Nematodes from the family Anisakidae are parasites that have a worldwide distribution (Anderson, 2000). The most widespread genera from this family are Anisakis and Contracaecum, which have similar life cycles. The third larval stage (L3) of Contracaecum is usually found in the body cavity, mesenteries and branchial chambers of a wide range of fish species (Norris & Overstreet, 1976). The adult stage is found in the intestine of piscivorous birds and mammals associated with fresh, brackish and seawater (Whitfield & Heeg 1977; Anderson, 2000). Anisakid larvae are the causative agent of anisakidosis in humans (Audicana et al., 2002; Audicana & Kennedy, 2008), particularly the species belonging to Anisakis, Contracaecum and Pseudoterranova (Oshima, 1987; Yagi et al., 1996). This disease has been reported worldwide, and it is endemic in Southeast Asia (Audicana & Kennedy, 2008; Mattiucci & Nascetti, 2008). The knowledge about the life cycle of these species and their geographical distribution is very poor (Shamsi, 2019). The identification of Contracaecum larvae from different fish hosts has attracted the interest of scientists in many countries in studying the ultrastructure and genetic characterisation of these larvae using different genetic markers, such as 28S rDNA, 18S rDNA, ITS-1, ITS-2, and mtDNA cox-2 (Garbin et al. 2013; Mattiucci et al. 2015; Younis et al., 2017; Zuo et al. 2018; Malviya et al., 2018). There are a few publications on the specific identification of the larval stages of Contracaecum in fishes from the world (Szostakowska & Fagerholm, 2007; Shamsi & Aghazadeh-Meshgi, 2011; Shamsi et al., 2017; Molnár et al., 2019; Pekmezci & Yardimci, 2019).
In Iraq, Contracaecum larvae were known principally from early work by Herzog (1969), Shamsuddin et al. (1971), Mhaisen (1986), Ali et al. (1987) and Khalifa et al. (1987). Recently there are 42 fish host species known for Contracaecum larvae in Iraq (Mhaisen, 2019). However, research to date has not yet investigated the specific identification of Contracaecum larva in fishes based on molecular approach in Iraqi waters, and this requiring further investigation. Previous studies have shown that specific identification of Contracaecum larvae is not possible based solely on morphological description (Shamsi et al., 2017). The present study is an investigation toward the molecular genetic characterization of Contracaecum nematodes in Iraq by using a combined molecular and ultra-morphological approach based on sequence data of well-identified adults and Contracaecum larval types in GenBank. This approach is useful for the reliable identification of Contracaecum larvae at the species level (Shamsi et al., 2011). The purpose of this study is to know the genetic characterisation and determine the Contracaecum larvae among freshwater fish in Iraq.
Sulaimani Province is located in the northeast of Iraq; it is situated between latitudes 35° 05’ and 36° 30’ and between longitudes ‘44° 25’ and 46° 20’. It is located close to the Iraqi-Iranian border. There are many water bodies in this province in addition to the two large rivers (the Lesser Zab River and the Sirwan River) that pass through this province (Fig. 1).
Fig. 1
Map of north Iraq showing the study area.
During the period from January to the end of December 2018, 1134 fish were collected and examined for infection with anisakid larvae (Contracaecum). The fishes comprised 13 species of Cyprinidae as follows: Acanthobrama marmid (n=20), Alburnus caeruleus (n=7), A. mossulensis (n=62), Arabibarbus grypus (n=123), Barbus lacerta (n=7), Capoeta trutta (n=222), C. umbla (n=161), Carasobarbus kosswigi (n=5), C. luteus (n=89), Carassius auratus (n=54), Chondrostoma regium (n=52), Cyprinion kais (n=10) and C. macrostomum (n=322). They were caught by a pulsed DC electro-shock device (SAMUS 1000). The fish were identified based on their morphometric and meristic characteristics (Coad, 2010) and the scientific names for fishes were identified according to Froese and Pauly (2019). The fish were immediately transported to the laboratory for parasitological examinations, where they were dissected from the ventral side. The body cavity, stomach, spleen, liver, kidneys, heart, muscles, swim bladder and gonads were all examined for anisakid cysts. The gastrointestinal tract was removed, from the rectum to the esophagus, opened longitudinally, and carefully examined under stereoscope (Amlacher, 1970). The cysts were collected and washed with physiological solution (saline solution 0.9 %) in a glass Petri dish. The cysts were opened under a stereoscope with the aid of a fine needle to release the Contracaecum larva, and then washed with saline solution and preserved. The prevalence and intensity of infection were calculated for each fish species based on Margolis et al. (1982). The infection level was calculated as prevalence (percentage of investigated fish infected) and mean intensity (mean number of Contracaecum larva per infected fish).
After opening the cysts, the Contracaecum larvae were washed with saline solution (0.9 %), fixed in hot 4 % formaldehyde solution (60°C) to relax the body, and preserved in 70 % ethanol. A small piece of the mid-body of each nematode was excised for molecular study (Shamsi & Aghazadeh-Meshgi, 2011), and the rest of the nematode was cleared with increasing concentrations of glycerine (5 %, 10 % and 50 %, each for 1 hour), followed by 100 % glycerine for 1 – 2 hours; they were then mounted in glycerine jelly (Moravec et al., 2009; Moravec & Yooyen, 2011). All measurements of the parasites were made with an Olympus ocular micrometre eyepiece and are given in millimetres. Photos were taken with a Sony Optical Steady Shot digital camera (model DSC-W570, 16.1 mega pixels). The morphology of detected third larval stage of the Anisakid were identified according to the key features and descriptions of Bykhovskaya-Pavlovskaya et al. (1962), Hoffman (1998), Anderson (2000) and Shamsi et al. (2011).
The larvae were removed from the cysts for scanning electron microscopy (SEM) study. The specimens were fixed in 4 % (v/v) hot formaldehyde solution (60°C), preserved in 70 % (v/v) ethanol, and post-fixed in 1 % osmium tetroxide (in phosphate buffer). The samples were then dehydrated by incubating in a graded series of acetone–ethanol concentrations ((1:1), (1.5 – 0.5) and abso
lute acetone, 15 min each) (Moravec et al., 2012). A critical-point method was used for drying by shaking the samples for 24 hr. in centrifuge to remove the acetone; they were then embedded on the targets and sputter-coated with gold (Moravec et al., 2009; Moravec & Yooyen, 2011). The specimens were examined using a FEI Quanta 400 scanning electron microscope at an accelerating voltage of 25 kV.
Genomic DNA was isolated from mid piece of individual larvae after being preserved directly in absolute ethanol (99 %). The genomic DNA was extracted by using a QIAamp® DNA Mini Kit with slight modifications. In brief, the mid piece of individual larval parasite digested for 1 – 3 h at 56 °C with proteinase K in ATL buffer and eluted in 50 μl of AE buffer (QIAamp® DNA Mini Kit).
The PCR was used to amplify the ITS-1, ITS-2, and COX-2 regions. The specific primer sets SS1F/NC13R, SS2F/NC2R (Shamsi et al., 2008) and 210F/211R (Nadler & Hudspeth, 2000) were used to amplify the two nuclear ribosomal markers (ITS-1 and ITS-2) and cytochrome oxidase-II (COX-2), respectively. The PCR reaction (in a volume of 30 μl) was performed in 25 mM Tris-HCl, pH 9.0 at 25°C and contained 50 mM KCl, 2 mM MgCl2, 0.1 mg/ml gelatin, 200 μM dATP, dGTP and dTTP, 100 μM [α32-P] dCTP (0.05 μCi/nmol), 12.5 μg of activated salmon sperm DNA, 10 pmol of each primer and 1.5 U Taq polymerase (Canvax Bio-tech, S.L.). The PCR reactions were carried out in a thermocycler (Applied Biosystems 2720, USA) using the following cycling instructions: 94°C for 5 min (initial denaturation), 35 cycles of 94°C, 30 sec (denaturation), 55°C, 30 sec (annealing), 72°C, 30 sec (extension) and a final extension of 72°C for 7 min, followed by holding at 4°C. Two microliters of genomic DNA (20 – 40 ng) in nuclease-free deionised distilled water were added to each PCR reaction. Samples with fish genomic DNA (extracted from muscle) were included in the PCR as negative controls; no amplicons were produced from these samples. Five microliters of each PCR product was examined on a 1.5 % w/v agarose gel, stained with DNA stain (Good ViewTM SBS Genetech Beijing, China) and photographed using a gel documentation system. A 1000 bp DNA ladder (Vivantis, Malaysia) was used. The amplicon were then purified using the EasyPure® Quick Gel Extraction Kit (TransGen Biotech), according to the manufacturer’s protocols. The resulting products were sent to Macrogen in South Korea for nucleotide sequence analysis by a dideoxy termination method using a Genetic Analyzer 3500 DNA sequencer (Applied Biosystems, USA) in both directions (forward and reverse) using the same PCR primers.
The resulted ITS-1, ITS-2 and COX-2 sequences (forwards) were compared with their complements (reverses) and then adjusted using online software tool (bioinformatics.org\sms\rev_comp.html) to obtain reverse complement. Then the resulted sequences were aligned to each other using multiple sequence alignment program by using the online software tool CLUSTALW (genome.jp/tools-bin/ clustalw) to get the most homologous sequences (one sequence). Subsequently, the obtained sequences were compared with previously published sequences data for identification by using the Basic Local Alignment Search Tool (BLAST) from GenBank database (http://www.ncbi.nlm.nih.gov/) (Altschul et al., 1990). Sequences of ITS-1 and ITS-2 from individual Contracaecum larvae in the different fish host species in the present study were aligned by using the online computer program CLASTALAW (https://www.genome.jp/tools-bin/clustalw) and then adjusted manually for searching nucleotide variations among Contracaecum larvae in different fish host species.
For the phylogenetic study, the sequence data of ITS-1, ITS-2 and COX-2 fragments obtained from Contracaecum larvae collected from all different fish host species were installed into the MEGA X version 10.7.1 software program (Kumar et al., 2018). To unify the length of the sequences, the common 447, 268 and 475 bp length of ITS-1, ITS-2 and COX-2 segments respectively were selected and used for phylogenetic analysis to determine the most appropriate sequence evolution model for the given data, treating gaps and missing data with the partial deletion option. The sequences were aligned using CLUSTALW alignment for constructing the trees of evolutionary development. The trees of all isolated species were constructed based on the Maximum Likelihood (ML) method and Tamura-Nei model (Tamura & Nei, 1993).
The care of experimental animals was consistent with Rebuplic of Iraq animal welfare laws, guidelines and policies approved by University of Sulaimani Local Ethics Committee (Permit reference number 122/2020). All fishes were collected from both Lesser Zab and Sirwan River drainages, with required permissions of the Directorate Police of Forest and Regional Sulaimani Province (reference number 1060/2018).
A total of 1134 fish were examined for parasitic larval Contracaecum nematodes. Only 17 fishes (belonged to five species) were infected with this parasite. The parasitic larvae were found in the intestinal wall, liver, ovaries, swim bladder, gallbladder and mesenteries of 1.49 % (17\1134) of fishes. The prevalence of Contracaecum larvae and the mean of intensity per infected fish varied among the different fish species (Table 1).
Prevalence of Contracaecum larva and mean of intensity among infected fish species.
| Host | Fish | Prevalence | Mean intensity | |
|---|---|---|---|---|
| Examined | Infected | % | ||
| Acanthobrama marmid | 20 | 7 | 35.00 | 5.57 |
| Arabibarbus grypus | 123 | 1 | 0.81 | 7.00 |
| Capoeta trutta | 222 | 2 | 0.90 | 3.00 |
| Carasobarbus luteus | 89 | 4 | 4.49 | 4.25 |
| Chondrostoma regium | 52 | 3 | 5.76 | 9.00 |
A total of 7 fish (prevalence 35 %, mean intensity 5.57) out of the 20 Acanthobrama marmid specimens examined were infected with Contracaecum larvae, which represented the highest prevalence among the collected fishes. While, only one fish (prevalence 0.81 %, mean intensity 7) out of 123 Arabibarbus grypus specimens examined was infected with this larva, which represented the smallest prevalence among the collected fishes in the present study. These results agree with Abdullah and Mhasein (2011), who recorded Contracaecum larvae in Acanthobrama marmid and Chondrostoma regium with prevalence of 16.6 % and 36.4 %, respectively, among ten fish species in the Lesser Zab River. This variation in the prevalence may be related to the temperature, water level, intensity of both the intermediate host and migratory bird (final host), and the types of food and feeding habits of the fish (Younis et al., 2017).
The present investigation shows that prevalence and intensity of Contracaecum larvae in the examined fishes are relatively low. This may be because only visual examination was used to isolate Contracaecum larvae in this study. The prevalence and number of larvae may have been higher than Contracaecum larvae found in the collected fishes in the present study if an incubation method had been used (Shamsi & Suthar 2016; Buchmann, 2007). Third larval stage of the anisakid nematode Contracaecum rudolphii Hartwich, 1964, are commonly infect a range of fish species and mainly cyprinids particularly in area where the final host (piscivorous birds such as cormorants) are found (Moravec, 1994).
Morphological examination and measurements were conducted by optical microscopy and identified that the Anisakid larvae in the present study were Contracaecum larvae (L3), as described by Moravec (2009). Additionally, there were no significant morphological differences among the larvae that were recorded in different fishes. The Contracaecum larvae were light brownish-yellow in colour. They had an elongated cylindrical body and short tail with conical and pointed tip. The bodies had fine, dense transverse striation of the cuticle. The larvae were encapsulated within a slender body, they have distinct boring tooth. Excretory pore situated anteriorly, cuticular striations were observed through the whole length of the body. The esophagus consisted of a long muscular part and a short glandular ventriculus. The esophageal caecum was extended posteriorly and the intestinal caecum was extended anteriorly. The intestine was filled with numerous small brownish granules, and the gonads and other parts of the reproductive system were not developed (Fig. 2).
Fig. 2
Photo micrograph of Contracaecum larva in Carasobarbus luteus. A - Anterior region of the larva (I00×); B - Posterior region of the larva (I00×); C - Mouth region of the larva (400×); D - Tail region of the larva (400×) an = anus; bt = boring tooth; er = cuticle ridges; eso = esophagus; esoc = esophageal caecum; int = intestine; intc = intestinal caecum; mo = mouth opening; p = papillae; rec = rectum; ta = tail.
The total length of the larvae was 3.5 – 11 mm, with a width of 0.10 – 0.25 mm. The esophagus length was 0.7 – 1.3 mm, the intestinal caecum length was 0.20 – 0.35 mm and the esophageal caecum length was 0.25 – 0.40 mm (Table 2). Since this parasite was in the larval stage and lacked the reproduction system, it was difficult to morphologically determine the exact classification at the species level. The specimens showed a close resemblance in both measurements and characteristics to those recorded by Moravec (2009) in Cyprinus carpio from the Czech Republic. In the microscopy studies, there were no significant morphological differences among the Contracaecum larvae in the five different fish species; the photomicrograph of the third larval stage of Contracaecum in Carasobarbus luteus is only an example (Fig. 2).
Comparison of measurements in Contracaecum larvae in different infected fish species in the present study (in millimeter).
| Host | Total length | Maximum width | Tail length | Rectum length | Boring teeth length | Esophagus length | Esophageal caeca length | Intestinal caeca length |
|---|---|---|---|---|---|---|---|---|
| A. marmid | 4.10 | 0.20 | 0.07 | 0.07 | 0.005 | 0.480 | 0.50 | 0.38 |
| A. grypus | 5.07 | 0.25 | 0.07 | 0.07 | 0.010 | 0.650 | 0.30 | 0.20 |
| C. trutta | 4.20 | 0.23 | 0.08 | 0.07 | 0.005 | 0.450 | 0.48 | 0.33 |
| C. luteus | 4.62 | 0.25 | 0.08 | 0.07 | 0.005 | 0.600 | 0.55 | 0.40 |
| C. regium | 7.75 | 0.35 | 0.08 | 0.09 | 0.010 | 0.600 | 0.60 | 0.45 |
The SEM study revealed a regular striation pattern in the cuticular striations of larvae from all fishes. The striations were narrow in the anterior of the larvae and in the pre-tail region, and they became gradually wider when extended posteriorly. There were no significant different ornamentations among the larvae. There were transverse striations between the striated cuticular rings (Fig. 3). The mouth was transverse in shape the lips are not developed and provided by four papillae, a well-defined boring tooth present. Excretory pore is anteriorly located just below the mouth. Anal opening is located near the posterior end. Tail is short. The body was free of any projections, such as a spine or papillae. The cloacal regions had no papillae because they were in the larval stage and not yet mature.
Fig. 3
Scanning electron micrograph of Contracaecum larva in Carasobarbus luteus. A - Anterior region of the larva; B - Head region of the larva; C - Tail region of the larva; D - Cuticle of the larva an = anus; bt = boring tooth; cu = cuticle; exp = excretory pore; mo = mouth opening; p = papillae; ta = tail.
The ultrastructural characteristics of the present specimens showed great similarity with the specimens (Contracaecum larva) of Younis et al. (2017), who collected them from different freshwater fishes in Lake Nasser, Egypt. Furthermore, the ultrastructural features of the present Contracaecum larvae were similar to those of Rahemo and Nawab Al-Din (2009), who used SEM to study larvae from Acanthobrama marmid collected in the Tigris River. The SEM study revealed no significant ultra-morphological differences among the Contracaecum larvae recorded in the present investigation in these five different fish species. The scanning electron photomicrograph of the third larval stage of Contracaecum in Carasobarbus luteus is used as an example (Fig. 3).
Agarose gels analyses demonstrated the same size for each ITS-1, ITS-2 and COX-2 region. The amplicons were ~530 bp, ~430 bp and ~630 bp for ITS-1, ITS-2, and COX-2 respectively, confirming that all sequences were of the same genus.
Alignment of the resulting sequences revealed that there was no significant variation in the ITS-1, ITS-2 and COX-2 regions, which indicated the presence of only one type of larva. Based on percentage identities of nucleotides from GenBank, the on-line BLAST tool showed the ITS-1 sequences obtained from larvae-infected A. marmid, A. grypus, C. trutta, C. luteus and C. regium matched 99.78 %, 100 %, 99.76 %, 100 % and 100 %, respectively to the previously reported reference gene sequences for the ITS-1 in Contracaecum rudolphii B (Zhang et al., 2009) from the stomachs of great cormorant Phalacrocorax carbo sinensis from the Guangzhou Zoo in Guangdong in China, which was examined previously and deposed in GenBank (FJ467618) (Zhang et al., 2009). The ITS-2 sequences obtained from larvae-infected A. marmid, A. grypus, C. trutta, C. luteus and C. regium matched 100 % to the previously reported reference gene sequences for the ITS-2 in Contracaecum rudolphii B (Li et al., 2005) from the stomachs of great cormorant Phalacrocorax carbo sinensis from the Venice Lagoon in northeastern Italy and from Monaci Lake in central Italy, which was examined previously and deposed in GenBank (AJ634786) (Li et al., 2005), and also matched 100 % to the previously reported reference gene sequences for the ITS-2 in Contracaecum rudolphii B from the same host from from the Guangzhou Zoo in Guangdong in China, which was examined previously and deposed in GenBank (FJ467618) (Zhang et al., 2009). The COX-2 sequences obtained from larvae-infected A. marmid matched 98.52 % to the previously reported reference gene sequences for the COX-2 in Contracaecum rudolphii B (Mattiucci et al., 2008) from the great cormorant Phalacrocorax carbo sinensis from Italy, which was examined previously and deposed in Gen-Bank (EF122203) (Mattiucci et al., 2008). The COX-2 sequences obtained from larvae-infected A. grypus, C. trutta and C. regium matched 100 %, 99.19 % and 100 % respectively to the Contracaecum rudolphii B from Phalacrocorax carbo sinensis which was examined previously and deposed in GenBank (EF513509). While, the COX-2 sequences obtained from larvae-infected C. luteus matched 99.79 % to the Contracaecum rudolphii B from the same host which was examined previously and deposed in Gen-Bank (EF558894) (Mattiucci et al., 2008).
The genetic characterisation of the parasite in the present study is available in the GenBank database; the ITS-1, ITS-2 and COX-2 sequences obtained were deposited in GenBank, and their accession numbers are stated in Table 3. The ITS-1, ITS-2 and COX-2 sequence analysis confirmed that third larval stage of Contracaecum (L3) parasitizing the Cyprinid fish (A. marmid, A. grypus, C. trutta, C. luteus and C. regium) from the present study belong to species C. rudolphii type-B, a parasite at the adult stage of the great cormorant Phalacrocorax carbo sinensis mainly from Italy water (Li et al., 2005; Mattiucci et al., 2008). ITS-1, ITS-2 and COX-2 markers may provide reliable evidence for specific species identification of Contracaecum larvae occurring in fish (Mattiucci et al., 2010). Therefore, the occurrence of C. rudolphii B larvae from Iraqi waters was also proved by molecular evidence inferred from the ITS-1, ITS-2 and COX-2 markers used in the present study. Furthermore, the sequences of ITS-1 and ITS-2 obtained from the collected larvae in different fish species were aligned with each other (the same gene). Pairwise comparisons of all nucleotides sequence among the Contracaecum larvae collected in the five different fish hosts revealed that only one nucleotide variation (0.2 %) in alignment position 161 for ITS-1 (Fig. 4) and there was no nucleotide variations for ITS-2 (Fig. 5).
Accession numbers provided by NCBI for the collected Contracaecum larvae in different fish hosts in the present study.
| Host of Contracaecum rudolphii | Accession numbers for ITS-1 sequences | Accession numbers for ITS-2 sequences | Accession numbers for COX-2 |
|---|---|---|---|
| Acanthobrama marmid | MN557376 | MN526259 | MN589997 |
| Arabibarbus grypus | MN557377 | MN563727 | MN589998 |
| Capoeta trutta | MN557378 | MN563728 | MN589999 |
| Carasobarbus luteus | MN557379 | MN563729 | MN590000 |
| Chondrostoma regium | MN557380 | MN563730 | MN590001 |
Fig. 4
Multiple sequence alignment for ITS-1 in C. rudolphii B in the five different fish species.
Fig. 5
Multiple sequence alignment for ITS-2 in C. rudolphii B in the five different fish species.
Fig. 6
Alignment of the ITS-1 and ITS-2 sequences representing genotype 1 (C. rudolphii B) from the present study and genotype 2 (C. rudolphii A) sequences have been deposited in GenBank under the accession numbers AJ634 782 and AJ634 785. Nucleotide differences between the aligned equences are indecated by having no asterisks.
Contracaecum rudolphii Hartwich, 1964 is a species complex it consist of several sibling species. C. rudolphii sensu lato (s.l.) named C. rudolphii A and C. rudolphii B (D’Amelio et al. 1990), they could also be differentiated from each other based on the ITS-1 and ITS-2 sequence data (Li et al. 2005). D’Amelio et al. (2007) indicated the existence of a third cryptic species of C. rudolphii complex (C. rudolphii C) in double-crested cormorants from west-central Florida based on PCR-RFLP and sequencing of the rrnS mitochondrial gene and nuclear ribosomal spacers. Shamsi et al. (2009) described two new sibling species of the C. rudolphii complex, C. rudolphii D from Phalacrocorax carbo and Contracaecum rudolphii E from Phalacrocorax varius in Australia, based on the ITS-1 and ITS-2 sequence data. Recently, D’Amelio et al. (2012) recorded new isolate of C. rudolphi complex (C. rudolphii F) from brown pelican Pelecanus occidentalis in the northern Gulf of Mexico.
Various studies demonstrated that internal transcribed spacers (ITS-1 and ITS-2) of the nuclear ribosomal DNA (rDNA) provide genetic markers for the accurate identification of a range of species of Ascaridoids. In addition, more studies indicated that sibling species can be differentiated based on the ITS sequences (Jacobs et al., 1997; Zhu et al., 2000; 2001; 2002). The ITS-1 sequences of the obtained larvae (C. rudolphii B) in the present study show 11 (2.46 %) nucleotide differences with the previously reported reference gene sequence for the ITS-1 in C. rudolphii A which was examined and deposed in GenBank (Accession number: AJ634782). While, ITS-2 shows 14 (5.22 %) nucleotide differences with the C. rudolphii A which previously reported reference gene sequence for the ITS-2 in C. rudolphii A which was examined and deposed in GenBank (Accession number: AJ634785) (Li et al., 2005) (Fig. 6). This clear genetic differentiation support previous sequence analyses (Li et al., 2005) that there are sequence differences (1.8 %) in the ITS-1 and (5.1 %) in ITS-2 between the sibling species of C. rudolphii A and B (Li et al., 2005). Extending these studies, we investigated that there is no significant sequence variation in the ITS-1 and ITS-2 within and among the larvae collected from different fish host in the present investigation. The molecular finding of the present investigation support that the present finding larvae belong to C. rudolphii type-B.
In the phylogenetic analysis, the sequence data aligned with the data sequences of ITS-1, ITS-2 and COX-2 form other different Contracaecum species (different genotypes) and Ascaris suum used as out group detected in GenBank (Table 4, 5). Phylogenetic analyses were conducted in MEGA X (Kumar et al., 2018). The evolutionary histories were inferred by using the Maximum Likelihood method and Tamura-Nei model (Tamura & Nei, 1993). The tree with the highest log likelihood (-6713.31, -3848.19 and -385797 for ITS-1, ITS-2 and COX-2 respectively) are shown (Fig. 7, 8, 9). Initial trees for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the Maximum Composite Likelihood (MCL) approach, and then selecting the topology with superior log likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site (next to the branches). The Contracaecum larvae from all different fish hosts were clustered in the same clade of Contracaecum rudolphii B. Moreover, the concatenated phylogenetic tree of the ITS-1, ITS-2 and COX-2 sequences using ML analyses indicated that Contracaecum larvae clades were distinct species by high bootstrap values (Fig. 7, 8, 9).
Fig. 7
Phylogenetic relationships between Contracaecum larvae from the present study and other Contracaecum species as inferred by maximum likelihood obtained from ITSl. Ascaris suum was used as outgroup.
Fig. 8
Phylogenetic relationships between Contracaecum larvae from the present study and other Contracaecum species as inferred by maximum likelihood obtained from ITS2. Ascaris suum was used as outgroup.
Fig. 9
Phylogenetic relationships between Contracaecum larvae from the present study and other Contracaecum species as inferred by maximum likelihood obtained from COX2. Ascaris suum was used as outgroup.
Contracaecum nematodes and specimens/accession number (ITS) of taxa used to build phylogenetic trees and the nucleotide (bp) differences with the Contracaecum larvae collected from the present study.
| Parasite | GeneBank accession no. (ITS) | No. of nucleotide differences (bp) | Host | Source |
|---|---|---|---|---|
| C. rudolphii A | ITS1 (AJ634782) | 9 | Phalacrocorax carbo sinensis | Li et al. (2005) |
| C. rudolphii B | ITS1 (AJ634783) | 1 | Phalacrocorax carbo sinensis | Zhang et al., 2009 |
| C. rudolphii D | ITS1 (FM210251) | 6 | Phalacrocorax varius | Shamsi et al. (2009b) |
| C. rudolphii E | ITS1 (FM210257) | 6 | Phalacrocorax varius | Shamsi et al. (2009b) |
| C. rudolphii F | ITS (JF424597) | 21 | Pelecanus occidentalis | D'Amelio et al. (2012) |
| C. ogmorhini | ITS1 (AJ291468) | 10 | Arctocephalus pusillus doriferus | Zhu et al. (2001) |
| C. eudyptulae | ITS1 (AJ007461) | 8 | - | Zhu et al.(unpublished) |
| C. chubutensis | ITS1 (HQ389546) | 13 | Phalacrocorax brasilianus | Garbin et al. (2011) |
| C. variegatum | ITS1 (MK424804) | 22 | Bird | Hbaiel & Mohammad (unpublished) |
| C. microcephalum | ITS2 (FM177537) | 14 | Anhinga melanogaster | Shamsi et al. (2009a) |
| C. septentrionale | ITS1 (AJ634784) | 24 | Phalacrocorax carbo sinensis | Li et al. (2005) |
| C. bioccai | ITS (JF424598) | 30 | Pelecanus occidentalis | D'Amelio et al. (2012) |
| C. radiatum | ITS (AY603529) | 35 | Leptonechotes weddlli | Kijewska et al. (2008) |
| C. osculatum | ITS (AB277825) | 77 | Arabesque greenling | Umehara et al. (2008) |
| C. multipapillatum | ITS1 (AM940056) | 146 | Pelecanus conspicillatus | Shamsi et al. (2008) |
| C. pyripapillatum | ITS1 (AM940062) | 141 | Pelecanus conspicillatus | Shamsi et al. (2008) |
| Contracaecum larva | ITS1 (MN557376) | - | Acanthobrama marmid | Present study |
| Contracaecum larva | ITS1 (MN557377) | - | Arabibarbus grypus | Present study |
| Contracaecum larva | ITS1 (MN557378) | - | Capoeta trutta | Present study |
| Contracaecum larva | ITS1 (MN557379) | - | Carasobarbus luteus | Present study |
| Contracaecum larva | ITS1 (MN557380) | - | Chondrostoma regium | Present study |
| Ascaris suum | ITS1 (AB110023) | - | Pig | Ishiwata et al. (2004) |
Contracaecum nematodes and specimens/accession number (COX-2) of taxa used to build phylogenetic trees and the nucleotide (bp) differences with the Contracaecum larvae collected from the present study.
| Parasite | GeneBank accession no. (COX-2) | No. of nucleotide differences (bp) | Host | Source |
|---|---|---|---|---|
| C. rudolphii A | EF122201 | 34 | Phalacrocorax carbo sinensis | Mattiucci et al. (2008) |
| C. rudolphii B | EF558894 | 1 | Phalacrocorax carbo sinensis | Mattiucci et al. (2008) |
| C. rudolphii C | EF014283 | 272 | Phalacrocorax auritus | D'Amelio et al. (2007) |
| C. rudolphii F | JF727879 | 40 | Pelecanus occidentalis | D'Amelio et al. (2012) |
| C. ogmorhini | MN624184 | 32 | Zalophus californianus | Mladineo et al. (under press) |
| C. chubutensis | HQ328504 | 46 | Phalacrocorax atriceps | Garbin et al. (2011) |
| C. microcephalum | EF122208 | 71 | Phalacrocorax pygmaeus | Mattiucci et al. (2008) |
| C. septentrionale | EF558898 | 60 | Phalacrocorax carbo carbo | Mattiucci et al. (2008) |
| C. bioccai | EF558899 | 50 | Pelecanus occidentalis | Mattiucci et al. (2008) |
| C. osculatum | KC412224 | 58 | Chionodraco hamatus | Santoro et al. (2013) |
| C. multipapillatum | AF179910 | 72 | - | Nadler & Hudspeth (2000) |
| C. micropapillatum | EU852350 | 70 | Pelecanus onocrotalus | Mattiucci et al. (2010) |
| C. austral | GQ847539 | 55 | Phalacrocorax brasilianus | Garbin et al. (2011) |
| C. pelagicum | EF122210 | 60 | Spheniscus magellanicus | Mattiucci et al. (2008) |
| Contracaecum larva | MN589997 | - | Acanthobrama marmid | Present study |
| Contracaecum larva | MN589998 | - | Arabibarbus grypus | Present study |
| Contracaecum larva | MN589999 | - | Capoeta trutta | Present study |
| Contracaecum larva | MN590000 | - | Carasobarbus luteus | Present study |
| Contracaecum larva | MN590001 | - | Chondrostoma regium | Present study |
| Ascaris suum | HQ704901 | - | swine | Liu et al. (2012) |
The first information on Contracaecum larvae from Iraqi freshwater fishes was provided by Herzog (1969) from ten fish species collected from different inland water bodies in Iraq. In the Kurdistan Region, larvae were recorded from Dukan Lake in Sulaimani Province by Abdullah and Rasheed (2004) in Arabibarbus grypus, Carasobarbus luteus, Chondrostoma regium, Cyprinion macrostomum, Cyprinus carpio, Luciobarbus barbulus, L. esocinus, L. kersin, L. subquincunciatus, L. xanthopterus, and Squalius lepidus. Thus far, a total of 21 host fish species are known for Contracaecum larvae in the Kurdistan Region of Iraq (Mhaisen & Abdullah, 2017). In addition, molecular identification of Contracaecum larvae in fish species have also not been studied, and there is still no specific identification of the Contracaecum species in fresh and marine water fish species in Iraq. Till now, morphological identification based on only optical microscopy has been used to identify larval stage of Contracaecum only at genus level in Iraq. Recently, 42 different fish species were known as hosts for Contracaecum sp. larvae in Iraq from north to south including marine water fish (Mhaisen, 2019). Therefore, the authors provide the first molecular evidence of Contracaecum infection in fish from Iraqi waters in the present study. Notably, the Contracaecum larvae can infect many types of fish in Iraq. This nonspecific infection ability towards a variety of different organs and fish species may lead to the infection of a variety of piscivorous birds and mammals in the region. Anisakidosis is a disease caused by the accidental ingestion of larval anisakid nematodes in raw fish. All fishes which infected with Contracaecum larvae in the present study are edible, particularly A. marmid (prevalence 35 %) in Sulaimani Province. This may affect human health in this region, because this fish is used by local people and other consumers as a food source.
It is noteworthy that the adult Contracaecum rudolphii sensu lato was recorded previously for the first time in Iraq in the digestive tract of the great black cormorant Phalacrocorax carbo from Baghdad province in Iraq (Al-Moussawi & Mohammad, 2011). Furthermore, three other species of this genus (Contracaecum) were reported in birds (final host) in Iraq, namely, C. microcephalum, C. multipapillatum and C. ovale (Habish, 1977; Al-Hadithi & Abdullah, 1991).
The Contracaecum larvae in these five fish species in Sulaimani Province were found to be Contracaecum rudolphii B. A study of the complete life cycle of these larvae in the laboratory is necessary to examination the morphology and ultrastructure of the adult stage (male and female).