On Dracograllus miguelitus sp. nov. (Nematoda: Draconematidae) from an inactive structure: insights into its taxonomy, biodiversity and ecology at hydrothermal vents

Study area and sampling collection

The Lucky Strike (LS) vent field is located in the northern part of the Mid-Atlantic Ridge (MAR), south of the Azores (Fig.1A), with a mean depth of 1,700 m (De Busserolles et al., 2009). LS consists of three volcanic cones that harbor over 25 active hydrothermal edifices surrounding a central lava lake (Humphris et al., 2002; Ondreas et al., 2009). Each active site-or edifice-is made of several smokers as well as patches of diffuse venting areas that extend in the periphery. To characterize the meiofaunal communities at the vent field scale, three habitat types were sampled: an active area, the periphery away from hydrothermal activity and a visually inactive edifice (Fig.1B).

The active habitat was located on the Montségur edifice (37°17.28′N, 32°16.53′W), in the southern region of LS, and consisted of cracks on a flat hydrothermal slab at the base of the edifice. The peripheral habitat was approximately 30 m from Montségur (37°17.28′N, 32°16.52′W), and covered by a few centimeters of sediments. Finally, the visually inactive structure, peripheral to the active Sintra edifice, lied about 400 m north of Montségur (37°17.48′N, 32°16.50′W), and consisted in an indurated sulfide structure covered at its base by a thin black layer of manganese oxy-hydroxides (Figs. 1C, 1D). Sampling was conducted during the 2018 Momarsat cruise (Cannat, 2018) using the suction sampler of the Remotely Operated Vehicle Victor6000. Neither the peripheral nor the inactive habitats exhibited visible hydrothermal activity or typical vent fauna. Once onboard, samples were sieved on 300 and 32 µm mesh sizes, and the fraction between 32–300 µm was preserved in 4% borax buffered formalin.

Sample preparation and image acquisition

Nematodes were extracted from the sediment by the use of colloidal silica (Ludox), with specific gravity of 1.39 (Pfannkuche & Thiel, 1988). Specimens were fixed in formalin, and after the (De Grisse, 1969) protocol, they were transferred to glycerol and mounted onto permanent slides (Somerfield & Warwick, 1996). Drawings and measurements were made using a light microscope Leica DM 2500 LED with the aid of a drawing tube and a Leica DMC 4500 camera.

For the fluorescent observations, a Zeiss Axio Imager.Z2 microscope equipped with an Colibri.7 light, an ORCAFlash4.OLT (Hamamatsu, Hamamatsu-city, Japan) camera and a Apotome.2 slider module (for optical sections) was used. Autofluorescence and Phloxine B stain (Exitation 561 nm, Emission 571 nm) were used to observe internal and external structures in 3D. Thus, four fluorescent channels were used: Blue-filter Zeiss 49 DAPI Ex. G365 nm, Em. 445/50 nm, Green-filter Zeiss 38 HE GFP Ex 470/40 nm, Em LP 515 nm, Orange-filter Zeiss 43HE dsREd Ex. 550/25 nm Em. 605/70 nm, Red-filter Zeiss 50Cy5 Ex. 640/30 Em. 690/50 nm. Combinations of one to five channels (with brightfield) were used for optical section, increased depth of field and 3D depending on the observations. Images were processed using Zeiss Zen Pro and Arivis 4D Pro software.

In one of the earliest reviews of Draconematidae, Allen & Noffsinger (1978) provided key recommendations regarding specimen measurements, morphological analysis, and species delimitation. Building on their guidance, this study incorporates the following recommendations and observations: (1) measurements on the CATs should be taken on the right side of the nematode, (2), the length of the swollen esophageal and cephalic regions should be measured from the anterior tip of the lip region to just posterior to the swollen esophageal region (in most Draconematidae, the body is constricted in this region), (3) the body diameter width should be measured at the widest part of the swollen esophageal region while (4) the rostral width should be measured at the base of the rostrum, just anterior to the first body annule. For comprehensive details about measurements and possible variations along developmental stages, see Allen & Noffsinger (1978) and Clasing (1980).

Nomenclatural acts

The electronic version of this article in Portable Document Format (PDF) will represent a published work according to the International Commission on Zoological Nomenclature (ICZN), and hence the new names contained in the electronic version are effectively published under that Code from the electronic edition alone. This published work and the nomenclatural acts it contains have been registered in ZooBank, the online registration system for the ICZN. The ZooBank LSIDs (Life Science Identifiers) can be resolved and the associated information viewed through any standard web browser by appending the LSID to the prefix http://zoobank.org/. The LSID for this publication is: (AA6564D7-6BA7-405E-94D3-B659E62B8BDB). The online version of this work is archived and available from the following digital repositories: PeerJ, PubMed Central SCIE and CLOCKSS.

SYSTEMATICS

Class CHROMADOREA Inglis, 1983

Subclass CHROMADORIA Pearse, 1942

Order Desmodorida De Coninck, 1965

Suborder Desmodorina De Coninck, 1965

Superfamily Desmodoroidea Filipjev, 1922

Family Draconematidae Filipjev, 1918

Diagnosis. (Emended from Leduc & Zhao (2016)): Body short, S-shaped, usually with more or less enlarged pharyngeal and mid-body region. Cuticle annulated except for the head capsule (helmet) and tail terminus. Annules sometimes with spines, minute vacuoles, or a longitudinal lateral field in mid-body region or tail region. Cephalic sensilla in three circles (6 + 6 + 4): six inner labial papillae, six outer labial setae, and four cephalic setae. Rostrum present (except in Dinetia). Amphideal fovea spiral to loop-shaped, rarely reduced or an internal longitudinal bar. CATs present, located dorsally on the cephalic capsule. Somatic setae arranged in eight longitudinal rows. Buccal cavity small to well developed, usually with a dorsal tooth, with or without subventral teeth. Pharynx cylindrical, dumbbell shaped, or with posterior bulb. Secretory–excretory system absent. At least the anterior-most PATs are arranged in four longitudinal rows, two subventrally and two ventrosublaterally, located on the posterior third of the body. PATs with or without differentiated tips, usually straight, rarely long and flexible. Copulatory apparatus with two spicules and trough-shaped gubernaculum. Three caudal glands extending beyond anus/cloaca.

Subfamily Draconematinae Filipjev, 1918

Diagnosis. (Emended from Decraemer, Gourbault & Backeljau (1997)): Pharynx dumbbell-shaped. Swollen anterior body region is usually conspicuous and short. CATs with clearly open tips, mainly blister-shaped, and enlarged bases of insertion, located on the rostrum. Cephalic acanthiform setae may be present on the helmet. Buccal cavity narrow, unarmed. PATs shorts and straights (except for Draconema trispinosum, characterized by some longer tubes), with bell-shaped tips. All PATs are anterior to cloaca (except in Dracograllus eira and Dracograllus minutus). Copulatory thorns are rare. Paravulvar setae present or not. Tail cylindro-conoid with numerous nodules. Non-annulated tail region, with different length between species.

Type genus. Draconema Cobb, 1913.

Genus Dracograllus Allen & Noffsinger, 1978

Diagnosis. (Emended from Leduc & Zhao (2016)): Draconematidae, Draconematinae. Swollen anterior body region 18–26% of total body length. Usually eight CATs on the helmet, rarely 10 to 15. Non-enlarged cuticle on the rostrum. Amphids lateral, usually loop-shaped, with branches usually of unequal length, rarely spiral. Precloacal copulatory thorns are usually absent. PATs all anterior to cloacal opening (except in Dracograllus eira Inglis, 1968 and Dracograllus minutus Decraemer, 1988), and usually extending posterior to anus in females (four exceptions). Paravulvar setae may be present.

Type species. Dracograllus cobbi Allen & Noffsinger, 1978.

Remarks on Dracograllus taxonomy. The genus Dracograllus is placed within the subfamily Draconematinae, along with Draconema, Paradraconema, and also the genus Tenuidraconema. On the other hand, the subfamily Prochaetosomatinae includes the deep-sea typical genera: Bathychaetosoma, Cephalochaetosoma, and Dinetia, as well as Prochaetosoma. This classification is based on characteristics such as the cylindrical shape of the pharyngeal sphincter with a terminal bulb, suggesting that the typical dumbbell-shaped pharynx was possibly secondarily lost in Tenuidraconema (Decraemer, Gourbault & Backeljau, 1997).

A molecular phylogenetic analysis of the family Draconematidae by Rho & Min (2011) revealed that the genus Dracograllus was the first lineage to diverge in the family, followed by five branching orders: Dracograllus—Megadraconema—Draconema—Paradraconema and Prochaetosoma. Consequently, the genus was excluded from the subfamily Draconematinae, as previously suggested. More recently, Leduc & Zhao (2016) examined the phylogenetic position of species within Desmodoroidea and confirmed the basal placement of Dracograllus within the Draconematidae, consistent with (Rho & Min, 2011) findings. Additionally, Leduc & Zhao (2016) found that sequences from Dracograllus, along with two other specimens of the genus, clustered with high posterior probability and bootstrap support, further reinforcing its basal position in the Draconematidae and providing new insights into the group’s evolutionary relationships.

Taxonomic issues. Taxonomic issues within Dracograllus include synonymy, redescriptions, and taxon transfers, often arising from descriptions based on immature specimens or discrepancies in the number of longitudinal PAT rows. For example, Dracograllus eira was originally described as Draconema eira by Inglis (1968), later synonymized with Dracograllus eira. Similarly, Chaetosoma falcatum Irwin-Smith, 1918 underwent multiple reclassifications before being recognized as Dracograllus falcatus Allen & Noffsinger, 1978. Another synonym is Tristicochaeta falcata Johnston, 1938. Since Allen & Noffsinger (1978), and now, the valid name is Dracograllus falcatus. See valid species and nomen nudum section.

Allen & Noffsinger (1978) first described Dracograllus filipjevi from holdfasts of kelps from Japan (Oarai, Ibaraki-ken, Honshu Island). It was characterized by: (1) larger body size (500–700 µm long), (2) the absence of cephalic acantiform setae, (3) the presence of longitudinally areolated body cuticle with dot-like punctations, (4) the presence of some cuticular collar in swollen pharyngeal region, and 9 sublateral and 8–9 subventral PAT in males, and 12–13 and 9–10 in females. Rho, Kim & Kim (2006) also found D. filipjev associated with calcareous algae in Daebo-ri, Guryongpo, Korea, at 3–5 m depth. However, the Korean specimens did not align well with the original description in the number of PATs in male with eight to nine tubes, compared to nine to 11 in the original description. Given that these characteristics are crucial for the taxonomy of Draconematidae, this discrepancy supported the redescription made by Rho, Kim & Kim (2006).

Analyzing the Draconematidae from Guryongpo (Daebo-ri, Korea), Rho & Min (2011) reported several species of the genus Dracograllus. However, these species are considered invalid, meaning they do not comply with certain taxonomic criteria required for formal recognition (see Article 16.1 of the International Code of Zoological Nomenclature). According to this article, every new name published after 1999 must clearly indicate its new status using specific terms such as ‘fam. nov.,’ ‘gen. nov.,’ ‘sp. nov.,’ ‘ssp. nov.,’ or an equivalent expression (e.g., ‘species nova,’ ‘new species’).

List of valid species.

Dracograllus antillensis Decraemer & Gourbault, 1986

Dracograllus chitwoodi Allen & Noffsinger, 1978

Dracograllus cobbi Allen & Noffsinger, 1978

Dracograllus cornutus Decraemer, 1988

Dracograllus demani Allen & Noffsinger, 1978 [Decraemer, 1988; Verschelde & Vincx, 1993]

Dracograllus eira (Inglis, 1968) Allen & Noffsinger, 1978; [Decraemer, 1988; Verschelde & Vincx, 1993]

Syn. Draconema eira Inglis, 1968

Dracograllus falcatum (Irwin-Smith, 1918)

Syn. Chaetosoma falcatum Irwin-Smith, 1918

Syn. Notochaetosoma falcatum (Irwin-Smith, 1918) Cobb, 1929

Syn. Drepanonema falcatum (Irwin-Smith, 1918) Cobb, 1933

Syn. Claparediella falcatum (Irwin-Smith, 1918) Filipjev, 1934

Syn. Draconema falcatum (Irwin-Smith, 1918) Kreis, 1938

Syn. Tristicochaeta falcata (Irwin-Smith, 1918) Johnston, 1938

Syn. Dracograllus filipjevi Allen & Noffsinger, 1978

Syn. Dracograllus gerlachi Allen & Noffsinger, 1978

Syn. Dracograllus gilbertae Verschelde & Vincx, 1993

Dracograllus grootaerti Decraemer, 1988

Dracograllus kreisi Allen & Noffsinger, 1978

Dracograllus laingensis Decraemer, 1988

Dracograllus mawsoni Allen & Noffsinger, 1978

Dracograllus minutus Decraemer, 1988 [Gourbault & Decraemer, 1992]

Dracograllus miguelitus sp. nov. Johnson da Silva et al., 2025

Dracograllus ngakei Leduc & Zhao, 2016

Dracograllus papuensis Decraemer, 1988

Dracograllus pusillus Decraemer, 1988

Dracograllus solidus (Gerlach, 1952) Allen & Noffsinger, 1978

Syn. Draconema solidum Gerlach, 1952

Dracograllus spinosus Decraemer, 1988

Dracograllus stekhoveni Allen & Noffsinger, 1978

Dracograllus timmi Allen & Noffsinger, 1978 [Gourbault & Decraemer, 1992]

Dracograllus trispinosum (Allen & Noffsinger, 1978) Decraemer, 1988

Syn. Dracotoranema trispinosum Allen & Noffsinger, 1978

Dracograllus trukensis Min et al., 2016

Dracograllus wieseri Allen & Noffsinger, 1978

Nomen nudum.

Dracograllus brevitubulus Rho & Kim, 2011 (unaccepted > nomen nudum)

Dracograllus geomunensis Rho & Kim, 2011 (unaccepted > nomen nudum)

Dracograllus gosanensis Rho & Kim, 2011 (unaccepted > nomen nudum)

Dracograllus jaewani Rho & Kim, 2011 (unaccepted > nomen nudum)

Dracograllus jongmooni Rho & Kim, 2011 (unaccepted > nomen nudum)

Dracograllus sungjooni Rho & Kim, 2011 (unaccepted > nomen nudum)

Dracograllus chiloensis Clasing, 1980 (uncertain > taxon inquirendum)

Description of Dracograllus miguelitus sp. nov.

(Table 1; Figs. 2–5; S1–S4)

Type material. All specimens are deposited in the Muséum National d’Histoire Naturelle de Paris, France. Male holotype, two juvenile paratypes and the female paratype in the inventory number MNNH-BN511-I1-L1-B. Two male paratypes and female paratypes in MNHN-BN511-I2-L1-A.

Other material. Other specimens are held in the collection of the Laboratoire Environnement Profond of the Biologie et Ecologie des Ecosystèmes marins Profonds research unit-Ifremer, Plouzané, France.

Etymology. The specific epithet is in honor of ‘Pedro Miguel’, nephew of the first author.

Type locality and habitat. Lucky Strike vent field-MAR. Samples were collected from a hard substratum covered by a thin layer of volcaniclastic sediment, on a visually inactive vent structure at 1,649 m depth. Environmental conditions exhibited background or slightly higher seawater temperature (i.e. 4.8–5.7 °C) and higher pH (i.e. 7.8–7.9) than the surrounding deep-sea but the activity was very low compared to active habitats where recorded temperature varied between 5.2 °C to 9.5 °C, reaching a maximum of 22.1 °C, and pH varying from 7.2 to 7.6.

Measurements. Table 1.

Holotype male. Habitus typical for the genus. A total of 612 µm long, swollen anterior body region representing 19% of total length (Figs. 2A, 2C). Amphid elongate loop-shaped with non-equal branch sizes, and with one more ventrally than another, amphideal fovea 7.1 µm (Figs. 2C and 3C). Helmet strongly cuticularized (Fig. 2D), with punctations and granular appearance in the lateral part (Fig. 3A). Annulation without ornamentation along the body, except for the tail tip and helmet, with minute punctations (Figs. 2C, 2H, 3D). Four CATs on the rostrum (22.5–26.0 µm long), arranged dorsally in two transverse rows, all with enlarged bases (Figs 2C, 3A, 3B). Some specimens exhibited depressions resembling CAT insertions; however, none of these depressions contained tubes. The setae in the cephalic region and along the body possess a cuticular collar at their insertion and alternation of short and long setae, this collar is projected outside of the cuticle, as a pedicel setae (PS), with 1.2–2.1 µm long (Figs. 2C, 3A, 3B), also in the paratype male (Fig. 6E). Cephalic and cervical region with eight longitudinal rows of setae on each side, between 26 and 32 µm long, and also some irregular minute setae (6–12 µm long) (Fig. 2C). Slender cervical region without lateral differentiation (Figs. 2C and 3A). Buccal cavity narrow, unarmed (Fig. 2B). Cardia short. Pharynx dumbbell-shaped with a weakly developed isthmus from the muscular posterior large endbulb (Figs. 2D, 3A, 3B). Intestine narrow, mostly cylindrical, with a granular appearance, gradually widening posteriorly and lying dorsally to the reproductive system (Figs. 3B–3D). Reproductive system with a single and outstretched anterior testis (monorchic) with a well-developed germinative region (Figs. 2G and 3B). Spicules 50.2 µm long, moderately arcuate (in some specimens more arcuate than in others), proximal region with an offset knob-like capitulum (Figs. 2E, 2F and 3F). Gubernaculum 13.6 µm long, lying parallel with the distal end of the spicules, with a minute distal and lateral wing-like expansions (Fig. 3F).

PATs weakly slender, with tongue-like tips (Figs. 2E, 3B, 3D and 3E), difficult to observe due to their thickness. All PATs located anterior to the cloacal opening (Figs. 2E, 3D). PATs are arranged in four longitudinal rows: two sublateral rows each with 10–12 PATs, and two subventral rows consisting of 10 PATs each one (Figs. 2E, 3D), intermingled setae are present between the PATs. Dorsal posterior part with small and irregular distributed setae between the setae following the rows of the body, more visible than in the ventral side, where only few and minute intermingled setae are present (Figs. 3D). Two pairs of setae are present in the annulated tail region. One pair of long setae in the last annules of the dorsal part of the tail, each setae with a collar at the base, close to each other (48–50 µm long), ventrally, one pair of shorter setae, with collar at the basis (11–12 µm long; Figs 2H, 4B-–4D). Additional five setae are present in the tail, in each somatic row of setae. One to two minute setae in the middle of the non-annulated tail tip (1.5 µm long), in some specimens, some of the setae were absent, but the minute insertion of them is visible.

Paratype female. Similar to males in most aspects, but differ in the following characteristics: greater length, with 765.5 µm long (748–788 µm long in paratypes female and 630–735 µm long in paratypes males; Fig 4A). Amphid also elongated loop-shaped, however slightly wider than in males. In addition, amphid in females is positioned more anteriorly than in males (Figs. 4C, 5B). Pedicel setae at both lateral and ventral side of the head, distributed for all body length and more developed than in males (2.5–3.2 µm long; Figs. 4C, 5B). Swollen pharyngeal region 18% of total body length. Some of the setae appear to be lost, with only the cuticular collars remaining, distributed irregularly and in smaller various sizes than in the head (Figs. 6A, 6B). Just some collars, without setae, are also present in the swollen head region and also on the helmet. Both the anterior and posterior regions of the pharynx vary between sexes. The anterior pharyngeal bulb in females has a diameter of 24–26 µm, and the posterior bulb has a diameter of 34–38 µm (compared to 20–22 µm long and 29–33 µm long respectively in males).

Reproductive system didelphic-amphidelphic with reflexed ovaries, both located ventrally relative to the intestine. Uterus filled with a mass of ovoid reproductive cells (Fig. 4F, circles). The region surrounding the vulvar aperture protrudes outward, with the cuticle giving a labial appearance (Fig. 4E). Two pairs of paravulvar setae present, one anterior and one posterior to the vulvar aperture, with length between 6.09–6.69 µm long. Also a setae emerging from the vulvar aperture (5.03 µm long; Figs. 5A and 5E). Well-developed contractor muscles in the vagina (Fig. 4F). PATs all anterior to the anus, more slender, but shorter than in males (56.8–58.8 vs 63.4–68.4 in males), with weakly developed bell shaped tips with a tongue-like valve. PATs arranged in four longitudinal rows: two sublateral rows each consisting of 13 adhesion tubes with intermingling and irregular somatic setae and two subventral rows of 13 adhesion adhesion tubes also with intermingling somatic setae (Fig. 4G). First SlAT on the females with 58.9 µm long and 62.5 on males. All PATs weakly broadened at insertion base. Tail gradually tapering posteriorly to a cylindro-conoidal non-annulated tail tip (Figs. 4H, 5A and 5D). One pair of long setae in the last annules of the dorsal part of the tail (Figs. 4H, circle and 5D). Each setae with a collar at the base and close to each other (50.44 µm long). Ventrally, there is one pair of short setae with also a collar at the base (11–13 µm long). Two pairs of setae are present on the last annules of the tail, each setae featuring a collar at its base and positioned close to one another, measuring 52–56 µm in length (Fig. 5D). Additionally, there is another pair of shorter setae, also with collars at their bases, measuring 12–14 µm in length (Figs. 4G, 4H, 5D). Five more setae are distributed along the tail, in similar size as in males, in each somatic row of setae. One or two minute setae (1.5 µm long) are located at the non-annulated tail tip. In some specimens, one of these setae (and also for those on the annulated tail) is absent, though its minute insertion point remains visible (Fig. 4H). Non-annulated tail tips are long, constituting 56–59% of total tail length. Caudal glands not evident.

Juveniles paratypes

Juvenile third stage. Body shape similar to adults. Body length 426.3 µm long, head diameter 19.3 µm long and a pharynx 75.3 µm long, with a minimally developed isthmus (Fig. 6A). Swollen region representing 24% of the total length. Amphideal fovea is smaller in both length (9.2 µm) and width (4 µm), circular and closed-shape (Fig. 6C). Several pedicel setae with 2.18 µm long, in some specimens as long as in some in adults (Fig. 6C). Two CATs in the dorsal region of the helmet, all at the level of the amphid (15.89–22.24 µm long) (Fig. 6C). Five PATs in both subventral and sublateral row (Figs. 6A, 6D). Tail slender, with the non-annulated tail tip corresponding to 50.7% of the tail length, similar to that in adults (Fig. 6D). The cuticle of the non-annulated tail tip end ornamented with minute punctations. All observed juveniles exhibit a globular appearance on the lateral sides of the body between the body wall and the cuticle, sometimes with brownish or yellowish coloration in the pharyngeal and anterior region.

Juvenile fourth stage. Body shape similar to adults. Body length 514.3 µm long, with head diameter 24.3 µm long. Swollen region representing 25.2% of the total length (Fig. 6B). Amphideal fovea 80.6 µm, circular and closed, similar to the third stage. Presence of pedicel setae with 2.82 µm long. Three CATs located on the helmet with the longer one and more evident with 21.2 µm long. Seven PATs on both sublateral and subventral row, first SlAT with 50.4 µm long and first SvAT with 44.3 µm long (Fig. 6B).

Diagnosis.

D. miguelitus sp. nov. is characterized by the presence of four CATs located in the dorsal side of the helmet at the level of the amphid. All CATs situated anterior to the cuticular annulations. Amphid elongate loop-shaped with different branch sizes in males and females and circular in juveniles. A collar is present at the base of some setae, as a pedicel-like structures. Paravulvar setae are present in two pairs, one anterior and one posterior to the vulvar aperture. Additionally, a minute setae is visible parallel to the vulvar aperture. In the males, PATs are arranged in four longitudinal rows: two sublateral rows each with 10–12 CATs, and two subventral rows consisting of 10 PATs each one. In the females, sublateral and subventral rows with 13 CATs each one. Two pairs of setae in the annulated part of the tail, one pair with long setae (40–50 µm and one pair with shorter setae (11–13 µm). The non-annulated tail tip corresponds to 40–44% in males, 56–59% in females, and 50–55% in juveniles.

Differential diagnosis and relationship

Dracograllus miguelitus sp. nov. is immediately distinguished from its congeners by possessing only four cephalic adhesive tubes (CATs) located at the level of the amphid, in contrast to species with six CATs (D. minutus), eight CATs (D. antillensis, D. chitwoodi, D. cobbi, D. demani, D. eira, D. filipjevi, D. gilbertae, D. grootaerti, D. kreisi, D. laingensis, D. mawsoni, D. ngakei, D. papuensis, D. pusillus, D. solidus, D. spinosus, D. timmi, D. trispinosum, D. trukensis, and D. wieseri), or more, such as D. cornutus (11 CATs), D. falcatus (12 CATs), D. gerlachi (13 CATs), and D. stekhoveni (14 CATs).

The absence of cuticular ornamentation further differentiates D. miguelitus sp. nov. from species with spines (D. antillensis, D. chitwoodi, D. grootaerti, D. minutus, and D. trukensis) or dot-like punctations (D. filipjevi, D. gerlachi, D. kreisi, D. pusillus, and D. trispinosum). The cuticle of D. miguelitus sp. nov., with the collars at the bases of its setae, resembles those observed in D. cobbi, D. mawsoni, D. filipjevi, and D. timmi, though the setae in these species are significantly smaller compared to those in D. miguelitus. The presence of paravulvar setae distinguishes D. miguelitus sp. nov. from several species, including D. chitwoodi, D. cobbi, D. cornutus, D. filipjevi, D. grootaerti, D. minutus, D. ngakei, D. pusillus, D. solidus, D. spinosus, D. stekhoveni, D. timmi, D. trispinosum, and D. trukensis, all of them lacking setae at the vulva.

Dracograllus miguelitus sp. nov. is geographically closest to D. demani and D. trispinosum but can be distinguished from these species by several morphological features. See Tables 2 and 3 for the morphometrical and descriptive comparison between all the valid species In addition to the number of cephalic adhesive tubes (CATs)—four in D. miguelitus sp. nov. compared to eight in both D. demani and D. trispinosum—the new species differs in the number of sublateral adhesive tubes (10–12 in D. miguelitus sp. nov. vs six in D. demani and 10 in D. trispinosum), subventral adhesive tubes (10 in both D. miguelitus sp. nov. and D. demani, but seven in D. trispinosum), and spicule length (50 µm in D. miguelitus sp. nov., compared to 37 µm in D. demani and 61 µm in D. trispinosum). Additionally, the non-annulated tail tip of D. miguelitus sp. nov. is longer (44% of body length in the male holotype and 56% in the female paratype) compared to the shorter tail tips in D. demani and D. trispinosum (32% and 26%, respectively). Morphometrical and descriptive comparison between all the valid species (Tables 2 and 3).

Comments on the imaging approach

Several challenges related to the study of marine nematodes have been discussed here, and we would like to emphasize one of the most important ones: the difficulty in observing and measuring their morphological structures. Therefore, it is crucial to investigate advanced imaging methods to facilitate identification and capture additional morphological features (Foulon, Malloci & Zeppilli, 2025 in press). The pioneering work of Zullini & Villa (2009) first documented the autofluorescence using confocal microscopy of free-living nematodes, with the redescription of Eutobrilus andrassyi. Other examples include the redescriptions of Craspodema reflectans (Cyatholaimidae) and Longicyatholaimus maldivarum (Cyatholaimidae) by Semprucci & Burattini (2015) and Semprucci et al. (2017), respectively. Additionally, an introduction to the application of confocal techniques for observing marine nematodes is provided in Semprucci et al. (2016). In our study, 3D fluorescence imaging has proven effective in several key aspects of identification, particularly for the Draconematidae family. We observed that various structures, that are difficult to study with traditional microscopic techniques, were analyzed with relative ease in our study. These included the insertion of the CATs, the cuticular ornamentation or annulations, and the number of rows of setae. Videos of the 3D fluorescence captures and additional pictures are available in the Supplemental Material (S1–S4).

Biodiversity, distribution and ecology

In terms of species diversity, Dracograllus is the largest genus in the Draconematidae family, with 26 valid species (Nemys, 2024). These species are distributed across a variety of marine environments, and despite their large distribution, the genus is predominantly associated with shallow, tropical and subtropical regions (Min et al., 2016). The distribution and general ecological characteristics of all Dracograllus valid species are presented in Table 4. The Dracograllus genus includes species distributed across the Pacific (18 species), Atlantic (nine species) and Indian (four species) oceans. Some species, such as D. eira, occur in multiple oceanic regions, underscoring their adaptability to diverse oceanic regimes. Distribution of all valid species, and species occurrences including non-identified Dracograllus specimens are provided in Fig. 7. The Pacific Ocean is the region where the highest number of Dracograllus species have been both recorded and described, likely reflecting a bias due to a more extensive sampling. Examples include D. cornutus, D. falcatus, D. filipjevi, D. gerlachi, D. grootaert, D. laingensis, D. mawsoni, D. minutus, D. papuensis, D. pusillus, D. spinosus, D. timmi, D. trukensis, and D. wieseri from a variety of habitats and environmental conditions, particularly in coastal regions. More recently, Leduc & Zhao (2016) described D. ngakei, a species from intertidal coarse sand and gravel sediments in New Zealand, including molecular and morphological data.

In the Atlantic Ocean, species such as D. antillensis, D. chitwoodi, and D. kreisi are typically found in shallow marine environments, often associated with sandy beaches on intertidal or subtidal zones. Decraemer & Gourbault (1986) found approximately 500 individuals of D. antillensis in samples from Guadeloupe, a notably high number for a single species, especially when compared to the abundances typically observed in Dracograllus and even within the broader Draconematidae family. Allen & Noffsinger (1978) described D. trispinosus at 20 m depth, revealing the species occurrence in subtidal zones.

The Indian Ocean hosts fewer described species, yet their habitats share similarities with those in other regions. For instance, D. demani has been reported in tidal coral sands along the Malindi coast and also, in similar sandy habitats in the Pacific Ocean. Likewise, D. eira is known from both the Atlantic Ocean, where it inhabits subtidal sandy zones, and the Indian ocean, where it has been recorded in mangrove-associated sediments, demonstrating its ability to thrive in a range of coastal habitats. D. solidus, another widespread species, has been documented in the Atlantic, Pacific, and Indian oceans, consistently associated with coarse sand in subtidal regions.

Only a few studies have investigated the spatial distribution of Dracograllus in deep-sea ecosystems. Four morphotypes were found at the summit of the GSM in the Atlantic, which is a flat plateau covering more than 1,400 km², with 293 to 511 m depth (Pfannkuche, Sommer & Kähler, 2000). They exhibit significant abundance compared to other Draconematidae genus, with non-overlapping occurrences between each morphotype. The specific sedimentary processes on the GSM (Levin & Nittrouer, 1987), combined with the erosion of old coral reefs, create a coarse sedimentary environment, with small and morphologically complex biogenic structures covering the substratum (Pasenau, 1971; Nellen, 1998). This wide variety of ecological niches would explain their high abundance and the co-occurrence of several species. Similarly, Zeppilli et al. (2014) reported several Dracograllus specimens at the Condor Seamount (CS), at 206 m depth, in the Azores archipelago (Northern Atlantic). The summit of this structure exhibited a highest species richness and dominance for several genus, clearly differing from the surrounding deep-sea habitats or along other CS habitats. The summit was the only area of the study where Dracograllus sp. 1 was present, a flat region, covered by biogenic structures such as sponge sediments or corals, as observed for most Dracograllus species. Similar results regarding Draconematidae species were obtained on other biogenic and sedimentary habitats (Willems et al., 1982; Ndaro & O’lafsson, 1999; Raes & Vanreusel, 2006; Raes et al., 2007; Raes, Decraemer & Vanreusel, 2008).

Draconematidae species were recently observed in samples collected from two deep hydrothermal vent fields, TAG and Snake Pit (SP) (Spedicato et al., 2020) located on the Mid-Atlantic Ridge. Dracograllus sp. was present in 50% of the SP samples, occurring in reddish sediments covered by polychaete tubes. In contrast, these features were absent or less evident at TAG, where only Cephalochaetosoma was recorded. Environmental conditions differed significantly between the vent fields. The total sulfur content in the sediment profiles (0–5 cm) was higher at SP than at TAG and the oxygen penetration about ten times lower at SP. High concentrations of sulfur can lead to death due to the inhibitory action of H₂S on cytochrome c oxidase, an essential enzyme for aerobic respiration. This mechanism blocks the electron transport chain, disrupting ATP production and resulting in metabolic collapse (Bagarinao, 1992). However, some nematode species have developed strategies to cope with sulfide toxicity, such as the oxidation of H₂S into elemental sulfur and its deposition in the epidermis, a process observed in Oncholaimus campylocercoides (Thiermann, Vismann & Giere, 2000). This ability to accumulate and later remove elemental sulfur may enable nematodes to colonize sulfide-rich environments, exploiting niches where most organisms cannot survive. Moreover, body elongation and a higher surface-to-volume ratio may help them cope with low O₂ levels in habitats with limited oxygen availability (Vanreusel et al., 2010b).

The non-overlapping distribution of Dracograllus species at deeper sites suggests that each species may have specific habitat requirements, influenced by both the nature and composition of the substratum, as well as the level of hydrothermal activity. The type-habitat of D. miguelitus sp. nov., is characterized by low hydrothermal influence compared to active sites at the Lucky Strike (LS) vent field (Chavagnac et al., 2018). However, residual venting activity is still present, evidenced by the presence of manganese oxide-hydroxide and high CH4 concentrations measured above the substratum in one of the samples. Environmental conditions, including sediments rich in sulfide minerals, can stimulate microbial communities, which are essential as primary producers in these deposits (Van Gaever et al., 2009). This creates a higher food resource availability and provides structural conditions suitable for the occurrence of Draconematidae species, including D. miguelitus sp. nov., classified as microbial feeders. In summary, the residual hydrothermal activity likely promotes microbial growth, a significant food source for bacterivores and microbial feeders like D. miguelitus sp. nov. and most Draconematidae species. Moreover, the presence of biogenic structures like microbial mats, appear to play a role in the distribution of Dracograllus species. These species are capable of using adhesive tubes to anchor to these structures and may also feed on them (Raes et al., 2007).

Prior to the description of D. miguelitus sp. nov., the Snake Pit species were the deepest-known representatives of the genus, found at depths between 3,480–3,570 m.

Biogeography and evolutionary perspectives

Several species of Dracograllus, and Draconematidae in general, have been found in only a few locations beyond their type habitats. However, nematologists agree that cosmopolitanism is common among various species and groups of marine nematodes (Decraemer, Gourbault & Helléouet, 2001). As reflected by Gad (2009), based on Draconematidae species, one important starting point to determine the origins of these species is to identify their closest relatives and where they occur. In fact, some of the closely related species of Dracograllus miguelitus sp. nov. (D. demani and D. minutus) inhabit coastal and sublittoral environments of the Mediterranean Sea. These Mediterranean regions could be the source of this species, as surface currents transport waters from Gibraltar toward the Azores and upper regions of the North Atlantic (Dietrich et al., 1975; Brenke, 2002). Drifting-buoy experiments have confirmed that Mediterranean water eddies travel westward from Gibraltar across the Atlantic (Richardson, 1996). Such westward flows also occur at approximately 900 m depth, facilitating the transport of fauna, including meiofauna, which may drift as eggs, juveniles, or adults attached to marine snow (Pingree, García-Soto & Sinha, 1999; Gad & Schminke, 2004). This may also be the case for some Prochaetosoma species, as congeners have also been recorded in the Mediterranean. Other potential source regions, such as the coasts of Mauritania or Morocco, remain unconfirmed due to the absence of Draconematidae records from these areas. If such transport occurred, it would also depend on Mediterranean water flows (Gad, 2009).

Studies by Gad (2009) on the Great Meteor Seamount, Zeppilli et al. (2014) on the Condor Seamount, and Spedicato et al. (2020) on the TAG and Snake Pit vent fields identified closely related species in various deep-sea habitats along the northern MAR. Fifteen Draconematidae species, spanning several genera—including Draconema, Paradraconema, Eudraconema, Prochaetosoma, Cephalochaetosoma, and Tenuidraconema—were recorded on the GMS plateau, located 500 km from the Lucky Strike vent field. Remarkably, fourteen of these species were new to science, including four Dracograllus morphotypes (Table 5). In the CS (~300 km from LS), a rich and exclusive nematode community was documented, with 35 species distributed across genera such as Akanthepsilonema, Apenodraconema, Bathychaetosoma, Dracograllus, and others. Similarly, Spedicato et al. (2020) observed several Draconematidae specimens from three genera: Cephalochaetosoma, Dinetia, and Dracograllus.

Both hydrothermal vents and seamounts can be considered true oases of life compared to the surrounding deep-sea environment (McClain et al., 2012), emphasizing their importance for biogeographic studies, particularly for taxa that exhibit some degree of habitat exclusivity, as observed in Draconematidae in the North Atlantic. Another intriguing aspect of Draconematidae in these regions is their morphological variability, which may reflect underlying biogeographic processes (Costello & Chaudhary, 2017). For example, in Dracograllus species from the GMS, individuals from the southern part of the plateau possess a fully divided cephalic capsule (helmet), whereas those from the northern part have a partially divided one (Gad, 2009). Additionally, there are variations in the number of SlATs and SvATs. Several other distinctive traits were reported, including the presence of eight strong spines around the vulva in Draconema sp. 1, a long and conical cephalic capsule in Cephalochaetosoma sp. 10, and extra-wide annules in the pharyngeal region of Prochaetosoma sp. 12. None of these distinctive traits were observed in Draconematidae species from LS (Tchesunov, 2015; W Johnson, 2025, unpublished data).

The intrageneric variation in the helmet among Dracograllus, along with the non-overlapping distributions of several Draconematidae genera and species across the CS, GMS, Snake Pit, and LS, and this may be related to an ongoing speciation process (George, 2004; Gad, 2004; Gad, 2009), similar to what was observed by George & Schminke (2002) and Gad & Schminke (2004) in copepods and macrofaunal species, respectively. In fact, when closely related species exhibit significant morphological variations within small geographic regions, it suggests that species may be arising through micro-allopatric speciation, where populations diverge due to localized environmental differences, leading to subtle, but sometimes crucialmorphological distinctions (Rundle & Nosil, 2005). As these populations adapt to specific ecological niches, genetic divergence and reproductive isolation may drive the emergence of new species, highlighting the importance of understanding local biodiversity and the environmental factors influencing species differentiation.

Given the known limitations of morphology-based taxonomy, such as cryptic diversity and convergent evolution, future studies integrating molecular markers, such as COI or 18S, will be crucial for validating the observed patterns and refining our understanding of species connectivity and dispersal (Palmer, 1988a, 1988b; De Ley et al., 2005; Bhadury et al., 2006; Derycke et al., 2010; Curini-Galletti et al., 2012; Ahmed et al., 2015; Martínez García et al., 2023). Despite these challenges, our findings, together with the limited existing data on species distributions, suggest that both oceanic currents and local conditions and adaptations may play a role in shaping Draconematidae distributions. This highlights the need for further interdisciplinary approaches to fully elucidate the evolutionary and ecological processes governing meiofaunal diversity in deep-sea environments.

Inactive vent structure remarks and conservation implications

Hydrothermal vents have been the focus of numerous ecological studies since their discovery in 1977. These investigations have significantly enhanced our understanding of the structure and dynamics of benthic communities and the role of environmental conditions at various spatial and temporal scales (Godet, Zelnio & Van Dover, 2011). These habitats are known for their unique biogeochemical characteristics, which include commercially valuable mineral resources such as iron, copper, and zinc (Van Dover, 2019). However, the prospect of mining these sites poses serious environmental threats, including permanent alterations in the local topography and removal of habitats (Boschen et al., 2013). Furthermore, mining could release toxic metals, disrupt ecological functions, and hinder the recruitment and recovery of sessile invertebrates, particularly in regions where hard substrata are limited (Van Dover, 2019).

While the fauna of active sites has been the focus of most vent studies, that of inactive sites is virtually unknown. Few studies report the presence of filter-feeders on the relief created by these mineral-rich mounds (Boschen et al., 2013; Van Dover, 2019). Moreover, it is suspected that these mineral deposits may host totally different communities than those found at active vents. This is supported by an eDNA study by Cowart et al. (2020) on the Lucky Strike vent field, which observed significantly higher diversity (OTUs—Operational Taxonomic Units) in both inactive and peripheral regions compared to active ones, as well as notably distinct communities among the active, inactive, and peripheral areas. While inactive vent systems differ from active ones, both of them face significant threats from deep-sea mining, with potentially severe consequences for biological communities and ecosystem functioning. This challenge is exacerbated by the limited knowledge on these habitats and their associated communities, particularly meiofaunal organisms, which are often overlooked in ecological studies. These knowledge gaps may hinder the development of effective management and conservation strategies (Menini et al., 2023).

By documenting the species present at inactive vents, researchers can better assess their ecological roles and connections with neighboring active systems. The discovery of Dracograllus miguelitus sp. nov. at an inactive vent structure exemplifies the biodiversity hidden in these understudied environments and highlights the urgent need for species-level research. Such findings are crucial for balancing conservation priorities with industrial ambitions, ensuring that management strategies are grounded in a comprehensive understanding of ecosystem dynamics and connectivity.

Dichotomous key to Dracograllus valid species

The dichotomous key was constructed based on previous studies (Allen & Noffsinger, 1978; Decraemer, Gourbault & Backeljau, 1997; Min et al., 2016). The complete list for the description of valid Dracograllus species is listed in the reference section.

• 1.

  Four CATs on rostrum D. miguelitus sp. nov.

  • -

    More than four CATs on rostrum2

• 2.

  Without sublateral cephalic acanthiform setae on rostrum3

  • -

    With one pair of bilateral cephalic acanthiform setae on mid-rostrum D. stekholveni

• 3.

  Males with 3–4 preanal corniform setae; 10–11 CATs. Females with 10 CATs: SlATn 18, SvATn 13–16 D. cornutus

  • -

    Males without preanal corniform setae; more CATs. Females with larger numbers of PATs4

• 4.

  Males with 7–8 short stiff setae in subventral rows just anterior to SvATl; spicules 39 µm long. Females with 24 SlAT, including 2 tubes posterior to the anus. Both sexes with a swollen pharyngeal region representing 22% of the total body length D. gerlarchi

  • -

    Males with 3–4 short stiff setae in subventral rows just anterior to SvATl; spicules 71 µm long. Females with 21 SlAT, including 3 tubes posterior to the anus. Swollen pharyngeal region 13–14% of total length D. falcatus

• 5.

  Six CATs on rostrum; males with 5 SlAT, 2–3 SvAT; total length 290 µm; spicules 18 µm D. minutus

  • -

    Eight CATs on the rostrum. Number of PAT higher in males; spicule typically long6

• 6.

  All CATs adjacent to or posterior to the amphideal fovea7

  • -

    All CATs anterior to the amphideal fovea D. eira

• 7.

  Males with preanal corniform setae; slender, conspicuously long and short SlAT alternating in both sexes D. trispinosum

  • -

    Males without preanal corniform setae; SlAT without alternating long and short tubes in both sexes8

• 8.

  Several somatic setae in the posterior body region with spiny cuticular insertion and non-annulated tail tip representing 59% of tail length D. spinosus

  • -

    Somatic setae with spiny insertion collar9

• 9.

  Some somatic setae pedicellate; pedicels 1–8 µm long10

  • -

    Somatic setae without pedicels13

• 10.

  Males with 5–9 SlAT; females with 6–12 SlAT, all anterior to the anus; 9–14 SvAT11

  • -

    Males with 12–24 SlAT; females with 15 SlAT (1 posterior to anus) and 16 SvATD. mawsoni

• 11.

  Males with 5–7 SlAT; females with 9–13 SvAT12

  • -

    Males with 9 SlAT; females with 14 SlAT D. cobbi

• 12.

  Males and females with pedicellate setae in ventrosublateral row just anterior to SlAT; spicules 45–53 µm; females with 6–8 SlAT D. demani

  • -

    Without pedicellate setae in ventrosublateral rows; spicules 36 µm; females with 12 SlAT D. kreisi

• 13.

  Annulated body cuticle without ornamentation14

  • -

    Annulated body cuticle ornamented with spines, dots, and vacuoles15

• 14.

  Amphids long, inverted U-shaped in both sexes; males with 10 SlAT, 11 SvAT; spicules 29 µm; females with 11–13 SlAT and 9–11 SvAT D. papuensis

  • -

    Amphids sexually dimorphic: loop-shaped in males, elongated unispiral in females; males with 7 SlAT, 11 SvAT; spicules 46 µm; females with 8–11 SlAT and same for SvAT D. solidus

• 15.

  Body cuticle with vacuolar and granular ornamentation D. wieseri

  • -

    Body cuticle ornamented with dots and spines16

• 16.

  Body annules ornamented with two rows of dots17

  • -

    Spiny ornamentation of the body cuticle18

• 17.

  Amphids long, oblique loop-shaped in females; tail slender (tail/abd = 5.6) D. chitwoodi

  • -

    Amphids inverted U-shaped in females; tail/abd = 3.9 D. timmi

• 18.

  Amphids long, inverted U-shaped, as long as the rostrum19

  • -

    Amphids short and wide, inverted U-shaped D. antillensis

• 19.

  Short body (L = 310 µm); faint rostrum ornamentation; body annules with minute spines; spicule 26 µm; males with 6 SvAT D. pusillus

  • -

    Body > 400 µm; spiny rostrum ornamentation; longer spicules; more than 6 SvAT in males20

• 20.

  Long swollen pharyngeal region; amphids inverted U-shaped in males and elongated unispiral in females D. laingensis

  • -

    Shorter, wider swollen pharyngeal region; amphids U-shaped in both sexes; spicules 68 µm D. grootaerti

• 21.

  Body annules with dot-like punctations; no anal flap; females with two pairs of paravulvar setae (anterior and posterior to vulva, 5–6 µm) D. filipjevi

  • -

    Anal flap present; females with different number/position of paravulvar setae22

• 22.

  Large spaces between body annules; females without paravulvar setae D. gilbertae

  • -

    Body annules closely spaced; females without paravulvar setae23

• 23.

  Body annules without ornamentation; paravulvar setae absent D. ngakei

  • -

    Paravulvar setae present; body annules with ornamentation24

• 24.

  Body annules with numerous ridges and spiny protrusions, denser in lateral fields. Males with large loop-shaped amphids with ventral branch longer than dorsal, extending to the first body annule D. trukensis

Abbreviations

The terminology used for the description and measurements was according to Min et al. (2016), Leduc & Zhao (2016) and the classical approach by de Man (1880). The abbreviations used are as follows:

L

body length

a

ratio body length/body maximum width

b

ratio body length/pharynx length

c

ratio body length/tail length

c’

ratio tail length/anal body diameter

C. m.

constrictor muscles of the vulvar region

abd

anal body diameter

Amphid from ant

distance from amphid to anterior end

Amphid. Length

amphideal length

Amphid. Width

amphideal width

Amphid./cbd (%)

amphideal length relative to the corresponding body diameter in %

CATs

cephalic adhesive tubes

CATl

length of cephalic adhesive tubes

CATn

number of cephalic adhesive tubes

Gub. Apoph. Length

gubernaculum apophysis length

Head diam.

head diameter

Max. body diam. Mb.

maximum body diameter at mid-body region

Max. body diam. Phar.

maximum body diameter at pharynx level

Min. body diam. Mb.

minimum body diameter at mid-body region

Non. ann. T. length

non-annulated tail tip length

Non. ann. T. length (%)

percentage of non-annulated tail tip relative to total tail length

Ov.

ovaries

PATs

posterior adhesive tubes

Phar. bulb diam. (ant.)

anterior pharyngeal bulb diameter

Phar. bulb diam. (post.)

posterior pharyngeal bulb diameter

Phar. length

pharynx length

PS

paravulvar setae

PS length

paravulvar setae length

SlAT

sublateral adhesive tubes

SvAT

subventral adhesive tubes

SlATn

number of sublateral adhesive tubes

SvATn

number of subventral adhesive tubes

1SlAT1

length of first sublateral adhesion tubes

1SvATl

length of first subventral adhesion tube

Spic

spicule

Spic. Length

spicule length (measured along the median line)

T. length

tail length

T. length/abd

ratio of tail length to anal body diameter

V. (%)

vulva position as percentage of total body length

V. b. diam.

vulvar body diameter

V. to ant.

distance from vulva to anterior end