Geographical distribution and phoretic associations of the viviparous nematode Tokorhabditis atripennis with Onthophagus dung beetles in Japan

To isolate T. atripennis, we collected dung beetles using various methods, including pit-hole traps, direct capture from within or beneath animal droppings, excavation of underground nests, and nocturnal searches among insects attracted to artificial light. For the pit-hole traps, we dug the soil surface and placed paper cups at a volume of 105 ml at ground level. Rotted fish meal was used as an attractant for dung beetles. Pit-hole traps were set up at four distinct locations: Kasugai campus of Chubu University, Kasugai, Japan (Coordinates, 35° 16′ 31.8″ N, 137° 00′ 58.6″ E; Date, 2016.6.5, 7.5, 11.5, 2017.5.8, 5.10, 5.26, 2020.11. 16 to 28), Ena Campus of Chubu University, Ena, Japan (Coordinates, 35° 25′ 45.6″ N, 137° 21′ 15.2″ E; Date, 2016.5.22), Ikuta Campus of Meiji University, Kawasaki, Japan (Coordinates, 35° 36′ 39.8″ N, 139° 32′ 55.8″ E; Date, 2021.5.14, 8.2 to 4, 9.16) and at the Meiji University Kurokawa Field Science Center, Kawasaki, Japan (Coordinates, 35° 36′ 31.5″ N, 139° 27′ 20.8″ E; Date, 2021.9.30, 10.20, 10.21). Details of the dung beetles collected using methods other than pit-hole traps are described in Table 1.

The collected beetles were individually stored in plastic cases with humid tissue paper until dissection. Before dissection, each beetle’s viability was confirmed, and they were observed under a stereo microscope for species and sex identification following the illustration reference, “Scarab beetles of Japan” (Kawai et al., 2008). Most beetles were able to survive for up to one week using this sampling method. However, if any beetles died, they were excluded from further study.

The dung beetles were dissected to verify the presence or absence of phoretically associated nematodes. We identified six distinct body parts of the nematode: 1) the entire body surface, 2) the dorsum of the wings, 3) the pronotum-elytron groove, 4) the male genitalia and testis, 5) ovary, and 6) the pronotum-front groove and the anterior regions for locating the phoretic nematodes (Fig. 1). We carefully rinsed the surface of the dung beetles with ion exchange water (IEW) on a Syracuse watch glass and looked for nematodes under a dissecting microscope. Subsequently, we dissected the beetle elytron and examined the dorsal area as well as the groove between the pronotum and the elytron in the presence of nematodes. Next, we removed and examined the male genitalia, testes, and ovaries for the presence of nematodes. Then, we detached the heads of dung beetles from the pronotum and observed the presence of nematodes.

Figure 1:

Figure 2:

Once nematodes were found during the dissection of dung beetles, we transferred them onto nematode growth medium (NGM) (Brenner, 1974), as well as onto the NGM + dog food medium (DFM: 20 g crushed dog food, 4 g agar and IEW were poured until the total volume became 200 ml, autoclaved and solidified) seeded with E. coli OP50 and incubated them at 25 °C (Hara et al., 1981; Ogura and Mamiya, 1989). Two to three days after being transferred to NGM or DFM plates, most free-living nematodes reached the adult stage, allowing us to distinguish their reproductive modes as either oviparous or viviparous. Viviparous nematodes were sterilized using a 10% SDS solution and established as laboratory strains, except for the strains of SHR that were just washed in IEW. Although oviparous nematodes were collected in this study, they were not identified as strains.

Genomic DNA were extracted from nematodes using the Qiagen DNeasy Blood and Tissue Kit (Qiagen, USA) or Direct PCR Lysis Reagent (Viagen Biotech, USA). The D1/D4 or D2/D3 extension segments of the 28S ribosomal RNA gene (LSU) (Kanzaki et al., 2021; Nunn, 1992) and a partial fragment of the 18S ribosomal RNA gene (SSU) (Floyd et al., 2005; Carta and Li., 2018) were amplified and sequenced using universal primers (Table S1). The obtained DNA fragments were purified from agarose gels with NucleoSpin^® Gel and PCR Clean-up (Macherey-Nagel, Germany) or FastGene Gel/PCRExtraction kit (NIPPON Genetics Co., Ltd, Japan). Samples were submitted to Hokkaido System Science Co. (Sapporo, Japan) or Macrogen Japan Corp. for sequencing from both strands using the same PCR primers. The sequences obtained were confirmed and edited manually using a Serial Cloner (ver. 2-6-1) and identified to the species level based on the results of the Basic Local Alignment Search Tool (BLAST) analysis. Isolated nematodes were considered potentially distinct species if they exhibited <99% sequence similarity to their nearest neighbors. Sequence data were deposited in the NCBI for Biotechnology Information GenBank database.

Significant differences in the percentages of dung beetles associated with viviparous nematodes among regions, beetle genera, and species were determined. First, a Fisher’s exact test was performed using the R package in Jupyter Lab. When significant differences were detected by the Fisher’s exact test, Tukey’s WSD was performed for multiple comparisons at P < 0.05.

A total of 615 dung beetles were collected from 12 prefectures and classified into 30 species across six genera (Table S2). Most of the nematodes associated with dung beetles were free-living nematodes with oviparous reproduction (data not shown). This was evident when eggs were laid on plates seeded with E. coli OP50 after reaching adulthood. However, a subset of the isolated nematodes did not lay eggs, but released larvae, leading us to classify them as exhibiting viviparous reproduction.

All isolated viviparous nematodes were successfully cultured on plates, genomic DNA was isolated, and sequences of the LSU and SSU genes were obtained. In total, 27 viviparous nematode strains were obtained from individual dung beetles of eight species found in six prefectures (Fig. 2; Table 2). Upon comparing their LSU and SSU gene sequences, it was evident that all genes in the 27 strains were identical to those of T. atripennis. The NCBI accession numbers assigned to the gene sequences of each strain are shown in Table S3.

Most dung beetles associated with T. atripennis were members of the genus Onthophagus (23/276). A few beetles belonging to the genera Caccobius (2/118) and Phelotrupes (2/85) were also associated with T. atripennis. The association rate differed significantly between Onthophagus and Phelotrupes (Tukey’s WSD, q=3.3145, wsd=0.05945). T. atripennis was not isolated from the beetles belonging to the genera Aphodius (n=103), Ataenius (n=8), or Copris (n=25) (Table 3). Across all beetle species, approximately 70% of T. atripennis detected in this study were isolated from two species, O. apicetinctus and O. atripennis, both of which belong the genus Onthophagus. When multiple comparisons were performed (Tukey’s WSD, q=4.286, P < 0.05), significant differences in association rates were observed among the three cohorts: Caccobius jessoensis and Onthophagus apicetinctus (wsd=0.1420), Phelotrupes laevistriatus and O. apicetinctus (wsd=0.1530), and O. atripennis and C. jessoensis (wsd=0.1139). Although there was no significant association between Onthophagus species and viviparous nematodes, the rates of two Onthophagus species, O. atripennis and O. apicetinctus, were notably elevated compared to other Onthophagus species (Table 3). This suggests a preference for viviparous nematodes with specific Onthophagus species as hosts.

During the dissection of dung beetles, we investigated the nematode-associated sites on the beetles’ bodies by categorizing them into six parts:1) entire body surface, 2) dorsum of the wings, 3) pronotum-elytron groove, 4) male genitalia and testis, 5) ovary, and 6) pronotum-front groove. In general, T. atripennis was primarily associated with 2) the dorsum of the wings and 3) the pronotum-elytron groove, with no instances of T. atripennis detected in 5) the ovaries (Table 4). In the Onthophagus genus, T. atripennis was observed on 2) the dorsum of the wings (10/24) and 3) the pronotum-elytron groove (11/24). Nematodes were detected on 1) the entire body surface (2/24) and 4) the male genitalia and testis. In Caccobius beetles, T. atripennis has been isolated from 2) dorsum of the wings (1/3), 3) pronotum-elytron groove (1/3), and 4) male genitalia and testes (1/3). In Phelotrupes beetles, T. atripennis was isolated from 1) the entire body surface (1/2) and 6) the groove between the pronotum and front (1/2).

In terms of individual prefectures, the highest association rate of T. atripennis was observed in Okinawa (11/84), followed by Miyazaki (1/8) and Kanagawa (5/90). The rates in the other three prefectures were all below 5%, as shown in Table 5. Among Onthophagus species, T. atripennis was detected in four prefectures: Aichi (4/45), Kanagawa (3/39), Nara (5/95), and Okinawa (11/60). Based on these findings, it appears that T. atripennis is distributed across Japan, wherever Onthophagus species are found.