Description of Panagrolaimus namibiensis n. sp. (Rhabditida: Panagrolaimidae), an Anhydrobiotic Nematode from the Namib Desert of Namibia

Soil samples were collected in the Namib Desert from beneath a common shrub, pencil-bush, Arthraerua leubnitziae (Kuntze) Schinz (Fig. 1). Soils were collected using plastic scoops from multiple locations under the shrub branches from a depth of 0 to 10 cm (litter was gently scraped to the side first). Soils were sieved in the field (2 mm mesh) to remove rocks and transferred to plastic bags. Samples were transported to the University of Richmond (Virginia, USA) and stored at 4°C. Nematodes were extracted over 72 h from soil using a Baermann funnel technique (Baermann, 1917) and subsequently transferred to uninoculated baby food agar (BFA) plates (Stock et al., 2001). Nematodes, and the bacteria they brought with them, flourished on the plates. From this initial culture, a subsequent culture was established from a single female nematode that was transferred to a new plate using a sterile platinum wire pick.

Figure 1:

A) Soil collection site for isolation of Panagrolaimus namibiensis n. sp. Soils were collected from under Arthraerua leubnitziae shrubs (inset). B) Map of Namibia showing location of soil collection site (blue dot). Red dots indicate other soils containing a Panagrolaimus nematode morphologically indistinct from P. namibiensis n. sp.

To sanitize the culture, we employed a 36-h surface sterilization technique (modified from Manzanilla-López and Ehlers, 2021). Nematodes were washed from culture plates with deionized water and pipetted into warm 1% water agar at the bottom of a petri dish. A second warm layer of 1% water agar containing antibiotic (0.3%, dihydrostreptomycin) and antifungal (0.03%, amphotericin B) solutions was added for the nematodes to migrate through over the ensuing 36 hours. The nematodes on the top of the plate were then aseptically washed onto a new BFA plate that was pre-inoculated with an 18 to 24 h broth of Escherichia coli OP50 (Brenner, 1974) in Tryptic Soy Broth. Plates were maintained at 15°C.

Nematodes were washed into a microcentrifuge tube with deionized water. Centrifugation was performed after each processing step to ensure maximum retention of the nematode sample. The nematodes were fixed in a cocktail of 4% paraformaldehyde and 2% glutaraldehyde in sodium cacodylate buffer, followed by a secondary fixation in 1% osmium tetroxide, and subsequently run through serial ethanol dehydration. The final suspension of nematodes in 100% ethanol was then placed in a styrofoam basket where the nematodes were critical-point dried, mounted, and sputter-coated. Images were taken using a JEOL JSM-IT700HR scanning electron microscope (JEOL USA, Inc., Peabody, MA). Cultured nematodes were induced into anhydrobiosis in relative humidity chambers on Supor^® membrane filters (Pall Corporation, Port Washington, NY) and then directly sputter-coated and imaged.

Nematodes were fixed with 5% formalin-glycerin (Seinhorst, 1962), mounted on temporary glass slides, and examined on a Zeiss Axiovert microscope under oil with a x63 ocular lens and a x1.6 Optovar cube to achieve x1000 magnification. Digital measurements of nematode characters were performed using ZEN 2.3 software (Carl Zeiss, Inc., White Plains, NY). Light images were collected using a Nikon Eclipse Ci DIC-equipped compound microscope (Nikon Instruments, Tokyo, Japan) and an Olympus Fluoview FV1200 equipped with a Q Imaging scientific CMOS camera (Olympus Microscope, Tokyo, Japan).

Line drawings were performed by digitally tracing the superimposition of multi-focal plane light images.

DNA was extracted from a whole, single, individual male and female of P. namibiensis n. sp. using the DNeasy Blood and Tissue Kit (Qiagen Inc., Valencia, CA) according to the manufacturer’s instructions. An approximately 700 base-pair fragment of the 5′ end of the 28S nuclear ribosomal large subunit (LSU), including the D2-D3 variable domains, was amplified and sequenced using forward primer Pd2a (5′ACAAGTACCGTGAGGGAAAGT3′) (Kelly Thomas, UNH, pers. comm.) and reverse primer D3B (5′TGCGAAGGAACCAGCTACTA3′) (Nunn, 1992). An approximately 600 base pair fragment of the 5′ end of the 18S nuclear ribosomal small subunit (SSU) was amplified and sequenced using forward primer pNem18SF (5′CGCGAATRGCTCATTACAACAGC3′) and reverse primer pNem18SR (5′GGGCGGTATCTGATCGCC3′) (Floyd et al., 2005). Polymerase chain reactions (PCR, 25 μl total reaction volume) were performed using 2 to 3 μl of DNA template, 12.5 μl of DreamTaq Green PCR master mix (ThermoFisher Scientific, Waltham, MA) and 1.5 μl of each primer at 10 μM concentration. The PCR thermocycling parameters included denaturation at 94°C for 5 min, followed by 35 cycles of 94°C for 30 s, 50 to 51°C (LSU) or 54°C (SSU) for 30 s, and 72°C for 1 min. A final extension period of 5 min at 72°C concluded the amplification. Prior to direct sequencing, PCR products were enzymatically treated with ExoSAP-It PCR Product Cleanup Reagent (ThermoFisher Scientific, Waltham, MA) to remove excess primers and dNTPs. Sequencing reactions were conducted by GENEWIZ/Azenta Inc. (South Plainfield, NJ) with the original PCR primers. Geneious Prime 2023.1.2 (https://www.geneious.com) was used for examination of electropherograms and contig assembly. Sequences obtained for the male and female Panagrolaimus namibiensis n. sp. have been deposited in GenBank as accession numbers PP692155, PP692156, PP69763, and PP697634.

Taxon inclusion for the LSU and SSU analyses aimed to match those included in Yadav et al. (2022) and Shatilovich et al. (2023), though some sequences were excluded due to being considerably shorter than the fragment sequenced for P. namibiensis n. sp. For both the LSU and SSU analyses, sequences of Acrobeles complexus Thorne 1925 (Cephalobidae Filipjev, 1934) and Caenorhabditis elegans (Maupas 1900) Dougherty 1953 N2 strain (Rhabditidae Örley, 1880) were included as outgroups. The SSU analysis included 63 total sequences, with two sequences representing a male and female of P. namibiensis n. sp. (Supplementary Table 1). Sequences were downloaded from GenBank that met the following criteria: unique (not identical), at least 500 base pairs in length, and including the 5′ end of the SSU locus. Forty-two published SSU sequences of Panagrolaimus spp., ten representatives of all available genera of Panagrolaimidae [as listed in the Nemys database (Nemys eds., 2024)], and one unidentified member of Panagrolaimidae (isolate DS129) were included in the alignment and phylogenetic analysis. Three members of Alloionematidae Chitwood & McIntosh, 1934, two members of Brevibuccidae Paramonov, 1956, and one member of Strongyloididae Chitwood & McIntosh, 1934 were also included. The LSU analysis included 36 total sequences in the alignment and phylogenetic analysis (Supplementary Table 1), again with two sequences representing a male and female of P. namibiensis n. sp. Thirty-four published LSU sequences were downloaded from GenBank that were unique, had at least 600 base pairs in length, and that included the D2-D3 variable domain. All the available sequences of Panagrolaimus (21) that met these criteria were included, as well as the same unidentified member of Panagrolaimidae (isolate DS129). Sequences from eleven members of other genera that met these criteria were also included: representatives of all available genera of Panagrolaimidae (Nemys eds., 2024), three members of Alloionematidae, two members of Brevibuccidae, and one member of Strongyloididae.

To obtain the nuclear ribosomal operon of P. kolymaensis, the genome assembly (GCA_028622995.1) was downloaded from NCBI and queried using publicly available Panagrolaimus sequences that collectively span most of the nuclear ribosomal operon (NCBI Nucleotide database accession numbers EU253569.1, LT837699.1, DQ285636.1) and the complete nuclear ribosomal operons of three other nematodes (Meloidogyne chitwoodi Golden, O’Bannon, Santo & Finley 1980 - ON496983.1, Longidorus macrosoma Hooper 1961 - AY580055.1, and Xiphinema americanum (sensu stricto) Cobb, 1913 - AY580056.1) with blastn 2.9.0+ with an e-value cutoff of 1e-100. The top ten hits according to bitscore (ranging from 4667 to 4673) were aligned to the query sequences using mafft 7.453 using the --ep 0 --genafapair options. These sequences exhibited low intragenomic variation, and thus a representative sequence was selected haphazardly.

For both the LSU and SSU analyses, multiple alignments were conducted using Clustal Omega 1.2.3 (Sievers et al., 2020) as implemented in Geneious Prime 2023.1.2 (https://www.geneious.com) using default parameters. MrBayes 3.2.7a x86_64 (Ronquist et al., 2012) was used for Bayesian inference of phylogenetic trees using the CIPRES Science Gateway portal (http://www.phylo.org). The default setting of the “4by4” nucleotide model was used as recommended by the manual for non-coding DNA sequences (Ronquist et al., 2020). Mr. Bayes allows the Markov chain Monte Carlo to sample across the entire general time-reversible (GTR) substitution model space (“Nst=mixed” command), rendering a priori model testing unnecessar y (Ronquist et al., 2020). This also allows the program to use a weighted average of the posterior probability of the best substitution models rather than using a single model (Ronquist et al., 2020). Default, uninformative prior probabilities were used for stationary state frequencies and substitution rates. Each analysis ran for ten million generations as that provided better chain convergence than one million or fewer generations, and the chains were sampled every 2,000 generations with burn-in set to 25%. Support for internal clades was shown by posterior probability (pp) values.

Nematodes used for anhydrobiosis experiments were taken from 10-day laboratory cultures. Nematodes were washed from culture plates using 1 ml sterile M9 buffer (Brenner, 1974), transferred to a glass tube, and vortexed gently to evenly distribute the nematodes. An aliquot of this solution containing approximately 100 to 300 nematodes was then transferred to the surface of a 0.45 µm Supor^® membrane filter (de Souza et al., 2017) on top of a 50-mm bottle-top Nalgene™ filter apparatus (Thermo Fisher Scientific, Waltham, MA). Gentle suction was applied to remove excess buffer solution. Immediately after vacuum filtration, membrane filters were each placed in a 50-mm petri dish base and transferred to a 100% relative humidity (RH) chamber for 24 hours. RH chambers consisted of 8-inch diameter airtight glass desiccators within which target RH levels were established (see below).

After an initial 24 h at 100% RH, nematodes on filters were subjected to one of the following three treatments. First, filters from a baseline treatment were immediately resuspended in 3 ml M9 buffer and examined to establish nematode viability at the start of the experiment. Second, after 24 h at 100% RH, a preconditioned treatment was moved to 97% RH. The nematodes remained at 97% RH for 72 h, were transferred to 85% RH for 24 h, and then to 75% RH for 24 h. After 24 h at 75% RH, the preconditioned nematodes were transferred to a desiccator at 0% RH for 24 h. Third, the direct exposure treatment was immediately transferred to 0% RH after 24 h at 100%. After 24 h at 0% RH, the preconditioned and direct exposure treatment nematodes were rehydrated by adding 3 ml M9 buffer to each petri dish, which were then transferred to 100% RH. Following 24 h of rehydration, the percentage of nematodes on each filter that were alive (regained motility) and dead (non-motile) was determined and used to calculate a percent survival. A 395 nm UV light was periodically flashed at the filter to stimulate motility of the worms (Rajasekharan et al., 2018). Each of the three treatments (baseline, preconditioned, and direct exposure) was replicated eight times, with the filters containing an average of 171 ± 32 nematodes.

One hundred percent RH was maintained by placing two 60-ml beakers of deionized water in the chambers. Other RH levels were achieved using 60 ml of saturated salt solutions (potassium sulfate for 97% RH, potassium chloride for 85% RH, and sodium chloride for 75% RH) or 5 g phosphorus pentoxide for 0% RH (Winston and Bates, 1960; Solomon et al., 1999). The efficacy of each salt solution at achieving and maintaining the desired RH was monitored using a hygrometer (Traceable^®, Cole-Parmer, Vernon Hills, IL). All RH chambers were established with fresh solutions for 24 h before any nematodes were added, and they were incubated at 25°C throughout the experiment.

Statistical analyses were performed with R version 4.3.1 (https://www.r-project.org) (R Development Core Team, 2023). Analysis of variance (ANOVA) was used to study differences in nematode survival among the treatment groups. Where effects were significant, means were compared using Tukey’s Honestly Significant Difference (HSD) multiple comparison procedure. Prior to ANOVA, the Shapiro-Wilk test was used to assess the percent survival for normality, and percent survival was arcsine (sqrt(x/100)) transformed.

Figs. 2–4.

Measurements, see Table 1.

Figure 2:

Panagrolaimus namibiensis n. sp. line drawings. A: Anterior body region. B: Anterior body end, showing stoma and rhabdia. C: Full body, female. D: Full body, male. E: Lateral field. F: Basal bulb, excretory pore. G: Posterior body end, female. H: Posterior body end, male.

Figure 3:

Panagrolaimus namibiensis n. sp. light micrographs. A: Anterior body region and pharynx. B: Lip region, stoma, stegostomal dorsal tooth (DT). C: Full body, female. D: Part of reproductive tract, female. E: Lateral field. F: Part of testis, male. G: Posterior body end, spicule (S), gubernaculum (G), male. H: Posterior body end, rectum, female.

Figure 4:

Panagrolaimus namibiensis n. sp. scanning electron microscopic images. A: Anterior body region. B: Lip region in frontal view. C: Full body, female. D: Excretory pore (EP), anterior deirids (AD). E: Anterior deirid (AD). F: Lateral field. G, H: Vulva. I: Anus, phasmid (P), female. J: Posterior end, posterior deirid (PD), female. K: Posterior end, spicule, pre- and post-cloacal genital papillae (GP), midventral papillae (MP), male.

Body straight, cylindrical, ventrally curved (greater in males) post-fixation. Cuticle finely annulated, thickest at mid-body. Prominent lateral fields containing two distinct raised bands (three incisures) with gradual post-cloacal and prephasmid taper. Lip region thinly cuticularized with six distinct rounded lip segments separated in pairs. Labial papillae pronounced. Amphid apertures inconspicuous. Stoma wide, pipe-like, panagrolaimoid, containing distinctive cheilostom, gymnostom, and stegostom. Both cheilostom and gymnostom contain cuticularized rhabdia, with gymnostom also being heavily sclerotized. Minute, hooked dorsal tooth protruding from stegostom. Pharyngeal sleeve conspicuous but variable in stoma overlap, covering 21 to 60% of total stoma length. Pharyngeal corpus cylindrical with long procorpus and marginally prominent, shorter metacorpus. Isthmus long and thin, clearly distinct from metacorpus. Nerve ring near midpoint of isthmus. Anterior deirids in line with excretory pore, lying toward terminus of pharynx at mid-bulb level. Basal bulb swollen and generally spheroid, with noticeable pharyngo-intestinal valve. Cardia conoid, surrounded by thick intestinal tissue. Body widest at mid. Conspicuous posterior deirid dorsal to lateral field and posterior to mid-body. Base of tail wide at anus, narrowing sharply past phasmids.

P. namibiensis n. sp. is named for the country in which it was found.

Panagrolaimus namibiensis n. sp. was collected from the Namib Desert gravel plains in Namib Naukluft National Park, Namibia, Africa from soils under the shrub Arthraerua leubnitziae (pencil-bush), GPS: 22°56′35.2″S 14°54′51.9″E.

Eight slides were deposited in the Nematode Collection at the United States Department of Agriculture, Beltsville, Maryland, USA, including a female holotype (slide T-799t) and paratypes (three females, slides T-8028p, T8039p, T-8030p, and four males, slides T-800t, T8031p, T8032p, T8033p). The LSID code of this publication is: urn:lsid:zoobank.org:pub:9FF9A841-DB31-4D51-9542-8BE9C991B833.

Main characteristics of P. namibiensis n. sp. include the presence of a hook-shaped stegostomal dorsal tooth and a distinct posterior deirid in both males and females. Additionally, the excretory pore is aligned with both anterior deirids and the middle of the basal bulb. Based on sharing morphological features [e.g., structure of the lip region (six distinct rounded, low lip segments separated in pairs), number of lateral incisures (three), and/or general tail shape (conoid)], the new species resembles Panagrolaimus artyukhovskii Blinova and Mishina, 1975, Panagrolaimus conophthori Massey, 1974, Panagrolaimus davidi Timm, 1971, Panagrolaimus detritophagus Fuchs, 1930, Panagrolaimus goodeyi Rühm, 1956, P. labiatus (Kreis, 1929) Andrássy, 1960, Panagrolaimus leperisini Massey 1974, Panagrolaimus orientalis Korenchenko, 1986, Panagrolaimus rigidus Schneider, 1866, Panagrolaimus subelongatus Cobb, 1914, and P. superbus. Based on phylogenetic analyses (see below), P. namibiensis n. sp. is most closely related to P. labiatus, P. kolymaensis, P. rigidus, and P. davidi. The new species can be distinguished from these morphologically similar and phylogenetically related species as follows:

From P. artyukhovskii by females having a longer body (717.7–976.2 vs. 710–770 μm), males having two pre-cloacal papillae within the range of the spicules (vs. one) and lack of dentiform thickenings in the metastom.

From P. conophthori by having labial papillae (absent in P. conophthori), three lateral incisures (vs. four), shorter average female length (897.8 vs. 960 μm), and absence of metarhabdion denticles.

From P. davidi by having a shorter average male reproductive tract (207.8 μm vs. 450 μm), a hooked-shaped tooth vs. a triangular tooth, eight total pairs of male genital papillae (vs. six), and two pre-cloacal pairs of genital papillae (vs. one).

From P. detritophagus by having lips fused in pairs (unfused lips in P. detritophagus), excretory pore opposite the basal bulb (vs. near the middle of the isthmus), and eight pairs of genital papillae (vs. seven).

From P. goodeyi by having a shorter cheilogymnostom relative to width (less than two times the width vs. greater than three times the width).

From P. kolymaensis by gonochorism as indicated by the presence of males in both laboratory cultures and field samples (P. kolymaensis is parthenogenic).

From P. labiatus by having a stegostomal dorsal tooth (absent in P. labiatus), an excretory pore at basal-bulb level (vs. the middle of the isthmus), and a broad tail lacking acute posterior elongation.

From P. leperisini by having rounded labial papillae that lack labial or cephalic setae (present in P. leperisini) and an excretory pore at basal-bulb level (vs. opposite the nerve ring).

From P. orientalis by having inconspicuous amphidial apertures (conspicuous in P. orientalis), a cuticularized cheilostom containing distinct rhabdia (absent in P. orientalis), a longer female body (717.7–976.2 vs. 500–700 μm), a non-elongated conical tail, and a non-rounded manubrium.

From P. rigidus by having a dorsal tooth, nonsetose labial papillae, a generally broader frame [e.g. a wider average lip width (8.5 vs. 5.7 μm), a wider average mid-body diameter (43.4 vs. 32.3 μm), and a wider anal-body diameter (23.4 vs. 17.5 μm)], and an excretory pore aligned with both anterior deirids and the middle of the basal bulb (vs. the anterior end of the bulb).

From P. subelongatus by having six distinctly segmented lips (vs. three), a significantly longer body in females (717.7–976.2 vs. 600–680 µm), and an excretory pore aligned with anterior deirids at the middle of the basal-bulb (vs. deirids and excretory pore near the anterior end of the bulb).

From P. superbus by shorter average female body length (897.8 vs. 1200 μm), and two pre-cloacal and one midventral genital papillae (vs. three pre-cloacal).

Both the SSU and LSU phylogenetic analyses confirmed the placement of P. namibiensis n. sp. within the Panagrolaimidae in strongly supported clades consisting of most of the Panagrolaimus species that were included in the analyses (Figs. 5 and 6). In the SSU analysis, P. namibiensis n. sp. was placed with full support (1.0 pp) as sister taxon to an undescribed species, Panagrolaimus sp. 4164 from an unknown location (Holterman et al., 2019) (Fig. 5). The clade formed by Panagrolaimus sp. 4164 and P. namibiensis n. sp. was well supported (0.98 pp) as sister to P. kolymaensis from Siberia (Shatilovich et al., 2023). More broadly, P. namibiensis was shown to be part of a maximally supported (1.0 pp) clade that included one of the included sequences of P. davidi (isolate AC Pd2), Panagrolaimus sp. isolate rigidus1, and P. labiatus. Twenty-seven members of Panagrolaimus were part of a moderately supported (0.79) clade that was nearly completely unresolved. While the majority of included members of Panagrolaimus (36 out of 42) formed a strongly supported clade (1.0 pp), the genus was shown to be non-monophyletic, as several members formed additional clades with taxa like Turbatrix aceti (Müller, 1783) Peters, 1927, Procephalobus sp. 1 WB-2008, Propanagrolaimus spp., and Halicephalobus spp.. One sequence of Panagrolaimus paetzoldi Goodey, 1963 isolate 452 was particularly distantly placed from the majority of Panagrolaimus taxa, forming a maximally supported (1.0 pp) clade with two other genera of Panagrolaimidae, Baujardia mirabilis Bert, De Ley, Segers, Van Driessche & De Ley, 2003 and Panagrellus redivivus (Linnaeus, 1767) Goodey, 1945. A different sequence identified in GenBank as P. paetzoldi was placed in a clade with Procephalobus sp. 1 WB-2008. The Panagrolaimidae family was shown to be non-monophyletic. Members of two other families, Cephalobidae (Acrobeles complexus) and Brevibuccidae (Tarantobelus arachnicida Abolafia & Peña-Santiago, 2018 and Plectonchus wyganti Massey, 1974) were placed in clades with members of Panagrolaimidae.

Figure 5:

Phylogenetic tree inferred by Bayesian analysis of SSU sequences showing the relationship of Panagrolaimus namibiensis n. sp. (highlighted in blue) to other members of Panagrolaimidae and other taxa. Support for internal clades is shown by posterior probability (pp) values placed next to nodes. Scale bar represents 0.21 estimated substitutions per site.

Figure 6:

Phylogenetic tree inferred by Bayesian analysis of LSU sequences showing the relationship of Panagrolaimus namibiensis n. sp. (highlighted in blue) to other members of Panagrolaimidae and other taxa. Support for internal clades is shown by posterior probability (pp) values placed next to nodes supported by greater than 0.5 pp. Scale bar represents 0.26 estimated substitutions per site.

The LSU analysis produced a somewhat different topology with improved resolution (Fig. 6). P. namibiensis n. sp. was fully supported (1.0 pp) as sister to P. labiatus SN14, and that clade was strongly supported (1.0 pp) as sister to two undescribed species from Mexican soils, Panagrolaimus sp. HMM550 and Panagrolaimus sp. HMM476 (Mejía-Madrid 2018), and Panagrolaimus sp. S91 from an unknown location. The majority of the remaining Panagrolaimus species (15 out of 22) formed an additional strongly supported clade (1.0 pp) with well-resolved relationships. P. kolymaensis was strongly supported (1.0 pp) as sister to all but two of the other included members of Panagrolaimus. Panagrolaimus was again shown to be nonmonophyletic as Panagrolaimus sp. DS127 was moderately supported as part of a clade with Turbatrix aceti, Halicephalobus spp., and two members of Alloionematidae, Rhabditophanes sp. KR3021 and Alloionema appendiculatum Schneider, 1859. Members of Brevibuccidae, Alloionematidae, and one Cephalobidae formed clades with members of Panagrolaimidae, rendering the family nonmonophyletic.

P. namibiensis n. sp. survived exposure to 0% RH in vitro (Fig. 7). Survival rates of nematodes varied among exposure treatments (ANOVA, significant treatment effect, F_{2, 15} =11.616, P = 0.0009, Fig. 7). Nematodes exposed to desiccation at 0% RH, with or without preconditioning, showed lower survival than the baseline nematodes that were not desiccated. However, there was no difference in the survival percentage between nematodes that were preconditioned and those that were not (Fig. 7).

Figure 7:

A) Panagrolaimus namibiensis n. sp. survival (% of population regaining motility) after exposure to three treatments. Baseline nematodes were maintained at 100% RH for 24 h. Pre-conditioned nematodes were slowly exposed to increasingly lower RH levels before 24 h at 0%. Direct exposure nematodes were directly exposed to 0% RH. Boxplots represent the interquartile range with lines representing the median value. Whiskers represent the minimum/maximum values, and circles are outliers. N = 8 replicates per treatment. Bars with different lowercase letters are statistically different from each other (Tukey’s HSD test). B) SEM of a coiled and anhydrobiotic P. namibiensis n. sp.