Morphological and molecular characterization of Acrobeloides saeedi Siddiqi, De Ley and Khan, 1992 (Rhabditida, Cephalobidae) from India and comments on its status

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Morphological and molecular characterization of Acrobeloides saeedi Siddiqi, De Ley and Khan, 1992 (Rhabditida, Cephalobidae) from India and comments on its status

Aasha Rana / Aashaq Hussain Bhat * / Suman Bhargava / Ashok Kumar Chaubey / Joaquín Abolafia

Keywords : 18S rDNA, 28S rDNA, Acrobeloides bodenheimeri , Acrobeloides gossypii n. syn., Acrobeloides ishraqi n. syn., Acrobeloides longiuterus , Acrobeloides maximus , Acrobeloides mushtaqi n. syn., description, ITS rDNA, taxonomy

Citation Information : Journal of Nematology. Volume 52, Pages 1-21, DOI: https://doi.org/10.21307/jofnem-2020-027

License : (CC-BY-4.0)

Received Date : 17-October-2019 / Published Online: 28-April-2020

ARTICLE

ABSTRACT

Two cultured populations of Acrobeloides saeedi are described from India. Morphologically and morphometrically this material agrees with other species of the Maximus-group (A. bodenheimeri, A. longiuterus, and A. maximus), especially with A. longiuterus. However, molecular studies based on 18 S, 28 S and ITS rDNA confirmed the Indian material is well differentiated from all of these species. According to this, A. saeedi is considered a valid taxon distinguished mainly from A. bodenheimeri by having dextral female reproductive system (vs sinistral), from A. longiuterus by having larger females (1.03-1.57 vs 0.57-0.88 mm) and from A. maximus by having seta-like labial processes (vs absent) and males as frequent as females (vs males very infrequent). Molecular and phylogenetic studies revealed the present specimens to be conspecific to undescribed Acrobeloides sp. population from Iran, and hence, both regarded to be conspecific to each other. In addition, other similar species are revised: Acrobeloides ishraqi is considered new junior synonym of A. saeedi, Acrobeloides mushtaqi is considered new junior synonym of A. bodenheimeri, while Acrobeloides gossypia is also considered junior synonym of A. saeedi.

Graphical ABSTRACT

Acrobeloides saeedi was described by Siddiqi et al. (1992) to erect the material previously described as Cephalobus litoralis (Akhtar, 1962; Andrássy, 1984) from Pakistan by Saeed et al. (1988). This last material, based only in two females was observed having morphology and morphometry somewhat different (Siddiqi et al., op. cit.) with respect to the type material of Paracephalobus litoralis described by Akhtar (1962) from Pakistan. Later, Khan and Hussain (1997) proposed the new genus Rafiqius to include A. saeedi and other morphological related species as A. bodenheimeri (Steiner, 1936; Thorne, 1937). This newly proposed genus was differentiated from Acrobeloides (Cobb, 1924) according to the morphology of the lip region, having seta-like processes at labial primary axils. However, the creation of this new genus was considered unjustified by De Ley et al. (1999). Unfortunately, none of these studies provided molecular study.

Figure 1:

Acrobeloides saeedi (isolate KMW) (Siddiqi et al., 1992) (line drawing). A: adult neck region; B: anterior end; C: female reproductive system; D: entire male; E: entire female; F: female posterior end; G: male posterior end; H: lateral field.

10.21307_jofnem-2020-027-f001.jpg
Figure 2:

Acrobeloides saeedi (Siddiqi et al., 1992) (light microscopy). A: neck (arrow pointing the excretory pore); B: stoma; C: intestinal cardiac part with bacteria; D: entire female; E, F: vagina region in lateral and ventral views, respectively (black arrows pointing the vaginal glands, white arrow pointing the postvulval uterine sac); G: vulva; H: entire male; I: female posterior end; J: male posterior end; K: testis.

10.21307_jofnem-2020-027-f002.jpg
Figure 3:

Acrobeloides saeedi (Siddiqi et al., 1992) (scanning electron microscopy). A-B: male lip region (arrows pointing the amphids); C-F: female lip region.

10.21307_jofnem-2020-027-f003.jpg
Figure 4:

Acrobeloides saeedi (Siddiqi et al., 1992) (scanning electron microscopy). A: cuticle at excretory pore level (arrow); B, C, F, G: male posterior end in left lateral (B, F) and ventral (C, G) views (black arrows pointing the genital papillae, white arrows pointing the phasmids); D: lateral fields (arrows pointing the longitudinal incisures); E: female posterior end (arrow pointing the phasmid).

10.21307_jofnem-2020-027-f004.jpg

With respect to the isolation of soil nematodes using the Galleria soil baiting technique of Bedding and Akhurst (1975), the insect associate nature of some Acrobeloides species has been previously reported (Azizoglu et al., 2016). Besides their insect associate nature, their infestation has also been observed with some mollusks, arthropods, and annelids (Grewal et al., 2003). Kraglund and Ekelund (2002) reported infestation of A. nanus (de Man, 1880; Anderson, 1968) in earthworm cocoons. Baquiran et al. (2013) studied the association of these nematodes with microbes and repeatedly observed the presence of three bacterial species in association with A. maximus (Thorne, 1925, 1937). Later, Thiruchchelvan et al. (2018) found a free-living nematode similar to A. longiuterus (Rashid and Heyns, 1990; Siddiqi et al., 1992) in Sri Lanka infecting crop pests. Additionally, Suman et al. (2020) collected other rhabditid species, Distolabrellus veechi Anderson, 1983, from soil samples using the insect baiting technique. Their involvement in soil nutrient cycle and soil mineralization is well evident and during these processes, they interact with many arthropods and other invertebrate species, which may be phoretic to pathogenic, thus may be important for their use in biological control programs.

During a survey of soil nematodes in Meerut, Uttar Pradesh, India, two isolates of Acrobeloides were obtained and were labeled as KMW and DH1. Study of the specimens of these two populations showed that they were conspecific to A. saeedi (Siddiqi et al., 1992). Detailed redescription of this species based in morphological and morphometrical data is provided. We also provided a high quality photographic documentation of important morphological characters of A. saeedi through light microscopy (LM) and scanning electron microscopy (SEM). Additionally, molecular data of this species based in the D2-D3 region of the 28 S rDNA, 18 S rDNA, and internal transcribed spacer (ITS) regions of rDNA genes are included to support the morpho-taxometrical studies. This is the first molecular study of this species and its first valid report from India.

Materials and methods

Nematode isolation, culture, and processing

Soil samples were collected from agricultural farmlands in Mawana, Meerut (28°9´N, 77°71´E, and elevation of 225 m), India, and were tested for the presence of nematodes. Nematode specimens were isolated from two soil samples by Galleria soil baiting technique and were designated as DH1 and KMW. The cadavers were transferred to white trap (White, 1927) after proper washing with double distilled water and sterilization with 1% NaOCl. The nematodes that emerge in white trap were harvested, and stored in 250 ml tissue culture flasks in incubator at 15°C as described by Bhat et al. (2019). For observations and morphometrics, third-stage juveniles (200) were injected to larvae of Galleria mellonella by Insulin Syringe 1 ml and larvae were killed within 36 hr at 27°C. The dead larvae were then transferred to white trap. The adult generations and third-stage juveniles were collected from white trap which emerge into water within six to seven days. These specimens were then killed with hot water, transferred to TAF (2% triethanolamine and 7% formaldehyde) for fixation. The fixed nematodes were processed to dehydrated glycerine as described by Seinhorst (1959) and mounted in pure glycerine on permanent glass-slides (Siddiqi, 1964).

Light microscopy (LM)

Nematode specimens were observed for morphological characters under phase contrast microscope (Nikon Eclipse 50i) and light microscope (Magnus MLX) while morphometric characters were measured with built-in software of the Nikon Eclipse 50i (Nikon DS–L1). Demanian indices (de Man, 1880) and other morphometrical ratios were calculated. Line drawings were made with the help of drawing tube attached to the Nikon microscope provided with differential interference contrast (DIC) optics. Images were taken with the Nikon microscope that was provided with DIC optics and Nikon Digital Sight DS-U1 camera. Micrographs were edited using Adobe® Photoshop® CS. The terminology used for the morphology of stoma and spicules follows the proposals by De Ley et al. (1995) and Abolafia and Peña-Santiago (2017a), respectively.

Scanning electron microscopy (SEM)

For the SEM, male and female generations were first fixed in TAF and then preserved in glycerine. Glycerine preserved specimens were used for SEM observations according to the Abolafia’s (2015) proctocol. They were hydrated in distilled water, dehydrated in a graded ethanol-acetone series, critical point dried with liquid CO2, mounted on SEM stubs and finally coated with gold. The mounts were examined with a Zeiss Merlin microscope (5 kV) (Zeiss, Oberkochen, Germany).

Molecular analyses

DNA extraction, amplification, and sequencing

DNA was extracted from pool of juveniles isolated from cadavers of Galleria mellonella infected with A. saeedi using Qiagen DNeasy® Blood and Tissue Kit (Qiagen, Hilden, Germany) (Bhat et al., 2017). Juveniles were first washed separately with Ringer’s solution followed by washing in PBS solution (Bhat et al., 2017, 2020). They were then transferred into a sterile Eppendorf tube (0.5 ml) and DNA was extracted following manufacturer’s instructions. The ITS region was amplified using the primers 18 S: 5´-TTG ATT ACG TCC CTG CCC TTT-3´ (forward) and 28 S: 5´-TTT CAC TCG CCG TTA CTA AGG-3´ (reverse) (Vrain et al., 1992). The 18 S rDNA fragment was amplified using primers NEM18SF: 5´-CGCGAATRGCTCATTACAACAGC-3´ (forward) and NEM18SR: 5´-GGGCGGTATCTGATCGCC-3´ (reverse) (Floyd et al., 2005). The flanking segment, D2-D3 regions of 28 S rDNA was amplified using primers D2F: 5´-CCTTAGTAACGGCGAGTGAAA-3´ (forward) and 536: 5´-CAGCTATCCTGAGGAAAC-3´ (reverse) (Nadler et al., 2006). The PCR master mix consisted of ddH2O 16.8 μl, 10× PCR buffer 2.5 μl, dNTP mix (10 mM each) 0.5 μl, 1 μl of each forward and reverse primers, dream taq green DNA polymerase 0.2 μl and 3 μl of DNA extract. The PCR profiles used was: 1 cycle of 94°C for 3 min followed by 40 cycles of 94°C for 30 sec, + 54°C for 30 sec for 18 S rDNA, 52°C for 30 sec for 28 S rDNA or 55°C for 30 sec for ITS rDNA, + 72°C for 60 sec, and a final extension at 72°C for 10 min. PCR was followed by electrophoresis (45 min, 100 V) of 5 μl of PCR product in a 1% TAE (Tris-acetic acid-EDTA) buffered agarose gel stained with ethidium bromide (Bhat et al., 2018; Aasha et al., 2019). All PCR-products were sequenced using ABI 3730 (48 capillary) electrophoresis instrument by Bioserve Pvt. Ltd (Hyderabad, India) and sequencing results were submitted to NCBI with accession numbers: MK935149 and MK935150 for 18 S of DH1 and KMW, respectively; MN101167 and MK935147 for 28 S of DH1 and KMW, respectively; MK935148 and MK935151 for ITS of DH1 and KMW, respectively.

Phylogenetic analyses

The sequences were edited and compared with those already present in GenBank using the basic local alignment search tool (BLAST) of the National Centre for Biotechnology Information (NCBI) (Altschul et al., 1990). An alignment of nematode samples together with sequences of related cephalobid species was produced for the LSU (D2-D3 rDNA), SSU, and ITS rDNA sequences using default Clustal W parameters in MEGA 6.0 (Kumar et al., 2016) and optimized manually in BioEdit (Hall, 1999). Pairwise distances were computed using MEGA 6.0 (Kumar et al., 2016). All characters were treated as equally weighted and gaps as missing data. Drilocephalobus sp. (AY284679) for the 18 S tree and Teratolobus sp. (KJ652552) for the 28 S tree were used as the out-group taxa and to root the trees. ITS tree was not included because too few sequences are available in the GenBank database for their comparisons. The base substitution model was evaluated using jModeltest 0.1.1 (Posada, 2008). Phylogenetic trees were elaborated using the Bayesian inference method as implemented in the program MrBayes 3.2.7 (Ronquist et al., 2012). The HKY + Γ (gamma distribution of rate variation with a proportion of invariable sites) model was selected. The selected model was initiated with a random starting tree and run with the Markov Chain Monte Carlo for 106 generations. The Bayesian tree was ultimately visualized using the FigTree program 1.4.3 (Rambaut, 2018).

Results and discussion

The morphological and morphometrical studies and molecular (D2-D3, 18 S and ITS rDNA) analyses confirmed the present strains KMW and DH1 as conspecific to A. saeedi (Siddiqi et al., 1992) and hence, described as the same. This is the first report of this species from Indian subcontinent.

Morphological characterization

A. saeedi (Siddiqi et al., 1992) (Figs. 1–4).

Material examined: 20 females, 21 males and 27 L3 juveniles in each KMW and DH1 populations (obtained from Galleria specimens from agricultural soils).

Measurements: see Tables 1 and 2.

Table 1.

Morphometric data for Acrobeloides saeedi KMW isolated from Galleria culture.

10.21307_jofnem-2020-027-t001.jpg
Table 2.

Morphometric data for Acrobeloides saeedi DH1 isolated from Galleria culture.

10.21307_jofnem-2020-027-t002.jpg

Female: Body is larger, 1.31 to 1.57 mm long, in the KMW population and smaller, 1.06 to 1.45 mm, in the DH1 population, more or less fusiform with a sudden narrowing behind the vulva, tapering anteriorly from mid-pharynx to lip region, fusiform, slightly arcuated ventrally and becomes open C shaped upon heat killing. Cuticle with annuli separated from each other by a narrow groove. Lateral fields with four alae limited by five longitudinal incisures ending at tail tip terminus, showing only three incisures after the phasmids. Lip region bears six inner labial papillae and four outer cephalic papillae. Lips are in pairs, with smooth margin; primary axils are “U”-shaped, usually with acute tip; secondary axils are “V”-shaped; guard processes are absent. Labial probolae is low, triangular in section, connected by tangential ridges. Amphidial apertures are pore like, oval. Oral opening is triangular leading into a narrow cephaloboid stoma bearing well-developed refringent rhabdia, cheilostom is short with bar-shaped cheilorhabdia, gymnostom is very short and stegostom is elongated with robust rhabdia. Pharynx is cephaloboid, divided in three regions: pharyngeal corpus is slightly fusiform, 2.7 to 3.1 times the isthmus length in KMW population while 3.7 to 5.4 times in case of DH1; isthmus is robust and basal bulb is spheroid with well-developed valvular apparatus. Excretory pore is located at isthmus level, at 60 to 89% of neck length, at 53 annuli; renette cells are just behind pharyngeal bulb. Hemizonid is present just anterior to the excretory pore. Deirids are present at basal bulb level, at 70 to 92% of neck length, at 48 annuli. Nerve ring surrounds the isthmus at metacorpus-isthmus junction or slightly posterior. Intestine with anterior end with thinner walls. Reproductive system is monodelphic, prodelphic: ovary well developed, with several lines of oocytes, with or without a double flexure at postvulval region; oviduct short; spermatheca well developed, 0.4 to 0.5 times longer than the body width; uterus is very long, divided in two parts only observed in young females, one distal tubular part and other proximal swollen part with thinner walls; in old females all length usually swollen containing 16 to 30 uterine eggs, 41 to 55 µm long and 24 to 35 µm wide; post-vulval uterine sac 0.7 to 0.9 times the body width; vagina is straight or slightly arcuate, 21 to 31% of body width; vulva ventral. Rectum is distinct, shorter than anal body width with three unicellular glands at its junction with the intestine. Anus is large, directed posteriorly. Tail is straight, conoid, truncated to slightly rounded terminus with 15 to 20 annuli ventrally. Phasmids are distinct pore like and located at 59 to 62% of tail length.

Male: Body is 0.81 to 1.16 mm long in the KMW population, and 0.80 to 1.14 mm long in the DH1 population, “J” shaped after heat killing with general morphology similar to female. Reproductive system is monorchic with testis ventrally reflexed anteriorly. Two deep latero-subventral grooves are extended from the sides of the cloacal apparatus approximately to the first preanal pair of the papillae. Genital papillae are in eight pairs, three pairs are pre-cloacal and five pairs are post-cloacal (two at mid tail length, one lateral at lateral field and one subventral, and three terminal, two subventral and one subdorsal), and one midventral papillae. Phasmids are well observed, located posterior to the anterior lateral papillae, at 67 to 70% of tail length. Spicules are long, broad and arcuate, larger than gubernaculum, with manubrium reduced, ventrally bent, rounded-elongate, calamus is conoid and lamina is slightly ventral curved with angular dorsal hump, long ventral velum and very thin rounded tip. Gubernaculum with manubrium-corpus is almost straight, well developed crura with acute tip. Tail is conoid, ventrally curved, with blunt terminus bearing a short fine mucro.

Third stage juvenile (L3): Body is robust, 0.62 to 0.70 mm long in the KMW population, and 0.40 to 0.64 mm in the DH1 population, elongate, straight or slightly curved at posterior end. Cuticle is almost smooth; lip region is similar to adult specimens. Stoma is narrow. Pharynx is clearly visible and differentiated into the three cephaloboid parts. Nerve ring surrounds the isthmus. Excretory pore is at isthmus level. Deirid is obscure. Cardia is reduced, surrounded by intestinal tissue. Rectum is 6 to 7% times the rectum width. Anus is prominent. Tail is conoid with an acute tip.

Diagnosis (of Indian populations)

The material examined of A. saeedi from India is characterized by having 1.06 to 1.57 mm in females and 0.80 to 1.16 mm in males, lateral field with five longitudinal incisures, lip region with six paired lips, smooth, primary and secondary axils lacking guard processes, labial probolae low, triangular in section and frontally flattened, stoma cephaloboid with rounded cheilorhabdia, pharynx cephaloboid with slightly swollen metacorpus, female reproductive system monodelphic-prodelphic, dextral, with spermatheca well developed and postvulval uterine sac slightly shorter than the body diam., female rectum shorter than anal body diam., female tail conoid with truncate to slightly rounded terminus (41-54 µm long, c = 22.0-33.0, c’ = 1.5-2.4), male tail conoid, ventral curved (32-40 µm long, c = 21.0-34.0, c’ = 1.2-2.2), spicules 41 to 54 µm long with reduced ventral bent manubrium and slightly humped lamina, gubernaculum 21 to 30 µm long.

Relationships

Both populations (KMW and DH1) examined now of A. saeedi from India agree well with the type material described by Siddiqi et al. (1992). Morphometric measurements were in close proximity to the Pakistani population described by Siddiqi et al. (1992) (Table 3).

Table 3.

Comparative morphometrics of females from populations of Acrobeloides maximus – group (all measurements in µm except L in mm).

10.21307_jofnem-2020-027-t003A.jpg10.21307_jofnem-2020-027-t003B.jpg

Additionally, A. saeedi resembles morphologically with A. bodenheimeri (Steiner, 1936; Thorne, 1937), A. longiuterus, and A. maximus (Tables 3 and 4). However, from A. bodenheimeri, the Indian populations can be distinguished on the basis of the position of the uterus with respect to the intestine which is dextral (right-handed) in present strains (KMW and DH1) and sinistral (left-handed) in A. bodenheimeri; postvulval uterine sac with shorter range (85-112 vs 45-132 µm), female body length with less range (1.03-1.57 vs 0.87-1.53 mm); pharyngeal basal bulb longer (31-53 vs 22-32 µm), nerve ring to anterior end more anterior (91-151 vs 113-174 µm), distance from anterior end to excretory pore shorter (112-157 vs 131-209 µm), distance from anterior end to deirid shorter (130-178 vs 212 µm), rectum shorter (12-32 vs 27-42 µm).

Table 4.

Comparative morphometrics of males from populations of Acrobeloides maximus – group (all measurements in µm except L in mm).

10.21307_jofnem-2020-027-t004A.jpg10.21307_jofnem-2020-027-t004B.jpg

From A. longiuterus described by Rashid and Heyns (1990) (redescribed by Abolafia and Peña-Santiago, 2017b, authors who synonymized it with A. camberenensis described by De Ley et al., 1990, 1999, its junior synonym), it can be distinguished by having longer body size of females (1.31-1.57 vs 0.65-0.86 mm), neck comparatively longer (168-208 vs 135-175 µm), longer isthmus (28-46 vs 14.5-19 µm), shorter phasmid to anus distance (26-39 vs 49-65 µm, longer tail (44-62 vs 37-45 µm), longer postvulval uterine sac (85-112 vs 75-101 µm) and Demanian indices. Males can be distinguished by longer size (0.81-1.16 vs 0.61-0.89 mm), comparatively longer neck (156-203 vs 143-171 µm), b ratio (4.7-7.0 vs 4.1-5.5 µm), c ratio (21-30 vs 16-21 µm), stoma (11-17 vs 11-12 µm), isthmus (28-50 vs 19-22 µm), nerve ring (28–41 vs 22–28 µm), neck size (156-203 vs 156-168 µm), mid-body diam. (40-73 vs 35-42 µm) and excretory pore position (112–165 vs 119-145 µm); while some measurements like pharyngeal corpus (81-108 vs 101-111 µm), nerve ring (88-129 vs 120-132 µm) and phasmid to anus (21-36 vs 50-64 µm) were comparatively shorter.

From A. maximus, Indian strains (KMW and DH1) can be distinguished by having lips lacking seta-like processes (vs bearing seta-like process at primary axils), pharyngeal metacorpus slightly fusiform (vs fusiform in Thorne (1925) but not well appreciated in Steiner (1936)), lateral field with five incisures (vs three according Smythe and Nadler (2006), being unknown in Thorne (1925) and Steiner (1936)), males as frequent as females (vs male rare or absent, presumably parthenogenetic females (Smythe and Nadler, 2006)), female tail terminus truncate (vs finely rounded). Although the size of the females of the Indian populations of A. saeedi are similar to A. maximus (1.31-1.57 (1.2-1.4) vs 1.2 mm) but they differed in Demanian indices.

Molecular characterization and its taxonomical implications

A. saeedi strains DH1 and KMW were molecularly characterized by ITS rDNA (901 bp, 938 bp), 18 S rDNA (894 bp, 895 bp) and flanking regions D2-D3 of rDNA (984 bp, 997 bp), respectively. The NBlast analysis of D2-D3, 18 S and ITS rDNA sequences of present specimens showed 100% similarity with D2-D3 (KY914573), 18 S (KY090631) and ITS (KY090632) rDNA sequences of Acrobeloides sp. ES-2017 isolate SMF3 from Iran. 18 S sequences of the present two strains do not show any nucleotide difference with each other and with Acrobeloides sp. ES-2017 present in the GenBank. ITS and D2-D3 sequences of DH1 do not show any nucleotide difference with Acrobeloides sp. ES-2017 (KY090632), however, together these regions show two and one nucleotide differences with KMW, respectively. According to this, the Acrobeloides material from Iran could be considered conspecific with A. saeedi.

On the other hand, A. saeedi was considered a probable junior synonym of A. maximus by De Ley et al. (1999) based on morphological data. However, the 18 S sequence alignment of present strains DH1 and KMW showed 21 bp differences with A. maximus (JQ237850), while 28 S sequence alignment showed 51 bp differences and three gaps with A. maximus (AF147067). ITS sequences of A. maximus are lacking. This shows that both species are not conspecific.

On the other hand also, A. saeedi displays some similar morphology with A. longiuterus, two almost undistinguished taxa. However, molecularly both are different. Our D2-D3 sequences of A. saeedi when aligned with only one available D2-D3 sequence (AF147069) of A. longiuterus (formerly A. camberenensis), it showed 38 bp differences. Also, alignment of ITS rDNA of present two strains DH1 and KMW with ITS rDNA of A. longiuterus (MG946132) from Sri Lanka showed 73 bp differences and 23 gaps. According to this, both taxa must be maintained separated.

With respect to A. bodenheimeri (AF202162), the sequence alignment of 18 S genes of present strains showed 22 bp differences. In the D2D3 expansion fragment of 28 S genes, 54 bp differences were observed in aligned data of present strains with DQ145625 (A. bodenheimeri) from USA. These confirm the present strains to be different from A. bodenheimeri.

Distance matrix analyses with other closely related populations of several Acrobeloides species were also carried out using above three genes studies. Thus, the 18 S rDNA sequences of DH1 and KMW are separated from those of other closely related species of Acrobeloides by 9 to 89 bp (Table 5). The D2-D3 segment of 28 S rDNA gene in the Indian isolates differed in 5 to 76 bp from other closely related species of Acrobeloides (Table 6).

Table 5.

Pairwise distances of the 18 S rDNA regions between present strains of Acrobeloides and already described species.

10.21307_jofnem-2020-027-t005.jpg
Table 6.

Pairwise distances of the D2D3 regions of 28 S rDNA regions between present strains of Acrobeloides and already described species.

10.21307_jofnem-2020-027-t006.jpg

All of these data showed that A. saeedi is molecularly different with respect to its more similar species, A. bodenheimeri, A. longiuterus, and A. maximus, and hence, it should be considered as valid species.

Phylogenetic analysis

The phylogenetic analyses of the present stains based on 18 S rDNA and flanking region D2-D3 segment of 28 S rDNA gene also supported the molecular data. Phylogenetic analyses based on 18 S rDNA sequences (Fig. 5) showed a clear monophyly of the group formed by the isolates DH1 and KMW and other undescribed Acrobeloides species ES-2017 from Iran, probably conspecific isolates within a highly supported (100%) clade and together formed a sister clade with other species of “maximus” group from different geographical regions, namely A. maximus and A. bodenheimeri. In D2-D3 rDNA tree (Fig. 6), present two strains DH1 and KMW formed a monophyletic group with Acrobeloides sp. ES-2017, and together formed sister clad with A. longiuterus (including A. camberenensis, its junior synonym (Abolafia and Peña-Santiago, 2017a, b) from USA. Here also, this pair was sister to the other two species of “maximus” group from different geographical regions, namely A. maximus and A. bodenheimeri. For the ITS rDNA region, there were not enough sequences within Acrobeloides genus to construct any useful phylogenetic tree or use it for comparisons. However, both resulting sequences were added to GenBank with accession numbers of KU721840 (KMW) and KU721841 (DH1) for future comparisons.

Figure 5:

Bayesian Inference tree from known and the newly sequenced Acrobeloides saeedi based on sequences of the 18 S rDNA region. Bayesian posterior probabilities (%) are given for each clade. Scale bar shows the number of substitutions per site.

10.21307_jofnem-2020-027-f005.jpg
Figure 6:

Bayesian Inference tree from known and the newly sequenced Acrobeloides saeedi based on sequences of the 28 S rDNA region. Bayesian posterior probabilities (%) are given for each clade. Scale bar shows the number of substitutions per site.

10.21307_jofnem-2020-027-f006.jpg

Taxonomical remarks

Acrobeloides strains DH1 and KMW obtained during the present study were conspecific to A. saeedi from Pakistan. Although they shared morphological similarities with A. longiuterus, A. maximus and A. bodenheimeri but some divergences were also found and displayed morphometrical differences (Tables 3 and 4). This is the first molecular study of this species and first valid report from India. ITS, 18 S, and D2-D3 rDNA studies confirm it to be different from morphologically closely related species of Acrobeloides. Molecular and phylogenetic studies based on the above three genes revealed the specimens studied now and the Acrobeloides population examined from Iran, could be conspecific.

On the other hand, Pervez (2011) described A. ishraqi as a new species from Uttar Pradesh, India. This author compared the specimens with A. bodenheimeri and A. arenicola, but did not compare it with its more similar species, A. saeedi, having identical morphology and morphometry. According to this, we considered both species as conspecific being A. ishraqi a junior synonym of A. saeedi.

Another species, described by Pervez (2011), A. mushtaqi (Pervez, 2011), was described from Uttar Pradesh, India. The author compared it with A. bodenheimeri and did not find very strong diagnostic characters to differentiate between them. However, their material does not have any important differences with respect to A. bodenheimeri. Although this author does not mention the position of the uterus with respect to the intestine (dextral or sinistral), the main character to distinguish A. bodenheimeri from other similar species, its morphology and morphometry agree with it and we considered A. mushtaqi as junior synonym of A. bodenheimeri.

Recently, Nahiyoon et al. (2019) described a new species, A. gossypii (Nahiyoon et al., 2019), from Pakistan. These authors described it using only morphological approaches and related their specimens only with A. bodenheimeri, but they did not compare it with its more similar species, A. saeedi, which has almost identical morphology and morphometry. Accordingly, we considered both species as conspecific being A. gossypii a junior synonym of A. saeedi.

Acknowledgements

The authors would like to thank to the research activity “PAIUJA 2019/2020: EI_RNM02_2019” of the University of Jaén, Spain. SEM pictures were obtained with the assistance of technical staff (Amparo Martínez-Morales) and equipment of “Centro de Instrumentación Científico-Técnica (CICT)” from University of Jaén. The first author would like to thank to the Department of Science and Technology, New Delhi India for providing financial assistance through DST WOS-A (SR/WOS-a/LS-1083/2014).

References


  1. Aasha, R. , Chaubey, A. K. and Bhat, A. H. 2019. Notes on Steinernema abbasi (Rhabditida: Steinernematidae) strains and virulence tests against lepidopteran and coleopterans pests. Journal of Entomology and Zoology Studies 7:954–964.
  2. 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.
  3. Abolafia, J. and Peña-Santiago, R. 2002. Nematodes of the order Rhabditida from Andalucı´a Oriental, Spain. The Genera Nothacrobeles Allen & Noffsinger, 1971 and Zeldia Thorne, 1937. Journal of Nematology 35:233–243.
  4. Abolafia, J. and Peña-Santiago, R. 2017a. On the identity of Chiloplacus magnus Rashid & Heyns, 1990 and C. insularis Orselli & Vinciguerra, 2002 (Rhabditida: Cephalobidae), two confusable species. Nematology 19:1017–1034, doi: 10.1163/15685411–00003104.
  5. Abolafia, J. and Peña-Santiago, R. 2017b. On the identity of Acrobeloides longiuterus (Rashid & Heyns, 1990) Siddiqi, De Ley & Khan, 1992 (Rhabditida: Cephalobidae). Nematology 19:817–820, doi: 10.1163/15685411–00003088.
  6. Akhtar, S. A. 1962. Paracephalobus (Nematoda: Cephalobidae) a new genus of soil inhabiting nematodes. Proceedings of Helminthological Society Washington 29:207–210.
  7. Altschul, S. F. , Gish, W. , Miller, W. , Myers, E. W. and Lipman, D. J. 1990. Basic local alignment search tool. Journal of Molecular Biology 215:403–410, doi: 10.1016/S0022-2836(05)80360-2.
  8. Anderson, R. V. 1968. Variation in taxonomic characters of a species of Acrobeloides (Cobb, 1924) Steiner and Buhrer, 1933. Canadian Journal of Zoology 46:309–320, available at: https://doi.org/10.1139/z68-048
  9. Andrássy, I. 1967. Die Unterfamilie Cephalobinae (Nematoda: Cephalobidae) und ihre Arten. Acta Zoologica Hungarica 13:1–37.
  10. Andrássy, I. 1984. Klasse Nematoda: Ordnungen Monhysterida, Desmoscolecida, Araeolaimid, Chromadorida, Rhabditida Akademie–Verlag, Berlin, 509.
  11. Azizoglu, U. , Karabörklü, S. , Ayvaz, A. and Yilmaz, S. 2016. Phylogenetic relationships of insect–associated free–living rhabditid nematodes from eastern mediterranean region of Turkey. Applied Ecology and Environmental Research 14:93–103.
  12. Baquiran, J. P. , Thater, B. , Sedky, S. , De Ley, P. , Crowley, D. and Orwin, P. M. 2013. Culture–independent investigation of the microbiome associated with the nematode Acrobeloides maximus . PLoS ONE 8:e67425, available at: https://doi.org/10.1371/journal.pone.0067425
  13. Bedding, R. A. and Akhurst, R. J. 1975. A simple technique for the detection of insect parasitic rhabditid nematodes in soil. Nematologica 21:109–110, doi: 10.1163/187529275X00419.
  14. Bhat, A. H. , Istkhar , Chaubey, A. K. , Půža, V. and San–Blas, E. 2017. First report and comparative study of Steinernema surkhetense (Rhabditida: Steinernematidae) and its symbiont bacteria from subcontinental India. Journal of Nematology 49:92–102.
  15. Bhat, A. H. , Chaubey, A. K. and Půža, V. 2018. The first report of Xenorhabdus indica from Steinernema pakistanense: co-phylogenetic study suggests co-speciation between X. indica and its steinernematid nematodes. Journal of Helminthology 92:1–10, doi: 10.1017/S0022149X17001171.
  16. Bhat, A. H. , Chaubey, A. K. , Shokoohi, E. and Mashela, P. W. 2019. Study of Steinernema hermaphroditum (Nematoda, Rhabditida) from the West Uttar Pradesh, India. Acta Parasitologica 64:720–737, doi: 10.2478/s11686-019-00061-9.
  17. Bhat, A. H. , Askary, T. H. , Ahmad, M. J , Suman, B. , Aasha, R. and Chaubey, A. K 2020. Description of Heterorhabditis bacteriophora (Nematoda: Heterorhabditidae) isolated from hilly areas of Kashmir Valley. Egyptian Journal of Biological Pest Control, (in press), available at: https://doi.org/10.1186/s41938-019-0197-6
  18. Bussau, V. C. 1991. Freilebende Nematoden aus Kustendunen und angrenzenden Biotopen der deutschen und danischen Kusten. 3. Dorylaimida. Zoologischer Anzeiger 226:33–63.
  19. Cobb, N. A. 1924. Amended characterization of the nemic genera Cephalobus and Acrobeles. Journal of Parasitology 11:108.
  20. De Ley, P. , Geraert, E. and Coomans, A. 1990. Seven cephalobids from Senegal (Nematoda: Rhabditida). Journal of African Zoology 104:287–304.
  21. De Ley, P. , van de Velde, M. C. , Mounport, D. , Baujard, P. and Coomans, A. 1995. Ultrastructure of the stoma in Cephalobidae, Panagrolaimidae and Rhabditidae, with a proposal for a revised stoma terminology in Rhabditida (Nematoda). Nematologica 41:153–182, doi: 10.1163/003925995X00143.
  22. De Ley, P. , Félix, M. A. , Frisse, L. M. , Nadler, S. A. , Sternberg, P. W. and Thomas, W. K. 1999. Molecular and morphological characterisation of two reproductively isolated species with mirror–image anatomy (Nematoda: Cephalobidae). Nematology 1:591–612, doi: 10.1163/156854199508559.
  23. de Man, J. G. 1880. Die einheimischen, frei in der reinen Erde und im süssen Wasser lebenden Nematoden. Tijdschrift van der Nederlandsche dierkundige Vereeniging 5:1–104.
  24. Floyd, R. M. , Rogers, A. D. , Lambshead, P. J. D. and Smith, C. R. 2005. Nematode specific PCR primers for the 18S small subunit rRNA gene. Molecular Ecology Notes 5:611–612, available at: https://doi.org/10.1111/j.1471-8286.2005.01009.x
  25. Grewal, P. S. , Grewal, S. K. , Tan, L. and Adams, B. J. 2003. Parasitism of molluscs by nematodes: types of associations and evolutionary trends. Journal of Nematology 35:46–56.
  26. 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.
  27. Khan, H. A. and Hussain, S. S. 1997. Biosystematics of Rafiqius saeedi (Siddiqi, Deley and Khan, 1992) Gen. N. Comb. (Nematoda: Cephalobidae) with observation on its life cycles. Pakistan Journal of Zoology 29:139–143.
  28. Kraglund, H. O. and Ekelund, F. 2002. Infestation of natural populations of earthworm cocoons by rhabditid and cephalobid nematodes. Pedobiologia 46:125–135, doi: 10.1078/0031-4056-00119.
  29. 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.
  30. Mehdizadeh, S. , Shokoohi, E. and Abolafia, J. 2013. Morphological and molecular characterisation of Panagrolaimus Fuchs, 1930 (Nematoda, Rhabditida, Panagrolaimidae) species from Iran. Russian Journal of Nematology 21:93–115.
  31. Nadler, S. A. , De Ley, P. , Mundo-Ocampo, M. , Smythe, A. B. , Stock, S. P. , Bumbarger, D. , Adams, B. J. , De Ley, T. , Holovachov, I. O. and Baldwin, J. G. 2006. Phylogeny of Cephalobina (Nematoda): molecular evidence for recurrent evolution of probolae and incongruence with traditional classifications. Molecular Phylogenetics and Evolution 40:696–711, doi: 10.1016/j.ympev.2006.04.005.
  32. Nahiyoon, A. A , Fayyaz, S. and Kazi, N. 2019. New and known nematodes associated with cotton plantation in Sindh, Pakistan. Pakistan Journal of Zoology 51:1309–1314, available at: http://dx.doi.org/10.17582/journal.pjz/2019.51.4.1309.1314
  33. 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, available at: http://doi.org/10.1080/03235408.2010.505363
  34. Posada, D. 2008. jModelTest: phylogenetic model averaging. Molecular Biology and Evolution 25:1253–1256, available at: http://dx.doi.org/10.1093/molbev/msn083
  35. Rambaut, A. 2018. FigTree 1.4.4 (computer program), available at: http://tree.bio.ed.ac.uk/software/figtree/
  36. Rashid, F. and Heyns, J. 1990. Chiloplacus and Macrolaimellus species from South West Africa/ Namibia (Nematoda: Cephalobidae). Phytophylactica 22:189–199.
  37. 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.
  38. Saeed, M. , Khan, S. A. , Khan, H. A. and Qamar, F. 1988. Nematodes associated with nurseries in Karachi, part I. Rose. Pakistan Journal of Scientific and Industrial Research 31:729–730.
  39. Seinhorst, J. W. 1959. A rapid method for the transfer of nematodes from fixative to anhydrous glycerin. Nematologica 4:67–69, available at: https://doi.org/10.1163/187529259X00381
  40. Siddiqi, M. R. 1964. Three new species of Dorylaimoides Thorne & Swanger, 1936, with a description of Xiphinema orbum n. sp. (Nematoda: Dorylaimoidea). Nematologica 9:626–634, available at: https://doi.org/10.1163/187529263X00737
  41. Siddiqi, M. R. , De Ley, P. and Khan, H. A. 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.
  42. Smythe, A. B. and Nadler, S. A. 2006. Molecular phylogeny of Acrobeloides and Cephalobus (Nematoda: Cephalobidae) reveals paraphyletic taxa and recurrent evolution of simple labial morphology. Nematology 8:819–836, doi: 10.1163/156854106779799178.
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FIGURES & TABLES

Figure 1:

Acrobeloides saeedi (isolate KMW) (Siddiqi et al., 1992) (line drawing). A: adult neck region; B: anterior end; C: female reproductive system; D: entire male; E: entire female; F: female posterior end; G: male posterior end; H: lateral field.

Full Size   |   Slide (.pptx)

Figure 2:

Acrobeloides saeedi (Siddiqi et al., 1992) (light microscopy). A: neck (arrow pointing the excretory pore); B: stoma; C: intestinal cardiac part with bacteria; D: entire female; E, F: vagina region in lateral and ventral views, respectively (black arrows pointing the vaginal glands, white arrow pointing the postvulval uterine sac); G: vulva; H: entire male; I: female posterior end; J: male posterior end; K: testis.

Full Size   |   Slide (.pptx)

Figure 3:

Acrobeloides saeedi (Siddiqi et al., 1992) (scanning electron microscopy). A-B: male lip region (arrows pointing the amphids); C-F: female lip region.

Full Size   |   Slide (.pptx)

Figure 4:

Acrobeloides saeedi (Siddiqi et al., 1992) (scanning electron microscopy). A: cuticle at excretory pore level (arrow); B, C, F, G: male posterior end in left lateral (B, F) and ventral (C, G) views (black arrows pointing the genital papillae, white arrows pointing the phasmids); D: lateral fields (arrows pointing the longitudinal incisures); E: female posterior end (arrow pointing the phasmid).

Full Size   |   Slide (.pptx)

Figure 5:

Bayesian Inference tree from known and the newly sequenced Acrobeloides saeedi based on sequences of the 18 S rDNA region. Bayesian posterior probabilities (%) are given for each clade. Scale bar shows the number of substitutions per site.

Full Size   |   Slide (.pptx)

Figure 6:

Bayesian Inference tree from known and the newly sequenced Acrobeloides saeedi based on sequences of the 28 S rDNA region. Bayesian posterior probabilities (%) are given for each clade. Scale bar shows the number of substitutions per site.

Full Size   |   Slide (.pptx)

REFERENCES

  1. Aasha, R. , Chaubey, A. K. and Bhat, A. H. 2019. Notes on Steinernema abbasi (Rhabditida: Steinernematidae) strains and virulence tests against lepidopteran and coleopterans pests. Journal of Entomology and Zoology Studies 7:954–964.
  2. 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.
  3. Abolafia, J. and Peña-Santiago, R. 2002. Nematodes of the order Rhabditida from Andalucı´a Oriental, Spain. The Genera Nothacrobeles Allen & Noffsinger, 1971 and Zeldia Thorne, 1937. Journal of Nematology 35:233–243.
  4. Abolafia, J. and Peña-Santiago, R. 2017a. On the identity of Chiloplacus magnus Rashid & Heyns, 1990 and C. insularis Orselli & Vinciguerra, 2002 (Rhabditida: Cephalobidae), two confusable species. Nematology 19:1017–1034, doi: 10.1163/15685411–00003104.
  5. Abolafia, J. and Peña-Santiago, R. 2017b. On the identity of Acrobeloides longiuterus (Rashid & Heyns, 1990) Siddiqi, De Ley & Khan, 1992 (Rhabditida: Cephalobidae). Nematology 19:817–820, doi: 10.1163/15685411–00003088.
  6. Akhtar, S. A. 1962. Paracephalobus (Nematoda: Cephalobidae) a new genus of soil inhabiting nematodes. Proceedings of Helminthological Society Washington 29:207–210.
  7. Altschul, S. F. , Gish, W. , Miller, W. , Myers, E. W. and Lipman, D. J. 1990. Basic local alignment search tool. Journal of Molecular Biology 215:403–410, doi: 10.1016/S0022-2836(05)80360-2.
  8. Anderson, R. V. 1968. Variation in taxonomic characters of a species of Acrobeloides (Cobb, 1924) Steiner and Buhrer, 1933. Canadian Journal of Zoology 46:309–320, available at: https://doi.org/10.1139/z68-048
  9. Andrássy, I. 1967. Die Unterfamilie Cephalobinae (Nematoda: Cephalobidae) und ihre Arten. Acta Zoologica Hungarica 13:1–37.
  10. Andrássy, I. 1984. Klasse Nematoda: Ordnungen Monhysterida, Desmoscolecida, Araeolaimid, Chromadorida, Rhabditida Akademie–Verlag, Berlin, 509.
  11. Azizoglu, U. , Karabörklü, S. , Ayvaz, A. and Yilmaz, S. 2016. Phylogenetic relationships of insect–associated free–living rhabditid nematodes from eastern mediterranean region of Turkey. Applied Ecology and Environmental Research 14:93–103.
  12. Baquiran, J. P. , Thater, B. , Sedky, S. , De Ley, P. , Crowley, D. and Orwin, P. M. 2013. Culture–independent investigation of the microbiome associated with the nematode Acrobeloides maximus . PLoS ONE 8:e67425, available at: https://doi.org/10.1371/journal.pone.0067425
  13. Bedding, R. A. and Akhurst, R. J. 1975. A simple technique for the detection of insect parasitic rhabditid nematodes in soil. Nematologica 21:109–110, doi: 10.1163/187529275X00419.
  14. Bhat, A. H. , Istkhar , Chaubey, A. K. , Půža, V. and San–Blas, E. 2017. First report and comparative study of Steinernema surkhetense (Rhabditida: Steinernematidae) and its symbiont bacteria from subcontinental India. Journal of Nematology 49:92–102.
  15. Bhat, A. H. , Chaubey, A. K. and Půža, V. 2018. The first report of Xenorhabdus indica from Steinernema pakistanense: co-phylogenetic study suggests co-speciation between X. indica and its steinernematid nematodes. Journal of Helminthology 92:1–10, doi: 10.1017/S0022149X17001171.
  16. Bhat, A. H. , Chaubey, A. K. , Shokoohi, E. and Mashela, P. W. 2019. Study of Steinernema hermaphroditum (Nematoda, Rhabditida) from the West Uttar Pradesh, India. Acta Parasitologica 64:720–737, doi: 10.2478/s11686-019-00061-9.
  17. Bhat, A. H. , Askary, T. H. , Ahmad, M. J , Suman, B. , Aasha, R. and Chaubey, A. K 2020. Description of Heterorhabditis bacteriophora (Nematoda: Heterorhabditidae) isolated from hilly areas of Kashmir Valley. Egyptian Journal of Biological Pest Control, (in press), available at: https://doi.org/10.1186/s41938-019-0197-6
  18. Bussau, V. C. 1991. Freilebende Nematoden aus Kustendunen und angrenzenden Biotopen der deutschen und danischen Kusten. 3. Dorylaimida. Zoologischer Anzeiger 226:33–63.
  19. Cobb, N. A. 1924. Amended characterization of the nemic genera Cephalobus and Acrobeles. Journal of Parasitology 11:108.
  20. De Ley, P. , Geraert, E. and Coomans, A. 1990. Seven cephalobids from Senegal (Nematoda: Rhabditida). Journal of African Zoology 104:287–304.
  21. De Ley, P. , van de Velde, M. C. , Mounport, D. , Baujard, P. and Coomans, A. 1995. Ultrastructure of the stoma in Cephalobidae, Panagrolaimidae and Rhabditidae, with a proposal for a revised stoma terminology in Rhabditida (Nematoda). Nematologica 41:153–182, doi: 10.1163/003925995X00143.
  22. De Ley, P. , Félix, M. A. , Frisse, L. M. , Nadler, S. A. , Sternberg, P. W. and Thomas, W. K. 1999. Molecular and morphological characterisation of two reproductively isolated species with mirror–image anatomy (Nematoda: Cephalobidae). Nematology 1:591–612, doi: 10.1163/156854199508559.
  23. de Man, J. G. 1880. Die einheimischen, frei in der reinen Erde und im süssen Wasser lebenden Nematoden. Tijdschrift van der Nederlandsche dierkundige Vereeniging 5:1–104.
  24. Floyd, R. M. , Rogers, A. D. , Lambshead, P. J. D. and Smith, C. R. 2005. Nematode specific PCR primers for the 18S small subunit rRNA gene. Molecular Ecology Notes 5:611–612, available at: https://doi.org/10.1111/j.1471-8286.2005.01009.x
  25. Grewal, P. S. , Grewal, S. K. , Tan, L. and Adams, B. J. 2003. Parasitism of molluscs by nematodes: types of associations and evolutionary trends. Journal of Nematology 35:46–56.
  26. 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.
  27. Khan, H. A. and Hussain, S. S. 1997. Biosystematics of Rafiqius saeedi (Siddiqi, Deley and Khan, 1992) Gen. N. Comb. (Nematoda: Cephalobidae) with observation on its life cycles. Pakistan Journal of Zoology 29:139–143.
  28. Kraglund, H. O. and Ekelund, F. 2002. Infestation of natural populations of earthworm cocoons by rhabditid and cephalobid nematodes. Pedobiologia 46:125–135, doi: 10.1078/0031-4056-00119.
  29. 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.
  30. Mehdizadeh, S. , Shokoohi, E. and Abolafia, J. 2013. Morphological and molecular characterisation of Panagrolaimus Fuchs, 1930 (Nematoda, Rhabditida, Panagrolaimidae) species from Iran. Russian Journal of Nematology 21:93–115.
  31. Nadler, S. A. , De Ley, P. , Mundo-Ocampo, M. , Smythe, A. B. , Stock, S. P. , Bumbarger, D. , Adams, B. J. , De Ley, T. , Holovachov, I. O. and Baldwin, J. G. 2006. Phylogeny of Cephalobina (Nematoda): molecular evidence for recurrent evolution of probolae and incongruence with traditional classifications. Molecular Phylogenetics and Evolution 40:696–711, doi: 10.1016/j.ympev.2006.04.005.
  32. Nahiyoon, A. A , Fayyaz, S. and Kazi, N. 2019. New and known nematodes associated with cotton plantation in Sindh, Pakistan. Pakistan Journal of Zoology 51:1309–1314, available at: http://dx.doi.org/10.17582/journal.pjz/2019.51.4.1309.1314
  33. 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, available at: http://doi.org/10.1080/03235408.2010.505363
  34. Posada, D. 2008. jModelTest: phylogenetic model averaging. Molecular Biology and Evolution 25:1253–1256, available at: http://dx.doi.org/10.1093/molbev/msn083
  35. Rambaut, A. 2018. FigTree 1.4.4 (computer program), available at: http://tree.bio.ed.ac.uk/software/figtree/
  36. Rashid, F. and Heyns, J. 1990. Chiloplacus and Macrolaimellus species from South West Africa/ Namibia (Nematoda: Cephalobidae). Phytophylactica 22:189–199.
  37. 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.
  38. Saeed, M. , Khan, S. A. , Khan, H. A. and Qamar, F. 1988. Nematodes associated with nurseries in Karachi, part I. Rose. Pakistan Journal of Scientific and Industrial Research 31:729–730.
  39. Seinhorst, J. W. 1959. A rapid method for the transfer of nematodes from fixative to anhydrous glycerin. Nematologica 4:67–69, available at: https://doi.org/10.1163/187529259X00381
  40. Siddiqi, M. R. 1964. Three new species of Dorylaimoides Thorne & Swanger, 1936, with a description of Xiphinema orbum n. sp. (Nematoda: Dorylaimoidea). Nematologica 9:626–634, available at: https://doi.org/10.1163/187529263X00737
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  42. Smythe, A. B. and Nadler, S. A. 2006. Molecular phylogeny of Acrobeloides and Cephalobus (Nematoda: Cephalobidae) reveals paraphyletic taxa and recurrent evolution of simple labial morphology. Nematology 8:819–836, doi: 10.1163/156854106779799178.
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