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The effects of the fungus Metarhizium anisopliae var. acridum on different stages of Lutzomyia longipalpis (Diptera: Psychodidae)
Acta Tropica 113 (2010) 214–220
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The effects of the fungus Metarhizium anisopliae var. acridum on different stages of Lutzomyia longipalpis (Diptera: Psychodidae) Sthenia Santos Albano Amóra a,∗ , Claudia Maria Leal Bevilaqua a,∗ , Francisco Marlon Carneiro Feijó b , Romeika Hermínia de Macedo Assunc¸ão Pereira b , Nilza Dutra Alves b , Fúlvio Aurélio de Morais Freire b , Michel Toth Kamimura c , Diana Magalhães de Oliveira c , Elza Áurea Luna-Alves Lima d , Marcos Fábio Gadelha Rocha a a
Programa de Pós-Graduac¸ão em Ciências Veterinárias, Universidade Estadual do Ceará, Fortaleza, Ceará 60740-000, Brazil Laboratório de Microbiologia Veterinária, Universidade Federal Rural do Semi-Árido, Mossoró, Rio Grande do Norte 59625-900, Brazil c Núcleo de Genômica e Bioinformática Tarsísio Pimenta, Universidade Estadual do Ceará, Fortaleza, Ceará 60740-000, Brazil d Departamento de Micologia, Centro de Ciências Biológicas, Universidade Federal de Pernambuco, Recife, Pernambuco 50741-040, Brazil b
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Article history: Received 19 May 2009 Received in revised form 22 October 2009 Accepted 24 October 2009 Available online 31 October 2009 Keywords: Biological control Vector Lutzomyia longipalpis Entomopathogenic fungus Metarhizium anisopliae var. acridum Visceral Leishmaniasis
a b s t r a c t The control of Visceral Leishmaniasis (VL) vector is often based on the application of chemical residual insecticide. However, this strategy has not been effective. The continuing search for an appropriate vector control may include the use of biological control. This study evaluates the effects of the fungus Metarhizium anisopliae var. acridum on Lutzomyia longipalpis. Five concentrations of the fungus were utilized, 1 × 104 to 1 × 108 conidia/ml, accompanied by controls. The unhatched eggs, larvae and dead adults previously exposed to fungi were sown to reisolate the fungi and analysis of parameters of growth. The fungus was subsequently identified by PCR and DNA sequencing. M. anisopliae var. acridum reduced egg hatching by 40%. The mortality of infected larvae was significant. The longevity of infected adults was lower than that of negative controls. The effects of fungal infection on the hatching of eggs laid by infected females were also significant. With respect to fungal growth parameters post-infection, only vegetative growth was not significantly higher than that of the fungi before infection. The revalidation of the identification of the reisolated fungus was confirmed post-passage only from adult insects. In terms of larvae mortality and the fecundity of infected females, the results were significant, proving that the main vector species of VL is susceptible to infection by this entomopathogenic fungus in the adult stage. © 2009 Elsevier B.V. All rights reserved.
1. Introduction Leishmaniasis is a zoonosis transmitted by vector rather complex because it involves several host species and sand fly vectors both the ecology and epidemiology of the disease are multifarious (Singh, 2006). The sand fly Lutzomyia longipalpis Lutz and Neiva (1912) is the primary vector for the etiological agent of Visceral Leishmaniasis (VL), Leishmania chagasi Cunha and Chagas (1937), which constitutes a serious public health problem in South America (Secundino et al., 2002). The VL control strategy is comprised of a combination of early detection, drug treatment, euthanasia of positive dogs, environmental management and chemical-based vector control. However, these measures have not been effective (Amóra et al., 2006). In the
∗ Corresponding authors. Tel.: +55 85 3101 9853; fax: +55 85 3101 9840. E-mail addresses: [email protected] (S.S.A. Amóra), [email protected] (C.M.L. Bevilaqua). 0001-706X/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.actatropica.2009.10.018
absence of an effective vaccine or an ideal drug treatment, the best method to interrupt any vector-borne disease is to reduce humanvector contact (Sharma and Singh, 2008). A search for appropriate vector control methods to augment the current limited arsenal of tools is required, and biological control is one promising avenue for this search (Scholte et al., 2007). Over the last decade, there have been major developments in biological, pesticide-based and integrated control strategies for mosquito vectors (Scholte et al., 2004). However, little information is available on the biological control of sand flies. In laboratory studies, the infection of sand flies with different microorganisms such as nematodes (Secundino et al., 2002), bacilli (Robert et al., 1998; Wahba et al., 1999; Wahba, 2000) and fungi (Warburg, 1991; Reithinger et al., 1997; Amóra et al., 2009) inhibited egg hatching and caused larval and adult death, but these results were variable. Meanwhile, there is little information on the effects parasites may produce on sand flies. In particular, entomopathogenic fungi have been highlighted as the main agent in insect control (Scholte et al., 2004). These
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pathogens have been shown to be an alternative to biological control method against many insects (Alves, 1998; Faria and Magalhães, 2001; Scholte et al., 2004), and they are consequently an additional tool for VL control, considering that one of the most effective control of VL is the vector control (Dye, 1996), reducing the use of chemical insecticides, resulting in benefits to humans and the environment (Amóra et al., 2009). In this context, Metarhizium anisopliae (Metsch.) Sorokin (Ascomycota: Hypocreales) is able to infect over 200 insect species (Alves, 1998), is considered to be a highly infective pathogenic fungus against grasshoppers (Goettel et al., 1995) and has been studied for the biological control of several other species of insects (Kanga et al., 2002; Bahiense et al., 2006; Scholte et al., 2007), including Diptera (Feijó et al., 2009). Specifically, M. anisopliae var. acridum [formerly Metarhizium flavoviride Gams and Rozsypal, but now reclassified (Driver et al., 2000)] is known for its effectiveness in the control of grasshoppers in Brazil and elsewhere (Faria and Magalhães, 2001). Currently, this fungus is commercialized under the trade name “Green Muscle® – Lubilosa” and “Green Guard® – Becker Underwood Pty Ltd.” for grasshopper (Acrididae) control in South Africa and Australia, respectively (Entz et al., 2005). In Brazil, field tests have been conducted (Magalhães et al., 2001). Despite its importance to agriculture, its pathogenicity in Diptera that are related to public health issues have not yet been sufficiently explored. However the efficacy results of M. anisopliae var. acridum on Diptera Chrysomya albiceps (Feijó et al., 2009) encourages new researches with this fungus on other Diptera. Also it is now known that this pathogen can be used to efficiently control Rhammatocerus schistocercoides Rehn (1906) (Orthoptera: Acrididae) in Brazil (Magalhães et al., 2001) Thus, the biological potential of M. anisopliae var. acridum makes this fungus a candidate for control of other Diptera, including sand flies. To this end, the present study aimed to evaluate the effect of M. anisopliae var. acridum on various stages of L. longipalpis development. 2. Materials and methods 2.1. L. longipalpis collection and identification L. longipalpis captured in the field were maintained in BOD incubators acclimatized to 27 ◦ C, 80% RH with a photoperiod of 12 h (Rangel et al., 1985). To promote oviposition, female sand flies were allowed to feed and obtain a blood meal from anesthetized hamsters for 2 h. Forty-eight hours post-feeding, the adult females were individualized in plastic pots measuring 4 cm in diameter and 4.5 cm in height that were internally coated with sterile plaster to maintain moisture. After oviposition, the females were dissected (Aransay et al., 2000) for identification (Galati, 2003). The hatched larvae were fed daily with a diet based on rabbit feces and crushed dried cassava leaves until they reached the pupa stage. After emergence, adults were transferred to nylon tulle cages measuring 20 cm3 in diameter, fed for 3 days with a glucose solution soaked in sterile cotton, and on the 4th day, an anesthetized hamster was offered to provide a blood meal for the females.
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2.3. L. longipalpis susceptibility to M. anisopliae var. acridum The bioassays, in which eggs, larvae and adults were treated with 5 fungal concentrations, were conducted with 2 control groups: 0.05% (v/v) sterile Tween 80 (negative control) (Feijó et al., 2008) and 196 g/ml of the pyrethroid, cypermethrin (positive control). The insecticide chosen as positive control is the compound used by the Brazilian Ministry of Health in the sand flies control (Ministério da and Saúde, 2006). Randomized treatments (21) were performed; 7 for each insect stage, with each treatment performed in 3 repetitions and each repetition in triplicate. Each repetition consisted of 30 samples totaling 630 individuals/ repetition. 2.3.1. Egg susceptibility The eggs were placed in plastic pots similar to those used in the maintenance of the colony. One of the fungal suspensions (3 ml), cypermethrin or Tween 80 was spilled on the walls and ground of the pots using a pipette. The containers were then stored in BOD incubators at 27 ◦ C, 80% RH with a photoperiod of 12 h. The egg hatching was observed daily and larval mortality was counted 8 days post-treatment, with the aid of stereomicroscope. 2.3.2. Larval susceptibility First stage larvae were selected to enable monitoring of four development phases of this stage after the infection. The larvae were placed, maintained and infected as described for the eggs. They were fed with the same diet used for the colony, and the larval mortality counts were conducted daily until all surviving larvae reached the pupal stage, with the aid of stereomicroscope. 2.3.3. Adult susceptibility Each repetition consisted of 15 males and 15 females used 48 h post-blood feeding in nylon cages, which were placed in plastic pots, as described previously. For infection, the insects were first chilled to −2 ◦ C for 5 min for immobilization. Immediately after, they were treated with 3 ml of one of the fungal suspensions, cypermethrin or Tween 80. The adult mortality was recorded daily to determine longevity and eggs counts laid by treated females were analyzed after the death of all adults when the pots were opened and observed in stereomicroscope. Larvae hatch born from these eggs were quantified 8 days post-infection to obtain egg-hatching rate. 2.4. M. anisopliae var. acridum growth and microscopic features post-passage in L. longipalpis
2.2. Preparation of M. anisopliae var. acridum inoculum
The unhatched eggs, larvae and dead adults were sterilized with 3 ml 70% ethanol, 3 ml 4% sodium hypochlorite and 3 ml sterile distilled water for 3 min each. The material was then seeded in PDA medium for fungal growth (Alves, 1998). The analysis of fungal growth parameters (conidiogenesis, vegetative growth, colony count and sporulation) were performed in triplicate, and microscopic aspects followed the methodology outlined in Feijó et al. (2007). These data were compared to the parameters observed before the fungal infection.
Metharizium anisopliae var. acridum inoculum was obtained from strain 291 (URM-3800), kindly provided by the Mycology Collection of the Department of Mycology, Universidade Federal de Pernambuco. The fungal cultures were kept in PDA medium (Potato Dextrose Agar, Vertec® ), maintained at 28 ◦ C and diluted to prepare concentrations of 1 × 108 , 1 × 107 , 1 × 106 , 1 × 105 and 1 × 104 conidia/ml in 0.05% (v/v) Tween 80. Conidia were quantified via direct counting using an optical microscope with a Neubauer chamber (Alves and Moraes, 1998).
2.4.1. Conidiogenesis A disc 5 mm in diameter was removed from a M. anisopliae var. acridum culture after 12 days of growth and transferred to a test tube containing 10 ml 0.05% (v/v) Tween 80. The suspension was shaken to separate the conidia, diluted and adjusted to 104 conidia/ml. From this suspension, 0.1 ml was spread-plated on PDA medium using a Drigalski spatula. The number of conidia was determined by counting under a light microscope at 16 h postinoculation. A total of 500 conidia per Petri dish were counted and
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categorized into 2 groups: germinated with having a germ tube in development, and not germinated. 2.4.2. Vegetative growth A disc 5 mm in diameter of M. anisopliae var. acridum was sown in the center of a Petri dish with PDA medium. Growth was measured at day 15. 2.4.3. Colony counting The same dilution of M. anisopliae var. acridum used for the conidiogenesis experiments was also used for colony counting. The dilution was spread on Petri dishes (0.1 ml), and the colonies were counted 3, 6, 9, 12 and 15 days post-inoculation. 2.4.4. Sporulation Using the same methodology used for the vegetative growth experiments, at 3, 6, 9, 12 and 15 days post-inoculation, 10 ml of 70% ethanol was added to 3 Petri dishes with growing fungus for 5 min, for the purpose of conidia inactivation and drying. The 70% ethanol (containing conidia) was retrieved from the surface of the Petri dish and placed into a sterile receptacle. Subsequently, Petri dishes were washed 9 times with 10 ml 0.05% (v/v) Tween 80, and between washes, the Tween 80 solution was removed from the plates and placed in the same container. The conidia were then quantified in a Neubauer chamber. 2.4.5. Microscopic aspects An aliquot of fungal culture was aseptically placed at 4 equidistant points on a Petri dish with PDA medium and covered with a sterile cover slip. These cultures were analyzed with an optical microscope after 24, 48, 72, 96 and 120 h. The fungal structures were stained with Amann blue and observed at 100×, 400× and 1000× magnification. 2.5. Genomic DNA extraction and PCR The genomic DNA of M. anisopliae var. acridum reisolated from infected L. longipalpis eggs, larvae and adults was extracted using an Invisorb® Spin Plant Mini Kit (Invitek, GmbH, Berlin-Buch), then resuspended in 100 l TE and stored at −20 ◦ C, according to the manufacturer’s recommendations. The primers Mac-ITS-spF (5 CTGTCACTGTTGCTTCGGCGGTAC3 ) and Mac-ITS-spR (5 CCCGTTGCGAGTGAGTTACTACTGC3 ) were designed based on the ITS1 and ITS2 regions of the rDNA sequence data for M. anisopliae var. acridum (Entz et al., 2005). PCR amplifications were performed in a total volume of 10 l, containing 1× buffer (20 mM Tris–HCl, pH 8.3, 50 mM KCl), 50 mM MgCl2 , 10 pM/l of each primer, 10 mM dNTP mix (InvitrogenTM ), 5 U/l Platinum Taq DNA polymerase (InvitrogenTM ) and 0.1 l of the target DNA. The negative control contained sterile ultrapure water in place of DNA and the positive control contained M. anisopliae var. acridum (291, URM-3800). The DNA amplification was performed in a Primus 96 HPL thermocycler (MWG Biotech, Inc.) programmed according to the recommendation of Entz et al. (2005): initial denaturation at 94 ◦ C × 5 min, 30 cycles of denaturing at 94 ◦ C × 1 min, annealing and extension combined at 72 ◦ C × 3 min. The amplicons were visualized on a 1% agarose gel containing 1 Kb Plus DNA ladder (InvitrogenTM ), and stained with ethidium bromide and subjected to transillumination with UV light (FB-TI-88).
3100 Genetic Analyzer (Applied Biosystems® ). All fragments were sequenced in both directions and data were processed by programs provided by the sequencer manufacturer. The electrophoretograms generated were stored as files of the Chroma program and the nucleotide sequences were submitted to GenBank. Data were then analyzed using BioEdit software (Hall, 1999) for the purpose of verifying the sequence quality. The Chroma program was used to transform the data output files into the FASTA format (default), giving quality values (0–99) for each nucleotide using algorithms to specify each peak’s intensity (height and width) generated by the electrophoretogram. The sequences were also subjected to BLAST (Basic Local Alignment Search Tool) (Altschul et al., 1990) to verify the similarities of the obtained fragment sequences to the sequences of other proteins in GenBank. 2.7. Statistical analysis The design was completely randomized for all experiments. The effects of fungal infection on egg hatching, larval mortality and adult longevity, as well as the data concerning fungal growth parameters, were normalized when necessary and submitted to ANOVA and Pearson correlation coefficient analyses. After analysis of variance, the means were compared using a Student–Newman–Keuls test, and the growth parameters were compared using Dunnett’s Method. For both, P < 0.05 (SigmaStat software 3.1). 3. Results 3.1. Susceptibility of L. longipalpis to entomopathogenic fungus At the highest concentrations, M. anisopliae var. acridum infection reduced the number of L. longipalpis eggs that hatched by 40% (H = 33.03, df = 6, P < 0.05), although its effect was lower than that of the positive control. There was a direct and significant correlation (r = 0.86) between fungal concentration and the inhibition of egg hatching (F = 22.80, df = 1, P < 0.001). An increase in mortality of the larvae born of these eggs was also observed, and an index of 60% was found at higher concentrations (H = 27.17, df = 6, P < 0.05) (Table 1). The fungus had a greater effect on larval mortality than the control, and the effect was inoculum-dependent reaching 87% (H = 36.04; df = 6, P < 0.05) (Table 1). The mean larval survival time was 26.51 ± 6.75 days for the treated group and 31.89 ± 4.48 for the negative control. However, only the positive control (which caused instantaneous death) had a significant effect on mean larval survival time (H = 29.92, df = 6, P < 0.05). The longevity of adults infected with the fungus was significantly reduced, on average by 7 days, but this effect did not overcome that of cypermethrin, which caused instantaneous death (F = 43.54, df = 6, P < 0.05). Egg hatching was reduced in fungus-infected females, especially at the highest concentration, which produced 85% efficacy (F = 53.57, df = 6, P < 0.05) (Table 1). There was a significant direct correlation (r = 0.49) between fungal concentration and inhibition of egg hatching (F = 13.61, df = 1, P < 0.05). 3.2. Growth and microscopic features of M. anisopliae var. acridum after passage in L. longipalpis
2.6. DNA sequencing The amplicons were purified by isopropanol/ethanol precipitation according to the manufacturer’s recommendations (Applied Biosystems® ) and sequenced using the BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystem® ) in an ABI PRISM®
M. anisopliae var. acridum was successfully reisolated postinfection only from L. longipalpis adults. The conidiogenesis of the reisolated fungus was significantly higher than that of the fungus before infection (F = 4.53, df = 3, P < 0.05), but vegetative growth was similar (F = 2.40, df = 3, P = 0.14) (Table 2).
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Table 1 The effect of M. anisopliae var. acridum on eggs, larvae and adults of L. longipalpis. Treatments (mean ± SD)
Eggs
Larvae
Hatching reduction (%) 1 × 104 conidia/ml 1 × 105 conidia/ml 1 × 106 conidia/ml 1 × 107 conidia/ml 1 × 108 conidia/ml Tween 80 0.05% Cypermethrin 196 g/ml
28.89 30.00 35.56 40.00 41.67 12.78 100.00
± ± ± ± ± ± ±
8.61b 3.85b 2.72a 8.43a 6.24a 9.05c 0.00d
Adults
Larval mortality (%)
Mortality (%)
18.29 ± 27.02 ± 35.11 ± 37.60 ± 49.83 ± 0.00 ± –
63.59 71.90 72.38 83.81 86.67 40.48 100.00
6.51b 2.67b 8.53a 16.54a 24.12a 0.00c
± ± ± ± ± ± ±
Longevity (days)
18.28c 5.39c 5.35c 13.11b 4.71a 7.31d 0.00e
9.57 6.57 6.29 6.86 7.29 8.71 0.00
± ± ± ± ± ± ±
1.51b 1.27a 1.38a 1.77a 0.76a 1.11b 0.00c
Hatching reduction (%) 53.69 ± 64.54 ± 72.30 ± 72.12 ± 84.76 ± 39.26 ± –
10.20c 1.88b 4.97b 3.11b 7.91a 6.68d
Means followed by the same lowercase letter in the same column are not significantly different (Student–Newman–Keuls test, P < 0.05).
Table 2 Conidiogenesis and vegetative growth of Beauveria bassiana reisolated from infected Lutzomyia longipalpis eggs, larvae and adults on PDA medium. Stages (mean ± SD)
Conidiogenesis (no.)* 16 h post-inoculation
Vegetative growth (cm)NS 15 days post-inoculation
Adults Control#
413.00 ± 36.77a 255.00 ± 7.00b
4.07 ± 0.98a 3.07 ± 0.29a
Means followed by the same lowercase letter in the same column are not significantly different (Dunnett’s Method, * (P < 0.05) and NS (P = 0.14). # Metharizium anisopliae var. acridum 291 (URM-3800) before infection of L. longipalpis.
The number of fungus colonies was significantly higher post-passage in adult sand flies (F = 13.05; df = 3, P < 0.05). Sporulation was also significantly higher post-passage, and it was also higher than the fungus before infection and increasing over the entire observation period (F = 158.50; df = 3, P < 0.001) (Table 3). M. anisopliae var. acridum microscopic structures were observed post-infection. Mycelium formation, anastomoses and apressoria were observed in the first 96 h, as were primordia from the conidiophores and young conidiophores along the hyphal axis (data not shown). There were no morphological differences between the fungus used for infection and the fungus reisolated from L. longipalpis adults.
Fig. 1. Amplification of M. anisopliae var. acridum DNA from L. longipalpis adults at different days post-infection with varying conidia/ml concentrations: lane Br: no DNA, lane C+: M. anisopliae var. acridum DNA, lane 1: 1 × 106 at 7 days, lanes 2–4: 1 × 107 at 1, 5 and 6 days, lanes 5–7: 1 × 108 at 1, 6 and 10 days, Kb: ladder.
4. Discussion 3.3. PCR and sequencing The eggs of insects are more resistant to infection than other developmental stages (Tanada and Kaya, 1992). Although fungal infection reduced egg hatching significantly and increased the mortality of larvae born from infected eggs, the effectiveness was around 40%. Superior results were observed when another entomopathogenic fungus, Beauveria bassiana (Bals.) Vuilleman (Deuteromycotina: Hyphomycetes), was tested on eggs of the same sand fly species reaching a 60% efficacy on egg hatching and 88% on the mortality of larvae born from infected eggs (Amóra et al., 2009). Additionally, another biocontrol study showed that aqueous suspensions of Bacillus sphaericus Meyer and Neide (1904) (Bacillales: Bacillaceae) inhibited Phlebotomus duboscqi Leveu-Nemaire (1906) and Sergentomyia schwetzi Adler, Theodor and Parrot (1929) (Diptera: Psychodidae) egg hatching by 95% (Robert et al., 1998).
M. anisopliae was reisolated from L. longipalpis adults infected with 1 × 106 , 1 × 107 and 1 × 108 conidia/ml 6 days post-infection. In all cases, M. anisopliae var. acridum DNA was amplified by PCR (Fig. 1). M. anisopliae var. acridum DNA could not be reisolated from eggs, larvae or adults infected with 1 × 104 and 1 × 105 conidia/ml, but it was possible to observe mycelial growth on insects infected with 1 × 107 and 1 × 108 conidia/ml. No PCR products were amplified from the control group. The DNA sequences obtained were approximately 394 bp in length. These were compared with other sequences deposited in GenBank (NCBI), and this confirmed that the reisolated fungal species was in fact a match to M. anisopliae var. acridum (accession number EF113338.1) with 100% homology (e-value 0.0).
Table 3 Colony counts and sporulation of M. anisopliae var. acridum (mean ± SD) reisolated from infected L. longipalpis adults at 3, 6, 9, 12 and 15 days after inoculation on PDA medium. Parameters
Stages
Colony counting (no. colony)
Adults Control#
3 days 8.50 ± 4.95a 0.00 ± 0.00b
Sporulation (no. conidia)
Adults Control#
12.62 ± 0.54a 4.06 ± 0.08b
6 days
9 days
846.00 ± 124.45a 276.00 ± 69.30b
40.50 ± 0.71a 7.00 ± 1.41b
12 days 62.50 ± 2.12a 0.50 ± 0.71b
15 days 17.00 ± 2.83a 0.00 ± 0.00b
40.11 ± 0.18a 26.27 ± 0.18b
53.38 ± 0.32a 59.16 ± 0.06b
148.48 ± 0.28a 115.19 ± 0.01b
164.66 ± 0.87a 161.40 ± 0.29b
Means followed by the same lowercase letter in the same column are not significantly different (Student–Newman–Keuls test, P < 0.05). # Metharizium anisopliae var. acridum 291 (URM-3800) before infection of L. longipalpis.
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The efficiency of M. anisopliae var. acridum at the concentration of 1 × 108 conidia/ml on larvae of L. longipalpis (87%) and Boophillus microplus Canestrini (1887) (Acari: Ixodidae) (97%) (Bahiense et al., 2006) was similar. But, was smaller than the use of B. bassiana in the same concentration on L. longipalpis larvae obtaining an efficiency of 100% of mortality (Amóra et al., 2009). Comparing the susceptibility of sand fly larvae to different biocontrols, the result of spraying fungus on L. longipalpis larvae was better than offering a diet contaminated with Bacillus thuringiensis var. israelensis Goldberg and Margalit (1977) (Bacillales: Bacillaceae) to Phlebotomus papatasi Scopoli (1763) larvae (Diptera: Psychodidae) (Wahba et al., 1999). Both of these had inoculum-dependent effects. However, the fungus was not reisolated, and similar to P. papatasi larvae infected with B. sphaericus, bacteraemia was not detected (Wahba, 2000). Biolarvicide application in the field is difficult due to the diversity of sand fly breeding habitat, thus its application appears to be limited to adult control. More research needs to be done to develop an efficient larvicide for vector control. Any reduction in mosquito longevity reduces the mean number of blood meals taken, and the probability of the vector acquiring a disease agent and subsequently transmitting it to other hosts is also reduced (Scholte et al., 2007). The longevity of L. longipalpis infected with M. anisopliae var. acridum was significantly lower than that of uninfected sand flies, except for sand flies infected at the concentration of 1 × 104 conidia/ml, whose longevity was greater than the negative control. These data are similar to the effect of other entomopathogenic fungi against P. papatasi, L. longipalpis (Warburg, 1991) and Aedes aegypti Linnaeus (1762) (Scholte et al., 2007) in that they show the highest mortality rates at around seven days. While the effect of M. anisopliae was tested on mites, the longevity of these insects was six days (Kanga et al., 2002). More recently another in vitro study with B. bassiana on adult L. longipalpis reduced the longevity of the insect to 4 days (Amóra et al., 2009). Although there was no significant decrease in infected female oviposition, there was a significant reduction in egg hatching. Similar data were obtained with the same fungus in grasshoppers (Blanford and Thomas, 2001) and B. bassiana in L. longipalpis (Amóra et al., 2009). The low efficiency of M. anisopliae var. acridum compared to other entomopathogenic fungi could be related to its primary specificity for Acridids (Blanford and Thomas, 2001). The reduction in longevity and fecundity may not decrease the vectorial capacity of adult L. longipalpis in the short term, but it could still be effective over the long term. It is possible that infected sand flies are more susceptible to factors such as predation and secondary infections by other pathogens, and infection might also decrease insect migration ability. Traditionally, the identification of the genus Metarhizium is based on the observation of morphological features on culture media and microscopic examination of spores and associated structures (Entz et al., 2005). Fungal growth studies, such as analysis of conidiogenesis, sporulation, colony counts and radial growth were conducted to assist entomopathogenic fungus characterization, as recommended by Almeida et al. (2005), because these parameters are important for defining the virulence of the fungal isolate (Liu et al., 2003). The rates of radial and vegetative growth are directly linked to the speed of infection in the host (Feijó et al., 2007), probably because the radial growth is used as a criterion of selection of mycoinsecticides (Yeo et al., 2003). However, in this study the vegetative growth of M. anisopliae var. acridum was slow, possibly explaining the failure to reisolate it from L. longipalpis eggs and larvae. Similar data were observed by Feijó et al. (2009) and by Amóra et al. (2009), but in the latter case the fungus utilized was B. bassiana although the fungus has been reisolated of all development phases of L. longipalpis. The sporulation, conidiogenesis and colony counts of M. anisopliae var. acridum was significantly higher after reisolation,
corroborating the results of Entz et al. (2005) and Amóra et al. (2009). However the data of sporulation and colony counts of M. anisopliae var. acridum re-isolated from C. albiceps were lower than control (Feijó et al., 2009). These data suggest a positive correlation between conidia production and time. Rapid sporulation may be an important criterion for the selection of fungal isolates because it helps epizootic spread (Mitchell, 2003). Conidiogenesis is another important parameter to consider, as the number of germinated conidia is directly proportional to isolate virulence. The conidiogenesis rate may be influenced by the storage form, the presence of nutrients and the mode of host exposure to fungus (Alves, 1998). Nevertheless, the studies with M. anisopliae var. acridum are still incipient (Feijó et al., 2009). In the analysis of the vegetative and reproductive structures present in M. anisopliae var. acridum before and after passage in adult L. longipalpis was observed the formation of mycelium in the first 48 h, anastomosis at 72 h, appressoria at 96 h and at 120 h abundant ramified conidiophores were also observed. This structures reisolated from adult did not differ from the control, these data are similar to the findings of Feijó et al. (2009). The anastomoses permit the intercommunication between the fungic segments, allowing the passage into cytoplasmatic organelles, including the nuclear migration (Messias and Azevedo, 1980). This phenomenon allows the recombination and genetic improvement in economically important fungi that do not present a sexual cycle (Alves, 1998). The appressoria, in turn, are adhesion sites and production of enzymes assisting the fungus to penetrate the cuticle of the insect target and nutrition (St. Leger et al., 1989). The colony morphology of M. anisopliae var. acridum was examined, producing dark yellow-green conidial masses, conidia typically are ovoid, similar to Brazilian (Feijó et al., 2009), Australian, Korean (Fernandes et al., 2009), Canadian (Entz et al., 2005) and Mexican strains (Milner et al., 2003). However it is important to emphasize that difficulties in identification are frequently encountered, and morphological differences can be exhibited under varying environmental and physiological conditions. The spore morphology varies within the same culture and between isolates of the genus Metarhizium. Conidia and blastospores can be of variable size and shape (Milner et al., 2003). Therefore, M. anisopliae var. acridum cannot be distinguished from other M. anisopliae varieties on the basis of spore size and shape (Fernandes et al., 2009). Thus, molecular tests should be performed (Lomer et al., 2001) that allow differentiation of fungal strains with similar morphological and cytological characteristics (Kouvelis et al., 2008). The failure to reisolate fungus from treated eggs and larvae prevented completion of the PCR, but its effect was observed statistically on L. longipalpis. The same situation was observed in a previous study that involved the histological examination of Lutzomyia young Feliciangeli and Murillo (1987) (Diptera: Psychodidae) infected with B. bassiana, where the histological sections were negative and no cuticle or tissue damage was observed, but the fungal mycelium was seen (Reithinger et al., 1997), similar to our observations. With respect to adult L. longipalpis, the PCR and sequencing proved that infection by the fungus had occurred at the two higher concentrations on different days post-infection. Comparative studies of nucleotide sequences of rDNA genes have provided significant data for phylogenetic and taxonomic analyses (Entz et al., 2005). As expected, the Mac-ITS-spF and Mac-ITS-spR primers successfully amplified a 394 bp DNA sequence from the total genomic DNA extracted from M. anisopliae var. acridum, similar to the results of Entz et al. (2005). In that same study, no other species of Metharizium, other fungal entomopathogens, or other microorganisms were detected by these primers, demonstrating their specificity (Entz et al., 2005).
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Although the effect of an entomopathogenic fungus is not immediate, it can minimize the environmental contamination risk inherent in chemical insecticides (Oliveira et al., 2003). At present, the potential risk of adverse effects of the fungus on human health when used indoors and/or in the vicinity of immunocompromised individuals is considered low due to the pathogen’s opportunistic nature (Scholte et al., 2007). Although some cases have been reported involving exclusively immunocompromised patients (Burgner et al., 1998; Revankar et al., 1999; Osorio et al., 2007) and only one case of feline rhinitis (Muir et al., 1998) and another of human keratitis (Jani et al., 2001) in healthy patient, all caused by M. anisopliae var. anisopliae, with effective treatment and without relapse. Thus, to determine sensitivity to these organisms is necessary to identify specific allergens to provide a basis for mitigation of these allergens causing biocontrol products safer (Westwood et al., 2005). The finding that L. longipalpis is susceptible to infection with an entomopathogenic fungus is the first step towards the final goal of using this fungus for VL vector control. However, there are many more scientific issues to confront, such as the screening of other fungal species and strains, finding the optimal dosages and application techniques. In particular, fungal persistence following application must be substantial to minimize the logistical challenges and cost of re-treatment (Scholte et al., 2007). Several entomopathogenic fungi are already in use for agricultural and veterinary pest control, demonstrating the feasibility of this biocontrol technique with respect to acceptability and applicability. Among the fungi currently used for that purpose stands B. bassiana, which recently demonstrated to be pathogenic on L. longipalpis (Amóra et al., 2009). The results presented here suggest a lack of pathogenicity of M. anisopliae var. acridum against L. longipalpis eggs and larvae, leading to the conclusion that this pathogen should not be used for the control of sand flies in their immature stages. However, in terms of adult mortality and infected female fecundity, the results were significant, proving that the primary vector species of VL is susceptible to entomopathogenic fungal infection in the adult stage. These results encourage further studies on the use of M. anisopliae var. acridum as a biocontrol agent. Acknowledgements We thank Dr. Nélio B. Morais, Richristi A. Silva, Raimundo Nonato de Sousa and Lindemberg Caranha (Secretaria de Saúde do Estado do Ceará), Ana Claudia B. Mendonc¸a and Sodré Rocha (Secretaria Municipal de Saúde de Mossoró) for assistance in sand fly field collections, Dr. Rui Sales Júnior, Dra. Celicina M.S.B. Azevedo (UFERSA). UFERSA administrative staff for logistic support. To Msc Lorena M.B. Oliveira for reviewing and improving the paper. As also the inhabitants of the studied areas for their patient and kindness. Financial support: Msc Amora has a grant from CAPES and Dr. Bevilaqua is a CNPq researcher. Ethical approval: Ceará State University, Committee of Ethics for the Use of Animals (process no. 07465297-4). References Almeida, J.C., Albuquerque, A.C., Luna-Alves Lima, E.A., 2005. Viabilidade de Beauveria bassiana (bals.) Vuill. reisolado de ovos, larvas e adultos de Anthonomus grandis (Boheman) (Coleoptera: Curculionidae) artificialmente infectado. Arq. Inst. Biol. São Paulo 72, 473–480. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403–410. Alves, S.B. (Ed.), 1998. Controle Microbiano de Insetos, 2nd ed. FEALQ, Piracicaba, São Paulo. Alves, S.B., Moraes, S.A., 1998. Quantificac¸ão de inóculo de patógenos de insetos. In: Alves, S.B. (Ed.), Controle Microbiano de Insetos, 2nd ed. FEALQ, Piracicaba, São Paulo, pp. 765–777.
219
Amóra, S.S.A., Santos, M.J.P., Alves, N.D., Gonc¸alves da Costa, S.C., Calabrese, K.S., Monteiro, A.J., Rocha, M.F.G., 2006. Fatores relacionados com a positividade de cães para leishmaniose visceral em área endêmica do Estado do Rio Grande do Norte, Brasil. Cienc. Rural 36, 1854–1859. Amóra, S.S.A., Bevilaqua, C.M.L., Feijó, F.M.C., Silva, M.A., Pereira, R.H.M.A., Silva, S.C., Alves, N.D., Freire, F.A.M., Oliveira, D.M., 2009. Evaluation of the fungus Beauveria bassiana (Deuteromycotina: Hyphomycetes), a potential biological control agent of Lutzomyia longipalpis (Diptera, Psychodidae). Biol. Control 50, 329–335. Aransay, A.M., Scoulica, E., Tselentis, Y., 2000. Detection and identification of Leishmania DNA within naturally infected sand flies by seminested PCR on minicircle kinetoplastic DNA. Appl. Environ. Microbiol. 66, 1933–1938. Bahiense, T.C., Fernandes, E.K.K., Bittencourt, V.R.E.P., 2006. Compatibility of the fungus Metarhizium anisopliae and deltamethrin to control a resistant strain of Boophilus microplus tick. Vet. Parasitol. 141, 319–324. Blanford, S., Thomas, M.B., 2001. Adult survival, maturation, and reproduction of the desert locust Schistocerca gregaria infected with the fungus Metarhizium anisopliae var. acridum. J. Invert. Pathol. 78, 1–8. Burgner, D., Eagles, G., Burgess, M., Procopis, P., Rogers, M., Muir, D., Pritchard, R., Hocking, A., Priest, M., 1998. Disseminated invasive infection due to Metarrhizium anisopliae in an immunocompromised child. J. Clin. Microbiol. 36, 1146–1150. Driver, F., Milner, R.F., Trueman, W.H., 2000. A taxonomic revision of Metarhizium based on a phylogenetic analysis of ribosomal DNA sequence data. Mycol. Res. 104, 134–150. Dye, C., 1996. The logic of Visceral Leishmaniasis control. Am. J. Trop. Med. Hyg. 55, 125–130. Entz, S.C., Johnson, D.L., Kawchuk, L.M., 2005. Development of a PCR-based diagnostic assay for the specific detection of the entomopathogenic fungus Metarhizium anisopliae var. acridum. Mycol. Res. 109, 1302–1312. Faria, M.R., Magalhães, B.P., 2001. O uso de fungos entomopatogênicos no Brasil. Biotecnolog. Cienc. Desenvolv. 22, 18–21. Feijó, F.M.C., Lima, P.M., Alves, N.D., Luna-Alves Lima, E.A., 2007. Comportamento e aspectos citológicos de Beauveria basssiana após passagem em ovo, larva e adulto de Chrysomya albiceps. Arq. Inst. Biol. 74, 349–355. Feijó, F.M.C., Lima, P.M., Melo, E.H.M., Maciel, M.V., Athayde, A.C.R., Alves, N.D., Luna-Alves Lima, E.A., 2008. Susceptibilidade de Chrysomya albiceps a Beauveria bassiana em condic¸ões de laboratório. Cienc. Anim. Bras. 9, 771–777. Feijó, F.M.C., Lima, P.M., Melo, E.H.M., Athayde, A.C.R., Luna-Alves Lima, E.A., 2009. Behavior and cytological aspects of Metarhizium anisopliae and Metarhizium flavoviride after passage in Chrysomya albiceps. Rev. Caatinga 22, 39–44. Fernandes, É.K.K., Keyser, C.A., Chong, J.P., Rangel, D.E.N., Miller, M.P., Roberts, D.W., 2009. Characterization of Metarhizium species and varieties based on molecular analysis, heat tolerance and cold activity. J. Appl. Microbiol. Epub ahead of print. doi:10.1111/j.1365-2672.2009.04422.x. Galati, E.A.B., 2003. Morfologia e Taxonomia. Morfologia, terminologia de adultos e identificac¸ão dos táxons da América. In: Rangel, E.F., Lainson, R. (Eds.), Flebotomíneos do Brasil. Fiocruz, Rio de Janeiro, pp. 53–175. Goettel, M.S., Jonshon, D.L., Inglis, G.D., 1995. The role of fungi in the control of grasshoppers. Can. J. Bot. 73, S71–S75. Hall, T., 1999. BioEdit Version 5.0.6. North Carolina State University, Department of Microbiology. Jani, B.R., Rinaldi, M.G., Reinhart, W.J., 2001. An unusual case of fungal keratitis: Metarrhizium anisopliae. Cornea 20, 765–768. Kanga, L.H.B., James, R.R., Boucias, D.G., 2002. Hirsutella thompsonii and Metarhizium anisopliae as potential microbial control agents of Varroa destructor, a honey bee parasite. J. Invert. Pathol. 81, 175–184. Kouvelis, V.N., Ghikas, D.V., Edgington, S., Typas, M.A., Moore, D., 2008. Molecular characterization of isolates of Beauveria bassiana obtained from overwintering and summer populations of Sunn Pest (Eurygaster integriceps). Lett. Appl. Microbiol. 46, 414–420. Liu, H., Skinner, M., Brownbrigde, M., Parker, B.L., 2003. Characterization of Beauveria bassiana and Metarhizium anisopliae isolates for management of tarnished plant bug, Lygus lineolaris (Himeptera, Miridae). J. Invert. Patholol. 82, 139–147. Lomer, C.J., Bateman, R.P., Johnson, D.L., Langewald, J., Thomas, M., 2001. Biological control of locusts and grasshoppers. Ann. Rev. Entomol. 46, 667–702. Magalhães, B.P., De Faria, M.R., Lecoq, M., Schmidt, F.G.V., Silva, J.B.T., Frazão, H.S., Balanc¸a, G., Foucart, A., 2001. The use of Metarhizium anisopliae var. acridum against the grasshopper Rhammatocerus schistocercoides in Brazil. J. Orthoptera Res. 10, 199–202. Messias, C.L., Azevedo, J.L., 1980. Parassexuality in the deuteromycete Metarhizium anisopliae. Trans. Br. Mycol. Soc. 75, 473–477. Milner, R.J., Lozano, L.B., Driver, F., Hunter, D., 2003. A comparative study of two Mexican isolates with an Australian isolate of Metarhizium anisopliae var. acridum—strain characterisation, temperature profile and virulence for wingless grasshopper, Phaulacridium vittatum. BioControl 48, 335–348. Ministério da Saúde. Secretaria de Vigilância em Saúde. Departamento de Vigilância Epidemiológica, 2006. Manual de vigilância e controle da leishmaniose visceral. Brasília: Ministério da Saúde, 122 pp. [cited on August 30, 2008]. Avaliable from: http://portal.saude.gov.br/portal/arquivos/pdf/manual leish visceral2006.pdf. Mitchell, J.K., 2003. Development of a submerged liquid sporulation medium for the potential smart weed bioherbicide Septaria polyganorum. Biol. Control 27, 293–299. Muir, D., Martin, P., Kendall, K., Malik, R., 1998. Invasive hyphomycotic rhinitis in a cat due to Metarhizium anisopliae. Med. Mycol. 36, 51–54.
220
S.S.A. Amóra et al. / Acta Tropica 113 (2010) 214–220
Oliveira, C.N., Neves, P.M.O.J., Kawazoe, L.S., 2003. Compatibility between the entomopathogenic fungus Beauveria bassiana and insecticides used in coffee plantations. Sci. Agric. 60, 663–667. ˜ F., Rodriguez-Tudela, Osorio, S., de la Cámara, R., Monteserin, M.C., Granados, R., Ona, J.L., Cuenca-Estrella, M., 2007. Recurrent disseminated skin lesions due to Metarrhizium anisopliae in an adult patient with acute myelogenous leukemia. J. Clin. Microbiol. 45, 651–655. Rangel, E.F., Souza, N.A., Wermelinger, E.D., Barbosa, A.F., 1985. Estabelecimento de colônia, em laboratório, de Lutzomyia intermedia Lutz & Neiva, 1912 (Diptera, Psychodidae, Phlebotominae). Mem. Inst. Oswaldo Cruz 80, 219–226. Reithinger, R., Davies, C.R., Cadena, H., Alexander, B., 1997. Evaluation of the fungus Beauveria bassiana as a potential biological control agent against phlebotomine sand flies in colombian coffee plantations. J. Invertebr. Pathol. 70, 131–135. Revankar, S.G., Sutton, D.A., Sanche, S.E., Rao, J., Zervos, M., Dashti, F., Rinaldi, M.G., 1999. Metarrhizium anisopliae as a cause of sinusitis in immunocompetent hosts. J. Clin. Microbiol. 37, 195–198. Robert, L.L., Perich, M.J., Schlein, Y., Jacobson, R.L., 1998. Bacillus sphaericus inhibits hatching of phlebotomine sand fly eggs. J. Am. Mosq. Control. Assoc. 14, 351–352. Scholte, E.-J., Knols, B.G.J., Samson, R.A., Takken, W., 2004. Entomopathogenic fungi for mosquito control: a review. J. Insect Sci. 4, 1–24. Scholte, E.-J., Takken, W., Knols, B.G.J., 2007. Infection of adult Aedes aegypti and Ae. albopictus mosquitoes with the entomopathogenic fungus Metarhizium anisopliae. Acta Trop. 102, 151–158. Secundino, N.F.C., Araújo, M.S.S., Oliveira, G.H.B., Massara, C.L., Carvalho, O.S., Lanfredi, R.M., Pimenta, P.F.P., 2002. Preliminary description of a new ento-
moparasitic nematode infecting Lutzomyia longipalpis sand fly, the vector of Visceral Leishmaniasis in the New World. J. Invert. Pathol. 80, 35–40. Sharma, U., Singh, S., 2008. Insect vectors of Leishmania: distribution, physiology and their control. J. Vector Borne Dis. 45, 255–272. Singh, S., 2006. New developments in diagnosis of leishmaniasis. Indian J. Med. Res. 123, 311–330. St. Leger, R.J.S., Butt, T.M., Goettel, M.S., Staples, R.S., Roberts, D.W., 1989. Production in vitro of appressoria by the entomopathogenic fungus Metarhizium anisopliae. Exp. Mycol. 13, 274–288. Tanada, Y., Kaya, H.K., 1992. Insect Pathology. Elsevier Science & Technology Books, Hardcover. Wahba, M.M., 2000. The influence of Bacillus sphaericus on the biology and histology of Phlebotomus papatasi. J. Egypt. Soc. Parasitol. 30, 315–323. Wahba, M.M., Labib, I.M., el Hamshary, E.M., 1999. Bacillus thuringiensis var. israelensis as a microbial control agent against adult and immature stages of the sand fly. Phlebotomus papatasi under laboratory conditions. J. Egypt. Soc. Parasitol. 29, 587–597. Warburg, A., 1991. Entomopathogens of phlebotomine sand flies, laboratory experiments and natural infections. J. Invertebr. Pathol. 58, 189–202. Westwood, G.S., Huang, S.W., Keyhani, N.O., 2005. Allergens of the entomopathogenic fungus Beauveria bassiana. Clin. Mol. Allergy 3, 1–8. Yeo, H., Pell, J.K., Alderson, P.G., Clark, S.J., Pye, B.J., 2003. Laboratory evaluation of temperature effects on the germination and growth of entomopathogenic fungi and on their pathogenicity to two aphid species. Pest Manage. Sci. 59, 156–165.
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The effects of the fungus Metarhizium anisopliae var. acridum on different stages of Lutzomyia longipalpis (Diptera: Psychodidae) Sthenia Santos Albano Amóra a,∗ , Claudia Maria Leal Bevilaqua a,∗ , Francisco Marlon Carneiro Feijó b , Romeika Hermínia de Macedo Assunc¸ão Pereira b , Nilza Dutra Alves b , Fúlvio Aurélio de Morais Freire b , Michel Toth Kamimura c , Diana Magalhães de Oliveira c , Elza Áurea Luna-Alves Lima d , Marcos Fábio Gadelha Rocha a a
Programa de Pós-Graduac¸ão em Ciências Veterinárias, Universidade Estadual do Ceará, Fortaleza, Ceará 60740-000, Brazil Laboratório de Microbiologia Veterinária, Universidade Federal Rural do Semi-Árido, Mossoró, Rio Grande do Norte 59625-900, Brazil c Núcleo de Genômica e Bioinformática Tarsísio Pimenta, Universidade Estadual do Ceará, Fortaleza, Ceará 60740-000, Brazil d Departamento de Micologia, Centro de Ciências Biológicas, Universidade Federal de Pernambuco, Recife, Pernambuco 50741-040, Brazil b
a r t i c l e
i n f o
Article history: Received 19 May 2009 Received in revised form 22 October 2009 Accepted 24 October 2009 Available online 31 October 2009 Keywords: Biological control Vector Lutzomyia longipalpis Entomopathogenic fungus Metarhizium anisopliae var. acridum Visceral Leishmaniasis
a b s t r a c t The control of Visceral Leishmaniasis (VL) vector is often based on the application of chemical residual insecticide. However, this strategy has not been effective. The continuing search for an appropriate vector control may include the use of biological control. This study evaluates the effects of the fungus Metarhizium anisopliae var. acridum on Lutzomyia longipalpis. Five concentrations of the fungus were utilized, 1 × 104 to 1 × 108 conidia/ml, accompanied by controls. The unhatched eggs, larvae and dead adults previously exposed to fungi were sown to reisolate the fungi and analysis of parameters of growth. The fungus was subsequently identified by PCR and DNA sequencing. M. anisopliae var. acridum reduced egg hatching by 40%. The mortality of infected larvae was significant. The longevity of infected adults was lower than that of negative controls. The effects of fungal infection on the hatching of eggs laid by infected females were also significant. With respect to fungal growth parameters post-infection, only vegetative growth was not significantly higher than that of the fungi before infection. The revalidation of the identification of the reisolated fungus was confirmed post-passage only from adult insects. In terms of larvae mortality and the fecundity of infected females, the results were significant, proving that the main vector species of VL is susceptible to infection by this entomopathogenic fungus in the adult stage. © 2009 Elsevier B.V. All rights reserved.
1. Introduction Leishmaniasis is a zoonosis transmitted by vector rather complex because it involves several host species and sand fly vectors both the ecology and epidemiology of the disease are multifarious (Singh, 2006). The sand fly Lutzomyia longipalpis Lutz and Neiva (1912) is the primary vector for the etiological agent of Visceral Leishmaniasis (VL), Leishmania chagasi Cunha and Chagas (1937), which constitutes a serious public health problem in South America (Secundino et al., 2002). The VL control strategy is comprised of a combination of early detection, drug treatment, euthanasia of positive dogs, environmental management and chemical-based vector control. However, these measures have not been effective (Amóra et al., 2006). In the
∗ Corresponding authors. Tel.: +55 85 3101 9853; fax: +55 85 3101 9840. E-mail addresses: [email protected] (S.S.A. Amóra), [email protected] (C.M.L. Bevilaqua). 0001-706X/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.actatropica.2009.10.018
absence of an effective vaccine or an ideal drug treatment, the best method to interrupt any vector-borne disease is to reduce humanvector contact (Sharma and Singh, 2008). A search for appropriate vector control methods to augment the current limited arsenal of tools is required, and biological control is one promising avenue for this search (Scholte et al., 2007). Over the last decade, there have been major developments in biological, pesticide-based and integrated control strategies for mosquito vectors (Scholte et al., 2004). However, little information is available on the biological control of sand flies. In laboratory studies, the infection of sand flies with different microorganisms such as nematodes (Secundino et al., 2002), bacilli (Robert et al., 1998; Wahba et al., 1999; Wahba, 2000) and fungi (Warburg, 1991; Reithinger et al., 1997; Amóra et al., 2009) inhibited egg hatching and caused larval and adult death, but these results were variable. Meanwhile, there is little information on the effects parasites may produce on sand flies. In particular, entomopathogenic fungi have been highlighted as the main agent in insect control (Scholte et al., 2004). These
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pathogens have been shown to be an alternative to biological control method against many insects (Alves, 1998; Faria and Magalhães, 2001; Scholte et al., 2004), and they are consequently an additional tool for VL control, considering that one of the most effective control of VL is the vector control (Dye, 1996), reducing the use of chemical insecticides, resulting in benefits to humans and the environment (Amóra et al., 2009). In this context, Metarhizium anisopliae (Metsch.) Sorokin (Ascomycota: Hypocreales) is able to infect over 200 insect species (Alves, 1998), is considered to be a highly infective pathogenic fungus against grasshoppers (Goettel et al., 1995) and has been studied for the biological control of several other species of insects (Kanga et al., 2002; Bahiense et al., 2006; Scholte et al., 2007), including Diptera (Feijó et al., 2009). Specifically, M. anisopliae var. acridum [formerly Metarhizium flavoviride Gams and Rozsypal, but now reclassified (Driver et al., 2000)] is known for its effectiveness in the control of grasshoppers in Brazil and elsewhere (Faria and Magalhães, 2001). Currently, this fungus is commercialized under the trade name “Green Muscle® – Lubilosa” and “Green Guard® – Becker Underwood Pty Ltd.” for grasshopper (Acrididae) control in South Africa and Australia, respectively (Entz et al., 2005). In Brazil, field tests have been conducted (Magalhães et al., 2001). Despite its importance to agriculture, its pathogenicity in Diptera that are related to public health issues have not yet been sufficiently explored. However the efficacy results of M. anisopliae var. acridum on Diptera Chrysomya albiceps (Feijó et al., 2009) encourages new researches with this fungus on other Diptera. Also it is now known that this pathogen can be used to efficiently control Rhammatocerus schistocercoides Rehn (1906) (Orthoptera: Acrididae) in Brazil (Magalhães et al., 2001) Thus, the biological potential of M. anisopliae var. acridum makes this fungus a candidate for control of other Diptera, including sand flies. To this end, the present study aimed to evaluate the effect of M. anisopliae var. acridum on various stages of L. longipalpis development. 2. Materials and methods 2.1. L. longipalpis collection and identification L. longipalpis captured in the field were maintained in BOD incubators acclimatized to 27 ◦ C, 80% RH with a photoperiod of 12 h (Rangel et al., 1985). To promote oviposition, female sand flies were allowed to feed and obtain a blood meal from anesthetized hamsters for 2 h. Forty-eight hours post-feeding, the adult females were individualized in plastic pots measuring 4 cm in diameter and 4.5 cm in height that were internally coated with sterile plaster to maintain moisture. After oviposition, the females were dissected (Aransay et al., 2000) for identification (Galati, 2003). The hatched larvae were fed daily with a diet based on rabbit feces and crushed dried cassava leaves until they reached the pupa stage. After emergence, adults were transferred to nylon tulle cages measuring 20 cm3 in diameter, fed for 3 days with a glucose solution soaked in sterile cotton, and on the 4th day, an anesthetized hamster was offered to provide a blood meal for the females.
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2.3. L. longipalpis susceptibility to M. anisopliae var. acridum The bioassays, in which eggs, larvae and adults were treated with 5 fungal concentrations, were conducted with 2 control groups: 0.05% (v/v) sterile Tween 80 (negative control) (Feijó et al., 2008) and 196 g/ml of the pyrethroid, cypermethrin (positive control). The insecticide chosen as positive control is the compound used by the Brazilian Ministry of Health in the sand flies control (Ministério da and Saúde, 2006). Randomized treatments (21) were performed; 7 for each insect stage, with each treatment performed in 3 repetitions and each repetition in triplicate. Each repetition consisted of 30 samples totaling 630 individuals/ repetition. 2.3.1. Egg susceptibility The eggs were placed in plastic pots similar to those used in the maintenance of the colony. One of the fungal suspensions (3 ml), cypermethrin or Tween 80 was spilled on the walls and ground of the pots using a pipette. The containers were then stored in BOD incubators at 27 ◦ C, 80% RH with a photoperiod of 12 h. The egg hatching was observed daily and larval mortality was counted 8 days post-treatment, with the aid of stereomicroscope. 2.3.2. Larval susceptibility First stage larvae were selected to enable monitoring of four development phases of this stage after the infection. The larvae were placed, maintained and infected as described for the eggs. They were fed with the same diet used for the colony, and the larval mortality counts were conducted daily until all surviving larvae reached the pupal stage, with the aid of stereomicroscope. 2.3.3. Adult susceptibility Each repetition consisted of 15 males and 15 females used 48 h post-blood feeding in nylon cages, which were placed in plastic pots, as described previously. For infection, the insects were first chilled to −2 ◦ C for 5 min for immobilization. Immediately after, they were treated with 3 ml of one of the fungal suspensions, cypermethrin or Tween 80. The adult mortality was recorded daily to determine longevity and eggs counts laid by treated females were analyzed after the death of all adults when the pots were opened and observed in stereomicroscope. Larvae hatch born from these eggs were quantified 8 days post-infection to obtain egg-hatching rate. 2.4. M. anisopliae var. acridum growth and microscopic features post-passage in L. longipalpis
2.2. Preparation of M. anisopliae var. acridum inoculum
The unhatched eggs, larvae and dead adults were sterilized with 3 ml 70% ethanol, 3 ml 4% sodium hypochlorite and 3 ml sterile distilled water for 3 min each. The material was then seeded in PDA medium for fungal growth (Alves, 1998). The analysis of fungal growth parameters (conidiogenesis, vegetative growth, colony count and sporulation) were performed in triplicate, and microscopic aspects followed the methodology outlined in Feijó et al. (2007). These data were compared to the parameters observed before the fungal infection.
Metharizium anisopliae var. acridum inoculum was obtained from strain 291 (URM-3800), kindly provided by the Mycology Collection of the Department of Mycology, Universidade Federal de Pernambuco. The fungal cultures were kept in PDA medium (Potato Dextrose Agar, Vertec® ), maintained at 28 ◦ C and diluted to prepare concentrations of 1 × 108 , 1 × 107 , 1 × 106 , 1 × 105 and 1 × 104 conidia/ml in 0.05% (v/v) Tween 80. Conidia were quantified via direct counting using an optical microscope with a Neubauer chamber (Alves and Moraes, 1998).
2.4.1. Conidiogenesis A disc 5 mm in diameter was removed from a M. anisopliae var. acridum culture after 12 days of growth and transferred to a test tube containing 10 ml 0.05% (v/v) Tween 80. The suspension was shaken to separate the conidia, diluted and adjusted to 104 conidia/ml. From this suspension, 0.1 ml was spread-plated on PDA medium using a Drigalski spatula. The number of conidia was determined by counting under a light microscope at 16 h postinoculation. A total of 500 conidia per Petri dish were counted and
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categorized into 2 groups: germinated with having a germ tube in development, and not germinated. 2.4.2. Vegetative growth A disc 5 mm in diameter of M. anisopliae var. acridum was sown in the center of a Petri dish with PDA medium. Growth was measured at day 15. 2.4.3. Colony counting The same dilution of M. anisopliae var. acridum used for the conidiogenesis experiments was also used for colony counting. The dilution was spread on Petri dishes (0.1 ml), and the colonies were counted 3, 6, 9, 12 and 15 days post-inoculation. 2.4.4. Sporulation Using the same methodology used for the vegetative growth experiments, at 3, 6, 9, 12 and 15 days post-inoculation, 10 ml of 70% ethanol was added to 3 Petri dishes with growing fungus for 5 min, for the purpose of conidia inactivation and drying. The 70% ethanol (containing conidia) was retrieved from the surface of the Petri dish and placed into a sterile receptacle. Subsequently, Petri dishes were washed 9 times with 10 ml 0.05% (v/v) Tween 80, and between washes, the Tween 80 solution was removed from the plates and placed in the same container. The conidia were then quantified in a Neubauer chamber. 2.4.5. Microscopic aspects An aliquot of fungal culture was aseptically placed at 4 equidistant points on a Petri dish with PDA medium and covered with a sterile cover slip. These cultures were analyzed with an optical microscope after 24, 48, 72, 96 and 120 h. The fungal structures were stained with Amann blue and observed at 100×, 400× and 1000× magnification. 2.5. Genomic DNA extraction and PCR The genomic DNA of M. anisopliae var. acridum reisolated from infected L. longipalpis eggs, larvae and adults was extracted using an Invisorb® Spin Plant Mini Kit (Invitek, GmbH, Berlin-Buch), then resuspended in 100 l TE and stored at −20 ◦ C, according to the manufacturer’s recommendations. The primers Mac-ITS-spF (5 CTGTCACTGTTGCTTCGGCGGTAC3 ) and Mac-ITS-spR (5 CCCGTTGCGAGTGAGTTACTACTGC3 ) were designed based on the ITS1 and ITS2 regions of the rDNA sequence data for M. anisopliae var. acridum (Entz et al., 2005). PCR amplifications were performed in a total volume of 10 l, containing 1× buffer (20 mM Tris–HCl, pH 8.3, 50 mM KCl), 50 mM MgCl2 , 10 pM/l of each primer, 10 mM dNTP mix (InvitrogenTM ), 5 U/l Platinum Taq DNA polymerase (InvitrogenTM ) and 0.1 l of the target DNA. The negative control contained sterile ultrapure water in place of DNA and the positive control contained M. anisopliae var. acridum (291, URM-3800). The DNA amplification was performed in a Primus 96 HPL thermocycler (MWG Biotech, Inc.) programmed according to the recommendation of Entz et al. (2005): initial denaturation at 94 ◦ C × 5 min, 30 cycles of denaturing at 94 ◦ C × 1 min, annealing and extension combined at 72 ◦ C × 3 min. The amplicons were visualized on a 1% agarose gel containing 1 Kb Plus DNA ladder (InvitrogenTM ), and stained with ethidium bromide and subjected to transillumination with UV light (FB-TI-88).
3100 Genetic Analyzer (Applied Biosystems® ). All fragments were sequenced in both directions and data were processed by programs provided by the sequencer manufacturer. The electrophoretograms generated were stored as files of the Chroma program and the nucleotide sequences were submitted to GenBank. Data were then analyzed using BioEdit software (Hall, 1999) for the purpose of verifying the sequence quality. The Chroma program was used to transform the data output files into the FASTA format (default), giving quality values (0–99) for each nucleotide using algorithms to specify each peak’s intensity (height and width) generated by the electrophoretogram. The sequences were also subjected to BLAST (Basic Local Alignment Search Tool) (Altschul et al., 1990) to verify the similarities of the obtained fragment sequences to the sequences of other proteins in GenBank. 2.7. Statistical analysis The design was completely randomized for all experiments. The effects of fungal infection on egg hatching, larval mortality and adult longevity, as well as the data concerning fungal growth parameters, were normalized when necessary and submitted to ANOVA and Pearson correlation coefficient analyses. After analysis of variance, the means were compared using a Student–Newman–Keuls test, and the growth parameters were compared using Dunnett’s Method. For both, P < 0.05 (SigmaStat software 3.1). 3. Results 3.1. Susceptibility of L. longipalpis to entomopathogenic fungus At the highest concentrations, M. anisopliae var. acridum infection reduced the number of L. longipalpis eggs that hatched by 40% (H = 33.03, df = 6, P < 0.05), although its effect was lower than that of the positive control. There was a direct and significant correlation (r = 0.86) between fungal concentration and the inhibition of egg hatching (F = 22.80, df = 1, P < 0.001). An increase in mortality of the larvae born of these eggs was also observed, and an index of 60% was found at higher concentrations (H = 27.17, df = 6, P < 0.05) (Table 1). The fungus had a greater effect on larval mortality than the control, and the effect was inoculum-dependent reaching 87% (H = 36.04; df = 6, P < 0.05) (Table 1). The mean larval survival time was 26.51 ± 6.75 days for the treated group and 31.89 ± 4.48 for the negative control. However, only the positive control (which caused instantaneous death) had a significant effect on mean larval survival time (H = 29.92, df = 6, P < 0.05). The longevity of adults infected with the fungus was significantly reduced, on average by 7 days, but this effect did not overcome that of cypermethrin, which caused instantaneous death (F = 43.54, df = 6, P < 0.05). Egg hatching was reduced in fungus-infected females, especially at the highest concentration, which produced 85% efficacy (F = 53.57, df = 6, P < 0.05) (Table 1). There was a significant direct correlation (r = 0.49) between fungal concentration and inhibition of egg hatching (F = 13.61, df = 1, P < 0.05). 3.2. Growth and microscopic features of M. anisopliae var. acridum after passage in L. longipalpis
2.6. DNA sequencing The amplicons were purified by isopropanol/ethanol precipitation according to the manufacturer’s recommendations (Applied Biosystems® ) and sequenced using the BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystem® ) in an ABI PRISM®
M. anisopliae var. acridum was successfully reisolated postinfection only from L. longipalpis adults. The conidiogenesis of the reisolated fungus was significantly higher than that of the fungus before infection (F = 4.53, df = 3, P < 0.05), but vegetative growth was similar (F = 2.40, df = 3, P = 0.14) (Table 2).
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Table 1 The effect of M. anisopliae var. acridum on eggs, larvae and adults of L. longipalpis. Treatments (mean ± SD)
Eggs
Larvae
Hatching reduction (%) 1 × 104 conidia/ml 1 × 105 conidia/ml 1 × 106 conidia/ml 1 × 107 conidia/ml 1 × 108 conidia/ml Tween 80 0.05% Cypermethrin 196 g/ml
28.89 30.00 35.56 40.00 41.67 12.78 100.00
± ± ± ± ± ± ±
8.61b 3.85b 2.72a 8.43a 6.24a 9.05c 0.00d
Adults
Larval mortality (%)
Mortality (%)
18.29 ± 27.02 ± 35.11 ± 37.60 ± 49.83 ± 0.00 ± –
63.59 71.90 72.38 83.81 86.67 40.48 100.00
6.51b 2.67b 8.53a 16.54a 24.12a 0.00c
± ± ± ± ± ± ±
Longevity (days)
18.28c 5.39c 5.35c 13.11b 4.71a 7.31d 0.00e
9.57 6.57 6.29 6.86 7.29 8.71 0.00
± ± ± ± ± ± ±
1.51b 1.27a 1.38a 1.77a 0.76a 1.11b 0.00c
Hatching reduction (%) 53.69 ± 64.54 ± 72.30 ± 72.12 ± 84.76 ± 39.26 ± –
10.20c 1.88b 4.97b 3.11b 7.91a 6.68d
Means followed by the same lowercase letter in the same column are not significantly different (Student–Newman–Keuls test, P < 0.05).
Table 2 Conidiogenesis and vegetative growth of Beauveria bassiana reisolated from infected Lutzomyia longipalpis eggs, larvae and adults on PDA medium. Stages (mean ± SD)
Conidiogenesis (no.)* 16 h post-inoculation
Vegetative growth (cm)NS 15 days post-inoculation
Adults Control#
413.00 ± 36.77a 255.00 ± 7.00b
4.07 ± 0.98a 3.07 ± 0.29a
Means followed by the same lowercase letter in the same column are not significantly different (Dunnett’s Method, * (P < 0.05) and NS (P = 0.14). # Metharizium anisopliae var. acridum 291 (URM-3800) before infection of L. longipalpis.
The number of fungus colonies was significantly higher post-passage in adult sand flies (F = 13.05; df = 3, P < 0.05). Sporulation was also significantly higher post-passage, and it was also higher than the fungus before infection and increasing over the entire observation period (F = 158.50; df = 3, P < 0.001) (Table 3). M. anisopliae var. acridum microscopic structures were observed post-infection. Mycelium formation, anastomoses and apressoria were observed in the first 96 h, as were primordia from the conidiophores and young conidiophores along the hyphal axis (data not shown). There were no morphological differences between the fungus used for infection and the fungus reisolated from L. longipalpis adults.
Fig. 1. Amplification of M. anisopliae var. acridum DNA from L. longipalpis adults at different days post-infection with varying conidia/ml concentrations: lane Br: no DNA, lane C+: M. anisopliae var. acridum DNA, lane 1: 1 × 106 at 7 days, lanes 2–4: 1 × 107 at 1, 5 and 6 days, lanes 5–7: 1 × 108 at 1, 6 and 10 days, Kb: ladder.
4. Discussion 3.3. PCR and sequencing The eggs of insects are more resistant to infection than other developmental stages (Tanada and Kaya, 1992). Although fungal infection reduced egg hatching significantly and increased the mortality of larvae born from infected eggs, the effectiveness was around 40%. Superior results were observed when another entomopathogenic fungus, Beauveria bassiana (Bals.) Vuilleman (Deuteromycotina: Hyphomycetes), was tested on eggs of the same sand fly species reaching a 60% efficacy on egg hatching and 88% on the mortality of larvae born from infected eggs (Amóra et al., 2009). Additionally, another biocontrol study showed that aqueous suspensions of Bacillus sphaericus Meyer and Neide (1904) (Bacillales: Bacillaceae) inhibited Phlebotomus duboscqi Leveu-Nemaire (1906) and Sergentomyia schwetzi Adler, Theodor and Parrot (1929) (Diptera: Psychodidae) egg hatching by 95% (Robert et al., 1998).
M. anisopliae was reisolated from L. longipalpis adults infected with 1 × 106 , 1 × 107 and 1 × 108 conidia/ml 6 days post-infection. In all cases, M. anisopliae var. acridum DNA was amplified by PCR (Fig. 1). M. anisopliae var. acridum DNA could not be reisolated from eggs, larvae or adults infected with 1 × 104 and 1 × 105 conidia/ml, but it was possible to observe mycelial growth on insects infected with 1 × 107 and 1 × 108 conidia/ml. No PCR products were amplified from the control group. The DNA sequences obtained were approximately 394 bp in length. These were compared with other sequences deposited in GenBank (NCBI), and this confirmed that the reisolated fungal species was in fact a match to M. anisopliae var. acridum (accession number EF113338.1) with 100% homology (e-value 0.0).
Table 3 Colony counts and sporulation of M. anisopliae var. acridum (mean ± SD) reisolated from infected L. longipalpis adults at 3, 6, 9, 12 and 15 days after inoculation on PDA medium. Parameters
Stages
Colony counting (no. colony)
Adults Control#
3 days 8.50 ± 4.95a 0.00 ± 0.00b
Sporulation (no. conidia)
Adults Control#
12.62 ± 0.54a 4.06 ± 0.08b
6 days
9 days
846.00 ± 124.45a 276.00 ± 69.30b
40.50 ± 0.71a 7.00 ± 1.41b
12 days 62.50 ± 2.12a 0.50 ± 0.71b
15 days 17.00 ± 2.83a 0.00 ± 0.00b
40.11 ± 0.18a 26.27 ± 0.18b
53.38 ± 0.32a 59.16 ± 0.06b
148.48 ± 0.28a 115.19 ± 0.01b
164.66 ± 0.87a 161.40 ± 0.29b
Means followed by the same lowercase letter in the same column are not significantly different (Student–Newman–Keuls test, P < 0.05). # Metharizium anisopliae var. acridum 291 (URM-3800) before infection of L. longipalpis.
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The efficiency of M. anisopliae var. acridum at the concentration of 1 × 108 conidia/ml on larvae of L. longipalpis (87%) and Boophillus microplus Canestrini (1887) (Acari: Ixodidae) (97%) (Bahiense et al., 2006) was similar. But, was smaller than the use of B. bassiana in the same concentration on L. longipalpis larvae obtaining an efficiency of 100% of mortality (Amóra et al., 2009). Comparing the susceptibility of sand fly larvae to different biocontrols, the result of spraying fungus on L. longipalpis larvae was better than offering a diet contaminated with Bacillus thuringiensis var. israelensis Goldberg and Margalit (1977) (Bacillales: Bacillaceae) to Phlebotomus papatasi Scopoli (1763) larvae (Diptera: Psychodidae) (Wahba et al., 1999). Both of these had inoculum-dependent effects. However, the fungus was not reisolated, and similar to P. papatasi larvae infected with B. sphaericus, bacteraemia was not detected (Wahba, 2000). Biolarvicide application in the field is difficult due to the diversity of sand fly breeding habitat, thus its application appears to be limited to adult control. More research needs to be done to develop an efficient larvicide for vector control. Any reduction in mosquito longevity reduces the mean number of blood meals taken, and the probability of the vector acquiring a disease agent and subsequently transmitting it to other hosts is also reduced (Scholte et al., 2007). The longevity of L. longipalpis infected with M. anisopliae var. acridum was significantly lower than that of uninfected sand flies, except for sand flies infected at the concentration of 1 × 104 conidia/ml, whose longevity was greater than the negative control. These data are similar to the effect of other entomopathogenic fungi against P. papatasi, L. longipalpis (Warburg, 1991) and Aedes aegypti Linnaeus (1762) (Scholte et al., 2007) in that they show the highest mortality rates at around seven days. While the effect of M. anisopliae was tested on mites, the longevity of these insects was six days (Kanga et al., 2002). More recently another in vitro study with B. bassiana on adult L. longipalpis reduced the longevity of the insect to 4 days (Amóra et al., 2009). Although there was no significant decrease in infected female oviposition, there was a significant reduction in egg hatching. Similar data were obtained with the same fungus in grasshoppers (Blanford and Thomas, 2001) and B. bassiana in L. longipalpis (Amóra et al., 2009). The low efficiency of M. anisopliae var. acridum compared to other entomopathogenic fungi could be related to its primary specificity for Acridids (Blanford and Thomas, 2001). The reduction in longevity and fecundity may not decrease the vectorial capacity of adult L. longipalpis in the short term, but it could still be effective over the long term. It is possible that infected sand flies are more susceptible to factors such as predation and secondary infections by other pathogens, and infection might also decrease insect migration ability. Traditionally, the identification of the genus Metarhizium is based on the observation of morphological features on culture media and microscopic examination of spores and associated structures (Entz et al., 2005). Fungal growth studies, such as analysis of conidiogenesis, sporulation, colony counts and radial growth were conducted to assist entomopathogenic fungus characterization, as recommended by Almeida et al. (2005), because these parameters are important for defining the virulence of the fungal isolate (Liu et al., 2003). The rates of radial and vegetative growth are directly linked to the speed of infection in the host (Feijó et al., 2007), probably because the radial growth is used as a criterion of selection of mycoinsecticides (Yeo et al., 2003). However, in this study the vegetative growth of M. anisopliae var. acridum was slow, possibly explaining the failure to reisolate it from L. longipalpis eggs and larvae. Similar data were observed by Feijó et al. (2009) and by Amóra et al. (2009), but in the latter case the fungus utilized was B. bassiana although the fungus has been reisolated of all development phases of L. longipalpis. The sporulation, conidiogenesis and colony counts of M. anisopliae var. acridum was significantly higher after reisolation,
corroborating the results of Entz et al. (2005) and Amóra et al. (2009). However the data of sporulation and colony counts of M. anisopliae var. acridum re-isolated from C. albiceps were lower than control (Feijó et al., 2009). These data suggest a positive correlation between conidia production and time. Rapid sporulation may be an important criterion for the selection of fungal isolates because it helps epizootic spread (Mitchell, 2003). Conidiogenesis is another important parameter to consider, as the number of germinated conidia is directly proportional to isolate virulence. The conidiogenesis rate may be influenced by the storage form, the presence of nutrients and the mode of host exposure to fungus (Alves, 1998). Nevertheless, the studies with M. anisopliae var. acridum are still incipient (Feijó et al., 2009). In the analysis of the vegetative and reproductive structures present in M. anisopliae var. acridum before and after passage in adult L. longipalpis was observed the formation of mycelium in the first 48 h, anastomosis at 72 h, appressoria at 96 h and at 120 h abundant ramified conidiophores were also observed. This structures reisolated from adult did not differ from the control, these data are similar to the findings of Feijó et al. (2009). The anastomoses permit the intercommunication between the fungic segments, allowing the passage into cytoplasmatic organelles, including the nuclear migration (Messias and Azevedo, 1980). This phenomenon allows the recombination and genetic improvement in economically important fungi that do not present a sexual cycle (Alves, 1998). The appressoria, in turn, are adhesion sites and production of enzymes assisting the fungus to penetrate the cuticle of the insect target and nutrition (St. Leger et al., 1989). The colony morphology of M. anisopliae var. acridum was examined, producing dark yellow-green conidial masses, conidia typically are ovoid, similar to Brazilian (Feijó et al., 2009), Australian, Korean (Fernandes et al., 2009), Canadian (Entz et al., 2005) and Mexican strains (Milner et al., 2003). However it is important to emphasize that difficulties in identification are frequently encountered, and morphological differences can be exhibited under varying environmental and physiological conditions. The spore morphology varies within the same culture and between isolates of the genus Metarhizium. Conidia and blastospores can be of variable size and shape (Milner et al., 2003). Therefore, M. anisopliae var. acridum cannot be distinguished from other M. anisopliae varieties on the basis of spore size and shape (Fernandes et al., 2009). Thus, molecular tests should be performed (Lomer et al., 2001) that allow differentiation of fungal strains with similar morphological and cytological characteristics (Kouvelis et al., 2008). The failure to reisolate fungus from treated eggs and larvae prevented completion of the PCR, but its effect was observed statistically on L. longipalpis. The same situation was observed in a previous study that involved the histological examination of Lutzomyia young Feliciangeli and Murillo (1987) (Diptera: Psychodidae) infected with B. bassiana, where the histological sections were negative and no cuticle or tissue damage was observed, but the fungal mycelium was seen (Reithinger et al., 1997), similar to our observations. With respect to adult L. longipalpis, the PCR and sequencing proved that infection by the fungus had occurred at the two higher concentrations on different days post-infection. Comparative studies of nucleotide sequences of rDNA genes have provided significant data for phylogenetic and taxonomic analyses (Entz et al., 2005). As expected, the Mac-ITS-spF and Mac-ITS-spR primers successfully amplified a 394 bp DNA sequence from the total genomic DNA extracted from M. anisopliae var. acridum, similar to the results of Entz et al. (2005). In that same study, no other species of Metharizium, other fungal entomopathogens, or other microorganisms were detected by these primers, demonstrating their specificity (Entz et al., 2005).
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Although the effect of an entomopathogenic fungus is not immediate, it can minimize the environmental contamination risk inherent in chemical insecticides (Oliveira et al., 2003). At present, the potential risk of adverse effects of the fungus on human health when used indoors and/or in the vicinity of immunocompromised individuals is considered low due to the pathogen’s opportunistic nature (Scholte et al., 2007). Although some cases have been reported involving exclusively immunocompromised patients (Burgner et al., 1998; Revankar et al., 1999; Osorio et al., 2007) and only one case of feline rhinitis (Muir et al., 1998) and another of human keratitis (Jani et al., 2001) in healthy patient, all caused by M. anisopliae var. anisopliae, with effective treatment and without relapse. Thus, to determine sensitivity to these organisms is necessary to identify specific allergens to provide a basis for mitigation of these allergens causing biocontrol products safer (Westwood et al., 2005). The finding that L. longipalpis is susceptible to infection with an entomopathogenic fungus is the first step towards the final goal of using this fungus for VL vector control. However, there are many more scientific issues to confront, such as the screening of other fungal species and strains, finding the optimal dosages and application techniques. In particular, fungal persistence following application must be substantial to minimize the logistical challenges and cost of re-treatment (Scholte et al., 2007). Several entomopathogenic fungi are already in use for agricultural and veterinary pest control, demonstrating the feasibility of this biocontrol technique with respect to acceptability and applicability. Among the fungi currently used for that purpose stands B. bassiana, which recently demonstrated to be pathogenic on L. longipalpis (Amóra et al., 2009). The results presented here suggest a lack of pathogenicity of M. anisopliae var. acridum against L. longipalpis eggs and larvae, leading to the conclusion that this pathogen should not be used for the control of sand flies in their immature stages. However, in terms of adult mortality and infected female fecundity, the results were significant, proving that the primary vector species of VL is susceptible to entomopathogenic fungal infection in the adult stage. These results encourage further studies on the use of M. anisopliae var. acridum as a biocontrol agent. Acknowledgements We thank Dr. Nélio B. Morais, Richristi A. Silva, Raimundo Nonato de Sousa and Lindemberg Caranha (Secretaria de Saúde do Estado do Ceará), Ana Claudia B. Mendonc¸a and Sodré Rocha (Secretaria Municipal de Saúde de Mossoró) for assistance in sand fly field collections, Dr. Rui Sales Júnior, Dra. Celicina M.S.B. Azevedo (UFERSA). UFERSA administrative staff for logistic support. To Msc Lorena M.B. Oliveira for reviewing and improving the paper. As also the inhabitants of the studied areas for their patient and kindness. Financial support: Msc Amora has a grant from CAPES and Dr. Bevilaqua is a CNPq researcher. Ethical approval: Ceará State University, Committee of Ethics for the Use of Animals (process no. 07465297-4). References Almeida, J.C., Albuquerque, A.C., Luna-Alves Lima, E.A., 2005. Viabilidade de Beauveria bassiana (bals.) Vuill. reisolado de ovos, larvas e adultos de Anthonomus grandis (Boheman) (Coleoptera: Curculionidae) artificialmente infectado. Arq. Inst. Biol. São Paulo 72, 473–480. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403–410. Alves, S.B. (Ed.), 1998. Controle Microbiano de Insetos, 2nd ed. FEALQ, Piracicaba, São Paulo. Alves, S.B., Moraes, S.A., 1998. Quantificac¸ão de inóculo de patógenos de insetos. In: Alves, S.B. (Ed.), Controle Microbiano de Insetos, 2nd ed. FEALQ, Piracicaba, São Paulo, pp. 765–777.
219
Amóra, S.S.A., Santos, M.J.P., Alves, N.D., Gonc¸alves da Costa, S.C., Calabrese, K.S., Monteiro, A.J., Rocha, M.F.G., 2006. Fatores relacionados com a positividade de cães para leishmaniose visceral em área endêmica do Estado do Rio Grande do Norte, Brasil. Cienc. Rural 36, 1854–1859. Amóra, S.S.A., Bevilaqua, C.M.L., Feijó, F.M.C., Silva, M.A., Pereira, R.H.M.A., Silva, S.C., Alves, N.D., Freire, F.A.M., Oliveira, D.M., 2009. Evaluation of the fungus Beauveria bassiana (Deuteromycotina: Hyphomycetes), a potential biological control agent of Lutzomyia longipalpis (Diptera, Psychodidae). Biol. Control 50, 329–335. Aransay, A.M., Scoulica, E., Tselentis, Y., 2000. Detection and identification of Leishmania DNA within naturally infected sand flies by seminested PCR on minicircle kinetoplastic DNA. Appl. Environ. Microbiol. 66, 1933–1938. Bahiense, T.C., Fernandes, E.K.K., Bittencourt, V.R.E.P., 2006. Compatibility of the fungus Metarhizium anisopliae and deltamethrin to control a resistant strain of Boophilus microplus tick. Vet. Parasitol. 141, 319–324. Blanford, S., Thomas, M.B., 2001. Adult survival, maturation, and reproduction of the desert locust Schistocerca gregaria infected with the fungus Metarhizium anisopliae var. acridum. J. Invert. Pathol. 78, 1–8. Burgner, D., Eagles, G., Burgess, M., Procopis, P., Rogers, M., Muir, D., Pritchard, R., Hocking, A., Priest, M., 1998. Disseminated invasive infection due to Metarrhizium anisopliae in an immunocompromised child. J. Clin. Microbiol. 36, 1146–1150. Driver, F., Milner, R.F., Trueman, W.H., 2000. A taxonomic revision of Metarhizium based on a phylogenetic analysis of ribosomal DNA sequence data. Mycol. Res. 104, 134–150. Dye, C., 1996. The logic of Visceral Leishmaniasis control. Am. J. Trop. Med. Hyg. 55, 125–130. Entz, S.C., Johnson, D.L., Kawchuk, L.M., 2005. Development of a PCR-based diagnostic assay for the specific detection of the entomopathogenic fungus Metarhizium anisopliae var. acridum. Mycol. Res. 109, 1302–1312. Faria, M.R., Magalhães, B.P., 2001. O uso de fungos entomopatogênicos no Brasil. Biotecnolog. Cienc. Desenvolv. 22, 18–21. Feijó, F.M.C., Lima, P.M., Alves, N.D., Luna-Alves Lima, E.A., 2007. Comportamento e aspectos citológicos de Beauveria basssiana após passagem em ovo, larva e adulto de Chrysomya albiceps. Arq. Inst. Biol. 74, 349–355. Feijó, F.M.C., Lima, P.M., Melo, E.H.M., Maciel, M.V., Athayde, A.C.R., Alves, N.D., Luna-Alves Lima, E.A., 2008. Susceptibilidade de Chrysomya albiceps a Beauveria bassiana em condic¸ões de laboratório. Cienc. Anim. Bras. 9, 771–777. Feijó, F.M.C., Lima, P.M., Melo, E.H.M., Athayde, A.C.R., Luna-Alves Lima, E.A., 2009. Behavior and cytological aspects of Metarhizium anisopliae and Metarhizium flavoviride after passage in Chrysomya albiceps. Rev. Caatinga 22, 39–44. Fernandes, É.K.K., Keyser, C.A., Chong, J.P., Rangel, D.E.N., Miller, M.P., Roberts, D.W., 2009. Characterization of Metarhizium species and varieties based on molecular analysis, heat tolerance and cold activity. J. Appl. Microbiol. Epub ahead of print. doi:10.1111/j.1365-2672.2009.04422.x. Galati, E.A.B., 2003. Morfologia e Taxonomia. Morfologia, terminologia de adultos e identificac¸ão dos táxons da América. In: Rangel, E.F., Lainson, R. (Eds.), Flebotomíneos do Brasil. Fiocruz, Rio de Janeiro, pp. 53–175. Goettel, M.S., Jonshon, D.L., Inglis, G.D., 1995. The role of fungi in the control of grasshoppers. Can. J. Bot. 73, S71–S75. Hall, T., 1999. BioEdit Version 5.0.6. North Carolina State University, Department of Microbiology. Jani, B.R., Rinaldi, M.G., Reinhart, W.J., 2001. An unusual case of fungal keratitis: Metarrhizium anisopliae. Cornea 20, 765–768. Kanga, L.H.B., James, R.R., Boucias, D.G., 2002. Hirsutella thompsonii and Metarhizium anisopliae as potential microbial control agents of Varroa destructor, a honey bee parasite. J. Invert. Pathol. 81, 175–184. Kouvelis, V.N., Ghikas, D.V., Edgington, S., Typas, M.A., Moore, D., 2008. Molecular characterization of isolates of Beauveria bassiana obtained from overwintering and summer populations of Sunn Pest (Eurygaster integriceps). Lett. Appl. Microbiol. 46, 414–420. Liu, H., Skinner, M., Brownbrigde, M., Parker, B.L., 2003. Characterization of Beauveria bassiana and Metarhizium anisopliae isolates for management of tarnished plant bug, Lygus lineolaris (Himeptera, Miridae). J. Invert. Patholol. 82, 139–147. Lomer, C.J., Bateman, R.P., Johnson, D.L., Langewald, J., Thomas, M., 2001. Biological control of locusts and grasshoppers. Ann. Rev. Entomol. 46, 667–702. Magalhães, B.P., De Faria, M.R., Lecoq, M., Schmidt, F.G.V., Silva, J.B.T., Frazão, H.S., Balanc¸a, G., Foucart, A., 2001. The use of Metarhizium anisopliae var. acridum against the grasshopper Rhammatocerus schistocercoides in Brazil. J. Orthoptera Res. 10, 199–202. Messias, C.L., Azevedo, J.L., 1980. Parassexuality in the deuteromycete Metarhizium anisopliae. Trans. Br. Mycol. Soc. 75, 473–477. Milner, R.J., Lozano, L.B., Driver, F., Hunter, D., 2003. A comparative study of two Mexican isolates with an Australian isolate of Metarhizium anisopliae var. acridum—strain characterisation, temperature profile and virulence for wingless grasshopper, Phaulacridium vittatum. BioControl 48, 335–348. Ministério da Saúde. Secretaria de Vigilância em Saúde. Departamento de Vigilância Epidemiológica, 2006. Manual de vigilância e controle da leishmaniose visceral. Brasília: Ministério da Saúde, 122 pp. [cited on August 30, 2008]. Avaliable from: http://portal.saude.gov.br/portal/arquivos/pdf/manual leish visceral2006.pdf. Mitchell, J.K., 2003. Development of a submerged liquid sporulation medium for the potential smart weed bioherbicide Septaria polyganorum. Biol. Control 27, 293–299. Muir, D., Martin, P., Kendall, K., Malik, R., 1998. Invasive hyphomycotic rhinitis in a cat due to Metarhizium anisopliae. Med. Mycol. 36, 51–54.
220
S.S.A. Amóra et al. / Acta Tropica 113 (2010) 214–220
Oliveira, C.N., Neves, P.M.O.J., Kawazoe, L.S., 2003. Compatibility between the entomopathogenic fungus Beauveria bassiana and insecticides used in coffee plantations. Sci. Agric. 60, 663–667. ˜ F., Rodriguez-Tudela, Osorio, S., de la Cámara, R., Monteserin, M.C., Granados, R., Ona, J.L., Cuenca-Estrella, M., 2007. Recurrent disseminated skin lesions due to Metarrhizium anisopliae in an adult patient with acute myelogenous leukemia. J. Clin. Microbiol. 45, 651–655. Rangel, E.F., Souza, N.A., Wermelinger, E.D., Barbosa, A.F., 1985. Estabelecimento de colônia, em laboratório, de Lutzomyia intermedia Lutz & Neiva, 1912 (Diptera, Psychodidae, Phlebotominae). Mem. Inst. Oswaldo Cruz 80, 219–226. Reithinger, R., Davies, C.R., Cadena, H., Alexander, B., 1997. Evaluation of the fungus Beauveria bassiana as a potential biological control agent against phlebotomine sand flies in colombian coffee plantations. J. Invertebr. Pathol. 70, 131–135. Revankar, S.G., Sutton, D.A., Sanche, S.E., Rao, J., Zervos, M., Dashti, F., Rinaldi, M.G., 1999. Metarrhizium anisopliae as a cause of sinusitis in immunocompetent hosts. J. Clin. Microbiol. 37, 195–198. Robert, L.L., Perich, M.J., Schlein, Y., Jacobson, R.L., 1998. Bacillus sphaericus inhibits hatching of phlebotomine sand fly eggs. J. Am. Mosq. Control. Assoc. 14, 351–352. Scholte, E.-J., Knols, B.G.J., Samson, R.A., Takken, W., 2004. Entomopathogenic fungi for mosquito control: a review. J. Insect Sci. 4, 1–24. Scholte, E.-J., Takken, W., Knols, B.G.J., 2007. Infection of adult Aedes aegypti and Ae. albopictus mosquitoes with the entomopathogenic fungus Metarhizium anisopliae. Acta Trop. 102, 151–158. Secundino, N.F.C., Araújo, M.S.S., Oliveira, G.H.B., Massara, C.L., Carvalho, O.S., Lanfredi, R.M., Pimenta, P.F.P., 2002. Preliminary description of a new ento-
moparasitic nematode infecting Lutzomyia longipalpis sand fly, the vector of Visceral Leishmaniasis in the New World. J. Invert. Pathol. 80, 35–40. Sharma, U., Singh, S., 2008. Insect vectors of Leishmania: distribution, physiology and their control. J. Vector Borne Dis. 45, 255–272. Singh, S., 2006. New developments in diagnosis of leishmaniasis. Indian J. Med. Res. 123, 311–330. St. Leger, R.J.S., Butt, T.M., Goettel, M.S., Staples, R.S., Roberts, D.W., 1989. Production in vitro of appressoria by the entomopathogenic fungus Metarhizium anisopliae. Exp. Mycol. 13, 274–288. Tanada, Y., Kaya, H.K., 1992. Insect Pathology. Elsevier Science & Technology Books, Hardcover. Wahba, M.M., 2000. The influence of Bacillus sphaericus on the biology and histology of Phlebotomus papatasi. J. Egypt. Soc. Parasitol. 30, 315–323. Wahba, M.M., Labib, I.M., el Hamshary, E.M., 1999. Bacillus thuringiensis var. israelensis as a microbial control agent against adult and immature stages of the sand fly. Phlebotomus papatasi under laboratory conditions. J. Egypt. Soc. Parasitol. 29, 587–597. Warburg, A., 1991. Entomopathogens of phlebotomine sand flies, laboratory experiments and natural infections. J. Invertebr. Pathol. 58, 189–202. Westwood, G.S., Huang, S.W., Keyhani, N.O., 2005. Allergens of the entomopathogenic fungus Beauveria bassiana. Clin. Mol. Allergy 3, 1–8. Yeo, H., Pell, J.K., Alderson, P.G., Clark, S.J., Pye, B.J., 2003. Laboratory evaluation of temperature effects on the germination and growth of entomopathogenic fungi and on their pathogenicity to two aphid species. Pest Manage. Sci. 59, 156–165.