Introduction
Species of Falcaustra Lane, 1915 (= Spironoura Leidy, 1856) (Cosmocercoidea, Kathlaniidae) have been reported in the digestive tract of fish, amphibians and reptiles (Bursey et al., Reference Bursey, Sung and Goldberg2012). Generally, findings on the life cycle of different species of Falcaustra are not adequate. Specifically, Bartlett & Anderson (Reference Bartlett and Anderson1985) described third-stage larvae of Falcaustra in tissues of a freshwater snail, whereas Moravec et al. (Reference Moravec, Huffman and Swim1995) recovered larvae of Falcaustra spp. from freshwater fish collected in Texas. Anderson (Reference Anderson2000) also suggested that, in general, kathlaniids develop to the third stage outside the host and then invade various invertebrates, which serve as paratenic hosts.
To date, there is little information about the phylogenetic position of the genus and only one gene sequence has been published (Hasegawa et al., Reference Hasegawa, Sato, Kai and Uchida2013). There are two reports of Falcaustra araxiana in the large intestine of the European pond turtle, Emys orbicularis, from Armenia and Iran, albeit without molecular analysis (Yamaguti, Reference Yamaguti1962; Bursey & Rivera, Reference Bursey and Rivera2009; Shayegh et al., Reference Shayegh, Rajabloo, Gholamhosseini, Mootabi Alavi, Salarian and Zolfaghari2016). In our previous study, F. araxiana was found in the large intestine of the European pond turtle (Shayegh et al., Reference Shayegh, Rajabloo, Gholamhosseini, Mootabi Alavi, Salarian and Zolfaghari2016). However, other helminths similar to F. araxiana in appearance, but different in size, were also discovered in gastric nodules. In the present study, helminths from the nodules and the large intestine of the European pond turtle have been described, as well as the phylogenetic position of F. araxiana using molecular analysis of the 18S rDNA gene, together with histopathological changes to host tissue.
Materials and methods
Sample preparation
Worms were recovered from the large intestine and gastric nodules of European pond turtles (E. orbicularis) as described previously by Shayegh et al. (Reference Shayegh, Rajabloo, Gholamhosseini, Mootabi Alavi, Salarian and Zolfaghari2016). The worms were then examined for morphological features according to the previous reports (Yamaguti, Reference Yamaguti1962). In order to determine probable similarities between helminths originating from gastric nodules and those from the large intestine, we examined primarily the dimensional differences in the recovered worms. In the next step, 18S rDNA gene analysis was conducted. For molecular examination two males and two females of F. araxiana recovered from the large intestine and two of each from the nodules were selected randomly. Formalin-fixed nodules were also used for histopathological studies.
Molecular examination
Genomic DNA from all collected samples was extracted and purified individually using the DNeasy Tissue Kit (Qiagen, Valencia, California, USA), according to the manufacturer's recommended protocol, and used as template DNA for the polymerase chain reaction (PCR). Briefly, to achieve the desired results, we used a 2–3 h incubation time for samples at 56°C with 180 μl tissue lysis buffer (ATL, Qiagen) and 20 μl (50 μg/ml) of proteinase K, vortexing every 30 min. After adding 200 μl of lysis buffer (AL, Qiagen) containing guanidine hydrochloride and 200 μl ethanol, the mixture was vortexed for 15 s and then added to a DNA-binding column and spun down for 1 min. The column was then washed several times using 500 μl of AW1 and AW2 buffers (Qiagen). The genomic DNA extract was diluted to a working concentration of 20 ng/μl, and 4 μl of it was used as a template in PCR. All extracted DNA was stored at −20°C until further processing.
Primers and PCR amplification
The 18S rDNA sequences (~1800 bp) were amplified by universal primers NC18SF1 (5′-AAAGATTAAGCCATGCA-3′) and NC5BR (5′- GCAGGTT CACCTACAGAT-3′) (Brianti et al., Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012). The following PCR conditions were applied to each assay: 50 mm KC1, 10 mm Tris–HCl (pH = 9.0), 1.5 mm MgCl2, 200 μm deoxynucleoside triphosphates (dNTPs), 20 pmol of each primer and 2 U Taq DNA polymerase (Fermentas, Burlington, California, USA) per 50-μl reaction, using 4 μl of the DNA extracted as the template. For amplification, the samples were cycled in a Bio-Rad thermocycler (Bio-Rad Laboratories, Hercules, California, USA). Cycling conditions included an initial DNA denaturation at 94°C for 5 min, followed by 40 cycles of denaturation at 94°C for 45 s, annealing at 50°C for 1 min and an extension at 72°C for 1 min. Sterile water was used as a negative control. The presence of amplicons and their size was assessed by electrophoresis of 5 μl of each reaction product in 1.5% (w/v) Tris–acetate/EDTA agarose gel and visualized by staining with ethidium bromide (final concentration of 0.5 μg/ml) under UV light. Images were captured on a computer and printed.
Amplified products were purified with a PCR product purification kit (BioNeer, Korea) and sequenced directly using a capillary DNA analyser (ABI 3730; Applied Biosystems, Foster City, California, USA) after sequencing reactions with a BigDye Terminator v3.1 cycle sequencing kit (Applied Biosystems). Forward and reverse nucleic acid sequence data were used to construct a continuous sequence of each inserted DNA. The 16S rDNA sequence obtained was compared to GenBank entries using the BLAST tool provided by the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/blast/Blast.cgi).
To determine the phylogenic position of F. araxiana, the sequence obtained for this parasite was compared with homologous sequences previously reported for other nematodes in Ascaridida. Creating multiple-sequence alignment was established using the Clustal W program in the MEGA 4.0 software for each queried DNA sequence (Tamura et al., Reference Tamura, Dudley, Nei and Kumar2007). Data sequences were also used for construction of the phylogenetic trees using maximum parsimony and neighbour-joining methods.
Histopathology
Tissues removed from the liver, stomach, intestines and spleen of the pond turtle were fixed in 10% neutral buffered formalin, embedded in paraffin, sectioned at 5 μm and stained with haematoxylin and eosin (H&E) for light microscopic examination.
Results
Morphological study
Different stages of nematodes recovered from the gastric nodules (fig. 1A) were previously identified as F. araxiana using morphological characteristics (Shayegh et al., Reference Shayegh, Rajabloo, Gholamhosseini, Mootabi Alavi, Salarian and Zolfaghari2016). Two stages of nematodes were identified, including larvae or pre-adults and adult worms with genitalia, some of which were in the nodules but the majority of adults occurred in the large intestine. All worms were cylindrical with truncated anterior and tapered posterior ends. The mouth was surrounded by three large vesiculated cephalic lips, each with a medial V-shaped indentation, giving a hexagonal appearance to the oral opening when viewed apically. The oesophagus possesses a posterior spherical bulb and anteriorly positioned isthmus. Males possess two elongate spicules with four pseudosuckers and a gubernaculum (fig. 1B). Pairs of caudal papillae showed a 6–6–10 + 1 pattern. There were four pseudosuckers in the posterior end of the body.
Fig. 1. (A) Nodules (arrowed) on the gastric surface of the European pond turtle (Emys orbicularis) caused by Falcaustra araxiana, scale bar: 500 μm. (B) Posterior end of male F. araxiana with protruding spicules (arrowed), gubernaculum (g) and four pseudosuckers (S), scale bar: 500 μm.
Molecular analysis
Using universal amplification as described, all parasites produced a fragment of approximately 1800 bp, which corresponds to the 18S rDNA gene.
Interestingly, sequence analysis showed that all parasite samples originating from different tissue sources were completely identical to each other based on the 18S rDNA sequence. GenBank accession numbers KM200715 and KM200716 were provided for worms separated from the large intestine and stomach, respectively. Comparative sequence analysis of Iranian F. araxiana sequences with other related taxa existing in the GenBank database demonstrated the highest homology (>99%) with Falcaustra catesbeianae and Paraquimperia africana. Unexpectedly, phylogenetic analysis of concatenated data showed that both species of F. araxiana and F. catesbianae (as the members of Cosmocercoidea) were located within the Ascaridoidea clade, whereas all other Cosmocercoidea members, i.e. Cosmocercoides sp., Raillietnema sp., Dacnitoides sp., Cruzia americana and Nemhelix bakeri, form a separated major clade (fig. 2).
Fig. 2. Phylogenetic analysis of the partial 18S rDNA gene sequences of Iranian F. araxiana (KM200715 and KM200716) and related genera and species, with the tree constructed using neighbour-joining methods and bootstrap percentage values given at the nodes.
According to the phylogenetic tree, the Iranian F. araxiana together with P. africana and F. catesbianae form a sister group for members of Seuratoidea (Truttaedacnitis truttae, Cucullanus baylisi and Cucullanus robustus). In addition, in accordance with the morphological classification, T. truttae, C. robustus and C. baylisi all belong to the superfamily Seuratoidea and are placed in a separate clade based on 18S rDNA analysis.
Histopathology
Small foci of haemorrhage were observed in the thickened mucosal surface of the small intestine, with free worms in the lumen. Also, firm nodules were visible on the gastric serosal and mucosal surfaces, with most of the nodules containing worms.
Histopathological examinations of the tissue sections of gastric nodules revealed granulomatous reactions on larvae, which were surrounded by necrotic debris, eosinophils, mononuclear cells and a few giant cells within a connective tissue capsule. No lesions were observed on the liver or spleen.
Discussion
Up to now, 95 species of Falcaustra have been identified from different hosts. Reptiles and amphibians are final hosts for these species (Anderson, Reference Anderson2000; Bursey & Rivera, Reference Bursey and Rivera2009); among them, freshwater turtles and lizards are common hosts for these parasites (Anderson, Reference Anderson2000). Despite the identification of many species of this parasite, the life cycle of the family Kathalaniidae (the Oxyascaridinae, with Oxyascaris and Pteroxyascaris) have not been completely identified (Anderson, Reference Anderson2000). Moravec et al. (Reference Moravec, Huffman and Swim1995) reported that freshwater fishes can act as paratenic hosts for Falcaustra spp. Also, third-stage larvae of Falcaustra wardi have been found in the tissue of the freshwater snail Lymnea stagnalis (Bartlett & Anderson, Reference Bartlett and Anderson1985).
In this study we found that both larvae and adults of F. araxiana were located in the gastric nodules, and phylogenetic analysis using 18S rDNA gene sequencing confirmed that the two types were similar, suggesting that nodule formation is part of the life cycle or possibly a survival strategy. In addition, larval stages of F. araxiana develop in the gastric mucosa before migrating to the large intestine.
Some nematodes, such as Oesophagostomum, have nodule formation in their life cycles (Soulsby, Reference Soulsby1982; Zajac & Conboy, Reference Zajac and Conboy2012). Nodular formation caused by hypobiotic or arrested larvae is seen in strongylid and ascarid nematodes (Zajac & Conboy, Reference Zajac and Conboy2012), such as Toxocara, Amplicaecum and Ophidascaris (Wharton, Reference Wharton1986; Anderson, Reference Anderson2000). Consistently, our study concludes that the nodule formation is a part of the life cycle of F. araxiana. Thus, we can argue that nodule formation is part of a strategy for survival of the parasite, which can be caused by arrested or normal larvae, but further studies are needed to confirm this issue.
Previous studies have focused on the morphology of the genus Falcaustra but few studies exist on gene sequencing and phylogeny (Hasegawa et al., Reference Hasegawa, Sato, Kai and Uchida2013). In the present study, BLAST searches and phylogenetic analysis showed a close relationship between F. araxiana and P. africana, a member of the Ascaridoidea (fig. 2). Because of the difficulties in studying turtle nematodes, associated with their morphological and biological peculiarities, most species of these parasites are poorly characterized (Anderson, Reference Anderson2000). It seems that their present classification system does not perfectly reflect phylogenetic relationships and a taxonomic revision of this nematode group, based on detailed morphology (including scanning electron microscopy and transmission electron microscopy), life cycle and molecular studies of individual species, is necessary.
Acknowledgements
Authors would like to thank Dr Majid Rostami Shahrbabaki, Dr Amin Gholamhosseini and Dr Ali Zolfaghari for their kind help. Special thanks to Professor Charles R. Bursey for his efforts and scientific support during this project.
Financial support
The researchers would like to thank the Research Council of Shiraz University and School of Veterinary Medicine, Shiraz University for financial and technical support of this study (Grant No. 87-GR-VT-47).
Conflict of interest
None.
