Skip to main content
NCBI home page
International Journal for Parasitology: Parasites and Wildlife logoLink to International Journal for Parasitology: Parasites and Wildlife
. 2021 Mar 17;14:341–354. doi: 10.1016/j.ijppaw.2021.03.006

Ophiotaenia echidis n. sp. (Cestoda: Proteocephalidae) from the saw-scaled viper, Echis carinatus sochureki Stemmler (Ophidia: Viperidae), one of the world's deadliest snakes, from the United Arab Emirates

Alain de Chambrier a, Philippe V Alves b, Rolf K Schuster c, Tomáš Scholz d,∗
PMCID: PMC8056141  PMID: 33898236

Abstract

Ophiotaenia echidis n. sp. (Cestoda: Proteocephalidae) is described from the intestine of one of the world's deadliest snakes, the saw-scaled viper Echis carinatus sochureki Stemmler (Ophidia: Viperidae) in the United Arab Emirates. The new species differs from other species of the non-monophyletic Ophiotaenia by the position of testes in two longitudinal lines on both sides of the uterus, and by the large size of an embryophore (diameter of 44–55 μm versus less than 40 μm in other species). Phylogenetic reconstructions based on lsrDNA and concatenated lsrDNA + COI datasets place the new species among proteocephalids from unrelated zoogeographical realms but mostly infecting venomous snakes. In all analyses, O. echidis n. sp. exhibited a strongly supported sister relationship with O. lapata Rambeloson, Ranaivoson et de Chambrier, 2012, a parasite of a pseudoxyrhophiid snake endemic to Madagascar. Despite a shared close evolutionary history between these taxa, morphological synapomorphies remain unclear, which impedes the erection of a new genus to accommodate them. A list of the 71 tapeworms of the former, non-monophyletic subfamily Proteocephalinae, parasitising snakes and lizards, including species inquirendae, and the phylogenetically closely related Thaumasioscolex didelphidis from opossum, with selected characteristics, is also provided, together with a checklist of helminth parasites reported from E. carinatus.

Keywords: Tapeworms, Onchoproteocephalidea, Phylogenetic relationships, Taxonomy, Morphology, Snakes, United Arab Emirates

Graphical abstract

Image 1

Open in a new tab

Highlights

  • •

    We describe the first proteocephalid cestode from the saw-scaled viper Echis carinatus.

  • •

    The new ‘Ophiotaenia’ is typified by the position of testes and embryophore size.

  • •

    The new species phylogenetically clusters with snake cestodes from unrelated regions.

  • •

    The transmission mechanism remains unknown but likely involves some arthropod prey.

1. Introduction

Ophiotaenia La Rue, 1911 is the most species-rich genus of proteocephalid cestodes (Onchoproteocephalidea), with a hundred species (88 according to de Chambrier et al., 2017 plus 12 taxa considered species inquirendae) parasitising reptiles and amphibians throughout the world (Rego, 1994; de Chambrier et al., 2017). Despite the overall morphological uniformity of most species of Ophiotaenia, the most comprehensive phylogenetic analysis (lsrDNA-based) for proteocephalids assessed by de Chambrier et al. (2015), revealed multiple colonisation events in snakes worldwide. These authors recognised three main clades containing species of Ophiotaenia. The generic name should be restricted to species of Clade O of de Chambrier et al. (2015), which includes the type species Ophiotaenia perspicua La Rue (1911) from North American colubrid snakes (La Rue, 1911, 1914), and also Ophiotaenia europaea Odening, 1963 from Palaearctic colubrids (de Chambrier et al., 2015).

Viperid snakes of the genus Echis Merrem, also known as saw-scaled vipers, are small to medium-sized animals widely distributed across xeric areas from Africa (countries north to the Equator) to India and Sri Lanka, including most regions of the Middle East (Pook et al., 2009); they exhibit a diversified diet usually corresponding to a particular species of the genus, e.g., vipers of the E. carinatus (Schneider) and E. pyramidum (Geoffroy Saint-Hilaire) groups prey mostly upon arthropods, whereas species of the E. coloratus Günther group feed almost exclusively on vertebrates (Barlow et al., 2009).

The World Health Organization (2020) classified most species of Echis within Category 1 of risk, i.e., of the highest medical importance, which is reasonable since these vipers are presumably responsible for the greatest number of snakebite deaths per year in Africa and also being important causes of mortality and morbidity in India (see Pook et al., 2009 and references therein). Despite the relevance of Echis spp. to human health, their cestode fauna is poorly known, with just a few reports in faunistic surveys (see Sharpilo, 1976; Gibson et al., 2005). Data on helminth parasites of the saw-scaled viper are also scarce and most of them were published by Soviet authors in the 1950's and 1960's (see Sharpilo, 1976). Since most papers were written in Russian and are not easily accessible, a checklist of helminths parasitising Echis carinatus is provided to facilitate availability of information largely published in local journals and proceedings (‘sbornik’).

In the present paper, a new species of Ophiotaenia from the saw-scaled viper, Echis carinatus sochureki Stemmler, in the United Arab Emirates is described and its phylogenetic position is discussed.

2. Materials and methods

2.1. Morphological data

Tapeworms were found in the small intestine of six saw-scaled vipers, Echis carinatus sochureki Kemmler found in different localities of the United Arab Emirates and examined from 2003 to 2020. This subspecies occurs in northern India, Bangladesh, southern Afghanistan, Pakistan, central Iran, southern Iraq and the United Arab Emirates, with an isolated population in southeastern Arabian Peninsula (Uetz et al., 2020). The nominotypical subspecies E. carinatus carinatus (Schneider) is limited in its distribution to peninsular India (Uetz et al., 2020). None of 33 other snakes examined was infected, namely one Echis omanensis Babocsay, five Cerastes gasperettii Leviton et Anderson (both Viperidae), ten Eryx jayakari Boulenger (Boidae), one Malpolon moilensis (Reuss), 13 Psammophis schokari (Forskål) (both Psammophiidae), and three Platyceps rhodorachis (Jan) (Colubriidae). A total of 58 tapeworms in different states of maturity were found, but some specimens were kept in tap water until their strobilae were totally relaxed. Therefore, only a few specimens collected in a fresh snake (host code No. UAE 4) fixed in 4% hot formalin were suitable for morphological evaluation (see below).

Tapeworms were stained with Mayer's carmine, dehydrated in an ethanol series, clarified by eugenol (clove oil) and mounted in Canada balsam as permanent preparations. For histology, pieces of strobila were embedded in paraffin, transversely sectioned at 12–15 μm intervals, stained with Weigert's hematoxylin, and counterstained with 1% eosin B (acidified with five drops of pure acetic acid for 100 ml solution) (see de Chambrier, 2001). Eggs were studied in distilled water. For scanning electron microscopy (SEM) observations, one scolex was dehydrated through a graded ethanol series, dried in hexamethyldisilazane, coated with gold (thickness of 10–20 nm) and examined in a JEOL JSM-740 1F scanning electron microscope at the Institute of Parasitology, Biology Centre of the Czech Academy of Sciences. All measurements in morphological description are given in micrometres unless otherwise indicated. Abbreviations used in description (usually if the number of measurements was >5) are: x = mean; n = number of measurements. Host and zoogeographical realm classifications follow Uetz et al. (2020) and Holt et al. (2013), respectively. Material studied is deposited in the Natural History Museum, Geneva, Switzerland (acronym MHNG-PLAT), and in the Helminthological Collection of the Institute of Parasitology, České Budějovice, Czech Republic (IPCAS).

A piece of another specimen (paragenophore from host UAE 03; MHNG-PLAT-120508) was used for DNA sequencing (courtesy of J. Brabec) of the large subunit nuclear ribosomal RNA (lsrDNA; D1–D3 domains) and the partial mitochondrial cytochrome c oxidase subunit I (COI) following the methodology outlined by de Chambrier et al. (2019). The sequences were assembled and inspected for errors using Geneious version R11 (Kearse et al., 2012), and submitted to GenBank (MW703700 – lsrDNA; MW703548 – COI); COI gene assembly was trimmed to the protein-coding region using the echinoderm translational code. Two alignments were created using the newly obtained sequences and selected members of the Proteocephalidae mostly corresponding to Clade K of de Chambrier et al. (2015) (these taxa were informed by a more comprehensive analysis of currently unpublished data) using default parameters of MAFFT (Katoh and Standley 2013) implemented in the Guidance2 web server (http://guidance.tau.ac.il/; Sela et al. 2015): (i) alignment including only the lsrDNA sequence data, and (ii) a concatenated alignment (lsrDNA + COI) including only those representatives with available sequences for both markers (see Table 1). Unreliable positions in the alignments were identified and removed using the Gblock web server (https://ngphylogeny.fr/; Dereeper et al., 2008) with less stringent settings.

Table 1.

Summary data for the isolates used in the molecular analyses sorted by zoogeographical realms. GenBank numbers in bold indicate sequences obtained as part of this study.

Region/Taxon Host species (family) [group] Country (isolate) Accession number
lsrDNA COI
Afrotropical
 Ophiotaenia ophiodex Causus maculatus (Viperidae) [R] Ivory Coast (MHNG-PLAT-25962) AJ388620 N/A
Australian
 Australophiotaenia gallardi Pseudechis porphyriacus (Elapidae) [R] Australia (NHMUK: PBI-518) KC786025 KC785990
 Australophiotaenia sp. 1 Antaresia maculosa (Pythonidae) [R] Australia (NHMUK: PBI-513) KC786024 KC785988
Madagascan
 Ophiotaenia lapata Madagascarophis colubrinus (Pseudoxyrhophiidae) [R] Madagascar (NHMUK: PBI-677) KC786021 KC785985
 Ophiotaenia sp. Compsophis sp. (Pseudoxyrhophiidae) [R] Madagascar (NHMUK: PBI-678) KC786023 KC785986
Nearctic
 Megathylacoides giganteuma Ictalurus punctatus (Ictaluridae) [T] USA (US 685a1) MT193860 MT193845
 Megathylacoides lamotheia Ictalurus furcatus (Ictaluridae) [T] USA (US 688-1) MT193867 MT193852
 Ophitaenia grandis Agkistrodon piscivorus (Viperidae) [R] USA (N/A) AJ388632 N/A
Neotropical
 Crepidobothrium gerrardii Boa constrictor (Boidae) [R] Peru (NHMUK: PBI-516) KC786018 N/A
 Ophiotaenia jararaca Bothrops jararaca (Viperidae) [R] Brazil (MHNG-PLAT-12393) AJ388607 N/A
Panamanian
 Thaumasioscolex didelphidis Didelphis marsupialis (Didelphidae) [M] Mexico (NHMUK: PBI-646) KC786019 KC785983
Oriental
 Macrobothriotaenia ficta Xenopeltis unicolor (Xenopeltidae) [R] Vietnam (NHMUK: PBI-491) KC786020 KC785984
 Ophiotaenia bungari Bungarus fasciatus (Elapidae) [R] Vietnam (NHMUK: PBI-500) KC786022 KC785987
Saharo-Arabian
 Ophiotaenia echidis n. sp. Echis carinatus sochureki (Viperidae) [R] United Arab Emirates (MHNG-PLAT-120508) MW703700 MW703548
Open in a new tab
a

Outgroups. Abbreviations: N/A – not available; MHNG-PLAT – Natural History Museum, Geneva, Switzerland; NHMUK – Natural History Museum, London, UK; PBI-No. – unique specimen ID, see http://www.tapewormdb.uconn.edu/index.php/parasites/molecular_search/; M – mammals; R – reptiles; T – teleosts.

Phylogenetic reconstructions were performed with the maximum likelihood (ML) criterion using the evolutionary models implemented in ModelFinder (Kalyaanamoorthy et al., 2017) within IQ-TREE (Trifinopoulos et al., 2016), based on the small sample size corrected Akaike Information Criterion (AICc). The models chosen were as follows: TIM3 + F + R2 for the lsrDNA dataset alone; K3P + I for 1st codon position and GTR + F + G4 for lsrDNA, 2nd and 3rd codon positions using the concatenated dataset. The ML trees were generated via IQ-TREE and clade supports were estimated with 5000 replicates of the ultrafast bootstrap (UFBoot – Minh et al., 2013) and an SH-aLRT test with 5000 replicates (Guindon et al., 2010). To avoid overestimation of UFBoot, we used a hill-climbing nearest neighbour interchange (NNI), as recently recommended by Hoang et al. (2018). Clades with support values of both UFBoot ≥95 and SH-aLRT ≥ 80 were considered strongly supported, while clades with only one of UFBoot ≥95 or SH-aLRT ≥ 80 were weakly supported; nodes with both UFBoot <95 or SH-aLRT < 80 were unsupported. All the above-mentioned analyses were run on the computational resource CIPRES (Miller et al., 2010).

3. Results

3.1. Ophiotaenia echidis n. sp. Fig. 1, Fig. 2, Fig. 3

Fig. 1.

Fig. 1

Open in a new tab

Scanning electron micrographs of Ophiotaenia echidis n. sp. from Echis carinatus sochureki, United Arab Emirates. A – subapical view of the scolex. B – frontal view of the scolex. C – detail of suckers. D–J – microtriches on the apex of the scolex, external (non-adherent) surface of suckers, upper rim, luminal surface and lower rim of suckers, between suckers and neck, respectively. Note: small black letters in A and B correspond to the figures showing higher magnification images of these surfaces.

Fig. 2.

Fig. 2

Open in a new tab

Ophiotaenia echidis n. sp. from Echis carinatus sochureki, United Arab Emirates. A – scolex, IPCAS C-876/1 (UAE 04). B – posterior part of proglottid, holotype, MHNG-PLAT-0137383, dorsal view. C – terminal genitalia with evaginated cirrus and vaginal sphincter, MHNG-PLAT-0137385, ventral view. D – cross section at level of anterior part of pregravid proglottid, MHNG-PLAT-0120507. E − egg with a trilayered embryophore, drawn in distilled water, MHNG-PLAT-0120507. F – cross section at level of the vagina, showing the vaginal sphincter, mature proglottid, MHNG-PLAT-0120507. G – terminal genitalia, C-876/1 (UAE 04 TS), dorsal view. H, I – eggs with a trilayered embryophore, drawn in distilled water, MHNG-PLAT-0120507. Abbrevations: ci: cirrus; cs: cirrus-sac; doc: dorsal osmoregulatory canal; ec: ejaculatory duct; em: trilayered embryophore; ilm: internal longitudinal musculature; ln: longitudinal nerve cord; loc: lateral osmoregulatory canal; Mg: Mehlis' glands; oc: secondary osmoregulatory canals; oe: outer envelope; on: oncosphere; ot: ootype; ov: ovary; sc: subtegumental cells; te: testes; tg: tegument; ud: uterine diverticula; up: uterine pore; us: uterine stem; ut: uterus; va: vas deferens; vc: vaginal canal; vd: vitelline duct; vf: vitelline follicles; voc: ventral osmoregulatory canal; vs: vaginal sphincter.

Fig. 3.

Fig. 3

Open in a new tab

Ophiotaenia echidis n. sp. from Echis carinatus sochureki, United Arab Emirates. A - pregravid proglottid, holotype, MHNG-PLAT-0137383, dorsal view. B – gravid proglottid, paratype, IPCAS C-876/1, ventral view. Abbrevations: cs: cirrus-sac; eci: everted cirrus; oc: osmoregulatory canal; ov: ovary; te: testes; ut: uterus; vc: vaginal canal; vf: vitelline follicles.

Type- and only host: Echis carinatus sochureki Stemmler, 1969 (Serpentes: Viperidae).

Type-locality: Houbara Breeding Centre (25.10418; 55.11283), Dubai, United Arab Emirates.

Other localities: Camel Reproduction Centre, Dubai (25.08752; 55.38819); Dubai Safari (25.17971; 55.45056); Emirates Industry for Camel Milk Products, Dubai (25.02290; 55.41492), all in the United Arab Emirates.

Prevalence: Six snakes of 20 examined (i.e., 30%) in March 2007, July 2014, July and December 2017, and June 2020 were positive.

Intensity of infection: A total of 58 tapeworms were found in six positive hosts, i.e., mean intensity of infection was 10 tapeworms/infected host; minimum four tapeworms in June 2020, maximum 21 specimens in December 2017.

Site of infection: Small intestine.

Distribution: United Arab Emirates.

Type-material: Holotype MHNG-PLAT-0137383, 18 slides (complete whole-mounted specimen on three slides and 15 slides with serial sections) and two slides (with serial sections) IPCAS C-876/1 from host field number UAE 04, Houbara Breeding Centre, collected in July 2017; one paratype MHNG-PLAT-0137384, 21 slides (one complete whole-mounted specimens on three slides and 18 slides with serial sections), and two slides (with serial sections) IPCAS C-876/1 field number UAE 04, Houbara Breeding Centre (July 2017); two paratypes, IPCAS C-876/1 (two complete whole-mounted specimens on three and two slides, field number UAE 04-T2a-3a), Houbara Breeding Centre (July 2017), all Emirates Industry for Camel Milk Products in Dubai, examined by R. Schuster in June 2020.

Other material: Vouchers MHNG-PLAT-120508 (paragenophores), 14 slides (seven slides with whole-mounted specimen and seven slides serial sections from host field number UAE 03, Dubai Safari; IPCAS C-876/1 (one whole-mounted specimen); MHNG-PLAT-88912, Houbara Breeding Centre (March 2007), one specimen without scolex, 17 slides (three whole-mount and 14 serial sections), field number UAE 01; MHNG-PLAT-120507, one specimen without scolex, 35 slides (four whole-mounts and 31 with serial sections) Houbara Breeding Centre, field number UAE 02; MHNG-PLAT-0137385, Houbara Breeding Centre (July 2017) (one whole-mounted specimens without scolex, on two slides and 23 slides serial sections) and MHNG-PLAT-0137386, Houbara Breeding Centre (July 2017) (one whole-mounted specimens without scolex, on three slides), field number UAE 04, all from E. carinatus.

Etymology: The specific name is derived from the generic name of the host.

Representative DNA sequences and phylogenetic relationships: one isolate (MHNG-PLAT-120508) yielded partial lsrDNA (MW703700; 1489 bp long) and COI (MW703548; 921 bp long) sequences. The final alignment of only lsrDNA dataset comprised 1012 positions including 124 parsimony informative sites, whereas the concatenated alignment comprised 1615 positions (1012 of lsrDNA and 603 of COI) including 250 parsimony-informative sites. The trees resulting from the ML analyses of both single and concatenated datasets showed a strongly supported sister relationship between O. echidis n. sp. and Ophiotaenia lapata Rambeloson, Ranaivoson et de Chambrier, 2012, from Madagascarophis colubrinus Schlegel (Pseudoxyrhophiidae) endemic to Madagascar (Fig. 4) (Rambeloson et al., 2012). The analysis of lsrDNA alone further shows these species within a polytomy (Clade Q) together with other proteocephalids from distantly related groups of snakes (families Elapidae, Pseudoxyrhophiidae, Pythonidae and Xenopeltidae) from unrelated zoogeographical realms (Afrotropical, Australian, Madagascan and Oriental). While clade K of de Chambrier et al. (2015) is well supported (posterior probability = 1) in their analysis, our results show only weak support for this group (Fig. 4) that encompasses, besides snake cestodes, the only proteocephalid from a mammal, Thaumasioscolex didelphidis Cañeda-Guzmán, de Chambrier et Scholz, 2001, from the common opossum (Didelphis marsupialis Linnaeus) in Neotropical Mexico (Veracruz).

Fig. 4.

Fig. 4

Open in a new tab

Maximum likelihood phylograms of the relationships of Ophiotaenia echidis n. sp. among selected Proteocephalidae inferred from partial lsrDNA (A), and concatenated lsrDNA + COI datasets (B). Colours of branch and species names correspond to the zoogeographical realms of the isolates as proposed by Holt et al. (2013) (see inset). Branch length scale bar indicates number of substitutions per site. Abbreviations: AUS, Australian; AFR, Afrotropical; MAD, Madagascan; NEA, Nearctic; NEO, Neotropical; PAN, Panamanian; ORI, Oriental; SAH, Saharo-Arabian; * indicates tapeworms from viperid snakes (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

3.2. Description (Fig. 1, Fig. 2, Fig. 3)

(Based on four entire specimens and part of another two worms)

Proteocephalidae. Large worms, 230–275 mm long, up to 1.2 mm wide, flattened dorsoventrally, with proglottids greatly elongated, up to 6.6 mm long. Strobila acraspedote, anapolytic, with 96–105 immature proglottids (up to appearance of spermatozoa in vas deferens), 5–6 mature proglottids (up to appearance of eggs in uterus), 3–5 pregravid proglottids (up to appearance of hooks in oncospheres), 105–118 proglottids in total. Immature proglottids wider than long to longer than wide (length: width ratio 1: 0.5–6.0); mature, pregravid and gravid proglottids much longer than wide. Terminal proglottids much longer than wide (Fig. 3), length: width ratio up to 1: 6.4. Tegument 7.5–10 thick.

Scolex spherical, aspinose (Fig. 1, Fig. 2A), 190–450 long (measured from anterior extremity to posterior margin of suckers; x = 235, n = 3), 400–725 (x = 475, n = 3) wide, wider than neck, with four uniloculate, spherical suckers (Fig. 1, Fig. 2A), 165–210 in diameter, representing 30–45% of scolex width; apical sucker or organ absent (Fig. 2A). Neck, i.e., unsegmented zone posterior to scolex to first recognisable proglottids elongate (up to 3.3 mm), up to 540 wide. Apex of scolex and luminal surface of suckers covered with capilliform filitriches (Fig. 1D, G); external (non-adherent) surface of suckers and between suckers covered with gladiate spinitriches (Fig. 1E, I); upper and lower rims of suckers and neck covered with gladiate spinitriches, interspersed with few capilliform filitriches (Fig. 1F, H, J).

Internal longitudinal musculature weakly developed, consisting of few narrow muscle fibres, more numerous along lateral margins of proglottid (Fig. 2D). Osmoregulatory canals run along vitelline follicles, sometimes overlapping them (Fig. 2B). Ventral canals, 10–20 in diameter, with numerous secondary canals directed externally. Dorsal canals thick-walled, 2.5–4 in diameter, situated alongside and median to vitelline follicles (Fig. 2C, F, G). Genital ducts run between osmoregulatory canals (Fig. 2F and G).

Testes spherical, 30–50 in diameter, 118–205 in number (x = 162, n = 20), in single layer and in two longitudinal columns on both sides of uterine stem, composed of single line of testes, situated at about 30% from lateral side of proglottids (Fig. 2, Fig. 3). Anteriorly, testes do not reach anterior margin of proglottids, starting at distance of 2–5% of proglottid length, i.e., slightly posterior to anterior-most vitelline follicles (Fig. 3). Posteriorly, testes never reach ovary, with terminal testes at distance of 9–13% of proglottid length (Fig. 2, Fig. 3).

Vas deferens strongly coiled, reaching almost to mid-line of proglottid, occupying very small area (Fig. 2, Fig. 3). Cirrus-sac ovoid, thick-walled, 220–290 long, i.e. 20–34% (x = 24%, n = 22) of proglottid width, 100–140 wide (Fig. 2G). Cirrus occupies 73–80% of length of cirrus-sac (Fig. 2C). Genital atrium narrow; genital pores irregularly alternating, slightly postequatorial to equatorial, situated at 49–61% (x = 56%, n = 21) of proglottid length (Fig. 3).

Ovary small, bilobed, 370–540 wide, occupying 52–60% (x = 57%, n = 20) of proglottid width (Fig. 2, Fig. 3); relative size of ovary, i.e., ratio of surface of ovary to surface of proglottid (see de Chambrier et al., 2012), 1.1–1.6% of proglottid size. Ovary length represents 2.5–4.2% of proglottid length. Vagina predominantly anterior (79%), rarely posterior (21%, n = 24) to cirrus-sac, lined with intensely stained cells in its terminal (distal) part and surrounded by small circular sphincter (diameter of 70–80) near genital atrium (Fig. 2C, F). Mehlis’ gland 70–100 in diameter, representing 8–13% of proglottid width.

Vitelline follicles oval, very small (30–60 long, 30–60 wide), arranged in two lateral, longitudinal columns on dorsal side of proglottid (Fig. 3), not interrupted on poral side dorsally at level of terminal genitalia (cirrus-sac and vagina – Fig. 2G). Follicles not reaching anterior margin of proglottids (Fig. 3); posteriorly, follicles may reach ovary, occupying porally 89–94% and aporally 88–92% of proglottid length, respectively.

Primordium of uterine stem ventral, with 47–55 (x = 50, n = 8) lateral branches (diverticula) on each side (Fig. 3B), never reaching posteriorly beyond ovarian isthmus (Fig. 2B). Formation of uterus of type 1 of de Chambrier et al. (2004) as follows: Uterine stem present as undifferentiated longitudinal median concentration of chromophilic cells in immature proglottids. In mature proglottids, uterine stem straight, occupying almost entire length of proglottids. Lumen appears in first mature proglottids. Lateral branches formed when eggs appear in uterus. Thin-walled lateral branches grow in pregravid and gravid proglottids (Fig. 3B), occupying up to 72% of proglottid width, and opening by elongate, slit-like uterine pore.

Eggs large, oval to spherical, with thin, hyaline outer envelope, up to 120 (n = 12) in diameter, somewhat collapsed in distilled water (Fig. 2E, H, I). Inner envelope consists of trilayered embryophore, with thick external layer, 44–53 in diameter, and nucleated envelope of irregular shape, 40–49 in diameter. Oncosphere surrounded by additional thick layer, 20–23 in diameter; oncospheres spherical, 11–12 in diameter, with 3 pairs of embryonic hooks 5–6 long (Fig. 2E, H, I).

3.3. Differential diagnosis

The new species is placed in Ophiotaenia (the former subfamily Proteocephalinae) because of the medullary position of the vitelline follicles, the unarmed scolex with uniloculate suckers and testes forming two separate fields (Rego, 1994). Based on molecular data, the new species belongs to Clade K of de Chambrier et al. (2015), which currently includes species of (non-monophyletic) Ophiotaenia (= Ophiotaenia sensu lato); Australophiotaenia de Chambrier, Beveridge and Scholz (2018); Macrobothriotaenia Freze, 1965; and Thaumasioscolex Cañeda-Guzmán, de Chambrier et Scholz, 2001 (de Chambrier et al., 2015).

The new species differs from all species of Ophiotaenia (sensu lato) by the position of testes in single lines alongside the uterus, and by a large embryophore (44–53 μm versus less than 40 μm) (see Table 2).

Table 2.

List of tapeworms of the former, non-monophyletic subfamily Proteocephalinae, parasitising snakes and lizards, including species inquirendae, and phylogenetically closely related Thaumasioscolex didelphidis from opossum with selected characteristics (see de Chambrier et al., 2017).

Species (clade/subclade)a Type host Country Total length (mm) Width of scolex (μm) Testis number Relative cirrus-sac lengthb Genital pore positionc Position of vagina Vaginal sphincter No. uterine diverticula Apical organ Ovary surfaced Diameter of embryo-phore Embryophore (no. layers)
Australophiotaenia amphiboluri (Nybelin, 1917) Pogona barbata Australia ? absent 127–152 10–14% 62–73% ant–post present 29–32 ? 6.7% 30–35 bilayered
A. gallardi (Johnston, 1911) (clade K & subclade Q) Pseudechis porphyriacus Australia 400 960 72–80 25% 50% anterior absent numerous present 3.2% ? trilayered
A. longmani (Johnston, 1916) (clade K & subclade Q; not included in our analyses – no lsrDNA data) Aspidites ramsayi Australia 94 800 ? 25% 50% anterior present 25–35 absent 2.3% 32–36 trilayered
A. mjobergi (Nybelin, 1917) Demansia psammophis Australia 150 700–850 200–300 ?35% 50% ant–post present 30–40 absent 4.7% 30–34 trilayered
A. striata (Johnston, 1914) Lialis burtonis Australia 33 330–465 64–83 13–17% 48–51% ant–post absent 18–27 absent 1.5% 27–30 ?
Crepidobothrium gerrardi (clade B) Boa constrictor Brazil 560 1820–2120 237–370 23–33% 35–53% ant–post present 18–35 present ? 17–23 ?
Macrobothriotaenia ficta (Meggitt, 1931) (clade K & subclade Q) Xenopeltis unicolor Burma 51–62 525–1290 43–63 34–48% 44–62% ant–post present 26–37 absent 4.1% 33–39 trilayered
Ophiotaenia adiposa Rudin, 1917f Bitis arietans Africa 300–400 55–600 170–220 20–25% 50% ant–post present 40–50 present 2.1% ? ?
O. agkistrodontis (Harwood, 1933)f Agkistrodon piscivorus USA small 850–950 90–110 20–25% 20–25% ant–post ? 25–30 absent 5.1% ? ?
O. andersoni Jensen, Schmidt et Kuntz, 1983f Trimeresurus stejnegeri Taiwan 130 220–450 42–116 40–45% 42–43% ? present about 50 present 2.1% 23–31 ?
O. arandasi Santos et Rolas, 1973 species inquirenda Erythrolamprus miliaris Brazil 80 390 70 41–42% 35–45% ant–post ? about 60 absent 5.1% ? ?
O. atretiumi (Devi, 1973) Atretium schistosum India 160–190 210–280 70–95 17–20% 45–59% ant–post absent 10–15 present 6.5–8.5% 15 ?
O. azevedoi de Chambrier et Vaucher, 1992f Bothrops jararaca Brazil 415 630–735 88–212 17–26% 40–55% ant–post present 45–61 present 1.9% 27–31 bilayered
O. barbouri Vigueras, 1934 Tretanorhinus variabilis Cuba 16–18 730 46–58 ? 33% posterior ? 18–22 absent 2.9% 26–28 ?
O. bungari de Chambrier, Binh et Scholz, 2012 (clade K & subclade Q) Bungarus fasciatus Vietnam 240 360–420 100–150 29–30% 29–48% ant–post present 50–65 present 3–3.3% 28–32 trilayered
O. calmettei (Barrois, 1898)f Bothrops lanceolatus Martinique 270–800 1000–1300 130–160 17–25% 50% posterior present 24–35 absent 2.2% ? ?
O. catzeflisi de Chambrier et Vaucher, 1992f Bothrops jararaca Brazil 124–240 990–1220 107–158 14–22% 40–54% ant–post present numerous present 2.1% 15–23 bilayered
O. chattoraji Srivastava, 1980 Naja naja India 24–26 300–360 140–160 18% 45–47% ? ? 10–26 present 2.9% 27–30 ?
O. congolensis (Southwell et Lake, 1939) Boaedon olivaceus Zaire 80 ? 65 25% 50% ant–post ? 15–20 ? 4.3% 15 ?
O. crotali Lopez-Neyra et Diaz–Ungria, 1958f Crotalus durissus Venezuela 520 absent 308–412 7–12 41–43% ? present .8–16 ? 2.8% 27–30 ?
O. crotaphopeltis Sandground, 1928 Crotaphopeltis tornieri Lake Tanganyika ? 210–260 94–98 19% 50% posterior ? 15–18 absent 3.8% 26 ?
O. dubinini Freze et Sharpilo, 1967 Coronella austriaca Ukraine 130 202–283 87–166 25% 33–(?66%) ant-post absent 20–26 absent 9.8% ? ?
Ophiotaenia echidis n. sp.f (clade K & subclade Q) Echis carinatus sochureki UAE 230–275 400–725 118–205 20–34% 49–61% ant–post present 47–55 absent 1.1–1.6% 44–55 trilayered
O. elapsoidaea Sandground, 1928 Elapsoidea guentherii Lake Tanganyika 150 1000–1100 100–125 30% 50% posterior ? present 48–55 absent 3.4% ? ?
O. elongata Fuhrmann, 1927 ‘a small snake’ Brazil 30–40 ? 26–44 ? ?50% ? ? 13–16 ? 2.5% 19 ?
O. europaea Odening, 1963 (clade O) Natrix natrix Europe 280–540 222–313 189–344 25–43% 33–66% posterior absent 28–73 absent 12.7% 18–22 ?
O. euzeti de Chambrier et Vaucher, 1992f Bothrops jararaca Brazil 45 300–310 116–141 25–34 24–42% ant–post present ? absent 2.2% ? ?
O. faranciae (MacCallum, 1921) Farancia abacura USA >180 500 390–420 ?25% 16–25% posterior ? 30–50 present 2.1% ? ?
O. fima (Meggitt, 1927) Amphiesma stolatum India ? 200 68–89 25–33% ?35% posterior ? 27–33 absent 3.4% ? ?
O. fixa (Meggitt, 1927) species inquirenda Amphiesma stolatum India 50 absent 71–94 20% 33–40% ant–post ? 20–24 ? 3.8% ? ?
O. flava Rudin, 1917 Coluber sp. Brazil 50–60 500–600 45–60 50% 20–40% ant–post absent ? absent 3.6% 28–30 ?
O. gabonica (Beddard, 1913)f Bitis gabonica Africa 380 300 130–170 20% ?43% posterior present 38–46 absent 2.8% 39 ?
O. georgievi de Chambrier, Ammann et Scholz, 2010 Leioheterodon geayi Madagascar 57 225–235 92–140 19–32% 44–56% ant–post present 23–28 absent 2.5–4.3% 31–35 trilayered
O. gilberti Ammann et de Chambrier, 2008 Thamnodynastes pallidus Paraguay 60–120 140–145 57–91 15–23% 42–50% ant–post present 28–41 present 3.7% 27–28 bilayered
O. grandisLa Rue, 1911f (clade K) Agkistrodon piscivorus USA 300–400 1000–1200 200–250 20–33% ?50% ant–post present 40–60 absent 2.1% 21–23 bilayered
O. greeri (Bursey, Goldberg et Kraus, 2006) Sphenomorphus aignanus Papua New Guinea 7–15 205–410 29–51 15–17% 38–47% anterior present 14–24 present 6.8% 15–18 ?
O. habanensis Freze et Rysavy, 1976 Tropidophis pardalis Cuba 57–67 360 31–51 >50% env.60% ? present 26–32 absent 2.7% 22–28 ?
O. hyalina Rudin, 1917 Coluber sp. (Liophis ?) Brazil 120 680–800 50–55 50% ?33% ant–post present ? absent 5.5% ? ?
O. indica Johri, 1955 Naja naja India 110–180 200–370 31–32 33% 50% ?anterior ? 15–17 absent 5.4% 24–30 ?
O. japonensis Yamaguti, 1935 Rhabdophis tigrinus Japan 400 330–500 90–130 ?25% 50% ant–post present 20–30 absent 1.9% 39 ?
O. jarara Fuhrmann, 1927f (clade K) Bothrops jarara Brazil 140 1100–1200 150 27–34% 50% anterior absent 27–32 ? absent 2.4% 23 bilayered
O. joanae de Chambrier et Paulino, 1997 Xenodon neuwiedii Brazil 140–250 480–790 147–210 14–25% 28–56% ant–post present 26–49 present 3.1% 26–30 bilayered
O. kuantanensis Yeh, 1956 Ophiophagus hannah Malaysia 100 2400 250–350 20% <50% ant–post ? 35–40 absent 5.1% 33 ?
O. lapata Rambeloson, Ranaivoson et de Chambrier, 2012 (clade K & subclade Q) Madagascarophis colubrinus Madagascar 295 190–280 89–170 19–25% 43–53 ant–post present 41–68 present 2.1–2.9% 34–39 trilayered
O. lactea (Leidy, 1855) species inquirenda Nerodia sipedon USA ? ? ? ? 50% ? ? ? absent ? ? ?
O. macrobothria Rudin, 1917 Micrurus corallinus Brazil 50 400–500 50–60 >50% 20–33% ant–post absent ? absent 4.4% ? ?
O. marenzelleri (Barrois, 1898)f Agkistrodon piscivorus USA 400 1200–2000 150–240 25–33% 50% anterior ? 20–30 absent 2.6% 20 ?
O. meggitti Hilmy, 1936fspecies inquirenda Atheris chlorechis Liberia 68 absent 86–104 33% >50% ant–post ? 35.00 ? 2.3% ? ?
O. micruricola (Shoop et Corkum, 1982) Micrurus diastema Mexico 259–290 720–760 121–169 14–20% 48–56% posterior absent 35–53 absent 3.2% 20–23 ?
O. monnigi Fuhrmann, 1924 Crotaphopeltis hotamboeia Africa 50 ? 80 10% 50% anterior present 50–57 ? 3.1% 25 bilayered
O. najae (Beddard, 1913) Naja tripudianse India 180 248–303 120–(165) 25–33% ?50% anterior absent 14–25 absent 3.5% 25 ?
O. nankingensis (Hsü, 1935) Ptyas dhumnades China 105–124 320 147–166 ?17% <50% ant–post present 36–40 absent 3.0% ? ?
O. nattereri (Parona, 1901) Coluber sp. Brazil 75–250 250 80–100 28–33% <50% ant–post present 15–30 absent ? 22–26 ?
O. nicolae Coquille et de Chambrier, 2008 Thecadactylus rapicauda Ecuador 230–515 325–340 142–204 21–33% 34–53% ant–post present 13–27 present 4.9% 32–36 bilayered
O. nigricollis Mettrick, 1963 Naja nigricollis Zimbabwe 170 300–310 176–210 ∼25% ∼35% ant–post ? 16–20 ? 6.3% 26–33 ?
O. nybelini Hilmy, 1936 Meizodon coronatus Africa 52 105 67–90 16–20% ?50% ant–post ? 25–40 absent 3.6% 25 ?
O. ophiodex Mettrick, 1960f (clade K & subclade Q) Causus rhombeatus Zimbabwe 210–270 790–1140 110–120 25% 45–54% ant-post present 30–42 absent ? 27–36 ?
O. paraguayensis Rudin, 1917 (clade N) Hydrodynastes gigas Paraguay 550–600 240 238–344 12–19% 27–39% ant–post present 20–36 absent 3.3% 21–24 ?
O. perspicua La Rue, 1911 (clade O; type species) Nerodia rhombifer USA 360 255–410 150–215 25–33% 33–50% anterior present 20–30 present 2.3% ? ?
O. phillipsi (Burt, 1937)f Trimeresurus trigonocephalus Sri Lanka 400–920 670–1400 170–230 ∼ 50% ∼50% posterior present 60–80 absent 2.1% 24–30 ?
O. racemosa (Rudolphi, 1819) Coluber sp. Brazil 160 540–650 80–120 ∼33% ∼33% ant–post present 40–50 ? 4.3% 24 ?
O. rhabdophidis (Burt, 1937) Amphiesma stolatum Sri Lanka 250 130–187 110–140 20–25% 33–50% ant–post ? 30–45 present 3.1% 22–27 ?
O. sanbernardinensis Rudin, 1917 (clade N) Helicops leopardinus Paraguay 100–120 230–250 70–102 ? 20–40% ant–post present 27–33 absent 5.0% 22–23 ?
O. sinensis Cheng et Lin, 2002 Rhabdophis tigrinus China 460–694 247–325 256–320 ∼25% ∼45% ? ? ? ? 2.5% ? ?
O. southwelliFreze, 1965f Causus rhombeatus Africa 90 1500 170–230 ? ∼50% ant–post ? 8–12 ? 3.4% 30 bilayered
O. spasskyi Freze and Sharpilo, 1967f Vipera berus Russia 70–90 295–355 179–271 20–25% ∼30% ant–post present 19–31 ? 9.1% ? ?
O. theileri Rudin, 1917 Naja haje Africa 300 400 160–310 20–25% ∼50% ant–post absent 35–40 absent 4.5% ? ?
O. trimeresuri (Parona, 1898)f Trimeresurus formosus (= T. sumatranus) Mentawai Islands 105 750 100–108 20–30% ∼50% ant–post present 20–30 absent 3.6% ? ?
O. variabilis Brooks, 1978 Nerodia cyclopion North America 300 170–200 77–253 22–33% 15–30% ant–post absent 25–45 absent 2.8% ? ?
O. wuyiensis Cheng, Yuguang et Zao He, 2007f Trimeresurus gramineus China 460–700 245–325 256–320 23% ∼45% ? ? ? ? 2.8% ? ?
O. zschokkei Rudin, 1917 Naja haje South Africa ∼550–600 400 160–200 20–25% ∼50% ant–post ? ? absent 6.4% ? ?
Thaumasioscolex didelphidis Cañeda-Guzmán, de Chambrier et Scholz, 2001 (clade K) Didelphis marsupialis Mexico 640–1045 655–1040 432–548 11–19% 31–44% ant–post present 12–22 absent 8.5% 30–33 bilayered
Open in a new tab
a

Clades – see de Chambrier et al. (2015); subclade Q – see the text.

b

Ratio of the cirrus-sac length to the proglottid width (in %).

c

Ratio of the genital pore position to the proglottid length (in %).

d

Ratio of the ovary surface to the proglottid surface (in %; see de Chambrier et al., 2012).

e

Naja tripudians is currently not recognised and it may be Naja kaouthia Lesson or Naja naja (Linnaeus) (Uetz et al., 2020).

f

Indicates species of Ophiotaenia from viperid snakes.

In addition, O. echidis n. sp. possesses a trilayered embryophore, which is present only in a few species of Ophiotaenia from snakes, such as Ophiotaenia lapata Rambeloson, Ranaivoson et de Chambrier, 2012 from Madagascarophis colubrinus (Schlegel), Ophiotaenia georgievi de Chambrier, Ammann et Scholz, 2010 from Leioheterodon geayi (Mocquard) (both host (Pseudoxyrphophiidae), and O. bungari de Chambrier, Binh et Scholz, 2012 from Bungarus fasciatus (Schneider) (Elapidae) (see de Chambrier et al., 2010; 2012; Rambeloson et al., 2012).

Ophiotaenia echidis differs from the closely related O. lapata (see Fig. 4) by the absence of an apical organ and by the larger size of the scolex (400–725 μm versus 190–280 μm). The new species can be differentiated from O. bungari by the position of the genital pore (at 49–61% of the proglottid length versus 29–48%) and by the smaller relative size of the ovary (1.1–1.6% versus 3.3%). Ophiotaenia echidis differs from O. georgievi by the number of uterine diverticula (47–55 versus 23–28) and smaller relative size of the ovary (1.1–1.6% versus 2.5–4.3%).

Eight species that belong to Clade Q in Fig. 4 (subclade of Clade K of de Chambrier et al., 2015) share the presence of a third embryophoric layer, in contrast to the three remaining species in the Clade K (Fig. 4), namely O. jarara (Fuhrmann, 1927), O. grandis La Rue, 1911, and T. didelphidis, which possess only a bilayered embryophore (Table 2). Interestingly, all but one species of Clade K listed above have an embryophore of a similar size (diameter always less than 34 μm), the only exception being the new species, O. echidis, with extraordinarily large embryophore (diameter 44–53 μm). The presence of a third layer in the embryophore is not common in the Proteocephalidae and has been observed only in some proteocephalids parasitising reptiles from different genera, such as Australophiotaenia, Cairaella Coquille et de Chambrier, 2008, Kapsulotaenia Freze, 1963, Macrobothriotaenia and Ophiotaenia from the Old World (Asia, Australia and Madagascar), and in Cairaella henrii Coquille et de Chambrier, 2008, a parasite of the polychrotid lizard Norops trachyderma (= Anolis trachyderma Cope) in Ecuador (Coquille and de Chambrier, 2008; Scholz et al., 2013; de Chambrier et al., 2018, 2020). The only proteocephalid parasitising amphibians that possesses a third embryophore layer is Ophiotaenia alessandrae Marsella et de Chambrier, 2008 from Hyla boans (= Boana boans (Linnaeus)) in Ecuador (Marsella and de Chambrier, 2008). This indicates that the presence of a trilayered embryophore is a homoplastic character that has evolved in several groups of proteocephalids parasitising tetrapods independently.

3.4. Checklist of helminth parasites of the saw-scaled viper, Echis carinatus

The present checklist is primarily based on data compiled by Sharpilo (1976) and Gibson et al. (2005), with a subsequent search of primary, especially literature written in Russian.

  • Cestoda – tapeworms

Cyclophyllidea: Anoplocephalidae Cholodkovsky, 1902

  • 1.

    Oochoristica fedtschenkoi Bogdanov et Markov, 1955

    This species was described from several species of viperid snakes, including E. carinatus, from Turkmenistan and Uzbekistan. The highest infection rate was reported for the saw-scaled viper (prevalence of 4.2%, mean intensity of infection 26, range 1–50 worms) (Bogdanov and Markov, 1955; Markov et al., 1970; Sharpilo, 1976).

  • 2.

    Oochoristica sindensis Farooq, Khanum et Ansar, 1983

    This cestode was described from the saw-scaled viper in Pakistan by Farooq et al. (1983).

Cyclophyllidea: Dipylidiidae Stiles, 1896

  • 3.

    Diplopylidium acanthotetra (Parona, 1898) – larvae

    Metacestodes of this tapeworm were found in the body cavity of E. carinatus in Turkmenistan (Markov et al., 1970). The most common intermediate host of D. acanthoptera is the Caspian bent-toed gecko, Tenuidactylus caspius (Eichmann), and its metacestodes were found in Algeria, Egypt, Israel, Iraq and Italy (Sharpilo, 1976).

  • 4.

    Diplopylidium noelleri (Skrjabin, 1924) – larvae

    Larvae of this cestode occur in the liver, intestinal wall and body cavity of reptiles of distantly related groups, such as geckoes, lacertids, agamas and colubrid snakes (Sharpilo, 1976). They were also found in E. carinatus from Turkmenistan (Markov et al., 1970).

  • 5.

    Joyeuxiella echinorhynchoides (Sonsino, 1889) – larvae

    Metacestodes are frequently found in the liver, intestinal wall and body cavity of a wide spectrum of reptiles, including viperid snakes (Sharpilo, 1976). Markov et al. (1970) found J. echinorhynchoides in E. carinatus from Turkmenistan.

Onchoproteocephalidea: Proteocephalidae La Rue, 1911

  • 6.

    Ophiotaenia sp.

    Markov et al. (1970) reported unidentified species of Ophiotaenia from E. carinatus in Turkmenistan. However, the authors did not provide morphological characterisation and illustrations, and no voucher specimens have been deposited. Therefore, it is impossible to compare tapeworms found in Middle Asia with those found in the United Arab Emirates by the present authors.

  • Acanthocephala – spiny-headed worms

Archiacanthocephala: Giganthorhynchidae Hamann, 1892

  • 7.

    Centrorhynchus aluconis (Müller, 1780) – larvae

    Cystacanths of this very common parasite of owls have also been found in E. carinatus from Turkmenistan (Markov et al., 1970).

Archiacanthocephala: Oligacanthorhynchidae Southwell et Macfie, 1924

  • 8.

    Centrorhynchus globocaudatus (Zeder, 1800) – larvae

    Markov et al. (1970) reported cystacanths of this parasite of owl and raptors from E. carinatus in Turkmenistan, but Sharpilo (1976) did not list this species in his survey of helminth parasites of reptiles in the former USSR.

  • 9.

    Centrorhynchus sp. – larvae

    Unidentified larvae (cystacanths) of another species of Centrorhynchus Lühe, 1911 were found in E. carinatus from Turkmenistan by Markov et al. (1970). Their identification is not possible because the authors did not provide any morphological data and illustrations.

  • 10.

    Oligacanthorhynchus sp. – larvae

    Cystacanths of unidentified species of Oligacanthorhynchus Travassos, 1915 were found in the body cavity and liver of various reptiles, including E. carinatus from Turkmenistan (Markov et al., 1970).

  • Nematoda – nematodes (roundworms)

Rhabditida: Ascarididae Blanchard, 1849

  • 11.

    Hexametra skrjabini Markov et Bogdanov, 1960

    This nematode was described from the small intestine and stomach of the spotted desert racer, Coluber karelini Brandt (= Platyceps karelini) (Colubridae), dwarf sand boa, Eryx miliaris (Pallas) (Boidae), and E. carinatus from Dagestan (Russia) and Turkmenistan (Markov and Bogdanov, 1960; Markov et al., 1970).

Rhabditida: Habronematidae Chitwood et Wehr, 1932

  • 12.

    Agamospirura sp. – larvae

    Markov et al. (1970) reported larvae of unidentified species of Agamospirura Henry et Sisoff, 1913 from E. carinatus in Turkmenistan.

Rhabditida: Pharyngodonidae Travassos, 1919

Rhabditida: Physalopteridae Railliet, 1893

  • 14.

    Abbreviata uzbekistana Bogdanov and Markov (1955)

    This parasite of the stomach, rarely of the oesophagus and intestine, was described by Bogdanov and Markov (1955). It has been found in several distantly related reptiles, such as agamas, geckoes and the saw-scaled viper from Dagestan (Russia), Turkmenistan and Uzbekistan (Sharpilo, 1976). Echis carinatus is probably postcyclic, not definitive host of this spirurine nematode.

In addition, larvae of the following nematodes unidentified to the genus or even family level were reported from E. carinatus: Acuaroidea gen. sp. in Turkmenistan (Velikanov, 1984); Omeiinae gen. sp. (Quimperiidae) in Turkmenistan (Markov et al., 1970). One of the present authors (R.K.S.) found the following helminths in the saw-scaled viper from the United Arab Emirates, but these records were presented only at an international conference (Schuster and Sivakumar, 2013): cestodes Joyeuxiella sp., most probably J. pasqualei (Diamare, 1893), and Diplopylidium noelleri (Cestoda); acanthocephalan larvae Centrorhynchus aluconis and Macracanthorhynchus catulinus Kostylew, 1927; and larvae of spirurine nematodes, most probably Physocephalus dromedarii Muschkambarova, 1967.

4. Discussion

Ophiotaenia is a composite genus pending taxonomic revision. Validity of its species should be re-assessed and poorly characterised taxa redescribed. However, this task is difficult because of the large number of species (88 according to de Chambrier et al., 2017 plus 12 species inquirendae) and difficulties in obtaining new, properly fixed material suitable for molecular studies. The generic name Ophiotaenia should be retained only for taxa of Clade O of de Chambrier et al. (2015), because it includes the type species O. perspicua from colubrid snakes in North America. Interestingly, species of this clade have proportionally larger ovaries than those in the other species of Ophiotaenia (Clades K and N of de Chambrier et al., 2015). Species in individual clades also differ in uterine development: Type 1 according to de Chambrier et al. (2004) is present in species of Clade K, whereas a Type 2 uterus occurs in species of Clades N and O (de Chambrier et al., 2015). It is clear that species of Clades K and N need to be allocated to other (new) genera, which also affects the new species described in the present paper.

However, there are two principal obstacles in erecting a new genus for the new species and the closely related O. lapata: (i) molecular data are available for a limited number of species of Ophiotaenia only; and (ii) the absence of obvious morphological synapomorphies of the new species and O. lapata, which would unequivocally differentiate them from the other taxa of Clade K. For example, the scoleces of most species of all but two genera are rather uniform; only species of monotypic Macrobothriotaenia and Thaumasioscolex have distinct scoleces (Cañeda-Guzmán et al., 2001; Scholz et al., 2013). In addition, some species of Australophiotaenia and M. ficta (Meggitt, 1931) have a trilayered embryophore, similar to O. echidis n. sp. (see de Chambrier et al., 2018). Therefore, the new species is provisionally placed in Ophiotaenia, even though this genus is not monophyletic (de Chambrier et al., 2015).

Clade K (updated in this study in Fig. 4) is composed almost exclusively of parasites of snakes, namely Boidae, Elapidae, Pseudoxyrhophiidae, Pythonidae, Viperidae (three of the 18 species of Ophiotaenia from viperids sequenced), and Xenopeltidae, throughout the world and the only proteocephalid parasitising a mammal, the common opossum (D. marsupialis), in Neotropical Mexico in the case of T. didelphidis. According to de Chambrier et al. (2015), species of Clade K may represent a relatively recent colonisation of unrelated groups of snakes in most continents or a trace of a very ancient colonisation of these snake hosts. Based on the most informative tree presented here (using only the lsrDNA dataset), the early colonisation of Clade K representatives in vertebrates (snakes and the common opossum) currently distributed across the Americas is hypothetised, followed by colonisation of snakes in the Afrotropical, Australian, Madagascan, Oriental, and Saharo-Arabian zoogeographical realms (see Fig. 4). Nevertheless, for proposing a more precise scenario of the evolution of this clade of Ophiotaenia, a broader spectrum of related taxa should be analysed, including diverse proteocephalids in Australian reptiles (de Chambrier et al., 2018), which might provide stronger statistical support for internal nodes.

Recently, de Chambrier et al. (2018) erected a new genus, Australophiotaenia, to accommodate species of Ophiotaenia from snakes in the Australian region, including A. gallardi (Johnston, 1911) from Clade K of de Chambrier et al. (2015). In the present analyses, two species of this genus, A. gallardi and Australophiotaenia sp. 1, grouped together (Fig. 4). Australophiotaenia is characterised by the following characteristics: a trilayered, thick-walled embryophore, a scolex with large, anteriorly directed suckers, an exclusively dorsal and paramuscular position of vitelline follicles (i.e., follicles may penetrate into the medulla between fibres of the inner longitudinal musculature), and a postequatorial to equatorial genital pore in most species (de Chambrier et al., 2018).

The present study supports the assumption about strict (oioxenous) host specificity of species of Ophiotaenia from reptilian hosts (see de Chambrier et al., 2010, 2017, 2020) because the new species was found only in E. carinatus sochukeri, but not in any of 33 other snakes of six species, including the congeneric Oman saw-scaled viper, Echis omanensis. It is worth noting that the latter species belongs to the E. coloratus group sensu Pook et al. (2009), species of which feed almost exclusively on vertebrates. In contrast, E. carinatus, which belongs to the E. carinatus group, chiefly feeds on scorpions and other arthropods (Barlow et al., 2009).

Information on proteocephalid life-cycles is scant (Scholz and de Chambrier 2003), but it is postulated that species of Ophiotaenia exhibit two types of life-cycle strategies, i.e., a three-host cycle including planktonic copepods (first intermediate host), fish or amphibian tadpoles (second intermediate host) and reptiles (definitive host), and a two-host cycle with a tissue phase of development in the final host and posterior migration to the intestine, with no involvement of intermediate or paratenic vertebrate host (Biserkov and Kostadinova, 1997). It remains unclear how O. echidis n. sp. is transmitted to its definitive host, but considering the similar diet of vipers of the E. carinatus and E. pyramidum groups (Barlow et al., 2009), new studies should focus on these potential hosts.

The taxonomic history of Echis has been convoluted, which has direct implications for human health, given that venom composition is highly variable among species resulting in several cases of a lack of antivenom cross-neutralisation (Pook et al. 2009). That having been said and considering the assumption of oioxenous host specificity, the fauna of proteocephalids of these venomous snakes may help to inform us about distinct evolutionary lineages of their hosts, which ultimately may be used to guide searches for specific antivenoms. In the most comprehensive phylogeny of viperid snakes by Alencar et al. (2016), Echis appeared as sister to Cerastes Laurenti, but species of the latter genus genus have never been found infected with tapeworms of Ophiotaenia (in the present study, we examined five specimens of Cerastes gasperettii but none was infected). The common European adder, Vipera berus (Linnaeus), which is the definitive host of Ophiotaenia spasskyi Freze et Sharpilo, 1967 in Europe, clustered with species of Echis in a weakly supported clade (Alencar et al., 2016). Therefore such comparison is not informative enough.

Literary search for data on the helminth parasites of the saw-scaled viper has revealed that most information was accumulated by Soviet researchers, especially G.S. Markov and O.P. Bogdanov, from Turkmenistan in the 1950's and 1960's. More recent data are scarce and include just a few reports (Farooq et al., 1983; Nama, 1984; Velikanov, 1984). The helminth fauna of E. carinatus includes 14 species and another two larvae of nematodes unidentified to the genus level. This fauna is composed of adult cestodes (3 spp.), their larvae (3 spp.), larval acanthocephalans (4 spp.), adult nematodes (3 spp.) and their larvae (3 spp.). Noteworthy is the complete absence of any trematode (Digenea) (Sharpilo, 1976; Gibson et al., 2005).

Declaration of competing interest

There is no conflict of interest.

Acknowledgements

Two anonymous reviewers provided insightful comments that help to improve the manuscript. The authors express their gratitude to Jan Brabec for providing unpublished sequence of the new species and preliminary assessing its phylogenetic position, and Roman Kuchta (both Institute of Parasitology, České Budějovice) for providing literature on the parasites of Echis carinatus and helpful suggestions. Thanks are also due to Janik Pralong, Gilles Roth (both Geneva) and Martina Borovková (České Budějovice) for technical help. This study was supported by the Institute of Parasitology (RVO 60077344) and CAPES (postdoc scholarship to PVA, grant 88887.508591/2020-00). We also acknowledge that the Laboratory of Electron Microscopy, IPCAS is supported by the Ministry of Education, Youth and Sports of the Czech Republic (project LM2015062 Czech-BioImaging).

References

  1. Alencar L.R.V., Quental T.B., Grazziotin F.G., Alfaro M.L., Martins M., Venzon M., Zaherb H. Diversification in vipers: phylogenetic relationships, time of divergence and shifts in speciation rates. Mol. Phylogenet. Evol. 2016;105:50–62. doi: 10.1016/j.ympev.2016.07.029. [DOI] [PubMed] [Google Scholar]
  2. Barlow A., Pook C.E., Harrison R.A., Wüster W. Coevolution of diet and prey specific venom activity supports the role of selection in snake venom evolution. Proc. Roy. Soc. B. 2009;276:2443–2449. doi: 10.1098/rspb.2009.0048. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Biserkov V., Kostadinova A. Development of the pleroceroid I of Ophiotaenia europaea in reptiles. Int. J. Parasitol. 1997;27:1513–1516. doi: 10.1016/s0020-7519(97)00138-0. [DOI] [PubMed] [Google Scholar]
  4. Bogdanov O.P., Markov G.S. Parasitic worms of some reptiles of Middle Asia. Izv. Akad. Nauk Uzb. SSR. 1955;8:59–68. (In Russian) [Google Scholar]
  5. Bogdanov O.P., Markov G.S. Venomous Animals of Middle Asia and their Venoms. Tashkent. 1960. Parasite fauna of venomous snakes of Middle Asia and Kazakhstan; pp. 38–45. (In Russian) [Google Scholar]
  6. Cañeda-Guzmán I.C., de Chambrier A., Scholz T. Thaumasioscolex didelphidis n. gen. and n. sp. (Cestoda: Proteocephalidae) from the black-eared opossum Didelphis marsupialis from Mexico, the first proteocephalidean tapeworm from a mammal. J. Parasitol. 2001;87:639–647. doi: 10.1645/0022-3395(2001)087[0639:TDNGNS]2.0.CO;2. [DOI] [PubMed] [Google Scholar]
  7. de Chambrier A. A new tapeworm from the Amazon, Amazotaenia yvettae n. gen., n. sp. (Eucestoda: Proteocephalidea) from the siluriform fishes Brachyplatystoma filamentosum and B. vaillanti (Pimelodidae) Rev. Suisse Zool. 2001;108:303–316. [Google Scholar]
  8. de Chambrier A., Zehnder M.P., Vaucher C., Mariaux J. The evolution of the Proteocephalidea (Platyhelminthes, Eucestoda) based on an enlarged molecular phylogeny, with comments on their uterine development. Syst. Parasitol. 2004;57:159–171. doi: 10.1023/B:SYPA.0000019083.26876.34. [DOI] [PubMed] [Google Scholar]
  9. de Chambrier A., Ammann M., Scholz T. First species of Ophiotaenia (Cestoda: Proteocephalidea) from Madagascar: O. georgievi n. sp., a parasite of the endemic snake Leioheterodon geayi (Colubridae) Folia Parasitol. 2010;57:197–205. doi: 10.14411/fp.2010.026. [DOI] [PubMed] [Google Scholar]
  10. de Chambrier A., Binh T.T., Scholz T. Ophiotaenia bungari n. sp. (Cestoda), a parasite of Bungarus fasciatus (Schneider) (Ophidia: Elapidae) from Vietnam, with comments on relative ovarian size as a new and potentially useful diagnostic character for proteocephalidean tapeworms. Syst. Parasitol. 2012;81:39–50. doi: 10.1007/s11230-011-9320-0. [DOI] [PubMed] [Google Scholar]
  11. de Chambrier A., Waeschenbach A., Fisseha M., Scholz T., Mariaux J. A large 28S rDNA-based phylogeny confirms the limitations of established morphological characters for classification of proteocephalidean tapeworms (Platyhelminthes, Cestoda) ZooKeys. 2015;500:25–59. doi: 10.3897/zookeys.500.9360. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. de Chambrier A., Scholz T., Mariaux J., Kuchta R. Onchoproteocephalidea I Caira, Jensen, Waeschenbach, Olson & Littlewood, 2014. In: Caira J.N., Jensen K., editors. Planetary Biodiversity Inventory (2008–2017): Tapeworms from Vertebrate Bowels of the Earth. University of Kansas, Natural History Museum, Special Publication No. 25; Lawrence: 2017. pp. 251–277. [Google Scholar]
  13. de Chambrier A., Beveridge I., Scholz T. Tapeworms (Cestoda: Proteocephalidae) of Australian reptiles: hidden diversity of strictly host-specific parasites. Zootaxa. 2018;4461:477–498. doi: 10.11646/zootaxa.4461.4.2. [DOI] [PubMed] [Google Scholar]
  14. de Chambrier A., Brabec J., Tran B.T., Scholz T. Revision of Acanthotaenia von Linstow, 1903 (Cestoda: Proteocephalidae), parasites of monitors (Varanus spp.), based on morphological and molecular data. Parasitol. Res. 2019;118:1761–1783. doi: 10.1007/s00436-019-06326-6. [DOI] [PubMed] [Google Scholar]
  15. de Chambrier A., Brabec J., Scholz T. Molecular data reveal unexpected species diversity of tapeworms of Australasian reptiles: revision of Kapsulotaenia (Cestoda: Proteocephalidae) Zootaxa. 2020;4869:529–561. doi: 10.11646/zootaxa.4869.4.4. [DOI] [PubMed] [Google Scholar]
  16. Coquille S.C., de Chambrier A. Cairaella henrii gen. n., sp. n., a parasite of Norops trachyderma (Polychrotidae), and Ophiotaenia nicoleae sp. n. (Eucestoda: Proteocephalidea), a parasite of Thecadactylus rapicauda (Gekkonidae), in Ecuador. Folia Parasitol. 2008;55:197–206. doi: 10.14411/fp.2008.027. [DOI] [PubMed] [Google Scholar]
  17. Dereeper A., Guignon V., Blanc G., Audic S., Buffet S., Chevenet F., Dufayard J.F., Guindon S., Lefort V., Lescot M., Claverie J.M., Gascuel O. Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic Acids Res. 2008;36:W465–W469. doi: 10.1093/nar/gkn180. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Farooq M., Khanum Z., Ansar R. Helminth parasites of fresh water tortoise and poisonous snake from Sind, Pakistan. Pakistan J. Zool. 1983;15:31–37. [Google Scholar]
  19. Freze V.I. vol. V. 1965. (Essentials of Cestodology). [Proteocephalata in fish, amphibians and reptiles.] Nauka, 538 pp. (In Russian: English translation: Israel Program of Scientific Translation, Jerusalem, 1969, Cat. No. 1853, v + 597pp.) [Google Scholar]
  20. Gibson D.I., Bray R.A., Harris E.A. (Compilers) 2005. Host-Parasite Database of the Natural History Museum.http://www.nhm.ac.uk/research-curation/scientific-resources/taxonomy-systematics/host-parasites/ London. accessed on December 2020. [Google Scholar]
  21. Guindon S., Dufayard J.-F., Lefort V., Anisimova M., Hordijk W., Gascuel O. New algorithms and methods to estimate maximum likelihood phylogenies: assessing the performance of PhyML 3.0. Syst. Biol. 2010;59:307–321. doi: 10.1093/sysbio/syq010. [DOI] [PubMed] [Google Scholar]
  22. Hoang D.T., Chernomor O., von Haeseler A., Minh B.Q., Vinh L.-S. UFBoot2: improving the ultrafast bootstrap approximation. Mol. Biol. Evol. 2018;35:518–522. doi: 10.1093/molbev/msx281. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Holt B.G., Lessard J.-P., Borregaard M.K., Fritz S.A., Araújo M.B., Dimitrov D., Fabre P.-H., Graham C.-H., Graves G.R., Jønsson K.A., Nogués-Bravo D., Wang Z., Whittaker R.J., Fjeldså J., Rahbek C. An update of Wallace's zoogeographic regions of the world. Science. 2013;339:74–78. doi: 10.1126/science.1228282. [DOI] [PubMed] [Google Scholar]
  24. Kalyaanamoorthy S., Minh B.Q., Wong T.K.F., von Haeseler A., Jermiin L.S. ModelFinder: fast model selection for accurate phylogenetic estimates. Nat. Methods. 2017;14:587–589. doi: 10.1038/nmeth.4285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Katoh K., Standley D.M. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol. Biol. Evol. 2013;30:772–780. doi: 10.1093/molbev/mst010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Kearse M., Moir R., Wilson A., Stones-Havas S., Cheung M., Sturrock S., Buxton S., Cooper A., Markowitz S., Duran C., Thierer T., Ashton B., Meintjes P., Drummond A. Geneious Basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics. 2012;28:1647–1649. doi: 10.1093/bioinformatics/bts199. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Markov G.S., Bogdanov O.P. Parasite fauna of reptiles of Turkmenistan. Tr. Inst. Biol. Akad. Nauk Turkm. SSR. 1956;4:260–277. (In Russian) [Google Scholar]
  28. Markov G.S., Bogdanov O.P. Helminths and ticks – parasites of snakes of Middle Asia. Uzb. Biol. Zhurnal. 1960;2:35–41. (In Russian) [Google Scholar]
  29. Markov G.S., Bogdanov O.P., Persianova L.A. Venomous Animals of Middle Asia and their Venoms. Tashkent. 1970. Helminth fauna of saw-scaled viper; pp. 46–60. (In Russian) [Google Scholar]
  30. Marsella C.M., de Chambrier A. Ophiotaenia alessandrae sp. n. (Eucestoda: Proteocephalidea), a parasite of Hyla boans (Anura: Hylidae) from Amazonia in Ecuador. Rev. Suisse Zool. 2008;115:553–563. [Google Scholar]
  31. Miller M.A., Pfeiffer W., Schwartz T. Proceedings of the Gateway Computing Environments Workshop. GCE; 2010. Creating the CIPRES Science Gateway for inference of large phylogenetic trees; pp. 1–8. 14 Nov. 2010. [Google Scholar]
  32. Minh B.Q., Nguyen M.A.T., von Haeseler A. Ultrafast approximation for phylogenetic bootstrap. Mol. Biol. Evol. 2013;30:1188–1195. doi: 10.1093/molbev/mst024. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Nama H.S. Faunistic survey of helminth parasites of vertebrates of Rajasthan with special reference to arid zone. Sci. Rev. Arid Zone Res. 1984;2:187–225. [Google Scholar]
  34. Pook C.E., Joger U., Stümpel N., Wüster W. When continents collide: phylogeny, historical biogeography and systematics of the medically important viper genus Echis (Squamata: Serpentes: Viperidae) Mol. Phylogenet. Evol. 2009;53:792–807. doi: 10.1016/j.ympev.2009.08.002. [DOI] [PubMed] [Google Scholar]
  35. Rambeloson V.R., Ranaivoson H.C., de Chambrier A. Ophiotaenia lapata sp. n. (Eucestoda: Proteocephalidea) from Madagascar: a parasite of the endemic snake Madagascarophis colubrinus (Colubridae) Rev. Suisse Zool. 2012;119:547–559. [Google Scholar]
  36. Rego A.A. Order Proteocephalidea Mola, 1928. In: Khalil L.F., Jones A., Bray R.A., editors. Keys to the Cestode Parasites of Vertebrates. CAB International; Wallingford: 1994. pp. 257–293. [Google Scholar]
  37. La Rue G.R. A revision of the cestode family Proteocephalidae. Zool. Anz. 1911;38:473–482. [Google Scholar]
  38. La Rue G.R. A revision of the cestode family Proteocephalidae. Ill Biol. Monogr. 1914;1(1–2):3–351. [Google Scholar]
  39. Scholz T., de Chambrier A. Taxonomy and biology of proteocephalidean cestodes: current state and perspectives. Helminthologia. 2003;40:65–77. [Google Scholar]
  40. Scholz T., de Chambrier A., Kuchta R., Littlewood D.T.J., Waeschenbach A. Macrobothriotaenia ficta (Cestoda: Proteocephalidea), a parasite of sunbeam snake (Xenopeltis unicolor): example of convergent evolution. Zootaxa. 2013;3640:485–499. doi: 10.11646/zootaxa.3640.3.12. [DOI] [PubMed] [Google Scholar]
  41. Schuster R.K., Sivakumar S. 24th Conference of the World Association for the Advancement of Veterinary Parasitology, 25–29 August 2013, Perth, Australia. 2013. Reptiles as paratenic hosts of feline parasites. [Google Scholar]
  42. Sela I., Ashkenazy H., Katoh K., Pupko T. GUIDANCE2: accurate detection of unreliable alignment regions accounting for the uncertainty of multiple parameters. Nucleic Acids Res. 2015;43:W7–W14. doi: 10.1093/nar/gkv318. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Sharpilo V.P. Naukova Dumka; Kiev: 1976. Parasitic Worms of Reptiles of the Fauna of the USSR. Systematics, Chorology, Biology; p. 288. (In Russian) [Google Scholar]
  44. Trifinopoulos J., Nguyen L.T., von Haeseler A., Minh B.Q. W-IQ-TREE: a fast online phylogenetic tool for maximum likelihood analysis. Nucleic Acids Res. 2016;44:W232–W235. doi: 10.1093/nar/gkw256. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Uetz P., Freed P., Hošek J., editors. The Reptile Database. 2020. http://www.reptile-database.org Available. [Google Scholar]
  46. Velikanov V.P. Larval forms of the Acuarioidea superfamily (Nematoda, Spirurata). Parasites of reptiles in Turkmenistan. Vestnik Zool. 1984;1984(1):3–8. (In Russian) [Google Scholar]
  47. World Health Organization Venomous snakes distribution and species risk categories. 2020. https://apps.who.int/bloodproducts/snakeantivenoms/database/default.htm Available.

Articles from International Journal for Parasitology: Parasites and Wildlife are provided here courtesy of Elsevier

RESOURCES