Prevalence of Gastrointestinal Parasitic Infections in Captive Snakes, Bangladesh

ABSTRACT

Background

Gastrointestinal (GI) parasitic infections are a common health concern in captive snakes, potentially affecting their well‐being. This study aimed to determine the prevalence and associated risk factors of helminth and protozoan infections in venomous and non‐venomous snakes.

Methods

A total of 174 faecal samples were collected from venomous and non‐venomous captive snakes housed at the Venom Research Centre of Chattogram, Bangladesh, between September 2024 and January 2025. Venomous snakes were housed individually in climate‐controlled terrariums and received biannual deworming; faecal samples were collected six months post‐treatment. In contrast, non‐venomous snakes were not dewormed and were kept either individually in climate‐controlled terrariums or group‐housed in unregulated terrariums. All collected faecal samples underwent routine coproscopy to identify the eggs and oocysts of helminths and protozoan parasites.

Results

Results demonstrated that the overall prevalence of GI parasitic infections was 39.1% (95% CI: 31.8–46.8). Helminths were significantly more prevalent (100%) than protozoa (8.8%). Non‐venomous snakes exhibited a higher prevalence of parasitic infections (70.8%) than venomous snakes (34%). Multiple infections were more common in non‐venomous snakes (76.5%) than in venomous snakes (47.1%). Nematodes were the most frequently detected helminths (91.2%), while Cestodes were found only in venomous snakes. Among venomous snakes, Ophiophagus hannah had the highest infection rate (66.7%), while among non‐venomous species, Python molurus and Ptyas mucosa showed the highest prevalence (80%). Risk factor analysis revealed that juveniles had significantly higher odds of infection (aOR = 2.55, p = 0.005), while snakes housed in controlled terrariums had lower infection risks (aOR = 0.143, p = 0.001).

Conclusion

This study provides the first record of GI parasitic infections in venomous and non‐venomous captive snakes in Bangladesh, highlighting the need for regular monitoring and appropriate therapeutic measures to mitigate their impact.

This study documented gastrointestinal parasitic infections in captive venomous and non‐venomous snakes in Bangladesh for the first time, finding an overall prevalence of 39.1%, with helminths more common than protozoa. Non‐venomous snakes and juveniles were most susceptible, while controlled terrarium housing reduced risk, highlighting the need for routine monitoring and preventive care.

1. Introduction

Snakes are an ecologically diverse group of reptiles, with over 3000 recognized species worldwide, about 600 of which are venomous, but only around 200 are medically significant to humans (Gopal et al. 2023). Bangladesh is home to more than 100 snake species, and clinical records of many remain scarce (Khan 2018; Khan and Chowdhury 2015). A recent study identified 65 venomous snake species where eight species (Daboia russelii, Bungarus caeruleus, Bungarus lividus, Bungarus niger, Naja naja, Bungarus walli, Naja kaouthia, Trimeresurus erythrurus) are recognized as medically important due to their frequent interactions with humans, and documented envenomation incidents (Al Haidar et al. 2023).

Venomous snakes play a critical role in antivenom production, advanced toxinology research and the discovery of novel biopharmaceuticals (de Souza et al. 2014), for these purposes, snakes are often collected from the wild and maintained in captivity. However, the transition of snakes from their wild habitats to captive conditions often results in substantial changes to their physiology, behaviour and overall health (Rajesh et al. 2015). Confinement, frequent handling, dietary changes and altered environmental parameters may induce chronic stress, weaken immune function and increase susceptibility to various infectious diseases, including viral, bacterial, fungal and parasitic infections (Fischer and Romero 2019; Van et al. 2018).

Gastrointestinal (GI) parasitic infections are among the most prevalent diseases in captive reptiles and considered one of their most significant health challenges (Ferdous et al. 2023; Akhila et al. 2018; Carrera‐Játiva et al. 2018; Adeniyi et al. 2015). In natural habitats, snakes often maintain an ecological balance with parasites, limiting disease manifestation (Barbosa et al. 2020; Moudgil and Singla 2015; Thawait et al. 2014). Many endoparasites including intestinal helminths, protozoa and pentastomids have been documented in both wild and captive snakes, which may serve as definitive, intermediate, paratenic or accidental hosts for these parasites (Nasiri et al. 2014). Snakes can harbour parasites for extended periods without noticeable clinical signs until external factors compromize their immune system (Patra et al. 2017; Rajesh et al. 2015; Reavill and Griffin 2014). In captivity, disruption of the natural host–parasite balance, combined with stressful conditions, facilitates parasite transmission, through contaminated substrates or faeces and eventual disease outbreaks, leading to reduced fitness and even death (Rajesh et al. 2015; Nasiri et al. 2014; Rataj et al. 2011). Understanding diversity of these parasites is vital for improving captive husbandry practices, preventing disease outbreaks and ensuring the health of captive animals.

In Bangladesh, the Venom Research Centre, Chattogram is the only facility that houses venomous snakes from various geographic regions of the country for venom extraction and other research (Rahman et al. 2024). Despite the recognized importance of parasite surveillance in reptiles, there is a complete lack of data on GI parasitic infections in captive snakes in the country. Therefore, the current study was undertaken to investigate the prevalence and diversity of GI parasitic infections in captive snakes providing baseline data to support improved health management and well‐being of captive snake populations.

2. Materials and Methods

2.1. Ethics Approval Statement

Before the research was conducted, Venom Research Centre authorities reviewed and approved all the procedures. The samples for the current study were opportunistically and non‐invasively taken from the snake terrarium without handling or disturbing the animals. No direct interaction, restraint or invasive procedures were involved during sample collection.

2.2. Study Site and Snake Husbandry

The study was conducted at the Venom Research Centre of Chattogram, Bangladesh, between September 2024 and January 2025. The centre housed over 400 venomous snakes representing eight species, collected from various regions of Bangladesh for venom extraction and research purposes. Additionally, the facility housed four non‐venomous snake species, which were kept for exhibition purposes and as prey for certain other snake species.

Both venomous and non‐venomous snakes were individually housed in a separate terrarium, with conditions carefully maintained at 27°C and 70% humidity; however, some non‐venomous snake species were occasionally group‐housed in shared terrariums without environmental control, resulting in natural fluctuations in temperature and humidity. Some enclosures had muddy flooring.

A dedicated staff member, supported by CCTV monitoring, oversaw individual faecal sample collection. Snake demographic and management data including age, sex, diet, housing type, terrarium conditions, deworming status and floor type were obtained from the centre's daily updated database. Snakes were categorized into two age groups: juvenile and adult, with the criteria for adulthood and juvenile varying by species (Table S1).

The feeding schedule for the snakes was carefully managed, varying by species and dietary requirements. Bungarus caeruleus, Bungarus niger, Naja naja, Naja kaouthia, Xenochrophis piscator, Ptyas mucosa, Python molurus and Coelognathus radiatus were fed every 7 days; while Trimeresurus erythrurus and Daboia russelii were fed every 10 days. Ophiophagus hannah required a longer interval and was fed every 20 days. Most snakes were fed mice except Python molurus (chicks), Xenochrophis piscator (primarily fed on fish) and Ophiophagus hannah (snakes).

Venomous snakes received a biannual deworming protocol consisting of topical application of ivermectin and oral administration of fenbendazole (70 mg/kg body weight), while the non‐venomous snakes were not treated. Faecal samples for this study were collected 6 months after the administration of anthelmintics. All snakes were closely monitored by zoologists and treated by veterinarians to ensure their health and well‐being.

2.3. Faecal Sample Collection and Preservation

A total of 174 faecal samples were collected from 11 different snake species. The distribution of samples was as follows: 20 samples from Bungarus caeruleus, 17 from Bungarus niger, 5 from Trimeresurus erythrurus, 20 from Daboia russelii, 38 from Naja naja, 47 from Naja kaouthia, 3 from Ophiophagus hannah, 4 from Xenochrophis piscator, 10 from Python molurus, 5 from Ptyas mucosa, 5 from Coelognathus radiatus. As the number of samples collected per species was not uniform, with some species represented by many individuals (e.g., Naja kaouthia) and others by only a few (e.g., Ophiophagus hannah, Xenochrophis piscator), this variation may introduce potential bias in estimating parasite prevalence. Larger sample sizes generally provide more reliable estimates, whereas smaller sample sizes may lead to either under‐ or over‐estimation of prevalence and reduce the comparability across species. Therefore, the prevalence values for species with limited samples should be interpreted with caution.

For each individual, approximately 5–10 g of individual, freshly voided faecal samples were collected from the terrarium floor early morning before routine cleaning. Samples were immediately transferred to clean labelled plastic containers and preserved in 10% formalin. They were then transported to the Parasitology Laboratory, Department of Pathology and Parasitology, Chattogram Veterinary and Animal Sciences University, Bangladesh, and stored at 4°C until analysis.

2.4. Examination of Faecal Samples

The samples were analysed using routine faecal tests (e.g., direct smear, flotation and sedimentation) to identify the morphological characteristics of eggs of helminths and oocysts of protozoan parasites. For performing direct smear, faecal samples were homogenized in water and the undigested materials were strained out. Then, a drop of faecal suspension was placed on a glass slide and examined it under a microscope. For the test tube flotation method, around 3 g of faeces were mixed with 50 mL of saturated sugar solution (flotation fluid). A strainer was used to remove coarse faecal materials from the resulting faecal suspension. The filtrate was then transferred into a test tube, and a coverslip was placed on top, touching the convex meniscus of the liquid. After 15 min, the coverslip was gently lifted, placed onto a glass slide, and examined under a microscope. For the simple sedimentation method, the faecal suspension was allowed to settle for about 15 min. The supernatant was discarded, and a drop of the sediment was taken onto a glass slide and examined under a microscope (Hendrix and Robinson 2022). Triplet smears were prepared for each specimen tested. A sample was considered ‘positive’ if at least one egg or oocyst was identified in the smear examined. However, parasite species were only identified up to the genus level and in some cases up to species level. Morphological features of eggs of helminths and oocysts of protozoa were identified through a comprehensive review of published literature (Mendoza‐Roldan et al. 2024; Mustafa et al. 2018; Rom et al. 2018; Patra et al. 2017; Vasaruchapong et al. 2017; Šlapeta et al. 2017; Rajesh et al. 2015; de Souza et al. 2014; Divers and Mader 2005; Chaiyabutr and Chanhome 2002) (Figure 1).

FIGURE 1.

Eggs and oocysts of helminths and protozoa identified in snake faeces.

2.5. Statistical Analysis

Microsoft Excel 2016 was used to compile and sort the raw data in a spreadsheet, and IBM SPSS Statistics (Version 26) was then used to perform data analysis. We performed descriptive statistics using percentages. To identify potential risk factors for GI parasitic infection, a Chi‐square test was performed to assess associations between the binary outcome variable and categorical risk factors. Univariable logistic regression was then conducted to estimate the odds ratios for each risk factor. Variables with a p value ≤ 0.20 in the univariable analysis were selected for inclusion in the multivariable logistic regression model. A more liberal threshold (p ≤ 0.20) was applied to minimize the risk of excluding potentially important predictors, recognizing that variables not strongly associated in univariable analysis may still act as significant predictors or confounders once adjusted for other factors in the multivariable model. In the final model, variables with a p value ≤ 0.05 were considered statistically significant. A single infection was defined as the detection of either one helminth egg or one protozoan oocyst in a sample. Multiple infections were considered when more than one helminth egg and/or protozoan oocyst was identified in a sample.

3. Result

3.1. Overall Prevalence of Gastrointestinal Parasitic Infections in Snakes

The overall prevalence of GI parasitic infections (helminths and protozoa) in snakes (both venomous and non‐venomous) was 39.1% (95% CI: 31.8–46.8, N = 174). Helminth infections (100%; 95% CI: 94.7–100) were significantly more common than protozoan infections (8.8%; 95% CI: 3.3–18.2) (Figure 2, Table S2). Among the helminths, Nematodes were the most frequently observed (91.2%; 95% CI: 81.8–96.7), followed by Cestodes (17.6%; 95% CI: 9.5–28.8) and Trematodes (1.5%; 95% CI: 0.7–9). Regarding infection patterns, single infections were less (41.2%; 95% CI: 29.4–53.8) than multiple infections which accounted for 58.8% (95% CI: 46.2–70.6) of cases in snakes (Table S2).

FIGURE 2.

Overall prevalence of helminths and protozoan infections in venomous and non‐venomous snake groups.

3.2. Prevalence of Gastrointestinal Parasitic Infections in Venomous and Non‐Venomous Snake Groups

GI parasitic infections (helminths and protozoa) were more prevalent in non‐venomous snakes (70.8%; 95% CI: 48.9–87.4) compared to venomous snakes (34%; 95% CI: 26.5–42.2) (Figure 2, Table S2). Interestingly, multiple infections were predominantly observed in non‐venomous snakes (76.5%; 95% CI: 50.1–93.2), whereas single infections were more common in venomous snakes (47.1%; 95% CI: 32.9–61.5). Helminth infections exhibited a higher overall prevalence compared to protozoal infections across both snake groups. While helminth infections were universally detected in both venomous (100%; 95% CI: 95.9–100) and non‐venomous snakes (100%; 95% CI: 81.5–100), protozoal infections were more prevalent in non‐venomous snakes (17.6%; 95% CI: 3.8–43.4). Among helminth parasites, Nematodes were the most frequently detected, with a prevalence of 94.1% (95% CI: 71.3–99.9) in non‐venomous snakes and 90.2% (95% CI: 78.6–96.7) in venomous snakes. Cestodes were identified exclusively in venomous snakes at a prevalence of 23.5% (95% CI: 12.8–37.5), whereas Trematodes were found only in non‐venomous snakes (Table S2).

In venomous snake species, GI parasitic infections (helminths and protozoa), were most prevalent in Ophiophagus hannah, with a prevalence of 66.7% (95% CI: 9.4–99.2), while the lowest prevalence was observed in Bungarus caeruleus at 5% (95% CI: 0.1–24.9) (Table 1). Multiple infections were highest in Naja kaouthia, with a prevalence of 75% (95% CI: 50.9–91.3). In contrast, no multiple infections were detected in Bungarus caeruleus and Trimeresurus erythrurus, although single infections were observed at 100% in both species. All venomous snake species that tested positive for GI parasitic infections harboured helminths (100%). However, protozoal infections were limited to Naja naja, Naja kaouthia, with the highest prevalence recorded in Naja naja at 14.3% (95% CI: 1.8–42.8) (Table 1). Among helminths, Nematodes were the most frequently detected, showing a prevalence of 100% across all venomous snake species. Cestodes were detected exclusively in Naja naja, Naja kaouthia and Daboia russelii, with a prevalence of 35.7%, 30% and 10% respectively. Trematodes were not detected in any venomous snake species (Table 1).

TABLE 1.

Overall prevalence of helminths and protozoan infections in venomous snakes.

		BC (N = 20)		BN (N = 17)		TE (N = 5)		DR (N = 20)		NN (N = 38)		NK (N = 47)		OH (N = 3)
Variables	Categories	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI
Presence of parasites	Yes	5 (1)	0.1–24.9	17.6 (3)	3.8–43.4	20 (1)	0.5–71.6	50 (10)	27.2–72.8	36.8 (14)	21.8–54	42.6 (20)	28.3–57.8	66.7 (2)	9.4–99.2
	No	95 (19)	75.1–99.9	82.4 (14)	56.6–96.2	80 (4)	28.4–99.5	50 (10)	27.2–72.8	63.2 (24)	46–78.2	57.4 (27)	42.2–71.7	33.3 (1)	0.8–90.6
Types of infections	Single	100 (1)	2.5–100	66.7 (2)	9.4–99.2	100 (1)	2.5–100	60 (6)	26.2–87.8	57.1 (8)	28.9–82.3	25 (5)	8.7–49.1	50 (1)	1.3–98.7
	Multiple	0	0–97.5	33.3 (1)	0.8–90.6	0	0–97.5	40 (4)	12.2–73.8	42.9 (6)	17.7–71.1	75 (15)	50.9–91.3	50 (1)	1.3–98.7
Types of parasites	Helminths	100 (1)	2.5–100	100 (3)	29.2–100	100 (1)	2.5–100	100 (10)	69.2–100	100 (14)	76.8–100	100 (20)	83.2–100	100 (2)	15.8–100
	Protozoa	0	0–97.5	0	0–70.8	0	0–97.5	0	0–30.8	14.3 (2)	1.8–42.8	5 (1)	0.1–24.9	0	0–84.2
Types of helminths	Cestode	0	0–97.5	0	0–70.8	0	0–97.5	10 (1)	0.3–44.5	35.7 (5)	12.8–64.9	30 (6)	11.9–54.3	0	0–84.2
	Nematode	100 (1)	2.5–100	100 (3)	29.2–100	100 (1)	2.5–100	90 (9)	55.5–99.7	71.4 (10)	41.9–91.6	100 (20)	83.2–100	100 (2)	15.8–100
	Trematode	0	0–97.5	0	0–70.8	0	0–97.5	0	0–30.8	0	0–23.2	0	0–16.8	0	0–84.2

Abbreviations: BC, Bungarus caeruleus; BN, Bungarus niger; DR, Daboia russelii; NN, Naja naja, NK, Naja kaouthia; OH, Ophiophagus hannah; TE, Trimeresurus erythrurus.

In non‐venomous snake species, GI parasitic infections were most prevalent in Python molurus and Ptyas mucosa, with a prevalence of 80%, while the lowest prevalence was observed in Coelognathus radiatus at 40% (Table 2). Multiple infections were highest in Python molurus, where all positive cases (100%; 95% CI: 63.1–100) involved multiple infections. In contrast, no multiple infections were detected in Coelognathus radiatus, although single infections were observed at 100%. All non‐venomous snake species that tested positive for parasitic infections harboured helminths, with a prevalence of 100%. However, protozoan infections were observed exclusively in Ptyas mucosa, with a prevalence of 75% (95% CI: 19.4–99.4). Among helminth parasites, Nematodes were the most commonly detected, showing the highest prevalence of 100% in Python molurus, Ptyas mucosa and Coelognathus radiatus species and the lowest prevalence in Xenochrophis piscator at 66.7%. Cestodes were not detected in any of the snake species. However, Trematodes were identified only in Xenochrophis piscator, with a prevalence of 33.3% (95% CI: 0.8–90.6) (Table 2).

TABLE 2.

Overall prevalence of helminths and protozoan infections in non‐venomous snakes.

		XP (N = 4)		PYM (N = 10)		PM (N = 5)		CR (N = 5)
Variables	Categories	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI	Prevalence	95% CI
Presence of parasites	Yes	75 (3)	19.4–99.4	80 (8)	44.4–97.5	80 (4)	28.4–99.5	40 (2)	5.3–85.3
	No	25 (1)	0.6–80.6	20 (2)	2.5–55.6	20 (1)	0.5–71.6	60 (3)	14.7–94.7
Types of infections	Single	33.3 (1)	0.8–90.6	0	0–36.9	25 (1)	0.6–80.6	100 (2)	15.8–100
	Multiple	66.7 (2)	9.4–99.2	100 (8)	63.1–100	75 (3)	19.4–99.4	0	0–84.2
Types of parasites	Helminths	100 (3)	29.2–100	100 (8)	63.1–100	100 (4)	39.8–100	100 (2)	15.8–100
	Protozoa	0	0–70.8	0	0–36.9	75 (3)	19.4–99.4	0	0–84.2
Types of helminths	Cestode	0	0–70.8	0	0–36.9	0	0–60.2	0	0–84.2
	Nematode	66.7 (2)	9.4–99.2	100 (8)	63.1–100	100 (4)	39.8–100	100 (2)	15.8–100
	Trematode	33.3 (1)	0.8–90.6	0	0–36.9	0	0–60.2	0	0–84.2

Abbreviations: CR, Coelognathus radiatus; PM, Ptyas mucosa; PYM, Python molurus; XP, Xenochrophis piscator.

3.3. Occurrence of Helminths and Protozoan Infections

Several genera of both helminths and protozoa were identified in the examined snake samples (Table 3). Among the helminths, Strongyloides sp. (38.82%) was the most frequently detected, followed by Kalicephalus sp. (20.59%) and Capillaria sp. (13.23%). Strongyloides sp. was recorded in Bungarus niger, Daboia russelii, Trimeresurus erythrurus, Naja kaouthia, Naja naja, Python molurus and Ptyas mucosa, whereas Kalicephalus sp. was found in Bungarus niger, Daboia russelii, Naja kaouthia, Python molurus and Coelognathus radiatus. Cestodes were mainly observed in Naja kaouthia, Naja naja and Daboia russelii. Polydelphis sp. was recorded only in the non‐venomous snake Python molurus. Among the protozoa, Caryospora sp. was detected in Naja kaouthia, Naja naja and Ptyas mucosa. Pentastomes were recorded only in Naja kaouthia. In addition, pseudoparasites including Syphacia muris, Aspicularis tetraptera and mice oxyurid eggs were recovered from snakes (e.g., Naja kaouthia, Naja naja, Bungarus caeruleus, Bungarus niger, Daboia russelii, Trimeresurus erythrurus and Coelognathus radiatus) that had been fed with laboratory mice.

TABLE 3.

Relative prevalence of helminths and protozoan infections among infected snakes (venomous and non‐venomous).

Helminths and protozoa	Infected snake species	Relative prevalence percentage (positive samples)
		Venomous snakes (51)	Non‐venomous snakes (17)	Overall (68)
Cestodes
Taeniid	DR	1.96 (1)	0	1.47 (1)
Hymenolepis nana	NK, NN	17.65 (9)	0	13.23 (9)
Hymenolepis diminuta	NN	1.96 (1)	0	1.47 (1)
Unknown tapeworm	NK	3.92 (2)	0	2.94 (2)
Nematodes
Capillaria spp.	NK	17.65 (9)	0	13.23 (9)
Strongyloides spp.	BN, DR, TE, NK, NN, PYM, PM	33.33 (17)	35.29 (6)	38.82 (23)
Rhabdias spp.	DR	1.96 (1)	0	1.47 (1)
Kalicephalus spp.	BN, DR, NK, PYM, CR	15.69 (8)	35.29 (6)	20.59 (14)
Strongylus spp.	DR, OH	5.88 (3)	0	4.41 (3)
Aspicularis tetraptera	BN, NK, NN	11.76 (6)	0	8.82 (6)
Dujardinascaris spp.	XP	0	11.76 (2)	2.94 (2)
Ophidascaris spp.	BC, NK	3.92 (2)	0	2.94 (2)
Oxyurid	NK, OH	9.80 (5)	0	7.35 (5)
Polydelphis spp.	PYM	0	35.29 (6)	8.82 (6)
Syphacia muris	DR, NK, NN	13.72 (7)	0	10.29 (7)
Unknown pinworm	NN	1.96 (1)	0	1.47 (1)
Mice oxyurid	BC, BN, DR, TE, NK, NN, CR	82.35 (42)	11.76 (2)	64.70 (44)
Trematodes
Unknown Trematode	CR	0	5.88 (1)	1.47 (1)
Protozoa
Caryospora spp.	NK, NN, PM	3.92 (2)	5.88 (1)	4.41 (3)
Eimeria spp.	PM	0	17.65 (3)	4.41 (3)
Sarcocystis spp.	NN	1.96 (1)	0	1.47 (1)
Pentastomes	NK	1.96 (1)	0	1.47 (1)

Note: This table reports prevalence among parasite‐positive snakes (relative prevalence). Percentages indicate the proportion of infected snakes harbouring each parasite species within the group. The overall prevalence of gastrointestinal parasites in the sampled population is reported separately in Table S2.

Abbreviations: BC, Bungarus caeruleus; BN, Bungarus niger; CR, Coelognathus radiatus; DR, Daboia russelii; NN, Naja naja, NK, Naja kaouthia; OH, Ophiophagus hannah; PM, Ptyas mucosa; PYM, Python molurus; TE, Trimeresurus erythrurus; XP, Xenochrophis piscator.

3.4. Risk Factors Associated With Gastrointestinal Parasite Infections in Snakes

The univariable analysis identified six potential risk factors associated with GI parasitic infections in snakes: age, terrarium conditions, food, sex, deworming, housing type and floor type (p ≤ 0.20). Subsequently, a multivariable logistic regression analysis revealed significant associations between age and terrarium conditions with the presence of GI parasitic infections. Regarding age, juveniles had 2.55 times higher odds of experiencing GI parasitic infections compared to adults (aOR = 2.55, 95% CI: 1.33–4.89, p = 0.005). As for terrarium conditions, snakes housed in an environmentally controlled terrarium had significantly lower odds of infection compared to those in uncontrolled conditions (aOR = 0.143, 95% CI: 0.04–0.46, p = 0.001). This indicates that keeping snakes in an environmentally controlled terrarium reduces the odds of infection by approximately 85.7% compared to an uncontrolled environment (Table 4).

TABLE 4.

Univariable and multivariable logistic regression analysis of risk factors associated with gastrointestinal parasitic infections in snakes.

Variables	Covariable	No. of snakes	No. of snake's positive for gastrointestinal parasite (%)	Univariable analysis		Multivariable analysis
				OR (95% CI)	p value	aOR (95% CI)	p value
Age	Adult	94	27 (28.72)	Reference	0.003	Reference	0.005
	Juvenile	80	41 (51.25)	2.61 (1.39–4.88)		2.55 (1.33–4.89)
Terrarium Conditions	Uncontrolled	19	15 (78.95)	Reference	0.001	Reference	0.001
	Environmental controlled	155	53 (34.19)	0.139 (0.04–0.44)		0.143 (0.04–0.46)
Food	Mice	157	55 (35.03)	Reference	0.029
	Snake	3	2 (66.67)	3.71 (0.33–41.83)
	Chick	10	8 (80)	7.42 (1.52–36.15)
	Fish	4	3 (75)	5.56 (0.56–54.76)
Sex	Male	84	31 (36.90)	Reference	0.57
	Female	90	37 (41.11)	1.19 (0.65–2.19)
Deworming	No	24	17 (70.83)	Reference	0.001
	Yes	150	51 (34)	0.212 (0.083–0.54)
Housing type	Group	14	11 (78.57)	Reference	0.005
	Individual	160	57 (35.62)	0.151 (0.04–0.56)
Floor type	Muddy	15	10 (66.67)	Reference	0.029
	Plastic sheet	159	58 (36.48)	0.287 (0.094–0.88)

Abbreviations: aOR, adjusted odds ratio; CI, confidence interval; OR, odds ratio.

4. Discussion

Parasites are among the most common pathogens affecting reptiles (e.g., snakes) in captivity, whether they are housed in zoos, breeding facilities, or kept as domestic pets (Wolf et al. 2014). These parasites can have significant negative impacts on the health and well‐being of the animals. Regular monitoring of GI parasitism, including both helminths and protozoa, is essential to ensure effective management and to maintain the overall health of captive snakes. GI parasitic infections have been reported in snakes across various regions worldwide (Halán and Kottferová 2021; Hallinger et al. 2020; Mustafa et al. 2018; Rom et al. 2018; Akhila et al. 2018; Patra et al. 2017; Rajesh et al. 2015; Ras‐Norynska and Sokol 2015; de Souza et al. 2014; Wolf et al. 2014; Chaiyabutr and Chanhome 2002). This study specifically documents the presence of GI parasitic infections in snakes within Bangladesh, contributing to the understanding of parasitism in reptiles in this region.

In this study, the overall prevalence of GI parasitic infections in snakes was found to be 39.1%, aligning with findings from earlier research (Rinaldi et al. 2012). However, reported prevalence rates vary across studies. Some investigations on captive snake populations have documented higher prevalence ranging from 70% to 80% (Akhila et al. 2018; de Souza et al. 2014; Chaiyabutr and Chanhome 2002), while others have reported lower rates, such as 46% (Patra et al. 2017). In contrast, studies focusing on particular snake species have shown wider variability, with infection rates ranging from 40% to 95% (Vasaruchapong et al. 2017; Ras‐Norynska and Sokol 2015). These discrepancies may be attributed to several factors, including differences in the number of specimens examined, species variation, dietary sources, geographic location and living conditions. Additionally, the variation could stem from the fact that some studies focused on captive snakes, while others examined wild or aquatic species, each with distinct environmental and physiological influences.

In current study, venomous snakes were maintained under a strict anthelmintic schedule, housed individually in separate terrariums and prevented from contact with soil. These measures likely contributed to the lower overall prevalence of GI parasitic infections observed in venomous snakes (34%). In contrast, non‐venomous snakes were not administered anthelmintics and were housed communally in species‐specific groups within terrariums featuring muddy floors. This communal housing facilitated the potential transmission of parasites among non‐venomous snakes, including self‐reinfection through contact with contaminated faeces (Rajesh et al. 2015; Rataj et al. 2011). As a result, the overall prevalence of GI parasitic infections (both helminths and protozoa) was significantly higher in non‐venomous snakes (70.8%). Factors such as husbandry practices, overcrowding and the presence of concurrent infections may contribute to the occurrence of protozoan infections in snakes (Rajesh et al. 2015). Rajesh et al. (2015) noted that protozoa (coccidia) are typically transmitted through the ingestion of sporulated oocysts from contaminated faeces or soil, highlighting the importance of proper hygiene and housing conditions in preventing parasitic infections. Discrepancies were noted when comparing these findings to previous studies, which reported a 75% prevalence of GI parasitic infections in venomous snakes and 70.6% in non‐venomous snakes (Chaiyabutr and Chanhome 2002). These variations may be attributed to differences in the use of anthelmintic treatments, hygiene standards in terrariums or other management practices.

Among the helminths, Nematodes exhibited the highest prevalence of infections in both venomous and non‐venomous snake groups, a finding consistent with previous studies (Patra et al. 2017; de Souza et al. 2014; Chaiyabutr and Chanhome 2002). The higher occurrence of Nematode infections can be attributed to the fact that many GI Nematodes of reptiles possess a direct (monoxenous) life cycle that does not require intermediate hosts, facilitating transmission in confined environments (Nasiri et al. 2014). Nematodes are easily transmitted through contaminated feed, water and soil, facilitating their spread (Mir et al. 2016). In contrast, Trematodes and most Cestodes depend on at least one intermediate host to complete their life cycle, making their transmission more complex (Mir et al. 2016; Nasiri et al. 2014). This reliance on intermediate hosts likely explains the lower occurrence of Trematode and Cestode infections in both snake groups. In this study, we detected Trematode infection in only one water snake (Xenochrophis piscator) sample. Water snakes primarily feed on fish, which are known to act as intermediate hosts for a variety of parasites, including Trematodes (Labony et al. 2020). Additionally, we identified Cestodes in this study, including Taeniid, Hymenolepis nana and Hymenolepis diminuta, at a prevalence of 17.6%. These Cestodes do not naturally infect snakes but are likely acquired through ingesting infected prey, making them spurious rather than host‐specific parasites (Mendoza‐Roldan et al. 2024).

During the faecal examination, Rhabdias spp. larvae‐containing eggs were identified in a sample from the venomous snake (Daboia russelii). Among the detected Nematodes, Strongyloides sp. was the most prevalent, with an overall occurrence of 38.82% in both venomous and non‐venomous snakes. Previous studies reported varying prevalence rates of Strongyloides sp., with Hallinger et al. (2020) documenting 2.6% and Akhila et al. (2018) reporting 26.5%. Although Rhabdias spp. eggs and Strongyloides sp. eggs appear similar, Rhabdias spp. eggs are larger. In this study, Rhabdias spp. eggs measured 60 µm × 85 µm, while Strongyloides sp. eggs ranged from 30–34 µm × 55–60 µm. These findings are consistent with previous studies (Vasaruchapong et al. 2017; Jacobson 2007). Additionally, snakes containing Rhabdias spp. eggs exhibited breathing difficulties, further confirming their identification. Rhabdias spp. infections can cause verminous pneumonia, leading to open‐mouth breathing and a frothy discharge from the nares (Šlapeta et al. 2017). On the other hand, Strongyloides sp. infections result in enteritis (Šlapeta et al. 2017).

In this study, Kalicephalus spp. were identified as the second most prevalent intestinal Nematode species, with an infection rate of 20.59%. These Nematodes are reported to infect snakes worldwide, with prevalence rates varying from 3.3% to 60% (Hallinger et al. 2020; Patra et al. 2017; Vasaruchapong et al. 2017; Chaiyabutr and Chanhome 2002). Kalicephalus Nematodes are blood‐feeding parasites that attach to the intestinal lining to access blood vessels. Infection with these Nematodes can range from mild enteritis to severe, even fatal, haemorrhagic‐necrotic gastroenteritis, often complicated by secondary bacterial infections (Hallinger et al. 2020; Kavitha et al. 2014). Kalicephalus spp. infections was found to be lower in venomous snakes (15.69%) compared to non‐venomous snakes (35.29%). This difference may be attributed to several factors, including suboptimal conditions in captivity, a lack of deworming practices and varying feeding habits (Patra et al. 2017).

Pentastomes are worm‐like arthropods with a unique life cycle and zoonotic potential. In our study, we detected pentastome eggs in the faecal sample of a venomous snake (Naja kaouthia). A previous study reported a 22.5% prevalence of pentastome eggs in snake faeces (Vasaruchapong et al. 2017). Adult pentastomes inhabit the lungs of snakes, where they release fertilized eggs into the respiratory secretions. The infected snakes then swallow these eggs, which are subsequently excreted in the faeces (Paré 2008).

In the current investigation, Ophidascaris sp. was identified from venomous snakes (Bungarus caeruleus and Naja kaouthia), while Polydelphis sp. was found exclusively in Python molurus, a non‐venomous snake, with a prevalence of 35.29%. These findings align with previous studies indicating that Ophidascaris sp. is more prevalent in venomous snakes, whereas Polydelphis sp. tends to be more abundant in non‐venomous snakes (Patra et al. 2017). Pseudoparasites can be detected in snake faeces during faecal examinations, including Syphacia muris, Hymenolepis nana, Hymenolepis diminuta, Aspicularis tetraptera, mice oxyurid eggs and mite eggs. These pseudoparasites do not cause clinical signs in snakes, as they originate from the prey, such as mice, rather than from the snakes themselves (Divers & Mader 2005). In our study, we identified pseudoparasites in 38.50% of total samples. This finding aligns with a previous study by Rinaldi et al. (2012), which reported a higher prevalence of pseudoparasites at 39.2%. In contrast, another study by Ellerd et al. (2022) documented a significantly lower prevalence pseudoparasites of 1.2%. Wild snakes are more likely to harbour pseudoparasites compared to those in captivity, primarily because they actively hunt and consume small prey in their natural habitat, which may carry such organisms (Mendoza‐Roldan et al. 2024).

Several protozoa were identified in the present study, including Caryospora sp., Eimeria sp. and Sarcocystis sp. The occurrence of these protozoa was documented in previous studies worldwide (Ras‐Norynska and Sokol 2015; de Souza et al. 2014). Both Caryospora sp. and Eimeria sp. have direct life cycles, involving a single host. Their oocysts are produced and excreted in faeces, continuing the transmission cycle. In contrast, Sarcocystis spp. follow an obligate two‐host cycle, typically involving a predator‐prey relationship, where the definitive host acquires the infection by consuming infected tissues of the intermediate host (Bezerra et al. 2023; Divers and Mader 2005). It is important to note that not all infections with coccidians result in disease. The term coccidiasis refers to the presence of coccidian parasites without clinical disease, whereas coccidiosis occurs when the infection leads to pathological effects. Many healthy individuals may shed large numbers of oocysts, indicating infection rather than disease (Divers and Mader 2005). Additionally, Sarcocystis spp. found in snakes have potential public health significance, highlighting the need for further study on their impact and transmission risks (Bezerra et al. 2023).

The current study identified age and terrarium conditions as significant risk factors associated with GI infections in captive snakes. Multivariable logistic regression analysis revealed that juveniles had 2.55 times higher odds of being infected compared to adults. This finding suggests that younger snakes may be more susceptible to parasitic infections, possibly due to their higher activity levels and broader dietary diversity, which expose them to a greater range of potential parasites. In contrast, larger, more mature snakes likely benefit from a more evolved immune system, resulting in fewer infections (Santoro et al. 2013; Capizzi et al. 2008). Additionally, snakes housed in environmentally controlled terrariums had 85.7% lower odds of infection compared to those in uncontrolled environments. GI parasite burden tends to be higher during warmer, more humid seasons, as increased humidity can prolong the survival of parasite eggs, while fluctuating temperatures may accelerate their development (Lee et al. 2022; Lettoof et al. 2020). This may explain the significantly higher infection risk observed in snakes housed in uncontrolled environments. Proper environmentally controlled terrariums, including regulated temperature, humidity and sanitation, likely reduce parasite survival and transmission by limiting favourable conditions for parasite persistence. Proper management of captive snakes requires regular health monitoring and preventive measures to control parasitic infections.

5. Conclusion

This study provides the first systematic assessment of GI parasitic infections in captive venomous and non‐venomous snakes in Bangladesh. Nematodes were identified as the most prevalent parasites, with infection patterns varying across snake species and husbandry conditions. Such evidence highlights that captivity does not eliminate parasite risks and, in some cases, may facilitate transmission. Although the sample size was limited, the evidence underscored the importance of routine parasitological screening and targeted deworming protocols for both snakes and their prey. Implementing such measures will improve captive snake health, welfare and management standards. Future research should include larger populations and wild counterparts to better understand parasite dynamics and associated conservation risks.

Author Contributions

Nazmul Hasan: investigation, methodology, data curation, formal analysis, visualization, software, writing –original draft preparation, writing – review and editing (equal). Sahriar Islam: investigation. Mahmudur Rahman: investigation. Abdul Auawal: investigation, resources, validation. Md. Mizanur Rahman: resources. Mohammed Noman: resources. Abdullah Abu Sayeed: resources. Mohammad Abdul Alim: conceptualization, methodology, funding acquisition, resources, project administration, supervision, validation, writing – review and editing (equal).

Funding

The authors have nothing to report.

Ethics Statement

Before the research was conducted, Venom Research Centre authorities reviewed and approved all the procedures. The samples for the current study were opportunistically and non‐invasively taken from the snake terrarium without handling or disturbing the animals. No direct interaction, restraint, or invasive procedures were involved during sample collection.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supplementary Table 1: Criteria for juvenile and adult stages in various snake species.

Supplementary Table 2: Overall prevalence of gastrointestinal parasitic infections in venomous and non‐venomous snake groups.

Data Availability Statement

Data files associating this research are available at https://figshare.com/s/14a2d81e0400c7ba489e