Compatibility of FPI with Natural Compounds Chitosan as a Case Study

FPI have to break their hosts outer barriers for infection. These barriers, as we have discussed (see Sect. 9.2.3) mostly include a chitin-protein structure (e.g. nematode egg-shell, insect cuticle). Furthermore chitin is the second most abundant polymer in nature after cellulose (Cohen-Kupiec and Chet 1998) and occurs in various organisms (e.g. crustaceans, insects, nematodes and most fungi). Chitin waste is an abundant by-product of the crustacean fishing industry worldwide. This waste product has been a target for the development of organic nematicides. The rationale behind this was that a chitin amendment to soil would enhance the chitinolytic microbiota, which would in turn be nematophagous, in view of the biochemical composition of nematode barriers. This resulted in the isolation of nematophagous organisms, including new species of bacteria (Spiegel et al. 1986, 1987, 1988). The experiments of soil organic amendments for biocontrol of soil-borne pathogens have produced various results with the development of composting, and their performance has improved (Trillas et al. 2002, 2006). However, addition of chitin rich waste to soil for nematode control resulted in a build up of ammonia in quantities which may turn to be phytotoxic (Rodriguez-Kabana et al. 1987; Carvajal and Rodriguez-Kabana 1998; Hallmann et al. 1999).

Chitosan is obtained by chitin deacetylation, and consists of polymers of b-1,4-glucosamine subunits, with molecular weight up to 400 kDa (Rabea et al. 2003). It can be produced by chemical or microbial/enzymatic treatments (Tsai et al 2002). Besides, fungi can turn chitin in their cell walls to chitosan. This is achieved by means of chitin deacetylase activity and may block host recognition and degradation of fungal cell walls by plant chitinases in biotrophic fungi (Deising et al. 1995). Commercial chitosan is mainly produced by chemical methods using chitin waste products from seafood industry (Kumar 2000).

Chitosan has several advantages over chitin for combination with biocontrol agents purposes, the main ones are a higher solubility and interesting biological properties. This polymer is no toxic to mammals (Dodane and Vilivalam 1998; Lee et al. 2004) and it elicits plant defence mechanisms (Benhamou et al. 1994; Lafontaine and Benhamou 1996; Ait Barka et al. 2004; Trotel-Aziz et al. 2006), but displays antibiotic activity against microorganisms, both bacteria (Liu et al. 2001b, 2004; Tikhonov et al. 2006) and fungi (Bell et al. 1998; Laflamme et al. 1999; Park et al. 2002; Pascencia-Jatomea et al. 2003; Bautista-Banos et al. 2006; Palma-Guerrero et al. 2008). We have investigated its compatibility with biocontrol agents and are extending these results to the biological control of plant parasitic nematodes.

The study of the fungitoxic effect of chitosan has mostly dealt with colony growth inhibition of plant-pathogenic fungi and associated ultrastructural changes in the hyphae (Laflamme et al. 1999). Much less is known about the effects of chitosan on spore germination. Palma-Guerrero et al. (2008), were the first in comparing the effect of chitosan on biocontrol fungi and plant-pathogenic fungi, considering both hyphal growth and spore germination. Great variations in tolerance to chitosan between the different fungi tested were obtained. Most plant pathogenic fungi tested were highly sensitive to chitosan, whereas nematophagous and ento-mopathogenic fungi were much less inhibited by chitosan. Mycoparasitic fungi were the only exception among the three types of biocontrol fungi tested, since they were as sensitive to chitosan as the plant pathogenic fungi. The low effect of chitosan on nematophagous and entomopathogenic fungi seemed to be related, at least partly, with their ability to degrade chitosan. These fungi showed the highest chitosan-degrading activity, according to the size of their substrate degradation halos observed in amended solid growth media.

Conidial germination was more sensitive to chitosan than hyphal growth. Conidia of nematophagous and entomopathogenic fungi were again the least sensitive to chitosan. For some isolates, germination in the presence of chitosan was similar to that of untreated controls. Furthermore, germination was not completely inhibited by increasing chitosan concentration. Only one P. chlamydosporia isolate (P.c. 4624) out of 9 tested showed the same sensitivity to chitosan as the plant pathogenic fungi tested. In fact this was the nematophagous fungus strain with the lowest chitosanase activity.

Nematophagous fungi showed the highest ability to degrade chitosan. This was confirmed for several P. chlamydosporia isolates tested, from worldwide origins. One possible explanation to the little effect of chitosan on the growth of nematophagous and entomopathogenic fungi may be that their chitosan degrading enzymes are more abundant and/or efficient. They could prevent diminish the toxic effect of chitosan to fungal cells. These fungi may even be using chitosan as nutrient in some cases. Although chitinolytic and chitosanolytic activities have been found in plant pathogenic and mycoparasitic fungi (Shimosaka et al. 1993; Nogawa et al. 1998), Palma-Guerrero et al. (2008) could not detect halos of chi-tosan degradation for the plant pathogenic fungi or oomycetes tested (except for V. dahliae). This would suggest low chitosanase activities for these organisms. The "special" chitosan degrading enzymes of nematophagous and entomopatho-genic fungi may be related with their multimodal lifestyle as explained above (Sect. 9.2.3). In fact both fungal groups have coevolved to degrade their host cuticles which contain chitin, a similar polymer to chitosan (Palma-Guerrero et al. 2010).

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