ABSTRACT
Morphological characteristics and DNA sequencing were used to identify plerocercoids of a Schistocephalus sp. infecting slimy sculpin (Cottus cognatus) from northern New Brunswick and plerocercoids of Ligula intestinalis infecting blacknose dace (Rhinichthys atratulus) in Fundy National Park (FNP, New Brunswick). To our knowledge, no previous publications documented either cestode from New Brunswick, Canada. Blacknose dace represent a new host record for L. intestinalis. Identifications were made based on the presence or absence of segmentation and sequencing partial nicotinamide adenine dinucleotide dehydrogenase subunit 1 (ND1; mitochondrial DNA) and/or partial cytochrome c oxidase subunit 1 (COI; mitochondrial DNA). Plerocercoids from blacknose dace in FNP were identified as Ligula intestinalis based on >99% nucleotide identity with COI for this species in the NCBI GenBank database. Plerocercoids in slimy sculpin from northern New Brunswick were identified as a Schistocephalus sp. based on high nucleotide identity with congenerics in the NCBI GenBank database. The absence of GenBank entries with sufficient high percent identity to our specimens, and potential species hybrids in this genus, prevents species-level identification of Schistocephalus sp. plerocercoids currently. The absence of previous documentation of these cestodes might reflect recent environmental change promoting the transmission of these parasites that can modulate host fish behavior, induce sterility of host fishes, and contribute to epizootics.
Cestodes are parasitic flatworms that infect a broad diversity of hosts, often requiring multiple obligate hosts during their lifecycle. Ligula and Schistocephalus are closely related genera of cestodes in the family Diphyllobothridae. Reproduction by adult Ligula and Schistocephalus occurs in the intestine of a piscivorous bird, and parasite eggs pass to the environment with feces from the bird definitive host (Hopkins and Smyth, 1951; Loot et al., 2001; Piasecki et al., 2004). Eggs release coracidium larvae in freshwater aquatic environments and develop into procercoid larvae within the hemocoel of freshwater copepod intermediate hosts following ingestion (Hopkins and Smyth, 1951; Loot et al., 2001). Infected copepods are consumed by planktivorous fish hosts in which procercoid larvae develop into precocious plerocercoids (Hopkins and Smyth, 1951; Loot et al., 2001; Piasecki et al., 2004). Infected fishes are then consumed by a piscivorous bird definitive host, including mergansers, kingfishers, gulls, ducks, herons, grebes, and terns (Hopkins and Smyth, 1951; Weekes and Penlington, 1986; Szalai et al., 1989; Loot et al., 2001). Plerocercoids of Ligula and Schistocephalus can alter the behavior of their intermediate host fish to increase the odds of fish predation by their bird definitive host (Loot et al., 2001; Barber et al., 2004), a phenomenon known as Parasite Increased Trophic Transmission (PITT; Lafferty, 1999). Plerocercoids of both species can also induce sterility of host fishes (Heins and Baker, 2010; Heins, 2017; Biswas and Ash, 2021), and epizootics caused by both parasites are reported (Kennedy et al., 2001; Heins and Ecke, 2012).
Ligula and Schistocephalus show broad geographic distribution across North America and infect a variety of fish. Ligula show low specificity for their intermediate fish host and infect a wide range of cyprinids, catastomids, and occasionally salmonids or percids (Orr, 1967; Weekes and Penlington, 1986; Szalai et al., 1989; Marcogliese and Cone, 1991). Schistocephalus was previously considered a specialist parasite of sticklebacks, but recent observations support additional Schistocephalus sp. infecting coastrange sculpin, slimy sculpin, and bullhead (Cottus spp.) as their second intermediate host (Chubb et al., 2006; French and Muzzall, 2008; Harmon et al., 2015). Species-level identification of cestode plerocercoids is difficult due to their lack of distinctive morphological features, making molecular analyses necessary (Van Steenkiste et al., 2015). Our research aimed to identify plerocercoids infecting small-bodied fish in New Brunswick using morphological and/or molecular techniques due to the absence of previous documentation of these important parasites in this province.
MATERIALS AND METHODS
Field collections and parasitological observations
Two cestode plerocercoids lacking segmentation were liberated from captured brook charr (Salvelinus fontinalis) or blacknose dace (Rhinichthys atratulus) during a fish survey in Wolfe Lake, Fundy National Park (FNP; New Brunswick, Canada) in November 2021. Plerocercoids were preserved in ethanol in advance of DNA extraction, PCR, and sequencing. One additional blacknose dace was subsequently observed with a plerocercoid actively emerging from the body cavity at this same sampling location later in 2021; this plerocercoid was not retained for specific identification. Targeted collections of blacknose dace from Wolfe Lake (45°39′39.276″N, 65°8′23.784″W) in November 2022 assessed prevalence of individuals with grossly distended abdomens, suggestive of plerocercoid infection. Fish lengths and weights were determined as were lengths and weights of plerocercoids retrieved from blacknose dace. Five representative plerocercoids from Wolfe Lake were subjected to DNA extraction for PCR and sequencing.
Slimy sculpin (n = 201) were collected from the Charlo and Quisibis River watersheds (New Brunswick, Canada), respectively, as part of research on fish health and contaminants in aquatic systems influenced by varying forest management activities in 2017 (White, 2020; Negrazis et al., 2022). Charlo River sampling sites included NBR2 (47°51′36.756″N, 66°34′18.984″W), NBR4 (47°1′49.932″N, 66°32′31.992″W), and NBR5 (47°51′14.616″N, 66°33′29.988″W), and the Quisibis River sampling site was NBE2 (47°24′41.724″N, 68°4′32.016″W). Fish were examined for plerocercoids, plerocercoid segment number was counted for fish with single plerocercoid infections, and plerocercoids were preserved in ethanol in advance of DNA extraction for PCR and sequencing.
Molecular identification
A small piece (∼5 mm) of each plerocercoid was cut from the posterior end and left to desiccate until the ethanol evaporated completely. DNA was extracted using a DNeasy Blood and Tissue DNA extraction kit following the manufacturer’s instructions (Qiagen, Toronto, Ontario, Canada). In instances where a fish host harbored multiple plerocercoids, only 1 plerocercoid was subjected to DNA extraction. Mitochondrial DNA from partial cytochrome c oxidase subunit 1 (COI) of each suspect Ligula was PCR amplified and sequenced using COI forward and COI reverse primers (Table I). Unlike Ligula, limited COI records are available in the GenBank database for Schistocephalus spp., prompting PCR amplification and sequencing of mitochondrial DNA for both partial Nicotinamide Adenine Dinucleotide dehydrogenase subunit 1 (ND1) gene and partial COI gene for suspect Schistocephalus specimens (Table I). Cestode DNA was amplified in 50 μl reaction volumes containing 0.25 μl of Taq DNA polymerase (OneTaq, New England Biolabs, Whitby, Ontario, Canada), 10 μl 5× PCR buffer, 2.5 μl dNTP mix (New England Biolabs), 1.25 μl of respective forward and reverse primers (10 μm; Integrated DNA Technologies, Coralville, Iowa; see Table I), 1 μl template DNA extracted from each plerocercoid, and nuclease-free water to a final volume of 50 μl. Negative control reactions contained an equivalent volume of nuclease-free H2O in place of template DNA. Samples were denatured at 94 C for 5 min and with 35 cycles of denaturation at 94 C for 30 sec, primer annealing at 48–52 C for 30 sec (see Table I), and extension by DNA polymerase at 68 C for 1 min, followed by a final extension of amplicons at 68 C for 5 min.
Agarose gel electrophoresis was used to assess PCR success (2% agarose dissolved in TAE containing SYBR Safe; Thermo Fisher Scientific, Burlington, Ontario, Canada). Twelve microliters of each PCR product were mixed with loading dye, and a 100-bp DNA ladder (New England Biolabs) was included in representative lanes for determination of PCR amplicon sizes. Gels were run at 125V until the dye front was ∼75% through the gel. Gels were visualized using SYBR Safe settings in a Gel Doc XR+ System (Bio-Rad; Mississauga, Ontario, Canada). Expected amplicon sizes for each primer set are included in Table I. DNA from successful PCR reactions was purified using a QIAquick PCR Purification Kit (Qiagen) and submitted for Sanger sequencing (Robarts Research Institute, London, Ontario, Canada) using respective forward and reverse primers (Table I). Consensus nucleotide sequences for each gene or specimen were created by merging the respective forward and reverse sequencing results. DNA consensus sequences were subjected to nucleotide BLAST at the National Center for Biotechnology Information (NCBI) to assess species with high nucleotide identity from previously deposited sequence data (Altschul et al., 1990). Nucleotide sequences of top BLAST species matches were retrieved for preparation of a phylogenetic tree to infer cestode relatedness. The respective COI and ND1 nucleotide sequences from plerocercoids were also entered into an open reading frame finder to acquire translated amino acid sequences (Stothard, 2000) to assess the influence of nucleotide polymorphisms among individual parasites from slimy sculpin and blacknose dace.
A maximum likelihood (ML) phylogenetic tree was estimated via the IQ-TREE server (Trifinopoulos et al., 2016) from a multiple alignment (Corpet, 1988) of partial COI (304 bp) mitochondrial DNA consensus nucleotide sequences for our specimens and paralogues from other cestode taxa retrieved from GenBank BLAST searches. The COI sequences of Dibothriocephalus dendriticus (GenBank AB623150) and D. ursi (GenBank AB605763) were used in our phylogenetic estimation because they belong to a distinct lineage within the same cestode family (Diphyllobothriidae) that was used as an outgroup in previous assessments of phylogenetic relationships between Ligula and Schistocephalus (Nazarizadeh et al., 2022). Our phylogenetic estimations considered the best-fitting nucleotide substitution model (HKY + F + I) identified by the ModelFinder tool available in IQ-TREE server options (Minh et al., 2020). Node support was evaluated with both 1,000 ultrafast bootstrap replicates and the Approximate Bayes test. Phylogenetic results were visualized using FigTree (http://tree.bio.ed.ac.uk/software/Figtree/).
RESULTS
Plerocercoids from blacknose dace in Wolfe Lake, Fundy National Park, New Brunswick, Canada
Both plerocercoids collected from Wolfe Lake (FNP) in 2021 were ∼26 cm in length before ethanol preservation (Fig. 1A) and lacked segmentation. Here 4.7% (6/127) of blacknose dace caught in Wolfe Lake (FNP) in 2022 presented with grossly distended abdomens and were consistent with the observation of infection in blacknose dace in 2021 (Fig. 1B). Dissection revealed 1–3 plerocercoids in each infected fish. Plerocercoids collected in 2022 were 12.1–25.5 cm before preservation, lacked external segmentation, and were transversely rugose in appearance. Cestodes were suspected as Ligula sp. based on the absence of segmentation (Fig. 1C). Blacknose dace harvested from Wolfe Lake (FNP) in 2022 ranged from 6.2 to 8.1 cm in length (average 6.7 SD ± 0.6) and 2.07–4.69 g (average 3.0, SD ± 0.8). Plerocercoid weights ranged from 0.51 to 1.89 g (average 1.02, SD ± 0.47) and represented 24.8 to 57.6% of total live fish weight (average 34.4%, SD ± 14.2%).
COI nucleotide sequences for all 5 specimens had >99.3% identity to each other, 99.3–100% nucleotide identity with Ligula intestinalis (GenBank KY552875), 90.6–91.2% identity with L. pavlovskii (GenBank KY552876), and 89.0–89.7% identity with L. alternans (GenBank KY552873). COI nucleotide sequences for plerocercoids from blacknose dace were deposited to GenBank under accession numbers OP913237–OP913241 (Table II). Plerocercoid specimens were deposited in the New Brunswick Museum, Saint John, New Brunswick, Canada (New Brunswick Museum catalog numbers NBM-GI-011728 to NBM-GI-011732; Table II).
Plerocercoids from slimy sculpin in northern New Brunswick, Canada
We observed 4% (6/150) and 2% (1/51) of slimy sculpin from the Charlo and Quisibis River watersheds with segmented plerocercoids within their body cavity, respectively (Fig. 1D). Intensity of infection ranged from 1 to 10 (average 2.4, SD ± 3.4) plerocercoids per infected fish. Fresh body weight was recorded for only 4 of 7 infected slimy sculpins. Weights of infected fish ranged from 1.9 to 4.4 g (average 3.0, SD ± 1.2), and individual plerocercoids ranged from 0.31 to 0.57 g (average 0.45, SD ± 0.13). Total plerocercoid mass represented 12.6 to 18.6% of the total live fish mass (average 14.9%, SD ± 2.5%). Fresh body length was not recorded for plerocercoids from slimy sculpins, but lengths were recorded after ethanol preservation. The body length of preserved plerocercoids ranged from 48 to 72 mm (average 59.6, SD ± 9.34), but shrinkage would have occurred due to preservation. All plerocercoid specimens from the Charlo and Quisibis River watersheds had external segmentation from anterior to posterior end (Fig. 1D). Segment number ranged from 143 to 177 (average 156, SD ± 13.1) for fish containing single plerocercoid infections. Segment number of plerocercoids was compatible with ranges reported for Schistocephalus spp. (Chubb et al., 2006).
Both COI and ND1 nucleotide sequences for each of 7 slimy sculpin plerocercoids were highly similar to each other (99.2–100% and 99.6–100% respectively), indicating low-level intraspecific variation in nucleotide sequence. One hundred percent sequence identity for all 7 plerocercoids at the amino acid level further supports that they are the same species. COI nucleotide and translated amino acid sequences from slimy sculpin plerocercoids show 84.9–85.7% and 92.4–93.3% identity with Schistocephalus solidus (GenBank KP865371) and 84.6–84.9% and 92.4–93.3% identity with S. pungitii (GenBank MW602516), respectively. ND1 nucleotide and translated amino acid sequences from slimy sculpin plerocercoids show 83.6–83.8% and 84.0% identity with S. solidus (GenBank KT326872), 82.2–82.4% and 84.6% identity with S. pungitii (GenBank KT326910) and 83.4–83.8% and 89.4% identity with S. cotti (GenBank KT326911), respectively. COI nucleotide sequences and ND1 nucleotide sequences for plerocercoids from slimy sculpin were deposited to GenBank under accession numbers OP927017–OP927023 and OP912668–OP912674 (Table II). Plerocercoid specimens were deposited in the New Brunswick Museum, Saint John, New Brunswick, Canada (New Brunswick Museum catalog numbers NBM-GI-011735 to NBM-GI-011739; Table II).
Phylogenetic relatedness of cestode plerocercoids from small-bodied fish in New Brunswick, Canada
The ML phylogenetic tree estimated using partial COI gene sequences for our plerocercoid specimens and top hits from GenBank nucleotide BLAST searches were consistent with our predictions based on morphological features of larval cestodes. The COI phylogenetic tree depicts parasites from slimy sculpin with 82% bootstrap support in a clade with Schistocephalus species and parasites from blacknose dace with 87% bootstrap support for relatedness with Ligula intestinalis in a clade containing congeneric Ligula species (Fig. 2). These data indicate that slimy sculpin from northern New Brunswick are infected with a Schistocephalus species not currently represented in the GenBank database, and blacknose dace from Wolfe Lake (FNP), New Brunswick, are infected with L. intestinalis.
DISCUSSION
Our research objective was to identify plerocercoids infecting slimy sculpin from the Charlo and Quisibis River watersheds in northern New Brunswick and from blacknose dace in Wolfe Lake, Fundy National Park, New Brunswick. Morphological evaluation narrowed presumptive identifications of plerocercoids to the genera Schistocephalus and Ligula. Development of degenerate primer sets that amplify partial COI and/or ND1 for congeneric species represented in GenBank enabled PCR, sequencing, and confirmation of a Schistocephalus sp. not represented in GenBank as infecting slimy sculpin and Ligula intestinalis infecting blacknose dace in New Brunswick.
Plerocercoids from Wolfe Lake in 2021 were liberated from either blacknose dace or brook charr co-housed live following collections and therefore were of unknown host origin. Wolfe Lake contains only brook charr and blacknose dace; no other fish species are present. Infection status of endemic brook charr was not assessed locally, but the absence of L. intestinalis infection in ∼20,000 brook charr in a study from Ontario where this parasite was endemic (Black and Fraser, 1984) suggests that brook charr are not typical intermediate hosts. This is compatible with the low prevalence (1/210) observed previously in Quebec brook charr (Choquette, 1948). Furthermore, Parks Canada staff subsequently observed blacknose dace with ruptured abdomens and other live blacknose dace with swollen abdomens, suggesting that they were the source host of plerocercoids liberated during co-housing of blacknose dace and brook charr in 2021. Our targeted collections of blacknose dace from Wolfe Lake with swollen abdomens in 2022 confirmed plerocercoid infection of this fish host. Blacknose dace are in the order Cypriniformes (Tan and Armbruster, 2018) and the family Leuciscidae with 3 congeneric species of dace in North America reported as permissive hosts for L. intestinalis (Gee, 1959; Dodge, 1993; Mpoame and Rinne, 1983; Muzzall et al., 1992). These prior observations of infection in several Rhinichthys sp., and both the generalist and opportunistic nature of L. intestinalis, support this first report of blacknose dace as a suitable fish intermediate host.
Ligula intestinalis plerocercoids grow large relative to fish host size and can cause distension of the abdomen to the point of tissue rupture (Dence, 1940; Arme and Owen, 1968). Ligula intestinalis plerocercoids are known to rupture through the abdominal wall or escape through the vent of fish intermediate hosts when conditions are no longer optimal for the parasite (Dence, 1958; Urdeş and Hangan, 2013). Dence (1958) suggests that plerocercoid liberation can be triggered when fish are maintained in water depleted of oxygen or in anesthetic or killing solutions. Furthermore, Ligula plerocercoids are infrequently observed “free-living” in water following liberation from host fish (Dence, 1940). Our observation of plerocercoid specimens liberated from their fish host upon initial discovery is compatible with previous observations (Dence, 1940, 1958; Urdeş and Hangan, 2013).
A wrinkled and unsegmented exterior surface occurs in plerocercoids of L. intestinalis because proglottids are neither separated nor well defined in this life stage (Smyth, 1947; Threadgold and Hopkins, 1981; Smyth, 1990). Ligula parasites present with external segmentation only after sexual maturity in the definitive bird host (Southwell, 1922) or following maturation in vitro (Smyth, 1947). Plerocercoids from Wolfe Lake lacked external segmentation, and our presumptive morphological identification is confirmed by partial COI nucleotide sequence showing 99.3% identity and 87% bootstrap support for relatedness with Nearctic Ligula intestinalis (Nazarizadeh et al., 2022) in a clade containing congeneric Ligula species.
Plerocercoids of Schistocephalus spp. have a fully segmented exterior surface with a prominent ridged appearance (Southwell, 1922; Hopkins and Smyth, 1951; Smyth, 1990). This feature led to the presumptive identification of our specimens from northern New Brunswick slimy sculpin as Schistocephalus. Three species of Schistocephalus are reported from intermediate host fishes across North America, including S. solidus, S. pungitii, and S. thomasi (Garoian, 1960; Holloway, 1984; Marcogliese, 1992), but S. thomasi is considered of uncertain status (Chubb et al., 2006; French and Muzzall, 2008). Identification to species level was often made based on a combination of fish host and plerocercoid segment counts, but ambiguity is resolved more recently via DNA sequencing (Chubb et al., 2006; Nishimura et al., 2011).
Schistocephalus solidus (Müller, 1776) is considered a specialist infecting threespine stickleback (Gasterosteus aculeatus) across North America (Reimchen, 1982; Holloway, 1984; LoBue and Bell, 1993). Morphologically distinct forms of S. solidus are described from threespine stickleback with plerocercoid segment counts ranging 48–100 and 99–138 for each of the 2 host phenotypes (Bakker and Sevenster, 1988; Chubb et al., 2006). One Schistocephalus sp. infecting wild ninespine stickleback is reported previously from southwestern New Brunswick (Smith and Kramer, 1987), S. pungitii (Dubinina, 1959) is reported from ninespine stickleback from Sable Island near the neighboring province of Nova Scotia (Marcogliese, 1995), and S. solidus is reported from ninespine stickleback in the neighboring province of Quebec (Coad and Power, 1973). Schistocephalus pungitii is generally regarded to infect ninespine stickleback, S. solidus infecting threespine stickleback, and with recent evidence supporting distinct clades of Schistocephalus sp. infecting both hosts (Nishimura et al., 2011). Evidence for hybridization between S. pungitii and S. solidus with hybrids infecting both hosts (Henrich et al., 2013) suggests further caution regarding definitive species-level identification.
Schistocephalus cotti is a distinct species described from European bullhead (Cottus gobio) in Finland with plerocercoids having 103–189 segments (Chubb et al., 2006). French and Muzzall (2008) identified plerocercoids from slimy sculpin in Lake Michigan as an undetermined Schistocephalus sp. Whereas plerocercoid segment number (n = 113) is compatible with S. cotti (see Chubb et al., 2006), and despite their slimy sculpin host being congeneric with European bullhead (both Cottus species), French and Muzzall (2008) cautioned against species-level identification due to past challenges with accurate Schistocephalus identification. Harmon et al. (2015) similarly concluded that only generic-level identification of Schistocephalus sp. plerocercoids was warranted for specimens from 2 Cottus species hosts (coastrange sculpins and slimy sculpins) in 1 Alaskan lake. Although plerocercoid segment counts from our slimy sculpin (143–177; average 156) were compatible with S. cotti (Chubb et al., 2006), sequencing of both partial ND1 and COI resulted in low sequence identity with S. cotti and all other Schistocephalus spp. available in GenBank. This supports an alternative species identification, potentially a new but currently undetermined species. Definitive identification awaits more widespread sequencing of Nearctic specimens due to the uncertain status of Schistocephalus spp. from this region (Chubb et al., 2006).
To our knowledge, no previous publications document Ligula intestinalis or this Schistocephalus sp. from New Brunswick, Canada. It remains to be determined whether these parasites are recent introductions or result from environmental change promoting their emergence. Despite outstanding species-level identification of the Schistocephalus sp., plerocercoids from species in both genera raise ecological concerns because they can induce sterility of host fishes (Heins and Baker, 2010; Heins, 2017; Biswas and Ash, 2021). Their influence on local fish populations remains to be determined, but epizootics induced by plerocercoids of both parasites are reported (Kennedy et al., 2001; Heins and Ecke, 2012). Future efforts will assess parasite distribution, host utilization, seasonality of infections, and reproductive consequences in local fishes to establish baseline data at this time of environmental change.
ACKNOWLEDGMENTS
The authors assert all applicable international, national, and/or institutional guidelines for the care and use of animals were followed. Fish collections were approved by the University of New Brunswick Animal Care and Use Committee Protocol no. 22041. Fish collections from Fundy National Park were further approved by Parks Canada (permit no. FNP-2022-44019). We thank Becky Graham, Allain Cassie, Sara Plant, and John Robinson of Parks Canada for assistance with fish collections from Wolfe Lake in Fundy National Park. This project was funded by the Natural Sciences and Engineering Research Council of Canada Collaborative Research and Development Grant with J. D. Irving, Ltd. (M.A.G.); discretionary research funds (M.S.D.); New Brunswick Innovation Foundation STEM and Social Innovation Awards (M.L.F. and K.R.D.); a University of New Brunswick Accelerated Masters Award (K.D.R.D.); and the Michael D. B. Burt Memorial Scholarship (K.D.R.D.). Additional graduate student financial support was provided by the University of New Brunswick via graduate research assistantships and graduate teaching assistantships (M.L.F. and K.D.R.D.).