungi

Figure 6. Parasitic (a, c, e, g) vs. endophytic (b, d, f, h) behaviour of nematophagous fungi. (a) Conidial trap of Drechslerella sp. (c) Mycelia of a Nematoctonus sp. showing an "hour glass" trapping device and clamp connections (arrows). (e) Nematode egg infected by Pochonia sp. (g) Hyphae and toxin-producing organ of Pleurotus sp. (b, d, f, h). Display of endophytic colonisation of barley cortex cells by the nematophagous fungi displayed on the left hand side of each picture. Scale bars: a = 25 fim; b, d, h = 15 fim; c = 2 fim; e = 10 fim; f = 30 fim; g = 1 fim. (a and c: C. Olivares-Bernabeu, unpublished; b, d, h: from Lopez-Llorca et al., 2006, courtesy of Springer; e: from Lopez-Llorca et al., 2002b, courtesy of Elesevier; f: from Bordallo et al., 2002, courtesy of the New Phytologist Trust; g: from Nordbring-Hertz et al., 1995, courtesy of IWF Wissen und Medien, Göttingen).

Figure 6. Parasitic (a, c, e, g) vs. endophytic (b, d, f, h) behaviour of nematophagous fungi. (a) Conidial trap of Drechslerella sp. (c) Mycelia of a Nematoctonus sp. showing an "hour glass" trapping device and clamp connections (arrows). (e) Nematode egg infected by Pochonia sp. (g) Hyphae and toxin-producing organ of Pleurotus sp. (b, d, f, h). Display of endophytic colonisation of barley cortex cells by the nematophagous fungi displayed on the left hand side of each picture. Scale bars: a = 25 fim; b, d, h = 15 fim; c = 2 fim; e = 10 fim; f = 30 fim; g = 1 fim. (a and c: C. Olivares-Bernabeu, unpublished; b, d, h: from Lopez-Llorca et al., 2006, courtesy of Springer; e: from Lopez-Llorca et al., 2002b, courtesy of Elesevier; f: from Bordallo et al., 2002, courtesy of the New Phytologist Trust; g: from Nordbring-Hertz et al., 1995, courtesy of IWF Wissen und Medien, Göttingen).

In cereal roots proceeding from soils naturally infested with the cereal cyst nematode Heterodera avenae and Pochonia spp., either the syncytia induced by the nematode and fungal hyphae could be detected inside the roots (Fig. 7c). Abundant sporulation of Pochonia spp. was also observed on the root surface (Fig. 7 a, b). The results at least indicate the possibility that nematode infection by the fungus may occurr inside roots, although so far this event has not been observed.

Actually, it is unknown whether endophytic colonization induces systemic resistance to nematodes and/or plant pathogens in plants. We have found that P. chlamydosporia could reduce growth of the plant pathogenic fungus Gaeumannomyces graminis var. tritici (take-all fungus, Ggt) in dual culture Petri dish and in growth tube experiments. In pot experiments P. chlamydosporia increased plant growth whether Ggt was present in the roots or not, suggesting a growth promoting effect by P. chlamydosporia (Monfort et al., 2005).

Endophytic rhizobacteria reducing plant-parasitic nematodes have been described (Hallmann, Quadt-Hallmann, Miller, Sikora, & Lindow, 2001), as well as the reduction of root knot nematodes by arbuscular mycorrhizal fungi (Waecke, Waudo, & Sikora, 2001). If this is true also in nematophagous fungi this will open up a new area of biocontrol using these fungi. The endophytic root colonization by egg-parasitic fungi, e.g. Pochonia spp., may provide them an opportunity to infect eggs of economically important endoparasitic nematodes (e.g. cyst and root-knot species) inside the roots and to reduce subsequent spread and roots infection by the second generation of juveniles.

Structures resembling trapping organs were observed in epidermal cells colonized by A. oligospora, and these may serve the purpose of trapping newly hatched juveniles escaping the roots. The ability to colonize plant roots may also be a survival strategy of these fungi and could explain soil suppressiveness to plant-parasitic nematodes in nature. The colonization of plant roots by nematophagous fungi is a new area of research that deserves in-depth investigations, not the least for biocontrol purposes and is presently underway in our laboratory.

4.3. Rhizosphere Dynamics and Biocontrol

The rhizosphere is a microecosystem in which roots release nutrients which in turn will affect microbes and their grazers. The former will modify these nutrients and could affect root and plant development. In this complex scenario, nematophagous fungi are both "hunters" and "hunted" since they predate on nematodes and can be affected, for instance, by myceliophagous species. It is tempting to use a combination of current non-destructive methods to analyse dynamics of the biotic component of the rhizosphere. Modification, or engineering, of the rhizosphere resource exchange could be vital for modifying the endophytic behaviour of nematophagous fungi. This may in turn affect their capability to control root diseases. Recently, microbiosensors, i.e. hybrids of soil sensors and

Nematophagous Fungi

Figure 7. Rhizosphere colonization by fungal egg parasites in nematode suppressive soils. (a) Profuse hyphal growth and sporulation (LTSEM) in oat rhizosphere. (b) Close-up of phialides and slimy conidia of Pochonia spp. (c) Field emission scanning electron microscopy (FESEM) of longitudinal section through a cereal root infected by the nematode Heterodera avenae, showing syncytia (S) and fungal colonization (arrowheads) in root cortex cells (a: Lopez-Llorca & Duncan, 1988; b,c: Lopez-Llorca & Claugher, unpublished).

Figure 7. Rhizosphere colonization by fungal egg parasites in nematode suppressive soils. (a) Profuse hyphal growth and sporulation (LTSEM) in oat rhizosphere. (b) Close-up of phialides and slimy conidia of Pochonia spp. (c) Field emission scanning electron microscopy (FESEM) of longitudinal section through a cereal root infected by the nematode Heterodera avenae, showing syncytia (S) and fungal colonization (arrowheads) in root cortex cells (a: Lopez-Llorca & Duncan, 1988; b,c: Lopez-Llorca & Claugher, unpublished).

molecular methods for rhizosphere studies, have been devised (Cardon & Gage, 2006). These are genetically engineered bioreporter bacteria which join reporter genes, e.g. GFP and Lux, with promoters induced by several rhizosphere conditions (starvation, contaminants, quorum sensing). These are timely approaches for global studies on general rhizosphere function in ecosystems. Some of these bioreporters are biocontrol bacteria. Biocontrol fungi, e.g. nematophagous, are next on the list.

4.4. Root Exudates

To this point it is clear that the biocontrol scenario of plant-parasitic nematodes by nematophagous fungi relies on a multitrophic interaction in which plant roots play an important role. There is also abundant scientific evidence that roots produce compounds (exudates) which mediate plant-plant and plant-microbe interactions (Bais, Weir, Perry, Gilroy, & Vivanco, 2006). The latter would also include plant-nematode (and other micro- and meso-fauna) interactions.

Root exudates are very diverse structurally and chemically, and vary among plant species, but above all they may influence a wide array of processes relevant to the biocontrol action of nematophagous fungi. Leaving aside the effect of root exudates on nematode feeding and colonization, these compounds can influence nutrient availability in the rhizosphere (e.g. siderophores). Root exudates can also elicit release of compounds which could act in root defence or mediate signalling processes.

Root exudates also mediate plant-microbe interactions. The role of flavonoids on the specificity of rhizobia-Leguminosae interactions is well established (Perret, Staehelin, & Broughton, 2000). These root exudates induce the expression of rhizobia Nod genes, which are then involved in the synthesis of Nod factors (lipochitino-oligosaccharides with diverse chemical modifications) that are recognized by the appropriate host plant.

Closer to nematophagous fungi, arbuscular mycorrhizal fungi (AMF) recognize the presence of a compatible host plant through root exudates. A sesquiterpene has been identified as a branch-inducing factor for AMF in legumes (Akiyama, Matsuzaki, & Hayashi, 2005). Hyphal morphogenesis is vital for successful AMF-root colonization. This aspect may also be important in nematophagous fungi.

Root exudates affect nematodes, especially microbivorous species. On the other hand, plant-parasitic nematodes increase production of root exudates (rhizodeposition). The quality of root exudates is also changed. C/N-ratio in particular can alter the trophic stage of the fungus Rhizoctonia solani and turn it into a root pathogen (Van Gundy, Kirkpatrick, & Golden, 1977). These effects of root exudation on nematophagous fungi remain largely unknown, but are worth investigating.

There are new evidences that tri-trophic webs can be established in the rhizosphere leading to benefits for the plant host. Plant roots produce exudates which attract nematodes (Green, 1971). These can act as vectors of rhizobia that are thus transferred to roots (Horiuchi, Prithiviraj, Bais, Kimball, & Vivanco,

2005). It is also known that nematodes are attracted to nematophagous fungi to various extents (Jansson & Nordbring-Hertz, 1979; Jansson, 1982a). The role of non-parasitic nematodes as vectors to inoculate nematophagous fungi or root endophytes in nature has not yet been investigated.

4.5. Detection and Quantification

It is vital to be able to detect and quantify biocontrol agents, e.g. nematophagous fungi, in soil and rhizospheres, in the period following their addition. Many techniques for this purpose have been too unspecific or difficult to perform (Jansson, 2001). Antibodies have been tried with little success due to cross-reactions with other fungi (Eren & Pramer, 1966). Molecular markers such as the GUS gene have been transformed to A. oligospora (Persmark, Persson, & Jansson, 1996b; Tunlid, Ahman, & Oliver, 1999) and the GFP gene has been transformed to P. chlamydosporia (Atkins, Mauchline, Kerry, & Hirsch, 2004). In the former case it was not possible to quantify the growth of the fungus in soil at sufficiently low levels (Persmark et al., 1996b). The problem with P. chlamydosporia was to obtain stable transformants. A possible solution could be to try Agrobacterium-mediated transformation (Michielse, Arentshorst, Ram, & Van den Hondel, 2005).

Another promising approach is to use PCR-based techniques in combination with fluorogenic probes (e.g. scorpions and beacons). Such methods using realtime PCR and primers based on ITS sequences of P. chlamydosporia and Paecilomyces lilacinus have recently been presented (Ciancio, Loffredo, Paradies, Turturo, & Finetti Sialer, 2005; Atkins, Clark, Pande, Hirsch, & Kerry, 2005).

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