https://orcid.org/0000-0002-2483-0100
Figures
Abstract
Bithyniids are freshwater snails that play a crucial role in the transmission of various parasitic trematodes of medical and veterinary importance. In this study, we explored the prevalence of cercarial trematode infections in bithyniid snails from Thailand and examined the species diversity of both the intermediate snail hosts and parasite larvae. A total of 688 bithyniid snails were collected from diverse natural habitats at 24 locations in 16 provinces across 5 regions of Thailand. The presence of larval trematode infections was examined using the cercarial shedding method. Both the collected snails and the emerging cercariae were identified at the species level using a combination of morphological and molecular techniques. The mitochondrial COI and 16S rDNA sequences of bithyniid snails, along with the ITS2 sequences of cercariae, were obtained via PCR amplification and sequencing. Three species of bithyniid snails were identified in this study: Bithynia funiculata, Bithynia siamensis siamensis, and Hydrobioides nassa. Among these species, B. s. siamensis exhibited the highest population density, followed by B. funiculata and H. nassa. The overall rate of cercarial infection in the bithyniid snails was relatively low, at 1.45%. H. nassa snails had the highest infection prevalence, at 11.11%, while B. s. siamensis had a prevalence of 1.39%. Only the morphological type of the xiphidiocercariae was detected. BLASTn searches in GenBank and phylogenetic trees based on xiphidiocercariae were used to classify the samples into four different families spanning two superfamilies of digenean trematodes. The genera Plagiorchis, Prosthogonimus, Paralecithodendrium, and cercaria of Renicolidae are reported for the first time in B. s. siamensis. Plagiorchis and Paralecithodendrium are significant genera of zoonotic trematodes. These findings indicate that B. s. siamensis and H. nassa can act as the first intermediate hosts for various parasitic trematodes in Thailand.
Citation: Dumidae A, Ardpairin J, Pansri S, Homkaew C, Nichitcharoen M, Thanwisai A, et al. (2025) Bithyniid snails (Gastropoda: Bithyniidae) infected with Xiphidiocercariae in Thailand include a new record of Bithynia siamensis siamensis as the intermediate host of Plagiorchis and Paralecithodendrium. PLoS ONE 20(2): e0317052. https://doi.org/10.1371/journal.pone.0317052
Editor: Hudson Alves Pinto, Universidade Federal de Minas Gerais, BRAZIL
Received: July 8, 2024; Accepted: December 19, 2024; Published: February 4, 2025
Copyright: © 2025 Dumidae et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the manuscript.
Funding: This study was financially supported by Naresuan University (NU), and National Science, Research and Innovation Fund (NSRF), Thailand (Grant No. R2566B046) and partially supported by Global and Frontier Research University Fund, Naresuan University (Grant number R2567C003). All funds were received by Dr. Apichat Vitta. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Digenean trematodes, also known as flukes, are a group of parasitic flatworms that are significant parasites of humans and domestic animals, leading to numerous public health and livestock issues [1,2]. Many trematodes, particularly food-borne trematodes, have been documented to cause diseases in humans across numerous countries worldwide [3]. Foodborne trematodiases are a major group of neglected tropical diseases, with more than 40 million people infected and more than 750 million at risk (>10% of the world’s population) [4,5]. Furthermore, more than 100 species of foodborne trematodes are known to infect humans [3]. In Southeast Asia, humans are exposed to at least 70 species of foodborne and waterborne trematodes, which include lung flukes, liver flukes, intestinal flukes, and blood flukes [6]. These infections typically occur in focal outbreaks and remain endemic in various regions worldwide, notably in Southeast Asia, including Thailand [3,7].
Freshwater snails play a crucial role as intermediate hosts for numerous trematode species [8]. In general, the life cycle of trematodes typically involves freshwater and marine snails as the intermediate host [9]. After egg hatching, the miracidium stage is released, initiating larval development within a suitable snail host. The larvae undergo asexual multiplication, producing many cercariae [10]. The cercariae larvae emerge from the first intermediate host snail and penetrate the second intermediate host. Within the second intermediate host, the cercariae encyst and develop into the infective metacercariae stage. Humans or animals ingest metacercariae as definitive hosts, allowing them to develop into adults and complete their life cycle [10]. Most trematodes typically utilize specific species of freshwater snails as their first intermediate hosts [11]. Therefore, the circulation of trematodes relies on the presence of intermediate hosts [12]. Various snail species, particularly those belonging to the family Bithyniidae, are known to be the first intermediate hosts for medically significant parasitic trematodes endemic to Thailand [13].
Freshwater snails of the family Bithyniidae were first documented in Europe and Asia in the early 1870s [14,15]. In Thailand, ten species and subspecies have been identified and are distributed across four genera: Bithynia, Hydrobioides, Wattebledia, and Gabbia [16]. Previous research has shown that bithyniid snails, particularly those in the genera Bithynia and Hydrobioides, act as the first intermediate hosts for several medically and veterinary important trematodes. These include plagiorchiids, heterophyids, lecithodendriids, echinostomes, gastrothylacids, schistosomes, and the carcinogenic liver fluke Opisthorchis viverrini, which pose serious threats to public health [1,13,17]. Currently, in Thailand, the genus Bithynia is classified into two species: Bithynia siamensis and B. funiculata. Bithynia siamensis is further categorized into two subspecies, B. s. siamensis and B. s. goniomphalos. However, the morphology of B. s. siamensis and B. s. goniomphalos resembles that of Hydrobioides nassa, leading to confusion when traditional morphological characters are used for identification and classification [18]. Therefore, molecular methods have been used for species identification in previous studies. Specifically, mitochondrial genes such as cytochrome c oxidase subunit I (COI) and 16S rDNA have been utilized to distinguish between Bithynia and Hydrobioides snails, as demonstrated by Kulsantiwong et al. (2013) [19] and Bunchom et al. (2021) [18,20].
However, despite the crucial role of bithyniid snails in the life cycle of trematodes, there have been relatively few reports about trematode infections in these snails. For over a decade, there have been no recent reports on the infection rate and classification of trematode cercariae from bithyniid snails across various regions of Thailand. The most recent research dates from 2008–2009, when Kulsantiwong et al. (2015) [21] conducted a survey on trematode infections in freshwater snails of the family Bithyniidae in Thailand. The reported infection rate was 3.15%, and they identified six morphological types of cercariae. Additionally, a study by Kiatsopit et al. (2016) [22], in which snails were collected from 2009–2014, revealed that 20 morphologically distinct types of cercariae were identified in B. s. goniomphalos from the northeast regions. Most recently, in 2019, Tapdara et al. (2022) [17] collected H. nassa from three regions (north, west, and central) of Thailand and reported a prevalence of cercarial infection of 5.57%, with five different morphological types detected. However, the infection status of cercariae trematodes in bithyniid snails has not yet been assessed in many provinces in Thailand.
The conventional method used to identify trematode cercariae relies on their morphological characteristics, which can be quite challenging due to morphological similarities. This method typically allows identification only at the superfamily or family level [8]. However, identifying cercariae at the genus and species levels is often difficult or even impossible [23]. Correctly identifying cercariae at the genus and species levels is essential. Molecular genetic techniques offer higher resolution for identifying trematodes in their larval stage. Sequences from the nuclear ribosomal internal transcribed spacer 2 (ITS2) region have been utilized for identifying various stages, including the cercariae, metacercariae, and adult stages, in both intermediate and definitive hosts. They are also used in phylogenetic relationship analyses [24,25].
Trematode infectious diseases are predominantly endemic in areas where intermediate snail hosts are distributed [26]. However, the world is currently facing issues with rapidly growing snail populations, particularly near lakes and dams, leading to the widespread transmission of parasitic diseases [27,28]. As a result, current studies on trematode larval stages in snail hosts have garnered increasing attention because data on cercarial infection in snails are essential for developing effective strategies for the prevention and control of trematode diseases [29]. We hypothesize that bithyniid snails may be infected with diverse groups of zoonotic trematode parasites. Therefore, this study aimed to assess the prevalence of cercarial trematode infections in bithyniid snails from Thailand and to explore the species diversity of both intermediate snail hosts and parasite larvae.
Materials and methods
Snail collection and identification
Bithyniid snails were randomly collected from 16 provinces across 5 geographical regions (central, eastern, northern, southern, and western) of Thailand (Fig 1 and Table 1). The samples were collected from their natural habitats, including paddy fields, canals, and ponds, using handpicking and scooping methods. The collected snails were placed in porous plastic bags with water and transported to the Department of Microbiology and Parasitology, Faculty of Medical Science, Naresuan University, Phitsanulok, Thailand. All snails were cleaned by washing with dechlorinated water and identified according to the standard morphological criteria of Brandt (1974) [14], Chitramvong (1992) [16], Bunchom et al. (2021) [18], and Kulsantiwong et al. (2022) [30]. Briefly, B. s. siamensis has a slender body shape, a narrow umbilicus, and a weak carina. The shell of B. funiculata is slightly larger, subovate-conic in shape, and varies in color from dull green to reddish-brown or olive-brown. It features a funnel-shaped umbilicus and a strong carina. The shell of H. nassa is small with transverse growth lines and spiral markings on its surface. The umbilicus is completely closed, while the outer part of the last whorl extends as a sinuous flange. The shell aperture has a slight incision at the middle of the basal lip. Additionally, molecular confirmation was conducted using mitochondrial DNA sequencing. The experiments involving invertebrate animals (snails) were approved by the Center for Animal Research at Naresuan University (Project Ethics No: NU-AQ640803).
Geographic locations (A) and environments of the bithyniid snail collection sites examined in this study, including paddy fields (B-D), natural canals (E), lotus ponds (F), and irrigation canals (G). The map was created using MapChart software’s free version (https://www.mapchart.net/terms.html#licensing-maps), under a CC BY license, with permission from Minas Giannekas (owner and creator of the map-making website mapchart.net).
Examination of cercaria infection
Trematode infection in bithyniid snails was assessed using the cercarial shedding technique [31]. Each snail was placed in a small plastic container (4 cm in diameter and 6.5 cm in height) filled with 20 ml of dechlorinated water. The container was covered with a small, perforated plastic lid to provide air ventilation and prevent the snails from escaping. The snails were exposed to natural light for 3–5 h during the day and kept at room temperature for 24 h [32]. Afterward, each container was daily examined for the presence of cercariae under a stereomicroscope for 7–10 days. The numbers of examined and infected snails were recorded to calculate the prevalence of infection. The living cercariae were primarily classified into types based on their morphological characteristics using light microscopy, according to standard identification methods [1,33]. Cercariae were photographed using an Olympus DP72 digital camera fitted to an Olympus BX53 microscope (Olympus Corporation, Japan). The cercariae were collected in a 1.5 ml sterile microcentrifuge tube and preserved at -20°C for DNA extraction. Afterward, the body of each snail that tested positive for cercariae was removed from its shell, sectioned into small pieces of body tissue (approximately 25 mg), and stored at -20°C for subsequent DNA analysis.
DNA extraction
Genomic DNA from cercariae and snail samples was extracted using a NucleoSpin® Tissue Kit (Macherey-Nagel, Duren, Germany) following the manufacturer’s protocol. The DNA was eluted with 30 μl of elution buffer for cercariae and 70 μl for snail samples. The DNA concentration was assessed using 0.8% agarose gel electrophoresis in Tris-Boric-EDTA (TBE) buffer at 100 V, and the DNA was stained with ethidium bromide. The genomic DNA was then stored at -20°C until use for PCR analysis.
Polymerase chain reaction (PCR) amplification and DNA sequencing
Cercariae DNA was amplified for the ITS2 region [34], while bithyniid DNA was amplified for COI [35] and 16S rDNA [36]. PCR amplification was performed in a total volume of 30 μl, consisting of 15 μl of OnePCR Ultra (Biohelix, New Taipei, Taiwan), 1.5 μl of each primer at 5 μM (0.25 μM), 9 μl of distilled water, and 3 μl of the extracted DNA template (20–200 ng). The amplified products were separated by gel electrophoresis at 100 V for 35 min using a 1.2% agarose gel stained with ethidium bromide. Subsequently, the PCR products were purified using the NucleoSpin® Gel and PCR Clean-Up Kit (Macherey-Nagel, Germany) according to the manufacturer’s instructions. The purified PCR products were then analyzed using a 1.2% agarose gel. Nucleotide sequencing was conducted by Macrogen, Korea. The details of the primers and PCR conditions used for the cercariae and snails used in this study are provided in Table 2.
Molecular identification and phylogenetic analysis
The forward and reverse DNA sequence chromatograms were manually edited and assembled using SeqMan II (DNASTAR, Madison, WI, USA). The nucleotide sequences were subjected to BLASTn searches to identify similarities with sequences from other taxa in the GenBank database (National Center for Biotechnology Information: NCBI). The partial sequences obtained in the study, along with similar sequences selected from GenBank, were aligned using CLUSTAL W and trimmed with MEGA version 7.0 [37].
The sequence data from both the trematode ITS2 sequences and the snail COI and 16S rDNA mtDNA sequences generated in this study, as well as sequences from related reference species retrieved from the GenBank database, were used to construct phylogenetic trees. Phylogenetic relationships were analyzed using both maximum likelihood (ML) and neighbor-joining (NJ) methods. ML phylogenetic trees were constructed using the Tamura 3-parameter model [38] for 16S rDNA and ITS2 sequences and the Tamura-Nei model [39] for COI sequences. Moreover, NJ trees were generated utilizing the Kimura 2-parameter model [40] with bootstrap support of 1,000 replications using the MEGA version 7.0 program. Though two methods were used to establish the phylogeny, only the ML tree was selected for presentation in this study because both methods revealed congruent topologies. Additionally, genetic distances of the species were estimated using MEGA version 7.0 software to assess and compare genetic variation.
Results
Cercarial infection in snails
In this study, a total of 688 bithyniid snails were collected from 24 locations in 16 provinces across 5 regions of Thailand. Based on morphological identification, the snail samples belonged to three species: Bithynia funiculata, Bithynia siamensis siamensis, and Hydrobioides nassa. B. s. siamensis was the most abundant species, accounting for 93.89% (646/688) of the samples, followed by B. funiculata at 4.79% (33/688) and H. nassa at 1.31% (9/688). B. s. siamensis was found across all five regions (central, eastern, northern, southern, and western). In contrast, B. funiculata was limited to the northern and western regions, and H. nassa was found only in the central and western regions.
Out of the 688 bithyniid snails, ten were found to be infected with trematode cercariae and were distributed across 4 locations in 4 provinces of Thailand, resulting in an overall prevalence of infection of 1.45%. H. nassa exhibited the highest prevalence of infection, at 11.11% (1/9), while B. s. siamensis displayed a prevalence of 1.39% (9/646). The greatest percentage of cercarial infection was detected in the snails collected from Songkhla Province (17.66%), followed by those collected from Nakhon Nayok (15.15%), Tak (14.29%), and Ayutthaya (0.48%) Provinces (Fig 1 and Table 1).
Morphology of cercariae
Based on morphological characteristics, the cercariae found in this study were identified as xiphidiocercariae. They have a small, oval-elongated body measuring 170–175 μm in length and 80–90 μm in width. The oral sucker, round and located at the anterior end, features a distinctive stylet at its center, measuring 12–15 μm in length, a defining characteristic of this type. The ventral sucker, situated in the middle of the body, is globular and smaller than the oral sucker. The tail is tapered and shorter than the body, measuring 75–96 μm in length (Fig 2).
Bootstrap values (≥50%) for ML (left) and NJ (right) analyses are indicated at branch nodes. Bold letters denote sequences obtained in this study. The black silhouettes represent the hosts of the trematode according to molecular data from GenBank. Aspidogaster ijimai was used as the outgroup.
Molecular analysis of cercariae
ITS2 was successfully amplified and sequenced from 24 xiphidiocercariae samples from 8 out of the 10 infected bithyniid snails. However, sequence amplification failed for cercariae from two B. s. siamensis strains from the provinces of Songkhla and Nakhon Nayok due to poor DNA quality. The ITS2 sequences (309–339 bp) of these xiphidiocercariae (GenBank accession nos. PP094634-PP094657) were identified as belonging to Xiphidiocercariae sp., Prosthogonimus sp., Paralecithodendrium sp., Plagiorchis sp., and renicolid sequences. The detailed results from the BLASTn analysis of the cercariae samples are shown in Table 3.
The ML phylogenetic tree constructed from the ITS2 xiphidiocercariae sequences, comprising 24 sequences from this study and 26 sequences downloaded from GenBank (Table 4), revealed the presence of four different families within two superfamilies (Plagiorchioidea and Microphalloidea) of digenean trematodes. Two sequences (BD1B291TAK1 and BD5B291TAK1) of xiphidiocercariae released from H. nassa in Tak Province were closely related to Prosthogonimus cuneatus, a parasite within the family Prosthogonimidae known to infect Indian peafowl (Pavo cristatus), wild ducks (Anas platyrhynchos), and blackbirds (Turdus merula), exhibiting a genetic divergence of 0.32%. Moreover, five sequences (BI1B641NYK1-BI4B641NYK1 and BK9B642NYK1) of Xiphidiocercariae released from B. s. siamensis in Nakhon Nayok Province belonged to a clade of trematodes within the family Plagiorchiidae. These sequences closely clustered with those of Plagiorchis sp. found in the caddisfly (Lepidostoma sp.), snail (Galba pervia), and mayfly (Ecdyonurus sp.), indicating a genetic divergence of 0.29%. Furthermore, two sequences (BF2B638NYK and BF5B638NYK1) and three sequences (BG1B639NYK1, BG3B639NYK1, and BG5B639NYK1) from the same province were grouped with Prosthogonimus sp., which infects the bird Falco peregrinus, and Paralecithodendrium sp. in bats from the families Prosthogonimidae and Lecithodendriidae, respectively, showing genetic divergences of 0.32% and 10.09%, respectively. Four sequences (BE1B408AYA1-BE3B408AYA1 and BE5B408AYA1) from Ayutthaya Province closely matched the renicolid trematodes in the family Renicolidae, infecting the snail Filopaludina sumatrensis, with no genetic divergence (0.0%). However, eight samples (BB1B288SKA2-BB5B288SKA2, BC2B290SKA2, BC4B290SKA2, and BC5B290SKA2) from Songkhla Province clearly clustered as Xiphidiocercariae sp., which were isolated from snails Indoplanorbis exustus and B. s. siamensis. These samples were closely clustered between the families Plagiorchiidae and Prosthogonimidae, with a genetic divergence of 0.36% between these two identical species (Fig 2).
Molecular identification of snails
The COI and 16S rDNA sequences of bithyniid snails infected with cercariae were analyzed via PCR amplification and sequencing. Eight out of the ten bithyniid snails infected with cercariae were successfully amplified and sequenced. Sequence amplification failed for two snails from Ayutthaya and Nakhon Nayok provinces due to poor DNA quality. The molecular identification of bithyniid snails based on the COI and 16S rDNA genes was consistent with the morphological identification. Seven sequences (GenBank accession nos. PP094598-PP094604) of 578 bp of the COI gene were obtained from B. s. siamensis. BLASTn results showed 100% similarity to B. s. siamensis from Thailand (GenBank accession nos. KY118649, KY118672, MW832442, and KY118653), while one sequence (GenBank accession no. PP094614) from H. nassa displayed 100% similarity with H. nassa from Thailand (GenBank accession no. MK640188). Additionally, the 16S rDNA sequences (367–369 bp) from 7 samples (GenBank accession nos. PP094615-PP094621) of B. s. siamensis in the present study showed 98.62–100% similarity to those of B. s. siamensis from Thailand (GenBank accession nos. MW305394 and MW305395). Moreover, one sample (GenBank accession no. PP094628) of H. nassa exhibited 100% homology to H. nassa from Thailand (GenBank accession no. MK629223).
Phylogenetic analysis of snails
Phylogenetic analyses of bithyniid snails based on COI and 16S rDNA revealed congruent clustering results. The ML phylogenetic tree of the COI (578 bp) and 16S rDNA (367–369 bp) sequences showed that our B. s. siamensis obtained from Songkhla and Nakhon Nayok provinces clustered with B. s. siamensis from Thailand, while H. nassa sequences from Tak Province grouped together with H. nassa sequences from Thailand (Fig 3).
Maximum likelihood phylogenetic tree based on COI (A) and 16S rDNA (B) sequences of bithyniid snails in Thailand, along with other sequences obtained from GenBank. Bootstrap values (≥50%) for ML (left) and NJ (right) analyses are indicated at branch nodes. Bold letters denote sequences obtained in this study. Filopaludina martensi, F. sumatrensis, and F. bengalensis were used as the outgroup.
Discussion
Bithyniid snails are common freshwater gastropods in Thailand and are found in various types of water bodies [20]. Overall, our study revealed that bithyniid snails were prevalent in most paddy fields. Morphological identification revealed three species, B. funiculata, B. s. siamensis, and H. nassa, with B. s. siamensis being the most abundant, followed by B. funiculata and H. nassa. These findings are consistent with previous research [1,19,20,56], which reported the wide distribution of B. s. siamensis across five regions (central, northern, northeastern, western, and southern) of Thailand. However, our study revealed a new occurrence of B. s. siamensis in the eastern region, which differs from previous reports.
In this study, we performed molecular analysis of mitochondrial COI and 16S rDNA sequences to identify the species of bithyniid snails harboring trematode cercariae in Thailand. In our study, B. s. siamensis was confirmed to have 100% identity for the COI sequence and 98.62–100% identity for the 16S rDNA sequence after BLASTn searching. Similarly, H. nassa showed 100% identity with H. nassa sequences from the GenBank database. Consistent with the phylogenetic analysis, the ML phylogenetic tree of the COI and 16S rDNA sequences obtained in our study showed that B. s. siamensis and H. nassa clustered with B. s. siamensis and H. nassa from Thailand, respectively. Our findings align with those of Bunchom et al. (2021) [18,20], who described the molecular identification of these two snail species in Thailand through the analysis of different mitochondrial COI and 16S rDNA markers. Furthermore, we observed that the morphologies of B. s. siamensis and H. nassa are very similar, potentially causing taxonomic confusion when traditional morphological characters are used for identification and classification [18]. Therefore, our results indicate that COI and 16S rDNA markers are effective for confirming and differentiating between B. s. siamensis and H. nassa [18–20].
In our study, we found that the overall rate of cercarial infection in bithyniid snails was 1.45%. This rate was lower than the 3.15% reported by Kulsantiwong et al. (2015) [21] for bithyniid snails in Thailand. Variations in cercarial infection rates in snails can be influenced by numerous factors, such as differences in collection times, seasons, locality, rainfall, temperature, water quality, availability of infected definitive hosts, type and number of parasites and intermediate hosts, and snail population density [57–60]. The low infection rate in our study may be due to reduced parasite presence, which limits contact between snails and miracidia [27,61]. Additionally, the limited diversity of secondary intermediate hosts and definitive hosts at the sampling sites could also contribute to the reduced infection rate [27,62]. Moreover, decreased human activities such as open-field defecation, farming, and livestock grazing have been associated with lower trematode infection rates in snails [27,63].
In the present study, only xiphidiocercariae were detected in H. nassa and B. s. siamensis. The morphology of the xiphidiocercariae observed was consistent with the characteristics reported for xiphidiocercariae collected from B. s. siamensis in Bangkok Province [33]. Previous research also reported xiphidiocercariae infection in H. nassa in Chiang Rai Province, Thailand [17]. Our results indicate new areas of xiphidiocercariae distribution in H. nassa in Tak Province and in B. s. siamensis in Ayutthaya and Nakhon Nayok Provinces. Xiphidiocercariae were the predominant type of cercaria found in H. nassa and B. s. siamensis in this investigation, consistent with previous studies indicating that xiphidiocercariae are the most frequently observed type in other bithyniid snails [21,22,24]. This cercaria type typically requires birds, reptiles, or mammals as definitive hosts to mature into adult intestinal flukes [41,42,47,64]. The high prevalence of xiphidiocercariae observed in this study may be attributed to the coexistence of definitive and intermediate hosts within the same ecosystem, which facilitates the completion of the parasite’s life cycle [27]. Another aspect to consider is that H. nassa and B. s. siamensis may be susceptible to xiphidiocercariae infection. This phenomenon was previously reported in the snail B. s. siamensis [30], indicating that B. s. siamensis is more susceptible to O. viverrini infection than is B. funiculata. Schmid-Hempel and Stauffer (1998) [65] highlighted that the susceptibility of hosts to parasites can increase when genetic variation within the host population decreases. Lee et al. (1995) [66] investigated trematode (Fasciola hepatica) infection in Orientogalba viridis and reported that this snail is highly susceptible to infection at the miracidia stage across various growth stages. These findings indicate a well-established relationship between the parasite and its snail host, suggesting that H. nassa and B. s. siamensis are susceptible to xiphidiocercariae infection. However, there is limited research investigating the dynamics of the relationship between bithyniid snails and xiphidiocercariae. Thus, the susceptibility of H. nassa and B. s. siamensis to xiphidiocercariae infection remains uncertain. Further studies are necessary to reach definitive conclusions on this matter.
The molecular identification of the cercariae was performed using ITS2 sequences, which were analyzed to construct a phylogenetic tree. The ML phylogenetic tree based on the xiphidiocercariae sequences classified them into four different families: Plagiorchiidae (including Plagiorchis sp.), Prosthogonimidae (Prosthogonimus sp. and Prosthogonimus cuneatus), Lecithodendriidae (Paralecithodendrium sp.), and Renicolidae (Renicolid). This cercarial morphotype has been shown to produce a wide variety of trematode species, a finding consistent with previous studies [33]. In this study, we report for the first time the identification of Plagiorchis sp., Prosthogonimus sp., Paralecithodendrium sp., and renicolid in B. s. siamensis, as well as Prosthogonimus cuneatus in H. nassa from Thailand. Among these species, Plagiorchis and Paralecithodendrium are known to cause diseases in humans [67,68], while the other species have veterinary significance [54,64].
Many species of Plagiorchis species are known to parasitize the intestines of vertebrates [69]. They have also been identified as potential pathogens in humans, causing plagiorchiasis, with 12 recorded cases mainly in Asia [67,69–71]. The source of Plagiorchis infection in humans is presumed to be freshwater fish, freshwater snails, and aquatic arthropods [72]. In Thailand, adult Plagiorchis sp. were first identified in four opisthorchiasis patients following praziquantel treatment between 1980 and 1985 in the northeast region [68]. Moreover, Paralecithodendrium species are parasitic trematodes found in fish, amphibians, birds, and mammals, predominantly bats [73]. Specifically, in Thailand, cercariae of this parasite have been reported in B. s. goniomphalos and Filopaludina martensi martensi snails [24,74]. Additionally, the infective metacercariae stage has been detected in insects of the order Odonata in the northeastern region of Thailand [75]. Notably, Paralecithodendrium molenkampi is a well-known species infecting human in Thailand [67]. In contrast, Prosthogonimus species are significant pathogenic parasites in wild birds and poultry, leading to mortality and defective egg formation [64]. These parasites utilize dragonfly nymphs or dragonflies as their second intermediate hosts [47]. Previous research has reported that B. s. goniomphalos is the first intermediate host of Prosthogonimus in Thailand [24]. Despite the absence of reported infections in wild birds or poultry in Thailand, the presence of Prosthogonimus poses a potential threat to these populations. Renicolid trematodes are commonly known as a group of parasites inhabiting the kidneys and ureters of aquatic birds and exerting a strong pathogenic effect on their hosts. The metacercariae of these parasites develop in second intermediate host freshwater snails and fishes [54]. However, data on their distribution and infectivity in Thailand are scarce, with reports of metacercariae found in F. martensi, F. sumatrensis, and Idiopoma umbilicata snails in Bangkok, Thailand [25].
Conclusion
In conclusion, our study documents the presence and variety of trematode larvae in bithyniid snails across Thailand. Our findings revealed that three species/subspecies of bithyniid snails, including B. funiculata, B. s. siamensis, and H. nassa, were distributed variably across five regions. Two species/subspecies of snails within this family act as intermediate hosts for significant medical and veterinary trematodes. This study also provides new information on the intermediate hosts of two trematode species affecting human public health and three species of veterinary significance. Given the public health risk posed by trematode parasites in bithyniid snails, particularly the presence of the zoonotic trematode species Plagiorchis sp. and Paralecithodendrium sp., these findings underscore the important role of these snails as intermediate hosts for trematode cercariae in Thailand.
Acknowledgments
We gratefully acknowledge the Department of Microbiology and Parasitology, Faculty of Medical Science, Naresuan University, for their invaluable support and access to facilities.
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