EDP Sciences logo
Submit your paper
Search
Menu
All issues Volume 30 (2023) Parasite, 30 (2023) 42 Full HTML
Open Access
Issue
Parasite
Volume 30, 2023
Article Number 42
Number of page(s) 23
DOI https://doi.org/10.1051/parasite/2023042
Published online 19 October 2023

© O.M. Amin et al., published by EDP Sciences, 2023

Licence Creative CommonsThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Introduction

Cystacanths and adults of Profilicollis spp. (Polymorphidae) are common parasites of marine crabs (Decapoda) and shore birds, respectively, mostly along the Pacific and Atlantic coasts of North and South America [25, 50, 51, 54, 56]. Three exceptions include: (i) Profilicollis formosus (Schmidt and Kuntz, 1967) Hoklova, 1974 from the freshwater crayfish Macrobrachium mammillodactylus (Thallwitz) and the domestic duck Anas platyrhynchos Lin. in Taiwan [65], (ii) Profilicollis major Lundström, 1942 originally described from Sweden [43] but since found in Maine [50] in scaup, ducks, scoters, and sea otters [62], and (iii) Profilicollis novaezelandensis Brockerhoff and Smales, 2002 originally described in New Zealand [13]. Profilicollis rancoensis n. sp. represents another exception, like the cystacanths, and has been described from a freshwater South American endemic decapod anomuran Aegla abtao Schmitt 1942. A peculiarity of the species of the genus Aegla, is that its origin is marine, and because the elevation of Andes was gradual, this allowed the present day Aeglas to adapt to less saline waters [63]. Some of these species were included in dichotomous keys by [12, 33, 52, 59] based on intraspecific variations in proboscis armature and length. The latter key was most comprehensive and Amin [12] proposed synonymizing Hexaglandula Petrochenko, 1950 and Subcorynosoma Hoklova, 1967 with Polymorphus Lühe, 1911.

Most of the studies related to the hosts to species of Profilicollis were reported from Pacific South America. Some species of Profilicollis may also infect vertebrates other than birds [69]. Profilicollis altmani (Perry, 1942) Van Cleave, 1947 can also successfully infect mammals, with potentially serious outcomes that may occasionally represent a public health hazard [46, 57, 69]. In California, United States morbidity and mortality of the southern sea otter were attributed to massive infections with P. altmani reaching 8,760 worms per animal, causing intestinal perforations, nutrient depletion, and mortality [46].

Our work contributes to available knowledge of the genus Profilicollis, already known to have wide host use and a potential high impact on animal and even human populations. Here, we document the known morphology of this parasite with observations of newly observed structures using scanning electron micrographs and color optical micrographs, provide comparative morphometrical information, analyze the chemistry of hooks and spines for the first time, and produce new molecular analysis of cystacanths of a new acanthocephalan species from Chile.

Materials and methods

Collections

Collections of the decapod Aegla abtao were made by Lake Ranco at Riñinahue river inlet of Ranco Lake, Los Lagos, Ranco Province, north Patagonia, Chile (40.3331285°S–72.239685°W) at 68 m a.s.l. by hand from logs and large rocks (Fig. Sup.). Once collected, the individuals were sexed, distinguishing females from males by the presence of four pairs of abdominal pleopods, which are used to carry fertilized eggs during the spawning period. An initial collection of 27 crustaceans was made in May 2022, in which we found 7 cystacanths from 4 Aeglas (3 males, 1 female). Then in September 2022, we collected 81 crustaceans, in which five individual hosts (4 males, 1 female) were found to be infected with 24 acanthocephalan cystacanths. A third field task was carried out on November 10: we collected 65 crabs, in which four hosts (2 males, 2 females) were found harbouring 6 cystacanths. Another collection of 16 cystacanths and 2 acanthellae was made on December 27, 2022 obtained from 8 hosts (3 males, 5 females), of which 13 specimens and the 2 acanthellae were sent to PCI. We sent a total of 7 specimens to our affiliate laboratory at Brigham Young University, Provo, Utah for SEM and Energy Dispersive X-ray analysis. The remaining 27 specimens (11 males and 16 females) were processed for microscopical studies. A total of 11 specimens were used for sequencing and molecular analysis at Sistematica Lab., Universidad Austral de Chile. A subsequent collection of 259 crabs on four field excursions yielded 53 cystacanths, which were studied.

Deposited material: Specimens were deposited in the University of Nebraska’s State Museum’s Harold W. Manter Laboratory (HWML) collection, Lincoln, Nebraska, United States.

Optical microscope images

Optical microscope images were acquired using a BH2 light Olympus microscope (Olympus Optical Co., Osachi-shibamiya, Okaya, Nagano, Japan) attached to an AmScope 1000 video camera (United Scope LLC, dba AmScope, Irvine, CA, USA), linked to an ASUS laptop.

Scanning electron microscopy (SEM)

Seven specimens that had been fixed and stored in 70% ethanol were subsequently unsheathed and processed for SEM, following standard methods (Lee, 1992). These methods included drying with a critical point dryer (Tousimis Autosamdri 931.GL; Tousimis, Rockville, MD, USA) and mounting on aluminium stubs using conductive double-sided carbon tape. Samples were coated with an approximately 20 nm layer of 80%–20% gold-palladium using a Quorum Q150T ES magnetron sputtering system (Quorum, Laughton, United Kingdom) equipped with a tilted rotating stage. Samples were placed and observed in a Helios Dual Beam Nanolab 600 Focussed Ion Beam/Scanning Electron Microscope (FIB/SEM) (FEI, Hillsboro, OR, USA). Scanning electron micrographs were acquired using the Helios SEM, with an accelerating voltage of 5 kV, and a probe current of 86 pA, at high vacuum, using and Everhart–Thornley secondary electron detector.

Focused Ion Beam (FIB) sectioning of hooks

The Helios Dual Beam FIB/SEM, mentioned above, is also equipped with a Focussed Ion Beam (FIB) column with a Ga+ Liquid Metal Ion Source (LMIS). The FIB allows for in situ site-specific sectioning of the samples by irradiating precise areas of the sample with a focussed beam of Ga+ ions, which sputter material away and erode the specific areas of the samples. To minimize sample damage due to FIB irradiation, the hooks of the acanthocephalans were centered on the stage while imaging with the SEM. Sectioning time is dependent on the volume of tissue removed, the nature and sensitivity of the exposed tissue, and ion accelerating voltage and current. For this work, the hooks of the acanthocephalans were sectioned using an ion accelerating voltage of 30 kV and a probe current of 2.7 nA in two steps: initial rough removal of bulk material, followed by more precise “cross-section cleaning” that resulted in a smoother exposed surface for SEM imaging and X-ray microanalysis. The hooks were sectioned perpendicular to the long axis at three locations, near the tip of the hook, near the middle of the hook, and near the base. Additional longitudinal sections were prepared near the middle of the hook, which required longer mill times due to the volume of material required for removal.

Energy Dispersive X-Ray analysis (EDXA)

The Helios Nanolab 600 is equipped with a TEAM Pegasus system with an Octane Plus detector (EDAX, Mahwah, NJ, USA). Energy Dispersive X-Ray (EDX) spectra were collected from the center at each of the exposed surfaces of the sectioned cuts. EDX spectra were collected using an electron accelerating voltage of 15 kV, and a probe current of 1.4 nA. Data collected included images of the sectioned surfaces, images of the displayed spectra, as well as the raw EDX collected data. Relative elemental weight-percentages were generated by the TEAM software. To maintain consistency within the TEAM software, and to ease comparison among various spectra, the following elements list was kept constant among all EDX scans: C, N, O, Na, Mg, Al, P, S, K, Ca, Fe, Au, Pd, and Ga (being aware that Au, Pd, and Ga were process elements used during sample preparation and milling).

DNA extraction, PCR and sequencing

DNA of parasites, obtained from body cavity of hosts, was extracted from entire individuals (N = 11). Genetic comparison and phylogenetic analyses were based on a fragment of 627 bp of the mitochondrial cytochrome oxidase I (COI) gene and 760 bp fragment of the partial 18S rDNA gene. The samples were digested overnight at 55 °C and genomic DNA was isolated using a commercial extraction kit (Wizard® Genomic DNA Purification Kit, Promega, Madison, WI, USA). A fragment of the COI gene was amplified using the primers detailed by Folmer [1622], following the protocol of Amin [67]. The same parasite individuals (N = 11) were used to acquire 18 S rDNA sequences. PCR was performed in 30 μL volume reactions containing 2 × red PCR premix (Ampliqon, Odense, Denmark), 20 pmol of each primer, 3 μL of extracted DNA, and primers MGF (5′–GATCGGGGAGGTAGTGACG–3′) and MGR (5′–ACCCACCGAATCAAGAAAGAG–3′). PCR conditions for 18S rDNA were those of Rodríguez et al. [58]. PCR products were analyzed on 1.5% agarose gel and visualized with a UV transluminator. Later, the PCR products were purified and sequenced using an external sequencing service (Macrogen, Inc., Seoul, South Korea). Finally, all new DNA sequences were edited using Codon-Code (Codon Code Aligner, Dedham, MA, USA) and deposited in GenBank (COI: OQ417136–OQ417146; 18S: OQ434741–OQ434748).

Phylogenetic analysis

The 11 new COI sequences were integrated into a matrix with a representative of each genus of the family Polymorphidae available in GenBank. A total of 41 sequences belonging to the family Polymorphidae were downloaded from GenBank (Table 1) and analyzed. In addition, we used as the outgroup the phylogenetically related species Plagiorhynchus cylindraceus and Centrorhynchus globocaudatis (Plagiorhynchidae), and Sphaerirostris lanceoides (Centrorhynchidae). As such, the analyzed matrix has a total of 55 sequences. A second matrix was made with 15 sequences of COI of each species of the Profilicollis available in GenBank and we used P. cylindraceus as the outgroup.

Table 1

Species of acanthocephalans of the Polymorphidae family, their hosts, locations and GenBank accession number of the sequences used in the phylogenetic analysis. ND = No data available. (–) = Not indicated.

Sample size for the partial 18S rDNA gene was slightly smaller than for COI, including eight sequences of the new species of Profilicollis and 27 sequences of members of the family Polymorphidae available in GenBank (Table 1). One sequence of Plagiorhynchus cylindraceus (Plagiorhynchidae family) was used as the outgroup (Table 1). As such, the analyzed 18S rDNA matrix had a total of 36 sequences. As with COI, a second matrix was created with 10 sequences of 18S rDNA of each species of the genus Profilicollis available in GenBank. One sequence of P. cylindraceus was used as the outgroup.

Each gene matrix was aligned separately using MAFFT v.7 software [37], allowing the program to choose the alignment strategy (L-ins-i). Four models were carried out: two to analyze the family Polymorphidae, and the other two to analyze the genus Profilicollis. To select the best-fitting model of molecular evolution, we used the proposed model tool in the IQ-Tree v1.6.12 program [35], which selected K3Pu+F+I+G4 and TIM3e+I+G4 for the COI and 18S matrix of the family Polymorphidae, respectively and HKY+F+G4 for COI and TPM3+F for 18S for the matrix of the genus Profilicollis. We used the maximum-likelihood (ML) approach implemented in IQ-Tree v1.6.12 [72]. For each matrix, we conducted the following analyses: a) Five runs modifying the strength of the perturbation parameter from 0.5 (default value) to 0.3, b) Five runs using the default value (0.5) for the strength of the perturbation parameter, and c) Five runs modifying the strength of the perturbation parameter from 0.5 (default value) to 0.7.

In all cases, the number of unsuccessful interactions to stop parameter (−nstop) value was changed from 100 (default value) to 1000. Trees with the highest likelihood score were chosen (family Polymorphidae: COI = log-likelihood: −11404.169, −pers 0.3 and −nstop 1000; 18S = log-likelihood: −6647.679, −pers 0.3 and −nstop 1000) (genus Profilicollis: COI = log-likelihood: −2599.057, −pers 0.3 and −nstop 1000; 18S = log-likelihood: −3574.388, −pers 0.5 and −nstop 1000). Support for the nodes was evaluated using two approaches: the aBayes test [11] and the ultrafast bootstrap procedure using 1000 replicates [31]. Finally, observed genetic p-distances (p) between haplotype and sample pairs were calculated in MEGA 7 [68].

Results

Of a total of 259 crabs examined (A. abtao), 21 were parasitized (prevalence [P] = 8.10%), yielded a total of 53 cystacanths. All were from the body cavity (mean abundance [MA: amount of parasites /number of hosts] = 0.2; mean intensity [MI: amount of parasites/number of infected hosts] = 2.5). The number of cystacanths per freshwater crab ranged from 1 to 11. Of the total number of crabs caught, 193 were male and 66 were female. The prevalence (P) in males was 4.2% vs. (P) in females 3.8%. The MA in males was 0.13, and 0.30 in the female cystacanths per host. The MI values were 3 and 2 cystacanths per parasitized host, respectively.

The microscopical study provided considerable information depicted in the line drawings (Figs. 1 –6). The scanning electron microscopy studies produced considerable new detail adding more information not readily available by the optical microscopy study. Our descriptive account of cystacanths is based on 27 cystacanths (11 males and 16 females) and 2 acanthellae measured and studied with optical microscopy from whole mounts (Figs. 1–6) and 7 cystacanths studied with SEM (Figs. 7–22).

thumbnail Figures 1–4

Cystacanths of Profilicollis rancoensis n. sp. from the freshwater crab, Aegla abtao Schmitt (Decapoda) in Chile. (1, 2) A male and female cystacanth. (3) One longitudinal row of proboscis hooks showing the variable root structures with different degrees of manubriation. (4) Profile of a trunk spine.

thumbnail Figures 5, 6.

Two late acanthellae at slightly different ages.

thumbnail Figures 7–10

SEM of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, northern Patagonia, Chile. (7) A montage of a whole specimen after excystation. (8) A lateral view of a proboscis showing the organization of hooks. Note that some hooks in the posterior circle appear to emanate from the neck. (9) A semi-apical view of a proboscis showing the smaller apical hooks and the external appearance of possible apical organ. (10) Posterior circles of hooks showing the insertion of some posterior-most hooks outside of the actual proboscis in the neck.

thumbnail Figures 11–16

SEM of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, Valdivia, Chile. (11–13) Anterior, middle and posterior hooks in profile. Note differences in size, shape, and curvature. (14–16) Anterior, middle and posterior spines; their shape is variable.

thumbnail Figures 17–22

SEM of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, Valdivia, Chile. (17) The posterior tip of the corrugated posterior trunk (in red). The drying process may have made the corrugation more evident. Manifestations of the various size and distribution of micropores at the tip of a hook (18), proboscis base (19), trunk spine (20), and middle and posterior trunk locations (21, 22).

Primary diagnostic characters of P. rancoensis cystacanths including trunk length and proboscis armature (Table 2) are compared with those of 9 other species of Profilicollis: adults of P. major from Europe and Maine [58, 68] in scaup, ducks, scoters, and sea otters [62], and Profilicollis novaezelandensis from New Zealand in oystercatcher [16]. Morphometric details of male and female cystacanths of P. rancoensis are noted in Table 3.

Table 2

Diagnostic characteristics of Profilicollis rancoensis n. sp. from the freshwater crab Aegla abtao Schmitt, 1942 (Decapoda) in Chile compared to the 9 other species in the genus.

Table 3

Comparative morphometrics of male and female cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, Valdivia, Chile.

Profilicollis rancoensis n. sp.

urn:lsid:zoobank.org:act:8821DEF9-2BFC-49E2-BB9E-5D0A2718F688

Host: The freshwater Pancora crab Aegla abtao Schmitt, 1942 (Crustacea: Decapoda)

Type locality: Riñinahue river inlet of Ranco Lake, Los Lagos, Chile (40.3331285°S–72.239685°W)

Site of infection: Body cavity cephalothorax

Type specimens: Cystacanths & 1 acanthella in HWML Parasitology Collection no. (HWML 216948–HWML 216950)

Etymology: The new species is named for the collection locality of the decapod intermediate host.

Remarks on cystacanths

Cystacanths of P. rancoensis n. sp. are distinguished from cystacanths of all other species of Profilicollis by a combination of the following characters: “In freshwater decapod, Aegla abtao from Chile. Proboscis with 14–15 rows of 6–7 hooks each reaching 105–110 μm. All hooks with slight anterior manubrial and strong simple roots; basal hook with small simple root and anterior manubrium. Lemnisci long, slender.” With the exception of P. formosus whose cystacanth develop in the freshwater knob-tooth prawn Macrobrachium mammillodactylus Thallwitz in Taiwan [65], all other species of Profilicollis utilize marine decapods as intermediate hosts (Table 2). A special note needs to address P. arcticus whose cystacanth hosts are yet to be identified. The adults of P. arcticus have been described and reported from sea ducks, i.e., king eider Somateria spectabilis Linn. and Pacific eider S. nigra Gray from the arctic coasts of North America, NW Canada, and Russia. The marine distribution of these shore ducks suggests that the intermediate hosts of cystacanths of P. arcticus may well be marine decapods as has been suggested by Nickol et al.’s [50] characterization of Profilicollis. In addition to being the only species of Profilicollis using a freshwater decapod as intermediate host, P. rancoensis is further distinguished from all other species of Profilicollis by the following key. The key includes only one major intermediate host of each species as there may be many in some species. Note that hooks attain their maximum size in cystacanths, which is usually comparable to that in adults of the same species. The detailed coverage in Table 2 does not include all the characters selected for the key. Diagnostic characters used were chosen among features common to both sexes of cystacanths and adults, especially proboscis armature. Reference to simple roots indicates normal posteriorly directed roots.

Key to cystacanths of species of Profilicollis

1. In freshwater prawns, Macrobrachium mammillodactylus from Taiwan. Proboscis with 12–15 rows of 7–9 hooks, each reaching 95–117 μm. Anterior hooks with strong simple roots; posterior hooks rootless. Lemnisci long, flat. ….………………………………………………………Profilicollis formosus

– In decapod crustaceans elsewhere. Proboscis armature and lemnisci variable………………………….…2

2. In freshwater decapod, Aegla abtao from Chile. Proboscis with 14–15 rows of 6–7 hooks, each reaching 105–110 μm. All hooks with slight anterior manubrial and strong simple roots; basal hook with small simple root and anterior manubrium. Lemnisci long, slender…………………….…..…….Profilicollis rancoensis n. sp.

– In marine decapods elsewhere. Proboscis armature and lemnisci variable………………………………….3

3. In Emerita analoga from Pacific and Atlantic North and South America. Posterior trunk with tail. Proboscis with 21–35 rows of 9–16 hooks, each reaching 51–70 μm. All hooks rooted with prominent anterior manubria. Lemnisci plump, usually as long as receptacle……………………………………………Profilicollis altmani

– Posterior trunk without tail. Proboscis with 14–22 rows of 6–10 hooks each. ………………………………..4

4. In Paragrapsus spp. from Brazil, Australia and Tasmania. Anterior trunk cylindrical. Proboscis with 17–21 rows of 7–8 hooks, each reaching 69–73 μm. Only anterior hooks with simple roots and no manubria. Lemnisci long, cylindrical, and slender………………………………..…….…Profilicollis sphaerocephalus

– Anterior trunk ovoid. Proboscis with 14–22 rows of 5–10 hooks each……………………………………… 5

5. In Hemigrapsus crenulatus from New Zealand. Proboscis with 14–16 rows of 5–8 hooks each reaching 85–110 μm. Hook tips recurved. Apical and posterior hooks rootless. Roots of larger anterior hooks simple, longer than blades. Lemnisci long, thick and cylindrical………………….……………….Profilicollis novaezealandensis

– Proboscis with more hook rows (15–22). Hook tips not recurved……………………………………………6

6. In Neohelice (=Chasmagnathus) granulata from Uruguay and Argentina. Proboscis with 16–22 rows of 7–9 hooks, each reaching 83–91 μm. Hooks similar but more slender posteriorly; only second anterior hook with short simple root. All other hooks slightly manubriated anteriorly with stubby roots. Lemnisci cylindrical, relatively longer than receptacle………………………………………………………Profilicollis chasmagnathi

– Proboscis hooks and roots dissimilar; hooks reaching 118 μm. Lemnisci variable………..………..….……7

7. In Hemigrapsus crenulatus & Helice crassa from Chile, New Zealand, South Shetland Islands. Proboscis with 18–22 rows of 6–9 hooks, each reaching 71–80 μm. Apical hook miniature. Anterior hooks with prominent simple roots and no manubria. Posterior hooks with abbreviated simple roots and well-developed manubria. Lemnisci cylindrical, thick, longer than receptacle, wider at middle…Profilicollis chasmagnathi*

– Proboscis with 15–22 rows of 7–10 hooks each. Shape of hooks, roots and lemnisci different………………8

8. Intermediate host unknown. In NW Canada and Russia. Proboscis with 22 rows of 7–8 hooks, each reaching 89–118 μm. Apical and basal hooks smallest, rootless. Anterior and middle hooks with simple roots. Posterior hooks more slender, gradually deceasing in size posteriorly, with rudimentary roots. Lemnisci pyriform, slightly longer than receptacle…………………………………………………………Profilicollis arcticus

– Proboscis with fewer hook rows (15–20) and different hook/root and lemnisci morphology……………..…9

9. In Hemigrapsus oregonensis and Carcinus maenas from North America, Scotland, and Eurasia. Proboscis with 15–20 rows of 7–8 hooks, each reaching 87–96 μm. Anterior and middle hooks with simple roots; posterior hooks with rudimentary roots. Lemnisci plump-bulbous, about as long as receptacle………………………………………………….…Profilicollis botulus

– In Cancer irroratus from Maine and from other intermediate hosts in Sweden and Europe. Proboscis with 16–20 rows of 7–10 hooks, each reaching 92–108 μm. All hooks without manubria and with simple roots except rootless basal hook. Lemnisci long, cylindrical, widest at middle……………..…Profilicollis major

* Second report of Profilicollis chasmagnathi from Hemigrapsus crenulatus was originally reported as Profilicollis antarcticus.

Description of cystacanths of Profilicollis rancoensis from Aegla abtao in Chile

With characters of the family Polymorphidae and genus Profilicollis. Body flattened dorso-ventrally, divided by constrictions into proboscis, long cylindrical neck, spinose anterior trunk, and ovoid posterior trunk (Figs. 1, 2). See comparative measurements and counts of species of Profilicollis in Table 2 and complete specific measurements of P. rancoensis in Table 3. Proboscis ovoid with blunt anterior end and 14–15 longitudinal rows of 6–7 hooks each. Hooks of similar shape (Fig. 3), gradually decreasing in length and diameter posteriorly from largest hooks (2nd to 4th). Basal hooks smallest with shortest roots and longest anterior manubrium. Some basal hooks appear to emanate from the neck and not directly from proboscis (Figs. 8, 10). All other hooks with slightly shorter roots and unremarkable anterior manubria (Fig. 3). Complete measurements of hooks in 1 longitudinal row in Table 3. Edge of hook tips with high level of sulfur and low levels of phosphorous and calcium and center of mid-hook showing an apparent opposite trend (Figs. 23, 24) and Table 4). Neck long widening at base. Proboscis receptacle with single-layered muscle wall extending from base of proboscis to about anterior third of spinose trunk. Primordia of cephalic ganglia detectable in receptacle just anterior to level of junction of neck with anterior trunk. Trunk with many electron-dense micropores (Figs. 21, 22) and many hypodermic nucleated cells appearing as round cysts. Anterior trunk with somewhat irregular circles of spines ending at trunk constriction. Spines pointed with broad base, dense cortical layer and spongy core (Fig. 4). Lemnisci long, slender, extending well beyond receptacle, often to level of posterior end of anterior trunk. In males, ovoid testes usually appear diagonally in anterior trunk (Fig. 1) but may also be less frequently found in posterior trunk or singly in both trunk segments at same time. One elemental tubular cement gland emerging from each testis posteriorly with rest of male reproductive system in posterior trunk remaining underdeveloped and undefined. Posterior trunk with corrugated texture on surface. In female cystacanths, parts of developing reproductive system barely distinguishable (Fig. 2). Inner subcutaneous muscle layer well developed in both trunk segments and fibrous connective tissue bundles join anterior and posterior trunks together (Fig. 1, 2).

thumbnail Figures 23–25

Energy-Dispersive X-ray Spectra (EDXS) of anterior hooks of Profilicollis rancoensis n. sp. cystacanths from Aegla abtao in Chile, acquired from the edge of a tip hook (23), the center of the middle section of the hook (24), and the center of the base of the hook (25), where the red inlays indicate the region from where the spectra were obtained. The spectra show the raw number of counts collected, and the elements C, O, P, S, Ca peaks highlighted. See Table 4 for EDXA numerical detail. Insets point to parts of hooks used to generate spectra.

Table 4

Elemental weight percent of Na, Mg, P, S, K, and Ca (as reported by TEAM EDXA software) of hooks and spines of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Chile, at 6 hook sites. Weight percentages of other elements (C, N, O, Al, Fe, Au, Pd, and Ga) are omitted.

Description of acanthellae of Profilicollis rancoensis from Aegla abtao in Chile

Our secondfourth collection of cystacanths on December 27 included two late acanthellae of slightly different ages measuring 2.25 × 0.75 and 2.75 × 0.6 mm, long × wide (Figs. 5, 6). The acanthellae were as large as the cystacanths and, like the cystacanths, were enveloped within membranous sheaths. Both acanthellae had conspicuously inverted invaginated proboscides with delicate sheaths and under-developed hooks. Single-layered proboscis receptacle prominent, with delicate sheaths, enclosing proboscides. Many mini nuclei reminiscent of the earlier giant hypodermal nuclei appeared in large number anteriorly in one specimen (Fig. 5). Each specimen had a developing or developed cercomer-like tail clearly absent in the cystacanths. The primordia (nuclei, fragments, or embryonic elements) of lemnisci, reproductive structures, trunk spines, cephalic ganglia or muscular layers in proboscis, receptacle or trunk were not evident.

Micropores

Micropores have been detected and documented from every single tissue studied including hooks (Fig. 18), proboscis (Fig. 19), spines (Fig. 20) and all regions of the trunk (Figs. 21, 22). It appears that every possible surface area in these cystacanths is dedicated to the differential absorption of nutrients that occurs through micropores.

Energy Dispersive X-Ray analysis (EDXA)

Energy Dispersive X-ray analysis was used to qualitatively assess the levels of various elements in the hard structures of the Acanthocephala, among other animals, to account for the hardness of structures such as the hooks and spines. As previously mentioned, EDX spectra were collected from the hooks at various surfaces exposed by the FIB milling, paying attention to the center (core) of the hook and the sheathing material (edge). Weight-percents for various metals (as reported by the TEAM software) are summarized in Table 4. Sample spectra and their typical collection areas are shown in Figure 23–25. Highest levels of sulfur were observed in the sheathing regions of the hooks (i.e., hook edges). The core regions of the hooks (i.e., base and center) contained mostly phosphorus and calcium, the two essential elements for hook structural support.

Molecular results

A total of 19 new sequences were successfully obtained (11 of the COI gene and 8 of the 18S gene). These novel genetic sequences of COI and 18S rDNA were 14.11% and 21.49% dissimilar, respective to those available in GenBank for P. botulus. Both phylogenetic trees gathered via ML and Bayesian inference (BI) for COI were congruent (Fig. 26A). The family Polymorphidae was recovered as shown to be monophyletic with high support only for the BI analysis and moderate support in the ML analysis (PP = 0.99; BS = 66). Within the clade of Polymorphidae, three main lineages were found (Fig. 26A). One lineage is a highly supported clade (PP = 1; BS = 97) and is formed by the genus Andracantha, Corynosoma and Bolbosoma (Fig. 26A). Within it, the genus Andracantha is found to be paraphyletic with respect to Corynosoma bullosum, while the other species of Corynosoma form a highly supported clade (PP = 0.97; BS = 89). The second main clade (PP = 1; BS 98) of Polymorphidae is formed by Southwellina hispida and a clade (PP = 1; BS = 100) formed by two of the three species of Polymorphus included in the analysis, P. brevis and Polymorphus sp. The last main lineage (PP = 0.98; BS 53) of Polymorphidae is formed by Profilicollis, Polymorphus trochus, Arhytmorhynchus, Tenuisoma, Pseudocorynosoma, Hexaglandula, and Ibirhynchus. Here, Profilicollis appears paraphyletic with respect to Polymorphus phippsi. This later species is sister, in a highly supported relationship (PP = 0.99; BS = 80), to the clade (PP = 1; BS = 100) formed by Profilicollis chasmagnathi and P. novaezelandensis. Profilicollis botulus is sister (PP = 1; BS = 99) to a highly supported clade (PP = 1; BS = 100) formed by all haplotypes of P. rancoensis (Fig. 27A).

thumbnail Figure 26

Genealogical relationships of haplotypes based on partial COI (A) and 18S (B) gene sequences of specimens of the family Polymorphidae recovered in a Bayesian inference analysis. Numbers next to nodes refer to support values. Bayesian posterior probability values are shown left, and Bootstrap proportions gathered in the Maximum Likelihood analysis (COI: Ln = −11404.169, 18S: Ln = −6647.679). GenBank accession numbers are included in the terminal labels.

thumbnail Figure 27

Genealogical relationships of haplotypes based on partial COI (A) and 18S (B) gene sequences of specimens of the genus Profilicollis recovered in a Bayesian inference analysis. Numbers next to nodes refer to support values. Bayesian posterior probability values are shown left, and Bootstrap proportions gathered in the Maximum Likelihood analysis (COI: Ln = −2599.057, 18S: Ln = −3574.388). GenBank accession numbers are included in the terminal labels. Animal silhouettes with different box colors to the right of the trees indicate the intermediate host (IH) of each species. Blue = A. abtao; Green = Emerita spp.; Orange = Varunidae; Red = Portunidae.

The sample of P. rancoensis shows extremely low genetic variation. On average, the analyzed haplotypes of this species differed by 0.3%. For example, 3 specimens shared the same haplotype. Haplotypes of P. rancoensis differed from that of P. botulus by 15%. Similarly, P. rancoensis haplotypes differed by 24% and 28% from P. chasmagnathi and P. novaezelandensis, respectively.

The resulting ML and BI trees for the 18S rDNA matrix are mostly congruent (Fig. 26B). The family Polymorphidae includes two lineages. One of these two lineages is formed by the genera Andracantha, Bolbosoma, Corynosoma, Arhytmorhynchus, Southwellina, Hexaglandula, Ibirhynchus, Polymorphus, and Pseudocorynosoma. This clade appears to be well supported, in particular in the IB analysis (PP = 0.99; BS = 78; Figs. 26A and 26B). The other main lineage is highly supported (PP = 1; BS = 99) and is formed by the genera Profilicollis and Polymorphus, which are not reciprocally monophyletic (Fig. 27B). Haplotypes of P. rancoensis form a highly supported clade (PP = 1; BS = 100). Profilicollis rancoensis forms a highly supported clade (PP = 1; BS = 93) with the species P. altmani, Polymorphus sp., and P. botulus. In turn, this clade is sister (PP = 1; BS = 99) to the clade (PP = 1; BS = 100) formed by P. minutus and P. obtusus, which fall in a strongly supported clade (Fig. 27B). The analyzed haplotypes of P. rancoensis differed on the average by 0.6%. In this case, two specimens shared the same haplotype. On average, haplotypes of P. rancoensis differed from those of P. botulus and of P. altmani by 3% and 2%, respectively.

Discussion

Concept and position of Profilicollis vs. Polymorphus based, in part, on Amin (1992)

The taxonomic status of Profilicollis has been in flux since Meyer [47] created the genus based on the long neck and spheroid proboscis and included 2 species, P. botulus and P. arcticus. The generic of features of Polymorphus, erected for one species, P. minutus (Goeze, 1782), by Lühe [42] included small body size, body wall nuclei, grid-like lacunar system, trunk spines anterior to constriction, radially symmetrical proboscis hooks which decreases in size anteriorly and posteriorly, double-walled proboscis receptacle with cephalic ganglion near its base or middle, long neck, moderately long lemnisci, testes behind one another, tubular cement glands, genital opening terminal without spines, and eggs with polar prolongation of middle membrane. Except for features like the long neck or the position of the testes, the above diagnosis remains largely valid today. As the number of species included in the genus increased, the generic concept continued to expand, i.e., see Southwell and MacFie [66], Travassos [71], and Thapar [70]. Many of the diagnostic features added by various authors were too restrictive to be of generic value as the interspecific variability within Polymorphus continued to increase with the discovery of more species [1, 52]. These restrictive diagnostic features are noted in the following few paragraphs.

In Meyer’s [47] system, Polymorphus was correctly placed in his family Polymorphidae and order (Class) Palaeacanthocephala, and included 14 species [48] all of which are presently recognized members of the genus, except P. magnus which has been synonymized with P. minutus. In his brief diagnosis, Meyer [48] restrictively referred to short compact bodies cylindrical or weakly ovoid proboscides, trunk spines anterior to mid constriction, tubular cement glands, string-like lemnisci, and numerous small nuclei in the body wall. Meyer [47] also established a new genus, Profilicollis, for two long-necked species Polymorphus botulus and Polymorphus arcticus (=Filicollis botulus Van Cleave, 1916 and Filicollis arcticus Van Cleave, 1920) with eggs with concentric membranes, but kept Filicollis Lühe, 1911 for Filicollis anatis (Schrank, 1788) and Filicollis sphaerocephalus (Bremser in Rudolphi, 1819). Witenberg [80] and later Van Cleave [74–76] synonymized Profilicollis with Polymorphus and Van Cleave [74] reassigned F. botulus and F. arcticus as well as F. sphaerocephalus and Filicollis altmani Perry, 1942 to Polymorphus [76]. Webster [79] recognized the diversity within Polymorphus and erected a new subgenus, Falsifilicollis for forms with spheroidal proboscis and slender elongate neck previously included in Filicollis. These included Polymorphus sphaerocephalus, Polymorphus altmani, Polymorphus kenti Van Cleave, 1947 and Polymorphus texensis Webster, 1948. Schmidt and Kuntz [6591] correctly pointed out that Profilicollis Meyer, 1931 has priority for the concept of Falsifilicollis Webster, 1948. Petrochenko [52] erected Filicollidae for Filicollis Lühe, 1911 including only F. anatis (Schrank, 1788) Lühe, 1911, and Parafilicollis Petrochenko 1956 including P. major, P. altmani, P. kenti (a junior synonym of P. altmani) and P. sphaerocephalus. His three major diagnostic features of Parafilicollis were the long neck, spheroidal proboscis, and eggs without polar prolongation of middle membrane. Petrochenko [52] placed Parafilicollis in Gigantorhynchidae Southwell and MacFie, 1925 based primarily on the egg shape. Schmidt and Kuntz [65] declared Parafilicollis as without status because Meyer [47] erected Profilicollis for the same concept. The shape of eggs was clearly shown to be of no diagnostic value at the generic level in Polymorphidae by Van Cleave [75], Schmidt and Kuntz [64, 65] and Amin [1]. Golvan [24] accepted Profilicollis (= Falsifilicollis Webster, 1948) as a subgenus diagnosed with spheroidal proboscis and long slender neck, and which included the same four species listed by Webster [79], but without indicating a subgenus for the other species of the genus. Yamaguti [83] synonymized Profilicollis with Polymorphus, and Parafilicollis with Webster’s, 1948 Falsifilicollis, which he amended and elevated to the generic status and included F. altmani, F. kenti, F. major, F. sphaerocephalus, and F. texensis. Yamaguti’s [83] diagnosis of Falsifilicollis was essentially the same as that of Petrochenko’s [52] of Parafilicollis with few exceptions. Falsifilicollis and Parafilicollis clearly occupy the same concept for which Meyer [47] originally erected Profilicollis. Golvan [24] accepted Profilicollis with four species (P. botulus, P. arcticus, P. altmani, and P. texensis; a junior synonym of P. altmani) and recognized 26 species in Polymorphus. These included P. kenti and P. major, which, however, perfectly fit Golvan’s [24] own concept of Profilicollis; Polymorphus remained without an assigned subgenus. Hoklova [33] recognized Profilicollis as an independent genus with seven species, in Filicollidae; she had declared Falsificollis and Parafilicollis as synonyms. She did not, however, include P. arcticus (Van Cleave, 1920) Van Cleave, 1937 in her concept of Profilicollis but placed it in Polymorphus (Polymorphus) [32, 33, 73]. Her most complete diagnosis of Polymorphus was more encompassing of the wide diversity within the genus but still suffered from some of the restrictive characterizations stated by earlier observers. Nickol et al. [50] reached Hoklova’s [33] conclusions and “reintroduced” Profilicollis as a full-fledged genus after it had been recognized as a subgenus by many observers, including Schmidt and Kuntz [65] and Amin [2].

We update below the Amin [2] list of valid species of Profilicollis to show their synonymies and history.

GENUS Profilicollis Meyer, 1931 [= Falsifilicollis Webster, 1948; Parafilicollis Petrochenko 1956]

  1. Profilicollis altmani (Perry, 1942) Van Cleave, 1947 [= Filicollis altmani Perry, 1942; Parafilicollis altmani (Perry, 1942) Petrochenko, 1956; Polymorphus bullocki Mateo, Cordova, Guzman, 1982; Profilicollis kenti (Van Cleave, 1947) Hoklova, 1974; Polymorphus kenti Van Cleave, 1947; Parafilicollis kenti (Van Cleave, 1947) Petrochenko, 1956; Falsificollis kenti (Van Cleave, 1947) Yamaguti, 1963 fide Nickol et al. 2002; Filicollis sphaerocephalus sensu Harrington and Pillbury, 1938 fide Tantaleán et al. 2005; Profilicollis texensis (Webster, 1948) Hoklova, 1974; Polymorphus (Falsificollis) texensis (Webster, 1948) Yamaguti, 1963 fide Nickol et al. [50]].

  2. Profilicollis antarcticus Zdzitowiecki, 1985.

  3. Profilicollis arcticus (Van Cleave, 1920) Meyer, 1932 [= Filicollis arcticus Van Cleave, 1920].

  4. Profilicollis botulus (Van Cleave, 1916) Witenberg, 1932 (type species) [= Filicollis botulus Van Cleave, 1916].

  5. Profilicollis chasmagnathi (Holcman-Spector, Mane-Garzon, Dei-Cas, 1978) [1] [= Falsifilicollis chasmagnathi Holcman-Spector, Mane-Garzon, Dei-Cas, 1978].

  6. Profilicollis formosus (Schmidt and Kuntz, 1967) [33] [= Polymorphus formosus Schmidt and Kuntz, 1967].

  7. Profilicollis major (Lundström, 1942) [33] [= Polymorphus major Lundström, 1942; Parafilicollis major (Lundström, 1942) [74]; Filicollis major Lundström, 1942; Falsificollis major (Lundström, 1942) [83].

  8. Profilicollis novaezelandensis Brockerhoff, Smales, 2002.

  9. Profilicollis rancoensis n. sp.

  10. Profilicollis sphaerocephalus (Bremser in Rudolphi, 1819) [76] [= Echinorhynchus sphaerocephalus Bremser in Rudolphi, 1819; Filicollis sphaerocephalus (Bremser in Rudolphi, 1819) [71]; Parafilicollis sphaerocephalus (Bremser in Rudolphi, 1819) [52]; Falsifilicollis sphaerocephalus (Bremser in Rudolphi, 1819) [83].

Identification key

Our key, (above) is the only one designed from cystacanths even though some characters such as proboscis armature apply equally well to adults. All other published keys were created for adults and included such characters as size and shape of eggs, size of testes, and number of cement glands. Our key below is the latest and most complete.

Hosts

The 87 extant species and subspecies of aeglid freshwater crabs restricted to South America are the only taxon within the infraorder Anomura (freshwater hermit crab Clibanarius fonticola) that has its life cycle entirely restricted to freshwater environments [61]. Aegla abtao is found in 410 km2 and 199 m deep Ranco Lake, the largest lake in the Los Rios Region, north Patagonia, Chile. The lake, once connected to the Pacific Ocean as an embayment, now lies between the Chilean Central Valley and the Andes, bordered by moraines around its western shore. The primary inflows of Ranco Lake are Calcurrupe, Caunahue, and Nilahue and the out flow is the Bueno River [14]. The crab is an omnivorous species in adults and there are records of its predatory behavior [38]. Irrespective of diet and food habits, crabs appear to become infected with acanthocephalan cystacanths by ingesting eggs that adhere to food sources (e.g., plant debris), which are then deposited in the water body by definitive bird hosts [45, 55, 56].

Micropores

The micropores of P. rancoensis n. sp. are associated with different proboscis and trunk regions corresponding to differential absorption of nutrients. We have reported micropores in a large number of acanthocephalan species [29] and in a few more since, and demonstrated tunneling from the cuticular surface into the internal crypts by TEM in Corynosoma strumosum (Rudolphi, 1802) Lühe, 1904 from the Caspian seal Pusa caspica (Gmelin) in the Caspian Sea (Figs. 19, 20 of Amin et al. [5]) and in Neoechinorhynchus personatus Tkach, Sarabeev, Shvetsova, 2014 from Mugil cephalus Linn. in Tunisia (Figs. 26, 29, 30 in Amin et al. [8]). Amin et al. [4] provided a summary of the structural-functional relationships of the micropores in various acanthocephalan species. Wright and Lumsden [82] and Byram and Fisher [15] reported that the peripheral canals of the micropores are continuous, with canalicular crypts constituting a huge increase in external surface area, implicated in nutrient uptake. Whitfield [81] estimated a 44-fold increase at a surface density of 15 invaginations per 1 μm2 of M. moniliformis tegumental surface.

Energy Dispersive X-Ray analysis (EDXA)

Elemental composition of hooks and spines was evaluated via EDX spectra, with sulfur (S), calcium (Ca) and phosphorus (P) being the prominent elements in various species of acanthocephalans [28, 29]. In P. rancoensis, sulfur is usually prominent at the tip and the outer edge of large hooks and calcium and phosphorus are major constituents in the base and middle of the hooks, where tension and strength are paramount for hook function. Sulfur is part of the prominent outer layer of most acanthocephalan hooks and is a major contributor to the hardening process of hooks. While EDX spectra and weight-percentages are both regarded as highly qualitative, EDX analysis appears to be indicative of specific species. For example, sodium, a rarely prominent element, is evident in EDX spectra of whole hooks of Microsentis wardae (Martin and Multani, 1966) and of Pallisentis nandai Sarkar, 1953 [10], as well as in egg shells of Neoechinorhynchus qatarensis Amin, Saoud, Alkuwari, 2002 [28]. Similarly, Moniliformis cryptosaudi from Iraq is morphologically identical to Moniliformis saudi Amin, Heckmann, Mohammed, Evans, 2016 from Saudi Arabia, and it was erected based primarily on its distinctly different EDXA [7], as a cryptic species.

The elemental analysis of cystacanths of only 3 other species of acanthocephalans have previously been studied. In cystacanths of Neoandracantha peruensis (Amin, Heckmann, 2017) from the ghost crab in Peru, hooks near the middle of the proboscis showed comparatively very low sulfur (2.50%, 6.20%, 0.32% weight-percent as reported by TEAM) at the tip, middle, and base of the hook edge, and the corresponding levels of phosphorus and calcium were considerably higher (reaching 14.79% and 36.92%) at the base of the hook edge [3, 28]. This pattern is nearly opposite to that of P. rancoensis cystacanths and emphasizes the characteristic EDXA diagnosis for each species. Anterior and middle hooks of cystacanths of Sphaerirostris picae (Rudolphi, 1819) Golvan, 1956 from lizards and hedgehogs in Ukraine had moderate levels of phosphorus and sulfur (4.41–6.03%) and slightly higher levels of calcium (9.61–10.83%); again, showing variation in EDXA that is species specific [10]. Cystacanths of Moniliformis kalahariensis [Meyer, 1931], from Blatella germanica Linn. in India, had characteristically very low levels of phosphorus, sulfur, and calcium in anterior, middle, and posterior hooks alike [10]. A detailed discussion of the biological significance of EDXA as a diagnostic tool is exemplified by the observation that populations of an acanthocephalan species will consistently have similar EDX analysis irrespective of host species or geography, even though comparative morphometrics of different populations of the same species usually vary with host species and geography, as presented in Amin et al. [9, 10].

Hosts and geography

Our molecular results corroborate that the cystacanth found in the freshwater crab Aegla abtao belongs to a new acanthocephalan species. The phylogenetic analysis showed that this new species falls into the genus Profilicollis, and the molecular analysis showed that P. rancoensis n. sp. exhibits a low level of genetic variation where recovered haplotypes differ on average 0.3% and 2% (COI and 18S, respectively). At the same time, P. rancoensis n. sp. is highly divergent from other species of Profilicollis included in the analysis (COI = 14% and 18S = 21% from P. botulus). These results are similar to those for the genus Profilicollis previously reported [9, 18, 54, 56], showing the influence of environmental variables on a specific host-parasite relationships, as well as that mobility of hosts is determinant in the strict parasite-intermediate host links [25, 41, 54, 56]. Regarding host habitat, environmental conditions may limit acanthocephalan species to particular crab hosts that are exposed to different habitats [41, 56, 67]. For example, P. altmani parasitized Emerita spp., while P. chasmagnathi inhabits varanid host, which belong to marine and estuarine environments, respectively [41, 54, 56]. In this study, A. abtao lives in freshwater environmental conditions, becoming the first Profilicollis species that lives in this host species and in these systems in South America. On the other hand, birds belonging to the order Charadriiformes have been reported as definitive hosts for parasites of this genus. Unfortunately, in our fieldwork, we did not find adult worms or the definitive host, although species such as kelp gulls Larus dominicanus, brown-hooded gulls Chroicocephalus maculipennis, and cormorants Phalacrocorax olivaceus (Order Suliformes) are typical inhabitants of these habitats, and crabs are key prey in their diet [41, 56]. Therefore, close parasite-intermediate host relationships are often ascribed to specific environmental conditions that vary among host species [41, 56, 67].

In general, there are several taxa in Polymorphidae that do not form monophyletic groups, such the genus Profilicollis [6, 9, 41, 54, 56]. These results indicate that the systematic positions of the Acanthocephala are still unstable, and the composition of the genera need to be revised. Although most groups have been proposed on the basis of morphological and morphometric characters, they need to be supported by an explicit phylogenetic approach [9, 17, 20, 52].

Author contributions

Amin: Identified the parasitic acanthocephalan, wrote the manuscript including the hosts and habitat component, and the key, and created the line drawings. Rodríguez: Field work, samples processing, collected the research material, co-wrote the molecular section, and edited the manuscript.Farrer & Rivera: Created the SEM images and co-wrote the EDXA section. Fierro & Garcés: Field work, samples processing. D’Elía: Generated and analyzed molecular data, co-wrote the molecular section. All authors read the final manuscript and accepted submission of this work for publication.

Acknowledgments

We would like to thank Alex González for his assistance with the laboratory work in Sistemática Lab. From Universidad Austral of Chile, and Erwin Lienlaf for his assistance in the fieldwork. We also thank INVASAL (ANID-Millennium Science Initiative–NCN16_034) and VIDCA–Universidad Austral de Chile and ANID_SIA #85220111. Additionally, we thank Electron Microscopy Facility (EMF) at Brigham Young University (BYU), Provo, Utah for providing the time, facilities, and expertise to produce the SEM images and perform the EDXA analysis. The Department of Biology at BYU has gratefully provided full support for the EMF operation. This project was supported, in part, by an institutional grant from the Parasitology Center, Inc., Scottsdale, Arizona, USA and supporting staff. CG was supported by Scholarship ANID-Subdirección de Capital Humano/Magíster Nacional/2021 – 22210061.

Supplementary material

thumbnail Figure S1

(A) Image of individual of the freshwater crab Aegla abtao. (B) = Field work and collection of A. abtao in Ranco Lake, northern Patagonia, Chile.

References

  1. Amin OM. 1992. Review of the genus Polymorphus Lühe, 1911 (Acanthocephala: Polymorphidae), with the synonymization of Hexaglandula Petrochenko, 1950 and Subcorynosoma Hoklova, 1967, and a key to the species. Qatar University of Science Journal, 12, 115–123. [Google Scholar]
  2. Amin OM. 2013. Classification of the Acanthocephala. Folia Parasitologica, 60, 273–305. [CrossRef] [PubMed] [Google Scholar]
  3. Amin OM, Heckmann RA. 2017. Neoandracantha peruensis n. gen. n. sp. (Acanthocephala, Polymorphidae) described from cystacanths infecting the ghost crab Ocypode gaudichaudii on the Peruvian coast. Parasite, 24, 40. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  4. Amin OM, Heckmann RA, Radwan NA, Mantuano JS, Alcivar MAZ. 2009. Redescription of Rhadinorhynchus ornatus (Acanthocephala: Rhadinorhynchidae) from skipjack tuna, Katsuwonus pelamis, collected in the Pacific Ocean off South America, with special reference to new morphological features. Journal of Parasitology, 95, 656–664. [CrossRef] [PubMed] [Google Scholar]
  5. Amin OM, Heckmann RA, Halajian A, El-Naggar AM. 2011. The morphology of an unique population of Corynosoma strumosum (Acanthocephala, Polymorphidae) from the Caspian seal, Pusa caspica, in the land-locked Caspian Sea using SEM, with special notes on histopathology. Acta Parasitologica, 56, 438–445. [Google Scholar]
  6. Amin OM, Rodríguez SM, Heckmann RA. 2019a. Morphological updates and molecular description of Heterosentis holospinus Amin, Heckmann, Ha, 2011 (Acanthocephala: Arhythmacanthidae) in the Pacific Ocean off Vietnam. Parasite, 26, 73. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  7. Amin OM, Heckmann RA, Sharifdini M, Albayati NY. 2019b. Moniliformis cryptosaudi n. sp. (Acanthocephala: Moniliformidae) from the Long-eared Hedgehog Hemiechinus auritus (Gmelin) (Erinaceidae) in Iraq; A Case of Incipient Cryptic Speciation Related to M. saudi in Saudi Arabia. Acta Parasitologica, 64, 195–204. [CrossRef] [PubMed] [Google Scholar]
  8. Amin OM, Heckmann RA, Sharifdini M, Rubtsova N, Chine HJ. 2020a. On the Neoechinorhynchus agilis (Acanthocephala: Neoechinorhynchidae) complex, with the description of Neoechinorhynchus ponticus n. sp. from Chelon auratus Risso in the Black Sea. Parasite, 27, 48. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  9. Amin OM, Rodríguez SM, Rubtsova N, Heckmann RA, Peña C, Castro T, Rivera F, D’Elía G. 2022a. A comparative assessment of the morphology of Profilicollis altmani (Acanthocephala: Polymorphidae) from crustaceans and shore birds in Peru, with special notes on hook elemental analysis (EDXA), SEM imaging, histopathology, and molecular profile. Parasite, 29, 9. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  10. Amin OM, Lisitsyna O, Heckmann RA. 2022b. Evaluating energy dispersive X-ray analysis (EDXA) as a diagnostic tool in acanthocephalan taxonomy as evidenced in Palaeacanthocephala and Archiacanthocephala. Systematic Parasitology, 100, 43–57. [Google Scholar]
  11. Anisimova M, Gil M, Dufayard J-F, Dessimoz C, Gascuel O. 2011. Survey of branch support methods demonstrates accuracy, power, and robustness of fast likelihood-based approximation schemes. Systematic Biology, 60, 685–699. [CrossRef] [PubMed] [Google Scholar]
  12. Anglade T, Randhawa HS. 2018. Gaining insights into de ecological role of New Zealand sole (Peltorhamphus novaezeelandiae) through parasites. Journal of Helminthology, 92, 187–196. [CrossRef] [PubMed] [Google Scholar]
  13. Brockerhoff AM, Smales LR. 2002. Profilicollis novaezelandensis n. sp. (Polymorphidae) and two other acanthocephalan parasites from shore birds (Haematopodidae and Scolopacidae) in New Zealand, with records of two species in intertidal crabs (Decapoda: Grapsidae and Ocypodidae). Systematic Parasitology, 52, 55–65. [CrossRef] [PubMed] [Google Scholar]
  14. Burford T. 2005. Chile: The Bradt Travel Guide. Bradt Travel Guides, Chile. ISBN 9781841620763. [Google Scholar]
  15. Byram JE, Fisher Jr. FM. 1973. The absorptive surface of Moniliformis dubius (Acanthocephala). 1. Fine structure. Tissue and Cell, 5, 553–579. [CrossRef] [Google Scholar]
  16. Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R. 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Molecular Marine Biology and Biotechnology, 3, 294–299. [PubMed] [Google Scholar]
  17. García-Varela M, Nadler SA. 2006. Phylogenetic relationships among Syndermata inferred from nuclear and mitochondrial gene sequences. Molecular Phylogenetics and Evolution, 40, 61–72. [CrossRef] [PubMed] [Google Scholar]
  18. García-Varela M, Pérez-Ponce de León G. 2008. Validating the systematic position of Profilicollis Meyer, 1931 and Hexaglandula Petrochenko, 1950 (Acanthocephala: Polymorphidae) using cytochrome c oxidase (COX 1). Journal of Parasitology, 94, 212–217. [CrossRef] [PubMed] [Google Scholar]
  19. García-Varela M, Pérez-Ponce de León G, de la Torre P, Cummings MP, Sarma SSS, Laclette JP. 2000. Phylogenetic relationships of Acanthocephala based on analysis of 18S ribosomal RNA gene sequences. Journal of Molecular and Evolution, 50, 532–540. [CrossRef] [PubMed] [Google Scholar]
  20. García-Varela M, Pérez-Ponce de León G, Aznar FJ, Nadler SA. 2009. Systematic position of Pseudocorynosoma and Andracantha (Acanthocephala, Polymorphidae) based on nuclear and mitochondrial gene sequences. Journal of Parasitology, 95, 178–185. [CrossRef] [PubMed] [Google Scholar]
  21. García-Varela M, Pérez-Ponce de León G, Aznar FJ, Nadler SA. 2011. Erection of Ibirhynchus gen. nov. (Acanthocephala: Polymorphidae), based on molecular and morphological data. Journal of Parasitology, 97, 97–105. [CrossRef] [PubMed] [Google Scholar]
  22. García-Varela M, Pérez-Ponce de León G, Aznar FJ, Nadler SA. 2013. Phylogenetic relationship among genera Polymorphidae (Acanthocephala), inferred from nuclear and mitochondrial gene sequences. Molecular Phylogenetics and Evolution, 68, 176–184. [CrossRef] [PubMed] [Google Scholar]
  23. Goedknegt MA, Havermans J, Waser AM, Luttikhuizen PC, Velilla E, Camphuysen KCJ, van der Meer J, Thieltges DW. 2017. Cross-species comparison of parasite richness, prevalence, and intensity in a native compared to two invasive brachyuran crabs. Aquatic Invasions, 12, 201–212. [CrossRef] [Google Scholar]
  24. Golvan YJ. 1960–1961. Le phylum des Acanthocephala. Troisieme note. La classe des Palaeacanthocephala (Meyer, 1931). Annales de Parasitologie Humaine et Comparée, 35, 138–165, 350–386, 713–723. [CrossRef] [EDP Sciences] [Google Scholar]
  25. Goulding TC, Cohen CS. 2014. Phylogeography of a marine acanthocephalan: lack of cryptic diversity in a cosmopolitan parasite of mole crabs. Journal of Biogeography, 41, 965–976. [CrossRef] [Google Scholar]
  26. Gregori M, Aznar FJ, Abollo E, Roura Á, González ÁF, Pascual S. 2012. Nycthiphanes couchii as intermediate host for the acanthocephalan Bolbosoma balaenae in temperate waters of the NE Atlantic. Diseases of Aquatic Organisms, 99, 37–47. [CrossRef] [PubMed] [Google Scholar]
  27. Hay E, Jorge F, Poulin R. 2018. The comparative phylogeography of shore crabs and their acanthocephalan parasites. Marine Biology, 69, 165–169. [Google Scholar]
  28. Heckmann RA, Amin OM, Standing MD. 2007. Chemical analysis of metals in Acanthocephalans using energy dispersive X-ray analysis (EDXA, XEDS) in conjunction with a scanning electron microscope (SEM). Comparative Parasitology, 74, 388–391. [Google Scholar]
  29. Heckmann RA, Amin OM, El Naggar AM. 2013. Micropores of Acanthocephala, a scanning electron microscopy study. Scientia Parasitologica, 14, 105–113. [Google Scholar]
  30. Hernández-Orts JS, Hernández-Mena DI, Pantoja C, Kuchta R, García NA, Crespo EA, Loizaga R. 2021. A visitor of tropical waters: first record of a Clymene dolphin (Stenella clymene) off the Patagonian coast of Argentina, with comments on diet and metazoan parasites. Frontiers in Marine Science, 8, 658975. [CrossRef] [Google Scholar]
  31. Hoang DT, Chernomor O, von Haeseler A, Minh BQ, Vinh LS. 2018. UFBoot2: improving the ultrafast bootstrap approximation. Molecular Biology and Evolution, 35, 518–522. [CrossRef] [PubMed] [Google Scholar]
  32. Hoklova IG. 1971a. Revision of the genus Polymorphus Lühe, 1911 (Acanthocephala: Polymorphidae). Trudi Gelmintol. Lab. Akad. Nauk SSSR, 21, 120–126. (In Russian) [Google Scholar]
  33. Hoklova IG. 1974. Revision of the genus Profilicollis Meyer, 1931 = syn. Falsificollis Webster, 1948 = Parafilicollis Petrotschenko, 1956 Acanthocephala: Filicollidae). Helminthologia, 15, 803–811. [Google Scholar]
  34. Kaito S, Sasaki M, Goto K, Matsusue R, Koyama H, Nakao M, Hasegawa H. 2019. A case of small obstruction due to infection with Bolbosoma sp. (Acanthocephala: Polymorphidae). Parasitology International, 68, 14–16. [CrossRef] [PubMed] [Google Scholar]
  35. Kalyaanamoorthy S, Minh BQ, Wong TKF, von Haeseler A, Jermiin LS. 2017. ModelFinder: fast model selection for accurate phylogenetic estimates. Nature Methods, 14, 587–589. [CrossRef] [PubMed] [Google Scholar]
  36. Kang J, Liang L. 2018. First report on cystacanths of Sphaerirostris lanceoides (Petrochenko, 1949) (Acanthocephala: Centrorhynchidae) from the Asiatic toad Bufo gargarizans Cantor (Amphibia: Anura) in China. Systematic Parasitology, 95, 447–454. [CrossRef] [PubMed] [Google Scholar]
  37. Katoh K, Standley DM. 2013. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Molecular Biology and Evolution, 30, 772–780. [CrossRef] [Google Scholar]
  38. Lara GP, Moreno CA. 1995. Efectos de la depredación de Aegla abtao (Crustacea, Aeglidae) sobre la distribución espacial y abundancia de Diplodon chilensis (Bivalvia, Hyriidae) en el lago Panguipulli, Chile. Revista Chilena de Historia Natural, 68, 123–129. [Google Scholar]
  39. Levy E, Rossin MA, Braicovich PE, Timi JT. 2020. Profilicollis chasmagnathi (Acanthocephala) parasitizing freshwater fishes: paratenicity and an exception to the phylogenetic conservatism of the genus? Parasitology Research, 119, 3957–3966. [CrossRef] [PubMed] [Google Scholar]
  40. Lisitsyna OI, Kudalei O, Spraker TR, Tkach VV, Smales LR, Kuzmina TA. 2019. Morphological and molecular evidence for synonymy of Corynosoma obtuscens Lincicome, 1943 with Corynosoma australe Johnston, 1937 (Acanthocephala: Polymorphidae). Systematic Parasitology, 96, 95–110. [CrossRef] [PubMed] [Google Scholar]
  41. Lorenti E, Rodríguez SM, Cremonte F, D’Elía G, Diaz JI. 2018. Life cycle of the parasite Profilicollis chasmagnathi (Acanthocephala) on the Patagonian coast of Argentina based on morphological and molecular data. Journal of Parasitology, 104, 479–485. [CrossRef] [PubMed] [Google Scholar]
  42. Lühe M. 1911. Acanthocephalen. Register der Acanthocephalen und parasitischen plattwurmer, geordnet nach ihren wirten. Die Susswasserfauna Deutschlands eine Exkursions fauna (Brauer). Gustav Fischer, Jena, 16, 1–116. [Google Scholar]
  43. Lundström A. 1942. Die Acanthocephalen Schwedens mit Austanhme der Fischacanthocephalen von Süswasserstandorten. C.W. Lindstrom: Lund. [Google Scholar]
  44. Malyarchuk B, Derenko M, Mikhailova E, Denisova G. 2014. Phylogenetic relationships among Neoechinorhynchus species (Acantocephala: Neoechynorhynchidae) from North-East Asia based on molecular data. Parasitology International, 63, 100–107. [CrossRef] [PubMed] [Google Scholar]
  45. Mateo E, Córdova R, Guzmán E. 1983. Polymorphus (Profilicollis) bullocki, acantocéfalo de la gaviota Larus belcheri. Contribución al conocimiento de su ciclo biológico. Boletín de Lima, 5, 67–71. [Google Scholar]
  46. Mayer KA, Dailey MD, Miller MA. 2003. Helminth parasites of the southern sea otter Enhydra lutris nereis in Central California: abundance, distribution and pathology. Diseases of Aquatic Organisms, 53, 77–88. [CrossRef] [PubMed] [Google Scholar]
  47. Meyer A. 1931. Die Acanthocephalen des arktischen Gebietes. Fauna Arctica (Roemer u. Schaudinn). Gustav Fischer, Jena, 6, 9–20. [Google Scholar]
  48. Meyer A. 1932. Acanthocephala, in Dr. H.G. Bronns Klassen und Ordnungen des Tier-Reichs, vol. 4. Akademische Verlagsgesellschaft MBH: Leipzig. p. 1–332. [Google Scholar]
  49. Near TJ, Garey JR, Nadler SA. 1998. Phylogenetic relationships of the Acanthocephala inferred from 18S ribosomal DNA sequences. Molecular Phylogenetic and Evolution, 10, 287–298. [CrossRef] [Google Scholar]
  50. Nickol BB, Heard RW, Smith NF. 2002. Acanthocephalans from crabs in the southeastern US, with the first intermediate hosts known for Arhythmorhynchus frassoni and Hexaglandula corynosoma. Journal of Parasitology, 88, 79–83. [CrossRef] [PubMed] [Google Scholar]
  51. Perrot-Minnot MJ, Cozzarolo MS, Amin OM, Barčák D, Bauer A, Filipović Marijić V, García-Varela M, Hernández-Orts JS, Le Yen TT, Nachev M, Orosová M, Rigaud T, Šariri S, Wattier R, Reyda F, Sures B. 2023. Hooking the scientific community on thorny-headed worms: interesting and exciting facts, knowledge gaps and perspectives for research directions on Acanthocephala. Parasite, 30, 23. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  52. Petrochenko VI. 1956. Acanthocephala of domestic and wild animals, vol 1. Moscow: Lzdat. Akad. Nauk SSSR. (English translation by Israel Program for Scientific Translations Ltd., 1971, 465 p.) [Google Scholar]
  53. Presswell B, Bennett JDL, Smales LR. 2020. Morphological and molecular characterisation of a new genus and species of acanthocephalan, Tenuisoma tarapungi n. g., n. sp. (Acanthocephala: Polymorphidae) infecting red-billed gulls in New Zealand, with a key to the genera of the Polymorphidae Meyer, 1931. Systematic Parasitology, 97, 25–39. [CrossRef] [PubMed] [Google Scholar]
  54. Rodríguez SM, D’Elía G. 2016. Pan-American marine coastal distribution of Profilicollis altmani based on morfometric and phylogenetic analysis of Cystacanths from the mole crab Emerita brasiliensis. Journal of Helminthology, 91, 371–375. [Google Scholar]
  55. Rodríguez SM, Valdivia N. 2017. Mesoscale spatiotemporal variability in a complex host-parasite system influenced by intermediate host body size. PeerJ, 5, e3675. [CrossRef] [PubMed] [Google Scholar]
  56. Rodríguez SM, D’Elía G, Valdivia N. 2017. The phylogeny and life cycle of two species of Profilicollis (Acanthocephala: Polymorphidae) in marine hosts off the Pacific coast of Chile. Journal of Helminthology, 91, 589–596. [CrossRef] [PubMed] [Google Scholar]
  57. Rodríguez SM, Burgos K, Gutiérrez B, Escares V, Byers JE. 2020. First report of foraging behavior on Pacific mole crabs (Emerita analoga) by dogs (Canis lupus familiaris) on the coast of Valdivia, Chile. Gayana, 84, 158–162. [CrossRef] [Google Scholar]
  58. Rodríguez SM, Amin OM, Heckmann RA, Sharifdini M, D’Elía G. 2022. Phylogeny and life cycles of the Archiacanthocephala with a note on the validity of Mediorhynchus gallinarum. Acta Parasitologica, 67, 369–379. [CrossRef] [PubMed] [Google Scholar]
  59. Rodrigues W, Hebling NJ. 1978. Estudos biológicos em Aegla perobae Hebling & Rodrigues, 1977 (Decapoda, Anomura). Revista Brasileira de Biologia, 38, 383–390. [Google Scholar]
  60. Santoro M, Palomba M, Gili C, Marcer F, Marchiori E, Mattiucci S. 2021. Molecular and morphological characterization of Bolbosoma balaenae (Acanthocephala: Polymorphidae), a neglected intestinal parasite of the fin whale Balaenoptera physalus. Parasitology, 148, 1239–1302. [Google Scholar]
  61. Santos S, Bueno SLS. 2019. Aeglidae: life history and conservation status of unique freshwater anomuran decapods. Boca Raton, USA: CRC Press. [CrossRef] [Google Scholar]
  62. Sasaki M, Katahira H, Kobayashi M, Kuramochi T, Matsubara H, Nakao M. 2019. Infection status of commercial fish with cystacanth larvae of the genus Corynosoma (Acanthocephala: Polymorphidae) in Hokkaido, Japan. International Journal of Food Microbiology, 305, 108256. [CrossRef] [PubMed] [Google Scholar]
  63. Schmitt W. 1942. The species of Aegla, endemic South American fresh-water crustaceans. Proceedings of the United States National Museum Smithsonian Institution, 91, 510–512. [Google Scholar]
  64. Schmidt GD, Kuntz RE. 1966. New and little known plagiorhynchid Acanthocephala from Taiwan and the Pescadores Islands. Journal of Parasitology, 52, 520–527. [CrossRef] [Google Scholar]
  65. Schmidt GD, Kuntz RE. 1967. Notes on the life cycle of Polymorphus (Profilicollis) formosus sp. n., and records of Arhythmorhynchus hispidus Van Cleave, 1925 (Acanthocephala) from Taiwan. Journal of Parasitology, 53, 805–809. [CrossRef] [Google Scholar]
  66. Southwell T, MacFie JWS. 1925. On a collection of Acanthocephala in the Liverpool School of Tropical Medicine. Annals of Tropical Medicine and Parasitology, 19, 141–184. [CrossRef] [Google Scholar]
  67. Steinauer ML, Nickol BB, Orti G. 2007. Cryptic speciation and patterns of phenotypic variation of a highly variable acanthocephalan parasite. Molecular Ecology, 16, 2097–4109. [Google Scholar]
  68. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. 2013. MEGA6: molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution, 30, 2725–2729. [CrossRef] [PubMed] [Google Scholar]
  69. Tantaleán M, Cárdenas J, Güere R. 2002. Profilicollis altmani (Perry, 1942) Van Cleave, 1947 (Acanthocephala) en el Perú. Con notas sobre la infección experimental de mamíferos marinos. Revista Peruana de Biología, 9, 49–51. [Google Scholar]
  70. Thapar GS. 1927. On Acanthogyrus n.g. from the intestine of the Indian fish Labeo rohita, with a note on the classification of the Acanthocephala. Journal of Helminthology, 5, 109–120. [CrossRef] [Google Scholar]
  71. Travassos L. 1926. Contribuicoes para o conhecimento da fauna helmintologica brasileira. XX. Revisao dos acantocefalos brasileiros. Parte II. Familia Echinorhynchide Hamann, 1893. Sub-fam. Centrorhynchinae Travassos, 1919. Memórias do Instituto Oswaldo Cruz, 19, 31–125. [CrossRef] [Google Scholar]
  72. Trifinoupoulus J, Nguyen LT, von Haeseler A, Minh BQ. 2016. W-IQ-TREE: a fast online phylogenetic tool for maximum likelihood analysis. Nucleic Acids Research, 44, 232–235. [Google Scholar]
  73. Van Cleave HJ. 1937. Status of the generic name Profilicollis of A. Meyer. Journal of Parasitology, 23, 563. [Google Scholar]
  74. Van Cleave HJ. 1937. Developmental stages in acanthocephalan life histories, in Papers in Helminthology published in commemoration of the 30 year Jubileum of the Scientific, Educational, and Social Activities of the Honoured Worker of Science K. J. Skrjabin, M.Ac.Sci and of the Fifteenth Anniversary of the All-Union Institute of Helminthology. Skriabin KI, Schulz RES, Gnyedina MP, Editors. All-Union Lenin Academy of Agricultural Science: Moscow. p. 739–743. [Google Scholar]
  75. Van Cleave HJ. 1939. A new species of the acanthocephalan genus Polymorphus and notes on the status of the name Profilicollis. Journal of Parasitology, 25, 129–131. [CrossRef] [Google Scholar]
  76. Van Cleave HJ. 1947. Analysis of distinctions between the acanthocephalan genera Filicollis and Polymorphus, with description of a new species of Polymorphus. Transactions of the American Microscopial Society, 66, 302–313. [CrossRef] [Google Scholar]
  77. Verweyen L, Klimpel S, Palm HW. 2011. Molecular phylogeny of the Acanthocephala (Class: Palaeacanthocephala) with a paraphyletic assemblage of the orders Polymorphida and Echinorhynchida. PLoS One, 6, e28285. [CrossRef] [PubMed] [Google Scholar]
  78. Waindok P, Lehnert K, Siebert U, Pawliczka I, Strube C. 2018. Prevalence and molecular characterization of Acanthocephala in pinnipedian of the North and Baltic Seas. International Journal for Parasitology: Parasites and Wildlife, 7, 34–43. [CrossRef] [Google Scholar]
  79. Webster JD. 1948. A new acanthocephalan from the sanderling. Transactions of the American Microscopial Society, 67, 66–69. [CrossRef] [Google Scholar]
  80. Witenberg G. 1932. Akanthocephalen-studien. II. Ueber das System der Akanthocephalen. Bolletino della Societa Zoologica Italiana (Naples), 3, 253–266. [Google Scholar]
  81. Whitfield PJ. 1979. The biology of parasitism: an introduction to the study of associating organisms. University Park Press: Baltimore, Maryland. [Google Scholar]
  82. Wright RD, Lumsden RD. 1969. Ultrastructure of the tegumentary pore-canal system of the acanthocephalan Moniliformis dubius. Journal of Parasitology, 55, 993–1003. [CrossRef] [Google Scholar]
  83. Yamaguti S. 1963. Acanthocephala, in Systema helminthum, vol. 5. Wiley Interscience: New York, 423 p. [Google Scholar]

Cite this article as: Amin OM, Rodríguez SM, Farrer S, Fierro P, Garcés C, Rivera F & D’Elía G. 2023. Review of the concept of Profilicollis Meyer, 1931 with a description of Profilicollis rancoensis n. sp. (Acanthocephala: Polymorphidae) from the freshwater crab, Aegla abtao Schmitt, 1942 (Decapoda: Anomura) in Chile, with a key to congeneric species. Parasite 30, 42.

All Tables

Table 1

Species of acanthocephalans of the Polymorphidae family, their hosts, locations and GenBank accession number of the sequences used in the phylogenetic analysis. ND = No data available. (–) = Not indicated.

In the text
Table 2

Diagnostic characteristics of Profilicollis rancoensis n. sp. from the freshwater crab Aegla abtao Schmitt, 1942 (Decapoda) in Chile compared to the 9 other species in the genus.

In the text
Table 3

Comparative morphometrics of male and female cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, Valdivia, Chile.

In the text
Table 4

Elemental weight percent of Na, Mg, P, S, K, and Ca (as reported by TEAM EDXA software) of hooks and spines of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Chile, at 6 hook sites. Weight percentages of other elements (C, N, O, Al, Fe, Au, Pd, and Ga) are omitted.

In the text

All Figures

thumbnail Figures 1–4

Cystacanths of Profilicollis rancoensis n. sp. from the freshwater crab, Aegla abtao Schmitt (Decapoda) in Chile. (1, 2) A male and female cystacanth. (3) One longitudinal row of proboscis hooks showing the variable root structures with different degrees of manubriation. (4) Profile of a trunk spine.
In the text
thumbnail Figures 5, 6.

Two late acanthellae at slightly different ages.
In the text
thumbnail Figures 7–10

SEM of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, northern Patagonia, Chile. (7) A montage of a whole specimen after excystation. (8) A lateral view of a proboscis showing the organization of hooks. Note that some hooks in the posterior circle appear to emanate from the neck. (9) A semi-apical view of a proboscis showing the smaller apical hooks and the external appearance of possible apical organ. (10) Posterior circles of hooks showing the insertion of some posterior-most hooks outside of the actual proboscis in the neck.
In the text
thumbnail Figures 11–16

SEM of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, Valdivia, Chile. (11–13) Anterior, middle and posterior hooks in profile. Note differences in size, shape, and curvature. (14–16) Anterior, middle and posterior spines; their shape is variable.
In the text
thumbnail Figures 17–22

SEM of cystacanths of Profilicollis rancoensis n. sp. from Aegla abtao in Ranco Lake, Valdivia, Chile. (17) The posterior tip of the corrugated posterior trunk (in red). The drying process may have made the corrugation more evident. Manifestations of the various size and distribution of micropores at the tip of a hook (18), proboscis base (19), trunk spine (20), and middle and posterior trunk locations (21, 22).
In the text
thumbnail Figures 23–25

Energy-Dispersive X-ray Spectra (EDXS) of anterior hooks of Profilicollis rancoensis n. sp. cystacanths from Aegla abtao in Chile, acquired from the edge of a tip hook (23), the center of the middle section of the hook (24), and the center of the base of the hook (25), where the red inlays indicate the region from where the spectra were obtained. The spectra show the raw number of counts collected, and the elements C, O, P, S, Ca peaks highlighted. See Table 4 for EDXA numerical detail. Insets point to parts of hooks used to generate spectra.
In the text
thumbnail Figure 26

Genealogical relationships of haplotypes based on partial COI (A) and 18S (B) gene sequences of specimens of the family Polymorphidae recovered in a Bayesian inference analysis. Numbers next to nodes refer to support values. Bayesian posterior probability values are shown left, and Bootstrap proportions gathered in the Maximum Likelihood analysis (COI: Ln = −11404.169, 18S: Ln = −6647.679). GenBank accession numbers are included in the terminal labels.
In the text
thumbnail Figure 27

Genealogical relationships of haplotypes based on partial COI (A) and 18S (B) gene sequences of specimens of the genus Profilicollis recovered in a Bayesian inference analysis. Numbers next to nodes refer to support values. Bayesian posterior probability values are shown left, and Bootstrap proportions gathered in the Maximum Likelihood analysis (COI: Ln = −2599.057, 18S: Ln = −3574.388). GenBank accession numbers are included in the terminal labels. Animal silhouettes with different box colors to the right of the trees indicate the intermediate host (IH) of each species. Blue = A. abtao; Green = Emerita spp.; Orange = Varunidae; Red = Portunidae.
In the text
thumbnail Figure S1

(A) Image of individual of the freshwater crab Aegla abtao. (B) = Field work and collection of A. abtao in Ranco Lake, northern Patagonia, Chile.
In the text

Current usage metrics show cumulative count of Article Views (full-text article views including HTML views, PDF and ePub downloads, according to the available data) and Abstracts Views on Vision4Press platform.

Data correspond to usage on the plateform after 2015. The current usage metrics is available 48-96 hours after online publication and is updated daily on week days.

Initial download of the metrics may take a while.