Rugopharynx australis (Nematoda: Strongyloidea) Infection in Captive Red Kangaroos (Osphranter rufus) in Bulgaria: A Case Report

Abstract

Faecal parasitological examination of three red kangaroos (Osphranter rufus) newly arrived at Sofia Zoo, Bulgaria, revealed infestation with nematode eggs. The animals were successfully treated with albendazole and ivermectin. Adult nematodes obtained after deworming the kangaroos were identified as gastric strongylids of the species Rugopharynx australis (Mönnig, 1926). The initially collected faecal samples were used to follow the development of the parasites, in particular, the viability of the larvae hatching from the eggs. Morphometric and molecular data as well as some epidemiological observations of the species in the present materials are provided. The study provides the first morphological description of the free-living, infective third-stage larvae of the species. It is also the first record of R. australis in captive-bred kangaroos on the European continent.

1. Introduction

Red kangaroos (Osphranter rufus) are marsupials native to Australia. They are resistant to extreme environmental conditions, especially drought, and are widespread in the inland areas of the continent [1]. Osphranter rufus is among the adaptable species that benefit from grazing activities in Australia, including improved pastures, artificial water points, and the removal of top predators, which poses an ecological challenge to their management [2]. Although ways are being sought to reduce the number of red kangaroos in natural conditions [3], the main aim regarding captive animals, who often live in zoos around the world, is to ensure maximum welfare. However, in the presence of disease, this is challenging. Parasitoses can have a profound impact on the health and productivity of captive animals, as captive husbandry practises have the potential to drastically alter pre-existing host–parasite relationships [4]. Thus, the balance between host and parasite that normally exists in free-ranging populations can be disrupted and lead to clinically visible diseases.

The study of diseases in captive animals is directly related to the possibilities of their successful treatment. On the other hand, comprehensive observations of the specific relationships between wild animals, humans, and domestic animals in zoological facilities and the resulting conditions for the spread of pathogens are of great importance, especially with regard to zoonotic pathogens. The monitoring of pathogens in captive animals is not only important for ecological studies, but also for the protection of public health as part of a One Health approach. Therefore, routine pathogen screening is recommended to optimise targeted treatments and indirect prophylaxis in all kinds of zoological facilities [5]. Against this background, the aim of the present work was to describe a case of infection with Rugopharynx australis (Mönnig, 1926) in captive red kangaroos living far from their natural habitat and to provide morphological and molecular data for the species based on the available material.

2. Materials and Methods

2.1. Sample Origin and Coprological Study

In April 2024, three red kangaroos newly arrived at Sofia Zoo, Bulgaria, from a Czech zoo were parasitologically examined. Two pooled faecal samples were subjected to coproscopic analysis. The first was taken from the floor of the transport container on the day of the animals’ arrival, the other from the bottom of the enclosure in which they were housed on the third day after their arrival. Parts of each faecal sample were examined for the presence of parasite forms using faecal flotation, faecal sedimentation, and the Vaida method within 24–48 h after collection as described in the veterinary manual by Kamburov et al. [6]. A sodium chloride solution with a specific gravity of 1.180 was used as the flotation fluid. The Vaida method was performed as follows: 2–3 faecal balls were placed on a microscope slide and covered with a few drops of warm (45 °C) water. After 30 min, the faecal balls were removed and the liquid was examined under the microscope for the presence of larvae.

The remaining parts of the faecal samples (RFSs) were kept for further observations. They were placed in glass laboratory boxes and initially stored at room temperature (20–25 °C) for 3 months. Thereafter, the RFSs were stored in a refrigerator at a temperature of 4 °C, where they remained for more than 7 months. During storage, the RFSs were moistened with tap water at irregular intervals. The stored faecal samples were examined for the presence of nematode larvae at irregular intervals using the Vaida method [6]. Before the samples stored in the refrigerator were analysed, they were removed from the refrigerator, moistened with tap water, and stored at room temperature for 24 h. During the entire storage period, there were periods when the RFSs were completely dry.

2.2. Animals Deworming and Nematodes Collection

After the animals were diagnosed with a nematode infection, they were treated simultaneously with two medications: Albendazole (Albenol—100 Oral^®, 100 mg albendazole in 1 mL)—1 mL per animal, PO, with a syringe directly into the mouth as a single dose; and ivermectin (Intermectin 100^®, at 10 mg/mL)—0.7 mL per animal as a single dose, SC. Dead adult nematode parasites were detected in the faeces of the kangaroos on the first and second day after deworming. The nematodes were collected in a saline solution, allowed to stand for 30–60 min, and then stored in 70% ethanol for further analysis. To determine the effect of deworming, fresh faecal samples were collected 12 days after treatment and examined using the coproovoscopic method of Fueleborn [6].

2.3. Morphological and Morphometric Study

A total of 12 adult male and 14 adult female specimens were cleared in lactophenol. Each of the cleared worms was placed in a drop of lactophenol between a microscope slide and a coverslip and subjected to morphometric processing. The morphological structures of the helminths were measured after being photographed with a Motic Images Plus 3.0 camera (Motic, Xiamen, China) connected to an Amplival microscope (Carl Zeiss AG, Oberkochen, Germany) and then analysed with the associated software. The morphometric data were processed using Excel (Microsoft, Redmond, WA, USA). The morphometry of eggs and larvae, 10 of each, was performed using the same computer software. Helminths were identified based on their host species and morphological characteristics [7,8]. Adult specimens, preserved in 70% ethanol or embedded in glycerol gelatin as microscopic permanent preparations, were deposited at the Institute of Experimental Morphology, Pathology, and Anthropology with Museum of the Bulgarian Academy of Sciences in Sofia, Bulgaria. The original photographs of eggs and larvae were also preserved.

2.4. DNA Extraction and PCR Amplification

The middle body parts of two male and two female specimens subjected to morphometric studies were cut off prior to processing and stored in 70% ethanol. Before starting the molecular analysis, the body parts of the nematodes were removed from the vials with alcohol and the DNA was extracted individually according to the protocol. Genomic DNA was extracted from each specimen using a NucleoSpin Tissue DNA extraction kit (Macherey-Nagel, Düren, Germany) according to the manufacturer’s instructions. The partial region of the small subunit ribosomal RNA gene (SSU), with complete regions of the internal transcribed spacer 1 (ITS 1), 5.8 ribosomal RNA, and the internal transcribed spacer 2 (ITS 2) along with the partial region of the large subunit ribosomal RNA gene (LSU) were amplified by PCR using two oligonucleotide primers: forward- NC16 (5′-AGT TCAATC GCA ATG GCT T-3′) and reverse- NC2 (5′-TTA GTT TCT TTT CCT CCG CT-3′) [9]. Polymerase chain reactions were conducted in a 50 μL reaction mixture containing 20 μL of molecular biology reagent water (Sigma-Aldrich, St. Louis, MO, USA), 25 μL of AccuStart II PCR ToughMix (×2 concentration) (Quantabio, Beverly, MA, USA), 1 μL of GelTrack Loading Dye (×50 concentration) (Quantabio), 1 μL of each primer (20 mM), and 2 μL of template DNA. Negative control with molecular biology reagent water was used. The amplification conditions were modified as follows: initial denaturation at 94 °C for 2 min followed by 35 cycles of denaturation at 94 °C for 40 s, annealing at 55 °C for 40 s, and primers extension at 72 °C for 40 s; and a final extension at 72 °C for 5 min. The PCR-positive products were purified using the NucleoSpin Gel and PCR Clean-up kit (Macherey-Nagel), and sequenced by Genomed S.A. (Warsaw, Poland). The sequences were then assembled into contigs using CodonCode Aligner version 8.0 (CodonCode Corporation, Centerville, MA, USA). The obtained nucleotide sequences were compared to the NCBI database of sequences using the basic local alignment search tool (BLAST) (http://www.ncbi.nlm.nih.gov/BLAST/) (accessed on 3 March 2025).

2.5. Phylogenetic Analysis

The phylogenetic analysis was performed based on the described sequenced DNA fragment using the newly generated sequences and matching sequences available in the NCBI database (

Table 1. Nucleotide sequences included in the phylogenetic analysis.

Species	GenBank no.	Species	GenBank no.
Rugopharynx australis	PQ526355	R. zeta	KJ776397
	PQ526356		KJ776398
	PQ526357		KJ776399
	PQ526358		KJ776401
	LN906947		KJ776402
	LN906948		LN906992
	LN906949		LN906993
	LN906950		LN906994
	LN906951	R. rho	LN906975
	LN906952		LN906976
	LN906953		LN906977
R. sigma	LN906985		LN906978
	LN906986		LN906979
	LN906987	R. alpha	LN906946
R. setonicus	LN906985

). The sequences were aligned using ClustalW software, version 2.0 [10] and the alignment was trimmed to the length of the shortest sequence (788 bp). Maximum likelihood (ML) phylogenies were inferred with Mega 6 [11] using the best-fit nucleotide substitution model (i.e., General Time Reversible, GTR+I+G) as indicated by jModelTest 2.1.4 [12]. Branch support was estimated using nonparametric bootstrap analyses based on 1000 replicates.

3. Results

3.1. Coprological Findings

Examination of the faecal samples within 24–48 h after collection by the flotation technique showed the presence of large nematode eggs containing motile larvae, and on the 8th day by the method of Vaida—the presence of free actively motile nematode larvae. Examination of fresh control faecal samples collected on day 12 after deworming of the animals showed no parasitic forms.

Examination of RFSs stored at room temperature (20–25 °C) revealed actively motile nematode larvae on the 8th, 16th, 30th, 45th, and 60th day, and 3rd month after faeces collection. Actively motile nematode larvae were also detected in the RFSs cooled to 4 °C at the end of the 1st, 2nd, 4th, 6th, and 7th months after storage in the refrigerator, a total of 10 months and 5 days after the initial collection of the samples.

3.2. Morphometric Characterisation

Eggs in faeces (24–48 h after faecal collection) have the following characteristics: elliptical shape and blunt ends; are 139–169 (151.7 ± 9.60) µm long; 71–82 (78.1 ± 3.60) µm wide; contain a developed enbrio; and are covered with a double-layered shell. Larvae in the eggs are large, transversely striated, and some have a distinguishable shell (Figure 1).

Third-stage larvae (8–30 days after faecal collection) have the following characteristics: fusiform, covered with two sheaths—the outer one—translucent and barely perceptible and the inner one—transversely grooved and clearly visible; length of the larvae with sheath is 642–465 (732 ± 88.33) µm, without the sheath is 415–706 (597.2 ± 91.50) µm; maximum width of 20–29 (26.5 ± 2.95) µm; oesophagus strongyliform, 110–183 (144.9 ± 26.96) µm long and 11–14 (13 ± 1) µm wide; intestine with clearly differentiated intestinal cells, 32–34 in number; excretory pore opens 116–139 (127.9 ± 7.61) µm from the anterior end, and genital primordium located 217–381 (313.5 ± 54.98) µm from the anterior end (Figure 2).

Adult nematodes have the following characteristics: they are spindle shaped; greyish-whitish in colour with a slightly transversely striated cuticle; several well-defined papillae around the mouth opening; pharynx cylindrical, longer than wide, with chitinised walls with transverse stripes (Figure 3a). Oesophagus elongate, with a slightly pronounced widening around the middle and with a bulbous widening at the posterior end; the excretory pore is located slightly posterior to the middle of the oesophagus (Figure 3b). The male specimens have a body length of 0.7–1.2 cm, a body width at the end of the oesophagus of 254–370 μm; the pharynx measures 39−50 × 24−31 μm; the oesophagus measures 830−1143 × 75−138 μm; and the excretory pore is 617–797 μm from the anterior end. Copulatory bursa are well developed, with one dorsal lobe and two symmetrical left and right lobes. The ventral rays of the bursa are of equal length and are strongly fused together; the lateral rays are also strongly fused together, giving the impression of a single whole; the anterolateral rays are shorter than the medio- and posterolateral ones (Figure 4a); the dorsal ray is divided into two massive branches, from each of which a lateral branch separates at a right angle (Figure 4b). Numerous small papillae are observed on the surface of the ventral and lateral lobes of the bursa (Figure 4a). Spicules are equal in size, 1702–2400 μm long, tubular, with short comb-like wings extending from their proximal ends (Figure 5). At the distal parts of the spicules, the wings are expanded and fused, forming an oval membrane connecting the spicule tips (Figure 4). Gubernaculum is difficult to recognise, elongated, and is 32–39 μm long (Figure 4). A heart-shaped chitinised structure is clearly visible medially to the spicules, near their distal ends (Figure 4). The genital cone is triangular in shape and clearly visible ventrally (Figure 4). The female specimens have a body length of 0.8–1.3 cm and their body width at the end of the oesophagus is 264–363 μm; the pharynx measures 41−53 × 24−33 μm; the oesophagus measures 1084−1268 × 111−155 μm; the excretory pore is 681–1003 μm from the anterior end. The vagina is modified into an ovijector with two sphincters leading into the two uterine branches (Figure 5). The vulva is 539–715 μm from the posterior end. The tail pointed is 410–677 μm long, and the anus opens in the front of a small bulge of the cuticle, at 436–560 μm from the tail tip (Figure 5). The morphometry of the male and female adult nematodes corresponded to that of R. australis.

3.3. Nucleotide Seuences

Four nucleotide sequences containing partial SSU, complete ITS1, complete 5.8 S rRNA, complete ITS2, and partial LSU were obtained for each of the worms (GenBank: PQ526355, PQ526356, PQ526357, PQ526358). The sequences varied in length from 804 to 821 base pairs (bp) and were 99.27% to 100% homologous to each other. Our isolates were compared with those of the most similar species of the genus Rugopharynx (

Table 2. Nucleotide sequence similarity between our sequences of Rugopharynx australis (GenBank: PQ526355-PQ526358) and the most similar species of the genus Rugopharynx according to the Basic Local Alignment Search Tool (BLAST).

Species	GenBank no.	Host	Country	Similarity (%)
Rugopharynx australis	LN906947	Macropus rufus	Australia	98.94–99.60
	LN906948	Macropus rufus	Australia	94.29–94.55
	LN906949	Macropus giganteus	Australia	99.47
	LN906950	Macropus giganteus	Australia	98.94–99.07
	LN906951	Macropus dorsalis	Australia	98.94–99.60
	LN906952	Macropus robustus	Australia	98.94–99.07
	LN906953	Macropus fuliginosus	Australia	94.03–94.29
Rugopharynx zeta	KJ776397	Not provided	Australia	94.88–95.14
	KJ776398	Not provided	Australia	95.01–95.26
	KJ776399	Not provided	Australia	95.01–95.26
	KJ776401	Not provided	Australia	95.49–95.75
	KJ776402	Not provided	Australia	94.41–95.63
	LN906992	Petrogale assimilis	Australia	93.80–93.93
	LN906993	Petrogale inornata	Australia	94.46–94.59
	LN906994	Petrogaleherberti	Australia	94.06–94.20
Rugopharynx rho	LN906975	Macropus eugenii	Australia	94.87–95.13
	LN906976	Macropus eugenii	Australia	94.87–95.13
	LN906977	Macropus fuliginosus	Australia	95.92–96.05
	LN906978	Macropus fuliginosus	Australia	92.26–95.13
	LN906979	Macropus irma	Australia	95.65–95.78
Rugopharynx sigma	LN906985	Thylogale thetis	Australia	92.63–92.76
	LN906986	Thylogale stigmatica	Australia	94.73–94.99
	LN906987	Thylogale stigmatica	Australia	94.60–94.86
Rugopharynx setonicis	LN906984	Setonix brachyurus	Australia	95.52–95.65

). They showed the highest nucleotide sequence homology to R. australis with a percentage identity between 94.03% and 99.60%. The degree of sequence similarity between our isolates and those of R. zeta, R. rho, R. setonicis, and R. sigma ranged from 92.26% to 96.05% (Table 2). These results confirmed the morphological identification of the species as R. australis.

Phylogenetic analysis of the partial SSU, complete ITS1, complete 5.8 S rRNA, complete ITS2, and partial LSU sequence data with R. alpha as the outgroup revealed six strongly supported clades (Figure 6). One clade contained R. australis, the second clade contained the taxa R. rho and R. setonicis, the third clade contained a single isolate of R. sigma, the fifth clade contained R. zeta, the sixth clade contained two isolates of R. sigma, and the seventh clade contained R. alpha. The first clade contains two subclades containing R. australis taxa. The sequences obtained in this study co-created the first subclade of R. australis taxa.

4. Discussion

Rugopharynx australis (Mönnig, 1926) (Rhabditida: Strongylidae) is a gastric nematode of marsupial mammals of the family Macropodidae. It is one of the most common and widespread nematode parasites of kangaroos and dominates the gastric helminth community of the red kangaroo [8,9]. It has been found in free-ranging kangaroo populations in all mainland states of Australia [8,13] as well as in a variety of wild macropodid hosts and has been recorded under synonymous names such as Pharyngostrongylus alpha, P. beta, and P. brevis [14]. The nematode has also been recorded in free-living Bennett’s wallabies (Macropus rufogriseus rufogriseus) in New Zealand [15] and in semi-free-living yellow-footed rock wallabies (Petrogale xanthopus) from the Itozu no mori Zoological Park (Kitakyushu, Fukuoka) in Japan [16].

The first description of R. australis was by Mönnig [17], who described it in materials from a red kangaroo that had died in a zoo in Pretoria, South Africa, shortly after its importation. Almost a century later, Beveridge et al. [8] redescribed the species by analysing a large quantity of nematodes collected from seven macropod species (Osphranter rufus, Macropus giganteus, Osphranter robustus, Macropus fuliginosus, Notamacropus dorsalis, Lagorchestes conspicillatus, and Petrogale xanthopus) from across Australia.

The comparison of the morphometric data for R. australis from the red kangaroo material (

Table 3. Measurements of Rugopharynx australis in materials from red kangaroos (Osphranter rufus) of different origins.

	Parameter	Origin/Reference
		Osphranter rufus, Sofia Zoo, Bulgaria [Present Data]	Osphranter rufus, Australia Beveridge et al. [8]	Osphranter rufus, Mönnig [17]
		Range (Mean ± SD)
Adult male nematodes (n = 12)	Body length (mm)	7–12 (9 ± 0.14)	7.3–10.5 (9.4)	9.2–10
	Body width at the end of the oesophagus (µm)	254–370 (282 ± 32.47)	290–330 (310) *	380 *
	Length of pharynx (µm)	39–50 (45 ± 3.45)	40–45 (44)	-
	Width of pharynx (µm)	24–31 (27 ± 2.19)	25–30 (28)	-
	Oesophagus length (µm)	830–1143 (1007 ± 84.87)	940–1080 (1010)	-
	Maximum esophagus width (µm)	75–138 (110 ± 16.57)	-	-
	Excretory pore—ant. end	617–797 (699 ± 57.59)	500–770 (620)	-
	Spicule length (µm)	1702–2400 (1953 ± 229.88)	1750–2050 (1860)	2020
	Gubernaculum length (µm)	32–39 (35 ± 2.45)	30 (30)	-
				-
Adult female nematodes (n = 14)	Body length (mm)	8–13 (11 ± 0.12)	7.6–9.4 (8.7)	12.5
	Body width at the end of the oesophagus (µm)	264–363 (332 ± 34.11)	360–420 (390) *	500 *
	Length of pharynx (µm)	41–53 (47 ± 4.57)	40–50 (47)	76
	Width of pharynx (µm)	24–33 (29 ± 2.67)	30–35 (33)	-
	Oesophagus length	1084–1268 (1168 ± 61.90)	980–1180 (1070)	1170
	Maximum esophagus width (µm)	111–155 (129 ± 12.01)	-	130
	Excretory pore—ant. end	681–1003 (784 ± 118.58)	600–710 (660)	840
	Ovijector/vagina total length (µm)	572–805 (672 ± 85.66)	650–800 (740)	520
	Vulva-tail tip distance (µm)	539–715 (640 ± 53.46)	550–760 (650)	-
	Anus-tail tip distance (µm)	436–560 (513 ± 43.06)	-	-
	Tail length (µm)	410–677 (553 ± 85.90)	400–600 (490)	560

n—number; * maximum body width.

) shows that the metrics of the nematodes from the Bulgarian population are generally higher and their diapason is larger than that of the Australian populations. The observation of these population differences is consistent with the statement that R. australis is a morphologically polymorphic species [8,14], and this finding was confirmed by the present results. Despite extensive studies on the morphology, taxonomy, and phylogeny of the species [8,9,14], to the best of the authors’ knowledge, there is no description of its life cycle and larval forms in the available literature. Drawing an analogy with the life cycle of another strongyloid nematode—Labiostrongylus eugenii (Johnston and Mawson, 1940) which infests macropods in Australia [18], we believe that the larval forms observed and described here are free-living, infective third-stage larvae of R. australis. Our observations showed that infective R. australis larvae are extremely resistant to different and changing environmental conditions. They maintained their vitality and activity for 95 days at 20–25 °C and a further 135 days at 4 °C, during which the kangaroos’ faeces were repeatedly desiccated. During the last 2 months of their stay in the refrigerator, for example, the RFSs were not moistened at all. This explains the wide distribution of the species among macropods in Australia [8], despite the specific and changing climate and the degradation of many ecosystems on the continent [19]. Evidently, parasites have evolutionary survival adaptations that allow the species to spread outside its natural range, as is the present case with the nematode population that develops in kangaroos born and raised in captivity on the European continent.

5. Conclusions

The study confirms that the parasitic nematode R. australis is a morphologically polymorphic species, and molecular analyses support its identification. The infective third-stage larvae of the parasite are extremely resistant to different and changing environmental conditions. Rugopharynx australis has the potential to successfully spread to captive kangaroo populations outside Australia. This should be considered in order to protect the health of animals kept in zoos around the world.