1. Please check and comment entries here.
Table of Contents

    Topic review

    Entomopathogenic Microorganisms in Animals Protection

    Subjects: Microbiology
    View times: 16
    Submitted by: Francesca Mancianti

    Definition

    The control of ectoparasites requests the development of novel strategies and, among them, the use of entomopathogenic microorganisms appears as a promising tool to achieve an eco-friendly approach. 

    1. Entomopathogenic Fungi

    Several fungal species are well suited to control arthropods, being able to cause epizootic infection and most of them infect their host by direct penetration through the arthropod’s tegument [1]. Most of the organisms are related to biological control of crop pests, but, more recently, have been applied to combat some livestock ectoparasites. 

    Entomopathogenic fungi (EPFs) have been identified by their growth onto insect cadavers and can be commercially produced to act as biopesticides. Species of Beauveria, Metarhizium, Lecanicillium and Isaria are relatively easy to mass produce [2]. One of the main concerns about their extensive employ would be related to their sensitivity to temperature as well as ultraviolet radiation [3] and to the presence of a suitable moisture degree, to allow the conidia to germinate [4]. On the other hand, EPFs seem to have a negligible risk of inducing resistance [5], despite their long-term persistence in the environment.

    2. Entomopathogenic Bacteria

    Among the entomopathogenic bacteria (EPBs), Bacillus thuringiensis showed the most relevant activity against arthropods. B. thuringiensis is a Gram-positive, rod-shaped, spore-forming bacterium, innocuous for humans, animals and plants. It can be isolated from different environments, such as soil, rhizosphere, phylloplane, freshwater, and grain dusts; furthermore, it can be found in invertebrates and insectivorous mammals [6]. Its entomopathogenic property is related to the production of highly biodegradable proteins. Its action to the insect pest relies on insecticidal toxin and an array of virulence factors [7]. B. thuringiensis produces, upon sporulation, insecticidal crystal inclusion formed by several proteins named Cry or Cyt proteins. These proteins have been proven to be toxic to insects belonging to the orders Lepidoptera, Dipteran, Coleoptera, Hymenoptera, Homoptera, Orthoptera and Mallophage [7]. B. thuringiensis, because of its known entomopathogenic activity, has been used worldwide for biological control against several agriculture pests for a long time.

    Besides B. thuringiensis, Lysinibacillus (formely Bacillus) sphaericus is employed for preventing and controlling pests. Both agents are the only commercial entomopathogenic bacteria that are produced using mass production techniques and sold in sufficient commercial quantities.

    3. Ticks

    Ticks are large-bodied bloodsucking, nonpermanent parasitic Acari, feeding exclusively on vertebrates. They are divided into three families, among which Ixodidae (hard ticks) represent an important concern for mammalian health, although some of them also feed on birds, and can be carried between continents. Ticks exert a direct damage, feeding on their host. Saliva and/or mouthpart penetration can induce a toxic reaction in hosts, such as tick paralysis [8][9], or allergic state in human patients [10]. Heavy tick infestation can cause severe anemia, considering that an adult female tick can feed up to 2.0 mL of blood from the vertebrate host [11]

    3.1. Fungi

    The control of ticks by entomopathogenic fungi has been widely studied and, differently from insects, tick eggs are sensitive [12]. Tick species differ in their behavior, range of hosts and life cycle, so also, their sensitivity in comparison to a fungal species is not the same [13]. Furthermore, ticks are reported to be more tolerant to EPFs than other arthropods, so the amount of the inocula for tick control purposes should be larger. Different stages of ticks would exhibit differences in sensitivity versus EPFs. R. sanguineus engorged females and unfed other stages appeared more prone to fungal infection with M. anisopliae and M. flavoviride [14]. Nymphs were reported as less sensitive when compared with other stages [13][15]. A slight difference of sensitivity to M. brunneum, between adults and nymphs of I. scapularis was also reported [16][17] and larvae are considered the most susceptible stage to EPFs [18]. EPFs active against different tick species are summarized in Table 1.

    Table 1. Entomopathogenic fungi (EPFs) active versus different tick species.
    Tick Species EPFs References
    Amblyomma americanum Beauveria bassiana [19]
    Amblyomma parvum Metarhizium anisopliae [20]
    Amblyomma variegatum Beauveria bassiana [21]
    Amblyomma variegatum Metarhizium anisopliae [21]
    Amblyomma variegatum M. anisopliae + B. bassiana [22]
    Boophilus microplus Beauveria bassiana [23]
    Boophilus microplus Metarhizium anisopliae [24][25][23][26][27][28]
    Boophilus sp. Fusarium sp.
    Metarhizium anisopliae
    [29]
    Dermacentor albipictus Beauveria bassiana [30]
    Dermacentor albipictus Metarhizium anisopliae [30]
    Dermacentor albipictus Metarhizium brunneum [30]
    Dermacentor marginatus Aspergillus fumigatus [31]
    Dermacentor marginatus Trichothecium roseum [32]
    Dermacentor reticulatus Isaria fumosorosea [33]
    Dermacentor reticulatus Beauveria bassiana [33]
    Dermacentor reticulatus Metarhizium anisopliae [33]
    Dermacentor reticulatus Metarhizium robertsii [33]
    Dermacentor sp. Beauveria bassiana [34]
    Dermacentor variabilis Metarhizium anisopliae [16]
    Dermacentor variabilis Beauveria bassiana [16]
    Dermacentor variabilis Scopulariopsis brevicaulis [35]
    Haemaphysalis longicornis Beauveria bassiana [36]
    Haemaphysalis qinghaiensis Metarhizium anisopliae [37]
    Haemaphysalis qinghaiensis Beauveria bassiana [37]
    Hyalomma anatolicum Beauveria bassiana [38]
    Hyalomma anatolicum Metarhizium anisopliae [38]
    Hyalomma anatolicum Paecilomyces lilacinus [38]
    Hyalomma lusitanicum Beauveria bassiana [39][40]
    Hyalomma scupense Aspergillus fumigatus [31]
    Ixodes dammini Aspergillus ochraceus [29]
    Ixodes dammini Metarhizium anisopliae [41]
    Ixodes ricinus Conidiobolus coronatus [42]
    Ixodes ricinus Aspergillus flavus [43]
    Ixodes ricinus Aspergillus fumigatus [43]
    Ixodes ricinus Aspergillus niger [34]
    Ixodes ricinus Aspergillus parasiticus [34]
    Ixodes ricinus Beauveria bassiana [29][33]
    Ixodes ricinus Beauveria brognardi [42]
    Ixodes ricinus Paecilomyces farinosus [42]
    Ixodes ricinus Paecilomyces fumosoroseus [34][42]
    Ixodes ricinus Penicillium insectivorum [43]
    Ixodes ricinus Trichothecium roseum [32]
    Ixodes ricinus Verticillium aranearum [42]
    Ixodes ricinus Verticillium lecanii [34][42]
    Ixodes ricinus Metarhizium anisopliae [33]
    Ixodes ricinus Metarhizium robertsii [33]
    Ixodes ricinus Isaria fumosorosea [33]
    Ixodes scapularis Metarhizium brunneum [16][17][18][44]
    Ixodes scapularis Metarhizium anisopliae [16]
    Ixodes scapularis Beauveria bassiana [16]
    Rhipicephalus annulatus Metarhizium brunneum [45]
    Rhipicephalus appendiculatus Aspergillus sp. [46]
    Rhipicephalus appendiculatus Fusarium sp. [46]
    Rhipicephalus appendiculatus Metarhizium anisopliae [47][21][46]
    Rhipicephalus appendiculatus Beauveria bassiana [47]
    Rhipicephalus appendiculatus M. anisopliae + B. bassiana [21]
    Rhipicephalus decoloratus Beauveria bassiana [48]
    Rhipicephalus microplus Metarhizium robertsii [49][50][51][52]
    Rhipicephalus microplus Beauveria bassiana [53][51][54][55][56][57][58][59]
    Rhipicephalus microplus Metarhizium anisopliae [53][51][60][56][57][58][59]
    Rhipicephalus microplus Paecilomyces lilacinus [53]
    Rhipicephalus microplus Isaria fumosorosea [61]
    Rhipicephalus microplus Isaria farinosa [61]
    Rhipicephalus microplus Purpurocillium lilacinus [61]
    Rhipicephalus sanguineus Aspergillus ochraceus [62]
    Rhipicephalus sanguineus Fusarium sp. [63]
    Rhipicephalus sanguineus Curvularia lunata [64]
    Rhipicephalus sanguineus Rhizopus thailandensis [64]
    Rhipicephalus sanguineus Rhizopus arrhizus [64]
    Rhipicephalus sanguineus Metarhizium anisopliae [13][14][15][16]
    Rhipicephalus sanguineus Metarhizium flavoviride [14]
    Rhipicephalus sanguineus Isaria fumosorosea [14]
    Rhipicephalus sanguineus Beauveria bassiana [16]

    3.2. Bacteria

    Some bacterial species have been demonstrated to be pathogenic for ticks; thus, they are considered useful for biological control. Among EPBs, B. thuringiensis is the most studied agent with activity against ticks [65] and is largely employed in commercial insecticide formulations. The pathogenic action of B. thuringiensis normally occurs after the ingestion of spores by ticks, and the crystalline inclusions containing insecticidal δ-endotoxins specifically interact with receptors in the insect midgut epithelial cells [66].

    4. Dermanyssus Gallinae

    The genus Dermanyssus comprises hematophagous mite species, parasites of birds. The taxonomy of species within the genus was not clearly defined, until now [67]. Dermanyssus gallinae (poultry red mite) is very common in layer houses and is considered as the most damaging to laying hens worldwide [68].  Although birds are first choice hosts, D. gallinae feed on humans and other mammals, too [69], and can act as a vector for several pathogens of poultry [70], as well as zoonotic agents [71].

    4.1. Fungi

    Entomopathogenic fungi have been assayed to control the mite population. B. bassiana, M. anisopliae, Trichoderma album, and P. fumosoroseus are the most studied fungal species. The use of fungal entomopathogens to control arthropod pests as biological agents would be suggested considering their easy direct penetration through arthropod tegument, the lack of induction of host resistance, the ability to horizontally transmit from fungus-infected to uninfected arthropods, mostly in moist environments [72] and potential damage to flies, lice, and other pests [73][74]. B. bassiana, P. fumosoroseus and M. anisopliae were proven to kill several red mites, when administered in high doses, with a variability depending on the isolate [75][76][77][78], being able to cause high mortality within 5 days [76]. Different strains of M. anisopliae have been successfully applied to control the mites, under laboratory conditions, demonstrating differences in pathogenicity with a dose- and time-dependent effect [79].

    4.2. Bacteria

    The use of B. thuringiensis has been proposed as an alternative control method to chemical acaricides against D. gallinae in integrated management programs. It has been observed that B. thuringiensis var. kurstaki is able to damage the cuticle of D. galllinae and cause the loss of mobility of this mite in a period of 24 h [80]. Moreover, Torres and Hernandez [81] observed a moderate mortality of D. gallinae from day 2 of application (66%), which increased up to 78% at 7 days at a concentration of 35 mg/mL. Similarly, a previous study by Mullens et al. [82] on the fowl mite Ornithonyssus sylviarum, revealed that this mite was susceptible to B. thuringiensis, and the authors concluded that the entomopathogen had potential for the development of a control preparation for direct application to poultry.

    5. Psoroptes sp.

    Psoroptes mites are non-burrowing Acharina, responsible for ear and body mange of herbivores. Psoroptes ovis severely impacts on animal health. It induces an exudative dermatitis in beef cattle and sheep which, when not treated, can lead affected animals to lose condition and, sometimes, to death [83][84].

    5.1. Fungi

    In a comparative in vitro study with Hirsutella thompsonii, M. anisopliae was highly pathogenic and suitable for the control of P. ovis [85]. These features were furtherly corroborated by observing the efficiency of the mold in producing fatal infections, as well as the infectiveness of 5-day-old cadavers of mites [86]. The higher infectivity of M. anisopliae in comparison with B. bassiana was assessed in vivo, too [87]. The strong parasite killing of M. anisopliae seems to be related to its ability to induce the oxidative damage of mites [88].

    5.2. Bacteria

    The in vitro acaricidal effect of B. thuringiensis on P. cuniculi has been demonstrated. The bacterium can induce histological alterations of this mite, such as the presence of dilated intercellular spaces in the basal membrane, membrane detachment of the peritrophic matrix and morphological alterations in columnar cells of the intestine [89].

    Similarly, the combined use of B. thuringiensis and ivermectin has been proposed by some authors to combat Psoroptes sp., in view of a potential synergistic or additive effect with the possibility of lowering the dose of ivermectin [89].

    6. Varroa destructor

    Varroa destructor is a parasite Mesostigmata mite, exerting a huge impact on beekeeping. It has become a global parasite, switching host onto Apis mellifera from Apis cerana. Varroasis is often a threat for colonies, when nearby colonies collapse [90].

    V. destructor has been reported to be susceptible to the entomopathogenic fungi, M. anisopliae, B. bassiana, Verticillium lecanii, Hirsutella spp. [29][91], Hirsutella thompsonii [92][93], B. bassiana [94] and M. anisopliae [95][96]. Clonostachys rosea (formerly Gliocladium roseum) is an Ascomycete, belonging to Hypocreales, widely distributed in soil, and provided by an endophytic ability in tissues from several plants. The mold produces conidia and chlamydospores.

    7. Zoonotic Potential of EPFs

    Bacillus thuringiensis has been associated to different human infections; it has been cultured from marginal and apical periodontitis, wounds, corneal ulcera and gastrointestinal infections in humans [97]. Even though this EPB is not considered as a traditional zoonotic agent, its presence in different forms of human infections suggests that, at least in immunocompromised patients, it could represent a risk. B. thuringiensis, similarly to B. cereus, produces several virulence factors potentially acting against mammalian cells, such as hemolysins and enterotoxins [98].

    For these reasons, the use of B. thuringiensis in pest control should be carried out with attention to avoid possible infections, mainly in operators.

    The entry is from 10.3390/biology10060479

    References

    1. Hajek, A.E.; Delalibera, I. Fungal pathogens as classical biological control agents against arthropods. BioControl 2010, 55, 147–158.
    2. Vega, F.E.; Goettel, M.S.; Blackwell, M.; Chandler, D.; Jackson, M.A.; Keller, S.; Koikeg, M.; Maniania, N.K.; Monzo´, N.A.; Ownley, B.H.; et al. Fungal entomopathogens: New insights on their ecology. Fungal Ecol. 2009, 2, 149–159.
    3. Roberts, D.W.; Campbell, A.S. Stability of entomopathogenic fungi. Misc. Publ. Entomol. Soc. Am. 1977, 10, 19–76.
    4. Fargues, J.; Goettle, M.S.; Smits, N.; Ouedraogo, A.; Vidal, C.; Lacey, L.A.; Lomer, C.J.; Rougier, M. Variability in suceptibility to simulated sunlightof conidia among isolates of entomopathogenic hyphomycetes. Mycopathologia 1996, 133, 171–181.
    5. Gao, T.; Wang, Z.; Huang, Y.; Keyhani, N.O.; Huang, Z. Lack of resistance development in Bemisia tabaci to Isaria fumosorosea after multiple generations of selection. Sci. Rep. 2017, 7, srep42727.
    6. Raymond, B.; Johnston, P.R.; Nielsen-LeRoux, C.; Lereclus, D.; Crickmore, N. Bacillus thuringiensis: An impotent pathogen? Trends Microbiol. 2010, 18, 189–194.
    7. Bravo, A.S.M. Bacillus thuringiensis: Mechanisms and use. Compr. Mol. Insect Sci. 2005, 6, 175–205.
    8. Padula, A.M.; Leister, E.M.; Webster, R.A. Tick paralysis in dogs and cats in Australia: Treatment and prevention deliverables from 100 years of research. Aust. Vet. J. 2019, 98, 53–59.
    9. Simon, L.V.; West, B.; McKinney, W.P. Tick Paralysis. In StatPearls [Internet]; StatPearls Publishing: Treasure Island, FL, USA, 2020.
    10. Nunen, A.S. Tick-induced allergies: Mammalian meat allergy and tick anaphylaxis. Med. J. Aust. 2018, 208, 316–321.
    11. Basu, A.K.; Charles, R. Ticks of Trinidad and Tobago—An Overview, 1st ed.; Academic Press: London, UK, 2017.
    12. Gindin, G.; Samish, M.; Zangi, G.; Mishoutchenko, A.; Glazer, I. The Susceptibility of Different Species and Stages of Ticks to Entomopathogenic Fungi. Exp. Appl. Acarol. 2002, 28, 283–288.
    13. Fernandes, É.K.; Bittencourt, V.R.; Roberts, D.W. Perspectives on the potential of entomopathogenic fungi in biological control of ticks. Exp. Parasitol. 2012, 130, 300–305.
    14. Samish, M.; Gindin, G.; Alekseev, E.; Glazer, I. Pathogenicity of entomopathogenic fungi to different develop-mental stages of Rhipicephalus sanguineus (Acari: Ixodidae). J. Parasitol. 2001, 87, 1355–1359.
    15. Cafarchia, C.; Immediato, D.; Iatta, R.; Ramos, R.A.N.; Lia, R.P.; Porretta, D.; Figueredo, L.A.; Dantas-Torres, F.; Otranto, D. Native strains of Beauveria bassiana for the control of Rhipicephalus sanguineus sensu lato. Parasites Vectors 2015, 8, 80.
    16. Kirkland, B.H.; Westwood, G.S.; Keyhani, N.O. Pathogenicity of entomopathogenic fungi Beauveria bassiana and Metarhizium anisopliae to Ixodidae species Dermacentor variabilis, Rhipicephalus sanguineus, and Ixodes scapularis. J. Med. Entomol. 2004, 41, 705–711.
    17. Bharadwaj, A.; Stafford, K.C., 3rd. Susceptibility of Ixodes scapularis (Acari: Ixodidae) to Metarhizium brunneum F52 (Hypocreales: Clavicipitaceae) using three exposure assays in the laboratory. J. Econ. Entomol. 2012, 105, 222–231.
    18. Fernandes, É.K.K.; Bittencourt, V.R.E.P. Entomopathogenic fungi against South American tick species. Exp. Appl. Acarol. 2008, 46, 71–93.
    19. Cradock, K.R.; Needham, G.R. Beauveria bassiana (Ascomycota: Hypocreales) as a management agent for free-living Amblyomma americanum (Acari: Ixodidae) in Ohio. Exp. Appl. Acarol. 2010, 53, 57–62.
    20. Garcia, M.V.; Rodrigues, V.D.S.; Monteiro, A.C.; Simi, L.D.; Higa, L.D.O.S.; Martins, M.M.; Prette, N.; Mochi, D.A.; Andreotti, R.; Szabó, M.P.J. In vitro efficacy of Metarhizium anisopliae sensu lato against unfed Amblyomma parvum (Acari: Ixodidae). Exp. Appl. Acarol. 2018, 76, 507–512.
    21. Kaaya, G.P.; Mwangi, E.N.; Ouna, E.A. Prospects for Biological Control of Livestock Ticks, Rhipicephalus appendiculatus and Amblyomma variegatum, Using the Entomogenous Fungi Beauveria bassiana and Metarhizium Anisopliae. J. Invertebr. Pathol. 1996, 67, 15–20.
    22. Maranga, R.O.; Kaaya, G.P.; Mueke, J.M.; Hassanali, A. Effects of combining the fungi Beauveria bassiana and Metarhiziumanisopliae on the mortality of the tick Amblyomma variegatum (ixodidae) in relation to seasonal changes. Mycopathologia 2005, 159, 527–532.
    23. Verissimo, C.J. Natural enemies of the cattle tick. Agropecu. Catarin. 1995, 8, 35–37.
    24. Leemon, D.; Turner, L.; Jonsson, N. Pen studies on the control of cattle tick (Rhipicephalus (Boophilus) microplus) with Metarhizium anisopliae (Sorokin). Vet. Parasitol. 2008, 156, 248–260.
    25. Frazzon, A.P.G.; Junior, I.D.S.V.; Masuda, A.; Schrank, A.; Vainstein, M.H. In vitro assessment of Metarhizium anisopliae isolates to control the cattle tick Boophilus microplus. Vet. Parasitol. 2000, 94, 117–125.
    26. Bittencourt, V.R.E.P.; Massard, C.L.; De Lima, A.F. Action of the fungus Metarhizium anisopliae on eggs and larvae of the tick Boophilus microplus. Rev. Univ. Rural Ser. Cienc. Vida 1994, 16, 41–47.
    27. Bittencourt, V.R.E.P.; Massard, C.L.; De Lima, A.F. Action of the fungus Metarhizium anisopliae on the freeliving phase of the life cycle of Boophilus microplus. Rev. Univ. Rural Ser. Cienc. Vida 1994, 16, 49–55. (In Portuguese)
    28. Correia, A.C.B.; Fiorin, A.C.; Monteiro, A.C.; Verissimo, C.J. Effects of Metarhizium anisopliae on the tick Boophilus microplus (Acari: Ixodidae) in stabled cattle. J. Invert. Pathol. 1998, 71, 189–191.
    29. Chandler, D.; Davidson, G.; Pell, J.K.; Ball, B.V.; Shaw, K.; Sunderland, K.D. Fungal Biocontrol of Acari. Biocontrol Sci. Technol. 2000, 10, 357–384.
    30. Sullivan, C.F.; Parker, B.L.; Davari, A.; Lee, M.R.; Kim, J.S.; Skinner, M. Evaluation of spray applications of Metarhizium anisopliae, Metarhizium brunneum and Beauveria bassiana against larval winter ticks, Dermacentor albipictus. Exp. Appl. Acarol. 2020, 82, 559–570.
    31. Lipa, J.J. Microbial control of mites and ticks. In Microbial Control of Insects and Mites; Burges, H.D., Hussey, N.W., Eds.; Academic Press: London, UK, 1971; pp. 357–373.
    32. Balazy, S.; Wisniewski, J.; Kaczmarek, S. Some noteworthy fungi occurring on mites. Bull. Polish Acad. Sci. Biol. Sci. 1987, 35, 199–224.
    33. Szczepańska, A.; Kiewra, D.; Plewa-Tutaj, K.; Dyczko, D.; Guz-Regner, K. Sensitivity of Ixodes ricinus (L., 1758) and Dermacentor reticulatus (Fabr., 1794) ticks to entomopathogenic fungi isolates: Preliminary study. Parasitol. Res. 2020, 119, 3857–3861.
    34. Samsináková, A.; Kálalová, S.; Daniel, M.; Dusbábek, F.; Honzáková, E.; Cerný, V. Entomogenous fungi associated with the tick Ixodes ricinus (L.). Folia Parasitol. 1974, 21, 39–48.
    35. Yoder, J.A.; Rodell, B.M.; Klever, L.A.; Dobrotka, C.J.; Pekins, P.J. Vertical transmission of the entomopathogenic soil fungus Scopulariopsis brevicaulis as a contaminant of eggs in the winter tick, Dermacentor albipictus, collected from calf moose (New Hampshire, USA). Mycologia 2019, 10, 174–181.
    36. Zhendong, H.; Guangfu, Y.; Zhong, Z.; Ruiling, Z. Phylogenetic relationships and effectiveness of four Beauveria bassiana sensu lato strains for control of Haemaphysalis longicornis (Acari: Ixodidae). Exp. Appl. Acarol. 2018, 77, 83–92.
    37. Ren, Q.; Chen, Z.; Luo, J.; Liu, G.; Guan, G.; Liu, Z.; Liu, A.; Li, Y.; Niu, Q.; Liu, J.; et al. Laboratory evaluation of Beauveria bassiana and Metarhizium anisopliae in the control of Haemaphysalis qinghaiensis in China. Exp. Appl. Acarol. 2016, 69, 233–238.
    38. Sun, M.; Ren, Q.; Guan, G.; Liu, Z.; Ma, M.; Gou, H.; Chen, Z.; Li, Y.; Liu, A.; Niu, Q.; et al. Virulence of Beauveria bassiana, Metarhizium anisopliae and Paecilomyces lilacinus to the engorged female Hyalomma anatolicum anatolicum tick (Acari: Ixodidae). Vet. Parasitol. 2011, 180, 389–393.
    39. Olmeda, A.S.; Pe´rez Sanchez, J.L.; Valcarcel, F.; Espada-Espada, N.; Garcıa- Rojo Lopez, B.; Cota-Guajardo, S.; Cutuli, M.T. Isolation of entomopathogenic fungi from Hyalomma lusitanicum tick, in Spain. In Proceedings of the Seventh Ticks and Tick-Borne Pathogens International Conference, Zaragoza, Spain, 28 August–2 September 2011.
    40. González, J.; Valcárcel, F.; Pérez-Sánchez, J.L.; Tercero-Jaime, J.M.; Cutuli, M.T.; Olmeda, A.S. Control of Hyalomma lusitanicum (Acari: Ixodidade) Ticks Infesting Oryctolagus cuniculus (Lagomorpha: Leporidae) Using the Entomopathogenic Fungus Beauveria bassiana (Hyocreales: Clavicipitaceae) in Field Conditions. J. Med Entomol. 2016, 53, 1396–1402.
    41. Zhioua, E.; Browning, M.; Johnson, P.W.; Ginsberg, H.S.; Lebrun, R.A. Pathogenicity of the entomopathogenic fungus Metarhizium anisopliae (Deuteromycetes) to Ixodes scapularis (Acari: Ixodidae). J. Parasitol. 1997, 83, 815.
    42. Kalsbeek, V.; Frandsen, F.; Steenberg, T. Entomopathogenic fungi associated with Ixodes ricinus ticks. Exp. Appl. Acarol. 1995, 19, 45–51.
    43. Cherepanova, N.P. Fungi which are met on ticks. Botanicnyi Zhurnal Kiev 1964, 49, 696–699.
    44. Behle, R.W.; Jackson, M.A.; Flor-Weiler, L.B. Efficacy of a Granular Formulation Containing Metarhizium brunneum F52 (Hypocreales: Clavicipitaceae) Microsclerotia Against Nymphs of Ixodes scapularis (Acari: Ixoididae). J. Econ. Entomol. 2013, 106, 57–63.
    45. Samish, M.; Rot, A.; Ment, D.; Barel, S.; Glazer, I.; Gindin, G. Efficacy of the entomopathogenic fungus Metarhizium brunneum in controlling the tick Rhipicephalus annulatus under field conditions. Vet. Parasitol. 2014, 206, 258–266.
    46. Mwangi, E.N.; Kaaya, G.P.; Essuman, S. Experimental infections of the tick Rhipicephalus appendiculatus with entomopathogenic fungi, Beauveria bassiana and Metarhizium anisopliae, and natural infections of some ticks with bacteria and fungi. J. Afr. Zool. 1995, 109, 151–160.
    47. Kaaya, G.P.; Hassan, S. Entomogenous Fungi as Promising Biopesticides for Tick Control. Exp. Appl. Acarol. 2000, 24, 913–926.
    48. Murigu, M.M.; Nana, P.; Waruiru, R.M.; Nga’Nga’, C.J.; Ekesi, S.; Maniania, N.K. Laboratory and field evaluation of entomopathogenic fungi for the control of amitraz-resistant and susceptible strains of Rhipicephalus decoloratus. Vet. Parasitol. 2016, 225, 12–18.
    49. De Paulo, J.F.; Camargo, M.G.; Coutinho-Rodrigues, C.J.B.; Marciano, A.F.; De Freitas, M.C.; Da Silva, E.M.; Gôlo, P.S.; Morena, D.D.S.; Angelo, I.D.C.; Bittencourt, V.R.E.P. Rhipicephalus microplus infected by Metarhizium: Unveiling hemocyte quantification, GFP-fungi virulence, and ovary infection. Parasitol. Res. 2018, 117, 1847–1856.
    50. Fiorotti, J.; Menna-Barreto, R.F.S.; Gôlo, P.S.; Coutinho-Rodrigues, C.J.B.; Bitencourt, R.O.B.; Spadacci-Morena, D.D.; Angelo, I.D.C.; Bittencourt, V.R.E.P. Ultrastructural and Cytotoxic Effects of Metarhizium robertsii Infection on Rhipicephalus microplus Hemocytes. Front. Physiol. 2019, 10, 654.
    51. Bernardo, C.C.; Barreto, L.P.; Silva, C.D.S.E.; Luz, C.; Arruda, W.; Fernandes, É.K. Conidia and blastospores of Metarhizium spp. and Beauveria bassiana s.l.: Their development during the infection process and virulence against the tick Rhipicephalus microplus. Ticks Tick-Borne Dis. 2018, 9, 1334–1342.
    52. Marciano, A.F.; Mascarin, G.M.; Franco, R.F.F.; Golo, P.S.; Jaronski, S.T.; Fernandes, É.K.; Bittencourt, V.R.E.P. Innovative granular formulation of Metarhizium robertsii microsclerotia and blastospores for cattle tick control. Sci. Rep. 2021, 11, 4972.
    53. Salas, A.F.; Alonso-Díaz, M.A.; Alonso-Morales, R.A.; Lezama-Gutiérrez, R.; Rodríguez-Rodríguez, J.C.; Cervantes-Chávez, J.A. Acaricidal activity of Metarhizium anisopliae isolated from paddocks in the Mexican tropics against two populations of the cattle tick Rhipicephalus microplus. Med. Vet. Entomol. 2016, 31, 36–43.
    54. Campos, R.; Boldo, J.; Pimentel, I.; Dalfovo, V.; Arajo, W.; Azevedo, J.; Vainstein, M.; Barros, N. Endophytic and entomopathogenic strains of Beauveria sp to control the bovine tick Rhipicephalus (Boophilus) microplus. Genet. Mol. Res. 2010, 9, 1421–1430.
    55. Rivera, A.P.T.; Cuadros, M.O.; Claros, B.P.; Ayola, S.C.P.; Romero, D.C.M. Efectividad de Beauveria bassiana (Baubassil®) sobre la garrapata común del ganado bovino Rhipicephalus microplus en el Departamento de la Guajira, Colombia. Rev. Argent. Microbiol. 2018, 50, 426–430.
    56. Bittencourt, V.R.E.P.; Peralva, S.L.F.S.; Viegas, E.C.; Alves, S.B. Avaliação do sefeitos do contato de Beauveria bassiana (Bals.) Vuill. como vose larvas de Boophilus microplus (Canestrini, 1887) (Acari:Ixodidae). Rev. Brasiliana Parasitol. Vetinaria 1996, 5, 81–84.
    57. Perinotto, W.; Angelo, I.; Golo, P.; Quinelato, S.; Camargo, M.; Sá, F.; Bittencourt, V. Susceptibility of different populations of ticks to entomopathogenic fungi. Exp. Parasitol. 2012, 130, 257–260.
    58. Webster, A.; Pradel, E.; Souza, U.A.; Martins, J.R.; Reck, J.; Schrank, A.; Klafke, G. Does the effect of a Metarhizium anisopliae isolate on Rhipicephalus microplus depend on the tick population evaluated? Ticks Tick-Borne Dis. 2017, 8, 270–274.
    59. Fernández-Salas, A.; Alonso-Díaz, M.A.; Alonso-Morales, R.A. Effect of entomopathogenic native fungi from paddock soils against Rhipicephalus microplus larvae with different toxicological behaviors to acaricides. Exp. Parasitol. 2019, 204, 107729.
    60. Nogueira, M.R.D.S.; Camargo, M.G.; Rodrigues, C.J.B.C.; Marciano, A.F.; Quinelato, S.; De Freitas, M.C.; Fiorotti, J.; De Sá, F.A.; Perinotto, W.M.D.S.; Bittencourt, V.R.E.P. In vitro efficacy of two commercial products of Metarhizium anisopliae s.l. for controlling the cattle tick Rhipicephalus microplus. Rev. Bras. Parasitol. Veterinária 2020, 29, e000220.
    61. Angelo, I.C.; Fernandes, É.K.; Bahiense, T.C.; Perinotto, W.M.S.; Golo, P.S.; Moraes, A.P.R.; Bittencourt, V.R.E.P. Virulence of Isaria sp. and Purpureocillium lilacinum to Rhipicephalus microplus tick under laboratory conditions. Parasitol. Res. 2012, 111, 1473–1480.
    62. Estrada-Peña, A.; González, J.; Casasolas, A. The activity of Aspergillus ochraceus (fungi) on replete females of Rhipicephalus sanguineus (Acari: Ixodidae) in natural and experimental conditions. Folia Parasitol. 1990, 37, 331–336.
    63. Lombardini, G. Biological and anatomical observations on Rhipicephalus sanguineus. Latr. Redia 1950, 35, 173–183.
    64. Casasolas-Oliver, A.; Estrada-Pena, A.; Gonzalez-Cabo, J. Activity of Rhizopus thailandensis, Rhizopus arrhizus and Curvularia lunata on reproductive efficacy of Rhipicephalus sanguineus (Ixodidae). In Modern Acaralogy; Dusbadek, E., Bukva, V., Eds.; Academia Prague and SPB Academic Publishing BV: Prague, Czech Republic, 1991; pp. 633–637.
    65. Fernández-Ruvalcaba, M.; Peña-Chora, G.; Romo-Martínez, A.; Hernández-Velázquez, V.; De Parra, A.B.; De La Rosa, D.P. Evaluation of Bacillus thuringiensis Pathogenicity for a Strain of the Tick, Rhipicephalus microplus, Resistant to Chemical Pesticides. J. Insect Sci. 2010, 10, 1–6.
    66. Bravo, A.; Gill, S.S.; Soberón, M. Mode of action of Bacillus thuringiensis Cry and Cyt toxins and their potential for insect control. Toxicon 2007, 49, 423–435.
    67. Roy, L.; Chauve, C. Historical review of the genus Dermanyssus Dugès, 1834 (Acari: Mesostigmata: Dermanyssidae). Parasite 2007, 14, 87–100.
    68. Sparagano, O. A nonexhaustive overview on potential impacts of the poultry red mite (Dermanyssus gallinae) on poultry production systems. J. Anim. Sci. 2020, 98, S58–S62.
    69. George, D.R.; Finn, R.D.; Graham, K.M.; Mul, M.F.; Maurer, V.; Moro, C.V.; Sparagano, O.A. Should the poultry red mite Dermanyssus gallinae be of wider concern for veterinary and medical science? Parasites Vectors 2015, 8, 178.
    70. Oh, S.-I.; Do, Y.J.; Kim, E.; Yi, S.W.; Yoo, J.G. Prevalence of poultry red mite (Dermanyssus gallinae) in Korean layer farms and the presence of avian pathogens in the mite. Exp. Appl. Acarol. 2020, 81, 223–238.
    71. Sparagano, O.A.E.; Ho, J. Parasitic Mite Fauna in Asian Poultry Farming Systems. Front. Vet. Sci. 2020, 7, 400.
    72. Kaaya, G.P.; Okech, M.A. Horizontal transmission of mycotic infection in adult tsetse, Glossina morsitans morsitans. Entomophaga 1990, 35, 46–57.
    73. Kaufman, P.E.; Reasor, C.; Donald, A.; Rutz, D.A.; Ketzis, J.K.; Arends, J.J. Evaluation of Beauveria bassiana applications against adult house fly, Musca domestica, in commercial caged-layer poultry facilities in New York state. BioControl 2005, 33, 360–367.
    74. Gindin, G.; Glazer, I.; Mishoutchenko, A.; Samish, M. Entomopathogenic fungi as a potential control agent against the lesser mealworm, Alphitobius diaperinus in broiler houses. BioControl 2009, 54, 549–558.
    75. Immediato, D.; Camarda, A.; Iatta, R.; Puttilli, M.R.; Ramos, R.A.N.; Di Paola, G.; Giangaspero, A.; Otranto, D.; Cafarchia, C. Laboratory evaluation of a native strain of Beauveria bassiana for controlling Dermanyssus gallinae (De Geer, 1778) (Acari: Dermanyssidae). Vet. Parasitol. 2015, 212, 478–482.
    76. Steenberg, T.; Kilpinen, O. Fungus infection of the chicken mite Dermanyssus gallinae. IOBC WPRS Bull. 2003, 26, 23–26.
    77. Kasburg, C.R.; Alves, L.F.A.; Oliveira, D.G.P.; Rohde, C. Activity of some Brazilian isolates of entomopathogenic fungi against the poultry red mite Dermanyssus gallinae De Geer (Acari: Dermanyssidae). Braz. J. Poult. Sci. 2016, 18, 457–460.
    78. De Oliveira, D.G.P.; Kasburg, C.R.; Alves, L.F.A. Efficacy of Beauveria bassiana against the poultry red mite, Dermanyssus gallinae (De Geer, 1778) (Mesostigmata: Dermanyssidae), under laboratory and hen house conditions. Syst. Appl. Acarol. 2020, 25, 895–905.
    79. Tavassoli, M.; Ownag, A.; Pourseyed, S.H.; Mardani, K. Laboratory evaluation of three strains of the entomopathogenic fungus Metarhizium anisopliae for controlling Dermanyssus gallinae. Avian Pathol. 2008, 37, 259–263.
    80. Roberts, D.W.; Leger, R.J.S. Metarhizium spp., Cosmopolitan Insect-Pathogenic Fungi: Mycological Aspects. Adv. Appl. Microbiol. 2004, 54, 1–70.
    81. Torres, E.C.; Hernández, J.F. Actividad acaricida de Bacillus thuringiensis sobre el acaro rojo de las aves, Dermanyssus gallinae. Revista Veterinaria 2018, 29, 128–132.
    82. Mullens, B.A.; Wills, L.E.; Rodriguez, J.L. Evaluation of ABG-6208 (Thuringiensin) for control of northern fowl mite, 1987. Insect Acar. Tests 1988, 13, 408–409.
    83. Bates, P. Inter- and intra-specific variation within the genus Psoroptes (Acari: Psoroptidae). Vet. Parasitol. 1999, 83, 201–217.
    84. Broek, A.V.D.; Huntley, J. Sheep Scab: The Disease, Pathogenesis and Control. J. Comp. Pathol. 2003, 128, 79–91.
    85. Smith, K.; Wall, R.; French, N. The use of entomopathogenic fungi for the control of parasitic mites, Psoroptes spp. Vet. Parasitol. 2000, 92, 97–105.
    86. Brooks, A.; Wall, R. Infection of Psoroptes mites with the fungus Metarhizium anisopliae. Exp. Appl. Acarol. 2001, 25, 869–880.
    87. Abolins, S.; Thind, B.; Jackson, V.; Luke, B.; Moore, D.; Wall, R.; Taylor, M. Control of the sheep scab mite Psoroptes ovis in vivo and in vitro using fungal pathogens. Vet. Parasitol. 2007, 148, 310–317.
    88. Gu, X.; Zhang, N.; Xie, Y.; Zheng, Y.; Chen, Y.; Zhou, X.; Li, X.; Zhong, Z.; He, R.; Yang, G. Metarhizium anisopliae CQMa128 regulates antioxidant/detoxification enzymes and exerts acaricidal activity against Psoroptes ovis var. cuniculi in rabbits: A preliminary study. Vet. Parasitol. 2020, 279, 109059.
    89. Dunstand-Guzmán, E.; Peña-Chora, G.; Hallal-Calleros, C.; Pérez-Martínez, M.; Hernández-Velazquez, V.M.; Morales-Montor, J.; Flores-Pérez, F.I. Acaricidal effect and histological damage induced by Bacillus thuringiensis protein extracts on the mite Psoroptes cuniculi. Parasites Vectors 2015, 8, 285.
    90. Nolan, K.S. Delaplane Distance between honeybee Apis mellifera colonies regulates populations of Varroa destructor at a landscape scale. Apidologie 2017, 48, 8–16.
    91. Shaw, K.E.; Davidson, G.; Clark, S.J.; Ball, B.V.; Pell, J.K.; Chandler, D.; Sunderland, K.D. Laboratory bioassays to assess the pathogenicity of mitosporic fungi to Varroa destructor (Acari: Mesostigmata), an ectoparasitic mite of the honeybee, Apis mellifera. Biol. Control. 2002, 24, 266–276.
    92. Peng, C.Y.S.; Zhou, X.; Kaya, H.K. Virulence and site of infection of the fungus, Hirsutella thompsonii, to the honey bee ectoparasitic mite, Varroa destructor. J. Invertebr. Pathol. 2002, 81, 185–195.
    93. Kanga, L.; James, R.; Boucias, D. Hirsutella thompsonii and Metarhizium anisopliae as potential microbial control agents of Varroa destructor, a honey bee parasite. J. Invertebr. Pathol. 2002, 81, 175–184.
    94. Meikle, W.G.; Mercadier, G.; Holst, N.; Girod, V. Impact of two treatments of a formulation of Beauveria bassiana (Deuteromycota: Hyphomycetes) conidia on Varroa mites (Acari: Varroidae) and on honeybee (Hymenoptera: Apidae) colony health. Exp. Appl. Acarol. 2008, 46, 105–117.
    95. Kanga, L.H.B.; Jones, W.A.; James, R.R. Field Trials Using the Fungal Pathogen, Metarhizium anisopliae (Deuteromycetes: Hyphomycetes) to Control the Ectoparasitic Mite, Varroa destructor (Acari: Varroidae) in Honey Bee, Apis mellifera (Hymenoptera: Apidae) Colonies. J. Econ. Entomol. 2003, 96, 1091–1099.
    96. Kanga, L.H.B.; Adamczyk, J.; Patt, J.; Gracia, C.; Cascino, J. Development of a user-friendly delivery method for the fungus Metarhizium anisopliae to control the ectoparasitic mite Varroa destructor in honey bee, Apis mellifera, colonies. Exp. Appl. Acarol. 2010, 52, 327–342.
    97. Helgason, E.; Caugant, D.A.; Olsen, I.; Kolstø, A.-B. Genetic Structure of Population of Bacillus cereus and B. thuringiensis Isolates Associated with Periodontitis and Other Human Infections. J. Clin. Microbiol. 2000, 38, 1615–1622.
    98. Kotiranta, A.; Lounatmaa, K.; Haapasalo, M. Epidemiology and pathogenesis of Bacillus cereus infections. Microbes Infect. 2000, 2, 189–198.
    More