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A new species of Atriophallophorus Deblock & Rosé, 1964 (Trematoda: Microphallidae) described from in vitro-grown adults and metacercariae from Potamopyrgus antipodarum (Gray, 1843) (Mollusca: Tateidae)

Published online by Cambridge University Press:  29 November 2019

I. Blasco-Costa*
Affiliation:
Natural History Museum of Geneva, PO Box 6434, CH-1211Geneva 6, Switzerland Department of Arctic and Marine Biology, UiT The Arctic University of Norway, Langnes, PO Box 6050, 9037Tromsø, Norway
K. Seppälä
Affiliation:
Department of Aquatic Ecology, Swiss Federal Institute of Aquatic Science and Technology (EAWAG), Dübendorf, Switzerland Institute of Integrative Biology, ETH-Zürich, Zürich, Switzerland Research Department for Limnology, University of Innsbruck, 5310Mondsee, Austria
F. Feijen
Affiliation:
Department of Aquatic Ecology, Swiss Federal Institute of Aquatic Science and Technology (EAWAG), Dübendorf, Switzerland Institute of Integrative Biology, ETH-Zürich, Zürich, Switzerland
N. Zajac
Affiliation:
Department of Aquatic Ecology, Swiss Federal Institute of Aquatic Science and Technology (EAWAG), Dübendorf, Switzerland Institute of Integrative Biology, ETH-Zürich, Zürich, Switzerland
K. Klappert
Affiliation:
Department of Aquatic Ecology, Swiss Federal Institute of Aquatic Science and Technology (EAWAG), Dübendorf, Switzerland Institute of Integrative Biology, ETH-Zürich, Zürich, Switzerland
J. Jokela
Affiliation:
Department of Aquatic Ecology, Swiss Federal Institute of Aquatic Science and Technology (EAWAG), Dübendorf, Switzerland Institute of Integrative Biology, ETH-Zürich, Zürich, Switzerland
*
Author for correspondence: I. Blasco-Costa, E-mail: isa.blasco.costa@gmail.com
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Abstract

The adult and metacercaria life stages of a new species of the microphallid genus Atriophallophorus Deblock & Rosé, 1964 are described from specimens collected at Lake Alexandrina (South Island, New Zealand). In addition to molecular analyses of ribosomal and mitochondrial genes, metacercariae of Atriophallophorus winterbourni n. sp. from the snail host Potamopyrgus antipodarum (Gray) were grown in vitro to characterize internal and external morphology of adults using light and scanning electron microscopy and histological techniques. Atriophallophorus winterbourni n. sp. is readily distinguishable from Atriophallophorus coxiellae Smith, 1973 by having a different structure of the prostatic chamber, sub-circular and dorsal to genital atrium, rather than cylindrical, fibrous, elongate and placed between the seminal vesicle and the genital atrium. The new species is most similar to Atriophallophorus minutus (Price, 1934) with regards to the prostatic chamber and the morphometric data, but possesses elongate-oval testes and subtriangular ovary rather than oval and transversely oval in A. minutus. Phylogenetic analyses including sequence data for A. winterbourni n. sp. suggested a congeneric relationship of the new species to a hitherto undescribed metacercariae reported from Australia, both forming a strongly supported clade closely related to Microphallus and Levinseniella. In addition, we provide an amended diagnosis of Atriophallophorus to accommodate the new species and confirm the sinistral interruption of the outer rim of the ventral sucker caused by the protrusion of the dextral parietal atrial scale at the base of the phallus.

Type
Research Paper
Copyright
Copyright © Cambridge University Press 2019

Introduction

A microphallid species reported as Microphallus sp. from New Zealand has been largely used as a model species in parasitology and evolutionary studies for more than three decades (e.g. Lively, Reference Lively1987; Dybdahl & Lively, Reference Dybdahl and Lively1998; Lively & Dybdahl, Reference Lively and Dybdahl2000; Jokela et al., Reference Jokela, Dybdahl and Lively2009; Gibson et al., Reference Gibson, Jokela and Lively2016a). Broad knowledge has accumulated on the role of this parasite in coevolutionary dynamics with the host populations. Particularly, in this case the coevolutionary dynamics are suggested to favour sexual reproduction by the host (i.e. ‘parasite hypothesis’ or ‘Red Queen hypothesis’ for maintenance of sex; Jaenike (Reference Jaenike1978); Hamilton (Reference Hamilton1980); Lively (Reference Lively2016)). However, basic knowledge such as the identity of the species, its genetic diversity and phylogenetic affinities are still lacking, with the consequences that it may be difficult to judge the source of heterogeneity in infection experiments and field surveys.

Microphallids are a diverse and cosmopolitan group of very small worms, typically found in the intestine of birds. Second intermediate hosts tend to be crustaceans, but several species present abbreviated life cycles where the metacercariae encyst within the mollusc first intermediate host. Their metacercariae are usually identical to the adult stage in the definitive host. In the definitive host, microphallids tend to mature in a few days and live only for several weeks (Galaktionov & Dobrovolskij, Reference Galaktionov and Dobrovolskij2003). This feature of microphallids makes them amenable to in vitro culture as a method to obtain the mature adults and be able to characterize and describe new species. Such an alternative is, nowadays, desirable, considering the limitations of obtaining permits to collect potential vertebrate definitive hosts, which are in many cases protected by national or international regulations (Blasco-Costa & Poulin, Reference Blasco-Costa and Poulin2017).

The aim of this study is to morphologically characterize and describe the adult and metacercaria life stages of this microphallid, which represents a new species of the genus Atriophallophorus Deblock & Rosé, 1964 instead of Microphallus Ward, 1901 as had been previously reported. Metacercariae of microphallid specimens from the snail host Potamopyrgus antipodarum (Gray) were grown in vitro and the adults were analysed with light microscopy, scanning electron microscopy (SEM) and histological techniques to characterize their internal and external morphology. In addition, this study provides an amended diagnosis of Atriophallophorus, molecular data for two nuclear and two mitochondrial markers of the new species and an evaluation of its relationship with other microphallids.

Material and methods

Specimens

Potamopyrgus antipodarum snails were collected using a kick net and snorkelling from two sites in Lake Alexandrina (South Island, New Zealand), site ‘Swamp’ (−43.962102, 170.441728) and site ‘JMS’ (−43.937199, 170.459495). Live metacercariae were dissected from four infected snails from each locality and allowed to excyst in Tyrode's salt solution (Sigma, Buchs, Switzerland) with pancreatin (3 mg/ml; Sigma) and penicillin–streptomycin–neomycin solution (8% v/v; Sigma) at 39°C. Excysted juvenile worms were washed twice with Tyrodes's salt solution supplemented with penicillin–streptomycin–neomycin solution (8% v/v). Then, juveniles from the same locality were pooled and transferred to culture medium of RPMI 1640 (Gibco, ThermoFisher Scientific, Waltham (MA) U.S.) supplemented with horse serum (20% v/v, Gibco), penicillin–streptomycin–neomycin solution (8% v/v), HEPES buffer (25 mm; Gibco) and Amphotericin B solution (0.25 mg/ml; Sigma). Juveniles were incubated at 39°C and cultured for up to three days (72 h). Fresh culture media was changed daily. Metacercariae right after hatching and in vitro-grown adults after 24 h, 48 h and 72 h of culturing were fixed in hot saline and preserved in 75% ethanol for later morphological examination. A subsample of specimens was preserved in hot formalin for examination using SEM and histological analysis. Additionally, five metacercariae from each site (from a pool of four snails each) were preserved in 100% ethanol for molecular analyses.

Morphological data

Metacercariae and adult specimens grown in vitro were stained using iron acetocarmine, dehydrated through a graded ethanol series, cleared in dimethyl phtalate and examined as permanent mounts in Canada balsam. Figures were made using a drawing tube mounted on a Zeiss light compound microscope at ×1600 magnification. Measurements of the specimens were taken from drawings at ×640 magnification. Five specimens were dehydrated in a graded ethanol series, critical point-dried and sputter-coated with gold for SEM examination using a Zeiss DSM 940A (Zeiss AG, Oberkochen, Germany) at an accelerating voltage of 5 kV. Three specimens were used for histology after dehydration through a graded ethanol series, followed by propylene oxide and immersed in Epon resin. Histological sections of 1 µm were made with an ultramicrotome Reichert-Jung Ultracut E (Leica Microsystems, Heerbrugg, Switzerland), stained with toluidine blue 0.75% and examined using the light microscope mentioned previously. All measurements in the text are in micrometres unless otherwise stated, and are given as the range followed by the mean ± standard deviation. Permanent mounts of the type material and SEM preparations are deposited in the Platyhelminthes collection of the Natural History Museum of Geneva, the Institute of Parasitology of the Academy of Sciences of the Czech Republic and the Otago Museum, Dunedin, New Zealand.

Molecular data

Four metacercariae isolates from Swamp and five from JMS originating from a pool of four infected P. antipodarum from each site were characterized molecularly. Due to the small size of these specimens, it was impossible to keep hologenophores of the sequenced specimens. Nonetheless, the specimens used for morphological description herein represent paragenophores and likely genetic clones of the sequenced material. Genomic DNA was extracted from ethanol-fixed metacercariae isolates in 200 µl of a 5% suspension of Chelex® in deionized water and containing 0.1 mg/ml proteinase K followed by incubation at 56°C for 5 h, boiling at 90°C for 8 min and centrifugation at 14,000 g for 10 min. Partial fragments were amplified of the large ribosomal subunit (28S) (1800 bp; primers U178F: 5′-GCA CCC GCT GAA YTT AAG-3′ and L1642R: 5′-CCA GCG CCA TCC ATT TTC A-3′; Lockyer et al., Reference Lockyer, Olson and Littlewood2003) and internal transcribed spacer 2 (ITS2) (500 bp; primers 3S: 5′-GTA CCG GTG GAT CAC GTG GCT AGT G-3′ and ITS2·2: 5′-CCT GGT TAG TTT CTT TTC CTC CGC-3′ (Morgan & Blair, Reference Morgan and Blair1995; Cribb et al., Reference Cribb, Anderson, Adlard and Bray1998)). Additionally, two fragments of mitochondrial genes (mt) were amplified: cytochrome oxidase subunit I (cox I) (1050 bp; primers JB3: 5′-TTTTTTGGGCATCCTGAGGTTTAT -3′ and microph_rev: 5′- AAT CAT GAT GCA AAA GG-3′ (Bowles et al., Reference Bowles, Hope, Tiu, Liu and McManus1993; newly designed)) and nicotinamide adenine dinucleotide dehydrogenase subunit 5 (NADH5) (744 bp; primers F2_micND5: 5′-CTTCAACCTTGGTTGCTGCC-3′ and R2_micND5: 5′-TCCCAACGAAACCTAAAACTGC-3′ (newly designed)).

Polymerase chain reaction (PCR) amplifications were performed in 20 µl reactions containing 2 µl of extraction supernatant (~10–20 ng of template DNA), 2× MyFi™ Mix (Bioline France, France; containing DNA polymerase, dNTPs, MgCl2 and enhancers at optimal concentrations) and 0.4 µm of each primer combination. Thermocycling conditions used for amplification of the rDNA regions followed Galaktionov et al. (Reference Galaktionov, Blasco-Costa and Olson2012). The following thermocycling profile was used for amplification of the mt cox I and NADH5 fragments: denaturation (95°C for 3 min); 38 cycles of amplification (94°C for 50 s, 52°C for 30 s and 72°C for 1 min); and 4 min extension step at 72°C. PCR amplicons were purified prior to sequencing using exonuclease I and shrimp alkaline phosphatase enzymes (Werle et al., Reference Werle, Schneider, Renner, Volker and Fiehn1994). Amplicons were cycle-sequenced from both strands using PCR primers and an internal primer for the 28S fragment (L1200R: 5′-GCA TAG TTC ACC ATC TTT CGG-3′; Littlewood et al., Reference Littlewood, Curini-Galletti and Herniou2000) at the commercial facility Macrogen (Amsterdam, The Netherlands). Contiguous sequences were assembled and edited using Geneious® (v. 8.1 Biomatters Ltd., Auckland, New Zealand) and submitted to GenBank (see accession numbers in table 1).

Table 1. List of taxa included in the phylogenetic analyses, GenBank accession numbers and references.

Molecular analyses

Newly generated sequences for the 28S rDNA and the ITS2 fragments were aligned in two independent datasets, together with the published sequences of other microphallids from GenBank (see accession numbers in table 1). The sequences were aligned using default parameters of MAFFT implemented in Geneious®, and the extremes of the alignment were trimmed to match the shortest sequences. The 28S dataset (1280 bp long) included 14 representative sequences of Microphallus spp., 12 of Maritrema spp., one each of Levinseniella, Longiductotrema and an unidentified microphallid of Kudlai et al. (Reference Kudlai, Cutmore and Cribb2015) and a sequence labelled as Microphallus fusiformis (which should be disregarded as a species of Microphallus; see Kudlai et al., Reference Kudlai, Cutmore and Cribb2015) retrieved from GenBank (table 1). Additionally, five sequences of species belonging to sister families of the Microphallidae – i.e. Lecithodendriidae, Pleurogenidae and Prosthogonimidae in the Microphalloidea – and three sequences of species in the Plagiorchioidea were retrieved from GenBank and included as outgroups. The ITS2 dataset (401 bp long) included ten representative sequences of Microphallus spp.; nine sequences of Maritrema; one each of Levinseniella, Probolocoryphe and Longiductotrema; and one of an unidentified microphallid of Kudlai et al. (Reference Kudlai, Cutmore and Cribb2015). The phylogenetic analyses were run on the two datasets individually under the maximum likelihood (ML) and Bayesian inference (BI) criteria, employing the nucleotide substitution model GTR+Γ. ML analyses were conducted using the program RAxML v. 8.2 (Stamatakis, Reference Stamatakis2014). All model parameters, bootstrap nodal support values (1000 repetitions) and an extended majority-rule consensus topology were estimated using RAxML. BI trees were constructed using MrBayes v. 3.2 (Ronquist et al., Reference Ronquist, Teslenko and van der Mark2012), running two independent Markov Chain Monte Carlo runs of four chains with standard settings for 107 generations and sampling tree topologies every 103 generation. Burn-in periods were set automatically to 25% generations, ensuring the remaining trees were obtained after values for standard deviation of split frequencies were <0.01. A majority-rule consensus topology and nodal support estimated as posterior probability values (Huelsenbeck et al., Reference Huelsenbeck, Ronquist, Nielsen and Bollback2001) were calculated from the remaining trees. All MrBayes and RAxML analyses were performed on the computational resource CIPRES (Miller et al., Reference Miller, Pfeiffer and Schwartz2010). Genetic divergences amongst taxa were calculated as uncorrected p-distances for each gene region using MEGA v. X (Kumar et al., Reference Kumar, Stecher, Li, Knyaz and Tamura2018).

Results

Family: Microphallidae Ward, 1901

Atriophallophorus Deblock & Rosé, 1964

Atriophallophorus winterbourni n. sp.

Taxonomic summary

Description of adult (figs 1a–c and 2a–h; table 2; supplementary table S1)

Fig. 1. Atriophallophorus winterbourni n. sp. (a) Illustration of the holotype, 24 h in vitro grown adult in ventral view. (b) Microphotographs of the terminal genitalia of the holotype using light microscopy. (c) Histological oblique section of a paratype at the level of the ventral sucker. (d) Microphotograph of an encysted metacercaria ex Potamopyrgus antipodarum (Gray). Abbreviations: c, caeca; ga, genital atrium; pc, prostatic chamber; ph, phallus; sv, seminal vesicle; vs, ventral sucker.

Fig. 2. Scanning electron micrographs of Atriophallophorus winterbourni n. sp. (a) Adult. (b) Adult with phallus protruded. (c) Palmate spines on the ventral surface of the body. (d) Detail of the oral sucker, arrows point at a gland opening and a sensory papilla surrounding the oral sucker. (e) Detail of the outer rim of the ventral sucker with spination and the parietal atrial scale at the basis of the phallus. (f) Outer rim of the ventral sucker interrupted sinistrally by the opening for the genital pore with arrows pointing at glands. (g) Tip of the phallus evaginated, which appears as a flower-like structure when invaginated. (h) Detail of the configuration of the protruded phallus and the sinistrally interrupted outer rim of the ventral sucker.

Table 2. Comparative metrical data for Atriophallophorus spp.

a Estimated from the published illustration. n, number of specimens measured; SD, standard deviation of the mean.

(Based on whole mounts of 29 gravid specimens grown in vitro culture for 24–72 h and SEM preparations. Measurements provided as range and mean ± standard deviation for the type series, variation associated with specimens from each age class (duration of culture) are provided separately in supplementary table S1.)

Body minute, triangular, often curved concave ventrally (body-width-to-length ratio 1:1.3–1.9 (1:1.6 ± 0.4)) with maximum width at posterior level of testes, 145–200 (167 ± 12) × 85–115 (100 ± 8). Tegument bears spines, glands and sensory papillae. Spines palmate, smooth, 5–9 prongs; present in lateral margins and alongside midline, separated by two narrow ventro-lateral regions devoid of spines, sparser towards posterior extremity on dorsal side; anterior forebody spines width 1.5–1.8, inter-spine space 0.6–0.8; mid-forebody spines width 1.9–2.2, inter-spines space 0.6–1; anterior dorsal spines width 2.2–2.3, inter-spines space 1.0–1.5; mid-dorsal spines width 1.5–1.8, inter-spines space 2.1–4.1; lateral posterior dorsal spines width 1.6–1.8, inter-spines space 1.2–1.5. Forebody 81–125 (97 ± 9) long, representing 52–63% (58 ± 2%) of body length. Glands and sensory papillae at anterior extremity, surrounding oral sucker (>14) and lateral margins of body.

Oral sucker subterminal, spherical, 21–29 (25 ± 2) × 21–29 (25 ± 2). Ventral sucker at two-thirds of body length, subspherical, complete, 24–30 (27 ± 1) × 22–33 (28 ± 2); outer rim crescent, interrupted sinistrally by genital pore, bearing spines and nine glands; oral-sucker-to-ventral-sucker-length ratio 1: 0.9–1.2 (1:1.1 ± 0.2), width ratio 1: 0.9–1.3 (1:1.1 ± 0.1). Pre-pharynx absent. Pharynx small, oval 11–17 (14 ± 2) × 11–15 (13 ± 1). Pharynx-length-to-oral-sucker-length ratio 1:1.5–2.4 (1:1.7 ± 0.2). Oesophagus 29–46 (36 ± 5) long. Intestinal bifurcation pre-equatorial, immediately anterior to seminal vesicle. Caeca as long as oesophagus, widely divergent, extend to anterior margin of testes.

Testes two, postovarian, symmetrical, lateral, somewhat diagonal, smooth, slightly elongate-oval, right testis 18–29 (25 ± 4) × 17–28 (20 ± 3); left testis 18–33 (25 ± 5) × 14–24 (19 ± 3). Seminal vesicle arcuate, transversely oval, intercaecal in mid-body, overlapping anterior margin of ventral sucker dorsally, 12–24 (17 ± 3) × 24–39 (32 ± 3). Seminal-vesicle-length-to-ventral-sucker-length ratio 1:1.1–2.2 (1:1.7 ± 0.3). Seminal proximal duct long, entering prostatic chamber (i.e. phallophorus). Prostatic chamber subspherical, with loose fibres, sinistral to ventral sucker, dorsal to genital atrium, reaching proximal part of seminal vesicle. Prostatic glands not observed. Ejaculatory duct sinuous, enters prostatic chamber, opens to small papillae dorso-sinistral in genital atrium, together with prostatic ducts running through to periphery of male duct. Phallus of ‘microphalloid’-type, glabrous, turgid, evaginable, with ejaculatory duct in axis, with large dextral triangular distal scale at the base. Genital atrium surrounding phallus, 16–24 (20 ± 3), genital pore large, sinistral to ventral sucker.

Ovary dextral to ventral sucker, pre-testicular, ventral to caeca, adjacent to or slightly overlapping ventral sucker laterally, subtriangular with large cells, 13–26 (21 ± 3) × 17–34 (25 ± 4). Oötype inter-testicular, slightly dextral posterior to ovary and ventral sucker. Mehlis' gland surrounding oötype posteriorly. Laurer's canal not observed. Uterus confined posterior to mid-level of ventral sucker, overlapping testes ventrally. Metraterm thin-walled, with widened opening into sinistral wall of genital atrium. Vitellarium in two compact clusters of follicles, disaggregated in older specimens (after 24 h cultured), para-, post-testicular or overlapping testes, converging into seminal receptacle next to oötype. Eggs few, large. Excretory vesicle obscured by vitellarium masses and eggs. Flame-cell formula not observed.

Description of metacercaria (fig 1d; table 2; supplementary table S1)

Overall form highly developed and consistent with adult anatomy, except for the absence of eggs.

(Measurements based on 18 encysted specimens; measurements provided as range followed by mean ± standard deviation.)

Metacercaria folded within small spherical, translucent cyst, 114−130 × 106−120 (123 × 113 ± 4). Cyst wall consisting of three or more hyaline layers, 5−6 (5 ± 0.5) thick. Metacercaria encysted in the gonads of P. antipodarum.

Genetic affinities

Two distinct genotypes for the 28S rRNA gene were obtained from the sequences of the nine specimens analysed. The two genotypes differed in two transitions at the nucleotide positions 281, a cytosine in the sequence of specimens originated from Swamp and a thymine in specimens from JMS and 1358, adenine and guanine, respectively. However, all specimens shared the same sequence for the ITS2 region and the mt cox I. Independent phylogenetic analyses of the 28S rDNA and ITS2 regions using BI and ML methods showed congruent results (figs 3 and 4), with the Microphallidae as monophyletic and A. winterbourni embedded within. A. winterbourni appeared as sister taxon to the sequence of an undescribed microphallid from Australia by Kudlai et al. (Reference Kudlai, Cutmore and Cribb2015). In the 28S phylogenetic tree (fig. 3), the sequence of M. fusiformis appeared closely related to A. winterbourni and the undescribed microphallid with strong nodal support, although considerably divergent. In all trees, Atriophallophorus appears closely related to Microphallus and Levinseniella, but the sister relationship among these three genera could not be established due to the low support of an internal node within the clade (figs 3 and 4).

Fig. 3. Phylogenetic relationships for representatives of the family Microphallidae, inferred by maximum likelihood analysis of 28S rDNA sequence data. The newly generated sequences are indicated in bold. Values on the branches correspond to posterior probabilities >0.95 followed by bootstrap support >60. Values below these thresholds were not reported.

Fig. 4. Phylogram for representatives of the family Microphallidae, inferred by maximum likelihood analysis of sequence data for the internal transcribed spacer 2 of the rRNA genes. The newly generated sequences are indicated in bold. Values on the branches correspond to posterior probabilities >0.95 followed by bootstrap support >60. Values below these thresholds were not reported.

Genetic divergence between A. winterbourni and the unidentified microphallid from Australia was 1.2–1.3% for the 28S and 3.2% for the ITS2, which are within the lower limit of the range observed among congeneric species of Microphallus (28S: 0.8–9.1%; ITS2: 1.0–9.9%) and Maritrema (28S: 0.6–9.1%; ITS2: 0.3–12.3%). The 28S sequences of A. winterbourni and the unidentified microphallid diverged 10.5–10.8% from the sequence of M. fusiformis, which fell within range of intergeneric distances in the 28S region of microphallids (MaritremaMicrophallus: 7.5–12.6%; MicrophallusLevinseniella: 5.5–10.0%; MaritremaLevinseniella: 9.1–11.4%; LongiductotremaLevinseniella: 7.7%; LongiductotremaMaritrema: 7.8–11.1%; LongiductotremaMicrophallus: 6.5–10.6%).

Sequence data for the COI marker resulted in identical sequences for all specimens (five from JMS and three from Swamp; one sample from swamp did not amplify and another one produced chromatograms with double peaks), whereas sequences for the NADH5 marker showed variability at the position 191, with specimens from Swamp having a thymine and specimens from JMS a cytosine.

Discussion

Species differentiation

Following the key to the Microphallidae provided by Deblock (Reference Deblock, Bray, Gibson and Jones2008), the new species described from in vitro grown adults conforms to the general morphology of the Microphallidae in having a typically very small body, longer than broad, densely covered with squamous spines, suckers well-separated and sub-equal, short digestive tract with divergent caeca not extending posteriorly beyond ventral sucker. Ovary pretesticular, in the opposite side of the body to the genital pore. Two testes, lateral and symmetrical, with male terminal genitalia intercaecal and anterior to ventral sucker. These specimens fit to the diagnosis of the supersubfamily Microphallidi Ward, 1901 by having the terminal genitalia free in the parenchyma, and the subfamily Microphallinae Ward, 1901 by presenting a genital atrium that closely envelops the phallus. However, the opening of the genital atrium and the presence of the scale at the base of the phallus of the new species were found to alter the outer rim of the ventral sucker. Whereas the latter feature is characteristic of the monotypic Endocotylinae, our specimens differ from the genus Endocotyle in lacking a connection between the cavity of the sucker and the genital pore, and presenting a scale at the base of a phallus of the ‘microphalloid-type’. Our specimens fall within the Microphallini tribe by having a fleshy and muscular phallus, and the opening of the metraterm on the sinistral wall of the genital atrium. The new species exhibits features consistent with the genus Atriophallophorus: body of triangular shape; male genital pouch absent; proximal ejaculatory ducts very long and entering a large prostatic chamber (described as ‘phallophorus’ by Deblock & Rosé (Reference Deblock and Rosé1964)), subcircular and similar in size to the ventral sucker and dorsal to the genital atrium, with long prostatic ducts running through the periphery of the male duct; genital atrium containing a phallus of the ‘microphalloid-type’ and, at its base, a large dextral parietal atrial scale often protruding from the genital pore; a metraterm that widens at its opening into the sinistral wall of the genital atrium; and few but large eggs.

Currently the genus contains only two species, A. minutus (Price, 1934) and A. coxiellae Smith, 1973. Atriophallophorus coxiellae was described from metacercariae infecting Coxiella badgerensis in a freshwater lake in Tasmania (Smith, Reference Smith1973). However, the metacercaria of A. winterbourni n. sp. is readily distinguishable from A. coxiellae by having a different structure of the prostatic chamber, sub-circular and dorsal to genital atrium, rather than cylindrical, fibrous, elongate and placed between the seminal vesicle and the genital atrium, as well as smaller pharynx and testes, and somewhat smaller oral sucker and ovary, which barely overlap the lower limit of the range of A. coxiellae.

The new species is most similar to A. minutus with regards to the prostatic chamber and the morphometric data. But A. minutus possesses oval testes and a transversely oval ovary, versus elongate-oval and subtriangular, respectively, in A. winterbourni. In addition, the original description of A. minutus, based on adult specimens from Aythya affinis from the Caribbean (Price, Reference Price1934), reported A. minutus had a shorter oesophagus than the new species, although this feature might vary greatly depending on the position of the specimen. Morphometric data provided by Stunkard (Reference Stunkard1958) for the redescription of A. minutus were based on adults experimentally grown in white mice from metacercariae infecting Hydrobia minuta and Amnicola limosa along the east coast of the USA. However, Stunkard's specimens were measured alive, whereas our measurements were taken from fixed, stained and mounted worms. Therefore, the morphometric data are not directly comparable. Furthermore, Stunkard noted that fixed specimens were slightly smaller than alive, which suggests that if the measurements of A. minutus were taken after mounting they would be slightly smaller than those of A. winterbourni. Nonetheless, live specimens of A. minutus also show smaller dimensions for several metrical features (body length, ventral sucker width and testes width) extending outside the lower range for A. winterbourni, and eggs three times more numerous and with size overlapping the lower range of variation of the new species.

Metrical data of A. minutus described by Deblock & Rosé (Reference Deblock and Rosé1964) from France supports that A. minutus differs from A. winterbourni. Whereas the specimens examined by Deblock & Rosé (Reference Deblock and Rosé1964) were of comparable size to the largest in our sample, their ovary width is smaller than that of A. winterbourni, and the ventral sucker width overlapped and extended below the lower range of variation of A. winterbourni. Atriophallophorus minutus also shows smaller values for the ratios oral sucker length to ventral sucker length and seminal vesicle length to ventral sucker length (estimated from the original illustrations) than A. winterbourni.

In vitro cultivation of the metacercaria and adult allowed us to observe developmental changes through time in the shape of the vitellarium, the seminal vesicle and testes (see also supplementary table S1). In metacercariae, the testes are well visible, the seminal vesicle is small, indistinct and quite empty, and the vitellarium forms two masses of tight follicles. After 24 h of growth, the adults are already gravid, testes become indistinct, the seminal vesicle is full and extended, and the follicles constituting the vitellarium masses start to disaggregate slightly. After 48 h or more, the testes are faintly visible, the seminal vesicle stays fully extended and the vitellarium follicles are drawn further apart in the posterior region of the body. When descriptions are based on specimens collected from the wild birds, this variation in the sample of specimens should be taken into account.

Previous researchers working on Atriophallophorus spp. have emphasized the difficulty of observing the features of specimens of such tiny size. Thus, it is highly recommended to describe new species with the support of genetic data and using a holistic biological approach (Blasco-Costa et al., Reference Blasco-Costa, Cutmore, Miller and Nolan2016a). Despite A. winterbourni showing subtle morphological differences from A. minutus, we consider them sufficient to distinguish the new species given the intrinsic difficulties of this group. Molecular results showed, convincingly, that specimens of A. winterbourni belong to a different microphallid genus to those already represented by sequence data. The new species appeared sister to an unidentified microphallid recovered from Posticobia brazieri (Smith) (Gastropoda, Tateidae) and Caridina indistincta Calman (Decapoda, Atyidae) in Australia by Kudlai et al. (Reference Kudlai, Cutmore and Cribb2015). These authors suggested that their specimens were likely closely related to (or even conspecific) with the material reported as Microphallus sp. ‘livelyi’ by Hechinger (Reference Hechinger2012), which is considered a junior synonym of A. winterbourni herein. Divergence between our sequences and those of the Australian microphallid of Kudlai et al. (Reference Kudlai, Cutmore and Cribb2015) suggests that they represent two distinct but congeneric species, as the authors anticipated.

The examination of specimens of A. winterbourni with SEM has allowed us to confirm a sinistral interruption of the outer rim of the ventral sucker caused by the opening of the genital pore and protrusion of the dextral parietal atrial scale, while light microscopy and histological sections show that, internally, the ventral sucker is complete. Deblock & Rosé (Reference Deblock and Rosé1964) mentioned the possibility of the scale being united to the rim of the ventral sucker (p. 229). However, this feature went unnoticed and was never confirmed by a later redescription or new species description for the genus, nor was it mentioned in the most recent diagnosis provided by Deblock (Reference Deblock, Bray, Gibson and Jones2008). Since the three species known for the genus so far are characterized by the presence of the large atrial scale, it is likely that in all three cases it has resulted in the same modification of the ventral sucker. Thus, we consider this feature of diagnostic value for the genus and amend herein the generic diagnosis of Atriophallophorus as follows.

Atriophallophorus Deblock & Rosé, 1964

Generic diagnosis

Body piriform or triangular, small (150–200 µm). Resembles Microphallus. Ventral sucker postequatorial, outer rim interrupted sinistrally by genital pore. Oesophagus medium or short. Caeca short, divergent, in mid-body. Testes symmetrical in hindbody. Male genital pouch absent. Seminal vesicle intercaecal in mid-body, ovoid; prostatic gland caps distal part of seminal vesicle; proximal ejaculatory duct very long, supported by envelope acting as large prostatic chamber, either: (i) subcircular with diameter of ventral sucker, dorsal to genital atrium (formation described as ‘phallophorus’ (apparently bearing phallus)) and with long prostatic duct running through to periphery of male duct inside phallus; or (ii) cylindrical, fibrous, elongate between seminal vesicle and genital atrium, adjacent to margin of ventral sucker; bundle of long prostatic ducts enter as far as mid-part of cylindrical prostatic chamber (not inside phallus). Phallus of ‘microphalloid’-type, more or less turgid, with ejaculatory duct in axis. Genital atrium present, envelopes phallus, with enormous dextral parietal atrial scale; genital pore sinistral to ventral sucker. Ovary dextral to ventral sucker. Uterus postcaecal, with few coils around testes. Metraterm long, with widened opening into sinistral wall of genital atrium. Eggs not numerous, relatively large. Vitellarium formed of two clusters of follicles, paratesticular and post-testicular in hindbody; vitelline ducts short, arched, post-testicular. Excretory vesicle short, Y-shaped, post-testicular. In intestine of birds (Anseriformes, Charadriiformes); cosmopolitan.

  • Type species. Atriophallophorus minutus (Price, 1934).

  • Synonym. Atriophallophorus samarae Deblock & Rosé, 1964.

  • Synonym lapsus calami. Atriophallus samarae Deblock & Rosé, 1964 (Fig. 1).

Life cycle and putative definitive hosts

Compared to other microphallid genera, the known species diversity of Atriophalophorus is quite low. So far, the four members of this genus (including the yet undescribed but molecularly characterized lineage from Australia of Kudlai et al. (Reference Kudlai, Cutmore and Cribb2015)) show an abbreviated life cycle with the absence of the cercarial stage. Furthermore, A. winterbourni also lacks daughter sporocyst parthenitae-bearing germ cells (or has a very reduced life span), so that germ balls and embryonic metacercariae are observed free in the visceral mass of the snail host and appear to develop directly into encysted metacercariae, as reported by Krist & Lively (Reference Krist and Lively1998).

Based on the experimental exposure of snails to the faeces of different waterfowl species from Lake Alexandrina, Osnas & Lively (Reference Osnas and Lively2011) concluded that the likely definitive hosts of A. winterbourni are mallard ducks (Anas platyrhynchos L.), grey ducks (Anas superciliosa Gmelin), their hybrids and the New Zealand scaup (Aythya novaeseelandiae Gmelin). Their conclusion agrees with the known distribution of both the putative hosts and A. winterbourni throughout the South Island. Furthermore, recent studies of the parasite fauna of mallards have discovered new species to science of microphallids and strigeids in New Zealand (Presswell et al., Reference Presswell, Blasco-Costa and Kostadinova2014; Blasco-Costa et al., Reference Blasco-Costa, Poulin and Presswell2016b). Altogether, these results highlight the still scarce knowledge on the parasites of birds, the most diverse group of vertebrates native to New Zealand, and the need for more biodiversity studies to address this gap.

Supplementary material

To view supplementary material for this article, please visit https://doi.org/10.1017/S0022149X19000993.

Acknowledgements

We thank Pilar Ruga Fahy from the University Medical Centre of the Pôle Facultaire de Microscopie Ultrastructurale at the University of Geneva for carrying out the histological sections and procedures, Janik Pralong (Natural History Museum of Geneva) for preparation of the permanent mounts and Dr André Piuz (Natural History Museum of Geneva) for his help with the SEM. We are also grateful to Dr Rodney Bray and Tomáš Scholz for discussions on the interpretation of the morphology, and one anonymous reviewer and the editor for very useful comments that helped us improve the manuscript.

Financial support

This work was supported by the Natural History Museum of Geneva, and indirectly by the Swiss National Science Foundation (IBC, grant number 31003A_169211/1).

Conflicts of interest

None.

Ethical standards

The authors assert that all procedures contributing to this work comply with the ethical standards of Switzerland and New Zealand and our institutional guides on the care and use of wild invertebrate animals.

References

Al-Kandari, WY and Al-Bustan, SA (2010) Molecular identification of Probolocoryphe uca (Sarkisian, 1957; Digenea: Microphallidae) from Kuwait Bay using ITS1 and ITS2 sequences. Parasitology Research 106, 11891195.CrossRefGoogle ScholarPubMed
Al-Kandari, WY, Al-Bustan, SA and Alnaqeeb, M (2011) Ribosomal DNA sequence characterization of Maritrema cf. eroliae Yamaguti, 1939 (Digenea: Microphallidae) and its life cycle. Journal of Parasitology 97, 10671074.CrossRefGoogle ScholarPubMed
Bankers, L and Neiman, M (2017) De novo transcriptome characterization of a sterilizing trematode parasite (Microphallus sp.) from two species of New Zealand snails. G3–Genes Genomes Genetics 7, 871880.Google ScholarPubMed
Bankers, L, Fields, P, McElroy, KE, Boore, JL, Logsdon, JM and Neiman, M (2017) Genomic evidence for population-specific responses to co-evolving parasites in a New Zealand freshwater snail. Molecular Ecology 26, 36633675.CrossRefGoogle Scholar
Blasco-Costa, I and Poulin, R (2017) Parasite life-cycle studies: a plea to resurrect an old parasitological tradition. Journal of Helminthology 91, 647656.CrossRefGoogle ScholarPubMed
Blasco-Costa, I, Cutmore, SC, Miller, TL and Nolan, MJ (2016a) Molecular approaches to trematode systematics: ‘best practice’ and implications for future study. Systematic Parasitology 93, 295306.CrossRefGoogle Scholar
Blasco-Costa, I, Poulin, R and Presswell, B (2016b) Species of Apatemon Szidat, 1928 and Australapatemon Sudarikov, 1959 (Trematoda: Strigeidae) from New Zealand: Linking and characterising life cycle stages with morphology and molecules. Parasitology Research 115, 271289.CrossRefGoogle Scholar
Bowles, J, Hope, M, Tiu, WU, Liu, X and McManus, DP (1993) Nuclear and mitochondrial genetic markers highly conserved between Chinese and Philippine Schistosoma japonicum. Acta Tropica 55, 217229.CrossRefGoogle ScholarPubMed
Cribb, TH, Anderson, GR, Adlard, RD and Bray, RA (1998) A DNA-based demonstration of a three-host life-cycle for the Bivesiculidae (Platyhelminthes: Digenea). International Journal for Parasitology 28, 17911795.CrossRefGoogle Scholar
Deblock, S (2008) The Microphallidae Ward, 1901. pp. 451492in Bray, RA, Gibson, DI and Jones, A (Eds) Keys to the Trematoda. Vol. 3. London, CAB International and The Natural History Museum.CrossRefGoogle Scholar
Deblock, S and Rosé, F (1964) Contribution a l'etudes des Microphallidae Travassos, 1920 (Trematoda) des oiseaux de France. VIII Creation du genre Atriophallophorus, parasite de canard sauvages. Bulletin de la Societe Zoologique de France 89, 225230.Google Scholar
Diaz, JI and Cremonte, F (2010) Development from metacercaria to adult of a new species of Maritrema (Digenea: Microphallidae) parasitic in the kelp gull, Larus dominicanus, from the Patagonian coast, Argentina. Journal of Parasitology 96, 740745.CrossRefGoogle ScholarPubMed
Dybdahl, MF and Krist, AC (2004) Genotypic vs. condition effects on parasite-driven rare advantage. Journal of Evolutionary Biology 17, 967973.CrossRefGoogle ScholarPubMed
Dybdahl, MF and Lively, CM (1996) The geography of coevolution: comparative population structures for a snail and its trematode parasite. Evolution 50, 22642275.CrossRefGoogle ScholarPubMed
Dybdahl, MF and Lively, CM (1998) Host-parasite coevolution: evidence for rare advantage and time-lagged selection in a natural population. Evolution 52, 10571066.CrossRefGoogle Scholar
Dybdahl, MF, Jokela, J, Delph, LF, Koskella, B and Lively, CM (2008) Hybrid fitness in a locally adapted parasite. American Naturalist 172, 772782.CrossRefGoogle Scholar
Fromme, AE and Dybdahl, MF (2006) Resistance in introduced populations of a freshwater snail to native range parasites. Journal of Evolutionary Biology 19, 19481955.CrossRefGoogle Scholar
Galaktionov, KV and Blasco-Costa, I (2018) Microphallus ochotensis sp nov (Digenea, Microphallidae) and relative merits of two-host microphallid life cycles. Parasitology Research 117, 10511068.CrossRefGoogle ScholarPubMed
Galaktionov, KV and Dobrovolskij, AA (2003) The biology and evolution of trematodes. p. 592. Dordrecht, The Netherlands, Kluwer Academic Publishers.CrossRefGoogle Scholar
Galaktionov, KV, Blasco-Costa, I and Olson, PD (2012) Life cycles, molecular phylogeny and historical biogeography of the ‘pygmaeus’ microphallids (Digenea: Microphallidae): widespread parasites of marine and coastal birds in the Holarctic. Parasitology 139, 13461360.CrossRefGoogle ScholarPubMed
Gibson, AK, Jokela, J and Lively, CM (2016a) Fine-scale spatial covariation between infection prevalence and susceptibility in a natural population. The American Naturalist 188, 114.CrossRefGoogle Scholar
Gibson, AK, Xu, JY and Lively, CM (2016b) Within-population covariation between sexual reproduction and susceptibility to local parasites. Evolution 70, 20492060.CrossRefGoogle Scholar
Gibson, AK, Delph, LF, Vergara, D and Lively, CM (2018) Periodic, parasite-mediated selection for and against sex. American Naturalist 192, 537551.CrossRefGoogle ScholarPubMed
Gilardoni, C, Etchegoin, J, Diaz, JI, Ituarte, C and Cremonte, F (2011) A survey of larval digeneans in the commonest intertidal snails from Northern Patagonian coast, Argentina. Acta Parasitologica 56, 163.CrossRefGoogle Scholar
Hamilton, WD (1980) Sex versus non-sex versus parasite. Oikos 35, 282290.CrossRefGoogle Scholar
Hechinger, RF (2012) Faunal survey and identification key for the trematodes (Platyhelminthes: Digenea) infecting Potamopyrgus antipodarum (Gastropoda: Hydrobiidae) as first intermediate host. Zootaxa 3418, 127.CrossRefGoogle Scholar
Hernandez-Orts, JS, Pinacho-Pinacho, CD, Garcia-Varela, M and Kostadinova, A (2016) Maritrema corai n. sp. (Digenea: Microphallidae) from the white ibis Eudocimus albus (Linnaeus) (Aves: Threskiornithidae) in Mexico. Parasitology Research 115, 547559.CrossRefGoogle Scholar
Hofmann, H, Blasco-Costa, I, Knudsen, R, Matthaei, CD, Valois, A and Lange, K (2016) Parasite prevalence in an intermediate snail host is subject to multiple anthropogenic stressors in a New Zealand river system. Ecological Indicators 60, 845852.CrossRefGoogle Scholar
Huelsenbeck, JP, Ronquist, F, Nielsen, R and Bollback, JP (2001) Bayesian inference of phylogeny and its impact on evolutionary biology. Science 294, 23102314.CrossRefGoogle ScholarPubMed
Jaenike, J (1978) A hypothesis to account for the maintenance of sex in populations. Evolutionary Theory 3, 191194.Google Scholar
Jokela, J and Lively, CM (1995) Spatial variation in infection by digenetic trematodes in a population of freshwater snails (Potamopyrgus antipodarum). Oecologia 103, 509517.CrossRefGoogle Scholar
Jokela, J, Lively, CM, Dybdahl, MF and Fox, JA (2003) Genetic variation in sexual and clonal lineages of a freshwater snail. Biological Journal of the Linnean Society 79, 165181.CrossRefGoogle Scholar
Jokela, J, Dybdahl, MF and Lively, CM (2009) The maintenance of sex, clonal dynamics, and host-parasite coevolution in a mixed population of sexual and asexual snails. The American Naturalist 174, S43S53.CrossRefGoogle Scholar
Kakui, K (2014) A novel transmission pathway: first report of a larval trematode in a tanaidacean crustacean. Fauna Ryukyuana 17, 1322.Google Scholar
King, KC, Delph, LF, Jokela, J and Lively, CM (2009) The geographic mosaic of sex and the red queen. Current Biology 19, 14381441.CrossRefGoogle ScholarPubMed
King, KC, Delph, LF, Jokela, J and Lively, CM (2011a) Coevolutionary hotspots and coldspots for host sex and parasite local adaptation in a snail-trematode interaction. Oikos 120, 13351340.CrossRefGoogle Scholar
King, KC, Jokela, J and Lively, CM (2011b) Parasites, sex, and clonal diversity in natural snail populations. Evolution 65, 14741481.CrossRefGoogle Scholar
Koskella, B and Lively, CM (2007) Advice of the rose: experimental coevolution of a trematode parasite and its snail host. Evolution 61, 152159.CrossRefGoogle ScholarPubMed
Koskella, B and Lively, CM (2009) Evidence for negative frequency-dependent selection during experimental coevolution of a freshwater snail and a sterilizing trematode. Evolution 63, 22132221.CrossRefGoogle Scholar
Koskella, B, Vergara, D and Lively, CM (2011) Experimental evolution of sexual host populations in response to sterilizing parasites. Evolutionary Ecology Research 13, 315322.Google Scholar
Krist, AC and Lively, CM (1998) Experimental exposure of juvenile snails (Potamopyrgus antipodarum) to infection by trematode larvae (Microphallus sp.): infectivity, fecundity compensation and growth. Oecologia 116, 575582.CrossRefGoogle Scholar
Krist, AC, Lively, CM, Levri, EP and Jokela, J (2000) Spatial variation in susceptibility to infection in a snail-trematode interaction. Parasitology 121, 395401.CrossRefGoogle Scholar
Krist, AC, Jokela, J, Wiehn, J and Lively, CM (2004) Effects of host condition on susceptibility to infection, parasite developmental rate, and parasite transmission in a snail-trematode interaction. Journal of Evolutionary Biology 17, 3340.CrossRefGoogle Scholar
Kudlai, O, Cutmore, SC and Cribb, TH (2015) Morphological and molecular data for three species of the Microphallidae (Trematoda: Digenea) in Australia, including the first descriptions of the cercariae of Maritrema brevisacciferum Shimazu et Pearson, 1991 and Microphallus minutus Johnston, 1948. Folia Parasitol (Praha) 62, 053.CrossRefGoogle ScholarPubMed
Kudlai, O, Cribb, TH and Cutmore, SC (2016) A new species of microphallid (Trematoda: Digenea) infecting a novel host family, the Muraenidae, on the northern Great Barrier Reef, Australia. Systematic Parasitology 93, 863876.CrossRefGoogle ScholarPubMed
Kumar, S, Stecher, G, Li, M, Knyaz, C and Tamura, K (2018) MEGA X: molecular evolutionary genetics analysis across computing platforms. Molecular Biology and Evolution 35, 15471549.CrossRefGoogle ScholarPubMed
Lagrue, C and Poulin, R (2008) Lack of seasonal variation in the life-history strategies of the trematode Coitocaecum parvum: no apparent environmental effect. Parasitology 135, 12431251.CrossRefGoogle ScholarPubMed
Lagrue, C, McEwan, J, Poulin, R and Keeney, DB (2007) Co-occurrences of parasite clones and altered host phenotype in a snail-trematode system. International Journal for Parasitology 37, 14591467.CrossRefGoogle Scholar
Levri, EP (1999) Parasite-induced change in host behavior of a freshwater snail: parasitic manipulation or byproduct of infection? Behavioral Ecology 10, 234241.CrossRefGoogle Scholar
Levri, EP and Fisher, LM (2000) The effect of a trematode parasite (Microphallus sp.) on the response of the freshwater snail Potamopyrgus antipodarum to light and gravity. Behaviour 137, 11411151.CrossRefGoogle Scholar
Levri, EP and Lively, CM (1996) The effects of size, reproductive condition, and parasitism on foraging behaviour in a freshwater snail, Potamopyrgus antipodarum. Animal Behaviour 51, 891901.CrossRefGoogle Scholar
Levri, EP, Dillard, J and Martin, T (2005) Trematode infection correlates with shell shape and defence morphology in a freshwater snail. Parasitology 130, 699708.CrossRefGoogle Scholar
Littlewood, DTJ, Curini-Galletti, M and Herniou, EA (2000) The interrelationships of Proseriata (Platyhelminthes: Seriata) tested with molecules and morphology. Molecular Phylogenetics and Evolution 16, 449466.CrossRefGoogle ScholarPubMed
Lively, CM (1987) Evidence from a New Zealand snail for the maintenance of sex by parasitism. Nature 328, 519.CrossRefGoogle Scholar
Lively, CM (1989) Adaptation by a parasitic trematode to local populations of its snail host. Evolution 43, 16631671.CrossRefGoogle ScholarPubMed
Lively, CM (2016) Coevolutionary epidemiology: disease spread, local adaptation, and sex. The American Naturalist 187, E77E82.CrossRefGoogle ScholarPubMed
Lively, CM and Dybdahl, MF (2000) Parasite adaptation to locally common host genotypes. Nature 405, 679.CrossRefGoogle ScholarPubMed
Lively, CM and Jokela, J (1996) Clinal variation for local adaptation in a host-parasite interaction. Proceedings of the Royal Society B-Biological Sciences 263, 891897.Google Scholar
Lively, CM and McKenzie, JC (1991) Experimental infection of a freshwater snail, Potamopyrgus antipodarum,with a digenetic trematode, Microphallus sp. New Zealand Natural Sciences 18, 5962.Google Scholar
Lively, CM, Dybdahl, MF, Jokela, J, Osnas, EE and Delph, LF (2004) Host sex and local adaptation by parasites in a snail-trematode interaction. American Naturalist 164, S6S18.CrossRefGoogle Scholar
Lively, CM, Delph, LF, Dybdahl, MF and Jokela, J (2008) Experimental test for a co-evolutionary hotspot in a host-parasite interaction. Evolutionary Ecology Research 10, 95103.Google Scholar
Lockyer, AE, Olson, PD and Littlewood, DTJ (2003) Utility of complete large and small subunit rRNA genes in resolving the phylogeny of the Neodermata (Platyhelminthes): implications and a review of the Cercomer theory. Biological Journal of the Linnean Society 78, 155171.CrossRefGoogle Scholar
McKone, MJ, Gibson, AK, Cook, D, Freymiller, LA, Mishkind, D, Quinlan, A, York, JM, Lively, CM and Neiman, M (2016) Fine-scale association between parasites and sex in Potamopyrgus antipodarum within a New Zealand lake. New Zealand Journal of Ecology 40, 330333.CrossRefGoogle Scholar
Miller, MA, Pfeiffer, W and Schwartz, T (2010), Creating the CIPRES Science Gateway for inference of large phylogenetic trees. In Proceedings of the Gateway Computing Environments Workshop (GCE), New Orleans, LA18.Google Scholar
Morgan, JAT and Blair, D (1995) Nuclear rDNA ITS sequence variation in the trematode genus Echinostoma: an aid to establishing relationships within the 37-collar-spine group. Parasitology 111, 609615.CrossRefGoogle ScholarPubMed
O'Dwyer, K, Blasco-Costa, I, Poulin, R and Faltynkova, A (2014) Four marine digenean parasites of Austrolittorina spp. (Gastropoda: Littorinidae) in New Zealand: morphological and molecular data. Systematic Parasitology 89, 133152.CrossRefGoogle ScholarPubMed
Osnas, EE and Lively, CM (2004) Parasite dose, prevalence of infection and local adaptation in a host-parasite system. Parasitology 128, 223228.CrossRefGoogle Scholar
Osnas, EE and Lively, CM (2006) Host ploidy, parasitism and immune defence in a coevolutionary snail-trematode system. Journal of Evolutionary Biology 19, 4248.CrossRefGoogle Scholar
Osnas, EE and Lively, CM (2011) Using definitive host faeces to infect experimental intermediate host populations: waterfowl hosts for New Zealand trematodes. New Zealand Journal of Zoology 38, 8390.CrossRefGoogle Scholar
Paczesniak, D, Adolfsson, S, Liljeroos, K, Klappert, K, Lively, CM and Jokela, J (2014) Faster clonal turnover in high-infection habitats provides evidence for parasite-mediated selection. Journal of Evolutionary Biology 27, 417428.CrossRefGoogle ScholarPubMed
Paczesniak, D, Klappert, K, Kopp, K, Neiman, M, Seppälä, K, Lively, CM and Jokela, J (2019) Parasite resistance predicts fitness better than fecundity in a natural population of the freshwater snail Potamopyrgus antipodarum. Evolution 73, 16341646.CrossRefGoogle Scholar
Presswell, B, Blasco-Costa, I and Kostadinova, A (2014) Two new species of Maritrema Nicoll, 1907 (Digenea: Microphallidae) from New Zealand: morphological and molecular characterisation. Parasitology Research 113, 16411656.CrossRefGoogle ScholarPubMed
Price, EW (1934) New trematode parasites of birds. Smithsonian Miscellaneous Collections 91, 16.Google Scholar
Ronquist, F, Teslenko, M, van der Mark, P, et al. (2012) MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic Biology 61, 539542.CrossRefGoogle ScholarPubMed
Smith, SJ (1973) Three new microphallid trematodes from Tasmanian birds. Papers and proceedings of the Royal Society of Tasmania 107, 197205.Google Scholar
Snyder, SD and Tkach, VV (2001) Phylogenetic and biogeographical relationships among some Holarctic frog lung flukes (Digenea: Haematoloechidae). Journal of Parasitology 87, 14331440.CrossRefGoogle Scholar
Soper, DM, King, KC, Vergara, D and Lively, CM (2014) Exposure to parasites increases promiscuity in a freshwater snail. Biology Letters 10, 20131091.CrossRefGoogle Scholar
Stamatakis, A (2014) RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30, 13121313.CrossRefGoogle ScholarPubMed
Stunkard, HW (1958) The morphology and life-history of Levinseniella minuta (Trematoda: Microphallidae). The Journal of Parasitology 44, 225229.CrossRefGoogle Scholar
Tkach, V, Pawlowski, J and Mariaux, J (2000) Phylogenetic analysis of the suborder Plagiorchiata (Platyhelminthes, Digenea) based on partial lsrDNA sequences. International Journal for Parasitology 30, 8393.CrossRefGoogle ScholarPubMed
Tkach, VV, Littlewood, DTJ, Olson, PD, Kinsella, JM and Swiderski, Z (2003) Molecular phylogenetic analysis of the Microphalloidea Ward, 1901 (Trematoda : Digenea). Systematic Parasitology 56, 115.CrossRefGoogle Scholar
Vergara, D, Lively, CM, King, KC and Jokela, J (2013) The geographic mosaic of sex and infection in lake populations of a New Zealand snail at multiple spatial scales. American Naturalist 182, 484493.CrossRefGoogle ScholarPubMed
Vergara, D, Jokela, J and Lively, CM (2014) Infection dynamics in coexisting sexual and asexual host populations: support for the Red Queen Hypothesis. American Naturalist 184, S22S30.CrossRefGoogle ScholarPubMed
Vergara, D, Fuentes, JA, Stoy, KS and Lively, CM (2017) Evaluating shell variation across different populations of a freshwater snail. Molluscan Research 37, 120132.CrossRefGoogle Scholar
Werle, E, Schneider, C, Renner, M, Volker, M and Fiehn, W (1994) Convenient single-step, one tube purification of PCR products for direct sequencing. Nucleic Acids Research 22, 43544355.CrossRefGoogle ScholarPubMed
Winterbourn, MJ (1974) Larval trematoda parasitising the New Zealand species of Potamopyrgus (Gastropoda : Hydrobiidae). Mauri Ora 2, 1730.Google Scholar
Figure 0

Table 1. List of taxa included in the phylogenetic analyses, GenBank accession numbers and references.

Figure 1

Fig. 1. Atriophallophorus winterbourni n. sp. (a) Illustration of the holotype, 24 h in vitro grown adult in ventral view. (b) Microphotographs of the terminal genitalia of the holotype using light microscopy. (c) Histological oblique section of a paratype at the level of the ventral sucker. (d) Microphotograph of an encysted metacercaria ex Potamopyrgus antipodarum (Gray). Abbreviations: c, caeca; ga, genital atrium; pc, prostatic chamber; ph, phallus; sv, seminal vesicle; vs, ventral sucker.

Figure 2

Fig. 2. Scanning electron micrographs of Atriophallophorus winterbourni n. sp. (a) Adult. (b) Adult with phallus protruded. (c) Palmate spines on the ventral surface of the body. (d) Detail of the oral sucker, arrows point at a gland opening and a sensory papilla surrounding the oral sucker. (e) Detail of the outer rim of the ventral sucker with spination and the parietal atrial scale at the basis of the phallus. (f) Outer rim of the ventral sucker interrupted sinistrally by the opening for the genital pore with arrows pointing at glands. (g) Tip of the phallus evaginated, which appears as a flower-like structure when invaginated. (h) Detail of the configuration of the protruded phallus and the sinistrally interrupted outer rim of the ventral sucker.

Figure 3

Table 2. Comparative metrical data for Atriophallophorus spp.

Figure 4

Fig. 3. Phylogenetic relationships for representatives of the family Microphallidae, inferred by maximum likelihood analysis of 28S rDNA sequence data. The newly generated sequences are indicated in bold. Values on the branches correspond to posterior probabilities >0.95 followed by bootstrap support >60. Values below these thresholds were not reported.

Figure 5

Fig. 4. Phylogram for representatives of the family Microphallidae, inferred by maximum likelihood analysis of sequence data for the internal transcribed spacer 2 of the rRNA genes. The newly generated sequences are indicated in bold. Values on the branches correspond to posterior probabilities >0.95 followed by bootstrap support >60. Values below these thresholds were not reported.

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A new species of Atriophallophorus Deblock & Rosé, 1964 (Trematoda: Microphallidae) described from in vitro-grown adults and metacercariae from Potamopyrgus antipodarum (Gray, 1843) (Mollusca: Tateidae)
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A new species of Atriophallophorus Deblock & Rosé, 1964 (Trematoda: Microphallidae) described from in vitro-grown adults and metacercariae from Potamopyrgus antipodarum (Gray, 1843) (Mollusca: Tateidae)
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A new species of Atriophallophorus Deblock & Rosé, 1964 (Trematoda: Microphallidae) described from in vitro-grown adults and metacercariae from Potamopyrgus antipodarum (Gray, 1843) (Mollusca: Tateidae)
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