Introduction
Loimoids are rare monocotylid monogeneans of marine elasmobranchs. Currently, three loimoid genera are considered valid: Loimos MacCallum, Reference MacCallum1917, Loimosina Manter, Reference Manter1944, and Loimopapillosum Hargis, Reference Hargis1955. Only five valid Loimos species are known globally, all of which are from the gills of Carcharhinus Blainville, 1816 or Rhizoprionodon Whitley, 1929 species, members of Carcharhinidae (Burhnheim Reference Burhnheim1972; Caballero and Bravo-Hollis Reference Caballero and Bravo-Hollis1961; Chauhan and Bhalerao Reference Chauhan and Bhalerao1945; Chauhan Reference Chauhan1954; Koratha Reference Koratha1955a, Reference Korathab; MacCallum Reference MacCallum1917; Manter Reference Manter1938; Tripathi Reference Tripathi1959). Although Carcharhinidae hosts are globally distributed, these known Loimos species originate from South, Central, and North America, from the eastern Pacific and western Atlantic coastlines, and off Myanmar and India, from the northern Indian Ocean. Of the invalid Loimos representations, an obscure literary account of an unidentified Loimos species from the gills of the teleost, Stellifer minor (Tschudi, 1846), a sciaenid from South America by Jara (Reference Jara1998), repeated in Luque et al. (Reference Luque, Cruces, Chero, Paschoal, Alves, Da Silva, Sanchez and Iannacone2016), is almost certainly a misidentification of a calceostomatid (Calceostomatidae Parona and Perugia, 1890). ‘Loimos parawilsoni Bravo-Hollis, 1970’ (Kohn et al. Reference Kohn, Cohen and Salgado-Maldonado2006; Mendoza-Garfia Reference Mendoza-Garfias, García-Prieto and Pérez-Ponce De León2017) is considered erroneous from lapsus, and the taxon Loimos polytesticularis Bashirullah & Khan, Reference Bashirullah1973 is a nomen nudum (Arthur and Ahmed Reference Arthur and Ahmed2002; Bashirullah Reference Bashirullah1973), although of particular interest because it was collected from an unusual host, Anoxypristis cuspidata (Latham, 1794). No Loimos species are known from Europe, Africa, or the entire Oceania region (including Australia). This is not for a lack of investigation. These regions are well known for their history of monocotylid research, and indeed, the senior author of the current work looked for them for 20 years off southern Africa, without success.
The most recent Loimos species was described more than 50 years ago, and very little information about these worms currently exists (Boeger et al. Reference Boeger, Kritsky, Domingues and Bueno-Silva2014). The phylogenetic position of Loimos was only recently presented in Vaughan et al. (Reference Vaughan, Hansen and Chisholm2024) based on a nonugen GenBank sequence of a single unidentified Loimos specimen from China, the only known Loimos sequence, which confirmed its relationship with Loimosina and within the Monocotylidae.
The current paper marks the first as part of exploratory biodiversity research into monogenean parasites of Australian cartilaginous fishes supported by the Department of Primary Industries, Queensland. A single dead Australian sharpnose shark, Rhizoprionodon taylori (Ogilby, 1915), was sampled from the Queensland Shark Control Program in September 2024 off the southern Great Barrier Reef. Live parasites were recovered from the gills, including an unidentified Loimos species, which is described herein as a new species. Both morphological and molecular data are provided, and the phylogenetic position of the new Loimos species is presented based on 28S rDNA sequences. Molecular confirmation is also provided for the host species.
Materials and methods
A single dead Rhizoprionodon taylori was collected on 20 September 2024, from drumline No. 23, Lammermoor, positioned off the Capricorn coastline, Central Queensland, Australia, operated as part of the Queensland Shark Control Program. The shark was dissected immediately on arrival at Central Queensland University, Rockhampton. Gill arches were removed and inspected individually in a bowl of fresh, filtered seawater (FFS), under an Olympus SZ60 dissection microscope. Live monogeneans were carefully removed from the gill lamellae using a pair of insulin syringes with 27-gauge x 13mm needles, carefully separating the haptor from the gill tissue. These monogeneans were transferred to a separate glass Petri dish containing FFS for live observations under the Olympus SZ60 fitted with a cold fibre-optic light source. Thereafter, the monogeneans were individually transferred to a glass microscope slide in a small drop of FFS, given time to orientate themselves against the glass, and the majority of the FFS was then removed by manipulating the capillary action of an angular piece of paper towel directed towards the side of the water droplet. Just prior to drying, a small circular glass coverslip (8 mm) was placed on top, providing the required amount of pressure for flattening. A small droplet of molecular-grade ethanol was introduced from one side of the coverslip, while simultaneously removing liquid from underneath the coverslip from the opposite side, using the capillary action of an angular piece of paper towel. This drew the ethanol through the coverslip, replacing any remaining FFS, while preserving the individual monogeneans flat. Once the monogeneans were opaque from the ethanol, the coverslip was carefully lifted. The monogeneans were rinsed with molecular-grade ethanol using a pipette and were then carefully removed from the glass microscope slide and transferred to a vial of molecular grade ethanol using a fine paint brush. Pectoral fin clippings were taken from the host and placed into a separate vial of molecular grade ethanol.
The majority of flat-preserved monogeneans were later rehydrated in freshwater and then stained using alum carmine, followed by dehydration using a graded ethanol series to absolute ethanol. Monogeneans were then cleared in cedarwood oil and mounted individually in Canada balsam on a glass microscope slide, under a glass coverslip. Mounted specimens were viewed under an Olympus CX21 or a Nikon Eclipse 200 compound light microscope fitted with a TrueChrome Metrix (Tuscan) microscope camera and phase-contrast and dark-field optics and measured using Fiji image analysis software (ImageJ version 1.54f; Schindelin et al. Reference Schindelin, Arganda-Carreras, Frise, Kaynig, Longair, Pietzch, Preibisch, Rueden, Saalfeld, Schmid, Tinevez, White, Hartenstein, Eliceiri and Tomancac2012), calibrated to a slide graticule, per microscope objective. Morphometrics are given in micrometres as the mean ± standard deviation, followed by the range and the number measured in parentheses. Where only two measurements are given for a character, only the mean and range are provided.
Three preserved monogeneans were processed for molecular characterisation. DNA from each specimen was extracted using an Arcturus PicoPure DNA extraction kit (Applied Biosystems) as per the manufacturer’s instructions. The partial large subunit of the ribosomal DNA (28S, rDNA) was amplified using primers ZX1 (forward: 5’-ACCCGCTGAATTTAAGCATAT-3’) and 1500R (reverse: 5’-GCTATCCTGAGGGAAACTTCG-3’) (Palm et al. Reference Palm, Waeschenbach, Olson and Littlewood2009; Van der Auwera et al. Reference Van der Auwera, Chapelle and De Wachter1994; see also in Lopez-Verdejo et al. Reference Lopez-Verdejo, Palomba, Crocetta and Santoro2023) following qPCR reaction and cycling conditions as published (Lopez-Verdejo et al. Reference Lopez-Verdejo, Palomba, Crocetta and Santoro2023). Host tissue was processed for molecular characterisation. Preserved tissue was processed following previously published methods (Wang et al. Reference Wang, Chen, Lin, Ai and Chen2016). Partial small subunits of the ribosomal DNA (12S and 16S) and the Cytochrome oxidase I (COI) gene regions were amplified using primers 12Sa (5’-AAACTGGGATTAGATACCCCACTAT-3’) and 12Sb (5’-TGAGGGTGACGGGCGGTGTGT-3’) (Kocher et al. Reference Kocher, Thomas, Meyer, Edwards, Pääbo, Villablanca and Wilson1989), 16sar (5’-CGCCTGTTTATCAAAAACAT-3’) and 16sbr (5’-CCGGTCTGAACTCAGATCACGT-3’) (Palumbi et al. Reference Palumbi, Martin, McMillan, Romano, Stice and Grabowsky1991), and FishF1 (5’-TCAACCAACCACAAAGACATTGGCAC-3’) and FishR1 (5’-TAGACTTCTGGGTGGCCAAAGAATCA-3’) (Ward et al. Reference Ward, Zemlak, Innes, Last and Hebert2005), respectively. Quantitative PCR reactions for 12S and 16S followed protocols as per Budd et al. (Reference Budd, Cooper, Le Port, Schils, Mills, Deinhart, Huerlimann and Strugnell2021), except we used 12.5 μL MyTaq Red mix (Bioline) instead of Environmental Mastermix 2.0. Cycling conditions for 12S and 16S followed Budd et al. (Reference Budd, Cooper, Le Port, Schils, Mills, Deinhart, Huerlimann and Strugnell2021). For COI, qPCR reactions contained 12.5 μL MyTaq Red mix, 0.5 μL of forward and reverse primers (both at 10 μM), 9.5 μL of Ultra Purified Deionised (DNase/RNase-Free) water (Invitrogen), and 2 μL of DNA (standardised at 10 ng/μL) for a final reaction of 25 μL. cycling conditions for COI involved an initial activation step of 3 min at 95°C, followed by 30 cycles of 30 sec at 95°C, 30 sec at 50°C, and 35 sec at 72°C, followed by a final extension of 3 min at 72°C. Purified amplicons for monogenean and host gene regions were submitted to AGRF for Sanger sequencing. Sequences for each monogenean specimen were curated using Geneious PRIME (2025.0.3) and aligned with representative monocotylid sequences (see Vaughan et al. Reference Vaughan, Hansen and Chisholm2024): Loimoinae, Loimopapillosum pascuali Chero, Cruces, Sáez, Oliveira, Santos & Luque, Reference Chero, Cruces, Sáez, Oliveira, Santos and Luque2021, Troglocephalinae Vaughan, 2024, Monocotylidae sp., Peruanocotyle pelagica Ruiz-Escobar, Torres-Carrera & Ramos-Sánchez, 2022 (Dasybatotreminae), and Capsala martinieri Bosc, 1811 (Capsalidae Baird, 1853; outgroup) using ClustalW. Best-fit partitioning models of molecular evolution were selected using the program MEGA (version 10.1.7) with the concatenated alignment created in Geneious. A consensus Maximum Likelihood phylogenetic tree was created with MEGA (partitions = 1, evolutionary model = GTI+G+I, bootstrap iterations = 100). Host sequences were curated using Geneious PRIME (2025.0.3) and taxonomic identity confirmed using BLAST at NCBI via Geneious.
All molecular sequences are accessioned in GenBank. The type series of the new species is deposited into the Australian Helminthological Collection (AHC) at the South Australian Museum in Adelaide, South Australia, Australia. A detailed series of high-resolution photomicrographs of Loimos scoliodoni (Manter, Reference Manter1938) taken by Professor Marcus Domingues of HWML 1424 (Harold W. Manter Laboratory of Parasitology Collection; two specimens on two slides) were used for comparative purposes.
Results
Monocotylidae Taschenberg, 1879
Loimoinae Price, Reference Price1936
Loimos MacCallum, Reference MacCallum1917
Loimos everinghami n. sp. (Figures 1 and 2)
Loimos everinghami n. sp.; A, whole mount, ventral view; B, detail of anterior head region; C, haptor, dorsal view; D, position and distribution of marginal hooklets; E, hamulus. Abbreviations: alg, anterolateral gland; avs, anterior ventral sucker (1 = anterior pair, 2 = posterior pair); eb, ejaculatory bulb; gdo, gland duct opening; ham, hamulus; i, intestinal caecum; m, mouth; mco, male copulatory organ; mh, marginal hooklet; o, ovary; oot, oötype; p, pharynx; pg, pharyngeal gland; sr, seminal receptacle; sv, seminal vesicle; t, testis; tvd, transverse vitelline duct; v, vagina; vd, vas deferens; vf, vitelline follicles; vp, vaginal pore. Scale bars: A = 500μm, B = 50μm, C, D = 100μm, E = 30μm.
Loimos everinghami n. sp.; A, reproductive system, ventral view; B, distal portion of male copulatory organ. Abbreviations: cr, crimped distal terminus of male copulatory organ; od, oviduct. All other abbreviations as for Figure 1. Scale bar = 100μm.
Description. Based on observations made from live specimens, and the flat-preserved whole-mounted holotype and 6 paratypes. Body slender and exceptionally extensible in live specimens, facilitated by 2 longitudinal bands of muscle running almost parallel to both branches of the intestinal caecum, for the length of the body proper. Total body (including haptor) 1201 ± 409.8(715–1935, n = 7) long, 256 ± 60.9(172–325, n = 7) at widest point (Figure 1A). Anterior head region rounded to conical, unlobed, containing 2 pairs of muscular, non-papillate ventral suckers, 1 pair positioned medially, the other more lateral and immediately posterior to medial pair (Figures 1A, B). Medial pair 25 (n = 2) in diameter, close together, posterior to anterior margin, separated by short medial septum (Figure 1B). Lateral pair 30 ± 6.6(23–39, n = 6) in diameter, wider apart and deeper internally (Figure 1B). Pair of indistinct gland duct openings located immediately anterior of antero-lateral margin of medial sucker pair (Figure 1B). Pair of anterolateral glands present on either side of mouth; gland ducts not observed (Figures 1A, B). Mouth ventral, subterminal. Large, ovoid muscular pharynx 121 ± 34.7(76–159, n = 7) long, 88 ± 26.8(58–126, n = 7) wide (Figure 1A). Pharyngeal glands present (Figure 1A). Non-diverticular intestinal caecum bifurcates immediately post-pharynx, both branches travel laterally initially before turning posteriorly, following length of body, ending blindly (non-confluent) in short peduncle of posterior body, anterior to haptor (Figure 1A). Muscular haptor ovoid, 179 ± 39.4(123–241, n = 7) long, 282 ± 51.1(218–349, n = 7) wide, aseptate, without 2-part attachment organ (TAO), without marginal membrane, without sclerotised accessory structures, with numerous radiating muscular ventral ridges (Figure 1A). Dorsal surface of haptor contains 2 pairs of delicate membranous flaps (Figure 1C). Fourteen very small (~6, n = 1) marginal hooklets distributed in muscular rim of haptor as illustrated (Figure 1D). Pair of hamuli with both roots embedded in muscular rim of posterior portion of haptor, hook point protruding (Figures 1A, C, D). Hamuli 35 ± 3.6(29–38, n = 7) long; well-developed superficial root similar in length to hamulus hook point; deep root short, narrow; similar to heterocotyline hamuli (Figure 1E). Ovary 115 ± 55.5(56–223, n = 7) long, 154 ± 53.9(79–244, n = 7) wide, medial, immediately pre-testicular; proximal portion irregularly and radially branched; distal portion a single anteriorly directed branch, does not loop right intestinal caecum, curves left as it narrows to form oviduct travelling dorsal to irregularly shaped seminal receptacle (Figures 1A, 2A). Seminal receptacle an irregular bipartite structure, 39 ± 10.1(30–54, n = 5) long, 92 ± 28.2(61–130, n = 5) wide in total dimension (Figures 1A, 2A). Oviduct connects with seminal receptacle, transverse vitelline duct, and duct leading from distal portion of vagina at connection between 2 halves of seminal receptacle (Figures 1A, 2A). Unarmed vagina 108 ± 21.4(89–142, n = 6) long, 35 ± 11.9(18–49, n = 6) wide, musculoglandular with thick walls, positioned ventral of left intestinal caecum, left of seminal vesicle (Figures 1A, 2A). Connecting duct from vagina initially narrow, increasing in diameter before narrowing again, connecting with seminal receptacle (Figures 1A, 2A). Vaginal pore indistinct, in line with concave arch of seminal vesicle (Figures 1A, 2A). Oötype indistinct, 54 (n = 1) long, 32 ± 7.6(25–40, n = 3) wide at widest point, dorsal to ejaculatory bulb (Figures 1A, 2A). Mehlis’ glands, common genital pore, eggs not observed. Vitellarium extends posteriorly from region either side of mid-pharynx, lateral and dorsal to caecal branches, and within post-testicular intercaecal space, ending just anterior to posterior caecal termini (Figure 1A). Eight or 9 transversely ovoid testes present, 29 ± 6.3(18–37, n = 7) long, 104 ± 33.8(63–155, n = 7) wide, with weakly irregular margins, tightly stacked one after the other, positioned medially, intercaecal (Figure 1A). Narrow vas deferens arises from left of anterior-most testis, passes dorsal or just left of ovary, dorsal to seminal receptacle, crossing connecting duct of vagina ventrally before becoming convoluted as it travels anteriorly along right side of vagina. Vas deferens curves initially left at or near vaginal pore, expands slightly in diameter anterior to vagina, curves medially right, narrows towards body mid-line, curves posteriorly to medially or sub-medially positioned botuliform seminal vesicle (Figures 1A, 2A). Narrow curved or straight duct from seminal vesicle connects with base of thick-walled, muscular ejaculatory bulb (Figures 1A, 2A). Ejaculatory bulb in 2 parts: a large bulbous proximal part, 66 ± 11.5(52–82, n = 6) long, 42 ± 11.1(29–56, n = 6) wide, followed by a distinct, narrow distal part, 66 ± 12.6(56–91, n = 6) long, 16 ± 3.5(12–20, n = 6) wide, gooseneck shaped or nearly straight, terminating at connection with proximal portion of male copulatory organ (Figures 1A, 2A). Male copulatory organ a thin-walled sclerotised straight tube, 250 ± 6.4(240–261, n = 7) long, 5 ± 0.7(4–6, n = 7) wide, with terminal distal section crimped (Figures 1A, 2B).
Type host. Rhizoprionodon taylori (Ogilby, 1915)
Type locality. Off Lammermoor, Central Queensland coast (southern Great Barrier Reef; 23°09’43.4"S, 150°47’16.4"E), Australia
Collection date. 20 September 2024
Microhabitat. Gill lamellae
Material deposited. AHC 37157 (holotype); AHC 37158–37163 (6 paratypes)
Etymology. Named in gratitude for Nathan Everingham, working with the Queensland Shark Control Program, who provided the host specimen
ZooBank registration. urn:lsid:zoobank.org:act:E1CD23E5-7F0C-4067-A2CA-759914E76E5E
Taxonomic remarks and differential diagnosis
Loimos species can generally be distinguished by the number of anterior ventral suckers present (a single pair or two pairs) and their sizes, the number of testes, the number of dorsal haptoral flaps, and the length of the male copulatory organ (Table 1). Loimos everinghami n. sp. together with L. salpinggoides MacCallum, Reference MacCallum1917 and L scitulus Burhnheim, Reference Burhnheim1972, have 2 pairs of anterior ventral suckers. Loimos scoliodoni (Manter, Reference Manter1938), L. secundus (Chauhan & Bhalerao, Reference Chauhan and Bhalerao1945), and L. winteri Caballero & Bravo-Hollis, Reference Caballero and Bravo-Hollis1961 all have only a single pair of anterior ventral suckers. Loimos everinghami n. sp. and L. scitulus are the only species recorded with 2 anterior ventral sucker pairs and 4 dorsal haptoral flaps. Loimos salpinggoides was not described with dorsal haptoral flaps, although these may still be present; however, this species only has a single testis (Price Reference Price1938). Loimos everinghami n. sp. is distinguished from L. scitulus by the number of testes, the sizes of the lateral pair of anterior ventral suckers, size of the haptor, and the lengths of the ejaculatory bulb and male copulatory organ (Table 1). Loimos everinghami n. sp. has 8 or 9 testes, where L. scitulus has 15. The lateral pair of anterior ventral suckers of L. scitulus are of unequal sizes, one measuring almost twice the size of its partner (see comments in Burhnheim Reference Burhnheim1972; Table 1), while those of L. everinghami n. sp. are of uniform size in each individual. The haptor of L. everinghami n. sp. is smaller than that of L. scitulus, where that of the latter is about 100 μm wider (Table 1). Burhnheim (Reference Burhnheim1972) also noted the musculoglandular nature of the 4 dorsal haptoral flaps in L. scitulus; however, these flaps in L. everinghami are extremely thin and delicate, without associated musculature, and are not glandular in nature. In live specimens, they appear as simple membranous extensions and likely function to anchor the haptor between two opposing gill lamellae. The ejaculatory bulb and male copulatory organ lengths are greater in L. everinghami, the latter with a range beyond the maximum length recorded for L. scitulus (Table 1). The distal terminus of the male copulatory organ of L. everinghami n. sp. is also notably crimped (Figure 2B), which appears to be unique; however, this requires confirmation by closely examining other Loimos species. Loimos everinghami n. sp. has a unique bipartite and irregularly shaped seminal receptacle (Figures 1A, 2A), whereas such an organ is not described for any other Loimos species.
Comparative data for all known Loimos species
Measurements in micrometres. 1Price (Reference Price1938) used for the data source because Emmett Price corrected original measurement and anatomical errors by MacCallum (Reference MacCallum1917), for the type material; 2measurement not specifying the inclusion or exclusion of haptor length; 3including haptor length; 4Price (Reference Price1938) stated ‘4000μ’ in error; 5given as diameter; 6extrapolated as total ‘combined cirrus and cirrus sac’ length measurements minus ‘cirrus’ length measurements (see Manter Reference Manter1938); 7Koratha (Reference Koratha1955b) provided width for length and vice versa, in error. Length follows the anterior-posterior axis of the body. Note on original host names: Carcharhinus brachyurus = Carcharhinus lamiella Jordan & Gilbert, 1882 (Caballero and Bravo-Hollis Reference Caballero and Bravo-Hollis1961); Rhizoprionodon terraenovae = Scoliodon terraenovae (Richardson, 1836) [Manter Reference Manter1938; Koratha Reference Koratha1955b] = Scolidoni terraenovae (Burhnheim Reference Burhnheim1972); Rhizoprionodon acutus = Scoliodon sorrakowah (Bleeker, 1853) [Chauhan and Bhalerao Reference Chauhan and Bhalerao1945]; ‘Scoliodon sp.’ of Tripathi (Reference Tripathi1959) is a likely synonym of Rhizoprionodon sp.
Marginal hooklets were not visible in all specimens due to the thick muscular rim of the haptor (compare Figures 1A, D). The marginal hooklets were only visible in the smallest specimens observed in this study, although all specimens were sexually mature with fully formed gonads.
Parasite sequences and phylogenetic analysis
Two partial 28S rDNA gene region sequences (845 bp) were successfully sequenced for L. everinghami n. sp. (GenBank numbers: specimen no. 1, PV335244; specimen no. 2, PV335243). Loimos everinghami n. sp. no. 1 and no. 2 showed 0.0737 and 0.0452 genetic divergence with Loimos sp. (Table 2), the smallest divergence across all the other representative Monocotylidae sequences (mean genetic divergence = 0.4539; see Supplemental file S1). Loimos everinghami n. sp. formed a well-supported clade (bootstrap = 99) with Loimos sp. (GenBank No.OM060238) sister to Loimosina wilsoni (Figure 3), creating a monophyletic clade of Loimoinae species sister to Loimopapillosum pascuali (MZ367713-14, Figure 3), as previously proposed within Monocotylidae (Vaughan et al. Reference Vaughan, Hansen and Chisholm2024).
Estimates of divergence between Loimos everinghami n. sp. and selected Monocotylidae sequences in this study
Analyses were conducted using the Tamura-Nei model (Tamura and Nei Reference Tamura and Nei1993); ambiguous positions were removed from each sequence. The complete analysis of 472 base pairs for all representative sequences in this study is available in supplementary file S1. Bold text highlights the estimated divergence between L. everinghami n. sp. and the nonugen Loimos sp. sequence from China.
Relative position of Loimos everinghami n. sp. within selected representatives of Monocotylidae based on a maximum likelihood phylogeny of the partial 28S rDNA gene region. Posterior probability values are presented by each respective branch. Scales refer to genetic distance based on the selected model of each analysis. Branch lengths reflect substitutions per site. Bolded sequences are from this study.
Host sequences
The host tissue COI sequence (GenBank No. PV291672; 688 bp) matched existing R. taylori COI sequences in GenBank (Supplemental file S2). Additional 12S (GenBank No. PV299280; 407 bp) and 16S (GenBank No. PV291675; 428 bp) sequences are also provided for R. taylori (see also Supplemental file S2).
Discussion
Loimos everinghami n. sp. is the first Loimos species described from Australia and the Oceania region. To the best of our knowledge, it is currently also the only known monogenean from this host species. The only other known helminth parasites of R. taylori are the cestodes, Doliobothrium sp. (Cutmore et al. Reference Cutmore, Bennett, Miller and Cribb2017) and Otobothrium carcharidis (Shipley & Hornell, 1906).
Loimoinae was erected under Monocotylidae by Price (Reference Price1936) for L. salpinggoides, based primarily on the aseptate haptor. MacCallum (Reference MacCallum1917) had originally included the type species under the old classification of Monticelli (Reference Monticelli1892), considering it under the sub-order Heterocotylea. Fuhrmann (Reference Fuhrmann1928) classified L. salpinggoides under the family Udonellidae Taschenberg, 1879, but the decision of Price (Reference Price1936) to remove it from this family was based on the presence of haptoral armature and a sclerotised male copulatory organ. Manter (Reference Manter1938) described Tricotyle scoliodonti Manter, Reference Manter1938, tentatively classifying it under Calceostomatidae, although acknowledging its possible relationship with Loimos. Manter (Reference Manter1944) later transferred this species to Loimos, thus relegating Tricotyle Manter, Reference Manter1938 as a junior synonym of Loimos. Chauhan and Bhalerao (Reference Chauhan and Bhalerao1945) proposed to transfer Loimoinae from Monocotylidae to Microbothriidae Price, Reference Price1936 based on the lack of haptoral septa, number of testes, the morphology of the ovary, and the presence of anterior ventral suckers in the head region. This was further discussed by Chauhan (Reference Chauhan1954), who opted not to establish this taxonomic treatment in the repeat description of L. secundus of the previous authors, retaining the genus within Loimoinae, given that Sproston (Reference Sproston1946) was not in agreement with their proposal. Indeed, Sproston (Reference Sproston1946) considered Loimos to be more similar to Calceostoma Van Beneden, 1858 than the microbothriids but agreed with Price (Reference Price1936).
It was Bychowsky (Reference Bychowsky1957) who elevated Loimoinae to family status, concluding that the inclusion of Loimos and Loimosina representing the subfamily Loimoinae within Monocotylidae was erroneous of Price (Reference Price1936, Reference Price1938) and also Manter (Reference Manter1944), and that there were similarities between these loimoids and representatives of Calceostomatidae that could not be ignored. Caballero and Bravo-Hollis (Reference Caballero and Bravo-Hollis1961) apparently accepted the proposal of Bychowsky (Reference Bychowsky1957), although they incorrectly classified Loimoinae Price, Reference Price1936 as the subfamily taxon of Loimoidae Bychowsky, Reference Bychowsky1957 in their taxonomic summary. McMahon (Reference McMahon1963) ignored the proposal of Bychowsky (Reference Bychowsky1957) and promoted Loimoinae as amended by Hargis (Reference Hargis1955) for Loimopapillosum dasyatis Hargis, Reference Hargis1955. The morphological phylogenetic analyses of Boeger and Kritsky (Reference Boeger and Kritsky1993, Reference Boeger, Kritsky, Littlewood and Bray2001) appeared to support the justification of Loimoidae Price, Reference Price1936. However, with later molecular analyses, Boeger et al. (Reference Boeger, Kritsky, Domingues and Bueno-Silva2014) demonstrated evidence for the rejection of Loimoidae, and its return as a subfamily of Monocotylidae, with sequences for Loimosina wilsoni Manter, Reference Manter1944 (see also Dalrymple et al. Reference Dalrymple, de Buron, Hill-Spanik, Galloway, Barker, Portnoy, Frazier and Boeger2022). Boeger et al. (Reference Boeger, Kritsky, Domingues and Bueno-Silva2014) retained Loimoidae pending further data from other loimoids. A new Loimopapillosum species, L. pascuali, was proposed by Chero et al. (Reference Chero, Cruces, Sáez, Oliveira, Santos and Luque2021), providing the first molecular sequences for a representative of this genus. These authors also formally reinstated Loimoinae under Monocotylidae. However, the phylogenies presented by Chero et al. (Reference Chero, Cruces, Sáez, Oliveira, Santos and Luque2021) and Dalrymple et al. (Reference Dalrymple, de Buron, Hill-Spanik, Galloway, Barker, Portnoy, Frazier and Boeger2022) indicated that Loimosina and Loimopapillosum are representatives of distinct clades. This was further confirmed by Vaughan et al. (Reference Vaughan, Hansen and Chisholm2024) who included the first Loimos species molecular sequence in a revised phylogeny of the Monocotylidae, demonstrating a close relationship between Loimos and Loimosina but separate from Loimopapillosum. However, that Loimos sequence (OM060238) is considered nonugen (see Roberts et al., Reference Roberts, Halanych, Arias, Curran and Bullard2018) because it lacks a verifiable morphological anchor and was never accompanied by a description, or details of the host from which it was collected in China. Interestingly, the estimated genetic divergence between this nonugen sequence and L. everinghami n. sp. is very low, between 0.0452 and 0.0737 (Table 2), where comparatively, Neoheterocotyle Hargis, Reference Hargis1955 species typically differ from between 0.1521 and 0.2427 (Supplemental file S1). This suggests that the nonugen sequence from China may either represent L. everinghami n. sp. or a very closely related congener. Additional data from China are required.
Loimoinae is included within the major group of monocotylids containing Dasybatotreminae Bychowsky, Reference Bychowsky1957 and Troglocephalinae, arising from a common ancestor (Vaughan et al. Reference Vaughan, Hansen and Chisholm2024). This ancestor likely possessed ventral pits in the anterior head region, which are present in modified forms in the species of Loimos, Loimosina, and of Troglocephalinae (see also comments by Boeger et al. Reference Boeger, Kritsky, Domingues and Bueno-Silva2014). In Loimos and Loimosina, these pits are modified with musculature to form anterior ventral suckers. This character is considered secondarily lost in Loimopapillosum and species of Dasybatotreminae (Vaughan et al. Reference Vaughan, Hansen and Chisholm2024). Mehracotyle insolita Neifar, Euzet & Ben Hassine, Reference Neifar, Euzet and Ben Hassine2002, Brancheocotyle imbricata Vaughan, Hansen & Chisholm, Reference Vaughan, Hansen and Chisholm2024, Scuticotyle cairae Vaughan, Hansen & Chisholm, Reference Vaughan, Hansen and Chisholm2024, and Troglocephalus rhinobatidis Young, Reference Young1967 (Troglocephalinae) all possess a bipartite seminal receptacle, and the ovary of M. insolita does not loop the right intestinal caecum (Neifar et al. Reference Neifar, Euzet and Ben Hassine2002; Vaughan et al. Reference Vaughan, Hansen and Chisholm2024; Young Reference Young1967). These characters are also shared by L. everinghami n. sp.
Three Rhizoprionodon species are known from Australia: R. acutus (Rüppell, 1837), R. oligolinx Springer, 1964, and R. taylori. Only R. taylori is endemic to Australia and Papua New Guinea but shares an overlapping distribution with R. acutus along the northern and eastern parts of Australia but separate from R. oligolinx (Gallo et al. Reference Gallo, Cavalcanti, Da Silva, Da Silva and Pagnoncelli2010). Rhizoprionodon taylori is considered closely related to R. oligolinx, which is also found off China and currently has no known associated monogenean records in the literature. Rhizoprionodon species are morphologically very similar to each other and are also similar to the closely related monotypic taxon Loxodon macrorhinus Müller & Henle, 1839, which has an overlapping distribution with all three Rhizoprionodon species off Australia. Consequently, the initial identity of the host species provided to us for L. everinghami n. sp. was L. marcrorhinus; however, our molecular investigation of host tissue provided confirmation of the species as R. taylori. The 12S and 16S sequences provided in the current study are also the first generated for this shark species.
Supplementary material
The supplementary material for this article can be found at http://doi.org/10.1017/S0022149X25100370.
Acknowledgements
We are grateful to Mr Nathan Everingham and his team for assisting us with host acquisitions from the QLD Shark Control Program. Thank you to Dr Tracey Scott-Holland from the Queensland Department of Primary Industries for supporting this research, to Dr Gemma Mann of the School of Access Education, CQUniversity for offering her vehicle to assist, and to Ms. Judy Couper of the School of Health, Medical and Applied Sciences for the use of the Nikon Eclipse. Thank you to the two anonymous reviewers for their insightful feedback.
Author contribution
DBV was awarded funding from an internal research grant (RSH7281) from Central Queensland University, supplemented by additional financial support from the School of Health, Medical and Applied Sciences and the Coastal Marine Ecosystems Research Centre. DBV designed the study, collected the samples, and performed the morphological investigation. ATG conducted the molecular investigation and phylogenetic analyses. DBV drafted the initial manuscript, and all authors revised and approved the final manuscript.
Competing interests
The authors declare that they have no known competing financial interests or personal relationships that could have influenced the work reported in this paper.
Ethical standard
The authors assert that all procedures contributing to this work comply with the ethical standards of the relevant national and institutional guides on the care and use of laboratory animals. This work was supported under the CQUniversity animal ethics number 24910, and the Queensland Department of Primary Industries general fisheries permit number 268691, both granted to DBV.
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