INTRODUCTION
Black flies (Diptera: Simuliidae) cause severe medical and veterinary problems worldwide. Simuliidae species are able to transmit parasites that can result in severe disease in humans and animals. In addition, their bites can cause allergic reactions and dermatitis in sensitized individuals, resulting in severe economic losses to tourism centers and negatively impacting animal production1–3. Black fly control remains a major public health challenge. Microsporidia are unicellular, eukaryotic organisms that are obligate, intracellular parasites with public health relevance4. Several studies have suggested a new classification for microsporidia as fungi, but Ebersberger5 stated that phylogenetic analysis did not support fungal characterization for this group.
Microsporidia are the most common black fly pathogens, although the species’ diversity, seasonal occurrence and transmission mechanisms remain poorly understood6,7. Infections caused by this agent are often chronic and non-lethal, but they can cause sub-lethal host effects, such as reduced fecundity, decreased life span and general loss of vigor8.
The objective of this study was to identify microsporidian species infecting Simulium (Chirostilbia) pertinax(Kollar, 1832) larvae from Caraguatatuba City, on the north coast of State of São Paulo, by molecular and morphological characterization.
The city’s economy greatly depends on tourism. Thus, the Simuliidae population plays an important role because black fly bites annoy visitors and have deleterious effects on the local economy. Monitoring and controlling black flies are essential to avoiding seasonal population outbreaks.
METHODS
Sampling and biological material processing
The sampling period was from May to August 2013, and the samples were collected from a stream in Caraguatatuba City, located on the north coast of the State of São Paulo, Brazil, which has a total area of 458,097km2 and had a population at that time of 100,8409. All of the larvae were held in aerated containers with water from the breeding site until examination. Tissues showing evidence of infection (whitish abdomens or whitish digestive tracts) were dissected in NaCl 0.9% solution, and fat bodies and adjacent tissues were removed10. Processed samples were frozen in 1.5ml tubes with 30µl of diethylpyrocarbonate (DEPC) (Invitrogen® Life Technologies, Carlsbad, CA, USA). Fresh smears of fat bodies were made, fixed with methanol for 5min and stained with 10% Giemsa in 7.4 pH buffer for 20min. The slides were washed in water and dried at 25°C overnight11 for further morphological analysis of spores.
Morphological analysis
The Nis Elements F 3.0 NIKON H550S software, with phase III objective scale 100X settings, was used for spore measurement. Morphological characterization was performed according to Sprague12.
Molecular assay
Molecular assays were performed with frozen tissues from infected larvae, and Aedes aegypti larvae infected with Edhazardia aedis were used as positive controls.
DNA extraction
Larvae exhibiting symptoms of infection had deoxyribonucleic acid (DNA) extracted using a viral DNA kit (QIAamp® viral RNA, Qiagen, Inc, Hilden, Germany). Healthy larvae (Figure 1A) were discarded. Tissue samples were processed with a proteinase K kit, incubated at 56°C for 2h and mixed every 20min. The supernatants were used to amplify the r16S ribosomal gene13.
Small subunit ribosomal gene (SSUrDNA) PCR (r16S)
Polymerase chain reaction (PCR) amplification was performed with 18f (CAC CAG GTT GAT TCT GCC) and 1492r (GGT TAC CTT GTT ACG ACT T), according to Vossbrinck et al.14.
The amplification products were visualized on 2% agarose gels, with positive and negative controls and a 100 bps ladder (Invitrogen® Life Technologies, Carlsbad, CA, USA), following electrophoresis.
Nucleotide sequencing
PCR products were purified with the Illustra GFX PCR DNA and Gel Band Purification Kit (GE Healthcare Limited, Little Chalfont, Buckinghamshire, UK) and were quantified with 2% agarose gel ethidium bromide staining, according to the Low DNA Mass Ladder (Invitrogen®) protocol. The products were sequenced using an ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction kit (PE Applied Biosystems), following the standard manufacturer protocols. The data were analyzed with the phred/phrap software, and the contigs were assembled with the cap3 software15.
Phylogenetic analysis
The analyses were performed using the Seaview software16. A phylogenetic tree was constructed, with reference sequences32–46 from Table 1(supplementary file), using the maximum likelihood method with the general time reversible (GTR) model of nucleotide substitution and gamma distribution (G) (GTR + G)17. The model was selected by the Modeltest software, version 3.0.618, and was optimized by the Seaview software. We calculated the bootstrap values with 1,000 replications to support the verification of branches in the topologies of the trees obtained, and bootstrap values greater than 70 were considered significant.
TABLE 1- Sequences and accession numbers used for phylogenetic analysis.
| Organism | Host | Geographic locale | Accession number |
|---|---|---|---|
| Amblyospora bracteata | Odagamia ornata | Czech Republic | AY09006832 |
| Antonospora scoticae | Andrena scotica | USA | AF024655* |
| Paranosema grylli | Gryllus bimaculatus | St. Petersburg, FL, USA | AY30532533 |
| Polydispyrenia simulii | Odagamia ornata | Czech Republic | AY09006932 |
| Weiseria palustris | Cnephia ornithophilia | USA | AF132544* |
| Nosema algerae | Anopheles stephensi | Illinois, USA | AF06906334 |
| Thelohania solenopsae | Solenopsis invicta | USA | AF03153828 |
| Janacekia debaisieuxi | Odagamia ornata | USA | AY09007035 |
| Hamiltosporidium magnivora | Daphnia magna | Russia | AJ302318.1* |
| Ichthyosporidium sp. | Leiostomus xanthurus | Not Informed | L3911031 |
| Glugea anomala | Gasterosteus aculeatus | Norway | AF044391.136 |
| Vavraia oncoperae | Wiseana spp. | New Zealand | X7411237 |
| Vavraia culicis | Aedes albopictus | USA | AJ25296129 |
| Endoreticulatus schubergi | Lymantria dispar | Switzerland | L3910931 |
| Vittaforma corneum | Homo sapiens | USA | L3911231 |
| Nucleospora salmonis | Oncorhynchus tshawytscha | Canada | U7817638 |
| Enterocytozoon bieneusi | Homo sapiens | USA | AF02465739 |
| Encephalitozoon cuniculi | Oryctolagus cuniculus | USA | Z19563.140 |
| Encephalitozoon intestinalis | Homo sapiens | USA | U0992941 |
| Encephalitozoon hellem | Homo sapiens | USA | L1907042 |
| Nosema bombycis | Bombyx mori | Switzerland | L3911131 |
| Vairimorpha necatrix | Malacosoma americanum | Not Informed | Y002664 |
| Nosema vespula | Species Unknown | USA | U11047* |
| Nosema apis | Apis mellifera | New Zealand | U97150.143 |
| Amblyospora ferocious | Psorophora ferox | Argentina | AY09006232 |
| Amblyospora criniferis | Aedes cernifera | Argentina | AY09006132 |
| Amblyospora stimuli | Diacyclops bicuspidatus | USA | AY09005032 |
| Amblyospora canadensis | Ochlerotatus canadensis | USA | AY09005632 |
| Amblyospora cinerei | Aedes cinereus | USA | AY09005732 |
| Amblyospora cinerei | Acanthacyclops vernalis | USA | AY09005932 |
| Amblyospora cinerei | Acanthacyclops vernalis | USA | AY09005832 |
| Amblyospora cinerei | Cyclops venustoides | USA | AY09006032 |
| Amblyospora connecticus | Ochlerotatus cantator | USA | AF025685* |
| Amblyospora excrucii | Ochlerotatus excrucians | USA | AY09004332 |
| Amblyospora stimuli | Aedes stimulans | USA | AF02768527 |
| Amblyospora excrucii | Acanthocyclops vernalis | USA | AY09004432 |
| Amblyospora khaliulini | Ochlerotatus communis | USA | AY09004532 |
| Amblyospora khaliulini | Acanthocyclops vernalis | USA | AY09004632 |
| Amblyospora khaliulini | Acanthocyclops vernalis | USA | AY09004732 |
| Amblyospora weiseri | Ochlerotatus cantans | USA | AY09004832 |
| Amblyospora stictici | Ochlerotatus sticticus | USA | AY09004932 |
| Edhazardia aedis | Aedes aegypti | Thailand | AF02768427 |
| Amblyospora sp. | Cyclops strenuus | Czech Republic | AY09005532 |
| Amblyospora californica | Culex tarsulis | USA | U6847344 |
| Amblyospora sp. | Culex nigripalpus | USA | AY09005332 |
| Amblyospora sp. | Culex salinarius | USA | U6847444 |
| Amblyospora salinaria | Culex salinarius | USA | AY32627032 |
| Culicospora magna | Culex restuans | USA | AY09005432 |
| Culicospora magna | Culex restuans | USA | AY32626932 |
| Intrapredatorus barri | Culex fuscanus | Norway | AY01335945 |
| Amblyospora indicola | Culex sitiens | India | AY09005132 |
| Amblyospora opacita | Culex territans | USA | AY09005232 |
| Hyalinocysta chapmani | Culiseta melanura | USA | AF48383746 |
| Hyalinocysta chapmani | Orthocyclops modestus | USA | AF48383846 |
| Culicosporella lunata | Culex pilosus | USA | AF02768327 |
| Parathelohania anophelis | Anopheles quadrimaculatus | USA | AF02768227 |
| Parathelohania obesa | Anopheles crucians | USA | AY09006532 |
| Trichotuzetia guttata | Cyclops vicinus | Czech Republic | AY32626832 |
| Hazardia milleri | Culex quinquefasciatus | Argentina | AY09006732 |
| Hazardia sp. | Anopheles crucians | USA | AY09006632 |
| Marsoniella elegans | Cyclops vicinus | Czech Republic | AY09004132 |
| Gurleya vavrai | Daphnia longispina | Finland | AF39452630 |
| Gurleya daphniae | Daphnia pulex | Austria | AF43932030 |
| Larssonia obtusa | Daphnia pulex | Sweden | AF39452730 |
| Berwaldia schaefernai | Daphnia galeata | Czech Republic | AY09004232 |
| Varimorpha sp. | Solenopsis richteri | USA | AF03153928 |
| Amblyospora sp. | Simulium sp. | UK | AJ25294929 |
USA: United States of America; FL:Florida; UK: United Kingdom.
RESULTS
A total of 1,574 S. pertinax larvae were examined. Eight larvae exhibited symptoms of microsporidian infection localized to the fat body (Figure 1B).
Morphological characterization indicated Polydispyreniaspp. infections in 7 larvae (Figure 2A), representing 87.5% of the infected larvae. Amblyospora sp. infection was observed in one larva (12.5% of the infected larvae) (Figure 2B). The prevalence of microsporidia parasitizing larvae of S. pertinax was 0.51%.
FIGURE 2- Phase-contrast microscopy of smear slides of Simulium pertinax infected by microsporidia. Sporophorous vesicle of Polydispyrenia sp. containing 32 mononuclear spores (A). Octospores of Amblyospora spp. containing 8 uninucleate spores each (B).
Polydispyrenia spp. infections were characterized by the presence of at least 32 mononuclear spores contained within a persistent sporophorous vesicle, with the spores measuring 6.9 ± 1.0 × 5.0 ± 0.7µm (n = 23). Similarly, Amblyospora spp. were characterized by the presence of eight uninucleate spores contained within a sporophorous vesicle, with the spores measuring 4.5 × 3.5µm (n = 12).
The PCR products targeting the 16S region and electrophoresis agarose gel analysis confirmed the presence of microsporidian DNA in 8 samples.
Six samples (Brazilian larvae) were found to be related to, but in a separate cluster (Figure 3) than, the Polydispyrenia simulii [GenBank: AY090069] and Caudospora palustris [GenBank: AF132544] reference sequences (with 100% bootstrapping). One sample (L2) was clustered with Amblyospora spp. [GenBank: AJ252949] with 100% bootstrapping. The Edhazardia aedis positive control (CONT+) taxon was clustered with Edhazardia aedis [GenBank: AF027684] with 100% bootstrapping.
FIGURE 3- Phylogenetic tree generated for microsporidia. Unrooted tree constructed with the maximum likelihood method using the general time reversible model of nucleotide substitution and gamma distribution (GTR + G), using Seaview software. The robustness of the phylogenetic groups was evaluated using 1,000 bootstrap replicates, and bootstrap values greater than 70 were considered significant.
DISCUSSION
Herein, we reported microsporidia parasitizing S. pertinax larvae in the State of São Paulo, with a prevalence of 0.51%. Araújo-Coutinho6 previously reported a 0.5-2.0% prevalence of microsporidia in S. pertinax in State of Rio de Janeiro. Our study showed a similar prevalence to that previously reported by Crosskey19 in other populations of black flies, with rates of up to 1%. Polydispyrenia spp. were the most prevalent parasitic species in S. pertinax from Caraguatatuba/SP in this study, while Amblyospora spp. showed a higher prevalence in Rio de Janeiro6. This difference could be explained by the small sample size, which prevented further analysis of the species population dynamics between S. pertinax from Rio de Janeiro and Caraguatatuba.
In this study, spores of the Polydispyrenia spp. measured 6.9 ± 1.0µm in length × 5.0 ± 0.7µm in width. Araújo-Coutinho6reported spores of a similar size for a Polydispyrenia sp. from S. pertinax that was ovocylindrical and measured 7.0 ± 0.6 × 4.9 ± 0.8µm. However, Castello-Branco and Andrade20 reported larger-sized spores measuring 8.3µm in length × 6.3µm in width for P. simulii from S. pertinax collected in State of São Paulo, Brazil. Sprague12 stated that the spore dimensions were 4.5 to 5.5µm × 2.5 to 3.5µm for P. simulii with the hosts listed as S. pertinax and S. perflavum from Brazil.
In this study, for Amblyospora spp. from Caraguatatuba, the spore measurement was 4.5µm in length × 3.5µm in width, similar to that found by Araújo-Coutinho6 for Amblyospora spp. infecting S. pertinax in the State of Rio de Janeiro. Both of these results were similar to those from Amblyospora bracteata and Amblyospora varians, described in black flies in North America and Europe21. According to Sprague12, the morphological similarity between species of microsporidia, particularly the spore measurements, makes identification difficult, and other methods are needed for identification. Such evidence indicates that spore dimension diversity is too variable; thus, molecular analysis could help in species identification.
Our sample, identified morphologically as Polydispyreniaspp., was grouped with the P. simulii and C. palustris clusters. This identification corroborated previous results22–26 regarding the phylogeny of these parasites.
The genera Parathelohania, Hazardia, Marsoniella, Gurleya, Larssonia, Berwaldia, Varimorpha, Amblyospora and the Amblyospora sp. from S. pertinax in this study form a separate group from the main Amblyospora cluster (Figure 3). Excluding the Varimorpha sp., which was characterized in an ant species, Solenopsis richteri (Forel, 1909), all genera in this group are parasites of aquatics hosts27–30.
Because the Amblyospora group is divided into two clades, corresponding to the hosts (Culex orAedes/Ochlerotatus)28, the aquatic group also demonstrated distinct phylogenetic characteristics according to the host. The genera that infect both Culex quinquefasciatus (Say, 1823) and crustaceans (Hazardia, Marsoniella, Gurleya, Larssonia and Berwaldia) are the main members of this clade. The genera that infect anopheline mosquitoes (Parathelohania), simulids (Amblyospora spp 3 in this study) and a species of ant (Varimorpha sp.), are more closely related to the aquatic group than to the main Amblyospora group. TheAmblyospora spp. in this study were clustered with Amblyospora sp. (AJ252949) from Simulium spp. from the Paleartic29,; confirming the morphological and molecular similarities between these 2 species.
Phylogenetic analysis with the 16S gene showed considerable distance between the Amblyospora spp., which infect simulids, and the main group of Amblyospora spp., which infects mosquitoes, indicating that these groups are not congeneric. The differences between taxonomic relationships, based on phylogenetic placement and classical morphological characteristics, could probably be explained by the possibility that some of these characteristics (diplokaryon, sporophorous vesicles, and meiosis) appear to have multiple origins31. Thus, molecular analysis of other genomic regions could improve the phylogenetic understanding of microsporidia. This work contributes to the phylogenetic analysis of microsporidia because it provides two genus sequences from these parasites.
