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Adaptive Radiation within Marine Anisakid Nematodes: A Zoogeographical Modeling of Cosmopolitan, Zoonotic Parasites

  • Thomas Kuhn,

    Affiliation: Biodiversity and Climate Research Centre (BiK-F, LOEWE), Medical Biodiversity and Parasitology; Senckenberg Gesellschaft für Naturforschung (SGN); Goethe-University (GO), Institute for Ecology, Evolution and Diversity, Frankfurt am Main, Germany

  • Jaime García-Màrquez,

    Affiliation: Departamento de Gestión Ambiental, Carbones del Cerrejón Limited, Bogotá, Colombia

  • Sven Klimpel

    sven.klimpel@senckenberg.de

    Affiliation: Biodiversity and Climate Research Centre (BiK-F, LOEWE), Medical Biodiversity and Parasitology; Senckenberg Gesellschaft für Naturforschung (SGN); Goethe-University (GO), Institute for Ecology, Evolution and Diversity, Frankfurt am Main, Germany

Adaptive Radiation within Marine Anisakid Nematodes: A Zoogeographical Modeling of Cosmopolitan, Zoonotic Parasites

  • Thomas Kuhn, 
  • Jaime García-Màrquez, 
  • Sven Klimpel
PLOS
x
  • Published: December 13, 2011
  • DOI: 10.1371/journal.pone.0028642

Abstract

Parasites of the nematode genus Anisakis are associated with aquatic organisms. They can be found in a variety of marine hosts including whales, crustaceans, fish and cephalopods and are known to be the cause of the zoonotic disease anisakiasis, a painful inflammation of the gastro-intestinal tract caused by the accidental consumptions of infectious larvae raw or semi-raw fishery products. Since the demand on fish as dietary protein source and the export rates of seafood products in general is rapidly increasing worldwide, the knowledge about the distribution of potential foodborne human pathogens in seafood is of major significance for human health. Studies have provided evidence that a few Anisakis species can cause clinical symptoms in humans. The aim of our study was to interpolate the species range for every described Anisakis species on the basis of the existing occurrence data. We used sequence data of 373 Anisakis larvae from 30 different hosts worldwide and previously published molecular data (n = 584) from 53 field-specific publications to model the species range of Anisakis spp., using a interpolation method that combines aspects of the alpha hull interpolation algorithm as well as the conditional interpolation approach. The results of our approach strongly indicate the existence of species-specific distribution patterns of Anisakis spp. within different climate zones and oceans that are in principle congruent with those of their respective final hosts. Our results support preceding studies that propose anisakid nematodes as useful biological indicators for their final host distribution and abundance as they closely follow the trophic relationships among their successive hosts. The modeling might although be helpful for predicting the likelihood of infection in order to reduce the risk of anisakiasis cases in a given area.

Introduction

Approximately 20,000 cases of human anisakidosis infections are reported every year from a wide range of coastal regions, primarily in Japan and Europe [1]. This zoonosis, named after a family of marine nematodes, the Anisakidae (“whale-, seal-, cod-, herringworms”), is the result of the ingestion of infectious third stage larvae (L3) in raw or undercooked marine fish products. The term anisakidosis designates infections caused by nematodes of the family Anisakidae, whereas the term anisakiasis includes all infections that are caused by members of the genus Anisakis [2]. Besides the major clinical symptoms such as abdominal pain, nausea, vomiting and fever, anisakidosis infections can be associated with an increased risk of allergic responses caused by even very small doses of nematode antigens [2]. Increased globalisation and fast growing international markets, the massive demand for exotic dishes such as sushi, sashimi, salted/smoked herring, and the increasing propensity not to overcook food, have led to an accumulation of infections even in countries that do not have traditions of consuming raw fish [2], [3]. Besides the seal worms Pseudoterranova decipiens and Contracaecum spp. (diseases: anisakidosis), species of the genus Anisakis (A. simplex complex, A. physeteris) are considered the most common cause of human anisakiasis infections [1]. The general life cycle of these parasites involves a broad range of invertebrates, (such as crustaceans and chaetognaths), teleost fish species and cephalopods, which act as intermediate or paratenic hosts. Marine mammals such as cetaceans (toothed and baleen whales) and, sometimes, pinnipeds (seals) serve as final hosts [4], [5]. Toothed and baleen whales of the families Delphinidae, Ziphiidae, Physeteridae and Kogiidae are considered the main definitive hosts, but members of the Balaenopteridae, Pontoporidae, Monodontidae, Phocoenidae and Neobalaenidae have also been parasitised [6]. The increased number of individuals resulting from stronger regulation to protect whale populations is thought to be another reason for the increase in anisakiasis infections in the last three decades [1], [2].

In spite of the fact that A. simplex s.l. is the most common anisakiasis cause, it is still unclear whether all Anisakis species can cause clinical symptoms in humans [1]. This is why the identification of species and knowledge about their distribution is of primary importance for understanding parasite epidemiology, especially for species of medical, veterinary and commercial importance [7].

The distribution of marine helminth parasites is influenced by a wide range of abiotic factors such as hydrographic and climate conditions. It is also influenced by transmission pathways that closely follow the trophic relationship between the parasites' final, intermediate and transport hosts. Thus the species range and, especially, feeding behaviour of the hosts has to be taken into account in order to understand the occurrence of anisakid nematodes and predict the likelihood of infections in a given area [5].

Most descriptions of zoogeographical distribution patterns rely on sampling data from regions where the occurrence of species has been recorded [8]. However, this type of information is scarce for many species and usually biased towards accessible areas or places of special interest [9][11]. In order to tackle systematic errors such as oversimplification (e.g. outline maps) or underestimation (e.g. dot maps) in the assessment of biogeographical maps, several techniques were developed that interpolate and project distribution ranges to places where species potentially exist but have not yet been sampled. Triangulation techniques, for example, have played an important role in conservation, and the results of applying the minimum convex polygon to a set of species collection records have been a crucial point in assessing the conservation status of species [12], [13]. However, this method has gone through a series of refinements that seek to avoid certain constraints and biases (e.g. overestimation, detection of disjunctive distribution). Burgmann and Fox [14] recommend the use of the α-hull- instead of the convex-hull-interpolation technique because it produces a more flexible outer polygon surface that is more resistant to bias and allows the outer polygon line to be broken into detailed and discrete hulls. More recently, Raedig et al. [11] developed a geometric conditional triangulation approach that is based on a specified interpolation distance and avoids the problem of excluding narrow endemic species from the analysis. The final distribution range of a particular species is constructed not only on those areas that are included within the interpolated triangles, but also on those that connect species occurrence locations within the given interpolation distance as well as areas where locations are isolated due to a lack of neighbouring occurrences.

Based on these developments we present here a methodology for interpolating occurrence locations for every described Anisakis spp. (Anisakis simplex (Rudolphi, 1809, det. Krabbe, 1878) (sensu stricto), A. pegreffii Campana-Rouget & Biocca, 1955, A. simplex C (Nascetti et al., 1986, Mattiucci et al., 1997), A. typica (Diesing, 1860), A. nascettii Mattiucci et al., 2009, A. ziphidarum Paggi et al, 1998, A. physeteris Baylis 1923, A. paggiae Mattiucci et al. 2005, A. brevispiculata Dollfus, 1966), which combines aspects of the α-hull algorithm and the geometric interpolation approach [11], [15][25].

Results and Discussion

A total of 373 Anisakis spp. larvae from 30 teleost and bony fish host species were collected during fieldwork in 21 different sampling areas and identified by sequence analyses of ribosomal internal transcribed spacers (ITS-1/2, 5.8S) (Table S1, Figure S1) (Genbank Accession number JN968593–JN968965). Sequence analyses identified five already described Anisakis species (A. simplex s.s., A. pegreffii, A. simplex C, A. typica, A. physeteris) (Table S2). We combined the sequence data obtained with occurrence reports (584 nematode individuals) taken from 53 revised publications [5], [6], [18][21], [23], [26][71] and, using a non-commercial geographical information system (QGIS) transferred them as presence data to a grid with a resolution of 1°×1° covering the globe. Based on the centroids of the quadrats, the α-hull was calculated for 25 different values ranging from 2 to 50 in a two-step interval using the alpha-hull package [72]. The final distribution of every species was visualized in a range map with a continuous colour gradient, in which the intensity of the colour increased with the probability of species occurrence in a certain area (Figure S2).

The results of our modeling strongly indicate the existence of species-specific distribution patterns of Anisakis spp. within different climate zones and oceans that are in principle congruent with those of their respective final hosts (Figure 1). To date, 35 marine mammals and more than 75 different bony and elasmobranch fish species have been verified by molecular methods as potential hosts for Anisakis spp., but several studies provide evidence that the members of the Anisakis species complexes differ in their ecology and final host preferences [5], [21], [28], [37], [41], [73]. We found that species that are considered to be phylogenetically closely related show similar distribution patterns due to major congruities in the parasites' ecology and host preferences.

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Figure 1. Modeled species range of every described Anisakis spp.

Colour intensity reflects the probability of Anisakis occurrence. Dark red shadings indicate molecular proven Anisakis records. First clade: Anisakis simplex s.s., A. pegreffii, A. simplex C (building the A. simplex complex), A. ziphidarum, A. nascettii and A. typica. Second clade: Anisakis paggiae, A. physeteris and A. brevispiculata (A. physeteris complex) belong to the second clade (phylogeny follows Klimpel et al. 2010) [45].

doi:10.1371/journal.pone.0028642.g001

The genus Anisakis comprises two major clades. Whereas the first clade includes the A. simplex complex (A. simplex s.s., A. pegreffii, A. simplex C), as well as A. typica and both sister-species (A. nascettii and A. ziphidarum), the second clade consists exclusively of the A. physeteris complex (A. brevispiculata, A paggiae, A. physeteris) [5], [6].

Species of the A. simplex complex are mainly distributed within the Atlantic Ocean and in the West and East Pacific where large populations of their final hosts (Delphinidae) occur (Figure 1). In the case of A. simplex s.s., the most common exciter of human anisakiasis infection in Japan [74], the distribution could be narrowed down to a northern hemisphere range within the West and East Atlantic and Pacific Ocean, between 20°N and far beyond the Arctic polar circle (80°N) (Figure 1). The species range of A. pegreffii extends from Mediterranean waters through the East Atlantic Ocean down to the Antarctic Peninsula, with additional records in Japanese and Chinese waters (Figure 1). Anisakis simplex C has a discontinuous range: along the Canadian/US east coast; at the southern tip of Africa; and between New Zealand and Australia. Occurrence has also been proven between South American and Antarctic waters (Figure 1). These three closely related species of the A. simplex complex mainly include oceanic delphinids (e.g. Short-beaked Common Dophin Delphinus delphis, Long-finned Pilot Whale Globicephala melas) as final hosts in their life cycle, which are known to form large populations in the Atlantic Ocean where they usually feed on pelagic fish and cephalopods. The specific distribution patterns shown by these anisakids are likely to be a consequence of dispersal through the faeces of infested oceanic delphinids. This contrasts with the distribution of A. typica, the most abundant species in the tropics and subtropics between 45°N and 25°S (Figure 1). Anisakis typica is a common parasite of various dolphin species of warmer temperate waters, such as the Tucuxi Sotalia fluviatilis, the Common Bottlenose Dolphin Tursiops truncatus and the Pantropical Spotted Dolphin Stenella attenuata. These are more usually associated with shallower waters near the coast [50].

The sister taxa A. ziphidarum and A. nascettii are in principle congruent (Figure 1). While both species are distributed in the East Atlantic Ocean near Madeira and the Moroccan coast, the West Pacific between New Zealand and South Australia, and the southern tip of Africa, A. ziphidarum has a slightly extended range with additional records in the West Atlantic Ocean (Figure 1). This broadly similar zoogeography is also mirrored in their ecology as they have major similarities regarding their host preferences. Whereas A. ziphidarum also includes Cuvier's Beaked Whale Ziphius cavirostris as potential final host in its life cycle, both species seem to prefer ziphiid whales of the genus Mesoplodon spp. Taking into account the collection sites of the definitive hosts so far recorded, Mattiucci et al. [21] suggested that the geographical distribution of these parasites is related to that of their final hosts.

The close genetic relationship of the three representatives of the A. physeteris complex (A. physeteris, A. paggiae, A. brevispiculata) is also reflected in a relatively homogeneous distribution throughout the central Atlantic Ocean (Figure 1). Very similar to A. ziphidarum and A. nascettii, these species are known to be host specific and broadly include sperm whales as final hosts. While A. physeteris is the only species of this complex that parasitizes the Sperm Whale Physeter macrocephalus, all of them show clear preferences for the kogiid whales, the Pygmy Sperm Whale Kogia breviceps and the Dwarf Sperm Whale Kogia sima, which are mainly distributed in the central Atlantic Ocean. Sperm whales typically inhabit deepwater habitats in the tropics and the temperate zones where they mostly feed in depths between 500–1200 m, primarily on cephalopods and, less frequently, on deep sea fish and crustaceans [6], [75].

Our results support preceding studies that propose that anisakid nematodes as useful biological indicators for their final host distribution and abundance as they closely follow the trophic relationships among their successive hosts. Klimpel et al. (2011) [6] identified for the first time A. paggiae in the Irminger Sea and indicated a more extended migration towards northern latitudes than could have been inferred from the distribution range of kogiid whales reported so far. Our modeling clearly demonstrates, that the distribution patterns of Anisakis spp. can be narrowed down to certain areas within climatic zones and oceans and are mainly influenced by the species ranges and feeding behaviours of their respective intermediate and mammalian final hosts.

The world's least developed countries are particularly reliant on fish, which provides 27% of their dietary protein intake [76]. These countries also produce 20% of the world's fish exports [76]. A broad knowledge of the distribution of Anisakis spp. is of particular importance for predicting the likelihood of infection in order to reduce the risk of anisakidosis cases in a given area.

Materials and Methods

Ethics Statement

An approval by a review board institution or ethics committee was not necessary, because all the fish in the current study were obtained in different locations from fishermen selling fresh fish for consumption or were collected during regularly fishery cruises.

Sample collection

A total of 373 Anisakis spp. larvae from 30 teleost and bony fish host species were collected during field phase from 21 different sampling areas (Table S1, Figure S1). Fish were identified according to Fishbase [77] and Gon and Heemstra [78]. Nematodes were extensively washed in 0.9% saline solution and identified morphologically to genus level according to Anderson (2000) and Moravec [79], [80]. All samples were fixed and stored in EtOH (abs.) prior to molecular examination.

PCR amplification and species identification

Genomic DNA was isolated and purified from individual Anisakis spp. larvae using a genomic DNA extraction kit (Peqlab Biotechnology GmbH, Erlangen, Germany) according to the instructions of the manufacturer. The rDNA region comprising the ITS-1, 5.8S, ITS-2 and flanking sequences ( = ITS+) was amplified was amplified using primers NC2 (5′-TTA-GTT-TCT-TTT-CCT-CCG-CT-3′) and TK1 (5′-GGC-AAA-AGT-CGT-AAC-AAG-GT-3′) [81]. Primer TK1 was manually designed and then synthesized by Eurofins MWG Operon (Ebersberg, Germany). PCR-reaction (50 µl) included 25 µl Master-Mix (Peqlab Biotechnology GmbH, Erlangen, Germany) containing dNTP, MgCl2, Buffer and Taq-Polymerase, 3 µl of each primer, 14ddH2O and 5 µl genomic DNA. Each PCR reaction were performed in a thermocycler (Peqlab, Germany) under the following conditions: after an initial denaturation at 95°C for 1 min, 40 cycles of 94°C for 45 sec (denaturation), 55°C for 45 sec (annealing), 72°C for 45 sec (extension), followed by a final extension at 72°C for 10 min. Samples without DNA were included in each PCR run. PCR products were examined on 1% agarose gels. A 100 bp ladder marker (peqGOLD, Erlangen, Germany) was used to estimate the size of the PCR products. To prepare the samples for the sequencing, PCR products were purified with Cycle-Pure Kit (Peqlab Biotechnology GmbH, Erlangen, Germany). Afterwards a total volume of 7 µl, including 2 µl primer (individually) and 5 µl of the PCR product (250 ng/µl) were sequenced by Seqlab (Goettingen GmbH, Germany). Both spacers and the 5.8S gene from each PCR product were sequenced, using primer TK1 (5′-GGC-AAA-AGT-CGT-AAC-AAG-GT-3′). The ITS-1, 5.8S and ITS-2 sequences were determined for all 373 Anisakis nematodes isolated from the 30 host species (Genbank Accession number JN968593–JN968965). For species identification, the obtained sequences were compared with those previously deposited for the same marker in the Genbank using the BLASTn algorithm (Table S2).

Zoogeographical interpolation

A database was built comprising the species names and the exact coordinates or geographical regions where they have been recorded. The spatial information were collected from the fieldwork and molecular analyses mentioned above as well as from literature review of 53 publications [5], [6], [18][21], [23], [26][71]. The geographical information contained in this database where transferred onto a grid of 1°×1° covering the globe in which species occurrences were overlaid and the corresponding quadrats marked as occurrences for each species in turn. This procedure was manually done using the non-commercial geographical information system Quantum GIS (QGIS, version 1.6.0) (Figure S2A) [82].

To estimate distribution ranges two steps were followed. First, the alpha-hull was calculated based on the centroids (i.e. points) of the quadrats where a species was recorded [14], [83]. For this, a Delauney triangulation between the centroids was applied and the average length of all lines of all triangles was calculated. The final hull (i.e. polygon) results from those centroids, which are connected by a line smaller than a multiple (alpha parameter) of the average line length (Figure S2B). This procedure was implemented using the ‘alphahull’ package [72] in the R statistical software [84]. Second, distribution ranges were defined as the corresponding quadrats intersecting the created polygons and also those quadrats with single occurrences that were not included within any of the polygons (Figure S2C) [11].

Since the size and number of polygons as well as the number of single occurrences depends on the value of the alpha parameter (i.e. an alpha of zero is equal to the set of single occurrences (dot map) and infinite alpha will encompass all points in one polygon (convex hull) [14], the distribution range of each species was calculated with 25 different alphas (i.e. a sequence of alphas ranging from 2 to 50 in a two-step interval). Results were summed up into a final distribution range map which is depicted in a continuous colour gradient. Darker colours represent areas where the probability of finding the species is relatively high and as the colours get lighter the uncertainties of an area being part of the species range increases (Figure S2D).

Supporting Information

Figure S1.

Sampling locations of anisakid nematodes for molecular species identification. Locations marked with red dots and respective locality abbreviations. Asterisks indicate multiple sampling sites. Abbreviations are listed in Table S1.

doi:10.1371/journal.pone.0028642.s001

(TIF)

Figure S2.

Zoogeographical interpolation approach in four steps. (A) geographical information where transferred onto a grid of 1°×1° covering the globe. Corresponding quadrats marked as occurrence for each species. (B) The final hull resulting from those centroids, which are connected by a line smaller than a multiple of the average line length (α-Parameter). (C) Distribution ranges were defined as the corresponding quadrats intersecting the created polygons. (D) Range was calculated for 25 different α-values ranging from 2 to 50 in a two-step interval. Results were visualized by a continuous colour gradient (Figure 1).

doi:10.1371/journal.pone.0028642.s002

(TIF)

Table S1.

Information on parasites sampling locations and hosts. Sampling locations of the 373 anisakid nematodes used for molecular analyses including abbreviation (Abb.), host and sample size (n). Asterisks mark multiple sampling sites with the same abbreviation (see Figure S1).

doi:10.1371/journal.pone.0028642.s003

(XLS)

Table S2.

Total numbers of identified anisakid nematodes including hosts and sampling location. For location abbreviations see Table S1. Asimss = Anisakis simplex s.s., Apeg = A. pegreffii, Atyp = A. typica, Aphy = A. physeteris.

doi:10.1371/journal.pone.0028642.s004

(XLS)

Acknowledgments

We thank S. Mattiucci and H. W. Palm for providing Anisakis specimen. We are grateful to A. Kijewska for additional data on locality records. We also thank M. Leigh for his comments on an early draft of the manuscript.

Author Contributions

Conceived and designed the experiments: TK JGM SK. Performed the experiments: TK JGM. Analyzed the data: TK JGM. Contributed reagents/materials/analysis tools: TK SK. Wrote the paper: TK JGM SK.

References

  1. 1. Hochberg NS, Hamer DH (2010) Anisakidosis: perils of the deep. Clin Infect Dis 51: 806–812.
  2. 2. Audicana MT, Kennedy MW (2008) Anisakis simplex: from obscure infectious worm to inducer of immune hypersensitivity. Clin Microbiol Rev 21: 360–379.
  3. 3. Chai JY, Murrell KD, Lymbery AJ (2005) Fish-borne parasitic zoonoses: status and issues. Int J Parasitol 35: 1233–1254.
  4. 4. Klimpel S, Palm HW, Rückert S, Piatkowski U (2004) The life cycle of Anisakis simplex in the Norwegian Deep (northern North Sea). Parasitol Res 94: 1–9.
  5. 5. Mattiucci S, Nascetti G (2008) Advances and trends in the molecular systematics of anisakid nematodes, with implications for their evolutionary ecology and the host-parasite co-evolutionary processes. Adv Parasit 66: 47–148.
  6. 6. Klimpel S, Kuhn T, Busch MW, Karl H, Palm HW (2011) Deep-water life cycle of Anisakis paggiae (Nematoda: Anisakidae) in the Irminger Sea indicates kogiid whale distribution in North Atlantic waters. Polar Biol 34: 899–906.
  7. 7. Anderson TJC, Blouin MS, Beech NR (1998) Population biology of parasitic nematodes: applications of genetic markers. Adv Parasitol 41: 219–283.
  8. 8. Gaston KJ (1991) How large is a species' geographic range? Oikos 61: 434–438.
  9. 9. Reddy S, Dávalos L (2003) Geographic sampling bias and its implications for conservation priorities in Africa. J Biogeogr 30: 1719–1727.
  10. 10. Kadmon R, Farber O, Danin A (2004) Effect of roadside bias on the accuracy of predictive maps produced by bio-climate models. Ecol Appl 14: 401–413.
  11. 11. Raedig C, Dormann FC, Hildebrandt A, Lautenbach S (2010) Reassassing neotropical angiosperm distribution patterns based on monographic data: a geometric interpolation approach. Biodivers Conserv 19: 1523–1546.
  12. 12. IUCN (2001) IUCN Red List Categories and Criteria version 3.1. (1994).Cambridge: IUCN Species Survival Commission. 13. IUCN.
  13. 13. IUCN Red List Categories and Criteria. Cambridge: IUCN Species Survival Commission.
  14. 14. Burgmann MA, Fox JC (2003) Bias in species range estimates from minimum convex polygons: implications for conservation and options for improved planning. Anim Conserv 6: 19–28.
  15. 15. Rudolphi CA (1819) Entozoorum synopsis cui accedunt mantesia duplex et indices locupletissimi. 811 p. Berolini.
  16. 16. Krabbe H (1878) Saelernes og Tandhvalernes Spolorme. Overs K Danske Vidensk Selsk Forh 1: 43–51.
  17. 17. Campana-Rouget Y, Biocca E (1955) A new species of Anisakis in a Mediterranean seal. Ann Parasitol Hum Comp 30: 477–480.
  18. 18. Nascetti G, Paggi L, Orecchia P, Smith JW, Mattiucci S, et al. (1986) Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North-East Atlantic. Int J Parasitol 16: 633–640.
  19. 19. Mattiucci S, Nascetti G, Cianchi R, Paggi L, Arduino P, et al. (1997) Genetic and ecological data on the Anisakis simplex complex with evidence for a new species (Nematoda, Ascaridoidea, Anisakidae). J Parasitol 83: 401–416.
  20. 20. Diesing KM (1861) Revision der Nematoden. Sitz-Ber D Akad D Wiss, Math-Nat KI Wien 42: 595–736.
  21. 21. Mattiucci S, Paoletti M, Webb SC (2009) Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: morphological description, genetic relationship between congeners and ecological data. Syst Parasitol 74: 199–217.
  22. 22. Paggi L, Nascetti G, Webb SC, Mattiucci S, Cianchi R, et al. (1998) A new species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae) from beaked whale (Ziphiidae): allozyme and morphological evidence. Syst Parasitol 40: 161–174.
  23. 23. Baylis HA (1923) An ascarid from the sperm whale. Ann Mag nat Hist Ser 9 11: 211–217.
  24. 24. Mattiucci S, Nascetti G, Dailey M, Webb SC, Barros N, et al. (2005) Evidence for a new species of Anisakis Dujardin, 1845: morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Syst Parasitol 61: 157–171.
  25. 25. Dollfus RP (1966) Helminthofaune de Kogia breviceps (Blainville, 1838) cétacé odontocéte. Ann Soc Sci Nat Charente-Marit 4: 3–6.
  26. 26. Abe N, Ohya N, Yanagiguchi R (2005) Molecular characterization of Anisakis pegreffii larvae in Pacific cod in Japan. J Helmithol 79: 303–306.
  27. 27. Abollo E, Gestal C, Pascual S (2001) Anisakis infestation in marine fish and cephalopods from Galician waters: an updated perspective. Parasitol Res 87: 492–499.
  28. 28. Còlon-Llavina MM, Mignucci-Giannoni AA, Mattiucci S, Paoletti M, Nascetti G, et al. (2009) Additional records of metazoan parasites from Caribbean marine mammals, including genetically identified anisakid nematodes. Parasitol Res 105: 1239–1252.
  29. 29. Cross MA, Collins C, Campbell N, Watts PC, Chubb JC, et al. (2007) Levels of intra-host and temporal sequence variation in a large CO1 sub-units from Anisakis simplex sensu stricto (Rudolphi 1809) (Nematoda: Anisakidae): implications for fisheries management. Mar Biol 151: 695–702.
  30. 30. Du C, Zhang L, Shi M, Ming Z, Hu M, et al. (2010) Elucidating the identity of Anisakis larvae from a broad range of marine fishes from the Yellow Sea, China, using a combined electrophoretic-sequencing approach. Electrophoresis 31: 654–658.
  31. 31. Dzido J, Kijewska A, Rockicka M, Światąlska-Koseda A, Rokicki J (2009) Report on anisakid nematodes in polar regions – preliminary results. Polar Science 3: 207–211.
  32. 32. Fang W, Xu S, Zhang S, Wang Y, Chen X, et al. (2010) Multiple primer PCR for the identification of anisakid nematodes from Taiwan Strait. Exp Parasitol 124: 197–201.
  33. 33. Farjallah S, Busi M, Mahjoub MO, Slimane BB, Paggi L, et al. (2008) Molecular characterization of larval anisakid nematodes from marine fish off the Moroccan and Mauritanian coasts. Parasitol Int 57: 430–436.
  34. 34. Farjallah S, Slimane BB, Busi M, Paggi L, Amor N, et al. (2008) Occurrence and molecular identification of Anisakis spp. from the North African coasts of Mediterranean Sea. Parasitol Res 102: 371–379.
  35. 35. Garcia A, Mattiucci S, Damiano S, Santos MN, Nascetti G (2011) Metazoan parasites of swordfish, Xiphias gladius (Pisces: Xiphiidae) from the Atlantic Ocean: implications for host stock identification. ICES J Mar Sci 68: 175–182.
  36. 36. Garcia A, Santos MN, Damiano S, Nascetti G, Mattiucci S (2008) The metazoan parasites of swordfish from Atlantic tropical-equatorial waters. J Fish Biol 73: 2274–2287.
  37. 37. Iñiguez AM, Santos CP, Paulo Vicente AC (2009) Genetic characterization of Anisakis typica and Anisakis physeteris from marine mammals and fish from the Atlantic Ocean off Brazil. Vet Parasitol 165: 350–356.
  38. 38. Karl H, Baumann F, Ostermeyer U, Kuhn T, Klimpel S (2011) Anisakis simplex (s.s.) larvae in wild Alaska salmons: no indication of post-mortem migration from viscera into flesh. Dis Aquat Org 94: 201–209.
  39. 39. Kijewska A, Dzido J, Rokicki J (2009) Mitochondrial DNA of Anisakis simplex s.s. as a potential tool for differentiating populations. J Parasitol 95: 1364–1370.
  40. 40. Kijewska A, Dzido J, Shukhgalter O, Rokicki J (2009) Anisakid parasites caught on the African Shelf. J Parasitol 95: 639–645.
  41. 41. Klimpel S, Kellermanns E, Palm HW (2008) The role of pelagic swarm fish (Myctophidae: Teleostei) in the oceanic life cycle of Anisakis sibling species at the Mid-Atlantic Ridge, Central Atlantic. Parasitol Res 104: 43–53.
  42. 42. Klimpel S, Busch MW, Sutton T, Palm HW (2010) Meso- and bathy-pelagic fish parasites at the Mid-Atlantic Ridge (MAR): Low host specificity and restricted parasite diversity. Deep-Sea Res 57: 596–603.
  43. 43. Klimpel S, Kellermanns E, Palm HW, Moravec F (2007) Zoogeography of fish parasites of the pearlside (Maurolicus muelleri), with genetic evidence of Anisakis simplex (s.s.) from the Mid-Atlantic Ridge. Mar Biol 152: 725–732.
  44. 44. Klimpel S, Palm HW, Busch MW, Kellermanns E (2008) Fish parasites in the bathyal zone: The halosaur Halosauropsis macrochir (Günther, 1878) from the Mid-Atlantic Ridge. Deep-Sea Res 55: 229–235.
  45. 45. Klimpel S, Busch MW, Kuhn T, Rohde A, Palm HW (2010) The Anisakis simplex complex off the South Shetland Islands (Antarctica): endemic populations versus introduction through migratory hosts. Mar Ecol-Prog Ser 403: 1–11.
  46. 46. Marques JF, Cabral HN, Busi M, D'Amelio S (2006) Molecular identification of Anisakis species from Pleuronectiformes off the Portuguese coast. J Helminthol 80: 47–51.
  47. 47. Mattiucci S, Nascetti G (2007) Genetic diversity and infection levels of anisakid nematodes parasitic in fish and marine mammals from Boreal and Austral hemispheres. Vet Parasitol 148: 43–57.
  48. 48. Mattiucci S, Abaunza P, Ramadori L, Nascetti G (2004) Genetic identification of Anisakis larvae in European hake from Atlantic and Mediterranean waters for stock recognition. J Fish Biol 65: 495–510.
  49. 49. Mattiucci S, Nascetti G, Bullini L, Orecchia P, Paggi L (1986) Genetic structure of Anisakis physeteris, and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology 93: 383–387.
  50. 50. Mattiucci S, Paggi L, Nascetti G, Portes Santos C, Costa G, et al. (2002) Genetic markers in the study of Anisakis typica (Diesing, 1860): larval identification and genetic relationship with other species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae). Syst Parasitol 51: 159–170.
  51. 51. Mattiucci S, Farina V, Campbell N, MacKenzie K, Ramos P, et al. (2008) Anisakis spp. larvae (Nematoda: Anisakidae) from Atlantic horse mackerel: Their genetic identification and use as biological tags for host stock characterization. Fish Res 89: 146–151.
  52. 52. Mattiucci S, Farina V, Garcia A, Santos MN, Mariniello L, et al. (2005) Metazoan parasitic infections of Swordfish (Xiphias gladius L., 1758) from the Mediterranean Sea and the Atlantic Gibraltar waters: Implications for Stock Assessment. Col Vol Sci Pap ICCAT 58: 1470–1482.
  53. 53. Mattiucci S, Paggi L, Nascetti G, Abollo E, Webb SC, et al. (2001) Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). Int J Parasitol 16: 633–640.
  54. 54. Mattiucci S, Paggi L, Nascetti G, Ishikura H, Kikuchi K, et al. (1998) Allozyme and morphological identification of Anisakis, Contracaecum and Pseudoterranova from Japanese waters (Nematoda, Ascaridoidea). Syst Parasitol 40: 81–92.
  55. 55. Murphy TM, Berzano M, O'Keeffe SM, Cotter DM, McEvoy , et al. (2010) Anisakid larvae in Atlantic salmon (Salmo salar L.) grilse and post-smolts: molecular identification and histopathology. J Parasitol 96: 77.82.
  56. 56. Nadler SA, D'Amelio S, Dailey MD, Paggi L, Siu S, et al. (2005) Molecular phylogenetics and diagnosis of Anisakis, Pseudoterranova, and Contracaecum from northern Pacific marine mammals. J Parasitol 91: 1413–1429.
  57. 57. Noguera P, Collins C, Bruno D, Pert C, Turnbull A, et al. (2009) Red vent syndrome in wild Atlantic salmon Salmo salar in Scotland is associated with Anisakis simplex sensu stricto (Nematoda: Anisakidae). Dis Aquat Organ 87: 199–215.
  58. 58. Palm HW, Damriyasa IM, Linda , Oka IBM (2008) Molecular genotyping of Anisakis Dujardin, 1845 (Nematoda: Ascaridoidea: Anisakidae) larvae from marine fish of Balinese and Javanese waters, Indonesia. Helminthologia 45: 3–12.
  59. 59. Pontes T, D'Amelio S, Costa G, Paggi L (2005) Molecular characterization of larval anisakid nematodes from marine fishes of Madeira by a PCR-based approach, with evidence for a new species. J Parasitol 91: 1430–1434.
  60. 60. Quiazon KM, Yoshinaga T, Ogawa K, Yukami R (2008) Morphological differences between larvae and in vitro-cultured adults of Anisakis simplex (sensu stricto) and Anisakis pegreffi (Nematoda: Anisakidae). Parasitol Int 57: 483–489.
  61. 61. Quiazon KM, Yoshinaga T, Santos MD, Ogawa K (2009) Identification of larval Anisakis spp. (Nematoda: Anisakidae) in Alaska pollock (Theragra chalcogramma) in northern Japan using morphological and molecular markers. J Parasitol 95: 1227–1232.
  62. 62. Skov J, Kania PW, Olsen MM, Lauridsen JH, Buchmann K (2009) Nematode infections of maricultured and wild fishes in Danish waters: a comparative study. Aquaculture 298: 24–28.
  63. 63. Suzuki J, Murata R, Hosaka M, Araki J (2010) Risk factors for human Anisakis infection and association between the geographic origins of Scomber japonicus and anisakid nematodes. Int J Food Microbiol 137: 88–93.
  64. 64. Umehara A, Kawakami Y, Araki J, Uchida A (2008) Multiplex PCR for the identification of Anisakis simplex sensu stricto, Anisakis pegreffii and the other anisakid nematodes. Parasitol Int 57: 49–53.
  65. 65. Umehara A, Kawakami Y, Araki J, Uchida A, Sugiyama H (2008) Molecular analysis of Japanese Anisakis simplex worms. Southeast Asian J Trop Med Public Health 39: 26–31.
  66. 66. Umehara A, Kawakami Y, Matsui T, Araki J, Uchida A (2006) Molecular identification of Anisakis simplex sensu stricto and Anisakis pegreffii (Nematoda; Anisakidae) from fish and cetacean in Japanese waters. Parasitol Int 55: 267–271.
  67. 67. Umehara A, Kawakami Y, Ooi HK, Uchida A, Ohmae H, et al. (2010) Molecular identification of Anisakis type I larvae isolated from hairtail fish off the coasts of Taiwan and Japan. Int J Food Microbiol 143: 161–165.
  68. 68. Valentini A, Mattiucci S, Bondanelli P, Webb SC, Mignucci-Giannone AA, et al. (2006) Genetic relationships among Anisakis species (Nematoda: Anisakidae) inferred from mitochondrial COX2 sequences, and comparison with allozyme data. J Parasitol 92: 156–166.
  69. 69. Zhang L, Hu M, Shamsi S, Beveridge I, Li H, et al. (2007) The specific identification of anisakid larvae from fishes from the Yellow Sea, China, using mutation scanning-coupled sequence analysis of nuclear ribosomal DNA. Mol Cell Probe 21: 386–390.
  70. 70. Zhu XQ, Gasser RB, Podolska M, Chilton NB (1998) Characterisation of anisakid nematodes with zoonotic potential by nuclear ribosomal DNA sequences. Int J Parasitol 28: 1911–1921.
  71. 71. Zhu XQ, Podolska M, Liu JS, Yu HQ, Chen HH, et al. (2007) Identification of anisakid nematodes with zoonotic potential from Europe and China by single-strand conformation polymorphism analysis of nuclear ribosomal DNA. Parasitol Res 101: 1703–1707.
  72. 72. Pateiro-López B, Rodríguez-Casal A (2010) Generalizing the convex hull of a sample: the R package alphahull. J Stat Softw 34: 1–28.
  73. 73. Iñiguez AM, Carvalho VL, Alves Motta MR, Sousa Nunes Pinheiro DC, Paulo Vicente AC (2011) Genetic analysis of Anisakis typica (Nematoda: Anisakidae) from cetaceans of the northeast coast of Brazil: new data on its definitive hosts. Vet Parasitol 178: 293–299.
  74. 74. Umehara A, Kawakami Y, Matsui T, Araki J, Uchida A (2007) Molecular identification of the etiological agent of the human anisakiasis in Japan. Parasitol Int 56: 211–215.
  75. 75. West KL, Walker WA, Baird RW, White W, Levine G, et al. (2009) Diet of pygmy sperm whales (Kogia breviceps) in the Hawaiian Archipelago. Mar Mammal Sci 25: 931–943.
  76. 76. Allison EH, Perry AL, Badjeck MC, Adger NW, Brown K, et al. (2009) Vulnerability of national economies to the impacts of climate change on fisheries. Fish Fisheries 10: 173–196.
  77. 77. Froese R, Pauly D, editors. (2011) Fishbase. Available from www.fishbase.org (accessed August, 2011).
  78. 78. Gon O, Heemstra PC, editors. (1990) Fishes of the Southern Ocean. Grahamstown: J.L.B. Smith Institute of Ichthyology. 462 p.
  79. 79. Anderson RC (2000) Nematode parasites of vertebrates - Their development and transmission. Wallingford: CAB International. 650 p.
  80. 80. Moravec F (1994) Parasitic nematodes of freshwater fishes of Europe. Praha: Academia. 473 p.
  81. 81. Zhu XQ, Gasser RB, Jacobs DE, Hung GC, Chilton NB (2000) Relationships among some ascaridoid nematodes based on ribosomal DNA sequence data. Parasitol Res 86: 738–744.
  82. 82. Quantum GIS Development Team (2011) Quantum GIS Geographic Information System. Open Source Geospatial Foundation Project (http://qgis.osgeo.org).
  83. 83. Edelsbrunner H, Kirkpatrick DG, Seidel R (1983) On the shape of a set of points in the plane. IEEE Trans Inform Theory IT 29: 551–559.
  84. 84. R Development Core Team (2009) R: a language and environment for statistical computing. Available from www.r-project.org.