First Record of Acrobeloides nanus (Cephalobidae: Rhabditida: Nematoda) from Korea

  • Kim, Taeho (Division of Environmental Science and Ecological Engineering, College of Life Sciences and Biotechnology, Korea University) ;
  • Kim, Jiyeon (Freshwater Biodiversity Research Division, Nakdonggang National Institute of Biological Resources) ;
  • Bae, Yeon Jae (Division of Environmental Science and Ecological Engineering, College of Life Sciences and Biotechnology, Korea University) ;
  • Park, Joong-Ki (Division of EcoScience, Ewha Womans University)
  • Received : 2016.08.18
  • Accepted : 2016.10.27
  • Published : 2016.10.31


Acrobeloides nanus (de Man, 1880) Anderson, 1968 belonging to the family Cephalobidae Filpijev, 1934 (Cephalobomorpha) is newly reported from South Korea. This species is distinguished from other Acrobeloides species by its low and blunt labial probolae, five lateral incisures with middle incisure extending to the tail tip, and bluntly rounded tail. In this study, details of morphological characters of A. nanus is described and illustrated based on optical and scanning electron microscopy. In addition, molecular sequence data of the D2-D3 region of 28S rDNA, 18S rDNA and mitochondria DNA cox1 region from this species are provided as DNA barcode sequences.


Genus Acrobeloides (Cobb, 1924) Thorne, 1937 are bacterial feeding nematodes and are widely distribute in various terrestrial environments such as forests (Háněl, 1999), sand dunes (Yeates, 1967), and agricultural land (Pervez, 2011). Species in this group have been studied extensively (Thorne, 1937; Brezeski, 1962; Anderson, 1965, 1968; Andrássy, 1984; Siddiqi et al., 1992). To date, only one Acrobeloides species (unidentified at the species level) has been reported in Korea (Kim et al., 2012).

Following a survey of several plots of farmland, A. nanus (de Man, 1880) Anderson, 1968, were isolated from soil samples from potato farms. In this paper, we provide details of a morphological characters and morphometrics for this species from optical microscope and scanning electron microscope (SEM) images. In addition, molecular sequence information of the D2-D3 region of the 28S rDNA, 18S rDNA, and mitochondrial DNA cox1 region from this species are provided as DNA barcode sequence data.



Nematode isolation and culture

Nematode specimens were extracted from potato farm soil from Hapcheon-gun, Gyeongsangnam-do, South Korea (GPS coordinates: 35°27′37.4″N, 128°00′19.6″E), using sieving and the Baermann funnel method. One individual nematode was transferred to a soil agar plate (25 mg/mL autoclaved soil, 5 μg/mL cholesterol, and 1% agar) and cultured at room temperature (18-20℃).

Fixation and morphological observation

For fixation, the nematode specimen was transferred to 2 mL water in a 15 mL tube, to which was added 4 mL of 80℃ TAF (2% triethanolamine and 7% formaldehyde). The fixed nematodes were processed to dehydrated glycerin using Seinhorst’s (1959) method and mounted in pure glycerin on permanent HS-slides (Shirayama et al., 1993). Morphological characters of nematode specimens were observed under an optical microscope (BX-51; Olympus, Tokyo, Japan) equipped with differential interference contrast, and morphometric characters were measured using a CoolSnap Photometrics color CCD digital camera (MP5.0-RTV-R-CLR-10; Photometrics, Tucson, AZ, USA) and the programQCapture Pro 5 (QImaging, Surrey, Canada).

Scanning electron microscope (SEM)

For SEM imaging, nematode specimens were fixed using TAF and maintained for a minimum of 24 h at room temperature. They were then transferred to a 4% aqueous osmium tetroxide solution and kept at 4℃ for 3 days for postfixation. Fixed nematode specimens were dehydrated through a 10%-100% pure ethanol series for 1 h. The samples were dried using a Hitachi HCP-2 critical point drier (Tokyo, Japan). Dried nematodes were mounted on copper/nickel tape and sputter coated with gold/palladium using an Eiko IB-3 ion coater (Tokyo, Japan). Morphological characters of the nematode specimens were observed with a Zeiss Ultra Plus SEM (Oberkochen, Germany) at 15 kV under high-vacuum conditions.

Molecular techniques and SEQUENCES analysis

Total genomic DNA from A. nanus was extracted using an Epicentre MasterPure DNA Purification Kit (Epicentre, Madison, WI, USA) following the manufacturer’s protocol. For amplification of the D2-D3 regions of 28S rDNA, 18S rDNA, and mitochondrial DNA cox1 fragments, polymerase chain reaction (PCR) was performed using universal primer sets (D2A [5′-ACAAGTACCGTGAGGGAAAGTTG-3′]/ D3B [5′-TCGGAAGGAACCAGCTACTA-3′]; De Ley et al., 1999 for D2-D3 region of 28S and 328-F [5′-TACCTG GTTGATCCTGCCAG-3′]/329-R [5′-TAATGATCCTTCC GCAGGTT-3′]; Adl et al., 2014 for 18S) and a nematodespecific primer set (Cepha_CO1_F [5′-ATGATTTTTTTTAT GGTGATGCC-3′]/Cepha_CO1_R [5′-ACTACAAAATATG TGTCATG-3′] for cox1 region) that was designed based on conserved regions of nematode mitochondrial genes. PCR reactions were performed in a total volume of 50 μL including 2 μL template DNA, 10 pmol of each primer, 10× Ex Taq buffer, 0.2 mM dNTP mixture, and 1.25 U of Taq polymerase (TaKaRa Ex Taq). PCR amplification conditions were as follows: initial denaturing step at 95℃ for 1 min, followed by 35 cycles of denaturation at 95℃ for 30 s, annealing at 50℃ for 30 s, and extension at 72℃ for 1 min (extended to 2 min for the 328/329 primer) followed by a final extension at 72℃ for 10 min. The amplified PCR products were purified using a QIAquick Gel Extraction Kit (Qiagen, Hilden, Germany) according to the manufacturer’s protocols. Big Dye Terminator Cycle-Sequencing (Applied Biosystems, Waltham, MA, USA) was used for sequencing the PCR-amplified fragments.

The obtained sequences of the D2-D3 region of 28S rDNA, 18S rDNA, and mtDNA cox1 region from the specimens were aligned with sequences of other acrobeloids available from GenBank, using Clustar X with default options (Thompson et al., 1997). Both ends of the aligned datasets were trimmed before sequence analyses.



1*Acrobeloides nanus (de Man, 1880) Anderson, 1968 (Table 1, Figs. 1, 2)

Material examined. 18♀♀, Korea: Gyeongsangnam-do, Hapcheon-gun, Gahoe-myeon, 26 Mar 2015, extracted by sieving and the Baermann funnel method from potato farm soil. Two specimens (slide Nos. NIBRIV0000326012 and NIBRIV0000326013) are deposited at the National Institute of Biological Resources, Republic of Korea. Sixteen specimens (slide Nos. 01010503001-01010503016) are deposited in the Animal Phylogenomics Laboratory, Ewha Womans University, Republic of Korea.

Measurements. See Table 1.

Table 1.All measurements are in μm and in the form mean±SD (range). aNumber of annules from the anterior end to the nerve ring. bNumber of annules from the anterior end to the excretory pore. cNumber of annules from the anterior end to the deirid. dNumber of annules from the anus to the tail end.

Description. Female: Body cylindrical, length 335.3-442.3 μm, ventrally curved after fixation (Fig. 1A). Cuticle annulated; annuli 1.6-2.2 μm wide and 0.5-0.8 μm thick at midbody. Lateral incisures varying in number along body length: three incisures at procorpus region, branching off from deirid into five incisures, and three incisures at anterior anus; two incisures fading out around phasmid; middle incisure extending to near tail end (Figs. 1A, 2B, D). Head region continuous with neck. Lip region 6.6-7.9 μm wide, triradiate symmetry with 6+4 papillae. Cephalic probolae absent; three straight, conical-rounded labial probolae present. Amphid openings present, transversely opening, and oval shaped (Figs. 1B, 2A). Stoma cephaloboid with length about 1.6-2 times lip region diameter; bar-shaped cheilorhabdions, with dorsal denticle on metastom. Pharyngeal corpus fusiform with swollen metacorpus, 3-5.2 times isthmus length. Isthmus narrower than corpus, distinctly demarcated from metacorpus. Basal bulb oval-shaped, with well-developed valves at middle part; cardia conoid, surrounded by intestinal tissue. Nerve ring located posterior of corpus to anterior isthmus region, 38-51 annuli from head, at 61.9%-69.5% of pharynx length. Excretory pore at posterior corpus to isthmus level, 41-54 annuli from anterior end, at 64.7%-73.7% of pharynx length. Deirid in lateral field at isthmus level, 47-59 annuli from anterior end, at 72.8%-81.1% of total neck length (Fig. 1B). Female reproductive system monodelpic-prodelpic. Vulva not protruding, vagina one-third of body width, postvulval sac rudimentary. Uterus tubular, 1.4-1.8 times body diameter. Spermatheca short (7.4-19.6 μm). Oviduct 1.5 times body width long, Ovary straight to posterior, sometimes with double flexure (n=2) (Figs. 1C, 2C). Rectum length 1.0-1.5 times anal body width. Tail conoid with rounded terminus, with 8-13 annules. Phasmids at 23.7%-43.3% of tail length (Figs. 1D, 2D).

Fig. 1.Acrobeloides nanus (de Man, 1880) Anderson, 1968. A, Entire female; B, Neck region; C, Female reproductive system; D, Female posterior region. Scale bars: A=50 μm, B-D=20 μm.

Fig. 2.Acrobeloides nanus (de Man, 1880) Anderson, 1968 (scanning electron microscopy). A, Head region; B, Lateral field at deirid region; C, Vulva; D, Tail region. Scale bars: A-C=1 μm, D=2 μm.

Male: Unknown.

Distribution. Australia (Bird et al., 1993), Brazil (Rashid et al., 1984), Canada (Anderson, 1968), Falkland Islands (Boström, 1996), South Georgia Island (Boström, 1996), Krakatau (Rashid et al., 1988), Malaysia (Boström, 1993), Korea (present study), Sweden (Boström and Gydemo, 1983).

Habitat. Soil sample in the potato farm.

Remarks. Anderson (1968) proposed that in A. nanus, there are intraspecific variations in some morphologies depending on environmental conditions, such as measurements, shape of labial probolae (low-rounded, knobbed, conoid, and apiculate) and tail (hemispherical, clavate, conoid-truncated, and conoid-rounded) and position of phasmid, nerve ring, excretory pore and deirid. The aforementioned morphological variability among A. nanus populations has also been reported from many geographic areas: Australia (Bird et al., 1993), Brazil (Rashid et al., 1984), Canada (Anderson, 1968), Falkland Islands (Boström, 1996), South Georgia Island (Boström, 1996), Krakatau (Rashid et al., 1988), Malaysia (Boström, 1993), and Sweden (Boström and Gydemo, 1983). The morphological characters of the specimens observed from the present study are within the range of intraspecific variation reported from other localities in earlier studies (Table 2).

Table 2.Morphometrics and morphological variability among Acrobeloides nanus (de Man, 1880) Anderson, 1968 populations

Identifying characteristics that distinguish A. nanus from A. buetschlii (de Man, 1884) Steiner and Buhrer, 1933 have long been debated. Anderson (1968) and Zell (1987) distinguished between A. nanus and A. buetschlii by the number of lateral incisures (five vs. three) and absence/presence of a postvulvar uterine branch (PUB). However, Rashid et al. (1984) reported three lateral incisures in A. nanus from a Brazil population. In addition, Bird et al. (1993), Boström (1993, 1996), and Rashid et al. (1984) refuted morphological differences (number of lateral incisures and absence or presence of PUB) between A. nanus and A. buetschlii.

Molecular sequence information. Molecular sequences deposited on GenBank: D2-D3 region in 28S rDNA (GenBank accession No. KX669640); 18S rDNA (GenBank accession No. KX669638); cox1 of mtDNA (GenBank accession No. KX669639).

Molecular information. The sequences of the D2-D3 region of 28S rDNA, 18S rDNA, and the partial cox1 gene of mitochondrial DNA were obtained from A. nanus (GenBank accession Nos. KX669640 [D2-D3 region of 28S rDNA], KX669638 [18S rDNA], and KX669639 [partial cox1 gene of mtDNA]) and compared with other acrobeloids available on GenBank. The cox1 sequences from other Acrobeloides species are not yet available on GenBank. We provide cox1 sequence data from A. nanus in this study for use in molecular barcoding.

The sequence of the D2-D3 region of 28S rDNA from A. nanus in this study is the same as A. thornei (DQ903083) and differs by one to five nucleotides from A. buetschlii (DQ903081; 3 bp), A. ellesmerensis (DQ145624; 3 bp), A. uberrinus (DQ903087; 3 bp), and A. nanus specimens from Jädras in Sweden (DQ903076; 1 bp), Cologne in Germany (EF417139; 1 bp), Sollentuna in Sweden (DQ903075; 2 bp), and Bourges in France (DQ903103; 5 bp). The sequence of the 18S rDNA from A. nanus in this study is the same as A. buetschlii (JQ957905), and differs by one or three nucleotides from A. nanus from an unknown location (DQ102707; 1 bp), A. apiculatus (AY284673; 1 bp) and A. thornei (JQ 957903; 3 bp). However, intraspecific variation of D2-D3 region sequences among some A. nanus populations is higher than interspecific variation among several Acrobeloides species. For example, the D2-D3 region sequences of A. ellesmerensis (DQ145624), A. uberrinus (DQ903078), and A. buetschlii (DQ903081) are identical, but D2-D3 sequences between different isolates of A. nanus (DQ903075 [Sweden], DQ903103 [France]) differ by 7 nucleotides. Also, the 18S rDNA sequences of A. nanus from an unknown location (DQ 102707) is distinguished by two base pairs from A. apiculatus (AY284673) and A. nanus in this study (KX669638). As described, molecular sequence data of A. nanus was the same or very similar to some other acrobeloids; however, its morphology clearly distinguishes A. nanus from A. thornei (with two lateral incisures, setose labial probolae and pointed tail), A. buetschlii (with three lateral incisures), A. ellesmerensis (with four lateral incisures with three extending to tail end, and setose labial probolae), A. uberrinus (with two to three incisures extending to tail end, and setose labial probolae), and A. apiculatus (with a pointed tail). In addition, earlier studies have reported that the D2-D3 region of 28S rDNA and the 18S sequence did not show clear resolution in their relationships among some species within Cephalobidae (Holterman et al., 2006; Nadler et al., 2006; Smythe and Nadler, 2006; Sonnenberg et al., 2007; Rybarczyk-Mydłowska et al., 2012). Therefore, the D2-D3 region of 28S rDNA and 18S rDNA should be used with great caution as molecular markers for species level identification of Acrobeloides species.


  1. Adl SM, Habura A, Eglit Y, 2014. Amplification primers of SSU rDNA for soil protists. Soil Biology and Biochemistry, 69:328-342.
  2. Anderson RV, 1965. Acrobeloides uberrinus n. sp., with a note on morphologic variation within soil and bacteria-reared populations. Proceedings of the Helminthological Society of Washington, 32:232-235.
  3. Anderson RV, 1968. Variation in taxonomic characters of a species of Acrobeloides (Cobb, 1924) Steiner and Buhrer, 1933. Canadian Journal of Zoology, 46:309-320.
  4. Andrássy I, 1984. Klasse Nematoda: Ordnungen Monhysterida, Desmoscolecida, Araeolaimid, Chromadorida, Rhabditida. Akademie-Verlag, Berlin, pp. 1-509.
  5. Bird AF, De Ley P, Bird J, 1993. Morphology, oviposition and embryogenesis in an Australian population of Acrobeloides nanus. Journal of Nematology, 25:607-615.
  6. Boström S, 1993. Some cephalobids from Ireland and Malaysia (Nematoda: Rhabditida). Afro-Asian Journal of Nematology, 3:128-134.
  7. Boström S, 1996. One new and two known nematode species from the Subantarctic Islands South Georgia and East Falkland Island. Fundamental and Applied Nematology, 19:151-158.
  8. Boström S, Gydemo R, 1983. Intraspecific variability in Acrobeloides nanus (de Man) Anderson (Nematoda, Cephalobidae) and a note on external morphology. Zoologica Scripta, 12:245-255.
  9. Brzeski M, 1962. Three new species of the genus Acrobeloides Cobb. (Nematoda, Cephalobidae). Bulletin de L’Académie Polonaise des Sciences, 10:335-339.
  10. De Ley P, Felix MA, Frisse LM, Nadler SA, Sternberg PW, Thomas WK, 1999. Molecular and morphological characterisation of two reproductively isolated species with mirror- image anatomy (Nematoda: Cephalobidae). Nematology, 1:591-612.
  11. de Man JG, 1880. Die einheimischen, frei in der feinen Erde und im süssen Wasser lebenden Nematoden. Vorläufiger Bericht und descriptive-systematischer Theil. Tijdschrift Nederlandsche Dierkundige Vereeniging, 5:1-104.
  12. Háněl L, 1999. Fauna of soil nematodes (Nematoda) in Trojmezná hora Reserve. Silva Gabreta, 3:89-94.
  13. Holterman M, van der Wurff A, van den Elsen S, van Megen H, Bongers T, Holovachov O, Bakker J, Helder J, 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.
  14. Kim DG, Park BY, Ryu YH, 2012. Soil Nematode fauna in Dokdo island of Korea. Research in Plant Disease, 18:381-386.
  15. Nadler SA, De Ley P, Mundo-Ocampo M, Smythe AB, Stock SP, Bumbarger D, Adams BJ, De Ley IT, Holovachov O, Baldwin JG, 2006. Phylogeny of Cephalobina (Nematoda): molecular evidence for recurrent evolution of probolae and incongruence with traditional classifications. Molecular Phylogenetics and Evolution, 40:696-711.
  16. Pervez R, 2011. Acrobeloides ishraqi sp. n. and Acrobeloides mushtaqi sp. n. (Nematoda: Rhabditida) from chickpea rhizosphere, Uttar Pradesh, India. Archives of Phytopathology and Plant Protection, 44:1438-1446.
  17. Rashid F, Geraert E, Sharma RD, 1984. Morphology, taxonomy and morphometry of some Cephalobidae (Nematoda: Rhabditida) from Brazil, with descriptions of two new genera and four new species. Nematologica, 30:251-298.
  18. Rashid F, Geraert E, Coomans A, Suatmadji W, 1988. Cephalobidae from the Krakatau region (Nematoda: Rhabditida). Nematologica, 34:125-143. 988X00224
  19. Rybarczyk-Mydłowska K, Mooyman P, van Megen H, van den Elsen S, Vervoort M, Veenhuizen P, van Doorn J, Dees R, Karssen G, Bakker J, Helder J, 2012. Small subunit ribosomal DNA-based phylogenetic analysis of foliar nematodes (Aphelenchoides spp.) and their quantitative detection in complex DNA backgrounds. Phytopathology, 102:1153-1160.
  20. Seinhorst JW, 1959. A rapid method for the transfer of nematodes from fixative to anhydrous glycerin. Nematologica, 4:67-69.
  21. Shirayama Y, Kaku T, Higgins RP, 1993. Double-sided microscopic observation of meiofauna using an HS-slide. Benthos Research, 44:41-44.
  22. Siddiqi MR, De Ley P, Khan HA, 1992. Acrobeloides saeedi sp. n. from Pakistan and redescription of A. bodenheimeri (Steiner) and Placodira lobata Thorne (Nematoda: Cephalobidae). Afro-Asian Journal of Nematology, 2:5-16.
  23. Smythe AB, Nadler SA, 2006. Molecular phylogeny of Acrobeloides and Cephalobus (Nematoda: Cephalobidae) reveals paraphyletic taxa and recurrent evolution of simple labial morphology. Nematology, 8:819-836.
  24. Sonnenberg R, Nolte AW, Tautz D, 2007. An evaluation of LSU rDNA D1-D2 sequences for their use in species identification. Frontiers in Zoology, 4:6.
  25. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG, 1997. The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Research, 25:4876-4882.
  26. Thorne G, 1937. A revision of the nematode family Cephalobidae Chitwood and Chitwood, 1934. Proceedings of the Helminthological Society of Washington, 4:1-16.
  27. Yeates GW, 1967. Studies on nematodes from Dune sands. 5. Acrobelinae. New Zealand Journal of Science, 10:527-547.
  28. Zell H, 1987. Nematoden eines Buchenwaldbodens 9. Die Cephaloben (Nematoda, Rhabditida). Carolinea, 45:121-134.