Introduction
Trematodes of the family Gorgoderidae Looss, 1899 comprise 3 sub-families: Anaporrhutinae Looss, 1901 (with seven genera infecting elasmobranchs and sea turtles), Degeneriinae Cutmore, Miller, Curran, Bennett & Cribb, Reference Cutmore, Miller, Curran, Bennett and Cribb2013 (with a single genus infecting deep-sea fishes), and Gorgoderinae Looss, 1899 (with four genera infecting marine and freshwater ray-finned fish and amphibians) (Cutmore et al. Reference Cutmore, Miller, Curran, Bennett and Cribb2013). Two- or three-host life cycles have been revealed only for some members of freshwater Gorgoderinae, with bivalves as first intermediate hosts, and insects, snails, crayfish, or amphibian tadpoles as second intermediate hosts (Campbell Reference Campbell, Bray, Gibson and Jones2008). A single record of an unidentified gorgoderid cercaria infecting a marine bivalve Lioconcha castrensis (Linnaeus) has been also reported (Bott and Cribb Reference Bott and Cribb2005).
The Munda round ray Urotrygon munda Gill (Myliobatiformes: Urotrygonidae) is a relatively poorly known species distributed throughout the eastern Pacific from southern Baja California (Mexico) to northern Peru at depths of 4–51 m (Robertson and Allen Reference Robertson and Allen2015), which feeds primarily in shallow soft bottoms on caridean decapods (Flores-Ortega et al. Reference Flores-Ortega, Godínez-Domínguez, González-Sansón, Rojo-Vázquez, Corgos and Morales-Jáuregui2011). According to Caira et al. (Reference Caira, Jensen and Barbeau2022) and Pollerspöck and Straube (Reference Pollerspöck and Straube2023), no parasite species has been reported in this host. Additionally, of the 62 marine trematode species reported from both coasts of Costa Rica, no trematode species have been reported in elasmobranchs (Solano-Barquero et al. Reference Solano-Barquero, Rojas and Cortés2023). Furthermore, the only Anaporrhutinae reported from this area was Plesiochorus cymbiformis (Rudolphi, 1819) Looss, 1901 in the olive ridley sea turtle Lepidochelys olivacea (Eschscholtz) from the Pacific coast of Costa Rica (Santoro and Morales Reference Santoro and Morales2007; Solano-Barquero et al. Reference Solano-Barquero, Rojas and Cortés2023).
During a parasitological survey of fish from the Pacific coast of Costa Rica, some individuals of an anaporrhutine species were found in the gill chambers of the Munda round ray. These proved to represent a morphologically distinct, previously unknown species of Anaporrhutum Brandes in von Ofenheim, Reference von Ofenheim1900, which is described herein, based on morphological, ultrastructural, and molecular characters, and phylogenetic analysis.
Materials and methods
Sample collection
On 07 April 2023, three individuals of the Munda round ray were obtained from off Playa Cuajiniquil (Guanacaste province) on the Pacific coast of Costa Rica using nets at benthic depths ranging from 5 to 10 m. They included two females, with total length (TL) and total weight (TW) of 52 cm and 917 g, and 50.9 cm and 798 g, respectively, and one male, with 23.8 cm TL and 231 g TW. Fish were obtained under the framework of a project of the Centro de Investigación en Ciencias del Mar y Limnología of the University of Costa Rica “Proyecto BioMar -ACG” (see Cortés and Joyce Reference Cortés and Joyce2020), aimed at studying the marine biodiversity of the Pacific coast of Costa Rica (permit no. ACG 019-2023), and a collaborative project between the Stazione Zoologica Anton Dohrn and the University of Costa Rica, aimed at studying the parasite biodiversity of elasmobranchs in the country.
Fish were refrigerated (4°C) and transferred to the laboratory, where they were studied within 6 h of fishing. During necropsy, eyes, skin, gills, mouth cavity, digestive tract (stomach and intestine), liver, heart, gonads, visceral cavity, mesenteries, and skeletal muscles of each fish were examined for metazoan parasites under a dissecting microscope (Axio Zoom V16, Zeiss, Switzerland) using the methods described in Santoro et al. (Reference Santoro, Bellisario, Tanduo, Crocetta and Palomba2022, Reference Santoro, Bellisario, Fernández-Álvarez, Crocetta and Palomba2023). When parasites were observed they were collected and washed in physiological saline solution and preserved in 70% ethanol for subsequent morphological and molecular analyses.
Morphological study
For light microscopy, trematodes were stained with Mayer’s acid carmine, dehydrated through a graded ethanol series, cleared in methyl salicylate, and mounted in permanent slides in Canada balsam. Measurements (in micrometres) are reported as range values with mean ± standard deviation in parentheses followed by the total number (n) of observations. Measurements were obtained using a compound microscope (Axio Imager M1, Zeiss, Switzerland) and a dissecting microscope (Axio Zoom V16) equipped with the ZEN 3.1 imaging system (Zeiss, Switzerland) Drawings were made with the aid of a XP PEN Deco 02 drawing tablet (Deco, Italy) and Adobe Illustrator and Adobe Photoshop software (Adobe Systems Inc., United States).
For scanning electron microscopy analysis, a specimen was also fixed overnight in 2.5% glutaraldehyde, then transferred to 40% ethanol (10 min), rinsed in 0.1 M cacodylate buffer, postfixed in 1% OsO4 for 2 h, and dehydrated in ethanol series, critical point dried, and sputter-coated with platinum. Observations were made using a JEOL JSM 6700F scanning electron microscope operating at 5.0 kV (JEOL, Italy).
Molecular analysis
For the molecular study, genomic DNA was extracted from a fragment of two 70% ethanol-preserved specimens using the Quick-gDNA Miniprep Kit (Zymo Research, California). The 28S nuclear ribosomal DNA region was amplified using the primers LSU5 (5’-TAGGTCGACCCGCTGAAYTTAAGC-3’) and 1500R (5’-GCTATCCTG AGGGAAACTTCG-3’) (Littlewood Reference Littlewood1994; Cutmore et al. Reference Cutmore, Bennett and Cribb2010); the ITS2 region was amplified using the primers 3S (5’-GGTACC GGTGGATCACGTGGCTAGTG-3’) and ITS2.2 (5’-CCTGGTTAGTTTCTTTTCCTCCGC-3’) (Cribb et al. Reference Cribb, Anderson, Adlard and Bray1998; Cutmore et al. Reference Cutmore, Bennett and Cribb2010). New specific primers also were designed for the 28S region to avoid any nonspecific amplifications; these were Anap28-F (5’-CAATGTGGTGTTYAGGTCGGTCTTC -3’) and Anap28-R (5’-CTGYCGCTCAWTRCYTGGT -3’). Polymerase chain reactions (PCRs) for both 28S and ITS2 regions were performed in a 25 μL volume containing 1.5 μL of each primer 10 mM, 3 μL of MgCl2 25 mM (Promega, Wisconsin), 5 μL of 5× buffer (Promega), 0.6 μL of dNTPs 10 mM (Promega), 0.2 μL of Go-Taq Polymerase (5U/μL) (Promega), and 2 μL of total DNA.
PCR cycling parameters for the 28S region were as follows: an initial 94°C for 2 min, followed by 40 cycles of 94°C for 30 sec, 58°C for 45 sec, 72°C for 2 min, and a final 72°C extension for 10 min. PCR cycling parameters for the ITS2 region were as follows: an initial single cycle of 95°C for 3 min, 45°C for 2 min, 72°C for 90 sec, followed by 4 cycles of 95°C for 45 sec, 50°C for 45 sec, 72°C for 90 sec, followed by 30 cycles of 95°C for 20 sec, 52°C for 20 sec, 72°C extension for 90 sec, and a final 72°C extension for 5 min.
PCR products were purified using Agencourt AMPure XP (Beckman Coulter, United States), following the standard manufacturer-recommended protocol. Clean PCR products were Sanger sequenced from both strands using an Automated Capillary Electrophoresis Sequencer 3730 DNA Analyzer (Applied Biosystems, United States) and the BigDye® Terminator v. 3.1 Cycle Sequencing Kit (Life Technologies, United States). The obtained contiguous sequences were assembled and edited using MEGAX v. 11 (Kumar et al. Reference Kumar, Stecher, Li, Knyaz and Tamura2018). Sequence identity was verified using the Nucleotide Basic Local Alignment Search Tool (BLASTn) (Morgulis et al. Reference Morgulis, Coulouris, Raytselis, Madden, Agarwala and Schaffer2008).
Phylogenetic analysis
Obtained sequences were cleaned and aligned with 28S and ITS2 sequences acquired from other members of the families Gorgoderidae and Allocreadiidae available in Genbank using MEGA10 (Kumar et al. Reference Kumar, Stecher, Li, Knyaz and Tamura2018) and the MUSCLE algorithm (Edgar Reference Edgar2004) (Table 1). Also for the ITS2, sequences of members of family Prostogonimidae were acquired (Table 1). Then, a Bayesian inference (BI) phylogenetic tree was built for the 28S and ITS2 alignments using the BEAST package v2.6 (Drummond and Rambaut Reference Drummond and Rambaut2007) with General Time Reversible with gamma distribution and Hasegawa-Kishino Yano nucleotide substitution models, as based in Bayesian inference criteria. A total of 108 Montecarlo Markov Chains and a burn-in of 10% were estimated. Effective sample sizes larger than 300 were confirmed for each parameter with Tracer v1.7.2 (Rambaut et al. Reference Rambaut, Drummond, Xie, Baele and Suchard2018). Finally, the converged tree was visualized with FigTree v1.4.3 (http://tree.bio.ed.ac.uk/software/figtree/) In addition, a maximum likelihood (ML) phylogenetic tree was reconstructed with the nucleotide substitution models mentioned above and 1000 bootstraps using MEGA10. The phylogenetic tree for the 28S fragment was rooted using Dicrocoelium dendriticum (Rudolphi 1819) Looss 1899 (AF151939), Paracreptotrematina limi Amin & Myer, 1982 (HQ833706), and Degeneria halosauri (Bell, 1887) Campbell 1977 (AY222257) as outgroups, while Provitellus chaometra Wee, Cutmore & Cribb, Reference Wee, Cutmore and Cribb2019 (MK501980) was added as the outgroup for the ITS2 phylogenetic tree.
Results
Description (see Figures 1–3)
ZooBank: urn:Isid:zoobank.org:act:991D0E3E-4C1E-4928-9510-69833DB834E1.
Family. Gorgoderidae Looss, 1899
Subfamily. Anaporrhutinae Looss, 1901
Anaporrhutum mundae Santoro, López-Verdejo, Angulo, Rojas, & Solano-Barquero, 2024 sp. nov. in Santoro, López-Verdejo, Angulo, Rojas, Cortés, Pacheco-Chaves & Solano-Barquero, 2024
Based on six mounted mature specimens studied by optical microscopy and one mature specimen studied by SEM. Body large, flat, oval, 5588–7152 (6624 ± 653.5; n = 7) long, 2782–5022 (3712 ± 904.1; n = 7) wide; length/width ratio 1:0.4–0.7 (0.5 ± 177.1; n = 7). Body divided into small forebody and large hindbody; forebody 1270–1782 (1425 ± 190.9; n = 7) long, representing 18–24% (21 ± 2.5; n = 7) of total body length (Figure 1). Tegument wrinkled, lacking spines. Stylet cavity large, dorsal to the oral sucker on the anterior end of the body; a large penetration gland opening is present on each side of this cavity (Figures 3B, 3D, 3E). Penetration gland openings small, symmetrically arranged ventral to the stylet cavity (Figure 3D). Oral sucker round, 547–705 (636 ± 62.1; n = 7) long, 612–743 (668 ± 48; n = 7) wide (Figures 3A, 3B). Penetration gland openings small, arranged in a ring around the mouth (Figures 3B, 3C). Prepharynx absent. Pharynx 315–410 (357 ± 33.1; n = 7) long, 408–479 (442 ± 28.9; n = 7) wide. Oesophagus short, 76–99 (87 ± 7.8; n = 7) long (straight line from posterior end of pharynx to intestinal bifurcation). Caeca bifurcation in forebody, slightly sinuous, lacking diverticula, terminating near to posterior margin of body (Figure 1). Genital pore median, opening in forebody, posterior to margin of pharynx and at the level of intestinal bifurcation (Figures 2A, 3A, 3F). Ventral sucker round, 547–658 (586 ± 41.9; n = 7) long, 523–742 (606 ± 87.6; n = 7) wide (Figure 3F). Oral sucker width to ventral sucker width ratio 1:0.8–1 (1:0.9 ± 288; n = 7). Testes pre-equatorial, consisting of two opposite groups of round to oval unlobed follicles prevalently intercaecal (Figure 1), right group 7–9 (8 ± 0.8; n = 7) in number, left group 7–11 (8 ± 1.8; n = 7) in number. Follicles highly variable in size, right group 109–186 (157 ± 29.6; n = 20) long, 92–309 (147 ± 54.6; n = 20) wide; left group 114–312 (175 ± 56.2; n = 20) long, 92–251 (138 ± 41.9; n = 20) wide. Seminal vesicle elongate, sinuous. Pars prostatica short. Ejaculatory duct short, opening into genital atrium. Cirrus-sac absent. Ovary pretesticular, median, irregularly shaped, 167–250 (212 ± 33.9; n = 6) long, 115–253 (162 ± 51.9; n = 6) wide (Figure 2B). Large seminal receptacle pre-ovarian, round to oval, 358–488 (436 ±48.9; n = 6) long, 229– 333 (278 ± 41.4; n = 6) wide. Mehlis’s gland globular, surrounding ootype, 86–106 (96 ± 8.7; n = 5) long, 78–112 (90 ± 14.7; n = 5) wide (Figure 2B). Vitellarium tubular, in two opposite, intercaecal, dendritic fields at level of ovary; fields consist of 4–5 main branching, winding tubes with finger-like projections branching from main tubes (Figures 1, 2B); dextral fields 264–703 (517 ± 158.6; n = 5) long, 316–528 (419 ± 87.2; n = 5) wide; sinistral fields 339–524 (438 ± 65.6; n = 6) long, 324–663 (532 ± 123.8; n = 6) wide. Uterus intercaecal, extending posteriorly to level of posterior limit of caeca, before turning extending anteriorly to reach the dorsal level of ventral sucker (Figure 1). Metraterm short, opening into genital atrium. Operculated eggs (from distal region of uterus), 47–66 (56 ± 5.7; n = 20) long, 22–33 (29 ± 2.7; n = 20) wide, with tick wall, 1.6–2.8 (2.2 ± 0.2; n = 20) wide (Figure 2C). Excretory vesicle H-shaped.
Figure 1. Drawing of Anaporrhutum mundae sp. nov., holotype ventrally mounted.
Figure 2. Drawing of Anaporrhutum mundae sp. nov., (A) paratype, ventral view of terminal genitalia; (B) holotype, ventral view of ovarian complex; (C) egg.
Figure 3. Scanning electron microscopy of Anaporrhutum mundae sp. nov. ventrally mounted: (A) anterior extremity showing the oral sucker, the genital pore, and the ventral sucker; (B) oral sucker showing the stylet cavity and the small penetration gland openings arranged in a ring around the mouth; (C) magnification of two penetration gland openings around the mouth; (D) magnification of the stylet cavity showing a large penetration gland opening on each side and smaller penetration gland openings ventral to the cavity; (E) magnification of a large penetration gland opening on the side of the stylet cavity; (F) magnification of ventral sucker and genital pore. White arrows indicate the stylet cavity; black arrows indicate the genital pore; white arrowheads indicate the small penetration gland openings; black arrowheads indicate the larger penetration gland openings.
Taxonomic summary
Type-host: Munda round ray Urotrygon munda Gill, 1863 (Myliobatiformes: Urotrygonidae).
Type-locality: Off Playa Cuajiniquil (10°56’04.38″N, 85°42’14.09″W), Guanacaste province, north Pacific coast of Costa Rica (collected on 07 April 2023).
Location in the host: Gill chambers.
Type-material: Holotype (MHNG-PLAT-0157452), two paratypes and an SEM preparation (MHNG-PLAT-0157451) in the Parasite Collection of the Natural History Museum of Geneva in Geneva (Switzerland); two paratypes in the Helminthological Collection of Costa Rica (CHCR-207-1; CHCR-207-2) at the Universidad de Costa Rica, San José (Costa Rica).
Prevalence and intensity: 2 (both females) of 3 individuals infected with 1 and 9 flukes.
Etymology: The new species is named after its host species.
Remarks
Currently, there are seven recognized species of Anaporrhutum. These are: A. albidum Brandes in Ofenheim, 1900, A. mantae Nagaty & Abdel-Aal, 1961, A. stunkardi Tandon, Reference Tandon1969, A. narayani Simha, Rao & Rao, Reference Simha, Rao and Rao1971, A. sinicum Cao, Reference Cao1990, A. torpedoense Khan & Begum, Reference Khan and Begum1991, and A. euzeti Curran, Blend & Overstreet, Reference Curran, Blend, Overstreet, Combes and Jourdane2003 (von Ofenheim Reference von Ofenheim1900; Nagaty and Abdel Reference Nagaty and Abdel Aal1961; Tandon Reference Tandon1969; Simha et al. Reference Simha, Rao and Rao1971; Cao Reference Cao1990; Khan and Begum Reference Khan and Begum1991; Curran et al. Reference Curran, Blend, Overstreet, Combes and Jourdane2003). Anaporrhutum mundae sp. nov. can be distinguished from all these species by having the genital pore opening at level of the intestinal bifurcation (whereas it opens just posterior to caecal bifurcation in all congeneric species). Moreover, it has a different location in the host (gill chambers vs pericardium and body cavity). Additionally, the new species can be distinguished from all other congeners except A. euzeti by having fewer testicular follicles per testis (7–11 vs 6–13 in A. euzeti, 12–25 in A. albidum, 13 in A. sinicum, 19–24 in A. mantae, 25–26 in A. stunkardi, 26 in A. narayani and 23–35 in A. torpedoensis) (von Ofenheim Reference von Ofenheim1900; Nagaty and Abdel Reference Nagaty and Abdel Aal1961; Tandon Reference Tandon1969; Simha et al. Reference Simha, Rao and Rao1971; Cao Reference Cao1990; Khan and Begum Reference Khan and Begum1991).
Anaporrhutum mundae sp. nov. most closely resembles A. euzeti, which was described from the Gulf of California (Mexico) in Myliobatis longirostris Applegate & Fitch, Narcine entemedor Jordan & Starks, Hypanus dipterurus (Jordan & Gilbert) and Urobatis maculatus Garman (Curran et al. Reference Curran, Blend, Overstreet, Combes and Jourdane2003). However, A. mundae sp. nov. is smaller than A. euzeti (5.5–7.1 vs 7.2–17.5 mm long), and it differs from A. euzeti in forebody shape, having a different location of the genital pore and having the vitellaria divided in two transversal tubular dendritic fields in the intercecal field vs follicular vitellaria divided in two opposing bunches medial or ventral to the caeca.
The ultrastructural characters observed in A. mundae sp. nov. using scanning electron microscopy analysis, i.e., a large stylet cavity dorsal to the oral sucker with a large penetration gland opening on each side of the cavity, small penetration gland openings ventral to the stylet cavity, and small penetration gland openings arranged in a ring around the mouth are suspected to be of taxonomic relevance. However, to date, there are no ultrastructural data for other species of Anaporrhutum for comparison.
Molecular and phylogenetic analyses
Two 28S rDNA sequences (899 bp and 901 bp) and two identical ITS2 (458 bp) were generated from two individuals of A. mundae sp. nov. Sequences were deposited in GenBank under the accession numbers PP133643 and PP354907 for the 28S and PP133644 for ITS2. The BLASTn search of the sequences obtained retrieved a percentage of identity of 93.8% and 93.6% for the 28S and 94.2% for ITS2 with the only sequences of the genus available in GenBank (KF013184: 1144 bp, 28S; KF013159: 437 bp, ITS2), corresponding to an unidentified Anaporrhutum sp. found in the brownbanded bamboo shark Chiloscyllium punctatum Müller & Henle in Australia.
Because BI and ML trees had similar topologies, only the BI trees of both 28S (Figure 4) and ITS2 (Figure 5) are shown. BI and ML phylogenetic tree separated 28S sequences according to their families and subfamilies. For instance, both sequences obtained from A. mundae sp. nov. clustered together and with another Anaporrhutum sp. (KF013184). In addition, the subfamily Anaporrhutinae formed three clusters: one with Anaporrhutum spp. and Staphylorchis cymatodes (Johnston, 1913) Travassos, 1922, a second group with Plesiochorus spp. Looss, 1901, and a third one with Nagmia spp. Nagaty, 1930. The subfamily Gorgoderinae included sequences available for Phyllodistomum spp. Braun, 1899, Gorgodera cygnoides (Zeder, 1800) Looss, 1899 and Cercaria duplicata (accepted as Phyllodistomum folium (Olfers, 1816) Braun, 1899). Sequences available for the family Allocreadiidae included Crepidostomum spp. Braun, 1900. All tree branches had good confidence with posterior probabilities (PP) larger than 0.726, except in the division of subfamilies Anaporrhutinae and Gorgoderinae, which had a PP = 0.449. However, the ML tree showed a value of 87 bootstraps in this same subdivision.
Figure 4. Bayesian inference phylogenetic tree for the 28S gene fragment of sequences derived from organisms of the families Gorgoderidae and Allocreadiidae. The tree was reconstructed using a General Time Reversible with gamma substitutions nucleotide substitution model. Line width and node circles are proportional to the posterior probabilities. The host group from which the organism was collected is indicated as a cartoon.
Figure 5. Bayesian inference phylogenetic tree for the ITS2 fragment of sequences derived from organisms of the families Gorgoderidae, Allocreadiidae, and Prostogonimidae. The tree was reconstructed using a Hasegawa-Kishino-Yano nucleotide substitution model. Line width and node circles are proportional to the posterior probabilities. The host group from which the organism was collected is indicated as a cartoon.
The phylogenetic BI and ML tree for the ITS2 sequences grouped sequences as in the 28S gene with high PP and bootstrap values. However, sequences of the subfamily Gorgoderinae were not available for the analysis. In addition, the low number of sequences available led to the formation of two clusters in the subfamily Anaporrhutinae in the BI tree: one with Plesiochorus spp. and a second group with S. cymatodes, Anaporrhutum spp. and Nagmia sp. In the ML tree, the two groups were also observed, but Nagmia sp. clustered with Plesiochorus spp. instead of Anaporrhutum spp. and S. cymatodes sequences.
Discussion
Members of the family Gorgoderidae infecting elasmobranchs belong to five genera within the subfamily Anaporrhutinae Loss, 1901. These are Anaporrhutum, Nagmia, Petalodistomum Johnston, 1913, Probolitrema Looss, 1902, and Staphylorchis Travassos, 1922 (Curran et al. Reference Curran, Blend, Overstreet, Combes and Jourdane2003; Cutmore et al. Reference Cutmore, Bennett and Cribb2010). In the most recent review of Gorgoderidae, most of genera of Anaporrhutinae were divided into a number of subgenera (see Campbell Reference Campbell, Bray, Gibson and Jones2008); however, because none of the subgenera proposed have received general acceptance by the scientific community (Cutmore et al. Reference Cutmore, Bennett and Cribb2010, Reference Cutmore, Miller, Curran, Bennett and Cribb2013), we follow in recognising them as full genera.
The specimens described here agree well with the diagnostic morphological characters of the genus Anaporrhutum as described by Curran et al. (Reference Curran, Blend, Overstreet, Combes and Jourdane2003) and Campbell (Reference Campbell, Bray, Gibson and Jones2008). In particular, the main characters used for their identification as belonging to Anaporrhutum were the caeca without diverticula, the testicular follicles traversing the caeca, the vitellaria medial or ventral to the caeca, and the H-shaped excretory vesicle.
Species of the genus Anaporrhutum had historically been erected based solely on their traditional morphological characters. Unfortunately, many of them are still poorly known and inadequately described. For instance, in the original descriptions of the species of Anaporrhutum, there is no mention of any tegumental structures. Only Curran et al. (Reference Curran, Blend, Overstreet, Combes and Jourdane2003), in the original description of A. euzeti, reported the occurrence of an ellipse-shaped opening pit at the anterior end of the oral sucker; however, no other information or figures were provided.
The present study of the tegumental surface of A. mundae, as revealed by scanning electron microscopy, allowed us to detect new morphological characters for a member of Anaporrhutinae that may be of systematic interest. Within the Gorgoderidae, these structures (i.e., the frontal pit, the stylet cavity, and the penetration gland openings) have been reported in some members of Gorgoderinae in which four types of presumed sensory papillae (here not found) were also described (Bakke and Bailey Reference Bakke and Bailey1987; Bakke and Hoole Reference Bakke and Hoole1988; Hoole et al. Reference Hoole, Mitchell and Mason1983; Mata-López and León-Règagnon Reference Mata-López and León-Règagnon2006). According to Hoole et al. (Reference Hoole, Mitchell and Mason1983), the frontal pit and stylet cavity in juvenile and adult gorgoderids might be a vestigial organ with no active function, although the function of the penetration gland openings remains enigmatic. The presence of penetration gland pores occurring anterior and lateral to the frontal pit have been described in metacercarial cysts and adults of Gorgoderina vitelliloba (Olsson, 1876) infecting Rana temporaria Linnaeus (Hoole et al. Reference Hoole, Mitchell and Mason1983). In Gorgoderina festoni Mata-López, 2005, penetration gland openings were arranged around the frontal pit (Mata-López and León-Règagnon Reference Mata-López and León-Règagnon2005). Moreover, Mata-López and León-Règagnon (Reference Mata-López and León-Règagnon2006), who studied six species of Gorgoderina Looss, 1902 infecting amphibians in Mexico, observed the occurrence of a frontal pit and/or stylet cavity in all species, but it varied in shape and size among species. In contrast, the penetration glands showing different arrangement patterns were observed only in G. diaster Lutz, 1926, G. parvicava Travassos, 1922, and G. megacetabularis Mata-López, León-Règagnon & Brooks, 2005. Comparative analyses on the detailed tegumental topography of gorgoderids of the genus Phyllodistomum clearly indicates that scanning electron microscopy is a pivotal tool to discriminate among closely related species (Pérez-Ponce de León et al. Reference Pérez-Ponce de León, Martínez-Aquino and Mendoza-Garfias2015a; Petkevičiūtė et al. Reference Petkevičiūtė, Zhokhov, Stunžėnas, Poddubnaya and Stanevičiūtė2020), because it is the only reliable method that can reveal the presence of small tegumental structures not easily visible under the light microscope. The comparative re-examination of species of Anaporrhutum using scanning electron microscopy can provide useful information in the taxonomic study of the group. This could also afford understanding of whether in this genus, the ultrastructural characters here described can be used as diagnostic characters.
Concerning the molecular characterisation, we presented the first 28S rDNA and ITS2 sequences for an identified species of Anaporrhutum. In particular, we also amplified the ITS2 rDNA because it has been used extensively in trematode taxonomy (including also Gorgoderidae), possessing sufficient variability to distinguish even closely related species (Nolan et al. Reference Nolan and Cribb2006; Cutmore et al. Reference Cutmore, Bennett and Cribb2010, Reference Cutmore, Miller, Curran, Bennett and Cribb2013). Unfortunately, at present only ITS2 and 28S rDNA sequences of four taxa of Anaporrhutinae from elasmobranchs are available in GenBank for comparison. These are those of unidentified species of Anaporrhutum and Nagmia from Chiloscyllium punctatum and Stegostoma tigrinum (Forster) respectively, and Staphylorchis cymatodes from Carcharhinus sorrah (Müller & Henle), Sphyrna lewini (Griffith & Smith), and C. punctatum, all from Australia (Cutmore et al. Reference Cutmore, Bennett and Cribb2010, Reference Cutmore, Miller, Curran, Bennett and Cribb2013), and Nagmia floridensis Markell, 1953 from Hypanus sabinus (Lesueur) and Rhinoptera bonasus (Mitchill) from the coasts of the United States of America (Olson et al. Reference Olson, Cribb, Tkach, Bray and Littlewood2003; Curran et al. Reference Curran, Tkach and Overstreet2006). The present phylogenetic analyses included only a relatively small subset of the 32 putative species of Anaporrhutinae listed in WoRMS (2023) thus far. Nevertheless, our results are congruent with those of Cutmore et al. (Reference Cutmore, Miller, Curran, Bennett and Cribb2013) who suggested that taxa infecting elasmobranchs host-switched into sea turtles. Unfortunately, due to the scarcity of molecular data from Anaporrhutinae, it is not currently possible to suggest other conclusions about the relationships among members of this subfamily.
Finally, the site of infection of A. munda was also of remarkable interest. Indeed, we found the new species in the gill chambers of U. munda, but most records of members of Anaporrhutum are from the body-cavity of elasmobranchs. Some exceptions are those reported by von Ofenheirn (1900) and Curran et al. (Reference Curran, Blend, Overstreet, Combes and Jourdane2003), who found A. albidum and A. euzeti in the pericardium of the ray Aetobatis narinari (Euphrasen) and the pericardium and body cavity of several other ray species, respectively, and Simha et al. (Reference Simha, Rao and Rao1971), who reported A. narayani from the buccopharingeal cavity of Mobula mobular (Bonnaterre). Based on the present findings, the Munda round ray represents a new host record and Costa Rica a new geographical record for Anaporrhutum species (see Solano-Barquero et al. Reference Solano-Barquero, Rojas and Cortés2023). Further, according to Caira et al. (Reference Caira, Jensen and Barbeau2022) and Pollerspöck and Straube (Reference Pollerspöck and Straube2023), A. mundae represents the first parasite described or reported for this host.
In conclusion, previous studies (Cutmore et al. Reference Cutmore, Bennett and Cribb2010, Reference Cutmore, Miller, Curran, Bennett and Cribb2013) and the present results show that the species of the genus Anaporrhutum need revision based on both molecular analysis and detailed morphological redescriptions (including light and scanning electron microscopy), emphasising the need to use the integrative taxonomy for accurate species identification and to provide insights into their poorly known life history.
Acknowledgements
We are thankful to Naidely Vidaurre, Gabryele Malcher, Pablo Rojas and Gilbert Ampie for their assistance during fish sampling, as well as to Alexa Rodriguez and Marialetizia Palomba for their technical support during the fish dissection.
Financial support
Fish sampling was partially supported by the “Proyecto BioMar-ACG (808-B9-508)” of the Centro de Investigación en Ciencias del Mar y Limnología, supported by the Guanacaste Dry Forest Conservation Fund and the University of Costa Rica. The parasitological study was supported by the Stazione Zoologica Anton Dohrn under a project between the Stazione Zoologica and the University of Costa Rica, which investigated the parasite biodiversity of elasmobranchs in the coastal waters of Costa Rica.
Competing interest
No potential conflict of interest was reported by the authors.
Ethical standard
The fish collection was authorized by the Ministerio de Ambiente y Energía and Sistema Nacional de Áreas de Conservacion of Costa Rica under the framework of a project of the Centro de Investigación en Ciencias del Mar y Limnología of the University of Costa Rica “Proyecto BioMar -ACG” (permit n. ACG 019-2023). Animal procedures were performed in full compliance with the ethical standards of the relevant national and institutional guides on the care and use for animal experimentation.
