- Open Access
A new look at the ventral nerve centre of Sagitta: implications for the phylogenetic position of Chaetognatha (arrow worms) and the evolution of the bilaterian nervous system
© Harzsch and Müller; licensee BioMed Central Ltd. 2007
- Received: 04 February 2007
- Accepted: 18 May 2007
- Published: 18 May 2007
The Chaetognatha (arrow worms) are a group of marine carnivores whose phylogenetic relationships are still vigorously debated. Molecular studies have as yet failed to come up with a stable hypothesis on their phylogenetic position. In a wide range of metazoans, the nervous system has proven to provide a wealth of characters for analysing phylogenetic relationships (neurophylogeny). Therefore, in the present study we explored the structure of the ventral nerve centre ("ventral ganglion") in Sagitta setosa with a set of histochemical and immunohistochemical markers.
In specimens that were immunolabeled for acetylated-alpha tubulin the ventral nerve centre appeared to be a condensed continuation of the peripheral intraepidermal nerve plexus. Yet, synapsin immunolocalization showed that the ventral nerve centre is organized into a highly ordered array of ca. 80 serially arranged microcompartments. Immunohistochemistry against RFamide revealed a set of serially arranged individually identifiable neurons in the ventral nerve centre that we charted in detail.
The new information on the structure of the chaetognath nervous system is compared to previous descriptions of the ventral nerve centre which are critically evaluated. Our findings are discussed with regard to the debate on nervous system organisation in the last common bilaterian ancestor and with regard to the phylogenetic affinities of this Chaetognatha. We suggest to place the Chaetognatha within the Protostomia and argue against hypotheses which propose a deuterostome affinity of Chaetognatha or a sister-group relationship to all other Bilateria.
- Ground Pattern
- Ventral Ganglion
- Vestibular Ganglion
- Soma Cluster
- Medial Bundle
The Chaetognatha (arrow worms) are bilaterally symmetrical marine carnivores and among the most abundant planktonic organisms. To date, about 120 described species are known worldwide from all vertical ranges of the ocean. Most of them are permanently pelagic but several epibenthic species are also known [1–3]. The chaetognaths range in length from 1–120 mm and are characterized by the presence of horizontally projecting fins and, at the anterior end, two groups of moveable, cuticular grasping spines used in capturing prey. Planktonic specimens are usually glassily transparent. Rapid bursts of swimming caused by dorso-ventral undulations alternate with phases during which the animals lie motionless and sink. Chaetognaths are hermaphroditic and develop directly so that newly hatched larvae display a body organisation that is in many aspects similar to the adult. Their phylogenetic affinities are controversial [4, 5]. Chaetognaths have traditionally been placed within the Deuterostomes mainly based on the differentiation of the archenteron seemingly resembling enterocoeli [reviewed in [1, 6–8]]. However, Kapp  emphasizes the phylogenetically isolated position of the Chaetognatha and designates them as incertae sedis. Nielsen [3, 9], on the other hand, unites the Chaetognatha together with the Rotifera and Gnathostomulida in the taxon Gnathifera thus suggesting a placement within the Protostomia. Important morphological characters in this debate are e. g. the coelomic epithelia and coelom formation [6, 10–12]. Chaetognath cleavage has traditionally been perceived as "radial" and thus as suggesting a deuterostome affinity [discussed in ]. However, a recent marking experiment of the first cleavage stages shows a spiral cleavage configuration of the four cell stage thereby suggesting a protostomian relationship . A position within the Protostomia is also supported by ultrastructural features of the brain  and by analyses of the genes that code for intermediate filament proteins [15, 16].
General information on the chaetognath morphology and anatomy has been summarised in the classical, histological contributions by Hertwig , Kuhl  and more recently in reviews by Goto and Yoshida , Bone and Goto , Kapp [1, 21], Nielsen , and Ax ; the most detailed review of their anatomy is probably that of Shinn . Chaetognaths have a complex nervous system that is largely epidermal. The general organisation of their nervous system has been examined by Bone and Pulsford , and Goto and Yoshida [[19, 24]; reviews [2, 20]; see Fig. 2A, B]. More specifically, the fine structure of the brain has received much attention [14, 19, 25, 26]. The layout of the neuromuscular innervation  and the ultrastructure of neuromuscular junctions  have been described. Much attention has also been focused on sensory organs such as ciliated receptor neurons [23, 29–32], the eyes [24, 33–35], and conjunction with this also on mechanisms of positive phototactical behaviours [36, 37]. Three studies have used immunohistochemical techniques to examine the distribution of serotonin and RFamide-like immunoreactive neurons [27, 38] and aspartate immunoreactivity  in the central and peripheral parts of the nervous systems.
Clearly, the phylogenetic position of arrow worms is unstable in both morphological and molecular studies. Yet, the recent molecular studies on mitochondrial genomes and ESTs seem to favor a position within or close to the Protostomia. Structure and development of the nervous system have always provided strong and important arguments in the discussion on metazoan phylogeny ("neurophylogeny") . Recent examples are the evolution from nerve nets to more centralised nervous systems [55, 56], the impact of apical organs, ciliary bands and serotonergic neurons on our understanding of bilaterian evolution [57–59], the fundamental phylogenetic relationships within the Arthropoda [60–62], a possible dorsoventral axis inversion during evolution towards the vertebrate CNS [55, 63, 64], or the structure of the ancestral bilaterian and chordate brain [65–69]. In the light of the conflicting hypotheses on chaetognath phylogeny, the current paper sets out to give a brief overview over the previous knowledge of the structure of the chaetognath nervous system and to explore the structure of the ventral ganglion in more detail than has been available so far. One main goal of our study was to add a new set of neuroanatomical characters to the discussion on the phylogenetic position of the Chaetognatha. Furthermore, with these data we want to contribute to the debate of how the nervous system of the last common ancester of the Bilateria may have looked like [56, 64, 68, 70].
Structure of the chaetognath central nervous system: current knowledge
A brief description of the layout of the adult arrow worm central nervous system will serve as the basis for illustrating our own results. It consists of six ganglia in the head, one unpaired ventral ganglion in the body, nerve tracts connecting these ganglia and peripheral nerves passing out of these ganglia [2, 19, 20] (Fig. 2). The ganglia in the head are the cerebral ganglion (the brain), a pair of vestibular ganglia, a pair of oesophageal ganglia, and a suboesophageal ganglion (Fig. 2A, B). The brain is located immediately below the surface epithelium of the head and consists of a neuropil core with numerous synapses surrounded by a layer of neuronal cell somata [14, 19, 20]. Connective nerve tracts are found between the cerebral and the vestibular, the cerebral and the ventral, and the vestibular and the oesophageal ganglia. A commissural nerve bundle behind the oesophagus connects the vestibular ganglia. The anterio-posteriorly running nerves which connect the cerebral and the ventral ganglia are called the main connectives, and those which connect the cerebral and the vestibular ganglia, the frontal connectives . Sensory organs associated with the brain are a pair of eyes [24, 33–35], a ciliated loop, the corona ciliata, localised in the dorsal part of the head [2, 20, 29–32] and the retrocerebral organ, a structure with an unknown putative sensory function .
The ventral ganglion is an elongate structure lying between the basement membrane and the epidermis. Two main connectives link it with the brain ganglia and two other nerve tracts continue caudally (Fig. 2A, B) [19, 23]. Similar to the brain, the ventral ganglion consists of a central fibrillar neuropil core, flanked by lateral clusters of cell bodies [2, 19, 20]. Along the length of the ventral ganglion a series of smaller nerves pass out radially (Fig. 2A, B) which branch in the periphery and form a dense ramifying plexus just external to the basement membrane to provide motor innervation to the body musculature and to innervate the ciliary fence receptors . The ventral ganglion controls swimming by initiating contractions of the body wall musculature and co-ordinating mechanosensory input from the numerous ciliary fence receptors in the epidermis [2, 20, 23]. The available descriptions of the lateral nerves that exit the ventral ganglion in closely related species of the genus Sagitta display a remarkable degree of variability over the last 125 years. Hertwig  described an irregular array of nerve fibres ("Nervenfibrillen") to emerge from the ventral ganglion (Fig. 2C). Kuhl , presenting a modified drawing from Burfield , depicts 12 distinct, stout nerve trunks to exit the ganglion on both sides (Fig. 2D). Yet, Goto and Yoshida  draw only six radial nerves on each side (Fig. 2B). In Shin , again, 12 bilaterally arranged radial nerves exit the ganglion on both sides in a nicely ordered way, much like the arrangement of the segmental nerves in the ventral nerve cord of an annelid or arthropod (Fig. 2E). In the same year, Duvert et al.  published a report in which they described 20 – 30 irregularly arranged aspartate-immunoreactive fibre bundles that pass out radially from the ventral ganglion on both sides and spread out diffusely and branch in the periphery, similar to the description already provided by Hertwig .
The structure of the ventral ganglion as revealed by the histochemical localization of actin and a nuclear marker
Tubulin labelling: the ventral ganglion as a condensation of the intraepidermal nerve plexus
An irregular array of ca. 20 – 30 tubulin-labelled fibres and fibre bundles emerges laterally from both sides of the ventral ganglion (Fig. 4C, D). These bundles contribute to and are confluent with the peripheral nerve plexus. We did not notice a regularly ordered spacing or distinct bilaterally symmetrical arrangement of these radial nerve bundles as earlier reports had indicated [2, 19]. Rather, our findings resemble the pattern of aspartate-immunoreactive fibre bundles that pass out from the ventral ganglion as described by Duvert et al. . Caudally, two thick bundles of tubulin-labelled fibres emerge from the ganglion to connect it to the intraepidermal nerve plexus in the posterior part of the animal (Fig. 4C). These caudal bundles are superficially similar to the radial fibre bundles, yet they contain more closely packed neurites. In summary, in tubulin-labelled preparations the ventral ganglion very much appears to be a condensed continuation of the peripheral intraepidermal nerve plexus.
Synapsin immunoreactivity: the ventral ganglion as a highly ordered structure
RFamide-like immunoreactivity: individually identifiable neurons
The RFamidergic neurons, all of which are unipolar, can be subdivided into two different populations: a first series of neurons with lateral somata (L1–4; small circles in Fig. 7) in the anterior third of the ganglion and a second series of slightly larger dorsal neurons (D1–5; large circles in Fig. 7) in the posterior two thirds of the ganglion the somata of which are located clearly more dorsally than those of the lateral neurons as is apparent in lateral views of the nervous system (Fig. 8F, G). Within the cell somata, the immunolabelled material is typically concentrated in a cap of cytoplasm opposite to the point of exit of the single neurite (Fig. 6G, 8D). The arrangement of the most anterior lateral neurons displayed some variation between individual specimens and in addition to the neurons L1 and L2 that we identified consistently in all six specimens that we analysed at the single-cell level, additional neurons are present in some specimens (small circles labelled with a question mark in Fig. 7). Contrary to L1 and L2, the neurons L3, L4, and D1–D6 were reliably present in all analysed specimens and hence represent typical examples for individually identifiable, bilateral symmetrically arranged neurons. The neurons D1–D5 have an identical morphology and appear to be serially repeated clones. Their neurites exit the soma in a medial direction to enter the neuropil core at a right angle to the anterior-posterior axis (Fig. 6G, 8B, C). The neurites of cells D1–D5 all cross over the lateral longitudinal bundle to join the intermediate bundle. The intermediate longitudinal bundle is composed of an inner and an outer portion (Fig. 8C). The neurites of cells D1–D5 (and also of D6; Fig. 8E) consistently contact the inner fibres of the intermediate bundle which is another indication of their serial identity (arrows in Fig. 8C). In some specimens, the presence of faintly immunolabelled material suggested the neurites of some D neurons to proceed even more medially to join the medial bundle. Screening these regions with high-magnification laser-scan microscopy did not provide conclusive evidence that this is the case (inset in Fig. 8D, arrow).
The ventral ganglion is a condensation of the intraepidermal nerve plexus
A conspicuous feature of the chaetognaths is that in addition to the central part of the nervous system, a peripheral part is present which is exclusively intraepidermal [2, 19, 20, 39] and which mediates motor innervation to the body musculature and innervates the ciliary fence receptors . By using anti-aspartate immunohistochemistry, Duvert et al.  visualized this extensive intraepidermal nerve plexuses both in the head region and in the trunk. In the trunk, muscle contraction seems to be controlled by this profuse intraepidermal network [23, 28, 67]. Compared to other Bilateria, an unusual feature of the arrow worm neuromuscular system is that in the trunk axonal varicosities lack specialized junctions and are separated from underlying muscles by a thick connective stratum. Acetylcholine is the major neuromuscular transmitter, which reaches muscle cells in the trunk by diffusion through the intervening body wall extracellular matrix [2, 20]. Acetylcholinergic fibres deliver their transmitter by boutons that in the trunk region terminate on the epidermis side of the connective stratum and therefore diffusely bath the muscle cells with acetylcholine [27, 39]. It should be noted, however, that in the head, nervous fibres which target oesophageal and somatic head muscles, have conventional nerve endings and neuromuscular junctions that display ultrastructural features similar to classic motor end plates [2, 28].
Our experiments on tubulin immunolocalization confirm and extend the observations on the structure of the peripheral plexus and ventral ganglion already published by Hertwig . We showed an irregular array of ca. 20 – 30 fibres and fibre bundles to emerge laterally from both sides of the ventral ganglion. These fibres pass out radially to spread out diffusely and branch in the periphery thus being a major source for the peripheral nerve plexus. Duvert et al.  described a similar irregular array of aspartate-immunoreactive fibre bundles to spread out from the ventral ganglion. These findings together contradict earlier reports that in Sagitta, six  or twelve [2, 18], distinct, bilaterally arranged radial nerves exit the ganglion on both sides in a serially ordered way. Our findings therefore defy any similarities of the chaetognath ventral ganglion with the internalized ventral nerve cords of e.g. annelids or arthropods that have segmentally arranged nerves projecting into the periphery. Rather, the arrow worm ventral ganglion appears to be a condensation of the intraepidermal nerve plexus. Hertwig  and Bone and Goto  observed numerous multipolar neurons to be embedded within the intraepidermal plexus. These neurons in the plexus may be the evolutionary precursors of the lateral clusters of cell bodies that flank the central fibrillar neuropil core of the ventral ganglion. During the emergence of Protostomia, the neurons in the plexus may have aggregated to form the centralized nerve centres that we observe in the Protostomia (Fig. 1).
Considering its role to coordinate sensory input from the fence receptors and efferent control of the trunk muscles for swimming behavior, Bone and Goto  proposed the ventral ganglion to be part of the central nervous system. From a histological point of view, the ventral ganglion can be seen as a condensation of the epidermal nerve plexus that displays a high degree of centralization. It represents a centralized, yet peripherally located center for complex sensory-motor integration. It is important to note that the chaetognath ventral ganglion is not internalized such as the subepidermal ganglia of Annelida, Arthropoda and Mollusca but remains in an intraepidermal position. It appears that, compared to other Protostomia, the Chaetognatha, by transforming a diffuse nerve net to a more centralized neuronal structure, followed their own distinct evolutionary pathway to generate a ventral nervous center in the trunk for sensory integration and motor control. Therefore, it may be appropriate to term this structure "ventral nerve centre" in order to stress the difference to ventral ganglia of other Protostomia. We suggest that this centralized nerve centre with its specific architecture and intraepidermal location is an autapomorphy of Chaetognatha.
The serial organization of the ventral nerve centre
Despite the suggested origin of the ventral nerve centre from the peripheral nerve net its central neuropil core displays a high degree of internal organization. Synapsin immunolocalization revealed a highly ordered system of serially arranged synaptic microcompartments in the ganglion core. This compartmentalization may anatomically be linked to a system of serially arranged transverse fibres that cross the neuropil core as reported by Bone and Pulsford . It does not have any equivalent in the ventral ganglia of other Protostomia [73–78] and can be considered another autapomorphy of the chaetognath ventral nerve centre. Our data on the immunolocalization of the neuropeptide RFamide provide further evidence for serially arranged nervous elements in the ventral nerve centre of Sagitta setosa and extend previous reports on the localization of this substance in Sagitta setosa  and Paraspadella gotoi . Most importantly, we provide evidence for serially arranged, individually identifiable neurons that can be homologized between different specimens. Although Goto et al.  (their Fig. 1d) did not map the pattern of RFamide-immunoreactive neurons in the ventral nerve centre of Paraspadella gotoi in detail, a comparison with our data nevertheless suggests that some of the L and D neurons may be evolutionary conserved across different chaetognath species. Papillon et al.  described the median Hox gene SceMed4 in embryos and early hatchlings of Spadella cephaloptera. This gene is expressed in two lateral stripes in the middle of the developing ventral nerve centre. The SceMed4 mRNA is localized in the bilateral soma clusters but not in the neuropil. These authors suggested that this gene may contribute to the diversity of neuronal subpopulations and to the establishment of distinct axon projection patterns . It would be interesting to explore the expression of this gene in the organism that we studied, Sagitta setosa, to see if the region of expression coincided e.g. with the anterior-posterior transition from the "L" to the "D" type of RFamidergic neurons that we describe in the current study.
Individually identifiable neurons (see  for a discussion of this concept) seem to be present in the nervous systems of all major taxa within the Protostomia: e.g. Arthropoda [60, 61], Annelida [74, 77, 78, 81–83], Nemathelminthes/Cycloneuralia [84, 85], basal Mollusca [73, 74, 86], Plathelminthes [87–89], and Gnathostomulida . Therefore, we suggest that developmental programmes generating neurons with an individual identity must be present in the ground pattern of the Protostomia. A serial arrangement of individual nerve cells cannot only be found in protostomians with typical segmentation such as annelids and arthropods [60, 77, 78], but also in unsegmented organisms such as Nematoda , Plathelminthes [87–89], Chaetognatha (present report), and in organisms the segmental organization of which is unclear such as basal Mollusca [73, 86, 91]. Our concepts of segmentation in Protostomia mainly rely on work done in annelids and arthropods (reviews e.g. [92, 93]). Budd  points out that "because segmentation is an evolutionary feature, it must have been acquired in a series of functional intermediates, and the sudden imposition of eusegmentation on a non-segmental precursor seems highly unlikely." In this view, the organization of individual organ systems such as the central nervous system (but not the entire organism) into serially repeated structures may be the starting point in an evolutionary trajectory from which segmentation as we see it in annelids and arthropods emerged .
Evolution of the bilaterian nervous system and the position of Chaetognatha
An intraepidermal nerve plexus is a prominent feature of many basal deuterostomes [55, 68]. It is present e.g. in enteropneusts , in urochordates such as tunicates , and in the basal chordate Amphioxus . An extensive intraepidermal nervous system also characterizes many Protostomia [3, 55]. Recent investigations on Annelida [77, 78, 81] and Onychophora (basal arthropods; Mayer and Harzsch, unpublished data) provide new evidence that, in addition to the internalized parts of the nervous system, the peripheral plexus is a prominent feature in these organisms that seem to have retained more motifs of a flatworm-like orthogonal nervous system than has been perceived before. It has recently been proposed that an epithelial (epidermal) nerve plexus (as is also present in Cnidaria; [97–99]) without concentrations into longitudinal cords characterizes the ground pattern of Bilateria [56, 64, 65] (Fig. 1). Yet, the presence of at least some individually identifiable neurons in basal deuterostomes such as tunicates (Urochordata; [100–102]), and the lancelet Amphioxus (Chordata; ) indicates that the potential to establish individual identities of the neurons in the plexus may not only be present in the ground pattern of Protostomia (see above) but may date back to the ground pattern of Bilateria (Fig. 1).
The evolution of the deuterostome nervous system is a field of intense research [56, 63–66, 95, 96, 103–107] but it is beyond the scope of this contribution to embark into this issue. For the Protostomia, Nielsen [3, 58] suggested that a perioral/circumesophageal brain in addition to the intraepidermal nerve plexus characterizes their ground pattern (Fig. 1). Such a circumoral brain can be recognized in most protostomian groups (e.g.  Fig. 12.2 therein; and [61, 71]) and is by Nielsen [3, 58] considered to be one of the key apomorphies of the Protostomia that may have evolved from a circumoral concentration of nerve cells around the mouth as present in Cnidaria. The brain components in Chaetognatha are also arranged in such a typical circumoral pattern (compare Fig. 2).
In summary, we suggest that in the ground pattern of Protostomia, the nervous system is characterized by the following features (see Fig. 1):
• An extensive intraepidermal plexus (plesiomorphic)
• The developmental programme to establish individual identities of neurons (plesiomorphic?)
• A circumoral brain ring (apomorphic)
• Most likely several longitudinal fibre tracts that are embedded within the peripheral plexus (apomorphic?)
• Ventral centralizations of the plexus that are linked to the brain by longitudinal tracts (apomorphic; "ventral" being the "chordin" expressing side in Protostomia; Lowe et al. 2006)
We conclude that the nervous system architecture of Chaetognatha including ultrastructural features  places them within the Protostomia (see also ). Phylogenetic affinities of the Chaetognatha to Deuterostomia in our view can be ruled out as well as molecular hypotheses that suggest a sister-group relationship to all other Bilateria [43, 49] and, to date, we consider a sister-group relationship of Chaetognatha and Protostomia [46, 50] to be unlikely. Based on the differences of the arrow worm brain to the collar-like shape of the typical cycloneuralian brain in Nematoda, we also believe that it is unlikely that Nematoda are the sister-group of Chaetognatha [42, 47]. More detailed analyses of the arrow worm ventral nerve centre and especially the brain including studies on neurogenesis will be necessary to explore any potential affinities of this group to specific taxa within the Protostomia.
Juvenile specimens of an unidentified species of the genus Sagitta were obtained from the coastal waters around the Mediterranean island Ibiza (Spain) in March 2006. A plankton net was towed across the surface waters of the Cala Llenya, Cala Vadella, Penyal de s'Aguila, and the Punta Grassio. Adult specimens of Sagitta setosa were obtained during a collection trip to the Biologische Anstalt Helgoland, German Bight http://www.awi-bremerhaven.de/BAH/ in August 2006. Specimens were obtained by horizontal (surface water samples) as well as by vertical (down to 20 meters depth) plankton hauls with the research vessel "MS Aade".
Histochemistry and immunohistochemistry
Specimens were fixed overnight at 4°C (or for 4 h, room temperature) in 4% paraformaldehyde (PFA) in phosphate buffer (PB; 0,1 M, pH 7.4). Histochemistry and immunohistochemistry were carried on free-floating whole mounts of adult specimens with fluorochrome-conjugated secondary antibodies using standard protocols. After fixation the tissues were washed in several changes of phosphate buffered saline (PBS) for at least 4 h, preincubated in PBS-TX (1% normal goat serum, 0,3% Triton X-100, 0,05% Na-acide) for 1 h and then incubated overnight in the following histochemical reagents and primary antibodies diluted in PBS-TX (room temperature):
• Phalloidin Alexa 488 (1:50; probe for actin; see e.g. ; Molecular Probes, obtained from MoBiTec, Göttingen, Germany)
• anti-acetylated alpha-tubulin from mouse (Sigma 1:100; see e.g. ).
• anti-FMRFamide from rabbit (1:1000; Diasorin; see e.g. ).
• anti-synapsin SYNORF 1 from mouse (1:10; ; antibody kindly provided by Prof. Dr. E. Buchner, Universität Würzburg)
For double labelling, combinations of these antisera were used, or the specimens were stained with the nuclear dye bisbenzimide (0.1%, 15 min. at room temperature; Hoechst H 33258), prior to the antibody incubations. Specimens were then washed for at least 2 h in several changes of PBS and subsequently incubated in secondary antibodies against mouse and rabbit proteins conjugated to the fluorochrome Alexa Fluor 488 (Molecular Probes, obtained by MoBiTec, Göttingen, Germany) for 4 hours. Finally the tissues were washed for at least 2 h in several changes of PBS and mounted in GelMount (Sigma). In control experiments, the omission of primary antibodies resulted in the absence of any neuronal labelling.
Digital images were obtained with a Zeiss Axioskop fitted with a CCD-1300B digital camera (Vosskühler GmbH) and processed with the Lucia Measurement 5.0 software package (Laboratory Imaging Ltd.). Alternatively, samples were scanned with a Leica TCS SP2 AOBS confocal laser-scanning microscope (Lichtmikroskopiezentrum, Institut für Zellbiologie und Biosystemtechnik, Universität Rostock). Those images are based on stacks of between 15 and 20 optical sections (single images are averages of four laser sweeps) of a z-series taken at intervals of 1 μm.
We would like to thank the team of the „Gastforschung“ at the Biologische Anstalt Helgoland for assistance in collecting Sagitta setosa. We gratefully acknowledge Prof. Dr. Dieter Weiß (Lichtmikroskopiezentrum, Institut für Zellbiologie und Biosystemtechnik, Universität Rostock) for providing access to a laser scanning microscope and Mr. Eik Hoffmann (Universität Rostock) for technical assistance. Our special thanks go to Mr. Jens Bünning (Universität Rostock), Mr. Rene Stüber (Dresden) and Mr. Christophe Ubbelohde (diving center H2O, Ibiza) for obtaining and sorting plankton samples. Prof. Dr. E. Buchner (Biozentrum, Universität Würzburg) generously provided samples of the SYNORF 1 antiserum. Our manuscript profited from stimulating discussion with Dr. Yvan Perez (Marseille). This study was supported by grant HA 2540/7-1 in the DFG focus programme SPP 1174 „Metazoan Deep Phylogeny“.
- Kapp H: Chaetognatha, Pfeilwürmer. Spezielle Zoologie. Teil 1. Wirbellose Tiere. Edited by: Westheide W, Rieger R. 1996, Stuttgart: Gustav Fischer Verlag, 757-762.Google Scholar
- Shinn GL: Chaetognatha. Microscopic Anatomy of Invertebrates, Hemichordata, Chaetognatha, and the Invertebrate Chordates. Edited by: Harrison FW, Ruppert EE. 1997, New York: Wiley-Liss Inc, 15: 103-220.Google Scholar
- Nielsen C: Animal evolution. 2001, Oxford: Oxford University PressGoogle Scholar
- Salvini-Plawen Lv: Systematic notes on Spadella and on the Chaetognatha in general. Z Zool Syst Evol. 1986, 24: 122-128.View ArticleGoogle Scholar
- Kapp H: Zum Ursprung der Chaetognathen – der aktuelle Stand von DNA-Analysen und morphologisch-anatomischer Forschung. Verh Dtsch Zool Ges. 1996, 89: 13-Google Scholar
- Ghiradelli E: Chaetognaths: two unsolved problems: the coelom and their affinities. Body Cavities: Function and Phylogeny. Selected Symposia and Monographs. UZI 8. Edited by: Lanzavecchia G, Valvassori R. 1995, Candia Carnevali MD, 167-185.Google Scholar
- Westheide W, Rieger R: Deuterostomia. Spezielle Zoologie. Teil 1. Wirbellose Tiere. Edited by: Westheide W, Rieger R. 1996, Stuttgart: Gustav Fischer Verlag, 755-756.Google Scholar
- Kapp H: Chatognatha, Pfeilwürmer. Spezielle Zoologie. Teil 1. Wirbellose Tiere. Edited by: Westheide W, Rieger R. 2007, München: Elsevier (Spektrum Akademischer Verlag), 898-903. 2Google Scholar
- Nielsen C: Proposing a solution to the Articulata-Ecdysozoa controversy. Zool Scr. 2003, 32: 475-482. 10.1046/j.1463-6409.2003.00122.x.View ArticleGoogle Scholar
- Hyman LH: Phylum Chaetognatha. The Invertebrates. 1959, New York: McGraw Hill Book Company, 5: 1-71.Google Scholar
- Welsch U, Storch V: Fine structure of the coelomic epithelium of Sagitta elegans. Zoomorphology. 1982, 100: 217-222. 10.1007/BF00311974.View ArticleGoogle Scholar
- Kapp H: The unique embryology of Chaetognatha. Zool Anz. 2000, 239: 263-266.Google Scholar
- Shimotori T, Goto T: Developmental fates of the first four blastomeres of the chaetognath Paraspadella gotoi: Relationship to protostomes. Develop Growth Differ. 2001, 43: 371-382. 10.1046/j.1440-169x.2001.00583.x.View ArticleGoogle Scholar
- Rehkämper G, Welsch U: On the fine structure of the cerebral ganglion of Sagitta (Chaetognatha). Zoomorphology. 1985, 105: 83-89. 10.1007/BF00312142.View ArticleGoogle Scholar
- Bartnik E, Weber K: Widespread occurrence of intermediate filaments in invertebrates: common principles and aspects of diversion. Eur J Cell Biol. 1989, 50: 17-33.Google Scholar
- Erber A, Riemer D, Bovenschulte M, Weber K: Molecular phylogeny of metazoan intermediate filament proteins. J Mol Evol. 1998, 47: 751-762. 10.1007/PL00006434.PubMedView ArticleGoogle Scholar
- Hertwig O: Die Chaetognathen. Mon Jena Z Med Naturw. 1880, 14: 196-311.Google Scholar
- Kuhl W: Chaetognatha. Klassen und Ordnungen des Tierreiches. Bd IV, Abt. IV, Buch 2, Teil 1. Edited by: Bronn HG. 1938, Leipzig: Akademische Verlagsgesellschaft M. B. H. Leipzig, 1-226.Google Scholar
- Goto T, Yoshida M: Nervous system in Chaetognata. Nervous Systems in Invertebrates. Edited by: Ali MA. 1987, Plenum Publishing Corporation, 461-481.View ArticleGoogle Scholar
- Bone Q, Goto T: The nervous system. The biology of chaetognaths. Edited by: Bone Q, Kapp H, Pierrot-Bults AC. 1991, Oxford: Oxford University Press, 18-31.Google Scholar
- Kapp H: Morphology and Anatomy. The Biology of Chaetognaths. Edited by: Bone Q, Kapp H, Pierrot-Bults AC. 1991, Oxford: Oxford University Press, 5-17.Google Scholar
- Ax P: Das System der Metazoa III. 2001, Göttingen: Spektrum Akademischer VerlagGoogle Scholar
- Bone Q, Pulsford A: The sense organs and ventral ganglion of Sagitta (Chaetognatha). Acta Zoologica (Stockh.). 1984, 65: 209-220.View ArticleGoogle Scholar
- Goto T, Yoshida M: Photoreception in Chaetognata. Photoreception and vision in invertebrates. Edited by: Ali MA. 1984, Plenum Publishing Corporation, 721-742.Google Scholar
- Scharrer E: The fine structure of the retrocerebral organ of Sagitta (Chaetognatha). Life Sci. 1965, 4: 923-926. 10.1016/0024-3205(65)90191-8.PubMedView ArticleGoogle Scholar
- Salvini-Plawen Lv: The epineural (vs. gastroneural) cerebral complex of the Chaetognatha. Z Zool Syst Evol. 1988, 26: 425-429.View ArticleGoogle Scholar
- Bone Q, Grimmelikhuijzen CLP, Pulsford A, Ryan KP: Possible transmitter functions of acetylcholine an an RFamide-like substance in Sagitta (Chaetognatha). Proc R Soc London B. 1987, 230: 1-14.View ArticleGoogle Scholar
- Duvert M, Barets AL: Ultrastructural studies of neuromuscular junctions in visceral and skeletal muscles of the chaetognath Sagitta setosa. Cell Tiss Res. 1983, 233: 657-669. 10.1007/BF00212233.View ArticleGoogle Scholar
- Horridge GA, Boulton PS: Prey detection by Chaetognatha via a vibration sense. Proc R Soc Lond B. 1967, 168: 413-419.View ArticleGoogle Scholar
- Feigenbaum DL: Hair-fan patterns in the Chaetognatha. Can J Zool. 1978, 56: 536-546.View ArticleGoogle Scholar
- Bone Q, Pulsford A: The arrangement of ciliated sensory cells in Spadella (Chaetognatha). J Mar Biol Ass UK. 1978, 58: 565-570.View ArticleGoogle Scholar
- Malakhov VV, Berezinskaya TL, Solovyes KA: Fine structure of sensory organs in chaetognaths: Ciliary fence receptors, ciliary tuft receptors and ciliary loop (in Russian). Invert Zool. 2005, 2: 67-77.Google Scholar
- Goto T, Yoshida M: Histochemical demonstration of a rhodopsin-like substance in the eye of the arrow-worm, Spadella schizoptera (Chaetognatha). Exp Biol. 1988, 48: 1-4.PubMedGoogle Scholar
- Goto T, Takasu N, Yoshida M: A unique photoreceptive structure in the arrowworms Sagitta crassa and Spadella schizoptera (Chaetognatha). Cell Tiss Res. 1984, 235: 471-478. 10.1007/BF00226941.View ArticleGoogle Scholar
- Goto T, Terazaki M, Yoshida M: Comparative morphology of the eyes of Sagitta (Chaetognatha) in relation to depth of habitat. Exp Biol. 1989, 48: 95-105.PubMedGoogle Scholar
- Goto T, Yoshida : Oriented light reactions of the arrow worm Sagitta crassa Tokioka. Biol Bull. 1981, 160: 419-430. 10.2307/1540849.View ArticleGoogle Scholar
- Goto T, Yoshida M: The role of the eye and CNS components in phototaxis of the arrow worm, Sagitta crassa Tokioka. Biol Bull. 1983, 164: 82-92. 10.2307/1541192.View ArticleGoogle Scholar
- Goto T, Katayama-Kumoi Y, Tohyama M, Yoshida M: Distribution and development of the serotonin-and RFamide-like immunoreactive neurons in the arrowworm, Paraspadella gotoi (Chaetognatha). Cell Tiss Res. 1992, 267: 215-222. 10.1007/BF00302958.View ArticleGoogle Scholar
- Duvert M, Savineau JP, Campistron G, Onteniente B: Distribution and role of aspartate in the nervous system of the chaetognath Sagitta. J Comp Neurol. 1997, 380: 485-494. 10.1002/(SICI)1096-9861(19970421)380:4<485::AID-CNE5>3.0.CO;2-Y.PubMedView ArticleGoogle Scholar
- Wada H, Satoh N: Details of the evolutionary history from invertebrates to vertebrates as deducted from the sequences of 18S rDNA. Proc Natl Acad Sci USA. 1994, 91: 1801-1804. 10.1073/pnas.91.5.1801.PubMed CentralPubMedView ArticleGoogle Scholar
- Telford MJ, Holland : The phylogenetic affinities of the chaetognaths: a molecular analysis. Mol Biol Evol. 1993, 10: 660-676.PubMedGoogle Scholar
- Halanych KM: Testing hypotheses of chaetognath origins: long branches revealed by 18S ribosomal DNA. Syst Biol. 1996, 45: 223-246. 10.2307/2413616.View ArticleGoogle Scholar
- Telford MJ, Holland PWH: Evolution of 28S ribosomal DNA in chaetognaths: duplicate genes and molecular phylogeny. J Mol Evol. 1997, 44: 135-144. 10.1007/PL00006130.PubMedView ArticleGoogle Scholar
- Littlewood TJ, Telford MJ, Clough KA, Rohde K: Gnathostomulida – an enigmatic metazoan phylum from both morphological and molecular perspectives. Mol Phylogenet Evol. 1998, 9: 72-79. 10.1006/mpev.1997.0448.PubMedView ArticleGoogle Scholar
- Zrzavý J, Mihulka S, Kepka P, Bezděk A: Phylogeny of the Metazoa based on morphological and 18S ribosomal DNA evidence. Cladistics. 1998, 14: 249-285.View ArticleGoogle Scholar
- Giribet G, Distel DL, Polz M, Sterrer W, Wheeler WC: Triploblastic relationships with emphasis on the Acoelomates and position of Ganthostomulida, Cycliophora, Plathelminthes and Chaetognatha: a combined approach to 18S rDNA sequences and morphology. Syst Biol. 2000, 49: 539-562. 10.1080/10635159950127385.PubMedView ArticleGoogle Scholar
- Peterson KJ, Eernisse DJ: Animal phylogeny and the ancestry of bilaterians: inferences from morphology and 18S rDNA gene sequences. Evol Dev. 2001, 3: 170-205. 10.1046/j.1525-142x.2001.003003170.x.PubMedView ArticleGoogle Scholar
- Mallat J, Winchell CJ: Testing the new animal phylogeny: first use of combined large-subunit and small sub-unit rRNA gene sequences to classify the protostomes. Mol Biol Evol. 2002, 19: 289-301.View ArticleGoogle Scholar
- Papillon D, Perez Y, Fasano L, Le Parco Y, Caubit X: Hox gene survey in the chaetognath Spadella cephaloptera: evolutionary implications. Dev Gen Evol. 2003, 213: 142-148.Google Scholar
- Helfenbein KG, Fourcade HM, Vanjani RG, Boore JL: The mitochondrial genome of Paraspadella gotoi is highly reduced and reveals that chaetognaths are a sister group to prostostomes. PNAS. 2004, 101: 10639-10643. 10.1073/pnas.0400941101.PubMed CentralPubMedView ArticleGoogle Scholar
- Papillon D, Perez Y, Caubit X, Le Parco Y: Identification of chaetognaths as protostomes is supported by the analysis of their mitochondrial genome. Mol Biol Evol. 2004, 21: 2122-2129. 10.1093/molbev/msh229.PubMedView ArticleGoogle Scholar
- Matus DQ, Copley RR, Dunn CW, Hejnol A, Eccleston H, Halanych KM, Martindale MQ, Telford MJ: Broad taxon and gene sampling indicate that chaetognaths are protostomes. Curr Biol. 2006, 16: R575-R576. 10.1016/j.cub.2006.07.017.PubMedView ArticleGoogle Scholar
- Marlétaz F, Martin E, Perez Y, Papillon D, Caubit X, Lowe CL, Freeman B, Fasano L, Dossat C, Wincker P, Weissenbach J, Le Parco Y: Chaetognath phylogenomics: a protostome with deuterostome-like development. Curr Biol. 2006, 16: R577-R578. 10.1016/j.cub.2006.07.016.PubMedView ArticleGoogle Scholar
- Paul DH: Neurophylogenist's view of decapod Crustacea. Bull Mar Sci. 1989, 45: 487-504.Google Scholar
- Holland ND: Early central nervous system evolution: an era of skin brains?. Nat Rev Neurosci. 2003, 4: 617-627. 10.1038/nrn1175.PubMedView ArticleGoogle Scholar
- Lowe CJ: Origins of the chordate central nervous system: insights from hemichordates. Evolution of nervous systems – A comprehensive reference. Non-mammalian vertebrates. Edited by: Kaas JH, Bullock TH. 2007, Oxford: Academic Press, 2: 25-38.Google Scholar
- Hay-Schmidt : The evolution of the serotonergic system. Proc R Soc Lond B. 2000, 267: 1071-1079. 10.1098/rspb.2000.1111.View ArticleGoogle Scholar
- Nielsen C: Larval and adult brains. Evol Dev. 2005, 7: 483-489. 10.1111/j.1525-142X.2005.05051.x.PubMedView ArticleGoogle Scholar
- Stach T: Comparison of the serotonergic nervous system among Tunicata: implications for its evolution within Chordata. Org Div Evol. 2005, 5: 15-24. 10.1016/j.ode.2004.05.004.View ArticleGoogle Scholar
- Harzsch S, Müller CHG, Wolf H: From variable to constant cell numbers: cellular characteristics of the arthropod nervous system argue against a sister-group relationship of Chelicerata and "Myriapoda" but favour the Mandibulata concept. Dev Gen Evol. 2005, 215: 53-68. 10.1007/s00427-004-0451-z.View ArticleGoogle Scholar
- Harzsch S: Neurophylogeny: architecture of the nervous system and a fresh view on arthropod phylogeny. Integr Comp Biol. 2006, 46: 162-194. 10.1093/icb/icj011.PubMedView ArticleGoogle Scholar
- Harzsch S: Architecture of the nervous system as a character for phylogenetic reconstructions: examples from the Arthropoda. Species, Phylogeny & Evolution.Google Scholar
- Meinertzhagen IA, Okamura Y: The larval ascidian nervous system: the chordate brain from its small beginnings. TINS. 2001, 24: 401-410.PubMedGoogle Scholar
- Lowe CJ, Teraski M, Wu M, Freeman RM, Runft L, Kwan K, Haigo S, Aronowicz J, Lander E, Gruber C, Kirschner M, Gerhart J: Dorsoventral patterning in hemichordates: insights into early chordate evolution. PLOS Biology 4. 2006, 291: 1603-1619.Google Scholar
- Lowe CJ, Wu M, Salic A, Evans L, Lander E, Stange-Thomann , Grubner CE, Gerhart J, Kirschner M: Anteroposterior patterning in hemichordates and the origins of the chordate nervous system. Cell. 2003, 113: 853-865. 10.1016/S0092-8674(03)00469-0.PubMedView ArticleGoogle Scholar
- Caòestro C, Bassham S, Postlethwait J: Development of the central nervous system in the larvacean Oikopleura dioica and the evolution of the chordate brain. Dev Biol. 2005, 285: 298-315. 10.1016/j.ydbio.2005.06.039.View ArticleGoogle Scholar
- Lichtneckert R, Reichert H: Insights into the urbilaterian brain: conserved genetic patterning mechanisms in insect and vertebrate brain development. Heredity. 2005, 94: 465-77. 10.1038/sj.hdy.6800664.PubMedView ArticleGoogle Scholar
- Lichtneckert R, Reichert H: Origin and evolution of the first nervous system. Evolution of nervous systems – A comprehensive reference. Theories, development, invertebrates. Edited by: Striedler GF, Rubenstein JLR. 2007, Oxford: Academic Press, Oxford, 1: 289-316.Google Scholar
- Reichert H: A tripartite organization of the urbilaterian brain: developmental genetic evidence from Drosophila. Brain Res Bull. 2005, 66: 491-494. 10.1016/j.brainresbull.2004.11.028.PubMedView ArticleGoogle Scholar
- Erwin DH, Davidson EH: The last common bilaterian ancestor. Development. 2002, 129: 3021-3032.PubMedGoogle Scholar
- Burfield ST: Sagitta . Proc Trans Liverpool Biol Soc. 1927, 41: 1-104.Google Scholar
- Duvert M, Salat C: Ultrastructural and cytochemical studies on the connective tissue of chaetognaths. Tiss Cell. 1990, 22: 865-878. 10.1016/0040-8166(90)90049-F.View ArticleGoogle Scholar
- Friedrich S, Wanninger A, Brückner M, Haszprunar G: Neurogenesis in the mossy chiton, Mopalia muscosa (Gould) (Polyplacophora): evidence against molluscan metamerism. J Morph. 2002, 253: 109-117. 10.1002/jmor.10010.PubMedView ArticleGoogle Scholar
- Stuart DK, Blair SS, Weisblat DA: Cell lineage, cell death, and the developmental origin of identified serotonin- and dopamine-containing neurons of the leech. J Neurosci. 1987, 7: 1107-1122.PubMedGoogle Scholar
- Wanninger A, Haszprunar G: The development of the serotonergic and FMRF-amidergic nervous system in Antalis entalis (Mollusca, Scaphopoda). Zoomorphology. 2003, 122: 77-85.Google Scholar
- Harzsch S: Ontogeny of the ventral nerve cord in malacostracan crustaceans: a common plan for neuronal development in Crustacea, Hexapoda, and other Arthropoda?. Arthr Struct Dev. 2003, 32: 17-38. 10.1016/S1467-8039(03)00008-2.View ArticleGoogle Scholar
- Orrhage L, Müller MCM: Morphology of the nervous system of Polychaeta (Annelida). Hydrobiologia. 2005, 535/526: 79-111. 10.1007/s10750-004-4375-4.Google Scholar
- Müller MCM: Polychaete nervous systems: ground pattern and variations – cLS microscopy and the importance of novel characteristics in phylogenetic analysis. Int Comp Biol. 2006, 46: 125-133. 10.1093/icb/icj017.View ArticleGoogle Scholar
- Papillon D, Perez Y, Fasano L, Le Parco Y, Caubit X: Restricted expression of a median Hox gene in the central nervous system of chaetognaths. Dev Gen Evol. 2005, 215: 369-373. 10.1007/s00427-005-0483-z.View ArticleGoogle Scholar
- Burrows M: The neurobiology of an insect brain. 1996, Oxford: Oxford University PressView ArticleGoogle Scholar
- Huang Y, Jellies J, Johansen KM, Johansen J: Development and pathway formation of peripheral neurons during leech embryogenesis. J Comp Neurol. 1998, 397: 394-402. 10.1002/(SICI)1096-9861(19980803)397:3<394::AID-CNE6>3.0.CO;2-Y.PubMedView ArticleGoogle Scholar
- Gilchrist LS, Klukas KA, Jellies J, Rapus J, Eckert M, Mesce KA: Distribution and developmental expression of octopamine-immunoreactive neurons in the central nervous system of the leech. J Comp Neurol. 1995, 353: 451-461. 10.1002/cne.903530312.PubMedView ArticleGoogle Scholar
- Brodfuehrer PD, Thorogood MSE: Identified neurons and leech swimming behavior. Prog Neurobiol. 2001, 63: 371-381. 10.1016/S0301-0082(00)00048-4.PubMedView ArticleGoogle Scholar
- White JG, Southgate E, Thomson JN, Brenner S: The structure of the nervous system of the nematode Caenorhabditis elegans. Phil Trans R Soc London B. 1986, 314: 1-340.View ArticleGoogle Scholar
- Walthall WW: Repeating patterns of motoneurons in nematodes: the origin of segmentation?. The nervous systems of invertebrates: an evolutionary and comparative approach. Edited by: Breidbach O, Kutsch B. 1995, Basel: Birkhäuser Verlag, 61-75.View ArticleGoogle Scholar
- Voronezhskaya EE, Tyurin SA, Nezlin LP: Neuronal development in larval chiton Ischnochiton hakodadensis (Mollusca: Polyplacophora). J Comp Neurol. 2002, 444: 25-38. 10.1002/cne.10130.PubMedView ArticleGoogle Scholar
- Halton DW, Gustafsson MKS: Functional morphology of the plathyhelminth nervous system. Parasitology. 1996, 113: 47-72.View ArticleGoogle Scholar
- Reuter M, Halton DW: Comparative neurobiology of Plathelminthes. Interrelationships of Plathyelminthes. Edited by: Littlewood DTJ, Bray RA. 2001, London: Taylor & Francis, London, 239-249.Google Scholar
- Reuter M, Mäntylä K, Gustafsson KS: Organization of the orthogon – main and minor nerve cords. Hydrobiologia. 1998, 383: 175-182. 10.1023/A:1003478030220.View ArticleGoogle Scholar
- Müller MCM, Sterrer W: Musculature and nervous system of Gnathostomula peregrina (Gnathostomulida) shown by phalloidin labeling, immunohistochemistry, and cLSM, and their phylogenetic significance. Zoomorphology. 2004, 123: 169-177.Google Scholar
- Jacobs DK, Wray CG, Wedeen CJ, Kostriken R, DeSalle R, Staton JL, Gates RD, Lindberg DR: Molluscan engrailed expression, serial organization, and shell evolution. Evol Dev. 2000, 2: 340-347. 10.1046/j.1525-142x.2000.00077.x.PubMedView ArticleGoogle Scholar
- Budd GE: Why are arthropods segmented. Evol Dev. 2001, 3: 332-342. 10.1046/j.1525-142X.2001.01041.x.PubMedView ArticleGoogle Scholar
- Scholtz G: The Articulata hypothesis – or what is a segment?. Org Div Evol. 2002, 2: 197-215. 10.1078/1439-6092-00046.View ArticleGoogle Scholar
- Benito J, Pardos F: Hemichordata. Microscopic anatomy of invertebrates, Hemichordata, Chaetognatha, and the invertebrate chordates. Edited by: Harrison FW, Ruppert EE. 1997, New York: Wiley-Liss Inc, 15: 15-101.Google Scholar
- Mackie GO, Burighel P: The nervous system in adult tunicates: current research directions. Can J Zool. 2005, 83: 151-183. 10.1139/z04-177.View ArticleGoogle Scholar
- Wicht H, Lacalli TC: The nervous system of Amphioxus: structure, development, and evolutionary significance. Can J Zool. 2005, 83: 122-150. 10.1139/z04-163.View ArticleGoogle Scholar
- Grimmelikhuijzen CJP: Antisera to the sequence Arg-Phe-amide visualize neuronal centralization in hydroid polyps. Cell Tiss Res. 1984, 241: 171-182. 10.1007/BF00214639.View ArticleGoogle Scholar
- Grimmelikhuijzen CJP, Spencer AN: FMRFamide immunoreactivity in the nervous system of the medusa Polyorchis penicillatus. J Comp Neurol. 1985, 230: 361-371. 10.1002/cne.902300305.View ArticleGoogle Scholar
- Anderson PAV, Moosler A, Grimmelikhuijzen CJP: The presence and distribution of Antho-RFamide-like material in scyphomedusae. Cell Tiss Res. 1992, 267: 67-74. 10.1007/BF00318692.View ArticleGoogle Scholar
- Meinertzhagen IA: Eutely, cell lineage, and fate within the ascidian larval nervous system: determinacy or to be determined?. Can J Zool. 2004, 83: 1-12.Google Scholar
- Meinertzhagen IA, Lemaire P, Okamura Y: The neurobiology of the ascidian tadpole larva: recent developments in an ancient chordate. Ann Rev Neurosci. 2004, 27: 453-485. 10.1146/annurev.neuro.27.070203.144255.PubMedView ArticleGoogle Scholar
- Imai JH, Meinertzhagen IA: Neurons of the ascidean larval nervous system in Ciona intestinalis: I. Central nervous system. J Comp Neurol. 2007, 501: 316-334. 10.1002/cne.21246.PubMedView ArticleGoogle Scholar
- Lacalli TC, Holland LZ: The developing dorsal ganglion of the salp Thalia democratica, and the nature of the ancestral chordate brain. Phil Trans R Soc Lond B. 1998, 353: 1943-1967. 10.1098/rstb.1998.0347.View ArticleGoogle Scholar
- Nielsen C: Origin of the chordate central nervous system – and the origin of chordates. Dev Gen Evol. 1999, 209: 196-205.View ArticleGoogle Scholar
- Lacalli TC: Apical organs, epithelial domains, and the origin of the chordate central nervous system. Am Zool. 1994, 34: 533-541.View ArticleGoogle Scholar
- Lacalli TC: Frontal eye circuitry, rostral sensory pathways and brain organization in amphioxus larvae: evidence from 3D reconstructions. Phil Trans R Soc Lond B. 1996, 351: 243-263. 10.1098/rstb.1996.0022.View ArticleGoogle Scholar
- Fritzsch B, Glover JC: Evolution of the deuterostome central nervous system: an intercalation of developmental patterning processes with cellular specification processes. Evolution of nervous systems – A comprehensive reference. Non-mammalian vertebrates. Edited by: Kaas JH, Bullock TH. 2007, Oxford: Academic Press, 2: 1-24.Google Scholar
- Vilpoux K, Sandeman R, Harzsch S: Early embryonic development of the central nervous system in the Australian crayfish and the Marbled crayfish (Marmorkrebs). Dev Gen Evol. 2006, 216: 209-223. 10.1007/s00427-005-0055-2.View ArticleGoogle Scholar
- Harzsch S, Anger K, Dawirs RR: Immunocytochemical detection of acetylated tubulin and Drosophila synapsin in the embryonic crustacean nervous system. Int J Dev Biol. 1997, 41: 477-484.PubMedGoogle Scholar
- Harzsch S, Dawirs RR: Development of neurons exhibiting FMRFamide-related immunoreactivity in the central nervous system of spider crab larvae (Hyas araneus L., Decapoda, Majidae). J Crust Biol. 1996, 16: 10-19. 10.2307/1548925.View ArticleGoogle Scholar
- Klagges BRE, Heimbeck G, Godenschwege TA, Hofbauer A, Pflugfelder GO, Reifegerste R, Reisch D, Schaupp M, Buchner S, Buchner E: Invertebrate synapsins: a single gene codes for several isoforms in Drosophila. J Neurosci. 1996, 16: 3154-3165.PubMedGoogle Scholar
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