- Open Access
The role of ventral and preventral organs as attachment sites for segmental limb muscles in Onychophora
© de Sena Oliveira et al.; licensee BioMed Central Ltd. 2013
Received: 12 August 2013
Accepted: 27 November 2013
Published: 5 December 2013
The so-called ventral organs are amongst the most enigmatic structures in Onychophora (velvet worms). They were described as segmental, ectodermal thickenings in the onychophoran embryo, but the same term has also been applied to mid-ventral, cuticular structures in adults, although the relationship between the embryonic and adult ventral organs is controversial. In the embryo, these structures have been regarded as anlagen of segmental ganglia, but recent studies suggest that they are not associated with neural development. Hence, their function remains obscure. Moreover, their relationship to the anteriorly located preventral organs, described from several onychophoran species, is also unclear. To clarify these issues, we studied the anatomy and development of the ventral and preventral organs in several species of Onychophora.
Our anatomical data, based on histology, and light, confocal and scanning electron microscopy in five species of Peripatidae and three species of Peripatopsidae, revealed that the ventral and preventral organs are present in all species studied. These structures are covered externally with cuticle that forms an internal, longitudinal, apodeme-like ridge. Moreover, phalloidin-rhodamine labelling for f-actin revealed that the anterior and posterior limb depressor muscles in each trunk and the slime papilla segment attach to the preventral and ventral organs, respectively. During embryonic development, the ventral and preventral organs arise as large segmental, paired ectodermal thickenings that decrease in size and are subdivided into the smaller, anterior anlagen of the preventral organs and the larger, posterior anlagen of the ventral organs, both of which persist as paired, medially-fused structures in adults. Our expression data of the genes Delta and Notch from embryos of Euperipatoides rowelli revealed that these genes are expressed in two, paired domains in each body segment, corresponding in number, position and size with the anlagen of the ventral and preventral organs.
Our findings suggest that the ventral and preventral organs are a common feature of onychophorans that serve as attachment sites for segmental limb depressor muscles. The origin of these structures can be traced back in the embryo as latero-ventral segmental, ectodermal thickenings, previously suggested to be associated with the development of the nervous system.
The body plan of the Onychophora, the putative sister group of the arthropods, displays a combination of segmental and non-segmental features [1, 2]. While parts of the integument, muscular and nervous systems do not show any segmentation, segmental organisation is evident, such as in the arrangement of nephridia [3, 4], ostia of the heart [5, 6], crural glands [7, 8], limbs with their muscles, apodemes and nerves [1, 9–12], and the so-called ventral organs [13–15]. The ventral organs are arguably one of the most controversial structures in Onychophora, as their development, anatomy, relationship to other structures and consequently their function remain unclear [1, 16–22]. Initially, the ventral organs were described as embryonic segmental, paired thickenings of the ventro-lateral ectoderm, which arise late in onychophoran development [23–25], but the term ventral organ has also been applied to segmental, cuticular structures that occur mid-ventrally between each leg pair in adult onychophorans [7, 8, 13–15, 26–28]. Whether the adult ventral organs are remnants of the embryonic thickenings [1, 8, 17, 23, 24] or whether these thickenings are transitory structures that disappear completely during development [19–21] is controversial.
Moreover, the role of the embryonic ventral organs is ambiguous , as they have been regarded either as rudiments of an ancient locomotory system comparable to the ventral ciliated areas of annelids , or as segmental anlagen of the nervous system or as rudimentary ganglia [19–22, 30]. Based on the last assumption, the embryonic ventral organs of onychophorans have been homologised with homonymous structures found in embryos of chelicerates and myriapods, in which they appear as paired, segmental epithelial vesicles that are involved in neurogenesis and incorporated into the segmental ganglia during development [2, 19, 20, 31, 32]. However, subsequent studies have demonstrated that the ventral organs are not associated with the nervous system in the onychophoran embryo as they arise after the presumptive nerve cords have formed [17, 29]. Thus, the homology of the onychophoran ventral organs to the homonymous structures in chelicerates and myriapods is unlikely [1, 17].
To further complicate matters, additional structures called “preventral organs” have been described from adults of a number of species of onychophorans [7, 13–15]. These structures are morphologically similar to the ventral organs, but they are smaller and located anterior to each ventral organ [14, 15]. Currently, neither the embryonic fate nor the relationship of the preventral organs to the embryonic and adult ventral organs of onychophorans is known. To clarify whether the ventral and preventral organs are a common feature of Onychophora, we analysed the anatomy of eight species of velvet worms, including representatives of the two major onychophoran subgroups, Peripatidae and Peripatopsidae. In addition, we have documented the embryonic fate of the ventral and preventral organs in embryos of Euperipatoides rowelli using in situ hybridization, histochemistry and immunocytochemical methods, in conjunction with confocal microscopy, to gain insights into their function.
Position and structure of the ventral and preventral organs
Morphogenesis of the ventral and preventral organs
During embryogenesis of the peripatopsid Euperipatoides rowelli, the ventral and preventral organs arise from single paired, segmental, ectodermal thickenings (Figure 2A–L). DNA labelling of embryos at subsequent developmental stages revealed that the thickenings appear early in development as segmentally repeated undulations of the ventrolateral ectoderm (Figure 2A–E). Initially, the thickenings of each body side are widely separated from each other by the ventral extraembryonic tissue, which is reduced during development, while the ventrolateral ectoderm of both sides fuses along the ventral midline (Figure 2A–F). After this fusion, the unitary anlagen of the ventral and preventral organs are recognisable as paired segmental structures that are separated by repeated transverse furrows along the body (arrows in Figure 2F). At this developmental stage, the paired thickenings, i.e., the anlagen of the ventral and preventral organs, occupy nearly the entire ventral body surface (Figure 2F, G).
Expression of Delta and Notch in the anlagen of the ventral and preventral organs
No evidence for a relationship of the ventral and preventral organs to the nervous system
Musculature associated with the ventral and preventral organs
The issue of the “ventral organs”: Homonymy does not necessarily imply homology
The term ventral organs has been commonly applied to paired segmental thickenings in the ventral ectoderm of the onychophoran embryo [1, 16, 19, 20, 23, 25]. These thickenings have been demonstrated to persist as midventral, segmental rudiments with an unknown function in adults [1, 24]. However, our developmental data from Euperipatoides rowelli show that each pair of segmental thickenings gives rise to two paired structures, i.e., the ventral and preventral organs, both of which persist in adults. Although the paired nature of the ventral and preventral organs is mostly evident in embryos of late developmental stages, it is still recognisable in adults, as these structures are subdivided in two halves by a longitudinal, sclerotized internal ridge. Based on these findings, it seems clear that neither the ventral and preventral organs nor their anlagen correspond to the embryonic structures called “ventral organs” in chelicerates and myriapods [2, 20, 31, 32]. While the ventral and preventral organs represent two paired structures per segment that persist in the epidermis of adult onychophorans, the homonymous “ventral organs” of chelicerates and myriapods arise as a single pair of vesicles per segment that are incorporated into each developing ganglion and disappear completely during development [2, 17, 19, 20, 31, 32].
Moreover, in contrast to the “ventral organs” of chelicerates and myriapods, neither the ventral and preventral organs of onychophorans nor their embryonic anlagen arise as segmental ectodermal invaginations. The only invaginations of the ventral ectoderm in the onychophoran embryo are the anlagen of the hypocerebral organs in the antennal segment, which entirely lose their connection to the epidermis during development and become associated with the ventral surface of the brain [17, 23–25]. The adult hypocerebral organs might be neurosecretory glands, homologous to the corpora allata of insects [21, 22, 34]. However, their serial homology to the ventral and preventral organs in onychophorans is uncertain  and there is no evidence for their homology to the embryonic “ventral organs” of chelicerates and myriapods, as they show an entirely different structure and fate . Thus, neither the hypocerebral organs nor the ventral and preventral organs of onychophorans are likely to be homologous to the “ventral organs” of chelicerates and myriapods.
Expression of Delta and Notch correlates with the embryonic origin of the ventral and preventral organs
Our gene expression data revealed two paired domains of Delta and Notch homologs in each embryonic segment of Euperipatoides rowelli, corresponding to regions that give rise to the ventral and preventral organs in this species. Exactly like the ventral and preventral organs, the posterior paired domain (corresponding to the anlagen of the ventral organ) is larger than the anterior domain (corresponding to the anlagen of the preventral organ). A similar pattern of expression was revealed in two previous studies (Figure 4B in ; Figure 4C in ) of the closely related species Euperipatoides kanangrensis. However, at the time when these studies [35, 36] were carried out, the striking spatio-temporal correlation of this expression pattern with the paired anlagen of the ventral and preventral organs was unknown, and so the authors assumed an exclusive function of Delta and Notch as proneural genes in these regions of the developing embryo. In our view at least two arguments speak against this assumption. First, most neurons have already been segregated from the neuroectoderm at stages when Delta and Notch are expressed in two paired domains [1, 17, 35, 36]. Second, the characteristic pattern of their domains, with a smaller anterior and a larger posterior pair, does not correspond to any known neural structure in onychophorans [10, 11, 29, 37–39]. We therefore suggest that the double-paired Delta and Notch domains specify regions of the ectoderm that give rise to the ventral and preventral organs rather than to the neurogenic tissue in the onychophoran embryo.
It is well known that apart from functioning as “proneural genes” [40, 41], Delta and Notch (Notch/Delta signalling) are involved in many other developmental processes, including cell death, cell division, and endocycle (DNA replication without an intervening mitosis) [42–46]. We have shown here that cell death and numerous cell divisions occur in the anlagen of the ventral and preventral organs in Euperipatoides rowelli. In addition, there is evidence that most of the cells in their anlagen might enter endocycle to increase biosynthetic activity . Therefore, it might well be that in this case the genes Delta and Notch govern these processes in the anlagen of the preventral and ventral organs rather than being involved in neurogenesis, as suggested previously [35, 36].
The ventral and preventral organs serve as attachment sites for the ventral limb depressor muscles
Although the function of the ventral and preventral organs in adult onychophorans has remained unknown, they were affiliated with the nervous system, as they were thought to be connected to the nerve cords via a pair of cell strands [16, 24]. Our data indeed show that each median commissure (as well as each ring commissure and each leg nerve) is accompanied by “glial” cells that might give the impression of cellular strands linking the ventral/preventral organs to each nerve cord. However, our detailed examination of complete series of histological and Vibratome sections, in conjunction with light, fluorescent and confocal microscopy, revealed no connections between the median commissures (which comprise neural tissue) and the ventral and preventral organs (which belong to the epidermis). Instead, they are clearly separated from each other by a thick layer of extracellular matrix and several layers of musculature and this spatial separation has also been reported from the onychophoran embryo (Figure 11C in ref. ).
In addition, we have shown herein that bundles of tracheal tubes are commonly associated with the ventral and preventral organs and they usually take a course towards each nerve cord by following the median commissures. This suggests that previous authors [16, 24] might have misinterpreted the cells associated with tracheal tubes, the only cellular structures that pass through the extracellular matrix in onychophorans, as tissue connections between the nerve cords and the ventral organs. Moreover, from the perspective of functional morphology, a cellular bridge between the nerve cords and the ventral and preventral organs is unlikely to exist in Onychophora for two reasons: First, most median commissures pass to the contralateral side in regions in which the ventral and preventral organs are lacking and, second, the ventral and preventral organs are segmental structures, whereas the median commissures are repeated in a non-segmental fashion along the body [1, 10, 11, 29]. Due to this lack of a corresponding spatial relationship between the ventral and preventral organs and the median commissures, it is unlikely that the ventral and preventral organs are in any way related to the nervous system.
Our data instead suggest that these structures are associated with the onychophoran leg musculature. According to our findings, the onychophoran ventral and preventral organs consist of epidermal cells that are covered by a thick cuticle, which forms hollow, sclerotized internal ridges. These ridges resemble typical apodemes of onychophorans and arthropods [12, 47]. Furthermore, phalloidin- rhodamine labelling revealed that the anterior leg depressor muscles (see ref.  for the nomenclature) attach to each preventral organ, whereas the posterior leg depressor muscles connect to each ventral organ, which also holds true for the depressor muscles of the slime papillae. The presence of two leg depressor muscles might explain why the ventral and preventral organs comprise two paired rather than a single paired structure in each leg-bearing segment. Based on these findings, we suggest that the ventral and preventral organs of onychophorans serve as attachment sites for the paired, extrinsic limb depressor muscles.
According to our findings, the ventral and preventral organs are a common feature of onychophorans, as they occur in representatives of Peripatidae and Peripatopsidae. During development, they arise from segmental thickenings of the embryonic ectoderm and persist in adults as sclerotized structures that serve as attachment sites for segmental limb depressor muscles. Whether the ventral and preventral organs are a derived feature of Onychophora or whether they are remnants of sclerotized, vaulted body rings described from fossil lobopodians  – putative stem-lineage representatives of Onychophora, Tardigrada, Arthropoda and/or Panarthropoda [49, 50] – is open for discussion.
Species of onychophorans studied and corresponding locality data
Epiperipatus sp. 1
Reserva Particular do Patrimônio Natural Estação Ambiental de Peti, 43°22′02.16224'' W, 19°53′33.44741'' S, 760 m, municipality of São Gonçalo do Rio Abaixo, Minas Gerais, Brazil
Epiperipatus sp. 2
PCH Porto das Pedras, 52°32′33.64'' W, 19°28′44.31'' S, 360 m, Mato Grosso do Sul state, municipality of Chapadão do Sul, Brazil
Epiperipatus biolleyi (Bouvier, 1902)
Los Juncos, 10°01′27.62'' N, 83°56′30.26'' W, 1760 m, Cascajal de Coronado, Province of San José, Costa Rica
Principapillatus hitoyensis Oliveira et al., 2012
Reserva Biológica Hitoy Cerere, 09°40′21.56'' N, 83°02′36.97'' W, 300 m, Province of Limón, region of Talamanca, Costa Rica
Plicatoperipatus jamaicensis (Grabham & Cockerell, 1892)
Ecclesdown, eastern slope of John Crow Mountains, eastern coast of the island, Jamaica (more precise data unavailable)
Euperipatoides rowelli Reid, 1996
Tallaganda State Forest, 35°26′ S, 149°33′ E, 954 m, New South Wales, Australia
Metaperipatus blainvillei (Gervais, 1837)
A forest near Lago Tinquilco, 39°09′ S, 71°42′ W, 815 m, IX Region de la Araucania, Chile
Metaperipatus inae Mayer, 2007
Forest near Contulmo, 38°01′ S, 73°11′ W, 390 m, VIII Region del Biobio, Chile
Stereomicroscopy and scanning electron microscopy
Adult specimens of all eight species studied (Table 1) were preserved in 70% ethanol and their ventral body surface was analysed with a stereomicroscope (Leica WILD M10, Leica Microsystems, Wetzlar, Germany) equipped with a digital camera (PCO AG SensiCam, Kelheim, Germany). For scanning electron microscopy, specimens of Epiperipatus biolleyi, Principapillatus hitoyensis, Plicatoperipatus jamaicensis, Metaperipatus blainvillei and Metaperipatus inae were fixed in 4% formaldehyde in PBS at room temperature. After several washes in water, the specimens were cut into suitable pieces, dehydrated in an ethanol series, dried in a CPD 030 Critical Point Dryer (BAL-TEC AG, Balzers, Liechtenstein), coated with gold in a SCD 040 Sputter Coater (BALZERS UNION, Balzers, Liechtenstein), and examined in a Quanta 200 Scanning Electron Microscope (FEI, Hillsboro, Oregon, USA). Embryos were dissected from some specimens after fixation and prepared in the same way as the body pieces from adults. Freshly moulted skins obtained from living specimens of Principapillatus hitoyensis, Metaperipatus blainvillei and Metaperipatus inae were spread on water surface according to Holliday , dehydrated in an ethanol series and processed further for scanning electron microscopy as described for the entire specimens.
Histology, Vibratome sectioning and light and confocal laser-scanning microscopy
For histological studies, specimens of Metaperipatus blainvillei were fixed in Bouin’s fluid as described previously [55, 56]. The specimens were dehydrated in an ethanol series, incubated in methylbenzoate and butanol, and embedded in Paraplast Tissue Embedding Medium (Kendall, Mansfield, MA, USA). Complete series of 5–7 μm thin sections were made with steel blades on a microtome (Reichert-Jung, 2050-supercut, Reichert Inc., Buffalo, NY, USA) and the sections were stained using Heidenhain’s  Azan staining method. The sections were then mounted on glass slides in Malinol and analysed under a light microscope (Leica Leitz DMR, Leica Microsystems) equipped with a digital camera (PCO AG SensiCam).
For Vibratome sectioning, specimens of Metaperipatus blainvillei, Euperipatoides rowelli, Principapillatus hitoyensis and Epiperipatus sp. 2 were cut in pieces and fixed overnight in 4% PFA in PBS at room temperature. The samples were then washed in several changes of PBS and either processed immediately or kept for several weeks in PBS containing 0.05% sodium azide at 4°C. Two embedding media were used for different body parts: (1) 6% agarose at 60°C, which was cooled down to room temperature as described previously , and (2) a 4:1 mixture of albumin/gelatine (3.75 g of albumin [Sigma-Aldrich, St. Louis, MO, USA, Grade II] in 10 ml distilled water; 0.5 g gelatine [Sigma-Aldrich, Type A] in 2.5 ml distilled water); the albumin/gelatine blocks were then fixed overnight in 10% PFA at 4°C and washed for 30 min in PBS at room temperature. The agarose and albumin/gelatine blocks were trimmed and sectioned into series of 100–200 μm thin sections with steel blades on a Vibratome (Vibratome Company, Saint Louis, USA). For morphological analyses of tracheal tubes, the sections were mounted on glass slides in 70% glycerine in PBS, covered with coverslips and imaged under a light microscope (Leica Leitz DMR) equipped with a digital camera (PCO AG SensiCam). For studies of myo- and neuroanatomy, the sections were processed for histochemistry and immunohistochemistry as described below, mounted on glass slides in Vectashield Mounting Medium (Vector Laboratories, Burlingame, CA) and analysed with the confocal laser-scanning microscopes Leica TCS STED (Leica Microsystems) and Zeiss LSM 510 META (Carl Zeiss MicroImaging GmbH, Jena, Germany).
DNA labelling and detection of fragmented DNA or cell death
Embryos of Euperipatoides rowelli stored in methanol were rehydrated in PBS and either labelled with one of the DNA-selective fluorescent dyes (Hoechst [=Bisbenzimide, H33258; Sigma-Aldrich; 1 μg/ml in PBS], SYBR® Green [Invitrogen, Carlsbad, CA, USA; 1:10,000 in PBS], Propidium Iodide [Roth, Karlsruhe, Germany; 1:3,000 in PBS], or RedDotTM2 [Biotium, Hayward, CA, USA; 1:250 in PBS]) for 1 h or used for the detection of fragmented DNA or cell death in conjunction with DNA labelling, as described previously . For this purpose, the embryos were incubated in 0.1 mol/L sodium citrate (pH 6.0) for 30 min at 70°C and rinsed in PBS containing 1% Triton1 X-100 (=PBS-TX; Sigma-Aldrich). Detection of apoptotic cells was carried out with the in situ Cell Death Detection Kit, TMR red (Roche, Mannheim, Germany). The embryos were placed in equilibration buffer for 10 min at room temperature and the buffer was replaced with label solution (450 μl) and enzyme solution (50 μl) according to the manufacturer’s protocol. After incubation on a nutator (3 h at 37°C), the embryos were counterstained with Bisbenzimide (Sigma-Aldrich) and mounted in Vectashield Mounting Medium (Vector Laboratories). For negative controls, the embryos were treated in the same way but without adding the enzyme. These embryos showed no nuclear labelling. For positive controls, the embryos were treated with DNase I recombinant, RNase-free (Roche), prior to detection of cell death. In these embryos, all nuclei were labelled. All embryos were analysed with the confocal laser-scanning microscopes Leica TCS STED (Leica Microsystems) and Zeiss LSM 510 META (Carl Zeiss MicroImaging GmbH) as described for the Vibratome sections.
Histochemistry and immunocytochemistry
Histochemical and immunocytochemical experiments on embryos and Vibratome sections of adult specimens of Euperipatoides rowelli were carried out as described previously [1, 17, 18, 58, 59]. Different fluorescent dyes and antisera were used either separately or in combination. As a general marker of neural structures, we used a monoclonal anti-acetylated α-tubulin antibody (Sigma-Aldrich; diluted 1:500 in PBS). As a mitosis marker, we applied a polyclonal anti-phospho-histone H3 antibody (=α-PH3; Upstate, Temecula, CA, USA; diluted 1:500 in PBS). For f-actin labelling, Vibratome sections and embryos were incubated overnight at room temperature in a solution containing phalloidin-rhodamine (Invitrogen) as described previously . Some Vibratome sections and embryos were counterstained with one of the DNA-selective fluorescent dyes described in the section “DNA labelling and detection of fragmented DNA or cell death”. In addition, to reveal “glial” cells, nerve cords were dissected from adult specimens of Euperipatoides rowelli, fixed overnight at room temperature in 4% PFA, rinsed several times in PBS and labelled with one of the DNA-selective dyes as described above. The Vibratome sections and embryos were mounted on glass slides or between two coverslips either in Vectashield Mounting Medium or in Vectashield Hard Set Mounting Medium (Vector Laboratories). The nerve cords were dehydrated in an ethanol series and mounted between two coverslips in methyl salicylate. All samples were analysed with the confocal laser-scanning microscopes Leica TCS STED (Leica Microsystems) and Zeiss LSM 510 META (Carl Zeiss MicroImaging GmbH).
Gene expression studies
RNA was isolated from embryos of Euperipatoides rowelli using TRIzol® Reagent (Invitrogen). Library preparation and assembly of the embryonic transcriptomes were performed as described by Hering et al. . To identify the Er-Delta and Er-Notch homologs, local tBLASTn searches were performed with the corresponding sequences from a closely related species of Onychophora, E. kanangrensis[35, 36]. The identified Er-Delta and Er-Notch sequences [GenBank accession numbers: KF322113 and KF322114, respectively] were used to design specific primers and to amplify the Er-Delta and Er-Notch fragments. The PCR products were cloned into the Escherichia coli strain KJ100 using pGEM®-T Vector System (Promega Corporation, Madison, WI, USA). Each vector was linearized and transcribed in vitro using DIG RNA Labelling Kit SP6/T7 (Roche, Mannheim, Germany).
For in situ hybridization, the embryos were rehydrated in PBST (PBS + 0.1% Tween-20; 5 min each). Pre-hybridization was carried out for 6 h at 60°C, after which the probe was added to the embryos and incubated overnight at 60°C. Excess probe was removed by several rinses in hybridization buffer, saline-sodium citrate buffer + 0.1% Tween-20 and PBST, after which the embryos were incubated for 3 h in a blocking solution (10% normal goat serum in PBST) at room temperature. The embryos were then incubated with the Anti-DIG-AP antibody (Roche; diluted 1:1000 in blocking solution) overnight at 4°C. After several washes in PBST, a NBT/BCIP staining solution (Roche) was added to the embryos and the staining reaction was kept in the dark. The embryos were checked every 10 to 15 min under a dissection microscope in dimmed light until staining was evident and the staining reaction was stopped by several washes with PBST. To denature the alkaline phosphatase, the embryos were post-fixed in 4% paraformaldehyde, transferred into 1.5 ml reaction tubes and stored at 4°C. After staining, embryos were either analysed under a light microscope (Leica Leitz DMR, Leica Microsystems) equipped with a digital camera (PCO AG SensiCam) or counterstained with the DNA-selective dye Bisbenzimide as described above (see the section “DNA labelling and detection of fragmented DNA or cell death”) and analysed with the confocal laser-scanning microscope Zeiss LSM 510 META (Carl Zeiss MicroImaging GmbH).
Confocal image stacks were processed with Leica AS AF v2.3.5 (Leica Microsystems) and Zeiss LSM IMAGE BROWSER v184.108.40.206 (Carl Zeiss MicroImaging GmbH). Optimal quality light, confocal and scanning electron micrographs were achieved by using Adobe (San Jose, CA, USA) Photoshop CS5.1. Final panels were designed with Adobe Illustrator CS5.1 and exported in the Tagged Image File Format.
The authors are thankful to the staff of the Instituto Nacional de Biodiversidad (INBio, Heredia, Costa Rica), the National System of Conservation Areas (SINAC, MINAE, Costa Rica), the Instituto Chico Mendes de Conservação da Biodiversidade (ICMBio, Brazil), the Instituto Estadual de Florestas (IEF-MG, Brazil) and the National Parks and Wildlife NSW and State Forests NSW (New South Wales, Australia) for providing permits. We gratefully acknowledge David Rowell, Paul Sunnucks, Paul Whitington, Klaus Wolf, Stefan Schaffer, Lars Hering, Franziska Franke, Sandra Treffkorn and Michael Gerth for their help with collecting the specimens, Susann Kauschke for technical support and maintaining the animals, Christine Martin for DNA labelling of nerve cords and Andreas Weck-Heimann for assisting with scanning electron microscopy at the Senckenberg Natural History Collection (Dresden, Germany). GM is thankful to Paul Whitington for his continuous support. This study was supported by a PhD fellowship of the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq: 290029/2010-4) to ISO. GM is a Research Group Leader supported by the Emmy Noether Programme of the German Research Foundation (DFG: Ma 4147/3-1).
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