Molecular Investigation Confirms Myotis Genus Bats as Common Hosts of Polychromophilus in Brazil

Abstract

Plasmodium spp. and some other blood parasites belonging to the order Haemosporida are the focus of many epidemiological studies worldwide. However, haemosporidian parasites from wild animals are largely neglected in scientific research. For example, Polychromophilus parasites, which are exclusive to bats, are described in Europe, Asia, Africa, and Oceania, but little is known about their presence and genetic diversity in the New World. In this study, 224 samples of bats from remaining fragments of the Atlantic Forest and Pantanal biomes, as well as urbanized areas in southern and southeastern Brazil, were analyzed for the presence of haemosporidian parasites by PCR of the mitochondrial gene that encodes cytochrome b (cytb). The PCR fragments of the positive samples were sequenced and analyzed by the Bayesian inference method to reconstruct the phylogenetic relationships between Polychromophilus parasites from bats in Brazil and other countries. Sequences from Brazilian lineages of Polychromophilus were recovered in a clade with sequences from Polychromophilus murinus and close to the one Polychromophilus sequence obtained in Panama, the only available sequence for the American continent. This clade was restricted to bats of the family Vespertilionidae and distinct from Polychromophilus melanipherus, a parasite species mainly found in bats of the family Miniopteridae. The detection of Polychromophilus and the genetic proximity to P. murinus were further confirmed with the amplification of two other genes (clpc and asl). We also found a Haemosporida parasite sequence in a sample of Noctilio albiventris collected in the Pantanal biome, which presents phylogenetic proximity with avian Haemoproteus sequences. Morphological and molecular studies are still needed to conclude and describe the Polychromophilus species in Brazilian Myotis bats in more detail and to confirm Haemoproteus parasites in bats. Nevertheless, these molecular results in Brazilian bats confirm the importance of studying these neglected genera.

1. Introduction

Bats harbor a large diversity of pathogens, causing emerging infectious diseases, including bacteria and viruses [1]. They also are hosts for protozoa, such as a variety of haemosporidian parasites (Apicomplexa: Haemosporida) [2]. The parasites belonging to the Order Haemosporida (Danilewsky, 1885) are characterized by being obligatorily heteroxenous, i.e., they need more than one host to complete their evolutionary cycle. The merogony stage of a haemosporidian occurs in vertebrate hosts (reptiles, birds, and mammals), the intermediate hosts, while the sporogony stage occurs in many species of hematophagous dipterans, the definitive hosts. Haemosporidian parasites are cosmopolitan organisms widely distributed on all continents, but their diversity and phylogenetic relationships are not well established [3].

Currently, more than 500 species of haemosporidians parasitizing different groups of vertebrate hosts have been described, and new species continue to be described [4,5]. In mammals, haemosporidian infections are mainly known from primates, rodents, ungulates, and bats [6]. Bats stand out for the diversity of haemosporidian genera that can parasitize them, demonstrating a well-established parasite-host-vector relationship [3,7,8,9].

Nine genera of the order Haemosporida have been described in bats in the Old World. Plasmodium Marchiafava and Celli, 1885, Polychromophilus Dionisi 1898, Hepatocystis Miller 1908, Nycteria Garnham and Heisch, 1953, Bioccala [10], Biguetiella [11], Dionisia [10], all from the Plasmodiidae family Mesnil, 1903; and Johnsprentia [12] and Sprattiella [13] from the Haemoproteidae family Doflein, 1916. The two new genera, Johnsprentia and Sprattiella, were only recently described, and only morphological data is available.

Polychromophilus is the most widely distributed parasite genus, mainly being found in insectivorous bats of the Miniopteridae and Vespertilionidae families in temperate areas of Europe and tropical areas in Africa, Southeast Asia, and Oceania [6]. In the New World, parasitism in bats by Haemosporida species is rare, with only one species described in South America, Polychromophilus deanei, found in Myotis nigricans in Pará State, Brazil [14]. In 2014, a Polychromophilus sequence was recovered from Myotis nigricans from Panama [15], and recently, Polychromophilus was reported for the first time by molecular detection in Brazilian bats [16]. Here, we aimed to screen for haemosporidian parasites in bats sampled in three Brazilian biomes, extending the survey for these parasites in the Neotropics. Moreover, we amplified the caseinolytic protease C (clpc) gene, present in the genome of the parasite’s apicoplast, and adenylosuccinate lyase (asl) nuclear gene, in addition to cytochrome b (cytb) to confirm the close phylogenetic relationship of Polychromophilus parasites in Brazil to P. murinus.

2. Materials and Methods

2.1. Sampling

Samples were collected from different species of bats belonging to the Atlantic Forest and Pantanal biomes from four states of different regions, including southern (Paraná/PR), central-western (Mato Grosso/MT), and southeastern (São Paulo/SP and Rio de Janeiro/RJ) Brazil (Figure 1 and Table 1). Specific data for each state are presented in the following items.

2.1.1. Paraná/PR

Brains from bats not identified to species (n = 96) from the Paraná State Reference Laboratory (LACEN) were acquired as part of the rabies virus circulation monitoring program. The collections were carried out between September 2019 and August 2020 in 29 different municipalities in the Paraná State, inserted in remnant fragments of the Atlantic Forest biome and urbanized areas. Samples were stored at −80 °C and thawed immediately prior to DNA/RNA extraction.

Eight blood samples from bats (six Desmodus rotundus and two Diphylla ecaudata, both family Phyllostomidae) were collected in EDTA-anticoagulated tubes by intracardiac puncture from sedated bats, between October and November 2015, in two caves in the municipality of Rio Branco do Sul, PR, which is located in a fragment of Atlantic Forest. Since they are hematophagous species, those bats were collected as part of the rabies monitoring program in herbivores (ruminants and horses) carried out by municipal and state public agencies. In addition, two non-hematophagous bats of the genus Molossus, family Molossidae, were collected for rabies surveillance in Curitiba city, as they were found in public areas nearby the Municipal Zoo of Curitiba, PR [17]. Blood samples were stored at −20 °C until processing.

2.1.2. Mato Grosso/MT and Rio de Janeiro/RJ

Bat blood samples stored on FTA^® Whatman^® cards (Whatman, Sigma-Aldrich, Darmstadt, Germany) were collected in Mato Grosso (17 samples) and Rio de Janeiro (four samples). Blood smears were also prepared for microscopical examination from Rio de Janeiro.

In Mato Grosso, samples were collected in October 2019 in the municipality of Poconé, in rural areas within the Pantanal biome. Species and families sampled: Glossophaga soricina, family Phyllostomidae (five samples); Molossus molossus, family Molossidae (five samples); Myotis cf. nigricans, family Vespertilionidae (four samples); Rhynchonycteris naso, family Emballonuridae (two samples); Noctilio albiventris, family Noctilionidae (one sample).

Samples from Rio de Janeiro were collected in October 2020 in the city of Rio de Janeiro, in a remnant of Atlantic Forest (Pedra Branca State Park), near Fiocruz Atlantic Forest Biological Station, a highly urbanized region in the central portion of the city, under severe anthropogenic pressure. Only bats of the species Artibeus lituratus, family Phyllostomidae were sampled.

2.1.3. São Paulo/SP

Blood samples from bats (n = 97) were collected in EDTA-anticoagulated tubes by intracardiac puncture from sedated bats. Certain specimens were euthanized using xylazine and ketamine, followed by inhalation of isoflurane. Subsequently, they were preserved in a 10% formaldehyde solution for future identification and storage at the Museum of Zoology of the University of São Paulo (MZUSP). From these specimens, liver (n = 30) and spleen (n = 12) were also collected.

Samples were collected from 2018 to 2021 in Legado das Águas reserve, the largest private reserve of Atlantic Forest in the country. It is inserted in the municipalities of Miracatu and Tapiraí, in the Ribeira Valley, south of the São Paulo State, located 122 km from the state capital, in the southern portion of the Serra do Mar ecological corridor. Legado das Águas covers 31,000 hectares contiguously connected to several other Conservation Units, contributing to an important ecological corridor between the coastal and inland areas of the southern region of São Paulo State. It is the largest continuous area of remaining Atlantic Forest that has suffered minimal human intervention, a factor attributed to the low demographic density and little economic development in the region.

Families and species sampled: family Phyllostomidae, Anoura caudifer (seven samples), Artibeus cinereus (two samples), Artibeus fimbriatus (three samples), Artibeus gnomus (one sample), Artibeus lituratus (nine samples), Artibeus obscurus (nine samples), Artibeus planirostris (three samples), Artibeus sp. (one sample), Carollia perspicillata (eighteen samples), Chrotopterus auritus (one sample), Desmodus rotundus (one sample), Ectophylla sp. (one sample), Lonchorhina aurita (one sample), Platyrrhinus lineatus (six samples), Platyrrhinus sp. (one sample), Rhinophylla pumilio (one sample), Sturnira lilium (seven samples), Thrachops cirrhosus (one sample), Uroderma bilobatum (one sample); family Emballonuridae, Pteropteryx sp. (three samples); family Molossidae, Molossus ater (one sample), Nyctinomops sp. (four samples); family Vespertilionidae, Eptesicus sp. (one sample), Myotis nigricans (eight samples), Myotis riparius (one sample), Myotis ruber (one sample), Myotis sp. (four samples). All these samples are also described in Table 1.

2.2. Ethics Statement

All animals and their tissue samples were collected and handled under appropriate authorizations by the Brazilian government. The project was authorized by SISBIO (Sistema de Autorização e Informação em Biodiversidade), ICMBio/MMA (Instituto Chico Mendes de Conservação da Biodiversidade/Ministério do Meio Ambiente), numbers 72790, 51714-1, and 19037-1. The study was approved by the Ethics in Use of Animals Committee, CEUA/SESA, of the Centro de Produção e Pesquisa de Imunobiológicos—CPPI/PR (approval number 01/2019 and date of approval 3 March 2020), CEUA/FIOCRUZ (approval number LM-6/18/2021 and date of approval 14 May 2018) and CEUA/SUCEN (approval number 09/2021 and date of approval 30 September 2021). Rio de Janeiro sampling was carried out under SisGen authorization A46B0E1.

2.3. Optical Microscopy Diagnosis

The four blood smears available for analysis acquired from bats in Rio de Janeiro were fixed with 100% methanol within 24 h of collection and stained with 10% Giemsa solution for one hour, up to 30 days after collection [3]. The smears were examined for approximately 15–20 min, viewing 100 fields at low magnification (400×) and 100 fields at high magnification (1000×) [3], using a Leica^® DM3000LED light microscope. The search for parasitic blood stages was carried out following previous morphological studies on haemosporidians in wild animals [3,8].

2.4. DNA Extraction

Brain tissue samples from Paraná State were extracted using the BioGene DNA/RNA Viral Kit (K204-4, Bioclin, Belo Horizonte, MG, Brazil), following the manufacturer’s instructions. For the blood (200 μL), the DNA was prepared according to the Illustra Kit Mini Genomic Blood Preparation Spin (GE Healthcare, Chalfont, St. Giles, UK), according to the manufacturer’s instructions.

Samples from Rio de Janeiro and Mato Grosso, stored on FTA^® Whatman^® cards (Whatman, Sigma-Aldrich, Darmstadt, Germany), were extracted using the commercial Wizard SV 96 Genomic DNA Purification System kit (PROMEGA^®, Madison, WI, USA), according to the manufacturer’s instructions.

For samples collected in the São Paulo State, genomic DNA was extracted from blood, liver and spleen tissues using PureLink™ Genomic DNA Mini Kit (Thermo Fisher Scientific, Waltham, MA, USA), according to the manufacturer’s instructions.

2.5. Molecular Detection of Haemosporidian Parasites

A fragment of ~1.1 kb (approximately 92% of the gene) from the mitochondrial cytb gene was amplified using a nested polymerase chain reaction (PCR), taking standard precautions to prevent cross-contamination of samples. The PCR reactions were conducted as described in [18] using primers DW2 and DW4 and 5 µL (50 ng) of genomic DNA in the first reaction. An aliquot of 1 µL of the PCR product was used as a template for a nested reaction with primers DW1 and DW6.

For clpc, a fragment of approximately 500 bp was amplified using a nested PCR, as described in [8], using primers clpcF and clpcR and 5 µL (50 ng) of genomic DNA in the first reaction. An aliquot of 1 µL of the PCR product was used as a template for a nested reaction with primers clpcF2 and clpcR2.

For asl, a fragment of approximately 240 bp was amplified using a nested PCR, as described in [8], using the primers aslF and aslR and 5 µL (50 ng) of genomic DNA in the first reaction. An aliquot of 1 µL of the PCR product was used as a template for a nested reaction with primers aslF2 and aslR2.

All PCR amplifications included two controls: one positive control (DNA sample with known Polychromophilus sp. infection) and one negative control (ultrapure water without DNA); the last one was included to check for possible contamination and false positives. All PCR products were evaluated by running 10 μL on 1% agarose gel.

PCR products were sequenced using BigDye^® Terminator v3.1 Cycle Sequencing Kit in ABI PRISM^® 3500 Genetic Analyzer (Applied Biosystems, Carlsbad, CA, USA) using nested PCR primers. For cytb, the oligonucleotides DW8 and DW3 were also used for sequencing [18]. The cytb, clpc or asl sequences were obtained and aligned with the sequences available in the GenBank^® database.

2.6. Phylogenetic Analysis

The phylogenetic relationships among the haemosporidian parasites were inferred using partial sequences of cytb (1116 bp) or the concatenated analysis of three genes, the mitochondrial cytochrome b gene (cytb, 725 bp), the nuclear adenylosuccinate lyase gene (asl, 206 bp), and the apicoplast caseinolytic protease C gene (clpc, 531 bp). Sequences acquired from the GenBank^® database used are shown in the phylogenetic trees. The alignment was obtained using the ClustalW algorithm [19] implemented in MEGAX software [20]. The phylogenetic reconstructions were performed using the Bayesian inference method implemented in MrBayes v3.2.0 [21]. The best evolution model was selected using MEGAX software [20]. The model GTR + G + I was considered to describe the substitution pattern the best (with the lowest Bayesian Information Criterion scores). Bayesian inferences were executed with two Markov Chain Monte Carlo searches of 8 million generations, with each sampling 1 of 300 trees. After a burn-in of 25%, the remaining 15,002 trees were used to calculate the 50% majority-rule consensus tree. The standard deviation of split frequencies was <0.01. The phylogenies were visualized using FigTree version 1.4.0 [22].

2.7. Host Species Identification

A fragment with ~650 bp from the mitochondrial cytochrome c oxidase (coi) gene was amplified using the universal primers LCO1490 and HCO2198 [23] and PCR protocol based on [24]. Amplified fragments were sequenced directly using the corresponding flanking primers. Sequences (at least 550 bp) were entered in the BOLD platform (Barcode of Life Data system) by the option “Species Level Barcode Records”. Sequences with >99% similarity were used for species identification. For sequences showing <98% identity, the neighbor-joining (NJ) tree of K2P distances showing intra and interspecific variation generated in BOLD was analyzed. Samples were considered identified if assigned to a monophyletic group of sequences corresponding to a single species.

3. Results

3.1. Host Species Identification

We used DNA barcodes and BOLD to identify bat species in the samples obtained from LACEN. Five of the 95 tested samples did not amplify (in two independent experiments). A total of 80 (84.2%) could be identified to the species level using DNA barcoding and BOLD, and 10 (10.5%) generated ambiguous identification and were identified just to genera (one Artibeus sp., one Eumops sp., four Molossus sp., one Sturnira sp., two Eptesicus sp. and one Myotis sp.). From the samples identified to species level, 50 samples showed >99% while 30 had <99% of similarity in the BOLD database. In the last case, the BOLD NJ tree was analyzed to consider the sample identified. Ten different species were obtained: 15 specimens were Vespertilionidae (two Myotis nigricans, one Myotis riparius, and 12 Eptesicus furinalis); 65 bats were Molossidae (23 Molossus rufus, 29 Molossus molossus, five Tadarida brasiliensis, three Promops nasutus, three Eumops glaucinus, one Molossops temminckii, and one Molossops neglectus).

3.2. Haemosporidian Parasites and Phylogeny

The percentage of positive samples by PCR was 2.67%, with six out of 224 bats infected (Table 1). No positive samples were obtained by microscopy. Positive samples were: two brain tissue samples of Myotis riparius (sample IDs 198 and 607), bats belonging to the Vespertilionidae family, collected in two municipalities in the Paraná State (Curitiba and São José dos Pinhais, respectively); one spleen tissue sample of Myotis ruber collected in Tapiraí-SP (sample ID 125); two samples, spleen and liver tissues respectively, of Myotis sp. from Miracatu-SP (sample IDs 138 and 141); and a blood sample of Noctilio albiventris (sample ID Bat17) that belongs to the Noctilionidae family. The latter species was collected near a lake in the Pantanal of Mato Grosso, municipality of Poconé (Figure 1). Interestingly, paired samples from bats ID 125, 138 and 141 obtained variable results, with liver samples being the worst for Polychromophilus detection (

Table 2. Bats examined in this study with positive PCR results in paired samples.

Sample ID	Blood Sample	Spleen Sample	Liver Sample
125	NA	Positive	Negative
138	Positive	Positive	Negative
141	Positive	NA	Positive

NA = not available.

).

Table 1. Bats examined in this study and the prevalence of haemosporidian infection.

Host Species	Locality	Year of Collection	Bats Examined	Bats Infected with Haemosporidians (%)
Vespertilionidae Gray, 1821
Myotis nigricans	Rio Bom, Toledo-PR	2019–2020	2	0
Myotis riparius	São José dos Pinhais, Curitiba-PR	2019–2020	2	2 (100%)
Eptesicus furinalis	Assis Chateaubriand, Curitiba, Foz do Iguaçu, Maringá, Ramilândia-PR	2019–2020	12	0
Eptesicus sp.	Curitiba-PR	2019–2020	2	0
Myotis sp.	Curitiba-PR	2019–2020	1	0
Myotis cf. nigricans	Poconé-MT	2019	4	0
Eptesicus sp.	Miracatu-SP	2018	1	0
Myotis nigricans	Tapiraí, Miracatu-SP	2018–2019	8	0
Myotis riparius	Miracatu-SP	2019	1	0
Myotis ruber	Tapiraí-SP	2019	1	1 (100%)
Myotis sp.	Miracatu-SP	2019	4	2 (50%)
Molossidae Gervais, 1856
Molossus sp.	Curitiba, Araruna-PR	2019–2020	4	0
Molossus sp.	Curitiba-PR	2015	2	0
Eumops sp.	Foz do Iguaçu-PR	2019–2020	1	0
Molossus rufus	Braganey, Cascavel, Céu Azul, Francisco Beltrão, Jacarezinho, Londrina, Mamborê, Maringá, Maripá, Sarandi, Telêmaco Borba, Vera Cruz do Oeste-PR	2019–2020	23	0
Molossus molossus	Araucária, Assis Chateaubriand, Cascavel, Curitiba, Foz do Iguaçu, Guapirama, Guaratuba, Mandaguaçu, Maringá, Paulo Frontin, Ramilândia, Sarandi-PR	2019–2020	29	0
Molossus molossus	Poconé-MT	2019	5	0
Tadarida brasiliensis	Curitiba, Imbituva, Mamborê-PR	2019–2020	5	0
Promops nasutus	Cascavel, União da Vitória-PR	2019–2020	3	0
Eumops glaucinus	Assis Chateaubriand, Foz do Iguaçu, Maringá-PR	2019–2020	3	0
Molossops temminckii	Foz do Iguaçu-PR	2019–2020	1	0
Molossops neglectus	Salto do Lontra-PR	2019–2020	1	0
Molossus ater	Tapiraí-SP	2018	1	0
Nyctinomops sp.	Tapiraí-SP	2019–2020	4	0
Phyllostomidae Gray, 1825
Sturnira sp.	Curitiba-PR	2019–2020	1	0
Artibeus sp.	Foz do Iguaçu-PR	2019–2020	1	0
Artibeus lituratus	Rio de Janeiro-RJ	2020	4	0
Desmodus rotundus	Rio Branco do Sul-PR	2015	6	0
Diphylla ecaudata	Rio Branco do Sul-PR	2015	2	0
Glossophaga soricina	Poconé-MT	2019	5	0
Anoura caudifer	Tapiraí-SP	2018	7	0
Artibeus cinereus	Tapiraí-SP	2018	2	0
Artibeus fimbriatus	Tapiraí, Miracatu-SP	2018	3	0
Artibeus gnomus	Tapiraí-SP	2018	1	0
Artibeus lituratus	Tapiraí-SP	2018	9	0
Artibeus obscurus	Tapiraí-SP	2018	9	0
Artibeus planirostris	Tapiraí-SP	2018	3	0
Artibeus sp.	Tapiraí-SP	2018	1	0
Carollia perspicillata	Tapiraí, Miracatu-SP	2018	18	0
Chrotopterus auritus	Tapiraí-SP	2018	1	0
Desmodus rotundus	Tapiraí-SP	2018	1	0
Ectophylla sp.	Tapiraí-SP	2018	1	0
Lonchorhina aurita	Miracatu-SP	2018	1	0
Platyrrhinus lineatus	Tapiraí-SP	2020	6	0
Platyrrhinus sp.	Tapiraí-SP	2020	1	0
Rhinophylla pumilio	Tapiraí-SP	2020	1	0
Sturnira lilium	Tapiraí-SP	2021	7	0
Thrachops cirrhosus	Tapiraí-SP	2021	1	0
Uroderma bilobatum	Miracatu-SP	2021	1	0
Emballonuridae Gervais, 1856
Rhynchonycteris naso	Poconé-MT	2019	2	0
Pteropteryx sp.	Tapiraí-SP	2020	3	0
Noctilionidae Gray, 1821
Noctilio albiventris	Poconé-MT	2019	1	1 (100%)
unknown	Curitiba, Paulo Frontin, Rolândia, Salto do Lontra-PR	2019–2020	5	0
TOTAL			224	6 (2.67%)

PR = Paraná State; MT = Mato Grosso State; RJ = Rio de Janeiro State; SP = São Paulo State.

Table 1. Bats examined in this study and the prevalence of haemosporidian infection.

Host Species	Locality	Year of Collection	Bats Examined	Bats Infected with Haemosporidians (%)
Vespertilionidae Gray, 1821
Myotis nigricans	Rio Bom, Toledo-PR	2019–2020	2	0
Myotis riparius	São José dos Pinhais, Curitiba-PR	2019–2020	2	2 (100%)
Eptesicus furinalis	Assis Chateaubriand, Curitiba, Foz do Iguaçu, Maringá, Ramilândia-PR	2019–2020	12	0
Eptesicus sp.	Curitiba-PR	2019–2020	2	0
Myotis sp.	Curitiba-PR	2019–2020	1	0
Myotis cf. nigricans	Poconé-MT	2019	4	0
Eptesicus sp.	Miracatu-SP	2018	1	0
Myotis nigricans	Tapiraí, Miracatu-SP	2018–2019	8	0
Myotis riparius	Miracatu-SP	2019	1	0
Myotis ruber	Tapiraí-SP	2019	1	1 (100%)
Myotis sp.	Miracatu-SP	2019	4	2 (50%)
Molossidae Gervais, 1856
Molossus sp.	Curitiba, Araruna-PR	2019–2020	4	0
Molossus sp.	Curitiba-PR	2015	2	0
Eumops sp.	Foz do Iguaçu-PR	2019–2020	1	0
Molossus rufus	Braganey, Cascavel, Céu Azul, Francisco Beltrão, Jacarezinho, Londrina, Mamborê, Maringá, Maripá, Sarandi, Telêmaco Borba, Vera Cruz do Oeste-PR	2019–2020	23	0
Molossus molossus	Araucária, Assis Chateaubriand, Cascavel, Curitiba, Foz do Iguaçu, Guapirama, Guaratuba, Mandaguaçu, Maringá, Paulo Frontin, Ramilândia, Sarandi-PR	2019–2020	29	0
Molossus molossus	Poconé-MT	2019	5	0
Tadarida brasiliensis	Curitiba, Imbituva, Mamborê-PR	2019–2020	5	0
Promops nasutus	Cascavel, União da Vitória-PR	2019–2020	3	0
Eumops glaucinus	Assis Chateaubriand, Foz do Iguaçu, Maringá-PR	2019–2020	3	0
Molossops temminckii	Foz do Iguaçu-PR	2019–2020	1	0
Molossops neglectus	Salto do Lontra-PR	2019–2020	1	0
Molossus ater	Tapiraí-SP	2018	1	0
Nyctinomops sp.	Tapiraí-SP	2019–2020	4	0
Phyllostomidae Gray, 1825
Sturnira sp.	Curitiba-PR	2019–2020	1	0
Artibeus sp.	Foz do Iguaçu-PR	2019–2020	1	0
Artibeus lituratus	Rio de Janeiro-RJ	2020	4	0
Desmodus rotundus	Rio Branco do Sul-PR	2015	6	0
Diphylla ecaudata	Rio Branco do Sul-PR	2015	2	0
Glossophaga soricina	Poconé-MT	2019	5	0
Anoura caudifer	Tapiraí-SP	2018	7	0
Artibeus cinereus	Tapiraí-SP	2018	2	0
Artibeus fimbriatus	Tapiraí, Miracatu-SP	2018	3	0
Artibeus gnomus	Tapiraí-SP	2018	1	0
Artibeus lituratus	Tapiraí-SP	2018	9	0
Artibeus obscurus	Tapiraí-SP	2018	9	0
Artibeus planirostris	Tapiraí-SP	2018	3	0
Artibeus sp.	Tapiraí-SP	2018	1	0
Carollia perspicillata	Tapiraí, Miracatu-SP	2018	18	0
Chrotopterus auritus	Tapiraí-SP	2018	1	0
Desmodus rotundus	Tapiraí-SP	2018	1	0
Ectophylla sp.	Tapiraí-SP	2018	1	0
Lonchorhina aurita	Miracatu-SP	2018	1	0
Platyrrhinus lineatus	Tapiraí-SP	2020	6	0
Platyrrhinus sp.	Tapiraí-SP	2020	1	0
Rhinophylla pumilio	Tapiraí-SP	2020	1	0
Sturnira lilium	Tapiraí-SP	2021	7	0
Thrachops cirrhosus	Tapiraí-SP	2021	1	0
Uroderma bilobatum	Miracatu-SP	2021	1	0
Emballonuridae Gervais, 1856
Rhynchonycteris naso	Poconé-MT	2019	2	0
Pteropteryx sp.	Tapiraí-SP	2020	3	0
Noctilionidae Gray, 1821
Noctilio albiventris	Poconé-MT	2019	1	1 (100%)
unknown	Curitiba, Paulo Frontin, Rolândia, Salto do Lontra-PR	2019–2020	5	0
TOTAL			224	6 (2.67%)

PR = Paraná State; MT = Mato Grosso State; RJ = Rio de Janeiro State; SP = São Paulo State.

Sequencing of the PCR amplicon revealed that sample ID 607 (M. riparius) was infected with Polychromophilus sp. It was not possible to identify the haemosporidian parasite in sample Bat17 (Noctilio albiventris) since an unprecedented sequence was obtained. Its closest sequence available on GenBank^® (KY653763) was obtained from Haemoproteus minutus, infecting Turdus merula, a passerine collected in Lithuania [25] with a 94% identity.

The cytb gene phylogenetic tree generated using reference sequences available in the GenBank^® database covering different haemosporidian genera from different hosts, as well as the sequences found herein are shown in Appendix A. Cytb-based phylogenetic analysis produced no conflict in any of the major nodes. All major genera and subgenera were recovered and represented in the phylogenetic tree by separate monophyletic clades (Figure 2). The results show eight clades within the order Haemosporida analyzed here. All Polychromophilus sequences from bats from different regions of the world were grouped into a monophyletic clade (posterior probability of 100) and consisted of six subclades (with posterior probabilities > 95), with all Polychromophilus found in Brazilian Myotis bats segregated in two of them (Figure 2).

The first Polychromophilus distinct subclade comprised two samples of Pipistrellus aff. grandidieri and Neoromicia capensis, both vespertilionid species from Guinea (KF159700 and KF159714). The second subclade contained the Polychromophilus sequences from Scotophilus kuhlii, a vespertilionid species from Thailand (MT750305, MT750307, and MT750308) (Figure 2). The third Polychromophilus subclade (posterior probability of 95) was composed of sequences of P. murinus from bats in Europe (Switzerland, Bulgaria), Madagascar, and Thailand. This subclade included a sequence of Polychromophilus sp. obtained in Rhinolophus sp. (Rhinolophidae) that stands out for not being part of the vespertilionids (Figure 2). The fourth subclade comprises sequences obtained in Eptesicus diminutus (MW984521) and Myotis ruber (OQ957064) from Brazil. The fifth subclade comprises the sequence of Polychromophilus obtained from M. nigricans from Panama and all other Brazilian sequences isolated from the genus Myotis. This subclade exclusively included Polychromophilus sequences from vespertilionids (including Brazilian ones). All P. melanipherus sequences from hosts of bats of the genus Miniopterus were distinctly separated into a subclade, confirming a clear separation of parasites from miniopterid and vespertilionid hosts (Figure 2).

The sequence of Bat17 (N. albiventris) clustered close to the subclade of Haemoproteus (Parahaemoproteus) spp., a specific genus of bird parasites, being positioned as a sister clade. Thus, although in the phylogenetic tree, the sequence obtained in bat was grouped with others from Haemoproteidae, it was not supported in a monophyletic clade.

The finding of Polychromophilus in Brazilian bats was confirmed with the amplification of the clpc gene, from the apicoplast of the parasite, in three samples (IDs 141, 198 and 607), presenting fragments of approximately 500 bp and also of the asl gene from the nuclear genome, in two samples (IDs 125 and 141), with 244 bp. Compared to sequences of the same target gene on GenBank^® for the genus Polychromophilus, the clpc sequences showed 97% similarities with the closest available sequences (LC715203 and LC715204), sequences from P. murinus described in Myotis macrodactylus, a bat collected in Japan [26].

A phylogenetic tree was generated with concatenated sequences from three genes: cytb, asl, and clpc (Figure 3), including the Polychromophilus from this study and all available sequences of this genus in the GenBank^® database (

Table A2. Mitochondrial gene cytochrome b (cytb), nuclear gene adenylosuccinate lyase (asl) and apicoplast gene caseinolytic protease C (clpc) sequences from Polychromophilus species used in phylogenetic analyzes and their respective GenBank^® accession numbers. Sequences from this study are highlighted in bold.

Host Species	Parasite Species	cytb	asl	clpc	Country of Source
Miniopterus schreibersii	Polychromophilus melanipherus	JN990708	-	JN990720	Switzerland
Miniopterus schreibersii	Polychromophilus melanipherus	JN990709	JN990726	JN990721	Switzerland
Miniopterus schreibersii	Polychromophilus melanipherus	JN990710	-	JN990722	Switzerland
Myotis daubentonii	Polychromophilus murinus	JN990712	JN990725	JN990723	Switzerland
Myotis daubentonii	Polychromophilus murinus	JN990713	-	JN990724	Switzerland
Miniopterus villiersi	Polychromophilus sp.	KF159699	-	KF159616	Guinea
Neoromicia capensis	Polychromophilus sp.	KF159681	-	KF159642	Guinea
Pipistrellus aff. grandidieri	Polychromophilus sp.	KF159714	-	KF159639	Guinea
Miniopterus natalensis	Polychromophilus melanipherus	KT750379	KT750646	KT750738	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750382	KT750633	-	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750380	KT750647	KT750740	Kenya
Miniopterus rufus	Polychromophilus melanipherus	KT750385	KT750637	KT750745	Kenya
Miniopterus rufus	Polychromophilus melanipherus	KT750386	-	KT750748	Kenya
Miniopterus sp.	Polychromophilus melanipherus	KT750387	KT750642	KT750749	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750377	KT750629	-	Kenya
Miniopterus africanus	Polychromophilus melanipherus	KT750375	KT750627	KT750734	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750400	KT750630	KT750737	Kenya
Miniopterus rufus	Polychromophilus melanipherus	KT750403	KT750636	KT750744	Kenya
Miniopterus rufus	Polychromophilus melanipherus	KT750404	KT750639	KT750746	Kenya
Miniopterus rufus	Polychromophilus melanipherus	KT750418	KT750641	-	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750376	KT750628	KT750735	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750401	KT750631	KT750739	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750402	KT750648	KT750742	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750406	-	KT750743	Kenya
Miniopterus natalensis	Polychromophilus melanipherus	KT750378	-	KT750736	Kenya
Miniopterus minor	Polychromophilus sp.	KT750388	KT750643	KT750750	Tanzania
Miniopterus minor	Polychromophilus sp.	KT750428	KT750644	KT750751	Tanzania
Miniopterus minor	Polychromophilus sp.	KT750429	KT750645	-	Tanzania
Miniopterus sp.	Polychromophilus sp.	KT750389	KT750552	KT750651	Mozambique
Miniopterus rufus	Polychromophilus sp.	KT750384	KT750635	-	Kenya
Miniopterus rufus	Polychromophilus sp.	KT750383	KT750634	-	Kenya
Miniopterus natalensis	Polychromophilus sp.	KT750381	KT750632	KT750741	Kenya
Miniopterus rufus	Polychromophilus sp.	KT750412	KT750638	-	Kenya
Miniopterus rufus	Polychromophilus sp.	KT750405	KT750640	KT750747	Kenya
Scotophilus kuhlii	Polychromophilus sp.	MT750307	-	MT750315	Thailand
Myotis macrodactylus	Polychromophilus murinus	LC668431	-	LC715204	Japan
Myotis macrodactylus	Polychromophilus murinus	LC668432	-	LC715203	Japan
Myotis macrodactylus	Polychromophilus murinus	LC668433	-	LC715205	Japan
Myotis riparius (ID 198)	Polychromophilus sp.	MW984519	-	OP503503	Brazil
Myotis riparius (ID 607)	Polychromophilus sp.	OP503502	-	OP503504	Brazil
Myotis ruber (ID 125)	Polychromophilus sp.	OQ957064	OQ957067	-	Brazil
Myotis sp. (ID 138)	Polychromophilus sp.	OQ957065	-	-	Brazil
Myotis sp. (ID 141)	Polychromophilus sp.	OQ957066	OQ957068	OQ957063	Brazil

, Appendix A). The tree topology of concatenated genes confirmed the separation of parasites from miniopterid and vespertilionid hosts, except for Neoromicia capensis, a vespertilionid species grouped with miniopterid hosts. The vespertilionid Polychromophilus subclade was divided into three branches: one with P. murinus sequences from Swiss bats (Myotis daubentonii), one with just a sequence from Myotis macrodactylus from Japan, and the third with all the Polychromophilus found in Brazilian bats and another two from Myotis macrodactylus from Japan.

4. Discussion

The study of haemosporidian parasites in bats can significantly contribute to understanding the evolution of these parasites in mammals since seven out of nine genera of this family occurring in bats are considered specific to these hosts [6]. Haemosporidians have been found mainly in Old World bats, except for Polychromophilus from vespertilionid bats: Myotis nigricans from Brazil [14], Myotis nigricans from Panama [15] and, more recently, in Myotis riparius, Myotis ruber and Eptesicus diminutus from Brazil [16].

This study extended the search for haemosporidian parasites in bats to two additional Brazilian areas, including the Pantanal biome. We found a low haemosporidian positivity rate prevalence (2.67%), consistent with our previous study (1.2%) [16]. It is important to note that 52% of the analyzed samples were obtained from tissues (brain, spleen or liver), sample sources that are not common in haemoparasite studies but confirmed its usefulness in the screening of Polychromophilus parasites since we obtained the same amount of positives found in the group of blood samples.

We hypothesize that the low positivity found in our studies is related to the number of samples collected from bats of the Myotis genus (10% in this study), which we believe to be the main host of Polychromophilus in Brazil. In fact, considering only the Myotis bats tested, we found 21% of positives. Of all the nine samples already found positive for Polychromophilus by molecular methods in Brazil (this study and [16]), only one was not within the Myotis species.

The four new Polychromophilus cytb sequences obtained in this study conserved the two nucleotides T (thymine) at positions 247 and 512 of the gene, which is also observed in other Brazilian isolates, but not in the sequence from Panama [16]. Future studies analyzing the cytb sequence of more isolates are needed to verify whether these SNPs are molecular markers of Brazilian Polychromophilus isolates.

The order Chiroptera corresponds to approximately one-quarter of the mammal species in the world [27]. In Brazil, there are nine families with 182 species [28]. The Brazilian families with their respective numbers of species are Emballonuridae (17), Phyllostomidae (94), Mormoopidae (4), Noctilionidae (2), Furipteridae (1), Thyropteridae (5), Natalidae (1), Molossidae (32) and Vespetilionidae (26) [28,29]. They inhabit the entire national territory and are distributed in the most diverse biomes and urban areas, occurring in the Amazon, Cerrado, Caatinga, Atlantic Forest, Pantanal, and Pampas [28,29,30,31,32,33]. To know the diversity of bat species tested in the present study, we used DNA barcoding to identify the bat species in samples with unknown species. The results showed that most of our samples come from the Phyllostomidae family (41.5%), followed by Molossidae (36.6%), Vespertilionidae (16.9%), and Noctilionidae plus Emballonuridae (2.6%), with 2.2% unidentified. Polychromophilus infection in Brazilian bats continues to be limited to just one family (Vespertilionidae). However, a Haemosporida sp. sequence was obtained from a Noctilionidae bat (Noctilio albiventris), a family with just one sample analyzed. It is important to note that there is one record of P. melanipherus in Emballonuridae (Taphozous melanopogon from Thailand) but no previous record of haemosporidian parasites in Molossidae, Phyllostomidae, and Noctilionidae families. Therefore, it is very likely that the prevalence of haemosporidian parasites was low in our study because the vast majority of samples analyzed were from species that are uncommon hosts for these parasites. Since molecular studies showed that 89% of Polychromophilus-positive samples in Brazil were from Myotis species, further studies are needed to confirm their host specificity and to determine if Myotis spp. are the primary hosts for Polychromophilus in the Neotropics.

The bat Noctilio albiventris has a wide geographic distribution, occurring practically throughout Latin America and almost the entire Brazilian territory. It has an insectivorous diet and is always related to humid forest habitats and environments close to rivers, lakes, or coastal marine habitats [33], making this species more susceptible to parasitic diseases transmitted by vectors available in the environment. Moreover, its involvement with dipteran ectoparasites has not been shown [33], reinforcing the possibility of transmission of Haemoproteidae by ceratopogonid dipterans of the genus Culicoides, known vectors of Haemoproteus (Parahaemoproteus) spp. in birds, as well as Hepatocystis in bats [3,7].

The generalist feeding preferences of vector species could provide opportunities for cross-species transmission of Haemoproteus between avian and bat hosts. In this case, the Haemosporida sp. parasite detected in Bat17 likely represents an abortive spill-over infection [3]. In fact, detecting DNA in the blood without the demonstration of parasites in blood smears does not necessarily indicate successful infection, being plausible that its development cannot be completed in bats.

The Haemosporida sp. sequence described here, with the closest sequence identity of 94% with Haemoproteus (Parahaemoproteus) minutus, is insufficient to identify this parasite as any of those previously described in bats or other animals. However, if this finding is not a spill-over, the parasite sequence position in the phylogenetic tree points to a parasite of the Haemoproteidae family. In fact, the Haemoproteidae family harbors genera of haemosporidian parasites that are exclusive to bats, such as Johnsprentia and Sprattiella, which have not been analyzed molecularly yet, and sequences are lacking for comparison.

A combination of morphological evaluation and molecular studies are needed to conclude and further describe the Polychromophilus parasite lineage, as well as the Haemosporida sp. found in Brazilian bats. Nevertheless, these results confirm the importance of studying these neglected haemosporidian parasites in bats in Brazil.