Description of the study area
Three locations were used for the study including Ikorodu, Ojota, and Badagry, all located within the Lagos metropolis in Southwestern Nigeria (Fig. 1). The study sites in these locations consisted of dumpsites and natural habitats. These locations were chosen because of the anthropogenic activities such as dumping of organic, inorganic, and toxic waste products within the vicinity and the presence of the natural habitat (less disturbed habitat) of the Amietophrynus regularis within the location. The first study station was Ikorodu dumpsite (06°35.8042'N, 03°34.8016'E) and natural habitat (06° 35′ 46.00″ N, 03° 34.5683′ E). The second was Badagry dumpsite (06° 25′ 42'' N, 02° 53′ 25″ E) and natural habitat (06°24′ 49″ N, 06° 53′ 52″ E). The third location was Ojota dumpsite (06° 35′ 40″ N, 03° 22′ 39″ E) and natural habitat (06° 34′ 47″ N, 03° 23′ 37″ E).
Sample collections
A total of 172 live Amietophrynus regularis (50 males, 122 females) were obtained randomly from the dumpsites and natural habitat of three different locations (Ikorodu, Ojota, and Badagry) from September 2019 to January 2021. They were collected with a sweep net or by hand. Samples were transported in well-aerated plastic containers to the Zoology department laboratory annex at the University of Lagos. On arrival to the laboratory, the morphometric features were obtained and the Amietophrynus regularis dissected. Selected organs (liver, lungs, and intestine) were collected for histopathology, and the blood smears for microscopy were carried out.
Morphometric assessment of the Amietophrynus regularis
Sex determination in the Amietophrynus regularis was conducted through physical observation of the throat area. The males are characterized by the dark or green throat, while the females are characterized by the white throat. Females also have coiled oviducts which are absent in males. A confirmatory determination was equally carried out on the reproductive system according to the description of Kobayashi et al. (2018).
The weight of each Amietophrynus regularis was determined to the nearest 0.01 g using a battery-powered digital Camry weighing balance (model EK-1A Series) before harvesting the organs. Using a vernier caliper, five measurements were taken to the nearest 0.1 cm. These included:
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Snout—urostyle length (SVL): This is the distance between the anterior tip of the snout and the posterior tip of the urostyle.
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Length of forelimb (LF): This is the distance between the posterior end of the humerus and the tip of the longest finger.
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Length of hind limb (LH): This is the distance between the tibial head and the tip of the fourth toe (which is the longest toe).
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Width across head (HW): This is the greatest width of the head at the level of the tympanum.
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Tympanic diameter (TD): This is the longitudinal distance between the outer margins of the tympanic annulus.
Laboratory analysis
Using sterile blades, the intestine of the Amietophrynus regularis specimens was eviscerated and placed in saline solution. The intestines were then dissected to obtain the enteric parasites. The lungs were also washed in saline solution. For further histopathological analysis, the intestine was preserved in separate sampling bottles containing Bouin's fluid. Thick and thin blood films were made on a clean glass slide for microscopy on blood parasites.
The recovered parasites were counted and recorded and then fixed in 70% alcohol. The enteric parasites collected from the Amietophrynus regularis were nematodes identified as Cosmocerca sp. and Amplicaecum africanum. The lung nematodes recovered were identified as Rhabdias spp. The identification procedure was carried out at the pathology laboratory of the Department of Veterinary Pathology, University of Ibadan, Nigeria. Parasite samples were then labeled according to the location and sex of Amietophrynus regularis collected.
Microscopy examination of blood and gastrointestinal samples
Blood specimens were obtained from clipped toes or the heart. Body fluid from the body cavity was also examined for the presence of parasites. The blood and body fluid specimens were first transferred to heparinized tubes. Thick and thin blood smears were prepared and left to air dry, and the thin film was fixed using absolute methanol. A working solution of Giemsa stain was prepared using the rapid method at 10% for 10 min after which it was flooded using buffered water at 7.0pH. The films were left to air dry and viewed using × 100 Objective with immersion oil. The direct examination method was used for the gastrointestinal samples. Intestinal samples were emulsified into normal saline and two drops were placed on the slide and a cover slip was placed on top. It was viewed using × 40 Objective.
Histopathology assessment
The intestinal tissues were placed in bottles containing Bouin’s fluid for 6 h, after which it was decanted and 10% buffered formalin was added to preserve the tissue. The tissues were routinely dehydrated in an ascending series of alcohol at 30 min intervals; they were then embedded in molten paraffin wax and allowed to solidify. The blocked tissues were sectioned at 4–5 microns processed and stained with hematoxylin and eosin (H&E) stains. The stained tissues were washed off in tap water, and the overstained ones were destained in 1% alcohol (Akinsanya et al. 2018). The tissues were mounted using DPX mountant and dried. Coverslips were then mounted over the sections and examined using a binocular dissecting microscope (American Optical Corporation, Model 570). The photomicrographs were taken with the aid of a Camera (INFINITY, 3-3URC 4.54 × 4.54 μm) in the pathology laboratory of the Department of Veterinary Pathology, the University of Ibadan, Nigeria.
Quality control and assurance
Glassware and dissecting kit used in this study were washed with detergent soaked in acetone and rinsed with tap water. They were then sealed with aluminum foil and autoclaved at 120 °C for 2 h. After drying and cooling, they were stored in a clean environment to prevent any accumulation of dust or other contaminants pending the next use. A single surgical blade was used per tissue sample, and thereafter, it was discarded safely. To avoid hand contamination of samples, sterile latex gloves and nose masks were used throughout the experimental session. For quality assurance analyte grade saline water was subjected to microbial and contamination analysis before use. All readings were taken in triplicate to minimize error.
Statistical analysis
The parasitological matrices: prevalence (%), mean intensity, and mean abundance, were analyzed and calculated as follows:
$${\text{Percentage}}\,{\text{prevalence}} = \frac{{{\text{Number}}\,{\text{of}}\,{\text{infected}}\,Amietophrynus\,regularis}}{{{\text{Number}}\,{\text{of}}\,Amietophrynus\,regularis\,{\text{examined}}}} \times 100$$
$${\text{Parasite}}\,{\text{abundance}} = \frac{{{\text{Number}}\,{\text{of}}\,{\text{collected}}\,{\text{parasites}}}}{{{\text{Number}}\,{\text{of}}\,Amietophrynus\,regularis\,{\text{examined}}}}$$
$${\text{Mean}}\,{\text{intensity}} = \frac{{{\text{Number}}\,{\text{of}}\,{\text{collected}}\,{\text{parasites}}}}{{{\text{Number}}\,{\text{of}}\,{\text{infected}}\,Amietophrynus\,regularis}}$$
The relationship between the prevalence of parasite infection and host factors such as the habitat of the host, sex, weight, and length was examined from data pooled from the three sampled locations within Lagos using Student’s t test analysis at a 5% level of significance. All statistical analyses were done using SPSS version 17 for Windows.