Nematophagous Fungus: Pochonia chlamydosporia and Duddingtonia flagrans in the Control of Helminths in Laying Hens (Gallus gallus domesticus) Genus Hy-line Brown - Evaluation and Effectiveness

The resistance to anthelmintics in poultry farming and the challenges with the restricted use of drugs in organic farms make the use of biological controllers an innovative bridge to verminosis control. This paper aims to evaluate the efficacy of the larvicidal fungus Duddingtonia flagrans and the ovicidal fungus Pochonia chlamydosporia in Hy-line Brown (Gallus gallus domesticus) layer farms. Both fungi were combined in the core of the feed administered to the birds of the treated groups. 28,000 birds were used and divided into three treated groups (TG) in which the feed containing the fungus was administered. The poultry house itself manufactures the birds’ feed; the formula was included in the nucleus at a dose of 100 grams per ton of feed for 7 months. The concentration was 105 chlamydospores of P. chlamydosporia and D. flagrans per gram of the formulation. The control group (CG) received regular food from the farm. The birds were separated into four sheds with 7,000 birds in each. The egg per gram of feces (EPG) testing was performed using fresh fecal samples collected from the sheds over six months. Weather data was collected during the experiment. There was a reduction in the EPG count into three treated groups. The most prevalent helminth was the genus Ascaris. The formulation tested shows little efficacy in this dosage.

Key points:

• The use of the fungus P. chlamydosporia and D. flagrans in organic farms;
• Use of biocontrol agents without chemicals in poultry;

The organic production systems farms have grown and gained market space in several countries. Aiming at the welfare of farm animals and the reduction of environmental impact, farms face a challenge: verminosis. Parasite control is affected by both system restrictions, outdoor time, and access to soil, intermediate hosts, and resistance to antiparasitic drugs [1]. Ascaridia infections are common in poultry and cause a range of damage. Welfare and health are affected, leading to diarrhea and dehydration, as well as decreased production, which negatively affects egg quality [1,2]. 

The most prevalent parasites comprise the nematode group, causing secondary diseases with high prevalence rates. The genera Ascaridia and Capillaria cause significant productive losses. Another common genus is Heterakis gallinarum, responsible for transmitting the Histomonas meleagridis agent, which has a mortality rate of approximately 35% in flocks, causing hyporexia, anorexia, and weight loss, leading to death [3,4]. Some genera of the Cestode class are found mainly in organic and free-range systems as they depend on the presence of intermediate hosts such as flies, earthworms, and others. The main cestode is Raillietina spp., responsible for anorexia and decreased egg production [4].

Resistance to anthelmintics is a global problem and aggravated by the incorrect use of chemotherapeutics, together with erroneous systematics in their administration [5]. The use of nematophagous fungi has obtained effective and promising results, associated or isolated in the control of verminosis. They are present in the environment, such as Duddingtonia flagrans (D. flagrans), which is classified as a larvicide by preying on larval stages through its three-dimensional network with adhesive hyphae. This species presents a great capacity to produce chlamydospores and conidia in its conidiophores. Remaining viable after passage through the gastrointestinal tract of animals, they develop in the fecal microenvironment, thus preying on and destroying the larvae. Due to the resistance of the chlamydospores, they can multiply in the environment and thus reduce the parasite load in the environment in the long term [5-7].

The fungus Pochonia chlamydosporia (P. chlamydosporia) is widely used as a biocontrol agent and classified as an opportunistic saprophytic ovicide; the presence of helminth eggs is not essential for its maintenance in the environment [8,9]. This species can pass through the gastrointestinal tract, colonize the fecal bolus, and, in the presence of helminth eggs, prey on them. With hyphae that use the apprehension mechanism to colonize the eggs by mechanical and enzymatic action of its proteases, thus destroying the eggs of the parasite [10,11]. Such enzymatic action is intrinsically linked to the protease-serine-alkaline present in the infection process, being responsible for the extenuation of the yolk membrane of the eggshell. This weakening also occurs due to the hydrolytic enzymatic action, the proteases and kinases that act in the degradation of eggs. Such action has been successfully tested in the field and in laboratory conditions, capable of destroying helminth eggs in their various embryonic stages [5,12].

The environment is an important factor in the efficiency of these fungi, since it interferes with the growth and dispersion of the mycelia. However, D. flagrans and P. chlamydosporia do not suffer limiting variations due to temperature when their applicability is aimed at parasite control [5,13]. Biological control using P. chlamydosporia has proven to be an excellent alternative for environmental prevention. Besides the action against nematodes, it also has mechanisms of action on paratenic hosts, intermediates, and vectors [5].

This experiment aimed to observe the efficacy of the two fungi in the control of gastrointestinal helminthiasis in laying hens in the organic management system.

The experiment was conducted on a private property in the municipality of Mogi-Guaçu, metropolitan region of Campinas, state of São Paulo, Brazil. The poultry farm is human-certified organic and does not use any type of chemotherapy for the treatment of helminths. Each group in the study corresponds to one poultry house, with the same age and approximate weight. The layers have access to the picketing area and are housed in sheds of 1,000 square meters; each shed holds 7,000 layers. The experimental groups were subdivided into three that received the treatment and a control group that received the feed formulation manufactured by the farm itself. Each group is composed of 7,000 birds at 46 weeks of age, all females, weighing approximately 1.8 kg each, of the hy-line brown breed. The experiment lasted 7 months, beginning in October 2021 and ending in May 2022, corresponding to the change from dry to rainy season.

The formulation was made from fungal cultures of D. flagrans and P. chlamydosporia using the fungi from the Mycology Collection of Parasitology Laboratory, Department of Veterinary, Federal University of Viçosa, MG. The mycelia were placed with rice grain, subsequently ground, forming rice bran. The final concentration was 105 [14-16]. The product is administered with the feed. The poultry house itself manufactures the chickens’ feed; the formula was included in the nucleus at a dose of 100 grams per ton of feed. No studies were found that establish the dose of associated fungi for laying hens. This was an initial dose in an attempt to establish the proper dosage. In this way, daily administration was continuous during the seven months of the experiment.

The main ingredients in the feed manufactured by the farm are, in ascending order: corn, soy bran, fine limestone, coarse limestone, bicalcium phosphate, salt, methionine, threonine, and bicarbonate. Those responsible for the farm only know the quantity order used do not pass on the percentage of ingredients. The feed is manufactured by the poultry every 15 days and offered together with the fungi for the treated groups. As it is an organic farm, the ingredients used cannot have chemical residues and are organic as well. 

The nucleus used in the chickens’ feed is composed of minerals, vitamins, and micronutrients. The use of any antimicrobial and artificial preservatives is not allowed by the regulatory policies [17]. The laying hens have access to the pasture area for 6 to 8 hours during the day. Each shed has a grazing area of just over a thousand square meters. Composed of grasses, shading, and fodder that are essential for the health and manifestation of the natural behavior of chickens.

During the seven months of the experiment, feces were collected every 15 days (except in January, due to excessive rainfall and lack of access). The feces collected from the barn and from the release pens, always preferring the freshest feces, thus forming a pool of feces. The samples analysis using the EPG determined by Gordon and Whitlock 1939 and modified by Lima 1989 [18,19]. The procedures follow the guidelines of the World Association for the Advancement of Veterinary Parasitology (WAAVP) [13]. The eggs were counted and identified for standardized averages and statistics prescribed by WAAVP. In order to identify and differentiate these eggs, according to Mattos, et al. 2019 [20], Ascaridia eggs have a thick and smooth wall, internally with granulation at one of the extremities. Heterakis eggs, on the other hand, are ovoid, with a smooth and thick shell. Furthermore, Ascaridia galli eggs are brown in color, and Heterakis gallinarum eggs have a double ovigerous [21].

The meteorological data collected during the experiment, as well as mortality and production data for each group. 

The means obtained and the data were submitted to Student’s t-test with a significance level of 5% or p > 0.005.

The EPG values over time its demonstrated in Figure 1. For the analysis of this data, the standardization of the arithmetic mean was used, considering the standard deviations. A reduction in the EPG values is noticed in March, April, and May. There is an increase in the EPG values in the control group for December.

The association of P. chlamydosporia and D. flagrans fungi showed no statistical difference when the treated groups were compared to the control group.

Pairwise comparisons showed a statistical difference between the treated groups. However, when comparing each treated group with the control group, there is no difference (p > 0.005). Table 1 shows the differential values between the groups when compared in pairs.

When compared to the control group, the treated groups showed no statistical difference. Such data may result from the short time of administration of the formulation, since the fungi act in the fecal microenvironment and disperse throughout the environment. As these layers are outdoors, the challenge is even greater, with higher chances of reinfection by the parasites, besides the possibility of greater contact with intermediate agents [21].

Comparing the production data with the mean EPG, no significant results were observed (p > 0.005) (Table 1). It is a fact that low-quality nutrition decreases productivity and affects the welfare and immunity of these animals with the increased exposure to various parasites present in organic farming [22].

The percentage of eggs found in the treated and control groups is shown in Figure 2. The EPG was performed using the method by Gordon and Whitlock, modified by Lima, and the identification was performed by the morphological characteristics of each egg. The eggs of the genus Ascaridia have a thick and smooth wall, internally with granulation at one end. The eggs of the genus Heterakis are ovoid, with a smooth and thick shell [20]. In addition, Ascaridia eggs are brown in color, and Heterakis eggs have a double ovigerous capsule [21]. Despite the identification of eggs, both genera were allocated as genus Ascaris to avoid any misinterpretation of the presented results, due to the identification difficulty in some samples. The higher prevalence of the Ascaris genus has a correlation with the indirect life cycle of both parasites [23-25].

The meteorological data were collected during the months of the experiment, and the temperatures and precipitation of the months were averaged. The experiment was conducted at the end of the dry season and the beginning of the rainy season. The average minimum temperature was 23.5 °C in October and the maximum was 30 °C in February. The month corresponding to the highest average precipitation is January with 67% and the lowest was April with 29% (Figure 3).

Other studies [23,24] have already performed the correlation between climate and D. flagrans, which has its greatest action in tropical climates with higher rainfall, similar conditions for the development of infective larvae; however, the most prevalent parasites in this study do not have free-living larval stages, except for S. avium. Probably one of the reasons why the results are not significant. Although P. chlamydosporia has ideal growing conditions with humidity and average temperature between 24 °C and 32 °C, its growth is generally not affected by the climatic conditions. Most probably that more time is needed for dispersal and action of the fungus on the eggs [24].

The comparison with mortality and EPG data did not show significant results (p > 0.005); only the G1 group was significant when compared to the CG (control - untreated group). This result can be justified by the age factor; the birds had an average of 46 weeks of age, and since the younger the bird, the more susceptible it is to parasitic mortality [25].

The results obtained corroborate those found by Fonseca, et al. [26], in which the authors obtained negative results using a formulation with P. chlamydosporia fungus to control nematodes in cattle. These results can be explained by strategies developed by the parasites themselves, such as the rapid transition from egg to larval stage, which impairs the contact of the fungus with the egg and, consequently, its mechanism of ovicidal action. The same may have occurred with the results found in this study, and although negative, the authors describe that the enzymatic mechanisms of P. chlamydosporia and adhesion in eggs it is demonstrated in several studies in the literature [27,28].

Rabbi, et al. in 2006 related a higher prevalence of A. galli in a study performed in Bangladesh. These authors indicate that the resistance of the eggs and their indirect life cycle increases the chances of infection. This fact corroborates the data found during this research since it belongs to the genus Ascaridia. [29] found a higher prevalence of A. galli in a study carried out in India and correlated a rise in backyards, ineffective sanitation of the environment, and malnutrition as a justification for this prevalence. Biological control can be an attractive alternative to reduce infective stages of helminths in grazing areas, with the advantage of the absence of chemical residues.

According to Thapa, et al. [30], in a study on the epidemiology and prevalence of helminths in several European countries, they found A. galli as the most prevalent parasite in organic farms. However, there was no positive epidemiological correlation between the parasitic loads of birds reared indoors or outdoors. This fact demonstrates how complex the infection process of these helminths is. This leads to the need for alternatives to control helminths.

The study carried out in Ethiopia by [31] found a high prevalence of helminths and corroborates the incidence of A. galli and H.gallinarum, differentiated by the higher percentage of H. gallinarum [31-33] had similar results, with the highest prevalence of H. gallinarum. Such data indicate that the location, type of management, sanitation, and biosecurity are linked to the parasites found. It should be considered that the different parasites found are also influenced by the method chosen for analysis and identification of eggs.

The present study was carried out in a commercial farm, making it impossible to perform necropsies, even of naturally occurring deaths. Raillietina tetragona was present in both cited studies [31-33]; however, there was no such finding in this study. Demonstrating that there is the possibility of controlling the parasite in the environment in which the present study was performed. Ovicidal fungi have their action mainly on helminths that have their cycle phases in the environment and persist in the egg phase, among which the Ascaridae can be mentioned [34].

H. gallinarum depends on the presence of intermediate hosts for infection. The life cycle of this parasite occurs through the ingestion of infective eggs carried by infected earthworms. His presence correlated with living in avian enclosures. S. avium infects the host by infection of the free-living larvae or by the infected egg. S. trachea is transmitted by ingestion of the intermediate host. It is most probable that the decrease of S. avium during the analysis is associated with the larvicidal action of D. flagrans. Observing the life cycle of these parasites, one realizes that free-range and organic farms are challenged with parasitic infections, as the soil is the habitat of the intermediate hosts and the parasites [29,31-33]. In agreement with [35], obtained in vitro large reduction in the percentage of the aquatic snail population Biomphalaria tenagophila was obtained using the fungus P. chlamydosporia on intermediate hosts of Schistosoma mansoni.

The decrease in the percentage of S. trachea found during the study period is linked to the ovicidal action of P. chlamydosporia. According to Fonseca, et al. 2023 [34], there are reports of the activity of P. chlamydosporia against bird helminth eggs. This activity is described both in vitro and in vivo under field conditions, with the use of P. chlamydosporia isolated or in association with another species of helminth fungus. As in the work of Braga, et al. 2011, which achieved a 64.1% reduction compared to the control group with the use of crude extract of P. chlamydosporia in the control of Ascaridia galli eggs [34]. This predatory action is shown in other studies in different species and parasite eggs [36,37].

This predatory action is also demonstrated in other studies in different species and parasite eggs, in addition to bird parasite eggs. Among these are cattle, horses, canines, felines, pigs, and humans ([13,43]. In these species, besides in vivo studies [5,38,39], there are several in vitro studies with the action of this fungus against eggs of Fasciola hepatica, Oxyuris equi, Eurytrema coelomaticum, Ascaris suum, Ancylostoma sp., among others [40]. This broad action demonstrates the ovicidal capacity of P. chlamydosporia and a promising commercial use.

As we did not have authorization from the farm to carry out the necropsy, the chosen method was the EPG, which is a valid method, widely used in many scientific papers consolidated in the literature. Most of the scientific groups in the world use this method, and the technique is used in almost every available study in helminthology. The study performed by [41] used Heterakis gallinarum to evaluate the effectiveness of EPG in relation to parasite load. The test proved efficient in fecal samples mixed daily, demonstrating a positive correlation between the real parasitic burden of the chickens and the values found in the EPG. This corroborates the choice of method in this present study.

Other studies [36,37] have already made the correlation between climate and D. flagrans, which has its greatest action in tropical climates with higher rainfall, similar conditions for the development of infective larvae; however, the most prevalent parasites in this study do not present free-living larval stages, except S. avium. This may be one reason why the results are not significant. Although P. chlamydosporia has ideal growth conditions with humidity and average temperature between 24 °C and 32 °C, its growth is generally not affected by climatic conditions. Most likely, more time is needed for dispersal and action of the fungus on the eggs [36].

The use of P. chlamydosporia and D. flagrans associated with this concentration showed low efficacy in the control of intestinal worms in poultry. We believe that longer-duration studies may have positive results, given the characteristics of the fungi. It is possible that with higher dosages of the fungi, better results could be achieved. The proposed dosage was low compared to other species. The need factor for organic farms is also of extreme importance, as they require alternatives without chemical residues and are in line with the principles of this type of production.

Despite the challenges encountered in organic farming, the welfare and demand for organic products are becoming a trend. With longer studies, nematophagous fungi will prove more effective, as there is a need for solutions that do not embrace chemotherapeutics and anthelmintic resistance to them. 

Despite the statistical differences between the treated groups, when compared to the control group, there was no significant efficacy of using Pochonia chlamydosporia and Duddingtonia flagrans in controlling helminths in hi-line brown layers from organic farms in this present dosage.

Updated conclusion

In this large-scale field study, the combined use of Pochonia chlamydosporia and Duddingtonia flagrans at the tested dosage did not demonstrate significant efficacy in controlling helminths in Hy-Line Brown laying hens under organic production conditions. Nonetheless, the trial provides valuable insights into the practical application of nematophagous fungi as an alternative to chemical anthelmintics.

The findings highlight both the potential and the challenges of implementing biological control in poultry farming. The replication of treatment across three sheds reinforces the reliability of the results, even though no statistical difference was observed compared to the control group. It is likely that higher dosages, longer treatment periods, or integrated strategies combining fungi with improved farm management could yield more promising outcomes.

Importantly, this study contributes to the limited body of knowledge on helminth control in commercial organic poultry systems, where chemical treatments are restricted. The use of nematophagous fungi represents a sustainable and residue-free strategy that aligns with the principles of organic farming and animal welfare.

Future research should focus on testing different concentrations, longer-term administration, and complementary management practices, as well as including microscopy-based documentation of parasite eggs. Such approaches will be essential for advancing biological control methods and offering effective, eco-friendly solutions to poultry farmers worldwide.

Declarators

Author contributions: Conceptualization, J.V.d.A. and I.A.F.; methodology, J.V.d.A and I.A.F.; software, I.S.V.; formal analysis, I.A.F; investigation, I.A.F; resources, I.A.F.; writing—original draft preparation, I.A.F; writing—review and editing, J.V.d.A, I.A.F, J. d.S.F and L.M.d.C. Supervision, J.V.A. All authors have read and agreed to the published version of the manuscript

Institutional review board statement: The experiment follows the ethics and animal welfare guidelines recommended by the Animal Use Ethics Committee (CEUA/UFV). Has been submitted, approved, and registered under number 26/2022 on 07/11/22.

Data availability statement: Available upon request from the corresponding author.

We are grateful as well to the UFV Veterinary Department, FAPEMIG, CNPQ, and CAPES for all support.

1. Thamsborg SM, Roepstorff A, Larsen M. Integrated and biological control of parasites in organic and conventional production systems. Vet Parasitol. 1999;84(3-4):169-186. Available from: https://doi.org/10.1016/s0304-4017(99)00035-7
2. Shifaw A, Feyera T, Walkden-Brown SW, Sharpe B, Elliott T, Ruhnke I. Global and regional prevalence of helminth infection in chickens over time: A systematic review and meta-analysis. Poult Sci. 2021;100(5):101082. Available from: https://doi.org/10.1016/j.psj.2021.101082
3. Cardozo SP, Yamamura MH. Parasites in poultry production systems in the colonial/backyard farming system in Brazil. Semin Cienc Agrar. 2004;25(1):63-74. Available from: https://doi.org/10.5433/1679-0359.2004v25n1p63
4. Zamboni R, Alberti TS, Venancio FR, Scheid HV, Brunner CB, Raffi MB, et al. Histomoniasis in chickens (Gallus gallus domesticus) from colonial farming systems in southern Brazil. Med Veterinária (UFRPE). 2021;15(4):370-375. Available from: http://dx.doi.org/10.26605/medvet-v15n4-3537
5. Araújo JV, Braga FR, Mendoza-de-Gives P, Paz-Silva A, Vilela VLR. Recent advances in the control of helminths of domestic animals by helminthophagous fungi. Parasitologia. 2021;1:168-176. Available from: https://doi.org/10.3390/parasitologia1030018
6. Vieira VD, Riet-Correa W, Vilela VLR, Medeiros MA, Batista JÁ, Melo LRB, et al. Control of gastrointestinal nematodes in sheep and financial analysis on a farm with irrigated rotational grazing system in the Brazilian semi-arid region. Pes Vet Bras. 2018;38(5). Available from: https://doi.org/10.1590/1678-5150-PVB-5400
7. Mota MA, Campos AK, Araújo JV. Biological control of helminth parasites of animals: current stage and future outlook. Pesq Vet Bras. 2003;23(3):93-100. Available from: https://doi.org/10.1590/S0100-736X2003000300001
8. Li S, Wang D, Gong J, Zhang Y. Individual and combined application of nematophagous fungi as biological control agents against gastrointestinal nematodes in domestic animals. Pathogens. 2022;11:172. Available from: https://doi.org/10.3390/pathogens11020172
9. Gams W, Zare R. A revision of Verticillium sect. Prostrata. III. Generic classification. Nova Hedwigia. 2001;73:329-337. Available from: https://doi.org/10.1127/nova.hedwigia/72/2001/329
10. Araújo JV, Mota MA, Campos AK. Biological control of helminth parasites of animals by nematophagous fungi. In: Congress of Veterinary Parasitology & Latin American Symposium on Rickettsioses, Ouro Preto, Brazil. Rev Bras Parasitol. 2004;13:165-170.
11. Braga F, Silva A, Carvalho R, Araújo J, Pinto P. Ovicidal activity of different concentrations of Pochonia chlamydosporia chlamydospores on Taenia taeniaeformis eggs. J Helminthol. 2011;85(1):7-11. Available from: https://doi.org/10.1017/s0022149x10000179
12. Lelis RT, Braga FR, Carvalho LM, Paula AT, Araújo JM, Fausto MC, et al. Effect of the fungus Pochonia chlamydosporia on Echinostoma paraensei (Trematoda: Echinostomatidae). Acta Tropica. 2014;139:88-92. Available from: https://doi.org/10.1016/j.actatropica.2014.07.006
13. Yazwinski TA, Chapman HD, Davis RB, Letonja T, Pote L, MaeS L, et al. WORLD ASSOCIATION FOR THE ADVANCEMENT OF VETERINARY PARASITOLOGY. World Association for the Advancement of Veterinary Parasitology (WAAVP) guidelines for evaluating the effectiveness of anthelmintics in chickens and turkeys. Vet Parasitol. 2003;116(2):159-173. Available from: https://doi.org/10.1016/s0304-4017(03)00264-4
14. Rodrigues JVF, Braga FR, Campos AK, Carvalho LM, Araujo JM, Aguiar AR, et al. Duddingtonia flagrans formulated in rice bran in the control of Oesophagostomum spp., an intestinal parasite of swine. Exp Parasitol. 2018;184:11-15. Available from: https://doi.org/10.1016/j.exppara.2017.11.001
15. Silva ME, Araújo JV, Braga FR, Borges LA, Soares FE, Lima WS, et al. Mycelial mass production of fungi Duddingtonia flagrans and Monacrosporium thaumasium under different culture conditions. BMC Res Notes. 2013;6:340-343. Available from: https://doi.org/10.1186/1756-0500-6-340
16. Dallemole-Giaretta R, Freitas LG, Caixeta LdB, Xavier DM, Ferraz S, Fabry CdFS. Production of chlamydospores of Pochonia chlamydosporia in different substrates. Ciência e Agrotecnologia. 2011;35. Available from: https://doi.org/10.1590/S1413-70542011000200012
17. BRASIL. Normative Instruction No. 64 of December 18, 2008. Technical regulation for organic animal and plant production systems and the lists of substances allowed for use in organic production systems for animals and plants. Official Gazette of the Federal Republic of Brazil. 19 Dec 2008;Section 1:21-26.
18. Gordon HM, Whitlock HV. A new technique for counting nematode eggs in sheep faeces. J Counc Sci Ind Res. 1999;12(1):50-52. Available from: https://www.cabidigitallibrary.org/doi/full/10.5555/19390800035
19. Lima WS. Dynamics of gastrointestinal parasitic nematode populations in beef cattle, some aspects of the parasite-host relationship, and the behavior of free-living stages in the Vale do Rio Doce region, MG, Brazil. PhD Thesis, Instituto de Ciências Biológicas, Universidade Federal de Minas Gerais. 1989.
20. Mattos PM, Rossato MR, Antonucci AM. Main parasites in industrial poultry (broilers, hens, and turkeys) - Literature review. Ver Bras Med Vet. 2019;32:1679-7353. Available from: http://faef.revista.inf.br/imagens_arquivos/arquivos_destaque/ssv7BzkR7dcYpml_2019-4-3-14-6-10.pdf
21. Boroviec BB, Gasparotto PHG, Filho JVD, Peixoto RMP, Viana GA, Rocha ASCM, et al. Occurrence of Ascaridia galli and Heterakis gallinarum in Numida meleagris (Helmeted Guineafowl) in the state of Rondônia, Brazil. Acta Sci Vet. 2020;48:487. Available from: https://www.cabidigitallibrary.org/doi/pdf/10.5555/20203378125
22. McDougald LR. Invited Minireview: Blackhead Disease (Histomoniasis) in Poultry: A Critical Review. Avian Dis. 2005;49(4):462–476. Available from: https://doi.org/10.1637/7420-081005r.1
23. Baptista AF. Parasitological profile in free-range chickens, Technical University of Lisbon. 2010. Available from: http://hdl.handle.net/10400.5/1843
24. Vita GF. Efficacy of the active principles of the medicinal plant Chenopodium ambrosioides Linnaeus, 1786 (Erva-de-Santa-Maria), in the control of endoparasites in Gallus gallus (Free-Range Chicken) and Coturnix japonica (Japanese Quail). Master's Thesis (Animal Biology) - Institute of Biological and Health Sciences, Universidade Federal Rural do Rio de Janeiro. 2013. Available from: https://rima.ufrrj.br/jspui/handle/20.500.14407/10806
25. Santiago DC, Homechin M, Silva JFV, Ribeiro ER, Gomes BC, Santoro PH. Selection of isolates of Paecilomyces lilacinus (Thom.) Samson to control Meloidogyne paranaensis in tomato. Rural. 2006;36(4). Available from: https://doi.org/10.1590/S0103-84782006000400003
26. Fonseca JD, Ferreira VM, Freitas SG, Vieira IS, Araújo JV. Efficacy of a fungal formulation with the nematophagous fungus Pochonia chlamydosporia in the biological control of bovine nematodiosis. Pathogens. 2022;11. Available from: https://doi.org/10.3390/pathogens11060695
27. Kerry BR, Hidalgo L. Application of Pochonia chlamydosporia in the integrated control of root-knot nematode on organically grown vegetable crops in Cuba. IOBC/WPRS Bull. 2004;27:123–126. Available from: https://www.cabidigitallibrary.org/doi/full/10.5555/20043147294
28. Braga FR, Araújo JV, Soares FEF, Tavela AO, Araujo JM, Carvalho RO, et al. Enzymatic analysis and in vitro ovicidal effect of Pochonia chlamydosporia and Paecilomyces lilacinus on Oxyuris equi eggs of horses. Biocontrol Sci Technol. 2012;22:685-696. Available from: https://doi.org/10.1080/09583157.2012.677807
29. Solanki JB, Kumar N, Varghese A, Thakre BJ, Puri G. Prevalence of Gastro-intestinal Parasitism in Poultry in and Around Navsari Area of South Gujarat. Int J Livest Res. 2020;3:28-30. Available from: https://www.researchgate.net/publication/344071585_Prevalence_of_Gastro-intestinal_Parasitism_in_Poultry_in_and_Around_Navsari_Area_of_South_Gujarat
30. Thapa S, Mejer H, Thamsborg SM, Lekfeldt JDS, Wang R, Jensen B, et al. Survival of chicken ascarid eggs exposed to different soil types and fungi. Appl Soil Ecol. 2017;121:143–151. Available from: https://doi.org/10.1016/j.apsoil.2017.10.001
31. Berhe M, Mekibib B, Bsrat A, Atsbaha G. Gastrointestinal Helminth Parasites of Chicken under Different Management Systems in Mekelle Town, Tigray Region, Ethiopia. J Vet Med. 2019;1307582. Available from: https://doi.org/10.1155/2019/1307582
32. Rabbi AKMA, Islam A, Majumder S, Anisuzzaman A, Rahman MH. Gastrointestinal helminth infection in different types of poultry. Bangl J Vet Med. 2006;4(1):13-18. Available from: https://doi.org/10.3329/bjvm.v4i1.1519
33. Ogbaje CI, Agbo EO, Ajanusi OJ. Prevalence of Ascaridia galli, Heterakis gallinarum and Tapeworm Infections in Birds Slaughtered in Makurdi Township. Int J Poult Sci. 2012;11:103-107. Available from: http://dx.doi.org/10.3923/ijps.2012.103.107
34. Fonseca JS, Castro LS, Carvalho LM, Soares FEF, Braga FR, Araújo JV. Nematophagous fungus Pochonia chlamydosporia to control parasitic diseases in animals. Appl Microbiol Biotechnol. 2023;1-10. Available from: https://doi.org/10.1007/s00253-023-12525-0
35. Castro LS, Martins IVF, Tunholi-Alves VM, Amaral LS, Pinheiro J, de Araújo JV, et al. Susceptibility of embryos of Biomphalaria tenagophila (Mollusca: Gastropoda) to infection by Pochonia chlamydosporia (Ascomycota: Sordariomycetes). Arch Microbiol. 2022;204:271. Available from: https://doi.org/10.1007/s00203-022-02894-x
36. Vieira IS, Oliveira ID, Freitas SG, Campos AK, de Araujo JV. Arthrobotrys cladodes and Pochonia chlamydosporia in the biological control of nematodiosis in an extensive bovine production system. Parasitology. 2020;147(6):699-705. Available from: https://doi.org/10.1017/S0031182020000098
37. Baudena MA, Chapman MR, Larsen M, Klei TR. Efficacy of the nematophagous fungus Duddingtonia flagrans in reducing equine cyathostome larvae on pasture in south Louisiana. Vet Parasitol. 2000;89:219-230. Available from: https://doi.org/10.1016/S0304-4017(00)00201-6
38. Carvalho RO, Araújo JV, Braga FR, Araujo JM, Alves CDF. Ovicidal activity of Pochonia chlamydosporia and Paecilomyces lilacinus on Toxocara canis eggs. Vet Parasitol. 2010;169:123–127. Available from: https://doi.org/10.1016/j.vetpar.2009.12.037
39. Braga FR, Araujo JM, Silva AR, Araújo JV, Carvalho RO, Soares FE, et al. Ovicidal activity of the enzymatic crude extract of the fungus Pochonia chlamydosporia on Ancylostoma sp. eggs. Rev Soc Bras Med Trop. 2011;44:116–118. Available from: https://doi.org/10.1590/S0037-86822011000100027
40. Braga FR, Araújo JV. Nematophagous fungi for biological control of gastrointestinal nematodes in domestic animals. Appl Microbiol Biotechnol. 2014;98:71–82. Available from: https://doi.org/10.1007/s00253-013-5366-z
41. Daş G, Savaş T, Kaufmann F, Idris A, Abel H, Gauly M. Precision, repeatability and representative ability of faecal egg counts in Heterakis gallinarum infected chickens. Vet Parasitol. 2011;183(1-2):87-94. Available from: https://doi.org/10.1016/j.vetpar.2011.07.005
42. Fernández SA, Henningsen E, Larsen M, Nansen P, Grønvold J, Søndergaard J. A new isolate of the nematophagous fungus Duddingtonia flagrans is a biological control agent against free-living larvae of horse strongyles. Equine Vet J. 1999;31(6):488–491. Available from: https://doi.org/10.1111/j.2042-3306.1999.tb03856.x