Introduction
Filariasis is defined as a group of diseases caused by parasitic nematodes belonging to the superfamily Filarioidea, which is classified into the families Filariidae and Onchocercidae, which are considered a relatively small group within the extensive phylum Nematoda (Anderson, Reference Anderson2000). Filarioid nematodes are transmitted by mosquitoes (Culicidae) as well as by black flies (Simuliidae) and other blood-feeding arthropods (Ludlam et al. Reference Ludlam, Jachowski and Otto1970; Anderson, Reference Anderson2000; McCall et al. Reference McCall, Genchi, Kramer, Guerrero and Venco2008; Lefoulon et al. Reference Lefoulon, Giannelli, Makepeace, Mutafchiev, Townson, Uni, Verocai, Otranto and Martin2017). In their vertebrate definitive hosts, these parasites produce microfilariae that can be found in the bloodstream, cutaneous tissues, or mucous membranes, becoming available for the infection of new competent arthropod vectors (Anderson, Reference Anderson2000; Otranto et al. Reference Otranto, Dantas-Torres and Breitschwerdt2009, Reference Otranto, Dantas-Torres, Brianti, Traversa, Petrić, Genchi and Capelli2013). Various genera and species of filarioid nematodes are known to have relevant medical and veterinary importance (Morales-Hojas et al. Reference Morales-Hojas, Post, Shelley, Maia-Herzog, Coscarón and Cheke2001). In humans, they are the primary etiological agents of lymphatic filariasis and river blindness, among other diseases (Anderson, Reference Anderson2000).
The genus Dirofilaria includes more than 20 species associated with mammalian definitive hosts, and most species use mosquitoes as intermediate hosts (Anderson, Reference Anderson2000). However, the most relevant species in veterinary medicine are Dirofilaria immitis and Dirofilaria repens. The former is responsible for causing a cardiopulmonary disease, commonly known as heartworm disease, which may lead to congestive right heart failure, potentially resulting in death. In contrast, in other parts of the world, such as the Middle East and particularly in Eastern European countries, Dirofilaria repens is the most endemic parasitic nematode (McCall et al. Reference McCall, Genchi, Kramer, Guerrero and Venco2008; Capelli et al. Reference Capelli, Genchi, Baneth, Bourdeau, Brianti, Cardoso, Danesi, Fuehrer, Giannelli, Ionică, Maia, Modrý, Montarsi, Krücken, Papadopoulos, Petrić, Pfeffer, Savić, Otranto, Poppert and Silaghi2018; Genchi and Kramer, Reference Genchi and Kramer2020). This species causes subcutaneous infections characterized by the presence of nodules (Albanese et al. Reference Albanese, Abramo, Braglia, Caporali, Venco, Vercelli, Ghibaudo, Leone, Carrani, Giannelli and Otranto2013). Although less pathogenic, these infections seem to have a higher zoonotic potential compared to those caused by D. immitis (Genchi and Kramer, Reference Genchi and Kramer2017). In Brazil, several species within the genus Dirofilaria have been reported, including D. immitis (Dantas-Torres et al. Reference Dantas-Torres, Figueredo, Sales, Miranda, Alexandre, Silva, Silva, Valle, Ribeiro, Otranto, Deuster, Pollmeier and Altreuther2020; Barbosa et al. Reference Barbosa, Nava, Neto, Dias, Silva, Mesquita, Sampaio, Barros, Farias, Silva, Crainey, Tadei, Koolen and Pessoa2023; Zanfagnini et al. Reference Zanfagnini, Bento, Malavazi, Souza and Pacheco2024), often associated with domestic and wild carnivores; Dirofilaria incrassata (Vicente et al. Reference Vicente, Rodrigues, Gomes and Pinto1997), D. repens (Noronha et al. Reference Noronha, Vicente and Pinto2002; Moraes et al. Reference Moraes, Silva, Magalhães-Matos, Albuquerque, Tebaldi, Mathias and Hoppe2017), Dirofilaria spectans (Vicente et al. Reference Vicente, Rodrigues, Gomes and Pinto1997; Noronha et al. Reference Noronha, Vicente and Pinto2002) and Dirofilaria striata (Lentz and Freitas, Reference Lentz and Freitas1937; Vicente et al. Reference Vicente, Rodrigues, Gomes and Pinto1997).
The genus Brugia includes around 10 species associated with mammal definitive hosts and mosquito vectors (Xie et al. Reference Xie, Bain and Williams1994). Out of these, Brugia malayi and B. timori are the most significant to public health, serving as the primary agents of lymphatic filariasis in South and Southeast Asia, as well as historically in Oceania. Infections by these parasites can lead to various clinical manifestations, particularly affecting the lymphatic system, such as lymphedema, hydrocele, fever, chills, lymphadenitis, skin ulcers, pain and tenderness (Anderson, Reference Anderson2000; Tan et al. Reference Tan, Fong, Mahmud, Muslim, Lau and Kamarulzaman2011; Gordon et al. Reference Gordon, Jones and McManus2018). Although cases of Brugia-induced lymphatic filariasis have been reported in North America, there is evidence suggesting underreporting (Elenitoba-Johnson et al. Reference Elenitoba-Johnson, Eberhard, Dauphinais, Lammie and Khorsand1996). In contrast, in Brazil, the disease in humans is primarily caused by Wuchereria bancrofti, phylogenetically closely related to Brugia (GOV.BR – https://www.gov.br/saude/pt-br/assuntos/saude-de-a-a-z/f/elefantiase). In the Americas, the following species from the genus Brugia have been reported: B. beaveri, B. guyanensis, B. lepori and other not fully characterized species (Ash and Little Reference Ash and Little1964; Orihel, 1964; Schlesinger et al. Reference Schlesinger, Dubois and Beaver1977; Eberhard Reference Eberhard1984; Beaver and Wong, Reference Beaver and Wong1988; Eberhard et al. Reference Eberhard, Telford III and Spielman1991; Moraes et al. Reference Moraes, Silva, Magalhães-Matos, Albuquerque, Tebaldi, Mathias and Hoppe2017; Kulpa et al. Reference Kulpa, Goldsmith and Verocai2023).
Understanding the health risks faced by wild animals is essential for preventing the emergence and spread of parasitic diseases that may threaten their conservation. Furthermore, other factors associated with anthropization, such as poaching, vehicle collisions on highways and habitat loss or destruction due to the expansion of agricultural and livestock activities, also pose significant threats (Sanderson et al. Reference Sanderson, Redford, Chetkiewicz, Medellin, Rabinowitz, Robinson and Taber2002; Rodden et al. Reference Rodden, Rodrigues, Bestelmeyer, Sillero-Zubiri, Hoffmann and Macdonald2004; Michalski and Peres, Reference Michalski and Peres2005; Paula et al. Reference Paula, Medici and Morato2008). This is particularly relevant in the neotropical region, where significant gaps in knowledge exist regarding the diversity of onchocercid species due to the wide range of available hosts and the diversity of biomes. Therefore, the present study aimed to investigate the molecular occurrence of filarioid nematodes in wild mammals of the Amazon Rainforest, Cerrado and Pantanal biomes in Brazil.
Materials and methods
Sampled species and study area
Six wildlife species were sampled from 2016 to 2023, overall, including 71 jaguars (Panthera onca; Carnivora; Felidae), 2 pumas (Puma concolor Carnivora; Felidae), 8 giant anteaters (Myrmecophaga tridactyla; Xenarthra; Myrmecophagidae), 5 maned wolves (Chrysocyon brachyurus; Carnivora; Canidae), 5 crab-eating foxes (Cerdocyon thous; Carnivora; Canidae) and 2 ocelots (Leopardus pardalis; Carnivora; Felidae). Whole blood with EDTA was collected from a total of 93 animals, of which 57 were free-living and 36 were kept in captivity. Animal capture was conducted with the assistance of a licensed and registered veterinarian technical team with the Regional Council of Veterinary Medicine and Animal Health. Jaguars and pumas were captured using foot-snare traps (Araujo et al. Reference Araujo, Deco-Souza, Morato, Crawshaw, Silva, Jorge-Neto, Csermak-jr, Bergo, Kantek, Miyazaki, Beisiegel, Tortaro, May-Junior, Silva, Leuzinger, Salomão-Jr and Paula2021), and the giant anteaters were captured by active search, followed by restraint with a catcher pole. Animals were sedated using the association of medetomidine (0·08–0·1 mg kg−1; IM) and ketamine (5 mg kg−1; IM) (Araujo et al. Reference Araujo, Deco-Souza, Morato, Crawshaw, Silva, Jorge-Neto, Csermak-jr, Bergo, Kantek, Miyazaki, Beisiegel, Tortaro, May-Junior, Silva, Leuzinger, Salomão-Jr and Paula2021; Araújo et al. Reference Araújo, Jorge-Neto, Salmão-Júnior, Silva, Zanella, Csermak-Júnior, Francisco, Deco-Souza and Pizzutto2023). The samples were collected in the municipalities of Rio Negro (19°44’19.7”S 54°98’11.8”W), Bodoquena (20°54’09.1”S 56°71’34.2”W), Aquidauana (20°45’18.2’’S 55°78’21.0’’W), Corumbá (19°00’80.3’’S 57°64’35.5’’W), Miranda (19°52’40.3’’S 57°02’90.1’’W), in the state of Mato Grosso do Sul (Cerrado and Pantanal biomes), as well as in the states of Amapá, on Maracá-Jipioca Island (2°04’92.3’’S 50°45’06.6’’W) (Amazon biome), and Goiás at the Onça-Pintada Institute (17°89’19.3’’S 53°00’81.4’’W) and in Emas National Park (17°92’18.4’’S 53°00’48.7’’W) (Cerrado biome) (Figure 1). Whole blood samples were stored at –80 °C until further processing.
Figure 1. Distribution of sampled wild mammals captured by the state of Brazil (AP, Amapá, GO, Goiás and MS, Mato Grosso do Sul). The map also depicts the different biomes from where samples were collected.
Molecular analyses
Genomic DNA was extracted from whole blood samples following a previously published protocol (Araujo et al. Reference Araujo, Ramos, Luiz, Péres, Oliveira, Souza and Russi2009). Subsequently, conventional PCR (cPCR) was performed on the extracted DNA samples to amplify the mammalian endogenous gene GAPDH (glyceraldehyde-3-phosphate dehydrogenase) (Kojima et al. Reference Kojima, Naraba, Miyamoto, Beppu, Aoki and Kawai2004) to verify the presence of amplifiable DNA. Samples that tested positive in this cPCR assay for mammalian endogenous genes were subjected to screening by cPCR assays for the detection of filarioid nematodes, through amplification of the 12S rDNA gene (Casiraghi et al. Reference Casiraghi, Bain, Guerrero, Martin, Pocacqua, Gardner, Franceschi and Bandi2004) and characterization with the cytochrome c oxidase subunit 1 (COI) gene (Casiraghi et al. Reference Casiraghi, Anderson, Bandi, Bazzocchi and Genchi2001).
All cPCR assays were performed using 2 μL of DNA (approximately 50 ng) in a mixture containing 1·5 U Taq DNA Polymerase (Ludwig, Alvorada, Rio Grande do Sul, Brazil), PCR buffer (10X PCR buffer – 10 mM Tris-HCl, pH 8·3, 50 mM KCl), 0·5 mM of deoxynucleotides (dATP, dTTP, dCTP and dGTP) (Cellco, São Carlos, São Paulo, Brazil), 1·5 mM of Magnesium Chloride (Ludwig, Alvorada, Rio Grande do Sul, Brazil), 0·5 μM of each primer (Invitrogen®, Carlsbad, California, USA), and ultrapure sterilized water (Ludwig, Alvorada, Rio Grande do Sul, Brazil) q.s.p. 25 μL. DNA samples from dogs infected with Cercopitifilaria bainae, from the study by Soares et al. (Reference Soares, Parolin, Mateus, Figueiredo, Rodrigues, Oliveira, Silva, Bacha, Ramos, Otranto, Tutija and Ramos2020), were used as positive controls. Ultrapure sterilized water (Ludwig, Alvorada, Rio Grande do Sul, Brazil) was used as a negative control.
The amplified products were separated by agarose gel electrophoresis 2% agarose gel stained with GelRed® Nucleic Acid Gel Stain (Biotium, San Francisco, California, USA), using an electric current of 100 V/150 mA for 50 min. Agarose gels were exposed to ultraviolet light (Easy doc 100, Bio Agency, São Paulo, São Paulo, Brazil).
Purification of amplicons and sequencing
PCR products were purified using ExoSAP-IT PCR Product Cleanup Reagent (Applied Biosystems™) and sequenced using the BigDye™ Terminator v3.1 Cycle Sequencing Kit (Thermo Fisher Scientific™, Waltham, MA, USA) and ABI PRISM 3130 DNA Analyzer (Applied Biosystems™, Foster City, CA, USA) (Sanger et al. Reference Sanger, Nicklen and Coulson1997), with the same primer pairs from cPCR.
The sequences obtained were subjected to quality screening using Geneious software (Kearse et al. Reference Kearse, Moir, Stones-Havas, Cheung, Sturrock, Buxton, Cooper, Markowitz, Duran, Thierer, Ashton, Meintjes and Drummond2012) to assess the quality of the electropherograms and obtain consensus sequences from the alignment of the direct and inverse sequences. Subsequently, the BLASTn analysis tool (Altschul et al. Reference Altschul, Gish, Miller, Myers and Lipman1990) was used to analyse the nucleotide sequences by comparing them with sequences previously deposited in the GenBank database (Benson et al. Reference Benson, Cavanaugh, Clark, Karsch-Mizrachi, Lipman, Ostell and Sayers2017).
Phylogenetic analyses
After an initial analysis with BLAST, a database was created containing all available sequences from GenBank for the sequenced regions and taxonomic groups (search conducted in October 2023). Exploratory alignments by genus were performed to exclude highly divergent sequences (likely identification errors) or incomplete sequences from subsequent analyses. Multiple sequence alignments for each locus were performed using MAFFT v.7 (Katoh and Standley, Reference Katoh and Standley2013), with the auto option, followed by manual inspection. The best nucleotide substitution model was selected using the Bayesian Information Criterion (BIC) in the jModelTest v.2.1.7 software (Darriba et al. Reference Darriba, Taboada, Doallo and Posada2012).
Phylogenetic trees were reconstructed using maximum likelihood analysis (ML) and Bayesian inference (BA). ML analyses were performed using RAxML (Stamatakis Reference Stamatakis2014), utilizing the GTRGAMMA model, with support values estimated from 1000 bootstrap pseudoreplicates. For Bayesian analysis (BA), we used the BEAST 2.6.7 package (Bouckaert et al. Reference Bouckaert, Vaughan, Barido-Sottani, Duchêne, Fourment, Gavryushkina, Heled, Jones, Kühnert, Maio, Matschiner, Mendes, Müller, Ogilvie, Plessis, Popinga, Rambaut, Rasmussen, Siveroni, Suchard, Wu, Xie, Zhang, Stadler and Drummond2019), employing the HKY substitution model, the strict molecular clock and the Yule Model. Our analysis consisted of 2 independent Markov Monte Carlo Chains (MCMC) runs, each covering 20 000 000 generations and sampling every 10 000. JmodelTest, RAxML and Beast analyses were performed on CIPRES Science Gateway (Miller et al. Reference Miller, Pfeiffer and Schwartz2010). The convergence was confirmed by inspecting the log probability in the Tracer 1.7 program (Rambaut et al. Reference Rambaut, Drummond, Xie, Baele and Suchard2018). Estimated effective sampling size values exceeding 200 indicated that the convergence was achieved. The maximum clade credibility tree was estimated with TreeAnnotator, part of the BEAST package. As burn-in, 10% of the first trees obtained were discarded. The trees were visualized and edited using the FigTree v1.4.4 software (http://tree.bio.ed.ac.uk/software/figtree/). Branches with bootstrap support ≥75% in the ML and a posterior probability ≥0·95 in BA analysis were considered strongly supported.
Pairwise ML distances (substitutions/site) were estimated in PAUP 4.0b10 (Swofford, Reference Swofford2002), with base frequencies calculated in JMODELTEST2 program (Guindon and Gascuel Reference Guindon and Gascuel2003; Darriba et al. Reference Darriba, Taboada, Doallo and Posada2012).
Results
Occurrence of filarioid nematodes in free-living and captive wildlife
Out of the 93 samples analysed, 90 (96·77%) tested positive for the mammalian endogenous gene GAPDH via cPCR. In the screening assays targeting filarioid nematodes, based on cPCR for the 12S rDNA gene, 14·44% (13/93) yielded positive results. These included 9·86% of jaguars (7/71), 50% of puma (1/2), 12·5% of giant anteaters (1/8), 50% of ocelots (1/2) and 60% of crab-eating foxes (3/5). Among the samples that tested positive for the 12S rDNA gene, 46% (6/13), including 42·85% of the jaguars (3/7), 100% of the puma (1/1), 100% of the giant anteaters (1/1) and 33·33% of the crab-eating foxes (1/3), also tested positive for the COI gene (Table 1). It is important to note that the high percentages observed in certain species are due to the low sample sizes. Only free-ranging animals tested positive for filarioid nematodes.
Table 1. List of animals that tested positive in the conventional PCR assays for the 12S rDNA and COI genes, along with information regarding identification (ID), species, sex, age group, location and sequence
GO, Goiás; MA, Maranhão; MS, Mato Grosso do Sul.
Exploratory sequence analyses
All 12S rDNA-amplified samples yielded high-quality sequences; however, we encountered difficulties in obtaining high-quality sequences for the COI region under the tested conditions. All sequences were deposited in the GenBank database and assigned the following accession numbers: OR852676-87, PQ699718 and PQ699178. In the results of the identity analyses conducted by BLASTn, the hit with the highest percentage of identity was considered for filarioid detection based on the 12S rDNA target gene (Table 2).
Table 2. Percentage of BLASTn-associated identity of sequences of the family Onchocercidae detected in free-ranging and wild-caught wild animals in Brazil
Phylogenetic analyses of 12S rDNA and COI genes
Based on the 12S rDNA gene sequences of the Onchocercidae family, the phylogenetic analyses grouped our sequences into 4 distinct lineages (Figure 2, Figure S1). Group Ia formed a sister clade to B. pahangi, while Group 1b formed a sister clade to Malayfilaria sofiani. The anteater sequence (Group Ic), while closely related to those in Group I, was more divergent and positioned as a sister group to this larger clade containing species within different genera: Malayfilaria (M. sofiani), Wuchereria (W. bancrofti) and Brugia (B. timori, B. malayi and B. pahangi).
Figure 2. Bayesian phylogenetic inference of filarioid nematodes detected in different wild mammals based on the 12S rDNA gene. The outgroup used was Filaria martis. Complete list of accession numbers found in Supplementary File 2.
All sequences from Group II clustered within the clade of D. immitis and D. striata (Figure 2). This group includes sequences from crab-eating foxes, an ocelot, and a puma. Also in Group II, 1 sample of puma (OR852682 #59) formed a sister clade with D. striata, represented by a sequence generated from a specimen isolated from a domestic cat from Florida, USA (MN635456). This puma Dirofilaria isolate appeared separate from the other puma sequences, which belong to Groups Ia and Ib.
Pairwise ML distances reinforced the patterns observed in the phylogenetic trees (Table 3).
Table 3. Pairwise maximum likelihood distances among 12S sequences of the family Onchocercidae, detected in free-living and captive wild animals in Brazil
Discussion
Filarioid nematodes belonging to different genera were detected in samples from 5 out of 6 mammalian species. Based on the data obtained in this study, the presence of Dirofilaria species with zoonotic potential was recorded in wild cats. Using the mitochondrial 12S rRNA gene as a molecular marker, DNA of Dirofilaria was detected in both an ocelot and a puma. However, the phylogenetic position of the sequence obtained from the ocelot could not be clearly determined using only the 12S marker. Despite this limitation, the scientific literature indicates that parasites of the genus Dirofilaria infect a wide range of hosts and have previously been reported in different mammal species in the Americas. The described hosts include the jaguarundi (Herpailurus yagouaroundi), the jaguar (P. onca), several species of sloths (Choloepus hoffmanni, Bradypus variegatus and B. tridactylus) and the maned wolf (C. brachyurus) (Diaz-Ungria, Reference Diaz-Ungria1973; Eberhard Reference Eberhard1978a, Reference Eberhard1978b; Trotti et al. Reference Trotti, Pampiglione and Rivasi1997; Noronha et al. Reference Noronha, Vicente and Pinto2002; Deem et al. Reference Deem, Bronson, Angulo, Acosta, Murray, Robbins, Giger, Rothschild and Emmons2012; Fagundes-Moreira et al. Reference Fagundes-Moreira, Bezerra-Santos, May-Junior, Berger, Baggio-Souza, Souza, Bilhalva, Reis, Wagner, Peters, Favarini, Albano, Sartorello, Rampim, Tirelli, Otranto and Soares2024). In contrast, the sequence obtained from the puma clustered in a sister clade to D. striata. Notably, a similar observation was recently reported in Texas, USA, where the use of an alternative molecular marker (COI) led to the identification of D. striata in a specimen of bobcat (Lynx rufus) (Ramos et al. Reference Ramos, Hakimi, Salomon, Busselman, Curtis-Robles, Hodo, Hamer and Verocai2024).
Interestingly, when analysing filarioid sequences that form Group Ia (Figure 2), we observed that 3 jaguar sequences were allocated into a single group, closely related to the B. pahangi. Nevertheless, these may represent a previously uncharacterized species of Brugia. The current knowledge on the biodiversity within the genus Brugia is rather scarce, in particular those distributed across the Americas and associated with wildlife (Moraes et al. Reference Moraes, Silva, Magalhães-Matos, Albuquerque, Tebaldi, Mathias and Hoppe2017; Kulpa et al. Reference Kulpa, Goldsmith and Verocai2023). A thorough molecular characterization of Brugia isolates from different hosts and geographic locations is warranted for a better understanding of Brugia diversity and phylogenetic relationships among species. The occurrence of Brugia species in the Americas highlights their broad geographical distribution and the diversity of hosts for these filarioid nematodes. In North America, Brugia sp. has been reported in a dog (Canis lupus familiaris) in Canada (Kulpa et al. Reference Kulpa, Goldsmith and Verocai2023). In the USA, Brugia beaveri is the most prevalent species, with infection records in raccoons (Procyon lotor), bobcats and possibly minks (Neogale vison). This wide variety of hosts suggests that B. beaveri has a remarkable ability to adapt to different ecological niches (Ash and Little, Reference Ash and Little1964; Beaver and Wong, Reference Beaver and Wong1988). Additionally, the identification of Brugia lepori in hares reinforces the hypothesis that lagomorphs play a significant role in the life cycle of these parasites in the region (Schlesinger et al. Reference Schlesinger, Dubois and Beaver1977; Eberhard et al. Reference Eberhard, Telford III and Spielman1991). In South America, the identification of Brugia guyanensis in ring-tailed coatis (Nasua nasua vittata) from Guyana (Orihel 1964), as well as the detection of Brugia sp. in ring-tailed coatis from Brazil (Moraes et al. Reference Moraes, Silva, Magalhães-Matos, Albuquerque, Tebaldi, Mathias and Hoppe2017), indicates that the region harbours a still underexplored diversity of filarioids.
When analysing sequences forming Group Ib (Figure 2), based on the 12S rDNA gene, the clustering of 4 Onchocercidae sequences from jaguars was observed, positioned as a sister group to the M. sofiani, a parasite of a rodent, the treeshrew (Tupaia glis) from Malaysia, closely related to Brugia and Wuchereria (Uni et al. Reference Uni, Udin, Agatsuma, Saijuntha, Junker, Ramli, Omar, Lim, Sivanandam, Lefoulon, Martin, Belabut, Kasim, Halim, Zainuri, Bhassu, Fukuda, Matsubayashy, Harada, Low, Chen, Suganuma, Hashim, Takaoka and Azirun2017). A recent study by Fagundes-Moreira et al. (Reference Fagundes-Moreira, Bezerra-Santos, May-Junior, Berger, Baggio-Souza, Souza, Bilhalva, Reis, Wagner, Peters, Favarini, Albano, Sartorello, Rampim, Tirelli, Otranto and Soares2024) also identified sequences of a filarioid nematode closely related to Malayfilaria in 2 jaguars (OR434083 and OR434084) in Brazil. This study characterized the COI gene, instead of the 12S gene targeted by us. A direct comparison was hindered as the cPCR targeting the COI gene for Group Ib samples was not successful. In light of this, the question arose as to whether these sequences represent the same species, as the samples were collected in the same region, the southern Pantanal of Mato Grosso do Sul.
Similarly, the identification of Group Ic, composed of the filarioid sequence (ID = 95·44% – OR852683 #9), was detected in a giant anteater from the municipality of Rio Negro, Mato Grosso do Sul, Brazil. However, the identification remained inconclusive, as in the phylogenetic analysis, the sequence was positioned separately from the larger clade containing the other sequences from Groups Ia and Ib. The ML distances between Group Ic and its closest lineages suggest a probable association with another genus. Therefore, the assignment of Group Ic to a specific genus also remains inconclusive.
Within the sequences obtained from puma, one (OR852682 #59) is substantially distinct from the others (Groups Ia and Ib), positioned as a sister group of D. striata. This pattern was confirmed by sequencing the COI region, which revealed a 6·3% divergence between the sequence of sample OR852682 #59 and that of D. striata found in a domestic cat from Florida, USA. This level of variation is notably high for intraspecific COI differences in Dirofilaria and other filarioid genera (Lefoulon et al. Reference Lefoulon, Giannelli, Makepeace, Mutafchiev, Townson, Uni, Verocai, Otranto and Martin2017; Kulpa et al. Reference Kulpa, Lefoulon, Beckmen, Allen, Malmberg, Crouse, Thompson, Benedict, Goldsmith, McCarthy, Jones, Yabsley, Crum, Kutz and Verocai2025). For instance, based on the available COI sequences in GenBank, the intraspecific divergence in D. immitis specimens does not exceed 1·5%, and in D. repens, it is limited to 1·1%. Hence, it is probable that the puma OR852682 #59 sample belongs to another Dirofilaria species closely related to D. striata, which is not yet represented in GenBank or could potentially represent a novel species.
Finally, Group II comprises samples from different hosts, including a canid (C. thous) and a felid (L. pardalis). This group comprises closely related and likely conspecific sequences, forming a sister clade to D. striata/ OR852682 #59 sequences. The average ML distance between Group II and D. striata was 0·0548 (0·0395–0·0934), while the average distance between Group II and puma 59 was 0·0568 (0·0532–0·0621). Therefore, these results suggest that the samples of Cerdocyon and Leopardus share the same Dirofilaria species, which is different from the one that infected the puma (Accession number: OR852682 #59).
Throughout the Americas, several species of the genus Dirofilaria have been reported from carnivore hosts through morphological, serological and molecular analyses. The canine heartworm, D. immitis has been found in a variety of wild carnivores, including gray foxes (Urocyon cinereoargenteus) (Simmons et al. Reference Simmons, Nicholson, Hill and Briggs1980; Carlson and Nielsen Reference Carlson and Nielsen1983; Hernández-Camacho et al. Reference Hernández-Camacho, Pineda-López, Guerrero-Carrillo, Cantó-Alarcón, Jones, Moreno-Pérez, Mosqueda-Gualito, Zmora-Ledesma and Camacho-Macías2016), raccoons (Procyon lotor) (Snyder et al. Reference Snyder, Hamir, Hanlon and Rupprecht1989; Ramos et al. Reference Ramos, Hakimi, Salomon, Busselman, Curtis-Robles, Hodo, Hamer and Verocai2024), black bears (Ursus americanus) (Crum et al. Reference Crum, Nettles and Davidson1978), ring-tailed coatis (Moraes et al. Reference Moraes, Silva, Magalhães-Matos, Albuquerque, Tebaldi, Mathias and Hoppe2017, Reference Moraes, Pollo and Hoppe2022), oncilla (Leopardus tigrinus) (Filoni et al. Reference Filoni, Pena, Gennari, Cristo, Torres and Catão-Dias2009), coyotes (Canis latrans) (Zinck et al. Reference Zinck, Priest, Shutler, Boudreau and Lloyd2021; Sobotyk et al. Reference Sobotyk, Nguyen, Negrón, Varner, Saleh, Hilton, Tomeček, Esteve-Gasent and Verocai2022), small-clawed Asian otters (Aonyx cinereus) (Upton et al. Reference Upton, Sobotyk, Edwards and Verocai2022) and wild felids (P. onca, L. pardalis and Leopardus guttulus) (Fagundes-Moreira et al. Reference Fagundes-Moreira, Bezerra-Santos, May-Junior, Berger, Baggio-Souza, Souza, Bilhalva, Reis, Wagner, Peters, Favarini, Albano, Sartorello, Rampim, Tirelli, Otranto and Soares2024). Dirofilaria cancrivori has been reported in crab-eating raccoon (Procyon cancrivorus) (Eberhard Reference Eberhard1978b), and Dirofilaria incrassata from the subcutaneous tissues of both ring-tailed and crab-eating raccoons (Vicente et al. Reference Vicente, Rodrigues, Gomes and Pinto1997; Guimarães et al. Reference Guimarães, Barros, Saddi, Cardoso, Vasconcelos and Ramos2023), but no molecular data are currently available for this species. Another Dirofilaria species has been reported from the subcutaneous tissue of coatis, but questionably diagnosed as D. repens, a zoonotic species normally found in the Old World (Noronha et al. Reference Noronha, Vicente and Pinto2002; Capelli et al. Reference Capelli, Genchi, Baneth, Bourdeau, Brianti, Cardoso, Danesi, Fuehrer, Giannelli, Ionică, Maia, Modrý, Montarsi, Krücken, Papadopoulos, Petrić, Pfeffer, Savić, Otranto, Poppert and Silaghi2018). While (N. nasua) Dirofilaria lutrae has been reported in the North American river otter (Lontra canadensis) (Orihel, 1965; Swanepoel et al. Reference Swanepoel, Cleveland, Olfenbuttel, Dukes, Brown, Brown, Surf, Tumlison and Yabsley2018), Dirofilaria spectans was found parasitizing the lungs and blood vessels of tayras (Eira barbara) (Noronha et al. Reference Noronha, Vicente and Pinto2002), Neotropical otters (Lontra longicaudis) (Noronha et al. Reference Noronha, Vicente and Pinto2002) and giant otters (Pteronura brasiliensis) (Vicente et al. Reference Vicente, Rodrigues, Gomes and Pinto1997). The felid-associated D. striata was observed parasitizing margay cats (Leopardus wiedii) and pumas (Puma concolor) (Lentz and Freitas Reference Lentz and Freitas1937; Forrester et al. Reference Forrester, Conti and Belden1985; Lamm et al. Reference Lamm, Roelke, Greiner and Steible1997; Vicente et al. Reference Vicente, Rodrigues, Gomes and Pinto1997), as well as bobcats (Orihel and Ash, Reference Orihel and Ash1964; Miller and Harkema, Reference Miller and Harkema1968; Ramos et al. Reference Ramos, Hakimi, Salomon, Busselman, Curtis-Robles, Hodo, Hamer and Verocai2024). Finally, Dirofilaria tenuis was detected in raccoons (P. lotor) in North America (Sauerman and Nayar, Reference Sauerman and Nayar1985; Isaza and Courtney, Reference Isaza and Courtney1988; Telford and Forrester, Reference Telford and Forrester1991; Richardson et al. Reference Richardson, Owen and Snyder1992; Pung et al. Reference Pung, Davis and Richardson1996; Hernández-Núñez et al. Reference Hernández-Núñez, Vidal-Martínez and Aguirre-Macedo2024), which has been recently molecularly characterized. The lack of molecular data in many of these studies presents a challenge, particularly in light of recent revisions of the taxonomic framework for filarioids (Ferri et al. Reference Ferri, Barbuto, Bain, Galimberti, Uni, Guerrero, Ferté, Bandi, Martin and Casiraghi2009; Lefoulon et al. Reference Lefoulon, Bain, Bourret, Junker, Guerrero, Cañizales, Kuzmin, Satoto, Cardenas-Callirgos, Lima, Raccurt, Mutafchiev, Gavotte and Martin2015). This limitation has contributed to a gap in such studies in the Americas, as well as a paucity of records in Brazilian wild animals.
The genus Dirofilaria can also infect non-carnivorous mammals, with records of the species Dirofilaria acutiuscula and D. striata associated with the family Tayassuidae (Molin Reference Molin1858). Furthermore, it is relevant to highlight the role of the superorder Xenarthra in infections caused by filarioid nematodes. The literature has described the occurrence of Dirofilaria freitasi, D. incrassata, and Dirofilaria macrodemos in the 3-toed sloth (Bradypus tridactylus) (Sandground, Reference Sandground1938; Mendonça, Reference Mendonça1948), as well as Dirofilaria panamensis in the two-toed sloth (Choloepus hoffmanni) (Sandground Reference Sandground1938; Eberhard Reference Eberhard1978a; Trotti et al. Reference Trotti, Pampiglione and Rivasi1997). Currently, there is no molecular data for any of these Dirofilaria species associated with xenarthran hosts. The diversity of hosts that a parasite can infect increases the risk of it reaching new hosts, including domestic animals and humans.
According to the IUCN Red List (www.iucnredlist.org), the jaguar and maned wolf are near threatened, the giant anteater is vulnerable, and the crab-eating fox and the ocelot are of least concern. Considering the free-living habits of the animals and the data from this study, it is likely that these animals serve as reservoirs of pathogens, given the lack of knowledge about the arthropod vectors responsible for transmitting filarioid nematode third-stage larvae to wild mammalian definitive hosts in different biomes across Brazil. Various biological and logistic factors impact studies on parasite biodiversity, including filarioid nematodes, of wildlife in Brazil, including the remoteness of field sites, opportunistic sampling, funding allocation and conservation status of multiple host species. Additionally, the scarcity of molecular data available for many of the known, valid species within the genus Dirofilaria further hinders species-level characterization of taxa found through molecular screening of archival blood and tissue samples, in the absence of adult specimens or microfilariae in fresh blood samples. Therefore, studies using integrative taxonomy are essential to confirm the taxonomic status of these parasites and describe potential new taxa.
Conclusion
Jaguars, puma, giant anteaters, crab-eating foxes and ocelots may play an important role as reservoirs for uncharacterized filarioid nematode species belonging to the family Onchocercidae. Currently, the impact of those infections on host health and their potential to infect companion animals and humans remains unknown. Furthermore, the fact that jaguars are infected by 2 different species of filarioid nematode species, likely belonging to distinct genera, highlights the complexity and significance of these parasites in the local fauna. Given the cryptic biodiversity of filarioid nematodes associated with Neotropical mammals, further studies are needed to assess their host and vector associations, geographic distribution and to elucidate their life cycle and pathogenicity.
Supplementary material
The supplementary material for this article can be found at https://doi.org/10.1017/S0031182025101042.
Data availability statement
Data supporting the conclusions of this study are included in the article. Generated sequences were submitted to the GenBank database under accession numbers: OR852676-87, PQ699718 and PQ699178.
Acknowledgements
We would like to express our sincere gratitude to everyone who contributed to the collection and provision of samples, and to the Reprocon Institute. This project would not have been possible without the collaboration and commitment of everyone involved.
Author contributions
The study was conceived and designed by M.S.S. and C.A.N.R. Samples were provided by M.C.C.d.S., G.R.d.A., A.C.C.-J. and T.d.D.S.A. Sample preparation and molecular analysis were carried out by M.S.S. and H.P.K.C. The paper was written by M.S.S., A.P.L., G.G.V., L.S.U. and C.A.N.R. All authors have read and agreed to the present version of the manuscript.
Financial support
This study was funded in part by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior – Brasil (CAPES) – Finance Code 001; the Fundação Universidade Federal de Mato Grosso do Sul – UFMS/MEC – Brazil; and the Reprocon Institute. This work was partially supported by CNPq, Conselho Nacional de Desenvolvimento Científico e Tecnológico – Brasil (Processo: 403623/2021-9) and MS Fundação de Apoio ao Desenvolvimento de Ensino, Ciência e Tecnologia (Processo: 83/024.003/2023).
Competing interests
The authors declare none.
Ethical approval
Biological samples from free-ranging and captive wild mammals were collected under the approval of the Ethics Committee on Animal Use (CEUA) and the Brazilian System of Authorization and Information on Biodiversity (SISBIO). Authorizations were granted for research projects conducted by the wildlife conservation and management group at the Federal University of Mato Grosso do Sul (CEUA protocols: 628/2014, 727/2015, 1035/2019, 1221/2022, and 1355/2024; SISBIO authorizations: 46031-4, 57293-3, 68167-1, 72633-8, and 757626), as well as the University of São Paulo (CEUA protocol: 74398-2; SISBIO: 6429170220).
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