First report on molecular identification of Anisakis simplex in Oncorhynchus nerka from the fish market, with taxonomical issues within Anisakidae

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First report on molecular identification of Anisakis simplex in Oncorhynchus nerka from the fish market, with taxonomical issues within Anisakidae

Alina E. Safonova / Anastasia N. Voronova * / Konstantin S. Vainutis

Keywords : Detection, Anisakidosis, Anisakis simplex , Oncorhynchus nerka , Systematics, Taxonomy, ITS, Skrjabinisakis , Peritrachelius

Citation Information : Journal of Nematology. Volume 53, Pages 1-10, DOI: https://doi.org/10.21307/jofnem-2021-023

License : (CC-BY-4.0)

Received Date : 17-November-2020 / Published Online: 26-March-2021

ARTICLE

ABSTRACT

Alive anisakids cause acute gastrointestinal diseases, and dead worms contained in food can provoke sensibilization and allergic reactions in humans. Detected in the purchased minced salmon Oncorhynchus nerka nematodes were identified as Anisakis simplex sensu stricto (Anisakidae). We found that recently published phylogenetic trees (reconstructed using different ribosomal and mitochondrial genetic markers) showed independent clusterization of species recognized in the A. simplex sensu lato species complex. This prompted us to undertake this full-fledged molecular genetics study of anisakids from Kamchatka with phylogenetic reconstructions (NJ/ML) and calculated ranges of interspecific and intergeneric p-distances using ITS1-5.8S-ITS2 sequences. We confirmed that molecular markers based on the ITS region of rDNA were able to recognize ‘pure’ specimens belonging to the cryptic species. We offer new insights into the systematics of anisakids. The genus Anisakis sensu stricto should include Anisakis simplex sensu stricto, Anisakis pegreffii, Anisakis berlandi, Anisakis ziphidarum, and Anisakis nascettii. Presumably, two genera should be restored in the structure of the subfamily Anisakinae: Skrjabinisakis for the species Anisakis paggiae, Anisakis brevispiculata, and Anisakis physeteris; and Peritrachelius for the species Anisakis typica. In addition, we provide the short annotated list of some genera of the family Anisakidae, including their diagnoses.

Graphical ABSTRACT

Commercial fish species Oncorhynchus keta (Walbaum, 1792), O. nerka (Walbaum, 1792), O. tshawytscha (Walbaum, 1792), O. kisutch (Walbaum, 1792), Gadus morhua (Linnaeus, 1758), Clupea harengus (Linnaeus, 1758), Merluccius merluccius (Linnaeus, 1758), anchovy (Engraulis sp.), sardine (Sardina sp.), Scomber sp. from the Atlantic and Pacific Oceans are potentially susceptible to infection by third-stage larvae (L3) of anisakids. Thus, up to 98% of the scomber and 94% of the cod (G. morhua) in the Asian food markets were infected (Setyobudi et al., 2011); almost 34% of fish (mackerel, hake, scomber) from the Atlantic and Mediterranean Sea are reservoir hosts for Anisakis spp. (Debenedetti et al., 2019). In the north of Primorsky Region (Russia) infection intensity by Anisakis spp. of chum salmon (O. keta) and herring (C. pallasii Valenciennes, 1847) is very high (95%) (Kravtsova et al., 2004; Rybnikova et al., 2009). Along with fish, the larvae of anisakids commonly parasitize the edible parts of squids. Larval stages of three species of the genus Anisakis (Dujardin, 1845): A. simplex (Rudolphi, 1809), A. pegreffii (Campana-Rouget and Biocca, 1955), and A. physeteris (Baylis, 1923) are the causative agents of human gastrointestinal anisakidosis (Karmanova, 2007). The whole anisakids and decomposed worms in thermally processed food can provoke severe allergic reactions range from hives and angioneurotic edema to anaphylactic shock (Audicana et al., 2002; Hoshino and Narita, 2011). Anisakidosis is a common disease in Holland, Germany, France, Italy, Spain, Greece, USA, countries of the Pacific coast of Latin America, China, Japan, Thailand, in the Russian Far East; Japan has the highest incidence (up to 1,000 cases per year) (Baird et al., 2014; Nieuwenhuizen and Lopata, 2014; Pravettoni et al., 2012; Yera et al., 2018). In Russia, the problem of anisakidosis remains insufficiently studied. There are several studies of the extensiveness and intensity of fish invasion (Pacific herring, chum salmon, pink salmon, cod, flounder, greenling, mackerel, hake, saury, sea bass, smelts) and the assessment of the viability of nematode larvae (Besprozvannykh et al., 2003; Kravtsova et al., 2004; Rybnikova et al., 2009; Vyalova, 2002). Four cases of A. simplex larvae were described in the human stomach (Kravtsova et al., 2004; Solovieva and Taran, 2000).

In current taxonomy there is a complex of species A. simplex sensu lato (Mattiucci et al., 1997; Nascetti et al., 1983, 1986) which is made up of the three sibling species: A. simplex sensu stricto, A. pegreffii, and A. berlandi (Mattiucci et al., 2014). But, according to the available literature data a clear division of species (A. simplex, A. pegreffii, and A. berlandi) has been repeatedly demonstrated on phylogenetic trees reconstructed using nuclear ITS rDNA (Pekmezci et al., 2014; Tunya et al., 2020) and mitochondrial cox2 gene (Anshary et al., 2014; Barcala et al., 2018; Mattiucci et al., 2009; Setyobudi et al., 2011; Valentini et al., 2006) DNA markers, and finally complete mitochondrial genomes (Yamada et al., 2017). Also A. typica (Diesing, 1860) is suspicious with its uncertain position on different phylogenetic reconstructions (Iñiguez et al., 2009; Sardella and Luque, 2016). Obviously, all this indicating the necessity of the generic revision. In our opinion, forming a species complex A. simplex sensu lato can lead to the loss of some critical information about the individual characteristics of the biology of parasites, and opportunities to influence the changes taking place in their ethology, pathogenesis, evolution, and divergence. A solid and clear taxonomic framework is necessary for examine the basic biology of the parasites, and establish the control system in epidemiology and medicine. Thus, the scope of this study was to accurately identify species of nematodes found in minced salmon O. nerka using the molecular genetics methods and reconstruct phylogenetic relationships within the family Anisakidae which was originally described by Skrjabin and Karokhin (1945) and Skrjabin and Mozgovoy (1973). In the present manuscript, according to Jägerskiöld (1894) we restore the validity of the genus Peritrachelius (Diesing, 1851) and according to Mozgovoy (1951) upgrade subgenus Skrjabinisakis to genus.

Materials and methods

Parasite material

Minced sockeye salmon (frozen) from the Kamchatka Peninsula in the amount of 1 kg was bought at the Vladivostok’s fish market. The organoleptic properties of the product were compromised: among the muscle fibers, inclusions of intestines and membranes were identified during the visual examination of the minced fish. Using a binocular microscope (Micromed MC2 Zoom 1CR) 12 worms (third-stage larvae of nematodes) were extracted and washed in distilled water, then fixed in 70% ethanol. The size of worms ranged from 5 to 15 mm.

PCR amplification

The total DNA was extracted using a QIAamp DNA Mini Kit (QIAGEN, Germany). The ITS1-5,8S-ITS2 region of rDNA was amplified using the classical polymerase chain reaction (PCR) method with specific primers 5′-CCGGGCAAAAGTCGTAACAA-3′ (AscITF) and 5′-ATATGCTTAAATTCAGCGGGT-3′ (R) (Naoki et al., 2010) and DreamTaq Green Master Mix (Thermo Scientific, Lithuania). Cycling conditions consist of a preliminary denaturation at 94°C for 2 min, followed by 30 cycles of denaturation at 94°C for 2 min, annealing at 49°C for 30 sec, elongation at 72°C for 1.5 min, and a final product extension at 72°C for 5 min. The amplification products were visualized on 1.5% agarose gel by ethidium bromide. Amplicons purification was conducted using exonuclease and alkaline phosphotase 1:3 (ExoSap-IT, Thermo Scientific). The PCR products were sequenced by the Sanger method using BigDye Terminator Cycle Sequencing Kit, and specific primers the same as for PCR. After the BigDye Terminator sequencing reaction products were cleaned to remove excess unlabeled dideoxynucleotides using 0.125 М EDTA, CH3COONa, and 96% C2H5OH. The nucleotide sequences were analyzed on an automatic ABI PRIZM 3130 gene analyzer (the center for collective use of FSC EATB FEB RAS). Assembly and alignment of the ITS1-5.8S-ITS rDNA sequences with similar nematode sequences from Genbank (Table 1) were performed in programs FinchTV (Geospiza Inc., Seattle, WA, USA) and MEGA7 (Kumar et al., 2016). Contiguous ITS1-5.8S-ITS2 sequences 863 bp in length were submitted to GenBank NCBI under accession numbers: MT192598, MT192599, MT250915, and MT250916. Genetic distances (p-distance; %) between sequences were calculated using the two-parameter Kimura model and the gamma distribution (0.5).

Table 1.

Sequences of the ITS1-5.8S-ITS2 rDNA region from GenBank used in this study.

10.21307_jofnem-2021-023-t001.jpg

The phylogenetic relationships between nematodes were reconstructed using Neighbor-Joining (NJ) method (Kimura 2-parameter + G) in program MEGA7 (Kumar et al., 2016) and maximum likelihood (ML) method based on the Hasegawa-Kishino-Yano model (chosen using the information criteria of Bayes (BIC)) in PhyML (Guindon et al., 2010) implemented on the ATGC online bioinformatics platform. The tree with the highest log likelihood was shown. Initial trees for the heuristic search were obtained by applying the BioNJ. A Gamma shape parameter was used to model evolutionary rate differences among sites (5 categories (+G, parameter = 0.8895)). The analysis involved 34 nucleotide sequences. There were a total of 877 positions in the final dataset. Ascaris lumbricoides (Linnaeus, 1758) from the family Ascarididae Baird, 1853 was used as an outgroup.

Results and discussion

Morphological analysis of the nematodes according to Mozgovoy (1953) showed that they are from the genus Anisakis. Molecular analysis detected that these nematodes belonged to the species A. simplex. All 12 sequences were identical. No fixed substitutions were found when compared nucleotide sequences of the ITS1-5.8S-ITS2 rDNA region of the studied samples with sequences of this species from different geographical localities (Table 1). Probably, the methods of nematode expansion are so effective that panmixia occurs at the whole-species level, which can be indirectly associated with anthropopression. A. simplex worms have the potential to affect fishing industries, which maintain food and economic stability. The detection of A. simplex helminths in minced fish increases the probability of infection of the human population, so there should be tight control over the distribution of such products in the food markets (Bao et al., 2017). Dead A. simplex larvae can save allergenic properties even after prolonged storage in frozen form (−20 ± 2°C for 11 months) (Rodríguez-Mahillo et al., 2010). WHO recognized A. simplex as the parasite with the largest number of known allergens 14 ‘Ani s’ proteins, demonstrated strong cross-reactivity (especially Ani s 2 (paramyosin) and Ani s 3 (tropomyosin)) to homologous proteins of other nematodes and invertebrates and resistant to pepsin (Aibinu et al., 2019; Baird et al., 2014).

In order to establish phylogenetic relationships among anisakids, the phylogenetic tree was reconstructed based on the sequences of the ITS1-5.8S-ITS2 region with indicated ranges of genetic p-distances (Fig. 1). Two clades according to the subfamilies Contracaecinae (Anisakinae Railliet and Henry, 1912; Mozgovoy and Shakhmatova, 1971) of the family Anisakidae were distinctly distinguished on the tree. Contracaecinae clade contains three species: Contracaecum osculatum (Rudolphi, 1802), C. rudolphii (Hartwich, 1964), and C. bioccai (Mattiucci et al., 2008).

Figure 1:

Phylogenetic relationships of family Anisakidae based on ITS1-5.8S-ITS2 rDNA sequences reconstructed by Maximum Likelihood (ML) and Neighbor-Joining (NJ) methods. Nodal support values are shown based on 1,000 bootstrap replicates (ML/NJ). The tree was drawn to scale, with branch lengths measured in the number of substitutions per site. Accession numbers are given for each species/strain at the end of each sequence. The arrows with generic names are indicating branches. Genetic distances between species and genera are indicated at the junction of the dotted lines. The scale bar represents the number of substitutions per site.

10.21307_jofnem-2021-023-f001.jpg

The Anisakinae clade branched into four distinct, reliably supported subclades. The first subclade (I) includes species of the genus Pseudoterranova (Mozgovoy, 1950). The species A. paggiae (Mattiucci, Nascetti, Dailey, Webb, Barros, Cianchi, and Bullini, 2005), A. brevispiculata (Dollfus, 1966), A. physeteris form the second subclade (II). The third subclade (III) is represented exclusively by the species A. typica. The species A. simplex, A. pegreffii, A. berlandi, A. nascettii (Mattiucci, Paoletti, and Webb, 2009), A. ziphidarum (Paggi, Nascetti, Webb, Mattiucci, Cianchi, and Bullini, 1998) make up the fourth subclade (IV). Within the IV subclade, nematodes are divided into three groups located on separate, well supported branches: (i) A. simplex, hybrid A. simplex×A. pegreffii, A. pegreffii, and A. berlandi; (ii) A. nascetti group; (iii) A. ziphidarum group.

When determine sibling species A. simplex, A. pegreffii, A. berlandi into a single complex A. simplex sensu lato Mattiucci et al. were based on analyses of data collected from allozyme loci (Mattiucci et al., 1986, 2005; Nascetti et al., 1983, 1986; Paggi et al., 1998). Probably, we should once again check the correctness of the assumption that the data on the polymorphism of enzyme loci obtained by electrophoretic methods can be considered typical for the genotype as a whole. Apparently, we are not able to estimate the overall frequency of polymorphism by simply extrapolating the data on the frequency of proteins polymorphism. To choose appropriate gene regions for resolving a particular systematic question among the organisms at a certain categorical level is still a difficult process. Such highly conserved nuclear markers as 18S and 28S rRNA genes are not able to divide the pure specimens of anisakids; moreover, there are not enough data on these markers to reconstruct the phylogenetic relationships of these worms between genera and within the family Anisakidae (Nadler et al., 2005, 2007). On the contrary, high copy and short transcribed rDNA spacers ITS1, ITS2 (Campbell et al., 1994; Hoste et al., 1995; Samson-Himmelstjerna et al., 1997) and the region spanning the ITS1, the 5.8S gene, and the ITS2 of the ribosomal DNA are suitable genetic markers for the identification of nematodes, in particular anisakid species regardless of their stage of development (D’Amelio et al., 2000; Jabbar et al., 2012, 2013). The genetic differentiation among cryptic species of the A. simplex complex is detectable in the ITS region of the rDNA, and this differentiation supports the validity of these species. According to our data, between the sequences of A. simplex and the hybrid form A. simplex × A. pegreffii, and similarly between A. pegreffii and A. simplex × A. pegreffii, genetic p-distances were the same – 0.1%. Genetic p-distances between the ITS1-5.8S-ITS2 regions of A. simplex and A. pegreffii were 0.3%. Genetic p-distances between A. simplex and A. berlandi (former A. simplex C), as well as between A. pegreffii and A. berlandi were 0.7%. Similar p-distances (0.2-0.6%) have been obtained by comparing the ITS sequences of the A. suum (Goeze, 1782) and A. lumbricoides – widely known pig parasites in China (Li et al., 2017). The researchers noted that there was no clear clustering on a phylogenetic tree reconstructed for ascarids between A. suum and A. lumbricoides. However, these worms were considered as the valid ones. The division of the A. suum and A. lumbricoides species has been a crucial factor in understanding the epidemiology of helminths and the possibility of developing methods for controlling parasitic infections for medicine and veterinary. The phylogenetic reconstruction by Tunya et al. (2020) showed a clear division of Anisakis spp. despite the fact that the distances between A. pegreffii and A. simplex were even less (0.1%) than those described in our study. The percentages of genetic differences vary within different worm (and other organisms’) taxa and there is no absolute ‘yardstick’ (Blasco-Costa et al., 2016).

First, we suggest that distances of 0.3 to 0.7% correspond to interspecific ranges in the structure of the family Anisakidae. Second, a powerful argument for the reconsideration of taxonomic structure of A. simplex species complex was phylogenetic analysis performed using the mitogenome sequences of A. pegreffii, A. simplex sensu stricto, and A. berlandi. Three sibling species were distinctly separated from each other and this was also strongly supported by bootstrap values (Yamada et al., 2017). Third, even gene loci coding metallopeptidase enables detection of fixed nucleotide positions (SNP) demonstration that A. pegreffii, A. simplex (s. s.), and A. berlandi are independent (Palomba et al., 2020). Fourth, biology of helminths cannot be the reason for determine A. simplex and A. pegreffii into one species complex. In the in vivo and in vitro studies A. simplex sensu stricto and A. pegreffii species had a differential pathogenic potential and the propensity to trigger allergic reactions (D’Amelio et al., 2020). A. simplex sensu stricto and A. pegreffii are able of hybridizing in the sympatric areas and co-infect the same fish host, but no fertile adult hybrids F1 A. simplex × A. pegreffii have been found (Aibinu et al., 2019; Mattiucci et al., 2016; Mladineo et al., 2017). Fifth, A. simplex and A. pegreffii were considered independent species by Mozgovoy (1953). So, at present, we suppose, that there is absolutely no reason to save the species complex Anisakis simplex sensu lato, because only the precise identification of parasites is essential for their distribution and epidemiology.

Interspecific relationships within the subclade II vary in the range of 4.9 to 6.2%, overlapping with genetic p-distances within species from the subclade IV – 0.3 to 8.0%. Pairwise comparison between species of these two subclades demonstrated a higher level of divergence more likely corresponded to the intergeneric ones – 10.4 to 16.4% (Fig. 1). This is also confirmed by the similar values of p-distances between the anisakids from IV, III, II subclades and species from the other genus Pseudoterranova (subclade I), which vary in the range of 10.8 to 18.6%. In addition to genetic data, the question about the morphological differences of the third-stage larvae of the species included in subclade II has already been raised, according to it, they belong to the second morphotype that distinguishes them from the species from IV and III subclades, comprising larvae of the first morphotype (Iñiguez et al., 2009). Based on these data, we suggest restoring the taxonomic status of the subgenus Skrjabinisakis (Mozgovoy, 1953) upgrading it to the genus level. Thus, this subgenus was first described in the genus Anisakis for species A. physeteris, A. skrjabini, and A. schupakovi (Mozgovoy, 1953). Then after Mozgovoy subgenus Skrjabinisakis supported by Japanese scientists based on the morphology of adults and larva (Oishi et al., 1970). Based on larval morphology and genetic data for A. physeteris and later for A. brevispiculata published by Mattiucci et al. (1986, 2001), respectively, the validity of the subgenus Skrjabinisakis was justified as well. Of these worms species status was confirmed using molecular genetics data only for A. physeteris (Iñiguez et al., 2009).

The species A. typica (subclade III) does not take an unambiguous position on phylogenetic trees reconstructed using various methods with respect to the other Anisakis spp. (Iñiguez et al., 2009; Sardella and Luque, 2016), which may suggest paraphyletic relationships among representatives of the genus Anisakis. The genetic distances for A. typica indicate its remarkable divergence from the other species of the subfamily Anisakinae. Thus, A. typica is very distant from: species of subclade II (17.8-19.7%); species of subclade IV (14.9-17.8%); species of genus Pseudoterranova (18.1-18.6%) (Fig. 1). Based on the intergeneric distances of Pseudoterranova species and A. typica, the latter should be considered in the distinct genus. It is known from the literature that this species was previously put in the genus Peritrachelius (Diesing, 1851; Jägerskiöld, 1894). Until 1882, this genus has been considered separate and included the species P. insignis (synonym for Anisakis insignis) (Dräsche, 1882). Peritrachelius was a subgenus of Ascaris with species Anisakis typica and Anisakis insignis (Jägerskiöld, 1893). Later Mozgovoy (1953) synonymized Peritrachelius with the genus Anisakis (Mozgovoy, 1953). Based on the genetic data analysis, we suggest restoring the genus Peritrachelius for Anisakis typica.

To sum up, the true genus Anisakis sensu stricto probably includes independent species: A. simplex sensu stricto, A. pegreffii, A. berlandi, A. ziphidarum, and A. nascettii. In the structure of the subfamily Anisakinae, for the first time, it was proposed to restore two genera: Skrjabinisakis including species A. paggiae, A. brevispiculata, and A. physeteris, and Peritrachelius – for A. typica.

Short annotated list of some genera of the family Anisakidae (Skrjabin and Karokhin, 1945)

The genus Anisakis (Dujardin, 1845)

Diagnosis: Ventriculus long, S-shaped or straight (juveniles), length is larger four or more times than width. Vulva located in the midbody or anteriorly or posteriorly to it. Spicules long, its length exceeds 1.5 mm.

Type species: Anisakis dussumierii (Beneden, 1870) Baylis, 1920 (sensu Mozgovoy, 1953).

Anisakis simplex (Rudolphi, 1809) Baylis, 1920

Synonyms: Ascaris simplex (Rudolphi, 1809), nec. A. simplex (Dujardin, 1845); Ascaris angulivalvus (Creplin, 1851); Anisaks salaris (Gmelin, 1790) Yamaguti, 1935.

Hosts: Balaenoptera acutirostrata (Lacepede, 1804), B. borealis (Lesson, 1828), B. musculus (Linnaeus, 1758), B. species; Delphinapterus leucas (Pallas, 1776); Delphinus delphis (Linnaeus, 1758), D. species; Eumetopias jubatus (Schreber, 1776); Hyperoodon rostratus (Van Beneden and Gervais, 1880); Lagenorhynchus albirostris (Gray, 1846), L. obscurus (Gray, 1828); Mesoplodon bidens (Sowerby, 1804); Monodon monoceros (Linnaeus, 1758); Phocaena phocaena (Linnaeus, 1758); Pseudoreca crassidens (Owen, 1846); Platanista gangetica (Lebeck, 1801); Globicephala melaena (Traill, 1809); Orcinus orca (Linnaeus, 1758); Stenella coeruleoalba (Meyen, 1833); Halichoerus grypus (Fabricius, 1791); Phoca vitulina (Linnaeus, 1758).

Localization: stomach, intestine, esophagus.

Distribution: Northern waters of the Atlantic Ocean and the Pacific Ocean; Baltic Sea.

Anisakis pegreffii (Campana-Rouget and Biocca, 1955)

Synonym: Anisakis simplex A of Nascetti, Paggi, Orecchia, Smith, Matticucci, and Bullini (1986).

Hosts: Monachus monachus (Hermann, 1779); Delphinus delphis (Linnaeus, 1758); Ziphius cavirostris (Cuvier, 1823); Tursiops truncates (Montagu, 1821).

Localization: digestive tract (first part of the intestine).

Distribution: the Mediterranean Sea (type locality – East coast of Sardinia, near Dorgali) and the waters of the Southern Atlantic Ocean.

Anisakis berlandi (Mattiucci, Cipriani, Webb, Paoletti, Marcer, Bellisario, Gibson and Nascetti, 2014)

Synonym: Anisakis simplex sp. C of Mattiucci et al. (1997).

Hosts: Pseudoreca crassidens (Owen, 1846); Ziphius cavirostris (Cuvier, 1823).

Localization: stomach.

Distribution: Southern waters of the Atlantic Ocean and Northern Pacific Ocean.

Anisakis nascettii (Mattiucci, Paoletti and Webb, 2009)

Synonyms: Anisakis sp. A of Pontes et al. (2005) and Iglesias et al. (2008); Anisakis sp. of Valentini et al. (2006).

Host: Mesoplodon grayi (Von Haast, 1876).

Localization: stomach.

Type locality: New Zealand coast, South Pacific.

Anisakis ziphidarum (Paggi, Nascetti, Webb, Mattiucci, Cianchi and Bullini, 1998)

Hosts: Mesoplodon mirus (True, 1913), M. layardii (Gray, 1865); Ziphius cavirostris (Cuvier, 1823).

Localization: stomach.

Distribution: the Mediterranean Sea and the Southern waters of the Atlantic Ocean.

The genus Skrjabinisakis (Mozgovoy, 1951) stat. n.

Diagnosis: Ventriculus short, straight, length is equal or nearly equal to width. Vulva located in the anterior fourth or third of the body. Spicules short, its length not exceeding 0.67 mm.

Type species: Skrjabinisakis physeteris (Baylis, 1923) comb. n.

Skrjabinisakis physeteris (Baylis, 1923) comb. n.

Synonym: Anisakis physeteris (Baylis, 1923); Anisakis skrjabini (Mozgovoy, 1949).

Hosts: Physeter catodon (Linnaeus, 1758), P. microcephalus (Linnaeus, 1758); Kogia breviceps (Blainville, 1838); K. sima (Owen, 1866); Ziphius cavirostris (Cuvier, 1823).

Localization: stomach.

Distribution: South-Eastern Africa; the Mediterranean Sea and the Atlantic Ocean.

Skrjabinisakis schupakovi (Mozgovoy, 1951)

Synonym: Anisakis species (Schupakov, 1936).

Hosts: Phoca caspica (Gmelin, 1788).

Localization: stomach.

Distribution: Caspian Sea (Chechen Island).

Skrjabinisakis paggiae (Mattiucci, Nascetti, Dailey, Webb, Barros, Cianchi and Bullini, 2005) comb. n.

Synonyms: Anisakis paggiae (Mattiucci, Nascetti, Dailey, Webb, Barros, Cianchi and Bullini, 2005).

Hosts: Kogia breviceps (Blainville, 1838); K. sima (Owen, 1866).

Localization: stomach.

Distribution: the Atlantic Ocean, particularly in its central part.

Skrjabinisakis brevispiculata (Dollfus, 1966) comb. n.

Synonyms: Anisakis brevispiculata (Dollfus, 1966).

Hosts: Kogia breviceps (Blainville, 1838); K. sima (Owen, 1866); Physeter microcephalus (Linnaeus, 1758).

Distribution: the Mediterranean Sea and the waters of the Central Atlantic Ocean.

Localization: stomach.

The genus Peritrachelius (Diesing, 1851)

Diagnosis (sensu Stiles and Hassall, 1899): Ventriculus medium-sized, S-shaped, length is larger nearly five times than width. Vulva located near the midbody. Spicules unequal, long: left – 3.0 mm, right – 0.9 mm length.

Type species: Peritrachelius typicus (Diesing, 1860; Jägerskiöld, 1894).

Peritrachelius typicus (Diesing, 1860; Jägerskiöld, 1894)

Synonyms: Anisakis alexandri (Hsü and Hoeppli, 1933); Anisakis tursiopis (Crusz, 1946); Conocephalus typicus (Diesing, 1860); Anisakis typica (Baylis, 1920; Diesing, 1860).

Hosts: Delphinus delphis (Linnaeus, 1758); Globicephalus melas (Flower, 1885); Phocaena phocaena (Linnaeus, 1758); Phoca sp.; Lagenorhynchus obscurus (Gray, 1828); Sotalia fluviatiles (Gervais and Deville, 1853); Stenella coeruleoalba (Meyen, 1833), S. attenuate (Gray, 1846), S. longirostris (Gray, 1828); Steno bredanensis G. Cuvier in Lesson, 1828; Lagenodelphis hosei (Fraser, 1956).

Distribution: Germany, South-Eastern Africa; warm waters of the Atlantic Ocean.

Localization: stomach.

Acknowledgements

The authors thank Dr. Besprozvannykh V. V. for valuable comments and reviewing drafts of this paper. The authors sincerely thank the fisheries engineer Perebeynos N.L. for consultations and valuable advice throughout this study.

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  7. Besprozvannykh, V. V. , Ermolenko, A. V. and Kamnev, V. D. 2003. Fish of the Peter the Great Bay - a source of human invasion by the nematode Anisakis simplex . Medical Parasitology and Parasitic Diseases 3:18–21, (in Russian).
  8. Blasco-Costa, I. , Cutmore, S. C. , Miller, T. L. and Nolan, M. J. 2016. Molecular approaches to trematode systematics: ‘best practice’ and implications for future study. Systematic Parasitology 93:295–306.
  9. Campbell, A. J. D. , Gasser, R. B. and Chilton, N. B. 1994. Differences in a ribosomal DNA sequence of Strongylus species allows identification of single eggs. International Journal for Parasitology 25:359–365.
  10. D’Amelio, S. , Lombardo, F. , Pizzarelli, A. , Bellini, I. and Cavallero, S. 2020. Advances in omic studies drive discoveries in the biology of Anisakid nematodes. Genes 11:801.
  11. D’Amelio, S. , Mathiopoulos, K. D. , Santos, C. P. , Pugachev, O. N. , Webb, S. C. , Picanço, M. and Paggi, L. 2000. Genetic markers in ribosomal DNA for the identification of members of the genus Anisakis (Nematoda: Ascaridoidea) defined by polymerase-chain-reaction-based restriction fragment length polymorphism. International Journal for Parasitology 30:223–226.
  12. Debenedetti, Á. L. , Madrid, E. , Trelis, M. , Codes, F. J. , Gil-Gómez, F. , Sáez-Durán, S. and Fuentes, M. V. 2019. Prevalence and risk of anisakid larvae in fresh fish frequently consumed in Spain: an overview. Fishes 4:1–16.
  13. Dräsche, R. F. 1882. Zur Charakteristik der Nematoden – Gattung Peritrachelius Diesing. (Tafel 12). Verhandlungen der Zoologisch-Botanischen Gesellschaft in Wien. Frueher: Verh.des Zoologisch-Botanischen Vereins in Wien.
  14. Guindon, S. , Dufayard, J. F. , Lefort, V. , Anisimova, M. , Hordijk, W. and Gascuel, O. 2010. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Systematic Biology 59:307–321.
  15. Hoshino, C. and Narita, M. 2011. Anisakis simplex-induced anaphylaxis. Journal of Infection and Chemotherapy 17:544–546.
  16. Hoste, H. , Chilton, N. B. , Gasser, R. B. and Beveridge, I. 1995. Differences in the second internal transcribed spacer (ribosomal DNA) between five species of Trichostrongylus (Nematoda: Trichostrongylidae). International Journal for Parasitology 25:75–80.
  17. Iñiguez, A. M. , Santos, C. P. and Vicente, A. C. P. 2009. Genetic characterization of Anisakis typica and Anisakis physeteris from marine mammals and fish from the Atlantic Ocean off Brazil. Veterinary Parasitology 165:350–356.
  18. Jabbar, A. , Fong, R. W. J. , Kok, K. X. , Lopata, A. L. , Gasser, R. B. and Beveridge, I. 2013. Molecular characterization of anisakid nematode larvae from 13 species of fish from Western Australia. International Journal of Food Microbiology 161:247–253.
  19. Jabbar, A. , Khoon, A. T. W. , Hui, T. X. , Schaeffner, B. C. , Jex, A. R. , Nolan, M. J. , Lopata, A. , Gasser, R. B. and Beveridge, I. 2012. Mutation scanning-based analysis of anisakid larvae from Sillago flindersi from Bass Strait, Australia. Electrophoresis 33:499–505.
  20. Jägerskiöld, L. 1894. Bidrag Till Kannedomen Om Nematoderna, Stockholm.
  21. Karmanova, I. V. 2007. On some species of parasites of salmonids of Kamchatka, potentially hazardous to human health. Bulletin of Kamchatka State Technical University 6:31–34 (in Russian).
  22. Kravtsova, V. O. , Tsarenko, S. S. , Filimonova, N. B. and Gagalchy, N. M. 2004. Anisakiasis in Primorsky Region. Pacific Medical Journal 3:5–7.
  23. Kumar, S. , Stecher, G. and Tamura, K. 2016. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Molecular Biology and Evolution 33:1870–1874.
  24. Li, K. , Luo, H. , Zhang, H. , Mehmood, K. , Shahzad, M. , Zhang, L. and Li, J. 2017. Analysis of the internal transcribed spacer region of Ascaris suum and Ascaris lumbricoides derived from free range Tibetan pigs. Mitochondrial DNA Part A 29:624–628.
  25. Mattiucci, S. , Paoletti, M. and Webb, S. C. 2009. Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: morphological description, genetic relationships between congeners and ecological data. Systematic Parasitology 74:199–217.
  26. Mattiucci, S. , Nascetti, G. , Bullini, L. , Orecchia, P. and Paggi, L. 1986. Genetic structure of Anisakis physeteris, and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology 93:383–38.
  27. Mattiucci, S. , Nascetti, G. , Dailey, M. , Webb, S. C. , Barros, N. B. , Cianchi, R. and Bullini, L. 2005. Evidence for a new species of Anisakis Dujardin, 1845: morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Systematic Parasitology 61:157–171.
  28. Mattiucci, S. , Acerra, V. , Paoletti, M. , Cipriani, P. , Levsen, A. , Webb, S. C. , Canestrelli, D. and Nascetti, G. 2016. No more time to stay ‘single’ in the detection of Anisakis pegreffii, A. simplex (s. s.) and hybridization events between them: a multi-marker nuclear genotyping approach. Parasitology 143:998–1011.
  29. Mattiucci, S. , Cipriani, P. , Webb, S. C. , Paoletti, M. , Marcer, F. , Bellisario, B. , Gibson, D. I. and Nascetti, G. 2014. Genetic and morphological approaches distinguishing the three sibling species of the Anisakis simplex species complex, with a species designation as Anisakis berlandi n. sp. for A. simplex sp. C (Nematoda: Anisakidae). The Journal of Parasitology 100:199–214.
  30. Mattiucci, S. , Paggi, L. , Nascetti, G. , Abollo, E. , Webb, S. C. , Pascual, S. , Cianchi, R. and Bullini, L. 2001. Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). International Journal for Parasitology 31:9–14.
  31. Mattiucci, S. , Paoletti, M. , Olivero-Verbel, J. , Baldiris, R. , Arroyo-Salgado, B. , Garbin, L. , Navone, G. and Nascetti, G. 2008. Contracaecum bioccai n. sp. from the brown pelican Pelecanus occidentalis (L.) in Colombia (Nematoda: Anisakidae): morphology, molecular evidence and its genetic relationship with congeners from fish-eating birds. Systematic Parasitology 69:101–121.
  32. Mattiucci, S. , Nascetti, G. , Cianchi, R. , Paggi, L. , Arduino, P. , Margolis, L. , Brattey, J. , Webb, S. , D’Amelio, S. , Orecchia, P. and Bullini, L. 1997. Genetic and ecological data on the Anisakis simplex complex with evidence for a new species (Nematoda Ascaridoidea, Anisakidae). The Journal of Parasitology 83:401–416.
  33. Mladineo, I. , Trumbić, Ž. , Radonić, I. , Vrbatović, A. , Hrabar, J. and Bušelić, I. 2017. Anisakis simplex complex: ecological significance of recombinant genotypes in an allopatric area of the Adriatic Sea inferred by genome-derived simple sequence repeats. International Journal for Parasitology 47:215–223.
  34. Mozgovoy, A. A. 1953. Fundamentals of nematodology. Ascaridates of animals and humans and diseases caused by them Publishing House of the Academy of Sciences of the USSR, Moscow (in Russian).
  35. Nadler, S. A. , Carreno, R. A. , Mejía , Madrid, H. , Ullberg, J. , Pagan, C. , Houston, R. and Hugot, J. P. 2007. Molecular phylogeny of clade III nematodes reveals multiple origins of tissue parasitism. Parasitology 134:1421–1442.
  36. Nadler, S. A. , D’Amelio, S. , Dailey, M. D. , Paggi, L. , Siu, S. and Sakanari, J. A. 2005. Molecular Phylogenetics and Diagnosis of Anisakis, Pseudoterranova, and Contracaecum from Northern Pacific Marine Mammals. The Journal of Parasitology 91:1413–1429.
  37. Naoki, A. , Yoshimura, Y. , Naoki, T. , Yamada, M. , Tatsuya, T. , Onishi, K. and Uchikawa, R. 2010. Ascariasis in Japan: Is pig-derived Ascaris infecting humans?. Japanese Journal of Infectious Diseases 63:447–458.
  38. Nascetti, G. , Paggi, L. , Orecchia, P. , Mattiucci, S. and Bullini, L. 1983. Two sibling species within Anisakis simplex (Ascaridida: Anisakidae). Parassitologia 25:306–307.
  39. Nascetti, G. , Paggi, L. , Orecchia, P. , Smith, J. W. , Mattiucci, S. and Bullini, L. 1986. Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North- East Atlantic. International Journal for Parasitology 16:33–40.
  40. Nieuwenhuizen, N. E. and Lopata, A. L. 2014. Allergic reactions to Anisakis found in fish. Current Allergy and Asthma Reports 14:455.
  41. Oishi, K. , Oka, S. and Josho, S. 1970. An introduction to food hygiene with regard to the Anisakis larva. Fisheries Research Board of Canada Translation Series No.1414, Fisheries Research Board of Canada Biological Station Nanaimo, BC.
  42. Paggi, L. , Nascetti, G. , Webb, S. C. , Mattiucci, S. , Cianchi, R. and Bullini, L. 1998. A new species of Anisakis Dujardin, 1845 (Nematoda, Anisakidae) from beaked whales (Ziphiidae): allozyme and morphological evidence. Systematic Parasitology 40:161–174.
  43. Palomba, M. , Paoletti, M. , Webb, S. and Mattiucci, S. 2020. A novel nuclear marker and development of an ARMS-PCR assay targeting the metallopeptidase 10 (nas 10) locus to identify the species of the Anisakis simplex (s. l.) complex (Nematoda, Anisakidae). Parasite 27:39.
  44. Pekmezci, G. Z. , Onuk, E. E. , Bolukbas, C. S. , Yardimci, B. , Gurler, A. T. , Acici, M. and Umur, S. 2014. Molecular identification of Anisakis species (Nematoda: Anisakidae) from marine fishes collected in Turkish waters. Veterinary Parasitology 201:82–94.
  45. Pravettoni, V. , Primavesi, L. and Piantanida, M. 2012. Anisakis simplex: current knowledge. European Annals of Allergy and Clinical Immunology 44:150–156.
  46. Rodríguez-Mahillo, A. I. , González-Muñoz, M. , de las Heras, C. , Tejada, M. and Moneo, I. 2010. Quantification of Anisakis simplex allergens in fresh, long-term frozen, and cooked fish muscle. Foodborne Pathogens and Disease 7:967–973.
  47. Rybnikova, I. G. , Pushnikova, G. M. and Shvetsova, L. S. 2009. Infestation of Pacific herring with nematodes larvae. Scientific Studies of Far Eastern State Technical Fisheries University (in Russian).
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  49. Sardella, C. J. R. and Luque, J. L. 2016. Morphological and molecular diagnostic of Anisakis typica and Anisakis brevispiculata of southeastern coast of Brazil. Brazilian Journal of Veterinary Medicine 38:87–98.
  50. Setyobudi, E. , Jeon, C. H. , Lee, C. H. , Seong, K. B. and Kim, J. H. 2011. Occurrence and identification of Anisakis spp. (Nematoda: Anisakidae) isolated from chum salmon (Oncorhynchus keta) in Korea. Parasitology Research 108:585–592.
  51. Skrjabin, K. I. and Karokhin, V. I. 1945. Reconstruction of the taxonomy of nematodes from the suborder Ascaridata Skrjabin, 1915. Scientific Reports USSR 48:318–320 (in Russian).
  52. Skrjabin, K. I. and Mozgovoy, A. A. 1973. Justification of a new superfamily of nematodes Ascaridioidea superfam. n. Parasitology, VII 6:518–520 (in Russian).
  53. Solovieva, G. F. and Taran, N. A. 2000. Two cases of revealing of Anisakis simplex (Ascaridina, Anisakidae) larvae in human stomach. TINRO (in Russian).
  54. Tunya, R. , Wongsawad, C. , Wongsawad, P. and Chai, J. Y. 2020. Morphological and molecular characteristics of Anisakis typica larvae in two species of Threadfin Bream, Nemipterus hexodonand and N. japonicus, from the Gulf of Thailand. The Korean Journal of Parasitology 58:15–25.
  55. Valentini, A. , Mattiucci, S. , Bondanelli, P. , Webb, S. C. , Mignucci-Giannone, A. A. , Colom-Llavina, M. M. and Nascetti, G. 2006. Genetic relationships among Anisakis species (Nematoda: Anisakidae) inferred from mitochondrial cox2 sequences, and comparison with allozyme data. The Journal of Parasitology 92:156–166.
  56. Vyalova, G. P. 2002. Distribution of Anisakis simplex larvae in the musculature of salmonids and peculiarities of their localization in the pink salmon population. TINRO News 131:430–438 (in Russian).
  57. Yamada, A. , Ikeda, N. and Ono, H. 2017. The complete mitochondrial genome of Anisakis pegreffii Campana-Rouget and Biocca, 1955, (Nematoda, Chromadorea, Rhabditida, Anisakidae) – clarification of mitogenome sequences of the Anisakis simplex species complex. Mitochondrial DNA Part B: Resources 2:240–241.
  58. Yera, H. , Frealle, E. , Dutoit, E. and Dupouy-Camet, J. 2018. A national retrospective survey of anisakidosis in France (2010-2014): decreasing incidence, female predominance and emerging allergic potential. Parasite 25:23.
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FIGURES & TABLES

Figure 1:

Phylogenetic relationships of family Anisakidae based on ITS1-5.8S-ITS2 rDNA sequences reconstructed by Maximum Likelihood (ML) and Neighbor-Joining (NJ) methods. Nodal support values are shown based on 1,000 bootstrap replicates (ML/NJ). The tree was drawn to scale, with branch lengths measured in the number of substitutions per site. Accession numbers are given for each species/strain at the end of each sequence. The arrows with generic names are indicating branches. Genetic distances between species and genera are indicated at the junction of the dotted lines. The scale bar represents the number of substitutions per site.

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REFERENCES

  1. Aibinu, I. E. , Smooker, P. M. and Lopata, A. L. 2019. Anisakis Nematodes in Fish and Shellfish – from infection to allergies. International Journal for Parasitology: Parasites and Wildlife 9:384–393.
  2. Anshary, H. , Sriwulan , Freeman, M. A. and Ogawa, K. 2014. Occurrence and molecular identification of Anisakis Dujardin, 1845 from marine fish in southern Makassar Strait, Indonesia. Korean Journal of Parasitology 52:9–19.
  3. Audicana, M. T. , Ansotegui, I. J. , de Corres, L. F. and Kennedy, M. W. 2002. Anisakis simplex: dangerous – dead and alive? Trends in Parasitology 18:20–25.
  4. Baird, F. J. , Gasser, R. B. , Jabbar, A. and Lopata, A. L. 2014. Foodborne anisakiasis and allergy. Molecular and Cellular Probes 28:167–174.
  5. Bao, M. , Pierce, G. J. , Strachan, N. J. C. , Martinez, C. , Fernandez, R. and Theodossiou, I. 2017. Consumers’ attitudes and willingness to pay for Anisakis-free fish in Spain. Fisheries Research 202:149–160.
  6. Barcala, E. , Ramilo, A. , Ortega, N. , Pico, G. , Abollo, E. , Pascual, S. and Muñoz, P. 2018. Occurrence of Anisakis and Hysterothylacium larvae in commercial fish from Balearic Sea (Western Mediterranean Sea). Parasitology Research 117:4003–4012.
  7. Besprozvannykh, V. V. , Ermolenko, A. V. and Kamnev, V. D. 2003. Fish of the Peter the Great Bay - a source of human invasion by the nematode Anisakis simplex . Medical Parasitology and Parasitic Diseases 3:18–21, (in Russian).
  8. Blasco-Costa, I. , Cutmore, S. C. , Miller, T. L. and Nolan, M. J. 2016. Molecular approaches to trematode systematics: ‘best practice’ and implications for future study. Systematic Parasitology 93:295–306.
  9. Campbell, A. J. D. , Gasser, R. B. and Chilton, N. B. 1994. Differences in a ribosomal DNA sequence of Strongylus species allows identification of single eggs. International Journal for Parasitology 25:359–365.
  10. D’Amelio, S. , Lombardo, F. , Pizzarelli, A. , Bellini, I. and Cavallero, S. 2020. Advances in omic studies drive discoveries in the biology of Anisakid nematodes. Genes 11:801.
  11. D’Amelio, S. , Mathiopoulos, K. D. , Santos, C. P. , Pugachev, O. N. , Webb, S. C. , Picanço, M. and Paggi, L. 2000. Genetic markers in ribosomal DNA for the identification of members of the genus Anisakis (Nematoda: Ascaridoidea) defined by polymerase-chain-reaction-based restriction fragment length polymorphism. International Journal for Parasitology 30:223–226.
  12. Debenedetti, Á. L. , Madrid, E. , Trelis, M. , Codes, F. J. , Gil-Gómez, F. , Sáez-Durán, S. and Fuentes, M. V. 2019. Prevalence and risk of anisakid larvae in fresh fish frequently consumed in Spain: an overview. Fishes 4:1–16.
  13. Dräsche, R. F. 1882. Zur Charakteristik der Nematoden – Gattung Peritrachelius Diesing. (Tafel 12). Verhandlungen der Zoologisch-Botanischen Gesellschaft in Wien. Frueher: Verh.des Zoologisch-Botanischen Vereins in Wien.
  14. Guindon, S. , Dufayard, J. F. , Lefort, V. , Anisimova, M. , Hordijk, W. and Gascuel, O. 2010. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Systematic Biology 59:307–321.
  15. Hoshino, C. and Narita, M. 2011. Anisakis simplex-induced anaphylaxis. Journal of Infection and Chemotherapy 17:544–546.
  16. Hoste, H. , Chilton, N. B. , Gasser, R. B. and Beveridge, I. 1995. Differences in the second internal transcribed spacer (ribosomal DNA) between five species of Trichostrongylus (Nematoda: Trichostrongylidae). International Journal for Parasitology 25:75–80.
  17. Iñiguez, A. M. , Santos, C. P. and Vicente, A. C. P. 2009. Genetic characterization of Anisakis typica and Anisakis physeteris from marine mammals and fish from the Atlantic Ocean off Brazil. Veterinary Parasitology 165:350–356.
  18. Jabbar, A. , Fong, R. W. J. , Kok, K. X. , Lopata, A. L. , Gasser, R. B. and Beveridge, I. 2013. Molecular characterization of anisakid nematode larvae from 13 species of fish from Western Australia. International Journal of Food Microbiology 161:247–253.
  19. Jabbar, A. , Khoon, A. T. W. , Hui, T. X. , Schaeffner, B. C. , Jex, A. R. , Nolan, M. J. , Lopata, A. , Gasser, R. B. and Beveridge, I. 2012. Mutation scanning-based analysis of anisakid larvae from Sillago flindersi from Bass Strait, Australia. Electrophoresis 33:499–505.
  20. Jägerskiöld, L. 1894. Bidrag Till Kannedomen Om Nematoderna, Stockholm.
  21. Karmanova, I. V. 2007. On some species of parasites of salmonids of Kamchatka, potentially hazardous to human health. Bulletin of Kamchatka State Technical University 6:31–34 (in Russian).
  22. Kravtsova, V. O. , Tsarenko, S. S. , Filimonova, N. B. and Gagalchy, N. M. 2004. Anisakiasis in Primorsky Region. Pacific Medical Journal 3:5–7.
  23. Kumar, S. , Stecher, G. and Tamura, K. 2016. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Molecular Biology and Evolution 33:1870–1874.
  24. Li, K. , Luo, H. , Zhang, H. , Mehmood, K. , Shahzad, M. , Zhang, L. and Li, J. 2017. Analysis of the internal transcribed spacer region of Ascaris suum and Ascaris lumbricoides derived from free range Tibetan pigs. Mitochondrial DNA Part A 29:624–628.
  25. Mattiucci, S. , Paoletti, M. and Webb, S. C. 2009. Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: morphological description, genetic relationships between congeners and ecological data. Systematic Parasitology 74:199–217.
  26. Mattiucci, S. , Nascetti, G. , Bullini, L. , Orecchia, P. and Paggi, L. 1986. Genetic structure of Anisakis physeteris, and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology 93:383–38.
  27. Mattiucci, S. , Nascetti, G. , Dailey, M. , Webb, S. C. , Barros, N. B. , Cianchi, R. and Bullini, L. 2005. Evidence for a new species of Anisakis Dujardin, 1845: morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Systematic Parasitology 61:157–171.
  28. Mattiucci, S. , Acerra, V. , Paoletti, M. , Cipriani, P. , Levsen, A. , Webb, S. C. , Canestrelli, D. and Nascetti, G. 2016. No more time to stay ‘single’ in the detection of Anisakis pegreffii, A. simplex (s. s.) and hybridization events between them: a multi-marker nuclear genotyping approach. Parasitology 143:998–1011.
  29. Mattiucci, S. , Cipriani, P. , Webb, S. C. , Paoletti, M. , Marcer, F. , Bellisario, B. , Gibson, D. I. and Nascetti, G. 2014. Genetic and morphological approaches distinguishing the three sibling species of the Anisakis simplex species complex, with a species designation as Anisakis berlandi n. sp. for A. simplex sp. C (Nematoda: Anisakidae). The Journal of Parasitology 100:199–214.
  30. Mattiucci, S. , Paggi, L. , Nascetti, G. , Abollo, E. , Webb, S. C. , Pascual, S. , Cianchi, R. and Bullini, L. 2001. Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). International Journal for Parasitology 31:9–14.
  31. Mattiucci, S. , Paoletti, M. , Olivero-Verbel, J. , Baldiris, R. , Arroyo-Salgado, B. , Garbin, L. , Navone, G. and Nascetti, G. 2008. Contracaecum bioccai n. sp. from the brown pelican Pelecanus occidentalis (L.) in Colombia (Nematoda: Anisakidae): morphology, molecular evidence and its genetic relationship with congeners from fish-eating birds. Systematic Parasitology 69:101–121.
  32. Mattiucci, S. , Nascetti, G. , Cianchi, R. , Paggi, L. , Arduino, P. , Margolis, L. , Brattey, J. , Webb, S. , D’Amelio, S. , Orecchia, P. and Bullini, L. 1997. Genetic and ecological data on the Anisakis simplex complex with evidence for a new species (Nematoda Ascaridoidea, Anisakidae). The Journal of Parasitology 83:401–416.
  33. Mladineo, I. , Trumbić, Ž. , Radonić, I. , Vrbatović, A. , Hrabar, J. and Bušelić, I. 2017. Anisakis simplex complex: ecological significance of recombinant genotypes in an allopatric area of the Adriatic Sea inferred by genome-derived simple sequence repeats. International Journal for Parasitology 47:215–223.
  34. Mozgovoy, A. A. 1953. Fundamentals of nematodology. Ascaridates of animals and humans and diseases caused by them Publishing House of the Academy of Sciences of the USSR, Moscow (in Russian).
  35. Nadler, S. A. , Carreno, R. A. , Mejía , Madrid, H. , Ullberg, J. , Pagan, C. , Houston, R. and Hugot, J. P. 2007. Molecular phylogeny of clade III nematodes reveals multiple origins of tissue parasitism. Parasitology 134:1421–1442.
  36. Nadler, S. A. , D’Amelio, S. , Dailey, M. D. , Paggi, L. , Siu, S. and Sakanari, J. A. 2005. Molecular Phylogenetics and Diagnosis of Anisakis, Pseudoterranova, and Contracaecum from Northern Pacific Marine Mammals. The Journal of Parasitology 91:1413–1429.
  37. Naoki, A. , Yoshimura, Y. , Naoki, T. , Yamada, M. , Tatsuya, T. , Onishi, K. and Uchikawa, R. 2010. Ascariasis in Japan: Is pig-derived Ascaris infecting humans?. Japanese Journal of Infectious Diseases 63:447–458.
  38. Nascetti, G. , Paggi, L. , Orecchia, P. , Mattiucci, S. and Bullini, L. 1983. Two sibling species within Anisakis simplex (Ascaridida: Anisakidae). Parassitologia 25:306–307.
  39. Nascetti, G. , Paggi, L. , Orecchia, P. , Smith, J. W. , Mattiucci, S. and Bullini, L. 1986. Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North- East Atlantic. International Journal for Parasitology 16:33–40.
  40. Nieuwenhuizen, N. E. and Lopata, A. L. 2014. Allergic reactions to Anisakis found in fish. Current Allergy and Asthma Reports 14:455.
  41. Oishi, K. , Oka, S. and Josho, S. 1970. An introduction to food hygiene with regard to the Anisakis larva. Fisheries Research Board of Canada Translation Series No.1414, Fisheries Research Board of Canada Biological Station Nanaimo, BC.
  42. Paggi, L. , Nascetti, G. , Webb, S. C. , Mattiucci, S. , Cianchi, R. and Bullini, L. 1998. A new species of Anisakis Dujardin, 1845 (Nematoda, Anisakidae) from beaked whales (Ziphiidae): allozyme and morphological evidence. Systematic Parasitology 40:161–174.
  43. Palomba, M. , Paoletti, M. , Webb, S. and Mattiucci, S. 2020. A novel nuclear marker and development of an ARMS-PCR assay targeting the metallopeptidase 10 (nas 10) locus to identify the species of the Anisakis simplex (s. l.) complex (Nematoda, Anisakidae). Parasite 27:39.
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