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. 2020 Apr 24;44(3):536–545. doi: 10.1007/s12639-020-01222-8

Occurrence and molecular identification of Anisakis larval type 1 (Nematoda: Anisakidae) in marketed fish in Egypt

Eman Mostafa 1, Marwa Omar 1, Shimaa S Hassan 2, Mohamed Samir 3,✉
PMCID: PMC7410882  PMID: 32801505

Abstract

Anisakidosis is a zoonotic infection caused by members of the family Anisakidae. The presence of anisakid larvae in fish poses risk for humans and dissuade consumers from purchasing infected products. Although fish constitute important component of Egyptian diet, the prevalence of anisakid larvae in marketed fish in Egypt is not well described. Furthermore, the species of anisakid larvae is not defined in most of the available studies due to the over reliance on morphological analyses. The aim of the current work was to assess the prevalence and intensity of anisakid larvae in three common marketed fish in Egypt (Atlantic herring, Mediterranean horse mackerel and Atlantic mackerel) and to determine the species of the isolated larvae using morphological and molecular methods. Light and scanning electron microscope (SEM) analyses revealed the details of the isolated larvae. However, partial sequencing of cytochrome oxidase subunite-1 (mt cox1) gene revealed that all larvae isolated from Atlantic herring and Mediterranean horse mackerel belonged to Anisakis simplex sensu stricto with prevalence of 87.1% and 83.3%, respectively, whereas Atlantic mackerel harbored Anisakis typica with a prevalence of 42.8%. The Mediterranean horse mackerel demonstrated the highest larval mean intensity (n = 20 larvae/infected fish). This study highlights the importance of these fish as potential reservoirs for human anisakiasis in Egypt and possibly in other coastal countries.

Electronic supplementary material

The online version of this article (10.1007/s12639-020-01222-8) contains supplementary material, which is available to authorized users.

Keywords: A. simplex, Sequencing, Atlantic mackerel, Mediterranean horse mackerel, Atlantic herring

Introduction

The World Health Organization (WHO) reported that around 56 million human parasitic infections are due to consumption of infected fish products (WHO 2012). Infection with anisakid nematode is one fish-borne parasitic infection, where the larvae are known to infect broad range of fish and are capable of infecting humans, causing severe pathologies (Pampiglione et al. 2002). Human infection with anisakid worm has been reported worldwide, notably in Egypt (Aibinu et al. 2019). The adult worm lives in the intestine of large marine mammals (e.g., dolphins and whales). When the gravid female releases eggs into water, the larvae hatch from eggs and are further eaten by crustaceans, which are then ingested by marine fish (Palm et al. 2017), where it resides mostly in visceral organs (Abou-Rahma et al. 2016; Bahlool et al. 2012). Humans acquire the infection accidentally by ingesting undercooked fish harboring anisakid L3 (Baptista-Fernandes et al. 2017). In humans, the larvae could induce inflammation in the stomach and/or intestine (Baptista-Fernandes et al. 2017), augment cancer formation (Garcia-Perez et al. 2015) or lead to allergy and hypersensitivity even if they were dead upon ingestion (Audicana et al. 2002; de Corres et al. 1996).

Anisakis larval type 1 include Anisakis simplex complex [encompassing Anisakis simplex sensu stricto (s.s), A. pegreffii and A. berlandi], A. typica, A. ziphidarum and A. nascettii, whereas Anisakis larval type II include A. paggiae, A. physeteris and A. brevispiculata (Mattiucci and Nascetti 2008; Mattiucci et al. 2009). Other recently identified species in South east Asia is Anisakis typica var. indonesiensis (Palm et al. 2017).

Anisakis larval type 1, particularly A. simplex, A. pegreffii and A. typica, have been isolated from a wide variety of fish all over the world (Costa et al. 2016), including mackerel (Eissa et al. 2018), European anchovy (Mladineo et al. 2012), European hake (Cipriani et al. 2015) and Atlantic herring (McGladdery 2011). In Egypt, the prevalence of Anisakis larvae exhibited variation depending on the fish host, which reached 36.6% in European hake (Abou-Rahma et al. 2016), 92.3% in mackerel (Abdel-ghany 2011), 23–72.8% in Mediterranean sand smelt (Abdel-ghany 2011; Samir et al. 2015), 70% in Orange spotted trevally (Arafa et al. 2009), 75% in greater lizard fish (Morsy et al. 2015), 19% in smoked herring and 42.8% in mackerel fish (Arafa et al. 2019). However, the prevalence of anisakid larvae in other commonly consumed fish in Egypt such as Atlantic herring (Clupea harengus), Mediterranean horse mackerel (Trachurus trachurus) and Atlantic mackerel (Scomber scombrus) has received much less attention. In addition, studies done on Anisakis larvae that were isolated from marketed fish in Egypt did not define the species of the isolated larvae (Abdel-ghany 2011; Samir et al. 2015), which is an important issue in determining the extent of risk for human consumers (Suzuki et al. 2010). When it comes to the molecular diagnosis of anisakid worm, mitochondrial cytochrome oxidase subunite-1 (mt cox1) gene has been frequently used as marker for species identification (Sohn et al. 2014; Song et al. 2019). Sequencing of this gene and analyzing the extent of its similarities among anisakid worm could lead to better determination of larvae species, which in this regard would be advantageous compared to using morphological assessments. The aim of the present study was to determine the occurrence and species of anisakid larvae found in fish hosts that are commonly sold in Egyptian markets. The morphology of the isolated larvae was described by both light and scanning electron microscope (SEM) and their species were determined by sequencing of mt cox1.

Material and methods

Fish sample collection and processing

Before sampling, sample size was estimated for each fish species based on an expected margin error (confidence interval) of 5%, confidence level at 95% and expected prevalence (true positive rate) of 40%. A correction for cluster bias (the bias generated when taking fish samples from the same market) was done by incorporating “dff” argument in the applied R function. This analysis was done using function “n.for.survey” in the package “epiDisplay”, within R software (v 3.6.2) (Chongsuvivatwong 2015). Four hundred and twelve marine fish belonging to three different fish species (Fig. S1) were collected from local fish markets in El-sharkia Province (Zagazig city), Egypt from March 2017 through December 2017. These included 140 Atlantic herring (Clupea harengus), 132 Mediterranean horse mackerel (Trachurus trachurus) and 140 Atlantic mackerel (Scomber scombrus). Fish collection was done through ~ 35 visits (1 visit/week). In each visit, ~ four fish from each species were collected. In some visits, less or more samples were collected due to market condition. All examined fish were imported. The examined Mediterranean horse mackerel and Atlantic mackerel fish were sold frozen, whereas the examined Atlantic herring fish were sold as smoked fish. Fish species identification was done by help of trained fish parasitologist at the Faculty of Veterinary Medicine, Zagazig University. For larval isolation, the collected fish were transferred on ice to the laboratory of Parasitology, Faculty of Medicine, Zagazig University. After making an incision from the anus to mouth opening, fish muscles, body cavities and viscera were separated. Quantitative assessment of larval occurrence in fish was done by calculating the prevalence (P) = number of infected fish/number of examined fish, mean intensity (MI) = total number of isolated larvae per fish species/number of infected fish belonging to this species and mean abundance (MA) = total number of isolated larvae per fish species/number of examined fish belonging to this species (Bush et al. 1997). All the collected larvae were processed for microscopic examination to differentiate type I (anisakid larvae) and type II larvae.

Morphological examination of isolated larvae using light microscope

The larvae isolated form each fish species were washed in saline solution for 30 min, fixed in alcohol-glycerin hot fixative (Garcia 1979), temporarily mounted in pure glycerin (Garcia 2007) and then were examined by ordinary light microscopy. All the larvae were assessed morphometrically and morphologically as previously described (Cannon 1977; Mattiucci et al. 2014; Shih 2004).

Scanning electron microscopic (SEM) examination of isolated larvae

To provide more insight into the larval fine details, specimens isolated from each fish type were examined using SEM as described previously (Mafra et al. 2015; Roongruangchai et al. 2012). Finally, larvae were examined and photographed using a JEOL JSM-6510 1v SEM at the Faculty of Agriculture, Mansoura University, Egypt.

Molecular characterization of the isolated anisakid larvae

Larvae were collected from each of the 3-fish hosts, pooled and washed 3-times by phosphate buffer saline (PBS) and then incubated with 500 ml PBS in a sterile 1.5 ml Eppendorf tubes at room temperature. For molecular characterization, randomly selected larvae from each fish were pooled into 6–7 pools, each contains ~ 100 identical larvae. The larvae pools were homogenized in PBS using metal-bar homogenizer (Wisetis, HG-15D, Wertheim, Germany). The subsequent steps were done for each pool. Genomic DNA was extracted from the homogenate using QIAamp DNA Mini Kit, Qiagen (Cat. Nr. 51304) according to the manufacture recommendation. The extracted DNA was quantified using Quantus™ Fluorometer system (Cat. Nr. E6150, Promega 2800 Woods Hollow Road· Madison, WI 53711-5399 USA) according to the manufacture instructions. To determine the species of the larvae, polymerase chain reaction (PCR) followed by sequencing was done. PCR technique was used to amplify a partial sequence (approximately 450 base pair) of mt cox1 gene. The used primer pair was as follow: Forward: 5′ TTTTTTGGGCATCCTGAGGTTTAT 3′, Reverse: 3′ TAAAGAAAGAACATAATGAAAATG 5′ (Bowles et al. 1993; Hu et al. 2001; Sohn et al. 2014). The primer pair was checked for specificity by running a real time PCR against a serially diluted cDNA samples and the melting curve values were determined. Amplification was conducted in Veriti 96-Well Thermal Cycle (Applied bioscience, cat No. 4375786). PCR products were run on 2% ethidium bromide-stained agarose gels and the positive bands were visualized by UV transilluminator (JY02, Cleaver, UK).

To sequence the amplified mt cox1 gene, the amplified PCR product was gel-purified using QIAquick PCR Purification Kit (Qiagen, Germany, cat. Nr. 28104) and directly sequenced in fluorescence-based cycle sequencer using BigDye™ Terminator v3.1 Cycle Sequencing Kit (Applied bioscience, USA, cat Nr. 4337455). The obtained nucleotide (nt) sequences were checked for quality using BioEdit sequence alignment editor (version 7.2.0) and Chromas (version 2.6.5) software. The generated nucleotide sequences for mt cox1 gene were deposited in NCBI database under accession numbers: MH363812, MH363813 and MH363814.

Phylogenetic and sequence identity analyses

For the sake of credibility and to exclude any bias due to lack of accuracy in morphological analyses, the cDNA isolated from each larvae pool was used for sequencing of cox1 gene to determine the species of the collected larvae via phylogenetic analyses. The nucleotide sequences of our larvae along with other 22 sequences of mt cox1 gene collected from NCBI database (Geer et al. 2010) and representing different species within genus Anisakis (Anisakidae family) were used to build the phylogenetic tree. The evolutionary history of the larvae was inferred using the BioNJ algorithm (Gascuel 1997), a variant of the Neighbor-Joining algorithm (Saitou and Nei 1987). The evolutionary distances were computed using the Kimura method (Kimura 1980). In order to assess the divergence and identity among the analyzed sequences, a Clustal W multiple alignment was done (Thompson et al. 1994) and the identity and divergence values were calculated among pairs of sequences. Divergence is calculated by comparing sequence pairs in relation to the phylogeny. This analysis was done using the MegAlign software in the Lasergene v 7.1.0 package (DNASTAR, Madison, Wisconsin, USA).

Results

Prevalence and intensity of isolated anisakid larvae and their morphological characteristics

The prevalence (P), mean intensity (MI) and mean abundance (MA) of anisakid larvae are shown in Table 1. The overall prevalence of anisakid larvae in Atlantic herring and Atlantic horse mackerel exceeded 80% and reached 42.8% in Atlantic mackerel. The monthly prevalence of anisakid larvae in Atlantic herring ranged from 71.4 to 100% in Mediterranean horse mackerel from 60 to 100% and in Atlantic mackerel from 13.3 to 61%. There was no strong variation in the prevalence of the anisakid larvae in the three examined fish in different months (Table S3). Out of 3722 isolated anisakid larvae, 1342 (36%) larvae were found in Atlantic herring, 2200 (59.6%) larvae were collected from Mediterranean horse mackerel and 180 (4.8%) larvae were collected from Atlantic Mackerel. Mediterranean horse mackerel contained the highest larval MI (20 larvae/infected fish) followed by Atlantic herring (11 larvae/infected fish) and Atlantic mackerel (3 larvae/infected fish) (Table 1). Most of anisakid larvae were found as encapsulated larvae in fish muscle and visceral organs including the peritoneum (Fig. S2). Using ordinary light microscope and SEM, all isolated larvae revealed features that resemble Anisakis larval type 1, particularly the presence of long ventriculus and mucron at the larval posterior end (Figs. 1 and 2 and Table S1).

Table 1.

Prevalence (P), mean intensity (MI) and mean abundance (MA) of isolated anisakid larvae in the examined fish

Fish host No. examined (n = 412) No. parasitized No. larvae isolated P MI MA
Atlantic herring (Clupea harengus) 140 122 1342 87.1% 11 9.5
Mediterranean horse mackerel (Trachurus mediterraneus) 132 110 2200 83.3% 20 16.6
Atlantic mackerel (Scomber scombrus) 140 60 180 42.8% 3 2.7
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The parameters of parasite burden were calculated as described in the “Material and method” section

Fig. 1.

Fig. 1

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Morphology of the isolated larvae. a Anterior end of A. simplex (s.s) isolated from Atlantic herring. b Anterior end of A. simplex (s.s) isolated from Mediterranean horse mackerel. c Anterior end of A. typica isolated from Atlantic mackerel. d Body of A. simplex (s.s) isolated from Atlantic herring. e Body of A. simplex (s.s) isolated from Mediterranean horse mackerel. f Body of A. typica isolated from Atlantic mackerel. g Posterior end of A. simplex (s.s) isolated from Atlantic herring. h Posterior end A. typica isolated from Atlantic mackerel. i Posterior end of A. typica isolated from Atlantic mackerel. P papillae, bt boring tooth, ep excretory pore, e eosophagus, ed excretory duct, ve ventriculus, mu mucron, ao anal opening, ag anal gland, ta transverse annulation

Fig. 2.

Fig. 2

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Scanning electron microscopy images of the isolated third-stage anisakid larvae. a Anterior end of A. typica isolated from Atlantic mackerel. b Posterior end of A. typica isolated from Atlantic mackerel. c Body of A. typica isolated from Atlantic mackerel. d Anterior end of A. simplex (s.s.) isolated from Mediterranean horse mackerel. e Posterior end of A. simplex (s.s) isolated from Mediterranean horse mackerel. f Posterior end of A. simplex (s.s.) isolated form Atlantic herring showing mucron. P papillae, bt boring tooth, ep excretory pore, ao anal opening, mu mucron, ta transverse annulation

Larvae isolated from Atlantic herring (Clupea harengus) and Mediterranean horse mackerel (Trachurus mediterraneus)

The L3 recovered from Atlantic herring and Mediterranean horse mackerel shared similar morphological features (Fig. 1a, b, d, e, g and Table S1). The body of the recovered L3 was cylindrical in shape. Body length ranged from 14.3 to 25.5 mm and body width ranged from 0.43 to 0.62 mm. The body covered with a rigid cuticle and annular transverse striations that start from cephalic to caudal end of the larvae (Figs. 1g, 2f). As shown in Figs. 1a, b and 2d, the cephalic end was flattened and carried four prominent labial papillae (two dorsolateral and two ventrolateral) surrounding the mouth opening and boring tooth. Excretory pore was prominent at the lateral part of the cephalic region. The esophagus extended downward the mouth opening and consisted of an anterior muscular part and a glandular long ventriculus (Fig. 1a). The esophago-ventricular junction was oblique (Fig. 1d, e). No intestinal caeca or diverticula were present. The caudal part of the larvae carried the characteristic mucron (length ranged from 0.018 to 0.03 mm) and anal glands opened into anal opening (Figs. 1g, h, 2e, f). The larval cuticle was irregularly wrinkled near the tail and had clear annular striations.

Larvae isolated from Atlantic mackerel (Scomber scombrus)

The L3 isolated from Atlantic Mackerel had a cylindrical body measuring 14.87–22 mm in length and 0.38–0.56 mm in width. The cuticle was transversely striated (Fig. 2c). The cephalic end was pointed (Figs. 1c, 2a) and had a characteristic boring tooth that was located near the ventral part of the dorsal lip. The excretory pore opened laterally into the excretory duct. The long ventriculus measured 0.47–0.75 mm in length and the esophago-ventricular junction was oblique (Fig. 1f). The tail was short, bluntly rounded and had a small mucron (length = 0.08–0.18 mm) (Figs. 1h, i, 2b). Anal glands opened into the anal opening. The larval cuticle was irregularly wrinkled near the tail and had clear annular striations (Figs. 1i, 2f).

Molecular identification of isolated anisakid larvae

Because the larvae within the 6–7 pools in each fish species revealed almost similar sequences of cox1 gene, we used 3 cox1 gene sequences of one larval pool from each fish host in this analyses. The phylogenetic analysis (Fig. 3) revealed that all the studied larvae belonged to type-1 anisakid larvae. Larvae isolated from Atlantic herring (MH363812) and Mediterranean horse mackerel (MH363813) were clustered close to each other (Similarity = 97.1%) and were classified as A. simplex (s.s.) because they were clustered close to other A. simplex (s.s.) larvae (e.g. Accession numbers: GQ132132.1, GQ132126.1, GQ132130.1 and JN786321.1). In support of this, larvae isolated from Atlantic herring (MH363812) showed the highest similarity to previously isolated A. simplex (s.s.) larvae GQ132132.1 (99.4%) and GQ132126 (99.4%). Similarly, larvae isolated from Mediterranean horse mackerel (MH363813) showed the highest similarity (97.4%) to the same previously mentioned A. simplex (s.s.). On the other hand, the L3 harvested from Atlantic Mackerel (MH363814) were classified as A. typica as they were positioned very close to previously isolated A. typica larvae (e.g., KF693786.1 and KF693785.1). The larvae isolated from Atlantic mackerel larvae exhibited the highest similarity to A. typica larvae KF693786.1 (99.3%) and KF356666.1 (98.6%) (Figs. 3, 4). The larvae isolated from Atlantic herring (MH363812) and Mediterranean horse mackerel (MH363813) showed an identity of 95 and 93%, respectively to that isolated from Atlantic mackerel (MH363814) (Fig. 4).

Fig. 3.

Fig. 3

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Phylogenetic relatedness of the study isolates (black box beside) to other 22 isolates belonging to anisakid family. This phylogeny was constructed based on analyzing partial sequences of the mt cox1 gene. The nucleotide sequences were retrieved from the NCBI database (https://www.ncbi.nlm.nih.gov/pubmed/) after performing a blast search

Fig. 4.

Fig. 4

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Sequence distance matrix showing the percentages of identity and divergence among the analyzed mt cox1 gene sequences. The study isolates are indicated with asterisks beside. Each of the sequences represent a pooled larvae from each fish host. The sequences were retrieved from the NCBI database (https://www.ncbi.nlm.nih.gov/pubmed/)

Discussion

The aim of the current study was to determine the occurrence of anisakis larvae in Atlantic herring, Mediterranean horse mackerel and Atlantic mackerel fish, three fish that are widely sold in Egyptian markets. A. simplex (s.s.) larvae were highly prevalent in examined Atlantic herring and Mediterranean horse mackerel, whereas A. typica dominated the larvae population in examined Atlantic mackerel, suggesting the importance of these fish as sources for human anisakiasis in Egypt.

The long ventriculus and presence of mucron in all examined larvae indicated that these larvae belonged to Anisakis larval type 1. Our morphological analyses support and agree with previous reports (Borges et al. 2012; Nada and El-Ghany 2011; Rocka 2004) as well as in concordance with the previously characterized Anisakis spp. (Listed in table S1). The data showed that larvae of A. simplex (s.s.) and A. typica have somehow overlapped morphometric features (Table S1), indicating that morphological analyses might not be a suitable tool for identifying larval species. This highlights the importance of reliance on advanced molecular tools (e.g., sequencing) in defining larvae species. Although A. simplex (s.s.) is a common fish parasite and is associated with human disease (Baptista-Fernandes et al. 2017; Li et al. 2015), studies done on the prevalence of such nematodes in Egyptian marketed fish, such as the ones studied here, still limited. In Egypt, Atlantic herring is a common fish diet that is consumed as a whole or in a form of salad. In the current study, we showed that prevalence of A. simplex (s.s.) in marketed Atlantic herring was 87.1% with a MI of 11 larvae/infected fish (Table 1), suggesting the importance of this fish as a source of human anisakiasis. Our estimated prevalence is far higher than that reported recently by Arafa et al. (Arafa et al. 2019), which stated that the prevalence of A. simplex (s.s.) in Herring was 19.05%. We assume that the low prevalence in the study done by Arafa et al. is due to his small sample size (n = 84 herring fish) compared to our sample size. Surprisingly, some studies done in Egypt reported high prevalence of anisakid larvae reaching 97.4% in smoked and 88.51% in frozen herring (Abdel-ghany 2011), yet the authors did not define the species of the larvae. The variation in prevalence of Anisakis larvae in fish might be attributed to differences in fish species and origin, detection method and other ecological factors (Bak et al. 2014). It is worth mentioning that most of the examined herring fish in Egypt are consumed as ready to eat smoked product, which poses more risk for humans, yet this depends on the efficiency of smoking process in eliminating the anisakid larvae. Higher prevalence (98–100%) was previously reported in various sizes of Norwegian spring spawning herring (Clupea harengus L.), which live in the northeast Atlantic water (Levsen and Lunestad 2010). In a different study, 15–60% of Clupea harengusL. has been shown to harbor A. simplex (s.s.) (Levsen et al. 2005). In the current study, the prevalence of A. simplex (s.s.) in Mediterranean horse mackerel was comparable to that of herring fish (83.3%) with higher larval MI. The high MI of A. simplex (s.s.) in examined Mediterranean horse mackerel highlights the importance of this fish as a source of infection for human. Indeed, previous studies reported that Mediterranean horse mackerel could harbor other Anisakid spp. (e.g., A. pegreffii) (Casti et al. 2017), with a prevalence reached 32.7% with much lower MI (MI = 3). It is worth mentioning that Trachurus trachurus (Atlantic horse mackerel), a closely related species to Mediterranean horse mackerel, could also harbor A. simplex (s.s.) larvae with a prevalence ranging from 35 (Aziz 2017) to 52.5% (Angelucci et al. 2011) and that MI of larvae in this fish host lies in the same range as the MI shown in our study (15.5–39.9 larvae/infected fish) (Abollo et al. 2001; Tantanasi et al. 2012). This suggests that Mediterranean or Atlantic stocks of Trachurus fish represent potential reservoirs of A. simplex (s.s.). It would be interesting to investigate the prevalence of A. simplex (s.s.) in various imported stocks of Trachurus fish that are commonly sold in Egyptian markets.

Fish belonging to genus Scomber (mackerel) are widely distributed and exhibit complex diversity (Zardoya et al. 2004). However, Atlantic mackerel (Scomber scombrus) is the most widely consumed Scomber fish in Egyptian market chain. Here, we provide first clue on the occurrence of A. typica in this fish that is imported to and sold in Egyptian markets. The prevalence of A. typica larvae in Atlantic mackerel was 42.8% and the MI was low (3 larvae/infected fish) (Table 1). Actually, Mackerel fish imported to Egypt is documented to harbor other type 1 anisakis larvae [e.g., A. simplex (s.s.) and A. pegreffii] as reported in Egypt by Arafa et al., wherein the prevalence of the larvae reached levels (42.86%) similar to ours (Arafa et al. 2019). This suggests the high probability of occurrence of A. simplex type 1 in marketed mackerel fish in Egyptian markets. Only a single report documented the presence of one A. typica larvae in Atlantic mackerel caught off the Mediterranean coast of Gabes city, Tunisia. (Farjallah et al. 2008). It is plausible to assume that the low MI of A. typica in Atlantic mackerel might be correlated to the lack of reports on this fish because of potential underestimation of the infection and ignoring some of the infected fish during the inspection process at various importation portals as suggested previously (Gazzonis et al. 2017). In the current study, we could not observe any specific pattern in the prevalence of anisakid larvae in the examined fish (Table S3). Several peaks and drops were observed in various months. This indicates that there is no seasonal occurrence of anisakid larvae.

While morphological analyses could provide initial clue about larval species, more advanced molecular tools could provide decisive identification of the species. In the current study, partial sequencing of mt cox1 gene enabled classifying the isolated anisakis larvae in spite of the similarities observed at the morphological level. Larvae isolated from Mediterranean horse mackerel and Atlantic herring were clustered with that of A. simplex (s.s.), and clearly separated from the larvae isolated from Atlantic mackerel (similarity ranged from 93 to 95%), which was classified as A. typica. It is interesting to observe that the larvae isolated from Atlantic herring (MH363812) and Mediterranean horse mackerel (MH363813) showed the highest similarity to the same A. simplex (s.s.) larvae that were previously deposited in NCBI (99.4% and 97.4% similarity to GQ132132.1 and GQ132126., respectively). Interestingly, one A. simplex (s.s.) isolate (JN78632.1.1), which clustered close to our Atlantic herring isolate was also isolated from Clupea harengus (Table S2).

In the current study, the sequences of the isolated A. typica showed the highest similarity to previously published sequences of A. typica (KF693786.1; 99.3% similarity and KF356669.1; 98.6% similarity), suggesting that the species parasitizing Atlantic mackerel is mostly A. typica. All the Atlantic mackerel and Mediterranean horse mackerel sold in Egypt are imported from fish stocks that live in and caught off Atlantic, pacific oceans and Mediterranean sea. It will now be important to consider the variability in prevalence of anisakid larvae in these imported fish hosts and how this threatens the human health.

Conclusion

From this study, it could be concluded that A. simplex (s.s.) larvae are highly prevalent in Atlantic herring and Mediterranean horse mackerel, whereas A. typica dominated Atlantic mackerel fish. It was also obvious that larval species are best defined using molecular tools such as sequencing. These data represent an important addition to the exciting reports on Anisakis larvae in fish in Egypt and are highly relevant to the public health threat posed by these larvae.

Electronic supplementary material

Below is the link to the electronic supplementary material.

12639_2020_1222_MOESM1_ESM.tif (461.6KB, tif)

Fig S1. Fish hosts examined in the present study. a. Atlantic Herring (Clupea harengus). b. Mediterranean horse mackerel (Trachurus mediterraneus). c. Atlantic mackerel (Scomber scombrus) (TIF 461 kb)

12639_2020_1222_MOESM2_ESM.tif (659.1KB, tif)

Fig S2. a. Mediterranean horse mackerel heavily infected with Anisakis simplex (s.s) L3 found encysted in back muscle (arrow). b. Anisakis simplex (s.s) found localized in the musculature of Atlantic herring fish (arrow) (TIF 659 kb)

Supplementary file3 (DOCX 21 kb) (21.4KB, docx)
Supplementary file4 (DOCX 17 kb) (17.9KB, docx)
Supplementary file5 (XLSX 10 kb) (10.7KB, xlsx)

Acknowledgements

The authors would like to thank Dr. Amany abdel-ghany (Parasitology Department, Faculty of Veterinary Medicine, Zagazig University) for helping in fish species identification.

Author contribution

MS conceptualized the study idea. MS, EM, MO, SH participated in study design. All authors participated in the experimental work, wrote the initial draft and revised the final version of the manuscript.

Funding

This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

Compliance with ethical standards

Conflict of interest

The authors declare that they have no competing interest.

Footnotes

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  1. Abdel-ghany A (2011) Parasitological and immunological studies on anisakid parasites. PhD, Zagazig, Egypt
  2. Abollo E, Gestal C, Pascual S. Anisakis infestation in marine fish and cephalopods from Galician waters: an updated perspective. Parasitol Res. 2001;87:492–499. doi: 10.1007/s004360100389. [DOI] [PubMed] [Google Scholar]
  3. Abou-Rahma Y, Abdel-Gaber R, Kamal Ahmed A. First record of Anisakis simplex third-stage larvae (Nematoda, Anisakidae) in European Hake Merluccius merluccius lessepsianus in Egyptian water. J Parasitol Res. 2016;2016:9609752. doi: 10.1155/2016/9609752. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Aibinu IE, Smooker PM, Lopata AL. 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]
  5. Angelucci G, et al. Prevalence of Anisakis spp. and Hysterothylacium spp. larvae in teleosts and cephalopods sampled from waters off. Sardinia J Food Protect. 2011;74:1769–1775. doi: 10.4315/0362-028X.JFP-10-482. [DOI] [PubMed] [Google Scholar]
  6. Arafa SZ, Al-Hoot AA, Hussein HS. Pathological and ultrastructural studies on anisakis simplex rudolphi-1809 infecting Carangoides bajad with special reference to intestinal maturation in puppies. J Egypt Soc Parasitol. 2009;39:607–616. [PubMed] [Google Scholar]
  7. Arafa WM, Hassan AHA, Mahrous LN, Abdel-Ghany AE, Aboelhadid SM. Occurrence and molecular characterization of zoonotic Anisakis simplex sensu stricto and Anisakis pegreffii larvae in retail-marketed fish. J Food Saf. 2019;39:e12682. doi: 10.1111/jfs.12682. [DOI] [Google Scholar]
  8. Audicana MT, Ansotegui IJ, de Corres LF, Kennedy MW. Anisakis simplex: dangerous—dead and alive? Trends Parasitol. 2002;18:20–25. doi: 10.1016/S1471-4922(01)02152-3. [DOI] [PubMed] [Google Scholar]
  9. Aziz EA. Anisakis simplex (Nematoda: Anisakidae) from horse mackerel (Trachurus trachurus) in Atlantic coast of Morocco. Asian Pac J Trop Dis. 2017;7:463–466. doi: 10.12980/apjtd.7.2017D6-439. [DOI] [Google Scholar]
  10. Bahlool QZ, Skovgaard A, Kania P, Haarder S, Buchmann K. Microhabitat preference of Anisakis simplex in three salmonid species: immunological implications. Vet Parasitol. 2012;190:489–495. doi: 10.1016/j.vetpar.2012.07.009. [DOI] [PubMed] [Google Scholar]
  11. Bak TJ, Jeon CH, Kim JH. Occurrence of anisakid nematode larvae in chub mackerel (Scomber japonicus) caught off. Korea Int J Food Microbiol. 2014;191:149–156. doi: 10.1016/j.ijfoodmicro.2014.09.002. [DOI] [PubMed] [Google Scholar]
  12. Baptista-Fernandes T, et al. Human gastric hyperinfection by Anisakis simplex: a severe and unusual presentation and a brief review. Int J Infect Dis IJID Off Publ Int Soc Infect Dis. 2017;64:38–41. doi: 10.1016/j.ijid.2017.08.012. [DOI] [PubMed] [Google Scholar]
  13. Borges JN, Cunha LFG, Santos HLC, Monteiro-Neto C, Santos CP. Morphological and molecular diagnosis of Anisakid nematode Larvae from Cutlassfish (Trichiurus lepturus) off the Coast of Rio de Janeiro, Brazil. PLoS ONE. 2012;7:e40447. doi: 10.1371/journal.pone.0040447. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Bowles J, Hope M, Tiu WU, Liu X, McManus DP. Nuclear and mitochondrial genetic markers highly conserved between Chinese and Philippine Schistosoma japonicum. Acta Trop. 1993;55:217–229. doi: 10.1016/0001-706X(93)90079-Q. [DOI] [PubMed] [Google Scholar]
  15. Bush AO, Lafferty KD, Lotz JM, Shostak AW. Parasitology meets ecology on its own terms: Margolis et al. revisited. J Parasitol. 1997;83:575–583. doi: 10.2307/3284227. [DOI] [PubMed] [Google Scholar]
  16. Cannon LRG. Some larval ascaridoids from south-eastern queensland marine fishes. Int J Parasitol. 1977;7:233–243. doi: 10.1016/0020-7519(77)90053-4. [DOI] [PubMed] [Google Scholar]
  17. Casti D, et al. Occurrence of nematodes of the genus Anisakis in Mediterranean and Atlantic Fish Marketed in Sardinia. Italian J Food Saf. 2017;6:6185. doi: 10.4081/ijfs.2017.6185. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Chongsuvivatwong V (2015) epiDisplay: epidemiological data display package. https://cranr-projectorg/web/packages/epiDisplay/index.html. Accessed 1 Dec 2018
  19. Cipriani P, et al. Genetic identification and distribution of the parasitic larvae of Anisakis pegreffii and Anisakis simplex (s. s.) in European hake Merluccius merluccius from the Tyrrhenian Sea and Spanish Atlantic coast: implications for food safety. Int J Food Microbiol. 2015;198:1–8. doi: 10.1016/j.ijfoodmicro.2014.11.019. [DOI] [PubMed] [Google Scholar]
  20. Costa A, et al. Survey on the presence of A. simplex s.s. and A. pegreffii hybrid forms in Central-Western Mediterranean Sea. Parasitol Int. 2016;65:696–701. doi: 10.1016/j.parint.2016.08.004. [DOI] [PubMed] [Google Scholar]
  21. de Corres FL, et al. Anisakis simplex induces not only anisakiasis: report on 28 cases of allergy caused by this nematode. J Investigat Allergol Clin Immunol. 1996;6:315–319. [PubMed] [Google Scholar]
  22. Eissa AE, Showehdi ML, Ismail MM, El-Naas AS, Abu Mhara AA, Abolghait SK. Identification and prevalence of Anisakis pegreffii and A. pegreffii × A. simplex (s.s.) hybrid genotype larvae in Atlantic horse Mackerel (Trachurus trachurus) from some North African Mediterranean coasts. Egypt J Aquat Res. 2018 doi: 10.1016/j.ejar.2018.02.004. [DOI] [Google Scholar]
  23. Farjallah S, et al. Occurrence and molecular identification of Anisakis spp. from the North African coasts of Mediterranean Sea. Parasitol Res. 2008;102:371–379. doi: 10.1007/s00436-007-0771-9. [DOI] [PubMed] [Google Scholar]
  24. Garcia LS. Diagnostic parasitology clinical laboratory manual. 2. St. Louis: C. V. Mosby Co.; 1979. [Google Scholar]
  25. Garcia LS. Diagnostic medical parasitology. 5. Santa Monica: American Society of Microbiology; 2007. [Google Scholar]
  26. Garcia-Perez JC, Rodríguez-Perez R, Ballestero A, Zuloaga J, Fernandez-Puntero B, Arias-Díaz J, Caballero ML. Previous exposure to the fish parasite Anisakis as a potential risk factor for gastric or colon adenocarcinoma. Medicine. 2015;94:e1699–e1699. doi: 10.1097/MD.0000000000001699. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Gascuel O. BIONJ: an improved version of the NJ algorithm based on a simple model of sequence data. Mol Biol Evol. 1997;14:685–695. doi: 10.1093/oxfordjournals.molbev.a025808. [DOI] [PubMed] [Google Scholar]
  28. Gazzonis AL, et al. Anisakis sp. and Hysterothylacium sp. larvae in anchovies (Engraulis encrasicolus) and chub mackerel (Scomber colias) in the Mediterranean Sea: molecular identification and risk factors. Food Control. 2017;80:366–373. doi: 10.1016/j.foodcont.2017.05.004. [DOI] [Google Scholar]
  29. Geer LY, et al. The NCBI BioSystems database. Nucleic Acids Res. 2010;38:D492–496. doi: 10.1093/nar/gkp858. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Hu M, Damelio S, Zhu X, Paggi L, Gasser R. Mutation scanning for sequence variation in three mitochondrial DNA regions for members of the Contracaecum osculatum (Nematoda: Ascaridoidea) complex. Electrophoresis. 2001;22:1069–1075. doi: 10.1002/1522-2683()22:6<1069::AID-ELPS1069>3.0.CO;2-T. [DOI] [PubMed] [Google Scholar]
  31. Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;16:111–120. doi: 10.1007/BF01731581. [DOI] [PubMed] [Google Scholar]
  32. Levsen A, Lunestad BT. Anisakis simplex third stage larvae in Norwegian spring spawning herring (Clupea harengus L.), with emphasis on larval distribution in the flesh. Veterinary Parasitol. 2010;171:247–253. doi: 10.1016/j.vetpar.2010.03.039. [DOI] [PubMed] [Google Scholar]
  33. Levsen A, Lunestad BT, 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]
  34. Li S-W, Shiao S-H, Weng S-C, Liu T-H, Su K-E, Chen C-C. A case of human infection with Anisakis simplex in Taiwan. Gastrointest Endosc. 2015;82(4):757–758. doi: 10.1016/j.gie.2015.03.1983. [DOI] [PubMed] [Google Scholar]
  35. Mafra C, Mantovani C, Borges JN, Barcelos RM, Santos CP. Morphological and molecular diagnosis of Pseudoterranova decipiens (sensu stricto) (Anisakidae) in imported cod sold. Brazil Revista brasileira de parasitologia veterinaria Braz J Veterinary Parasitol Orgao Oficial do Colegio Brasileiro de Parasitologia Veterinaria. 2015;24:209–215. doi: 10.1590/S1984-29612015037. [DOI] [PubMed] [Google Scholar]
  36. Mattiucci S, Nascetti G. Advances and trends in the molecular systematics of anisakid nematodes, with implications for their evolutionary ecology and host-parasite co-evolutionary processes. Adv Parasitol. 2008;66:47–148. doi: 10.1016/s0065-308x(08)00202-9. [DOI] [PubMed] [Google Scholar]
  37. Mattiucci S, Paoletti M, Webb SC. Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: morphological description, genetic relationships between congeners and ecological data. Syst Parasitol. 2009;74:199–217. doi: 10.1007/s11230-009-9212-8. [DOI] [PubMed] [Google Scholar]
  38. Mattiucci S, et al. 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]
  39. McGladdery SE. Anisakis simplex (Nematoda: Anisakidae) infection of the musculature and body cavity of atlantic herring (Clupea harengus harengus) Can J Fish Aquat Sci. 2011;34:1312–1317. doi: 10.1139/f86-164. [DOI] [Google Scholar]
  40. Mladineo I, Šimat V, Miletić J, Beck R, Poljak V. Molecular identification and population dynamic of Anisakis pegreffii (Nematoda: Anisakidae Dujardin, 1845) isolated from the European anchovy (Engraulis encrasicolus L.) in the Adriatic Sea. Int J Food Microbiol. 2012;157:224–229. doi: 10.1016/j.ijfoodmicro.2012.05.005. [DOI] [PubMed] [Google Scholar]
  41. Morsy K, Bashtar AR, Mostafa N, El Deeb S, Thabet S. New host records of three juvenile nematodes in Egypt: Anisakis sp. (Type II), Hysterothylacium patagonense (Anisakidae), and Echinocephalus overstreeti (Gnathostomatidae) from the greater lizard fish Saurida undosquamis of the Red Sea. Parasitol Res. 2015;114:1119–1128. doi: 10.1007/s00436-014-4285-y. [DOI] [PubMed] [Google Scholar]
  42. Nada MS, El-Ghany AM. Anisakid nematodes in marine fishes. J Am Sci. 2011;7:1000–1005. [Google Scholar]
  43. Palm HW, et al. Anisakis (Nematoda: Ascaridoidea) from Indonesia. Dis Aquat Organ. 2017;123:141–157. doi: 10.3354/dao03091. [DOI] [PubMed] [Google Scholar]
  44. Pampiglione S, et al. Human Anisakiasis in Italy: a report of 11 new cases. Pathol Res Pract. 2002;198:429–434. doi: 10.1078/0344-0338-00277. [DOI] [PubMed] [Google Scholar]
  45. Rocka A. Nematodes of the Antarctic fishes. Warsaw: Polish Polar Research; 2004. [Google Scholar]
  46. Roongruangchai J, Tamepattanapongsa A, Roongruangchai K. Stereo and scanning electron microscopic studies of the third stage larvae of Anisakis simplex. Southeast Asian J Trop Med Public Health. 2012;43:287–295. [PubMed] [Google Scholar]
  47. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406–425. doi: 10.1093/oxfordjournals.molbev.a040454. [DOI] [PubMed] [Google Scholar]
  48. Samir M, Amin MA, Hassan AO, Merwad AM, Awadallah MAI. Research note. Prevalence, protein analysis and possible preventive measures against zoonotic anisakid larvae isolated from marine Atherina fish. Helminthologia. 2015;52:375–383. doi: 10.1515/helmin-2015-0060. [DOI] [Google Scholar]
  49. Shih HH. Parasitic helminth fauna of the cutlass fish, Trichiurus lepturus L., and the differentiation of four anisakid nematode third-stage larvae by nuclear ribosomal DNA sequences. Parasitol Res. 2004;93:188–195. doi: 10.1007/s00436-004-1095-7. [DOI] [PubMed] [Google Scholar]
  50. Sohn WM, Kang JM, Na BK. Molecular analysis of Anisakis type I larvae in marine fish from three different sea areas in Korea. Korean J Parasitol. 2014;52:383–389. doi: 10.3347/kjp.2014.52.4.383. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Song H, Jung BK, Cho J, Chang T, Huh S, Chai JY. Molecular identification of Anisakis larvae extracted by gastrointestinal endoscopy from health check-up patients in Korea. Korean J Parasitol. 2019;57:207–211. doi: 10.3347/kjp.2019.57.2.207. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Suzuki J, Murata R, Hosaka M, Araki J. Risk factors for human Anisakis infection and association between the geographic origins of Scomber japonicus and anisakid nematodes. Int J Food Microbiol. 2010;137:88–93. doi: 10.1016/j.ijfoodmicro.2009.10.001. [DOI] [PubMed] [Google Scholar]
  53. Tantanasi J, Diakou A, Tamvakis A, Batjakas IE. Anisakis spp. burden in Trachurus trachurus. Helminthologia. 2012;49:16–20. doi: 10.2478/s11687-012-0003-4. [DOI] [Google Scholar]
  54. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 1994;22:4673–4680. doi: 10.1093/nar/22.22.4673. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. WHO . Soil-transmitted Helminths. Geneva: World Health Organization; 2012. [Google Scholar]
  56. Zardoya R, et al. Differential population structuring of two closely related fish species, the mackerel (Scomber scombrus) and the chub mackerel (Scomber japonicus), in the Mediterranean Sea. Mol Ecol. 2004;13:1785–1798. doi: 10.1111/j.1365-294X.2004.02198.x. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

12639_2020_1222_MOESM1_ESM.tif (461.6KB, tif)

Fig S1. Fish hosts examined in the present study. a. Atlantic Herring (Clupea harengus). b. Mediterranean horse mackerel (Trachurus mediterraneus). c. Atlantic mackerel (Scomber scombrus) (TIF 461 kb)

12639_2020_1222_MOESM2_ESM.tif (659.1KB, tif)

Fig S2. a. Mediterranean horse mackerel heavily infected with Anisakis simplex (s.s) L3 found encysted in back muscle (arrow). b. Anisakis simplex (s.s) found localized in the musculature of Atlantic herring fish (arrow) (TIF 659 kb)

Supplementary file3 (DOCX 21 kb) (21.4KB, docx)
Supplementary file4 (DOCX 17 kb) (17.9KB, docx)
Supplementary file5 (XLSX 10 kb) (10.7KB, xlsx)

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