Redescription and phylogenetic analysis of the type species of the genus Panagrellus Thorne, 1938 (Rhabditida, Panagrolaimidae), P. pycnus Thorne, 1938, including the first SEM study

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

31
Reader(s)
42
Visit(s)
0
Comment(s)
0
Share(s)

SEARCH WITHIN CONTENT

FIND ARTICLE

Volume / Issue / page

Related articles

Redescription and phylogenetic analysis of the type species of the genus Panagrellus Thorne, 1938 (Rhabditida, Panagrolaimidae), P. pycnus Thorne, 1938, including the first SEM study

Joaquín Abolafia * / Matteo Vecchi

Keywords : 18S rDNA, 28S rDNA, Description, Molecular analysis, Morphology, Phylogeny, SEM, Tarantobelinae n. subfam., Taxonomy

Citation Information : Journal of Nematology. Volume 53, Pages 1-20, DOI: https://doi.org/10.21307/jofnem-2021-080

License : (CC-BY-4.0)

Received Date : 25-April-2021 / Published Online: 01-October-2021

ARTICLE

ABSTRACT

The identity of Panagrellus pycnus, the type species of the genus Panagrellus, is discussed after studying specimens from a cultured population collected in Italy that fits the original material of the species. A new characterization is consequently provided as follows: body 0.93–1.32 mm long, lip region continuous with the adjoining body, stoma with gymnostom very reduced, pharynx with not swollen metacorpus, neck 161–203 µm long, excretory pore at level of the metacorpus, post-vulval uterine sac 99–162 µm long or 2.6–3.8 times as long as the body diameter divided in a short tubular proximal part and a long swollen distal part, vulva post-equatorial (V = 63–69), female tail conical elongate with acute terminus (133–170 µm, c = 6.8–8.1, c’ = 4.9–7.0), male tail conical elongate with acute terminus (104–137 µm, c = 7.8–10.9, c’ = 3.6–5.1), and spicules 70–81 µm long having angular hook-like and very curved ventrad lamina ending in a spatulate tip with a refringent forked axis. The evolutionary relationships of this species and the genus Panagrellus, as derived from the analyses of 18S and 28S rDNA fragments, are discussed. Additionally, the phylogenetic relationships among the members of the infraorder Panagrolaimomorpha is studied, being the genus Tarantobelus transferred to the family Panagrolaimidae and the new subfamily Tarantobelinae n. subfam. is proposed to accommodate it.

Graphical ABSTRACT

Panagrellus Thorne, 1938 is an infrequent genus belonging to the family Panagrolaimidae Thorne, 1937 (proposed as subfamily by Thorne (1937) and erected to family by Paramonov (1956)), and includes 15 species (Abolafia et al., 2016; Ivanova et al., 2018). This genus is characterized by having usually ventral curved spicules with hooked manubrium and bifurcate lamina tip. Its type species, P. pycnus, was described and succinctly illustrated by Thorne (1938) on the base of male and female specimens collected in slime secreted by wounds from the trunk of Great Plains cottonwood tree in Utah, USA. Later, Goodey (1943) transferred this species to the new erected genus Turbator Goodey, 1943 (currently junior synonym of Panagrellus together to Tylorhabdus Sukul, 1971). Andrássy (1958, 2005) and Varga (1958) reported this species from organic matter undergoing fermentation in Hungary. Hechler (1971a) revised the Thorne’s material and redescribed P. pycnus providing some line illustrations of the stoma and of the male and female posterior ends. This species is distinguished from its congeners by having spicules with angular hook-shaped manubrium and lamina very thin lacking dorsal hump and bearing spatulate terminus having forked refringent axis.

Currently, only five species of the genus [P. ceylonensis Hechler, 1971b, P. dubius Sanwal, 1960, P. levitatus, Ivanova, Perfilieva and Spiridonov, 2018, P. redivivoides Goodey, 1943 and P. redivivus (Linné, 1767) Goodey, 1945] have been characterized molecularly and used for phylogenetic analyses.

In the present paper, a cultured population of P. pycnus collected in Italy and used as food for carnivorous tardigrades is studied providing new LM and SEM illustrations and molecular data.

Materials and methods

Nematode extraction and culture

The nematodes were initially obtained from rotting pears on the ground in Borgata Città, Bologna, Italy (44°34′42.7″N 11°10′26.6″E; 27th October 2019; leg. Matteo Vecchi). Other than nematodes, the fruits were heavily colonized by Drosophila spp. and Nitidulidae beetles. Fruit pulp was spread on Yeast-Sucrose Agar (Yeast extract 1%, Sucrose 2%, Agar 1%) plates and after 1 week of incubation at 21°C a single gravid female was handpicked with a loop to start an isofemale line. Nematodes were mass cultured at 21°C in 0.3 L plastic containers on a substrate composed of 19 g of whole-grain wheat breakfast cereal (Weetabix®), 3 g of dry brewer’s yeast and 60 mL of distilled water.

Nematode processing

The specimens were killed by heating, fixed in a 70% ethanol solution, transferred to pure glycerin following the Siddiqi’s (1964) technique, and mounted on glass microscope slides with the glycerine-paraffin method (de Maeseneer and d’Herde, 1963) somewhat modified using hot liquid paraffin.

Light microscopy (LM)

Observations were made and measurements were taken using a Nikon Eclipse 80i (Nikon, Tokyo, Japan) microscope with a drawing tube (camera lucida) attached to it. Demanian indices and other ratios were calculated according to de Man (1881). Pictures were taken with a Nikon microscope equipped with differential interference contrast (DIC) optics and an associated Nikon Digital Sight DS-U1 camera. Micrographs were combined using Adobe® Photoshop® CS. The terminology used for the morphology of stoma and spicules follows De Ley et al. (1995) and Abolafia and Peña-Santiago (2017), respectively.

Scanning electron microscopy (SEM)

Specimens preserved in glycerine were selected and prepared for observation under SEM according to Abolafia (2015). They were cleaned in distilled water, dehydrated in a graded ethanol-acetone series, critical point dried, coated with gold, and observed with a Zeiss Merlin microscope (5 kV) (Zeiss, Oberkochen, Germany).

Molecular analyses

DNA extraction, PCR, and sequencing

Nematode DNA was extracted from single individuals using a modified DNA extraction and PCR assays described by Castillo et al. (2003) and Archidona-Yuste et al. (2016). The specimens were cut in small pieces using a sterilized dental needle on a clean slide with 18 ml of TE buffer (10 mM Tris-Cl + 0.5 mM EDTA; pH 9.0), transferred to a microtube and adding 2 μ l proteinase K (700 μ g/ml‒1) (Roche, Basel, Switzerland), and stored to –80°C within 15 min (for several days) until processing. The microtubes were incubated at 65°C (1 hr), then at 95°C (15 min). For DNA amplification, 3 μ l of the extracted DNA was transferred to a microtube containing: 0.6 μ l of each primer (10 mM), 3 μ l Master Mix Taq DNA Polymerase (5x Hot FirePol Blend Master Mix) and ddH2O to a final volume of 20 μ l. The primers used for amplification of the region of 18S rRNA gene were the forward primer SSU F_04 (5′-GCTTGTCTCCAAAGATTAAGCC-3′) and the reverse primer SSU R_26 (5′-CATTCTTGGCAAATGCTTTCG-3′) (Blaxter et al., 1998). The primers used for amplification of the D2-D3 region of 28S rRNA gene were the forward primer D2A (5′-ACAAGTACCGTGAGGGAAAGTTG-3′) and the reverse primer D3B (5′-TCGGAAGGAACCAGCTACTA-3′) (De Ley et al., 1999; Nunn, 1992). PCR cycle conditions were as follows: one cycle of 94°C for 15 min, followed by 35 cycles of 94°C for 45 sec + annealing temperature of 55°C for 45 sec + 72°C for 45 sec, and finally one cycle of 72°C for 5 min. After DNA amplification, 5 μ l of product was loaded on a 1% agarose gel in 0.5% Tris-acetate-EDTA (40 mM Tris, 20 mM glacial acetic acid and 2 mM EDTA; pH = 8) to verify the amplification using an electrophoresis system (Labnet Gel XL Ultra V–2, Progen Scientific, London, UK). The bands were stained with 1.25 µl RedSafe (20,000x) previously added to the agarose gel solution (25 ml). The sequencing reactions of the PCR products were performed at Sistemas Genómicos (Paterna, Valencia, Spain) according the Sanger et al. (1977) method. The DNA sequences obtained for P. pycnus (MZ656001 for the 18S rDNA and MZ656000 for the 28S rDNA) and Tarantobelus arachnicida Abolafia and Peña-Santiago, 2018 (MZ655998–MZ655999 for the 18S rDNA and MZ656002–MZ656003 for the 28S rDNA) were submitted to the GenBank database.

Phylogenetic analyses

For phylogenetic relationships, the analyses were based on 18S and 28S rDNA fragments. The newly obtained sequences were manually edited using BioEdit 7.2.6 (Hall, 1999) and aligned with other 18S and 28S rRNA gene sequences representative of Panagrolaimomorpha and closely related taxa available in GenBank (accession numbers available in Supplementary Table 1) with MAFFT ver. 7 (Katoh and Toh, 2008; Katoh et al., 2002) with the G-INS-i method (thread = 4, threadtb = 5, threadit = 0, reorder, adjustdirection, anysymbol, maxiterate = 1,000, retree 1, globalpair input). Alignments ends were trimmed using MEGA7 (Kumar et al., 2016) up to 1,851 and 1,195 bp for 18S and 28S respectively. The best-fit models of nucleotide substitution used for the phylogenetic analysis were selected using jModelTest 2.1.10 (Darriba et al., 2012). Sequences were concatenated with the R package ‘concatipede’ v1.0.0 (Vecchi and Bruneaux, 2021). The phylogenetic tree was generated with Bayesian inference method using MrBayes 3.2.6 (Ronquist et al., 2012). Plectus aquatilis was used as outgroup for the Rhabditida phylogenetic tree. The phylogenetic analyses were initiated with a random starting tree and run with the Markov Chain Monte Carlo (MCMC) (Larget and Simon, 1999) for 2 x 107 generations. The posterior tree distributions trace plots and ESS were checked with Tracer (Rambaut et al., 2018). The tree was visualized and saved with FigTree 1.4.4 (Rambaut, 2018).

Results

Panagrellus pycnus Thorne, 1938 (Figs. 1–4)

Figure 1:

Panagrellus pycnus Thorne, 1938 (line drawing). (A) Neck; (B) Anterior end; (C) Female reproductive system; (D) Entire female; (E) Female posterior end; (F) Lateral field; (G) Male posterior end; (H) Entire male.

10.21307_jofnem-2021-080-f001.jpg
Figure 2:

Panagrellus pycnus Thorne, 1938 (light microscopy). (A) Neck (anterior white arrow pointing the excretory pore, posterior white arrow pointing the hemizonid, black arrow pointing the deirid); (B) Anterior end; (C) Vagina; (D) Female reproductive system; (E) Female posterior end (arrow pointing the phasmid); (F) Lateral field; (G) Entire female; (H) Male posterior end (black arrows pointing genital papillae, GP; mid-ventral papillae, MP; white arrow pointing the phasmid, ph); (I) Entire male.

10.21307_jofnem-2021-080-f002.jpg
Figure 3:

Panagrellus pycnus Thorne, 1938 (light microscopy). (A–E) Morphological variability of spicules and gubernaculum.

10.21307_jofnem-2021-080-f003.jpg
Figure 4:

Panagrellus pycnus Thorne, 1938 (scanning electron microscopy). A–C: Lip region in lateral, ventral and frontal views, respectively (arrows pointing the amphids); D–F: Stomatal denticles; G: Lateral field; H: Excretory pore; I: Deirid; J: Vagina; K, Q: Spicules tip in lateral and dorsal views, respectively; L, M: Female posterior end in lateral and ventral views, respectively; N: Male phasmid; O, P: Male posterior end in lateral and ventral views, respectively (black arrows pointing the genital papillae, white arrow pointing the phasmid); R: Mid-ventral papillae (arrow).

10.21307_jofnem-2021-080-f004.jpg

= Turbator pycnus (Thorne, 1938) Goodey, 1943

Material examined: 10 females and 10 males obtained from culture.

Measurements

Measurements are provided in Table 1.

Table 1.

Morphometrics of Panagrellus pycnus Thorne, 1938 obtained from culture.

10.21307_jofnem-2021-080-t001.jpg

Description

Adult

Moderately slender to slender nematodes of small size, 0.93–1.32 mm long. Upon fixation, habitus nearly straight in females or somewhat curved ventral, J-shaped, in males. Cuticle 1–2 µm thick, bearing minute transverse striations, with annuli 1–2 µm wide. Lateral fields occupying 10–20% of mid-body diameter, with four longitudinal incisures or three alae. Lip region continuous with the adjacent body: lips six, separate, slightly rounded, with protruding, rounded labial and cephalic sensilla; primary and secondary axils with similar morphology, primary ones slightly deeper. Oral opening large, with smooth margin. Amphids very small, oval, located at middle length of lateral lips. Stoma panagrolaimoid, 0.7–1.1 times the lip region diameter long: cheilostom large, slightly wider than long, with strongly refringent, bar-shaped rhabdia, posteriorly thicker; gymnostom very reduced, with small rhabdia; stegostom funnel-shaped with poorly refringent rhabdia, metastegostom bearing dorsal acute rhabdia (dorsal tooth). Pharynx also panagrolaimoid: pharyngeal corpus robust, subcylindrical, 2.2–4.0 times the isthmus length, with procorpus and metacorpus not well discernible; isthmus comparatively thin; basal bulb ovoid, with both valvular apparatus and posterior haustrulum well developed. Cardia small, surrounded by intestinal tissue. Nerve ring at 70–85% of neck length from the anterior end, surrounding the anterior part of isthmus. Excretory pore at 52–75% of neck length, at level of the posterior part of metacorpus. Hemizonid located at level of isthmus. Deirids 78–90% of neck length, at level of isthmus-bulb junction. Intestine without distinct specializations, but with slightly thinner walls at cardiac part; intestinal lumen with rest of diatom frustules.

Female

Reproductive system monodelphic-prodelphic. Ovary very long, lacking flexure at post-vulval region, having oocytes arranged in several rows at its distal part and then in only one rows at its proximal part. Oviduct short, slightly longer than the body diameter developing a scarcely discernible spermatheca at its proximal part. Uterus very long, 6.8–10.9 times as long as body diameter, tubular, frequently including uterine eggs (20–33 × 28–54 µm) inside in different stages of development. Post-vulval uterine sac well developed, 2.6–3.8 times as long as the body diameter, with very thin walls, frequently poorly discernible, with proximal part short, tubular, and distal part large, swollen. Vagina extending inwards to 31–43% of body diameter, sigmoid. Vulva slightly protruding. Rectum short, 0.9–1.3 times the anal body width; three small gland-like cells are distinguishable around the intestine-rectum junction. Tail conical-elongate with acute terminus. Phasmids located at 34–40% of tail length from anus.

Male

Reproductive system monorchic, with testis reflexed ventrad anteriorly. Spicules paired and symmetrical: manubrium ventrally bent, angular hook-shaped, reduced calamus, and very curved ventrad lamina lacking dorsal hump, with well-developed ventral velum, narrower at its middle length, and spatulate tip in lateral view having a refringent forked axis. Gubernaculum well developed, slightly curved, about 0.4 times of the spicules length, with thin corpus. Three small gland-like cells are distinguishable around the beginning of the cloaca. Tail conical, slightly curved ventrad, conoid anterior to phasmid and them filiform. Genital papillae seven pairs, tree pre-cloacal (GP1 and GP2 subventral, GP3 lateral) and four post-cloacal: two pairs subventral (GP5, GP6) and one subdorsal (GP4) at the middle of tail length and one subdorsal (GP7) at beginning of the filiform part. One mid-ventral adcloacal papilla (MP) present. Phasmids at 32 to 50% of tail length from the cloacal aperture, close to genital papillae GP7.

Remarks

The material examined in this study agrees well with the type population of P. pycnus described by Thorne (1938), especially in the morphology of the spicules although, unfortunately, Thorne (op. cit.) did not provide their measurements. Apparently, according to the drawing illustrations, the author did not observe the post-vulval uterine sac which is now know to be very large but (in the specimens examined in this study) its walls are very thin and sometimes not very well discernible. Also, the vagina structure was not illustrated with precision. Later, Hechler (1971a) described lectotype specimens of this species and observed the presence of post-vulval uterine sac but it was not described or illustrated. The main characters to identify this species, the spicules morphology (with manubrium having dorsal angular side and ventral hook-like side; Fig. 4) and morphometry (56–61 vs 54–70 µm, measured as a straight line, or chord, connecting the spicule manubrium with the lamina tip) agree perfectly with P. pycnus, while the gubernaculum is slightly longer (29–32 vs 25–27 µm). Unfortunately, most of the measurements were not provided in the previous records of the species (Table 2).

Table 2.

Comparative morphometrics of Panagrellus pycnus Thorne, 1938.

10.21307_jofnem-2021-080-t002.jpg

On the other hand, the most similar species to P. pycnus is P. leperisini Massey, 1974, with which it could be confused by having similar spicules morphology. However, P. pycnus presents larger body (0.80–1.4 mm in Thorne’s description, 0.90–1.41 in Hechler’s description and 0.93–1.32 mm in the present paper vs 0.74–0.97 mm), lip region wider (wider than the adjacent part of body vs narrower) and longer spicules (70–81 vs 60 µm).

Diagnosis

Panagrellus pycnus is characterized by having 0.93–1.32 mm long body, lip region continuous with the adjoining body, lips separated six having rounded sensilla, amphids small, stoma with gymnostom very reduced, pharynx with not swollen metacorpus, neck 161–203 µm long, excretory pore at level of the metacorpus, female reproductive system monodelphic-prodelphic with post-vulval uterine sac 99–162 µm long or 2.6–3.8 times as long as the body diameter divided in a short tubular proximal part and a long swollen distal part, vulva post-equatorial (V = 63–69), female tail conical elongate with acute terminus (133–170 µm, c = 6.8–8.1, c′ = 4.9–7.0), male tail conical elongate with acute terminus (104–137 µm, c = 7.8–10.9, c′ = 3.6–5.1), spicules 70–81 µm long having angular hook-shaped and very curved ventrad lamina ending in a spatulate tip with a refringent forked axis, and gubernaculum 29–32 µm long.

Differential diagnosis

The Panagrellus species are easily differentiated by the size (Table 3) and morphology (Fig. 5) of the spicules (Abolafia et al., 2016). Three main groups are distinguished according the morphology of the spicules. The first group, (Fig. 5B–D) with more simple spicules, includes three species [P. dorsobidentatus (Rühm, 1956) Baker, 1962, P. ludwigi (de Man, 1910) Goodey, 1945 and P. ventrodentatus (Heindl-Mengert, 1956) Baker, 1962] having robust spicules with irregular truncated manubrium, probably a plesiomorphic character. The second group (Fig. 5E–I), with five species [P. japonicus (Yokoo and Ota, 1961) Andrássy, 1984 P. levitatus P. nepenthicola (Menzel, 1922) Goodey, 1945, P. redivivoides (Goodey, 1943) Goodey, 1945 and, probably, P. ulmi Abolafia, Alizadeh and Khakver, 2016], presents ventrally curved spicules with rounded manubrium, straight or slightly ventral bent, and lamina with variable bifurcated tip. The third group (Fig. 5J–P), composed by seven species [P. ceylonensis, P. dubius, P. filiformis (Sukul, 1971) Andrássy, 1984 P. leperisini, P. pycnus, P. redivivus and P. silusioides Tsalolikhin, 1965], have ventrally curved spicules with ventral bent manubrium and lamina with well-developed bifurcated tip, probably an apomorphic character. Between this last group, the curvature degree of the manubrium is a diagnostic character. Thus, spicules with spirally curved manubrium appear in P. ceylonensis (with scarcely sigmoid lamina and short bifurcated terminus), P. dubius (with very curved lamina and long bifurcated terminus) and P. filiformis (with curved lamina while the terminus was not well illustrated). Manubrium poorly ventrally bent appears in P. redivivus and P. silusioides, having scarcely to very bent manubrium respectively, and slightly developed dorsal hump at lamina, while hook-shaped manubrium appears in some P. ceylonensis (narrow manubrium and poorly curved lamina), P. leperisini and P. pycnus (both species with wide manubrium and ‘C’-like lamina), more ventrally curved in P. ceylonensis, while P. pycnus is more anteriorly curved than P. leperisini.

Table 3.

Length of the spicules (measured in arc) of the Panagrellus species with hooked manubrium.

10.21307_jofnem-2021-080-t003.jpg
Figure 5:

Morphology of the spicules and gubernacula of the species of the genera Baujardia Bert, Tandingan De Ley, Van Driessche, Segers and De Ley, 2003 and Panagrellus Thorne, 1938. (A) Baujardia; (B–D) Panagrellus species with truncated manubrium; (E–I) Panagrellus species with rounded or conoid manubrium; (E–P) Panagrellus species with curved or hook-shaped manubrium (c = calamus, dh = dorsal hump, l = lamina, m = manubrium, v = velum, t = tip).

10.21307_jofnem-2021-080-f005.jpg

Molecular characterization and phylogenetic position

One 928 bp 18S rDNA sequence (GenBank accession number MZ656001) and one 766 bp 28S rDNA sequence (GenBank accession number MZ656000) were obtained for P. pycnus. After the molecular analysis, P. pycnus exhibits a high level of rDNA similarity with other Panagrellus species (Fig. 6).

Figure 6:

Bayesian Inference tree showing the phylogenetic position of Panagrellus pycnus and its related taxa based on a concatenated 18S and 28S rDNA regions. Bayesian posterior probabilities (pp) are given for each clade. Nodes with pp < 0.70 were collapsed. When pp = 1, its value is not shown. Scale bar shows the number of substitutions per site.

10.21307_jofnem-2021-080-f006.jpg

Discussion

On the identity of Panagrellus pycnus and other related species

The morphology and morphometry of the material examined now agree with the type population of Panagrellus pycnus described by Thorne (1938) and redescribed by Hechler (1971a, b). The main characters to distinguish this material from other species are the size of the spicules (70–81 µm length and reaching the GP1) and the presence of a manubrium with angular dorsal side and hook-like ventral side. Two species, P. ceylonensis and P. leperisini, are similar to P. pycnus, but the former species differs in size (70–81 µm reaching the GP1 vs 81–89 µm length reaching the GP1 in P. ceylonensis and 56–62 µm length not reaching the GP1 in P. leperisini) and morphology (lamina ventrally more curved at first third vs homogeneously curved but ventrally convex at its mid-length in P. ceylonensis and slightly ventrally more curved anterior and posteriorly in P. leperisini).

Other species have very similar spicules and could be confused with P. pycnus. For example, some populations of P. pycnus could have been confused with P. redivivus. Thus, de Man (1914, as Anguillula silusiae), Goodey (1943), Hechler (1970, 1971a, Zullini (1982) and Ferris (2009) described several populations of Panagrellus agreeing with P. redivivus, showing specimens with spicules that do not reach the pre-cloacal GP1 and possessing a dorsal hump with a slight ventral bend to the calamus. However, the material examined by Corrêa de Carvalho and Álvares Corrêa (1953) and Rühm (1956) does not agree well with R. redivivus as observed Hechler (1971a); thus, the material described by Corrêa de Carvalho and Álvares Corrêa (op. cit.) shows spicules reaching the GP1 (vs not reaching the GP1 in P. redivivus, see de Man, 1914 as P. silusiae (Aubertot, 1925; Goodey, 1922; Hechler, 1971a) having almost angular hook-like manubrium; on the other hand, the material described by Rühm (op. cit.) is very similar to P. ceylonensis agreeing in measurements, excretory pore at basal bulb level and spicules slightly sigmoid with narrow hooked manubrium.

On the other hand, Stock and Nadler (2006) characterized morphological and molecularly three species of the genus Panagrellus. However, there are some discrepancies about their identity. With respect to the morphology of the illustrated spicules, the spicule named as ‘P. ceylonensis’ (see Fig. 3B) is more similar to P. redivivus and P. silusioides by having ventral bent manubrium and wide velum; the spicule named as ‘P. redivivus’ (see Fig. 3D) agrees with P. ceylonensis (although it is unusually short, 65 µm long) and P. parasiticus (junior synonym of P. redivivus) by having spirally curved manubrium and narrow velum; and the spicule named as ‘P. dubius’ (see Fig. 3E), agreeing with this species by having long bifurcated terminus but also it is similar to P. leperisini by the size and by having angular hook-like manubrium. However, other morphological and morphometrical characters are necessary to correctly identify these species.

On the phylogenetic position of Panagrellus pycnus in the genus Panagrellus

The material examined now of Panagrellus pycnus is phylogenetically related with other sequenced species of the genus Panagrellus. The species included in this genus are characterized by having very short gymnostom with minute rhabdia, post-vulval uterine sac differentiated in a short tubular proximal part and a large swollen distal part, and female and male tails conoid-elongate. An important character of the spicules is the ventral curvature and hook-shaped manubrium. There are two primary clades in the rDNA phylogenetic tree of Panagrellus (Fig. 6). One includes the species P. levitatus and P. redivivoides and is characterized by the more robust spicules with rounded manubrium, like other genera of the family Panagrolaimidae. The second clade includes P. ceylonensis, P. dubius, P. pycnus and P. redivivus and is characterized by thinner and elongate spicules, increasing the curvature of the manubrium from slightly ventrally curved to hook-like.

Phylogenetic position of the genus Panagrellus and its related genera

Panagrellus is recovered as paraphyletic (Fig. 6) with Baujardia Bert, Tandingan De Ley, Van Driessche, Segers and De Ley, 2003 located inside the genus. Indeed, Baujardia is very similar to Panagrellus, only differentiated from it by the morphology of the stoma, slightly longer in Baujardia. Unfortunately, only a sequence based on 18S rDNA was obtained of Baujardia mirabilis Bert, Tandingan De Ley, Van Driessche, Segers and De Ley, 2003, the only species of the genus, leading to the possible consideration of the synonymy of Baujardia.

With respect to the subfamily Panagrellinae Andrássy, 1976 containing the genera Baujardia and Panagrellus, is shown as monophyletic. This subfamily was synonymized with Baujardinae Andrássy, 2005 by Abolafia et al. (2016) and the present molecular analysis agree with these authors.

Other subfamilies of the family Panagrolaimidae as Medibullinae Siddiqi, 1993 [tranferred to Panagrolaimidae by Abolafia and Peña-Santiago (2018), including Shahnema Siddiqi, 2014] Tricephalobinae Andrássy, 1976 (including Halicephalobus Timm, 1956 and Turbatricinae Goodey, 1943 (including Turbatrix Peters, 1927) are shown as monophyletic.

Conversely, the subfamily Panagrolaiminae is shown as polyphyletic. Four genera belonging to this subfamily, Panagrobelus Thorne, 1939, Panagrolaimus Fuchs, 1930 Procephalobus Steiner, 1934 and Propanagrolaimus Andrássy, 2005 appear not directly related, unlike what was proposed by Andrássy (1984, 2005). However, additional morphological and molecular studies are necessary to confirm the relationships between these genera and, after that, it could be suitable to reestablish the subfamily Procephalobinae Paramonov, 1956 to include Procephalobus and Propanagrolaimus, among others.

Nevertheless, the genus Macrolaimus Maupas, 1900 currently belonging to the family Chambersiellidae Thorne, 1937 subfamily Macrolaiminae Sanwal, 1971, appears related with Panagrolaimus. Unfortunately, not enough species have been sequenced to clarify its phylogenetic position. To this respect, Kitagami et al. (2019) sequenced a species identified as Panagrolaimus sp. (LC382049), however, it agrees with some species of the genus Macrolaimus according the stoma morphology being showed in the present phylogenetic tree belonging to this last genus (see Supplementary Fig. S1 in Kitagami et al., op. cit.).

On the other hand, the genera Plectonchus Fuchs, 1930 and Tarantobelus Abolafia and Peña-Santiago, 2018 proposed as belonging to the family Brevibuccidae Paramonov, 1956 by Andrássy (1976, 1984, 2005) and Abolafia and Peña-Santiago (2018), respectively, appear not closely related with this family in this new study. According to this, both genera are transferred to the family Panagrolaimidae. Regarding the genus Plectonchus (with post-equatorial vulva and without post-vulval uterine sac) appears related to the panagrolaimid genus Panagrobelus (with equatorial vulva and very short post-vulval uterine sac); however with only limited 18S rDNA sequence in support of the genus, there is a clear need to obtain detailed morphological and molecular data, without which the genus should be considered incertae sedis until to obtain a detailed morphological-molecular study of its species. Secondly, the genus Tarantobelus appears forming part of a well-supported, separate clade which is proposed now as a new subfamily, Tarantobelinae n. subfam., clearly differentiated from other subfamilies, especially by having short and robust pharyngeal isthmus and vulva post-equatorial. This new subfamily is composed by the only genus Tarantobelus.

Taxonomic account

Tarantobelinae n. subfam.

Diagnosis: small body with cuticle nearly smooth, lateral field inconspicuous, lip region with six separated lips, each bearing a small cuticular flap topping it, panagrolaimoid stoma with well-developed gymnostom having broad and refringent rhabdia, panagrolaimoid pharynx with isthmus robust and slightly longer than basal bulb, excretory pore situated at level of isthmus, female reproductive system monodelphic-prodelphic, length of post-vulval sac less than the corresponding body diameter, vulva post-equatorial and distinctly protruding, female tail conical with acute tip, male tail conical with a long and thin mucro, spicules curved ventrad with rounded manubrium, and thick gubernaculum.

Table S1.

GenBank accession numbers of the species used in the phylogenetic tree.

10.21307_jofnem-2021-080-t0s1.jpg

Acknowledgments

Financial support was received from the Research Support Plans ‘PAIUJA 2019/2020: EI_RNM02_2019’ and ‘PIUJA 2021/2022: EI_RNM02_2021’ of the University of Jaén, Spain. SEM pictures were obtained with the assistance of technical staff (Amparo Martínez-Morales and Alba N. Ruiz-Cuenca) and equipment of ‘Centro de Instrumentación Científico-Técnica (CICT)’ from University of Jaén. MV was supported by the Academy of Finland (Fellowship #314219 to Sara Calhim, University of Jyvaskyla).

References


  1. Abolafia, J. 2015. A low-cost technique to manufacture a container to process meiofauna for scanning electron microscopy. Microscopy Research and Technique 78:771–776, doi: 10.1002/jemt.22538.
  2. Abolafia, J. and Peña-Santiago, R. 2017. On the identity of Chiloplacus magnus Rashid and Heyns, 1990 and C. insularis Orselli and Vinciguerra, 2002 (Rhabditida: Cephalobidae), two confusable species. Nematology 19:1017–1034, doi: 10.1163/15685411–00003104.
  3. Abolafia, J. and Peña-Santiago, R. 2018. Morphological and molecular characterization of Tarantobelus arachnicida gen. n., sp. n. (Nematoda, Rhabditida, Brevibuccidae), a parasitic nematode of tarantulas. Journal of Helminthology 92:491–503, doi: 10.1017/S0022149X17000566.
  4. Abolafia, J. , Alizadeh, M. and Khakvar, R. 2016. Description of Panagrellus ulmi sp. n. (Rhabditida, Panagrolaimidae) from Iran, and comments on the species of the genus and its relatives. Zootaxa 4162:245–267, doi: 10.11646/zootaxa.4162.2.3.
  5. Andrássy, I. 1958. Szabadonélő fonálférgek (Nematoda libera). In the series: Magyarország állatvilága, 3. Kötot, 1, Nemathelminthes-Archipodiata. 1. Akadémiai Kiadó (Fauna Hungariae, 36), Budapest: 362 pp.
  6. Andrássy, I. 1976. Evolution as a basis for the systematization of nematodes. Pitman Publishing, London: 288 pp.
  7. Andrássy, I. 1984. Klasse Nematoda (Ordnungen Monhysterida, Desmoscolecida, Araeolaimida, Chromadorida, Rhabditida). In the series: Bestimmungsbücher zur Bodenfauna Europas, No. 9. Berlin, Deutschland, Akademie Verlag: 509 pp.
  8. Andrássy, I . 2005. Free-living nematodes of Hungary (Nematoda errantia). Volume 1. In the series: Pedozoologica Hungarica, No. 3. Hungarian Natural History Museum, Budapest, 518 pp.
  9. Archidona-Yuste, A. , Navas-Cortés, J. A. , Cantalapiedra-Navarrete, C. , Palomares-Rius, J. E. and Castillo, P. 2016. Unravelling the biodiversity and molecular phylogeny of needle nematodes of the genus Longidorus (Nematoda: Longidoridae) in olive and a description of six new species. PLoS ONE 11:e0147689, doi: 10.1371/journal.pone.01447689.
  10. Aubertot, M. 1925. Nématodes d’Alsace. Observations sur l’Anguillule de la bière (Anguillula silusiae de Man 1914). Bulletin de la Association Philomathique d’Alsace et de Lorraine 6:333–342.
  11. Baker, A. D. 1962. Check lists of the nematode superfamilies Dorylaimoidea, Rhabditoidea, Tylenchoidea and Aphelenchoidea Leiden: Brill, p. 261.
  12. Bert, W. , Tandingan De Ley, I. , van Driessche, R. , Segers, H. and De Ley, P. 2003. Baujardia mirabilis gen. n., sp. n. from pitcher plants and its phylogenetic position within Panagrolaimidae (Nematoda: Rhabditida). Nematology 5:405–420, doi: 10.1163/156854103769224395.
  13. Blaxter, M. L. , De Ley, P. , Garey, J. R. , Liu, L. X. , Scheldeman, P. , Vierstraete, A. , Vanfleteren, J. R. , Machey, L. Y. , Dorris, M. , Frisse, L. M. , Vida, J. T. and Thomas, W. K. 1998. A molecular evolutionary framework for the phylum Nematoda. Nature 392:71–75, doi: 10.1038/32160.
  14. Castillo, P. , Vovlas, N. , Subbotin, S. A. and Troccoli, A. 2003. A new root-knot nematode, Meloidogyne baetica n. sp. (Nematoda: Heteroderidae), parasitizing wild olive in Southern Spain. Phytopathology 93:1093–1102, doi: 10.1094/PHYTO.2003.93.9.1093.
  15. Corrêa de Carvalho, J. and Álvares Corrêa, M. O. 1953. A ocorrência de nematoides em massa de tomate. Revista do Instituto Adolfo Lutz 13:37–44.
  16. Darriba, D. , Taboada, G. L. , Doallo, R. and Posada, D. 2012. jModelTest 2: more models, new heuristics and parallel computing. Nature Methods 9:772, doi: 10.1038/nmeth.2109.
  17. De Ley, P. , Felix, A. M. , Frisse, L. M. , Nadler, S. A. , Sternberg, P. W. and Thomas, W. K. 1999. Molecular and morphological characterization of two reproductively isolated species with mirror-image anatomy (Nematoda: Cephalobidae). Nematology 1:591–612, doi: 10.1163/156854199508559.
  18. De Ley, P. , van de Velde, M. C. , Mounport, D. , Baujard, P. and Coomans, A. 1995. Ultrastructure of the stoma in Cephalobidae, Panagrolaimidae and Rhaditidae, with a proposal for a revised stoma terminology in Rhabditida (Nematoda). Nematologica 41:153–182, doi: 10.1163/003925995X00143.
  19. de Maeseneer, J. and d’Herde, J. 1963. Méthodes utilisées por l'étude des anguillules libres du sol. Revue d’Agriculture 16:441–447.
  20. de Man, J. G. 1881. Die einheimischen, frei in der reinen Erde und im süssen Wasser lebenden Nematoden. Vorläufiger Bericht und descriptivsystematischer Theil. Tijdschrift van der Nederlandsche dierkundige Vereeniging 5:1–104.
  21. de Man, J. G. 1910. Beiträge zur Kenntnis der in dem weißen Schleimfluß der Eichen lebeden Anguilluliden, nebst Untersuchungen über den Bau des Essigälchens und der Gattung Anguillula Ehrb. Zoologische Jahrbüch Systematik 29:359–394.
  22. de Man, J. G. 1914. Anguillula silusiae de Man, eine neue in der sogenannten Bierfilzen lebende Art der Gattung Anguillula Ehrb. Annales de la Sociéte Royale Zoologique et Malacologique de Belgique 48 (Vol. Jubilaire): 1–9.
  23. Ferris, H. 2009. The beer mat nematode, Panagrellus redivivus: A study of the connectedness of scientific discovery. Nematode Morphology and Systematics 12:19–25.
  24. Fuchs, G. 1930. Neue an Borken- und Rüsselkäfergebundene Nematoden, halbparasitische und Wohnungseinmieter. Freilebende Nematoden aus Moos und Walderde in Borken-und Rüsselkäfergängen. Zoologische Jahrbücher (Systematik) 59:505–646.
  25. Goodey, T. 1922. The eel-worm in paper-hangers’ paste [Anguillula rediviva (Linnaeus, 1767), Stiles and Hassall, 1905. Annals and Magazine of Natural History (Series 9) 10:297–307, doi: 10.1080/00222932208632783.
  26. Goodey, T. 1943. On the systematic relationship of the vinegar eelworm, Turbatrix aceti and its congeners, with a description of a new species. Journal of Helminthology 21:1–9, doi: 10.1017/S0022149X00031825.
  27. Goodey, T. 1945. A note on the subfamily Turbatricinae and the genus Turbator Goodey, 1943. Journal of Helminthology 21:69–70, doi: 10.1017/S0022149X00031953.
  28. 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, doi: 10.1021/bk–1999-0734.ch008.
  29. Hechler, H. C. 1970. Reproduction, chromosome number, and postembryonic development of Panagrellus redivivus (Nematoda: Cephalobidae). Journal of Nematology 2:355–361.
  30. Hechler, H. C. 1971a. Taxonomic notes on four species of Panagrellus Thorne (Nematoda, Cephalobidae). Journal of Nematology 3:227–237.
  31. Hechler, H. C. 1971b. Panagrellus ceylonensis n. sp. (Nematoda, Cephalobidae). Journal of Nematology 3:246–250.
  32. Heindl-Mengert, H. 1956. Die Nematodenfauna im Schleimfluss lebender Laubbaüme. Sitzungberichte der Physikalisch Medizinischen Sozietät Erlangen 77:158–176.
  33. Ivanova, E. , Perfilieva, K. and Spiridonov, S. 2018. Panagrellus levitatus sp. n. (Rhabditida: Panagrolaimidae), a nematode suppressing Drosophila melanogaster in laboratory cultures. Nematology 20:285–297, doi: 10.1163/15685411-00003141.
  34. Katoh, K. and Toh, H. 2008. Recent developments in the MAFFT multiple sequence alignment program. Briefings in Bioinformatics 9:286–298, doi: 10.1093/bib/bbn013.
  35. Katoh, K. , Misawa, K. , Kuma, K. and Mitaya, T. 2002. MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Research 30:3059–3066, doi: 10.1093/nar/gkf436.
  36. Kitagami, Y. , Kanzaki, N. , Tanikawa, T. and Matsuda, Y. 2019. Free-living nematodes associated with pine cones of Pinus thunbergii and P. taeda at Japanese coastal and inland forest sites. Nematology 21:389–400, doi: 10.1163/15685411-00003221.
  37. Kumar, S. , Stecher, G. and Tamura, K. 2016. MEGA7: Molecular Evolutionary Genetics Analysis Version 7.0 for bigger datasets. Molecular Biology and Evolution 33:1870–1874, doi: 10.1093/molbev/msw054.
  38. Larget, B. and Simon, D. L. 1999. Markov chain Monte Carlo algorithms for the Bayesian analysis of phylogenetic trees. Molecular Biology and Evolution 16:750–759, doi: 10.1093/oxfordjournals.molbev.a026160.
  39. Linné, C. von 1767. “Systema naturae sive regna tria naturae, secundum classes, ordines, genera, species, cum characteribus, differentiis, synonymis, locis”, Vol. 1. Part 2, 12th Ed. Laurentii Salvii, Holmiae 795; [pp. 533–1327].
  40. Massey, C. L. 1974. Biology and taxonomy of nematode parasites and associates of bark beetles in the Unites States. Agriculture Handbook, 446 (Forest Service, U.S. Department of Agriculture):1–233.
  41. Maupas, E. F. 1900. Modes et formes de reproduction des nématodes. Archives de Zoologie Expérimentale et Générale 8:463–624.
  42. Menzel, R. 1922. Beiträge zur kenntnis der Mikrofauna von Niederländisch-ost-Indien. Treubia 3:116–126.
  43. Nunn, G. B. 1992. Nematode molecular evolution. An investigation of evolutionary patterns among nematodes based upon DNA sequences. Ph.D. dissertation, University of Nottingham, 228 pp.
  44. Paramonov, A. A. 1956. K revizii sistemy Rhabditat rastenii [On the revision of the Rhabditat system of plants] (In Russian). Trudy Gelmintologicheskoi Laboratorii Akademiya Nauk SSSR 8:77–111.
  45. Peters, B. G. 1927. On the nomenclature of the vinegar eelworn. Journal of Helminthology 5:133–142.
  46. Rambaut, A. 2018. Figtree, a graphical viewer of phylogenetic trees, version 1.4.4. Institute of Evolutionary Biology, University of Edinburgh, Edinburgh, available at: https://github.com/rambaut/figtree/releases/tag/v1.4.4.
  47. Rambaut, A. , Drummond, A. J. , Xie, D. , Baele, G. and Suchard, M. A. 2018. Posterior summarization in Bayesian phylogenetics using Tracer 1.7. Systematic Biology 67:901, doi: 10.1093/sysbio/syy032.
  48. Ronquist, F. , Teslenko, M. , van der Mark, P. , Ayres, D. L. , Darling, A. , Höhna, S. , Larget, B. , Liu, L. , Suchard, M. A. and Huelsenbeck, J. P. 2012. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic Biology 61:539–542, doi: 10.1093/sysbio/sys029.
  49. Rühm, W. 1956. Die Nematoden der Ipiden. Parasitologische Schriftenreihe 6:1–437.
  50. Sandground, J. H. 1939. Cephalobus parasiticus n. sp. and “Pseudostrongyloidiasis” in Macaca irus mordax . Parasitology 31:132–137.
  51. Sanger, F. , Nicklen, S. and Coulson, A. R. 1977. DNA sequencing with chain-terminating inhibitors. Proceedings of the National Academy of Sciences U.S.A 74:5463–5467, doi: 10.1073/pnas.74.12.5463.
  52. Sanwal, K. C. 1960. Panagrellus dubius n. sp. (Nematoda, Turbatricinae Goodey, 1943), from frass of beetle Sternochetus lapathi (L.), with remarks on redescriptions of Anguillula rediviva (L., 1767). Canadian Journal of Zoology 38:1041–1046, doi: 10.1139/z60-108.
  53. Sanwal, K. C. 1971. Geraldius n. gen., Macrolaiminae n. subfam., with a revision of the subfamilies and genera of Chambersiellidae (Nematoda). Canadian Journal of Zoology 49:965–967, doi: 10.1139/z71-147.
  54. Siddiqi, M. R. 1964. Studies on Discolaimus spp. (Nematoda: Dorylaimidae) from India. Zeitschrift für zoologische Systematik und Evolutionsforschung 2:174–184, doi: 10.1111/j.1439–0469.1964.tb00720.x.
  55. Siddiqi, M. R. 1993. Nematodes of tropical rainforests: 2. Five new genera and eight new species of cephalobs. Afro-Asian Journal of Nematology 3:212–25.
  56. Siddiqi, M. R. 2014. Descriptions of Shahnema typicum gen. n., sp. n. and S. cabii gen. n., sp. n. (Diplogastrina: Shahnematidae fam. n.). International Journal of Nematology 24:153–160.
  57. Steiner, G. 1934. Procephalobus mycophilus n. g., n. sp. (Cephalobidae), a nematode living in the sclerotia of the fungus Balansia claviceps . Proceedings of the helminthological Society of Washington 1:54–56.
  58. Steiner, G. 1936. Opuscula miscellanea nematologica, III. Proceedings of the Helminthological Society of Washington 3:16–22.
  59. Stock, S. P. and Nadler, S. A. 2006. Morphological and molecular characterisation of Panagrellus spp. (Cephalobina: Panagrolaimidae): taxonomic status and phylogenetic relationships. Nematology 8:921–938, doi: 10.1163/156854106779799277.
  60. Sukul, N. C. 1971. New free-living nematodes from the thermal spring of Tantloya, Bihar. Indian Journal of Helminthology 23:135–145.
  61. Thorne, G. 1937. A revision of the nematode family Cephalobidae Chitwood and Chitwood, 1934. Proceedings of the Helminthological Society of Washington 4:1–16.
  62. Thorne, G. 1938. Notes on free-living and plant-parasitic nematodes. IV. Proceedings of the Helminthological Society of Washington 5:64–65.
  63. Thorne, G. 1939. Notes on free-living and plant-parasitic nematodes. V. Proceedings of the helminthological Society of Washington 6:30–32.
  64. Timm, R. W. 1956. Marine nematodes from the Bay of Bengal. I. Phasmidea. Journal of the Bombay Natural History Society 54:87–90.
  65. Tsalolikhin, S. J. 1965. Novaja saprobiotischeskaja nematoda Panagrellus silusioides sp. n. Vestnik Leningradskogo Gosudarstvennogo Universiteta 21:152–153.
  66. Varga, M. 1958. Panagrellus pycnus. Európában eddig nem jelzett fonálféregfaj első jelentkezése. Élelmezési Ipar 12:74–80.
  67. Vecchi, M. and Bruneaux, M. 2021. Concatipede: an R package to concatenate fasta sequences easily, Available at: http://doi.org/10.5281/zenodo.5130604.
  68. Yokoo, T. and Ota, Y. 1961. On the variations of dimensions within soil nematodes. III. Dimensions of the Brevibucca japonica n. sp. found in the decayed fruit of pear in Japan, with descriptions of this new species. Agricultural Bulletin of Saga University 12:149–156.
  69. Zullini, A. 1982. Nematodi (Nemata). In the series: Guide per il riconoscimento delle specie animali delle acque interne italiane. Volume 17. Consiglio Nazionale delle Ricerche, Rome: 117 pp.
XML PDF Share

FIGURES & TABLES

Figure 1:

Panagrellus pycnus Thorne, 1938 (line drawing). (A) Neck; (B) Anterior end; (C) Female reproductive system; (D) Entire female; (E) Female posterior end; (F) Lateral field; (G) Male posterior end; (H) Entire male.

Full Size   |   Slide (.pptx)

Figure 2:

Panagrellus pycnus Thorne, 1938 (light microscopy). (A) Neck (anterior white arrow pointing the excretory pore, posterior white arrow pointing the hemizonid, black arrow pointing the deirid); (B) Anterior end; (C) Vagina; (D) Female reproductive system; (E) Female posterior end (arrow pointing the phasmid); (F) Lateral field; (G) Entire female; (H) Male posterior end (black arrows pointing genital papillae, GP; mid-ventral papillae, MP; white arrow pointing the phasmid, ph); (I) Entire male.

Full Size   |   Slide (.pptx)

Figure 3:

Panagrellus pycnus Thorne, 1938 (light microscopy). (A–E) Morphological variability of spicules and gubernaculum.

Full Size   |   Slide (.pptx)

Figure 4:

Panagrellus pycnus Thorne, 1938 (scanning electron microscopy). A–C: Lip region in lateral, ventral and frontal views, respectively (arrows pointing the amphids); D–F: Stomatal denticles; G: Lateral field; H: Excretory pore; I: Deirid; J: Vagina; K, Q: Spicules tip in lateral and dorsal views, respectively; L, M: Female posterior end in lateral and ventral views, respectively; N: Male phasmid; O, P: Male posterior end in lateral and ventral views, respectively (black arrows pointing the genital papillae, white arrow pointing the phasmid); R: Mid-ventral papillae (arrow).

Full Size   |   Slide (.pptx)

Figure 5:

Morphology of the spicules and gubernacula of the species of the genera Baujardia Bert, Tandingan De Ley, Van Driessche, Segers and De Ley, 2003 and Panagrellus Thorne, 1938. (A) Baujardia; (B–D) Panagrellus species with truncated manubrium; (E–I) Panagrellus species with rounded or conoid manubrium; (E–P) Panagrellus species with curved or hook-shaped manubrium (c = calamus, dh = dorsal hump, l = lamina, m = manubrium, v = velum, t = tip).

Full Size   |   Slide (.pptx)

Figure 6:

Bayesian Inference tree showing the phylogenetic position of Panagrellus pycnus and its related taxa based on a concatenated 18S and 28S rDNA regions. Bayesian posterior probabilities (pp) are given for each clade. Nodes with pp < 0.70 were collapsed. When pp = 1, its value is not shown. Scale bar shows the number of substitutions per site.

Full Size   |   Slide (.pptx)

REFERENCES

  1. Abolafia, J. 2015. A low-cost technique to manufacture a container to process meiofauna for scanning electron microscopy. Microscopy Research and Technique 78:771–776, doi: 10.1002/jemt.22538.
  2. Abolafia, J. and Peña-Santiago, R. 2017. On the identity of Chiloplacus magnus Rashid and Heyns, 1990 and C. insularis Orselli and Vinciguerra, 2002 (Rhabditida: Cephalobidae), two confusable species. Nematology 19:1017–1034, doi: 10.1163/15685411–00003104.
  3. Abolafia, J. and Peña-Santiago, R. 2018. Morphological and molecular characterization of Tarantobelus arachnicida gen. n., sp. n. (Nematoda, Rhabditida, Brevibuccidae), a parasitic nematode of tarantulas. Journal of Helminthology 92:491–503, doi: 10.1017/S0022149X17000566.
  4. Abolafia, J. , Alizadeh, M. and Khakvar, R. 2016. Description of Panagrellus ulmi sp. n. (Rhabditida, Panagrolaimidae) from Iran, and comments on the species of the genus and its relatives. Zootaxa 4162:245–267, doi: 10.11646/zootaxa.4162.2.3.
  5. Andrássy, I. 1958. Szabadonélő fonálférgek (Nematoda libera). In the series: Magyarország állatvilága, 3. Kötot, 1, Nemathelminthes-Archipodiata. 1. Akadémiai Kiadó (Fauna Hungariae, 36), Budapest: 362 pp.
  6. Andrássy, I. 1976. Evolution as a basis for the systematization of nematodes. Pitman Publishing, London: 288 pp.
  7. Andrássy, I. 1984. Klasse Nematoda (Ordnungen Monhysterida, Desmoscolecida, Araeolaimida, Chromadorida, Rhabditida). In the series: Bestimmungsbücher zur Bodenfauna Europas, No. 9. Berlin, Deutschland, Akademie Verlag: 509 pp.
  8. Andrássy, I . 2005. Free-living nematodes of Hungary (Nematoda errantia). Volume 1. In the series: Pedozoologica Hungarica, No. 3. Hungarian Natural History Museum, Budapest, 518 pp.
  9. Archidona-Yuste, A. , Navas-Cortés, J. A. , Cantalapiedra-Navarrete, C. , Palomares-Rius, J. E. and Castillo, P. 2016. Unravelling the biodiversity and molecular phylogeny of needle nematodes of the genus Longidorus (Nematoda: Longidoridae) in olive and a description of six new species. PLoS ONE 11:e0147689, doi: 10.1371/journal.pone.01447689.
  10. Aubertot, M. 1925. Nématodes d’Alsace. Observations sur l’Anguillule de la bière (Anguillula silusiae de Man 1914). Bulletin de la Association Philomathique d’Alsace et de Lorraine 6:333–342.
  11. Baker, A. D. 1962. Check lists of the nematode superfamilies Dorylaimoidea, Rhabditoidea, Tylenchoidea and Aphelenchoidea Leiden: Brill, p. 261.
  12. Bert, W. , Tandingan De Ley, I. , van Driessche, R. , Segers, H. and De Ley, P. 2003. Baujardia mirabilis gen. n., sp. n. from pitcher plants and its phylogenetic position within Panagrolaimidae (Nematoda: Rhabditida). Nematology 5:405–420, doi: 10.1163/156854103769224395.
  13. Blaxter, M. L. , De Ley, P. , Garey, J. R. , Liu, L. X. , Scheldeman, P. , Vierstraete, A. , Vanfleteren, J. R. , Machey, L. Y. , Dorris, M. , Frisse, L. M. , Vida, J. T. and Thomas, W. K. 1998. A molecular evolutionary framework for the phylum Nematoda. Nature 392:71–75, doi: 10.1038/32160.
  14. Castillo, P. , Vovlas, N. , Subbotin, S. A. and Troccoli, A. 2003. A new root-knot nematode, Meloidogyne baetica n. sp. (Nematoda: Heteroderidae), parasitizing wild olive in Southern Spain. Phytopathology 93:1093–1102, doi: 10.1094/PHYTO.2003.93.9.1093.
  15. Corrêa de Carvalho, J. and Álvares Corrêa, M. O. 1953. A ocorrência de nematoides em massa de tomate. Revista do Instituto Adolfo Lutz 13:37–44.
  16. Darriba, D. , Taboada, G. L. , Doallo, R. and Posada, D. 2012. jModelTest 2: more models, new heuristics and parallel computing. Nature Methods 9:772, doi: 10.1038/nmeth.2109.
  17. De Ley, P. , Felix, A. M. , Frisse, L. M. , Nadler, S. A. , Sternberg, P. W. and Thomas, W. K. 1999. Molecular and morphological characterization of two reproductively isolated species with mirror-image anatomy (Nematoda: Cephalobidae). Nematology 1:591–612, doi: 10.1163/156854199508559.
  18. De Ley, P. , van de Velde, M. C. , Mounport, D. , Baujard, P. and Coomans, A. 1995. Ultrastructure of the stoma in Cephalobidae, Panagrolaimidae and Rhaditidae, with a proposal for a revised stoma terminology in Rhabditida (Nematoda). Nematologica 41:153–182, doi: 10.1163/003925995X00143.
  19. de Maeseneer, J. and d’Herde, J. 1963. Méthodes utilisées por l'étude des anguillules libres du sol. Revue d’Agriculture 16:441–447.
  20. de Man, J. G. 1881. Die einheimischen, frei in der reinen Erde und im süssen Wasser lebenden Nematoden. Vorläufiger Bericht und descriptivsystematischer Theil. Tijdschrift van der Nederlandsche dierkundige Vereeniging 5:1–104.
  21. de Man, J. G. 1910. Beiträge zur Kenntnis der in dem weißen Schleimfluß der Eichen lebeden Anguilluliden, nebst Untersuchungen über den Bau des Essigälchens und der Gattung Anguillula Ehrb. Zoologische Jahrbüch Systematik 29:359–394.
  22. de Man, J. G. 1914. Anguillula silusiae de Man, eine neue in der sogenannten Bierfilzen lebende Art der Gattung Anguillula Ehrb. Annales de la Sociéte Royale Zoologique et Malacologique de Belgique 48 (Vol. Jubilaire): 1–9.
  23. Ferris, H. 2009. The beer mat nematode, Panagrellus redivivus: A study of the connectedness of scientific discovery. Nematode Morphology and Systematics 12:19–25.
  24. Fuchs, G. 1930. Neue an Borken- und Rüsselkäfergebundene Nematoden, halbparasitische und Wohnungseinmieter. Freilebende Nematoden aus Moos und Walderde in Borken-und Rüsselkäfergängen. Zoologische Jahrbücher (Systematik) 59:505–646.
  25. Goodey, T. 1922. The eel-worm in paper-hangers’ paste [Anguillula rediviva (Linnaeus, 1767), Stiles and Hassall, 1905. Annals and Magazine of Natural History (Series 9) 10:297–307, doi: 10.1080/00222932208632783.
  26. Goodey, T. 1943. On the systematic relationship of the vinegar eelworm, Turbatrix aceti and its congeners, with a description of a new species. Journal of Helminthology 21:1–9, doi: 10.1017/S0022149X00031825.
  27. Goodey, T. 1945. A note on the subfamily Turbatricinae and the genus Turbator Goodey, 1943. Journal of Helminthology 21:69–70, doi: 10.1017/S0022149X00031953.
  28. 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, doi: 10.1021/bk–1999-0734.ch008.
  29. Hechler, H. C. 1970. Reproduction, chromosome number, and postembryonic development of Panagrellus redivivus (Nematoda: Cephalobidae). Journal of Nematology 2:355–361.
  30. Hechler, H. C. 1971a. Taxonomic notes on four species of Panagrellus Thorne (Nematoda, Cephalobidae). Journal of Nematology 3:227–237.
  31. Hechler, H. C. 1971b. Panagrellus ceylonensis n. sp. (Nematoda, Cephalobidae). Journal of Nematology 3:246–250.
  32. Heindl-Mengert, H. 1956. Die Nematodenfauna im Schleimfluss lebender Laubbaüme. Sitzungberichte der Physikalisch Medizinischen Sozietät Erlangen 77:158–176.
  33. Ivanova, E. , Perfilieva, K. and Spiridonov, S. 2018. Panagrellus levitatus sp. n. (Rhabditida: Panagrolaimidae), a nematode suppressing Drosophila melanogaster in laboratory cultures. Nematology 20:285–297, doi: 10.1163/15685411-00003141.
  34. Katoh, K. and Toh, H. 2008. Recent developments in the MAFFT multiple sequence alignment program. Briefings in Bioinformatics 9:286–298, doi: 10.1093/bib/bbn013.
  35. Katoh, K. , Misawa, K. , Kuma, K. and Mitaya, T. 2002. MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Research 30:3059–3066, doi: 10.1093/nar/gkf436.
  36. Kitagami, Y. , Kanzaki, N. , Tanikawa, T. and Matsuda, Y. 2019. Free-living nematodes associated with pine cones of Pinus thunbergii and P. taeda at Japanese coastal and inland forest sites. Nematology 21:389–400, doi: 10.1163/15685411-00003221.
  37. Kumar, S. , Stecher, G. and Tamura, K. 2016. MEGA7: Molecular Evolutionary Genetics Analysis Version 7.0 for bigger datasets. Molecular Biology and Evolution 33:1870–1874, doi: 10.1093/molbev/msw054.
  38. Larget, B. and Simon, D. L. 1999. Markov chain Monte Carlo algorithms for the Bayesian analysis of phylogenetic trees. Molecular Biology and Evolution 16:750–759, doi: 10.1093/oxfordjournals.molbev.a026160.
  39. Linné, C. von 1767. “Systema naturae sive regna tria naturae, secundum classes, ordines, genera, species, cum characteribus, differentiis, synonymis, locis”, Vol. 1. Part 2, 12th Ed. Laurentii Salvii, Holmiae 795; [pp. 533–1327].
  40. Massey, C. L. 1974. Biology and taxonomy of nematode parasites and associates of bark beetles in the Unites States. Agriculture Handbook, 446 (Forest Service, U.S. Department of Agriculture):1–233.
  41. Maupas, E. F. 1900. Modes et formes de reproduction des nématodes. Archives de Zoologie Expérimentale et Générale 8:463–624.
  42. Menzel, R. 1922. Beiträge zur kenntnis der Mikrofauna von Niederländisch-ost-Indien. Treubia 3:116–126.
  43. Nunn, G. B. 1992. Nematode molecular evolution. An investigation of evolutionary patterns among nematodes based upon DNA sequences. Ph.D. dissertation, University of Nottingham, 228 pp.
  44. Paramonov, A. A. 1956. K revizii sistemy Rhabditat rastenii [On the revision of the Rhabditat system of plants] (In Russian). Trudy Gelmintologicheskoi Laboratorii Akademiya Nauk SSSR 8:77–111.
  45. Peters, B. G. 1927. On the nomenclature of the vinegar eelworn. Journal of Helminthology 5:133–142.
  46. Rambaut, A. 2018. Figtree, a graphical viewer of phylogenetic trees, version 1.4.4. Institute of Evolutionary Biology, University of Edinburgh, Edinburgh, available at: https://github.com/rambaut/figtree/releases/tag/v1.4.4.
  47. Rambaut, A. , Drummond, A. J. , Xie, D. , Baele, G. and Suchard, M. A. 2018. Posterior summarization in Bayesian phylogenetics using Tracer 1.7. Systematic Biology 67:901, doi: 10.1093/sysbio/syy032.
  48. Ronquist, F. , Teslenko, M. , van der Mark, P. , Ayres, D. L. , Darling, A. , Höhna, S. , Larget, B. , Liu, L. , Suchard, M. A. and Huelsenbeck, J. P. 2012. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic Biology 61:539–542, doi: 10.1093/sysbio/sys029.
  49. Rühm, W. 1956. Die Nematoden der Ipiden. Parasitologische Schriftenreihe 6:1–437.
  50. Sandground, J. H. 1939. Cephalobus parasiticus n. sp. and “Pseudostrongyloidiasis” in Macaca irus mordax . Parasitology 31:132–137.
  51. Sanger, F. , Nicklen, S. and Coulson, A. R. 1977. DNA sequencing with chain-terminating inhibitors. Proceedings of the National Academy of Sciences U.S.A 74:5463–5467, doi: 10.1073/pnas.74.12.5463.
  52. Sanwal, K. C. 1960. Panagrellus dubius n. sp. (Nematoda, Turbatricinae Goodey, 1943), from frass of beetle Sternochetus lapathi (L.), with remarks on redescriptions of Anguillula rediviva (L., 1767). Canadian Journal of Zoology 38:1041–1046, doi: 10.1139/z60-108.
  53. Sanwal, K. C. 1971. Geraldius n. gen., Macrolaiminae n. subfam., with a revision of the subfamilies and genera of Chambersiellidae (Nematoda). Canadian Journal of Zoology 49:965–967, doi: 10.1139/z71-147.
  54. Siddiqi, M. R. 1964. Studies on Discolaimus spp. (Nematoda: Dorylaimidae) from India. Zeitschrift für zoologische Systematik und Evolutionsforschung 2:174–184, doi: 10.1111/j.1439–0469.1964.tb00720.x.
  55. Siddiqi, M. R. 1993. Nematodes of tropical rainforests: 2. Five new genera and eight new species of cephalobs. Afro-Asian Journal of Nematology 3:212–25.
  56. Siddiqi, M. R. 2014. Descriptions of Shahnema typicum gen. n., sp. n. and S. cabii gen. n., sp. n. (Diplogastrina: Shahnematidae fam. n.). International Journal of Nematology 24:153–160.
  57. Steiner, G. 1934. Procephalobus mycophilus n. g., n. sp. (Cephalobidae), a nematode living in the sclerotia of the fungus Balansia claviceps . Proceedings of the helminthological Society of Washington 1:54–56.
  58. Steiner, G. 1936. Opuscula miscellanea nematologica, III. Proceedings of the Helminthological Society of Washington 3:16–22.
  59. Stock, S. P. and Nadler, S. A. 2006. Morphological and molecular characterisation of Panagrellus spp. (Cephalobina: Panagrolaimidae): taxonomic status and phylogenetic relationships. Nematology 8:921–938, doi: 10.1163/156854106779799277.
  60. Sukul, N. C. 1971. New free-living nematodes from the thermal spring of Tantloya, Bihar. Indian Journal of Helminthology 23:135–145.
  61. Thorne, G. 1937. A revision of the nematode family Cephalobidae Chitwood and Chitwood, 1934. Proceedings of the Helminthological Society of Washington 4:1–16.
  62. Thorne, G. 1938. Notes on free-living and plant-parasitic nematodes. IV. Proceedings of the Helminthological Society of Washington 5:64–65.
  63. Thorne, G. 1939. Notes on free-living and plant-parasitic nematodes. V. Proceedings of the helminthological Society of Washington 6:30–32.
  64. Timm, R. W. 1956. Marine nematodes from the Bay of Bengal. I. Phasmidea. Journal of the Bombay Natural History Society 54:87–90.
  65. Tsalolikhin, S. J. 1965. Novaja saprobiotischeskaja nematoda Panagrellus silusioides sp. n. Vestnik Leningradskogo Gosudarstvennogo Universiteta 21:152–153.
  66. Varga, M. 1958. Panagrellus pycnus. Európában eddig nem jelzett fonálféregfaj első jelentkezése. Élelmezési Ipar 12:74–80.
  67. Vecchi, M. and Bruneaux, M. 2021. Concatipede: an R package to concatenate fasta sequences easily, Available at: http://doi.org/10.5281/zenodo.5130604.
  68. Yokoo, T. and Ota, Y. 1961. On the variations of dimensions within soil nematodes. III. Dimensions of the Brevibucca japonica n. sp. found in the decayed fruit of pear in Japan, with descriptions of this new species. Agricultural Bulletin of Saga University 12:149–156.
  69. Zullini, A. 1982. Nematodi (Nemata). In the series: Guide per il riconoscimento delle specie animali delle acque interne italiane. Volume 17. Consiglio Nazionale delle Ricerche, Rome: 117 pp.

EXTRA FILES

COMMENTS