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. 2021 Mar 16;23:e00120. doi: 10.1016/j.fawpar.2021.e00120

Occurrence and abundance of zoonotic nematodes in snapper Chrysophrys auratus, a popular table fish from Australian and New Zealand waters

Md Shafaet Hossen a,b,⁎, Skye Wassens c, Shokoofeh Shamsi a
PMCID: PMC8010209  PMID: 33817358

Abstract

In Australia and New Zealand (NZ), snapper Chrysophrys auratus is known for delicate mild flavoured flesh and is a favoured species to serve raw as sashimi or in sushi. The diet of snapper includes a variety of intermediate hosts of larval nematodes, and as a result, snapper has potential to become highly infected with zoonotic/non-zoonotic nematodes. The aims of this study were to survey nematodes in snapper from Australia and New Zealand waters and to identify nematode species using combined morphological and molecular methods. The zoonotic potential of nematodes identified in this study are discussed. A total of 112 snapper were purchased from the Sydney fish market, New South Wales, Australia. Fish were dissected and only the visceral content and digestive tract were examined for nematode infection. Parasites were initially identified by the microscopic method as four different types belonging to the families Anisakidae (Anisakis types I & III, and Terranova type II) and Cucullanidae (Dichelyne spp.). All Anisakidae nematodes were at infective stages. Species-level identification was actualised through sequencing of the internal transcribed spacer (ITS–1, 5.8S, ITS–2) regions. The Anisakis types I & III were confirmed as Anisakis pegreffii and A. brevispiculata, respectively of which A. pegreffii is considered globally as a zoonotic nematode. The specific identification of Terranova type II and Dichelyne spp. was not possible as no comparable sequence data were available in GenBank. The phylogenetic tree clustered Anisakis types I & III with A. pegreffii and A. brevispiculata, respectively; Terranova type II sequences as a separate clade with previously identified larval and adult Terranova and Pseudoterranova species. Based on phylogenetic analyses the present Cucullanid specimens were assigned herein as Dichelyne cf. pleuronectidis, and an unknown species Dichelyne sp. 1. This study represents the first host record globally for zoonotic Anisakid nematodes in this popularly consumed table fish and a new region record for D. cf. pleuronectidis and Dichelyne sp. 1. Further investigation is required, using more comprehensive parasite detection and recovery methods, to assess the health risk these nematodes may pose to human and fish health in Australia/NZ.

Keywords: Anisakidae, Cucullanidae, Fish, Zoonotic nematodes, Seafood safety, Public health

Highlights

  • •

    First evidence of zoonotic Anisakis pegreffii infection in snapper Chrysophrys auratus.

  • •

    First evidence of A. brevispiculata and Terranova type II infection in snapper.

  • •

    Snapper was the verified host for Dichelyne cf. pleuronectidis and Dichelyne sp. 1.

  • •

    Infective stage of A. pegreffii was the highest prevalent nematode found in snapper.

1. Introduction

The snapper Chrysophrys auratus (Perciformes: Sparidae) is important to Australian recreational, commercial (Kailola et al., 1993; Norriss and Crisafulli, 2010), and charter industries. Snapper is also an important food species for indigenous Australians (FRDC, 2019) and is a revered fish within their spirituality and traditional knowledge (Prober et al., 2011). It is considered one of the nation's most important and iconic fish (Fowler et al., 2017; Norriss and Crisafulli, 2010) and is a popular and well-known table species that returns high prices at the retail level (FRDC, 2019). Australia/New Zealand (NZ), as multicultural nations, have embraced a range of imported cultural cuisines, including sushi and sashimi. According to the Kobe Jones Blog (Kobe Jones, 2019) snapper in Australia is considered suitable for sashimi and is regularly recommended by Australian chefs as compatible to use in raw fish dishes (Allen, 2019; Kwong, 2015).

Snapper is distributed in warm to temperate Indo-Pacific waters which occur off Australia/NZ (Paulin, 1990). Snapper is a long-lived predatory fish (Norriss and Crisafulli, 2010), and feed predominantly on crustaceans and small fish which are the intermediate hosts of many nematode species (Godfriaux, 1969; Gregori et al., 2015; Hurst, 1984). As a result, snapper have the potential to become highly parasitised. There have been no contemporary studies using combined morphological and molecular tools to accurately describe nematode parasites of snapper in Australian/NZ waters (Table 1). The most detailed parasitological examination of snapper nematodes was conducted by Johnston and Mawson (1945) and Sharples and Evans, 1995a, Sharples and Evans, 1995b, Sharples and Evans, 1995c in Australia and NZ waters, respectively. Both studies relied on morphological methods for nematode identification. However, few morphological markers of taxonomic significance for reliable species identification of larval nematodes are available (McManus and Bowles, 1996). During the last two decades, the application of molecular tools for the identification of nematodes has greatly advanced their taxonomy. Therefore, the primary aim of this study was to investigate the potential of snapper from the waters of Australia/NZ to be infected with nematode parasites. The secondary aim was to accurately identify these nematode parasites using combined morphological and molecular tools.

Table 1.

Previous records of nematodes identified from the snapper Chrysophrys auratus in Australia and New Zealand.

Nematode Microhabitat Family Location Reference
Anisakis sp. larva Encapsulated on viscera, mesenteries, and peritoneum of the body cavity Anisakidae Hauraki Gulf (Okakari Point and Kawau Bay), NZ Sharples and Evans, 1995a, Sharples and Evans, 1995b
Cucullanus sp. Intestine Cucullanidae Hauraki Gulf (Okakari Point and Kawau Bay), NZ Sharples and Evans, 1995a, Sharples and Evans, 1995b
Philometra lateolabracis Gonads Philometridae Hauraki Gulf (Okakari Point and Kawau Bay), NZ Sharples and Evans, 1995a, Sharples and Evans, 1995b
P. lateolabracis – Philometridae NZ Hine and Anderson (1981)
Anisakis sp. larva – Anisakidae NZ Brunsdon (1956)
Dichelyne cnidoglanis – Cucullanidae NZ Brunsdon (1953)
Hysterothylacium sp. – Raphidascarididae NZ Brunsdon (1953)
Dichelyne sheardi – Cucullanidae Outer harbour, SA Johnston and Mawson (1949)
Dichelyne sheardi – Cucullanidae Glenelg, SA Johnston and Mawson (1945)
Echinocephalus uncinatus larva Mesentery or omentum Gnathostomatidae Glenelg, SA Johnston and Mawson (1945)
Anisakis larval type⁎ – Anisakidae Glenelg and Cape Jervis, SA Johnston and Mawson (1945)
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‘–’ indicates no information available; Abbreviations: NZ = New Zealand, SA = South Australia.

⁎

Anisakis marina (= Capsularia marina; Stomachus marinus) has been reported from various hosts across Australian coasts (Johnston and Mawson, 1944, Johnston and Mawson, 1945, Johnston and Mawson, 1949). In these reports “marina” mostly refers to larval stage of the nematode. Therefore, A. marina is not considered a valid taxon but regarded as Anisakis larval type. Both Capsularia and Stomachus have later been synonymised with Anisakis (Shamsi, 2014).

2. Materials and methods

2.1. Fish collection

A total of 112 fish were purchased from the Sydney fish market, New South Wales (NSW), Australia. The fish had been sourced from three separate localities: off the coast of NSW (n = 44; 11/10/2018), off the coast of NZ ((n = 20; 28/07/2018), and (n = 30; 16/09/2019)), and an unknown location (n = 20; 29/08/2018). Fish were transferred to the Parasitology Laboratory of Charles Sturt University, Wagga Wagga Campus in an insulated ice-filled box.

2.2. Parasite collection

All fish from each batch were examined on the day of arrival at the University. Fish were dissected and examined for the presence of nematodes according to the method described in Shamsi and Suthar (2016b) using both visual examination and incubation method to ensure maximum recovery of nematodes. Firstly, the surface of all inner organs was thoroughly inspected for the presence of nematodes under a dissecting microscope (Leica EZ4 Stereo Microscope, China). Encysted nematodes were removed and placed in sterile Eppendorf tubes containing 70% ethanol. The alimentary canal was then split from mouth to anus and other internal organs gently separated and placed in a petri dish containing a small amount of ambient temperature water before examining again under a dissecting microscope for the presence of parasites. The internal organs were placed in containers with water and after vigorous shaking were left to incubate overnight at room temperature. Splitting the digestive tract and separating internal organs allows trapped nematodes an opportunity to be released into the water and provides greater parasite recovery. All collected parasites were washed in ambient temperature physiological saline and preserved in 70% ethanol and stored at room temperature for further morphological and genetic study. Laboratory temperature was kept at a constant 25 °C.

2.3. Morphological examination

A small piece from the mid-body of each nematode was excised for molecular study and the rest of the body (head and tail) were cleared with lactophenol for morphological study. This study was conducted through a microscope (Upright Motorized Microscope ECLIPSE Ni-E, Nikon, Japan) fitted with a computer screen. Anisakid nematodes were initially identified to genus level using morphology and morphometry of anterior and posterior ends, oesophagus, ventriculus, the position of excretory pore and nerve ring (Murata et al., 2011; Shamsi and Suthar, 2016a). Cucullanid nematodes were morphologically identified based on the presence of anterior pseudobuccal capsule, spicules, number and organisation caudal papillae, precloacal ventral sucker, and tail according to instructions in Li et al. (2014); Moravec et al. (2019); Yamaguti (1935); Yamaguti (1941). The morphometric and meristic characteristics of systematic importance were measured directly with an eyepiece micrometre (BX-43 Olympus Microscope, Olympus Corporation, Japan). All measurements were recorded in both micrometres and millimetres as the mean, followed by the range in parentheses. A dash (−) indicates that measurements could not be made or were not available. All drawings were made to scale with the aid of a drawing tube of the compound microscope (BX-43 Olympus Microscope, Olympus Corporation, Japan). The prevalence, mean intensity, and mean abundance of nematodes were calculated according to Bush et al. (1997).

2.4. Sequencing

Genomic DNA from nematodes was extracted by DNeasy Blood & Tissue Kits (QIAGEN, Germany) and eluted by 40 μl of elution buffer. A volume of 25 μl PCR reaction was conducted to amplify the whole ITS (ITS–1, 5.8S, and ITS–2) regions of Anisakid nematodes using the primer sets of forward (SS1): 5′–GTTTCCGTAGGTGAACCTGCG–3′ and reverse (NC2): 5′–TTAGTTTCTTTTCCTCCGCT–3′. The cycling condition was followed according to Hossen and Shamsi (2021); Shamsi et al. (2020). A similar volume (25 μl) of PCR for the Cucullanid nematodes was conducted to amplify the whole ITS regions using another primer set, which included forward ITS–F: 5′–CCTAACAAGCCTCAACGGGTG–3′ and reverse ITS–R1: 5’–GCATACGAACTGAGAGCAGCG–3′ with the cyclic conditions of initial 95 °C for 2 min, then 95 °C for 30 s, 60 °C for 45 s, 72 °C for 1 min × 40 cycles following extension at 72 °C for 10 min and finally at 4 °C. An aliquot (3 μl) of each amplicon from both Anisakid and Cucullanid nematodes was examined on a 1.5% w/v agarose gel after staining with GelRed™ and photographed using a gel documentation system.

Representative samples were chosen from each group to send for sequencing to the Australian Genome Research Facility (AGRF). Identical primers set as for PCR were used to prepare the samples to send for sequencing. Sequence data including chromatogram were observed initially through Sequence Scanner software (Applied Biosystems® Genetic Analysers). Subsequently, sequences were aligned by MUSCLE (in MEGA v. 7) (Kumar et al., 2016) and then adjusted manually wherever necessary. Evolutionary analyses were conducted in MEGA v. 7 (Kumar et al., 2016).

2.5. Construction of phylogenetic tree

Two phylogenetic trees were constructed (based on two groups identified: Anisakids and Cucullanids) from the sequences generated in this study along with the representative sample sequences from GenBank (Table 2). GenBank sequences were sorted based on the BLAST searches and from the available publications. The phylogenetic relationships among specimens were calculated by the Bayesian method using MrBayes v 3.2 (Ronquist and Huelsenbeck, 2003). The GTR + G model was applied for both trees as suggested by jModelTest 2 (Darriba et al., 2012). Dichelyne romani (GenBank accession: KP699576) and Hysterothylacium aduncum (GenBank accession: KY909270) were used as outgroups for Anisakid and Cucullanid nematodes, respectively. For the construction of both phylogenetic trees, the sample frequencies were set at 1000, and calculated for 10,00,000 generation until the p value reached <0.01. After the mcmc run, the first 30% samples were discarded, and the sum command was used to summarise the phylogenetic trees. The phylogenetic tree was visualised using Figtree v 1.4.3 (Rambaut, 2014).

Table 2.

Details of the specimens used to construct the phylogenetic trees.

Nematode specimen GenBank accession number Host scientific name Host common name Geographical origin of the specimen Reference
Anisakis pegreffii AY821740 Lissodelphis borealis Northern right whale dolphin Drakes Beach, California, USA Nadler et al. (2005)
A. simplex S. S. AY826723 Trachurus trachurus Atlantic horse mackerel Cantabrian Sea, Spain Nadler et al. (2005)
A. typica AY826724 Stenella longirostris Spinner dolphin Coast of Brazil Nadler et al. (2005)
A. physeteris AY826721 Physeter catodon Sperm whale Tyrrhenian Sea, Italy Nadler et al. (2005)
A. brevispiculata MK325199 Kogia breviceps Pygmy sperm whale Southeast of Melbourne, Australia Shamsi et al. (2019b)
A. berlandi MK325187 K. breviceps Pygmy sperm whale Southeast of Melbourne, Australia Shamsi et al. (2019b)
A. paggiae MK325218 K. breviceps Pygmy sperm whale Southeast of Melbourne, Australia Shamsi et al. (2019b)
A. nascettii JQ912692 Mesoplodon grayi Gray's beaked whale Off the NZ coast Mattiucci et al. (2014)
A. ziphidarum JQ912691 Ziphius cavirostris Cuvier's beaked whale Off the South African coast Mattiucci et al. (2014)
Contracaecum rudolphii D FM210251+FM210261 Phalacrocorax carbo Great cormorant NSW and Victoria, Australia Shamsi et al. (2009a)
C. rudolphii E FM210257+FM210269 Phalacrocorax varius Pied cormorant NSW and Victoria, Australia Shamsi et al. (2009a)
C. pyripapillatum AM940062+AM940066 Pelecanus conspicillatus Australian pelican Victoria, Australia Shamsi et al. (2008)
C. multipapillatum D AM940056+AM940060 Pelecanus conspicillatus Australian pelican Victoria, Australia Shamsi et al. (2008)
C. bancrofti EU839566+FM177880 Pelecanus conspicillatus Australian pelican Victoria, Northern Territory, NSW, Australia Shamsi et al. (2009b)
C. microcephalum FM177524+FM177528 Phalacrocorax melanoleucos Little pied cormorant Lara and Healesville, Victoria, Australia Shamsi et al. (2009b)
C. variegatum FM177531+FM177541 Anhinga melanogaster and Pelecanus conspicillatus Australian darter and Australian pelican Melbourne, Victoria. Shamsi et al. (2009b)
C. eudyptulae FM177550+FM177578 Eudyptula minor Little penguin Victoria, Australia Shamsi et al. (2009b)
C. ogmorhini FM177542+FM177549 Arctocephalus pusillus doriferus and A. forsteri Australian and New Zealand fur seals Victoria, Australia Shamsi et al. (2009b)
Mawsonascaris vulvolacinata MK476521 Pastinachus ater Cowtail stingray Queensland, Australia Shamsi et al. (2019c)
Terranova type I MT635348 Platycephalus bassensis Sand flathead NSW, Australia Hossen et al. (2021)
Pulchrascaris australis MK890747 Sphyrna lewini Scalloped hammerhead shark Off Cairns, Australia Shamsi et al. (2020)
Terranova type II MT635350 Platycephalus richardsoni Tiger flathead NSW, Australia Hossen et al. (2021)
T. pectinolabiata MK542878 S. mokarran Great hammerhead shark NSW, Australia Shamsi et al. (2019a)
Pseudoterranova azarasi AJ413973+AJ413974 Eumetopias jubatus Steller sea lion Iwani, Japan Zhu et al. (2002)
P. bulbosa AJ413970+AJ413971 Erignathus barbatus Bearded seal Newfoundland, Canada Zhu et al. (2002)
P. cattani AJ413982+AJ413984 Otaria byronia South American sea lion Concepcion, Chile Zhu et al. (2002)
P. decipiens AJ413967+AJ413968 Phoca vitulina Harbour seal Newfoundland, Canada Zhu et al. (2002)
P. krabbei AJ413965+AJ413980 Halichoerus grypus Gray seal Froya Island, Norway Zhu et al. (2002)
Raphidascaris acus AY603537 Anguilla anguilla European eel Vistula Lagoon, Poland Kijewska et al. (2008)
Dichelyne pleuronectidis KF470872–83 Pleuronichthys cornutus Ridged-eye flounder East China Sea Li et al. (2014)
D. romani KP699576 Notacanthus chemnitzii Snub-nosed spiny eel Northeast Atlantic Isbert et al. (2015)
D. szidati MK131263 Acanthistius patachonicus Argentine sea bass Argentina Unpublished
Hysterothylacium aduncum KY909270 Peltorhamphus novaezeelandiae New Zealand sole Off the coast of Otago, NZ Anglade and Randhawa (2018)
H. australe HE862216+HE862223 Seriola lalandi Yellowtail amberjack Port Augusta, South Australia Shamsi (2016)
H. brucei HE862222+HE862230 Kajikia audax Striped marlin Nelson Bay, NSW, Australia Shamsi (2016)
H. kajikiae HE862220+HE862226 Kajikia audax Striped marlin Nelson Bay, NSW, Australia Shamsi (2016)
Anisakis type I MT791088–103 Chrysophrys auratus Snapper Australia and NZ Present study with voucher numbers 169, 177, 231–2, 232–1, 235, 236–1, 239, 246–1, 247, 252–1, 258–1, 285, 291, 420, 427, and 440
Anisakis type III MT791104 C. auratus Snapper Australia Present study with voucher number 413
Terranova type II MT791105–06 C. auratus Snapper NZ Present study with voucher numbers 168 and 252–5
Dichelyne cf. pleuronectidis MT791107–10 C. auratus Snapper Australia and NZ Present study with voucher numbers 91–4, 93–1, 281–1, and 283
Dichelyne sp. 1 MT791111 C. auratus Snapper Australia Present study with voucher number 282
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Abbreviations: NZ = New Zealand, NSW = New South Wales.

NB: Single GenBank accession indicates the whole ITS (ITS–1, 5.8S, ITS–2) sequence. Two ITS sequences connected with ‘+’ represent ITS-1 and ITS-2 sequences, respectively.

3. Results

3.1. Morphological identification of nematodes

A total of four different nematode morphotypes belonging to the families Anisakidae and Cucullanidae were identified in this study. All Anisakid nematodes were identified as infective L3 larvae of Anisakis types I & III (Figs. 1a–d), and Terranova type II (Fig. 1e–f). Larvae were found in the digestive tracts, gonads, and liver. All Cucullanid nematodes were extracted only from the digestive system of the examined fish and were morphologically identified as Dichelyne (Cucullanellus) spp. which included larvae and adults (Figs. 2a–l). Among all nematode morphotypes identified in this study, Anisakis type I occurred in all samples/batch and had the highest prevalence of infection. The second most prevalent nematode morphotype was identified as Dichelyne spp. Anisakis type III and Terranova type II larvae were the least prevalent nematode morphotypes identified in this study. The general epidemiological data are represented in Table 3.

Fig. 1.

Fig. 1

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Morphology of Anisakid nematodes identified from snapper Chrysophrys auratus. a) Anterior end and b) Posterior end of Anisakis type I; c) Anterior end and d) Posterior end of Anisakis type III; e) Anterior end and f) Posterior end of Terranova type II.

Fig. 2.

Fig. 2

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Morphology of Cucullanid nematodes identified as Dichelyne (Cucullanellus) spp. from snapper Chrysophrys auratus. a) Anterior end and b) Posterior end of mature male Dichelyne cf. pleuronectidis; c) Anterior end and d) Posterior end of gravid female Dichelyne cf. pleuronectidis; e) Anterior end and f) Posterior end of gravid female Dichelyne sp. 1; g) A typical pseudobuccal capsule; h–j) Posterior end of immature female specimens; k) Posterior end of immature male specimen; l) Eggs.

Table 3.

Occurrence and abundance of nematodes in snapper Chrysophrys auratus examined in the present study.

Source of fish (number examined) Nematode Number of fish infected Range in infected fish Prevalence (%) Total number of parasites found Mean intensity Mean abundance
SFM, NSW, Australia Date: 29-08-2018 (n = 20) Anisakis pegreffii⁎ 3 1–2 15 4 1.33 ± 0.59 0.20
Dichelyne cf. pleuronectidis 4 1–1 20 4 1 ± 0.61 0.20
Dichelyne sp. 1⁎ 1 1–1 5 1 1 ± 0.62 0.05
Off the coast of NSW, Australia Date: 11-10-2018 (n = 44) Anisakis pegreffii 3 1–1 7 3 1 ± 0.66 0.10
Anisakis brevispiculata⁎ 1 1–1 2 1 1 ± 0.67 0.03
Dichelyne cf. pleuronectidis 4 1–4 9 8 2 ± 0.69 0.18
Off the coast of NZ Date: 28-07-2018 (n = 20) Anisakis pegreffii 2 1–1 10 2 1 ± 0.75 0.10
D. cf. pleuronectidis 3 1–1 15 3 1 ± 0.78 0.15
Terranova type II⁎ 1 1–1 5 1 1 ± 0.81 0.05
Off the coast of NZ Date: 16-09-2019 (n = 28) Anisakis pegreffii 9 1–6 32 25 2.78 ± 0.86 0.89
D. cf. pleuronectidis 3 1–1 11 3 1 ± 0.78 0.11
Terranova type II 1 1–1 4 1 1 ± 0.82 0.04
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Abbreviations: SFM = Sydney Fish Market, NSW = New South Wales, NZ = New Zealand.

⁎

Asterisk indicates the new host records of these nematodes.

3.2. Molecular identification

Representative samples from each morphotype were subjected to sequencing to confirm the specific identity of nematodes as detailed below.

Sixteen specimens belonging to Anisakis type I were subjected to sequencing. The length of ITS regions was 857 bp long and identical. Our sequences were 100% identical to those belonging to adult A. pegreffii (accession number AY821740) identified from the Northern right whale dolphin Lissodelphis borealis in the Drakes Beach, California, USA (Nadler et al., 2005). A single specimen belonging to Anisakis type III had the ITS sequence of 804 bp long and was 100% identical with the adult A. brevispiculata (accession number MK325199) reported from a pygmy sperm whale Kogia breviceps in the Southeast of Melbourne, Australia (Shamsi et al., 2019b).

Two specimens belonging to Terranova type II were subjected to sequencing. The ITS sequences of both specimens were 893 bp long and identical. There was no identical or highly similar sequence available in the GenBank.

Five specimens (voucher numbers 91–4, 93–1, 281–1, 283, and 282) belonging to Dichelyne spp. which included larva, mature males, and gravid females were subjected to sequencing. The ITS sequences of four specimens were 930 bp long and showed 0–0.30% nucleotide variability. A search in GenBank showed 99% similarity with D. pleuronectidis (accession numbers KF470872–KF470883) identified from ridged-eye flounder Pleuronichthys cornutus in the East China Sea (Li et al., 2014). The length of the ITS sequence for the other specimen (voucher number 282; a gravid female Cucullanid) was 813 bp long which had no closest similarity with the registered GenBank sequences. The pairwise comparison between the first four ITS sequences and the later, generated in the present study, revealed a substantial nucleotide variability at 0–24.10% and considered herein as interspecific genetic variation (Fig. 5).

Fig. 5.

Fig. 5

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The ITS (ITS-1, 5.8S, ITS-2) sequence alignments of the present Dichelyne specimens and closely related species from GenBank. Sample's information is provided in Table 2. The dots represent identical bases and dashes indicate alignment gaps. The numbers at the right of alignments indicate the alignment position.

3.3. Phylogenetic analyses of the nematodes

The Bayesian inference phylogenetic tree clustered Anisakid nematodes found in this study with the members of Anisakidae identified previously in the literature (Fig. 3A). Anisakis type I independently grouped with members of A. pegreffii and Anisakis type III with A. brevispiculata demonstrating 100% posterior probabilities, respectively. Terranova type II larva found in the present study clustered into a separate clade and revealed a clear distinction with previously identified larval and adult Terranova and Pseudoterranova specimens in Australian waters and elsewhere with 100% posterior probability value. The Cucullanid nematodes found in the present study grouped with closely related GenBank Dichelyne specimens. Four sequences explored in this study clustered with the D. pleuronectidis with 100% posterior probability value (Fig. 3B). A single sequence obtained from a gravid female Cucullanid was isolated from the existing GenBank Dichelyne species with a 100% posterior probability value. The isolated specimen in this study was assigned herein as Dichelyne species 1 (Fig. 3B).

Fig. 3.

Fig. 3

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Phylogenetic relationship among the nematodes identified from snapper Chrysophrys auratus and those closely related species in GenBank (see Table 2 for details) inferred using Bayesian method with ITS (ITS-1, 5.8S, ITS-2) sequences. A) Phylogenetic tree for Anisakid nematodes; B) Phylogenetic tree for Dichelyne spp. nematodes. *indicates the ITS sequences generated in this study. Bayesian posterior probability values (%) were shown on the node.

4. Discussion

This study confirmed the presence of A. pegreffii, A. brevispiculata, Terranova type II, D. cf. pleuronectidis, and Dichelyne sp. 1 infection in snapper from coastal waters of NSW and NZ. This is the first time that infectious stage larvae of the zoonotic A. pegreffii and potentially zoonotic Terranova type II have been identified in this species of snapper. New host records for A. brevispiculata, D. cf. pleuronectidis, and Dichelyne sp. were also established.

The individual prevalence of Anisakid and Cucullanid nematodes were 18% and 13%, respectively and for zoonotic A. pegrefii, the overall prevalence was 15%. This differs from Sharples and Evans (1995a) where Anisakis sp. was found to be rare with limited prevalence and low abundance. In the same study, Cucullanus sp. was reported moderately abundant in snapper. Differences between the present study and Sharples and Evans (1995a) may be due to the epidemiological profile of nematodes changing over time and differences in genera identified in the previous study. A further investigation with a greater sample size and conducted at different time points is required to clarify nematode infection trends in snapper.

In the present study, Anisakis larval types were morphologically identified as types I & III (Figs. 1a–d). In morphological identification, Anisakis type I could not be distinguished from larvae of A. pegreffii, A. simplex S. S., A. berlandi, and A. typica (Mattiucci et al., 2018; Murata et al., 2011). Similarly, Anisakis type III could be the larvae of A. brevispiculata, A. physeteris, and A. paggiae (Mattiucci et al., 2018; Murata et al., 2011). Previously, Anisakis sp. larva was found from the same host in Australia (Johnston and Mawson, 1945) and NZ (Sharples and Evans, 1995b, Sharples and Evans, 1995c). However, specific identification of the larva was uncertain at the time due to a lack of molecular techniques. The phylogenetic tree showed a clear distinction between Anisakis types I & III found in this study and grouped them with previously identified adult A. pegreffii and A. brevispiculata, respectively (Fig. 3A). Therefore, the present study confirms snapper as a host of A. pegreffii and A. brevispiculata.

Another Anisakid nematode morphologically identified in this study was Terranova type II. Terranova types I and II have been previously identified and reported from other marine fish in Australia and New Caledonia (Hossen et al., 2021; Jabbar et al., 2012; Shamsi et al., 2018a; Hossen and Shamsi, 2019; Shamsi et al., 2018b; Shamsi and Suthar, 2016a) as Pulchrascaris australis and T. pectinolabiata, respectively (Shamsi et al., 2019a; Shamsi et al., 2020). Although, the morphological and morphometric data of the present specimens either partially or completely matched with previously identified Terranova larval types (Table 4) the molecular data did not match with existing sequences registered in GenBank and revealed 20.70–20.90% nucleotide variability (Fig. 4).

Table 4.

Comparative measurements of Terranova larval types, for specimens found in the present study and previous studies.

Present study Jabbar et al. (2012) Shamsi et al. (2018a) Shamsi et al. (2016a)
Larval type Terranova type II Terranova type I Terranova type II Terranova type I Terranova type II Terranova type II
Locality Off the coast of NZ Lizard Island in the Great Barrier Reef, QLD, Australia Province Sud, New Caledonia Off Australian coasts including NSW, VIC, QLD, SA, and WA
Number of specimens observed 02 10 10 08 10 10
Body length 7.09 (6.8–7.38) – – 9.92 (7.5–12.55) 6.63 (5.42–8.30) 6.60 (3.00–9.00)
Maximum body width 0.24 (0.22–0.25) – – 0.25 (0.19–0.32) 0.23 (0.18–0.28) 0.24 (0.18–0.28)
Oesophagus length 0.92 (0.90–0.93) 1.16 (1.08–1.65) 0.94 (0.78–1.10) 1.00 (0.7–1.28) 0.85 (0.73–1.03) 0.88 (0.40–1.14)
Ratio of oesophagus length to body length (%) 12.98 (12.60–13.24) – – 10.08 (7.06–12.90) 12.82 (11.01–15.54) 14.30 (9.50–26.50)
Intestinal caecum length 0.88 (0.85–0.90) 1.52 (0.73–2.08) 0.72 (0.60–0.87) 1.23 (0.60–1.70) 0.68 (0.61–0.85) 0.71 (0.50–0.90)
Ventriculus length 0.57 (0.51–0.62) 1.32 (0.95–1.96) 0.38 (0.35–0.45) 1.09 (0.35–1.45) 0.34 (0.29–0.38) 0.38 (0.24–0.54)
Nerve ring to anterior end 0.29 (0.28–0.30) 0.30 (0.25–0.33) 0.28 (0.26–0.36) 0.27 (0.2–0.33) 0.25 (0.22–0.32) 0.37 (0.22–0.72)
Tail length 0.19 (0.18–0.20) 0.18 (0.16–0.20) 0.15 (0.12–0.18) 0.13 (0.11–0.17 0.13 (0.09–0.15) 0.13 (0.12–0.14)
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All measurements are given in millimetres; mean followed by range in parentheses. ‘–’ indicates no measurements/data available; Abbreviations: NSW=New South Wales, VIC=Victoria, SA = South Australia, WA = Western Australia, QLD = Queensland, NZ = New Zealand.

Fig. 4.

Fig. 4

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The ITS (ITS-1, 5.8S, ITS-2) sequence alignments of the present Terranova specimens and closely related species from GenBank. Sample's information is provided in Table 2. The dots represent identical bases and dashes indicate alignment gaps. The numbers at the right of alignments indicate the alignment position.

Very little is known about Terranova larval types compared to other Anisakid nematodes (Moravec and Justine, 2020). The larvae belonging to the genera Pulchrascaris, Terranova, and Pseudoterranova look morphologically similar and are all referred to as Terranova larval types (Shamsi and Suthar, 2016a). The species belonging to the later genus are considered to be the second most common zoonotic nematode causing human illness after Anisakis spp. (Arizono et al., 2011; McClelland, 2002; Menghi et al., 2020; Torres et al., 2007; Weitzel et al., 2015). Although, Terranova type A larva has been reported for two cases of human infection in Korea (Lee et al., 1985; Seo et al., 1984), however this requires further investigation (Moravec and Justine, 2020). The phylogenetic tree grouped Terranova type II sequences obtained in the present study independently from the registered sequences for Ascaridoid nematodes larvae and adults in GenBank with 100% posterior probability value (Fig. 3A). The new sequences did not cluster with earlier larval and adult Terranova and Pseudoterranova species identified in the Australian waters and abroad. Therefore, the specific identification of the present Terranova type II requires clarification. Further parasitological examination with greater sample size is required to explore more genetic information for the specific identification of Terranova larval infection in snapper.

Although other nematodes (Dichelyne spp.) found in this study are not of zoonotic significance, they impact fish health and can significantly damage the intestinal wall and disrupt functional nutrient absorption leading to deficits in host growth, health and death (Dunn et al., 1983; Li et al., 2014; Rezaei et al., 2013). Until now, two Dichelyne (Cucullanellus) spp. have been reported from the Australian waters, D. sheardi from the silver spot Chironemus maculosus and snapper C. auratus as well as D. cnidoglanis from the estuarine catfish Cnidoglanis macrocephalis (Johnston and Mawson, 1944, Johnston and Mawson, 1945, Johnston and Mawson, 1949). In NZ, a single species D. cnidoglanis has been reported (Brunsdon, 1953; ex. from Sharples and Evans in 1995c) with reports of unidentified species as Cucullanus (not Dichelyne species) from snapper (Sharples and Evans, 1995c). The morphometric and meristic data suggest our specimens are different from those previously described from the Australian and NZ waters.

Globally, a total of three Dichelyne (Cucullanellus) spp. have been identified from the Sparid hosts (Isbert et al. (2015), D. adriaticus (Petter, 1974); D. jialaris (Moravec et al., 2018); and D. pleuronectidis (Li et al., 2014; Yamaguti, 1935; Yamaguti, 1941) of which our specimens most closely resembled D. pleuronectidis (Table 5). However, specimens in our study were different from previous descriptions in Li et al. (2014); Yamaguti (1935) and Yamaguti (1941) with respect to: i) markedly thick cuticle with transverse striations; ii) muscular oesophagus expanded at the anterior end to form a distinct pseudobuccal capsule; iii) anterior oesophageal region being much-expanded/wider than the posterior region and clearly distinguishable. The pairwise genetic comparison between the four ITS sequences generated in this study and those available in GenBank demonstrated 0–0.60% nucleotide variability (Fig. 5) with D. pleuronectidis thus the Dichelyne nematode identified in this study was assigned as D. cf. pleuronectidis. Further investigation is required to reach a solid conclusion if the observable morphological variations in D. cf. pleuronectidis found in the present specimens are due to geographical differences of host origin. In the present study, a single gravid female Dichelyne species herein named as Dichelyne sp. 1 remains unidentified due to the lack of comparable mature male specimens and revealed 0.10–24.40% nucleotide variability with those sequences registered in GenBank (Fig. 5). The phylogenetic tree clustered four (out of five) Dichelyne nematode sequences obtained in the present study with the sequences registered for D. pleuronectidis in GenBank. A single sequence from the present study grouped in isolation with 100% posterior probability value. The isolated specimen was herein named as Dichelyne sp. 1 (Fig. 3B). Further genetic analyses are required, for the morphologically identified Dichelyne nematodes, to verify their specific position.

Table 5.

Comparative measurements of Dichelyne cf. pleuronectidis and Dichelyne sp. 1 from snapper Chrysophrys auratus, for specimens collected in the present study and previous studies.

Present study Present study Li et al. (2014) ⁎Yamaguti (1935) ⁎Yamaguti (1941)
Parasite Dichelyne cf. pleuronectidis Dichelyne sp. 1 Dichelyne pleuronectidis Dichelyne pleuronectidis Dichelyne pleuronectidis
Host Snapper Chrysophrys auratus Snapper C. auratus Ridged-eye flounder Pleuronichthys cornutus Three Pleuronectids: Pleuronichthys cornutus, Paralichthys olivaceus, Pseudorhombus cinnarnoneus Snapper Pagrosomus unicolor (Syn. C. auratus)
Locality Australia: SFM Australia: SFM China: The East China Sea Japan: Toyama Bay; the Inland Sea; and Mutu Bay Japan: The Inland Sea
New Zealand: Off the coast of NZ
Specimen type (number measured) Mature male (n = 3) Immature male (n = 3) Gravid female
(n = 2)		 Immature female
(n = 5)		 Gravid female
(n = 1)		 Mature male (n = 15) Gravid female (n = 15) Male (n = not specified) Female (n = not specified) Male (n = not specified) Female (n = not specified)
Body length (mm) 4.76 (4.13–5.63) 3.14 (3.00–3.38) 7.22 (6.63–7.80) 5.56 (2.8–7.43) 4.03 6.67 (5.49–7.94) 7.40 (5.80–8.70) 3.15–8.00 5.50–11.00 3.70–4.40 4.00–11.00
Maximum body width 460 (380–500) 207 (180–250) 985 (670–1300) 556 (220–900) 450 328 (225–421) 391 (294–471) 175–350 300–650 270–320 260–700
Oesophagus length 800 (700–850) 523 (450–640) 975 (850–1100) 776 (500–950) 520 809 (735–882) 915 (833–980) 500–910 800–1120 600–700 740–1000
Maximum oesophagus width 173 (150–200) 100 (80–140) 210 (200−220) 184 (120−220) 100 170 (147–196) – 75–250 125–270 120–130 110–240
Minimum oesophagus width 73 (60–80) 55 (40–75) 90 (80–100) 76 (40–90) 40 107 (88–137) – – – – –
Ratio of oesophagus length to body length (%) 17 (15–21) 17 (15–19) 13 (13–14) 14 (12–18) 13 12 (10–15) 13 (11–15) – – – –
Pseudobuccal capsule length 290 (240–350) 215 (180–275) 365 (330–400) 310 (190–400) 200 70 (49–88) 64 (49–78) – – – –
Pseudobuccal capsule width 217 (160–260) 157 (120−200) 270 (240–300) 232 (190–270) 130 83 (69–108) 97 (78–118) 110–220 150–200 135–155 150–240
Intestinal caecum length 250 (200−300) 180 (150–180) 350 (320–380) 153 (80–200) 130 239 (98–394) 262 (69–415) 180–450 110–530 320–380 200–500
Intestinal caecum width 100 (80–100) 50 (40–50) 130 (110–150) 123 (40–200) 50 58 (39–69) 58 (49–69) – – – –
Excretory pore to anterior end 480 (430–530) – 700 (590–810) 723 (670–820) – 529 (501–588) 646 (508–784) 550–1450 800–1550 600
Nerve ring to anterior end 433 (400–500) 253 (230−300) 495 (470–520) 456 (250–580) 300 347 (314–392) 380 (323–392) 200–400 280–500 260–300 300–430
Deirids to anterior end 650 (520–750) 410 (330–450) 900 (850–950) 845 (500–1050) – 750 (559–902) 783 (539–902) – – – –
Ventral precloacal sucker length 215 (200−230) – – – – – – – – – –
Ventral precloacal sucker width 130 (110–150) – – – – – – – – – –
Ventral precloacal sucker to cloaca 377 (350–400) – – – – 487 (412–539) – 510–650 – – –
Ventral precloacal sucker to posterior end 570 (530–600) – – – – – – – – – –
Spicule length 933 (900–1000) – – – – 975 (735–1176) – 630–1030 890–930
Ratio of spicule length to total body length (%) 20 (17–24) – – – – 15 (12–17) – – – – –
Gubernaculum length 40 (40–45) – – – 44 (40–54) – 33–48 – – –
Number of precloacal papillae (pairs) 3 – – – – 3 – – – – –
Number of paracloacal papillae (pairs) 4 – – – – 4 – – – – –
Number of postcloacal papillae (pairs) 3 – – – – 3 – – – – –
Total number of caudal papillae (pairs) 10 – – – – 10 – 11 – – –
Tail length 173 (150–200) 165 (150–180) 270 (250–290) 310 (210–460) 130 186 (157–206) 268 (225–323) 130–180 200–280 150–160 200–250
Phasmids to posterior end 233 (200–250) – 400 (350–450) 423 (370–450) – – 127 (118–147) – – – –
Eggs count – – ~100–400 – ~100 – 45 – – – –
Eggs length – – 80 (70–80) – 70 – 62 (59–69) – 63–84 – 66–84
Eggs width – – 40 (40–50) – 40 – 46 (39–49) – 39–46 – 39–45
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All measurements are given in micrometres unless otherwise stated; mean followed by range in parentheses. ‘–’ indicates no measurement/data available; Abbreviations: SFM = Sydney Fish Market, NZ = New Zealand.

⁎

Some of the measurements have been converted into micrometres.

The zoonotic potential of all Anisakis spp. are not yet clear (Aibinu et al., 2019; Mattiucci et al., 2018). In most cases nematode larvae infecting humans are often damaged on removal (Mattiucci et al., 2013) are identified morphologically or based on the assumption (Mattiucci and D'Amelio, 2014). Given that a range of genera within the family can frequently infect humans, it is likely that all members of the Anisakis genus are potentially zoonotic. Therefore, further research on the specific identification of these larvae in human cases of infection is required.

According to Bao et al. (2017a) anisakiasis (human infection caused by Anisakis nematodes) is an emerging zoonosis which is underestimated globally. These zoonotic nematodes are medically important and cause intestinal (Kojima et al., 2013), and less commonly extra-intestinal anisakiasis as well as allergic reactions (Mattiucci et al., 2017). Anisakis pegreffii has increasingly been identified in human cases of allergic (Mattiucci et al., 2013), intestinal (Moschella et al., 2004), and extra-intestinal anisakiasis (Guardone et al., 2018; Bao et al., 2019).

Identification of zoonotic and/or potentially zoonotic larvae from a popular Australian/NZ table fish such as the iconic snapper, considered a suitable species for consuming raw, is of concern for human health (Shamsi, 2020). According to Shamsi and Sheorey (2018), anisakidosis (human infection caused by the larval Anisakid nematodes) in Australia is of emerging importance and may correspond with the adoption of novel cuisines including the popular sushi and sashimi. Very recently, the first human anisakiasis, caused by Anisakis spp. larvae was diagnosed in NZ after consumption of a sushi rolls (Beig et al., 2019).

All anisakid nematodes found in this study were viable infectious third-stage larvae and may cause human illness if accidentally consumed in raw or partially cooked fish (Bao et al., 2017a; Bao et al., 2018; Buchmann and Mehrdana, 2016; Cipriani et al., 2016; Caldeira et al., 2021; D'Amico et al., 2014). Anisakis pegreffii as the dominant species in the present study and recent reports of other infected fish in the Asia Pacific region may support a southward shift in parasite distribution (Palm et al., 2017; Chen et al., 2018; Zhang et al., 2018). Anisakis pegreffi has been identified as the dominant species in pelagic blue mackerel Scomber australasicus (Taiwanese waters) (Chen and Shih, 2015), pelagic scombrid bullet tuna Auxis rochei (Indonesian waters) (Palm et al., 2017), white spotted Conger Conger myriaster (South China Sea) (Chen et al., 2018) and the yellow goosefish Lophius litulon (East China Sea) (Zhang et al., 2018). Further monitoring of fish in Australian waters, therefore, seems warranted.

Further research is required to determine the human health risks using a greater sample size covering both a greater temporal and spatial range. In addition, more advanced methods of parasite isolation should be considered in future studies as examining only the fish viscera in the present study limited the strength of the results. Candling of fish fillets is recommended in Codex Alimentarius ‘Code of practice for fish and fishery products’ for identification of parasites (Codex Alimentarius, 2020) in fish musculature. However, this method is less effective in detection of Anisakis spp. and darker Pseudoterranova spp. larvae (Levsen et al., 2005; McGladdery, 1986; Petrie et al., 2007; Mercken et al., 2020a). Candling combined with pressing may be more effective to detect nematode parasites in the musculature of fish (Karl and Leinemann, 1993). Levsen et al. (2005) found that in blue whiting with an average thickness of 11 mm, the detection of nematodes using UV light was only 10–15% however Gómez-Morales et al. (2018) found the UV press method had a high sensitivity for detection of nematodes in fish musculature and viscera. Artificial pepsin digestion of fish musculature is also recommended in the Codex fishery code of practice for recovery of parasites in high-risk fish species (Codex Alimentarius, 2020) and fillets (Mercken et al., 2020b). This results in the total destruction of the fish and for commercial purposes is impractical. However, with the optimisation of this technique described in Llarena-Reino et al. (2013) the pepsin digestion method in the experimental setting would be an effective, low-cost, and accessible alternative to advanced methods such as Magnetic Resonance Imaging (MRI) described in Bao et al. (2017b). Press method candling in combination with artificial digestion of fish fillets in future studies will provide a more accurate indication of the potential of snapper fillets to be infected with zoonotic nematodes larvae.

Also, of importance is the potential for cross-contamination as a human health concern. Anisakis simplex has been identified as an important hidden allergen in food (Anibarro et al., 2007) and Bao et al. (2019) considers all Anisakis spp. as potential and important food allergens. As A. pegreffii has been identified as the causative agent in cases of human allergic anisakiasis the potential for cross-contamination of viable larvae from viscera onto fillets should be considered as a human health concern.

Seafood borne parasitic disease in Australia/NZ is little recognised or acknowledged. The Australia New Zealand Food Standards Code (ANZFSC, 1997) has not included information on fish borne parasites in local fish. The ‘The Compendium of Microbiological Criteria for Food (2018)’ from Food Standards Australia and NZ in Appendix I (FSANZ, 2018) mentioned parasites as a possible pathogenic microorganism that can cause foodborne illness only once.

In conclusion, snapper sourced from the waters of Australia and NZ was identified infected with zoonotic, potentially zoonotic, and non-zoonotic nematodes. Further research into zoonotic nematodes in snapper using advanced detection methods will identify if current Australian food safety regulations regarding zoonotic or potentially zoonotic nematodes in fish and fishery products in Australia/NZ require updating. Fish is an excellent source of lean protein and contains many beneficial fatty acids essential for early development as well as eye, brain and cardiovascular health (Aadland et al., 2015). The identification of zoonotic and potentially zoonotic parasites should not be a deterrent to regular consumption of this healthy protein. If intended for consumption raw or lightly processed, it is recommended that fish be frozen at ≤ − 20 °C for a minimum period of one day (EC, 2004). Adequately cooking, where the internal temperature of fish reaches ≥55 °C, for a minimum period of 5 min will largely negate the risks to human health (De Marval et al., 2013).

Animal ethics

Not applicable.

Declaration of Competing Interest

Md. Shafaet Hossen, Sky Wassens, and Shokoofeh Shamsi declare that they have no conflict of interest.

Acknowledgment

Md. Shafaet Hossen (MSH) is the recipient of the Australian Research Training Program Scholarship through Charles Sturt University. MSH acknowledges Dr. Di Barton, Dr. Xiaocheng (Diego) Zhu, and Ms. Michelle Williams for their support during this study.

Contributor Information

Md. Shafaet Hossen, Email: mhossen@csu.edu.au.

Shokoofeh Shamsi, Email: sshamsi@csu.edu.au.

References

  1. Aadland E.K., Lavigne C., Graff I.E., Eng Ø., Paquette M., Holthe A., Mellgren G., Jacques H., Liaset B. Lean-seafood intake reduces cardiovascular lipid risk factors in healthy subjects: results from a randomized controlled trial with a crossover design1,2. Am. J. C. Nutr. 2015;102:582–592. doi: 10.3945/ajcn.115.112086. [DOI] [PubMed] [Google Scholar]
  2. Aibinu I.E., Smooker P.M., Lopata A.L. Anisakis nematodes in fish and shellfish-from infection to allergies. Int. J. Parasitol. Parasites Wildl. 2019;9:384–393. doi: 10.1016/j.ijppaw.2019.04.007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Allen A. Goldband Snapper Sashimi with Soy, Rice Wine Vinegar & Finger Limes. 2019. https://www.sbs.com.au/food/recipes/goldband-snapper-sashimi-soy-rice-wine-vinegar-finger-limes Accessed on 15 January 2019.
  4. Anglade T., Randhawa H.S. Gaining insights into the ecological role of the New Zealand sole (Peltorhamphus novaezeelandiae) through parasites. J. Helminthol. 2018;92:187–196. doi: 10.1017/S0022149X17000323. [DOI] [PubMed] [Google Scholar]
  5. Anibarro B., Seoane F.J., Mugica M.V. Involvement of hidden allergens in food allergic reactions. J Investig Allergol Clin Immunol. 2007;17:168–172. [PubMed] [Google Scholar]
  6. ANZFSC . 1997. Australia and New Zealand Food Standards Code. Food standards code. Accessed on 16 January 2019. [Google Scholar]
  7. Arizono N., Miura T., Yamada M., Tegoshi T., Onishi K. Human infection with Pseudoterranova azarasi roundworm. Emerg. Infect. Dis. 2011;17:555. doi: 10.3201/eid1703.101350. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Bao M., Pierce G.J., Pascual S., González-Muñoz M., Mattiucci S., Mladineo I., Cipriani P., Bušelić I., Strachan N.J. Assessing the risk of an emerging zoonosis of worldwide concern: anisakiasis. Sci. Rep. 2017;7:43699. doi: 10.1038/srep43699. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Bao M., Strachan N.J., Hastie L.C., MacKenzie K., Seton H.C., Pierce G.J. Employing visual inspection and Magnetic Resonance Imaging to investigate Anisakis simplex s.l. infection in herring viscera. Food Control. 2017;75:40–47. [Google Scholar]
  10. Bao M., Pierce G.J., Strachan N.J., Martínez C., Fernández R., Theodossiou I. Consumers' attitudes and willingness to pay for Anisakis-free fish in Spain. Fish. Res. 2018;202:149–160. [Google Scholar]
  11. Bao M., Pierce G.J., Strachan N.J., Pascual S., González-Muñoz M., Levsen A. Human health, legislative and socioeconomic issues caused by the fish-borne zoonotic parasite Anisakis: challenges in risk assessment. Trends Food Sci. Technol. 2019;86:298–310. [Google Scholar]
  12. Beig J., Lane R.J., Lane M.R. Gastric anisakiasis: a rare cause of abdominal pain. Intern. Med. J. 2019;49:129–130. doi: 10.1111/imj.14194. [DOI] [PubMed] [Google Scholar]
  13. Brunsdon R.V. 1953. A Systematic Study of the Nematodes Parasitic in New Zealand, with an Account of the Life History and Culture of Stomachus marinus (Linn., 1776) Unpublished M.Sc. thesis, lodged in the Library, Victoria University of Wellington. (Ex. from Sharples and Evans, 1995) [Google Scholar]
  14. Brunsdon R.V. 1956. Studies on Nematode Parasites of New Zealand Fishes. A Systematic and Parasitological Study of the Nematodes Occurring in New Zealand Marine and Fresh-water Fishes, Including Biological Studies on the Genus Anisakis Dujardin 1845. Unpublished Ph.D. thesis, lodged in the Library, Victoria University of Wellington. (Ex. from Sharples and Evans, 1995) [Google Scholar]
  15. Buchmann K., Mehrdana F. Effects of anisakid nematodes Anisakis simplex (s.l.), Pseudoterranova decipiens (s.l.) and Contracaecum osculatum (s.l.) on fish and consumer health. Food Waterborne Parasitol. 2016;4:13–22. [Google Scholar]
  16. Bush A.O., Lafferty K.D., Lotz J.M., Shostak A.W. Parasitology meets ecology on its own terms. J. Parasitol. 1997;83:575–583. [PubMed] [Google Scholar]
  17. Caldeira A.J.R., Alves C.P.P., Santos M.J. Anisakis notification in fish: an assessment of the cases reported in the European Union Rapid Alert System for Food and Feed (RASFF) database. Food Control. 2021;124 [Google Scholar]
  18. Chen H.X., Zhang L.P., Gibson D.I., Lü L., Xu Z., Li H.T., Ju H.D., Li L. Detection of ascaridoid nematode parasites in the important marine food-fish Conger myriaster (Brevoort) (Anguilliformes: Congridae) from the Zhoushan Fishery, China. Parasit. Vectors. 2018;11:274. doi: 10.1186/s13071-018-2850-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Chen H.-Y., Shih H.-H. Occurrence and prevalence of fish-borne Anisakis larvae in the spotted mackerel Scomber australasicus from Taiwanese waters. Acta Trop. 2015;145:61–67. doi: 10.1016/j.actatropica.2015.02.011. [DOI] [PubMed] [Google Scholar]
  20. Cipriani P., Acerra V., Bellisario B., Sbaraglia G.L., Cheleschi R., Nascetti G., Mattiucci S. Larval migration of the zoonotic parasite Anisakis pegreffii (Nematoda: Anisakidae) in European anchovy, Engraulis encrasicolus: implications to seafood safety. Food Control. 2016;59:148–157. [Google Scholar]
  21. Codex Alimentarius . Food and Agriculture Organisation of the United Nations and World Health Organisation; Geneva, Switzerland: 2020. Code of Practice for Fish and Fishery Products, Code of Practice for Fish and Fishery Products; pp. 1–372. [Google Scholar]
  22. D'Amico P., Malandra R., Costanzo F., Castigliego L., Guidi A., Gianfaldoni D., Armani A. Evolution of the Anisakis risk management in the European and Italian context. Food Res. Int. 2014;64:348–362. doi: 10.1016/j.foodres.2014.06.038. [DOI] [PubMed] [Google Scholar]
  23. Darriba D., Taboada G.L., Doallo R., Posada D. jModelTest 2: more models, new heuristics and parallel computing. Nat. Methods. 2012;9:772. doi: 10.1038/nmeth.2109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. De Marval F., Gottstein B., Weber M., Wicht B. Imported diphyllobothriasis in Switzerland: molecular methods to define a clinical case of Diphyllobothrium infection as Diphyllobothrium dendriticum, August 2010. Euro Surveill. 2013;18:20355. [PubMed] [Google Scholar]
  25. Dunn I.J., Russell L.R., Adams J.R. Cecal histopathology caused by Truttaedacnitis truttae (Nematoda: Cucullanidae) in rainbow trout, Salmo gairdneri. Int. J. Parasitol. 1983;13:441–445. doi: 10.1016/s0020-7519(83)80006-x. [DOI] [PubMed] [Google Scholar]
  26. EC Regulation (EC) No 853/2004 of the European Parliament and of the Council of 29 April 2004 laying down specific hygiene rules for on the hygiene of foodstuffs. 2004. https://eur-lex.europa.eu/LexUriServ/LexUriServ.do?uri=OJ:L:2004:139:0055:0205:en:PDF O. J. E. U. 30.4.2004 (L 139/55), 151. Retrieved from.
  27. Fowler A.J., Huveneers C., Lloyd M.T. Insights into movement behaviour of snapper (Chrysophrys auratus, Sparidae) from a large acoustic array. Mar. Freshw. Res. 2017;68:1438–1453. [Google Scholar]
  28. FRDC . 2019. Status of Australian Snapper – Briefing Notes. Version one: July 2019. [Google Scholar]
  29. FSANZ . Kingston ACT; Australia: 2018. Food Standards Australia New Zealand: Compendium of Microbiological Criteria for Food. [Google Scholar]
  30. Godfriaux B.L. Food of predatory demersal fish in Hauraki Gulf: 1: food and feeding habits of snapper. N. Z. J. Mar. Freshwat. Res. 1969;3:518–544. [Google Scholar]
  31. Gómez-Morales M.A., Castro C.M., Lalle M., Fernández R., Pezzotti P., Abollo E., Pozio E., Trial T.R. UV-press method versus artificial digestion method to detect Anisakidae L3 in fish fillets: comparative study and suitability for the industry. Fish. Res. 2018;202:22–28. [Google Scholar]
  32. Gregori M., Roura Á., Abollo E., González Á.F., Pascual S. Anisakis simplex complex (Nematoda: Anisakidae) in zooplankton communities from temperate NE Atlantic waters. J. Nat. Hist. 2015;49:755–773. [Google Scholar]
  33. Guardone L., Armani A., Nucera D., Costanzo F., Mattiucci S., Bruschi F. Human anisakiasis in Italy: a retrospective epidemiological study over two decades. Parasite. 2018;25 doi: 10.1051/parasite/2018034. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Hine P.M., Anderson C.D. Diseases of the gonads and kidneys of NZ snapper Chrysophrys auratus Forster (Family Sparidae) In: Fowler M.E., editor. Wildlife Diseases of the Pacific Basin and Other Countries. Academic Press; London: 1981. pp. 166–170. [Google Scholar]
  35. Hossen M.S., Shamsi S. Zoonotic nematode parasites infecting selected edible fish in New South Wales. Australia. Int. J. Food Microbiol. 2019;308:108306. doi: 10.1016/j.ijfoodmicro.2019.108306. [DOI] [PubMed] [Google Scholar]
  36. Hossen M.S., Wassens S., Shamsi S. Integrative species delimitation and community structure of nematodes in three species of Australian flathead fishes (Scorpaeniformes: Platycephalidae) Parasitol. Res. 2021;120:461–480. doi: 10.1007/s00436-020-06802-4. [DOI] [PubMed] [Google Scholar]
  37. Hurst R.J. Marine invertebrate hosts of New Zealand Anisakidae (Nematoda) N. Z. J. Mar. Freshwat. Res. 1984;18:187–196. [Google Scholar]
  38. Isbert W., Montero F.E., Carrassón M., González-Solís D. Dichelyne (Cucullanellus) romani n. sp.(Nematoda: Cucullanidae) in notacanthid fishes from the Northeast Atlantic and Western Mediterranean. Syst. Parasitol. 2015;91:35–47. doi: 10.1007/s11230-015-9556-1. [DOI] [PubMed] [Google Scholar]
  39. Jabbar A., Asnoussi A., Norbury L.J., Eisenbarth A., Shamsi S., Gasser R.B., Lopata A.L., Beveridge I. Larval anisakid nematodes in teleost fishes from Lizard Island, northern Great Barrier Reef. Australia. Mar. Freshw. Res. 2012;63:1283–1299. [Google Scholar]
  40. Johnston T.H., Mawson P.M. Remarks on some parasitic nematodes from Australia and New Zealand. Trans. R. Soc. S. Aust. 1944;68:60–66. [Google Scholar]
  41. Johnston T.H., Mawson P.M. Some parasitic nematodes from South Australian marine fish. Trans. R. Soc. S. Aust. 1945;69:114–117. [Google Scholar]
  42. Johnston T.H., Mawson P.M. Some nematodes from Australian hosts, together with a note on Rhabditis allgeni. Trans. R. Soc. S. Aust. 1949;73:63–71. [Google Scholar]
  43. Kobe Jones A Guide to Making Sashimi at Home. 2019. https://www.kobejones.com.au/a-guide-to-making-sashimi-at-home Accessed on 10 January 2019.
  44. Kailola P.J., Williams M.J., Stewart P.C., Reichelt R.E., McNee A., Grieve C. Bureau of Resource Sciences, Department of Primary Industries and Energy, Fisheries Research and Development Corporation; Canberra, Australia: 1993. Australian Fisheries Resources. [Google Scholar]
  45. Karl H., Leinemann M. A fast and quantitative detection method for nematodes in fish fillets and fishery products. J. Food Saf. Food Qual. 1993;44:124–125. [Google Scholar]
  46. Kijewska A., Czarna A., Fernandez M., Zdzitowiecki K., Rokicki J., Wrobel B. Analysis of 5.8 S rDNA and internal transcribed spacer 1 (ITS1) sequences of ascaridoid nematodes: phylogenetic signal and hypothesis testing. Genes Genom. 2008;30:291–306. [Google Scholar]
  47. Kojima G., Usuki S., Mizokami K., Tanabe M., Machi J. Intestinal anisakiasis as a rare cause of small bowel obstruction. Am. J. Emerg. Med. 2013;31:1422.e1–1422.e2. doi: 10.1016/j.ajem.2013.05.015. [DOI] [PubMed] [Google Scholar]
  48. Kumar S., Stecher G., Tamura K. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 2016;33:1870–1874. doi: 10.1093/molbev/msw054. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Kwong K. Sashimi with Chilli-Lime Dressing. 2015. https://www.delicious.com.au/recipes/snapper-sashimi-chilli-lime-dressing/ab5de775-2fd5-4084-94f8-be258b2e17e1 Accessed on 29 January 2019.
  50. Lee A.H., Kim S.M., Choi K.Y. A case of human infection with the larva of Terranova type A. Korean J. Pathol. 1985;19:463–467. [Google Scholar]
  51. Levsen A., Lunestad B.T., Berland B. Low detection efficiency of candling as a commonly recommended inspection method for nematode larvae in the flesh of pelagic fish. J. Food Prot. 2005;68:828–832. doi: 10.4315/0362-028x-68.4.828. [DOI] [PubMed] [Google Scholar]
  52. Li L., Du L.-Q., Xu Z., Guo Y.-N., Wang S.-X., Zhang L.-P. Morphological variability and molecular characterisation of Dichelyne (Cucullanellus) pleuronectidis (Yamaguti, 1935) (Ascaridida: Cucullanidae) from the flatfish Pleuronichthys cornutus (Temminck & Schlegel) (Pleuronectiformes: Pleuronectidae) in the East China Sea. Syst. Parasitol. 2014;87:87–98. doi: 10.1007/s11230-013-9456-1. [DOI] [PubMed] [Google Scholar]
  53. Llarena-Reino M., Piñeiro C., Antonio J., Outeriño L., Vello C., González Á.F., Pascual S. Optimization of the pepsin digestion method for anisakids inspection in the fishing industry. Vet. Parasitol. 2013;191:276–283. doi: 10.1016/j.vetpar.2012.09.015. [DOI] [PubMed] [Google Scholar]
  54. Mattiucci S., D'Amelio S. Anisakiasis. In: Bruschi F., editor. Helminth Infections and their Impact on Global Public Health. Springer; Vienna, Austria: 2014. pp. 325–365. [Google Scholar]
  55. Mattiucci S., Fazii P., De Rosa A., Paoletti M., Megna A.S., Glielmo A., De Angelis M., Costa A., Meucci C., Calvaruso V. Anisakiasis and gastroallergic reactions associated with Anisakis pegreffii infection, Italy. Emerg. Infect. Dis. 2013;19:496. doi: 10.3201/eid1903.121017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Mattiucci S., Cipriani P., Webb S.C., Paoletti M., Marcer F., Bellisario B., Gibson D.I., Nascetti G. Genetic and morphological approaches distinguish 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) J. Parasitol. 2014;100:199–214. doi: 10.1645/12-120.1. [DOI] [PubMed] [Google Scholar]
  57. Mattiucci S., Colantoni A., Crisafi B., Mori-Ubaldini F., Caponi L., Fazii P., Nascetti G., Bruschi F. IgE sensitization to Anisakis pegreffii in Italy: comparison of two methods for the diagnosis of allergic anisakiasis. Parasite Immunol. 2017;39 doi: 10.1111/pim.12440. [DOI] [PubMed] [Google Scholar]
  58. Mattiucci S., Cipriani P., Levsen A., Paoletti M., Nascetti G. Molecular epidemiology of Anisakis and anisakiasis: an ecological and evolutionary road map. In: Rollinson D., Stothard R., editors. Advances in Parasitology. Elsevier; London, UK: 2018. pp. 93–263. [DOI] [PubMed] [Google Scholar]
  59. McClelland G. The trouble with sealworms (Pseudoterranova decipiens species complex, Nematoda): a review. Parasitology. 2002;124:183. doi: 10.1017/s0031182002001658. [DOI] [PubMed] [Google Scholar]
  60. McGladdery S.E. Anisakis simplex (Nematoda: Anisakidae) infection of the musculature and body cavity of Atlantic herring (Clupea harengus harengus) Can. J. Fish. Aquat. Sci. 1986;43:1312–1317. [Google Scholar]
  61. McManus D.P., Bowles J. Molecular genetic approaches to parasite identification: their value in diagnostic parasitology and systematics. Int. J. Parasitol. 1996;26:687–704. doi: 10.1016/0020-7519(96)82612-9. [DOI] [PubMed] [Google Scholar]
  62. Menghi C.I., Gatta C.L., Arias L.E., Santoni G., Nicola F., Smayevsky J., Degese M.F., Krivokapich S.J. Human infection with Pseudoterranova cattani by ingestion of “ceviche” in Buenos Aires, Argentina. Rev. Argent. Microbiol. 2020;52:118–120. doi: 10.1016/j.ram.2019.06.005. [DOI] [PubMed] [Google Scholar]
  63. Mercken E., Van Damme I., Serradell A., Gabriël S. Presence of Anisakidae in commercial fish species imported into the Belgian food markets: a systematic review and meta-analyses. Int. J. Food Microbiol. 2020;318:108456. doi: 10.1016/j.ijfoodmicro.2019.108456. [DOI] [PubMed] [Google Scholar]
  64. Mercken E., Van Damme I., Vangeenberghe S., Serradell A., De Sterck T., Lumain J.P.L., Gabriël S. Ascaridoids in commercial fish: occurrence, intensity and localization in whole fish and fillets destined for the Belgian market. Int. J. Food Microbiol. 2020;327:108657. doi: 10.1016/j.ijfoodmicro.2020.108657. [DOI] [PubMed] [Google Scholar]
  65. Moravec F., Justine J.-L. Erection of Euterranova n. gen. and Neoterranova n. gen. (Nematoda, Anisakidae), with the description of E. dentiduplicata n. sp. and new records of two other anisakid nematodes from sharks off New Caledonia. Parasite. 2020;27:58. doi: 10.1051/parasite/2020053. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Moravec F., Nagasawa K., Madinabeitia I. Redescription of Dichelyne (Cucullanellus) jialaris (Nematoda: Cucullanidae), a parasite of seabreams (Perciformes: Sparidae) in East Asia. Acta Parasitol. 2018;63:802–807. doi: 10.1515/ap-2018-0097. [DOI] [PubMed] [Google Scholar]
  67. Moravec F., de Buron I., González-Solís D. Redescription of three species of nematodes (Nematoda) parasitising fishes in the USA, with a key to the species of Dichelyne Jägerskiöld, 1902 parasitic in freshwater and brackish-water fishes of North America. Syst. Parasitol. 2019;96:79–94. doi: 10.1007/s11230-018-9829-6. [DOI] [PubMed] [Google Scholar]
  68. Moschella C.M., Mattiucci S., Mingazzini P., De Angelis G., Assenza M., Lombardo F., Monaco S., Paggi L., Modini C. Intestinal anisakiasis in Italy: case report. J. Helminthol. 2004;78:271–273. doi: 10.1079/joh2004237. [DOI] [PubMed] [Google Scholar]
  69. Murata R., Suzuki J., Sadamasu K., Kai A. Morphological and molecular characterization of Anisakis larvae (Nematoda: Anisakidae) in Beryx splendens from Japanese waters. Parasitol. Int. 2011;60:193–198. doi: 10.1016/j.parint.2011.02.008. [DOI] [PubMed] [Google Scholar]
  70. Nadler S.A., D'Amelio S., Dailey M.D., Paggi L., Siu S., Sakanari J.A. Molecular phylogenetics and diagnosis of Anisakis, Pseudoterranova, and Contracaecum from northern Pacific marine mammals. J. Parasitol. 2005;91:1413–1430. doi: 10.1645/GE-522R.1. [DOI] [PubMed] [Google Scholar]
  71. Norriss J.V., Crisafulli B. Longevity in Australian snapper Pagrus auratus (Sparidae) J. R. Soc. West. Aust. 2010;93:129. [Google Scholar]
  72. Palm H.W., Theisen S., Damriyasa I.M., Kusmintarsih E.S., Oka I.B.M., Setyowati E.A., Suratma N.A., Wibowo S., Kleinertz S. Anisakis (Nematoda: Ascaridoidea) from Indonesia. Dis. Aquat. Org. 2017;123:141–157. doi: 10.3354/dao03091. [DOI] [PubMed] [Google Scholar]
  73. Paulin C.D. Pagrus auratus, a new combination for the species known as “snapper” in Australasian waters (Pisces: Sparidae) N. Z. J. Mar. Freshwat. Res. 1990;24:259–265. [Google Scholar]
  74. Petrie A., Wootten R., Bruno D., MacKenzie K., Bron J. Food Standards Scotland; Aberdeen, UK: 2007. A Survey of Anisakis and Pseudoterranova in Scottish Fisheries and the Efficacy of Current Detection Methods FSAS Project S14008. FSAS Project S14008. [Google Scholar]
  75. Petter A.J. Essai de classification de la famille des Cucullanidae. Bulletin du Muséum National d'Histoire Naturelle, Paris, 3 Series No. 255. Zoologie. 1974;177:1469–1490. [Google Scholar]
  76. Prober S.M., O'Connor M.H., Walsh F.J. Australian Aboriginal peoples' seasonal knowledge: a potential basis for shared understanding in environmental management. Ecol. Soc. 2011;16:12. [Google Scholar]
  77. Rambaut A. FigTree v1.4.2, a Graphical Viewer of Phylogenetic Trees. 2014. http://tree.bio.ed.ac.uk/software/figtree Accessed on 31 January 2019.
  78. Rezaei S., Pazooki J., Sharifpour I., Masoumian M. Histopathological observations in Neogobius bathybius (Actinopterygii: Gobiidae) infected by Dichelyne minutus (Nematoda: Cucullanidae) in the Caspian Sea, Iran. Turk. J. Zool. 2013;37:329–333. [Google Scholar]
  79. Ronquist F., Huelsenbeck J.P. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics. 2003;19:1572–1574. doi: 10.1093/bioinformatics/btg180. [DOI] [PubMed] [Google Scholar]
  80. Seo B.S., Chai J.Y., Lee S.H., Hong S.T., Seo J.W., Noh S.H. A human case infected by the larva of Terranova type A in Korea. Korean J. Parasitol. 1984;22:248–252. doi: 10.3347/kjp.1984.22.2.248. [DOI] [PubMed] [Google Scholar]
  81. Shamsi S. Recent advances in our knowledge of Australian anisakid nematodes. Int. J. Parasitol. Parasites Wildl. 2014;3:178–187. doi: 10.1016/j.ijppaw.2014.04.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  82. Shamsi S. Morphometric and molecular descriptions of three new species of Hysterothylacium (Nematoda: Raphidascarididae) from Australian marine fish. J. Helminthol. 2016;91:1–12. doi: 10.1017/S0022149X16000596. [DOI] [PubMed] [Google Scholar]
  83. Shamsi S. Seafood-borne parasites in Australia: human health risks, fact or fiction? Microbiol. Australia. 2020;41:33–37. [Google Scholar]
  84. Shamsi S., Sheorey H. Seafood-borne parasitic diseases in Australia: are they rare or underdiagnosed? Intern. Med. J. 2018;48:591–596. doi: 10.1111/imj.13786. [DOI] [PubMed] [Google Scholar]
  85. Shamsi S., Suthar J. Occurrence of Terranova larval types (Nematoda: Anisakidae) in Australian marine fish with comments on their specific identities. PeerJ. 2016;4 doi: 10.7717/peerj.1722. [DOI] [PMC free article] [PubMed] [Google Scholar]
  86. Shamsi S., Suthar J. A revised method of examining fish for infection with zoonotic nematode larvae. Int. J. Food Microbiol. 2016;227:13–16. doi: 10.1016/j.ijfoodmicro.2016.03.023. [DOI] [PubMed] [Google Scholar]
  87. Shamsi S., Gasser R., Beveridge I., Alizadeh Shabani A. Contracaecum pyripapillatum n. sp. (Nematoda: Anisakidae) and a description of C. multipapillatum (von Drasche, 1882) from the Australian pelican, Pelecanus conspicillatus. Parasitol. Res. 2008;103:1031–1039. doi: 10.1007/s00436-008-1088-z. [DOI] [PubMed] [Google Scholar]
  88. Shamsi S., Norman R., Gasser R., Beveridge I. Genetic and morphological evidences for the existence of sibling species within Contracaecum rudolphii (Hartwich, 1964) (Nematoda: Anisakidae) in Australia. Parasitol. Res. 2009;105:529–538. doi: 10.1007/s00436-009-1424-y. [DOI] [PubMed] [Google Scholar]
  89. Shamsi S., Norman R., Gasser R., Beveridge I. Redescription and genetic characterization of selected Contracaecum spp. (Nematoda: Anisakidae) from various hosts in Australia. Parasitol. Res. 2009;104:1507–1525. doi: 10.1007/s00436-009-1357-5. [DOI] [PubMed] [Google Scholar]
  90. Shamsi S., Chen Y., Poupa A., Ghadam M., Justine J.-L. Occurrence of anisakid parasites in marine fishes and whales off New Caledonia. Parasitol. Res. 2018;117:3195–3204. doi: 10.1007/s00436-018-6018-0. [DOI] [PubMed] [Google Scholar]
  91. Shamsi S., Steller E., Chen Y. New and known zoonotic nematode larvae within selected fish species from Queensland waters in Australia. Int. J. Food Microbiol. 2018;272:73–82. doi: 10.1016/j.ijfoodmicro.2018.03.007. [DOI] [PubMed] [Google Scholar]
  92. Shamsi S., Barton D.P., Zhu X. Description and characterisation of Terranova pectinolabiata n. sp.(Nematoda: Anisakidae) in great hammerhead shark, Sphyrna mokarran (Rüppell, 1837), in Australia. Parasitol. Res. 2019;118:2159–2168. doi: 10.1007/s00436-019-06360-4. [DOI] [PubMed] [Google Scholar]
  93. Shamsi S., Spröhnle-Barrera C., Hossen M.S. Occurrence of Anisakis spp. (Nematoda: Anisakidae) in a pygmy sperm whale Kogia breviceps (Cetacea: Kogiidae) in Australian waters. Dis. Aquat. Org. 2019;134:65–74. doi: 10.3354/dao03360. [DOI] [PubMed] [Google Scholar]
  94. Shamsi S., Dang M., Zhu X., Nowak B. Genetic and morphological characterization of Mawsonascaris vulvolacinata n. sp. (Nematoda: Anisakidae) and associated histopathology in a wild caught cowtail stingray, Pastinachus ater. J. Fish Dis. 2019;42:1047–1056. doi: 10.1111/jfd.13016. [DOI] [PubMed] [Google Scholar]
  95. Shamsi S., Barton D.P., Zhu X. Description and genetic characterisation of Pulchrascaris australis n. sp. in the scalloped hammerhead shark, Sphyrna lewini (Griffin & Smith) in Australian waters. Parasitol. Res. 2020:1–14. doi: 10.1007/s00436-020-06672-w. [DOI] [PubMed] [Google Scholar]
  96. Sharples A.D., Evans C.W. Metazoan parasites of the snapper, Pagrus auratus (Bloch & Schneider, 1801), in New Zealand: 1. Prevalence and abundance. N. Z. J. Mar. Freshwat. Res. 1995;29:195–201. [Google Scholar]
  97. Sharples A.D., Evans C.W. Metazoan parasites of the snapper, Pagrus auratus (Bloch & Schneider, 1801), in New Zealand: 2. Site-specificity. N. Z. J. Mar. Freshwat. Res. 1995;29:203–211. [Google Scholar]
  98. Sharples A.D., Evans C.W. Taxonomy of the metazoan parasites of the snapper Pagrus auratus in New Zealand: 2. Endoparasites. N. Z. J. Zool. 1995;22:163–174. [Google Scholar]
  99. Torres P., Jercic M.I., Weitz J., Dobrew E., Mercado R. Human Pseudoterranovosis, an emerging infection in Chile. J. Parasitol. 2007;93:440–443. doi: 10.1645/GE-946R.1. [DOI] [PubMed] [Google Scholar]
  100. Weitzel T., Sugiyama H., Yamasaki H., Ramirez C., Rosas R., Mercado R. Human infections with Pseudoterranova cattani nematodes, Chile. Emerg. Infect. Dis. 2015;21:1874. doi: 10.3201/eid2110.141848. [DOI] [PMC free article] [PubMed] [Google Scholar]
  101. Yamaguti S. Studies on the helminth fauna of Japan. Part 9. Nematodes of fishes, I. Jpn. J. Zool. 1935;6:337–386. doi: 10.1017/s0022149x00017788. [DOI] [PubMed] [Google Scholar]
  102. Yamaguti S. Studies on the helminth fauna of Japan. Part 33. Nematodes of fishes, II. Jpn. J. Zool. 1941;9:343–396. doi: 10.1017/s0022149x00017788. [DOI] [PubMed] [Google Scholar]
  103. Zhang K., Xu Z., Chen H.-X., Guo N., Li L. Anisakid and raphidascaridid nematodes (Ascaridoidea) infection in the important marine food-fish Lophius litulon (Jordan) (Lophiiformes: Lophiidae) Int. J. Food Microbiol. 2018;284:105–111. doi: 10.1016/j.ijfoodmicro.2018.08.002. [DOI] [PubMed] [Google Scholar]
  104. Zhu X.Q., D'Amelio S., Palm H.W., George-Nascimento M., Paggi L., Gasser R. SSCP-based identification of members within the Pseudoterranova decipiens complex (Nematoda: Ascaridoidea: Anisakidae) using genetic markers in the internal transcribed spacers of ribosomal DNA. Parasitology. 2002;124:615–623. doi: 10.1017/s0031182002001579. [DOI] [PubMed] [Google Scholar]

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