Morphological Re-Description and 18 S rDNA Sequence Confirmation of the Pinworm Aspiculuris tetraptera (Nematoda, Heteroxynematidae) Infecting the Laboratory Mice Mus musculus

Publications

Share / Export Citation / Email / Print / Text size:

Journal of Nematology

Society of Nematologists

Subject: Life Sciences

GET ALERTS DONATE

ISSN: 0022-300X
eISSN: 2640-396X

DESCRIPTION

349
Reader(s)
880
Visit(s)
0
Comment(s)
0
Share(s)

SEARCH WITHIN CONTENT

FIND ARTICLE

Volume / Issue / page

Related articles

VOLUME 50 , ISSUE 2 (September 2018) > List of articles

Morphological Re-Description and 18 S rDNA Sequence Confirmation of the Pinworm Aspiculuris tetraptera (Nematoda, Heteroxynematidae) Infecting the Laboratory Mice Mus musculus

Rewaida Abdel-Gaber * / Fathy Abdel-Ghaffar / Saleh Al Quraishy / Kareem Morsy / Rehab Saleh / Heinz Mehlhorn

Keywords : Laboratory mice, Pinworms, Aspiculuris species, Morphological description, Molecular study

Citation Information : Journal of Nematology. Volume 50, Issue 2, Pages 117-132, DOI: https://doi.org/10.21307/jofnem-2018-026

License : (PUBLISHER)

Published Online: 03-September-2018

ARTICLE

ABSTRACT

Aspiculuris tetraptera is a heteroxynematid nematoda infecting most of the laboratory animals, occasionally mice which represent the mostly used animal for biological, medical, and pharmacological studies. The present study aimed to investigate the prevalence of nematode parasites infection in the laboratory mice Mus musculus in Egypt. Morphologically, this oxyurid possessed four distinct cephalic papillae on a cephalic plate, with three small rudimental lips carrying two sessile poorly developed labial papillae and one pair of amphidial pores. Esophagus divided into cylindrical corpus and globular bulb. Distinct cervical alae interrupted at the level of esophago–intestinal junction forming an acute angle. At the caudal end, twelve caudal papillae in male worms while an ovijector apparatus opening and a vulva surrounded by protruded lips in females were observed. The general morphological criteria include this nematode with other Aspiculuris species which were compared in the present study. Molecular characterization based on 18SSU rDNA sequencing performed to confirm the taxonomic position of this species and to documents the morphological data. Sequence alignment detects a percent of identity up to 88.0% with other Heteroxynematidae species. Phylogenetic analysis showed that the present recorded is a putative sister taxon to A. tetraptera recorded in a previous study. The SSU rDNA sequence has been deposited in the GenBank under the accession no. MG019400.

Graphical ABSTRACT

Pinworms are routinely found in animals from modern animal facilities, even in facilities free of viral and bacterial diseases that affect mice (Jacoby and Lindsey, 1998; Zenner and Regnault, 2000; Behnke et al., 2015). Oxyurids are also a common parasite of Muroidea (Rodentia) (Singleton et al., 1993; Pisanu et al., 2001). Syphacia obvelata and Aspiculuris tetraptera are oxyurid nematodes which are cosmopolitan monoxenous parasites that are transmitted through the ingestion of embryonated eggs (Stojcevic et al., 2004; Robles and Navone, 2010; Khalil et al., 2014). Mice may be concurrently infected with both species of pinworms (Jacobson and Reed, 1974; Taffs, 1976; Nicklas et al., 1984; Gonçalves et al., 1998; Zenner, 1998; Agersborg et al., 2001). The common incidence of infection by both pinworm species can be explained by their preference for slightly different sites of the gastrointestinal tract. As such, these species do not compete directly for resources; they are able to maintain simultaneous infections (Pinto et al., 2001; Bazzano et al., 2002). In concurrent infections, there may be higher numbers of A. tetraptera worms because their longer lifespan permits the accumulation of parasites in their hosts (Scott and Gibbs, 1986). The prevalence of pinworms in an infected rodent population depends on many factors, including gender, age, strain, immune status, and the concentration of parasite ova in the environment.

The genus Aspiculuris was established by Nitzsch (1821) and later re-described by Schulz (1924) from Mus musculus. Many species of Aspiculuris have been reported worldwide (Hugot, 1980; Inglis et al., 1990; Falcón-Ordaz et al., 2010). Species of Aspiculuris were separated by Quentin (1975) into two groups based on the shape of their cervical alae. Nematodes that display interrupted cervical alae with pointed posterior ends belong to the first group. Those in the second group possess a rounded posterior end to the cervical alae. Aspicularis tetraptera are common oxyurids belonged to the first group. This species has been described in the cecum and colon of M. musculus in different regions, such as Tunisia, Iran, Venezuela, Europe, Siberia, China, Japan, Unite States, and Egypt (Hugot, 1980; Neifer et al., 1991; Durden et al., 2000; Perec-Matysiak et al., 2006; Mahmoud et al., 2009; Abdel-Gaber, 2016), and to a lesser extent, it has been recorded in other hosts, such as Cricetus, Rattus, Apodemus, Microtus, Arctomys, Jaculus, Clethrionomys, and Peromyscus in the same regions (Mathies, 1959; Sasa et al., 1962). Quentin (1975) indicated this species in Central Africa in Mastomys, Praomys, and Thamnomys.

Species of A. tetraptera are characterized by their medium or small size, the presence of three lips, absence of buccal capsule, and presence of esophagus with a well-developed single bulb located at its posterior end (Bazzano et al., 2002; Perec-Matysiak et al., 2006; Malsawmtluangi and Tandon, 2009, Li et al., 2016). Recently, morphological identification requires the use of molecular characteristics for accurate identification and validation; these characteristics are common in nematoda systematics (Mc-Manus and Bowles, 1996; Semenova et al., 1996; Gasser, 2001; Jones et al., 2012; Chaudhary et al., 2016; Curtis et al., 2017).

Therefore, the present study reported the natural prevalence, morphological, and morphometric characteristics, in addition to molecular analysis of ribosomal DNA gene sequences of the recovered oxyurid pinworm infecting the laboratory mouse M. musculus to clarify the taxonomic status and phylogenetic position of this parasite species within Heteroxynematidae.

Materials and Methods

Animal collection and parasitological examination

Fifty specimens of adult laboratory mice (Muridae: M. musculus) reared at the Animal House at Zoology Department, Faculty of Science, Cairo University, Cairo, Egypt; were randomly collected between December 2016 and September 2017. The collected mice were transported alive to the Laboratory of Parasitology Research for parasitological examination. Mice were anesthetized and killed according to the ethical rules for handling experimental animals. Mice were examined for any external signs of infection. After dissection, internal organs were removed from the rodent and examined for any parasitic infections. Isolated worms were fixed in 70% ethanol and subsequently clarified with lactophenol for morphological identification, in accordance with standard reference keys by Pinto et al. (2001). Prevalence of parasitic infection (number of infected mice/total number of mice hosts examined ×100) of M. musculus was calculated according to Bush et al. (1997). Illustrations of adult specimens were prepared with the aid of a microscope Leica DM 2500, LAS software (3.8) and Corel Draw X4® software. Measurements were based on 20 adult worm species; data were taken in millimeters and are presented as a range followed by the arithmetic mean ± SD in parentheses.

Molecular analysis

DNA extraction, polymerase chain reaction amplification, and sequencing

gDNA was extracted from ethanol-preserved samples using DNeasy tissue kit© (Qiagen, Hilden, Germany) following the manufacturer’s instructions. The DNA was stored in 50 μl of TE buffer at −20°C until further use. DNA concentration and purity were determined spectrophotometrically by measuring absorbance at wavelengths of 260 and 280 nm. PCR amplification was performed in a final volume of 25 µl, containing 3 μl of genomic DNA, 2.5 μl of 10X Taq polymerase buffer, 10 pmol of each primer, 100 μM of each dNTP (Finnzymes Products), and 1.5 U of Taq DNA polymerase (Finnzymes Products). The partial ribosomal 18S gene was amplified using the primer Nem 18SF (5′-CGC GAA TRG CTC ATT ACA ACA GC-3′) and Nem 18SR (5′-GGG CGG TAT CTG ATC GCC-3′) designed by Floyd et al. (2005). Polymerase chain reaction (PCR) consisted of an initial denaturation step at 94°C for 3 min, followed by 35 cycles of 1 min at 94°C, 1 min at 50°C, followed by 1 min at 72°C, and finally, post-PCR extension was carried out for 7 min at 72°C. All PCR products were verified on 1% agarose gel in ×1 Tris–acetate–EDTA (TAE) stained with 1% ethidium bromide visualized with UV transilluminator, and bands with predicted size were purified using Pure LinkTM Quick Gel Extraction Kit (Invitrogen) following the manufacturer’s instructions. Amplicons were sequenced (in both directions) using the BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems Thermo Fisher Scientific, Waltham, MA, USA) with the 310 Automated DNA Sequencer (Applied Biosystems, MA, USA) using the same primers for annealing.

Sequence alignment and phylogenetic analysis

BLAST search was carried out to identify related sequences on NCBI database. Sequences were aligned directly using CLUSTAL-X multiple sequence alignment (Thompson et al., 1997) and compared with previously recorded data from GenBankTM to analyze intra-specific differences. GenBank accession numbers of additional sequences utilized in the analyses were as follows: Cosmocerca japonica (LC052782), Parascaris equorum (JN617987), Enterobius vermicularis (HQ646164), Cucullanus extraneus (KT192060), Pseudanisakis rajae (JN392470), Skrjabinema kamosika (AB699691), Aspiculuris dinniki (KT175736), Aspiculuris tianjinensis (KT175733), and A. tetraptera (KT175725, KT175728, KT175729, KJ143615, KJ143618, KJ143617, KJ143616, EF464551, and EU263107) as shown in Table (1). The alignment was corrected manually using the alignment editor of software BIOEDIT 4.8.9 (Hall 1999). Phylogenetic calculations were performed with PAUP 4.0b10 (Swofford, 2000). The data were analyzed with maximum parsimony (neighbor-interchange [CNI] level 3, random addition trees 100). Additionally, neighbor-joining was calculated by using MEGALIGN package (DNASTAR, Windows version 3.12e).

Table 1

Nematoda species used in the phylogenetic analysis of Aspiculuris tetraptera in the present study.

10.21307_jofnem-2018-026-t001.jpg

Results

A total of 28 out of 50 (56.0%) specimens of laboratory mice M. musculus were found to be naturally infected with an oxyurid nematoda. The recovered parasite species were found in the cecum and upper colon of the infected host mice.

Description

In general, the body of the recovered worms was small, cylindrical in shape, and covered by a transversely striated cuticle. Head was bulb-like; mouth opening was surrounded by three less developed lips, one pair of lateral epaulettes, one pair of amphidial pores, and two pairs of large sub-median cephalic papillae. Mouth opening leads to the buccal cavity, followed by pharynx, esophagus, and long intestine opening exteriorly by an anal opening in females and cloacal opening in males. Anterior part of esophagus was club-shaped followed by well-developed bulb. Body in both sexes has distinct cervical alae, beginning immediately posterior to the anterior end of the cephalic vesicle. Cervical alae abruptly interrupted at the level of esophago–intestinal junction, forming an acute angle. Anterior end of the body has prominent, elaborate inflated region, forming cephalic vesicle (Fig. 1A–G, Tables 2,3).

Table 2

Main morphological features and measurements of male Aspiculuris tetraptera compared with previous studies.

10.21307_jofnem-2018-026-t002.jpg
Table 3

Main morphological features and measurements of female Aspiculuris tetraptera compared with previous studies.

10.21307_jofnem-2018-026-t003.jpg

Male worm (based on 10 mature specimens)

Body length was 2.23–3.29 (2.79 ± 0.1) mm with maximum width 0.16–0.20 (0.18 ± 0.1) mm. Cephalic vesicle was 0.06–0.09 (0.07 ± 0.001) mm long by 0.05–0.08 (0.06 ± 0.001) mm wide. Esophagus measured 0.32–0.40 (0.39 ± 0.1) mm long by 0.05–0.09 (0.07 ± 0.01) mm wide; while, the whole esophagus with bulb reached 0.13–0.17 (0.15 ± 0.1) mm long by 0.04–0.07 (0.05 ± 0.01) mm wide. Cervical alae began at 0.015–0.018 (0.017 ± 0.001) mm from the anterior end and measured about 0.21–0.29 (0.25 ± 0.001) mm long with recurved terminal ends by 0.029–0.038 (0.031 ± 0.001) mm wide. Nerve ring and excretory pore are located at 0.065–0.082 (0.078 ± 0.001) mm and 0.392–0.547 (0.491 ± 0.03) mm from the anterior end, respectively. Narrow lateral alae of the body end located at the beginning of the caudal alae are extended from the level of cloaca and surrounded the entire end of the body, bending ventrally at its tip as a vesicular swelling of the cuticle. Cloaca opening is located at 0.09–0.11 (0.10 ± 0.01) from the posterior extremity of the body. Testes are flexed over the anterior third of the intestine. Gubernaculum and spicules were absent. Posterior end with 12 caudal papillae included one pair precloacal, two pairs adcloacal, one pair postcloacal, two median papillae postcloacal, one behind the other, and a further posterior pair midway between cloaca and end of the tail. Tail, with blunt end, measured 0.11–0.14 (0.12 ± 0.1) mm long.

Fig. 1

A–G, Line drawings of different body parts of Aspiculuris tetraptera. A, Lateral view of female worm with mouth opening surrounded by three lips with cephalic papillae (CP) and amphids (AM), muscular esophagus (E), esophageal bulb region (EOB), intestine (IN), rectum (R) with rectal gland (RG), anal opening (AN) and ending with a long tapered tail (T). Note, transverse annulated (TA) cuticle, and the genital system characterized with a uterus filled with numerous eggs (EG), ovijector apparatus (OA), vagina (VA) and vulval opening (VU) surrounded by two fleshy vulval lips (VL). B, Lateral view of male worm with mouth opening surrounded by three lips with cephalic papillae (CP) and amphids (AM), followed by muscular esophagus (E), esophageal bulb region (EOB), intestine (IN), rectum (R) with rectal gland (RG), anal opening (AN), and ending with a long tapered tail (T). Note, transverse annulated (TA) cuticle, and the genital system with testes (TE), cloacal opening (CO) surrounded by precloacal papillae (PCP), adcloacal papillae (ACP), postadcloacal papillae (PACP), median postadcloacal papillae (MPACP), and posterior papillae (PP). C–G, High magnifications of: C, Face view of anterior extremity of female worm mouth opening (MO) surrounded by three lips (L) with cephalic papillae (CP) and amphids (AM) with cervical alae (CA). D, Face view of anterior extremity of male worm mouth opening (MO) surrounded by three lips (L) with cephalic papillae (CP) and amphids (AM) with cervical alae (CA). E, Ovejector region (OA) of female showing vulva opening (VU), two fleshy vulval lips (VL), muscular vagina (VA), and eggs collected (EG) from uterus. F, Posterior end of male worm showing the cloacal opening (CO) with caudal papillae of precloacal papillae (PCP), adcloacal papillae (ACP), postadcloacal papillae (PACP), median postadcloacal papillae (MPACP), and posterior papillae (PP). G, Morula (M) surrounded by egg shell (ES).

10.21307_jofnem-2018-026-f001.jpg

Female worm is larger than that of the male (based on 10 mature specimens)

Body length was 2.9–3.4 (3.1 ± 0.1) mm long with maximum width was 0.19–0.23 (0.20 ± 0.01) mm. Cephalic vesicle reached about 0.078–0.083 (0.082 ± 0.001) mm long by 0.106–0.130 (0.123 ± 0.01) mm wide. Esophagus measured 0.30–0.34 (0.32 ± 0.01) mm long and 0.14–0.16 (0.15 ± 0.01) mm wide; while, esophagus with bulb reached about 0.10–0.13 (0.11 ± 0.01) mm long and 0.05–0.09 (0.07 ± 0.01) mm wide. Nerve ring and excretory pore located at 0.078–0.090 (0.085 ± 0.002) mm and 0.564–0.780 (0.680 ± 0.02) mm from the anterior end, respectively. Cervical alae with recurved terminal end was 0.27–0.29 (0.26 ± 0.01) mm long. Distance from the anterior end to the beginning of cervical alae was 0.021–0.026 (0.024 ± 0.001) mm. Vulva was preequatorial, surrounded by protruded lips, and situated at 1.112–1.630 (1.406 ± 0.03) mm from the anterior extremity of the body. Ovejector apparatus measured about 0.29–0.38 (0.32 ± 0.01) mm long. Muscular vagina proceeded forward for a short distance then turned backward joining uterus filled with eggs. Two ovaries flexed over the proximal part of the intestine. Anal pore located at 0.32–0.39 (0.37 ± 0.01) mm from the posterior end of the body. Tail with blunt tip measured 0.30–0.42 (0.39 ± 0.01) mm long. Eggs were unoperculated, smooth, filled by morula and measured 0.04–0.06 (0.05 ± 0.01) mm long and 0.02–0.04 (0.03 ± 0.01) mm wide.

Taxonomic summary

Parasite name: Aspiculuris tetraptera (Nitzsch, 1821; Family: Heteroxynematidae (Skrjabin and Schikhobalova, 1948)).

Host: Laboratory mice M. musculus (Linnaeus, 1758; Family: Muridae).

Mode of transmission: Ingestion of embryonated eggs in feces, or in contaminated food and water, or bedding.

Morbidity and mortality: Infected laboratory mice were generally symptomless externally.

Site of infection: Cecum and upper colon of infected host mice.

Prevalence and intensity: 28 out of 50 (56.0%) examined individuals were infected, with a total number of 120 nematodes.

Material deposition: Voucher specimens were deposited at museum in Zoology Department, Faculty of Science, Cairo University, Cairo, Egypt.

Molecular analysis

A sequence of 840 bp was deposited in GenBank under accession no. MG019400 with a GC content of 42.26%, for SSU rDNA gene sequences of the present oxyurid species. Pairwise comparison of the isolated gDNA sequence of the present parasite species with a range of other Spirurina species and genotypes revealed a unique sequence. The calculated identity between this novel sequence and those retrieved from GenBank demonstrated a high degree of similarity, up to 88.0% (Table 1). Comparison of the nucleotide sequences and divergence showed that SSU rDNA of the present oxyruid species had the highest blast scores with a small number of nucleotide differences with other A. tetraptera species under the following accession numbers: EU263107, EF464551, KJ143616, KJ143617, KJ143618, KJ143615, KT175729, KT175728, KT175725, and KT175725; A. dinniki (acc. no. KT175736); A. tianjinensis (acc. no. KT175733); E. vermicularis (acc. no. HQ646164); and S. kamosika (acc. no. AB699691).

Phylogenetic analysis led to the construction of a neighbor-joining tree, constructed with partial sequences, which showed that Spirurina species consistently formed two major clades (Fig. 2). The first one represented the most related families in order Oxyurida, including families Heteroxynematidae and Oxyuridae with sequence similarity ranging between 99.0 and 91.0%. The second one was represented by four families Ascarididiae (P. equorum JN617987), Anisakidae (P. rajae JN392470), Cucullanida (C. extraneus KT192060), and Cosmocercidae (C. japonica LC052782), belonging to the order Ascaridia with sequence similarity ranging between 90.0 and 89.0%. This sequence in conjunction with existing data, suggested the placement of this oxyurid species within family Heteroxynematidae. The present species was deeply embedded in the genus Aspiculuris, and is closely related to other Aspiculuris species, especially to other previously described A. tetraptera as a putative sister taxon.

Fig. 2

Phylogenetic tree generated by neighbor-joining analyses of the partial SSU rDNA sequence of Aspiculuris tetraptera and oxyurid species with the strongest BLAST matches and in part with some ascarid species. GenBank accession numbers are given after the species names. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Oxyurid parasite examined in the present study is bolded.

10.21307_jofnem-2018-026-f002.jpg

Discussion

Laboratory animal models, especially rodents of the family Muridae, constitute important links in the food chains within the ecosystems they inhabit (Rosas, 1997, Gonçalves et al., 1998). These animals are often in contact with humans and domestic animals and can transmit various parasitic species (Bazzano et al., 2002). In conventional animal facilities, rodent colonies are frequently infected with helminth parasites, or become infected during the experimental period (Sato et al., 1995, Rehbinder et al., 1996). Oxyurids are cosmopolitan nematoda parasites of public health importance (Khalil et al., 2014). The order Oxyurida includes three families namely Oxyuridae (Cobbold, 1864); Pharyngodonidae (Travassos, 1919); and Heteroxynematidae (Skrjabin and Schikhobalova, 1948). Nematodes from the genera Syphacia and Aspiculuris are common parasitic pinworms of rodents all over the world (Adamson, 1994; Robles and Navone, 2007; Millazzo et al., 2010; Sotillo et al., 2012; Verma et al., 2013; Weyand et al., 2016; Zarei et al., 2016; Stewart et al., 2017).

Based on morphological characters, the oxyurid species described here showed the characteristic features of the genus Aspiculuris, including the presence of four distinct cephalic papillae lying on the cephalic plate, and three small rudimental lips that carry two sessile poorly developed labial papillae. According to the present results, A. tetraptera naturally infect the laboratory mice M. musculus, which is consistent with data reported by Hugot (1988), Mohd Zain et al. (2012) Pakdel et al. (2013), and Panti-May et al. (2015) who stated that species of the genus Aspiculuris belonging to the family Heteroxynematidae are characterized as parasites of Muroidea. Based on the stated characters herein, the present species was identified as A. tetraptera and recorded in 56.0% of the examined specimens representing a higher prevalence. This is in agreement with data obtained by Bluszcz et al. (1987), Bazzano et al. (2002), Izdebska and Rolbiecki (2006), Klimpel et al. (2007), Kataranovski et al. (2008), and Baird et al. (2012), who reported that mice are naturally infected with different oxyurid species, with 9.0 to 75.0% infection range.

The present parasite species was compared morphologically and morphometrically with other Aspiculuris species as shown in Tables 2 and 3, and exhibited strong similarities to those reported in other studies by Yamaguti (1935), Hugot (1980), Pinto et al. (1994), Landaeta-Aqueveque et al. (2007), Khalil et al. (2014), and Abdel-Gaber and Fol (2015). Only a few differences in measurements of the different body parts were observed. However, this species differs from other Aspiculuris species in the structure of the cephalic region, length of esophagus, position of nerve ring, excretory pore and vulva opening, number and arrangement of cloacal papillae in males, and size of eggs in females. The presently described parasite species resembled A. tetraptera and Aspiculuris huascaensis by having cervical alae ending at the mid-length of the esophageal bulb with the same number of caudal papillae. However, it can be distinguished from A. huascaensis by having a single sessile pre-cloacal papilla located between two cuticular folds and slightly anteriorly to the cloaca, whereas A. tetraptera lack both the sessile pre-cloacal papilla and two cuticular folds. In addition, males of A. tetraptera have a double pedunculate papilla immediately posterior to the cloaca, and the anterior most papilla located between the caudal folds of the tail is double, while, those of A. huascaensis lack a doubled medial papilla associated with the cloaca and a simple anterior most papillae was recorded. These data were consistent with a previous report by Falcón-Ordaz et al. (2010). In agreement with Ashour (1980), Pinto et al. (1994), and Abdel-Gaber and Fol (2015), 12 papillae were present in A. tetraptera; conversely, Schulz (1924) reported 10 and Yamaguti (1935) and Falcón-Ordaz et al. (2010) reported 14 papillae. In addition, the division of the dorsal and both subventral lips of A. tetraptera males are unique. Chitwood and Chitwood (1950) noted a similar division of the dorsal lip only of A. ackerti with sexual dimorphism of this character in male worms and no tendency for separation in the female specimens.

Aspiculuris is one of five subgenera listed by Akhtar (1955) in which the cephalic bulb and lateral alae are present, the cervical alae end in a sickle shaped margins, but the cervical and lateral alae are not continuous, as stated by Petter and Quentin (2009). The same results were obtained in the present study on the recovered worms, which have well-developed cervical alae extending into cephalic vesicle, and poorly marked cuticular striations. Furthermore, the present described parasite species were similar to other species of Aspiculuris, such as Aspiculuris dinnicki (Schulz, 1924); Aspiculuris schulzi (Popov and Nasarova, 1930); Aspiculuris azerbaidjanica (Tarzhimanova, 1969); Aspiculuris arianica (Erhadová-Kotrlá and Daniel, 1970); and Aspiculuris witenbergi (Quentin, 1975); with cervical alae that are abruptly interrupted with the pointed posterior ends and forming an acute angle toward the anterior. However, it differs from Aspiculuris kazakstanica (Nasarova and Sweschikova, 1930); Aspiculuris americana (Erickson, 1938); Aspiculuris lahorica (Akhtar, 1955); Aspiculuris pakistanica (Akhtar, 1955); Aspiculuris africana (Quentin, 1966); Aspiculuris tschertkowi (Tarzhimanova, 1969); Aspiculuris rysavyi (Kotrla and Daniel, 1970); and Aspiculuris versterae (Hugot, 1980), since the posterior end of the cervical alae of those species does not form an acute angle, and the caudal alae of males is not close to the tip. In addition, with the exception of A. tschertkowi, which has 16 caudal papillae, the remaining species have a smaller number of papillae than A. huascaensis, varying from 4 to 11 versus 12 papillae.

Due to close morphological similarities, molecular phylogenetic approaches have been used extensively in association with traditional morphological techniques as reliable methods for confirmation of accurate identification, and differentiation between pinworms infecting laboratory rodents (Jacobs et al., 1997; Zhu et al., 1998; Vermund and Wilson, 2000; Morales-Hojas et al., 2001; Nakano et al., 2006; Li et al., 2007; Zhu et al., 2007; Chang et al., 2009). In the present study, a nuclear rDNA region of the recovered parasite species was amplified using the species-specific primers Nem 18SF/Nem 18SR, designed by Floyd et al. (2005). It is apparent that, the phylogenetic tree based on nuclear SSU rDNA sequences estimated in this study supported strongly the higher taxonomic groups of both orders: Oxyurida (representing the two main families Oxyuridae and Heteroxynematidae) and Ascaridia (representing four families, namely Cosmocercidae, Cucullanidae, Anisakidae, and Ascarididiae). These results are in agreement with data obtained by Blaxter et al. (1998) who reported that clade III of the full dataset of the nematoda phylogeny was represented by all members of the suborder spirurina and clustered into four classical orders of Ascaridia, Oxyurida, Rhigonematida, and Spirurida. Anderson (2000) proposed that Ascaridia and Spirurida were sister groups, which in turn, were more closely related to Strongylida than a group consisting of Oxyurida plus Rhigonematida. Subsequent analyses of SSU rDNA sequences strongly supported the monophyly of clade III taxa with bootstrap values for the clade exceeding 95.0% (De Ley and Blaxter, 2002; Bert et al., 2006; Holterman et al., 2006; Wijová et al., 2006; Qiu et al., 2016; Ribas et al., 2017).

Khalil et al. (2014) reported that the order Oxyurida incorporates three main families, Oxyuridae (Cobbold, 1864); Pharyngodonidae (Travassos, 1919); and Heteroxynematidae (Skrjabin and Schikhobalova, 1948); which was consistent with the results of the current study. In addition, Petter and Quentin (2009) included Syphacia obvelata and Syphacia muris of the genus Syphacia with 22 genera in the family Oxyuridae, and included the genus Aspiculuris together with seven further genera in the subfamily Heteroxynematinae belonging to the family Heteroxynematidae. This was consistent with the present findings indicating that Oxyuridae species, represented by the genus Aspiculuris, is monophyletic in origin, supporting the taxonomic position of the present Aspiculuris species, which is deeply embedded in the genus Aspiculuris with a close relationship with other described species of A. tetraptera as a more related sister taxon.

Conclusion

Recent field studies have provided useful tools for the rapid identification and phylogenetic analysis of pinworms infecting laboratory rodents. The 18S rDNA gene of A. tetraptera yielded a unique sequence that confirms the taxonomic position within the family Heteroxynematidae.

Compliance with ethical standards

All procedures contributing to this work comply with the ethical standards of the relevant national guides on the care and use of laboratory animals and have been approved and authorized by Institutional Animal Care and Use Committee (IACUC) in Faculty of Science, Cairo University, Egypt (No. CU/I/S/19/16).

Conflict of interest

The authors have declared that they have no conflict of interest regarding the content of this article.

Acknowledgements

The authors are thankful to Faculty of Science, Cairo University, Cairo, Egypt, and extend their appreciation to The Deanship of Scientific Research at King Saud University for funding this work through research group no (RG-002).

References


  1. Abdel-Gaber, R.. 2016. Syphacia obvelata (Nematoda, Oxyuridae) infecting laboratory mice Mus musculus (Rodentia, muridae): Phylogeny and host-parasite relationship. Parasitology Research 115 3: 975-985.
    [CROSSREF]
  2. Abdel-Gaber, R., and Fol, M.. 2015. Aspicularis tetrapetra (Nematoda, Heteroxynematidae) of laboratory mice Mus musculus (Rodentia, Muridae): A potential risk of zoonotic infection for researchers. Ciencia e Tecnica Vitivinicola 30 8: 125-136.
  3. Adamson, M.L.. 1994. Evolutionary factors influencing the nature of parasite specificity. Parasitology 109: S85-S95.
    [CROSSREF]
  4. Agersborg, S.S., Garza, K.M., and Tung, K.S.. 2001. Intestinal parasitism terminates self-tolerance and enhances neonatal induction of auto-immune disease and memory. European Journal of Immunology 31: 851-859.
    [CROSSREF]
  5. Akhtar, S.A.. 1955. On nematoda parasites of rats and mice of Lahore, with some remarks on the genus Aspiculuris Schulz, 1924 and two news species of the genus. Pakistan Journal of Scientific Research 7: 104-111.
  6. Anderson, R.C.. 2000. Nematoda parasites of vertebrates: Their development and transmission, 2nd ed.., CABI Publishing, New York.
    [CROSSREF]
  7. Araujo, P.. 1965. Aspiculuris artigasi n. sp. (Nematoda: Oxyuridae) em Mus musculus. Memórias do Instituto de Butantan 32 1: 101-108.
  8. Ashour, A.A.. 1980. Ultrastructural and other studies on intestinal nematodes of small mammals from Egypt. Thesis, Ain Shams University, Cairo, Egypt.
  9. Baird, S.J.E., Ribas, A., Macholan, M., Albrecht, T., Pialek, J., and Gouyde Bellocq, J.. 2012. Where are the wormy mice? A re-examination of hybrid parasitism in the European house mouse hybrid zone. Evolution 66 9: 2757-2772.
    [CROSSREF]
  10. Bazzano, T., Restel, T.I., Pinto, R.M., and Gomes, D.C.. 2002. Patterns of infection with nematodes Syphacia obvelata and Aspicularis tetraptera in conventionally maintained laboratory mice. Memórias do Instituto Oswaldo Cruz 97: 847-853.
    [CROSSREF]
  11. Behnke, J.M., Barnard, C.J., Bajer, A., Bray, D., Dinmore, J., and Frake, K.. 2001. Variation in the helminth community structure in bank voles (Cletrionomys glareolus) from three comparable localities in the Mazury Lake District region of Poland. Parasitology 123: 401-414.
    [CROSSREF]
  12. Behnke, J.M., Stewart, A., Bajer, A., Grzybek, M., Harris, P.D., Lowe, A., Ribas, A., Smales, L., and Vandegrift, K.J.. 2015. Bank voles (Myodes glareolus) and house mice (Mus musculus musculus; M. m. domesticus) in Europe are each parasitized by their own distinct species of Aspiculuris (Nematoda, Oxyurida). Parasitology 142 12: 1493-1505.
    [CROSSREF]
  13. Bert, W., Messiaen, M., Manhout, J., Houthoofd, W., and Borgonie, G.. 2006. Evolutionary loss of parasitism by nematodes? Discovery of a free-living filaroid nematoda. The Journal of Parasitology 92: 645-647.
    [CROSSREF]
  14. Blaxter, M.L., De Ley, P., Garey, J.R., Liu, L.X., Scheldeman, P., and Vierstraete, A.. 1998. A molecular evolutionary framework for the phylum Nematoda. Nature 392: 71-75.
    [CROSSREF]
  15. Bluszcz, A., Blaski, M., Sabesta, R., and Szilman, P.. 1987. Helmintofauna drobnych gryzoni (Rodentia) kilku miejscowosci okrêgu katowickiego [Helminth fauna of small rodents (Rodentia) of several localities from the environment of Katowice]. Acta Biologica Szegediensis 6: 127-129.
  16. Bush, A.O., Lafferty, K.D., Lotz, J., and Shostak, A.W.. 1997. Parasitology meets ecology on its own terms: Margolis et al. revised. Journal of Parasitology 83 4: 575-583.
    [CROSSREF]
  17. Chang, T.K., Liao, C.W., and Haung, Y.C.. 2009. Prevalence of Enterobius vermicularis infection among preschool children in kinder gardens of Taipei City, Taiwan in 2008. Korean Journal of Parasitology 47: 185-187.
    [CROSSREF]
  18. Chaudhary, A., Goswami, U., and Singh, H.S.. 2016. Molecular characterization of Nippostrongylus brasiliensis (Nematoda: Heligmosomatidae) from Mus musculus in India. Korean Journal of Parasitology 54 6: 743-750.
    [CROSSREF]
  19. Chitwood, B.G., and Chitwood, M.B.. Eds), 1950. An Introduction to Nematology, Monumental Printing Co, Baltimore, MA.
  20. Cobbold, T.S.. 1864. Entozoa: an introduction to the study of helminthology, more particularly to the internal parasites of man. Quoted from Petter and Quentin (1976), p. 508.
  21. Curtis, R.C., Murray, J.K., Campbell, P., Nagamori, Y., Molnar, A., and Jackson, T.A.. 2017. Interspecies variation in the susceptibility of a wild-derived colony of mice to pinworms (Aspiculuris tetraptera). Journal of the American Association for Laboratory Animal Science 56 1: 42-46.
  22. De Ley, P., and Blaxter, M.. 2002. Systematic position and phylogeny. in Lee, D.L.. Ed.), The Biology of Nematodes, Taylor and Francis, London, pp. 1-30.
  23. Durden, L.A., Hu, R., Oliver, J.H., and Cilek, J.E.. 2000. Rodent ectoparasites from two locations in northwestern Florida. Journal of Vector Ecology 25: 222-228.
    [PUBMED]
  24. Erhadová-Kotrlá, B., and Daniel, M.. 1970. Parasitic worms of small mammals from the mountain regions of the Eastern Hindu Kush. Folia Parasitologica 17: 201-216.
  25. Erickson, A.B.. 1938. Parasites of some Minnesota Cricetidae and Zapodidae and a host catalogue of helminth parasites of native American mice. American Midland Naturalist 20: 575-589.
    [CROSSREF]
  26. Falcón-Ordaz, J., Pulido-Flores, G., and Monks, S.. 2010. New species of Aspiculuris (Nematoda: Heteroxynematidae), parasite of Mus musculus (Rodentia: Muridae), from Hidalgo, Mexico. Revista Mexicana de Biodiversidad 81: 669-676.
    [CROSSREF]
  27. Floyd, R.M., Rogers, A.D., Lambshead, J.D., and Smith, C.R.. 2005. Nematoda-specific PCR primers for the 18S small subunit rRNA gene. Molecular Ecology Notes 5: 611-612.
    [CROSSREF]
  28. Gasser, R.B.. 2001. Identification of parasitic nematodes and study of genetic variability using PCR approaches. in Kennedy, M.W., and Harnett, W.. Eds), Parasitic Nematodes: Molecular Biology, Biochemistry and Immunology, CABI Publishing, London, pp. 53-82.
  29. Gonçalves, L., Pinto, R.M., Vicente, J.J., Noronha, D., and Gomes, D.C.. 1998. Helminth parasites of conventionally maintained laboratory mice – II. Inbred strains with an adaptation of the anal swab technique. Memorias Do Instituto Oswaldo Cruz 93: 121-126.
    [CROSSREF]
  30. Hall, T.A.. 1999. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series 41: 95-98.
  31. Holterman, M., Van Der Wurff, A., Van Den Elsen, S., Van Megen, H., Bongers, T., and Holovachov, O.. 2006. Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown clades. Molecular Biology and Evolution 23: 1792-1800.
    [CROSSREF]
  32. Hugot, J.P.. 1980. Sur le genre Aspiculuris Schulz, 1924 (Nematoda, Heteroxynematidae), oxyures parasites de Rongeurs Muroidea. Bulletin du Museum National d´Histoire Naturelle 2: 723-735.
  33. Hugot, J.P.. 1988. Les nematodes Syphaciinae parasites de Rongeurs et de Lagomorphes. Taxonomie. Zoogeographie_Evolution. Memoires du Museum national d’Histoire Naturelle, Paris, Serie A, Zoologie 141: 1-153.
  34. Inglis, W.G., Harris, E.A., and Lewis, J.W.. 1990. A new species of the nematoda genus Aspiculuris Schulz, 1924 from Aethomys namaquensis (Mammalia: Rodentia) in the Kruger National Park, South Africa. Systematic Parasitology 17: 231-236.
    [CROSSREF]
  35. Izdebska, J.N., and Rolbiecki, L.. 2006. Correlation between the occurrence of mites (Demodex spp.) and nematodes in house mice (Mus musculus Linnaeus, 1758) in the Gdañsk urban agglomeration. Biology Letters 43 2: 175-178.
  36. Jacobs, D.E., Zhu, X.Q., Gasser, R.B., and Chilton, N.B.. 1997. PCR based methods for the identification of potentially zoonotic ascaridoid parasites of the dog, fox and cat. Acta Tropica 68: 191-200.
    [CROSSREF]
  37. Jacobson, R.H., and Reed, N.D.. 1974. The thymus dependency of resistance to pinworm infection in mice. Journal of Parasitology 60: 976-979.
    [CROSSREF]
  38. Jacoby, R.O., and Lindsey, J.R.. 1998. Health care for research animals is essential and affordable. FASEB Journal 11: 609-614.
    [CROSSREF]
  39. Jones, R., Brown, D.S., Harris, E., Jones, J., Symondson, W.O.C., and Bruford, M.W.. 2012. First record of Neoxysomatium brevicaudatum through the non-native invasive sampling of Anguis fragilis: Complementary morphological and molecular detection. Journal of Helminthology 86: 125-129.
    [CROSSREF]
  40. Kataranovski, D., Vukićević-Radić, O.D., Kataranovski, M., Radović, D.L., and Mirkov, I.I.. 2008. Helminth fauna of Mus musculus Linnaeus 1758 from the suburban area of Belgrade, Serbia. Archives of Biological Sciences 60 4: 609-617.
    [CROSSREF]
  41. Khalil, A.I., Lashein, G.H., Morsy, G.H., and Abd El-Mottaleb, D.I.. 2014. Oxyurids of wild and laboratory rodents from Egypt. Life Science Journal 11 3: 94-107.
  42. Klimpel, S., Förster, M., and Günter, S.. 2007. Parasite fauna of the bank vole Chletrionomys glareolus in an urban region of Germany: Reservoir of zoonotic metazoan parasites?. Parasitology Research 102: 69-75.
    [CROSSREF]
  43. Kohn, A., and Macedo, B.. 1984. First record of Aspiculuris tetraptera (Nitzsch, 1821) (Nematoda: Oxyuroidea) and Dollfusentis chandleri (Golvan, 1969) (Acanthocephala: Illiosentidae) in Haemulon sciurus (Shaw 1803) (Pisces: Pomadasyidae). Annales De Parasitologie Humaine Et Comparee 59 5: 477-482.
    [CROSSREF]
  44. Kruidenier, J.J., and Mehra, K.N.. 1959. Aspiculuris ackerti n. sp. (Nematoda: Oxyuridae) from the wood rats of Arizona. Proceedings of the Helminthological Society of Washington 26 2: 147-150.
  45. Landaeta-Aqueveque, C.A., Robles, M.D.R., and Cattan, P.E.. 2007. The community of gastrointestinal helminths in the house mouse, Mus musculus, in Santiago, Chile. Parasitología latinoamericana 62: 165-169.
    [CROSSREF]
  46. Li, C.L., Du, X.Y., Gao, J., Wang, C., Guo, H.G., Dai, F.W., Sa, X.Y., An, W., and Chen, Z.W.. 2016. Phylogenetic analysis of the Mongolian gerbil (Meriones unguiculatus) from China based on mitochondrial genome. Genetics and Molecular Research 15 3: 1-13.
  47. Li, M.W., Lin, R.Q., Chen, H.H., Sani, R.A., Song, R.A., and Song, H.Q.. 2007. PCR tools for the verification of the specific identity of ascaridoid nematodes from dogs and cats. Molecular and Cellular Probes 21: 349-354.
    [CROSSREF]
  48. Linnaeus, C.. 1758. Systema naturae per regna tria naturae, secundum classes, ordines, genera, species, cum characteribus, differentiis, synonymis, locis. Tomus I. Editio decima, reformata. Impensis Direct. Laurentii Salvii, Holmiae.
  49. Liu, B., Bu, Y., and Zhang, L.. 2012. A new species of Aspiculuris Schulz, 1924 (Nematoda, Heteroxynematidae) from the gray-sided vole, Clethrionomys rufocanus (Rodentia, Cricetidae), from Tianjin, China. Acta Parasitologica 57: 311-315.
    [CROSSREF]
  50. MacArthur, J.A., and Wood, M.. 1978. Control of oxyurids in mice using thiabendazole. Laboratory Animals 12: 141-143.
    [CROSSREF]
  51. Mahmoud, A.E., Attia, R.A.H., Eldeek, H.E.M., Abdel Baki, L., and Oshaish, H.A.. 2009. Oxyurid nematodes detected by colonoscopy in patients with unexplained abdominal pain. Parasitologists United Journal 2 2: 93-102.
  52. Malsawmtluangi, C., and Tandon, V.. 2009. Helminth parasite spectrum in rodent hosts from bamboo growing areas of Mizoram, north-east India. Journal of Parasitic Diseases 33 1–2: 28-35.
    [CROSSREF]
  53. Mathies, A.W.J.. 1959. Certain aspects of the host-parasite relationship of Aspiculuris tetraptera, a mouse pinworm. I. Host specificity and age resistance. Experimental Parasitology 8: 31-38.
    [CROSSREF]
  54. Mc-Manus, D.P., and Bowles, J.. 1996. Molecular genetic approaches to parasite identification: Their value in diagnostic parasitology and systematics. International Journal of Parasitology 26: 687-704.
    [CROSSREF]
  55. Millazzo, C., Ribasa, A., Casanova, J.C., Cagnin, M., Geraci, F., and Di Bella, C.. 2010. Helminths of the brown rat (Rattus norvegicus) (Berkenhout, 1769) in the city of Palermo, Italy. Helminthology 47 4: 238-240.
    [CROSSREF]
  56. Miller, G.E., and Schmidt, G.D.. 1982. Helminths of bushytailed wood rats, Neotoma cinerea subspp. from Colorado, Idaho, and Wyoming. Proceedings of the Helminthological Society of Washington 49: 109-117.
  57. Mohd Zain, S.N., Behnke, J.M., and Lewis, J.W.. 2012. Helminth communities from two urban rat populations in Kuala Lumpur, Malaysia. Parasit. Vectors 5 p. 47.
    [CROSSREF]
  58. Morales-Hojas, R., Post, R.J., Shelley, A.J., Maia-Herzog, M., Coscaron, S., and Cheke, R.A.. 2001. Characterization of nuclear ribosomal DNA sequences from Onchoncerca volvulus and Mansonella ozzardi (Nematoda: Filaroidea) and development of a PCR-based method for their detection in skin biopsies. International Journal of Parasitology 31: 169-177.
    [CROSSREF]
  59. Nakano, T., Okamoto, M., Ikeda, Y., and Hasegawa, H.. 2006. Mitochondrial cytochrome c oxidase subunit 1 gene and nuclear rDNA regions of Enterobius vermicularis parasitic in captive chimpanzees with special reference to its relationship with pinworms in humans. Parasitology Research 100: 51-57.
    [CROSSREF]
  60. Nasarova, Y.A., and Sweschnikova, N.M.. 1930. Sur la connaissance des vers parasites des Rongeurs du Kazakhstan. Review of Microbiology and Epidemiology of Parasites 9 1: 101-104.
  61. Neifer, S., Kremsner, P.G., Weinig, M., Harms, G., Sahlmüller, G., and Bienzle, U.. 1991. Interferon-gamma treatment in mice experimentally infected with Trichinella spiralis. Parasitology Research 77 5: 437-442.
    [CROSSREF]
  62. Nicklas, W., Le Corre, R., and Graw, J.. 1984. Experiences with fenbendazole in the treatment of oxyuriasis in an experimental animal colony. Berliner Und Munchener Tierarztliche Wochenschrift 97: 21-24.
    [PUBMED]
  63. Nitzsch, C.L.. 1821. Ascaris. Allg. Encycl. d. wissensch. v. Künste (Ersch und Gruber), Leipzig, 6: 44-49.
  64. Pakdel, N., Naem, S., Rezaei, F., and Chalehchaleh, A.A.. 2013. A survey on helminthic infection in mice (Mus musculus) and rats (Rattus norvegicus and Rattus rattus) in Kermanshah, Iran. Veterinary Research Forum 4 2: 105-109.
  65. Panti-May, J.A., Hernández-Betancourt, S.F., Rodríguez-Vivas, R.I., and Robles, M.R.. 2015. Infection levels of intestinal helminths in two commensal rodent species from rural households in Yucatan, Mexico. Journal of Helminthology 89: 42-48.
    [CROSSREF]
  66. Perec-Matysiak, A., Okulewicz, A., Hildebrand, J., and Zalesny, G.. 2006. Helminth parasites of laboratory mice and rats. Wiadomosci Parazytologiczne 52: 99-102.
  67. Petter, A.J., and Quentin, J.C.. 2009. Keys to the genera of the Oxyuroidea. in Anderson, R.C., Chabaud, A.G., and Willmott, S.. Eds), CIH keys to the nematoda parasites of vertebrates, Commonwealth Agricultural Bureaux, England, pp. 1-30.
  68. Pinto, R.M., Goncalves, L., Noronha, D., and Gomes, D.C.. 2001. Worm burdens in outbred and inbred laboratory rats with morphometric data on Syphacia muris (Yamaguti, 1935) Yamaguti, 1941 (Nematoda: Oxyuroidea). Memórias do Instituto Oswaldo Cruz 96: 133-136.
    [CROSSREF]
  69. Pinto, R.M., Vicente, J.J., Noronha, D., Gonçalves, L., and Gomes, D.C.. 1994. Helminth parasites of conventionally maintained laboratory mice. Memórias do Instituto Oswaldo Cruz 89: 33-40.
    [CROSSREF]
  70. Pisanu, B., Chapuis, J.L., and Durette-Desset, M.C.. 2001. Helminths from introduced small mammals on Kerguelen, Crozet, and Amsterdam Islands. Journal of Parasitology 85 5: 1205-1208.
    [CROSSREF]
  71. Popov, N., and Nazarova, J.. 1930. Neue art Parasitischer wurmer (Oxyuridae). Vestnik Mikrob. Epid. Parazitol. Saratow 9 1: 105-108.
  72. Qiu, J.H., Lou, Y., Zhang, Y., Chang, Q.C., Liu, Z.X., Duan, H., Guo, D.H., Gao, D.Z., Yue, D.M., and Wang, C.R.. 2016. Sequence variability in internal transcribed spacers of nuclear ribosomal DNA among isolates of the oxyurid nematodes Syphacia obvelata and Aspiculuris tetraptera from mice reared in laboratories in China. Journal of Helminthololy 90 1: 81-85.
    [CROSSREF]
  73. Quentin, J.C.. 1966. Oxyures de Muridae africains. Annales De Parasitologie Humaine Et Comparee 41 5: 443-452.
    [CROSSREF]
  74. Quentin, J.C.. 1975. Essai de classification des oxyures Heteroxynematidae. Mémoires du Muséum National Histoire Naturelle, Zoologie 94: 51-96.
  75. Rehbinder, C., Baneux, P., Forbes, D., Van Herck, H., Niclas, W., and Rugaya, Z.Y.. 1996. FELASA – Recommendations for the health monitoring of mouse, rat, hamster, gerbil, guinea pig and rabbit experimental units. Laboratory Animal Science 30: 193-208.
    [CROSSREF]
  76. Ribas, A., Diagne, C., Tatard, C., Diallo, M., Poonlaphdecha, S., and Brouat, C.. 2017. Whipworm diversity in West African rodents: A molecular approach and the description of Trichuris duplantieri n. sp. (Nematoda: Trichuridae). Parasitology Research 116 4: 1265-1271.
    [CROSSREF]
  77. Robles, M.R., and Navone, G.T.. 2007. A new species of Syphacia (Nematoda: Oxyuridae) from Oligoryzomys nigripes (Rodentia: Cricetidae) in Argentina. Parasitology Research 101 4: 1069-1075.
    [CROSSREF]
  78. Robles, M.R., and Navone, G.T.. 2010. Redescription of Syphacia venteli Travassos 1937 (Nematoda: Oxyuridae) from Nectomys squamipes in Argentina and Brazil and description of a new Syphacia from Melanomys caliginosus in Colombia. Parasitology Research 106 5: 1117-1126.
    [CROSSREF]
  79. Rosas, G.A.. 1997. Diagnóstico: Parasitosis intestinal por Aspiculuris tetraptera. Animals Experimentation 2: 9-11.
  80. Sasa, M., Tanaka, H., Fukui, M., and Takata, A.. 1962. Internal parasites of laboratory animals. in Harris, R.J.C.. Ed.), The Problems of Laboratory Animal Disease, Academic Press, London; New York, pp. 195-214.
  81. Sato, Y., Ooi, H.K., Nonaka, N., Oku, Y., and Kamiya, M.. 1995. Antibody production in Syphacia obvelata infected mice. Journal of Parasitology 8: 559-562.
    [CROSSREF]
  82. Schulz, R.E.S.. 1924. Oxyuridae of Armenian mice. Reportes Tropical Institute Armenia 2: 41-51.
  83. Scott, M.E., and Gibbs, H.C.. 1986. Long-term population dynamics of pinworms (Syphacia obvelata and Aspiculuris tetraptera) in mice. Journal of Parasitology 72: 652-662.
    [CROSSREF]
  84. Semenova, S.K., Romanova, E.A., and Pyskov, A.P.. 1996. Genetic differentiation of helminthes on the basis of data of polymerase chain reaction using random primers. Genetics 32 2: 304-309.
  85. Singleton, G.R., Smith, A.L., Shellam, G.R., Fitzgerald, N., and Muller, W.J.. 1993. Prevalence of viral antibodies and helminthes in field populations of house mice (Mus domesticus) in southeastern Australia. Epidemiology and Infection 110: 399-417.
    [CROSSREF]
  86. Skrjabin, K.I., and Schikhobalova, N.D.. 1948. Quoted from Petter and Quentin (1976).
  87. Sotillo, J., Trelis, M., Cortés, A., Luz Valero, M., Sánchezdel Pino, M., and Esteban, J.G.. 2012. Proteomic analysis of the pinworm Syphacia muris (Nematoda: Oxyuridae), a parasite of laboratory rats. Parasitology International 61 4: 561-564.
    [CROSSREF]
  88. Stewart, A., Lowe, A., Smales, L., Bajer, A., Bradley, J., Dwuznik, D., Franssen, F., Griffith, J., Stuart, P., Turner, C., Zalesny, G., and Behnke, J.M.. 2017. Parasitic nematodes of the genus Syphacia Seurat, 1916 infecting Muridae in the British Isles, and the peculiar case of Syphacia frederici. Parasitology 23: 1-12.
  89. Stojcevic, D., Mihljevic, Z., and Marnculic, A.. 2004. Parasitological survey of rats in rural regions of Croatia. Veterinary Medicine – Czech 49 3: 70-74.
    [CROSSREF]
  90. Swofford, D.L.. 2000. PAUP*. Phylogenetic Analysis using Parsimony (*and other Methods), Version 4, Sinauer Associates, Sunderland, MA.
  91. Taffs, L.F.. 1976. Pinworm infection in laboratory rodents: a review. Laboratory Animals 10: 1-13.
    [CROSSREF]
  92. Tarzhimanova, R.A.. 1969. New nematodes of the genus Aspiculuris from Rodents. Azerbaidzhanskogo Nauchno. Issledovatel’skogo Instituta Meditsinkoi Parasitologii Tropicheskoi Meditsiny im. S.M. Kirova 7: 302-306.
  93. Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., and Higgins, D.G.. 1997. The CLUSTAL-X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Research 25: 4876-4882.
    [CROSSREF]
  94. Travassos, L.. 1919. Quoted from Petter and Quentin (2009).
  95. Verma, S., Gaherwal, S., Prakash, M.M., and Kanhere, R.R.. 2013. Anthelmintic efficacy of Ocimum sanctum against Syphacia muris in mice. Acta Parasitologica 4 1: 24-28.
  96. Vermund, S.H., and Wilson, C.M.. 2000. Pinworm (Enterobius vermicularis). Seminars in Pediatric Infectious Diseases 11: 252-256.
    [CROSSREF]
  97. Weyand, N.J., Ma, M., Phifer-Rixey, M., Taku, N.A., Rendon, M.A., Hockenberry, A.M., Kim, W.J., Agellon, A.B., Biais, N., Suzuki, T.A., Goodyer-Sait, L., Harrison, O.B., Bratcher, H.B., Nachman, M.W., Maiden, M.C., and So, M.. 2016. Isolation and characterization of Neisseria musculi sp. nov., from the wild house mouse. International Journal of Systematic and Evolutionary Microbiology 66 9: 3585-3593.
    [CROSSREF]
  98. Wijová, M., Moravec, F., Horák, A., and Lukeš, J.. 2006. Evolutionary relationships of Spirurina (Nematoda: Chromadorea: Rhabditida) with special emphasis on dracunculoid nematodes inferred from SSU rRNA gene sequences. International Journal for Parasitology 36: 1067-1075.
    [CROSSREF]
  99. Yamaguti, S.. 1935. Studies on the helminth fauna of Japan. Part 13. Mammalian nematodes. Japanese Journal of Zoology 6: 433-457.
  100. Zarei, Z., Mohebali, M., Heidari, Z., Davoodi, J., Shabestari, A., Motevalli Haghi, A., Khanaliha, K., and Kia, E.B.. 2016. Helminth Infections of Meriones persicus (Persian Jird), Mus musculus (House Mice) and Cricetulus migratorius (Grey Hamster): A cross-sectional study in Meshkin-Shahr District, Northwest Iran. Iranian Journal of Parasitology 11 2: 213-220.
    [PUBMED]
  101. Zenner, L.. 1998. Effective eradication of pinworms (Syphacia muris, Syphacia obvelata and Aspiculuris tetraptera) from a rodent breeding colony by oral anthelmintic therapy. Laboratory Animals 32: 337-342.
    [CROSSREF]
  102. Zenner, L., and Regnault, J.P.. 2000. Ten-year long monitoring of laboratory mouse and rat colonies in French facilities: A retrospective study. Laboratory Animals 34: 76-83.
    [CROSSREF]
  103. Zhu, X.Q., Amelio, S.D., Gasser, R.B., Yang, T.B., Paggi, L., and He, F.. 2007. Practical PCR tools for the delineation of Contracaecum rudolphii A and Contracaecum rudolphii B (Ascaridoidea: Anisakidae) using genetic markers in the nuclear ribosomal DNA. Molecular Cellular Probes 21: 97-102.
    [CROSSREF]
  104. Zhu, X.Q., Jacobs, D.E., Chilton, N.B., Sani, R.A., Cheng, N., and Gasser, R.B.. 1998. Molecular characterization of Toxocara variant from cats in Kuala Lumpur, Malaysia. Parasitology 117: 155-164.
    [CROSSREF]
XML PDF Share

FIGURES & TABLES

Fig. 1

A–G, Line drawings of different body parts of Aspiculuris tetraptera. A, Lateral view of female worm with mouth opening surrounded by three lips with cephalic papillae (CP) and amphids (AM), muscular esophagus (E), esophageal bulb region (EOB), intestine (IN), rectum (R) with rectal gland (RG), anal opening (AN) and ending with a long tapered tail (T). Note, transverse annulated (TA) cuticle, and the genital system characterized with a uterus filled with numerous eggs (EG), ovijector apparatus (OA), vagina (VA) and vulval opening (VU) surrounded by two fleshy vulval lips (VL). B, Lateral view of male worm with mouth opening surrounded by three lips with cephalic papillae (CP) and amphids (AM), followed by muscular esophagus (E), esophageal bulb region (EOB), intestine (IN), rectum (R) with rectal gland (RG), anal opening (AN), and ending with a long tapered tail (T). Note, transverse annulated (TA) cuticle, and the genital system with testes (TE), cloacal opening (CO) surrounded by precloacal papillae (PCP), adcloacal papillae (ACP), postadcloacal papillae (PACP), median postadcloacal papillae (MPACP), and posterior papillae (PP). C–G, High magnifications of: C, Face view of anterior extremity of female worm mouth opening (MO) surrounded by three lips (L) with cephalic papillae (CP) and amphids (AM) with cervical alae (CA). D, Face view of anterior extremity of male worm mouth opening (MO) surrounded by three lips (L) with cephalic papillae (CP) and amphids (AM) with cervical alae (CA). E, Ovejector region (OA) of female showing vulva opening (VU), two fleshy vulval lips (VL), muscular vagina (VA), and eggs collected (EG) from uterus. F, Posterior end of male worm showing the cloacal opening (CO) with caudal papillae of precloacal papillae (PCP), adcloacal papillae (ACP), postadcloacal papillae (PACP), median postadcloacal papillae (MPACP), and posterior papillae (PP). G, Morula (M) surrounded by egg shell (ES).

Full Size   |   Slide (.pptx)

Fig. 2

Phylogenetic tree generated by neighbor-joining analyses of the partial SSU rDNA sequence of Aspiculuris tetraptera and oxyurid species with the strongest BLAST matches and in part with some ascarid species. GenBank accession numbers are given after the species names. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Oxyurid parasite examined in the present study is bolded.

Full Size   |   Slide (.pptx)

REFERENCES

  1. Abdel-Gaber, R.. 2016. Syphacia obvelata (Nematoda, Oxyuridae) infecting laboratory mice Mus musculus (Rodentia, muridae): Phylogeny and host-parasite relationship. Parasitology Research 115 3: 975-985.
    [CROSSREF]
  2. Abdel-Gaber, R., and Fol, M.. 2015. Aspicularis tetrapetra (Nematoda, Heteroxynematidae) of laboratory mice Mus musculus (Rodentia, Muridae): A potential risk of zoonotic infection for researchers. Ciencia e Tecnica Vitivinicola 30 8: 125-136.
  3. Adamson, M.L.. 1994. Evolutionary factors influencing the nature of parasite specificity. Parasitology 109: S85-S95.
    [CROSSREF]
  4. Agersborg, S.S., Garza, K.M., and Tung, K.S.. 2001. Intestinal parasitism terminates self-tolerance and enhances neonatal induction of auto-immune disease and memory. European Journal of Immunology 31: 851-859.
    [CROSSREF]
  5. Akhtar, S.A.. 1955. On nematoda parasites of rats and mice of Lahore, with some remarks on the genus Aspiculuris Schulz, 1924 and two news species of the genus. Pakistan Journal of Scientific Research 7: 104-111.
  6. Anderson, R.C.. 2000. Nematoda parasites of vertebrates: Their development and transmission, 2nd ed.., CABI Publishing, New York.
    [CROSSREF]
  7. Araujo, P.. 1965. Aspiculuris artigasi n. sp. (Nematoda: Oxyuridae) em Mus musculus. Memórias do Instituto de Butantan 32 1: 101-108.
  8. Ashour, A.A.. 1980. Ultrastructural and other studies on intestinal nematodes of small mammals from Egypt. Thesis, Ain Shams University, Cairo, Egypt.
  9. Baird, S.J.E., Ribas, A., Macholan, M., Albrecht, T., Pialek, J., and Gouyde Bellocq, J.. 2012. Where are the wormy mice? A re-examination of hybrid parasitism in the European house mouse hybrid zone. Evolution 66 9: 2757-2772.
    [CROSSREF]
  10. Bazzano, T., Restel, T.I., Pinto, R.M., and Gomes, D.C.. 2002. Patterns of infection with nematodes Syphacia obvelata and Aspicularis tetraptera in conventionally maintained laboratory mice. Memórias do Instituto Oswaldo Cruz 97: 847-853.
    [CROSSREF]
  11. Behnke, J.M., Barnard, C.J., Bajer, A., Bray, D., Dinmore, J., and Frake, K.. 2001. Variation in the helminth community structure in bank voles (Cletrionomys glareolus) from three comparable localities in the Mazury Lake District region of Poland. Parasitology 123: 401-414.
    [CROSSREF]
  12. Behnke, J.M., Stewart, A., Bajer, A., Grzybek, M., Harris, P.D., Lowe, A., Ribas, A., Smales, L., and Vandegrift, K.J.. 2015. Bank voles (Myodes glareolus) and house mice (Mus musculus musculus; M. m. domesticus) in Europe are each parasitized by their own distinct species of Aspiculuris (Nematoda, Oxyurida). Parasitology 142 12: 1493-1505.
    [CROSSREF]
  13. Bert, W., Messiaen, M., Manhout, J., Houthoofd, W., and Borgonie, G.. 2006. Evolutionary loss of parasitism by nematodes? Discovery of a free-living filaroid nematoda. The Journal of Parasitology 92: 645-647.
    [CROSSREF]
  14. Blaxter, M.L., De Ley, P., Garey, J.R., Liu, L.X., Scheldeman, P., and Vierstraete, A.. 1998. A molecular evolutionary framework for the phylum Nematoda. Nature 392: 71-75.
    [CROSSREF]
  15. Bluszcz, A., Blaski, M., Sabesta, R., and Szilman, P.. 1987. Helmintofauna drobnych gryzoni (Rodentia) kilku miejscowosci okrêgu katowickiego [Helminth fauna of small rodents (Rodentia) of several localities from the environment of Katowice]. Acta Biologica Szegediensis 6: 127-129.
  16. Bush, A.O., Lafferty, K.D., Lotz, J., and Shostak, A.W.. 1997. Parasitology meets ecology on its own terms: Margolis et al. revised. Journal of Parasitology 83 4: 575-583.
    [CROSSREF]
  17. Chang, T.K., Liao, C.W., and Haung, Y.C.. 2009. Prevalence of Enterobius vermicularis infection among preschool children in kinder gardens of Taipei City, Taiwan in 2008. Korean Journal of Parasitology 47: 185-187.
    [CROSSREF]
  18. Chaudhary, A., Goswami, U., and Singh, H.S.. 2016. Molecular characterization of Nippostrongylus brasiliensis (Nematoda: Heligmosomatidae) from Mus musculus in India. Korean Journal of Parasitology 54 6: 743-750.
    [CROSSREF]
  19. Chitwood, B.G., and Chitwood, M.B.. Eds), 1950. An Introduction to Nematology, Monumental Printing Co, Baltimore, MA.
  20. Cobbold, T.S.. 1864. Entozoa: an introduction to the study of helminthology, more particularly to the internal parasites of man. Quoted from Petter and Quentin (1976), p. 508.
  21. Curtis, R.C., Murray, J.K., Campbell, P., Nagamori, Y., Molnar, A., and Jackson, T.A.. 2017. Interspecies variation in the susceptibility of a wild-derived colony of mice to pinworms (Aspiculuris tetraptera). Journal of the American Association for Laboratory Animal Science 56 1: 42-46.
  22. De Ley, P., and Blaxter, M.. 2002. Systematic position and phylogeny. in Lee, D.L.. Ed.), The Biology of Nematodes, Taylor and Francis, London, pp. 1-30.
  23. Durden, L.A., Hu, R., Oliver, J.H., and Cilek, J.E.. 2000. Rodent ectoparasites from two locations in northwestern Florida. Journal of Vector Ecology 25: 222-228.
    [PUBMED]
  24. Erhadová-Kotrlá, B., and Daniel, M.. 1970. Parasitic worms of small mammals from the mountain regions of the Eastern Hindu Kush. Folia Parasitologica 17: 201-216.
  25. Erickson, A.B.. 1938. Parasites of some Minnesota Cricetidae and Zapodidae and a host catalogue of helminth parasites of native American mice. American Midland Naturalist 20: 575-589.
    [CROSSREF]
  26. Falcón-Ordaz, J., Pulido-Flores, G., and Monks, S.. 2010. New species of Aspiculuris (Nematoda: Heteroxynematidae), parasite of Mus musculus (Rodentia: Muridae), from Hidalgo, Mexico. Revista Mexicana de Biodiversidad 81: 669-676.
    [CROSSREF]
  27. Floyd, R.M., Rogers, A.D., Lambshead, J.D., and Smith, C.R.. 2005. Nematoda-specific PCR primers for the 18S small subunit rRNA gene. Molecular Ecology Notes 5: 611-612.
    [CROSSREF]
  28. Gasser, R.B.. 2001. Identification of parasitic nematodes and study of genetic variability using PCR approaches. in Kennedy, M.W., and Harnett, W.. Eds), Parasitic Nematodes: Molecular Biology, Biochemistry and Immunology, CABI Publishing, London, pp. 53-82.
  29. Gonçalves, L., Pinto, R.M., Vicente, J.J., Noronha, D., and Gomes, D.C.. 1998. Helminth parasites of conventionally maintained laboratory mice – II. Inbred strains with an adaptation of the anal swab technique. Memorias Do Instituto Oswaldo Cruz 93: 121-126.
    [CROSSREF]
  30. Hall, T.A.. 1999. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series 41: 95-98.
  31. Holterman, M., Van Der Wurff, A., Van Den Elsen, S., Van Megen, H., Bongers, T., and Holovachov, O.. 2006. Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown clades. Molecular Biology and Evolution 23: 1792-1800.
    [CROSSREF]
  32. Hugot, J.P.. 1980. Sur le genre Aspiculuris Schulz, 1924 (Nematoda, Heteroxynematidae), oxyures parasites de Rongeurs Muroidea. Bulletin du Museum National d´Histoire Naturelle 2: 723-735.
  33. Hugot, J.P.. 1988. Les nematodes Syphaciinae parasites de Rongeurs et de Lagomorphes. Taxonomie. Zoogeographie_Evolution. Memoires du Museum national d’Histoire Naturelle, Paris, Serie A, Zoologie 141: 1-153.
  34. Inglis, W.G., Harris, E.A., and Lewis, J.W.. 1990. A new species of the nematoda genus Aspiculuris Schulz, 1924 from Aethomys namaquensis (Mammalia: Rodentia) in the Kruger National Park, South Africa. Systematic Parasitology 17: 231-236.
    [CROSSREF]
  35. Izdebska, J.N., and Rolbiecki, L.. 2006. Correlation between the occurrence of mites (Demodex spp.) and nematodes in house mice (Mus musculus Linnaeus, 1758) in the Gdañsk urban agglomeration. Biology Letters 43 2: 175-178.
  36. Jacobs, D.E., Zhu, X.Q., Gasser, R.B., and Chilton, N.B.. 1997. PCR based methods for the identification of potentially zoonotic ascaridoid parasites of the dog, fox and cat. Acta Tropica 68: 191-200.
    [CROSSREF]
  37. Jacobson, R.H., and Reed, N.D.. 1974. The thymus dependency of resistance to pinworm infection in mice. Journal of Parasitology 60: 976-979.
    [CROSSREF]
  38. Jacoby, R.O., and Lindsey, J.R.. 1998. Health care for research animals is essential and affordable. FASEB Journal 11: 609-614.
    [CROSSREF]
  39. Jones, R., Brown, D.S., Harris, E., Jones, J., Symondson, W.O.C., and Bruford, M.W.. 2012. First record of Neoxysomatium brevicaudatum through the non-native invasive sampling of Anguis fragilis: Complementary morphological and molecular detection. Journal of Helminthology 86: 125-129.
    [CROSSREF]
  40. Kataranovski, D., Vukićević-Radić, O.D., Kataranovski, M., Radović, D.L., and Mirkov, I.I.. 2008. Helminth fauna of Mus musculus Linnaeus 1758 from the suburban area of Belgrade, Serbia. Archives of Biological Sciences 60 4: 609-617.
    [CROSSREF]
  41. Khalil, A.I., Lashein, G.H., Morsy, G.H., and Abd El-Mottaleb, D.I.. 2014. Oxyurids of wild and laboratory rodents from Egypt. Life Science Journal 11 3: 94-107.
  42. Klimpel, S., Förster, M., and Günter, S.. 2007. Parasite fauna of the bank vole Chletrionomys glareolus in an urban region of Germany: Reservoir of zoonotic metazoan parasites?. Parasitology Research 102: 69-75.
    [CROSSREF]
  43. Kohn, A., and Macedo, B.. 1984. First record of Aspiculuris tetraptera (Nitzsch, 1821) (Nematoda: Oxyuroidea) and Dollfusentis chandleri (Golvan, 1969) (Acanthocephala: Illiosentidae) in Haemulon sciurus (Shaw 1803) (Pisces: Pomadasyidae). Annales De Parasitologie Humaine Et Comparee 59 5: 477-482.
    [CROSSREF]
  44. Kruidenier, J.J., and Mehra, K.N.. 1959. Aspiculuris ackerti n. sp. (Nematoda: Oxyuridae) from the wood rats of Arizona. Proceedings of the Helminthological Society of Washington 26 2: 147-150.
  45. Landaeta-Aqueveque, C.A., Robles, M.D.R., and Cattan, P.E.. 2007. The community of gastrointestinal helminths in the house mouse, Mus musculus, in Santiago, Chile. Parasitología latinoamericana 62: 165-169.
    [CROSSREF]
  46. Li, C.L., Du, X.Y., Gao, J., Wang, C., Guo, H.G., Dai, F.W., Sa, X.Y., An, W., and Chen, Z.W.. 2016. Phylogenetic analysis of the Mongolian gerbil (Meriones unguiculatus) from China based on mitochondrial genome. Genetics and Molecular Research 15 3: 1-13.
  47. Li, M.W., Lin, R.Q., Chen, H.H., Sani, R.A., Song, R.A., and Song, H.Q.. 2007. PCR tools for the verification of the specific identity of ascaridoid nematodes from dogs and cats. Molecular and Cellular Probes 21: 349-354.
    [CROSSREF]
  48. Linnaeus, C.. 1758. Systema naturae per regna tria naturae, secundum classes, ordines, genera, species, cum characteribus, differentiis, synonymis, locis. Tomus I. Editio decima, reformata. Impensis Direct. Laurentii Salvii, Holmiae.
  49. Liu, B., Bu, Y., and Zhang, L.. 2012. A new species of Aspiculuris Schulz, 1924 (Nematoda, Heteroxynematidae) from the gray-sided vole, Clethrionomys rufocanus (Rodentia, Cricetidae), from Tianjin, China. Acta Parasitologica 57: 311-315.
    [CROSSREF]
  50. MacArthur, J.A., and Wood, M.. 1978. Control of oxyurids in mice using thiabendazole. Laboratory Animals 12: 141-143.
    [CROSSREF]
  51. Mahmoud, A.E., Attia, R.A.H., Eldeek, H.E.M., Abdel Baki, L., and Oshaish, H.A.. 2009. Oxyurid nematodes detected by colonoscopy in patients with unexplained abdominal pain. Parasitologists United Journal 2 2: 93-102.
  52. Malsawmtluangi, C., and Tandon, V.. 2009. Helminth parasite spectrum in rodent hosts from bamboo growing areas of Mizoram, north-east India. Journal of Parasitic Diseases 33 1–2: 28-35.
    [CROSSREF]
  53. Mathies, A.W.J.. 1959. Certain aspects of the host-parasite relationship of Aspiculuris tetraptera, a mouse pinworm. I. Host specificity and age resistance. Experimental Parasitology 8: 31-38.
    [CROSSREF]
  54. Mc-Manus, D.P., and Bowles, J.. 1996. Molecular genetic approaches to parasite identification: Their value in diagnostic parasitology and systematics. International Journal of Parasitology 26: 687-704.
    [CROSSREF]
  55. Millazzo, C., Ribasa, A., Casanova, J.C., Cagnin, M., Geraci, F., and Di Bella, C.. 2010. Helminths of the brown rat (Rattus norvegicus) (Berkenhout, 1769) in the city of Palermo, Italy. Helminthology 47 4: 238-240.
    [CROSSREF]
  56. Miller, G.E., and Schmidt, G.D.. 1982. Helminths of bushytailed wood rats, Neotoma cinerea subspp. from Colorado, Idaho, and Wyoming. Proceedings of the Helminthological Society of Washington 49: 109-117.
  57. Mohd Zain, S.N., Behnke, J.M., and Lewis, J.W.. 2012. Helminth communities from two urban rat populations in Kuala Lumpur, Malaysia. Parasit. Vectors 5 p. 47.
    [CROSSREF]
  58. Morales-Hojas, R., Post, R.J., Shelley, A.J., Maia-Herzog, M., Coscaron, S., and Cheke, R.A.. 2001. Characterization of nuclear ribosomal DNA sequences from Onchoncerca volvulus and Mansonella ozzardi (Nematoda: Filaroidea) and development of a PCR-based method for their detection in skin biopsies. International Journal of Parasitology 31: 169-177.
    [CROSSREF]
  59. Nakano, T., Okamoto, M., Ikeda, Y., and Hasegawa, H.. 2006. Mitochondrial cytochrome c oxidase subunit 1 gene and nuclear rDNA regions of Enterobius vermicularis parasitic in captive chimpanzees with special reference to its relationship with pinworms in humans. Parasitology Research 100: 51-57.
    [CROSSREF]
  60. Nasarova, Y.A., and Sweschnikova, N.M.. 1930. Sur la connaissance des vers parasites des Rongeurs du Kazakhstan. Review of Microbiology and Epidemiology of Parasites 9 1: 101-104.
  61. Neifer, S., Kremsner, P.G., Weinig, M., Harms, G., Sahlmüller, G., and Bienzle, U.. 1991. Interferon-gamma treatment in mice experimentally infected with Trichinella spiralis. Parasitology Research 77 5: 437-442.
    [CROSSREF]
  62. Nicklas, W., Le Corre, R., and Graw, J.. 1984. Experiences with fenbendazole in the treatment of oxyuriasis in an experimental animal colony. Berliner Und Munchener Tierarztliche Wochenschrift 97: 21-24.
    [PUBMED]
  63. Nitzsch, C.L.. 1821. Ascaris. Allg. Encycl. d. wissensch. v. Künste (Ersch und Gruber), Leipzig, 6: 44-49.
  64. Pakdel, N., Naem, S., Rezaei, F., and Chalehchaleh, A.A.. 2013. A survey on helminthic infection in mice (Mus musculus) and rats (Rattus norvegicus and Rattus rattus) in Kermanshah, Iran. Veterinary Research Forum 4 2: 105-109.
  65. Panti-May, J.A., Hernández-Betancourt, S.F., Rodríguez-Vivas, R.I., and Robles, M.R.. 2015. Infection levels of intestinal helminths in two commensal rodent species from rural households in Yucatan, Mexico. Journal of Helminthology 89: 42-48.
    [CROSSREF]
  66. Perec-Matysiak, A., Okulewicz, A., Hildebrand, J., and Zalesny, G.. 2006. Helminth parasites of laboratory mice and rats. Wiadomosci Parazytologiczne 52: 99-102.
  67. Petter, A.J., and Quentin, J.C.. 2009. Keys to the genera of the Oxyuroidea. in Anderson, R.C., Chabaud, A.G., and Willmott, S.. Eds), CIH keys to the nematoda parasites of vertebrates, Commonwealth Agricultural Bureaux, England, pp. 1-30.
  68. Pinto, R.M., Goncalves, L., Noronha, D., and Gomes, D.C.. 2001. Worm burdens in outbred and inbred laboratory rats with morphometric data on Syphacia muris (Yamaguti, 1935) Yamaguti, 1941 (Nematoda: Oxyuroidea). Memórias do Instituto Oswaldo Cruz 96: 133-136.
    [CROSSREF]
  69. Pinto, R.M., Vicente, J.J., Noronha, D., Gonçalves, L., and Gomes, D.C.. 1994. Helminth parasites of conventionally maintained laboratory mice. Memórias do Instituto Oswaldo Cruz 89: 33-40.
    [CROSSREF]
  70. Pisanu, B., Chapuis, J.L., and Durette-Desset, M.C.. 2001. Helminths from introduced small mammals on Kerguelen, Crozet, and Amsterdam Islands. Journal of Parasitology 85 5: 1205-1208.
    [CROSSREF]
  71. Popov, N., and Nazarova, J.. 1930. Neue art Parasitischer wurmer (Oxyuridae). Vestnik Mikrob. Epid. Parazitol. Saratow 9 1: 105-108.
  72. Qiu, J.H., Lou, Y., Zhang, Y., Chang, Q.C., Liu, Z.X., Duan, H., Guo, D.H., Gao, D.Z., Yue, D.M., and Wang, C.R.. 2016. Sequence variability in internal transcribed spacers of nuclear ribosomal DNA among isolates of the oxyurid nematodes Syphacia obvelata and Aspiculuris tetraptera from mice reared in laboratories in China. Journal of Helminthololy 90 1: 81-85.
    [CROSSREF]
  73. Quentin, J.C.. 1966. Oxyures de Muridae africains. Annales De Parasitologie Humaine Et Comparee 41 5: 443-452.
    [CROSSREF]
  74. Quentin, J.C.. 1975. Essai de classification des oxyures Heteroxynematidae. Mémoires du Muséum National Histoire Naturelle, Zoologie 94: 51-96.
  75. Rehbinder, C., Baneux, P., Forbes, D., Van Herck, H., Niclas, W., and Rugaya, Z.Y.. 1996. FELASA – Recommendations for the health monitoring of mouse, rat, hamster, gerbil, guinea pig and rabbit experimental units. Laboratory Animal Science 30: 193-208.
    [CROSSREF]
  76. Ribas, A., Diagne, C., Tatard, C., Diallo, M., Poonlaphdecha, S., and Brouat, C.. 2017. Whipworm diversity in West African rodents: A molecular approach and the description of Trichuris duplantieri n. sp. (Nematoda: Trichuridae). Parasitology Research 116 4: 1265-1271.
    [CROSSREF]
  77. Robles, M.R., and Navone, G.T.. 2007. A new species of Syphacia (Nematoda: Oxyuridae) from Oligoryzomys nigripes (Rodentia: Cricetidae) in Argentina. Parasitology Research 101 4: 1069-1075.
    [CROSSREF]
  78. Robles, M.R., and Navone, G.T.. 2010. Redescription of Syphacia venteli Travassos 1937 (Nematoda: Oxyuridae) from Nectomys squamipes in Argentina and Brazil and description of a new Syphacia from Melanomys caliginosus in Colombia. Parasitology Research 106 5: 1117-1126.
    [CROSSREF]
  79. Rosas, G.A.. 1997. Diagnóstico: Parasitosis intestinal por Aspiculuris tetraptera. Animals Experimentation 2: 9-11.
  80. Sasa, M., Tanaka, H., Fukui, M., and Takata, A.. 1962. Internal parasites of laboratory animals. in Harris, R.J.C.. Ed.), The Problems of Laboratory Animal Disease, Academic Press, London; New York, pp. 195-214.
  81. Sato, Y., Ooi, H.K., Nonaka, N., Oku, Y., and Kamiya, M.. 1995. Antibody production in Syphacia obvelata infected mice. Journal of Parasitology 8: 559-562.
    [CROSSREF]
  82. Schulz, R.E.S.. 1924. Oxyuridae of Armenian mice. Reportes Tropical Institute Armenia 2: 41-51.
  83. Scott, M.E., and Gibbs, H.C.. 1986. Long-term population dynamics of pinworms (Syphacia obvelata and Aspiculuris tetraptera) in mice. Journal of Parasitology 72: 652-662.
    [CROSSREF]
  84. Semenova, S.K., Romanova, E.A., and Pyskov, A.P.. 1996. Genetic differentiation of helminthes on the basis of data of polymerase chain reaction using random primers. Genetics 32 2: 304-309.
  85. Singleton, G.R., Smith, A.L., Shellam, G.R., Fitzgerald, N., and Muller, W.J.. 1993. Prevalence of viral antibodies and helminthes in field populations of house mice (Mus domesticus) in southeastern Australia. Epidemiology and Infection 110: 399-417.
    [CROSSREF]
  86. Skrjabin, K.I., and Schikhobalova, N.D.. 1948. Quoted from Petter and Quentin (1976).
  87. Sotillo, J., Trelis, M., Cortés, A., Luz Valero, M., Sánchezdel Pino, M., and Esteban, J.G.. 2012. Proteomic analysis of the pinworm Syphacia muris (Nematoda: Oxyuridae), a parasite of laboratory rats. Parasitology International 61 4: 561-564.
    [CROSSREF]
  88. Stewart, A., Lowe, A., Smales, L., Bajer, A., Bradley, J., Dwuznik, D., Franssen, F., Griffith, J., Stuart, P., Turner, C., Zalesny, G., and Behnke, J.M.. 2017. Parasitic nematodes of the genus Syphacia Seurat, 1916 infecting Muridae in the British Isles, and the peculiar case of Syphacia frederici. Parasitology 23: 1-12.
  89. Stojcevic, D., Mihljevic, Z., and Marnculic, A.. 2004. Parasitological survey of rats in rural regions of Croatia. Veterinary Medicine – Czech 49 3: 70-74.
    [CROSSREF]
  90. Swofford, D.L.. 2000. PAUP*. Phylogenetic Analysis using Parsimony (*and other Methods), Version 4, Sinauer Associates, Sunderland, MA.
  91. Taffs, L.F.. 1976. Pinworm infection in laboratory rodents: a review. Laboratory Animals 10: 1-13.
    [CROSSREF]
  92. Tarzhimanova, R.A.. 1969. New nematodes of the genus Aspiculuris from Rodents. Azerbaidzhanskogo Nauchno. Issledovatel’skogo Instituta Meditsinkoi Parasitologii Tropicheskoi Meditsiny im. S.M. Kirova 7: 302-306.
  93. Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., and Higgins, D.G.. 1997. The CLUSTAL-X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Research 25: 4876-4882.
    [CROSSREF]
  94. Travassos, L.. 1919. Quoted from Petter and Quentin (2009).
  95. Verma, S., Gaherwal, S., Prakash, M.M., and Kanhere, R.R.. 2013. Anthelmintic efficacy of Ocimum sanctum against Syphacia muris in mice. Acta Parasitologica 4 1: 24-28.
  96. Vermund, S.H., and Wilson, C.M.. 2000. Pinworm (Enterobius vermicularis). Seminars in Pediatric Infectious Diseases 11: 252-256.
    [CROSSREF]
  97. Weyand, N.J., Ma, M., Phifer-Rixey, M., Taku, N.A., Rendon, M.A., Hockenberry, A.M., Kim, W.J., Agellon, A.B., Biais, N., Suzuki, T.A., Goodyer-Sait, L., Harrison, O.B., Bratcher, H.B., Nachman, M.W., Maiden, M.C., and So, M.. 2016. Isolation and characterization of Neisseria musculi sp. nov., from the wild house mouse. International Journal of Systematic and Evolutionary Microbiology 66 9: 3585-3593.
    [CROSSREF]
  98. Wijová, M., Moravec, F., Horák, A., and Lukeš, J.. 2006. Evolutionary relationships of Spirurina (Nematoda: Chromadorea: Rhabditida) with special emphasis on dracunculoid nematodes inferred from SSU rRNA gene sequences. International Journal for Parasitology 36: 1067-1075.
    [CROSSREF]
  99. Yamaguti, S.. 1935. Studies on the helminth fauna of Japan. Part 13. Mammalian nematodes. Japanese Journal of Zoology 6: 433-457.
  100. Zarei, Z., Mohebali, M., Heidari, Z., Davoodi, J., Shabestari, A., Motevalli Haghi, A., Khanaliha, K., and Kia, E.B.. 2016. Helminth Infections of Meriones persicus (Persian Jird), Mus musculus (House Mice) and Cricetulus migratorius (Grey Hamster): A cross-sectional study in Meshkin-Shahr District, Northwest Iran. Iranian Journal of Parasitology 11 2: 213-220.
    [PUBMED]
  101. Zenner, L.. 1998. Effective eradication of pinworms (Syphacia muris, Syphacia obvelata and Aspiculuris tetraptera) from a rodent breeding colony by oral anthelmintic therapy. Laboratory Animals 32: 337-342.
    [CROSSREF]
  102. Zenner, L., and Regnault, J.P.. 2000. Ten-year long monitoring of laboratory mouse and rat colonies in French facilities: A retrospective study. Laboratory Animals 34: 76-83.
    [CROSSREF]
  103. Zhu, X.Q., Amelio, S.D., Gasser, R.B., Yang, T.B., Paggi, L., and He, F.. 2007. Practical PCR tools for the delineation of Contracaecum rudolphii A and Contracaecum rudolphii B (Ascaridoidea: Anisakidae) using genetic markers in the nuclear ribosomal DNA. Molecular Cellular Probes 21: 97-102.
    [CROSSREF]
  104. Zhu, X.Q., Jacobs, D.E., Chilton, N.B., Sani, R.A., Cheng, N., and Gasser, R.B.. 1998. Molecular characterization of Toxocara variant from cats in Kuala Lumpur, Malaysia. Parasitology 117: 155-164.
    [CROSSREF]

EXTRA FILES

COMMENTS