Printer Friendly

In-vitro predatory activity of nematophagous fungi from Costa Rica with potential use for controlling sheep and goat parasitic nematodes.

In livestock production systems located in tropical and subtropical regions of the world, parasitic diseases have emerge as a major cause of losses in livestock productivity due to increased morbidity and mortality of the animals, reduced levels of production and productivity, reproductive disorders and high controls cost, among others (FAO 2003, Minglian et al. 2004, Araujo et al. 2006, Su et al. 2007). Gastrointestinal nematodes (GIN) in small ruminants include Haemonchus contortus, Telodorsagia circumcincta, Trichostrongylus spp., Nematodirus spp. and Cooperia spp. The proportion of each of these nematodes in small ruminants populations varies according to their geographical location; the highest prevalence exists in tropical humid areas, given that these conditions favor the development of eggs and larval states throughout the year (Chan drawathani 2004, Fleming et al. 2006). Due to the damage caused by the organisms, producers have been forced to invest significant resources trying to minimize the negative impact in their herds. For this reason, anthelmintics are highly utilized, often as the only strategy for the control of GIN, which has resulted in the development of antihelmintic resistance (FAO 2003, Fleming et al. 2006, Vargas 2006, Varady et al. 2007).

Given the acquired resistance by GIN to anthelmintics, biological control appears to be a feasible and effective alternative (Vargas 2006). One of the main natural enemies of nematodes in the soil are the nemathophagous fungi, which can be identified from soil samples and other substrates like domestic animal feces. These fungi are classified into four main groups: endoparasitic, opportunistic, toxic and predatory (Duddington 1955, Barron 1977, Minglian et al. 2004, Li et al. 2005, Su et al. 2007). Many of these fungi have been investigated worldwide, especially Arthrobotrys oligospora, Monacrosporium spp. and Duddingtonia flagrans (Larsen 2000).

This study seeks to determine the potential of Costa Rican strains of nematophagous fungi as an effective tool for GIN control in sheep and goat population in the country.


Samples: Samples were taken from 51 different farms throughout the provinces of Costa Rica, distributed as follows: 12 in Alajuela, 17 in Cartago, 2 in Guanacaste, 2 in Heredia, 7 in Limon, 3 in Puntarenas and 8 in San Jose (Tables 1-2).

The nature of the samples included soils of different crops (potatoes, tomatoes, banana, ornamentals, squash and coffee), animal feces (cattle, sheep, goats and horses), soil, dead forest leaves and plants with signs of illness caused by nematodes (Table 3). Collected samples were placed into plastic bags properly labeled with the date, elevation, and geographical coordinates (as provided by a global positioning system) and were then taken to the Phytoprotection Laboratory of the Specialized National Center of Organic Agriculture, where they were held under refrigeration until processed.

Nematode larvae: GIN suspensions were used to stimulate the production of traps in the isolates, in slide culture for taxonomic identification, in the predation assay and in the viability assay. The GIN larvae were obtained from animals naturally infected by means of the fecal culture technique (Zajac & Conboy 2006). Mixed cultures of infective larvae (L3) of Haemonchus contortus and Cooperia spp. (90% and 10% respectively), were used for the isolation, slide culture and viability tests. For the predation assay, the suspension used corresponded to 100% infective larvae of H. contortus.

Isolates: In the laboratory, samples were processed using three different techniques to extract fungi from soil: sprinkled (Barron 1977), soil dilution (Villalba 2006) and moist chamber (Delgado et al. 2001).

For the sprinkling method, the plates were placed in an incubator with an average temperature of 28 [degrees]C, 80% humidity and artificial light. The material was checked on a daily basis from the fifth day with a stereoscope. As soon as the presence of trapped nematodes was identified, the fungus was isolated and purified.

In the soil dilution, 0.5ml of the solutions (1:10, 1:100 and 1:1000) were cultivated in triplicate over selective media and placed on Petri dishes in advance. Potato-dextrose-agar (PDA Difco) with antibiotic (Oxoid Chloramphenicol selective supplement), acid PDA (1ml of citric acid at 10% per every 100ml of culture medium) and Rose Bengal (Oxoid Rose-Bengal Chloramphenicol Agar+Oxoid Chroramphenicol selective supplement) were used.

Plates were held in an incubator with an average temperature of 28 [degrees]C, 80% humidity and artificial light. Cultures were examined 72 hours after planting; the growing colonies were analyzed under the microscope (at 40X) and those that showed conidia and mycelium similar to those described for PNF were purified.

Vessels with humidity chambers were kept for 15 days in a room at 27-29 [degrees]C, 75-80% humidity and provided 12 hours of artificial light. The plates were monitored on a daily basis in order to observe mycelium development. The mature fungi were examined under a microscope at 40X. Those with conidia and mycelium similar to those described for PNF were purified (Orozco 2005).

For isolated fungi purification, conidia were collected with an inoculation loop and transferred to PDA with antibiotic (PDAatb) in order to eliminate bacteria. Then, conidia were transferred to acid PDA (PDAac) to facilitate the growth of contaminant saprophytic fungi. After achieving cleansing of PNF, isolates were transferred to PDA for storage.

Taxonomic identification: For taxonomic identification, the slide culture technique was used (Orozco 2005). The slides were observed under a microscope in order to identify characteristics and size of each conidia. According to shape and disposition of spores, the genus and species were identified using identification keys (Delgado et al. 2001).

pH of samples: The pH value was determined in each soil sampling.

Predation assay: All isolated strains were evaluated on capture percentages. Conidia were cultivated in water-agar (WA) plates, and 96 hours after fungus cultivation, they were added a suspension of approximately 100 GIN (six replicates per strain plus two controls using only larvae to evaluate if there was any contaminant nematophagous fungi) (Park et al. 2002, Araujo et al. 2004, Gonzalez et al. 2005, Elosegui 2006). Plates were then incubated at 28[degrees]C, 80% humidity with artificial light. After 40 hours of incubation larvae were classified into the following four categories: trapped (those fixed or in motion that were in contact with traps), free (those moving continuously all over the plate), coiled (curled, immobile) and dead (those that were rigid, in a straight position and not in contact with traps) were counted and individually observed under the stereoscope. The trapped larvae percentage of the total counted larvae was calculated through an equation used by Gonzalez et al. (2005).

Chlamydospore production: Cultures were performed with isolated fungi in four different agar media: Potato Dextrose Agar (PDA), Corn Meal Agar (CMA), Potato-Carrot Agar (PCA) and Malt Extract Agar (MEA). These cultures were incubated for 7 days at 28 [degrees]C, 50% humidity and artificial light. Plates were then held at room temperature and in darkness for 6 to 8 weeks. Cultures positive for chlamydospores production were placed into suspensions using the Ojeda et al. (2005) technique. In order to evaluate the production of chlamydospores for each isolated fungus, another own methodology was applied: a small sample from the center of cultures of six weeks of age was taken and examined under the microscope. The following criteria were used:
-     No chlamydospores were observed.

+     A total of 1 to 5 chlamydospores on the
      slide, and they were hard to find.

++    Easy to find chlamydospores; however,
      there were not too many.

+++   There were plenty of chlamydospores,
      some were even forming chains.

Digestibility assay: The samples were taken to the Animal Food Science Laboratory at the University of Costa Rica, where in vitro digestibility assays described by Tilly & Terry (1963) were performed under respective modifications regarding the substitution of fodder for chlamydospores (Orozco 2005). This assay was performed on fungi strains that formed the largest number of chlamydospores; in addition, the assay was performed on the A. conoides strain, which this has been reported as an acceptable candidate for BC.

Viability assay: This test was performed to determine if any chlamydospore had survived the in vitro digestibility assay. This determination was made by a conidia germination test: chlamydospore reference samples that had been subjected to the artificial digestive process were plated onto Petri dishes containing WA, a suspension of GIN was then added (Park et al. 2002, Araujo et al. 2004, Gonzalez et al. 2005, Elosegui 2006).

The statistical analysis included a comparison of the variables using the non-parametric Kruskal Wallis-test, based on a qualitative scale. Data analysis was performed using Infostat (2002). The statistical significance of the variables was tested at a 0.05 level of significance.


Fungal isolation: From the 51 sampled farms, 24 PNF of the Arthrobotrys and Candelabrella genera were isolated (Table 4). Additionally, 1 strain of Beauveria sp., 1 of Clonostachys sp., 1 of Lecanicillium sp., and 13 strains of Trichoderma sp were isolated.

From the isolated PNF, 20 were obtained by the soil sprinkling technique (p<0.001) vs. 2 by soil dilution and 2 by the humidity chamber. Strains of Beauveria sp., Clonostachys sp., Lecanicillium sp. and Trichoderma sp. were isolated by the soil dilution technique.

Taxonomic identification: Of the isolated PNF, 13 were identified as Arthrobotrys oligospora, 1 as A. conoides, 1 as A. dactyloides and 9 as Candelabrella musiformis.

pH of samples: The pH level of the samples fell within a wide range; the most acidic was 5.2 (SCh1), while the most alkaline was of 9.9 (VELG), creating an average of 7.8. In soils from which PNF were isolated, the lowest pH was 5.6 (SD) and the highest 7.5 (SFL) (Fig. 1).

The following strains showed an extremely slow growth in PDAac: SZa (A. dactyloides; pH 7.1), SM (C. musiformis; pH 7.6), SFL (A. oligospora; pH 7.5) and VLV (A. oligospora; pH 6.7).

Predation Assay: The strain that demonstrated the highest predacious ability was SAPt (C. musiformis) with 96.6%, while the one that showed the lowest capture percentage was TC (A. oligospora) with 9.1% (Fig. 2).



Capture percentage of the A. oligospora strains varied widely between 9.1% in the TC strain and 93.4% in the VSL strain, with an average capture ability of 68.7% (Fig. 2). The predacious capability of C. musiformis strains was otherwise more homogeneous, with a smaller range that varied from 70.8% of SCT strain to 96.6% of SAPt strain and a capture average of 86.1%.

Since just one strain of A. conoides (64.9%) and one of A. dactyloides (89.7%) were isolated, it was not possible to carry out comparisons among different strains.

The controls containing only larvae had a mortality rate of 1.6% and 4% of coiled larvae.

Despite the wide range of predation rates, statistical analysis showed no significant differences between the average catch (p=0.25).

Production of chlamydospores: Out of 24 isolated PNF, 23 were positive for the formation of resistance structures (Figs. 3-4). Only the TC (A. oligospora) strain did not produce them.

Generally, C. musiformis strains showed a higher production of chlamydospores than Arthrobotrys genus strains (Table 5); however, statistical analysis showed no significant differences (p=0.10) among the four species.

Regarding culture media, PCA was the most effective, followed by PDA, CMA (most effective for conidia formation) and MEA (p<0.01). On MEA media, fungi presented highly dispersed micelial growth and exhibited very limited chlamydospore and conidia formation (Table 5).


Viability assay: Out of 13 fungi that underwent the in vitro digestibility assay, 6 were positive for growth in WA plates added with larvae. Of these fungi, three corresponded to the C. musiformis species (ET, SCS, SM) and the other three to A. oligospora (VLV, COr, SBI) (Table 6).


Isolates and taxonomic identification: It was expected that most of PNF could be isolated using the soil sprinkling technique described by Barron (1977). Nevertheless, the use of other techniques favored and accelerated the growth of saprophyte fungi such as Rhizopus, Fusarium, Phoma, Aspergillus and Penicillium, since they are facultative saprophytic microorganisms. In addition, due to spacing, saprophyte fungi prevented the growth of PNF. Mota et al. (2003), states that some species of PNF develop traps as a result of external stimulus (physiological stress, presence of nematodes or their excreta), while others develop spontaneously and proved to be more dependant on nematodes as a source of nutrients. This explains why few PNF strains were obtained by using the soil dilution technique. In this technique, fungi are never stressed regarding the culture media (nutrients, humidity, time, etc.) and they are not stimulated with nematodes.


Growth of endoparasitic nematophagous fungi is even slower than that of predators, and the soil sprinkling technique proved ineffective in isolating them. Furthermore, another possible reason why endoparasitic nematophagous fungi were not isolated, is that PNFs have been reported to be more aggressive, quickly exhausting nutrient sources (nematodes), leaving others without media to grow and sporulate (Larsen 2000). Out of the purpose of this study, Barron (1977) described a Baermann funnel technique for the isolation of these fungi, but this methodology was not employed as we were interested in PNFs.

Conversely only PNF from plants with observed signs of nematode-induced disease could be isolated by the humidity chamber technique. According to Delgado et al. (2001) and Cabezas (2004), the purpose of this technique is to give favorable humidity conditions in order to reactivate and accelerate fungi growth in the sample.

When using soil and feces samples, certain structures similar to A. oligospora conidia were seen under the microscope, but when trying to isolate them, faster growing saprophyte fungi grew (mainly Fusarium and Aspergillus), possibly because they were more abundant than the PNFs. On the contrary, in the case of plants, there was a nematode population already established, providing the PNF with additional sources of nutrients, enabling them to colonize faster and more prolifically than other saprophytes.

The taxonomic identification of PNF was carried out based on the observation of fungi morphological characteristics presented when trapping nematodes, rather than in pure cultures where traps are usually non-existent. Furthermore, the dimensions of the conidia and the morphology of the conidiophore can be altered considerably (Cooke & Godfrey 1964, Haard 1968, Van Oorschot 1985). The fungus with the highest number of isolates was A. oligospora, followed by C. musiformis, which was already expected. Barron (1977), states that fungi that form adhesive networks are ubiquitous and very aggressive and thus predominant in soil samples. He also mentions that A. oligospora is the most common isolated PNF, which coincides with Orozco's (2005) study of Costa Rican PNF. Orozco obtained a similar result in the isolation of these two species. In the aforementioned study, A. oligospora, C. musiformis and Dactyllela sp. were isolated. Contrary to Orozco's (2005) results, in this study Dactyllela sp. was not isolated and only one strain of A. conoides and A. dactyloides were isolated. The above-mentioned discrepancy could be due to two situations: 1. human error: since the conidiophores of these fungi are similar to those of other fungi that grow in Petri dishes and they could have gone unnoticed; 2. These Arthrobotrys and Dactyllela spp. species might be not as abundant in Costa Rica, making difficult to isolate them (Orozco 2005).

Regarding altitude and life zone, there is no similar information in the consulted literature in order to make comparisons. Nevertheless, it may be concluded that there is PNF presence at various altitudes and life zones, which reveals the versatility of these fungi.

pH in soils: pH and temperature are two key parameters for fungi growth manipulation, sporulation and saprophytic ability (Hajieghrari et al. 2008). Madigan et al. (2000) states that fungi, as a group, tend to tolerate more acid environments than bacteria and that most of them grow properly in a pH of 5.0 or lower. On the contrary, the PNF in this study were isolated mainly from soil pH of 6.0 to 7.5. Strains such as SZa (A. dactyloides; pH 7.1), SM (C. musiformis; pH 7.6), SFL (A. oligospora; pH 7.5) and VLV (A. oligospora; pH 6.6) showed an extremely slow growth in PDAac. However, PNF such as A. oligospora, C. musiformis, A. oviformis, Monacrosporum megalosporum and Dactylaria parvispora show an extracellular serine-protease, similar to subtilisine, responsible for the immobilization of trapped nematodes and impoverishing their cuticle to introduce hyphae (Tunlid & Jansson 1991, Minglian et al. 2004, Kanda et al. 2008, Nagee et al. 2008). Acid pH affects the transcription of genes that codify said enzyme, its optimal pH activity is between 6.0 and 8.0 (Tunlid & Jansson 1991, Kanda et al. 2008). The expression of this serine protease is related to the physiology and the pathogenicity of the PNF. If this enzyme is not produced, fungi cannot pass from their saprophytic phase to the predacious mode, and may even be non-viable, because a cuticle peptide is released by the nematodes as a reaction to the serine protease, which promotes the formation of traps (Ahman et al. 2002, Kanda et al. 2008).

Predation assay: The transformation of PNF from saprophytic to a pathogenic state is determined by the presence and the quantity of nematodes, as well as the nutritional conditions of the medium: as the amount negative affect on the predacious ability, there is an amount of nutrients increases, mainly in A. oligospora (Morgan et al. 1997). Due to this fact, the assay was performed in WA, and although the amount of nematodes affects the change from saprophytic to pathogenic state in a directly proportional way, it was considered that 100 was a relatively good amount of larvae and it facilitated the final count in the assay. This explains the high predacious percentage in the majority of the strains: in a medium where the nutrients are limited, nematodes become an important source of nitrogen for growth and providers of essential substances such as amino acids and vitamins (Morgan et al. 1997, Gonzalez et al. 2005, Migunova & Byzov 2005).

On the other hand, it was noticed that some strains (TC, PC, 3OSL, 1SD, COr and VAl) demonstrated a slow growth before the larvae were added. Park et al. (2002) gives a four-day period for incubation of fast growth fungi, and a seven-day period for the slow growing fungi. However, this methodology was not utilized, which could have influenced the low capture percentages in those strains. In this case, the added larvae were extracted from an ovine sample. Orozco (2005) reported that certain Costa Rican strains of A. oligospora and C. musiformis as well, showed differences in the percentage of trapping larvae depending on their origin. Thus, according to Orozco (2005), the A. oligospora strains showed a higher tendency to capture larvae of ovine nematode and presented a homogeneous predatory capability, while the C. musiformis strains preferred larvae of goat nematodes and their predacious ability was very heterogeneous. The results in this study differ from Orozco's (2005), since capture percentages of larvae of ovine nematodes by the C. musiformis strains were higher than the A. oligospora strains (which showed a more heterogeneous capture percentage); this result resembles what Park et al. (2002) described.

The differences reported by several researchers, indicate that the predacious abilities of the PNF species can vary according to the region and environmental factors where the strains are isolated (Graminha et al. 2005a).

According to Gonzalez et al. (2005), the temperature can affect the development of the traps in A. conoides and A. oligospora; consequently, it can affect the trapping percentage of fungi. In the case of A. oligospora strains, it was noticed that both strains coming from cold zones, and the ones from the hot zones showed very different capture percentages, so it cannot be stated that the change of temperature for this specie had an influence on the test. On the other hand, only one strain of A. conoides was found; therefore, no comparisons could be made. Given that this species is reported by many as an excellent predator, it is possible that the change in temperature between its natural environment and the laboratory's had affected the development of its traps and prevented obtaining results as good as those reported by Graminha et al. (2005a), of 99.3% nematode capture.

In the case of A. dactyloides, there is little information about the predacious capability regarding animal nematodes; according to Nunez (2002), this fungus is closely related to roots of plants. Consequently, they are in a favorable position for trapping phytoparasitic nematodes. In studies carried out with Meloidogyne incognita (a root-knot nematode of various plant species), a capture rate of up to 81.8% of juveniles is reported (Kumar & Singh 2006), which is not far from the 89.7% obtained in the test done with larvae of sheep in this study. It was noted that the two strains with the lowest capture (TC and PC) were isolated from plants infected by nematodes. Gomes et al. (2000), states that free-living larvae are preferred by PNF above NGI of ruminants.

The only fungus isolated from ovine feces (C. musiformis 1OSL) had a good capture percentage; however, it was not as good as expected. The best three strains were isolated in places where there were no species of ovine: a national park, equine pastures and pastures for dairy cows. Araujo et al. (2006) indicates that PNF are not specific for a particular genus of nematodes of the same animal species.

Production of chlamydospores, digestibility and viability assays: Chlamydospores, resistance spores which survive in unfavorable environmental conditions, are found in many of the PNF. Their production is the result of different factors in the culture: incubation time, temperature and medium utilized (Park et al. 2002).

From the culture media used in this study, the PCA contained the lowest amount of digestible carbohydrates (Carris 2009); this would explain why the highest amounts of chlamydospores were formed in it. On the other hand, according to Carris (2009), the majority of fungi grow really well in PDA, but this medium can be very rich, and the excessive mycelia growth can affect sporulation. In this case, the stress factor could be the limited space in the Petri dish or the amount of nutrients: even though the medium is very rich, the growth of the fungus is extremely accelerated. In the case of MEA and CMA, these cultivation media are reported as weak media (as compared to PDA) that can be utilized for cultivation of soil fungi and that show a balance between the growth of mycelia and sporulation (Carris 2009).

During this research, the piece used to introduce humidity in the incubator for cultivating fungi broke down. In order to avoid delaying the project, vessels with distilled water were introduced into the chamber to provide humidity and continue with the cultivations. It was determined, using a hygrometer, that the humidity was held at 50%; however, there was not enough space in the incubator for more vessels, for this reason, it was decided to work with that percentage of humidity. This variation in the humidity caused all incubated fungi in that moment to form chlamydospores, which presented a great opportunity to perform this assay.

In this study, 95.8% of the isolated strains formed chlamydospores (23 out of 24). This result was not expected; in the work done by Orozco (2005) with Costa Rican strains, only strains of species of C. musiformis produced a big quota of chlamydospores, while strains of species A. oligospora did not. Park et al. (2002) reports this last species as producers of resistance structures and mentions that fungi that grow under dry conditions are prone to survive due to the formation of chlamydospores. The count was established by means of plus signs (+) in order to determine the best strains according to production of chlamydospores.

Out of the 23 strains that form chlamydospores, 14 underwent the digestibility assay. Out of these 14 strains, 13 were chosen for their good performance regarding production of resistance spores and the remaining strain was chosen for being the only A. conoides one. Haard (1968) describes that in this species, chlamydospores are abundant in old cultures; however, this strain did not prove to be a good producer of spores of resistance. Instead, it was chosen for the digestibility assay.

The six positive strains in the viability assay were divided between the A. oligospora and the C. musiformis species. Three of them (VLV, ET and COr) were isolated from pastures where the feces are frequently deposited. Animals then consume the spores along with the grass, as a result, these strains could even adapt to adverse conditions of the gastro-intestinal tract of animals, which can be interpreted as the positive result in the assay (Orozco 2005). A. conoides did not survive the digestibility assay; this coincides with Graminha et al. (2005b) who obtained the same results in his investigation of A. conoides and C. musiformis.

Regarding A. dactyloides, the strain that underwent the digestibility assay could not be re-isolated in the viability test. Unfortunately, there is no information in the literature consulted about survival of chlamydospores of this fungus species in similar assays in order to carry out any kind of comparison. Even so, Graminha et al. (2005b) indicates that chlamydospores might not survive due to the adverse conditions to which they were subjected, such as: acids, pH, temperature or competition with micro biota.

Regardless of the obtained results, it is not recommended to reject the strains that did not survive the digestibility assay, since Flores et al. (2003) assures that in animals, fungi can be protected by intestinal content. It should be kept in mind that the intestinal content of the ruminants is rich in cellulose, a compound resistant to the action of gastric acids, which can become a key element in the protection of spores, preventing the acids from working directly on and degrading them. This type of protection does not exist for in vitro tests, thus the reaction mixture is in direct contact with fungi, favoring its easy degradation (Orozco 2005). Consequently, Araujo et al. (2006) considers that when choosing a PNF, one should take into account not only in vitro tests, but also field tests. When carrying out field tests, PNF species from the same locations where they will be applied should be favored for use in BC of parasites.

Among these microorganisms, Candelabrella musiformis, due to its easy isolation, good predatory capacity and ability to form chlamydospores in a humidity common in the country, appears to be the most promising fungi for use as a biological control agent in Costa Rica.


We would like to thank Jorge Sanchez and Adrian Martinez, University of Costa Rica, for their collaboration during the digestibility assay. Miguel Obregon, who helped identifying fungi. The CNEAO-INA, especially to Carmen Duran and Efrain Munoz, who provided financial support. Felipe Torres, Armando Aguilar, Ivan Rodriguez, Ramon Camara and Nadia Ojeda for inspiring and guiding this investigation.

Received 25-I-2010.

Corrected 18-VII-2010.

Accepted 20-VIII-2010.


Ahman, J., T. Johansson, M. Olsson, P.J. Punt, C.A.M.J.J. van den Hondel & A. Tunlid. 2002. Improving the pathogenicity of a nematode-trapping fungus by genetic engineering of a subtilisin with nematotoxic activity. Appl. Environ. Microb. 68: 3408-3415.

Araujo, J.V., R.C.L. Assis, A.K. Campos & M.A. Mota. 2004. Atividade in vitro dos fungus nematofagos dos generos Arthrobotrys, Duddingtonia e Monacrosporium sobre nematoides trichostrongilideos (Nematoda: Trichostrongyloidea) parasitos gastrointestinais de bovinos. Rev. Bras. Parasitol. Vet. 13: 67-71.

Araujo, J.V., R.C.L. Assis, A.K. Campos & M.A. Mota. 2006. Efeito antagonico de fungos predadores dos generos Monacrosporium, Arthrobotrys e Duddingtonia sobre larvas infectantes de Cooperia sp. e Oesophagostomum sp. Arq. Bras. Med. Vet. Zootec. 58: 373-380.

Barron, G.L. 1977. The nematode destroying fungi: topics in mycobiology, no. 1. Canadian Biological, Ontario, Canada.

Bolanos, R., V. Watson & J. Tosi. 2005. Mapa ecologico de Costa Rica (Zonas de Vida, segun el sistema de clasificacion de zonas de vida del mundo de L.R. Holdridge). Escala 1:750 000. Centro Cientifico Tropical, San Jose, Costa Rica.

Carris, L. 2009. General mycology. (Downloaded: May 29, 2009,

Cabezas, O. 2004. Diagnostico de enfermedades en plantas. (Downloaded: October 16, 2008, www.senasa. tingo_maria/diagnostico_enfermedades_plantas.pdf).

Chandrawathani, P. 2004. Problems in the control of nematode parasites of small ruminants in Malaysia: resistance to anthelmintics and the biological control alternative. Ph.D thesis, Swedish University of Agricultural Sciences, Uppsala, Swedish.

Cooke, R.C. & B.E.S. Godfrey. 1964. A key to nematode-destroying fungi. T. Brit. Mycol. Soc. 47: 61-74.

Delgado, A.E., A.J. Pineiro & L.M. Urdaneta. 2001. Hongos coprofilicos mitosporicos del Estado Zulia, Venezuela. (Downloaded: April 29, 2009, dfRed. jsp?iCve=61412202).

Duddington, C.L. 1955. The friendly fungi: a new approach to the eelworm problem. Faber and Faber, London, United Kingdom.

Elosegui, O. 2006. Metodos artesanales de produccion de bioplaguicidas a partir de hongos entomopatogenos y antagonistas (Downloaded: November 6, 2007, METODOS%20ARTESANALES% 20DE%20PRODUCCI%C3%93N%20DE%20BIOPLAGUICIDAS.pdf).

FAO (Food and Agriculture Organization). 2003. Resistencia a los antiparasitarios: estado actual con enfasis en America Latina. Direccion de Produccion y Sanidad Animal de la FAO. Estudio FAO, Produccion y Sanidad Animal 157, Rome, Italy.

Fleming, S.A., T. Craig, R.M. Kaplan, J.E. Miller, C. Navarre & M. Rings. 2006. Anthelmintic resistance of gastrointestinal parasites in small ruminants. J. Vet. Intern. Med. 20: 435-444.

Flores, J., D. Herrera, P. Mendoza de Gives, E. Liebano, V.M. Vazquez & M.E. Lopez. 2003. The predatory capability of three nematophagous fungi in the control of Haemonchus contortus infective larvae in ovine faeces. J. Helminthol. 77: 297-303.

Gomes, A.P.S., M.L. Ramos, R.S. Vasconcellos, J.R. Jensen, M.C.R. Vieira & J.V. Araujo. 2000. In vitro activity of brazilian strains of the predatory fungi Arthrobotrys spp. on free-living nematodes and infective larvae of Haemonchus placei. Mem. Inst. Oswal do Cruz 95: 873-876.

Gonzalez, R., P. Mendoza de Gives, G. Torres, C. Becerril, E. Ortega & O. Hernandez. 2005. Estudio in vitro de la capacidad depredadora de Duddingtonia flagrans contra larvas de nematodos gastrointestinales de ovinos de pelo. Tec. Pecu. Mex. 43: 405-414.

Graminha, E.B.N., A.J. Costa, G.P. Oliveira, A.C. Monteiro & S.B.S. Palmeira. 2005a. Biological control of sheep parasite nematodes by nematode-trapping fungi: in vitro activity and after passage through the gastrointestinal tract. World J. Microb. Biot. 21:717-722.

Graminha, E.B.N., A.C. Monteiro, H.C. Silva, G. Pereira & A.J. Costa. 2005b. Controle de nematoides parasitos gastrintestinais por Arthrobotrys musiformis em ovinos naturalmente infestados mantidos em pastagens. Pesq. Agropec. Bras. 40: 927-933.

Hajieghrari, B., M. Torabi, M.R. Mohammadi & M. Davari. 2008. Biological potantial of some Iranian Trichoderma isolates in the control of soil borne plant pathogenic fungi. Afr. J. Biotechnol. 7: 967-972.

Haard, K. 1968. Taxonomic studies on the genus Arthrobotrys corda. Mycologia 60: 1140-1159.

InfoStat. 2002. InfoStat version 1.1. Grupo InfoStat, FCA, Universidad Nacional de Cordoba, Cordoba, Argentina.

Kanda, S., T. Aimi, S. Kano, S. Ishihara, Y. Kitamoto & T. Morinaga. 2008. Ambient pH signaling regulates expression of the serine protease gene (spr1) in pine wilt nematode-trapping fungus, Monacrosporium megalosporum. Microbiol. Res. 163: 63-72.

Kumar, D. & K.P. Singh. 2006. Assessment of predacity and efficacy of Arthrobotrys dactyloides for biological control of root knot disease of tomato. J. Phyto pathol. 154: 1-5.

Larsen, M. 2000. Prospects for controlling animal parasitic nematodes by predacious micro fungi. Parasitology 120: 121-131.

Li, Y., K.D. Hyde, R. Jeewon, L. Cai, D. Vijaykrishna & K. Zhang. 2005. Phylogenetics and evolution of nematode-trapping fungi (Orbiliales) estimated from nuclear and protein coding genes. Mycologia 97: 1034-1046.

Madigan, M.T., J.M. Martinko & J. Parker. 2000. Brock: Biologia de los Microorganismos. Prentice Hall Iberia, Madrid, Espana.

Migunova, V.D & B.A. Byzov. 2005. Determinants of trophic modes of the nematophagous fungus Arthrobotrys oligospora interacting with bacterivorous nematode Caenorhabditis elegans. Pedobiologia 49: 101-108.

Minglian, Z., M. Minghe & Z. Keqin. 2004. Characterization of a neutral serine protease and its full-length cDNA from the nematode-trapping fungus Arthrobotrys oligospora. Mycologia 96: 16-22.

Morgan, M., J.M. Behnke, J.A. Lukas & J.F. Peberdy. 1997. In vitro assessment of the influence of nutrition, temperature and larval density on trapping of the infective larvae of Heligmosomoides polygyrus by Arthrobotrys oligospora, Duddingtonia flagrans and Monacrosporium megalosporum. Parasitology 115: 303-310.

Mota, M.A., A.K. Campos & J.V. Araujo. 2003. Controle biologico de helmintos parasitos de animais: estagio atual e perspectivas futuras. Pesq. Vet. Bras. 23: 93-100.

Nagee, A., A. Acharya, A. Shete, P.N. Mukhopadhyaya & B.A. Aich. 2008. Molecular characterization of an expressed sequence tag representing the cuticle-degrading serine protease gene (PII) from the nematophagous fungus Arthrobotrys oviformis by differential display technology. Genet. Mol. Res. 7: 1200-1208.

Nunez, A.E. 2002. Aislamiento y evaluacion de hongos nematofagos asociados a quistes de Globodera rostochiensis (Woll.) en la region del Cofre de Perote. Master thesis, Universidad de Colima, Colima, Mexico.

Ojeda, N.F., P. Mendoza de Gives, J.F.J. Torres, R.I. Rodriguez & A.J. Aguilar. 2005. Evaluating the effectiveness of a Mexican strain of Duddingtonia flagrans as a biological control agent against gastrointestinal nematodes in goat faeces. J. Helminthol. 79: 151-157.

Orozco, M. 2005. Aislamiento y caracterizacion de hongos nematofagos como potenciales controladores biologicos de nematodos gastrointestinales para la produccion animal. Tesis. Posgrado, Universidad Nacional, Heredia, Costa Rica.

Park, J.O., W. Gams, M. Scholler, E.L. Ghisalberti & K. Sivasithamparam. 2002. Orbiliaceous nematode-trapping fungi and related species in Western Australia and their biological activities. Australasian Mycolo gist 21: 45-52.

Su, H., Y. Hao, M. Mo & K. Zhang. 2007. The ecology of nematode-trapping hyphomycetes in cattle dung from three plateau pastures. Vet. Parasitol. 144: 293-298.

Tilly, J.M. & R.A. Terry. 1963. A two-stage technique for the in vitro digestion of forage crops. J. Br. Grassl. Soc.18: 104-111.

Tunlid, A. & S. Jansson. 1991. Proteases and their involvement in the infection and immobilization of nematodes by the nematophagous fungus Arthrobotrys oligospora. App. Environ. Microb. 57: 2868-2872.

Van Oorschot, C.A.N. 1985. Taxonomy of the Dactylaria complex. A review of Arthrobotrys and allied genera. Stud. Mycol. 26: 61-96.

Varady, M., P. Cudekova & J. Corba. 2007. In vitro detection of benzimidazole resistance in Haemonchus contortus: egg hatch test versus larval development test. Vet. Parasitol. 149: 104-110.

Vargas, C.F. 2006. FAMACHA(c) Control de Haemonchosis en caprinos. Agronomia Mesoamericana 17: 79-88.

Villalba, V. 2006. Manual de laboratorio: Aspectos moleculares de la fitopatologia. Ingenieria en Biotecnologia, Escuela de Biologia, Instituto Tecnologico de Costa Rica, Cartago, Costa Rica.

Zajac, A.M. & G.A. Conboy. 2006. Veterinary Clinical Parasitology. Blackwell, New York, USA.

Natalia Soto-Barrientos (1), Jaqueline de Oliveira (1), Rommel Vega-Obando (2), Danilo MonteroCaballero (3), Bernardo Vargas (4), Jorge Hernandez-Gamboa (1) & Claudio Orozco-Solano (2)

(1.) Catedra de Parasitologia y Enfermedades Parasitarias, Escuela de Medicina Veterinaria, Universidad Nacional, Heredia, Costa Rica;,,

(2.) Centro Nacional Especializado en Agricultura Organica, Instituto Nacional de Aprendizaje, Cartago, Costa Rica;,

(3.) Catedra Salud de Hato, Escuela de Medicina Veterinaria, Universidad Nacional, Heredia, Costa Rica;

(4.) Programa de Posgrado en Ciencias Veterinarias Tropicales, Escuela de Medicina Veterinaria, Universidad Nacional, Heredia, Costa Rica;
Number of farms sampled
according to altitude

Altitude     Number of
(m.a.s.l.)     farms

0-500            17
501-1000         5
1001-1500        10
1501-2000        12
2001-2500        3
2501-3000        1
>3000            3

Number of farms sampled according
to life zone

Life zones *           Number of

Tropical                  1
  dry forest
Premontane moist          7
Lower montane             4
  moist forest
Tropical                  3
  moist forest
Premontane moist          1
  forest, basal belt
Tropical moist            1
  forest, premontane
  belt transition
Tropical moist            1
  forest, perhumid
Lower montane             10
  wet forest
Premontane wet            7
Premontane wet            7
  forest, basal belt
Tropical wet forest       1
Premontane wet            1
  forest, rainforest
Montane wet forest        1
Tropical wet              1
  forest, premontane
Montane rainforest        2
Lower montane             2
Premontane                1

* according to L.R. Holdrige
(Bolanos et al. 2005).

Origin of samples and code

Origin                                  Sample type            Code

Alajuela, San Carlos, Cedral,   Soil and pasture from           ECe
  Finca Los Vega                equine faeces
Alajuela, Grecia, Cariblanco,   Soil and fallen leaves         SCaB
  Refugio Nacional de Vida
Bosque Alegre, Laguna Hule
Alajuela, Los Chiles            Farm soil without management   SCh1
Alajuela, Los Chiles            Farm soil without management   SCh2
Alajuela, San Carlos, Aguas     Pasture soil and dairy cow      VAZ
  Zarcas                        feces
Alajuela, Sabanilla,            Pasture soil and dairy cow      VF
  Fraijanes, Finca La Rosalia   feces
Alajuela, San Carlos, Pital,    Pasture soil, sheep and         VOJ
  Finca La Josefina             dairy cow feces
Alajuela, Poas, San Juan        Pasture soil and dairy cow      VLV
  Norte, Finca La Vistada       feces
Alajuela, San Carlos,           Pasture soil and dairy cow      VM
  Monterrey                     feces
Alajuela, Naranjo               Pasture soil and dairy cow      VN
Alajuela, Poas, Finca La        Pasture soil and dairy cow     VPo1
  Rosela                        feces
Alajuela, Poas, Finca La        Pasture soil and dairy cow     VPo2
  Carmela                       feces
Cartago, Finca La Flor          Pasture soil and dairy goat     CF
Cartago, Turrialba, Juan        Pasture soil and dairy goat     CJV
  Vinas, Finca Ricardo Flores   feces
Cartago, Oreamuno, Cipreses,    Pasture soil and dairy goat     COr
  Finca Los Cipreses            feces
Cartago, faldas del volcan      Nematode diseased potato        PC
  Irazu, antiguamente finca
Cartago, Oreamuno, La           Soil and fallen leaves from     SBI
  Chinchilla, CNEAO-INA         forest
Cartago, Turrialba, Santa       Coffee soil                     SCT
Cartago, Paraiso, Orosi         Ornamental plant soil           SOC
Cartago, Oreamuno, Parque       Soil and fallen leaves from     SVI
  Nacional Volcan Irazu         forest
Cartago                         Nematode infected tomato        TC
Cartago, Alvarado, Capellades   Pasture soil and dairy cow      VAl
Cartago, Finca Cabeza Vaca      Pasture soil and dairy cow      VCV
Cartago, Llano Grande,          Pasture soil and dairy cow     VELG
  Hacienda Retes                feces
Cartago, Oreamuno, Paso         Pasture soil and dairy cow      VO
  Ancho, Finca La Cuesta        feces
Cartago, Alvarado, Pacayas      Pasture soil and dairy cow      VPa
Cartago, Finca Santa Rosa       Pasture soil and dairy cow      VSR
Cartago, Turrialba, Santa       Pasture soil and dairy cow      VT1
  Cruz                          feces
Cartago, Turrialba              Pasture soil and dairy cow      VT2
Guanacaste, Canas, Finca Rio    Pasture soil and feces of       OCC
  Lajas                         sheep and goats
Guanacaste, Canas               Soil and feces of fattening     VCa
                                cattle pasture
Heredia, Barreal, UNA --        Soil and pasture from equine   ECMV
  Escuela de Medicina           y caprine feces
Heredia, Barva, Santa Lucia,    Vermicompost, soil from        VOSL
  Finca Experimental de la      mulberry plantation and
  UNA                           feces of dairy cows and
Limon, Talamanca, Amubri        Soil and pasture from equine    ET
Limon                           Banana soil                    SBL1
Limon                           Banana soil                    SBL2
Limon, Siquirres, Cimarrones    Farm soil without management   SCiL
Limon, Siquirres, Cerro         Farm soil without management    SCS
Limon, Siquirres, La Francia    Farm soil without management    SFL
Limon, Guapiles                 Farm soil with organic          SG
Puntarenas, Buenos Aires,       Soil and fallen leaves from    SAPt
  Biolley, Parque               forest
  Internacional La Amistad
Puntarenas, Montes de Oro,      Farm soil without management    SM
Puntarenas, Puerto Jimenez      Farm soil without management    SZS
San Jose, Uruca, INA            Soil and feces from goats      CINA
San Jose, Zona de los Santos,   Soil and fallen leaves from     SPQ
  Parque Nacional Los           forest
San Jose, Dota, Santa Maria     Coffee soil                     SD
San Jose, Tarbaca               Farm soil without management    STa
San Jose, Sabanilla,            Soil and fallen leaves from    SUCRD
  Instalaciones deportivas      forest
  de la UCR
San Jose, Zapote                Squash soil                     SZa
San Jose, Coronado              Pasture soil and dairy cow      VCo
San Jose, Rancho Redondo,       Pasture soil and dairy cow      VSo
  Finca La Socola               feces

pH of the fungal isolate source and number of fungi isolated
by province, altitude and life zone


                               Arthrobotrys   Arthrobotrys
                                 conoides     dactyloides

pH            Least              5.6 (SD)      7.1 (SZa)
              Maximum            5.6 (SD)      7.1 (SZa)
              Average            5.6 (SD)      7.1 (SZa)

Province      Alajuela              0              0
              Cartago               0              0
              Guanacaste            0              0
              Heredia               0              0
              Limon                 0              0
              Puntarenas            0              0
              San Jose              1              1

Altitude      0-500                 0              0
(m.a.s.l.)    501-1000              0              0
              1001-1500             0              1
              1501-2000             1              0
              2001-2500             0              0
              2501-3000             0              0
              >3000                 0              0

Life zone *   Premontane            0              1
                moist forest
              Lower                 0              0
                moist forest
              Tropical moist        0              0
              Lower montane         1              0
                wet forest
              Premontane wet        0              0
              Premontane            0              0
                wet forest,
                basal belt
              Premontane wet        0              0
              Montane wet           0              0
              Montane               0              0
              Lower montane         0              0


                               Arthrobotrys   Candelabrella
                                oligospora     musiformis

pH            Least              5.6 (SD)       5.9 (SCT)
              Maximum           7.5 (SFL)       7.6 (SM)
              Average              6.6             6.7

Province      Alajuela              1               1
              Cartago               5               2
              Guanacaste            0               0
              Heredia               4               1
              Limon                 1               3
              Puntarenas            0               2
              San Jose              2               0

Altitude      0-500                 1               5
(m.a.s.l.)    501-1000              1               1
              1001-1500             5               1
              1501-2000             4               1
              2001-2500             0               1
              2501-3000             1               0
              >3000                 1               0

Life zone *   Premontane            1               5
                moist forest
              Lower                 0               2
                moist forest
              Tropical moist        2               0
              Lower montane         1               2
                wet forest
              Premontane wet        1               0
              Premontane            2               1
                wet forest,
                basal belt
              Premontane wet        1               0
              Montane wet           0               1
              Montane               0               1
              Lower montane         1               1

* according to L.R. Holdrige (Bolanos et al. 2005).

Chlamydospores production by the predatory nematophagous fungi
isolated (1)

Code        Fungi        CMA (2)   PDA (2)   PCA (2)   MEA (2)

ECe     C. musiformis       -         -        + +        -
VLV     A. oligospora      + +       + +        +        + +
COr     A. oligospora       +        + +        +        + +
PC      A. oligospora      + +       + +       + +        +
SBI     A. oligospora      + +      + + +     + + +       +
SCT     C. musiformis       -        + +        -         -
TC      A. oligospora       -         -         -         -
VAl     C. musiformis     + + +     + + +     + + +       -
VSR     A. oligospora       -        + +      + + +      + +
1OSL    C. musiformis       +        + +      + + +       +
2OSL    A. oligospora       -         -         +         -
3OSL    A. oligospora       -         -        + +        -
VSL     A. oligospora       +         +         +         -
MSL     A. oligospora      + +        +         -         -
ET      C. musiformis       +       + + +     + + +       -
SCL     C. musiformis     + + +     + + +     + + +       +
SCS     C. musiformis     + + +       +        + +        +
SFL     A. oligospora       -        + +       + +        -
SAPt    C. musiformis      + +        +         -         -
SM      C. musiformis     + + +     + + +     + + +       +
CINA    A. oligospora       +         +        + +        +
1SD     A. oligospora       -         -       + + +       -
2SD     A. conoides         +         -         -         +
SZa     A. dactyloides      +        + +      + + +       +

(1.) (-) no chlamydospores were observed; (+) indicated that there
were 1 to 5 chlamydospores total in the sample taken and they
were hard to find; (++) it was easy to find chlamydospores,
however, there were not too many; (+++) there were plenty of
chlamydospores; some were even forming chains.

(2.) CMA=corn-meal agar; PDA=potato-dextrose agar; PCA=potato-carrot
agar; MEA=malt-extract agar.

Summary data of the nematophagous fungi positive to the viability

Code      Fungus               Origin            Altitude

VLV    A. oligospora   Poas, Alajuela              2500
COr    A. oligospora   Oreamuno, Cartago           1700
SBI    A. oligospora   La Chinchilla, Cartago      1453
ET     C. musiformis   Talamanca, Limon             90
SCS    C. musiformis   Siquirres, Limon            358

SM     C. musiformis   Miramar, Puntarenas         340

Code      Fungus             Life zone (1)          pH    Capture

VLV    A. oligospora   Lower montane wet forest     6.7   78.8
COr    A. oligospora   Lower montane moist forest   7.1   79.5
SBI    A. oligospora   Lower montane moist forest   6.6   72.3
ET     C. musiformis   Tropical moist forest        7.4   94.2
SCS    C. musiformis   Premontane wet forest,       6.1   88
                         basal belt transition
SM     C. musiformis   Tropical moist forest        7.6   92.5

(1.) according to L.R. Holdrige (Bolanos et al. 2005).
COPYRIGHT 2011 Universidad de Costa Rica
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2011 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Soto-Barrientos, Natalia; de Oliveira, Jaqueline; Vega-Obando, Rommel; Montero- Caballero, Danilo; V
Publication:Revista de Biologia Tropical
Article Type:Report
Date:Mar 1, 2011
Previous Article:Plasma concentration of progesterone and 17[beta]-estradiol of black-rumped agouti (dasyprocta prymnolopha) during the estrous cycle.
Next Article:Distribucion, parametros poblacionales y dieta de Astropecten marginatus (Asteroidea: Astropectinidae) en el Atlantico venezolano.

Terms of use | Privacy policy | Copyright © 2020 Farlex, Inc. | Feedback | For webmasters