Population Dynamics of a Nematophagous Fungus Lecanicillium muscarium, and Root Knot Nematode, Meloidogyne incognita to Assess the Disease Pressure and its Management

Manzoor Hussain*, Miloslav Zouhar and Pavel Ryšánek

Department of Plant Protection, Faculty of Agrobiology, Food and Natural Resources, Czech University of Life Sciences, Prague, Kamýcká 961/129, 165 00 Praha 6, Suchdol, Czech Republic

ABSTRACT

A series of experiments were conducted in a greenhouse to determine the population dynamics of the root knot nematode, Meloidogyne incognita, and the nematophagous fungus, Lecanicillium muscarium, on tomato. The nematode population densities were measured by determining the number of galls and egg masses, juveniles and eggs per root system, and reproduction factor. Four initial populations (Pi) (500, 1000, 1500, and 2000 eggs) mixed with antagonistic fungal conidia levels (1×103, 1×104, 1×105, 1×106) were used during the experiments. The roots were stained with Phloxine B, the egg-masses were quantified, the root systems were rated for galling, and the egg masses were measured on a 0 to 5 scale, where 0 = no gall or egg masses, 1 = 1-2, 2 = 3-10, 3 = 11-30, 4 = 31-100, and 5 = >100 galls or egg masses per root system. The nematode reproduction rate (Pf/Pi, where Pf = final nematode population / initial nematode population) decreased proportionately with the increased initial fungal conidial Pi. Foliage growth was directly related to fungal Pi and inversely to nematode Pi. Our results showed that the higher Pi of L. muscarium was associated with the lower Pf of M. incognita. Foliage growth increased with increased fungus inoculum.

Article Information

Received 23 July 2016

Revised 06 August 2016

Accepted 21 August 2016

Available online 15 December 2016

Authors’ Contributions

MH designed experiments, collected data and wrote article, MZ and PR advised the study and proofread article.

Key words

Lecanicillum muscarium, Meloidogyne incognita, Reproduction factor, Tomato, Nematode population.

* Corresponding author: manzoor.sahi@gmail.com

0030-9923/2017/0001-0207 $ 9.00/0

Copyright 2017 Zoological Society of Pakistan

DOI: http://dx.doi.org/10.17582/journal.pjz/2017.49.1.207.214

 

Introduction

Plant parasitic nematodes constitute the most abundant and successful animal phylum (Boucher and Lambshed, 1994) and cause great economic losses to agricultural crops and forestry worldwide (Sasser and Freckman, 1987; Siddiqui and Mahmood, 1996; Li et al., 2007; Zasada et al., 2008; Anwar and McKenry, 2010; Renčo et al., 2012; Renčo, 2013). Among plant parasitic nematodes, the root knot nematodes (Meloidogyne spp.) are considered the most dominant and extensive plant pathogens, and they attack a wide range of host plants including field crops, vegetables, fruit trees and ornamentals (Regaieg et al., 2010). In total, 2000 host plant species are prone to nematode infection, which causes an approximate 5% loss worldwide (Hussey and Janssen, 2002). The damage to the global crop production due to root knot nematodes is estimated to be approximately US$ 80 billion annually (Rodriguez and Canullo, 1992; Li et al., 2007). Root knot nematodes are sedentary endoparasites, polyphagous and considered a silent threat to agriculture. Meloidogyne species such as M. incognita and M. javanica are considered the most limiting factors for vegetative growth as well as harvest yield. The nematode second stage juvenile penetrates into plant roots and after a cascade of changes leads to successful infection by generating giant cells that ultimately result in root galls (Sharon et al., 2001). The galls reduce plant nutrient absorption, causing the plant with impaired growth to become stunted and chlorotic (Ellis et al., 2008). Root knot nematodes are not easy to control as they have a wide range of hosts, very short generation times, high production rate and endoparasitic nature (Manzanilla-Lopez et al., 2004). Concerns exist for the ecological and human health hazards as well as the breakage of resistance in cultivars due to the emergence of new pathogen races, and the usage of notorious chemicals is being restricted (Zuckerman and Esnard, 1994). Scientists are struggling to find other ways to get rid of these lethal chemicals that are carcinogenic.

The use of beneficial or antagonistic microorganisms is an alternative strategy to reduce the consumption of chemical pesticides for minimizing the parasitic nematode population on a threshold level (Stirling, 1991; Berg et al., 2005). Several fungi have been identified to reduce the nematode densities in soil that exhibit a range of antagonistic activities including a production of compounds that have nematicidal properties or parasitization of nematodes through prey devices or traps (adhesive knobs or nets, constricted and non-constricted rings) (Barron, 1977; Bird and Herd, 1995; Kerry, 2000; Lopez-Llorca and Jansson, 2006; Zouhar et al., 2013). Various studies have reported the production of nematicidal compounds by the fungi that become active against plant parasitic nematodes when they come in contact with each other (Anke et al., 1995; Anke, 2010; Hallmann and Sikora, 1996; Anke and Sterner, 1997; Chen et al., 2000; Meyer et al., 2000, 2004). The natural association among soil dwelling plant parasitic nematodes and fungi together in the soil rhizosphere keeps the nematode population low through the production of naturally produced toxic compounds and metabolites by fungi (Siddiqui and Mehmood, 1996).

Lecanicillium muscarium (ex. Verticillium lecanii) (Gams and Zare, 2001) is widely known as an entomopathogenic fungus (Cuthbertson et al., 2010) that also possesses mycoparasitic and nematicidal properties against food spoilage (Fenice et al., 1998a) and plant diseases (Ownley et al., 2010; Goettel et al., 2008). The fungus also has the capability to produce chitinolytic enzymes, including chitinases, glucanases and proteases (Fenice and Gooday, 2006; Fenice et al., 1998a, 2012) that have been shown to be more efficient than Trichoderma harzianum (Fenice et al., 1998b) against plant pathogens.

Lecanicillium spp. successfully parasitizes the eggs of M. incognita (Gan et al., 2007; Nguyen et al., 2007) and the females, cysts and eggs of Heterodera glycines under both lab and greenhouse conditions (Meyer and Meyer, 1996). Moreover, the studies claim that immature J2 are more susceptible to Lecanicillium spp. (Chen and Chen, 2003; Irving and Kerry, 1986; Kim and Riggs, 1991). To establish a successful infection to their hosts, L. muscarium conidia adhere to the host cuticle through mucilage and germinate, penetrate and produce blastospores inside the nematode eggs and J2. The fungus also produces various toxic secondary metabolites that induce resistance in plants in order to overcome other pathogens (Hirano et al., 2008).

In our previous studies we demonstrated that L. muscarium was able to reduce the population level of Meloidogyne hapla and Heterodera Schachtii under both lab and greenhouse conditions. The objective of the present study was to examine 1) whether the concentration of L. muscarium directly affects the nematode population and 2) which concentration best reduces the infestation of nematodes in the soil.

 

Materials and Methods

 

A nematode culture was prepared on a susceptible tomato variety, and the eggs were extracted from the roots using 0.05% NaOCl. The extracted eggs were gently washed with tap water to remove NaOCl (Hussey and Barker, 1973). M. incognita was identified by morphological characteristics (Eisenback, 1985). The desired inoculum density was prepared by stirring the egg suspension in distilled water. The inoculum density was prepared as 100 eggs per ml of water. Lecanicillium muscarium previously isolated from the egg mass of Meloidogyne incognita in Turkey was cultured and processed for conidia production. Fungus was maintained on potato dextrose agar (PDA). Conidia were obtained in deionized water containing 0.03­% Tween 80 and filtered through four layers of sterile cheesecloth to remove mycelium (Güçlü et al., 2010). Conidia were counted and standardized using a hemocytometer under a compound microscope.

The experiment was carried out with a susceptible variety of tomato i.e., Beril. The seeds of the tomato were surface sterilized in 0.05% NaOCl for 1 minute before nursery growth. After three weeks of being in the nursery, the plants were shifted to pots with a 5 x 6 cm dimension containing 500 cm3 of sterilized sandy loam soil (62% sand, 18% silt, and 20% clay). The pots were placed in a completely randomized design with five replications on a bench in a green house. The pots were irrigated after two-day intervals throughout the period of the study. The daily temperature ranged between 25-28°C. After planting, fungus conidia with different densities (1×103, 1×104, 1×105, and 1×106) were pipetted onto each pot. One week later, nematode eggs of different concentration (500, 1000, 1500 and 2000 eggs) were inoculated near the root zone of the plant in each pot. The plants without fungus inoculum served as the control.

After 60 days, the plants were removed from the pots and washed in water carefully. The washed roots were blotted on paper, dried, and weighed.

Data collection

The data were calculated for plant growth (top fresh weight and root weight) and nematode reproduction parameters (number of galls and egg masses, number of eggs per root system, second-stage juveniles (J2) per 100 cc of soil, and reproduction factor).

After counting the galls, the whole root system was stained with a 0.005% “Phloxine B” (Holbrook et al., 1983) solution for 30 minutes to facilitate the counting of egg masses. The eggs were obtained from the roots using a 0.5% NaOCl solution passed through a sieve with a pore size of 74 and 25 mm (Hussey and Barker, 1973). The total number of eggs and juveniles were extracted from the soil of each individual plant from their respective pots following the Whitehead and Hemming Tray Method (Whitehead and Hemming, 1965).

The root system was rated for galling and egg mass presence on a 0 to 5 scale (Lamberti, 1971), where 0 = no gall or egg masses, 1 = 1-2, 2 = 3-10, 3 = 11-30, 4 = 31-100, and 5 = >100 galls or egg masses per root system.

The extracted eggs were rinsed thoroughly in tap water and then counted at 40X magnification. Nematode reproduction was assessed by calculating the nematode reproduction rate as Pf / Pi, where Pi = initial inoculum level, and Pf = final population at harvest.

Statistical analysis

The experiment was repeated once in a greenhouse. All the data from two experiments were subjected to factorial analysis using Statistica 10.4 software.

 

Results

 

In our greenhouse trials approximately 60 days after nematode infestation, the initial population densities of nematophagous fungus, L. muscarium and M. incognita were regressed with the final population of M. incognita using the combined data from two consecutive experiments (Table I). The initial population densities of L. muscarium were significantly associated with the final population densities of M. incognita in terms of the second stage juveniles (J2) per root system, J2 per cm3 of soil, gall and egg masses

Table I.- Effect of graded concentration of nematophagous fungus, Lecanicillium muscarium on different inoculum densities of root knot nematode, M. incognita with respect to nematode reproduction factors.

Conc. of L. muscarium	Inoculum densities of M. incognita
	500	1000	1500	2000
Galls root system*
0	25.00 e	53.00 d	74.80 b	104.00 a
1 × 103	13.80 g	20.00 f	27.00 e	58.00 c
1 × 104	8.60 j-l	11.00 h-j	11.60 gh	21.40 f
1 × 105	5.20m-o	7.40 k-m	9.00 i-k	11.20 hi
1 × 106	2.80 o	4.00 no	6.20 l-n	7.00 k-m
			LSD = 2.43
Egg masses root system
0	21.20 e	45.80 c	64.80 b	87.20 a
1 × 103	9.60 hi	15.20 g	17.80 f	31.60 d
1 × 104	7.20 j-l	9.60 hi	10.40 h	14.00 g
1 × 105	3.80 mn	7.60 i-k	8.20 h-j	8.00 ij
1 × 106	2.00 no	4.20 mn	5.60 k-m	5.00 lm
			LSD =2.35
Eggs root system
0	7764.0 e	15140.0 c	22600 b	32200 a
1 × 103	4135.0 g	5388.0 f	5680.0 f	9766.0 d
1 × 104	2080.0ij	2997.0 h	3027.0 h	4182.0 g
1 × 105	1330.0lm	2046.0 j	1886.0 jk	2463.6 i
1 × 106	734.00 n	1004.8 mn	1184.0lm	1524.0 kl
			LSD = 416.02
Eggs/g of root
0	1494.2 d	2076.6 c	2455.6 b	2851.2 a
1 × 103	1038.6 e	912.20 f	785.80 g	981.20 e
1 × 104	711.80hi	720.20 h	564.80 k	604.00 jk
1 × 105	601.80jk	654.40 ij	550.40 k	403.20 lm
1 × 106	372.80m	463.80 l	395.80 m	362.20 m
			LSD = 63.95
J2 root system
0	4726.0 e	9265.0 c	14595 b	17995 a
1 × 103	1431.0 j	1947.0 h	2839.0 f	4904.0 d
1 × 104	617.00 l	1071.0 k	1581.0 i	2218.0 g
1 × 105	361.00no	345.00 no	971.00 k	1078.0 k
1 × 106	192.00 p	305.00 op	468.00mn	518.00 lm
				LSD =145.25

J2 cc of soil
0	945.00 e	1853.0 c	2919.0 b	3599.0 a
1 × 103	286.20 j	389.40 h	567.80 f	980.80 d
1 × 104	123.40 l	214.20 k	316.20 i	443.60 g
1 × 105	72.20 no	101.00 lm	194.20 k	215.60 k
1 × 106	39.20 p	61.00 op	93.60 mn	103.60 lm
			LSD =27.03
Pf/Pi**
0	24.980 a	24.405 b	24.797 a	25.098 a
1 × 103	11.132 c	7.3350 d	5.6793 e	7.3350 d
1 × 104	5.3940 e	4.0680 f	3.0720 g	3.2000 g
1 × 105	3.3820 g	2.3910 h	1.9047 i	1.7708 i
1 × 106	1.8520 i	1.3098 j	1.1013 j	1.0210 j
			LSD =0.31

Data are mean of ten replications; Means within a column followed by the same letter are not significantly different according to Least Significant Difference Test P = 0.05.

*Gall and egg mass indices: 0-5 scale; where 0, no galls or egg masses; 1, 1-2 galls or egg masses; 2, 3-10 galls or egg masses; 3, 11-30 galls or egg masses; 4, 31-100 galls or egg masses; 5, >100 galls or egg masses per root system (Lamberti, 1971).

**RF, Reproduction factor whereas Pf is final nematode population density divided by initial nematode population density (Pi).

 

per root system, eggs per root system, eggs per gram of root, and reproduction factor. As the fungal population densities increased, the nematode population level decreased, which is clear evidence of fungal parasitic activities on nematodes. Maximum parasitism was noted at a higher concentration of fungus conidia (1×106).

The minimum number of eggs and J2 in the soil was recovered even at higher initial population densities of nematodes (Table I). Likewise, L. muscarium caused a significant increase in root shoot length and fresh and dry shoot weight compared to control plants (Table II). Top fresh weight (g) significantly improved with increased initial population densities of fungus (Table II), whereas fresh root weight decreased gradually with an increased fungus concentration.

The maximum top fresh weight was obtained when the fungal concentration was 1×106 compared to the control and other treatments in the experiment, while the minimum was recorded at concentration of 1×103. Similarly, the response of the roots in terms of induction of the root galls and egg production to M. incognita infection was directly proportionate to the Pi levels of the nematode and inversely proportional to L. muscarium. The minimum root weight was obtained at a higher level of fungal conidial suspension (Table I).

Moreover, L. muscarium showed no negative impact on plant growth. During our observations, the fresh root weight (g) and top fresh weight (g) gradually improved with increasing L. muscarium concentrations (Table II). The maximum plant growth parameters (root weight and top fresh weight) were obtained at the highest level of fungal conidia, whereas the minimum parameters were noted at a lower level. These studies demonstrate that there is a strong association of L. muscarium with the nematode population and plant growth.

 

Discussion

 

We posit that the growth and yield of vegetables might be enhanced by incorporating population densities of L. muscarium or in combination with other nematophagous fungi. The soil with higher population densities of L. muscarium had a lower final nematode population, ultimately resulting in positive effects on plant growth factors (root shoot lengths and shoot weight) (Yang et al., 2012). The number of galls, egg masses, J2 population, and eggs were significantly decreased (P=0.05) with increasing L. muscarium conidia (Table I). This decrease in the final nematode population was due to a higher mass of conidia with a successful antagonistic reaction towards M. incognita eggs as well as J2 (Zhang et al., 2008). Previously, during in-vitro observations, L. muscarium produced higher densities of conidia on potato dextrose agar (PDA) media under room temperature (data not published) since conidia are the main propagule of infection. Spore attachment, germination and enzyme activity are important factors for fungal virulence (Kim et al., 2014). L. muscarium was believed to produce mucilage matrix, which help fungus conidia stick to nematode eggs and J2 to successfully penetrate germination of fine hyphae (Veenhuis et al., 1985). Additionally, plant roots also provide a biological environment with facilitation of nutrients to nematode and fungi (Curl and Truelove, 1986)

 

Table II.- Effect of graded concentration of nematophagous fungus, Lecanicillium muscarium on different inoculum densities of root knot nematode, M. incognita with respect to plant growth parameters

Conc. of L. muscarium	Inoculum densities of M. incognita
	0	500	1000	1500	2000
Fresh root weight (g)
0	4.16 j	5.20 i	7.30 e	9.20 c	11.30 a
1 × 103	6.66 g	3.98 j	5.90 h	7.23 ef	9.96 b
1 × 104	7.62 d	2.92 l	4.17 j	5.37 i	6.99 f
1 × 105	9.20 c	2.21 m	3.13 l	3.47 k	6.16 h
1 × 106	11.28 a	1.97 n	2.17 m	2.99 l	4.24 j
					LSD =0.2715
Top fresh weight (g)
0	171.60ij	132.60 o	97.40 q	72.40 r	52.20 s
1 × 103	204.40d	161.20 k	143.60 m	137.80 n	128.00 p
1 × 104	235.00c	174.80hi	162.60 k	155.80 l	146.80 m
1 × 105	246.40b	191.40 f	175.80 h	169.76 j	184.78 g
1 × 106	272.40a	203.80 d	199.24 e	185.80 g	204.20 d
					LSD =0.3181

 

Data are mean of ten replications; Means within a column followed by the same letter are not significantly different according to Least Significant Difference Test P = 0.05.

 

and niches for their survival. The aggressiveness of L. muscarium could be correlated to a higher production of chitinolytic enzymes (generally called chitinases) in a wide range of temperatures (5-30°C) with an optimum temperature of 25°C. (Fenice et al., 1996, 1997, 2012; Khan et al., 2004) The cuticle of nematode eggs and J2 are mainly comprised of proteins and chitin (Blaxster and Robertson, 1998). L. muscarium is more powerful and efficient than Trichoderma harzianum, which is commercialized as a strong mycoparasitic and entomopathogenic agent exhibit scarce activity even in cold environments (Malathrakis and Kritsotaki, 1992; Fenice et al., 1998a), low or higher moisture (Jackson et al., 1985; Kope et al., 2008) and broad humidity levels (Kope et al., 2008). Furthermore, studies showed that L. muscarium also plays a vital role in activating the plant defensive system by triggering induced resistance (Hirano et al., 2008). This could be another reason to build the plant confidence level against pathogens. A gradual increase of fungal propagules enhanced plant growth by limiting nematode reproduction factors, which might be due to endophytic colonization of the roots by the fungus, as examined under microscope. Our study validates the results of Trifonova et al. (2009). These authors isolated three fungi viz Fusarium oxysporum, Gliocladium roseum and Verticillium chlamydosporium from infected M. incognita females in southern Bulgaria and found that these fungi decreased the number of eggs from 7.6% to 23.5%. Similarly, Dababat and Sikora (2007) investigated Fusarium oxysporum and found that it inhibited juvenile penetration, gall formation and egg mass production in tomato plants and that a double dose of the fungus had more precise results comparatively. According to Kiewnik and Sikora (2006), nematophagous fungi Paecilomyces lilacinus decreased the number of egg masses by 74%, the root galling by 66% and the final population of nematodes in the roots by 71% compared to the control (inoculated treatment).

 

Conclusion

 

The above discussion reports that Meloidogyne spp. could be managed by increasing the biocontrol concentration in the soil. By this means we could spare the environment and underground water from notorious chemicals that cause pollution and carcinogenic effects. Scientists must try to find alternative management strategies so that farmers may easily benefit from these strategies. Our findings show that the magnitude of the nematode reproduction rate, development of root galls, production of egg masses, eggs and J2 population are inversely related to levels of the fungus “Pi” . Our results also revealed that L. muscarium could be a potential biocontrol agent against plant parasitic nematodes by protecting the roots from extensive colonization.

 

Acknowledgement

 

The authors thank Czech University of Life Sciences, Prague for financial support. The work has been supported by CIGA project number PROJ201500056.

 

Conflict of interest statement

We declare that we have no conflict of interest.

 

References

 

Anke, H., 2010. Insecticidal and nematicidal metabolites from fungi. In: The Mycota: Industrial applications (ed. M. Hotrichter). Springer- Verlag, Berlin, pp. 10.

Anke, H., Stadler, M., Mayer, A. and Sterner, O., 1995. Secondary metabolites with nematicidal and antimicrobial activity from nematophagous fungi and ascomycetes. Can. J. Bot., 73: S932-S939. http://dx.doi.org/10.1139/b95-341

Anke, H. and Sterner, O., 1997. Nematicidal metabolites from higher fungi. Curr. Org. Chem., 1: 361-374.

Anwar, S.A. and McKenry, M.V., 2010. Incidence and reproduction of Meloidogyne incognita on vegetable crop genotypes. Pakistan J. Zool., 42: 135-141.

Anwar, S.A., Zia, A., Hussain, M. and Kamran, M., 2007. Host suitability of selected plants to Meloidogyne incognita in the Punjab, Pakistan. Int. J. Nematol., 17: 144-150.

Barron, G.L., 1977. The nematode-destroying fungi. Canadian Biological Publications, Guelph, Ont., pp. 140.

Berg, G., Zachow, C., Lottmann, J., Gotz, M., Costa, R. and Smalla, K., 2005. Impact of plant species and site on rhizosphere associated fungi antagonistic to Verticillium dahlia Kleb. Appl. environ. Microbiol., 71: 4203-4213. http://dx.doi.org/10.1128/AEM.71.8.4203-4213.2005

Bird, J. and Herd, R.P., 1995. In vitro assessment of two species of nematophagous fungi (Arthrobotrys oligospora and Arthrobotrys flagrans) to control the development of infective cyathostome larvae from naturally infected horses. Vet. Parasitol., 56: 181-187. http://dx.doi.org/10.1016/0304-4017(94)00663-W

Blaxster, M.L. and Robertson, W.M., 1998. The Cuticle. In: Free living and plant parasitic nematodes (eds. R.N. Perry and D.J. Wright). CAB International, Wallingford, UK, pp. 25-48

Boucher, G. and Lambshead, P.J.D., 1994. Ecological biodiversity of marine nematodes in samples from temperate, tropical, and deep-sea regions. Conserv. Biol., 9: 1594-1604. http://dx.doi.org/10.1046/j.1523-1739.1995.09061594.x

Chen, S.Y. and Chen, F.J., 2003. Fungal parasitism of Heterodera glycines eggs as influenced by egg age and pre-colonization of cysts by other fungi. J. Nematol., 35: 271-277.

Chen, S.Y., Dickson, D.W. and Mitchell, D.J., 2000. Viability of Heterodera glycines exposed to fungal filtrates. J. Nematol., 32: 190-197.

Curl, E.A. and Truelove, B., 1986. The Rhizosphere. Springer-Verlag, Berlin, pp. 288. http://dx.doi.org/10.1007/978-3-642-70722-3

Cuthbertson, A.G.S., Blackburn, L.F., Northing, P., Luo, W., Cannon, R.J.C. and Walters, K.F.A., 2010. Chemical compatibility testing of the entomopathogenic fungus Lecanicillium muscarium to control Bemisia tabaci in glasshouse environment. Int. J. environ. Sci. Technol., 7: 405-409. http://dx.doi.org/10.1007/BF03326150

Dababat, A. and Sikora, R.A., 2007. Importance of application time and inoculum density of Fusarium oxysporium 162 for biological control of Meloidogyne incognita on Tomato. Nematropica, 37: 276-275.

Eisenback, J.D., 1985. Detailed morphology and anatomy of second stage juveniles, males, and females of the genus Meloidogyne (root knot nematode). In: An advanced treatise on Meloidogyne, Vol. 1. Biology and control (eds. J.N. Sasser and C.C. Carter). North Carolina State University Graphics, Raleigh.

Ellis, S.D., Boehm, M.J. and Rhodes, L.H., 2008. Nematode diseases of plants. Extension Fact Sheet from the Department of Agriculture and Natural Resources. The Ohio State University, USA, pp. 1-3.

Fenice, M., Selbmann, L., Di Giambattista, R., Petruccioli, M. and Federici, F., 1996. Production of extracellular chitinolytic activities by a strain of the antarctic entomogenous fungus Verticillium cfr. lecanii. In: Chitin enzymology Volume 2 (ed. R.A.A. Muzzarelli). Atec Edizioni. Grottammare, Italy, pp. 285-292.

Fenice, M., Selbmann, L., Zucconi, L. and Onofri, S., 1997. Production of extracellular enzymes by Antarctic fungal strains. Polar Biol., 17: 275-280. http://dx.doi.org/10.1007/s003000050132

Fenice, M., Leuba, J.L. and Federici, F., 1998a. Chitinolytic enzyme activity of Penicillium janthinellum P9 in bench-top bioreactor. J. Ferment. Bioeng., 86: 620-623. http://dx.doi.org/10.1016/S0922-338X(99)80020-8

Fenice, M., Barghini, P., Selbmann, L. and Federici, F., 2012. Combined effects of agitation and aeration on the chitinolytic enzymes production by Antarctic fungus Lecanicillium muscarium CCFEE 5003. Microb. Cell Fact., 11: 12. http://dx.doi.org/10.1186/1475-2859-11-12

Fenice, M. and Gooday, G.W., 2006. Mycoparasitic actions against fungi and Oomycetes by strain (CCFEE 5003) of the fungus Lecanicillium muscarium isolated in Continental Antarctica. Ann. Microbiol., 56: 1-6. http://dx.doi.org/10.1007/BF03174961

Fenice, M., Selbmann, L., Di Giambattista R. and Federici, F., 1998b. Chitinolytic activity at low temperature of an Antarctic strain (A3) of Verticillium lecanii. Res. Microbiol., 149: 289-300. http://dx.doi.org/10.1016/S0923-2508(98)80304-5

Gams, W. and Zare, R., 2001. A revision of Verticillium sect. Prostrata. III. Generic classification. Nova Hedwigia, 72: 329-337.

Gan, Z.W., Yang, J.K., Tao, N., Liang, L.M., Mi, Q., Li, J. and. Zhang, K.Q., 2007. Cloning of the gene Lecanicillium psalliotae chintinase LpChi1 and identification of its potential role in the biocontrol of root knot nematode Meloidogyne incognita. Appl. Microbiol. Biotechnol., 76: 1309-1317. http://dx.doi.org/10.1007/s00253-007-1111-9

Goettel, M.S., Koike, M., Kim, J.J., Aiuchi, D., Shinya, R. and Brodeur, J., 2008. Potential of Lecanicillium spp. for management of insects, nematodes and plant diseases. J. Inverteb. Pathol., 98: 256-261. http://dx.doi.org/10.1016/j.jip.2008.01.009

Güçlü, Ş., Ak, K., Eken, C., Akyol, H., Sekban, R., Beytut, B. and Yildirim, R., 2010. Pathogenicity of Lecanicillium muscarium against Ricania simulans. Bull. Insectol., 63: 243-246.

Hallmann, J. and Sikora, R.A., 1996. Toxicity of fungal endophyte secondary metabolites to plant-parasitic nematodes and soil-borne plant pathogenic fungi. Eur. J. Pl. Pathol., 102: 155-162. http://dx.doi.org/10.1007/BF01877102

Hirano, E., Koike, M., Aiuchi, D. and Tani, M., 2008. Pre-inoculation of cucumber roots with Verticillium lecanii (Lecanicillium muscarium) induces resistance to powdery mildew. Res. Bull. Obihiro Univ., 29: 82-94.

Holbrook, C.C., Knauft, D.A. and Dickson, D.W., 1983. A technique for screening peanut for resistance to Meloidogyne arenaria. Pl. Dis., 67: 957-958. http://dx.doi.org/10.1094/PD-67-957

Hussain, M., Kamran, M., Singh, K., Zouhar, M., Ryšanek, P. and Anwar, S.A., 2016. Response of selected okra cultivars to Meloidogyne incognita. Crop Prot., 82: 1-6. http://dx.doi.org/10.1016/j.cropro.2015.12.024

Hussey, R.S. and Barker, K.R., 1973. Comparison of methods for collecting inocula of Meloidogyne spp. including a new technique. Pl. Dis. Rep., 57: 1025-1028.

Hussey, R.S. and Janssen, G.J.W., 2002. Root-knot nematodes: Meloidogyne species. In: Plant resistance to parasitic nematodes. (eds. J.L. Starr, Cook, R. and J. Bridge). CAB International, United Kingdom, pp. 43-70. http://dx.doi.org/10.1079/9780851994666.0043

Irving, F. and Kerry, B.R., 1986. Variation between strains of the nematophagous fungus, Verticillium chlamydosporium Goddard. II. Factors affecting parasitism of cyst nematode eggs. Nematologica, 32: 474-485. http://dx.doi.org/10.1163/187529286X00345

Jackson, C.W., Heale, J.B. and Hall, R.A., 1985. Traits associated with virulence to the aphid Macrosiphoniella sanborni in eighteen isolates of Verticillium lecanii. Annls. appl. Biol., 105: 39-48.

Kerry, B.R., 2000. Rhizosphere interactions and exploitation of microbial agents for the biological control of plant-parasitic nematodes. Annu. Rev. Phytopathol., 38: 423-441. http://dx.doi.org/10.1146/annurev.phyto.38.1.423

Khan, A., Williams, K.L. and Nevalainen, H.K.M., 2004. Effects of Paecilomyces lilacinus protease and chitinase on the eggshell structures and hatching of Meloidogyne javanica juveniles. Biol. Contr., 31: 346-352. http://dx.doi.org/10.1016/j.biocontrol.2004.07.011

Kiewnick, S. and Sikora, R.A., 2006. Biological control of the root-knot nematode Meloidogyne incognita by Paecilomyces lilacinus strain 251. Biol. Contr., 38: 179-187. http://dx.doi.org/10.1016/j.biocontrol.2005.12.006

Kim, D.G. and Riggs, R.D., 1991. Characteristics and efficacy of a sterile hyphomycete (ARF18), a new biocontrol agent for Heterodera glycines and other nematodes. J. Nematol., 23: 275-282.

Kim, J.J., Xie, L., Han, J.H. and Lee, S.Y., 2014. Influence of additives on the yield and pathogenicity of conidia produced by solid state cultivation of an Isaria javnica isolate. Mycobiology, 42: 346-352. http://dx.doi.org/10.5941/MYCO.2014.42.4.346

Kope, H.H., Alfaro, R.I. and Lavallée, R., 2008. Effects of temperature and water activity on Lecanicillium spp. conidia germination and growth, and mycosis of Pissodes strobe. Biocontrol, 53: 489-500. http://dx.doi.org/10.1007/s10526-007-9087-z

Li, G., Zhang, K., Xu, J., Dong, J. and Liu, Y., 2007. Nematicidal substances from fungi. Recent Pat. Biotechnol., 1: 1-22. http://dx.doi.org/10.2174/187220807782330165

Lopez-Llorca, L.V. and Jansson, H.B., 2006. Fungal parasites of invertebrates: multimodal biocontrol agents. In: Exploitation of fungi (eds. G.D. Robson, P. VanWest and G.M. Gadd). Cambridge University Press, Cambridge, pp. 310-335.

Malathrakis, N.E. and Kritsotaki, O., 1992. Effect of substrate, temperature and time of application on the effectiveness of three antagonistic fungi against Botrytis cinerea. In: Recent advances in Botrytis research Wageningen. Pudoc Scientific Publishers, pp. 187-191.

Manzanilla-Lopez, R.H., Kenneth, E. and Bridge, J., 2004. Plant diseases caused by nematodes. In: Nematology- advances and perspectives. Nematode management and utilization Volume II (eds. Z.X. Chen, S.Y Chen and D.W. Dickson). CABI publishing. Cambridge, MA, pp. 637-716. http://dx.doi.org/10.1079/9780851996462.0637

Meyer, S.L.F., Hufttel, R.N., Liu, X.Z., Humber, R.A., Juba, J. and Nitao, J.K., 2004. Activity of fungal culture filtrates against soybean cyst nematode and root-knot nematode egg hatch and juvenile motility. Nematology, 6: 23-32. http://dx.doi.org/10.1163/156854104323072883

Meyer, S.L.F., Massoud, S.I., Chitwood, D.J. and Roberts, D.P., 2000. Evaluation of Trichoderma virens and Burkhoderia cepacia for antagonistic activity against root knot nematode, Meloidogyne incognita. Nematology, 2: 871-879. http://dx.doi.org/10.1163/156854100750112815

Meyer, S.L.F. and Meyer, R.J., 1996. Greenhouse studies comparing strains of the fungus Verticillium lecanii for activity against the nematode Heterodera glycines. Fund. appl. Nematol., 19: 305-308.

Nguyen, N.V., Kim, Y.J., Oh, K.T., Jung, W.J. and Park, R.D., 2007. The role of chitinase from Lecanicillium antillanum B-3 in parasitism to root-knot nematode Meloidogyne incognita eggs. Biocontr. Sci. Techn., 17: 1047-1058. http://dx.doi.org/10.1080/09583150701668658

Ownley, B.H., Gwinn, K.D. and Vega, F.E., 2010. Endophytic fungal entomopathogens with activity against plant pathogens: ecology and evolution. Biocontrol, 55: 113-128. http://dx.doi.org/10.1007/s10526-009-9241-x

Quesenberry, K.H., Baltensperger, D.D., Dunn, R.A., Wilcox, C.J. and Hardy, S.R., 1989. Selection for tolerance to root-knot nematodes in red clover. Crop Sci., 29: 62-65. http://dx.doi.org/10.2135/cropsci1989.0011183X002900010014x

Regaieg, H., Ciancio, A., Horrigue, N.H., Brasso, G. and Rosso, L., 2010. Effects of culture filtrates from the nematyophagous fungus Verticillium leptobactrum on viability of the root-knot nematode Meloidogyne incognita. World J. Microbiol. Biotechnol., 26: 2285-2289. http://dx.doi.org/10.1007/s11274-010-0397-4

Renčo, M., 2013. Organic amendments of soil as useful tools of plant parasitic nematodes control Review Article. Helminthologia, 50: 3-14. http://dx.doi.org/10.2478/s11687-013-0101-y

Renčo, M., Sasanelli, N., Papajová, I., Maistrello, L., 2012. The nematicidal effect of chestnut tannin solutions on the potato cyst nematode Globodera rostochiensis (Woll.) Behrens. Helminthologia, 2: 108-114 .

Rodriguéz-Kábana, R. and Canullo, G.H., 1992. Cropping systems for the management of phytonematodes. Phytoparasitica, 20: 211-224. http://dx.doi.org/10.1007/BF02980843

Sasser, J.N. and Freckman, D.W., 1987. A world perspective on Nematology: The role of society. In: Vistas on nematology (eds. J.A. Veech and D.W. Dickson) SON Inc., Hyattsville, MD, pp. 7-14.

Sharon, E., Bar-Eyal, M., Chet, I., Herrera- Estrella, A., Kleifeld, O. and Spiegel, Y., 2001. Biological control of the root knot nematode Meloidogyne javanica by Trichoderma harzianum. Phytopathology, 91: 687-693. http://dx.doi.org/10.1094/PHYTO.2001.91.7.687

Siddiqui, Z.A. and Mahmood, I., 1996. Biological control of plant parasitic nematodes by fungi: A review. Bioresour. Technol., 58: 229-239. http://dx.doi.org/10.1016/S0960-8524(96)00122-8

Stirling, G.R., 1991. Biological control of plant parasitic nematodes: Problems, progress and prospects. CAB International, Wallingford, UK.

Trifonova, Z., Karadjova, J. and Georgieva, T., 2009. Fungal parasites of the root-knot nematodes Meloidogyne spp. in southern Bulgaria. Est. J. Ecol., 58(1): 47-52. http://dx.doi.org/10.3176/eco.2009.1.05

Veenhuis, M., Nordbring-Hertz, B. and Harder, W., 1985. An electron microscopical analysis of capture and initial stages of penetration of nematodes by Arthrobotrys oligospora. Antonie van Leeuwenhoek, 51:385-98. http://dx.doi.org/10.1007/BF02275043

Whitehead, A.G and Hemming, J.R., 1965. A comparison of some quantitative methods of extracting small vermiform nematodes from soil. Annls. appl. Biol., 55: 25-38.

Yang, J., Benecke, S., Jeske, D.R., Rocha, F.S., Becker, J.S., Timper, P., Becker, J.O. and Borneman, J., 2012. Population dynamics of Dactylella oviparasitica and Heterodera schachtii: toward a decision model for sugar beet planting. J. Nematol., 44: 237-244.

Lamberti, F., 1971. Results of nematicidal control on levantin tobacco in the province of Lecce. Il Tabacco, 738: 5-10 (In Italian).

Zasada, I., Avendano, F., Li, Y.C., Logan, T., Melakeberhan, H., Koenning, S.R. and Tylka, G.L., 2008. Potential of an alkaline-stabilized biosolid to manage nematodes: case studies on soybean cyst and root-knot nematodes. Pl. Dis., 92: 4-13. http://dx.doi.org/10.1094/PDIS-92-1-0004

Zhang, L., Yang, J., Niu, Q., Zhao, X., Ye, F., Liang, L. and Zhang, K.Q., 2008. Investigation on the infection mechanism of the fungus Clonostachys rosea against nematodes using the green fluorescent protein. Appl. Microbiol. Biotechnol., 78: 983-990. http://dx.doi.org/10.1007/s00253-008-1392-7

Zouhar, M., Douda, O., Nováková, J., Doudová, E., Mazáková, J., Wenzlová, J., Ryšánek, P. and Renčo, M., 2013. First report about the trapping activity of Stropharia rugosoannulata acanthocytes for Northern root knot nematode. Helminthologia, 2: 127-131.

Zuckerman, B.M. and Esnard, J., 1994. Biological control of plant nematodes: current status and hypothesis. Jap. J. Nematol., 24: 1-13. http://dx.doi.org/10.3725/jjn1993.24.1_1