Neuromuscular study of early branching Diuronotus aspetos (Paucitubulatina) yields insights into the evolution of organs systems in Gastrotricha

Bekkouche, Nicolas; Worsaae, Katrine

doi:10.1186/s40851-016-0054-3

Research article
Open access
Published: 22 September 2016

Neuromuscular study of early branching Diuronotus aspetos (Paucitubulatina) yields insights into the evolution of organs systems in Gastrotricha

Zoological Letters volume 2, Article number: 21 (2016) Cite this article

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Abstract

Background

Diuronotus is one of the most recently described genera of Paucitubulatina, one of the three major clades in Gastrotricha. Its morphology suggests that Diuronotus is an early branch of Paucitubulatina, making it a key taxon for understanding the evolution of this morphologically understudied group. Here we test its phylogenetic position employing molecular data, and provide detailed descriptions of the muscular, nervous, and ciliary systems of Diuronotus aspetos, using immunohistochemistry and confocal laser scanning microscopy.

Results

We confirm the proposed position of D. aspetos within Muselliferidae, and find this family to be the sister group to Xenotrichulidae. The muscular system, revealed by F-actin staining, shows a simple, but unique organization of the trunk musculature with a reduction to three pairs of longitudinal muscles and addition of up to five pairs of dorso-ventral muscles, versus the six longitudinal and two dorso-ventral pairs found in most Paucitubulatina. Using acetylated α-tubulin immunoreactivity, we describe the pharynx in detail, including new nervous structures, two pairs of sensory cilia, and a unique canal system. The central nervous system, as revealed by immunohistochemistry, shows the general pattern of Gastrotricha having a bilobed brain and a pair of ventro-longitudinal nerve cords. However, in addition are found an anterior nerve ring, several anterior longitudinal nerves, and four ventral commissures (pharyngeal, trunk, pre-anal, and terminal). Two pairs of protonephridia are documented, while other Paucitubulatina have one. Moreover, the precise arrangement of multiciliated cells is unraveled, yielding a pattern of possibly systematic importance.

Conclusion

Several neural structures of Diuronotus resemble those found in Xenotrichula (Xenotrichulidae) and may constitute new apomorphies of Paucitubulatina, or even Gastrotricha. In order to test these new evolutionary hypotheses, comparable morphological data from other understudied gastrotrich branches and a better resolution of the basal nodes of the gastrotrich phylogeny are warranted. Nonetheless, the present study offers new insights into the evolution of organ systems and systematic importance of so-far neglected characters in Gastrotricha.

Background

Gastrotricha are small, often sub-millimetre, interstitial worms, ubiquitously found in most aquatic environments with a long debated phylogenetic position [1–3]. They were first considered closely related to various meiofaunal, protostome groups, such as rotifers (Trochelminthes [4]), kinorhynchs (Nematorhyncha [5]) or Gnathostomulida (Neotrichozoa [6, 7]). Later, molecular phylogenies placed them within Spiralia with uncertain affinities; within the debated group Platyzoa, comprising Gastrotricha, Platyhelminthes and Gnathifera [1, 8, 9]. Recent phylogenomic studies propose a sister group relationship between Platyhelminthes and Gastrotricha [3, 10]. However, the controversy over the phylogenetic position of Gastrotricha masks other problems within the group. Indeed, compared to the diversity and omnipresence of these animals, relatively few phylogenetic and detailed morphological studies have been conducted on this group and the evolution of, e.g., nervous system, muscular system and nephridia is unresolved [11–13]. Its diversity also remains largely unexplored, as exemplified by the recent erection of the family Hummondasyidae (Macrodasyida) in 2014 [14] and the genera Thaidasys in 2015 [15] and Bifidochaetus in 2016 [16].

Diuronotus [17] is another recently described gastrotrich genus (2005), comprising two described species: Diuronotus aspetos Todaro et al. [17] and, Diuronotus rupperti Todaro et al. [17], and one undescribed species Diuronotus sp. [18–20], transferred from Halichaetonotus [17]. These are all found in marine interstitial environments of the North Atlantic; D. aspetos from Greenland [17] and Germany [2, 21], D. rupperti from Denmark [17] and Diuronotus sp. from North Carolina, USA [18]. Diuronotus was placed in Muselliferidae (Paucitubulatina, Chaetonitida) next to Musellifer [22] with which it shares the presence of a ciliated so-called ‘muzzle’ (or snout) and specific ultrastructural traits of scales and sperm [23].

Gastrotricha are divided into two main taxa: the supposedly monophyletic Macrodasyida and the possibly paraphyletic Chaetonotida, divided further into the Multitubulatina, (consisting of one genus, Neodasys, and possessing multiple adhesive glands) and the diverse Paucitubulatina (possessing generally only two adhesive tubes) [24]. Muselliferidae, belonging to Paucitubulatina, is the possible sister group to all remaining Paucitubulatina according to morphological [22, 25] and molecular [26, 27] studies. However, Paps and Riutorts [28] propose an alternative topology in which Xenotrichulidae is positioned as sister group of the remaining Paucitubulatina, and Muselliferidae being the sister group to Chaetonotidae. Kieneke et al. [29] find Proichthydiidae as sister group to the remaining Paucitubulatina, with Muselliferidae forming a clade together with Xenotrichulidae sister group to other Paucitubulatina. These different topologies overall suggest a key position of Muselliferidae within Gastrotricha, emphasizing the importance of this family for understanding the evolution of Gastrotricha. Indeed, some features of Muselliferidae, namely its marine habitat and well-developed hermaphroditism, are thought to be plesiomorphic character traits of Chaetonotida. However, detailed morphological studies on this family are lacking, most likely due to the paucity of these animals and their late discovery [26, 30].

Recently, a series of papers employing confocal laser scanning microscopy (CLSM) described the detailed muscular arrangement of several Paucitubulatina, namely Musellifer [22], Xenotrichulidae [22, 31], Chaetonotidae [22, 32], and Dasydytidae [11, 33], and notably, the helicoidal musculature, proposed to be a gastrotrich synapomorphy [34]. These recent reports were used to infer the plesiomorphic arrangement of the musculature of Gastrotricha as constituted by two ventro-lateral longitudinal muscles surrounded by outer circular muscles, and longitudinal splanchnic muscles surrounded by helicoidal and intestinal circular muscles [2]. In Paucitubulatina, the longitudinal muscles appear to be more numerous, and the outer circular muscles, if present, are incomplete and consist of dorso-ventral muscles [2]. These dorso-ventral or semi-circular muscles are found in marine chaetonotids [22], but are often missing or highly reduced in freshwater chaetonotids [11, 22, 33] emphasizing the importance of studying the marine Diuronotus in order to resolve their evolution and contribute to the broader understanding of muscular evolution within Gastrotricha.

To date, only one confocal study on Xenotrichula has described the nervous system of a member of Paucitubulatina in detail [12], while it has been extensively described for Multitubulatina (Neodasys) [13] and in several Macrodasyida with combined immunohistochemistry and CLSM (e.g., [35, 36]), or transmission electron microscopy (TEM) [37]. One of the conclusions of the Xenotrichula study [13] is the low structural variation of the nervous system within Gastrotricha, which always comprises a bilobed brain with a ventral commissure, a pair of anteriorly projecting longitudinal nerves, a pair of ventro-lateral nerve cords along the trunk, and a terminal commissure. These features were also interpreted as ancestral conditions of Gastrotricha in Kieneke and Schmidt-Rhaesa (2015) [2, 12]. Yet only a single Paucitubulatina, Xenotrichula, was considered for this state reconstruction. Moreover, substantial variation exists, such as the presence of an additional ventral nerve in Oregodasys cirratus Rothe & Schmidt-Rhaesa [38] and dorsal nerves in Xenodasys riedli (Schöpfer-Sterrer, 1969) [39, 40], or additional trunk commissures in Dactylopodola and Oregodasys cirratus. These studies highlight the unexplored diversity of gastrotrich nervous systems, which may be especially relevant in the diverse group of Paucitubulatina, in which only one study of the nervous system has been reported so far [13] and given the total lack of data on Muselliferidae.

Several studies have described the ultrastructure and repartition of protonephridia in Gastrotricha, with a few of them addressing species of Paucitubulatina [41, 42]. It has been suggested that members of Paucitubulatina always possess one pair of trunk protonephridia [41], although again, data on Muselliferidae are lacking. Each nephridium was found to encompass two monociliated terminal cells with coaxial cilia, a long canal cell, and a nephridopore cell [2].

In order to enhance our understanding of the evolution of major organ systems within Gastrotricha we acquired new morphological data on Diuronotus aspetos, using CLSM techniques and immunohistochemistry to describe the detailed arrangement of the musculature, nervous system, and ciliation. To assess the previously proposed relationship of D. aspetos within Muselliferidae, we analyzed the phylogenetic position within Chaetonotida, using molecular data. In this phylogenetic context, the morphology of Diuronotus is compared and discussed relative to other Chaetonotida, and Gastrotricha in general, leading to the proposition of several new homologies.

Methods

Collection

For Diuronotus aspetos, the samples were taken with a mini van Veen grab from shallow water (3–6 m water depth) of Flakkerhuk (69°38.63’N 51°51.13’W), Disko Island, West Greenland. All specimens were collected during the Arctic summer in August 2013. Sediment was well-sorted sand of fine to medium grain size. The specimens have been extracted with MgCl₂ narcotization and decantation.

For DNA, specimens of Xenotrichula sp. have been sampled in Ystad, Sweden (55°26.28’N 13°55.44’E) in subterranean environments on a beach with fine to medium-sized sand, and extracted with MgCl₂ narcotization and decantation. Marine Aspidiophorus sp. were sampled from cultures of Dinophilus gyrociliatus from Copenhagen University, where they are contaminants and unfortunately of unknown origin.

Sequence acquisition

Total genomic DNA was obtained from whole specimens using the Qiagen DNeasy Blood & Tissue Kit (Qiagen Inc., Valencia, CA, USA) following the manufacturer’s protocol, except for performing the DNA elution in 160 μL of AE buffer in order to increase the final DNA concentration.

Polymerase chain reactions (PCR) using Sanger based markers were prepared to a final volume of 25 μL with 12.5 μL of GoTaq® Green Master Mix (Promega Corporation, Madison, WI, USA), 1 μL of each primer (10 μM concentration), 10–8.5 μL of Milli-Q water (adjusted to amount of DNA template), and 0.5–2 μL of DNA template. Reaction mixtures were heated in a Bio-Rad G1000 Thermal Cycler at 94 °C for three minutes, followed by 35–40 cycles (primer specific) of 94 °C for 30 s, specific primer pair annealing temperatures for 30 s, and an extension at 68 °C for 45 s (unless indicated otherwise), and a final extension phase of 5 min at 72 °C. The COI primer set dgLCO1490/dgHCO2198 [43] was run with two cycling steps, both variable in temperature. COI annealing temperatures were 45 °C for 45 s and 51 °C for 45 s, respectively with extensions of 30s. Overlapping fragments of the small 18S rDNA (ca. 1800 bp) were obtained using paired primers corresponding to fragments 1 and 3 of the 18S rDNA [44]: (1) 18S1f/18S5R (ca. 900 bp) and (3) 18Sa2.0/18S9r (ca. 800 bp) both overlapping. Both primer sets (1) and (3) had annealing temperatures set to 49 °C. The 28S primer set used was 28SD3/28SG758 [45, 46] with an annealing temperature of 53 °C.

All newly generated and integrated sequences were deposited in the GenBank ® database with the following accession numbers KX531001 to KX531007 (Table 1).

Table 1 Sequences used for the phylogenetic reconstruction with their corresponding GenBank accession number. See material and methods for further information on the acquisitions of the sequences of Diuronotus aspetos

Full size table

Phylogenetic analysis

Sequences were cleaned on BioEdit [47], and a consensus has been realized from the reverse and forward sequences. Sequences were blasted on NCBI [48]. In parallel, COI, 18S and 28S of Diuronotus aspetos were found from its transcriptome [3], using Blastall from NCBI. Sangers and transcriptome acquired COI, 18S and 28S genes were aligned and compared, showing low quality and short length of Sangers sequences. Consequently, COI and 28S of the transcriptome were kept, while a consensus of 18S from the transcriptome and the Sangers sequencing was done having an identical overlapping segment. This hybrid approach was possible since the specimens used for the transcriptome and the Sanger sequences came from the same sample. Sequences of Aspidiophorus sp. and Xenotrichula sp. were added to the dataset. Sequences of other gastrotrichs acquired from GenBank, based on the tree proposed by Kånneby et al. 2012 [49] were added, selecting sequences from each genus (except Bifidochaetus [16]) for which sequences were not available at the time of the analysis), and representing the shortest and deepest branches possible. Sequences from Kånneby et al. [26] for Musellifer, Kånneby and Todaro [50] for Neogosseidae, and Todaro et al. [51] for the macrodasyidans, outgroups have additionally been collected from GenBank. Subsequently, the sequences were aligned gene per gene with Muscle in Seaview [47], checked by hand, and the three genes were concatenated with Sequence Matrix [52]. Finally, this dataset was analyzed with Bayesian inference in MrBayes 3.2.6 [53] under the model GTR + I + Γ. The gamma shape parameter, the substitution rates, the proportion of invariable sites, and the character state frequencies were all unlinked. The dataset was partitioned according to each gene and by codon position for COI and analyzed with four MCMC chains for each run, for 30,000,000 generations. Chains were sampled every 1000^th generations and the burn-in was set to 25 %. Convergence of the two runs as well as analysis quality was ascertained by checking the log likelihood graphs, the average standard deviation of split frequencies, and the model fit with Tracer [54].

Immunohistochemistry and CLSM

Specimens were anesthetized with isotonic magnesium chloride and fixed in 3.7 % paraformaldehyde in phosphate buffered saline (PBS) for 1 to 2 h at room temperature (RT), followed by six rinses in PBS and storage in PBS with 0.05 % NaN₃. Triple or quadruple stainings were performed for the investigation of muscular, nervous, glandular and ciliary systems, including F-actin staining (Alexa Fluor 488-labelled phalloidin, INVITROGEN, Carlsbad, USA), DNA-staining (405 nm fluorescent DAPI) and antibodies against neurotransmitters and tubulinergic elements (monoclonal mouse anti-acetylated α-tubulin (SIGMA T6793, St. Louis, USA), polyclonal chicken anti acetylated α-tubulin (SAB3500023-100UG), polyclonal rabbit anti-serotonin (5-HT, SIGMA S5545) and anti-FMRF-amide (IMMUNOSTAR 20091, Hudson, USA)). Prior to adding the primary antibody-mix, the samples were pre-incubated with 1 % PTA (PBS + 1 % Triton-X, 0.05 % NaN3, 0.25 % BSA, and 5 % sucrose) for 1 h. Samples were incubated over night at RT in primary antibodies mixed 1:1 with glycerol (in a final 1:400 concentration). Subsequently, specimens were rinsed in PBS six times and incubated with the secondary antibodies conjugated with fluorochromes over night at RT (mixed 1:1 with glycerol; 1:400 goat anti-mouse labeled with CY5 (JACKSON IMMUNO-RESEARCH, West Grove, USA, 115-175-062), 1:400 goat anti-mouse labeled with TRITC (JACKSON IMMUNO-RESEARCH, West Grove, USA, 115-175-062), 1:400 goat anti-rabbit labeled with TRITC (SIGMA T5268), and 1:200 goat anti-chicken labeled with Dylight (JACKSON IMMUNO-RESEARCH, West Grove, USA, 103-495-1550)). They were rinsed in PBS five times and one time in 1 % PTA and pre-incubated for 60 min in Alexa Fluor 488-labeled phalloidin (0.33 M in 1 % PTA). Thereafter, specimens were rinsed in PBS (without NaN₃) and mounted in Fluoromount-G with DAPI (SOUTHERN BIOTECHNOLOGY ASSOCIATES, Inc., Alabama, USA) or Vectashield with DAPI (VECTOR LABORATORIES, Burlingame, USA). The specificity of the antibodies was tested by examining specimens, where either the primary or secondary antibodies were omitted. Chicken anti acetylated α-tubulin staining did not produce satisfying results and is therefore not shown in this study (Sigma SAB3500023-100UG).

The mounted specimens were scanned using an Olympus Fluoview FV-1000 confocal laser scanning microscope (of K. Worsaae, University of Copenhagen, Denmark), with the acquired z-stacks of scans being either projected into 2D-images or analyzed three-dimensionally using IMARIS 7.1 (BITPLANE SCIENTIFIC SOFTWARE, Zürich, Switzerland). This software package was also used to conduct the measurements presented in the following text according to the conventions introduced by Hummon et al. [55], i.e. position in the body is given in units (U) as a relative measurements to total body length, measured from anterior to posterior. Schematic hand drawings and plate setup were done with Adobe Illustrator CS6 and image adjustments conducted in Adobe Photoshop CS6.

Results

Phylogeny

The tree (Fig. 1) shows a fully supported sister group relationship (100 % posterior probability (PP)) between the monophyletic Muselliferidae (100 % PP) and Xenotrichulidae (100 % PP), herein collectively called “group A”. Within Muselliferidae the genus Musellifer (100 % PP) (represented only by Musellifer delamarei (Renaud-Mornant [56]) and Musellifer reichardti Kånneby et al. [26]) is found to be the sister group to Diuronotus aspetos. Group A is found next to “group B”, together constituting the Paucitubulatina (100 % PP), with group B showing a monophyletic Dasydytidae (100 % PP) and Neogosseidae (100 % PP) nested within “Chaetonotidae”, the latter hereby becoming paraphyletic. Supports are high in all nodes of the tree, except for some of the numerous inner nodes of group B.

Musculature

The body wall musculature consists of several pairs of longitudinal muscles, numerous dorso-ventral muscles, a thin helicoidal musculature, semi-circular and complete circular muscles, as well as pharyngeal musculature. The pharyngeal musculature is especially dense and has an organization typical of chaetonotid gastrotrichs, as described below in more detail (Figs. 2 and 3).

Radial muscles

The pharynx, sensu stricto, is formed by three rows of dense radial pharyngeal muscles (rpm, Figs. 2d, e, m and 3c, d), and extend to U26 (units are calculated as length from anterior end, relative to total length, see material and methods). The radial muscles are cross-striated and each of them presents three to six Z-discs, which are less numerous anteriorly. The myoepithelial nuclei of the pharynx have a distinctive folded and elongated shape (mn, Fig. 2m). The pattern and repartition of these nuclei seems to be specific and corresponding nuclei could be found in the same position in different specimens.

Helicoidal muscles

Helicoidal muscles (hm, Figs. 2n and 3a, b, d, e) are very thin (0.5–1.2 μm) and limited to the anterior half of the specimen. It is difficult to confirm the presence of the helicoidal muscles closest to the pharynx due to the strong signal of other pharyngeal muscles (dashed lines in Fig. 3c-d). In a few locations along the midline of the pharynx very faint diagonal fibers were observed, suggesting that the helicoidal musculature is present along the entire pharyngeal region. Distinct helicoidal muscles are found extending from the midgut/pharynx junction at U26 until U42, enveloping the dorsal longitudinal muscle, but not the ventral or ventro-lateral longitudinal muscles.

Longitudinal musculature

Three longitudinal muscles span the entire body length: a pair of ventral longitudinal muscles (vlm, Figs. 2 and 3), a pair of ventro-lateral longitudinal muscles (vllm, Figs. 2 and 3), and a pair of dorsal longitudinal muscles (dlm, Figs. 2 and 3). The ventral longitudinal muscle bundle splits several times in a pattern described below for the different body regions.

Pharyngeal region

Several longitudinal muscles are present along the pharynx. Some are limited to the pharyngeal region, while others are the continuity of the body longitudinal muscles mentioned above. Two sets of muscles are strictly limited to the pharyngeal region: a pair of lateral and a pair of dorsal muscles. The lateral pharyngeal longitudinal muscle (lplm, Figs. 2d, e and 3c, d) extends adjacent to the pharynx along its entire length. The pharyngeal dorsal longitudinal muscles (pdlm, Figs. 2d, e, m, o and 3b-d) extend close to the pharyngeal midline along its total length, ventral to the dorsal longitudinal muscles. Moreover, several somatic and splanchnic longitudinal muscles supply the pharyngeal region.

The paired ventral longitudinal muscle (vlm, Figs. 2 and 3) originating in the head splits along the pharynx into a complex pattern (see Fig. 3b). One of its branches extends more laterally and splits into several sub-branches, supplying the lateral sides of the head.

The paired ventro-lateral longitudinal muscle (vllm, Figs. 2 and 3) lines the pharynx until reaching the head, where it bifurcates at U7, one branch extending ventro-laterally and the other dorso-laterally. Each of them subsequently splits into several minor branches, supplying the lateral sides of the head. These muscles together with the antero-lateral branch of the ventral longitudinal muscle (vlm), and the head diagonal muscle (hdm, see below) all supply the antero-lateral part of the head, and in addition to anchoring the longitudinal muscles for overall body contraction, they may function separately in contraction of the head (Figs. 2j and 3a, c).

The paired dorsal longitudinal muscle (pdlm) spans the anterior-most extremity of the pharynx.

Trunk region

Three main longitudinal muscles, i.e. ventral, lateral and dorsal, are supplying the trunk. The paths of the lateral and dorsal longitudinal muscles are relatively straight throughout the body. However, just posterior to the pharynx, the dorsal longitudinal muscle lines the intestine (Figs. 2f and 3c, d, e), while it runs closer to the dorsal body wall more posteriorly (Figs. 2g, h and 3f, g). The ventral longitudinal muscle splits into three muscle bundles at the anterior trunk. Two of these branches run in parallel mid-ventrally along the trunk (lvlm, mvlm, Figs. 2a, c, f-h, p and 3a, b, f-h), whereas the third branch extends dorso-laterally and supplies the dorso-lateral sides of the body until meeting the median-most branch at U86 (dvlm, Figs. 2a, g, l and 3a, b, f).

Posterior region

The median branch of the longitudinal ventral muscle splits posteriorly into two bundles at U86. One very short (8 μm) portion medially supplies the posterior part of the adhesive gland of the posterior tube, while another longer branch extends into the primary tube, supplying it for approximately two-thirds of its length to U97 (Fig. 3a). Additionally, the lateral branches of the ventral and lateral longitudinal muscles also extend along the primary tube. The dorsal longitudinal muscle supplies the anterior third of the secondary tube.

Diagonal muscles

The head diagonal muscle (hdm, Figs. 2j, p and 3a, b, c) forms a V-shape with two medially joined branches. The median part of the muscle is situated in the midline of the body in the posterior region of the head while the two extremities extend to the antero-lateral region of the head.

At the dorso-posterior pharynx, a pair of pharyngeal dorsal diagonal muscles decussate (pddm, Figs. 2p and 3b, d), ventral to the pharyngeal dorsal longitudinal muscle and the dorsal longitudinal muscle. Although their orientation is similar to that of the helicoidal muscles, their exclusively dorsal extension and their greater width differ significantly from those of the helicoidal muscles.

Three diagonal muscles are found in the furca, extending from one side to the contralateral one. The two anterior muscles (tmd, Figs. 2a, b, h, i, k and 3a, b, g, h) extend from the midline at U86, halfway to the primary and the secondary tube, respectively. The posterior diagonal muscle (pdm, Figs. 2i, k, and 3a, b, h) extends from U89 laterally into the first third of the secondary tube.

Circular muscles

Pharyngeal circular muscles (pcm 2E,M,O,P and 3A,B,C,D) are present around the pharynx. These muscles are numerous (ca. 110 in one specimen), positioned proximal to each other, and 1–1.50 μm thick with increasing diameter towards the posterior region.

Two sphincters are present at each extremity of the pharynx: one anterior pharyngeal sphincter (aps, Figs. 2b, o, p and 3a, b) located just posterior to the mouth, and one posterior pharyngeal sphincter (pps, Figs. 2m, o, p and 3a, b) marking the transition between the intestine and the pharynx. The anterior sphincter is smaller, being 1 μm thick and has a diameter of 17 μm, while the posterior sphincter is more prominent with 8 μm thickness and a diameter of 23 μm.

Supplementary circular muscles of the adhesive glands (cmag, Figs. 2h, i, k and 3a, b, g, h) are present in the tubes, forming a muscular layer around the large adhesive glands (ag, Fig. 2k). They thereby supply two cavities, one for each tube. The muscular layer surrounding the primary tube is smaller than the one surrounding the secondary tube, and both structures are connected. This suggests that both tubes are supplied by a single set of glands controlled by muscles. Three adjacent nuclei are found within the layer of circular muscles at the base of the gland (agn, Fig. 2k), near the anus, around U88.

Semi-circular muscles

Ventrally opened semicircular muscles (scm, Figs. 2l and 3a, b, f, g) are present in the posterior part of the specimen, but do not extend into the tubes. They originate ventrally, from each side of the body, and extend to the dorsal side. From there, they project to the contralateral side, external to the longitudinal musculature. They are more numerous in the posterior region, anterior to the furca, where they are separated by 2–5 μm. Semicircular muscles are less numerous and spaced further apart (5–8 μm) in the anterior region of the ovary. They appear to supply only the ovary region; their contraction probably reduces the body diameter and may be involved in the movement/release of eggs.

Dorso-ventral muscles

Numerous thin muscles (1–2 μm in diameter) traverse the entire trunk dorso-ventrally (dvm, Figs. 2a–c, e-h, k, l, n and 3a, b, e-g). These dorso-ventral muscles are spaced approximately 5 μm apart in the region between U18 and U95. In this region, two pairs are found laterally in transverse sections of the pharyngeal region (one external and one more internal pair, dvm, Figs. 2d, e and 3a, b). This number increases more posteriorly in the trunk, where up to five pairs of dorso-ventral muscles can be detected (dvm, Figs. 2g, k, l, and 3a, b, f). The dorso-ventral muscles extend dorso-ventrally between the different longitudinal muscles and the ciliary bands in various combinations. However, they are never found external to the ventro-lateral longitudinal muscles or between the pair of dorsal longitudinal muscles.

Nervous system

The nervous system of Diuronotus aspetos is described from acetylated α-tubulin-like immunoreactivity (LIR, Figs. 4, 5 and 6), serotonin-LIR (Fig. 7) and FMRF-amide-LIR (Fig. 8) (all different LIR of the head region are summarized in Fig. 9). Similar to previously investigated Gastrotricha, the nervous system consists of paired nerve cords, which originate from a bilobed dorsal brain, and extend posteriorly. In the following section, previously described and undescribed structures are detailed, such as: i) multiple pairs of longitudinal nerve projections in the head (danp, dlpn, hln, Figs. 6a, b, e, j and 9a, b); ii) paired anterior ventro-median nerves (avmn, Fig. 6a, d and 9b); iii) dorsal nerves posterior to the brain (hdpn, Figs. 6a, h and 9a); iv) paired ventro-lateral nerve cords (vlnc, Figs. 6b-d, f, g, k and 9b); v) paired posterior nerves, projecting into the adhesive tubes (nppt, Fig. 6k), vi) bilobed, dorsal brain with three commissures (main neuropil and anterior ventral and dorsal commissure, together forming a nerve ring encircling the pharynx) (np and anr, Figs. 6a-c, e, h-j and 9a, b); vii) two pairs of ganglia along the nerve cord: one anterior and one terminal (pgg and ang, Figs. 6b, f, k and 9b); viii) four ventral trunk commissures (spc, tvc, pac and pco, Figs. 6b, d, g, k and 9b); and ix) a pharyngeal nervous system, consisting of three longitudinal nerves (one per pharyngeal row of radial muscles) and supplementary minor nerves (Figs. 4 and 5). Additionally, the serotonin-LIR and FMRF-amide-LIR gave very detailed results, allowing us to collect precise data on the number, position and connection of perikarya (Figs. 7, 8 and 9).

Acetylated α-tubulin-like immunoreactivity (acetylated α-tubulin-LIR)

Acetylated α-tubulin-LIR provides information on most neurites, cilia as well as other portions of cytoskeletons of the cells. However, not all minor neurites of the nervous system are traced and the description focuses on the central nervous system and sensory structures.

Stomatogastric nervous system

The stomatogastric nervous system, confined to the pharynx, consists mainly of three main longitudinal nerves: a dorsal (dpn, Figs. 4a, d–h, j–n and 5a, b, e) and two ventro-lateral nerves (vpn, Figs. 4b, f–h, l–n and 5e), extending basally along the midline of each row of radial muscles (Fig. 4). The nerves are closely related to three structures: kinocilia, anterior pharyngeal glands and a pharyngeal canal system. The dorsal pharyngeal nerve (dpn, Figs. 4a, d–h, k–o and 5a, b, e) originates at the mouth, where it supplies a buccal nerve ring (bnr, Figs. 4a, b and 5a), encircling the mouth (probably innervating the anterior sphincter (aps, Figs. 2b, o, p and 3a, b) opening and closing the mouth). At U4, two anterior diagonal pharyngeal nerves (adpn Figs. 4a and 5a) originate from the dorsal nerve, extend antero-laterally to the anterior edges of the pharynx and medially join back the dorsal nerve. At U3, one pharyngeal dorso-ventral nerve (pdvn, Figs. 4a, b and 5a) originates from each of the anterior diagonal nerve, and supply ventrally a pharyngeal gland longitudinal nerve (plgn, Figs. 4b and 5c) innervating an anterior pharyngeal gland (apg, Figs. 4a, c, i and 5c) (which opens into the mouth). On the right side of the specimen, a dorso-anterior pharyngeal canal nerve (dpcn, Figs. 4a, d, j and 5a) extends posteriorly from the anterior diagonal nerve, possibly supplying the asymmetric dorsal pharyngeal canal (dpc, Figs. 4a, f, g, l, m and 5e). At U9 a pharyngeal nerve ring (pnr, Figs. 4a, b, e, k and 5b, d) supplying the ventral and dorsal nerves is present. A pair of paramedian dorsal pharyngeal nerves (dpnp, Figs. 4a, f, l and 5e) originates from the dorsal nerve at U19 and extends in a parallel fashion on each side, to fuse again with the dorsal nerve at U26 (Figs. 4a, g, m and 5e). The dorsal nerve extends more posteriorly where it innervates a two-celled pharyngeal posterior cluster (ppc, Figs. 4a and 5i) at the posterior margin of the pharynx. Two nerves extend the terminal part of the ventro-lateral pharyngeal sections: the lateral gland longitudinal nerve (plgn, Figs. 4a and 5c), and a median kinocilium longitudinal nerve (plkn, Figs. 4b, c and 5g), with the latter supplying a mouth and a pharyngeal kinocilia, respectively, at U1 and U6 (mk and pk, Figs. 4b-d, j, i and 5c, g). The gland and kinocilium nerves originate at U7 from an elongated ventro-lateral pharyngeal ganglion (vlpg, Figs. 4 and 5h, f), consisting of three nuclei and extending from U7 to U12 (probably integrating the signal collected by the two kinocilia and responsible for the putative terminal gland secrete release). Moreover, the ganglion seems to be related to the ventro-lateral pharyngeal canal (vlpc, Figs. 4a, b, e-g, k-n and 5f, i) described below. The ventral pharyngeal nerves (vpn, Figs. 4b, f, g, l, m and 5d) (supplied by perikarya at U15 and U19) extend from the ganglion, until U28.

Due to the unknown nature of the canal system and its main acetylated α-tubulin-LIR (as well as a weak FMRF-amide-LIR), it is described in this nervous system section. It consists of radially flattened cavities, sometimes asymmetrical, extending longitudinally within the pharynx (Figs. 4a, b, e, f, k, l and 5d, e). Six pharyngeal canals extend the pharynx: i) the unpaired right ventro-anterior pharyngeal canal (avrc, Figs. 4b, e, f, k, l and 5d) extending from U6 to U26; ii–iii) The paired ventro-lateral pharyngeal canals (vlpc, Figs. 4a, b, e-h, k-n and 5f, i), extending from U7 (at the level of the pharyngeal ganglia (vlpg, Figs. 4a and 5h, f)) to U30, and merging dorso-posteriorly; iv-v) the paired ventro-posterior pharyngeal canals extending from U24 for the left one (lpvc, Figs. 4b, g, h, m, n and 5d) to U28 and from U26 for the right one (rpvc, Figs. 4b, g, h, m, n and 5d) to U28; vi) the dorsal pharyngeal canal (dpc, Figs. 4a, f, g, l, m and 5e) extending along the right side from U6 to U15, then reaching the midline and bifurcating in two symmetrical branches, following the paramedian dorsal nerves of the pharynx between U19 and U27. Few nuclei are embedded in the pharyngeal canal system (Fig. 4a, b, g).

Central nervous system

The neuropil (np, Figs. 6a, c, e, h, j and 9a) is 14 μm thick and its center is positioned at U15. One 3 μm broad nerve extends antero-medially from the neuropil and branches laterally to form a dorsal and a ventral commissure at U12 and U9, respectively, together constituting an anterior nerve ring (anr, Figs. 5d, 6a-c, i, j and 9a, b). At the dorsal section of the anterior nerve ring (anr, Fig. 9), the acetylated α-tubulin-LIR is relatively weak, and the commissure consists of two transverse (anterior FMRF-amide-like-immunoreactive (LI-reactive) and posterior serotonin-LI-reactive) nerves, which eventually fuse dorso-laterally, forming the lateral sections of the anterior nerve ring. One anterior and one posterior short longitudinal nerves (cpn, Figs. 6b, i and 9b) extend from the ventral portion of the anterior nerve ring, innervating two median ciliary patches (described below). The neuropil supplies ventrally a pair of anterior ventro-median nerves (avmn, Figs. 6b, d, and 9b) extending between the anterior nerve ring and the post-pharyngeal ganglion posteriorly (pgg, Figs. 6b, f and 9b). It extends parallel to the pharyngeal median ciliated cell (pmcc, Fig. 10b, g), probably innervating it. Two pairs of dorso-median anterior nerves projections (danp, Figs. 6a, j and 9a) and two pairs of dorso-lateral anterior nerve projections (dlnp, Figs. 6a, j and 9a) extend from the anterior nerve ring and the lateral sides of the neuropil, respectively, projecting anteriorly. One pair of head lateral nerves (hln, Figs. 6a, b, e, j and 9a, b) extends from the lateral sides of the neuropil and bifurcates, posteriorly supplying a cell with a large and diffuse nucleus (possibly a gland cell (lgcb, Figs. 6a, e and 9a)), and anteriorly forming a nerve projection. Each of these anterior nerve projections probably innervates head sensory organs. Dorso-laterally, the posterior sides of the neuropil supply the ventro-lateral nerve cord (vlnc, Figs. 6b, c, d, f, g, j, k and 9b) of D. aspetos, which extends along the entire length of the specimen adjacent to the lateral longitudinal ciliary bands and the ventro-lateral longitudinal muscle, probably innervating these two structures. Two head dorso-posterior nerve (hdpn, Figs. 5e, 6a, h and 9a), extending along the pharynx, eventually supply the post-pharyngeal ganglion. They may innervate the anterior portion of the dorsal longitudinal muscle. A pair of head diagonal nerves (hdn, Figs. 6a, h and 9a) originates dorso-laterally of the neuropil, decussate dorsal to the pharynx at U22, and each extend ventro-laterally to a single perikaryon at U23. Comparison across specimens suggests that the position of these diagonal nerves corresponds to the position of the pharyngeal dorsal diagonal muscle (pddm, Figs. 2p and 3b, d), which it probably innervates. At U27 and U29, two thin nerves originate from the anterior ventro-median nerve, and form together at U28 a sub-pharyngeal commissure (spc, Figs. 6b, d and 9b). At U50, anterior to the testis, a thin trunk ventral commissure (tvc, Fig. 6g) is present. At U84, an anal ganglion (ang, Fig. 6k) of 6–8 cells supplies a pre-anal commissure (pac, Fig. 6k). Posterior to the anus, at U87, the two ventro-lateral nerve cords form the posterior commissure (pco, Fig. 6c, k), from which two nerve projections of the primary tube (nppt, Fig. 6c, k) extend.

Serotonin-like immunoreactivity (serotonin-LIR)

The nervous system shown by serotonin-LIR consists of a dorsal neuropil, the anterior nerve ring, anterior and posterior projections, the two ventro-lateral nerve cords, one posterior commissure as well as several perikarya.

Three main commissures (an anterior (sacn), a median (smcn) and a posterior (spcn)) in the brain neuropil show serotonin-LIR (Fig. 7a, d, e) as well as an isolated patch postero-median to the neuropil (spp, Fig. 7a, d, e). Three longitudinal nerves showing serotonin-LIR are found in the brain: i) the median-most brain nerve (smbn, Fig. 7a, d), ii) the paramedian brain nerve (spbn, Fig. 7a, d), and iii) the lateral brain nerve (slbn, Fig. 7a, d). Four very thin lateral nerves of the posterior commissure of the neuropil (slpn, Fig. 7a, d) form complex connections with the other nerves of the brain as well as to the dorso-median perikaryon (sdmp, Fig. 7a, c, d, e). A postero-lateral nerve node (spln, Fig. 7a, d, e) is present postero-laterally to the neuropil, being formed by the merging of several nerves, and supplies the ventro-lateral nerve cord (slnc, Fig. 7b, c, f). One dorso-lateral perikaryon (sdlp, Fig. 7a, c, d, e) supplies the postero-lateral nerve node, with a short nerve. The median-most brain nerve extends anteriorly from the posterior of the neuropil until U6 (Fig. 7a). The lateral brain nerve is short and extends from the postero-lateral nerve node, being supplied by some of the commissures of the neuropil (Fig. 7a). The paramedian brain nerve originates from the postero-lateral nerve node and supplies the serotonin-like-LI-reactive anterior nerve ring (sanr, Fig. 7a-d), subsequently extending more posteriorly as an anterior nerve projection until U4 (Fig. 7a, d). The anterior nerve ring consists dorsally of two and ventrally of one serotonin-LI-reactive nerves (sanr, Fig. 7a, b). The ventro-lateral nerve cord, consisting of two serotonergic-LI-reactive neurites, extends ventrally to supply a serotonin-LI-reactive para-pharyngeal cluster (sppc, Figs. 7b-e and 9b) with three perikarya, and extends to the posterior end forming the posterior commissure. Additionally, single serotonin-LI-reactive perikarya of the post-pharyngeal ganglion and of the anal ganglion are present respectively at U33 and U93 (spog, and spag, Fig. 7b, c, f) as well as nerve projections of the primary tube (snpt, Fig. 7c, f).

FMRF-amide-like immunoreactivity (FMRF-amide-LIR)

The FMRF-amide-LI-reactive nervous system consists of the brain neuropil, the anterior nerve ring, the anterior ventro-median nerve, the ventro-lateral nerve cord, the sub-pharyngeal commissure and the posterior commissure. Different parts of the nervous system show varying immunoreactivity intensities, as illustrated in Fig. 8.

The neuropil (fnp, Fig. 8a, c, d, f, g) consists of four connectives: two anterior and two posterior, supplied by several posterior and lateral perikarya. Antero-laterally to the neuropil, one pair of perikarya supplies a very short dorso-lateral anterior nerve projections (fpp, Fig. 8a, d, f, g) (corresponding to the base of the acetylated α-tubulin-LI-reactive projections (dlnp; Figs. 6a, j and 9a)). Additionally, three lateral perikarya of the brain (flpb, Fig. 8a, f, g) and a pair of dorso-posterior clusters of the brain (fdpc, Figs. 8a, f, g and 9a) with three perikarya, are present. Comparisons between differently stained specimens, and use of DAPI, enabled us to infer that the postero-median cell of the FMRF-amide-LI-reactive dorso-posterior cluster corresponds to the serotonin-LI-reactive dorso-posterior perikarya (sdmp, Fig.7; fdpc, Figs. 8a and 9a). Anteriorly, the neuropil supplies the anterior ventro-median nerve (fvmn, Figs. 8b, i and 9b), also supplied by two ventro-lateral perikarya of the brain (fvpb, Fig. 8a, e, i) and a ventral perikaryon of the anterior nerve ring (fvpr, Fig. 8b, i). The nerve ring (fanr, Fig. 8a-d, g, h, j) is supplied by one anterior and one posterior unpaired dorso-median perikarya (fpar, Fig. 8a, d, f, h). Ventro-posterior to the neuropil, a pair of tricellular clusters also supply the ventro-median nerve (fvnc, Fig. 8n, i), which extends further posterior until the two FMRF-amide-LI-reactive perikarya of the post-pharyngeal ganglion (fppg, Fig. 8b, c, k). The paired ventro-lateral nerve cords (flnc, Fig. 8b, c, i, k, l) is supplied by the postero-lateral part of the neuropil and by three anterior perikarya (fapn, Fig. 8b, c, e, i) (two anterior and one posterior, separated by 8 μm). The FMRF-amide-LI-reactive sub-pharyngeal commissure (fspc, Fig. 8b, i), ventro-lateral nerve cord, posterior commissure (fpco, Fig. 8c, l), and nerve projections of the primary tube (fnpt, Fig. 8c, l) follow the description of the acetylated α-tubulin-LIR.

Ciliation

The locomotor ciliation consists of a dense ventro-anterior ciliated area and two thin ciliated bands, which are extending to the posterior part of the specimen at U87 (Fig. 10c). This general pattern supports the original description of Todaro et al. [17], although numerous details can be added. CLSM allowed the identification of individual multiciliated cells and determination of their precise pattern.

Dorsally, the muzzle is covered by two transverse rows of multiciliated cells. The anterior row consists of three pairs of relatively small head dorso-anterior ciliated cells (3 μm, hacc, Fig. 10a, e, f), while the posterior row is constituted by a pair of larger head postero-lateral dorsal ciliated cells (hpcc, Fig. CA,E) and a head dorso-median ciliated cell (hmcc, Fig. CA,E) of similar size (6 μm). The pattern of the head lateral ciliated cells (hlcc, Fig. 10a, f) could not be resolved in details due to the dorso-ventral mounting of the specimen. However, at least four cells at the dorso-lateral level are present, and probably the same number at the ventro-lateral level.

The ventral head bears 20 multiciliated cells organized in four paired longitudinal rows and one median row, containing 2,2,3,2,2,2,3,2,2 cells (Fig. 10b). Posterior to the head, ventral to the pharynx, from U9, only two adjacent rows of multiciliated cells are present on each side, containing one large pharyngeal median ciliated cell (pmcc, Fig. 10b, g, 45 μm long) and three pharyngeal lateral ciliated cells (plcc, Fig. 10b, g, h, 20–35 μm long), respectively. Posterior to the pharynx, only one paired lateral row of cells is present, which extends until the posterior trunk, as described originally [17].

The two ventro-median ciliary patches at U7 and U11 (acp, pcp, Fig. 10b, h, i, position of patches measured from the center) are innervated by two short diffuse 5 μm wide longitudinal nerves (cpn, Figs. 6b, i and 7b), joining perpendicularly the anterior nerve ring. Each patch also shows an acetylated α-tubulin-LI-reactive positive ring around the ciliated area. The divergent morphology of these anucleate multiciliated cells and their close relation to the nervous system suggest that they could be sensory structures.

Several sensoria are scattered along the body (ss, Fig. 10d, j), and two pairs of pharyngeal sensory cilia (mk and pk, Figs. 3a-d, i, j, 4c, g and 10i) are located in the pharyngeal region (see the nervous system section for further details).

Two pairs of nephridia are found along the body (Fig. 10d): the anterior pair is situated ventro-laterally and the posterior pair is located dorso-laterally, relatively close to the midline. The anterior pair of protonephridia (apn, Fig. 10d, j) is situated anterior to the testis at U42 and the cilia are 20 μm long. The posterior pair of protonephridia (ppn, Fig. 10d, k) is situated at U74 with 15 μm long cilia. Each nephridium seems to possess two straight coaxial cilia (c, c’, Fig. 10j, k), thus resembling the general paucitubulatinian protonephridia with its two adjacent monociliated terminal cells projecting into a non-ciliated canal cell and ending with a nephridiopore epidermal cell [2, 41]. The canal cell and the nephridiopore cell have not been stained, which is why information on the orientation and opening of protonephridia in D. aspetos is lacking.