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All issues Volume 30 (2023) Parasite, 30 (2023) 23 Full HTML
Open Access
Review
Issue
Parasite
Volume 30, 2023
Article Number 23
Number of page(s) 18
DOI https://doi.org/10.1051/parasite/2023026
Published online 22 June 2023

© M-J. Perrot-Minnot 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: acanthocephalan origins and article outline

Acanthocephala is a small monophyletic group of heteroxenous parasites consisting exclusively of endoparasites of arthropods and vertebrates. Commonly referred to as thorny-headed worms, this group comprises almost 1300 species according to the most recent estimate published 10 years ago [6]. Long considered a separate phylum, Acanthocephala is recognized as a sister group to Seisonidea (Hemirotifera) within the Syndermata [106, 156, 158, 181, 182]. Since the first description of an acanthocephalan by Redi in 1684 and their naming by Rudolphi in 1802 [5], research on Acanthocephala has continuously fueled collective knowledge and questioning about parasite evolution and ecology. Their remarkable features stimulate the work of an international community of researchers in fields as diverse as integrative taxonomy and phylogeography, the evolution of parasitic life cycles, the ecology and physiology of host-parasite interactions, community ecology, environmental parasitology including ecotoxicology, and behavioral ecology. Part of this community meets every four years in a dedicated workshop. Following the 10th Acanthocephalan workshop held in August 2022, we wrote this opinion article to provide an insight about the knowledge gaps and a perspective on the future of acanthocephalan research. Key priorities in acanthocephalan parasites research have been categorized into three main areas: (i) Phylogeny, integrative taxonomy, and cytogenetics; (ii) Ecology and evolution of acanthocephalan life cycles, transmission, host exploitation strategies and host range; and (iii) Environmental issues related to global changes, including ecotoxicology. For each one, the most promising ideas and developments are presented based on selected case studies. Our purpose is not to cover all research gaps and future topics on acanthocephalans but rather highlight topics that are key in our opinion and/or that should attract interest from parasitologists with different expertise (Fig. 1).

thumbnail Figure 1

Graphical abstract of the various concepts outlined in this paper.

Box 1 A brief overview of Acanthocephalan origins

Acanthocephalans are thought to have appeared during the Cambrian along with a rich and abundant marine arthropod fauna [34], switching from the free-living/epizoic lifestyle of their syndermatan ancestors to a parasitic lifestyle in aquatic arthropods [156, 182]. They subsequently evolved complex life cycles by integrating vertebrates as definitive hosts very early in their evolution. A complex life cycle is thus considered ancestral to the entire group [34, 75], with no known reversion, as predicted by the fitness benefits brought about by complex life cycle, in particular long ones [17]. Acanthocephalans probably inherited traits relevant for an endoparasitic lifestyle from their shared epizoitic (ectoparasitic?) rotifer ancestors. Among these, epidermal crypts and an invertible body apex, linked to feeding and moving habits of Hemirotifera, may have been essential for the evolution of anchoring structures and nutrient uptake in these parasites lacking an intestinal system ([156] and ref. therein). The diversification of acanthocephalan life cycles seems characterized by both constraints, mainly driven by their association with mandibulate intermediate hosts (crustaceans, myriapods and insects), and evolutionary flexibility, especially towards their definitive vertebrate hosts [118]. Acanthocephalans have colonized freshwater and terrestrial environments from marine ones, and infect the major groups of vertebrates (fish, amphibians, reptiles, birds, and mammals) [27, 67, 118]. A thorough review of the biology of Acanthocephala can be found in the eponym seminal book of Crompton and Nickol (1985) [39], which awaits an update almost 40 years later (and more than 1700 articles published on acanthocephalans since).

Phylogeny, integrative taxonomy, and diversity

Systematic and taxonomic studies of acanthocephalans have been enhanced by the use of an expanded toolkit in the last three decades including molecular tools, as evidenced in the gradual increase in published studies integrating molecular data with morphological data since the late 1990s. However, this trend is not matched with an increase in the number of acanthocephalan taxonomists. We explain below what we think is needed to advance the scientific study of Acanthocephala systematics and evolution, by focusing on the topics of phylogenetic analyses in the area of high-throughput sequencing, and of integrative taxonomy (including non-molecular tools for advanced morphological and cytogenetic studies).

Higher level systematics and new prospects for comparative analysis within Acanthocephala

Studying the evolution of key features of Acanthocephala including morphological traits and life cycle relies on a robust phylogeny to apply comparative methods. Acanthocephala has been morphologically classified into 4 classes: Archiacanthocephala, with four orders (Aporhynchida, Gigantorhynchida, Oligacanthorhynchida and Moniliformida), Eocanthocephala with two orders (Gyracanthocephala and Neoechinorhynchida), Palaeacanthocephala with three orders (Echinorhynchida, Polymorphida and Heteramorphida) and Polyacanthocephala with one order (Polyacanthorhynchida) [5, 6, 26]. The first molecular studies based on 18S rRNA (18S) gene sequencing and a few species representing each order supported monophyly for the three major lineages that were tested, i.e., classes Archiacanthocephala, Eoacanthocephala and Palaeacanthocephala (see [61, 118]). The most recent mitogenomic studies further supported monophyly for these three major classes, but also positioned the class Polyacanthocephala within Eoacanthocephala [56, 62, 147, 181]. In addition, phylogenetic relationships remain unresolved at some order (paraphyly of Echinorhynchida), family (paraphyly of Rhadinorhynchidae) and genus levels, despite some recent efforts [56, 57, 81, 89, 114, 178, 181]. It is therefore necessary to generate larger amounts of genomic data and broader taxonomic sampling in all four major lineages to provide a robust phylogenomic framework.

In the last few decades, substantial advances in low-cost, high-throughput sequencing techniques have led to an enormous leap in phylogenomic studies of helminth parasites [82]. Currently, annotated mitogenomes of twenty-three acanthocephalan species are publicly available in which archiacanthocephalans and eoacanthocephalans remain grossly underrepresented [56]. Although mitochondrial phylogenomics offers a much higher resolution based on 12–34 genes [56, 114, 147, 181], it may also come with possible inference biases due to accelerated substitution rates and compositional heterogeneity [56, 156], and should therefore be combined with nuclear genes. The first whole genome of an acanthocephalan, Pomphorhynchus laevis, was generated by Mauer et al. (2020) [105] and was recently used to explore the phylogenomic relationships within Rotifera-Acanthocephala [106]. Draft genomes of additional species of acanthocephalans are currently being assembled [150]. High throughput phylogenomics based on Expressed Sequence Tags (EST) data [182], target enrichment of “ultraconserved elements” (UCEs) and/or Anchored Hybrid Enrichment (AHE) to design a large probe set of hundreds of loci could be implemented in the future (see Karin et al. [85] for a review of these phylogenomics methods). Using transcriptomic (EST) data, Wey-Fabrizius et al. (2014) [182] explored the evolutionary relationships within the Syndermata, in which transcriptomes of one eoacanthocephalan (Paratenuisentis ambiguus) and two palaeacanthocephalans (Echinorhynchus truttae and P. laevis) were included. Targeted sequence capture of UCEs has recently been applied to address schistosomatid diversification [46]. Together with extensive taxonomic sampling, high-throughput phylogenomics should provide a robust phylogeny to apply comparative methods in the future. This macroevolutionary view could shed light on host conservatism versus host switching over evolutionary time, across host taxonomic groups, and habitat [146, 180]. It will also allow for testing of evolutionary scenarios for several unique features of acanthocephalans in relation to their life cycle. More specifically, the diversification of anchoring structures and of egg and cystacanth morphology, and the evolution of phenotypic manipulation (see below), may relate to the intermediate and definitive host taxonomy, habitat, behavior and physiology [50, 128, 136, 181]. Such a comparative analysis relies on robust phylogenetic reconstruction and enhanced taxon sampling, as emphasized here.

Integrative taxonomy and phylogeography at species-genus levels: molecular and non-molecular tools

There are now various cases in which DNA sequence data are incorporated into integrative taxonomic and phylogeographic studies of acanthocephalans at the species-genus level. Molecular data have been instrumental in investigating taxonomic status at the genus level (for instance, [89, 114, 127, 146, 180]), and in delineating species as exemplified in several cases of cryptic species or synonyms [101, 104, 127, 141, 142, 160, 162], provided that it complements thorough morphological investigation. A great example of this is the series of studies on species of the eoacanthocephalan genus Neoechinorhynchus Stiles and Hassall, 1905 and its relatives in Middle America [7, 58, 132–134, 145]. These studies have helped increase knowledge about diversity and variability in this unwieldy genus, but also illustrated how both morphological and molecular data must be combined to reveal the presence of cryptic species [104, 131]. The paraphyly of the genus Neoechinorhynchus revealed by molecular data [130] is calling for a new taxonomic revision, a rather difficult task because no compelling morphological characters have been detected yet. More generally, revisionary actions need to eventually be taken in the light of molecular data in order to bring the classification of the Acanthocephala into the 21st Century, as done for other helminth groups such as cestodes. Moving forward, we also urge researchers to use a shared and larger set of genes to maximize comparison across studies, groups, and continents for taxonomic purposes, but also to strengthen phylogenetic resolution. For example, there is a mismatch between markers used for Neoechinorhynchus in species studies in Asia where the 18S rRNA gene and cytochrome c oxidase subunit 1 (COI) are used [103], and in the Americas where the 28S rRNA gene and Internal Transcribed Spacer (ITS) are favored (e.g. [130]). Whereas 28S and ITS provided a good phylogenetic signal that enabled reliable delimitation of species and genera from diverse families of acanthocephalans [86, 145], the mitochondrial COI gene has been used to successfully define, recognize, delineate, and better understand intraspecific variation among the acanthocephalans [3, 101, 127, 131, 145, 176]. The 18S has been proven informative for taxonomic and phylogenetic studies on acanthocephalans, often in association with 28S, ITS and/or COI (more than 80 references). More recently, COI has also been used widely in phylogeographic studies with acanthocephalans of the genera Profilicollis, Leptorhynchoides, Corynosoma, Neoechinorhynchus, Floridosentis, Southwellina, Hexaglandula, and Pomphorhynchus [59, 60, 69, 127, 131, 145, 154, 160]. Therefore, all four markers classically used (18S, 28S, ITS, COI) are useful for systematic analysis on acanthocephalans, and more so when utilized all together (see e.g. [176]).

A variety of other, non-molecular tools have been used to study acanthocephalan taxonomy. Our recommendation is to expand the use of these complementary tools to classic morphological and molecular investigations. Scanning electron microscopy (SEM) enables parasitologists to observe detailed structures such as the proboscis, hooks, spines and pores on the surface of the body of acanthocephalans [10, 146]. Another tool, complementary to the description of proboscis ultrastructure by SEM and Transmission Electron Microscopy (TEM), is energy dispersive X-ray analysis (EDXA). EDXA provides information on part of the macroelemental composition of hooks (e.g., calcium, sulfur, and phosphorus), functionally related to hook hardness vs. flexibility (see [8, 10]). These tools could inform on structural-functional relationships of unique features of acanthocephalans, mainly anchoring structures involved in attachment and micropores for nutrient uptake in the intestinal wall of their definitive hosts [10, 13].

Cytogenetics is another non-molecular tool that may be helpful in taxonomy. Such studies aim to characterize the organization of the genome and its changes. The karyotype is one of the basic cytogenetic characteristics of all species, and karyological data can reveal interspecific differences and similarities that are not obvious at the molecular or morphological level (i.e., polyploidy, aneuploidy, different chromosomal rearrangements, and presence of supernumerary B chromosomes). Yet, the cytogenetics of Acanthocephala is still an under-researched area. The studies that have been done on this group of parasites are incomplete and focus primarily on classical karyology, i.e., description of diploid chromosome number with little attention to chromosome morphology, from Giemsa-stained metaphase plates or histological sections. So far, only 12 acanthocephalan species have been karyotyped (~1% of the known species), whose chromosome numbers vary between 5 and 16 and whose sex-determining mechanisms are XX in females and X0 in males (M. Orosová, unpublished data). Due to the sparse data available for acanthocephalans, the application of karyotype data for taxonomic purpose is still very limited. However, there are some interesting examples of the usefulness of basic karyotype data in other parasite groups such as trematodes and cestodes [121, 122, 129, 161]. It is important to note that different environmental conditions may lead to intra-specific variation in karyotype, an issue that calls for careful use of this tool in taxonomic studies. For instance, B chromosomes were identified in a population of Acanthocephalus lucii from a heavily polluted water reservoir in eastern Slovakia, and a possible link between unfavorable environmental conditions and the occurrence of B chromosomes was suggested [159]. Complementary to karyotyping, the mapping of different repetitive DNA sequences on chromosomes by fluorescence in situ hybridization can provide reliable chromosomal markers that reveal some species-specific differences and could be useful in solving taxonomic and evolutionary problems also in Acanthocephala. This was demonstrated in a comparative cytogenetic study in the genus Pomphorhynchus, where the species-specific location of 18S loci distinguished the two sibling species P. laevis and Pomphorhynchus tereticollis [21]. Interestingly, telomeric repeats are not yet known within acanthocephalans. Fluorescence in situ hybridization with different telomeric probes failed to identify the chromosome end composition [22], suggesting either an as yet unknown telomeric repeat sequence or loss and replacement by other mechanisms of telomere maintenance. Genome sequencing might help in targeting this issue. In recent years, in the era of molecular phylogeny, bioinformatic analysis and genomics, it may seem that cytogenetics is an outdated technique. However, this is not the case as sequence data alone cannot provide insight into genome architecture and the role that genome architecture plays in genome plasticity [41]. Chromosomes are crucial in understanding speciation events, characterization, and often detection of cryptic species or subpopulations with specific chromosomal contents [120]. Thus, there is an urgent need to extend the limited knowledge of chromosomes in acanthocephalans. Sampling for DNA analysis is routine, now it is time to make the collection of material for karyological analysis routine as well. Finally, great care should be taken to disentangle intraspecific and interspecific variation in each of the above listed techniques in order to allow species delimitation in taxonomic studies.

Diversity database: DNA barcode, OTU identification and good practice in vouchers deposition

Central to moving forward research on Acanthocephala is the enhancement of DNA barcode data and museum collections. Twenty years after barcoding was proposed as a basis for molecular classification of operational taxonomic units (OTUs) [138, 139], an increased effort to broaden taxonomic coverage for acanthocephalans in COI barcoding is needed to catalyze taxonomic investigation. Difficulties in OTU identification in metabarcoding approaches and ecological studies occur mostly because currently, only a minority of described acanthocephalan species has been sequenced: as for today, 116 acanthocephalan species and 186 Barcode Index Numbers (BINs) are registered in the publicly available database (BOLD system, Feb 26, 2022), i.e., approximately 91% of known acanthocephalan species are not yet barcoded. In addition, the development of a comprehensive database of reference sequences, derived from accurate identifications of vouchers based on morphological grounds, is critical for OTU identification in metabarcoding approaches. It is complementary to the deposition of type, morphological and molecular voucher specimens in recognized, permanent and publicly accessible repositories, concomitantly to local repositories. This practice of “biobanking” [12] has steadily increased in recent years, with researchers depositing hologenophores (i.e., the acanthocephalan specimen from which molecular data was generated) or paragenophores (i.e., conspecific specimen vouchers collected from the same hosts, at the same time and locality as the molecular specimen) in a number of international repositories. Unfortunately, the deposition of additional molecular vouchers (i.e., whole worms, or parts thereof, preserved in 95–100% ethanol) to natural history collections that possess biobank repositories is not standard practice among acanthocephalan researchers. Therefore, we strongly advocate the use and deposition of morphological vouchers of molecular specimens and cross-referenced molecular vouchers as common practice in future acanthocephalan molecular studies, to ensure future taxonomic or ecological studies and the generation of new molecular data, including DNA barcodes.

Box 2 -omics tools, phylogeny and integrative taxonomy in acanthocephalans: past and future

  • High-throughput phylogenomics and broader taxonomic coverage should now be considered to provide robust phylogenetic relationships in Acanthocephala and a reliable phylogenetic framework for the comparative analysis of acanthocephalan features in relation to endoparasitic lifestyle and host diversity.

  • Modern -omics approaches should generate novel data on the molecular mechanisms involved in their adaptation to both invertebrate and vertebrate hosts from their syndermatan ancestors, in particular anchoring structures, nutrient acquisition strategy and immune evasion mechanisms.

  • Integrative taxonomy so far relied on COI, ITS and 28S, 18S genes. Optimal synergy of molecular and morphological data, as well as non-molecular tools, will help to clarify species delimitation and to support taxonomic diagnosis.

  • Barcoding and cytogenetics are lagging, with only 9% and 1% of species analyzed, respectively. It limits modern taxonomic investigation based on cross-check of morphological identification through barcode analysis, and the exploration of existing issues such as karyotype evolution, gene duplication/inversion, and telomere evolution.

  • Strengthening the infrastructure of this research on Acanthocephala requires an open access database and the systematic deposition of molecular and morphological vouchers in curated museums, and an increased effort in barcoding.

Ecology and evolution of acanthocephalan life cycles, exploitation and transmission strategy, and host range: filling knowledge gaps by taking multidisciplinary approaches

Acanthocephalan life cycles often include several possible intermediate and definitive hosts [27, 67, 87]. Both host availability and parasite plasticity in host exploitation shape life cycle, host range and transmission strategy of acanthocephalans at ecological and evolutionary timescales. Their dynamic changes more specifically include the frequency of occurrence of host switching or host acquisition, of paratenic host incorporation or of post-cyclic transmission (i.e., “When ingested as adults within their definitive hosts some acanthocephalans survive and parasitise the predator” [119]). Documenting these changes allows us to address the associated changes in virulence towards intermediate and definitive hosts, in parasite development and reproduction, and in host manipulation. It thereby allows for testing of general predictions about the costs and benefits in evolving complex life cycles [17]. We underscore here three topics with particularly notable knowledge gaps in the life cycle and host exploitation strategy of acanthocephalans (i) the incomplete resolution of life cycles, mainly due to a lack of records of intermediate hosts; (ii) the fragmentary understanding of host manipulation by acanthocephalans as part of their transmission and host exploitation strategies (physiology, immunity), based on a small number of species and only occasional interest in the underlying mechanisms; and (iii) the understudied factors contributing to plasticity in host range and specialization, such as ecological opportunities brought about by biological invasion and changes in trophic network.

Life cycle of acanthocephalans: fragmentary knowledge

Knowledge of acanthocephalan life cycles has historically been growing thanks to parasitological surveys of macroparasites (helminths) on one or few focal definitive host species, mainly fish, waterbirds, amphibians, and reptiles. Their ecology has been thoroughly reviewed by Kennedy, 2006 [87]. Yet, as for most parasitic groups, this knowledge is still fragmentary. In fact, despite an increasing number of species descriptions in various clades of acanthocephalans, in most cases, intermediate hosts are often unknown or only inferred from knowledge of closely related parasite species (e.g. [9, 37, 68, 157]), especially for marine acanthocephalans [27]. Elucidating life cycles is difficult because larval and juvenile stages of acanthocephalans (acanthella, cystacanth) are hardly detectable and/or difficult to identify using morphology, and because of low prevalence of infection in intermediate hosts. This knowledge gap is unfortunate for trophic-network ecologists in particular [20], for whom understanding each species-species interaction is critical [113]. Since acanthocephalan life cycles involve trophic transmission and often include several intermediate/paratenic hosts, any gap in knowledge of host-parasite relationship will indeed lead to a misunderstanding of trophic networks. For example, the well-studied P. laevis was seen as a parasite without paratenic hosts [87] until the discovery of high prevalence of cystacanths in the body cavity of minnows (Phoxinus phoxinus) and gudgeons (Gobio gobio) [111]. Since these extra-intestinal cystacanths are then infectious for chubs (Squalius cephalus; Sures, pers. comm.), a well-established definitive host, minnows (and presumably gudgeons) are paratenic hosts in the P. laevis life cycle [111], revealing new nodes in the trophic networks in European rivers. Other studies have been dedicated to finding new host species by using morphological techniques (e.g., [140]), but such studies are time-consuming and sometimes not fully reliable.

Molecular techniques can reduce search times and improve the accuracy of finding intermediate or paratenic hosts. For instance, DNA barcoding can be used to link larval stages of acanthocephalans in intermediate hosts to adults in definitive hosts, and to confirm or rule out the role of target species as intermediate hosts [3, 90]. The detection of acanthocephalans in various habitats or hosts could also rely on environmental DNA (eDNA) metabarcoding. This latter technique opens practical, fast and sensitive avenues to characterize parasite diversity from specific hosts or in a locality by screening eDNA samples, i.e., from soil, sediment, filtered water, feces, tissue, etc. [14]. It has already generated novel information for biodiversity assessment of helminth parasites [45, 148] and new host-parasite associations. Acanthocephalan-like OTUs have already been detected in metabarcoding screening of gut and fecal samples from wild hosts. For example, de Vos et al. [179] detected acanthocephalans (most probably belonging to Bolbosoma) in fecal samples of blue whales from the Northern Indian Ocean using a dietary DNA metabarcoding approach. Later, Elsaied et al. [47] recorded a Neoechinorhynchus-like OTU in the gut of Nile tilapia (Oreochromis niloticus) from Lake Nasser, Egypt. Recently, Verkuil et al. [177] detected unidentified acanthocephalans in fecal samples of Pied Flycatcher (Ficedula hypoleuca) from the Netherlands. Metabarcoding could therefore be used as a monitoring tool of acanthocephalans in a wide range of habitats, to detect new acanthocephalan–host associations, while reducing the need for complex host necropsies for monitoring purposes and for well trained taxonomists. However, it calls for an increased effort to generate DNA barcode data and identification in acanthocephalans, because currently only a minority of described acanthocephalan species has been sequenced (see above).

Obtaining new data on the intermediate hosts of acanthocephalans (more generally on their life cycle) is one thing, but sharing this knowledge in an open way is another. There is a need for an open database including information on host range (definitive, intermediate or paratenic), but also morphological and molecular information (see above), for several reasons. First, it would make it easier to share this knowledge in the community of researchers on acanthocephalans. Second, at an applied level, parasites can have a significant impact on commercial aquaculture, by lowering growth, reproduction and survival rates of infected hosts [155]. Therefore, any advance in understanding the life cycle of parasites can help control them. Third, this kind of information is important for trophic-network ecologists. Such a database should be part of or connected to the diversity database (see above) to collate information on DNA barcodes (BINs), OTUs and morphological characteristics, with ecological features including host range (definitive, intermediate or paratenic) and habitat. The data already available in books (e.g., [87]), review papers [27, 67], or in existing databases [16] should be transferred to an open online database and updated. This should also facilitate the identification, correction and communication of acanthocephalan species misidentifications as exemplified in the case of P. tereticollis and P. laevis confusion (see also Reier et al., 2020 [142] for another example). Thanks to COI and ITS identification of Pomphorhynchus sp. and the use of eDNA to detect and identify amphipod species, P. tereticollis was shown to be the species present in rivers in the United Kingdom and infecting Gammarus fossarum, while it was thought to be P. laevis and Gammarus pulex for over 70 years [18, 73, 127]. This confusion extended to definitive hosts: fish reported as definitive hosts of two “strains’’ of P. laevis in the UK [76, 77, 88] were actually infected with P. tereticollis and were in fact species known to be the preferred hosts of P. tereticollis in continental Europe [11, 124]. The value of a database to centralize important updates on host range and/or acanthocephalan identifications (or misidentifications) is also exemplified for freshwater fish in North America. Since the major treatise by Hoffman, the 2nd edition of which was published in 1999 [78], there have been many changes to species identifications and subsequent works on genera such as Neoechinorhynchus, as mentioned above, but there has been no centralized document or resource to publish these changes [152], which should be done in the future.

Addressing flexibility in life cycle and host specificity in the context of global changes

Flexibility in life cycle and the associated changes in host range can be addressed using cases of species introductions and climate change, two major causes of species range expansion. Alien host and acanthocephalan co-introductions can lead to local host capture by alien acanthocephalan species (spillover effect) or introduced host capture by local acanthocephalans and resulting spillback effect on the local host. A co-introduction process occurs when a host-parasite pair is introduced into a new geographic area. This phenomenon is not less likely to be successful in heteroxenous parasites compared to monoxenous ones, despite the constraint of using at least two obligate hosts [17, 102]. Acanthocephala is a potentially useful model group: many species are aquatic (marine or continental) and use crustaceans as intermediate hosts, two features associated with a high rate of biological invasions [63, 74, 94]. The case of co-introduction in the Danube and Rhine River systems of a Pomphorhynchus species with the Ponto-Caspian round goby Neogobius melanostomus, is an excellent case-study [40, 48, 53, 79, 141]. The invasive goby acts as a paratenic host in the invaded area where local fish species (the European barbel, Barbus barbus and the chub, Squalius cephalus) act as definitive hosts. The prior introduction and establishment in the Danube and Rhine River of its amphipod intermediate host, Dikerogammarus villosus, has probably initiated and facilitated the co-introduction of P. laevis (bosniacus) [79].

A second research avenue consists in monitoring the response of acanthocephalans to climate change, using a longitudinal study design to monitor the changes in host range, parasite transmission, and basic epidemiological parameters (acanthocephalan prevalence, intensity, fecundity, etc.). Climate-induced changes in host-parasite interactions are manifold. Changes in temperature can alter acanthocephalan development times [91, 174], and acidification and dissolved oxygen concentrations could also impact parasite transmission, by altering intermediate host behavior [183]. Combined with the effects of warming and hypoxia on invertebrate immunity [31, 32, 80, 95], climate change can therefore deeply affect the acanthocephalan life cycle, epidemiology, and local host range. Addressing these issues is probably very challenging considering the complexity of individual-, population- and community-level responses to climate change [91]. This has already been advocated in a review on the consequences of Arctic warming on the diversity, circulation and transmission of helminths in Arctic coastal ecosystems [55]. Such a challenge will require cooperation among parasitologists involved in long-term and ecosystem-specific parasitological monitoring.

Host exploitation strategies: immunity and nutrient uptake

At the level of host-parasite interactions, host exploitation and transmission strategies have been the focus of intensive research these past 70 years (reviewed in [38, 39, 50, 172]), but based on a handful of acanthocephalan species. Despite numerous studies thoroughly reviewed by Crompton (1970) [38], Schmidt (1985) [149], and Taraschewski (2000) [172], host exploitation strategies by acanthocephalans still raise puzzling questions, particularly regarding immune evasion and energy uptake and allocation, in both intermediate and definitive hosts.

Circumventing or depressing host immune defense is as vital as nutrient uptake to any endoparasite. In the intermediate host, the early step of development involves the acquisition of a “capsule” or “acellular envelop” by the acanthocephalan larvae. The function and origin of this outer envelope were the subject of studies in the 1960s to 1990s (reviewed by Crompton (1970) [38], Schmidt (1985) [149] and Taraschewski (2000) [172]). Its function is probably to evade the recognition mechanism of the host, as evidenced in the lack of healthy hemocyte accumulation on developing acanthellae and on cystacanths. Such protection relies on yet unidentified properties of this outer envelope, possibly based on the acquisition and incorporation of host proteins thereby masking parasite antigens [172]. However, this protective function has not been unambiguously established yet and certainly warrants further investigation. In addition, there is still controversy on the origin of this outer envelope [43, 172], that may be resolved by considering temporal alterations in its constitution. Made initially from the connective tissue surrounding the intestine upon penetration of the acanthor into the arthropod hemocoel, it may be progressively replaced during acanthella development by membranous material of parasite origin together with residues of disintegrated hemocytes [172], a hypothesis that needs further testing. The alteration of host immune system by acanthocephalan acanthella and cystacanth also leads to immunosuppression in the intermediate host (reviewed in [50, 172]). Immunosuppression in the few host-acanthocephalan species investigated was evidenced by lower humoral immunity and cellular immunity at the cystacanth stage (prophenoloxidase system and hemocyte concentration respectively [35, 143], reviewed in [50] and [172]). The proximate mechanisms underlying these alterations have not yet been studied, and the fitness outcome has barely ever been estimated. The molecular triggers and the timing of immunosuppression during larval development in the intermediate host are not yet known, but likely involve acanthocephalan excretory–secretory products that impair or inhibit both cellular and humoral effectors of the intermediate host immune system. In terms of fitness outcome, the evolutionary significance of immunosuppression in the intermediate host is not clear [36, 50]. Indeed, while immunosuppression may benefit the parasite by facilitating immune evasion and/or by saving host energy for its own growth/survival, it may also compromise the survival of infected hosts in response to secondary infection with other pathogens [35, 36]. Lastly, the neuropsychoimmune hypothesis of manipulation, suggesting a functional link between immunosuppression of the intermediate host and behavioral manipulation [1] (see below), is still left unexplored. At the adult stage, infection with acanthocephalan elicit a well characterized inflammatory response in the intestinal tract of fish definitive hosts (reviewed in Dezfuli et al. (2016) [42]). Histopathological observations of fish intestines revealed local response at the point of parasite attachment, with mucous cell hyperplasia and hypertrophy, and the accumulation of immune cells in response to mechanical damage and sometimes a fibrous capsule of inflammation tissue around the deeply penetrating proboscis [42]. An interesting perspective would be to address the link between intestinal inflammation and the dysbiosis recently evidenced in fish infected with P. laevis [33], since the alteration of gut microbiota may subsequently affect host metabolism and intestinal immune system, and ultimately host fitness.

As for parasite development and immune defense, nutrient uptake was the focus of active research in the 1960s to 1990s, and renewed interest in this topic has been brought about by new tools, more specifically isotope studies and transcriptomic studies. Most of these studies focused on definitive hosts. Using stable isotope analysis of carbon and nitrogen as a tool to unravel trophic relationships, it emerged that adult acanthocephalans are depleted in the heavier nitrogen isotope 15N, which shows that they feed mainly on metabolites provided by the host [84, 115]. The pattern of reduced δ15N (ratio of 15N/14N) when compared to definitive host tissues can also be found in cestodes [15, 64] but is in contrast to nematodes for example [115]. Accordingly, there are clear differences with respect to the trophic interaction between different parasite taxa and their hosts. Taxa actively feeding on host tissues, such as adult nematodes and monogeneans [64], can be clearly differentiated from acanthocephalans and cestodes that behave as absorptive feeders. Transcriptomic analyses in adult acanthocephalans revealed several genes involved in energy metabolism and carotenoid uptake especially under anaerobic conditions (fermentation) [105, 151]. Importantly, definitive host response to intestinal acanthocephalans involves coordinate responses from the enteric neural, endocrine, and immune systems (see Bosi et al. (2022) [23] for a review in fish), which relies on molecular crosstalk between host and parasite molecular effectors and modulators. Detailed transcriptomics and proteomics analyses should be encouraged in the future in a broader range of acanthocephalan species, to unravel the physiological/molecular pathways of nutrient acquisition in adult acanthocephalans and the molecular basis of inflammation, but also the impact of infection on gut microbiota [33], parasite resource acquisition and allocation in optimal and sub-optimal hosts [151], and ultimately the fitness for both the acanthocephalan and its host. This should also contribute to understand the evolution of acanthocephalan virulence towards their definitive hosts, as under some environmental conditions, acanthocephalans might actually be beneficial to their definitive hosts (cf. third section below).

Manipulation as an adaptive transmission strategy: opening new paths

Parasites with a complex life cycle are under strong selection pressure to evolve transmission strategies that bridge the gap between successive obligate hosts, one of which is intermediate host manipulation [17, 50]. Acanthocephalans are excellent model organisms for the study of intermediate host manipulation ([50] and references therein). The effects of acanthocephalans on their intermediate hosts’ behaviors, life-history, morphology, and physiology (reviewed in [39, 50, 87]) may make them more likely to increase their chances to be transmitted to their definitive host. In their meta-analysis, Fayard et al. [50] confirmed that larval stages of acanthocephalans generally induce significant changes in taxis (increased habitat overlap with predator), responses to predator stimuli, immunity (immunosuppression), reproduction (castration) and coloration (increased conspicuousness), in their intermediate hosts. Most importantly, predation by suitable definitive hosts is enhanced during the cystacanth (mature, transmissible) stage of at least five palaeacanthocephalan species, providing further evidence that these phenotypic alterations are adaptive for the parasite, which is rarely demonstrated in other groups of putatively manipulating parasites [137]. However, this knowledge is still fragmentary, mainly due to two biases or gaps in research: species coverage and proximate mechanisms.

First, there is a general bias in the study of phenotypic manipulation towards a handful of acanthocephalan species. This considerably limits the comparative analysis of manipulation across the taxa [50]. The issue at stake is the relative importance of convergence versus homology in the evolution of manipulation. Under the evolutionary convergence hypothesis, ecological constraints (mainly definitive host habitat use and diet) led to similar strategies of intermediate host manipulation. Under the homology hypothesis, similar host exploitation strategies, including manipulation, reflect shared ancestry and likely constrain host switching/acquisition. This chicken and egg conundrum was raised some time ago [29], but remains to be addressed. A broader identification of acanthocephalan intermediate hosts and accurate characterization of phenotypic alterations – or even better, of their underlying molecular mechanisms – followed by ancestral state reconstruction along a robust phylogeny, represents a colossal task. However, it could allow us to decipher which manipulation strategies are convergences and when the homologous ones originated.

Second, knowledge of the mechanisms underlying behavioral alterations in intermediate hosts is still in its infancy, and mainly restricted to candidate neuromodulatory pathways. For instance, the serotonergic pathway is likely involved in behavioral alterations induced by P. laevis in the amphipod host G. pulex [126, 170, 171]. In Gammarus roeseli individuals infected by the bird parasite Polymorphus minutus, negative geotaxis may be triggered by infection-induced hypoxia or hypoxia-signaling, as suggested by the reversal of geotaxis in uninfected individuals injected with lactate, and in the elevated brain lactate concentration in P. minutus-infected individuals compared to uninfected ones [125]. In these studies, changes in taxis, and thereby habitat preferences, were mimicked by the pharmacological manipulation of candidate (neuro-) physiological pathways. The corresponding alteration in brain chemistry (serotonin, lactate) of infected gammarids confirmed the involvement of these pathways. In addition, the striking and efficient ways for acanthocephalans to get transmitted is not only to change habitat preferences but also to alter anti-predatory behaviors. Therefore, any study on the proximate mechanisms of manipulation leading to increased acanthocephalan transmission to definitive hosts should investigate pathways modulating antipredator behavior.

These studies addressed the mechanisms underlying manipulation by acanthocephalans by targeting candidate neuromodulatory or physiological systems of the host. The prospects and limitations of such approaches have been reviewed, highlighting the complexity of these systems and the fact that they still show a host’s physiological response to manipulative parasites rather than evidencing the actual nature and targets of parasite excretion-secretion products [123]. In future studies, two complementary approaches may be taken: the proteomic analysis of excretion-secretion products of acanthocephalan larval stages, as done in trematode species [70, 72], and the quantification of gene expression in both the intermediate host brain and the larval stage of the acanthocephalan. This could enable screening for candidate proteins involved in host-parasite crosstalk, providing insights into the exact pathways involved, and identifying parasite genes that may be under natural selection for manipulation. Whole genome and transcriptome sequencing of P. laevis adults [105] provided a very useful tool to tackle this quest, which needs to be reproduced at the acanthella and cystacanth stages. In this perspective, the fact that acanthocephalans seem to follow the switcher paradigm [83] provides an original and unique opportunity in the study of host manipulation by trophically-transmitted parasites. The switcher paradigm proposes that manipulative parasites first decrease predation risk of the infected intermediate host at larval stages not yet infective to definite hosts, and then enhance predation by definitive hosts when the parasite larva is infective ([44]; reviewed in [50]). The comparison of acanthellae and cystacanths transcriptomes combined to extended phenotype characterization would give the opportunity to pinpoint potential genes - or their pleiotropic regulation linked to behavioral and immune alteration induced in the intermediate host - thereby deciphering the molecular basis of manipulation switch.

Box 3 Addressing the complexity of acanthocephalan life cycles

  • Intermediate hosts of most acanthocephalan species are unknown, which biases our understanding of trophic networks. DNA (meta-)barcoding has the potential to facilitate the detection of parasites in environmental DNA and the identification of larval stages in intermediate hosts.

  • Variable flexibility in acanthocephalan life cycle provides opportunities to test hypotheses regarding the effect of co-introductions of alien host-parasite pairs on local communities.

  • Climate changes alter host-parasite interactions, with likely cascading effects at population and community levels, especially as these parasites affect predator-prey interactions.

  • Parasitologists need an online database gathering morphological descriptions, molecular data, and ecological information such as geographic distributions and host ranges of thorny-headed worms.

  • An integrative approach to the immediate mechanisms of alterations in host energy metabolism and immune response in intermediate and definitive host is needed, relying on proteomic analysis of excretion-secretion products and transcriptomic analysis at different stages of parasite development.

  • Although excellent model organisms for the study of host behavioral manipulation, mechanisms thereof were rarely investigated and are far from elucidated, including the link between immune evasion/immunosuppression and behavioral manipulation.

Environmental aspects – interactions between acanthocephalans and pollutants

Given the complex interactions between infection with parasites and exposure to pollutants, environmental parasitology has developed as a discipline focusing on the potential role of parasites as indicators of environmental health as well as on the combined effects of parasites and pollutants on the health of the hosts [153, 166]. Acanthocephalans have been widely recognized as potential indicators of pollutant accumulation [167], and these parasites might lead to host detoxification and subsequent (beneficial) effects on host health or physiology [112]. The mechanisms involved in host detoxification processes, as well as the effects of pollutants on development and transmission of acanthocephalans, are largely unknown and should therefore be of increased research interest in the future.

Acanthocephalans as accumulation indicators of pollutants

Acanthocephalans are able to accumulate a variety of metals at higher levels than their hosts (reviewed in [116, 165, 166]). Even though metals are not detected in the host or in established free-living bioindicators such as bivalves, evaluation of metal occurrence in the environment is still facilitated by metal accumulation in acanthocephalans in their definitive hosts [167, 168]. A similar pattern was reported for polychlorinated biphenyl (PCB) accumulation in a perch-acanthocephalan system [24]. Metabolizable organic compounds such as phthalates and polycyclic aromatic hydrocarbons (PAHs) are also accumulated in the acanthocephalan at higher concentrations than in the chub host [112]. This observation might be due to an ability of the acanthocephalan Pomphorhynchus sp. to metabolize organic compounds or to take up metabolites formed in the host [112].

Moreover, the presence of contaminants in acanthocephalans is evidence not only of the presence of substances in the host’s environment, but also of their biological availability in general [166]. The substances detected in the body of acanthocephalans must be considered biologically available because they had to pass through the teguments and membranes of the parasite, since acanthocephalans lack a digestive tract. This unique substance uptake mechanism, which can be found not only in acanthocephalans but also in cestodes, makes such gutless taxa promising indicators for studies on the bioavailability of (nano)particles. If particulate substances or the elements that form them can be detected in Acanthocephala, as shown for particulate platinum-group metals [164, 184], they must pass through several biological membranes and are thus bioavailable. On the other hand, if filter-feeding organisms such as bivalves are used to study the uptake of (nano)particles, it remains unclear whether these elements are only adsorbed on the gill filaments or are present in the gut contents without having been taken up in a biological sense. Acanthocephalans could help to fill this gap and provide a better understanding of the biological availability of (nano)particles and other pollutants in ecosystems. In addition to metallic nanoparticles, this could also be a promising approach to study the uptake, accumulation, and effects of nano- and microplastic particles in the aquatic environment.

Recently, acanthocephalans have been included in physiologically-based toxicokinetic models simulating metal accumulation in fish-acanthocephalan systems [96, 97], which are based on the conceptual model developed by Sures et al. (1999) [168]. In these models, acanthocephalans are considered a sink for metals in the host intestine, thereby reducing metal availability to fish organs [167, 168]. These models provide a mechanistic explanation on the substantial variations in the partitioning of metals in fish-acanthocephalan systems [51, 52, 117, 135, 167, 173]. Accordingly, the variations in the accumulation potential in acanthocephalans among metals are not attributed to any metal-specific uptake mechanisms, but to the availability of the metals in the host intestine [97]. In particular, the accumulation of metals in acanthocephalans depends on metal-specific organotropism and cellular fate [97]. Similar models for the accumulation of organic contaminants by acanthocephalans [98] are currently lacking but would be highly desirable.

Effects of acanthocephalans on physiological responses of the host

The accumulation of pollutants in the host might lead to various physiological responses at different organization levels, from the synthesis of stress molecules and protective enzymes to a disruption of physiological homeostasis and mortality. Most research performed so far on combined effects of pollutants and acanthocephalans refers to larval acanthocephalans mainly in their intermediate crustacean hosts [71]. This is interesting in that larval stages are not able to accumulate contaminants to the same degree as adult Acanthocephala [169] and therefore do not generally lead to lower contaminant concentrations in infected intermediate hosts compared to uninfected conspecifics. However, infection-inhibited accumulation of metals in gammarids has been described (although metal accumulation in the cystacanths was much lower compared to the gammarid host), which might reduce adverse effects on the host [66]. Obviously, the mechanisms leading to altered metal accumulation in intermediate versus definitive hosts are completely different and deserve further mechanistic studies.

Also, the effects of concurrent contamination and infection on the physiology of the corresponding intermediate hosts show wide variations. For example, the acute toxicity of Cd to G. pulex was not affected by infection with the acanthocephalan P. laevis [108]. The results of several other studies showed that twice as many amphipods infected with cystacanths of P. laevis or P. minutus died compared to uninfected conspecifics when exposed to low metal concentrations [54, 107–109]. In contrast, in the study of Sures and Radszuweit (2007) [163], the mortality of P. minutus-infected G. roeseli was reduced following Pd exposure compared to uninfected individuals. Similar observations with the same parasite−host system during Cd exposure were made by Gismondi et al. (2012) [66] for the mortality of infected male gammarids. Such contradictory findings may be the result of complex combined effects of acanthocephalan infection and exposure to contaminants on host physiological homeostasis, which could be modulated primarily by infection with acanthocephalans. The mechanisms responsible for the different survival rates of infected and uninfected gammarids in the presence of concomitant exposure are completely unknown and therefore represent an exciting area of future research.

Various results on the influence of exposure to pollutants on antitoxic responses have been reported. Stress proteins (also termed “heat shock proteins”) are synthesized by organisms in response to different environmental stressors such as exposure to metals, organic pollutants, ultraviolet radiation, temperature changes, and osmolarity or hypoxia/anoxia [100]. The induction of heat shock proteins in the gammarids G. roeseli and G. fossarum in response to metal (Pd and Cd) exposure was modulated by the infection with cystacanths of the acanthocephalan P. minutus [54, 163]. Such modulation of the hosts’ repair and detoxification systems might enhance adverse effects of environmental pollution on the hosts. Similarly, the synthesis of metallothioneins, a specific biomarker for metal exposure [4], in the gammarids was enhanced by the infection with acanthocephalans, possibly via the influence on the transcription of the metallothionein gene [65]. However, such infection enhanced cell damage as evidenced by an increase in malondialdehyde concentration, a product of metal-induced lipoperoxidation, in the gammarids G. roeseli [65]. In summary, infection with larval acanthocephalans lowers the efficiency of the overall defense system in gammarids exposed to metals [65]. Another explanation for the more severe cell damage in the infected gammarids could be a high energy investment in antioxidant defense and in the parasite’s metabolism [65], which was supported by substantial decreases in lipid and glycogen contents [30, 65]. Infection with acanthocephalans might affect metal tolerance of the host also via other mechanisms. For example, increased sensitivity of gammarids infected with cystacanths to Cd was assumed to result from parasite-induced alterations of the host’s feeding rates (e.g., [25, 110]).

Compared to the combined pollutant and parasite effects on the intermediate host, information on their effects on definitive hosts is rather limited. However, interesting results have recently been published. Infection with the acanthocephalan Tenuisentis niloticus stimulated the activities of reduced glutathione and catalase in the fish host [2]. Another example is the lower oxidative stress and damage induced by emerging organic pollutants (e.g. PAHs, PBs, OCPs, PBDE, DEET) in acanthocephalan-infected chubs in comparison with that in uninfected conspecifics [112]. In addition to such effects on the molecular responses to pollutants, infection with parasites might affect the microbial community in the definitive host [33]. In particular, the European chub infected by Pomphorhynchus sp. hosted fewer bacterial communities than uninfected conspecifics [33]. These examples demonstrate the relevance of interactive effects between pollutants and adult acanthocephalans, which also seems to be a valuable area for future research activities. Moreover, the relationship between acanthocephalan-inhibited oxidative stress and the acanthocephalan-inhibited internal concentration of pollutants in the definitive host is still not addressed. An understanding of this link might provide us with a mechanistic understanding on the combined effects of pollutants and acanthocephalans on the definitive host.

Effects of environmental stressors on development and transmission of acanthocephalans

Compared with what we know about the influence of Acanthocephala on pollutant accumulation, our knowledge about the effects of environmental pollution on acanthocephalan development is largely limited and results are conflicting. Lewis et al. (2003) [99] suggested that acanthocephalans can respond quickly to environmental changes. For example, the abundance of the acanthocephalan Pandosentis aff. iracundus was negatively correlated with the concentrations of ammonia and dissolved oxygen [92]. However, the influence of dissolved oxygen concentration should be carefully interpreted because of: (i) the correlation between oxygen concentration and water temperature; and (ii) the impact of water temperature on acanthocephalan reproduction [87]. Similarly, acanthocephalans might be adversely affected by metals [93]. Other studies reported a higher abundance of acanthocephalans at sites with higher organic/nutrient loads and eutrophication levels, respectively (see e.g., [49, 175], which was assumed to be linked to a higher pollution tolerance of intermediate and definitive hosts. Accordingly, acanthocephalans might be directly affected by pollutants or indirectly via impacts on the host [28]. A correlation between the occurrence of acanthocephalan parasites and the abundance of intermediate hosts unraveled by Fanton et al. (2022) [49], is in contrast to a study that reported no significant effects of environmental stressors, such as pollution on acanthocephalans [19]. Such contradictory findings suggest a lack of thorough understanding of the effects of environmental stressors on the development and transmission of acanthocephalans. Therefore, future research needs to address the mechanisms of action for single acanthocephalan species, also considering their host specificity and the preferences of their hosts for the environmental conditions.

Box 4 Acanthocephalans at the forefront of research in environmental parasitology

  • Detailed studies on the uptake, accumulation, and potential effects of organic pollutants on acanthocephalan-host relationships should complement current knowledge.

  • Acanthocephalans could serve as indicators of particulate pollutants such as microplastics and nanoparticles.

  • Interdisciplinary and mechanistic studies elucidating processes of host detoxification by acanthocephalans as wells as their potentially beneficial effects on the physiology and health of their definitive hosts are highly desired.

  • Omics approaches should be used to obtain information on the effects of pollutants on parasites.

Conclusion

Research on the Acanthocephala illustrates the variety of topics of investigation arising from their unique features. To frame this research into a robust phylogenetic/phylogenomic framework, the taxonomic and genetic coverage of current phylogenies should be broadened. It would contribute not only to the systematic resolution of this group, but also to the study of the evolution of life cycles (transitions between marine, freshwater and terrestrial environments, type and diversity of intermediate and definitive hosts, patterns of phenotypic manipulation, etc.), as well as the evolution of genome architecture (repeated sequences, B chromosomes, karyotypes, and telomere maintenance). For most of these issues, data are still scarce and often restricted to a handful of species, as evidenced in the fragmentary knowledge about acanthocephalan cytogenetics, life cycles, or phenotypic manipulation. Yet, we hope that acanthocephalan research will continue to be an active area in the future and attract researchers from other fields. The study of acanthocephalans has long been undertaken by expert scientists with diversified skills, scientific questioning, and approaches. Most of them have interests in other parasite groups (noticeably helminths) and/or in the ecology/evolution of their hosts (mainly fish and crustaceans), thereby nurturing multidisciplinary research. However, the perennial challenges that remain are the low number of experts and the perception that this group has little impact on human and domestic animal health. These challenges have limited the extent of scientific input and the scope of this research compared to other parasite taxonomic groups.

Parasites are now perceived as important components of ecosystems, with impacts on trophic networks and nutrient flows, and have an acknowledged value as bioindicators of anthropogenic perturbations. Considering the present-day issues of biodiversity conservation and ecosystem stability/resilience in the face of anthropogenic changes, research on acanthocephalans should not lag behind because of a lack of appeal and funding. We hope to have convinced our readers of the excellent opportunities provided by acanthocephalans as a model system to many fundamental and applied aspects of the evolution and ecology of host-parasite interactions. Tools necessary to collectively improve our effectiveness and visibility include the enhancement of cross-referenced molecular voucher specimens in museum collections together with morphological vouchers, and the creation of an online open database gathering molecular barcoding, morphospecies and ecological (life-history and life cycle) data. This will contribute to promoting and coordinating research efforts in integrative taxonomy and in the ecology and evolution of the Acanthocephala.

a

Equal contribution.

Acknowledgments

The 10th Acanthocephalan Workshop took place in Dijon, France, from August 28 to August 31, 2022. We wish to thank Isaure De Buron, Halima Jmii Chine, Liang Li, Anna Marková, Bahram Sayyaf Dezfuli, and Rui-Jia Yang (participants in the workshop) for interesting and stimulating discussions leading to the identification of future perspectives in the research on Acanthocephala. We would also like to thank two anonymous reviewers for their feedback on previous versions of the manuscript. Financial support for the workshop was provided by the hosting Institution/Laboratory: Université de Bourgogne (BQR2022) and Biogéosciences, UMR 6282 CNRS uB.

References

  1. Adamo SA. 2013. Parasites: evolutions neurobiologists. Journal of Experimental Biology, 216, 3–10. [CrossRef] [PubMed] [Google Scholar]
  2. Akinsanya B, Ayanda IO, Fadipe AO, Onwuka B, Saliu JK. 2020. Heavy metals, parasitologic and oxidative stress biomarker investigations in Heterotis niloticus from Lekki Lagoon, Lagos, Nigeria. Toxicology Reports, 7, 1075–1082. [CrossRef] [PubMed] [Google Scholar]
  3. Alcántar-Escalera FJ, García-Varela M, Vázquez-Domínguez E, Pérez-Ponce de León G. 2013. Using DNA barcoding to link cystacanths and adults of the acanthocephalan Polymorphus brevis in central Mexico. Molecular Ecology Resources, 13, 1116–1124. [PubMed] [Google Scholar]
  4. Amiard J-C, Amiard-Triquet C, Barka S, Pellerin J, Rainbow P. 2006. Metallothioneins in aquatic invertebrates: their role in metal detoxification and their use as biomarkers. Aquatic Toxicology, 76, 160–202. [CrossRef] [Google Scholar]
  5. Amin OM. 1985. Classification, in Biology of the Acanthocephala. Crompton DWT, Nickol BB, Editors. University Press: Cambridge. p. 27–72. [Google Scholar]
  6. Amin OM. 2013. Classification of the Acanthocephala. Folia Parasitologica, 60, 273–305. [CrossRef] [PubMed] [Google Scholar]
  7. Amin OM. 2002. Revision of Neoechinorhynchus Stiles & Hassall, 1905 (Acanthocephala: Neoechinorhynchidae) with keys to 88 species in two subgenera. Systematic Parasitology, 53, 1–18. [CrossRef] [PubMed] [Google Scholar]
  8. Amin OM, Lisitsyna OI, Heckmann RA. 2022. 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]
  9. Amin OM, Heckmann RA, Ha NV. 2014. Acanthocephalans from fishes and amphibians in Vietnam, with descriptions of five new species. Parasite, 21, 53. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  10. Amin OM, Heckmann RA. 2022. SEM study of hooks in the Acanthocephala with emphasis on structural-functional relationships. Zoodiversity, 56(4). [Google Scholar]
  11. Andreou D, Antognazza CM, Williams CF, Bradley H, Reading AJ, Hardouin EA, Stewart JR, Sheath D, Galligar A, Johnson E, Britton JR. 2020. Vicariance in a generalist fish parasite driven by climate and salinity tolerance of hosts. Parasitology, 147, 1658–1664. [CrossRef] [PubMed] [Google Scholar]
  12. Astrin JJ, Zhou X, Misof B. 2013. The importance of biobanking in molecular taxonomy, with proposed definitions for vouchers in a molecular context. Zookeys, 30, 67–70. [CrossRef] [Google Scholar]
  13. Aznar FJ, Hernández-Orts JS, Raga JA. 2018. Morphology, performance and attachment function in Corynosoma spp. (Acanthocephala). Parasites & Vectors, 11, 633. [CrossRef] [PubMed] [Google Scholar]
  14. Bass D, Stentiford GD, Littlewood DTJ, Hartikainen H. 2015. Diverse applications of environmental DNA methods in parasitology. Trends in Parasitology, 31, 499–513. [CrossRef] [PubMed] [Google Scholar]
  15. Behrmann-Godel J, Yohannes E. 2015. Multiple isotope analyses of the piketapeworm Triaenophorus nodulosus reveal peculiarities in consumer-dietdiscrimination patterns. Journal of Helminthology, 89, 238–243. [CrossRef] [PubMed] [Google Scholar]
  16. Benesh DP, Lafferty KD, Kuris A. 2017. A life cycle database for parasitic acanthocephalans, cestodes, and nematodes. Ecology, 98, 882. [CrossRef] [PubMed] [Google Scholar]
  17. Benesh DP, Parker G, Chubb JC. 2021. Life-cycle complexity in helminths: What are the benefits? Evolution, 75, 1936–1952. [CrossRef] [PubMed] [Google Scholar]
  18. Blackman RC, Constable D, Hahn C, Sheard AM, Durkota J, Hänfling B, Handley LL. 2017. Detection of a new non-native freshwater species by DNA metabarcoding of environmental samples – first record of Gammarus fossarum in the UK. Aquatic Invasions, 12, 177–189. [CrossRef] [Google Scholar]
  19. Blanar CA, Munkittrick KR, Houlahan J, MacLatchy DL, Marcogliese DJ. 2009. Pollution and parasitism in aquatic animals: a meta-analysis of effect size. Aquatic Toxicology, 93, 18–28. [CrossRef] [Google Scholar]
  20. Blasco-Costa I, Poulin R. 2017. Parasite life-cycle studies: a plea to resurrect an old parasitological tradition. Journal of Helminthology, 91, 647–656. [CrossRef] [PubMed] [Google Scholar]
  21. Bombarová M, Marec F, Nguyen P, Špakulová M. 2007. Divergent location of ribosomal genes in chromosomes of fish thorny-headed worms, Pomphorhynchus laevis and Pomphorhynchus tereticollis (Acanthocephala). Genetica, 131, 141–149. [CrossRef] [PubMed] [Google Scholar]
  22. Bombarova M, Vitkova M, Špakulová M, Koubková B. 2009. Telomere analysis of platyhelminths and acanthocephalans by FISH and Southern hybridization. Genome, 52, 897–903. [CrossRef] [PubMed] [Google Scholar]
  23. Bosi G, Maynard BJ, Pironi F, Sayyaf Dezfuli B. 2022. Parasites and the neuroendocrine control of fish intestinal function: an ancient struggle between pathogens and host. Parasitology, 149, 1842–1861. [CrossRef] [PubMed] [Google Scholar]
  24. Brázová T, Hanzelová V, Miklisová D. 2012. Bioaccumulation of six PCB indicator congeners in a heavily polluted water reservoir in Eastern Slovakia: tissue-specific distribution in fish and their parasites. Parasitology Research, 111, 779–786. [CrossRef] [PubMed] [Google Scholar]
  25. Brown AF, Pascoe D. 1989. Parasitism and host sensitivity to cadmium: an acanthocephalan infection of the freshwater amphipod Gammarus pulex. Journal of Applied Ecology, 26, 473–487. [CrossRef] [Google Scholar]
  26. Bullock W. 1969. Morphological features as tools and as pitfalls in acanthocephalan systematics, in Problems in Systematics of Parasites, Schmidt G.D. University Park Press: Baltimore. p. 9–24. [Google Scholar]
  27. de Buron I, Golvan YJ. 1986. Les hôtes des Acanthocephales I - Les Hôtes intermédiaires. Annales de Parasitologie Humaine et Comparée, 61, 581–592. [CrossRef] [EDP Sciences] [Google Scholar]
  28. Carlson CJ, Burgio KR, Dougherty ER, Phillips AJ, Bueno VM, Clements CF, Castaldo G, Dallas TA, Cizauskas CA, Cumming GS, Doña J, Harris NC, Jovani R, Mironov S, Muellerklein OC, Proctor HC, Getz WM. 2017. Parasite biodiversity faces extinction and redistribution in a changing climate. Science Advances, 3, e1602422. [Google Scholar]
  29. Cézilly F, Thomas F, Médoc V, Perrot-Minnot MJ. 2010. Host-manipulation by parasites with complex life cycles: adaptive or not? Trends in Parasitology, 26, 311–317. [CrossRef] [PubMed] [Google Scholar]
  30. Chen H-Y, Grabner DS, Nachev M, Shih H-H, Sures B. 2015. Effects of the acanthocephalan Polymorphus minutus and the microsporidian Dictyocoela duebenum on energy reserves and stress response of cadmium exposed Gammarus fossarum. PeerJ, 3, e1353. [CrossRef] [PubMed] [Google Scholar]
  31. Cheng W, Liu CH, Hsu JP, Chen JC. 2002. Effect of hypoxia on the immune response of giant freshwater prawn Macrobrachium rosenbergii and its susceptibility to pathogen Enterococcus. Fish and Shellfish Immunology, 13, 351–365. [CrossRef] [Google Scholar]
  32. Cheng W, Wang L-U, Chen J-C. 2005. Effect of water temperature on the immune response of white shrimp Litopenaeus vannamei to Vibrio alginolyticus. Aquaculture, 250, 592–601. [CrossRef] [Google Scholar]
  33. Colin Y, Molbert N, Berthe T, Agostini S, Alliot F, Decencière B, Millot A, Goutte A, Petit F. 2022. Dysbiosis of fish gut microbiota is associated with helminths parasitism rather than exposure to PAHs at environmentally relevant concentrations. Scientific Reports, 12, 11084. [CrossRef] [PubMed] [Google Scholar]
  34. Conway Morris S, Crompton D. 1982. The origins and evolution of the Acanthocephala. Biological Reviews of the Cambridge Philosophical Society, 57, 85–115. [CrossRef] [Google Scholar]
  35. Cornet S, Franceschi N, Bauer A, Rigaud T, Moret Y. 2009. Immune depression induced by acanthocephalan parasites in their intermediate crustacean host: consequences for the risk of super-infection and links with host behavioural manipulation. International Journal for Parasitology, 39, 221–229. [CrossRef] [PubMed] [Google Scholar]
  36. Cornet S, Sorci G. 2010. Parasite virulence when the infection reduces the host immune response. Proceedings of the Royal Society of London Biological Sciences, 277, 1929–1935. [CrossRef] [PubMed] [Google Scholar]
  37. Costa Fernandes VS, Amin O, Borges JN, Santos CP. 2019. A New Species of the Acanthocephalan Genus Filisoma (Cavisomidae) from Perciform Fishes in Rio de Janeiro, Brasil. Acta Parasitologica, 64, 176–186. [CrossRef] [PubMed] [Google Scholar]
  38. Crompton DWT. 1970. An ecological approach to acanthocephalan physiology. Camb. Monogr. exp. Biol. No. 17. Cambridge University Press: London, p. 125. [Google Scholar]
  39. Crompton DWT, Nickol BB. 1985. Biology of the Acanthocephala. Cambridge University Press: Cambridge. p. 519. [Google Scholar]
  40. David GM, Staentzel C, Schlumberger O, Perrot-Minnot MJ, Beisel JN, Hardion L. 2018. A minimalist macroparasite diversity in the round goby of the Upper Rhine reduced to an exotic acanthocephalan lineage. Parasitology, 145, 1020–1026. [CrossRef] [PubMed] [Google Scholar]
  41. Deakin JE, Potter S, O’Neill R, Ruiz-Herrera A, Cioffi MB, Eldridge MD, Fukui K, Marshall Graves JA, Griffin D, Grutzner F, Kratochvíl L, Miura I, Rovatsos M, Srikulnath K, Wapstra E, Ezaz T. 2019. Chromosomics: Bridging the gap between genomes and chromosomes. Genes, 10, 627. [CrossRef] [PubMed] [Google Scholar]
  42. Dezfuli BS, Bosi G, DePasquale JA, Manera M, Giari L. 2016. Fish innate immunity against intestinal helminths. Fish and Shellfish Immunology, 50, 274–287. [CrossRef] [Google Scholar]
  43. Dezfuli BS, Simoni E, Duclos L, Rossetti E. 2008. Crustacean-acanthocephalan interaction and host cell-mediated immunity: parasite encapsulation and melanization. Folia Parasitologica, 55, 53–59. [CrossRef] [PubMed] [Google Scholar]
  44. Dianne L, Perrot-Minnot MJ, Bauer A, Gaillard M, Léger E, Rigaud T. 2011. Protection first then facilitation: A manipulative parasite modulates the vulnerability to predation of its intermediate host according to its own developmental stage. Evolution, 65, 2692–2698. [CrossRef] [Google Scholar]
  45. Douchet P, Boissier J, Mulero S, Ferté H, Doberva M, Allienne J, Toulza E, Bethune K, Rey O. 2022. Make visible the invisible: Optimized development of an environmental DNA metabarcoding tool for the characterization of trematode parasitic communities. environmental DNA, 4, 627–641. [CrossRef] [Google Scholar]
  46. Ebbs ET, Loker ES, Bu L, Locke SA, Tkach VT, Devkota R, Flores VR, Pinto HA, Brant SV. 2022. Phylogenomics and diversification of the Schistosomatidae based on Targeted Sequence Capture of Ultra-Conserved Elements. Pathogens, 11, 769. [CrossRef] [PubMed] [Google Scholar]
  47. Elsaied HE, Soliman T, Abu-Taleb HT, Goto H, Jenke-Kodam H. 2019. Phylogenetic characterization of eukaryotic and prokaryotic gut flora of Nile tilapia, Oreochromis niloticus, along niches of Lake Nasser, Egypt, based on rRNA gene high-throughput sequences. Ecological Genetics and Genomics, 11, 100037. [CrossRef] [Google Scholar]
  48. Emde S, Rueckert S, Palm HW, Klimpel S. 2012. Invasive Ponto-Caspian amphipods and fish increase the distribution range of the acanthocephalan Pomphorhynchus tereticollis in the River Rhine. PLoS One, 7, e53218. [CrossRef] [PubMed] [Google Scholar]
  49. Fanton H, Franquet E, Logez M, Cavalli L, Kaldonski N. 2022. Acanthocephalan parasites reflect ecological status of freshwater ecosystem. Science of the Total Environment, 838, 156091. [CrossRef] [Google Scholar]
  50. Fayard M, Dechaume-Moncharmont FX, Wattier R, Perrot-Minnot MJ. 2020. Magnitude and direction of parasite-induced phenotypic alterations: a meta-analysis in acanthocephalans. Biological Reviews, 95, 1233–1251. [CrossRef] [PubMed] [Google Scholar]
  51. Filipović Marijić V, Vardić Smrzlić I, Raspor B. 2013. Effect of acanthocephalan infection on metal, total protein and metallothionein concentrations in European chub from a Sava River section with low metal contamination. Science of the Total Environment, 463, 772–780. [CrossRef] [Google Scholar]
  52. Filipović Marijić V, Vardić Smrzlić I, Raspor B. 2014. Does fish reproduction and metabolic activity influence metal levels in fish intestinal parasites, acanthocephalans, during fish spawning and post-spawning period? Chemosphere, 112, 449–455. [CrossRef] [PubMed] [Google Scholar]
  53. Francová K, Ondračková M, Polačik M, Jurajda P. 2011. Parasite fauna of native and non-native populations of Neogobius melanostomus (Pallas, 1814) (Gobiidae) in the longitudinal profile of the Danube River. Journal of Applied Ichthyology, 27, 879–886. [CrossRef] [Google Scholar]
  54. Frank SN, Godehardt S, Nachev M, Trubiroha A, Kloas W, Sures B. 2013. Influence of the cestode Ligula intestinalis and the acanthocephalan Polymorphus minutus on levels of heat shock proteins (HSP70) and metallothioneins in their fish and crustacean intermediate hosts. Environmental Pollution, 180, 173–179. [CrossRef] [Google Scholar]
  55. Galaktionov KV. 2017. Patterns and processes influencing helminth parasites of Arctic coastal communities during climate change. Journal of Helminthology, 91, 387–408. [CrossRef] [PubMed] [Google Scholar]
  56. Gao J-W, Yuan X-P, Wu H, Xiang C-Y, Xie M, Song R, Chen Z-Y, Wu Y-A, Ou D-S. 2022. Mitochondrial phylogenomics of Acanthocephala: nucleotide alignments produce long-branch attraction artefacts. Parasites & Vectors, 15, 376. [CrossRef] [PubMed] [Google Scholar]
  57. García-Varela M, Andrade-Gómez L. 2021. First steps to understand the systematics of Echinorhynchidae Cobbold, 1876 (Acanthocephala), inferred through nuclear gene sequences. Parasitology International, 81, 102264. [CrossRef] [PubMed] [Google Scholar]
  58. García-Varela M, García-Prieto L, Rodríguez RP. 2011. Molecular identification and first description of the male of Neoechinorhynchus schmidti (Acanthocephala: Neoechinorhynchidae), a parasite of Trachemys scripta (Testudines) in México. Parasitology International, 60, 433–439. [CrossRef] [PubMed] [Google Scholar]
  59. García-Varela M, López-Jiménez A, González-García MT, Sereno-Uribe AL, Andrade-Gómez L. 2023. Contrasting the population genetic structure of a specialist (Hexaglandula corynosoma: Acanthocephala: Polymorphidae) and generalist parasite (Southwellina hispida) distributed sympatrically in Mexico. Parasitology, 150, 348–358. [CrossRef] [Google Scholar]
  60. García-Varela M, Masper A, Crespo EA, Hernández-Orts JS. 2021. Genetic diversity and phylogeography of Corynosoma australe Johnston, 1937 (Acanthocephala: Polymorphidae), an endoparasite of otariids from the Americas in the northern and southern hemispheres. Parasitology International, 80, 102205. [CrossRef] [PubMed] [Google Scholar]
  61. García-Varela M, Pérez-Ponce de León G, de la Torre P, Cummings MP, Sarma S, Laclette JP. 2000. Phylogenetic relationships of Acanthocephala based on analysis of 18S ribosomal RNA gene sequences. Journal of Molecular Evolution, 50, 532–540. [CrossRef] [PubMed] [Google Scholar]
  62. Gazi M, Kim J, García-Varela M, Park C, Littlewood DTJ, Park J-K. 2016. Mitogenomic phylogeny of Acanthocephala reveals novel class relationships. Zoologica Scripta, 45, 437–454. [CrossRef] [Google Scholar]
  63. Giari L, Fano EA, Castaldelli G, Grabner D, Sures B. 2020. The ecological importance of amphipod–parasite associations for aquatic ecosystems. Water, 12, 2429. [Google Scholar]
  64. Gilbert BM, Nachev M, Jochmann MA, Schmidt TC, Köster D, Sures B, Avenant-Oldewage A. 2020. You are how you eat. Differences in trophic levels of parasites infecting a single host according to stable isotopes. Parasitology Research, 119, 1393–1400. [CrossRef] [PubMed] [Google Scholar]
  65. Gismondi E, Beisel J-N, Cossu-Leguille C. 2012. Polymorphus minutus affects antitoxic responses of Gammarus roeseli exposed to cadmium. PLoS One, 7, e41475. [CrossRef] [PubMed] [Google Scholar]
  66. Gismondi E, Cossu-Leguille C, Beisel J-N. 2012. Acanthocephalan parasites: help or burden in gammarid amphipods exposed to cadmium? Ecotoxicology, 21, 1188–1193. [CrossRef] [PubMed] [Google Scholar]
  67. Golvan YJ, de Buron I. 1988. Les hôtes des Acanthocéphales II - Les Hôtes définitifs. 1. Poissons. Annales de Parasitologie Humaine et Comparée, 63, 349–375. [CrossRef] [EDP Sciences] [Google Scholar]
  68. Gomes APN, Olifiers N, Souza JGR, Barbosa HS, D’Andrea PS, Maldonado A. 2015. A new acanthocephalan species (Archiacanthocephala: Oligacanthorhynchidae) from the crab-eating fox (Cerdocyon thous) in the Brazilian Pantanal wetlands. Journal of Parasitology, 101, 74–79. [CrossRef] [PubMed] [Google Scholar]
  69. Goulding TC, Sarah Cohen C. 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]
  70. Gourbal BEF, Guillou F, Mitta G, Sibille P, Théron A, Pointier JP, Coustau C. 2008. Excretory-secretory products of larval Fasciola hepatica investigated using a two-dimensional proteomic approach. Molecular and Biochemical Parasitology, 161, 63–66. [CrossRef] [PubMed] [Google Scholar]
  71. Grabner D, Sures B. 2019. Amphipod parasites may bias results of ecotoxicological research. Diseases of Aquatic Organisms, 136, 121–132. [CrossRef] [Google Scholar]
  72. Guillou F, Roger E, Moné Y, Rognon A, Grunau C, Théron A, Mitta G, Coustau C, Gourbal BEF. 2007. Excretory-secretory proteome of larval Schistosoma mansoni and Echinostoma caproni, two parasites of Biomphalaria glabrata. Molecular and Biochemical Parasitology, 155, 45–56. [CrossRef] [PubMed] [Google Scholar]
  73. Harris M. 2020. The life cycle of the parasite Pomphorhynchus tereticollis in reference to 0+ cyprinids and the intermediate host Gammarus spp. in the UK, PhD Thesis, Bournemouth University. [Google Scholar]
  74. Hatcher MJ, Dick JTA, Bojko J, Stentiford GD, Stebbing P, Dunn AM. 2019. Infection and invasion: study cases from aquatic communities, in Wildlife Disease Ecology. Wilson K, Fenton A, Tompkins D, Editors. Cambridge University Press. p. 262–295. [Google Scholar]
  75. Herlyn H, Piskurek O, Schmitz J, Ehlers U, Zischler H. 2003. The syndermatan phylogeny and the evolution of acanthocephalan endoparasitism as inferred from 18S rDNA sequences. Molecular Phylogenetics and Evolution, 26, 155–164. [CrossRef] [PubMed] [Google Scholar]
  76. Hine PM, Kennedy CR. 1974. Observations on the distribution, specificity and pathogenicity of the acanthocephalan Pomphorhynchus laevis (Muller). Journal of Fish Biology, 6, 521–535. [CrossRef] [Google Scholar]
  77. Hine PM, Kennedy CR. 1974. The population biology of the acanthocephalan Pomphorhynchus laevis (Muller) in the River Avon. Journal of Fish Biology, 6, 665–679. [CrossRef] [Google Scholar]
  78. Hoffman GL. 1999. Parasites of North American freshwater fishes, 2nd edn. Cornell University Press: Ithaca, New York. [CrossRef] [Google Scholar]
  79. Hohenadler MAA, Nachev M, Thielen F, Taraschewski H, Grabner D, Sures B. 2018. Pomphorhynchus laevis: An invasive species in the river Rhine? Biological Invasions, 20, 207–217. [CrossRef] [Google Scholar]
  80. Holman JD, Burnett KG, Burnett LE. 2004. Effects of hypercapnic hypoxia on the clearance of Vibrio campbellii in the Atlantic blue crab, Callinectes sapidus Rathbun. Biological Bulletin, 206, 188–196. [CrossRef] [PubMed] [Google Scholar]
  81. Huston DC, Cribb TH, Smales LR. 2020. Molecular characterisation of acanthocephalans from Australian marine teleosts: proposal of a new family, synonymy of another and transfer of taxa between orders. Systematic Parasitology, 97, 1–23. [CrossRef] [PubMed] [Google Scholar]
  82. International Helminth Genomes Consortium. 2019. Comparative genomics of the major parasitic worms. Nature Genetics, 51, 163–174. [CrossRef] [PubMed] [Google Scholar]
  83. Iritani R, Sato T. 2018. Host-manipulation by trophically transmitted parasites: The switcher-paradigm. Trends in Parasitology, 34, 934–944. [CrossRef] [PubMed] [Google Scholar]
  84. Kamiya E, Urabe M, Okuda N. 2020. Does atypical 15N and 13C enrichment in parasites result from isotope ratio variation of host tissues they are infected? Limnology, 21, 139–149. [CrossRef] [Google Scholar]
  85. Karin BR, Gamble T, Jackman TR. 2020. Optimizing phylogenomics with rapidly evolving long exons: Comparison with anchored hybrid enrichment and ultraconserved elements. Molecular Biology and Evolution, 37, 904–922. [CrossRef] [PubMed] [Google Scholar]
  86. Keidel L, García-Varela M, Brener B, de León GP-P, Santos CP. 2019. Integrative taxonomy reveals a new species of Dollfusentis (Acanthocephala: Illiosentidae), in Orthopristis ruber (Osteichthyes: Haemulidae) from Rio de Janeiro, Brazil. Parasitology International, 71, 132–142. [CrossRef] [PubMed] [Google Scholar]
  87. Kennedy CR. 2006. Ecology of the Acanthocephala, 1st edn. Cambridge University Press. [CrossRef] [Google Scholar]
  88. Kennedy C, Bates R, Brown A. 1989. Discontinuous distributions of the fish acanthocephalans Pomphorhynchus laevis and Acanthocephalus anguillae in Britain and Ireland: an hypothesis. Journal of Fish Biology, 34, 607–619. [CrossRef] [Google Scholar]
  89. Kita Y, Hiruta SF, Sasaki M, Kajihara H. 2023. Systematic position of the genus Metacanthocephalus Yamaguti, 1959 (Palaeacanthocephala: Echinorhynchida) inferred from molecular evidence, with a redescription of Metacanthocephalus ovicephalus (Zhukov, 1960). Parasitology International, 94, 102731. [CrossRef] [PubMed] [Google Scholar]
  90. Koch RW, Shannon RP, Detwiler JT, Bolek MG. 2021. Molecular identification of juvenile Neoechinorhynchus spp. (Phylum: Acanthocephala) infecting ostracod and snail hosts provides insight into acanthocephalan host use. Journal of Parasitology, 107, 739–761. [Google Scholar]
  91. Labaude S, Rigaud T, Cézilly F. 2015. Host manipulation in the face of environmental changes: Ecological consequences. International Journal for Parasitology: Parasites and Wildlife, 4, 442–451. [CrossRef] [Google Scholar]
  92. Lacerda A, Roumbedakis K, Junior JB, Nuñer A, Petrucio M, Martins M. 2018. Fish parasites as indicators of organic pollution in southern Brazil. Journal of Helminthology, 92, 322–331. [CrossRef] [PubMed] [Google Scholar]
  93. Lafferty K. 1997. Environmental parasitology: what can parasites tell us about human impacts on the environment? Parasitology Today, 13, 251–255. [CrossRef] [Google Scholar]
  94. Lagrue C. 2017. Impacts of crustacean invasions on parasite dynamics in aquatic ecosystems: A plea for parasite-focused studies. International Journal for Parasitology: Parasites and Wildlife, 6, 364–374. [CrossRef] [Google Scholar]
  95. Le Moullac G, Soyez C, Saulnier D, Ansquer D, Avarre JC, Levy P. 1998. Effect of hypoxic stress on the immune response and the resistance to vibriosis of the shrimp Penaeus stylirostris. Fish and Shellfish Immunology, 8, 621–629. [CrossRef] [Google Scholar]
  96. Le TTY, Garcia MR, Nachev M, Grabner D, Balsa-Canto E, Hendriks AJ, Sures B. 2018. Development of a PBPK model for silver accumulation in chub infected with acanthocephalan parasites. Environmental Science & Technology, 52, 12514–12525. [CrossRef] [PubMed] [Google Scholar]
  97. Le TTY, Kiwitt G, Nahar N, Nachev M, Grabner D, Sures B. 2022. What contributes to the metal-specific partitioning in the chub-acanthocephalan system? Aquatic Toxicology, 247, 106178. [CrossRef] [Google Scholar]
  98. Le TTY, Rijsdijk L, Sures B, Hendriks AJ. 2014. Accumulation of persistent organic pollutants in parasites. Chemosphere, 108, 145–151. [CrossRef] [PubMed] [Google Scholar]
  99. Lewis J, Hoole D, Chappell LH. 2003. Parasitism and environmental pollution: parasites and hosts as indicators of water quality. Parasitology, 126, S1–S3. [CrossRef] [Google Scholar]
  100. Lewis S, Handy R, Cordi B, Billinghurst Z, Depledge M. 1999. Stress proteins (HSP’s): methods of detection and their use as an environmental biomarker. Ecotoxicology, 8, 351–368. [CrossRef] [Google Scholar]
  101. Lisitsyna OI, Kudlai 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]
  102. Lymbery AJ, Morine M, Kanani HG, Beatty SJ, Morgan DL. 2014. Co-invaders: The effects of alien parasites on native hosts. International Journal for Parasitology: Parasites and Wildlife, 3, 171–177. [CrossRef] [Google Scholar]
  103. Malyarchuk B, Derenko M, Mikhailova E, Denisova G. 2014. Phylogenetic relationships among Neoechinorhynchus species (Acanthocephala: Neoechinorhynchidae) from North-East Asia based on molecular data. Parasitology International, 63, 100–107. [CrossRef] [PubMed] [Google Scholar]
  104. Martínez-Aquino A, Reyna-Fabián ME, Rosas-Valdez R, Razo-Mendivil U, Pérez-Ponce de León G, García-Varela M. 2009. Detecting a complex of cryptic species within Neoechinorhynchus golvani (Acanthocephala: Neoechinorhynchidae) inferred from ITSs and LSU rDNA gene sequences. Journal of Parasitology, 95, 1040–1047. [CrossRef] [PubMed] [Google Scholar]
  105. Mauer K, Hellmann SL, Groth M, Fröbius AC, Zischler H, Hankeln T, Herlyn H. 2020. The genome, transcriptome, and proteome of the fish parasite Pomphorhynchus laevis (Acanthocephala). PLoS One, 15, e0232973. [CrossRef] [PubMed] [Google Scholar]
  106. Mauer KM, Schmidt H, Dittrich M, Fröbius AC, Hellmann SL, Zischler H, Hankeln T, Herlyn H. 2021. Genomics and transcriptomics of epizoic Seisonidea (Rotifera, syn. Syndermata) reveal strain formation and gradual gene loss with growing ties to the host. BMC Genomics, 22, 604. [CrossRef] [PubMed] [Google Scholar]
  107. McCahon CP, Poulton MJ. 1991. Lethal and sub-lethal effects of acid, aluminium and lime on Gammarus pulex during repeated simulated episodes in a Welsh stream. Freshwater Biology, 25, 169–178. [CrossRef] [Google Scholar]
  108. McCahon C, Brown A, Pascoe D. 1988. The effect of the acanthocephalan Pomphorhynchus laevis (Müller 1776) on the acute toxicity of cadmium to its intermediate host, the amphipod Gammarus pulex (L.). Archives of Environmental Contamination and Toxicology, 17, 239–243. [CrossRef] [Google Scholar]
  109. McCahon C, Brown A, Poulton MJ, Pascoe D. 1989. Effects of acid, aluminium and lime additions on fish and invertebrates in a chronically acidic Welsh stream. Water, Air, and Soil Pollution, 45, 345–359. [Google Scholar]
  110. Médoc V, Piscart C, Maazouzi C, Simon L, Beisel J-N. 2011. Parasite-induced changes in the diet of a freshwater amphipod: field and laboratory evidence. Parasitology, 138, 537–546. [CrossRef] [PubMed] [Google Scholar]
  111. Médoc V, Rigaud T, Motreuil S, Perrot-Minnot M-J, Bollache L. 2011. Paratenic hosts as regular transmission route in the acanthocephalan Pomphorhynchus laevis: potential implications for food webs. Naturwissenschaften, 98, 825–835. [CrossRef] [PubMed] [Google Scholar]
  112. Molbert N, Alliot F, Leroux-Coyau M, Médoc V, Biard C, Meylan S, Jacquin L, Santos R, Goutte A. 2020. Potential benefits of acanthocephalan parasites for chub hosts in polluted environments. Environmental Science & Technology, 54, 5540–5549. [CrossRef] [PubMed] [Google Scholar]
  113. Morton DN, Lafferty KD. 2022. Parasites in kelp-forest food webs increase food-chain length, complexity, and specialization, but reduce connectance. Ecological Monographs, 92, e1506. [CrossRef] [PubMed] [Google Scholar]
  114. Muhammad N, Ahmad MS, Li L, Zhao Q, Ullah H, Zhu X-Q, Ma J. 2020. Mitochondrial DNA dataset suggest that the genus Sphaerirostris Golvan, 1956 is a synonym of the genus Centrorhynchus Lühe, 1911. Parasitology, 147, 1149–1157. [CrossRef] [PubMed] [Google Scholar]
  115. Nachev M, Jochmann MA, Walter F, Wolbert JB, Schulte M, Schmidt TC, Sures B. 2017. Understanding trophic interactions in host-parasite associations using stable isotopes of carbon and nitrogen. Parasites & Vectors, 10, 90. [CrossRef] [PubMed] [Google Scholar]
  116. Nachev M, Sures B. 2016. Environmental parasitology: Parasites as accumulation bioindicators in the marine environment. Journal of Sea Research, 113, 45–50. [CrossRef] [Google Scholar]
  117. Nachev M, Zimmermann S, Rigaud T, Sures B. 2010. Is metal accumulation in Pomphorhynchus laevis dependent on parasite sex or infrapopulation size? Parasitology, 137, 1239–1248. [CrossRef] [PubMed] [Google Scholar]
  118. Near TJ, Garey JR, Nadler SA. 1998. Phylogenetic relationships of the Acanthocephala inferred from 18S ribosomal DNA sequences. Molecular Phylogenetics and Evolution, 10, 287–298. [CrossRef] [PubMed] [Google Scholar]
  119. Nickol BB. 1985. Epizootiology, in Biology of the Acanthocephala. Crompton DWT, Nickol BB, Editors. University Press: Cambridge, pp 307–346. [Google Scholar]
  120. de Oliveira EA, Bertollo LAC, Yano CF, Liehr T, de Bello Cioffi M. 2015. Comparative cytogenetics in the genus Hoplias (Characiformes, Erythrinidae) highlights contrasting karyotype evolution among congeneric species. Molecular Cytogenetics, 8, 1–10. [CrossRef] [PubMed] [Google Scholar]
  121. Orosová M, Králová-Hromadová I, Bazsalovicsová E, Špakulová M. 2010. Karyotype, chromosomal characteristics of multiple rDNA clusters and intragenomic variability of ribosomal ITS2 in Caryophyllaeides fennica (Cestoda). Parasitology International, 59, 351–357. [CrossRef] [PubMed] [Google Scholar]
  122. Orosová M, Marková A, Marec F, Barčák D, Brázová T, Oros M. 2022. New cytogenetic data on Caryophyllaeus laticeps and Paracaryophyllaeus gotoi, parasites of evolutionary interest. Parasitology, 149, 1094–1105. [CrossRef] [PubMed] [Google Scholar]
  123. Perrot-Minnot MJ, Cézilly F. 2013. Investigating candidate neuromodulatory systems underlying parasitic manipulation: Concepts, limitations and prospects. Journal of Experimental Biology, 216, 134–141. [CrossRef] [PubMed] [Google Scholar]
  124. Perrot-Minnot MJ, Guyonnet E, Bollache L, Lagrue C. 2019. Differential patterns of definitive host use by two fish acanthocephalans occurring in sympatry: Pomphorhynchus laevis and Pomphorhynchus tereticollis. International Journal for Parasitology: Parasites and Wildlife, 8, 135–144. [CrossRef] [Google Scholar]
  125. Perrot-Minnot MJ, Maddaleno M, Cézilly F. 2016. Parasite-induced inversion of geotaxis in a freshwater amphipod: A role for anaerobic metabolism? Functional Ecology, 30, 780–788. [CrossRef] [Google Scholar]
  126. Perrot-Minnot MJ, Sanchez-Thirion K, Cézilly F. 2014. Multidimensionality in host manipulation mimicked by serotonin injection. Proceedings of the Royal Society B: Biological Sciences, 281, 20141915. [CrossRef] [PubMed] [Google Scholar]
  127. Perrot-Minnot MJ, Špakulová M, Wattier R, Kotlík P, Düşen S, Aydoğdu A, Tougard C. 2018. Contrasting phylogeography of two Western Palaearctic fish parasites despite similar life cycles. Journal of Biogeography, 45, 101–115. [CrossRef] [Google Scholar]
  128. Pfenning-Butterworth AC, Sparkes TC. 2023. Evolutionary history and host ecology determine acanthocephalan egg shape. Evolutionary Biology, 50, 137–145. [CrossRef] [Google Scholar]
  129. Petkevičiūtė R. 1996. A chromosome study of Schistocephalus solidus (Müller, 1776) (Cestoda: Pseudophyllidea). Systematic Parasitology, 33, 183–186. [CrossRef] [Google Scholar]
  130. Pinacho-Pinacho CD, Hernández-Orts JS, Sereno-Uribe AL, Pérez-Ponce de León G, García-Varela M. 2017. Mayarhynchus karlae ng, n. sp. (Acanthocephala: Neoechinorhynchidae), a parasite of cichlids (Perciformes: Cichlidae) in southeastern Mexico, with comments on the paraphyly of Neoechynorhynchus Stiles & Hassall, 1905. Systematic Parasitology, 94, 351–365. [CrossRef] [PubMed] [Google Scholar]
  131. Pinacho-Pinacho CD, García-Varela M, Sereno-Uribe AL, de León GP-P. 2018. A hyper-diverse genus of acanthocephalans revealed by tree-based and non-tree-based species delimitation methods: Ten cryptic species of Neoechinorhynchus in Middle American freshwater fishes. Molecular Phylogenetics and Evolution, 127, 30–45. [CrossRef] [PubMed] [Google Scholar]
  132. Pinacho-Pinacho CD, Pérez-Ponce de León G, García-Varela M. 2012. Description of a new species of Neoechinorhynchus (Acanthocephala: Neoechinorhynchidae) a parasite of Dormitator latifrons from Southwestern Mexico based on morphological and molecular characters. Parasitology International, 61, 634–644. [CrossRef] [PubMed] [Google Scholar]
  133. Pinacho-Pinacho CD, Sereno-Uribe AL, García-Varela M. 2014. Morphological and molecular data reveal a new species of Neoechinorhynchus (Acanthocephala: Neoechinorhynchidae) from Dormitator maculatus in the Gulf of Mexico. Parasitology International, 63, 763–771. [CrossRef] [PubMed] [Google Scholar]
  134. Pinacho-Pinacho CD, Sereno-Uribe AL, Pérez-Ponce de León G, Garcia-Varela M. 2015. Checklist of the species of Neoechinorhynchus (Acanthocephala: Neoechinorhynchidae) in fishes and turtles in Middle-America, and their delimitation based on sequences of the 28S rDNA. Zootaxa, 3985, 98–116. [CrossRef] [PubMed] [Google Scholar]
  135. Podolska M, Polak-Juszczak L, Nadolna-Ałtyn K. 2016. Host condition and accumulation of metals by acanthocephalan parasite Echinorhynchus gadi in cod Gadus morhua from the southern Baltic Sea. Marine Pollution Bulletin, 113, 287–292. [CrossRef] [PubMed] [Google Scholar]
  136. Poulin R. 2007. Investing in attachment: evolution of anchoring structures in acanthocephalan parasites. Biological Journal of the Linnean Society, 90, 637–645. [CrossRef] [Google Scholar]
  137. Poulin R, Maure F. 2015. Host manipulation by parasites: A look back before moving forward. Trends in Parasitology, 31, 563–570. [CrossRef] [PubMed] [Google Scholar]
  138. Ratnasingham S, Hebert PDN. 2007. BOLD: The Barcode of Life Data system (http://www.barcodinglife.org). Molecular Ecology Notes, 7, 355–364. [CrossRef] [PubMed] [Google Scholar]
  139. Ratnasingham S, Hebert PDN. 2013. A DNA-based registry for all animal species: The Barcode Index Number (BIN) system. PLoS One, 8, e66213. [CrossRef] [PubMed] [Google Scholar]
  140. Rauque C, Flores V, Semenas L. 2022. Pseudocorynosoma enrietti (Molfi & Freitas Fernandes, 1953) (Acanthocephala: Polymorphidae) from Patagonia (Argentina): life cycle, localities, and new host records. Journal of Helminthology, 96, e38. [CrossRef] [PubMed] [Google Scholar]
  141. Reier S, Sattmann H, Schwaha T, Harl J, Konecny R, Haring E. 2019. An integrative taxonomic approach to reveal the status of the genus Pomphorhynchus Monticelli, 1905 (Acanthocephala: Pomphorhynchidae) in Austria. International Journal for Parasitology: Parasites and Wildlife, 8, 145–155. [CrossRef] [Google Scholar]
  142. Reier S, Sattmann H, Schwaha T, Fuehrer H-P, Haring E. 2020. Unravelling the hidden biodiversity – the establishment of DNA barcodes of fish parasitizing Acanthocephala Koehlreuther, 1771 in view of taxonomic misidentifications, intraspecific variability and possible cryptic species. Parasitology, 147, 1499–1508. [CrossRef] [PubMed] [Google Scholar]
  143. Rigaud T, Moret Y. 2003. Differential phenoloxidase activity between native and invasive gammarids infected by local acanthocephalans: differential immunosuppression? Parasitology, 127, 571–577. [CrossRef] [PubMed] [Google Scholar]
  144. Rosas-Valdez R, Morrone JJ, García-Varela M. 2012. Molecular phylogenetics of Floridosentis Ward, 1953 (Acanthocephala: Neoechinorhynchidae) parasites of mullets (Osteichthyes) from Mexico, using 28S rDNA sequences. Journal of Parasitology, 98, 855–862. [CrossRef] [PubMed] [Google Scholar]
  145. Rosas-Valdez R, Morrone JJ, Pinacho-Pinacho CD, Domínguez-Domínguez O, García-Varela M. 2020. Genetic diversification of acanthocephalans of the genus Floridosentis Ward 1953 (Acanthocephala: Neoechinorhynchidae), parasites of mullets from the Americas. Infection, Genetics and Evolution, 85, 104535. [CrossRef] [PubMed] [Google Scholar]
  146. Ru SS, Yang RJ, Chen HX, Kuzmina TA, Spraker TR, Li L. 2022. Morphology, molecular characterization and phylogeny of Bolbosoma nipponicum Yamaguti, 1939 (Acanthocephala: Polymorphidae), a potential zoonotic parasite of human acanthocephaliasis. International Journal for Parasitology: Parasites and Wildlife, 18, 212–220. [CrossRef] [Google Scholar]
  147. Sarwar H, Zhao W, Kibet CJ, Sitko J, Nie P. 2021. Morphological and complete mitogenomic characterisation of the acanthocephalan Polymorphus minutus infecting the duck Anas platyrhynchos. Folia Parasitologica, 68, 1–9. [CrossRef] [Google Scholar]
  148. Scheifler M, Ruiz-Rodríguez M, Sanchez-Brosseau S, Magnanou E, Suzuki MT, West N, Duperron S, Desdevises Y. 2019. Characterization of ecto- and endoparasite communities of wild Mediterranean teleosts by a metabarcoding approach. PLoS One, 14, e0221475. [CrossRef] [PubMed] [Google Scholar]
  149. Schmidt GH. 1985. Development and life cycles, in Biology of the Acanthocephala. Crompton DWT, Nickol BB, Editors. University Press: Cambridge. p. 273–305. [Google Scholar]
  150. Schmidt H, Mauer K, Glaser M, Dezfuli BS, Hellmann SL, Silva Gomes AL, Butter F, Wade RC, Hankeln T, Herlyn H. 2022. Identification of antiparasitic drug targets using a multi-omics workflow in the acanthocephalan model. BMC Genomics, 23, 677. [CrossRef] [PubMed] [Google Scholar]
  151. Schmidt H, Mauer K, Hankeln T, Herlyn H. 2022. Host-dependent impairment of parasite development and reproduction in the acanthocephalan model. Cell & Bioscience, 12, 75. [CrossRef] [PubMed] [Google Scholar]
  152. Scholz T, Choudhury A. 2014. Parasites of freshwater fishes in North America: Why so neglected? Journal of Parasitology, 100, 26–45. [CrossRef] [PubMed] [Google Scholar]
  153. Selbach C, Mouritsen KN, Poulin R, Sures B, Smit NJ. 2022. Bridging the gap: aquatic parasites in the One Health concept. Trends in Parasitology, 38, 109–111. [CrossRef] [PubMed] [Google Scholar]
  154. Sereno-Uribe AL, López-Jiménez A, González-García MT, Pinacho-Pinacho CD, Ríos RM, García-Varela M. 2022. Phenotypic plasticity, genetic structure and systematic position of Neoechinorhynchus emyditoides Fisher, 1960 (Acanthocephala: Neoechinorhynchidae): a parasite of emydid turtles from the Nearctic and Neotropical regions. Parasitology, 149, 991–1002. [CrossRef] [Google Scholar]
  155. Shinn AP, Pratoomyot J, Bron JE, Paladini G, Brooker EE, Brooker AJ. 2015. Economic costs of protistan and metazoan parasites to global mariculture. Parasitology, 142, 196–270. [CrossRef] [PubMed] [Google Scholar]
  156. Sielaff M, Schmidt H, Struck TH, Rosenkranz D, Mark Welch DB, Hankeln T, Herlyn H. 2016. Phylogeny of Syndermata (syn. Rotifera): mitochondrial gene oder verifies epizoic Seisonidea as sister to endoparasitic Acanthocephala within monophyletic Hemirotifera. Molecular Phylogenetics and Evolution, 96, 79–92. [CrossRef] [PubMed] [Google Scholar]
  157. Smales L. 2014. The genus Rhadinorhynchus (Acanthocephala: Rhadinorhynchidae) from marine fish in Australia with the description of four new species. Acta Parasitologica, 59, 721–736. [CrossRef] [PubMed] [Google Scholar]
  158. Sørensen MV, Giribet G. 2006. A modern approach to rotiferan phylogeny: combining morphological and molecular data. Molecular Phylogenetics and Evolution, 40, 585–608. [CrossRef] [PubMed] [Google Scholar]
  159. Špakulová M, Králová-Hromadová I, Dudiňák V, Reddy P. 2002. Karyotype of Acanthocephalus lucii: the first record of supernumerary chromosomes in thorny-headed worms. Parasitology Research, 88, 778–780. [CrossRef] [PubMed] [Google Scholar]
  160. Špakulová M, Perrot-Minnot MJ, Neuhaus B. 2011. Resurrection of Pomphorhynchus tereticollis (Rudolphi, 1809) (Acanthocephala: Pomphorhynchidae) based on new morphological and molecular data. Helminthologia, 48, 268–277. [CrossRef] [Google Scholar]
  161. Stanevičiũté G, Kiseliené V. 2001. Chromosome studies of Ichthyocotylurus platycephalus (Creplin, 1825) Odening 1969 with description of triploid variant and comparative karyology of the genus Ichthyocotylurus. Parasite, 8, 137–145. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  162. Steinauer M, Nickol B, Ortí G. 2007. Cryptic speciation and patterns of phenotypic variation of a highly variable acanthocephalan parasite. Molecular Ecology, 16, 4097–4109. [CrossRef] [PubMed] [Google Scholar]
  163. Sures B, Radszuweit H. 2007. Pollution-induced heat shock protein expression in the amphipod Gammarus roeseli is affected by larvae of Polymorphus minutus (Acanthocephala). Journal of Helminthology, 81, 191–197. [CrossRef] [PubMed] [Google Scholar]
  164. Sures B, Thielen F, Baska F, Messerschmidt J, Von Bohlen A. 2005. The intestinal parasite Pomphorhynchus laevis as a sensitive accumulation indicator for the platinum group metals Pt, Pd, and Rh. Environmental Research, 98, 83–88. [CrossRef] [PubMed] [Google Scholar]
  165. Sures B, Nachev M. 2022. Effects of multiple stressors in fish: how parasites and contaminants interact. Parasitology, 149, 1822–1828. [CrossRef] [PubMed] [Google Scholar]
  166. Sures B, Nachev M, Selbach C, Marcogliese DJ. 2017. Parasite responses to pollution: what we know and where we go in “Environmental Parasitology”. Parasites & Vectors, 10, 65. [CrossRef] [PubMed] [Google Scholar]
  167. Sures B, Reimann N. 2003. Analysis of trace metals in the Antarctic host-parasite system Notothenia coriiceps and Aspersentis megarhynchus (Acanthocephala) caught at King George Island, South Shetland Islands. Polar Biology, 26, 680–686. [CrossRef] [Google Scholar]
  168. Sures B, Siddall R, Taraschewski H. 1999. Parasites as accumulation indicators of heavy metal pollution. Parasitology Today, 15, 16–21. [CrossRef] [Google Scholar]
  169. Sures B, Taraschewski H. 1995. Cadmium concentrations in two adult acanthocephalans, Pomphorhynchus laevis and Acanthocephalus lucii, as compared with their fish hosts and cadmium and lead levels in larvae of A. lucii as compared with their crustacean host. Parasitology Research, 81, 494–497. [CrossRef] [Google Scholar]
  170. Tain L, Perrot-Minnot MJ, Cézilly F. 2006. Altered host behaviour and brain serotonergic activity caused by acanthocephalans: Evidence for specificity. Proceedings of the Royal Society B: Biological Sciences, 273, 3039–3045. [CrossRef] [PubMed] [Google Scholar]
  171. Tain L, Perrot-Minnot MJ, Cézilly F. 2007. Differential influence of Pomphorhynchus laevis (Acanthocephala) on brain serotonergic activity in two congeneric host species. Biology Letters, 3, 69–72. [CrossRef] [Google Scholar]
  172. Taraschewski H. 2000. Host-parasite interactions in acanthocephala: A morphological approach. Advances in Parasitology, 46, 1–179. [CrossRef] [PubMed] [Google Scholar]
  173. Thielen F, Zimmermann S, Baska F, Taraschewski H, Sures B. 2004. The intestinal parasite Pomphorhynchus laevis (Acanthocephala) from barbel as a bioindicator for metal pollution in the Danube River near Budapest, Hungary. Environmental Pollution, 129, 421–429. [CrossRef] [Google Scholar]
  174. Tokeson JPE, Holmes JC. 1982. The effects of temperature and oxygen on the development of Polymorphus marilis (Acanthocephala) in Gammarus lacustris (Amphipoda). Journal of Parasitology, 68, 112–119. [CrossRef] [Google Scholar]
  175. Valtonen ET, Holmes JC, Koskivaara M. 1997. Eutrophication, pollution and fragmentation: effects on parasite communities in roach (Rutilus rutilus) and perch (Perca fluviatilis) in four lakes in central Finland. Canadian Journal of Fisheries and Aquatic Sciences, 54, 572–585. [CrossRef] [Google Scholar]
  176. Vardić Smrzlić I, Valić D, Kapetanović D, Dragun Z, Gjurčević E, Ćetković H, Teskeredžić E. 2013. Molecular characterisation and infection dynamics of Dentitruncus truttae from trout (Salmo trutta and Oncorhynchus mykiss) in Krka River, Croatia. Veterinary Parasitology, 197, 604–613. [Google Scholar]
  177. Verkuil YI, Nicolaus M, Ubels R, Dietz MW, Samplonius JM, Galema A, Kiekebos K, de Knijff P, Both C. 2022. DNA metabarcoding quantifies the relative biomass of arthropod taxa in songbird diets: Validation with camera-recorded diets. Ecology and Evolution, 12, e8881. [CrossRef] [PubMed] [Google Scholar]
  178. 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]
  179. de Vos A, Faux CE, Marthick J, Dickinson J, Jarman SN. 2018. New determination of prey and parasite species for northern Indian Ocean blue whales. Frontiers in Marine Science, 5, 104. [CrossRef] [Google Scholar]
  180. Wayland MT, Vainio JK, Gibson DI, Herniou EA, Littlewood DT, Väinölä R. 2015. The systematics of Echinorhynchus Zoega in Müller, 1776 (Acanthocephala, Echinorhynchidae) elucidated by nuclear and mitochondrial sequence data from eight European taxa. Zookeys, 26, 25–52. [CrossRef] [Google Scholar]
  181. Weber M, Wey-Fabrizius AR, Podsiadlowski L, Witek A, Schill RO, Sugár L, Herlyn H, Hankeln T. 2013. Phylogenetic analyses of endoparasitic Acanthocephala based on mitochondrial genomes suggest secondary loss of sensory organs. Molecular Phylogenetics and Evolution, 66, 182–189. [CrossRef] [PubMed] [Google Scholar]
  182. Wey-Fabrizius AR, Herlyn H, Rieger B, Rosenkranz D, Witek A, Welch DBM, Ebersberger I, Hankeln T. 2014. Transcriptome data reveal syndermatan relationships and suggest the evolution of endoparasitism in Acanthocephala via an epizoic stage. PLoS One, 9, e88618. [CrossRef] [PubMed] [Google Scholar]
  183. Zheng C, Zhao Q, Li E, Zhao D, Sun S. 2022. Role of hypoxia in the behaviour, physiology, immunity and response mechanisms of crustaceans: A review. Reviews in Aquaculture, 14, 676–687. [CrossRef] [Google Scholar]
  184. Zimmermann S, Von Bohlen A, Messerschmidt J, Sures B. 2005. Accumulation of the precious metals platinum, palladium and rhodium from automobile catalytic converters in Paratenuisentis ambiguus as compared with its fish host, Anguilla anguilla. Journal of Helminthology, 79, 85–89. [CrossRef] [PubMed] [Google Scholar]

Cite this article as: Perrot-Minnot M-J, Cozzarolo C-S, Amin O, Barčák D, Bauer A, Filipović Marijić V, García-Varela M, Servando Hernández-Orts J, Yen Le 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.

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