Development of the nervous system in Phoronopsis harmeri (Lophotrochozoa, Phoronida) reveals both deuterostome- and trochozoan-like features
© Temereva and Wanninger; licensee BioMed Central Ltd. 2012
Received: 21 February 2012
Accepted: 6 June 2012
Published: 24 July 2012
Inferences concerning the evolution of invertebrate nervous systems are often hampered by the lack of a solid data base for little known but phylogenetically crucial taxa. In order to contribute to the discussion concerning the ancestral neural pattern of the Lophotrochozoa (a major clade that includes a number of phyla that exhibit a ciliated larva in their life cycle), we investigated neurogenesis in Phoronopsis harmeri, a member of the poorly studied Phoronida, by using antibody staining against serotonin and FMRFamide in combination with confocal microscopy and 3D reconstruction software.
The larva of Phoronopsis harmeri exhibits a highly complex nervous system, including an apical organ that consists of four different neural cell types, such as numerous serotonin-like immunoreactive flask-shaped cells. In addition, serotonin- and FMRFamide-like immunoreactive bi- or multipolar perikarya that give rise to a tentacular neurite bundle which innervates the postoral ciliated band are found. The preoral ciliated band is innervated by marginal serotonin-like as well as FMRFamide-like immunoreactive neurite bundles. The telotroch is innervated by two neurite bundles. The oral field is the most densely innervated area and contains ventral and ventro-lateral neurite bundles as well as several groups of perikarya. The digestive system is innervated by both serotonin- and FMRFamide-like immunoreactive neurites and perikarya. Importantly, older larvae of P. harmeri show a paired ventral neurite bundle with serial commissures and perikarya.
Serotonin-like flask-shaped cells such as the ones described herein for Phoronopsis harmeri are found in the majority of lophotrochozoan larvae and therefore most likely belong to the ground pattern of the last common lophotrochozoan ancestor. The finding of a transitory paired ventral neurite bundle with serially repeated commissures that disappears during metamorphosis suggests that such a structure was part of the “ur-phoronid” nervous system, but was lost in the adult stage, probably due to its acquired sessile benthic lifestyle.
KeywordsNeurogenesis Evolution Phylogeny Ventral nerve cord Last common lophotrochozoan ancestor
Although the exact phylogenetic position of Phoronida is still a matter of ongoing debate (and in some recent studies even their monophyly is questioned, e.g., [1, 2]), all recent molecular analyses agree in their inclusion within the protostome superclade Lophotrochozoa [3–5]. Mainly based on a proposed homologous feeding organ, the lophophore, Phoronida has been traditionally aligned with Ectoprocta and Brachiopoda to form the monophyletic Lophophorata [6, 7]. This view is supported by a shared radial type of cleavage  (although some unpublished observations claim that spiral cleavage may be present in some phoronids), but otherwise there is little morphological support for such an assemblage, mainly due to the high disparity of the adult bodyplans of these phyla. The same is true for the proposed lophophorate-spiralian alignment, where autapomorphies are hard to come by, one of them may be the occurrence of a larval apical organ with serotonin-like immunoreactive flask-shape cells . On the other hand, lophophorates share some embryological features with the seemingly distantly related deuterostomes, such as the above-mentioned radial cleavage [8, 10], the (sometimes questioned) three coelomic compartments , and an upstream ciliary filter feeding system [12, 13].
The nervous system is often considered highly conserved between major animal taxa and is therefore often used for morphology-based phylogenetic inferences . While an impressive amount of data on the development of the nervous system has recently become available for numerous lophotrochozoan subtaxa, phoronid neurogenesis is still comparatively little known, with most studies dealing with the description of the neuroanatomy of more or less mature larvae rather than with the development of the nervous system as such [15–21]. There are exceptions to the rule, however, but the few studies on phoronid neural development are restricted to one species belonging to the genus Phoronis and are not detailed enough to allow for answers concerning crucial issues such as the exact cellular composition and arrangement of the apical organ, thus rendering comparisons with other lophophorates, or even spiralians, difficult. Accordingly, a reconstruction of the neural anatomy of the larva of the last common lophophorate (and therefore lophotrochozoan) ancestor remains highly speculative. The unexpected recent finding of a paired ventral neurite bundle in the actinotroch larva of Phoronopsis harmeri similar to most spiralians may provide evidence for such a neural system in the phoronid as well as lophotrochozoan groundplan. Such a scenario would imply, however, secondary loss of this paired ventral nervous system in adult phoronids (probably as a result of a sessile lifestyle), since they do not exhibit a comparable neural structure. In order to broaden the database on phoronid neurogenesis and larval neuroanatomy and to contribute to the discussion concerning the groundpattern of the phoronid and lophotrochozoan neural bauplan, we investigated the development of the serotonin-like and FMFRamide-like immunoreactive nervous system during larval development of Phoronopsis harmeri.
Description of major developmental stages
Mid-gastrula (approximately 30–37 hps): this stage is 140 μm long and 100 μm wide. The archenteron is rounded (Figure 1D). The blastopore is flask-shaped with swollen anterior and narrow posterior portions (Figure 1C). At this stage, the anterior mesodermal precursor resembles a large mass of cells in front of and on both sides of the archenteron (Figure 1D).
Late gastrula I (approximately, 40–47 hps): this stage has an elongated archenteron with a cone-shaped posterior portion (Figure 1E). It represents the anlage of the future midgut. The blastopore closes and becomes tear-shaped with a narrow posterior portion (Figure 1F). The volume of the blastocoel decreases in the posterior part of the embryo. Cells of the anterior coelomic precursor often form processes that pass to the apical plate, and the preoral coelom forms in the anterior part of the embryo (Figure 1G).
Late gastrula II (approximately 48–52 hps): the body shape of this stage differs from that of the previous stage in that the mouth becomes deeper and the precursor of the preoral lobe forms (Figure 1H). The latter is an epidermal fold that is formed by two epidermal layers above the mouth. On the lateral and ventral sides of the posterior portion of the body, the postoral ciliated band, which is a belt of thick epithelium, appears. At this stage, the midgut contacts the surface of the epidermis of the posterior body part.
Preactinotrocha (approximately, 55–63 hps.): At this stage, the proctodaeum appears but the larva is still lecithotrophic (Figure 1I). The larva has a well-developed esophageal muscle, the preoral coelom, and the postoral ciliated band, which does not yet contain motionless latero-frontal cilia.
Young actinotrocha (approximately, 63–70 hps.): This stage has a well-developed preoral lobe (Figure 1J). The preoral lobe is the large anterior portion of the larva and has a large blastocoel, which is crossed by processes of mesodermal cells. The latero-frontal cilia on the tentacular ridge appear and the larva starts to feed. On both sides of the anus, small epidermal invaginations arise, which form the protonephridial tubes.
5- to 6-day-old actinotrocha: The larva has large blastocoels in the hyposphere, a short trunk under the tentacular ridge, and well-developed two protonephridia with several terminal cells (Figure 1K).
13-day-old actinotrocha: Specimens feed, increase in size, and retain nutrients in the cells of the stomach and hindgut (Figure 1L). There are three pairs of protrusions along the tentacular ridge, the primordial tentacles.
Actinotrocha with three pairs of tentacles (approximately 24 days after spawning): The larva has a well-developed trunk with a terminal telotroch around the anus (Figure 1M). The ventral pair of tentacles is the longest, and the dorso-lateral pair is the shortest. The trunk is occupied by a large trunk coelom. On the ventral body side under the tentacles, an area of thick epidermis appears which forms the primordium of the metasomal sac.
Although the apical tuft and the apical plate form in the early gastrula, perikarya do not differentiate and connect to the sensory cells of the apical plate until the mesodermal muscle cells appear, which is in the mid-gastrula stage.
Serotonin-like immunoreactive nervous system
In the late gastrula I (Figure 2E), the basal processes of serotonin-like immunoreactive perikarya form a neuropil in the center of the apical plate (Figure 2F). Nerve fibers occupy the basal portion of the apical plate and are in close contact with the mesodermal cells (Figure 2F). At this stage, the shape of perikarya does not change but the number increases with age and is seven or nine in the late gastrula II (Figure 2G). The arrangement of the perikarya forms a horseshoe-like pattern along the anterior edge of the apical plate. The apical portion of the cell becomes longer (Figure 2J). The neuropil becomes rounded and larger (Figure 2J). At this stage, and usually within the apical organ, there are two perikarya which are situated very closely adjacent to each other and form a cluster with two apical parts, two nuclei, and one process (Figures 2H, 3C).
The number of perikarya in the apical organ increases and the preactinotrocha has 12 perikarya (Figures 2K, L, 3D, E). The apical organ also increases in size and the perikarya are arranged in a wide horseshoe-like pattern with a large neuropil in the center (Figure 3D, E).
In the 5-day-old actinotroch (Figure 4K), additional serotonin-like immunoreactive neurites appear along the edge of the preoral lobe. They form two rows, which constitute the anterior and posterior marginal neurite bundles (Figure 4L). Each anterior marginal nerve passes along the marginal muscle of the preoral lobe and contains 4–5 bipolar serotonin-like immunoreactive perikarya. The posterior marginal neurite bundle is stained more intensely than the anterior marginal neurite bundle and connects with 12–15 large serotonin-like immunoreactive perikarya (Figure 4L). The anterior and posterior margin neurite bundles merge at the dorsal ends of the preoral lobe. Dorso-lateral neurite bundles emerging from the neuropil increase in diameter, pass along the tentacular ring, and form contact on the ventral side, forming the tentacular nerve ring (Figure 4L). The number of monopolar perikarya within the apical organ does not increase, but the shape of the perikarya does (Figure 4M). In the 5-day-old actinotroch, monopolar perikarya have a long basal process, which can reach 6 μm in length. The basal part of the perikaryon is wide and rounded (Figure 4N) and the nucleus is located here. The apical part of the cell resembles a collar and stains intensely. A serotonin-like immunoreactive cell with one basal process, two rounded nuclei (basal and apical), a small apical part, and a cilium (Figure 4O, P) can be found among other perikarya. Within the apical organ, there are bipolar or multipolar perikarya, which are located among the fibers of the neuropil (Figure 4Q); these perikarya have a large and irregularly shaped nucleus and they always occupy the anterior and lateral edge of the neuropil (Figure 4Q). Two bipolar cells can usually be found near the apical organ (Figure 4M). These cells are associated with the dorso-lateral branches of the tentacular neurite bundle and are located on both sides of the apical organ. A round nucleus is located in the center of the perikaryon.
The 24-day-old actinotroch has a weak marginal neurite bundle with several perikarya (Figure 5C) and a prominent median neurite bundle, which extends from the apical organ to the edge of the preoral lobe along its median line (Figure 8E, H, J, K). The median neurite bundle arises from the left group of multipolar perikarya (Figure 8F, J). In some larvae, a group of weakly stained perikarya is found in the center of the median neurite bundle (Figure 8H). This group consists of three small perikarya that contribute to the frontal organ. In some larvae, there is only one perikaryon instead of three (Figure 8B). Two dorso-lateral branches of the tentacular neurite bundle arise from the dorso-lateral parts of the neuropil where the multipolar perikarya are situated (Figure 8J, K). Near the apical organ, each dorso-lateral neurite bundle branches into several neurite bundles, which fuse on the dorsal side (Figure 8B, E). Here, the dorso-lateral neurite bundles are connected to each other via a dorsal commissure (Figure 8B, I). Along the whole length, the tentacular neurite bundle is formed by 7–10 neurites (Figure 8I). The tentacular neurite bundle runs under the tentacular ridge and forms a loop along the abfrontal side of each tentacle. The oral nerve ring is composed of thin circular neurites and several perikarya, which usually stain weakly (Figure 8B). These perikarya are located in the oral field epithelium, under the mouth. The oral nerve ring does not contact the tentacular neurite bundle or the apical organ, as stated previously . Thin neurites form a cylinder, which matches the shape of the esophagus (Figures 3K, 8B). In the 24-day-old larva, the trunk and the telotroch have formed (Figures 3L8B). The telotroch is innervated by two serotonin-like immunoreactive neurite bundles: the inner and the external one. They form two circles in the posterior end of the body (Figure 3L). Thin nervous fibers pass along the lateral and dorsal sides of the larval trunk (Figure 3K).
FMRFamide-like immunoreactive nervous system
In the preactinotroch, the thin and interrupted FMRFamide-immunoreactive neurites appear along the edge of the preoral lobe (Figure 9C). Interestingly, the first FMRFamide reactive neurite runs along the edge of the preoral lobe, whereas the serotonin-like immunoreactive neurite extends from the apical organ to the tentacular ridge.
The first perikarya with FMRFamide-like immunoreactivity occur in the young actinotroch. Perikarya are situated on the dorso-lateral side of the preoral lobe behind the neuropil (Figures 5D, 9D, 10C). Usually, two or three perikarya appear simultaneously. These are large bipolar cells, which connect to the neuropil via thin neurites (Figure 9D). In young actinotrochs, several thin neurites originate around the mouth and along the medio-ventral line of the oral field (Figure 9D). These neurites are very weak (Figure 10C).
After several hours, new FMRFamide-immunoreactive elements appear in young actinotrochs (Figure 9E, F). These are the perikarya of the oral field epidermis, perikarya in the midgut, and radial neurites in the preoral lobe (Figures 9E-K, 10D). In young actinotroch larvae, the apical organ consists of bipolar and monopolar perikarya. The latter is column-shaped with wide apical and basal parts (Figure 9G). The basal part contains the nucleus; the apical part is always brightly stained. The basal part of the perikaryon forms a process that passes into the neuropil. Bipolar perikarya are located distant from the neuropil and connect to it via long anterior processes (Figure 9I). The posterior process runs along the dorso-lateral side of the body and forms the primary tentacular neurite bundle. At this stage, the paired ventral neurite bundle contains six bipolar perikarya, which contact each other via thin longitudinal neurites. These bipolar perikarya form the longitudinal neurite bundles which pass from the oral ring to the tentacular ridge (Figure 9H). Second, two groups of latero-ventral perikarya, each consisting of two or three perikarya (the lower ventro-lateral perikarya), are located near the tentacular ridge (Figure 9E). Third, two large perikarya arise on the ventro-lateral side near the site of contact between the preoral lobe and the hyposphere (the upper ventro-lateral perikarya) (Figure 9F, J). These perikarya connect to the marginal neurite bundle via the thin precursors of the ventro-lateral neurites (which is a part of the marginal neurite bundle and which continues into the oral field) (Figure 9J). Perikarya in the midgut are located near the pyloric sphincter; they are bipolar and triangular with a wide basal part that forms two processes that extend around the midgut (Figure 9K). Radial neurites of the preoral lobe are concentrated along its midline and run along the radial muscles (Figure 10D). At this stage, two to five radial neurites with median perikarya can be detected in the preoral lobe.
In 6-day-old larvae, new perikarya appear adjacent to the paired ventral neurite bundle and in the epidermis of the preoral lobe (Figures 5E, 10E, 11F). In the latter case, most of the perikarya form two main neurites: one passes into the neuropil and the other connects to the marginal neurite bundle (Figure 11G). In some cases, the perikarya of the preoral lobe form more than two neurites, which spread into the preoral lobe epithelium (Figure 11G). The perikarya of the ventral neurite bundles form two longitudinal rows, which contact each other via thin commissures (Figure 11F, H). There are at least two commissures between the paired perikarya of the two longitudinal rows (Figures 10G, 11F, H). A new and large perikaryon appears near the mouth, in the epidermis of the oral field (Figure 10F). At this stage, the upper ventro-lateral perikarya form thin longitudinal ventro-lateral neurites, which contact the lower ventro-lateral perikarya (Figures 10F, 11F).
In 24-day-old larvae, the FMRFamide-like immunoreactive nervous system is highly complex and contains numerous thin neurites (Figures 5F, 10H, I, 12B). The most prominent nervous element is the apical organ. It contains monopolar and bipolar or multipolar perikarya (Figure 12C). Monopolar perikarya are triangular with a narrow apical part and a wide basal part; the latter produces one basal process that extends into the neuropil. As before, the bipolar or multipolar perikarya form two dorso-lateral groups that contain between three and seven perikarya (Figure 12C). The dorso-lateral parts of the tentacular neurite bundle arise from the dorsal groups of the bipolar or multipolar perikarya of the apical organ. The tentacular neurite bundle passes along the tentacular ridge (Figure 10I). On the dorsal side, the tentacular neurite bundle contains upper and lower branches (Figure 10I). On the median line of the preoral lobe, the median neurite bundle appears (Figure 10J). It contains several neurites and perikarya. The latter are concentrated in the central part of the median neurite bundle (Figure 12D). Radial and circular neurites and two marginal neurite bundles innervate the preoral lobe. Both marginal neurite bundles split into numerous thin neurites where the preoral lobe merges with the hyposphere. The anterior marginal neurite bundle continues into ventro-lateral neurites of the oral field, whereas the posterior marginal neurite bundle passes into the lateral neurites of the oral field (Figure 10I). Thus, there are two pairs of groups of neurites that pass from the preoral lobe to the tentacular neurite bundle and that innervate the lateral and ventro-lateral sides of the oral field. The ventral side of the oral field is innervated by the paired ventral neurite bundle, which contains 15–17 bipolar perikarya (Figure 12E). Some of these perikarya are paired, but commissures are not present. The ventral neurite bundles connect to the oral ring. The latter is very complex and contains numerous neurites and perikarya. The neurites spread along the esophagus and form the cylindrical nerve net with a long dorsal side and a short ventral side (Figure 10H). There are six large perikarya on the dorsal side of the esophagus, whereby three are situated on the dorsal wall of the cardiac sphincter (Figure 10K). At this stage, there are two large perikarya, which are located in the epidermis of the oral field near the mouth (Figure 10I, K). These FMRFamide-reactive perikarya form two or three processes, some of which are in contact with the oral nerve ring (Figure 10K; see also ). The number of perikarya in the midgut increases to as many as 8–10 (Figure 12F). They form a circle around the midgut near the pyloric sphincter. The telotroch is innervated by the nerve ring, which passes along its external perimeter (Figure 10I). The trunk and dorsal body region are innervated by thin neurites (Figure 10I).
The structure of the larval apical organ in lophotrochozoans and deuterostomes
The phoronid larval nervous system was first described on the basis of immunocytochemistry by Hay-Schmidt , who analyzed individual stages of early embryos and young larvae of Phoronis vancouverensis. According to these and our data, the first perikarya appear in the apical plate. The differentiation of the first perikarya in the apical organ is a common feature for numerous – but not all – lophotrochozoans ([24–27]; but see [28–30]). As is the case in other lophotrochozoans, the initial serotonin-like immunoreactive perikarya in the apical organ are flask-shaped in phoronid embryos. They possess a thin basal process and are retained during the entire larval development. The organization of the apical organ becomes more complex with age, as is indicated by the dramatic increase in the number of cells that constitute the apical organ.
In Phoronopsis harmeri larvae, four types of perikarya form the apical organ: monopolar and bipolar (or multipolar) serotonin-like immunoreactive as well as monopolar and bipolar (or multipolar) FMRFamide-like immunoreactive. Within the apical organ, only the monopolar serotonin-like immunoreactive perikarya are flask-shaped. In young larva, there are 20–25 monopolar serotonin-like immunoreactive perikarya, which form a horseshoe-shaped (U-shaped) field with two dorsal branches. These cells are monociliated, flask-shaped, and their basal processes pass to the centre of the apical organ and form the neuropil. Bipolar (or multipolar) serotonin-like immunoreactive perikarya and some of their processes form the posterior-most layer of the apical organ. Their perikarya form two (left and right) groups with 5–7 perikarya, which are located directly under the monopolar perikarya. In 24-day-old larva, the neuropil, which is composed of the neurites of the bipolar (or multipolar) serotonin-like immunoreactive perikarya, is subdivided into two portions (left and right). The monopolar FMRFamide-like immunoreactive perikarya form one group each on the left and right side of the apical organ, respectively. Bipolar or multipolar FMRFamide-like immunoreactive perikarya are strongly stained and form two dorso-lateral groups, which occupy the most dorsal area of the apical organ. Groups of bipolar or multipolar FMRFamide-like immunoreactive perikarya connect to the tentacular FMRFamide-like immunoreactive neurite bundle and appear earlier than monopolar FMRFamide-like immunoreactive perikarya.
Hay-Schmidt  found three types of perikarya in the apical organ of Phoronis muelleri. These are monopolar and bipolar or multipolar serotonin-like immunoreactive perikarya as well as monopolar FMRFamide-like immunoreactive perikarya. In early larvae of P. vancouverensis, the apical organ consists of two types of perikarya: monopolar and bipolar or multipolar serotonin-like immunoreactive cells . Interestingly, the apical organ neither contains monopolar nor bipolar (multipolar) FMRFamide-like immunoreactive cells. In Phoronis pallida, two types of serotonin-like immunoreactive perikarya were found in the apical organ: bipolar ciliate and nonciliate .
Taken together, the data available show that the apical organs of phoronid larvae have a much more complex architecture than that of other lophotrochozoans. It consists of at least four cell types and contains approximately 30–50 serotonin-like immunoreactive and 16–20 FMRFamide-like immunoreactive perikarya. The majority of the serotonin-like immunoreactive cells consist of monopolar or (presumably) bipolar flask-shaped perikarya, which form a U-shaped mass.
In most lophophotrochozoan larvae, the apical organ consists of only a few serotonin-like immunoreactive flask-shaped cells (annelids: [28–31]; molluscs: [32–34]; polyclad flatworms: ; ectoprocts: [36, 37]; brachiopods: [9, 38]; see  for review). Such a simple apical organ can therefore be regarded as a basal feature of spiralian protostomes [24, 27, 39]. However, there are deviations from this general pattern. The high degree of complexity of the apical organ of entoproct and polyplacophoran larvae has been considered one of several apomorphies of monophyletic Entoprocta-Mollusca (i.e., Tetraneuralia; ). On the other hand, some polychaete annelids or ectoprocts seem to lack flask-shaped cells in the apical organ altogether [40, 41].
The organization of the apical organ of lecithotrophic brachiopod larvae differs from that of the swimming planktotrophic juveniles of glottiid brachiopods. While the latter have numerous, probably non-flask-shaped cells, the former have four or more (eight?) serotonin-like immunoreactive flask-shaped cells [9, 38, 42], again suggesting that these cells belong to the groundpattern of Lophotrochozoa.
In the deuterostome larvae investigated so far (enteropneust tornariae and echinoderm larvae), the apical organ consists of more than ten or fifteen serotonin-like immunoreactive perikarya, the shape of which probably being typically flask-shaped [43–45].
Accordingly, two scenarios concerning the evolution of complexity in invertebrate apical organs appear principally possible. Either, the last common ancestor of protostomes and deuterostomes had a simple apical organ and its complexity evolved three times independently: once in the entoproct-molluscan (tetraneuralian) lineage, once in phoronids, and once in the deuterostomes, or a complex apical organ was present in the last common ancestor and has been retained in the three above-mentioned clades, while it was subsequently reduced multiple times independently in the remaining spiralians.
Comparison of the overall neural architecture of phoronid larvae with other lophotrochozoan and deuterostomian larvae
In young P. harmeri larvae, the serotonin-like immunoreactive nervous system consists of four perikarya-containing subsets: the apical organ, a group of six to eight perikarya distributed along the edge of the preoral lobe, frontal organ, and a group containing the oral nerve ring with ventro-lateral perikarya, which connect to the ventral neurite bundles. Perikarya of the preoral lobe interconnect with the serotonin-like immunoreactive marginal neurite bundles of the preoral lobe and can not be recognized in late stages. However, in P. muelleri larvae, these perikarya are retained in late stages and are concentrated along the midline of the edge of the preoral lobe (note that the serotonin-like immunoreactive marginal neurite bundle is absent in P. muelleri larvae; ).
In older P. harmeri larva, a fourth serotonin-like immunoreactive perikaryon appears in the frontal organ. It contains several perikarya which connect to the serotonin-like immunoreactive median neurite bundle of the preoral lobe. The frontal organ has been described for other phoronid larvae as well  and contains serotonin-like immunoreactive perikarya . The median neurite bundle of the preoral lobe is found in all phoronid larvae studied so far, except for young stages of P. vancouverensis. In the larva of P. harmeri, this neurite bundle emerges from the left group of serotonin-like immunoreactive bipolar (or multipolar) perikarya.
P. harmeri is the only phoronid larva to date for which a paired ventral neurite bundle has been described (, herein). It consists of two longitudinal neurite bundles, each being associated with several bipolar (or multipolar), paired perikarya. The ventral neurite bundles are interconnected by serially repeated commissures. Such a ventral nervous system is common for many larval (and/or) adult spiralians, and although the number of the ventral neurite bundles may vary between and even within various phyla, a paired ventral neurite bundle with serially arranged commissures is usually considered basal for lophotrochozoans . Accordingly, the larva of P. harmeri appears to have retained this neural structure, while it was lost in the adults as well as in adults and larvae of the other phoronid species investigated so far. At the same time, the paired ventral neural bundle of P. harmeri larvae differs from the one of typical spiralian larvae (for details see ).
In all phoronid larvae studied to date, the serotonin-like immunoreactive tentacular neurite bundle is a prominent structure. In P. harmeri larvae, two dorso-lateral branches of the tentacular neurite bundle are connected each other via a dorsal commissure, which seems to be lacking in other phoronid species. According to some data , there are two tentacular neurite bundles: one minor (extends along tentacles over them) and one main bundle (extends along tentacles under them).While in phoronid larvae the tentacular neurite bundle forms contact with the apical organ, the apical organ never directly connects to the neurite bundles that innervate the ciliated bands in spiralian trochophores [25, 29, 30]. In ambulacrarian larvae (chinoderms and hemichordates), however, the apical organ is also connected to the neurite bundles of ciliated bands, as is the case in the phoronids [45–48]. Moreover, according to some data [47, 48], the apical organ of echinoderms arises as a bilaterally symmetrical nerve plexus, which is generated by neurite bundles of ciliated bands. In ambulacraria, the apical organ arises after or simultaneously with the neurite bundles and perikarya associated with the ciliary bands [44, 47], while in actinotroch larvae, serotonin-like immunoreactive neurites innervating the ciliated bands are formed after the establishment of the apical organ. Accordingly, the left and right branches of the apical organ give rise to the two lateral tentacular neurite bundles of the phoronid larva. The telotroch in phoronid larvae is innervated by two (in P. harmeri) or one (in P. muelleri) serotonin-like immunoreactive circular neurite bundles.
Thus, in all actinotrochs studied, all ciliated bands are innervated by prominent serotonin-like immunoreactive neurite bundles, some of them (e.g., the marginal neurite bundle) containing serotonin-like immunoreactive perikarya. Scattered perikarya associated with neurites that innervate ciliated bands are known from larvae of enteropneust and echinoderm deuterostomes [44, 45, 49, 50].
The larval FMRFamide-like immunoreactive nervous system is very complex in P. harmeri. In this species, the FMRFamide-like immunoreactive nervous system appears to be associated with the major muscle systems. The first neurites appear in the epidermis of the apical plate and then along the edge of the preoral lobe (marginal neurite bundle) in young actinotrochs and is retained in late stages. In older actinotrochs, there are two FMRFamide-like immunoreactive marginal neurite bundles [16, 17], herein. These bundles extend into the oral field and furcate into individual neurites, which ventrolaterally connect to two groups of perikarya. In most previous reports [16, 19, 21], the FMRFamide-like immunoreactive system of actinotrochs does not seem to contain any perikarya even in the apical organ. However, in P. harmeri, there are five main groups of perikarya, which are located in the oral field (three groups), the preoral lobe, and the midgut. Perikarya in the oral field were found in P. muelleri larvae. They are multiplied and form the layer along the upper border of the tentacular ring . In larvae of P. harmeri and P. muelleri, there are perikarya along the medio-ventral side of the oral field. In young larvae of P. harmeri, these perikarya are part of the paired ventral neurite bundle. In older larvae, the commissures interconnecting the ventral neurite bundles were not found . Another group of perikarya is formed by perikarya which are scattered in the epidermis of the preoral lobe , herein. Neurites of these perikarya are associated with the radial muscles of the preoral lobe. Perikarya in the midgut were described for the first time in P. harmeri. The FMRFamide-like immunoreactive tentacular neurite bundle was found in all actinotrochs studied to date [16, 17, 19, 21]. It runs under the tentacles and has two pairs of dorsal branches in P. harmeri. The FMRFamide-like immunoreactive nerve ring around the telotroch is described for the first time herein.
The digestive tract of the actinotroch larva is associated with serotonin-like immunoreactive and FMRFamide-like immunoreactive perikarya and neurites (for details see ). The most prominent nervous elements were found in the esophagus (oral nerve ring), midgut (FMRFamide-reactive perikarya and circular neurites), and the proctodaeum of actinotroch larvae. Some bilaterian planktotrophic larvae have a nerve ring around the mouth (esophagus), but the innervation of other parts of the digestive tract is still unclear. In trochozoan planktotrophic larvae, there is an oral nerve ring, which connects to apical organ via esophageal connectives [25, 28–30]. Some deuterostome larvae have serotonin-like immunoreactive, FMRFamide-like immunoreactive, and SALFamidergic perikarya and neurites in the esophagus, stomach, and pylorus [43, 45]. These prominent neural features are directly correlated with the long planktotrophic life in the water column of these organisms.
P. harmeri has the most complex larval serotin- and FMRFamide-like immunoreactive nervous systems of all phoronids studied to date. The gross anatomy of the nervous system of the actinotroch larva combines characteristics of both lophotrochozoan and deuterostome larvae. On the one hand, the phoronid apical organ consists of flask-shaped serotonin-like immunoreactive monopolar perykaria, which most likely constitute a lophotrochozoan apomorphy . On the other hand, the complexity of the phoronid apical organ, which contains more than 40 serotonin-like immunoreactive perikaria, resembles that of some deuterostome larvae. The distant relationship of phoronids and deuterostomes suggests independent origin of these complex apical organs, although a complex apical organ at the base of the protostome-deuterostome split with subsequent independent simplification in most spiralians cannot be ruled out. The finding of a paired ventral neurite bundle with serially arranged commissures suggests that such a neural feature was part of the ancestral phoronid – and most likely also the ancestral lophotrochozoan – bodyplan, which was secondarily lost in adult phoronids, probably in connection with the acquired sedentary lifestyle.
Adult Phoronopsis harmeri were collected from May to June 2010 in Coos Bay, Oregon, USA, from intertidal sandy sediments. Fertilized eggs, which were extracted from reproductive females by opening the trunk, were kept in glass beakers containing filtered sea water; the temperature of the egg suspension was maintained at 13-14°С by keeping the beaker partially submerged in running sea water on a laboratory bench. Under these conditions, embryos developed normally. Within 15 min of exposure to sea water, two polar bodies were formed and cleavage commenced. Stages of development were monitored with a stereo microscope. The apical tuft and large apical plate became visible 16 h after formation of the polar bodies. This stage was determined as the initiation of neurogenesis. Larval cultures had a density of two larvae per 3–4 ml filtered sea water. Larvae were fed mixtures of Rhodomonas lens and Chaetoceros calcitrans and 75% of the sea water was changed every 2 days. Consecutive stages from the 16-h-old coeloblastula to the 6-tentacled larvae (24 days old) were photographed from living specimens with a Leica DFC 400 camera mounted on an Olympus BX51 microscope equipped with DIC optics.
At 4 h intervals (up to the early actinotrocha), specimens were prepared for immunocytochemistry and confocal laserscanning microscopy (CLSM). Embryos and larvae were narcotised in 0.34 M MgCl2 (Fisher Scientific, Pittsburgh, PA, USA), fixed for 60 min in a 4% paraformaldehyde (Electron Microscopy Science, Hatfield, PA, USA) solution in filtered sea water, and washed (three times x 15 min) in phosphate buffer (pH 7.4) (Fisher Scientific) with Triton X-100 (0.1%) (Fisher Scientific, Pittsburgh, PA, USA) and 0.1% albumine bovine (PBT/BSA) (Sigma-Aldrich, St. Louis, MO, USA). Nonspecific binding sites were blocked with 5% normal donkey serum (Jackson ImmunoResearch, Newmarket, Suffolk, UK) in PBT/BSA for 2 h at room temperature (RT). Subsequently, the specimens were washed in PBT/BSA, the larvae were transferred into primary antibody (either rabbit anti-serotonin or rabbit anti-FMRFamide, ImmunoStar, Hudson, WI, USA) solutions 1:800 in PBT/BSA and incubated overnight at 4 °C. Specimens were washed (three times x 15 min) in PBT/BSA and then exposed to the secondary antibody, donkey anti-rabbit-Alexa Fluor 488 (Invitrogen, Grand Island, NY, USA), at a dilution of 1:1000 for 2 h at RT in the dark. Then, the specimens were washed in PBT/BSA and incubated in a mixture of rhodamine-conjugated phalloidin (1:50) (Fisher Scientific, Pittsburgh, PA, USA) and Hoechst (1:1000) (Fisher Scientific, Pittsburgh, PA, USA) for 1 h at RT in the dark. In the following, they were washed in PBS (three times x 15 min), mounted on a cover glass covered with poly-L-lysine (Sigma-Aldrich, St. Louis, MO, USA), and embedded in Murray Clear or Vectashield (Vector Laboratories Inc., Burlingame, CA, USA). Specimens were viewed with an Olympus confocal microscope (OIMB, OR, USA). Z-projections were generated using the programme Image J version 1.43. Three-dimensional reconstructions were generated using Amira version 5.2.2 software (Bitplane, Zurich, Switzerland).
Since the larval intestine shows strong background signal, it may produce fluorescence on different channels, and some of our micrographs contain nonspecific staining of the intestine. We therefore conducted two experiments to discern autofluorescence of the intestine from true immunocytochemical signal. We treated the larvae according to the protocol described above but omitted either the primary or the secondary antibodies. In both cases, we detected autofluorescence of the intestine on 488, 594 and 633 channels; the samples were devoid of any other signal, however (data not shown).
This research was supported in part by grants to ET from the Russian Foundation for Basic Research (# 11-04-00690), the Russian Ministry of Education and Science (#P727, # 02.740.11.0875), and Grant of the President of Russia (# MD-2892.2011.4). ET is very grateful to her fiend Svetlana Maslakova for hosting her at the Oregon Institute of Marine Biology in 2010 and for providing resources and training, especially in immunohistochemistry and confocal microscopy. ET is grateful to Thomas Schwaha and Stephan Handschuh for helping her with the Amira program. We thank B. Jaffee for help with the English language.
- Cohen B, Weydmann A: Molecular evidence that phoronids are a subtaxon of brachiopods (Brachiopoda: Phoronata) and that genetic divergence of metazoan phyla began long before the early Cambrian. Organisms Divers Evol. 2005, 5: 253-273. 10.1016/j.ode.2004.12.002.View ArticleGoogle Scholar
- Santagata S, Cohen B: Phoronid phylogenetics (Brachiopoda; Phoronata): evidence from morphological cladistics, small and large subunit rDNA sequences, and mitochondrial cox1. Zool J Linn Soc. 2009, 157: 34-50. 10.1111/j.1096-3642.2009.00531.x.View ArticleGoogle Scholar
- Halanych KM, Bacheller JD, Aguinaldo AM, Liva SM, Hillis DM, Lake JA: Evidence from 18 S ribosomal DNA that the lophophorates are protostome animals. Science. 1995, 267: 1641-1643. 10.1126/science.7886451.PubMedView ArticleGoogle Scholar
- Dunn CW, Hejnol A, Matus DQ, Pang K, Browne WE, Smith SA, Seaver E, Rouse GW, Obst M, Edgecombe GD, Sørensen MV, Haddock ShD, Schmidt-Rhaesa A, Okusu A, Kristensen RM, Wheeler WC, Martindale MQ, Giribet G: Broad phylogenomic sampling improves resolution of the animal tree of life. Nature. 2008, 452: 745-749. 10.1038/nature06614.PubMedView ArticleGoogle Scholar
- Hejnol A, Obst M, Stamatakis A, Ott M, Rouse GW, Edgecombe GD, Martinez P, Baguñà J, Bailly X, Jondelius U, Wiens M, Müller WEG, Seaver E, Wheeler WC, Martindale MQ, Giribet G, Dunn CW: Assessing the root of bilaterian animals with scalable phylogenomic methods. Proc Biol Sci. 2009, 276: 4261-4270. 10.1098/rspb.2009.0896.PubMedPubMed CentralView ArticleGoogle Scholar
- Hyman LH: Phoronida. The invertebrates. Volume 5. Smaller coelomate groups. Edited by: Boell EJ. 1959, McGraw-Hill, New York, 228-274.Google Scholar
- Siewing R: Das Archicoelomatenkonzept. Zool Jahrb Anat Ontog. 1980, 103: 439-482.Google Scholar
- Temereva EN, Malakhov VV: Embryogenesis in phoronids. Invert Zool. 2012, 9 (1): 1-39.Google Scholar
- Altenburger A, Wanninger A: Neuromuscular development in Novocrania anomala: evidence for the presence of serotonin and a spiralian-like apical organ in lecithotrophic brachiopod larvae. Evol Dev. 2010, 12 (1): 16-24. 10.1111/j.1525-142X.2009.00387.x.PubMedView ArticleGoogle Scholar
- Temereva EN, Malakhov VV: Embryogenesis and larval development of Phoronopsis harmeri Pixell, 1912 (Phoronida): dual origin of the coelomic mesoderm. Invert Reprod Dev. 2007, 50: 57-66.View ArticleGoogle Scholar
- Temereva EN, Malakhov VV: Organization of the epistome in Phoronopsis harmeri (Phoronida) and consideration of the coelomic organization in Phoronida. Zoomorphol. 2011, 130: 121-134. 10.1007/s00435-011-0126-z.View ArticleGoogle Scholar
- Nielsen C: Animal phylogeny in the light of the trochaea theory. Biol J Linn Soc. 1985, 25: 243-299. 10.1111/j.1095-8312.1985.tb00396.x.View ArticleGoogle Scholar
- Nielsen C: Trochophora larvae: cell-lineages, ciliary bands and body regions. 2. Other groups and general discussion. J Exp Zool (Mol Dev Evol). 2005, 304B: 401-447. 10.1002/jez.b.21050.View ArticleGoogle Scholar
- Harzsch S: Neurophylogeny: architecture of the nervous system and a fresh view on arthropod phyologeny. Integr Comp Biol. 2006, 46: 162-195. 10.1093/icb/icj011.PubMedView ArticleGoogle Scholar
- Hay-Schmidt A: The nervous system of the actinotroch larva of Phoronis muelleri (Phoronida). Zoomorphol. 1989, 108: 333-351. 10.1007/BF00312274.View ArticleGoogle Scholar
- Hay-Schmidt A: Distribution of catecholamine containing, serotonin-like and neuropeptide FMRFamide-like immunoreactive neurons and processes in the nervous system of the actinotroch larva of Phoronis muelleri (Phoronida). Cell Tiss Res. 1990, 259: 105-118. 10.1007/BF00571435.View ArticleGoogle Scholar
- Hay-Schmidt A: Catecholamine-containing, serotonin-lake and FMRFamide-like immunoreactive neurons and processes in the nervous system of the early actinotroch larva of Phoronis vancouverensis (Phoronida): distribution and development. Can J Zool. 1990, 68 (7): 1525-1536. 10.1139/z90-226.View ArticleGoogle Scholar
- Lacalli TC: Structure and organization of the nervous system in the actinotroch larva of Phoronis vancouverensis. Phil Trans Roy Soc L. 1990, 327: 655-685. 10.1098/rstb.1990.0104.View ArticleGoogle Scholar
- Santagata S: Structure and metamorphic remodeling of the larval nervous system and musculature of Phoronis pallida (Phoronida). Evol Dev. 2002, 4: 28-42. 10.1046/j.1525-142x.2002.01055.x.PubMedView ArticleGoogle Scholar
- Santagata S: Larval development of Phoronis pallida (Phoronida): implications for morphological convergence and divergence among larval body plans. J Morphol. 2004, 259: 347-358. 10.1002/jmor.10205.PubMedView ArticleGoogle Scholar
- Santagata S, Zimmer RL: Comparison of the neuromuscular system among actinotroch larvae: systematic and evolutionary implication. Evol Dev. 2002, 4: 43-54. 10.1046/j.1525-142x.2002.01056.x.PubMedView ArticleGoogle Scholar
- Temereva EN: Ventral nerve cord in Phoronopsis harmeri larvae. J Exp Zool (Mol Dev Evol). 2012, 318B: 26-34. 10.1002/jez.b.21437.View ArticleGoogle Scholar
- Temereva EN: The digestive tract of actinotroch larvae (Lophotrochozoa, Phoronida): anatomy, ultrastructure, innervations, and some observations of metamorphosis. Can J Zool. 2010, 88 (2): 1149-1168. 10.1139/Z10-075.View ArticleGoogle Scholar
- Hay-Schmidt A: The evolution of the serotonergic nervous system. Proc R Soc L. 2000, 267: 1071-1079. 10.1098/rspb.2000.1111.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
- Wanninger A: Comparative lophotrochozoan neurogenesis and larval neuroanatomy: recent advances from previously neglected taxa. Acta Biol Hung. 2008, 59 (Suppl): 127-136.PubMedView ArticleGoogle Scholar
- Wanninger A: Shaping the things to come: ontogeny of lophotrochozoan neuromuscular systems and the tetraneuralia concept. Biol Bull. 2009, 216: 293-306.PubMedGoogle Scholar
- Voronezhskaya EE, Tsitrin EB, Nezlin LP: Neuronal development in larval Polychaete Phyllodoce maculate (Phyllodocidae). J Comp Neurol. 2003, 455: 299-309. 10.1002/cne.10488.PubMedView ArticleGoogle Scholar
- McDougall C, Chen W-Ch, Shimeld SM, Ferrier Dek: The development of the larval nervous system, musculature and ciliary bands of Pomatoceros lamarckii (Annelida): heterochrony in polychaetes. Front Zool. 2006, 3 (16): 10.1186/1742-9994-3-16.Google Scholar
- Nezlin LP: The golden age of comparative morphology: laser scanning microscopy and neurogenesis in trochophore animals. Russ J Dev Biol. 2010, 41 (6): 381-390. 10.1134/S1062360410060056.View ArticleGoogle Scholar
- Hay-Schmidt A: The larval nervous system of Polygordius lacteus Scheinder 1868 (Polygordiidae, Polychaeta): immunocytochemical data. Acta Zool. 1995, 76: 121-140. 10.1111/j.1463-6395.1995.tb00987.x.View ArticleGoogle Scholar
- Dickinson AJG, Croll RP: Development of the larval nervous system of the gastropod Ilyanassa obsoleta. J Comp Neurol. 2003, 466: 197-218. 10.1002/cne.10863.PubMedView ArticleGoogle Scholar
- Voronezhskaya EE, Nezlin LP, Odintsova NA: Neuronal development in larval mussel Mytilus trossulus (Mollusca: Bivalvia). Zoomorphol. 2008, 127: 97-110. 10.1007/s00435-007-0055-z.View ArticleGoogle Scholar
- Kristof A, Klussmann-Kolb A: Neuromuscular development of Aeolidiella stephanieae Valdéz, 2005 (Mollusca, Gastropoda, Nudibranchia). Front Zool. 2010, 7 (5): http://www.frontiersinzoology.com/content/7/1/5,Google Scholar
- Rawlinson KA: Embryonic and post-embryonic development of the polyclad flatworm Maritigrella crozieri; implications for the evolution of spiralian life history traits. Front Zool. 2010, 7 (12): http://www.frontiersinzoology.com/content/7/1/12,Google Scholar
- Nielsen C, Worsaae K: Structure and occurrence of cyphonautes larvae (Bryozoa, Ectoprocta). J Morphol. 2010, 2010 (271): 1094-1109.View ArticleGoogle Scholar
- Santagata S: Evolutionary and structural diversification of the larval nervous system among marine bryozoans. Biol Bull. 2008, 215: 3-23. 10.2307/25470679.PubMedView ArticleGoogle Scholar
- Santagata S, Resh C, Hejnol A, Martindale MQ, Passamaneck YJ: Development of the larval anterior neurogenic domains of Terebratalia transversa (Brachiopoda) provides insights into the diversification of larval apical organs and the spiralian nervous system. EvoDevo. 2012, 3 (3): 10.1186/2041-9139-3-3.Google Scholar
- Altenburger A, Wanninger A: Comparative larval myogenesis and adult myoanatomy in the rhynchonelliform (articulate) brachiopods Argyrotheca cordata, A. cistellula, and Terebratalia transversa. Front Zool. 2009, 6 (3): 14-Google Scholar
- Wanninger A, Koop D, Degnan BM: Immunocytochemistry of the larval nervous system of Triphyllozoon mucronatum (Ectoprocta: Gymnolaemata: Cheilostomata) and its fate during metamorphosis. Zoomorphol. 2005, 124: 161-170. 10.1007/s00435-005-0004-7.View ArticleGoogle Scholar
- Brinkmann N, Wanninger A: Larval neurogenesis in Sabellaria alveolata reveals plasticity in polychaete neural patterning. Evol Dev. 2008, 10: 606-618. 10.1111/j.1525-142X.2008.00275.x.PubMedView ArticleGoogle Scholar
- Altenburger A, Martinez P, Wanninger A: Homeobox gene expression in Brachiopoda: the role of Not and Cdx in bodyplan patterning, neurogenesis, and germ layer specification. Gene Expres Patt. 2011, 11: 427-436. 10.1016/j.gep.2011.07.001.View ArticleGoogle Scholar
- Beer AJ, Moss C, Thorndyke M: Development of serotonin-like and SALMFamide-like immunoreactivity in the nervous system of the sea urchin Psammechinus miliaris. Biol Bull. 2001, 200 (3): 268-280. 10.2307/1543509.PubMedView ArticleGoogle Scholar
- Nielsen C, Hay-Schmidt A: Development of the Enteropneust Ptychodera flava: ciliary bands and nervous system. J Morphol. 2007, 268: 551-570. 10.1002/jmor.10533.PubMedView ArticleGoogle Scholar
- Nezlin LP, Yushin VV: Structure of the nervous system in the tornaria larva of Balanoglossus proterogonius (Hemichordata: Enteropneusta) and its phylogenetic implications. Zoomorphol. 2004, 123: 1-13. 10.1007/s00435-003-0086-z.View ArticleGoogle Scholar
- Temereva EN: New data on distribution, morphology and taxonomy of phoronid larvae (Phoronida, Lophophorata). Invert Zool. 2009, 6 (1): 47-64.Google Scholar
- Byrne M, Nakajima Y, Chee FC, Burke RD: Apical organs in echinoderm larvae: insights into larval evolution in the Ambulacraria. Evol Dev. 2007, 9: 432-445. 10.1111/j.1525-142X.2007.00189.x.PubMedView ArticleGoogle Scholar
- Burke RD: Deuterostomy neuroanatome and the body plan paradox. Evol Dev. 2011, 13 (1): 110-115. 10.1111/j.1525-142X.2010.00460.x.PubMedView ArticleGoogle Scholar
- Murabe N, Hatoyama H, Hase S, Komatsu MK, Burke RD, Kaneko H, Nakajima Y: Neural architecture of the brachiolaria larva of the starfish, Asterina pectinifera. J Comp Neur. 2008, 509: 271-282. 10.1002/cne.21742.PubMedView ArticleGoogle Scholar
- Dupont S, Thorndyke W, Thorndyke MC, Burke RD: Neural development of the brittlestar Amphiura filiformis. Dev Genes Evol. 2009, 219: 159-166. 10.1007/s00427-009-0277-9.PubMedView ArticleGoogle Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.