Urochordate Nervous Systems
Summary and Keywords
Urochordates are chordate siblings that comprise the following marine invertebrates: the sessile Ascidiaceae, or sea squirts; planktonic Larvacea; and the pelagic salps, doliolids, and pyrosomes (collectively the Thaliacea), each more beautiful than the next. Tadpole larvae of ascidians and adult larvaceans both have a body plan that is chordate, with a notochord and dorsal, tubular nervous system that forms from a neural plate. Thalaciacea have a ganglion developed from a tubular structure, which has been compared to the vertebrate mes-metencephalic region, and while salps have well developed eyes, other anterior brain components are absent, and the connections within their central nervous system, as well as the neurobiology of other Thaliacea are all little reported. The ascidian tadpole larva is extensively reported, especially in the model species Ciona intestinalis, as is the caudal nerve cord in the larvacean Oikopleura dioica.
Chordate features that share proposed homology with vertebrate features include ciliary photoreceptors that hyperpolarize to light, descending decussating motor pathways that resemble Mauthner cell pathways, coronet cells in the ascidian larva and saccus vasculosus of fishes, the neural canal’s Reissner’s fiber; secondary mechanoreceptors that resemble hair cells; and ascidian bipolar cells that are like dorsal root ganglion cells.
Urochordates, or tunicates, had until recently been considered a subphylum of all chordate animals that include vertebrates, one of three chordate sibling groups of deuterostomes, together with echinoderms and hemichordates (Figure 1). Collectively, chordates have tadpole-type larvae with a notochord and a hollow, dorsal, tubular central nervous system (CNS) that arises from a neural plate, and the tadpole larval form of ascidians, the major class of tunicate, is a powerful reminder of the chordate ancestry of this group. A recent revision (Satoh, Rokhsar, & Nishikawa, 2014) suggests that urochordates are to be considered one of three phyla of a superphylum Chordata, while urochordates themselves comprise three groups: the sessile Ascidiaceae, or sea squirts, with tadpole larvae in many species having a body form and CNS that is chordate (Katz, 1983; Meinertzhagen & Okamura, 2001; Nicol & Meinertzhagen, 1991; Satoh, 1994); the planktonic larvaceans or Appendicularia, with a pelagic life style and an adult body that is likewise chordate in form (Glover & Fritzsch, 2009; Olsson, 1986; Søviknes, Chourrout, & Glover, 2005a, 2005b); and the pelagic salps, doliolids, and pyrosomes that have been far less well studied. All three groups are marine, and many are gelatinous zooplankton, marine planktonic herbivores with short generation times that can reach high population densities (Alldredge & Madin, 1982). Their transparency and size make many features of their internal organization visible by advanced optical methods, and accessible to a wide range of traditional experimental methods, but this requires ready access to an accommodating marine station, one that favors studies that can be initiated spontaneously, whenever local availability provides them with abundant specimens. Exceptions are provided by larvaceans, which are held in continuous culture in several laboratories worldwide (Bouquet, Spriet, Troedsson, Otterå, Chourrout, & Thompson, 2009; Martí-Solans, Ferrández-Roldán, Godoy-Marín, Badia-Ramentol, Torres-Aguila, & Rodríguez-Marí, 2015), and ascidians, which can be both collected, and cultured in the lab with relative ease (Joly, Kano, Matsuoka, Auger, Hirayama, & Satoh, 2007), but which may be fertile only in summer months.
The evolutionary relationships between these groups have frequently been revisited (Figure 1) for deuterostomes as a whole (Cameron, Garey, & Swalla, 2000), for larvaceans (Delsuc, Brinkmann, Chourrout, & Philippe, 2006), and within ascidians, in particular (Swalla, Cameron, Corley, & Garey, 2000). Urochordates are the claimed closest living relatives of vertebrates (Delsuc et al., 2006) and are rapidly evolving (Berná & Alvarez-Valin, 2014; Tsagkogeorga, Turon, Galtier, Douzery, & Delsuc, 2010). A close proximity exists between tunicate and vertebrate genomes (Delsuc et al., 2006). This suggests that the metameric segmentation formerly considered to unify cephalochordates with vertebrates might represent an ancestral feature that has been secondarily lost in tunicates, with attendant reductions also in their respective nervous systems.
Evolutionary considerations are a backdrop to all studies on ascidians (Meinertzhagen, Lemaire, & Okamura, 2004), and we are reminded that nothing about any nervous system makes sense except in the light of evolution (Dobzhansky, 1973)—neither ascidian nervous systems that are chordate in plan, with a dorsal neural tube, nor those that are not, mostly those possessed by other stages of the life cycle. Even so, the quest to cast any and all ascidian neural characters as the harbinger of a vertebrate is one that does seem rather unjust treatment of urochordate special features as a whole, no less than is the obsessive search for all family likenesses among humans.
Reports on the adult nervous system, especially the neural ganglion in Ciona, are quite separate from those on the larva, which has been the subject of extensive developmental studies, most directed at the events, patterning genes, and underlying genetic regulation of early neural development (Hudson, 2016; Meinertzhagen et al., 2004). However, recent studies have made efforts to reveal conserved components between the larval and adult nervous systems (Horie, Shinki, Ogura, Kusakabe, Satoh, & Sasakura, 2011).
This article summarizes, augments, and updates previous ones on urochordate nervous systems (Figure 2), in particular those by Burighel and Cloney (1997) on larval forms and adults, by Bone (1998) on pelagic tunicates (pyrosomes, salps, doliolids, and appendicularians), by Meinertzhagen et al. (2004) on larval ascidians, and by Mackie and Burighel (2005) on the nervous systems in adult tunicates, and with most attention given to morphological findings. It is paralleled by a recent useful primer on the reproductive and general biology of urochordates, their habits, and ecology (Holland, 2016) and by a masterful summary on the developmental neurobiology of the ascidian larva (Hudson, 2016).
Details of the Groups
Reports on the three different groups of urochordates are very unevenly spread, based in large measure on the ease with which specimens can be obtained. The most studied are sessile ascidians that cannot swim away, especially Ciona intestinalis and, in Japan, the commercial species Halocynthia roretzi, with—among larvaceans—the abundant cosmopolitan species Oikopleura dioica. Accounts on Ciona species are lumped together as Ciona intestinalis unless an alternative species; in fact, many studies derive from the closely related Ciona robusta (Brunetti, Gissi, Pennati, Caicci, Gasparini, & Manni, 2015). Next to these, the organization of the nervous system in three forms of pelagic thaliaceans, the salps, pyrosomes, and doliolids, are reported in valuable summaries (Bone, 1998) but have received only sporadic attention (Piette & Lemaire, 2015).
The Tadpole Larva
The CNS of the ascidian tadpole larva is essentially an epithelial tube, which lacks stratification and highly differentiated tracts, and instead comprises circuits of chiefly unbranched neurons connected mostly by axo-axonal connections. The larval CNS has been closely studied in Ciona intestinalis, with most studies providing data on the larval tissues and brain regions from light microscopy. Early studies notably by Grave (1921), were initially updated at cellular level (Katz, 1983; Nicol & Meinertzhagen, 1991), with individual neurons now identified from transmitter immunocytochemistry (Brown, Nishino, Bone, Meinertzhagen, & Okamura, 2005), the expression of neurotransmitter reporter constructs (Horie, Nakagawa, Sasakura, Kusakabe, & Tsuda, 2010; Takamura, Minamida, & Okabe, 2010), or developmental genes (Stolfi & Levine, 2011).
A dedicated study from semi-thin section light microscopy identified about 330 cells in the CNS of four sibling larvae of Ciona intestinalis (Nicol & Meinertzhagen, 1991) and indicated the distribution of these along the length of the tubular CNS. They populate three rather poorly demarcated regions. From rostral to caudal, these are the sensory vesicle, separated by a neck region from the visceral ganglion that contains the motor neurons and that, at a level roughly corresponding to the rostral tip of the notochord, gives way to the caudal nerve cord. The two major regions of the CNS were sensibly renamed, the sensory vesicle as the brain vesicle (Stolfi, Wagner, Taliaferro, Chou, & Levine, 2011) and the visceral ganglion as the motor ganglion (Horie, Sakurai, Ohtsuki, Terakita, Shichida, Usukura et al., 2008), usages adopted here.
The CNS is essentially epithelial in character and comprises neurons and ciliated ependymal cells. An early distinction between these two types was based on the cytological appearance of cells in semi-thin section light microscopy (Nicol & Meinertzhagen, 1991). This is now known to be not entirely reliable, and new definitive criteria from dense reconstruction of electron microscopy (EM) of serial sections identify neurons as having either an axon or a presynaptic site. A total of 177 neurons were reported in a single reconstructed larva (Ryan, Lu, & Meinertzhagen, 2016). The dense reconstruction of all cells within the reconstructed CNS means that no cell can hide or go undetected, and the inventory of cells is therefore comprehensive. Ependymal cells are ciliated cells abutting the neural canal that lack an axon; glial cells, that are neither neurons nor ependymal cells, as so defined, are lacking, as also is myelin (Ryan et al., 2016). Even with these clear criteria, however, assigning the cells to particular morphological categories is still problematic in a few cases. For example, not all neurons with synapses have axons.
A range of morphological classes has been reported in larvae based on a simple library of neurons viewed by light microscopy (Imai & Meinertzhagen, 2007a, 2007b). They are clearly visible in the CNS of, for example, larvae that hatch from precleavage embryos previously transfected by electroporation with a plasmid containing green fluorescent protein (GFP) driven by the promoter of the synaptotagmin gene (Imai & Meinertzhagen, 2007a). Morphological classes in the CNS identified by this method include the following: prominent eminens neurons, large posterodorsal interneurons of the brain vesicle, two large ventroposterior interneurons of the brain vesicle, photoreceptors of the ocellus, and putative antenna cells of the otolith. In the motor ganglion, at least four subtypes of motor neuron have also been identified, together with an ovoid cell that may innervate distal tail muscle cells and contrapelo cells with ascending projections, unique among motor ganglion neurons. Caudally lie somata of two types of planate neurons, the first reported neurons in the caudal nerve cord (Imai & Meinertzhagen, 2007a), which have been confirmed in serial-section electron microscopy (ssEM) series to be short projecting midtail motor neurons, and bipolar tail neurons (Ryan et al., 2016).
Neurons of the peripheral nervous system (PNS) have also been identified by the same methods and include anchor cells, at least eight in each apical adhesive papilla, with axons in the papillar nerves that project to the CNS brain vesicle (Imai & Meinertzhagen, 2007b). Two nerve bundles project from each papilla, suggesting the existence of at least two subpopulations of papillar neurons. Each bundle fasciculates in a stereotyped pattern with axons of the rostral trunk epidermal neurons (RTENs). The RTENs have an elaborate arbor of ciliated dendrites within the tunic (Yokoyama, Hotta, & Oka, 2014), suggesting that each has an extended sensorial field. Two subpopulations of apical trunk epidermal neurons (ATEN), anterior and posterior, have been distinguished, and like the RTENs, these neurons extend ciliated dendritic arbors into the tunic (Yokoyama et al., 2014). Two additional types of tail neuron, dorsal and ventral caudal epidermal neurons, as well as a novel bipolar interneuron, were all also identified by these authors. These neurons likewise extend cilia into the tunic of the fin, that turn as sensory dendrites to extend along the rostro-caudal axis. Together, these ciliated sensory dendrites overlap to then constitute the so-called ascidian dendritic network in the tunic (ASNET; Terakubo, Nakajima, Sasakura, Horie, Konno, Takahashi et al., 2010; Yokoyama et al., 2014).
Many of these details are confirmed and some amended, while others are reported for the first time in a study using ssEM (Ryan et al., 2016). This has provided the following clarifications of data from light microscopy:
(1) There are two eminens neurons of the anterior peripheral pathway, not one, and these are now known to have direct synaptic input from PNS neurons, but differing in their individual connections.
(2) The large posterodorsal interneurons of the brain vesicle number more than two, and co-populate a region with a number of other types of relay neurons.
(3) The photoreceptors constitute three major groups, and each contains a variety of morphological types.
(4) Associated with those morphological types are three additional neuronal types, a photoreceptor tract neuron and two vacuolated anterior neurons terminating within the photoreceptor tract.
(5) There are indeed only two antenna cells of the otolith, not three as previously postulated (Imai & Meinertzhagen, 2007a), and these also differ in their connections and their spatial relationships with the otolith pigment cell through two otolith-associated ciliated cells.
(6) In the motor ganglion at least some of the identified cell types are not motor neurons and include one pair of decussating interneurons, three pairs of descending interneurons, and seven cells called contrapelo cells, which are ascending motor ganglion interneurons receiving input from the PNS.
(7) Five motor neuron types, representing five pairs of neurons, including the caudal neurons with frondose endplates, also exist in the region of the motor ganglion.
(8) Planate neurons, now known to be midtail motor neurons in the caudal nerve cord.
Synapses are marked at presynaptic sites by small clear vesicles 30–60 nm in diameter clustered exclusively at some presynaptic sites, or accompanied by larger electron-lucent vesicles (70–110 nm) at other synapses. Yet other synapses also have dense-core vesicles, large (110–140 nm, medium, or small). Synapses are sometimes unpolarized, with vesicles on both sides of the synaptic cleft, a specialized form of synaptic reciprocity that has been reported in coelenterates and pulmonate mollusks (Meinertzhagen, 2017), and is probably a basal feature.
Many other features reported from EM for sensory neurons in the larva of Diplosoma macdonaldi (Torrence, 1986; Torrence & Cloney, 1982, 1983) and Styela plicata (Torrence, 1986) parallel these for Ciona (Dilly, 1964; Eakin & Kuda, 1971; Ryan, 2015; Stanley MacIsaac, 1999).
The physiology of oscillatory swimming has received considerable attention in many species of tailed craniates, especially lampreys (e.g., Grillner, Wallén, Brodin, & Lansner, 1991). A recently reported connectome for the tadpole larva of Ciona intestinalis reveals motor pathways with similarities to those of the lamprey, including crossed inhibitory pathways and descending pathways from decussating neurons (ddNs) that are a chordate feature. Homology with reticulospinal Mauthner cells was first suggested by light microscopy (Takamura et al., 2010) but has since been validated only from these cells’ synaptic connections (Ryan, Lu, & Meinertzhagen, 2017).
The neurobiology of the tadpole larva in Ciona has captured most recent attention, but it should be remembered that many important details are known for other species, notably the larval sensory system of Diplosoma (Torrence & Cloney, 1982, 1983; reviewed in Torrence & Cloney, 1988).
Nielsen (2000) inclines to the view that direct development in marine invertebrates is secondary, and that a life cycle built around planktotrophic larvae and metamorphosis, as in ascidians, is ancestral among metazoans. Development in Ciona and Clavelina reveals that the stomodaeum/mouth is situated at the same side as the neural tube, which then ought to be described as ventral (Veeman, Newman-Smith, El-Nachef, & Smith, 2010). During the transition to its sessile adult, the premetamorphic larva settles and the neural tube is resorbed along with the tail, the anterior end then differentiating into two parts: the neural gland, which forms from the neurohypophyseal duct, and the ganglion proper, with a cortex of neurons surrounding a central neuropile (Berrill, 1955; Horie et al., 2011; Mackie & Burighel, 2005). Thus the CNS is transformed from a chordate pattern to one more typical of an invertebrate. The patterns of neurotransmitter expression, both for GABA (Bollner, Beesley, & Thorndyke, 1993a) and substance P- and cholecystokinin (Bollner, Beesley, & Thorndyke, 1993b) have been documented during this transition, as also during regeneration (Bollner, Beesley, & Thorndyke, 1992).
Except for the caudal nerve cord, most of the larval CNS is retained during metamorphosis and forms adult CNS cells; most of the larval neurons themselves disappear, and only subsets of cholinergic motor neurons and glutamatergic neurons are retained (Horie et al., 2011). The neurons of the neck give rise to motor neurons of the adult branchial basket. Ependymal cells of the larval CNS contribute to the construction of the adult CNS, thus differentiating into neurons; they are consequently said to be stem-cell-like (Horie et al., 2011).
Considered by some as possibly rather boring, the nervous system of the adult is adapted to its sessile lifestyle no less than the dorsal tubular CNS of its progenitor is to the motile larva and its need for swimming. Based on earlier summaries (Bone & Mackie, 1982; Burighel & Cloney, 1997), Mackie and Burighel (2005) comprehensively review the adult CNS for ascidians as a whole. The present account serves mostly only to update these or highlight topics of special interest.
Most remarkable is the regenerative capacity of the CNS. In C. intestinalis adult, the cerebral ganglion can regenerate completely within a few weeks after ablation (Bollner, Howalt, Thorndyke, & Beesley, 1995), but the regenerative ability decreases with age (Jeffery, 2014). This neurogenerative capacity has led to the proposal that there are neurogenic placodal-like, or proto-placodal regions in larvae (Manni, Agnoletto, Zaniolo, & Burighel, 2005; Abitua, Gainous, Kaczmarczyk, Winchell, Hudson, Kamata et al., 2015). The behavior of Chelyosoma productum and Corella inflata (Ascidiacea) has been studied in normal and deganglionated animals (Mackie & Wyeth, 2000). Chelyosoma productum lives for over a year after deganglionation, during which time the ganglion does not regenerate. After ganglion removal, electrophysiological responses to stimulation and spontaneous activity continue to be transmitted through the body wall and branchial sac, but spread is slow, decremental, and facilitative. In regions showing conduction, cholinesterase histochemistry, and tubulin and gonadotropin-releasing hormone immunolabeling show no evidence of a peripheral nerve net. Motor neuron cell bodies lie entirely within the ganglion or its major roots, and their terminal branches intermingle to form netlike arrays. Apparently, innervation surviving deganglionation is composed of either interconnected motor nerve terminals, interconnected sensory neurites, or some combination of both of these (Mackie & Wyeth, 2000).
Three monoclonal antibodies detect different components of the adult CNS, the expression of which occurs in the same sequence in regeneration as in post-metamorphic development (Bollner et al., 1995).
Primary cultures of neurons from adult (Moss, Beesley, Thorndyke, & Bollner, 1998) and larval (Zanetti, Ristoratore, Francone, Piscopo, & Brown, 2007) Ciona intestinalis have been reported. Among the latter, light microscopy and electrophysiology discriminate photoreceptors and neurons of different types. However, neurite sprouting among some of the latter does not reflect the condition of most neurons in vivo, which lack extensive neurites (Imai & Meinertzhagen, 2007a; Ryan et al., 2016).
The larvacean body is flattened, with a trunk from the ventral side of which emerges the flattened tail. Species such as Oikopleura dioica are pelagic plankton that maintain an actively motile tail throughout the life cycle. A striking general feature is the constancy of cell lineage and consequently of adult cell number. The CNS in Oikopleura is sparsely populated, with about 130 neurons in O. dioica (Mikhaleva, Kreneisz, Olsen, Glover, & Chourrout, 2015), 65 to 75 of which are in the anterior (so-called cerebral) ganglion, and 25 to 30 in the caudal ganglion, with a further 30 in the caudal nerve cord (Søviknes et al., 2005b; Søviknes & Glover, 2007). Earlier counts are reported for O. longicauda (Martini, 1909); glia are not reported.
The rostral region of the brain is largely bilaterally symmetrical with three pairs of brain nerves (Olsson, Holmberg, & Lilliemarck, 1990), but a clear distinction between axons that are sensory and those that are motor has not always been made, and future studies will need to address this issue carefully:
(1) Nerve 1 (Bollner, Holmberg, & Olsson, 1986) is paired and sensory. The nerve of each side arises from the ventral sense organ below the mouth and runs around the pharynx to the brain. In Oikopleura longicauda the ventral sense organ comprises 30 cells (Martini, 1909). Further details are from Bollner et al. (1986) for Oikopleura dioica. Each cell of the ventral sense organ bears a long, slender modified cilium that extends from an apical pocket in the cell, and has a rod-shaped ending in an opening in the epithelium, thus resembling a vertebrate olfactory receptor. The sensory cells together project 15 axons in the Nerve 1 of each side of the pharynx to the brain. Each nerve terminates in a bulb-like swelling that contains four interneuron cell bodies belonging to three different types (Bollner et al., 1986). It is suggested that the ventral sense organ is chemosensory and its receptors similar to vertebrate olfactory sensory receptors, providing a possible “urochordate” counterpart to craniate olfactory organs (Bollner et al., 1986).
(2) Nerve 2. The second pair of cranial nerves connects between the anterior part of the brain and the pharynx and lower lip. Synaptic contacts are ultrastructurally ambiguous, and although reported as if efferent, the 1 µm diameter axon could also, or even alternatively, be sensory. The axons of left and right nerve 2’s each arise from a bipolar soma situated on the left side of the so-called forebrain (Olsson et al., 1990, his Figure 11) and extend rostrally to contact rows of ciliated sensory neurons (Olsson et al., 1990), so-called Tast-Zellen (Martini, 1909). Each axon branches to form a dorsal ramus and a somewhat thicker ventral ramus. Right and left dorsal branches connect to two specialized epidermal cells on the upper lip, and ventral branches follow the lateral pharynx walls ventrorostrally, making frequent contacts with a row of ciliated cells along their course. They then abut ciliated cells at the margin of the lower lip. The bipolar cell bodies lie asymmetrically, the left one bulging to the left, overlying the soma of the right. Caudally, both neurons send axons that travel in the brain to the hindbrain, where they make axo-axonic contacts with the common fiber from nerve 3 (n 3 branchiales communis of Olsson et al., 1990). During their further trajectory in a caudal direction, the two axons also make presynaptic contacts with the perikaryon of a midbrain neuron, which, in its turn, receives connections from a second neuron. From these morphological descriptions, the polarity of conduction in the axon, or whether this is indeed polarized, is unclear and begs further analysis.
(3) Nerve 3 extends from the hind part of the brain and innervates the ciliated cells that drive the water current out through the right and left stigmata (Martini, 1909). This pair of nerves exhibits marked left/right asymmetry, the left containing three axons, one that is agranular and two having dense granules; whereas the right one contains only two, one agranular and the other having dark granules. The three axons of the left nerve diverge at the level of the ciliated stigma ring, and the two of the right nerve also split. Varicosities formed by the terminal of the n3 branch on the surface of the gill opening, suggesting possible involvement in the beating of cilia (Onuma, Isobe, & Nishida, 2016).
Further to these, additional axons of apparently secretomotor neurons project from the dorsal surface of the anterior-most region of the forebrain (Olsson et al., 1990). On the brain’s right side, the ciliary funnel forms a connection with the pharynx (Onuma et al., 2016). A gravity-sensing organ in the dorsal sensory vesicle contains a statolith connected by a shaft to the brain wall, as well as several axon bundles in the inner membrane thought to be sensory cilia and sensory cells (Onuma et al., 2016).
The fibrogen cell, a large, dorsal, anterior ependymal cell within the cerebral ganglion of Oikopleura secretes Reissner’s fiber into the neural canal (Holmberg & Olsson, 1984).
Caudally, two nerves (Fol, 1872)—nervi recurrentes caudae—traced by Langerhans (1877) in Oikopleura, run from the caudal ganglion to bristle-bearing cells in the epidermis, one on each side of the trunk, at a site near the tail’s attachment to the ventral surface of the trunk. These cells and their synaptic connections to the nerves were later described from EM (Bone & Ryan, 1979), and the trajectories of the nerves in the caudal ganglion reported (Holmberg, 1986). Anatomically, they constitute a sensory system that shares some features with the caudal sensory system in the larva of the ascidian Diplosoma macdonaldi (Torrence & Cloney, 1982) but in Oikopleura are secondary mechanoreceptor neurons, compared with those of ascidians, which are primary mechanoreceptors. Mechanoreception in both cases is inferred from the cells’ structure. Clusters of a few cells occur at intervals, forming ganglia from which multiple neural processes extend as they do from the fascicle of the nerve cord (Onuma et al., 2016).
Probably, the 25 to 30 neurons of the caudal ganglion near the base of the tail in Oikopleura contain the central pattern generator for swimming, because the detached tail can generate swimming movements similar to those seen in free-floating intact animals (Bone & Mackie, 1975). Although bouts of rhythmic swimming are driven by the caudal ganglion, each caudal muscle cell is separately innervated by two sets of motor nerves, as well as being coupled to its neighbors; moreover, the external epithelium is excitable and linked to the caudal ganglion by axons from central cells (Bone, 1985). Thus mechanical stimulation of the epithelium evokes receptor potentials followed by action potentials and by bouts of rapid swimming (Bone, 1985).
Living animals are very transparent, so that special labeling methods are not needed to visualize many features of the CNS and neurons. Not only are some sensory structures visible in vivo, but so, too, is the motor innervation of the caudal muscle cells in Oikopleura. The innervation is visible as elaborate corymbiform terminal endplates. These provide a special opportunity in neurobiology, as previously reported in the literature (Bone & Mackie, 1975; Flood, 1975), and would repay closer examination in vivo, for example, to examine the anatomical stability of neuromuscular junctions at different ages or under different physiological conditions.
Transgenic techniques are not routinely applicable to Oikopleura; other techniques, such as RNA interference (RNAi) and morpholino antisense oligos are used for gene knockdown experiments (Omotezako, Nishino, Onuma, & Nishida, 2013) and more recently, DNA interference (DNAi; Omotezako, Onuma, & Nishida, 2015).
The few studies reporting features of nervous systems in pyrosomes and doliolids have been summarized by Bone (1998); the eyes of salps have rightfully attracted attention, especially their anatomy (Metcalf & Johnson, 1905) and the electrophysiology of their photoreceptors. Salps have a complex life cycle with an obligatory alternation of generations between a sexual, aggregate blastozooid, the chain stage, and an asexual, solitary oozooid stage (Brien, 1948). Photoreceptors hyperpolarize to light (Gorman, McReynolds, & Barnes, 1971; McReynolds & Gorman, 1975), like those of the ascidian tadpole larva Ciona (Gorman et al., 1971) and all known vertebrate photoreceptors (Lamb, 2013). The CNS is both short and compact, and the salp neural ganglion resembles that of the adult ascidian (Metcalf & Bell, 1932), suggesting that thaliaceans arose from sessile ascidians (Piette & Lemaire, 2015).
Some reports have identified coronal and cupular sensory organs among the Thaliacea. However, hair cells are absent in Thalia democratica, which possesses excitable epithelia and utilizes a muscular feeding mechanism, and these appear to have undergone secondary loss (Caicci, Gasparini, Rigon, Zaniolo, Burighel, & Manni, 2013). In an interesting juxtaposition, thaliacians have serotonin-like immunoreactive (5-HT-ir) neurons (Valero-Gracia, Marino, Crocetta, Nittoli, Tiozzo, & Sordino, 2016), whereas these are lacking in other adult tunicates. Some comparative details on doliolid nervous systems are provided in Godeaux and Harbison (2003), who report the presence of rounded dorsal neural ganglia with associated nerves in several species. Some aspects of the number and arrangement of nerves, including their projections to muscles, differ between species and stages. Most species also possess a neural gland, or remnant of the larval neural tube that opens to the pharynx, but in some species, a cellular thread to the ciliated funnel is present in its place (Godeaux & Harbison, 2003). In the cerebral ganglion of adult salpid and doliolid species examined, 5-HT-ir neurons are symmetrically distributed (Valero-Gracia et al., 2016), but are absent in the ciliated funnel.
Specific Neuron Systems
Eyes and Visual Systems
Evolution. Rods are reasoned to have evolved from cones at least 420 million years ago (Mya) to confer dim-light vision (Asteriti, Grillner, & Cangiano, 2015). As a result, lampreys have rod and cone photoreceptors with differences similar to those in jawed fishes. Ciona has only a single rhodopsin (Kusakabe, Kusakabe, Kawakami, Satou, Satoh, & Tsuda, 2001) that is related to those of lampreys, compatible with: the much earlier divergence of ascidians (approximately 610 Mya; Erwin, Laflamme, Tweedt, Sperling, Pisani, & Peterson, 2011); the common origin of all chordate eyes and their rhodopsins; and also with the presence of ascidian larval ocelli having ciliary photoreceptors (Eakin & Kuda, 1971; Barnes, 1974) that, like those of their sibling vertebrates (Lamb, 2013), hyperpolarize to light (Gorman et al., 1971). Salp photoreceptors also hyperpolarize to light (Gorman et al., 1971) but, importantly, are rhabdomeric and derived from the ganglion, and thus possibly of independent origin.
Some species of adult ascidians have various simple photoreceptors and eyespots. These presumably mediate light-sensitive spawning behavior (Svane & Young, 1989), but apparently none has been investigated by EM, nor are their pathways known; these are not even necessarily via the cerebral ganglion. A single study (Haffner, 1933) on the regeneration of the siphon and of the eight inverted ocelli illustrates general principles of developmental pattern regulation, and possibly the normal development of ocelli. Thus, new siphons arise from slits made in the body wall of old adults; if the slit is large enough a complete set of eight ocelli is produced; if it is smaller, then fewer than eight. Double slits arranged orthogonally produce siphons with supernumerary ocelli. In all cases, new siphons arise from any point along the length of the old cloacal siphon but from the oral siphon only, distal to the level of the peripharyngeal band (Flimmerschlinge); ocelli proximal to the slit opening arise before ones distal to it. Supernumerary ocelli also arise from incisions near the original ocelli themselves. Histological details of the regenerative formation of ocelli are curious. Pigment cells originating from mesenchymal blood cells are said to wander into the area around the incision and form the pigment cells for the ocelli. Receptor cells for the latter arise from the ocellar anlage, a local ectodermal thickening.
The most highly developed eyes are found in salps (Bullock, 1965); doliolids and larvaceans lack described eyes or photoreceptors, but thaliaceans have photoreceptors that differentiate secondarily from the nervous system (Gorman et al., 1971).
The eyes of salps are understandably but undeservedly neglected. The classical studies on eye structure are those of Goppert (1893) and Redikorzew (1905). A single, brief ultrastructural report in Salpa (Barnes, Gorman, & McReynolds, 1970) indicates that the photoreceptors are microvillous and, surprisingly, that these hyperpolarize to light (Gorman et al., 1971), uniquely so—it appears—amongst rhabdomeric eyes. Important early reports of salp eye development come from the studies of Metcalf (1893a, b) and Metcalf and Johnson (1905). In the adult, there is a single horseshoe-shaped eye in the asexual, solitary oozooid form of Salpa (Figure 3); instead there are five eyes in the sexual, aggregate chain form of the life cycle, a large dorsal one and two pairs of smaller eyes on the surface of the cerebral ganglion (Metcalf, 1893a). Asexual reproduction provides the opportunity to examine the eyes and nervous systems of isogenic individuals at different ages in a macroscopic parental host. Regeneration is reportedly absent (Brien, 1948).
The dorsal unpaired eye of the asexual chain form (about 0.4 mm wide) is larger than that in the sexual chain form, in which early development is reported from Metcalf (1893a, 1893b) on Cyclosalpa and provides interesting details. The single inverted eye arises from a crescent-shaped optic ridge, pushed up from the dorsal surface of the cerebral ganglion (Figure 3). The crescent shape has an anteriorly directed opening, and the ridge enlarges along with the ganglion. Later on, the cells elongate and become columnar, and are flanked on their undersides by intermediate cells (possibly with a supportive function) and pigment cells. Later, the receptors elongate further and differentiate a thicker layer of staining at their basal margins (probably the elaborated membrane lamellae observed by Barnes et al., 1970). The cells of the ridge, which already have axons passing into the neuropile of the ganglion, gradually rise up from the ganglion surface, still applied closely to the overlying ectoderm, to form a thickened optic disc. The disc rises asymmetrically, touching the ganglion at its anterior edge and eventually rolls right over like a flipped penny. This suggests that the flip forms the original fiber projection into a chiasma although this has not been described. The original posterior edge of the disc, now apical in position, curls over, and from it arise everted photoreceptors (Figure 3). These seem to differentiate somewhat later than the inverted receptors of the original anterior margin of the disc. The antiparallel orientation of receptor neurons perhaps reflects only the trivial consequence of the folded topography of the originally planar disc, but this is not explicitly suggested by Metcalf. Also produced at the folded apical edge of the disc is a mass of cells that later develops into the secondary eye, which is also everted. Pathways followed by the three optic nerves are not entirely clear from Metcalf, but apparently, all could fasciculate in a unified coherent projection; a topographic organization remains to be demonstrated. There is apparently no account of the two pairs of other smaller eyes in the chain individual, which resemble the single eye of the adult solitary individual.
From Metcalf’s account, it therefore appears that the dorsal eye of salps is neither ectodermal nor homologous with that of other (uro)chordates, being derived from a portion of the nervous system more posterior than that from which eye outgrowth occurs in chordates or with which the ocellus is associated in the ascidian larva. Pressing problems for future studies are the nature of the projections, threefold or unified, to the neuropile of the ganglion, and identification of photoreceptor interneurons. Metcalf describes the latter as arising after the inner cells of the ganglion mass degenerate to produce a hollow cortex surrounding a neuropile, like a characteristic invertebrate. It is not known whether the photoreceptors also found scattered in the ganglion itself are modified neurons or share the same developmental origins as those of the eyes. The chief obstacles to further experimental work, the sporadic occurrence and fragility of animals, are however considerable.
The ascidian larval ocellus, or eyespot, has been extensively studied. EM observations on Amaroucium (Barnes, 1974), Ciona and Distaplia (Eakin & Kuda, 1971) illustrate a small population of cells (15–20 receptor cells, 1 pigment-cup cell, and 3 lens cells in Amaroucium). These numbers and cell types have been revised in more recent studies on Ciona (Horie, Orii, & Nakagawa, 2005; Ryan et al., 2016), revealing three populations of photoreceptors: 23 Type I photoreceptors with outer segments projecting into the ocellus pigment cup, 7 Type II photoreceptors with outer segments projecting into the neural canal adjacent to the ocellus pigment, and 7 vacuolated Type III photoreceptors located ventro-posteriorly, with outer segments also projecting into the neural canal. Glycogen granules provide a gradient of refractive index in the lens cells (Eakin & Kuda, 1972). The receptors are of modest size (maximum diameter 20 µm), of the ciliary type, and arranged in an outer segment (Barnes, 1971; Barnes et al., 1970; Gorman et al., 1971), and with slender axons (0.7–1.3 µm diameter) that form a single optic nerve terminating in the posterior brain vesicle (Barnes, 1974). They hyperpolarize to light (Gorman et al., 1971) and thus resemble the photoreceptors of other chordate groups (Lamb, 2013). In other families, larvae of molgulids lack an ocellus but nevertheless have extra-ocellar photoreception (Torrence, 1983), while in styelids, the larvae of budding colonial and most other forms, which also lack an ocellus, photoreception is served by a replacement receptor formed in association with the statocyte, the latter thus constituting a compound sense organ or photolith (Garstang, 1928). The position of the photoreceptor in styelid larvae lies in the ventral wall of the brain vesicle, anterior to the statocyte, while the position of ocelli in larvae of other families lies in the latero-dorsal wall of the vesicle. The light-sensitive organ in styelids is considered a new acquisition (Grave & Riley, 1935; Torrence, 1983) with a different developmental origin from the ocellus.
Photoreceptor development has been studied from specimens recovered at different stages from brood pouches (Barnes, 1974) and that therefore lack developmental times. Shortly after the pigmented eye cup first appears the photoreceptors become polarized, developing an apical neck next to the young pigment-cup cell, and a basal body from which the sensory cilium arises, and, abutting the basal lamina, a basal region which later grows an axon. Subsequently, the neck progressively constricts and the cilium elaborates balloon-shaped membrane evaginations over the basal body, to form the lamellae of an outer segment in chordate manner. Microvilli also arise concurrently but from extraciliary plasmalemma and are not necessarily photoreceptive. Microvilli emanate only from the pigment-cup cell in Ciona and Distaplia (Eakin & Kuda, 1971). Outer segment lamellae grow symmetrically around the ciliary axis, unlike vertebrate receptors in which they arise along one edge of the cilium. The lamellae are extracellular and thus topographically equivalent in their organization to those in vertebrate cone outer segments. Junctions resembling chemical synapses are found both upon and between the bases of developing photoreceptors; these are lacking in the mature ascidian larval photoreceptor and must therefore be transient (Barnes, 1974). Recent analysis in fact reveals many synapses between photoreceptor axons and terminals in a hatched larva of Ciona (Ryan et al., 2016).
Larval ocelli also change after hatching. The position and shape of the pigment cup cell and the diameter and density of its pigment granules, as well as the number and length of photoreceptor lamellae all gradually change up until full differentiation is attained at 3.5 h in Ciona (Kajiwara & Yoshida, 1985). These changes roughly parallel the time course of onset of swimming photoresponsiveness and positive phototaxis, at 1.5 h in Ciona, and the later change to negative phototaxis at 3.5 h post-hatching (Kajiwara & Yoshida, 1985). Subsets of the photoreceptors express reporters for different neurotransmitters, some for glutamate (Horie et al., 2008), others for GABA (Zega, Biggiogero, Groppelli, Candiani, Oliveri, Parodi et al., 2008; Horie, unpublished as cited in Horie et al., 2010).
Gravity Sensing Receptors
Statocyte organs, one-celled gravity receptors, are present in adult appendicularians (Onuma et al., 2016), and larvae of ascidians (Eakin & Kuda, 1971; Ohtsuki, 1991; Torrence, 1986; Vorontsova, Nezlin, & Meinertzhagen, 1997). Some doliolid species also possess a statocyst, whereas others have lost a statolith remnant (Godeaux & Harbison, 2003). Eakin and Kuda (1971) discuss the terminological differences for these presumed gravity receptors. Their organization appears to be relatively conserved at least between ascidian species, in which a single statolith organ is accompanied by two ciliated accessory cells and two neurons (Eakin & Kuda, 1971; Ryan et al., 2016), which project axons to the posterior brain vesicle (Imai & Meinertzhagen, 2007a; Ryan et al., 2016; Torrence, 1986). A previous postulate for the presence of three statocyte neurons (Imai & Meinertzhagen, 2007a), has not been upheld from electron microscopy.
Coronet cells having bulbous ciliary protrusions, have been identified in appendicularians (Olsson, 1975) and ascidians (Eakin & Kuda, 1971), and although a close comparison between the coronet cells of tunicates and fishes has long been proposed from structural grounds, as emphasized by their common name (Eakin & Kuda, 1971; Olsson, 1975), their structure in fact differs from craniate coronet cells, each cell in tunicates having only a single sensory protrusion that is itself a modified cilium (Olsson, 1975; Ryan et al., 2016). Their contribution to the neural network of ascidian larvae has remained uncertain until the report of Ryan et al. (2016). Within the neural canal, the presumed sensory structures of these coronet cells—their bulbous protrusions, are interspersed with cilia from other cells (Konno, Kaizu, Hotta, Horie, Sasakura, Ikeo et al., 2010; Ryan et al., 2016). Recent ssEM analysis have revealed that the additional cilia belong to newly described neurons of the dorsal left brain vesicle that send their axons in a nerve bundle to the posterior brain vesicle, in a similar arrangement to that found in the saccus vasculosus of fish (Ryan et al., 2016). This same analysis revealed that the coronet cells themselves form synapses containing exclusively dense core vesicles onto each other, nearby neurons, and the basal lamina of the CNS. These neurons contain dopamine (Moret, Christiaen, Deyts, Blin, Joly, & Vernier, 2005), but it is unclear what neurotransmitter, if any, is actually used by their ciliated neuronal counterparts, the coronet-associated neurons. Both, however, feed into neurons of the brain vesicle that relay synaptic information from a variety of sensory systems to the motor systems (Ryan et al., 2016).
The neuromuscular organization is known in detail chiefly for the ascidian larva Ciona intestinalis (Imai & Meinertzhagen, 2007a; Ryan et al., 2016) and the larvacean Oikopleura dioica (Søviknes & Glover, 2007). Both have morphologically distinctive endplates, first clearly depicted after transfection with GFP in the Ciona larva (Imai & Meinertzhagen, 2007a) and in the larvacean Oikopleura dioica (Martini, 1909). In the latter, beautiful corymbiform end plates (Bone & Mackie, 1975), are visible in the living animal by means of differential interference optics. In Doliolum circular muscle bands are obliquely striated. Each fiber is multinucleate, and on their atrial and external faces are nerve terminals containing electron-lucent vesicles approximately 50 nm in diameter, features that are compared with the structure of other tunicate muscle fibers (Bone & Ryan, 1974).
Adult ascidians possess a range of sensory receptor cells based on ciliated neurons, and ascidian mechanoreceptors offer an important parallel with counterparts of the vertebrate acousticolateralis system (Manni, Mackie, Caicci, Zaniolo, & Burighel, 2006; Rigon, Stach, Caicci, Gasparini. Burighel, & Manni, 2013). Some modern reviews provide morphological data on adult ascidians (Mackie & Burighel, 2005). For example, various mechanoreceptor organs (cupular organs, capsular organs, cupular strands) are found in the atrial cavity of some solitary Enterogona (Bone & Ryan, 1978; Mackie & Singla, 2003, 2004) that are thought to sense local water movements or, in some cases, near-field vibrations. They may allow the adult to regulate water flow through the branchial sac. Two forms of mechanoreceptor complex in adult ascidians exemplify an arrangement of primary and secondary mechanoreceptors. The first is the cupular organ, and the second the coronal organ. The cupula receptor organs contain neurons with cell bodies in the periphery that send axons into the cerebral ganglion, i.e. that are primary sensory cells (Bone & Mackie, 1982). In addition, a newly reported second organ, the coronal organ, occurs in the inner wall of the oral siphon of several ascidian species belonging to the order Pleurogona (Burighel, Lane, Gasparini, Tiozzo, Zaniolo, & Carnevali, 2003; Manni et al., 2006). This organ, reported for Botryllus, Botrylloides, and Styela species, is composed of ciliated receptor cells, which, like vertebrate hair cells, lack axons and are innervated by the axons of secondary sensory neurons in the cerebral ganglion and arranged in a row like the vertebrate lateral line system. The sensory coronal cells exhibit an apical apparatus composed of a cilium accompanied or flanked by rod-like microvilli (stereovilli) but show some variation in the arrangement of their apical specializations, and in Styela plicata bear cilia situated to one side of a crescent-shaped bundle of stereovilli that are graded in length, the longest ones in the middle, thus resembling hair cells of the vertebrate acousticolateralis system, a proposed homologue. Both afferent and efferent synapses occur between hair cells and the neurites contacting their bases. These secondary sensory cells thus resemble those in vertebrates.
In addition to those of the adult, the larvae of ascidians have their own mechanosensory neurons. Most are primary sensory neurons, and many terminate in the CNS (Ryan et al., 2016), and some go on to contribute to the adult mechanosensory network (Abitua et al., 2015).
Cellular Composition of Urochordate Brains
Structural Features of All Neurons
Two cardinal differences between invertebrate and vertebrate brains are, first, that the somata of invertebrate neurons dwell in a cortex or rind surrounding the neuropile, to which they contribute neurites; and, second, that the vast majority of invertebrate neurons lack myelin. These differences have far-reaching consequences for the organization of different brains. A correlate of soma position, the axosomatic placement of synapses in vertebrates, is often assumed to be a canonical vertebrate feature, but it is not invariably so. In those urochordate neurons for which records exist, most synaptic contacts are axo-axonal, and fewer synapses are axo-somatic or axo-dendritic (Olsson et al., 1990; Ryan et al., 2016).
The second special case is myelin, which in vertebrates originated in chondrichthian ancestors (Bullock, Moore, & Fields, 1984) and together with jaws, increased size, increased swimming speed and the possession of a more complex nervous system made the seas a whole lot less safe for invertebrate prey. Early chordates have naked axons: myelin is absent in ascidian larvae (Katz, 1983), the lancelet (Lacalli & Kelly, 2002), and in the hagfish and lampreys (Bullock et al., 1984).
Cell Numbers, Ultrastructure, and Classes
The constancy of cell number was shown, in the early 1900s, in the larvaceans Oikopleura and Fritillaria (e.g., Martini, 1909), in which it was related to the fixed cell lineage and small number of cleavage generations typical of radial cleavage in urochordate embryos, and named eutely (Martini, 1909).
Three tadpole larvae of Ciona intestinalis, the sibling progeny of a single mating, have been reported to exhibit a CNS complement of 331–339 cells (Nicol & Meinertzhagen, 1991), thus with little variation. Of these larvae, a fourth, unrelated larva had similar cell numbers, including 177 neurons, each with an axon, that comprise at least 25 different types (Imai & Meinertzhagen, 2007a; Ryan et al., 2016) and 52 subtypes (Ryan et al., 2016), although some of the latter are not discrete, clearly distinguishable by all morphological features. Adult ascidian nervous systems have an order of magnitude more cells, in the range 102–103 (numbers given in Mackie & Burighel, 2005); whereas larvaceans are more simple, with only 130 or so neurons (Mikhaleva et al., 2015). Martini (1909) gives 87 for Fritillaria, and Søviknes and Glover (2007) give further details for Oikopleura dioica.
Urochordates with chordate-like nervous systems have neurons with simple morphologies reported in the CNS of both the tadpole larva of Ciona intestinalis (Imai & Meinertzhagen, 2007a; Ryan et al., 2016) and the larvacean Oikopleura dioica (Olsson et al., 1990). In larval Ciona, for example, individual neurons are mostly monopolar, but up to 42% have a soma with a single dendrite or with few dendrites, and an axon that usually has a clear terminal (Ryan et al., 2016). Their simple branching patterns are presumed to correlate with the relative paucity of connections possible among so few neurons. However, to examine this question will require an analysis of the brains from different species having a range of cell numbers.
A recent report on the CNS of the tadpole larva of Ciona lists an anterior brain vesicle with 10 types of neurons and 19 subtypes, a posterior brain vesicle with 7 types of neurons and 14 subtypes, and a motor ganglion with 4 types (including two types of motor neurons totally five pairs in all) and 6 subtypes (Ryan et al., 2016). These types are morphological, and the advent of numerous alternative, especially transcriptional methods, as for the vertebrate retina (e.g., Seung & Sümbül, 2014) seems destined to confirm these as well as distinguish additional subtypes.
As a possible correlate of their extreme reductions in overall cell number, urochordate brains from forms with a chordate body plan exhibit a marked sidedness, with left- and right-side neurons specialized for particular pathways and functions. The right-side ocelli and left-side coronet cells are clear cases in most ascidian larvae, but there are many other examples, both in the CNS of larval Ciona (Ryan et al., 2016) and in the larvacean Oikopleura (Olsson et al., 1990). The CNS exhibits a notable sidedness, with more cells on the left side than on the right side. The overall cell complement, including neurons, ependymal cells, and accessory cells, is closely similar (left: 125; right: 129; midline 46), but there are more sensory neurons and pigment cells on the right side and more interneurons on the left. Left and right differences are therefore mostly a question of the differential assignment of cell fates between neuron pairs. Antenna relay neurons project mainly to right-side motor interneurons, and photoreceptor relay neurons to left. Many of the motor ganglion’s synaptic connections occur on only one side, or are more numerous on one side than the other. In the ventral motor ganglion most neurons are by contrast paired, including five pairs of motor neurons and four pairs of interneurons.
The CNS in Oikopleura exhibits a remarkable sidedness, with left and right branches of a bifid neuron in nerve 3 (Olsson et al., 1990), similar to the bifid axon of one of two antenna relay neurons in the larval brain of Ciona, having an axon that splits to form long collaterals terminating at different depths in the motor ganglion (Ryan et al., 2016). It is not clear whether such cases reflect the condensation of two ancestral cells or the division during evolution of the axon from a single cell, to yield one with two axons
Excluding the cases just considered, most neurons have a unitary axon, although various examples of anaxonal neurons exist, for example coronet cells in the larval CNS of Ciona (Ryan et al., 2016). Possibly associated with their small numbers, urochordate neurons may have axons with collaterals, while others may have the bifid axons mentioned above.
The smallest axon sizes in larval Ciona intestinalis are 0.3–1 µm in diameter (Ryan et al., 2016), and giant fibers are lacking, presumably associated with the small size of the animal body and short conduction distances. Most axons form synapses, and are both pre- and postsynaptic at such sites.
Axons usually lack strict fasciculation. In the larval brain of Ciona the axons are too few to exhibit strict fasciculation, and do not even strictly bundle together (Ryan et al., 2016), with some exceptions, particularly for sensory neuron classes in the brain vesicle. In the larvacean Oikopleura, a brain nerve comprising a single axon terminates on two types of touch receptors in the oral region (Olsson et al., 1990). The two nerves are the dendrites of two perikarya in the forebrain and are the master neurons for ciliary reversal in the stigmata, which is a two-neuron reflex. They form axo-axonal synapses with one motor neuron in the midbrain, the command neuron for ciliary reversal.
Neurons in the larva of Ciona, the only urochordate that has been extensively sampled, form chemical synapses. In a single larva the 177 neurons form on average about 38 CNS synapses per neuron, about a quarter of which are neuromuscular synapses, and are augmented by about 18% of this number of gap junctions (Ryan et al., 2016). Most synapses are axo-axonal, some bidirectional, having synaptic vesicles situated on either side of the synaptic cleft, while about 14% are polyadic, having usually two postsynaptic elements per presynaptic release site; yet other sites are substrates for reciprocal or serial synaptic motifs (Ryan et al., 2016).
Given the prevalence of axo-axonic synapses, and the presence in both ascidians and larvaceans of neurons having few or nonexistent dendrites, neurons in these two urochordate groups violate the law of dynamic polarization as proposed for vertebrates in which dendrites alone should be postsynaptic. As a result, it seems safest to refer to all extensions of urochordate neurons simply as neurites, including those arising from the soma. We may suspect that branching simplicity results from each neuron’s few input or output partners, although this has not been analyzed, and would require a comparative study among species with differing wiring complexities.
This enigmatic, immunoreactive strand runs the length of the neural canal of the tubular nerve cord in all chordate groups investigated: the ascidians Halocynthia (Numakunai, Ishikawa, & Hirai, 1965), Amaroucium and Diplosoma (Olsson, 1972); the larvacean Oikopleura (Olsson, 1962); and the cephalochordate Branchiostoma (Olsson & Wingstrand, 1954). In the mammalian brain the subcommissural organ secretes glycoproteins that aggregate to form Reissner’s fiber that binds and clears away monoamines, presumably those released by the brain (Caprile, Hein, Rodríguez, Montecinos, & Rodríguez, 2003). Its function in urochordate brains may be similar, but is unproven.
Glia lack clear acknowledgment in urochordate brains. Basal groups generally lack clear glia, or at any rate lack reported forms, despite some people ascribing the term glia to ependymal cells of the neural tube. Nevertheless, glia seem to have had an early, parallel evolution to the neurons they surround, even if basal taxa such as urochordates lack convincing examples (Hartline, 2011). Historically, glia have mostly been identified by their exclusion from criteria, such as the presence of an axon or of presynaptic sites, used to recognize neurons. But there are many exceptions albeit these may be minor. The ascidian tadpole larva in Ciona has some CNS cell types that are not clearly neuronal or ependymal—that is, that lack either an axon or structural features of presynaptic sites (Ryan, 2015; Ryan et al., 2016). Glia have been tabulated for various invertebrate groups by Roots (1978) and by Radojcic and Pentreath (1979), and recently updated by additional molecular criteria (Hartline, 2011). Given these uncertainties, new glial types may well come to light.
Connectomes, Circuits, and Networks
A complete connectome has recently been reported for a single tadpole larva in Ciona intestinalis (Ryan et al., 2016), obtained by dense reconstruction from ssEM. Despite its numerical simplicity the connectome nevertheless reveals remarkable network complexity, constituting a so-called small-world network, characterized by highly connected local subnetworks linked by fewer long-range connections (Bezares-Calderón & Jékely, 2016; Watts & Strogatz, 1998).
Overall, the network is highly bidirectional, with more feedback loops in 2-node motifs than would be predicted from a random network. Such motifs have been proposed to provide robust dynamical stability (Prill, Iglesias, & Levchenko, 2005). This trend of bidirectionality holds true in both 3- and 4-node motifs as well, with the number of fully connected motifs (those in which all components are reciprocally connected) much higher than would be expected by chance. These types of motif generally indicate that a network contains well-connected groups (“cliques”) and exhibits a great deal of pathway redundancy. This is a common trait of biological networks that are small and highly connected, with few neurons coordinating complex behaviors.
The numbers of synaptic contacts provide a measure of pathway strength, with presynaptic sites ranging from less than 10 to greater than 300 per neuron for the 177 CNS neurons of the tadpole larva of Ciona intestinalis (Ryan et al., 2016). Some circuit details are also reported for the CNS of the larvacean Oikopleura dioica (Olsson et al., 1990), but these lack information on pathway strengths and are far more sparsely identified than those from the larval brain of Ciona, making the two hard to reconcile with each other.
Functional Aspects of Connectome Networks
Some indication of the polarity of transmission within connectome networks can be gained from reporter-driven expression of GFP, for ChAT or VAChT (acetylcholine), vGAT and GAD (GABA), or TH (DOPA and dopamine) in larval ascidians and in situ expression of mRNA for choline acetyltransferase (ChAT) in the larvacean Oikopleura. These provide evidence for the neurotransmitter phenotype, but indicate only the reporter for a gene required for transmitter synthesis or transport, and neither the expression of the gene nor its translation nor the presence of the neurotransmitter itself, and certainly not the release or physiological action of the latter. Such proxy evidence for a neurotransmitter (X) does not entitle reference to the cell in question providing that evidence as being “Xergic,” as frequently appears in the literature.
More direct evidence comes from the presence of the neurotransmitter itself, for example, for GABA in Ciona (Brown et al., 2005) and Oikopleura (Bollner, Storm-Mathisen, & Ottersen, 1991; Søviknes et al., 2005a), or 5-HT, for example in Phallusia (Pennati, Groppelli, Sotgia, Zega, Pestarino, & De Bernardi, 2001) and the peripheral neurons of the papillae in Clavelina (Pennati, Groppelli, De Bernardi, Mastrototaro, & Zega, 2009), but immunocytochemical methods are capricious, at least in larval ascidians, many antibodies failing to yield any signal at all (Vorontsova et al., 1997), possibly because their tiny reserve depletes during fixation, so that absence of evidence does not provide evidence for absence. In certain cases, the genome may lack the corresponding synthetic enzyme for a particular neurotransmitter, as, for example, histamine and adrenaline in Ciona (Dehal, Satou, Campbell, Chapman, Degnan, De Tomaso et al., 2002). In Ciona, some dopamine cells also accumulate serotonin via a functional serotonin transporter (Razy-Krajka, Brown, Horie, Callebert, Sasakura, Joly et al., 2012).
Functionally, the most useful information comes from studies that reveal the expression not of a neurotransmitter but of its receptor. This can yield information on both the polarity and dynamics of transmission, and thus the conductance change in the postsynaptic neuron, and is therefore most valuable in modeling the functions of anatomically identified circuits. Genes for some receptors are lacking in the genome for Ciona intestinalis, including D1 and D2 receptors (Kamesh, Aradhyam, & Manoj, 2008), but genes for glutamate, GABA, and glycine receptors, muscarinic and nicotinic acetylcholine receptors, and those potentially encoding 5-HT and ADR receptors, are all present. Of these, receptors for GABA (Zega et al., 2008); acetylcholine (Nishino, Baba, & Okamura, 2011), glycine (Nishino, Okamura, Piscopo, & Brown,, 2010), serotonin, and dopamine (Razy-Krajka et al., 2012) have been investigated either pharmacologically, or using genetic reporters. Confirmation of these data comes with the prospect, still unrealized, of single-cell transcriptome studies that would reveal the postsynaptic receptors expressed in target neurons, providing not only evidence for the neurotransmitter released at the presynaptic neuron but also the physiology of its transmission.
The nervous systems of urochordates, especially the CNS of the ascidian larva, are widely and often unfairly used as a basis for insights to the ancestral condition of the vertebrate brain, as if they had no other significance, and as if urochordates were the ancestors rather than siblings of chordates. A recurrent theme in all urochordate biology is therefore the extent to which especially ascidian and larvacean features adumbrate those of vertebrates. In some cases, the similarities are structurally remarkable, as is true between the coronet cells of ascidians and the saccus vasculosus of the fish caudal hypothalamus, or Reissner’s fiber. In others, coincidence cannot be excluded, as, for example, in the presence of cholinergic motor neurons. In yet other cases similarities may have been assumed too readily, as may be the case in inferring ancestral homology between the motor neurons in the ascidian larva and the vertebrate spinal cord, when at least those of the anterior motor ganglion in Ciona share more in common with fish reticulospinal neurons than neurons of the spinal cord (Ryan, 2015).
Abitua, P. B., Gainous, T. B., Kaczmarczyk, A. N., Winchell, C. J., Hudson, C., Kamata, K., et al. (2015). The pre-vertebrate origins of neurogenic placodes. Nature, 524, 462–465.Find this resource:
Alldredge, A. L., & Madin, L. P. (1982). Pelagic tunicates: Unique herbivores in the marine plankton. BioScience, 32, 655–663.Find this resource:
Asteriti, S., Grillner, S., & Cangiano, L. (2015). A Cambrian origin for vertebrate rods. elife, 4.Find this resource:
Barnes, S. N. (1971). Fine structure of the photoreceptor and cerebral ganglion of the tadpole larva of Amaroucium constellatum (Verril) (subphylum: Urochordata; class: Ascidiacea). Zeitschrift für Zellforschung mikroskopische Anatomie, 117(1) 1–16.Find this resource:
Barnes, S. N. (1974). Fine structure of the photoreceptor of the ascidian tadpole during development. Cell and Tissue Research, 155, 27–45.Find this resource:
Barnes, S. N., Gorman, A. L. F., & McReynolds, J. S. (1970). Fine structure and intracellular responses of photoreceptors of a pelagic tunicate. Salpa, Biological Bulletin, 139(2), 414.Find this resource:
Berná L., & Alvarez-Valin, F. (2014). Evolutionary genomics of fast evolving tunicates. Genome Biology and Evolution, 6, 1724–1738.Find this resource:
Berrill, N. J. (1955). The origin of vertebrates. London: Oxford University Press.Find this resource:
Bezares-Calderón, L. A., & Jékely, G. (2016). Think small. elife, 5.Find this resource:
Bollner, T., Beesley, P. W., & Thorndyke, M. C. (1992). The pattern of substance P- and cholecystokinin-like immunoreactivity during regeneration of the neural complex in the ascidian Ciona intestinalis. Journal of Comparative Neurology, 325, 572–580.Find this resource:
Bollner, T., Beesley, P. W., & Thorndyke, M. C. (1993a). Distribution of GABA-like immunoreactivity during post-metamorphic development and regeneration of the central nervous system in the ascidian Ciona intestinalis. Cell and Tissue Research, 272, 553–561.Find this resource:
Bollner, T., Beesley, P. W., & Thorndyke, M. C. (1993b). Substance P- and cholecystokinin-like immunoreactivity during post-metamorphic development of the central nervous system in the ascidian Ciona intestinalis. Cell and Tissue Research, 272, 545–552.Find this resource:
Bollner, T., Holmberg, K., & Olsson, R. (1986). A rostral sensory mechanism in Oikopleura dioica (Appendicularia). Acta Zoologica, 67(4), 235–241.Find this resource:
Bollner, T., Howalt, S., Thorndyke, M. C., & Beesley, P. W. (1995). Regeneration and post-metamorphic development of the central nervous system in the protochordate Ciona intestinalis: A study with monoclonal antibodies. Cell and Tissue Research, 279, 421–432.Find this resource:
Bollner, T., Storm-Mathisen, J., & Ottersen, O. P. (1991). GABA-like immunoreactivity in the nervous system of Oikopleura dioica (Appendicularia). The Biological Bulletin, 180(1), 119–124.Find this resource:
Bone, Q. (1985). Locomotor adaptations of some gelatinous zooplankton. Symposium of the Society for Experimental Biology, 39, 487–520.Find this resource:
Bone, Q. (1998). Nervous system, sense organs, and excitable epithelia. In Q. Bone (Ed.), The biology of pelagic tunicates (pp. 55–80). Oxford: Oxford University Press.Find this resource:
Bone, Q., & Mackie, G. O. (1975). Skin impulses and locomotion in Oikopleura (Tunicata: Larvacea). Biological Bulletin, 149, 267–286.Find this resource:
Bone, Q., & Mackie, G. O. (1982). Urochordata. In G. A. B. Shelton (Ed.), Electrical conduction and behaviour in “simple” invertebrates (pp. 473–535). Oxford: Clarendon.Find this resource:
Bone, Q., & Ryan, K. P. (1974). On the structure and innervation of the muscle bands of Doliolum (Tunicata: Cyclomyaria). Proceedings of the Royal Society of London, Series B Biological Sciences, 187, 315–327.Find this resource:
Bone, Q., & Ryan, K. P. (1978). Cupular sense organs in Ciona (Tunicata”: Ascidiacea). Journal of Zoology (London), 186, 417–429.Find this resource:
Bone, Q., & Ryan, K. P. (1979). The Langerhans receptor of Oikopleura (Tunicata: Larvacea). Journal of the Marine Biological Association of the United Kingdom, 59(1), 69–75.Find this resource:
Bouquet, J. M., Spriet, E., Troedsson, C., Otterå, H., Chourrout, D., & Thompson, E. M. (2009). Culture optimization for the emergent zooplanktonic model organism Oikopleura dioica. Journal of Plankton Research, 31(4), 359–370.Find this resource:
Brandenburger, J. L., Woollacott, R. M., & Eakin, R. M. (1973). Fine structure of eyespots in tornarian larvae (Phylum: Hemichordata). Zeitschrift für Zellforschung und mikroskopische Anatomie, 142(1), 89–102.Find this resource:
Brien, P. (1948). Embranchement des tuniciers: morphologie et reproduction. In P.‑P. Grassé (Ed.), Traité de Zoologie (Vol. 11, pp. 554–894). Paris: Masson.Find this resource:
Brown, E. R., Nishino, A., Bone, Q., Meinertzhagen, I. A., & Okamura, Y. (2005). GABAergic synaptic transmission modulates swimming in the ascidian larva. European Journal of Neuroscience, 22, 2541–2548.Find this resource:
Brunetti, R., Gissi, C., Pennati, R., Caicci, F., Gasparini, F., & Manni, L. (2015). Morphological evidence that the molecularly determined Ciona intestinalis type A and type B are different species: Ciona robusta and Ciona intestinalis. Journal of Zoological Systematics and Evolutionary Research, 53, 186–193.Find this resource:
Bullock, T. H. (1965). Chaetognatha, Pogonophora, Hemichordata, and Chordata Tunicata. In T. H. Bullock & G. A. Horridge (Eds.), Structure and function in the nervous systems of invertebrates (Vol. II, pp. 1559–1592). San Francisco: W. H. Freeman.Find this resource:
Bullock, T. H., Moore, J. K., & Fields, R. D. (1984). Evolution of myelin sheaths: Both lamprey and hagfish lack myelin. Neuroscience Letters, 48(2), 145–148.Find this resource:
Burighel, P., & Cloney, R. A. (1997). Urochordata: Ascidiacea. In F. W. Harrison & E. E. Ruppert (Eds.), Microscopic anatomy of invertebrates. Vol. 15: Hemichordata, Chaetognatha, and the Invertebrate Chordates (pp. 221–347). New York: Wiley-Liss.Find this resource:
Burighel, P., Lane, N. J., Gasparini, F., Tiozzo, S., Zaniolo, G., Carnevali, M. D., et al. (2003). Novel, secondary sensory cell organ in ascidians: In search of the ancestor of the vertebrate lateral line. Journal of Comparative Neurology, 461, 236–249.Find this resource:
Caicci, F., Gasparini, F., Rigon, F., Zaniolo, G., Burighel, P., & Manni, L. (2013). The oral sensory structures of Thaliacea (Tunicata) and consideration of the evolution of hair cells in Chordata. Journal of Comparative Neurology, 521, 2756–2771.Find this resource:
Cameron, C. B., Garey, J. R., & Swalla, B. J. (2000). Evolution of the chordate body plan: New insights from phylogenetic analyses of deuterostome phyla. Proceedings of the National Academy of Sciences of the United States of America, 97, 4469–4474.Find this resource:
Caprile, T., Hein, S., Rodríguez, S., Montecinos, H., & Rodríguez, E. (2003). Reissner fiber binds and transports away monoamines present in the cerebrospinal fluid. Brain Research: Molecular Brain Research, 110(2), 177–192.Find this resource:
Dehal, P., Satou, Y., Campbell, R. K., Chapman, J., Degnan, B., De Tomaso, A., et al. (2002). The draft genome of Ciona intestinalis: Insights into chordate and vertebrate origins. Science, 298, 2157–2167.Find this resource:
Delsuc, F., Brinkmann, H., Chourrout, D., & Philippe, H. (2006). Tunicates and not cephalochordates are the closest living relatives of vertebrates. Nature, 439, 965–968.Find this resource:
Dilly, N. (1964). Studies on the receptors in the cerebral vesicle of the ascidian tadpole, 2. The ocellus. Journal of Cell Science, 105(1), 13–20.Find this resource:
Dobzhansky, T. (1973). Nothing in biology makes sense except in the light of evolution. American Biology Teacher, 35(2), 125–129.Find this resource:
Eakin, R. M., & Kuda, A. (1971). Ultrastructure of sensory receptors in ascidian tadpoles, Zeitschrift für Zellforschung und mikroskopische Anatomie, 112(3), 287–312.Find this resource:
Eakin, R. M., & Kuda, A. (1972). Glycogen in lens of tunicate tadpole (Chordata: Ascidiacea). Journal of Experimental Zoology, 180, 267–270.Find this resource:
Eakin, R. M., & Westfall, J. A. (1962). Fine structure of photoreceptors in Amphioxus. Journal of Ultrastructure Research, 6(5–6), 531–539.Find this resource:
Erwin, D. H., Laflamme, M., Tweedt, S. M., Sperling, E. A., Pisani, D., & Peterson, K. J. (2011). The Cambrian conundrum: Early divergence and later ecological success in the early history of animals. Science, 334, 1091–1097.Find this resource:
Flood, P. R. (1973). Ultrastructural and cytochemical studies on the muscle innervation in Appendicularia, Tunicata. Journal de microscopie, 18, 317–326.Find this resource:
Flood, P. R. (1975). Scanning electron microscope observations on the muscle innervation of Oikopleura dioica Fol (Appendicularia, Tunicata) with notes on the arrangement of connective tissue fibers. Cell and Tissue Research, 164, 357–369.Find this resource:
Fol, H. (1872). Études sur les appendiculaires du détroit de Messine, Memoires de la Société de physique et d’histoire naturelle de Genève, 21, 445–499.Find this resource:
Garstang, W. (1928). The morphology of the Tunicata, and its bearings on the phylogeny of the Chordata. Quarterly Journal of Microscopical Science, 72, 51–187.Find this resource:
Glover, J. C., & Fritzsch, B. (2009). Brains of primitive chordates. In L. R. Squire (Ed.), Encyclopedia of neuroscience (pp. 439–448). Amsterdam: Elsevier.Find this resource:
Godeaux, J. E. A., & Harbison, G. R. (2003). On some pelagic doliolid tunicates (Thaliacea, Doliolida) collected by a submersible off the eastern North American coast. Bulletin of Marine Science, 72, 589–612.Find this resource:
Goppert, R. (1893). Untersuchungen uber das Sehorgan der Salpen. Morphologische Jahrbücher Abteilung für Anatomie, 19, 250–294.Find this resource:
Gorman, A. L. F., McReynolds J. S., & Barnes, S. N. (1971). Photoreceptors in primitive chordates: Fine structure, hyperpolarizing receptor potentials, and evolution, Science, 172, 1052–1054.Find this resource:
Grave, C. (1921). Amaroucium constellaturn (Verrill). II. The structure and organization of the tadpole larva. Journal of Morphology, 36, 71–101.Find this resource:
Grave, C., & Riley, G. (1935). Development of the sense organs of the larva of Botryllus schlosseri. Journal of Morphology, 57, 185–211.Find this resource:
Grillner, S., Wallén, P., Brodin, L., & Lansner, A. (1991). Neuronal network generating locomotor behavior in lamprey: Circuitry, transmitters, membrane properties, and simulation. Annual Review of Neuroscience, 14, 169–199.Find this resource:
Haffner, K. von (1933). Die überzähligen Siphonen und Ocellen von Ciona intestinalis L. (Experimentell-morphologische Untersuchungen), Zeitschrift für wissenschaftliche Zoologie, 143, 16–52.Find this resource:
Hartline, D. K. (2011). The evolutionary origins of glia. Glia, 59, 1215–1236.Find this resource:
Holland, L. Z. (2016). Tunicates. Current Biology, 26, R146–R152.Find this resource:
Holmberg, K. (1986). The neural connection between the Langerhans receptor cells and the central nervous system in Oikopleura dioica (Appendicularia). Zoomorphology, 106, 31–34.Find this resource:
Holmberg, K., & Olsson, R. (1984). The origin of Reissner’s fiber in an appendicularian. Oikopleura dioica, Videnskabelige Meddelelser Naturhistorisk Forening i København, 145, 43–52.Find this resource:
Horie, T., Nakagawa, M., Sasakura, Y., Kusakabe, T. G., & Tsuda, M. (2010). Simple motor system of the ascidian larva: Neuronal complex comprising putative cholinergic and GABAergic/glycinergic neurons. Zoological Science, 27(2), 181–190.Find this resource:
Horie, T., Orii, H., & Nakagawa, M. (2005). Structure of ocellus photoreceptors in the ascidian Ciona intestinalis larva as revealed by an anti-arrestin antibody. Journal of Neurobiology, 65, 241–250.Find this resource:
Horie, T., Sakurai, D., Ohtsuki, H., Terakita, A., Shichida, Y., Usukura, J., et al. (2008). Pigmented and nonpigmented ocelli in the brain vesicle of the ascidian larva. Journal of Comparative Neurology, 509, 88–102.Find this resource:
Horie, T., Shinki, R., Ogura, Y., Kusakabe, T. G., Satoh, N., & Sasakura, Y. (2011). Ependymal cells of chordate larvae are stem-like cells that form the adult nervous system. Nature, 469, 525–528.Find this resource:
Hudson, C. (2016). The central nervous system of ascidian larvae. WIRE’s Developmental Biology, 5.Find this resource:
Imai, J., & Meinertzhagen, I. A. (2007a). Neurons of the ascidian larval nervous system in Ciona intestinalis: I. Central nervous system. Journal of Comparative Neurology, 501, 316–334.Find this resource:
Imai, J., & Meinertzhagen, I. A. (2007b). Neurons of the ascidian larval nervous system in Ciona intestinalis: II. Peripheral nervous system. Journal of Comparative Neurology, 501, 335–352.Find this resource:
Jeffery, W. R. (2014). Closing the wounds: One hundred and twenty five years of regenerative biology in the ascidian Ciona intestinalis. Genesis, 53(1), 48–65.Find this resource:
Joly, J. S., Kano, S., Matsuoka, T., Auger, H., Hirayama, K., Satoh, et al. (2007). Culture of Ciona intestinalis in closed systems. Developmental Dynamics, 236, 1832–1840.Find this resource:
Kajiwara, S., & Yoshida, M. (1985). Changes in behavior and ocellar structure during larval life of solitary ascidians. Biological Bulletin, 169, 565–577.Find this resource:
Kamesh, N., Aradhyam, G. K., & Manoj, N. (2008). The repertoire of G protein-coupled receptors in the sea squirt Ciona intestinalis. BMC Evolutionary Biology, 8, 129.Find this resource:
Katz, M. J. (1983). Comparative anatomy of the tunicate tadpole, Ciona intestinalis. Biological Bulletin, 164(1), 1–27.Find this resource:
Konno, A., Kaizu, M., Hotta, K., Horie, T., Sasakura, Y., Ikeo, K., et al. (2010). Distribution and structural diversity of cilia in tadpole larvae of the ascidian Ciona intestinalis. Developmental Biology, 337(1), 42–62.Find this resource:
Kusakabe, T., Kusakabe, R., Kawakami, I., Satou, Y., Satoh, N., & Tsuda, M. (2001). Ci-opsin1, a vertebrate-type opsin gene, expressed in the larval ocellus of the ascidian Ciona intestinalis. FEBS Letters, 506(1), 69–72.Find this resource:
Lacalli, T. C., & Kelly, S. J. (2002). Floor plate, glia and other support cells in the anterior nerve cord of amphioxus larvae. Acta Zoologica, 83(2), 87–98.Find this resource:
Lamb, T. D. (2013). Evolution of phototransduction, vertebrate photoreceptors and retina. Progress in Retinal and Eye Research, 36, 52–119.Find this resource:
Langerhans, P. (1877). Zur Anatomie der Appendicularien. Monatsberichte der Königlichen Preussische Akademie des Wissenschaften zu Berlin, 561–566.Find this resource:
Mackie, G. O., & Burighel, P. (2005). The nervous system in adult tunicates: Current research directions. Canadian Journal of Zoology, 83(1), 151–183.Find this resource:
Mackie, G. O., & Singla, C. L. (2003). The capsular organ of Chelyosoma productum (Ascidiacea: Corellidae): A new tunicate hydrodynamic sense organ. Brain Behavior and Evolution, 61(1), 45–58.Find this resource:
Mackie, G. O., & Singla, C. L. (2004). Cupular organs in two species of Corella (Tunicata Ascidiacea). Invertebrate Biology, 123(3), 269–281.Find this resource:
Mackie, G. O., & Wyeth, R. C. (2000). Conduction and coordination in deganglionated ascidians. Canadian Journal of Zoology, 78, 1626–1639.Find this resource:
Manni, L., Agnoletto, A., Zaniolo, G., & Burighel, P. (2005). Stomodeal and neurohypophysial placodes in Ciona intestinalis: Insights into the origin of the pituitary gland. Journal of Experimental Zoology B Molecular Developmental Evolution, 304B(4), 324–339.Find this resource:
Manni, L., Mackie, G. O., Caicci, F., Zaniolo, G., & Burighel, P. (2006). Coronal organ of ascidians and the evolutionary significance of secondary sensory cells in chordates. Journal of Comparative Neurology, 495(4), 363–373.Find this resource:
Martini, E. (1909). Studien über die Konstanz histologischer Elemente. I. Oikopleura longicauda, Zeitschrift für wissenschaftliche Zoologie, 92, 563–626.Find this resource:
Martí-Solans, J., Ferrández-Roldán, A., Godoy-Marín, H., Badia-Ramentol, J., Torres-Aguila, N. P., Rodríguez-Marí, et al. (2015). Oikopleura dioica culturing made easy: A low-cost facility for an emerging animal model in EvoDevo, Genesis, 53(1), 183–193.Find this resource:
McReynolds, J. S., & Gorman, A. L. F. (1975). Hyperpolarizing photoreceptors in the eye of a primitive chordate Salpa democratica. Vision Research, 15, 1181–1186Find this resource:
Meinertzhagen, I. A. (2017). Morphology of invertebrate neurons and synapses. In J. H. Byrne (Ed.), Handbook of invertebrate neurobiology. Oxford: Oxford University Press.Find this resource:
Meinertzhagen, I. A., Lemaire, P., & Okamura, Y. (2004). The neurobiology of the ascidian tadpole larva: Recent developments in an ancient chordate. Annual Review of Neuroscience, 27, 453–485.Find this resource:
Meinertzhagen, I. A., & Okamura, Y. (2001). The larval ascidian nervous system: The chordate brain from its small beginnings. Trends in Neuroscience, 24(7), 401–410.Find this resource:
Metcalf, M. M. (1893a). On the eyes, subneural gland, and central nervous system in Salpa. Zoologische Anzeiger, 16, 6–10.Find this resource:
Metcalf, M. M. (1893b). The eyes and subneural gland of Salpa. In W. K. Brooks (Ed.), The genus Salpa, IV, Memoirs from the biological laboratory of the Johns Hopkins University 2 (pp. 305–372). Baltimore: Publication agency of the Johns Hopkins University.Find this resource:
Metcalf, M. M., & Bell, M. M. (1932). The Salpidae: A taxonomic study. In Papers on collections gathered by the “Albatross” Philippine expedition, 1907–1910 (pp. 5–191). Washington, DC: U.S. Government Printing Office.Find this resource:
Metcalf, M. M., & Johnson, M. E. (1905). The anatomy of the eyes and neural glands in the aggregated forms of Cyclosalpa dolichosoma-virgula and Salpa punctata. Biological Bulletin, 9(4), 195–212.Find this resource:
Mikhaleva, Y., Kreneisz, O., Olsen, L. C., Glover, J. C., & Chourrout, D. (2015). Modification of the larval swimming behavior in Oikopleura dioica, a chordate with a miniaturized central nervous system by dsRNA injection into fertilized eggs. Journal of Experimental Zoology, Part B, Molecular Developmental Evolution, 324, 114–127.Find this resource:
Moret, F., Christiaen, L., Deyts, C., Blin, M., Joly, J. S., & Vernier, P. (2005). The dopamine-synthesizing cells in the swimming larva of the tunicate Ciona intestinalis are located only in the hypothalamus-related domain of the sensory vesicle. European Journal of Neuroscience, 21), 3043–3055.Find this resource:
Moss, C., Beesley, P. W., Thorndyke, M. C., & Bollner, T. (1998). Preliminary observations on ascidian and echinoderm neurons and neural explants in vitro. Tissue and Cell, 30, 517–524.Find this resource:
Nicol, D., & Meinertzhagen, I. A. (1991). Cell counts and maps in the larval central nervous system of the ascidian Ciona intestinalis (L.). Journal of Comparative Neurology, 309, 415–429.Find this resource:
Nielsen C. (2000). The origin of metamorphosis. Evolution and Development, 2(3), 127–129.Find this resource:
Nishino, A., Baba, S. A., & Okamura, Y. (2011). A mechanism for graded motor control encoded in the channel properties of the muscle ACh receptor. Proceedings of the National Academy of Science, 108, 2599–2604.Find this resource:
Nishino, A., Okamura, Y., Piscopo, S., & Brown, E. R. (2010). A glycine receptor is involved in the organization of swimming movements in an invertebrate chordate. BMC Neuroscience, 11(6), 1–12.Find this resource:
Numakunai, T., Ishikawa, M., & Hirai, E. (1965). Changes of the structures stainable with modified Gomori’s aldehyde-fuchsine method in the tadpole larvae of the ascidian, Halocynthia roretzi (V. Drasche), relating to tail resorption. Bulletin of the Marine Biological Station of Asamushi, Tohôku University, 12, 161–172.Find this resource:
Ohtsuki, H. (1991). Sensory organs in the cerebral vesicle of the ascidian larva, Aplidium sp.—an SEM study. Zoological Science, 8(2), 235–242.Find this resource:
Olsson, R. (1962). Reissner’s fiber apparatus in its most primitive condition. General and Comparative Endocrinology, 2, 617–618.Find this resource:
Olsson, R. (1986). Basic design of the chordate brain. In T. Uyeno, R. Arai, T. Taniuchi, & K. Matsuura (Eds.). Indo-Pacific fish biology (pp. 86–93). Tokyo: Ichthyological Society of Japan.Find this resource:
Olsson, R. (1972). Reissner’s fiber in ascidian tadpole larvae. Acta Zoologica, 53, 17–21.Find this resource:
Olsson, R. (1975). Primitive coronet cells in the brain of Oikopleura (Appendicularia, Tunicata). Acta Zoologica, 56, 155–161.Find this resource:
Olsson, R., Holmberg, K., & Lilliemarck, Y. (1990). Fine structure of the brain and brain nerves of Oikopleura dioica (Urochordata, Appendicularia). Zoomorphology, 110, 1–7.Find this resource:
Olsson, R., & Wingstrand, K. G. (1954). Reissner’s fiber and the infundibular organ in Amphioxus: Results obtained with Gomori’s chrome alum haematoxylin. Årbok for Universitetet i Bergen, 14, 1–14.Find this resource:
Omotezako, T., Nishino, A., Onuma, T. A., & Nishida, H. (2013). RNA interference in the appendicularian Oikopleura dioica reveals the function of the Brachyury gene. Development, Genes and Evolution, 223(4), 261–267.Find this resource:
Omotezako, T., Onuma, T. A., & Nishida, H. (2015). DNA interference: DNA-induced gene silencing in the appendicularian Oikopleura dioica. Proceedings of the Royal Society, B Biological Sciences, 282.Find this resource:
Onuma, T. A., Isobe M., & Nishida, H. (2016). Internal and external morphology of adults of the appendicularian Oikopleura dioica: An SEM study. Cell and Tissue Research, 367(2), 213–227.Find this resource:
Pennati, R., Groppelli, S., De Bernardi, F., Mastrototaro, F., & Zega, G. (2009). Immunohistochemical analysis of adhesive papillae of Clavelina lepadiformis (Müller, 1776) and Clavelina phlegraea (Salfi, 1929) (Tunicata, Ascidiacea). European Journal of Histochemistry, 53(1), 25–34.Find this resource:
Pennati, R., Groppelli, S., Sotgia, C., Zega, G., Pestarino, M., & De Bernardi, F. (2001). Serotonin localization in Phallusia mammillata larvae and effects of 5-HT antagonists during larval development. Development Growth and Differentiation, 43, 647–656.Find this resource:
Piette, J., & Lemaire, P. (2015). Thaliaceans, the neglected pelagic relatives of ascidians: A developmental and evolutionary enigma. Quarterly Review of Biology, 90(2) 117–145.Find this resource:
Prill, R. J., Iglesias, P. A., & Levchenko, A. (2005). Dynamic properties of network motifs contribute to biological network organization. PLoS Biology, 3(11), e343.Find this resource:
Radojcic, T., & Pentreath, V. W. (1979). Invertebrate glia. Progress in Neurobiology, 12, 115–179.Find this resource:
Razy-Krajka, F., Brown, E. R., Horie, T., Callebert, J., Sasakura, Y., Joly, J.‑S., et al. (2012). Monoaminergic modulation of photoreception in ascidian: Evidence for a proto-hypothalamo-retinal territory. BMC Biology, 10, 45.Find this resource:
Redikorzew, W. (1905). Über das Sehorgan der Salpen. Gegenbaurs Morphologisches Jahrbuch, 34, 204–239.Find this resource:
Rigon, F., Stach, T., Caicci, F., Gasparini, F., Burighel, P., & Manni, L. (2013). Evolutionary diversification of secondary mechanoreceptor cells in Tunicata. BMC Evolutionary Biology, 13, 112.Find this resource:
Roots, B. I. (1978). A phylogenetic approach to the anatomy of glia. In G. Schoffeniels, L. Franck, D. B. Hertz, & D. B. Tower (Eds.), Dynamic properties of glial cells (pp. 45–54). New York: Pergamon.Find this resource:
Ryan K. (2015). The connectome of the larval brain of Ciona intestinalis (L.) (PhD thesis). Dalhousie University, Halifax, Nova Scotia.Find this resource:
Ryan, K., Lu, Z., & Meinertzhagen, I. A. (2016). The CNS connectome of a tadpole larva of Ciona intestinalis highlights sidedness in the brain of a chordate sibling. eLife 5.Find this resource:
Ryan, K., Lu, Z., & Meinertzhagen, I. A. (2017). Circuit homology of decussating pathways in the Ciona larval CNS and the vertebrate startle-response pathway. Current Biology, 27, 1–8.Find this resource:
Satoh, N. (1994). Developmental biology of ascidians. Cambridge, U.K.: Cambridge University Press.Find this resource:
Satoh, N., Rokhsar, D., & Nishikawa, T. (2014). Chordate evolution and the three-phylum system. Proceedings of the Royal Society Biological Sciences, 281(1794).Find this resource:
Seung, H. S., & Sümbül, U. (2014). Neuronal cell types and connectivity: Lessons from the retina. Neuron, 83, 1262–1272.Find this resource:
Søviknes, A. M., Chourrout, D., & Glover, J. C. (2005a). Development of putative GABAergic neurons in the appendicularian urochordate Oikopleura dioica. Journal of Comparative Neurology, 490, 12–28.Find this resource:
Søviknes, A. M., Chourrout, D., & Glover, J. C. (2005b). Development of the caudal nerve cord, motoneurons, and muscle innervation in the appendicularian urochordate Oikopleura dioica. Journal of Comparative Neurology, 503, 224–243.Find this resource:
Søviknes, A. M., & Glover, J. C. (2007). Spatiotemporal patterns of neurogenesis in the appendicularian Oikopleura dioica. Developmental Biology, 311, 264–275.Find this resource:
Stanley MacIsaac S. (1999). Ultrastructure of the visceral ganglion in the ascidian larva Ciona intestinalis: Cell circuitry and synaptic distribution (Master’s thesis). Dalhousie University Halifax, Nova Scotia.Find this resource:
Stolfi, A., & Levine, M. (2011). Neuronal subtype specification in the spinal cord of a protovertebrate. Development, 138, 995–1004.Find this resource:
Stolfi, A., Wagner, E., Taliaferro, J. M., Chou, S., & Levine, M. (2011). Neural tube patterning by Ephrin, FGF and Notch signaling relays. Development, 138, 5429–5439.Find this resource:
Svane, I., & Young, C. M. (1989). The ecology and behavior of ascidian larvae. Oceanography and Marine Biology: An Annual Review, 27, 45–90.Find this resource:
Swalla, B. J., Cameron, C. B., Corley, L. S., & Garey, J. R. (2000). Urochordates are monophyletic within the deuterostomes. Systematic Biology, 49(1), 52–64.Find this resource:
Takamura, K., Minamida, N., & Okabe, S. (2010). Neural map of the larval central nervous system in the ascidian Ciona intestinalis. Zoological Science, 27(2), 191–203.Find this resource:
Terakubo, H. Q., Nakajima, Y., Sasakura, Y., Horie, T., Konno, A., Takahashi, H., et al. (2010). Network structure of projections extending from peripheral neurons in the tunic of ascidian larva. Developmental Dynamics, 239, 2278–2287.Find this resource:
Torrence, S. A. (1983). Ascidian larval nervous system: Anatomy, ultrastructure and metamorphosis. PhD thesis (p. 178). University of Washington, Seattle.Find this resource:
Torrence, S. A. (1986). Sensory endings of the ascidian static organ (Chordata, Ascidiacea). Zoomorphology, 106, 61–66.Find this resource:
Torrence, S. A., & Cloney, R. A. (1982). Nervous system of ascidian larvae: Caudal primary sensory neurons. Zoomorphology, 99, 103–115.Find this resource:
Torrence, S. A., & Cloney, R. A. (1983). Ascidian larval nervous system: Primary sensory neurons in adhesive papillae. Zoomorphology, 102, 111–123.Find this resource:
Torrence, S. A., & Cloney, R. A. (1988). Larval sensory organs of ascidians. In M. F. Thompson, R. Sarojini, & R. Nagabhushanam (Eds.), Marine biodeterioration (pp. 151–164). New Delhi: Oxford & IBH Publishing.Find this resource:
Tsagkogeorga, G., Turon, X., Galtier, N., Douzery, E. J., & Delsuc, F. (2010). Accelerated evolutionary rate of housekeeping genes in tunicates. Journal of Molecular Evolution, 71(2), 153–167.Find this resource:
Valero-Gracia, A., Marino, R., Crocetta, F., Nittoli, V., Tiozzo, S., & Sordino, P. (2016). Comparative localization of serotonin-like immunoreactive cells in Thaliacea informs tunicate phylogeny. Frontiers in Zoology, 13, 45. .Find this resource:
Veeman, M. T., Newman-Smith, E., El-Nachef, D., & Smith, W. C. (2010). The ascidian mouth opening is derived from the anterior neuropore: Reassessing the mouth/neural tube relationship in chordate evolution. Developmental Biology, 344, 138–149.Find this resource:
Vorontsova, M. N., Nezlin, L. P., & Meinertzhagen, I. A. (1997). Nervous system of the larva of the ascidian Molgula citrina (Alder and Hancock, 1848). Acta Zoologica, 78(3), 177–185.Find this resource:
Watts, D. J., & Strogatz, S. H. (1998). Collective dynamics of “small-world” networks. Nature, 393, 440–442.Find this resource:
Yokoyama, T. D., Hotta, K., & Oka, K. (2014). Comprehensive morphological analysis of individual peripheral neuron dendritic arbors in ascidian larvae using the photoconvertible protein Kaede. Developmental Dynamics, 243, 1362–1373.Find this resource:
Zanetti, L., Ristoratore, F., Francone, M., Piscopo, S., & Brown, E. R. (2007). Primary cultures of nervous system cells from the larva of the ascidian Ciona intestinalis. Journal of Neuroscience Methods, 165(2), 191–197.Find this resource:
Zega, G., Biggiogero, M., Groppelli, S., Candiani, S., Oliveri, D., Parodi, M., et al. (2008). Developmental expression of glutamic acid decarboxylase and of gamma-aminobutyric acid type B receptors in the ascidian Ciona intestinalis. Journal of Comparative Neurology, 506(3), 489–505.Find this resource: