Background

The family Tabanidae, which belongs to the order Diptera, comprises approximately 4400 species [1]. These insects are of great medical and veterinary interest due to their persistent and irritating bites and their ability to transmit 32 pathogens, including viruses, bacteria, helminths, and protozoa, such as Trypanosoma spp. [1].

Trypanosoma evansi (Steel, 1885) is the etiological agent of surra, a disease that affects a variety of domestic and wild mammals. Trypanosoma evansi is distributed in Africa, Asia, and South America and causes important economic losses to livestock farming. Surra is of great concern because of the absence of pathognomonic signs, including fever, anemia, loss of body weight, low production of animal protein, nervous signs, abortion, cachexia, and death, with or without more particular signs related to the host species [2]. Trypanosoma evansi, which is mechanically transmitted by tabanids and is widespread in regions of Brazil and the rest of South America, causes outbreaks of surra in horses [3].

The focus of most of the published literature on tabanid species from Brazil and the rest of South America is their taxonomy [4,5,6,7], ecology [8, 9] and behavior [10, 11] in different biomes, and molecular tools were not used for identification or bioprospecting in these studies. More specifically, few studies have addressed tabanids in southern Brazil [12,13,14,15,16,17], and only one reported the molecular detection of a strain of a trypanosomatid, Trypanosoma kaiowa, which is associated with the crocodilian clade of Trypanosoma [18]. Cases of trypanosomosis caused by T. evansi [19] in that region have been described in which the involvement of tabanids was assumed. There are no data on the detection of T. evansi in tabanid flies in South America. This is, to the best of our knowledge, the first study to describe the molecular detection of T. evansi DNA in Dichelacera alcicornis (Wiedemann, 1828) and Dichelacera januarii (Wiedemann, 1819), both of which are widely distributed in various biomes of South America [20] and may be related to outbreaks of surra.

Methods

Study site

This study was carried out in the municipality of Lages (27°48′57″S, 50°19′33″ W), Santa Catarina State, Brazil, which has a mixed rainforest terrain and altitude of 930 m above mean sea level.

The Köppen climate classification of the municipality is Cfb, i.e. it has a temperate oceanic climate. The average temperature in the coldest month is above 0 °C, the average annual temperature is below 22 °C, and at least 4 months have an average temperature above 10 °C, with no significant difference in precipitation between the seasons [21].

Sample collection

Tabanids were collected weekly from February 2018 to February 2019 from two rural properties located in different parts of the region in which especially cattle and horses are reared.

The collections were made once a week at the same location on each property, close to lakes and rivers, for 3 h, from 3 p.m. until 6 p.m., using a contained horse from each property as bait throughout the collection period. Tabanids that landed on the horses were carefully captured with a glass tube (4.5 cm diameter, 9 cm length) [14]. The flies were placed in individual plastic bottles using a SECTAB device [22]. The collected insects were transported to the Laboratory of Hemoparasites and Vectors (Lages, Santa Catarina, Brazil) and killed with chloroform in flasks.

Taxonomic identification of tabanids

Taxonomic identification was performed according to the taxonomic keys described by Fairchild and Philip [23], and Dr. Inocêncio de Sousa Gorayeb (Museu Paraense Emílio Goeldi, Belém, Brazil) verified the identification of the species. The collected specimens were deposited at the Entomology Museum of the Federal University of Fronteira Sul, Laranjeiras do Sul campus, Paraná, Brazil.

DNA extraction

The extraction of DNA from the mouthparts was initially carried out for the three most abundant species of tabanid. After these analyses, DNA was also extracted from the other species of Dichelacera captured, D. januarii.

The flies were washed twice with 70% ethanol solution and twice with sterile distilled water. The mouthparts were removed with the aid of sterile fine scissors under an entomological magnifying glass. The mouthparts were stored in 1.5-mL microtubes in Tris–NaCl–ethylenediaminetetraacetic acid (ETDA) buffer (10 mM Tris base, 200 mM NaCl and 50 mM EDTA), and stored frozen at -80 °C until DNA extraction.

For DNA extraction, the mouthparts were placed in a 1.5-mL microtube and macerated with an appropriate sized pistil. The DNA sample was extracted once with phenol (pH 7.8), once with phenol:chloroform (1:1), and once with chloroform:isoamyl alcohol (24:1). The DNA of the sample was then precipitated using sodium acetate (pH 6.0) and absolute ethanol and resuspended in 50 µL Tris–EDTA buffer (10 mM Tris–HCl, 1 mM EDTA) [24].

Polymerase chain reaction amplification

Flies were individually screened for the presence of Trypanosoma evansi and Trypanosoma vivax using specific oligonucleotide primers, RoTat 1.2 forward 5′GCGGGGTGTTTAAAGCAATA3′ and RoTat 1.2 reverse 5′ATTAGTGCTGCGTGTGTTCG3′ for T. evansi [24], and TviSL1 5′GCTCTCCAATCTTAACCCTA3′ and TviSL2 5′GTTCCAGGCGTGCAAACGTC3′ for T. vivax [25]. The first two primers amplified a 205-base pair (bp) fragment and the second two a 210-bp fragment. Polymerase chain reaction (PCR) was conducted in a 400-µL reaction mixture comprising 323.5 µL deionized water, 10.5 µL MgCl2, 1.75 µL Taq DNA polymerase, 7 µL deoxyribonucleotide triphosphate mix (10 mM), 35 µL of 10× Taq DNA polymerase buffer, 10.5 µL of each forward primer, 7 µL of each reverse primer, and 5 µL of the template. Trypanosoma evansi DNA was obtained from parasites purified from the blood of experimentally infected albino rats. Trypanosoma vivax DNA was obtained from purified parasites of experimentally infected sheep (approved by the Animal Experimentation Ethics Committee of Universidade do Estado de Santa Catarina) and was used as a positive control. Nuclease-free water was added to the PCR mix instead of a DNA sample as a negative control.

PCR was performed using an automated DNA thermal cycler (Biocycler). The amplification conditions were: initial denaturation at 94 °C for 3 min followed by 35 denaturation cycles at 94 °C for 30 s, annealing at 62 °C for 30 s, primer extension at 72 °C for 1 min, and a final extension at 72 °C for 4 min. The final phase of the PCR included cooling the samples to 10 °C. The PCR products were visualized on a 1% agarose gel stained with ethidium bromide.

Sequencing and Basic Local Alignment Search Tool

The PCR amplicons were purified using the QIAGEN Gel Purification Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s protocol. Sequencing was conducted using the BigDye Terminator Cycle Sequencing Kit according to the manufacturer’s protocol (Applied Biosystems, Carlsbad, CA). The eluent was loaded into a 96-well plate which was placed into an ABI Prism 3500 Genetic Analyzer (Applied Biosystems).

Each DNA sample was purified according to the following protocol: 50 µL sample DNA was added to a mixture containing 5 µL of 3 M sodium acetate, 125 µL of 100% ethanol, and 2 µL glycogen (20 mg/ml) and placed in a freezer at − 80 °C for 1 h. Following centrifugation at 12,000 g for 45 min at 4 °C, the pellet formed was washed once with 75% ethanol and centrifuged for another 15 min at 75,000 g at 4 °C. The mixture was then dried in a SpeedVac at 20–25 °C for 30 min and resuspended in 20 µL ultrapure water (Milli-Q).

The retrieved gene sequences were edited using BioEdit software [26]. The nucleotide Basic Local Alignment Search Tool (BLASTn) was used (www.ncbi.nlm.nih.gov/blast/) to confirm the sequences obtained from the PCR analysis. Gene sequences with match scores of 80–100% similarity were considered significant.

Results

A total of 523 female tabanids were collected from February 2018 to February 2019, specifically in February, March, April, November and December 2018, and January and February 2019. There was no evidence of tabanids in the other months of the collection period.

Individuals of 14 species and seven genera were collected (Table 1). Dichelacera alcicornis was the most abundant species, representing 52.77% (276) of the total, followed by Chrysops fusciapex (Lutz, 1909a) (17.97%, 94) and Chrysops patricia (Pechuman, 1953) (10.70%, 56), similar to the results of previous studies [11, 14]. The least abundant species were Tabanus fuscus (Wiedemann, 1819), Tabanus nebulosus (De Geer, 1776) and Acanthocera kroeberi (Fairchild, 1939), and represented 0.19% of the total.

Table 1 Tabanid species and numbers of individuals caught from February 2018 to February 2019 in Lages, Santa Catarina, Brazil
Full size table

Samples of the three most abundant species, D. alcicornis (Additional file 1: Fig. 1), C. patricia and C. fusciapex, were analyzed. Samples of D. januarii (Additional file 2: Fig. 2) were also examined so that both species of Dichelacera that had been captured were included in the analyses. The PCR amplicons were positive for T. evansi using the 205-bp RoTat 1.2 gene. Sequencing and analysis of the amplicons demonstrated that the sequence obtained corresponded to the T. evansi variable surface glycoprotein (accession no. MZ209177.1). It was present in the feeding parts of D. alcicornis (Da1) with 96% identity at site 1 and in D. alcicornis and D. januarii (Da 2) with 99% identity at site 2 (Fig. 1). There was no amplification of the T. vivax splice leader gene for any of the samples. The sequences were deposited in GenBank (respective accession numbers OM971942 and OM971943).

Fig. 1
figure 1

Multiple sequence alignment between the polymerase chain reaction amplified sequences of DNA from the feeding apparatus of Dichelacera alcicornis (Da1) and Dichelacera januarii (Da2) and the variant surface glycoprotein sequence of T. evansi deposited in GenBank

Full size image

For C. patricia and C. fusciapex there was no amplification of the T. evansi RoTat 1.2 gene or T. vivax TviSL gene. The presence of DNA of other trypanosomatids was not evaluated due to the focal objective of the work.

Discussion

The few studies that exist on the seasonality of the Tabanidae in southern Brazil show D. alcicornis to be one of the most abundant species of this family in the region [6, 9, 10], as also found in the present study. The circulation of T. evansi in southern Brazil has already been demonstrated in several studies [14, 27,28,29], but this is the first one, to the best of our knowledge, in which the DNA of this parasite has been detected in the mouthparts of members of the Tabanidae. The sequenced PCR products showed high identity with the deposited sequences in GenBank, which is considered evidence of the presence of T. evansi DNA in the mouthparts of the tabanids examined here.

Two recent studies carried out in Brazil (in the Pantanal region and on the Coastal Plain of the state of Rio Grande do Sul) identified Trypanosoma kaiowa in tabanids using an insect dissection method and the observation of internal organs under an optical microscope [30] and through molecular detection [18]. Protozoan parasites have been detected in tabanids from other continents, such as Africa (particularly South Africa and Zambia), and included Trypanosoma congolense, Trypanosoma theileri, and Trypanosoma evansi [31]. In Europe (Poland), four species of tabanids (Haematopota pluvialis, Tabanus bromius, Tabanus maculicornis, and Tabanus distinguendus) were found to be infected by trypanosomatids of the subgenus Megatrypanum, of which Trypanosoma theileri is a member, and the occurrence of a trypanosomatid in T. maculicornis and T. distinguendus was described in that study for the first time [32]. Chrysops laetus and Dichelacera tetradelta infected with T. theileri were recently described for northern Brazil [33]. Our data, are, to the best of our knowledge, the first molecular confirmation of the presence of T. evansi DNA in the feeding apparatus of D. alcicornis and D. januarii, and support previous findings.

Although reservoirs play a central role in the maintenance and expansion of T. evansi infections, transmission depends on several ecological and epidemiological characteristics, such as the presence of competent vector species and susceptible mammalian hosts. To the best of our knowledge, there have been no studies on the vectorial capacity of the genera and species examined in the present study. However, a study has demonstrated the transmission of T. evansi by species of the genus Stomoxys under laboratory conditions [34]. A mathematical model has been developed that demonstrates the conditions necessary for the successful mechanical transmission of pathogens by tabanids [35]. Thus, the development and use of molecular detection approaches could help improve the identification of disease-causing agents and their tabanid vectors, in addition to facilitating the mapping of the circulation of these agents.

Conclusion

This is, to the best of our knowledge, the first report of T. evansi DNA in South American tabanids.