Introduction
Freshwater snails are relevant for public and veterinary health, as several species act as first and second intermediate hosts for helminths and parasites that cause various food-borne trematode diseases in humans and animals (Chai et al. Reference Chai, Murrell and Lymbery2005, Reference Chai, Shin, Lee and Rim2009; Madsen and Hung Reference Madsen and Hung2014). In Thailand, several freshwater snail species are reported to be infected by larval trematodes, with high prevalence noted in northern and northeastern regions (Burch and Lohachit Reference Burch and Lohachit1983; Sri-Aroon Reference Sri-Aroon2011; Woodruff and Upatham Reference Woodruff and Upatham1993). However, studies specifically addressing the potential for larval trematode infection in northern Thailand, especially in Chiang Rai Province, remain limited. Recent studies have documented larval trematode infections across various snail species in various ecosystems, with particular emphasis on paddy fields in this region (Chantima et al. Reference Chantima, Suk-ueng and Kampan2018a, Reference Chantima, Phromkhong, Sonyod, Chaichana, McCormick, Mungthaisong, Khittasangka, Chomchalow, Walsh and Lisec2018b).
In northern Thailand, rice is generally cultivated once or twice per year. Farmers typically grow rice during the rainy season, with some extending cultivation into the post-rainy season in areas with irrigation access. Following the harvest, the dry season brings unfavourable conditions for freshwater snails in rice paddies. During this period, snails face extreme environmental stresses, particularly drought. To survive, they burrow into the mud and enter a state of aestivation, a physiological process like hibernation. With the arrival of spring rains and monsoon flooding, snails recolonize their habitats and reproduce. Aestivation is a crucial survival strategy for snails facing extreme temperatures and desiccation, particularly in tropical environments. This adaptation is essential not only for their survival, but it also influences the dynamics of trematode transmission (Osborne and Wright Reference Osborne and Wright2018; Rubaba et al. Reference Rubaba, Chimbari and Mukaratirwa2016). The timing of aestivation in relation to infection significantly impacts the development of trematodes within their snail hosts. A study by Badger and Oyerinde (Reference Badger and Oyerinde1996) demonstrated that Biomphalaria pfeifferi snails that aestivate immediately after infection produce cercariae in greater numbers compared to those that delay aestivation. Furthermore, aestivation can prolong cercarial development, which in turn affects the transmission potential of trematodes in rice paddies. Notably, shorter aestivation durations lead to lower mortality rates for the snail intermediate host of Opisthorchis viverrini, Bithynia siamensis goniomphalos (Chaiyasaeng et al. Reference Chaiyasaeng, Pechdee, Sereewong, Suwannatrai, Laha and Tesana2019).
Aestivation serves as a crucial adaptive mechanism for snails; however, it also poses challenges for trematode transmission. Understanding these dynamics is essential for effectively managing snail populations and controlling trematode-related diseases in agricultural settings. Therefore, this study aimed to investigate the diversity and survival of aestivating snails in dry-season rice paddies in Chiang Rai Province, Thailand, with a specific focus on their role as intermediate hosts for trematodes. The findings of this study may enhance our understanding of the ecology of aestivating snails and its implications for public health regarding parasitic infections in agricultural landscapes.
Materials and methods
Study area and sampling localities
The study area in Chiang Rai Province, northern Thailand, within the North Mekong River basin of Thailand, has a tropical monsoon climate with distinctive seasons. The mean annual rainfall of this area varies from 1,200 mm to 2,000 mm (Eastham et al. Reference Eastham, Mpelasoka, Mainuddin, Ticehurst, Dyce, Hodgson, Ali and Kirby2008). Eight sampling sites were established across four districts: Mae Lao, Mueang Chiang Rai, Phan, and Wiang Chai, based on the availability of potential intermediate snail hosts, site accessibility, and the tendency of snails to aestivate after the rice harvest during the dry season (February to May) in endemic areas (Chantima et al. Reference Chantima, Suk-ueng and Kampan2018a, Reference Chantima, Phromkhong, Sonyod, Chaichana, McCormick, Mungthaisong, Khittasangka, Chomchalow, Walsh and Lisec2018b; Chantima and Rika Reference Chantima and Rika2020). Geographic coordinates of sampling sites were recorded using a GPS device or determined as accurately as possible from a map. Sampling sites were then digitally mapped on a dot-by-dot basis on a public domain map (GISTDA 2011; Github 2024). Final maps were compiled using Photoshop CS6 (Adobe Systems Inc., San José, CA, USA). The coordinates of these localities are presented in Table 1 and Figure 1.
Table 1. List of localities where snails were collected
Figure 1. Map of study area, showing the sampling localities and the districts where snails were collected as indicated in Table 1.
Snail and soil sampling
Snail sampling was conducted once a month during three sampling periods from March to May 2020, coinciding with the dry season. The selected sampling sites were located at the dried end of a paddy field. Snails buried in the soil were collected at two depth intervals: from the ground surface to 5 cm and from 5 to 10 cm. A quadrat measuring 0.36 m2 (0.6 m square) was used to define the sampling area. All snails found within the quadrat were brought to the laboratory for further analysis. Species identification was carried out using conchological methods based on the taxonomic keys of Brandt (Reference Brandt1974) and Upatham et al. (Reference Upatham, Sornmani, Kitikoon, Lohachit and Burch1983). The snails were subsequently examined for survival and trematode infections.
Soil samples were collected in the same quadrat as the snail sampling. At each site, three replicate samples were randomly collected from a 0.36 m2 area using a metal shovel. Approximately 200 g of sediment per replicate was gathered from each site. Following collection, the samples were sealed in labelled bags and transported to the laboratory for analysis. The soil properties, focusing on pH, electrical conductivity, soluble salts concentration, and cation or anion concentration in soil extract, were determined at each sampling site using the methods by the U.S. Salinity Laboratory Staff (1954) and Sparks et al. (Reference Sparks, Page, Helmke, Loeppert, Soltanpour, Tabatabai, Johnston and Sumner1996).
Assessment of aestivating snail species abundance and diversity
To evaluate the abundance and diversity of aestivating snail species, several indices were employed. First, the richness index, specifically Margalef’s richness index (D), was utilized and calculated using the following formula:
$$ D=\frac{S-1}{\ln n} $$
where S was the total number of species and n was the total number of individuals (Clarke and Warwick Reference Clarke and Warwick1994; Washington Reference Washington1984).
Second, Shannon’s diversity index (H) was calculated using the following formula:
$$ H=-{\sum}_{i=1}^n{p}_i\ln {p}_i $$
where pi was the proportion of the total count arising from the i-th species. A higher value indicated a greater number of species with similar abundances, whereas a lower value indicated low diversity that was dominated by one or a few species (Clarke and Warwick Reference Clarke and Warwick1994; Hill et al. Reference Hill, Fasham, Tucker, Shewry and Shaw2005).
Third, species evenness (J′), quantifies the distribution of individuals across different species within an ecosystem. This metric is commonly represented by Pielou’s evenness index, which is calculated using the following formula:
$$ J^{\prime }=\frac{H}{\ln S} $$
where H referred to Shannon’s diversity index. It values between 0 and 1; values closer to zero represented uneven populations that were dominated by one species, while values closer to 1 represented even populations that were comprised of several species with similar abundances (Brewer Reference Brewer1994; Clarke and Warwick Reference Clarke and Warwick1994; Hillebrand Reference Hillebrand, Jorgensen and Fath2008).
Examination for survival of aestivating snail
The survival of aestivating snails was evaluated by reactivating them in water for a duration of 24 hours. The shells of snails were categorized into three distinct groups: 1) live snails that resumed activity upon immersion in water, 2) dead snails, which either lacked soft tissues or had an operculum, and 3) old shells, indicative of snails that died during the prior dry season, distinguished by their white coloration or discoloration and erosion (Figure 2). The counts of dead and surviving snails, excluding old shells, were recorded to calculate survival rates.
Figure 2. Photographs of aestivating snail, Filopaludina martensi martensi: (a) live snail; (b) dead snail; (c) old shell with erosion.
Examination for trematode infection
The examination of snail infections was conducted using shedding and crushing methods. For the shedding method, individual snails were placed in 15 ml bottles filled halfway with dechlorinated tap water and left for 24 hours to facilitate the release of cercariae. After this period, the snails were exposed to artificial light for approximately 3 hours to check for the shedding of cercariae. Each snail was individually screened over a period of 48 hours. Prior to the crushing method, the snails were anesthetized by freezing. The shells of the snails were then broken, and the entire body was crushed in a Petri dish. Following this, larval trematode infections were detected by microscopically examining the crushed tissue. The identification of larval trematodes was based on their morphological characteristics, as described by Schell (Reference Schell1962) and Yamaguti (Reference Yamaguti1975), and counts were performed according to the species of snail. Additionally, the prevalence and intensity of infections were assessed, with infection intensity specifically determined by counting only the metacercariae.
Statistical analysis
Statistical analyses were conducted using SPSS v. 29.0.2. Two sets of analyses were performed. The first set employed basic descriptive statistics to summarize soil properties, with values presented as mean±S.D. Descriptive statistics were also used to summarize the prevalence and intensity of trematode infections by snail species. The second set involved correlation tests, where Pearson’s correlation was applied to examine the relationship between soil properties and the survival of aestivating snails. Statistical significance was defined as p < 0.05.
Results
Aestivating snail abundance and diversity
We collected and examined a total of 1,159 aestivating snails from various species belonging to the families Bithyniidae, Viviparidae, Ampullariidae, Thiaridae, Planorbidae, and Lymnaeidae. Based on conchological assessments, nine species were identified (Figure 3): Bithynia funiculata (1.6%), B. siamensis siamensis (72.4%), Filopaludina martensi martensi (8.4%), F. sumatrensis polygramma (6.9%), F. doliaris (7.0%), Pomacea canaliculata (2.8%), Melanoides tuberculata (0.3%), Indoplanorbis sp. (0.6%), and Lymnaea sp. (<0.1%) (Table 2). Most of the aestivating snails collected belonged to the Bithyniidae, with B. s. siamensis being the most abundant and widespread species, present at all sampling sites. Quantification of species richness and diversity revealed low values of 0.932±0.375 (D range: 0.314–1.431) and 0.792±0.572 (H range: 0.072–1.741), respectively. The evenness index was also low (J′ = 0.497±0.306; J′ range: 0.066–0.865). However, some sampling sites exhibited higher values, with J′ approaching 1 (J′ > 0.8).
Figure 3. Shell morphology of aestivating snails collected from rice paddies during the dry season: (a) Bithynia funiculata; (b) B. siamensis siamensis; (c) Filopaludina martensi martensi; (d) F. doliaris; (e) F. sumatrensis polygramma; (f) Pomacea canaliculata; (g) Melanoides tuberculata; (h) Lymnaea sp.; (i) Indoplanorbis sp. Scale bar: 1 cm.
Table 2. Snail fauna, survival, and mortality (percent) of aestivating snails*
* The old shells were excluded from the survival and mortality rate calculations.
Survival and mortality rates of aestivating snails
Aestivating snails were found only within a depth of 0 to 5 cm from the ground surface. None were found 5–10 cm from the ground surface. After reactivating the snails in water, they were categorized into three groups: live snails (674), dead snails (322), and old shells (163). The overall survival and mortality rates, excluding the old shells, were 67.7% and 32.3%, respectively (Table 2). The survival rate of aestivating snail species ranged from 5.9% to 81.5%, with B. s. siamensis exhibiting the highest survival rate. In contrast, the mortality rate for all species, except B. s. siamensis, was greater than 70%. Specifically, M. tuberculata, Indoplanorbis sp., and Lymnaea sp. had a 100% mortality rate, meaning that no individuals of these species survived the aestivation period. This 100% mortality rate reflects the complete loss of individuals within these species, with all individuals perishing during aestivation.
Relationship between soil properties and survival of aestivating snails
There was considerable variation in the recorded soil properties among the sampling sites. Soil pH varied from 4.01 to 6.10 (mean 4.88±0.75), while electrical conductivity varied between 0.0061 and 0.3248 ds/cm (mean 0.1063±0.1087). Soluble salts concentration in soil extracts also fluctuated, with values from 3.904 to 207.872 mg/l (mean 68.048±69.579), Cation or anion concentration in soil extracts ranged from 0.0061 and 0.3248 mg/l (mean 0.1063±0.1087). The analysis of soil properties in relation to the survival of aestivating snails revealed a positive relationship between electrical conductivity, soluble salt concentrations, and cation/anion concentrations in soil extracts with the survival of viviparid snails, including F. m. martensi, F. s. polygramma, and F. doliaris (p < 0.05). However, no significant correlations were observed for other snail species (B. funiculata, B. s. siamensis, and P. canaliculata) (p > 0.1).
Trematode infections in aestivating snails
The investigation of trematode infections in 674 aestivating snails from dry-season rice paddies revealed that none of the snails shed cercariae after being activated in water for 48 hours. Among the total snails examined, 8.46% (57/674) were infected with larval trematodes. Four types of larval trematodes were identified in this study, including three different kinds of metacercariae: echinostome metacercaria, Thapariella anastomusa metacercaria, and an unidentified metacercaria, as well as rediae (Figure 4).
Figure 4. Morphotypes of metacercariae and redia recovered from aestivating snail: (a) An isolated echinostome metacercaria showing head collar with collar spines (arrowhead); (b) An isolated unidentified metacercaria; (c) Redia stained with 0.5% neutral red; (d) Thapariella anastomusa metacerariae were free, not encysted in the tissue of snails (arrowhead); (e) An isolated T. anastomusa metacercaria stained with hematoxylin.
The infection rates and the number of metacercariae recovered from various snail species are presented in Table 3. Echinostome metacercariae, which were predominantly found clumping together in the pericardial sac of the infected snails, were the most commonly identified, with an overall prevalence rate of 6.37% (43/674). Throughout the survey, a total of 224 echinostome metacercariae were recovered from bithyniid and viviparid snails, including B. s. siamensis, F. m. martensi, F. s. polygramma, and F. doliaris. The infection rates for individual species ranged from 5.24% to 100%, and the intensity of infection varied from 3.00 to 28.33 metacercariae per infected snail, with an average intensity of 5.20 metacercariae per infected snail. Metacercariae of T. anastomusa were found co-infecting with echinostome metacercariae, and were recovered from a single snail species, F. doliaris. Out of 674 snails examined, 4 (0.59%) were infected, with a mean intensity of 2.25 metacercariae per infected snail. Additionally, an unidentified metacercaria co-infection with echinostome metacercariae was detected in B. s. siamensis, exhibiting a prevalence of 0.3% (2/674). Rediae were also observed in this snail species, with a prevalence of 2.07% (14/674).
Table 3. Prevalence and intensity of trematode infections in aestivating snails
Discussion
This study focused on investigating the diversity of aestivating snails and their potential role in the transmission of trematodes in dry-season rice paddies in Chiang Rai Province, Thailand. We identified nine aestivating snail species across eight locations with most belonging to the family Bithyniidae. B. s. siamensis was the most frequently encountered species, consistent with the previous findings highlighting its abundance in rice paddies and its broad physiological and ecological tolerance (Pratumchart et al. Reference Pratumchart, Suwannatrai, Sereewong, Thinkhamrop, Chaiyos, Boonmars and Suwannatrai2019; Suwannatrai et al. Reference Suwannatrai, Suwannatrai, Haruay, Piratae, Thammasiri, Khampoosa, Kulsantiwong, Prasopdee, Tarbsripair, Suwanwerakamtorn, Sukchan, Boonmars, Malone, Kearney and Tesana2011). In contrast, species such as M. tuberculata, Indoplanorbis sp., and Lymnaea sp. were less common, likely due to habitat preferences favouring stream environments (Chantima et al. Reference Chantima, Suk-ueng and Kampan2018a, Reference Chantima, Lekpet, Butboonchoo and Wongsawad2020; Krailas et al. Reference Krailas, Namchote, Koonchornboon, Dechruksa and Boonmekam2014).
Variability in snail diversity among sites was linked to the different agricultural practices, particularly chemical applications, which may have molluscicidal effects (Min et al. Reference Min, Wang, Liu, Yuan, Guo, Zhu, Chai, Zhang and Li2020; Qiu et al. Reference Qiu, Li, Zhu, Xia, Xiao, Huang and Wang2020). Seasonal changes in rice paddies, including paddy preparation and harvesting, also influence snail populations. The golden apple snail, P. canaliculata, was notably scarce, contrasting with its widespread abundance in other rice-growing regions of Thailand (Chaichana and Sumpan Reference Chaichana and Sumpan2015; Ng et al. Reference Ng, Annate, Jeratthitikul, Sutcharit, Limpanont and Panha2020). Since its introduction in the 1980s, this species has significantly impacted rice cultivation (Chanyapeth and Achawakhom Reference Chanyapeth and Achawakhom1998), yet local factors may limit its proliferation in our study area.
In our study, four snail species, B. s. siamensis, F. m. martensi, F. s. polygramma, and F. doliaris, were infected with larval trematodes. These species, common in northern Thailand’s agricultural areas, are important vectors of trematode transmission to humans and animals (Chantima et al. Reference Chantima, Chai and Wongsawad2013; Chantima et al. Reference Chantima, Suk-ueng and Kampan2018a; Chantima and Rika Reference Chantima and Rika2020; Phalee et al. Reference Phalee, Phalee and Wongsawad2018). Given their widespread distribution in rice paddies, managing aestivating snails requires an integrated approach that considers both their ecological role and their significance in parasitic transmission.
During the dry season, the snails encounter extreme environmental conditions. They must endure aestivation and burrowing into the mud, which is a physiological process like hibernation. However, aestivation occurs in times of drought and renders respiration difficult. This study found that almost all collected aestivating snails belonged to the family Bithyniidae, with B. s. siamensis as the most common species found at a soil depth of 0–5 cm. Our findings indicate that snails could bury themselves in depths up to 5 cm, in contrast to Brockelman et al. (Reference Brockelman, Upatham, Viyanant, Ardsungnoen and Chantanawat1986) and Chaiyaseang et al. (Reference Chaiyasaeng, Pechdee, Sereewong, Suwannatrai, Laha and Tesana2019), who noted that rapid habitat desiccation prevents snails from preparing for aestivation in deeper soil. The average survival rate of aestivating snails over a three-month period was high (67.7%), whereas Brockelman et al. (Reference Brockelman, Upatham, Viyanant, Ardsungnoen and Chantanawat1986) reported a survival rate of under 10% during a 16-month dry period in a non-irrigated area. Our study, however, focused on a three-month aestivating period, yielding a survival rate exceeding 60%. This finding is consistent with the report by Chaiyaseang et al. (Reference Chaiyasaeng, Pechdee, Sereewong, Suwannatrai, Laha and Tesana2019), which also indicated that bithyniid snails (B. s. goniomphalos) exhibited a higher survival rate during a shorter dry period of four months in irrigated rice plantations. Consequently, rice cultivation during the dry season in irrigated areas may enhance the survival of snail hosts, resulting in an increase in snail populations (Wang et al. Reference Wang, Ho, Feng, Namsanor and Sithithaworn2015). However, results from other studies have demonstrated that effective water management in irrigation schemes can reduce snail populations (Chandiwana et al. Reference Chandiwana, Taylor, Chimbari, Ndhlovu, Makura, Bradley and Gondo1998), highlighting the importance of manipulating snail habitats in combination with mass drug administration programs to more effectively reduce trematode transmission (Fenwick et al. Reference Fenwick, Webster, Bosque-Oliva, Blair, Fleming, Zhang, Garba, Stothard, Gabrielli and Clements2009; Wang et al. Reference Wang, Chen, Guo, Zeng, Hong, Xiong, Wu, Wang, Wang and Xia2009).
The ecology of aestivating snails is potentially influenced by land use practices, particularly in rice paddies. These environments undergo dynamic changes through different stages: irrigation, full growth, complete drying, and occasional burning throughout the rice planting seasons. Farmers in northern Thailand generally cultivate rice once or twice annually. After harvesting, the most common method for land clearing and rice straw removal is open burning. Dead snails have been found in rice paddy sampling locations where burning has occurred. This practice may impact snail populations and disrupt the trematode life cycle, consequently affecting the transmission of associated diseases. However, open burning does not completely eradicate snail intermediate hosts in rice paddies. Some snail species can burrow deep into the soil (Chaiyaseang et al. Reference Chaiyasaeng, Pechdee, Sereewong, Suwannatrai, Laha and Tesana2019) or re-enter paddies from adjacent habitats via irrigation systems (Wang et al. Reference Wang, Feng, Sithithaworn, Feng and Petney2011, Reference Wang, Ho, Feng, Namsanor and Sithithaworn2015).
The study on the relationship between soil properties and the survival of aestivating snails revealed a significant positive correlation with the survival of viviparid snails. Conversely, no significant relationship was observed for bithyniid snails. A previous study indicated that salinity is the most important environmental variable affecting the density and distribution of Bithynia intermediate host snails of O. viverrini in the Khorat basin (Suwannatrai et al. Reference Suwannatrai, Suwannatrai, Haruay, Piratae, Thammasiri, Khampoosa, Kulsantiwong, Prasopdee, Tarbsripair, Suwanwerakamtorn, Sukchan, Boonmars, Malone, Kearney and Tesana2011). Currently, studies are underway to determine whether differences in the survival of snail intermediate hosts related to soil properties correspond to variations in trematode infection prevalence. However, due to the limited number of soil property parameters analysed, no conclusive results regarding the association between soil properties and the survival of aestivating snails could be established. Future investigations should focus on collecting a broader array of soil property parameters to achieve adequate data for analysing the survival of aestivating snails.
This study investigated trematode infections in aestivating snails from rice paddies during the dry season. Over 48 hours, no cercarial shedding was observed, aligning with Chaiyaseang et al. (Reference Chaiyasaeng, Pechdee, Sereewong, Suwannatrai, Laha and Tesana2019) who also found no shedding after reactivating aestivating snails. The absence of cercarial shedding could be due to several factors. First, the dry season induces aestivation in snails, slowing metabolism and potentially interfering with parasite development and cercarial release. Second, the stress of limited nutrients and prolonged aestivation may lead to higher mortality rates in infected snails, delaying parasite development or causing parasite death. It is also possible that both the snails and trematodes enter dormancy during aestivation, with parasites ceasing cercarial production while the snails are metabolically inactive. Alternatively, the snails may have already reached an advanced stage of infection, with metacercariae encysted before the dry season, and cercarial shedding may have occurred earlier. These hypotheses suggest that environmental stress and metabolic changes during aestivation play a critical role in the dynamics of trematode infections, highlighting the need for further research to understand the timing and environmental influences on parasite development.
Although no cercarial infections were found, this study identified four types of larval trematodes: echinostome metacercariae, T. anastomusa metacercariae, unidentified metacercariae, and rediae in bithyniid and viviparid snails. Previous reports have also found echinostome metacercariae in these snail families in Thailand (Burch and Lohachit Reference Burch and Lohachit1983; Chantima et al. Reference Chantima, Chai and Wongsawad2013, Reference Chantima, Suk-ueng and Kampan2018a, Reference Chantima, Phromkhong, Sonyod, Chaichana, McCormick, Mungthaisong, Khittasangka, Chomchalow, Walsh and Lisec2018b; Chantima and Rika Reference Chantima and Rika2020; Mard-arhin et al. Reference Mard-arhin, Prawang and Wongsawad2001). Additionally, these snail species have been reported as second intermediate hosts for Echinostomatidae in various areas of Southeast Asia (Chai et al. Reference Chai, Sohn, Na and De2011; Madsen and Hung Reference Madsen and Hung2014).
Metacercariae of T. anastomusa were found exclusively in F. doliaris, consistent with previous reports in viviparid snails (Filopaludina spp.) from Thailand (Chantima et al. Reference Chantima, Suk-ueng and Kampan2018a, Reference Chantima, Phromkhong, Sonyod, Chaichana, McCormick, Mungthaisong, Khittasangka, Chomchalow, Walsh and Lisec2018b; Chantima and Rika Reference Chantima and Rika2020; Phalee et al. Reference Phalee, Phalee and Wongsawad2018). Earlier studies also recorded T. anastomusa in Bellamya bengalensis (Viviparidae) in India (Agrawal et al. Reference Agrawal, Pandey and Kiran2002; Rai and Pande Reference Rai and Pande1967; Srivastava Reference Srivastava1953), with its adult stage found in the oesophagus of birds (Agrawal Reference Agrawal1958; Prudhoe Reference Prudhoe1957). The occurrence of T. anastomusa in multiple snail species suggests a broader host range, with birds likely serving as definitive hosts. Since birds commonly consume these snails, they may play a key role in its transmission and geographic spread. Further research is needed to clarify its host specificity, life cycle, and potential avian hosts in the region.
This study recorded a low number of infected aestivating snails with larval trematodes. Previous research has shown that the prevalence of larval trematode infections in snails is closely linked to snail population density (Lively Reference Lively2001). Recent studies suggest that anthropogenic factors such as pollution, habitat modification, and the introduction of non-native species significantly influence larval trematode infections in freshwater snails. These factors can alter the dynamics of trematode infections within snail populations, as well as affect the availability of potential hosts or environmental conditions in their habitats. (Bachtel et al. Reference Bachtel, Rittenhouse, Sandland and Koop2019; Kuris and Lafferty Reference Kuris and Lafferty1994). Understanding these interactions is crucial for assessing the ecological implications of trematode infections in freshwater ecosystems. To gain further insights into the relationship between larval trematodes and their snail intermediate hosts during the aestivated state, additional experimental studies are needed. Such studies should focus on controlling the timing of infection and aestivation. It is also recommended that future research explore the experimental aestivation of snails, both with and without parasitic infections, to enhance our understanding of this phenomenon.
Conclusion
This study focused on the diversity of aestivating snails and their potential role in the transmission of trematodes in rice paddies during the dry season. Our findings highlight that burrowing behaviour played a crucial role in the survival of bithyniid snails, leading to high abundance of aestivating individuals. Additionally, we identified various larval trematodes in bithyniid and viviparid snails, suggesting that aestivation may facilitate the persistence of trematode life cycles in seasonal rice paddy ecosystems. Given the high survival rate of aestivating snails, recolonization after the rainy season may contribute to the continued transmission of trematodes. While habitat modification and targeted snail control have been proposed as potential strategies for trematode management, their effectiveness in seasonal rice paddy ecosystems remains uncertain. Future research should prioritize field-based experimental studies on aestivation dynamics and their impact on snail population regulation and trematode transmission.
Acknowledgements
The authors gratefully acknowledge the Environmental Science and Technology Program and Biological Science Program, Faculty of Science and Technology, Chiang Rai Rajabhat University for providing facilities.
Competing interest
The authors declare that there are no conflicts of interest.
Ethical standard
We followed the guidelines for animal care in the International Guiding Principles of Biomedical Research Involving Animals (Council for International Organizations of Medical Sciences: CIOMS) including the relevant document (U1-03137-2559 to KC; U1-09340-2564 to KS).
