Home » Volumes » Volume 47 September/Octuber 2014 » Description of microsporidia in simulids: molecular and morphological characterization of microsporidia in the larvae of Simulium pertinax Kollar (Diptera: Simuliidae)

Description of microsporidia in simulids: molecular and morphological characterization of microsporidia in the larvae of Simulium pertinax Kollar (Diptera: Simuliidae)

Isabel Maria Vicente Guedes de Carvalho[1] Artur Trancoso Lopo de Queiroz[2] Rosiane Brito de Moraes[4] Helio Benites Gil[3] Rafael Alves[5] Andréa de Barros Pinto Viviani[6] James John Becnel[7] Carlos José Pereira da Cunha de Araujo-Coutinho[4]

[1]Laboratório de Parasitologia, Instituto Butantan, São Paulo, SP [2]Laboratório de Imunoparasitologia, Centro de Pesquisa Gonçalo Moniz, Fundação Oswaldo Cruz, Salvador, BA [3]Disciplina de Infectologia, Universidade Federal de São Paulo, São Paulo, SP [4]Laboratório de Entomologia Médica, Superintendência de Controle de Endemias, São Paulo, SP [5]Departamento de Medicina, Disciplina de Gastroenterologia, Universidade Federal de São Paulo, São Paulo, SP [6]Laboratório de Simulídeos, Superintendência de Controle de Endemias, Caraguatatuba, SP [7]United States Department of Agriculture, Mosquito and Fly Research Unit, Gainesville, Florida, USA

DOI: 10.1590/0037-8682-0165-2014



Microsporidia constitute the most common black fly pathogens, although the species’ diversity, seasonal occurrence and transmission mechanisms remain poorly understood. Infections by this agent are often chronic and non-lethal, but they can cause reduced fecundity and decreased longevity. The objective of this study was to identify microsporidia infecting Simulium (Chirostilbia) pertinax (Kollar, 1832) larvae from Caraguatatuba, State of São Paulo, Brazil, by molecular and morphological characterization.


Larvae were collected at a single point in a stream in a rural area of the city and were kept under artificial aeration until analysis. Polydispyrenia spp. infection was characterized by the presence of at least 32 mononuclear spores measuring 6.9 ± 1.0 × 5.0 ± 0.7µm in persistent sporophorous vesicles. Similarly, Amblyospora spp. were characterized by the presence of eight uninucleate spores measuring 4.5 × 3.5µm in sporophorous vesicles.


The molecular analysis confirmed the presence of microsporidian DNA in the 8 samples (prevalence of 0.51%). Six samples (Brazilian larvae) were related to Polydispyrenia simulii and Caudospora palustrisreference sequences but in separate clusters. One sample was clustered with Amblyospora spp. Edhazardia aedis was the positive control taxon.


Samples identified as Polydispyrenia spp. and Amblyospora spp. were grouped with P. simulii and Amblyospora spp., respectively, corroborating previous results. However, the 16S gene tree showed a considerable distance between the black fly-infecting Amblyospora spp. and the mosquito-infecting spp. This distance suggests that these two groups are not congeneric. Additional genomic region evaluation is necessary to obtain a coherent phylogeny for this group.

Key words: Microsporidae; Amblyospora spp; Polydispyrenia spp; Phylogenetic analysis


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 production13. 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.


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.

FIGURE 1-  Simulium pertinax larvae. Healthy larvae with normal coloration of the integument (A). Larvae with symptoms of microsporidian infection in the fat bodies (B). 

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 sequences3246 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.


Nucleotide sequences and accession numbers

The nucleotide sequences obtained in this work were submitted to the GenBank nucleotide sequences databank under the following accession numbers GenBank: KC855552-KC855557 (L1_L6); and GenBank: KC855558 (L2).


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. 


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 results2226 regarding the phylogeny of these parasites.

The genera ParathelohaniaHazardiaMarsoniellaGurleyaLarssoniaBerwaldiaVarimorphaAmblyospora 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 hosts2730.

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.


The authors declare that there is no conflict of interest.


Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) 2012/23947-0


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Received: July 18, 2014; Accepted: October 9, 2014

Address to: Dr. Carlos José Pereira da Cunha de Araújo-Coutinho. Laboratório de Entomologia Médica/SUCEN. Rua Cardeal Arcoverde no 2878, 05408-003 São Paulo, SP, Brasil. Phone: 55 11 3032-2228. e-mail: cjpcacoutinho@gmail.com