NEW GENUS AND SPECIES OF CYCLOCOELIDAE STOSSICH, 1902 (PLATYHELMINTHES: DIGENEA) INFECTING THE NASOPHARYNGEAL CAVITY OF CANADA GOOSE, BRANTA CANADENSIS (ANSERIFORMES: ANATIDAE) FROM WESTERN ALABAMA

While surveying the parasites of birds associated with western Alabama aquaculture ponds raising channel catfish, Ictalurus punctatus Rafinesque, 1818 (Siluriformes, Ictaluridae), we discovered several large trematodes infecting the nasopharyngeal cavity of a Canada goose, Branta canadensis (Linnaeus, 1758) (Anseriformes: Anatidae). We describe these specimens as a new species of Cyclocoelidae Stossich, 1902 (Digenea) and propose a new genus for the new species. Cyclocoelidae includes >120 species that mature in the airways, air sacs, nasopharyngeal cavity, hypothalamus, liver, and body cavity of birds (Kossack, 1911; Witenberg, 1923, 1926; Bashkirova, 1950; Dubois, 1959; Kanev et al., 2002). Dronen and Blend (2015) published the most recent revision of the family and accepted 6 subfamilies and 22 genera.

An infected Canada goose was shot dead at an aquaculture farm (32°29′22.9″N, 87°36′47.0″W) in Hale County, Alabama (U.S. Department of Agriculture [USDA] Aquatic Nuisance Species Permit), transferred to personnel of the Southeastern Cooperative Fish Parasite and Disease Laboratory, transported to Auburn University in a cooler of ice, and dissected. The goose was identified as a Canada goose by having a black head with white cheeks and chinstrap, black neck, tan breast, and brown back (Dunn, 1987). A total of 4 live trematode specimens were removed from the nasopharyngeal cavity of this goose and placed in physiological saline. Specimens intended for morphology were heat killed on glass slides using a butane hand lighter under little or no coverslip pressure, fixed in 10% neutral buffered formalin, rinsed with distilled water, stained in Van Cleave’s hematoxylin with several drops of Ehrlich’s hematoxylin, dehydrated through a graded series of EtOHs, made basic at 70% EtOH with lithium carbonate and butyl-amine, dehydrated in absolute EtOH and xylene, cleared with clove oil, and permanently mounted on glass slides using Canada balsam (Dutton et al., 2019). The resulting whole mounts were examined and illustrated with the aid of an Olympus BX53 (Olympus, Tokyo, Japan) with differential interference contrast (DIC) optical components and a drawing tube, and a Ken-A-Vision X1000 Microprojector (Ken-A-Vision, Raytown, Missouri). Measurements were obtained with a calibrated ocular micrometer (as straight lines along the course of each duct) and are reported in micrometers (μm) as the range followed by the mean, +/− standard deviation, and sample size in parentheses. Types of the new species were deposited in the National Museum of Natural History’s Invertebrate Zoology Collection (Smithsonian Institution, USNM Collection Nos. 1683864–1683866). Classification and anatomical terms for cyclocoelids follow Dronen and Blend (2015).

Total genomic DNA (gDNA) was extracted from 2 cut portions of 1 EtOH-preserved specimen using DNeasy™ Blood and Tissue Kit (Qiagen, Valencia, California) as per the manufacturer’s protocol with 1 exception; the proteinase-K incubation period was extended overnight, and 100 μl of elution buffer was used to increase the final DNA concentration. The 28S, 18S, ITS2, and CO1 genes were amplified using the primer set and PCR amplifications according to Urabe et al. (2020). DNA sequencing was performed by Genewiz, Incorporated (South Plainfield, New Jersey). Sequence assembly and analysis of chromatograms were performed with Geneious version 2022.0.2 (http://www.geneious.com). All nucleotide sequence data were deposited in GenBank (OQ780427, OQ780428, OQ780430, OQ780431, OQ802836, OQ821763, OQ821977). The phylogenetic analyses included 2 identical sequences of the new species, all available 28S, 18S, and ITS2 sequences from Cyclocoelidae and Typhlocoelidae Harrah, 1922, expanding on the work of Olson et al. (2003), members from Schistosomatidae Stiles and Hassall, 1898, and a haploporid comprising the outgroups. Sequences were aligned with the multiple alignment tool using a fast Fourier transform (MAFFT) (Katoh and Standley, 2013) and trimmed to the length of the shortest sequence (1,179 [28S] base pairs). JModelTest 2 version 2.1.10 was implemented to perform statistical selection of the best-fit nucleotide substitution models based on the Bayesian information criterion (BIC) (Darriba et al., 2012). Aligned sequences were reformatted (from .fasta to .nexus) using the web application ALTER (Glez-Peña et al., 2010) to run Bayesian inference (BI). BI was performed in MrBayes version 3.2.5 (Ronquist and Huelsenbeck, 2003) using substitution model averaging (“nst-mixed”) and gamma distribution to model rate heterogeneity. Defaults were used in all other parameters. Three independent runs with 4 Metropolis-coupled chains were run for 5,000,000 generations, sampling the posterior distribution every 1,000 generations. Convergence was checked using Tracer v1.6.1 (Rambaut et al., 2014) and the “sump” command in MrBayes: All runs appeared to reach convergence after discarding the first 25% of generation as burn-in. A majority rule consensus tree of the post–burn-in posterior distribution was generated with the “sumt” command in MrBayes. The inferred phylogenetic tree was visualized using FigTree v1.4.4 (Rambaut et al., 2014) and further edited for visualization purposes with Adobe Illustrator (Adobe Systems).

Anativermis n. gen.
(Figs. 1–3)

Body large, slightly dorsoventrally flat, aspinose, broadest in the anterior body half, rounded anteriorly, more sharply tapered posteriorly. Mouth subterminal, unaccompanied by a demonstrable muscular complex (oral sucker absent). Ventral sucker absent. Pharynx well developed, immediately posterior to the mouth (hence, a so-called “prepharynx” is absent). Esophagus short, extending posteriad directly from pharynx before connecting to cecal bifurcation. Ceca elongate, sinuous, uniting in posterior body extremity to form cyclocoel. Testes spheroid (not transverse), having smooth margins (lacking lobes), in posterior quarter of body; anterior testis lateral to the midline, abutting respective cecum; posterior testis medial, abutting cyclocoel. Genital pore immediately postpharyngeal. Ovary spherical, lacking lobes, intertesticular, nearly abutting posterior testis. Vitellarium asymmetrical, dorsal to ceca, distributing from immediately posterior to cecal bifurcation posteriad to level of ovary, discontinuous posteriorly. Uterus extensively convoluted, dorsal to ceca, spanning breadth of body; uterine loops extending lateral to ceca and filling space between ceca and respective body margin for nearly entire body length, terminating at the level of posterior testis. Parasites of nasopharyngeal cavity of birds.

Body broadest in the anterior body half. Testes spheroid, lacking lobes, in posterior quarter of body; anterior testis lateral to the midline, abutting respective cecum; posterior testis medial, abutting cyclocoel. Genital pore immediately postpharyngeal. Ovary nearly abutting posterior testis. Vitellarium asymmetrical, distributing from immediately posterior to cecal bifurcation posteriad to level of ovary, discontinuous posteriorly. Uterine loops extend lateral to ceca and fill space between ceca and respective body margin for nearly the entire body length.

Anativermis normdroneni n. sp.

urn:lsid:zoobank.org:act:D2FD2C55-0EB5-4855-969D-DD88B75B9ACD.

Anativermis; “Anati” for the host family Anatidae and “vermis” for worm.

Anativermis normdroneni n. sp.
(Figs. 1–6)

Body large, 18,500–23,800 (21,125 ± 2,650; 3) long, 3,900–5,100 (4,667 ± 666; 3) in maximum width, broadest in anterior half of the body, 4–6× (5 ± 1; 3) longer than wide, posterior end tapering. Mouth 200–250 (227 ± 25; 3) long, 370–420 (388 ± 28; 3) wide. Pharynx strongly muscular, 500–575 (535 ± 38; 3) long or 58–59% (58% ± 1%; 2) of esophagus length, 550–625 (592 ± 38; 3) wide or 6× wider than maximum esophagus width. Esophagus short, 900–1,000 (950 ± 71; 2) long, 100 (100 ± 0; 2) wide (Fig. 2). Intestinal bifurcation 820–1,370 (1,147 ± 289; 3) or 3–7% (6% ± 2%; 3) of body length from anterior body end. Cecal bifurcation to cyclocoel 16,525–22,625 (19,600 ± 3,050; 3) long or 89–95% (93% ± 3%; 3) of body length.

Testes nearly in line, not forming a strongly triangular pattern with ovary; anterior testis 1,250–1,350 (1,283 ± 58; 3) long or 5–7% (6% ± 1%; 3) of body length, 950–1,250 (1,083 ± 153; 3) wide or 21–25% (23% ± 2%; 3) of body width at level of ovary, 74–95% (82% ± 11%; 3) of posterior testis width; intertesticular space 2,075–2,375 (2,233 ± 151; 3) long or 10–12% (11% ± 1%; 3) of body length; posterior testis 1,350–1,625 (1,450 ± 152; 3) long or 6–8% (7% ± 1%; 3) of body length, 1,000–1,700 (1,350 ± 350; 3) wide or 26–33% (29% ± 4%; 3) of body width at level of ovary, 310–625 (462 ± 158; 3) or 1–3% (2% ± 1%; 3) length from end of posterior testis to posterior extremity (Figs. 1, 2). Anterior vas efferens emanating from ventral surface of anterior testis, extending anteriad 13,825–16,200 (14,925 ± 1,197; 3) or 66–80% (71% ± 8%; 3) of body length, 30–50 (40 ± 10; 3) wide; posterior vas efferens emanating from ventral surface of posterior testis, extending anteriad 17,325–19,700 (18,342 ± 1,224; 3) or 82–97% (87% ± 9%; 3) of body length, 30–50 (40 ± 10; 3) wide, converging with anterior trunk of vas efferens in anterior 1/5 of body; vas deferens (visible in paratype USNM 1683865) 1,640 long, 50 wide. Cirrus sac oblong, 1,150–1,300 (1,200 ± 87; 3) long or 5–6% (6% ± 0%; 3) of body length, 350–550 (440 ± 101; 3) wide or 28–49% (38% ± 10%; 3) of body width at level of genital pore; cirrus aspinose, 550–680 (623 ± 57; 3) long or 48–56% (52% ± 4%; 3) of cirrus sac length, 50–100 (67 ± 29; 3) wide or 4–9% (6% ± 3%; 3) of cirrus sac width (Figs. 1, 2); internal seminal vesicle coiled, 680–800 (727 ± 64; 3) long or 59–62% (61% ± 1%; 3) of cirrus sac length, 350–500 (410 ± 79; 3) wide or 90–100% (94% ± 5%; 3) of cirrus sac width.

Ovary slightly longer than wide (not transverse), 700–800 (740 ± 53; 3) long or 3–4% (4% ± 1%; 3) of body length, 640–700 (680 ± 35; 3) wide or 14–16% (15% ± 1%; 3) of body width; postovarian space, 2,050–2,500 (2,342 ± 253; 3) or 11–12% (11% ± 1%; 3) of body length. Oviduct diminutive, 100–150 (117 ± 29; 3) long, 15–20 (17 ± 3; 3) wide, anterior to transverse vitelline duct (Fig. 3). Oötype indiscernible in stained, whole-mounted specimens. Mehlis’s gland present around oviduct. Laurer’s canal not observed. Vitellarium asymmetrical, comprising a series of interconnected, continuous, small, irregularly shaped masses of follicles, distributed along ceca, follicles 100–120 (110 ± 10; 3) long, 50–60 (53 ± 6; 3) wide; dextral vitellarium begins posterior to cecal bifurcation, 4,625–5,375 (4,958 ± 382; 3) or 23–25% (24% ± 1%; 3) from anterior body end, terminating 2,375–2,775 (2,583 ± 201; 3) or 11–14% (12% ± 2%; 3) from posterior body end; sinistral vitellarium begins posterior to cecal bifurcation, 2,250–3,000 (2,683 ± 388; 3) or 11–15% (13% ± 2%; 3) from anterior body end, terminating 1,875–2,500 (2,183 ± 313; 3) or 8–12% (10% ± 2%; 3) from posterior body end; transverse vitelline duct 1,400–2,250 (1,933 ± 465; 3) in breadth, 55–100 (75 ± 23; 3) wide; primary vitelline collecting duct 150–200 (175 ± 25; 3) long, 60–70 (65 ± 5; 3) wide, inserting into oviduct ventrally. Uterus 225,500–297,500 (257,833 ± 36,566; 3) long, individual coils 200–375 (292 ± 88; 3) wide; eggs filling lumen of uterus, oblong, 120–140 (127 ± 12; 3) long, 50–60 (53 ± 6; 3) wide; empty/hatched miracidia with eyespots in distal portion of uterus; metraterm absent. Common genital pore 750–1,100 (908 ± 177; 2) or 4–6% (4% ± 1%; 2) of body length from anterior end, 100–140 (120 ± 28; 2) in diameter (Fig. 1).

Excretory vesicle a vacuous chamber that can either expand (seemingly 1 large lobe) or contract (appearing multilobed) depending on condition of specimen; excretory pore, terminal 250–550 (423 ± 155; 3) wide (Fig. 3).

Branta canadensis (Linnaeus, 1758) (Anseriformes: Anatidae), Canada goose.

An aquaculture pond (32°29′22.9″N, 87°36′47.0″W) in Hale County, Alabama.

Holotype (USNM 1683864); paratypes (USNM 1683865–1683866); 28S, 18S, ITS2, and COI sequences (GenBank nos. OQ780427, OQ780428, OQ780430, OQ780431, OQ802836, OQ821763, OQ821977).

Nasopharyngeal cavity.

The single Canada goose examined was infected with 4 specimens of A. normdroneni.

urn:lsid:zoobank.org:act:B56CA8DA-527D-48CD-A4CF-908E21F1D5E5.

The specific epithet normdroneni honors the late Professor Norman Obert Dronen (Texas A&M University, College Station, Texas) for his extensive contributions to the taxonomy of cyclocoelids.

We refrain from assigning our new species to a cyclocoelid subfamily because at present the most morphologically appropriate subfamily (Hyptiasminae Dollfus, 1948) is paraphyletic and no nucleotide sequence exists for the type species of the type genus of Hyptiasminae (Hyptiasmus arcuatus [Brandes, 1892] Kossack, 1911). Using the subfamily key in Dronen and Blend (2015), the new species was assigned to Hyptiasminae Dollfus, 1948 by having an intertesticular ovary that is nearly in a straight line with the testes, which can be tandem to nearly tandem. Using the generic key in Dronen and Blend (2015), the new species was assigned to Morishitium by having a postpharyngeal genital pore and a vitellarium that is not confluent posteriorly. Anativermis resembles Morishitium by having testes that are spheroid with smooth margins and located in the posterior quarter of the body, an anterior testis that is lateral to the midline and abuts the respective cecum, a posterior testis that is medial (testes diagonal) and abuts the cyclocoel, a genital pore that is immediately postpharyngeal, and a vitellarium that is discontinuous posteriorly. However, the new genus differs from Morishitium and is unique among all other cyclocoelid genera by having the combination of a body that is broadest in the anterior body half, a posterior body end that is more sharply tapered than the anterior body end, an ovary that nearly abuts the posterior testis, a vitellarium that is asymmetrical and distributes from the area immediately posterior to the cecal bifurcation posteriad to approximately the level of the ovary, and uterine loops extending dorsolateral to the ceca and filling the space between the ceca and the respective body margin for nearly the entire body length.

The 28S sequences of the new species were identical and comprised 1,249 and 1,236 nucleotides. The 28S phylogenetic analysis contained 1,179 aligned nucleotides and recovered the new species sister to the clade comprising Neohaematotrephus arayae Zamparo, Brooks, Causey and Rodriguez, 2003 (86 nucleotide differences with A. normdroneni) and Cyclocoelum mutabile (Zeder, 1800) (95 nucleotide differences with A. normdroneni). Our results agree with Tkach et al. (2016), who recovered the same topology and suggested that Typhlocoelidae could be a junior subjective synonym of Cyclocoelidae. The 18S sequence of the new species was 1,770 nucleotides, and the phylogenetic analysis comprised 471 aligned nucleotides. The new species was recovered in the 18S tree as a sister to Morishitium grusi (Kocan, Waldrup, Ramakka, and Iverson, 1982) Dronen and Blend, 2015 (4 nucleotide differences with A. normdroneni) and within a polytomy. Based on nucleotide evidence, Sitko et al. (2016) reassigned Cyclocoelum obscurum Leidy, 1887 to Harrahium Witenberg, 1926, as Harrahium obscurum (Leidy, 1887) Sitko and Heneberg, 2016. They also asserted that the sequence ascribed to C. mutabile (AJ287494) by Cribb et al. (2001) was misidentified. They identified that sequence as belonging to Harrahium sp., because that sequence was distinct and unrelated to their sequence of C. mutabile (KU877900). There is reportedly no voucher specimen for the sequence of “C. obscurum” by Cribb et al. (2001), making it a nonugen as per Roberts et al. (2018), that is, a GenBank sequence that is unaccompanied by robust morphological evidence or a museum-curated voucher specimen that underpins its taxonomic identity. The ITS2 sequences of A. normdroneni were identical and comprised 1,030 and 1,036 nucleotides, and phylogenetic analysis contained 492 aligned nucleotides. The new species was recovered sister to the clade comprising Harrahium tringae (Brandes, 1892), Dronen and Blend, 2015 (102 nucleotide differences), and H. obscurum (97 nucleotide differences) and within a polytomy. COI sequences of the new species comprised 631 nucleotides. The new species was 88% similar to C. mutabile, but a phylogenetic analysis using that marker was moot, because too few cyclocoelid CO1 sequences exist to conduct a meaningful analysis. All phylogenetic analyses recovered the new species as a distinct lineage that shares a common ancestor with at least Morishitium and Cyclocoelum, the only genera of cyclocoelids represented by 18S, 28S, and ITS2 sequences. Noteworthy is that Morishitium is paraphyletic in the 18S tree, with congeners M. grusi and M. polonicum in different clades (not sharing a recent common ancestor), suggesting further that paraphyletic Hyptiasminae needs revision.

Numerous authors have called attention to the fact that Cyclocoelidae needs revision (Sitko et al., 2016; Tkach et al., 2016; López-Jiménez et al., 2018; Khan et al., 2019; Urabe et al., 2020; Assis et al., 2021; Díaz et al., 2022). Using morphological comparisons across genera, Dronen and Blend (2015) provided a robust and critical review of the family, treating several subfamilies that had been recovered as paraphyletic in previous phylogenetic analyses. Dronen et al. (2017) stated that Cyclocoelidae needs additional nucleotide information more than any other trematode family. Tkach et al. (2016) and Assis et al. (2021) synonymized Typhlocoelidae with Cyclocoelidae but did not reconcile the monophyly or taxonomic status of the subfamilies.

Additional nucleotide, phylogenetic, and life cycle information (especially including robust taxonomic identifications of the snail hosts) is needed in this group to test the monophyly of the cyclocoelid subfamilies further and to revise the family. These features are likely informative regarding the natural history of the group. For example, Cyclocoelum, Harrahium, and Hyptiasmus Kossack, 1911 are phylogenetically related and have similar life cycles (Sitko et al., 2006,; 2016; Dronen and Blend, 2015). However, obtaining the life cycle for additional species is complicated because these trematodes exploit a broad phylogenetic spectrum of molluscan hosts, that is, species of Lymnaeidae Rafinesque, 1815; Planorbidae, Rafinesque, 1815; Physidae, Fitzinger, 1833. Cyclocoelids undergo asexual reproduction in freshwater pulmonate snails or in xerothermic snails that serve as both first and second intermediate hosts. Miracidia, each containing a redia, hatch, in utero or water, and are either attached to or ingested by susceptible snail hosts. However, if not ingested, the miracidium attaches to the epithelium of the snail before injecting a single redia, which produces additional rediae and cercariae within primarily the connective tissue of the snail digestive gland but also in the head–foot and albumin gland (Taft, 1975; McKindsey and McLaughlin, 1995). The tailless cercaria encysts in the same snail individual before it develops into a metacercaria that is then ingested by a bird (Taft, 1975; Dronen and Blend, 2015). A study documenting the life cycle of the new species is underway.

We thank Tommy Graeter (USDA APHIS Wildlife Services, Greensboro, Alabama) for helping collect the goose; Steve Curran (Southeastern Cooperative Fish Parasite and Disease Laboratory, Auburn, Alabama) for mounting the worms; Anna Phillips, Chad Walter, Kathryn Ahlfeld, and William Moser (Department of Invertebrate Zoology, USNM, Smithsonian Institution, Washington, D.C.) for accepting our museum specimens. This study was supported by the Southeastern Cooperative Fish Parasite and Disease Project, the U.S. Fish and Wildlife Service (Department of Interior), National Sea Grant (National Oceanic and Atmospheric Administration), USDA (National Institute of Food and Agriculture), Federal Aid in Sport Fish Restoration (Alabama Department of Conservation and Natural Resources, Inland and Marine Resources Divisions), and the Alabama Agricultural Experiment Station (Auburn University, College of Agriculture).