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date: 17 November 2018

Development of the Nervous System of Invertebrates

Abstract and Keywords

The complex architecture of the nervous system is the result of a stereotyped pattern of proliferation and migration of neural progenitors in the early embryo, followed by the outgrowth of nerve fibers along rigidly controlled pathways, and the formation of synaptic connections between specific neurons during later stages. Detailed studies of these events in several experimentally amenable model systems indicated that many of the genetic mechanisms involved are highly conserved. This realization, in conjunction with new molecular-genetic techniques, has led to a surge in comparative neurodevelopmental research covering a wide variety of animal phyla over the past two decades. This chapter attempts to provide an overview of the diverse neural architectures that one encounters among invertebrate animals, and the developmental steps shaping these architectures.

Keywords: neuroanatomy, neuroectoderm, neural progenitor, neural patterning, neurogenetics, neural signaling, axonal pathfinding, neural connectivity

Neurons, structurally and functionally defined as cells with elongated processes that are specialized for the “repeated conduction of an excited state from receptor sites or other neurons to effectors or other neurons” (Bullock & Horridge, 1965, Vol. 1, p. 6), can be found in all metazoa, with the exception of placozoa and porifera. In the simplest (and presumably the primitive) state, neurons are more or less evenly distributed and form a “nerve net” associated with the body wall, as well as the intestinal tract and other inner organs (Fig. 1A). A nerve net can be found in cnidarians and ctenophores, as well as many bilaterian animals, including acoelomorphs, lophotrochozoa [platyhelminths (Fig. 1B), mollusks, bryozoans, chaethognaths], and deuterostomes (echinoderms, hemichordates; Bullock & Horridge, 1965; Grimmelikhuijzen & Westfall, 1995; Sakaguchi, Mizusina, & Kobayakawa, 1996; Mackie, 2004; Eichinger & Satterlie, 2014). In most of these cases, the nerve net is formed by sensory neurons, modified epithelial cells integrated into the epidermal epithelium, and ganglion cells, which lie underneath the epidermis (Fig. 1C). Similarly, ganglion cells and sensory cells form a nerve net in the intestinal wall of many animals.

Added to the nerve net are conglomerates of neurons that can take the shape of rounded (“ganglia”) or elongated structures (“nerve rings,” nerve cords). Such conglomerates are already present in many cnidarians, where a nerve ring surrounds the mouth opening or the base of tentacles, and ganglia of considerable size are connected to sensory organs, such as the rhopalia (Mackie, 2004; Skogh et al., 2006; Garm et al., 2007; Nakanishi et al., 2009). In most bilaterian taxa, sensory organs accumulate in the front part of the animal, and conglomerates of neurons that receive and process sensory input are clustered around these sensory structures, forming a brain. The brain, in addition to nerve cords extending throughout the body, forms the central nervous system (CNS; Fig. 1B, D). The nerve net and/or peripherally located sensory neurons, as well as nerve fibers commuting between the CNS and the periphery, represent the peripheral nervous system (PNS; Fig. 1B, D). Peripheral neurons are also associated with the gut and inner organs. This part of the PNS is called the “visceral,” “autonomic,” or “stomatogastric” nervous system (Reuter & Gustafsson, 1985; Csoknya et al., 1992; Hartenstein, 1997). In many animal taxa, neurons of the CNS and PNS are accompanied by a second class of cells, called glia (Hartline, 2011; Verkhratsky & Butt, 2013). Glial cells form sheath-like processes around neuronal cell bodies and nerve fibers, or extend branched processes that interact with terminal nerve fibers and synapses (Fig. 1E). They also play a pivotal role as phagocytes during development and in the mature nervous system.

Development of the Nervous System of InvertebratesClick to view larger

Figure 1 Structural elements of the invertebrate nervous system. (A) Line drawing of cnidarian polyp, showing multipolar neurons arranged in a basiepithelial nerve net. (B) Line drawing of nervous system of platyhelminth Bdelloura candida, illustrating peripheral nerve net, brain, and nerve cords. (C) Schematic cross-section of bodywall of cnidarian, depicting outer layer (epidermis), inner layer (gastrodermis), sensory neurons, and ganglion cells. (D) Drawing of nervous system of late Drosophila embryo (dorsolateral view, anterior to the left). Shown are brain and ventral nerve cord (central nervous system; red) and peripheral neurons (blue) innervating small sensory organs (sensilla). (E) Schematic cross section of ventral nerve cord of late Drosophila embryo, depicting ganglionic organization of central nervous system, with outer cell body cortex and inner neuropil, surrounded by glia. Neuropil is formed by neurites of interneurons and motor neurons (red), and axonal arborizations of peripheral sensory neurons (blue). (F1–F3) Schematic cross sections of central nervous system, illustrating the three configurations in which neurons and their processes are arranged in relationship to the bodywall. (Panels A and B from Hanström, 1968, with permission)

In regard to the structure and development of the nervous system, it is helpful to distinguish between three different configurations, basiepithelial, subepithelial, and invaginated, in which neurons are arranged (Fig. 1F). In a basiepithelial (also called basiepidermal) nervous system, neuronal cell bodies and nerve fiber bundles are arranged in one or more layers at the base of the epidermis, separated from the interior of the animal by a basement membrane (Fig. 1F1). In a subepithelial (subepidermal) nervous system, neurons have moved out of the epidermal layer and form ganglia or nerve chords within the body cavity, separated from the epidermis by a basement membrane. In these subepithelial ganglia and chords, neuronal cell bodies form an outer layer (cell body rind or cortex) that encloses a central mass of neuronal processes (axons, dendrites) and synapses, called the neuropil (Fig. 1E, F2). Invaginated nervous systems are also located within the body cavity, but they differ fundamentally from subepithelial nervous systems in regard to the arrangement of neuronal cell bodies and neuropil. At the core of an invaginated ganglion or cord is an epithelial vesicle or tube, whose apical surface faces an inner lumen. Neurons are arranged as layers surrounding the tube, and neuronal processes are directed further outward, forming additional layers around the cell body layers (Fig. 1F3). Basiepithelial and subepithelial nervous systems are distributed widely over the three major groups of bilaterians, the lophotrochozoa, ecdysozoa, and deuterostomia. Among the former, chaethognaths, gastrotrichs, and most lophophorates possess a basiepithelial nervous system; in platyhelminths, nemerteans, mollusks, and annelids, the nervous system is subepithelial. Among ecdysozoa, arthropod taxa all have subepithelial nervous systems, whereas most cycloneuralian groups (e.g., nematodes) possess basiepithelial nervous systems. Basiepithelial nerve nets and nerve chords also predominate in deuterostomes (echinoderms, hemichordates). Invaginated nervous systems are restricted to deuterostomes (urochordates, cephalochordates, vertebrates).

Synopsis of the Steps of Neural Development

Early Neurogenesis: Origin and Proliferation of Neural Progenitors

The nervous system is formed by neural progenitor cells that can be recognized by their structure and proliferation pattern, in addition to the expression of specific genes, in the early embryo. Neural progenitors are specified within the ectoderm that lines the outer surface of the embryo and, in some cases, the endoderm in the interior of the embryo. First, conserved signaling mechanisms and transcriptional regulators (“neural determinants”) define specific domains within the ectoderm (“neuroectoderm”) that have the potential to generate neural progenitors (Fig. 2A, A1). Detailed studies carried out in the few existing genetic model systems, in addition to first glances at gene expression and function in a wider range of different animal taxa, suggest that many of the genetic factors that are active in the neurectoderm to control the specification and proliferation of neural progenitors are highly conserved. At an initial stage, transcriptional regulators of the SoxB family are instrumental in defining populations of cells that will become neural progenitors. SoxB and other members of the Sox family are ubiquitously expressed in the neurectoderm of many animals (Sasai, 2001); other early transcriptional regulators, such as Six3/6, Nkx2.1/2.2, Pax6, and Pax3/7, are restricted to specific locations in the neurectoderm (e.g., Six3/6 to anterior location, Nkx2.1/2.2 to medial location; Arendt et al., 2016). These genes, which typically become active in parts of the ectoderm around gastrulation, trigger neurogenic potential, but at the same time inhibit neural differentiation, thereby maintaining the cells in which they are expressed in a proliferative state (Sasai, 2001; Bylund et al., 2003; Elkouris et al., 2011). Conserved signaling pathways, in particular Wnt/Wnt antagonists (Niehrs, 2010) and BMP/BMP antagonist, operate to set the expression pattern of SoxB and other early neural determinants. In many animals, from cnidarians to representatives of all bilaterian clades, Wnt signaling is activated around the blastopore/posterior pole, where it specifies the endomesoderm. Wnt also initiates a gradient of anti-neurogenic potential along the anteroposterior axis, which concentrates neurogenic potential closer to the anterior pole, where it is counteracted by Wnt antagonists (Darras et al., 2011; Marlow et al., 2013; Sinigaglia et al., 2013; Fig. 2A1). In bilaterian animals, a second gradient of neurogenic potential, oriented perpendicularly to that established by Wnt/Wnt antagonists, is established by the BMP signaling pathway (Angerer et al., 2011; Arendt et al., 2016); in parallel, these signaling pathways define the size, shape, and regional subdivision of the neurectoderm (Fig. 2A2).

Development of the Nervous System of InvertebratesClick to view larger

Figure 2 Steps in neural development. Panels of left column (A–F) show schematic sections of developing embryos in chronological order, illustrating the sequence of events by which the nervous system is assembled. Adjacent drawings (A1–F3) highlight pertinent aspects of developmental mechanisms, from the specification of the neuroectoderm (A1 and A2), selection of neural progenitors (B1 and B2), segregation and proliferation of neural progenitors (C1–C4, D1–D5), to aspects of axonal pathfinding (E1–E3) and target selection (F1–F3). (E1) This illustrates two pioneer neurons, N1 and N2. N2 expresses receptors for a repulsive signal (red) that forms a gradient from medial to lateral; as a result, the axon of N2 follows a pathway far from the midline. (E2 and E3) These show three groups of follower neurons (a, b, c) expressing differential affinities for, and fasciculating with, the “labeled” pioneer tracts (A, B, C). (F1 and F2) These show four neurons (blue, red, green, orange) with tiling dendrites. Arriving axon (bright red) forms exuberant projection which are selectively removed (“pruning”). (F3) This illustrates the concept of self-avoidance, where branches of the same neuron express “self-repulsive” membrane proteins (e.g., Dscam isoform); attractive membrane receptors and ligands allow for recognition of specific targets.

Following the establishment of the neurectoderm, members of the Achaete-Scute family (ASH) and Atonal family (ATO) of basic Helix-Loop-Helix (bHLH) transcription factors (“proneural genes”) are upregulated in specific groups of neurectodermal cells (proneural clusters), which subsequently initiate the developmental program of neural progenitors (Quan & Hassan, 2005; Powell & Jarman, 2008; Hartenstein & Wodarz, 2013; Huang et al., 2014; Fig. 2B, B1). In addition, proneural genes activate an inhibitory feed-back loop (“lateral inhibition”), mediated by the Notch signaling pathway, that spatially and temporally restricts the formation of neural progenitors (Kageyama et al., 2009; Hartenstein & Wodarz, 2013; Fig. 2B2). Proneural genes, in addition to the other early expressed transcription factors, also control the type of neuron generated at the neurectodermal location where these genes are expressed. In many bilaterians, these genes set up an orthogonal system of mediolateral and anteroposterior domains, each one giving rise to specific types of neurons (Fig. 2A2).

Neural progenitors appear within the neurectoderm as isolated, individual cells or clusters of contiguous cells Fig. 2C). In animals with basiepithelial nervous systems, neural progenitors remain within the surface neuroepithelium (Fig. 2C1). To give rise to a subepithelial nervous system, progenitors move into the interior of the embryo by a process of delamination, ingression, or invagination (Hartenstein & Stollewerk, 2015; Fig. 2C2–4). Invaginated nervous systems are formed when the entire neurectoderm, or large parts of it, are internalized and subsequently maintain their epithelial integrity as neural vesicles/tubes. The neuroepithelial cells develop into neural progenitors, producing neurons that assemble around the surface of the invaginated tube.

The central nervous system is rich in cell number (thousands to millions in most invertebrates), and to reach this cell number neural progenitors typically have to undergo an extended phase of proliferation (Fig. 2D). Proliferation can follow a pattern of symmetric divisions, followed by exit from the cell cycle and differentiation (Fig. 2D1); or a pattern of stem cell–like, asymmetric divisions where, with each division, a neural progenitor renews itself and generates a second daughter cell that then directly differentiates (Fig. 2D2) or acts as an “intermediate progenitor” (Hartenstein & Stollewerk, 2015; Fig. 2D3). Intermediate progenitors do not self-renew but perform a fixed number of divisions before they exit the cell cycle and differentiate (Noctor et al., 2007; Homem et al., 2015; Kang & Reichert, 2015). In some cases, notably insects where early neurogenesis has been investigated in detail for the genetic model system Drosophila, neural progenitors (“neuroblasts”) follow a stem cell–like pattern of asymmetric divisions that is highly invariant, resulting in so-called fixed lineages (Fig. 2D4, 5). In a fixed lineage, the later fate of a neuron or glial cell depends on its exact position within the family tree of the given progenitor. Neurons born during the first round of division may evolve a different structure and function than neurons born later rounds. These differential fate decisions depend to a large part on intrinsic determinants, that is, genetic factors activated in the progenitor in a fixed temporal order and then handed down to the progeny in that same order (Pearson & Doe, 2004; Kao & Lee, 2010; Li et al., 2013; Fig. 2D4, 5). Alternatively, extrinsic factors, signals expressed in cells outside the progenitor, act on the progeny born during a certain time interval and set their fate.

Axon Pathway Formation

Structural differentiation of neurons begins with the extension of elongated processes (neurites) called axons and dendrites. The term “axon,” as opposed to “dendrite,” refers to the neurite conducting impulses away from the cell body; dendrites carry activity toward the cell body. This distinction, generally clear-cut in vertebrate neurons, does not exist for most invertebrate neurons, where a single neurite extends from the cell body into the neuropil, where it splits into multiple dendritic and axonal branches. In arthropods, nematodes, and annelids, where neurite pathway formation has been studied most actively, neurons of the CNS typically emit a single neurite, for which the term “axon” is most commonly used. Axons assemble into bundles (nerve tracts or fascicles in the CNS; nerves in the PNS; Fig. 2E). Axons follow invariant, genetically determined pathways to reach their target. In most systems analyzed so far, neurons do not differentiate simultaneously. Typically, a small subset of neurons located at strategic positions within the nervous system differentiates at an early stage. Neurites of these cells (“pioneer neurons”) lay down a network of connections that form a more or less complete “blueprint” of the nerve tracts of the mature nervous system (Bate, 1976; Singer et al., 1979; Bentley & Keshishian, 1982; Goodman & Doe, 1993; Raper & Mason, 2010; Fig. 2E). Axons of neurons that are born and differentiate at later time points (“follower neurons”) grow along the preexisting pioneer tracts. The growth of pioneer axons is directed by cues presented by the neuroepithelium and the epidermis, as well as cell bodies of neurons and glia. Follower axons, on the other hand, simply have to select, among the pioneer tracts they encounter, the proper one and then grow on the membranes of this tract to reach their preordained target.

Signals from the microenvironment surrounding the extending axon control its trajectory. These signals can act as attractants, prompting axon growth toward the signaling source, or as repellants, making the axon avoid the source (Araujo & Tear, 2003; Evans, 2016; Fig. 2E1). Signals can be membrane-bound molecules that require close contact between the growing axon and the source, or they could act from a distance as diffusible factors. In some cases, specific cells (“guidepost cells”; Palka et al., 1992; Fig. 2E1), which could be other neurons or glial cells, have been identified as the principal source of guidance cues for particular axons.

The molecular machinery that receives the guidance cues and controls where an axon grows rests in the highly specialized tip of the elongating axon, called the growth cone (Gallo & Letourneau, 2004; Geraldo & Gordon-Weeks, 2009; Gomez & Letourneau, 2014). The growth cone has multiple fine, microfilament-filled processes called filopodia. Filopodia are short-lived structures that are constantly sent out and then quickly withdrawn. The growth cone with its filopodia has been compared to a probe that samples its environment, recognizes distinct cues, and moves accordingly, pulling the rest of the axon behind it. Guidance cues and their receptors, as well as signal transducers that control growth cone motility, form part of signaling pathways that are also highly conserved among the model organisms (including vertebrate and invertebrate) where they have been studied. Filopodial probing movement is driven by actin polymerization/ depolymerization and the dynamic formation of microfilament/ membrane attachments. A group of widely conserved actin-bound GTPases, forming the Rac/Rho family of proteins, are instrumental in orchestrating directed filopodial movement. Rac/Rho kinases are activated by external signals via receptor tyrosine kinase (RTK) pathways.

Conceptually, it makes sense to distinguish between two types of signaling mechanisms. First, there are localized adhesion (“recognition”) molecules expressed in highly specific patterns by axons and their guidance structures/targets which ensure that only the proper contacts or connections are formed. This concept (the “labeled pathway hypothesis”; Goodman et al., 1983; Grenningloh & Goodman, 1992; Fig. 2E2, 3) addresses scenarios where, for example, different neurons (“A,” “B,” “C”) encounter separate pioneer tracts or targets (“a,” “b,” “c”). If the pairs A and a, B and b, and C and c express different (homophilically interacting) adhesion molecules, they would form exclusive contacts; A would only interact with a, B with b, and C with c. Secondly, there exist more globally acting, possibly diffusible signals that are distributed in mediolateral and dorsoventral gradients (Fig. 2E1). These signals create a “Cartesian coordinate system” of positional information in which axons find their adequate position and target.

Most of the signaling systems characterized in recent studies belong to this second category. Among them are the highly conserved Ephrin, Semaphorin, Netrin, and Slit/Robo pathways (Bashaw & Klein, 2010; Evans, 2016). Ephrins are either secreted or membrane-bound ligands which act as repellants on axons. There is good evidence that ephrin-mediated ras activation in axons has a direct effect on the growth cone cytoskeleton, causing growth cone collapse (Gallo & Letourneau, 2004). Another family of guidance cues that act as repellants are the semaphorins and their receptors (plexins), membrane-bound proteins characterized by a long conserved extracellular domain (Negishi et al., 2005). The netrin and slit/robo system controls the orientation of extending axons relative to the midline, determining which axons cross, and at what distance to the midline a given axon grows. Netrins are bifunctional (attractive and repulsive) signals expressed in neurons and/or other cell types along the CNS midline. Depending on the type of netrin receptor it expresses, a given neuron turns toward or away from the midline (Round & Stein, 2007; Ogura et al., 2012). Slit is also expressed is by cells along the midline of the animal; Slit receptors, Ig-like proteins of the Roundabout (Robo) subfamily, are found in populations of axons that are prevented from approaching and crossing the midline (Dickson & Gilestro, 2006).

Neuronal Connectivity

Neurite bundles form a framework that defines the overall structure of the nervous system. Where neurites come together, synaptic connections can form. However, out of the large number of connections that are possible based on neurite contacts, only a small fraction actually forms. In other words, when reaching its “target zone” and contacting numerous potential synaptic targets, a growth cone has to select a small subset of these targets (Fig. 2F). Again, specific molecular interaction between growth cones and their postsynaptic targets is essential to order the highly invariant pattern of connectivity. In part, the same molecular factors that already guided axons on their way remain active to control axon–target connections. Many other neuronal “recognition systems” are of importance, most of which remain to be characterized. A number of general principles that govern the phase of axon–target interactions in invertebrates and vertebrates alike could be learned from studies in the model systems.

Neurons generally do not form synaptic contacts with themselves (“self-avoidance”; Kise & Schmucker, 2013; Zipursky & Grueber, 2013; Fig. 2F3). In Drosophila, the Dscam1 locus possesses several variable exons which allow for the formation of many thousands of different isoforms of the Dscam protein. Highly diversified expression of these isoforms plays a role in preventing synaptic contacts among branches of the same neuron (Hattori et al., 2008). More generally, neurons of a given class innervating a common volume of the neuropil, or area of the body wall, often avoid each other, with the result that arborizations of these neurons form nonoverlapping “tiles” (Grueber & Sagasti, 2010; Lefebvre et al., 2015; Fig. 2F2). Tiling behavior, mediated by Dscam, Semaphorins, or other signaling systems, plays an important role in the visual neuropils and sensory receptors in flies (see later discussion).

In many cases where the establishment of neuronal connectivity was analyzed, two phases were distinguished. During an early phase, a neuron typically forms widespread, exuberant connections with groups of neurons (or other targets) that include the ultimate target, but also additional cells to which connections are later lost (Fig. 2F1). During the early phase, which is independent of neural activity, an “unfocused” or “unrefined” pattern of connections is formed. Subsequently, in a second phase that requires neural activity, the projection pattern is refined by selectively removing exuberant connections (“pruning”; Low & Cheng, 2006; Fig. 2F2). Examples of exuberant projections followed by pruning are best studied in vertebrate models but are also common in invertebrates (e.g., Murphey, 1986; Low & Cheng, 2006; Riccomagno & Kolodkin, 2015).

Cnidaria and Ctenophora

Cnidaria represent a large phylum that consist of two sister clades, Anthozoa (sea anemones and corals) and Medusozoa (jellyfishes), which are further divided into Staurozoa, Scyphozoa, Cubozoa, and Hydrozoa (Collins et al., 2006). Cnidarian embryos develop into a worm-shaped larva, called planula, that possess a single axis with an anterior (aboral) pole and a posterior (oral) pole. Neurons form a basiepithelial nerve net underlying the epidermis and, in some species, the gastrodermis. Neurons are concentrated closer to the anterior pole, and in some species a distinct agglomeration of presumed sensory neurons at the anterior tip has been distinguished as “apical organ” (Chia & Koss, 1979; Martin, 1992; Gröger & Schmid, 2001; Nakanishi et al., 2008; Piraino et al., 2011; Nakanishi et al., 2012; Fig. 3A). After larval settling, animals undergo metamorphosis, which involves a reversal of the body axis. The oral pole becomes the “head” of the developing polyp. The head forms tentacles, which are innervated by a new nervous system that includes a nerve net and a nerve ring surrounding the mouth and tentacle bases (Nakanishi et al., 2008, 2009; Galliot et al., 2009; Fig. 3B). In many cnidaria, the polyp stage is followed by a medusa stage. Medusae also possess a diffuse nerve net, in addition to neuronal concentrations (“ganglia”) associated with the rhopalia (receptor complexes for light and gravity found in Scyphozoa and Cubozoa) or a nerve ring surrounding the umbrella (many Hydrozoans).

Development of the Nervous System of InvertebratesClick to view larger

Figure 3 Essential characteristics of nervous system architecture in Cnidaria (A: planula larval stage; B: polyp) and Acoela (C), represented as schematic sagittal sections. For explanation of graphic elements see key at upper right of figure. (D–L) Key events of early neurogenesis of cnidarians (D–F: Nematostella vectensis; G–I: Hydra vulgaris) and Acoela (J–L). The upper panel of each set (D, G, J) shows a schematic sagittal section of an early embryo at the onset of neurogenesis. Territories or cell populations with neurogenic potential are rendered in blue. The middle part of each set (E, H, K) shows a schematic cross section of ectoderm at a stage when neural progenitors appear and proliferate; panels at bottom (F, I, L) schematically depict organization of nervous system at late embryonic or larval stages. For explanation of graphic symbols, see key at top right. (M–O) Horizontal confocal sections of planula larva of cnidarian Aurelia sp. (M), juvenile of nemertodermatid Meara stichopi (N; from Borve & Hejnol, 2014; with permission), and juvenile of acoel Symsagittifera roscoffensis (O). Subsets of neurons are labeled by antibody against peptide FMRFamide (M and N) or tyrosinated tubulin (O). Scale bar: 20 μm.

In the anthozoan Nematostella vectensis, scattered cells or small cell clusters expressing neural markers appear already in the early embryo in both the ectoderm and endoderm (Fig. 3D, E). These neural progenitors generate the three main neural lineages, intraepithelial sensory neurons, basiepithelial neurons (ganglion cells), and nematocytes (stinging cells; Nakanishi et al., 2012; Rentzsch et al., 2017). Experimental studies confirmed that the canonical “proneural cassette” (Sox, bHLH, Notch) is active in the embryo and planula to control the number and distribution of neurons in Nematostella (Layden et al., 2012; Richards & Rentzsch, 2014, 2015; Rentzsch et al., 2017). In scyphozoans (e.g., the moon jelly Aurelia sp.), neurons expressing differentiation markers such as tyrosinated tubulin or peptide transmitters (e.g., RFamide, GLWamide) were first detected in the anterior planula ectoderm, where they form bipolar epithelial cells with basally elongating growth cones. Growth cones split and form T-shaped axons directed anteriorly and posterior along the longitudinal axis of the planula (Nakanishi et al., 2008; Fig. 3F, M). In the late planula, neurites lengthen, and circular fibers are added to the longitudinal ones, completing an orthogonal nerve net. Neurons are continuously added from within the ectoderm. At the anterior pole, neurites of epithelial RFamide-ir and GLWamide-ir neurons form a dense apical nerve plexus, similar to the apical organ observed in larvae of many bilaterian animals (Nakanishi et al., 2008; Piraino et al., 2011; Fig. 3M). Different types of neurons in the apical organ and elsewhere in the planula show specific patterns of distribution and neurite projections (Piraino et al., 2011).

After larval settling, which occurs at its anterior tip, the planula metamorphoses into a polyp. The ectoderm and endoderm around the mouth opening grow into the oral disk and the anlagen of tentacles. These give rise to neurons in a manner similar to that observed in the planula (Nakanishi et al., 2008). Most neurons are epithelial (sensory) cells with T-shaped axons that extend along the longitudinal axis of the tentacles or form circular fibers surrounding the mouth opening (Fig. 3B). In the final step toward the formation of the medusa, the body column of the polyp lengthens and splits into segments, with each segment giving rise to an immature medusa (ephyra). The ephyra regrows tentacles and complex sensory organs (rhopalia) located at the tentacle bases. The formation of the neuronal ganglia associated with the rhopalia is accompanied by the expression of gene cassettes (e.g., Otx, Pit1, Brn1; Nakanishi et al., 2010) that also play a role in early neurogenesis of the bilaterian CNS.

In some Hydrozoans (e.g., the freshwater polyp Hydra vulgaris), early neurogenesis follows a mechanism different from the one described earlier. Rather than segregating from the ectoderm, neural cells are formed by interstitial cells, a population of proliferative stem cell that arises in the embryonic endoderm and comes to lie between the ectoderm and endoderm of the body column (Fig. 3G). Interstitial cells proliferate throughout the lifetime of the polyp in an asymmetric, stem cell–like mode (Bode, 1996; David, 2012; Gold & Jacobs, 2013). Interstitial cells are not “dedicated” neural progenitors; aside from neurons, they also give rise to secretory cell types and to germ line cells. The neuronal precursors and committed progenitors derived from interstitial cells migrate throughout the body and into the tentacles, where they insert into the epidermis as sensory neurons and nematocytes, or differentiate as basiepithelial ganglion cells (Fig. 3H, I). Despite this apparently different manner of generating a nervous system, the underlying genetic mechanism (i.e., proneural cassette) appears to be similar to the one seen in other cnidarians and in bilaterians in general (Grens et al., 1995; Käsbauer et al., 2007).

Ctenophora (comb jellies) used to be linked with Cnidaria as Coelenterata. Similar to cnidarians, ctenophores have a basiepithelial nerve net (also called subepithelial or polygonal nerve net; Hernandez-Nicaise, 1991; Jager et al., 2011), formed by sensory neurons and ganglion cells. Condensations of nerve cells occur at the aboral pole (apical organ, surrounding statocyst) and along the longitudinal swimming plates (comb rows), rows of epidermal cells bearing specialized cilia whose beat propels the animal forward. According to several recent studies, ctenophores represent an independent phylum that branched off at the very base of the Metazoa. Since placozoa and porifera (sponges), positioned in between ctenophores and cnidarians based on molecular studies, lack a nervous system, ctenophores might have evolved a nervous system independently of cnidarians and bilaterians (Ryan et al., 2013; Moroz et al., 2014; Moroz, 2014). This interpretation is supported by the finding that many genes controlling the development and function of neurons, including members of the proneural cassette, or neurotransmitters, are either absent or strongly modified. Very little is known concerning the development of the ctenophore nervous system. In a recent study utilizing global markers for muscle and nerve development (Norekian & Moroz, 2016), the origin of muscle fibers and a subset of ganglion cells forming the polygonal nerve net was documented. Surprisingly, neurons differentiated much later than muscle, which sets ctenophores apart from cnidarians, where both muscle fibers and neurites appear concomitantly.

Xenacoelomorpha

The clade Xenacoelomorpha, which includes the Acoelomorpha (Acoela and Nemertodermatida) and the Xenoturbellida, is currently considered as the sister group to the remaining Bilateria (Hejnol et al., 2009; Srivastava et al., 2014; Cannon et al., 2016). Xenacoelomorphs are simple worms found almost exclusively in marine environments. The nervous system consists of a basiepithelial nerve net, in addition to local condensations of sensory and centralized neurons. In nemertodermatids and acoels, a “frontal organ” or “frontal glandular complex” has been described, consisting of the external openings of gland cells surrounded by bundles of sensory dendrites (Klauser et al., 1986; Smith & Tyler, 1986; Ehlers, 1992). Acoels have a more or less compact brain, where cell bodies of sensory neurons of the frontal organ, as well as ganglion cells, assemble around a central neuropil (for Neochildia fusca and Symsagittifera roscoffensis; Ramachandra et al., 2002; Bery et al., 2010; Semmler et al., 2010; Martinez et al., 2017; Fig. 3C). The acoel brain includes a statocyst and, in many species, simple eyes. Bilaterally symmetric nerve cords, typically including a dorsal, lateral, and ventral pair, extend backward from the brain throughout the body. In the brain and nerve cords, cell bodies and processes of neurons are intermingled with those of other cell types (muscle, gland). A basement membrane or glial sheath separating tissue layers in other bilaterian groups is absent.

Neural development has been followed in an acoel (Neochildia fusca: Ramachandra et al., 2002; Bailly et al., 2013) and a nemertodermatid (Meara stichopi; Børve & Hejnol, 2014), but no specific, early expressed molecular probes have been used so far, and the genetic mechanism underlying early neurogenesis is unknown. Following cleavage, the acoelomorph embryo forms a mass of cells, the “embryonic primordium,” in which various cell types differentiate “in situ,” without overt prior migration or germ layer formation akin to gastrulation or neurulation in other bilaterians. Cells at the surface of the embryonic primordium become the epidermis; cells into the center differentiate into neurons, muscle cells, and gland cells (Fig. 3J–L). Differentiating nerve cells, labeled by global markers such as anti-tyrosinated tubulin, extend neurites bundles that coalesce into the longitudinal and commissural tracts of the brain and nerve cords (Fig. 3N, O). Intermingled with these are muscle cells whose processes assemble into the regular, layered pattern of longitudinal, oblique, and circular muscles.

Inner cells that do not differentiate and maintain their proliferative activity throughout late embryogenesis and adult life form a conspicuous, pluripotent stem cell population that might be compared to the interstitial cells of hydrozoans described in the previous section. Among bilaterians, these stem cells, called neoblasts, were only described in the Acoela (Gschwentner et al., 2001; de Mulder et al., 2009) and in platyhelminths (Ladurner et al., 2000; Baguñà, 2012). Neoblasts are motile cells that spread out throughout the body and align themselves with the epidermis and neuromuscular system, constantly adding to or replacing differentiated cell types. Neoblasts are also responsible for the high regenerative capacity that has been observed in acoela and platyhelminths (Saló & Baguñà, 1985; Reddien, 2013; Rink, 2013; Sprecher et al., 2015).

Lophotrochozoa

According to our current understanding of metazoan phylogeny, the tree of bilaterian animals beyond the Xenacoelomorpha has the three branches Lophotrochozoa, Ecdysozoa, and Deuterostomia. Most bilaterian phyla have found a stable place on one of these three branches. Included in the Lophotrochozoa are two major polyphyletic groups, the Spiralia and Lophophorata, in addition to several other phyla, some of which (e.g., Rotifera, Gnathostomulida, Micrognathozoa) are folded into a taxon called Gnathifera (Erwin et al., 2011; Laumer et al., 2015; Kocot et al., 2017). Gnathifera and platyhelminths (flatworms, considered to be spiralians) branch at the base of the lophotrochozoan tree, followed by the remaining large polyphyla of lophophorates (entoprocta, ectoprocta, phoronida, brachiopoda) and spiralians (nemertea [ribbon worms], annelids, mollusks). Lophophorates are mostly sessile, worm-shaped animals that carry at their anterior end a complex array of tentacles used for filter feeding, called the lophophore. Spiralians share the spiral mode of cleavage. Most lophophorates and spiralians also form larvae, called trochophore larvae, that have many structural characters in common.

With the exception of some representatives of the platyhelminths and annelids, lophotrochozoa have not been investigated at great length in regard to the developmental processes that shape the structure of their nervous systems. On the other hand, many studies carried out during the past two decades have started to shed light on the neural architecture of lophotrochozoan larvae and were used to draw inferences about the evolution of the invertebrate nervous system. I will in the following provide a brief overview of the diverse nervous systems found among the different lophotrochozoan clades. In the second section of this subchapter, nervous system development of the platyhelminths, an early branch of the lophotrochozoan tree, will be discussed. The third section summarizes important aspects of lophotrochozoan larval neuroanatomy and neurodevelopment. The last section will take a look at neural development in two different clades of annelids, the polychaetes and leeches.

Neural Architecture in Lophotrochozoa

Platyhelminthes

Platyhelminthes are a large and diverse group of free-living or parasitic worms. The flatworm nervous system—and the body structure in general—resembles that of the acoela. Flatworms have a ciliated epidermis innervated by a basiepithelial nerve net; in addition, they possess a subepithelial brain and bilaterially symmetric sets of nerve cords extending along the dorsal, lateral, and ventral side of the body (Bullock & Horridge, 1965; Ehlers, 1985; Reuter & Gustafsson, 1985; Hartenstein, 2016; Fig. 4A, B). Circular commissures interconnect the longitudinal cords at more or less regular intervals; commissures and cords form a conspicuous orthogonal scaffold around the body, called the orthogon. Ciliated sensory neurons are integrated in the epidermis or form part of the brain (Ehlers, 1985; Rieger et al., 1991). As in acoels, a high concentration of sensory cells located in the brain extends dendrites to the anterior tip of the animal, where they accompany an array of glandular cells (frontal glandular complex; Ehlers, 1992). Flatworms also have simple eyes closely associated to the brain. Neuromuscular contacts are made by muscle processes projecting toward the brain and nerve cords, rather than the other way round. This condition, unusual from the perspective of the vertebrate nervous system where motor neurons have long peripheral processes toward muscles, is not only found in platyhelminthes but in many other invertebrate phyla as well.

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Figure 4 Essential characteristics of nervous system architecture in representative phyla of lophotrochozoa, shown as schematic sagittal sections. (A and B) Platyhelminthes (flat worms); (C and D) Gastrotricha (hairybacks); (E and F) Bryozoa (moss animals); (G and H) Mollusca; (I and J) Annelida; (K and L) Chaetognatha (arrow worms).

Gnathifera and Gastrotricha

The assembly of small phyla known as gnathifera (Gnathostomulida, Micrognathozoa, Rotifera, Acanthocephala) includes mostly microscopic, marine worms which possess cuticular specializations (“jaws”) associated with their pharynx or foregut. Another basally branching clade of lophotrochozoans with affinity to the gnathiferans is the gastrotrichs. All of these animals differ from acoelomorphs and platyhelminths in having a through-gut with an anterior mouth and posteror anus, and lacking a ciliated epidermis. Gnathifera contain very small numbers of neurons (tens to a few hundreds). Clusters of neurons in the head form a subepithelial or basiepithelial brain that surrounds the mouth. In gnathostomulids and micrognathozoans, neurons of the trunk typically form several paired cords (Schmidt-Rhaesa, 2016; Bekkouche & Worsaae, 2016). Gastrotrichs also possess multiple longitudinal nerve cords (Hochberg & Litvaitis, 2003; Schmidt-Rhaesa & Rothe, 2016; Fig. 4C, D). Trunk neurons are clustered in several ganglia in rotifers and acanthocephalans (Clement & Wurdak, 1991; Hochberg, 2009, 2016; Semmler Le, 2016).

Lophophorates

Four phyla, including Phoronida (horseshoe worms), Ectoprocta (Bryozoa: mossy animals), Brachiopoda (lamp shells), and Entoprocta (Kamptozoa: goblet worms), have a circular array of ciliated tentacles (lophophore) surrounding the mouth, and a U-shaped gut. However, despite these similarities in structure and lifestyle (sessile filter feeders), lophophorates are a polyphyletic assembly (Dunn et al., 2008; Hejnol et al., 2009; Erwin et al., 2011; Kocot et al., 2017). In lophophorates, the majority of neurons are grouped around the mouth and in the tenctacles. Phoronids have a basiepithelial nerve net that is condensed into a basiepithelial nerve ring at the base of the lophophore, and a longitudinal tract toward the posterior of the body column (Hanström, 1968; Santagata, 2002; Temereva & Tsitrin, 2014). Bryozoa also possess a peripheral nerve net, a paired cerebral ganglion (brain) located close to the mouth, and bilateral nerve tracts projecting out of the brain around the pharynx. Anteriorly, the brain extends two long cords (“ganglionic horns”) toward the lophophore (Hanström, 1968; Schwaha & Wanninger, 2012; Gruhl & Schwaha, 2016; Fig. 4E, F). The ganglionic horns emit many regularly spaced radial nerves projecting into the tentacles of the lophophore. The brain of bryozoans appears to develop from an invagination of the ectoderm. Thus, neurons are arranged around the wall of a vesicle with an inner lumen (Gruhl & Bartolomaeus, 2008; Weber et al., 2014). These are the so far only observations supporting the idea that invagination of (part of) the neurectoderm forms part of the repertoire of morphogenetic mechanisms involved in early neurogenesis in Lophotrochozoa. The nervous system of brachiopods and entoprocts is formed by subepidermal ganglia. In brachiopods, a paired cerebral ganglion and subesophageal ganglion form a ring around the pharynx. Several nerves extend from these ganglia toward the digestive tract, the tentacles, and stalk (Hanström, 1968; Santagata, 2011; Lüter, 2016). Entoprocts possess only one dumbell-shaped ganglion located close to the pharynx (Hanström, 1968; Schwaha et al., 2010) from which nerves radiate into the tentacles of the lophophore.

Nemertea

Nemerteans (or nemertines; “ribbon worms”) are errant marine worms that are characterized by a retractible proboscis used for prey capture. The brain of nemerteans consists of a paired supraesophageal and subesophageal ganglion. The trunk is innervated by a pair of ventral cords and two pairs of smaller lateral and dorsal cords; circular commissures interconnect the cords. The nemertean nervous system, similar to that of annelids and mollusks reviewed later in this chapter, possesses layers of glial cell process that surround the brain and the nerve cords at its outer surface, and also separate the cortex from neuropil (Hanström, 1968; Beckers et al., 2011, 2013; Beckers & von Döhren, 2016). One also observes in nemerteans a considerable increase in volume and complexity of the brain neuropil, which may reflect the existence of complex sensory organs located in the head (frontal organ, cerebral organ, eyes).

Annelida

Annelids are segmented worms and include many diverse clades of motile and sessile forms, among them the polyphyletic polychaetes, the echiurids, sipunculids, pogonophores, and clitellates (which include the terrestrial earthworms and the leeches). The central nervous system of annelids is formed by a subepithelial brain located in front of the mouth (the prostomium), and metamerically arranged ganglia located along the ventral side of the body (ventral nerve cord; Bullock & Horridge, 1965; Hanström, 1968; Fig. 4G, H). The gut is innervated by the stomatogastric nervous system. Many annelids, notably polychaetes, have large, complex sensory organs, including the palps, antennae (tentacles), eyes, and nuchal organs (Bullock & Horridge, 1965; Mill, 1978; Golding, 1992; Purschke, 2016; Fig. 4G). Accordingly, the brain and ventral ganglia have a large, complex neuropil accompanied by glial layers. The brain is compartmentalized into discrete domains (forebrain or procerebrum; midbrain or mesocerebrum; and hindbrain or metacerebrum) and possesses structured compartments, such as the central optic neuropil and the mushroom body (Hanström, 1968; Heuer & Loesel, 2008).

Mollusca

Mollusks includes clades of worm-like animals lacking a shell (aplacophorans), groups with simple, plate-like shells (monoplacophora, polyplacophora), and the more derived clades with complex shell structure (gastropods, bivalves, cephalopods, scaphopods). The aplacophorans and mono/polyplacophorans (Fig. 4J), by many considered basal among the mollusks, have a neural architecture resembling that of Platyhelminthes, with a basiepithelial nerve net, a subepithelial brain surrounding the mouth (“cerebro-buccal ring”), and two paired nerve cords, called the lateral pallio-visceral cord and the ventral pedal cord (Moroz et al., 1994; Shigeno et al., 2007; Sigwart & Sumner-Rooney, 2016). Two paired nerves extend from the brain to the foregut (esophageal ganglion) and muscles of the radula (the toothed “tongue” used to shred ingested food). In the more derived mollusk clades (gastropods, bivalves, and cephalopods (squid and octopus), the nerve cords are condensed into several compact ganglia; in addition, a peripheral diffuse nerve net is still present (Hanström, 1968; Wanninger, 2016; Voronezhskaya & Croll, 2016). Gastropods and bivalves have a brain (cerebral ganglion) that emits two paired of nerve cords, or connectives (Fig. 4I). The ventral (pedal) connective projects to the pedal ganglion, which is associated with the muscular “foot” of gastropods and bivalves. The lateral, pallio-visceral cord innervates three paired ganglia, the pleural ganglia (associated with the mantle cavity), branchial ganglia (gills), and visceral ganglia (intestinal tract). The stomatogastric nervous system consists of tracts connecting the brain to ganglia innervating the pharynx and foregut (buccal ganglia). In cephalopods, the ganglia of the head and trunk are fused into one large “brain” that surpasses many vertebrate brains in terms of cell number and complexity. In addition, intrabrachial ganglia, connected to the pedal ganglion, control movement of the arms (Wollesen, 2016). Prominent optic lobes with millions of neurons have evolved in conjunction with the large image-forming eyes. As mentioned for the annelids, the brain of mollusks has structured neuropil compartments stabilized by glial layers (Hanström, 1968; Elekes, 2000; Wollesen, 2016).

Chaetognatha

Chaetognaths (arrow worms) are predatory worm-like animals found in the plankton. They are usually placed at the base of the lophotrochozan tree, but their position is uncertain. Chaetognaths have a basiepithelial nervous system, with a supraesophageal brain and vestibular ganglion, connected to a large ventral ganglion (Fig. 4K, L). These ganglia can be seen as local condensations of a diffuse basiepithelial nerve net (Rehkämper & Welsch, 1985; Goto et al., 1992; Harzsch & Müller, 2007). Recent studies (Rieger et al., 2011; Perez et al., 2013) have also documented the pattern of neural progenitors, which appear as a highly ordered array at the apical surface of a ventral neurectoderm; postmitotic neuronal precursors are given off to the basal side, similar to what has been demonstrated for annelids.

Neural Development in Platyhelminthes

Most flatworms are direct developers in which embryonic development produces hatchlings (juveniles) that closely resemble the adult in body structure. As described for the Acoela, the primordium of the brain becomes visible at around 50% of embryonic development within an unstructured mass of cells that forms without the gastrulation movements characteristic of other bilaterian animals. The defining steps of early neurogenesis, including specification and proliferation of neural progenitors and their genetic control mechanisms, have not been elucidated yet. Structurally, the brain primordium is shaped as a bilobed condensation of several hundred cell bodies in the interior of the anterior half of the embryo (Younossi-Hartenstein et al., 2000, 2001; Younossi-Hartenstein & Hartenstein, 2000a, 2000b; Hartenstein & Jones, 2003; Morris et al., 2004, 2007; Cardona et al., 2005; Monjo & Romero, 2015). It is not known whether the neurons forming the nerve cords, or the peripheral nerve plexus, migrate out of this anterior mass, or originate at other (more posterior) positions. At around 60% of development, differentiating nerve cells emit nerve fibers into the center of the brain primordium where they form a small commissural neuropil; long axons leave the neuropil and form the nerve cords (Fig. 5A, B). In the species Mesostoma lingua (a member of the derived clade Rhabdocoela) and Macrostomum lignano (a more basally branching flatworm), elongating axons assemble into a ventral and a dorsal nerve cord. The thick ventral cords connect the brain with the nerve ring that surrounds the pharynx. Nerve cords typically grow along strands of differentiating muscle fibers, which suggests that reciprocal interactions between neurons and myoblasts play an important role in patterning the neuromuscular system (Reiter et al., 1996; Younossi-Hartenstein et al., 2000; Fig. 5C2). Furthermore, the phenomenon of early differentiating pioneer neurons that lay down a blueprint of the nervous system at an early stage (see earlier) has been documented for Mesostoma (Younossi-Hartenstein et al., 2000). Here, three paired clusters of 1–3 neurons each, labeled by the antibody against acetylated tubulin from 60% of development onward, pioneer the longitudinal cords (Fig. 5C1–3); two clusters of neurons that appear slightly later than the cord pioneers project across the midline and pioneer the commissural neuropil of the brain. These cells are also in close contact with the eyes, which make their appearance at around 75% of development. At this stage, synapses become ultrastructurally apparent in the neuropil and peripheral plexus, and muscle-based movement of the embryo begins.

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Figure 5 Neural development in Platyhelminthes. (A and B) Photographs of late-stage embryo of polyclad Imogine mcgrathi (A) and Macrostomum lignano (B) labeled with basic fuchsin (nuclei; red). Both panels represent lateral view, anterior to the left, dorsal up. The cortex and neuropil of the brain are in focus; also visible are the ventral nerve cord, muscle, and epidermis. (C1–C3) Schematic depiction of neural development in the rhabdocoel flatworm Mesostoma lingua. All panels show ventral view of embryo (C1: stage 5; C2: stage 6; C3: stage 7; staging after Younossi-Hartenstein et al., 2000). During stage 5, three clusters of acTub-positive pioneer neurons form in the anterior brain cortex, surrounding the incipient commissural neuropil. Clusters dn1 and dn2 pioneer the dorsal nerve cord; vn, the ventral nerve cord. AcTub-positive neurons also start to appear in the pharynx. Additional pioneer clusters (umn, pco) are added during stage 6 (C2 and C3); axons extend into the brain commissure and nerve cords, where they follow longitudinal arrays of muscle precursors. (D) Contribution of neoblasts to embryonic and postembryonic growth. Schematic sagittal sections of flatworm embryo (based on analysis of macrostomid Macrostomum lignano; Morris et al., 2007). At the early stage (upper left), inner mass of proliferating cells generates the primordia of the brain, muscle, and pharynx. Cells of these primordia become postmitotic at midstage of embryogenesis (bottom left). The only cells that do not differentiate and maintain proliferation are the progenitors of neoblasts. These cells contribute additional cells to all organ primordia during late embryogenesis and postembryonically. (E and F) Horizontal sections of anterior part of juveniles of Macrostomum lignano. In (E) the base analog 5-Ethyl-2′-Deoxyuridine (EdU) was applied over a period of 2 hours, after which animals were fixed. EdU-positive neoblasts appear as deeply located, isolated cells posterior of the brain. In (F), the EdU pulse was chased over a 7-day period; EdU-positive nuclei are now found in cells of the epidermis, muscle, and brain, derived from neoblasts. (G) Development of the photoreceptors and their projection in the planarian flatworm Dugesia japonica 3 days and 7 days after brain ablation (from Yamamoto & Agata, 2011, with permission). Photoreceptors were labeled with anti-Arrestin (green). (H) Expression of Slit (magenta) and Netrin (green) in midline territory of regenerating brain 2 days (top) and 3 days (bottom) after ablation. (From Yamamoto & Agata, 2011, with permission)

Neoblasts, introduced in a previous section for the acoels, form a population of proliferating, pluripotent stem cells that become prominent in the later half of embryonic development and persist throughout adulthood. Neoblasts continuously add to the number of cells; pulse-chase experiments with the proliferation marker Bromo-deoxyuridine (BrdU) in developing flatworms showed that, within days, BrdU-positive cells, initially part of the neoblast pool, become incorporated into the brain as differentiated neurons (Nimeth et al., 2004; Fig. 5D–F). Likewise, neoblasts contribute to the late embryonic and postembryonic growth of all other organ systems (Fig. 5D–F). Species of the clade Tricladida (Planaria; Dugesia japonica, Schmidtea mediterranea, Schmidtea polychroa) have become widely used as model systems to study neoblasts and regeneration. Studies in these triclads have also provided the reagents to investigate the molecular mechanisms that control neural development in Platyhelminthes (Hill & Petersen, 2015; Davies et al., 2017). Inventories of gene expression reveal the same signaling pathways and transcriptional regulators that were previously characterized for different genetic model systems. For example, the planarian Slit homolog is expressed along the midline of the nervous brain primordium (Fig. 5G, H). Slit RNAi knockdown experiments demonstrated that normally bilateral structures, such as the eyes, collapse in a single cluster in the midline (Cebrià & Newmark, 2005; Yamamoto & Agata, 2011). Likewise, knockdown of a Robo receptor resulted in path-finding defects of visual axons that normally cross the midline.

One class of flatworms, Polycladida, shows spiral cleavage and forms a type of larva resembling the ciliated primary larvae characteristic of other spiralians, and lophophorates, discussed in the following section. Polyclad larvae, called Goette’s larvae or Müller’s larvae, possess an anterior head with simple eyes, a frontal glandular complex, and a brain; they possess a posterior-ventral mouth, surrounded by epidermal lobes featuring bands of cells with long cilia (Fig. 6A). Two nerve tracts (dorsal and ventral connective) project from the brain into the lobes, and toward a nerve ring surrounding the pharynx. Sensory (epithelial) neurons and axons are also associated with the ciliated bands in the epidermal lobes surrounding the mouth (Lacalli, 1982, 1983). Neurogenesis has been studied for the species Imogene mcgrathi (Younossi-Hartenstein & Hartenstein, 2000b) and Maritigrella crozieri (Rawlinson, 2010), using antibodies against acetylated tubulin, and specific neurotransmitters (FMRF, serotonin). In Imogine, following gastrulation, at around 50% embryonic development, the primordium of the brain (which also includes nonneuronal cell types) appears as a cluster of 80–100 cells underneath the anterior ectodermal layer. As mentioned earlier for Mesostoma lingua, small, discrete clusters of pioneer neurons can be recognized from 60% development onward (Fig. 6A. They pioneer the commissure of the brain, as well as the dorsal and ventral connective. Slightly later, neurons start to differentiate within the ciliary bands, pharynx, and posterior body.

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Figure 6 The nervous system of lophotrochozoan larvae (trochophores). (A–D) Schematic side views (ventral to the left, anterior up) of larvae of a polyclad platyhelminth (A), polychaete annelid (B), mollusk (C), and bryozoan (D). Spatial pattern of neuronal cell bodies and neurite bundles are shown in red. Typical distribution of early differentiating (pioneer) neurons indicated by purple spheres. (E and F) Schematic ventral view of lophotrochozoan at larval stage (E) and after metamorphosis (F). Parts of the larval nervous system (apical organ of the brain, neurite bundles along ciliary bands; shown in red) are lost during metamorphosis, whereas other elements (e.g., cerebral ganglia, ventral nerve cord; rendered in green) are integrated into the postmetamorphic nervous system (after Nielsen, 2015, with permission). The semantic relationship between brain and apical organ is clarified at top of (A and B): the apical organ consists of intraepithelial (“flask-shaped”; often serotonergic) neurons associated with a tuft of elongated cilia. In most trochophores, these cells produce basal axons forming a small commissural neuropil, and they are presumed to be sensory neurons. The apical organ, which in most cases is lost during metamorphosis, is typically accompanied by bilateral “cerebral ganglia,” which nucleate the adult brain. (G–I) Frontal glandular complex in Platyhelminthes (Macrostomum lignano; G and H) and Acoela (Convoluta pulchra; I). (G and H) Horizontal confocal sections of head where glands (G) and neurons (H) are labeled; (I) represents a frontal view of the head, based on electron microscopy, showing profiles of gland necks (turquois), sensory dendrites (red), and epidermal cells (white; after Klauser et al., 1986, with permission). The frontal glandular complex is formed by the long necks of gland cells that project through the brain and end at the anterior tip of the animal (G and I). Gland necks are accompanied by apical dendrites of sensory neurons whose cell bodies are located in the brain (H and I).

Neural Architecture and Development in Lophotrochozoan Larvae

Members of most lophotrochozoan clades form a type of planctonic larva, called trochophore (Nielsen, 2005; Richter et al., 2016). These larvae, similar to to the polyclad larvae discussed in the previous section, have belts of ciliated cells, involved in locomotion and feeding, at characteristic positions. Trochophores have an anterior-ventral mouth and (in most cases) a posterior anus (Fig. 6B–D). A conspicuous ciliated ring, the prototroch, flanks the mouth anteriorly; many types of trochophores also develop a postoral ciliated ring (metatroch) and a ring surrounding the anus (telotroch). The anterior part of the trochophore, located in front of the prototroch, is called epimere; the region behind it represents the hypomere. The epidermis carrying the prototroch and/or metatroch is often folded into lobes, or tentacles, giving the larva of different taxa a number of different shapes (and names).

The nervous system of the trochophore larvae investigated for most lophophorates and spiralians resembles the one described earlier for the polyclad larva and typically includes four major elements (Fig. 6B–E). These are an apical brain, located near the tip of the epimere; axon tracts and sensory neurons associated with the ciliated bands; one or more longitudinal tracts extending between epimere and hypomere; and ring-shaped axon tracts, accompanied by neurons, surrounding the mouth and anus. Given the small size of the larvae, overall neuron numbers are small (tens to hundreds), and most neurons are intra/basiepithelial, although subepithelial neurons have been described for the brain of several taxa (e.g., Phoronida: Temereva & Wanninger, 2012). Despite the small number of neurons, numerous different transmitters were detected, including serotonin, catecholamines, FMRF, leu-enkephalin, and others.

The distribution of serotonergic neurons in the trochophore nervous system has been studied in great detail, and it has been used to infer phylogenetic relationships among lophotrochozoans and deuterostomians (Hay-Schmidt, 2000; Mollusca: Dickinson et al., 2000; Kreiling et al., 2001; Redl et al., 2014; Annelida: Müller & Westheide, 2002; Brinkmann & Wanninger, 2008; Fischer et al., 2010; Bryozoa: Santagata, 2008; Phoronida: Temereva & Wanninger, 2012; Brachiopoda: Altenburger & Wanninger, 2010; Nemertea: Hindinger et al., 2013; von Döhren, 2016). Serotonin is expressed in small clusters of neurons in the brain of virtually all bilaterian animals. In trochophores, as well as larvae of deuterostomes (see later), serotonergic neurons, located in small structure called the apical organ, are among the first neurons to differentiate. They form a long tuft of cilia and presumably act as sensory neurons. The apical organ is a transient, larval structure, which (along with various other larval neurons and organs) undergoes cell death during metamorphosis. The apical organ is distinguished from the bilaterally adjacent brain lobes (cerebral lobes, cerebral ganglia), which start to form in the trochophore slightly later than the apical organ, and which persist as the brain into the adult stage (Fig. 6E, F). It is difficult to determine how the apical organ, as defined for trochophore larvae, relates to the apical organ of cnidarian planulae, or polyclad (platyhelminth) larvae discussed earlier. The planula apical organ is also sensory and transient in nature; it consists, among others, of ciliated intraepithelial neurons whose axons form a small neuropil (Chia & Koss, 1979; Nakanishi et al., 2008; Piraino et al., 2011). In polyclad larvae, the apical organ (as defined and described by Lacalli [1983]) is formed by specialized epidermal cells that extend a tuft of cilia (but have no axons), accompanied by a circular array of gland cells that surround the tuft. An apical “tangle” of neurites formed by clusters of neurons of the brain underlying the apical organ is in contact with the apical organ and may conduct sensory information. This arrangement of cells is reminiscent of the “frontal glandular complex” (Klauser et al., 1986; Ehlers, 1992; Fig. 8G–I) that exists in many platyhelminths, as well as acoelomorphs. One might speculate that an apical, sensory organ predates the bilateria, is retained in lophotrochozoa and deuterostomia, and was modified, adding glandular elements, in aceolomorpha and platyhelminths.

In cases where the trochophore nervous system has been reconstructed at different stages, neurons are added sequentially, with some neurons appearing early (“pioneer neurons”), followed by later differentiating neurons that add to the tracts laid down earlier. Often, individual neurons located in the brain/apical organ and near the posterior pole of the larva appear first, and their axons grow toward each other, forming a longitudinal connective (e.g., Brinkmann & Wanninger, 2008; Fischer et al., 2010; Redl et al., 2014). Chains of nerve cells along the ciliated rings or lobes make their appearance slightly later and form axon bundles that connect with the apical organ or the connectives. The distribution of the early-appearing “pioneers” can vary among different members of the same clade; for example, in trochophores of the polychaete Platynereis dumerilii, the first, serotonin-IR-positive neurons to appear are located in the brain and posterior ventral nerve cord, followed by neurons in the anterior nerve cord and ciliated ring (Fischer et al., 2010); in Sabellaria alveolata, the first serotonin-IR-positive neurons appear in the prototroch and pharyngeal nerve ring (Brinkmann & Wanninger, 2008). Often, the pattern of tracts laid down in the larva approximates the adult architecture. In annelid trochophores, a prominent ventral pair of cords is built, which later becomes incorporated into the ventral ganglionic chain of the adult nervous system (Nielsen, 2015; Fig. 6E, F). Molluskan larvae (e.g., Redl et al., 2014) form two pairs of longitudinal cords, matching the adult pedal and visceral cords (see earlier). Interestingly, some entoproct larvae also show four cords, which is taken as evidence that these animals and mollusks could represent a common taxon, called “tetraneuralia” (Wanninger, 2009). Trochophores of other lophophorates (ectoprocts, phoronids, brachiopods) typically possess one paired cord, even though these are then lost during metamorphosis. Nemerteans have a peculiar trochophore-like larva, called pilidium, with an apical organ, chains of sensory neurons/axons along the ciliated margin of the head, and a nerve ring surrounding the mouth. Of these elements, only the oral nerve ring is retained in the adult, which develops from a set of adult primordia grouped around the gut of the larva; all other elements of the larval nervous system undergo cell death (“catastrophic metamorphosis”; Hindinger et al., 2013).

Neural Development in Annelids

The steps of early neurogenesis, as well as later events of axonal extension and pathfinding, have been studied in recent papers for the polychaete annelids Platynereis dumerilii (Denes et al., 2007) and Capitella sp. (Meyer & Seaver, 2009), and the leeches Helobdella robusta and Theromyzon rude (Stuart et al., 1989; Ramírez et al., 1995; Shankland, 1995; Shain et al., 1998; Shain et al., 2004; Zhang & Weisblat, 2005). In embryos of P. dumerilii, the neuroectoderm becomes delineated when an anterior and a posterior-ventral domain of the ectoderm express SoxB and other transcriptional regulators that form part of the proneural cassette (Denes et al., 2007; Simionato et al., 2008; Kerner et al., 2009; Demilly et al., 2013; Fig. 7A–B). The neuroectoderm contains rapidly proliferating neural progenitors. At later stages, postmitotic cells segregate from the neuroectoderm and assemble a second, deeper layer of neural precursors that lose expression of proneural genes and, in turn, activate genes that promote neural differentiation (e.g., prospero, elav; Simionato et al., 2008; Fig. 7C–F).

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Figure 7 Neural development in Annelida. (A–D) Key events of early neurogenesis in polychaete annelid. Upper panel (A) represents schematic sagittal section of an early embryo at the onset of neurogenesis; (B and C) show schematic cross sections of neurectoderm (blue; B), proliferating neural progenitors (purple; C) and neural primordium (D; postmitotic precursor in orange; differentiating neurons in red). (E and F) Horizontal confocal section (E) and digitally tilted cross sections (E’ and F) of neurectoderm of polychaete Platynereis dumerilii, labeled with probe against proneural gene neurogenin (red), neural precursor/neuron marker Elav (green in E and E’) and differentiated neuron marker synaptotagmin (Syt; green in F; from Simionato et al., 2008, with permission). (G) Early neurogenesis in leech Helobdella robusta. Bodywall and ganglia of ventral nerve cord are proliferated from a set of posteriorly located stem cells (teloblasts; bottom of schematic drawing). Most neurons derive from the N-teloblast (N), whose lineage is shown. Aside from the N-teloblast, other teloblasts (e.g., O) produce mostly epidermis and small numbers of neurons. Teloblasts sequentially bud off segmental founder cells (blast cells), whereby one pair of blast cells is responsible for one segment. Blast cells give rise to a few neural progenitors (colored; middle of diagram). Neural progenitors divide in a fixed pattern, producing clones of neurons that tile into one ganglion (top). Time axis (in hours after birth of teloblast) is shown at left of diagram. (From Zhang & Weisblat, 2005, with permission). (H) Sequential formation of nerve tracts and peripheral nerves in leech Theromyzon rude. Nerves were traced based on labeling with antibody against acetylated-Tubulin (black). Due to sequential generation of neurons (see G), ganglia at the top of drawing are formed early and are therefore more mature at the snapshot shown than ganglia at the bottom. Time axis at the right of drawing gives hours after birth of the neural progenitors. The first longitudinal tracts of the ventral nerve cord appear between 50 and 55 hr (“F,” “C”); peripheral nerves appear in the order medioanterior nerve (MA, pioneered by the cf3 neuron; 55–58 hr), anterior (AA; 55–58 hr), posterior (PP; 60–65 hr), ultraposterior (UP; 70 hr), and dorsoposterior (120 hr) (from Shain et al., 2004, with permission). (I) Horizontal confocal section of three consecutive ganglia of embryonic leech ventral nerve cord labeled with anti-Acetylated tubulin (red). Longitudinal tracts and peripheral nerves are visible. On left side, O-teloblast was injected, thereby visualizing segmentally repeated clones of epidermal cells and some neurons (green). On right side, O-teloblast was ablated, resulting in loss of few, specific neural elements, including posterior nerve (PP; truncated nerve indicated by arrow) (from Shain et al., 2004, with permission). (J) Schematic diagram of leech ventral ganglion, illustrating pattern of discrete longitudinal tracts (V, DL, DM, M) formed by sensory afferents (from Zipser et al., 1989, with permission). (K) Flow chart of axonal pathfinding of sensory afferents in leech (after Song & Zipser, 1995, with permission). Sensory axons form fasciculated peripheral neurite bundles (left). After entering the central nervous system (i.e., the segmental ganglion), sensory afferents defasciculate (middle). Subseqently, they sort according to origin and/or modality, forming specific central tracts (right). Experimental interference with specific membrane glycoproteins utilized for differential adhesion (“recognition”) disturbs fasciculation or sorting of sensory axons (bottom of diagram).

The exact pattern of proliferation of neural progenitors has not yet been reconstructed for polychaete embryos. In particular, it is not clear whether progenitors generate fixed lineages. A strictly controlled (“precise”) pattern of asymmetric divisions, resulting in fixed lineages, could easily explain the formation of a neural architecture that features invariant, uniquely identifiable cells, an architecture that is often encountered in invertebrates, including mollusks and annelids (Bullock & Horridge, 1965; Koester & Kandel, 1977; Zipser, 1982). Asymmetric neural progenitor divisions and fixed lineages form a key element of neurogenesis in many ecdysozoans (e.g., arthropods; see later). This pattern of neurogenesis has also been documented clearly for leeches, but it may be much more widespread among annelids and other lophotrochozoans. Thus, fixed lineages characterize the early (cleavage) divisions in all spiralian embryos, implying that the genetic mechanism that is required for the precise inheritance of intrinsic cell fate determinants from mother cell to daughter cells is in place. Larval structures, such as the apical organ, have been followed back to (invariant) blastomeres (Nielsen, 2015). In leech embryos, fixed lineage patterns of proliferation can be followed much further, into the nervous system. In these animals, a bilateral set of four large, posteriorly located blastomeres, called teloblasts, generate the trunk of the worm in a highly ordered anterior-to-posterior sequence (Stent, 1985; Weisblat, 2007). The N-teloblasts are responsible for most of the neurons of the ventral ganglia. N-teloblasts bud off pairs of so-called N-blast cells; each pair of N-blast cells then divides in an asymmetric, fixed pattern to form the neurons of one segmental ganglion (Zhang & Weisblat, 2005; Fig. 7G).

Later steps of neural development, in particular axonal pathfinding and target selection, have also been studied in the ventral ganglia of leech embryos. Each ganglion has approximately 400 neurons, and precise maps of these neurons have been generated (Muller et al., 1981). Motor neurons grow out in a fixed order to sequentially pioneer the different peripheral nerves of each segment (Shain et al., 2004; Fig. 7H, I). A small number of sensory neurons with specific modalities are located within the ventral ganglia. The majority of sensory neurons, sensing touch, chemical stimuli, and light have cell bodies associated with the epidermis of each segment, and project their axons toward the ventral ganglion via several compact nerves (Fig. 7J). Sensory neurons located along discrete positions within the epidermis follow an invariant birth order, with neurons aligned along the center appearing first (Stewart Macagno & Zipser, 1985). Subsets of sensory axons express different membrane epitopes, allowing for precise developmental reconstruction and experimental manipulations (Hockfield & McKay, 1983; Macagno et al., 1983). Upon reaching the neuropil, sensory axons defasciculate and sort out into different bundles that project as separate entities along the longitudinal axis of the ganglion (Fernandez, 1978; Zipser et al., 1989, 1994; Peinado et al., 1990; Fig. 7J). We here encounter a principle of neural architecture that applies to most, if not all, instances of large and complex nervous systems: neurons forming different classes, based on location, sensory modality, or various functional properties, define discrete, nonoverlapping axon tracts and neuropil domains.

Experimental studies confirmed the pivotal role of membrane adhesion proteins for the ordered projection and connectivity of leech sensory neurons. For example, by perturbing the function of Lan3-2, a glycosylated form of the leech neural cell adhesion molecule (NCAM) homolog expressed early on all peripheral sensory neurons, the initial formation of a cohesive peripheral nerve was disturbed (Zipser, 1995; Huang et al., 1997). At later stages, when sorting of axons with different modalities into discrete bundles occured, subsets of the NCAM expressing neurons changed their glycosylation state and could be differentially visualized and perturbed by other antibodies. Such perturbation experiments blocked the formation of the central bundles (Song & Zipser, 1995; Fig. 7K). Netrins and their receptors, another highly conserved family of proteins with an established role in controlling midline crossing of axons (see earlier), are also expressed in the leech nervous system (Gan et al., 1999; Aisemberg et al., 2001).

Deuterostomia

The second large grouping of bilaterians, deuterostomes, show a number of characters in early development and adult structure that sets them apart from the lophotrochozaoans discussed earlier, including the type of cleavage and gastrulation, as well as the formation of mesoderm and gills. Deuterostomes consist of two major supraphyla, the ambulacraria (echinoderms and hemichordates) and chordates (cephalochordates, tunicates, and vertebrates). Echinoderms include errant animals, the asteroids (starfish; Fig. 8A, B) and holothuroids (sea cucumbers), and sessile forms with tentacles, the crinoids (sea lillies). Hemichordates are comprised of the enteropneusts (acorn worms; Fig. 8C, D), large, errant worms, and the sessile colony-forming pterobranchs (sea angels). Echinoderms and many hemichordates form small planktonic larvae that have fundamental characteristics (including elements of the nervous system) in common with the lophotrochozoan trochophores discussed in a previous section. Among the chordates, urochordates (also called tunicates) are mainly sessile filter feeders. Urochordate larvae are motile tadpoles (Fig. 8F, G) with close resemblance to vertebrate larvae. Some urochordates (i.e., the appendicularians) retain a tadpole-like body plan during the adult phase; others (i.e., the ascidians [sea squirts]) undergo metamorphosis where much of the larval body is disposed of, and the adult develops from imaginal primordia. Cephalochordates (lancets) include a small number of fish-like animals with similarity to tunicate larvae (Fig. 8H, I).

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Figure 8 Nervous system architecture in the Deuterostomia, shown as schematic sagittal sections. (A and B) Echinodermata (starfish, sea urchins, sea cucumbers, sea lilies); (C and D) Hemichordata (acorn worms); (F and G) Urochordata (tunicata; sea squirts, salps, larvaceans); (H and I) Cephalochordata (lancelets). (E) Schematic representation of brain of hemichordate Saccoglossus kowalevskii (left), cephalochordate Branchiostoma floridae (center), and urochordate Botryllus schlosseri (right; dorsal view). Expression domains of Otx, Pax2/5/8, and Hox complex demarcate region considered to be homologous to the vertebrate midbrain-hindbrain boundary (red bar) (after Lacalli, 2001, with permission).

Neural Architecture of Deuterostomia

The subdivision of deuterostomes into two distinct clades, Ambulacraria and Chordata, is reflected noticeably in the architecture of the nervous system. Echinoderms and hemichordates have a diffuse basiepithelial nerve net, with ganglion cells and sensory neurons scattered over different parts of the body surface (Fig. 8A, C). A nerve net surrounding the intestine has been described for echinoderms (García-Arraras et al., 2001). In addition to the nerve net, ambulacrarians feature condensed basiepithelial cords, often associated with tubular invaginations of the epithelium. In echinoderms (e.g., sea urchins, starfish) these include a circum-oral nerve ring and radial nerve cords along the arms (Cobb & Stubbs, 1981; Mashanov et al., 2007; Fig. 8A). Neurons of the radial cords develop in association with invaginated epithelial tubes (Märkel & Röser, 1991; Mashanov et al., 2007). In the worm-shaped hemichordates, there is a longitudinal dorsal and a ventral cord. Within a middle part of the body, called the mesosome or collar, the dorsal cord has invaginated and forms a tube, with both nerve cells and throughgoing neurite bundles clustered around the basal (outer) surface of the tube (Nomaksteinsky et al., 2009; Kaul & Stach, 2010; Lowe et al., 2015; Fig. 8C). It has long been speculated that the dorsal cord of the collar may constitute the evolutionary forerunner of the neural tube in chordates. Thus, the anteriormost Hox genes (Hox1, 2), which mark the hindbrain in vertebrates, are expressed in the boundary region between collar and tail in the enteropneust Saccoglossus kowalevskii (Lowe et al., 2003; Pani et al., 2012; Fig. 8E). Engrailed (En), whose expression domain forms a stripe between the hindbrain and the midbrain in vertebrates, at the vertebrate midbrain-hindbrain boundary, is found in the posterior collar of Saccoglossus. Otx, one of the genes defining the forebrain and midbrain in vertebrates, appears in the entire collar region.

The similarity with vertebrates in regard to neural architecture is even more pronounced in the larvae of urochordates and in cephalochordates (Fig. 8F, G, H, I). None of these shows diffuse nerve nets; instead, the central nervous system is restricted to the neural tube, a simple invaginated basiepithelial nervous system extending along the dorsal side of the animal. The neural tube has an epithelial layer with an inner lumen, and relatively small numbers of neurons and their processes flanking the basal surface of the tube. Along the anterior-posterior axis, the neural tube of urochordate larvae is divided into three domains, the anterior sensory vesicle, followed by the visceral ganglion, and the spinal cord (Nicol & Meinertzhagen, 1991; Sorrentino et al., 2000; Lacalli, 2001; Imai & Meinertzhagen, 2007a; Fig. 8E, E). Simple photoreceptors and ciliated mechanoreceptors, thought to be forerunners of the eyes and inner ear in vertebrates (Caicci et al., 2007), are integrated into the epithelial wall of the sensory vesicle. Its sensory nature, as well as the expression of a characteristic set of genes, suggests that the anterior vesicle corresponds to the fore/midbrain of vertebrates (Wada et al., 1998; Cañestro et al., 2005; Lacalli, 2006; Fig. 8E). The posteriorly adjacent visceral ganglion, which expresses genes indicative of the hindbrain in vertebrates, is formed by a bilateral cluster of neurons whose axons form a pair of tracts extending posteriorly on either side of the the spinal cord. The spinal cord contains no neuronal cell bodies. In addition to the CNS, ascidian larvae (and adults) have a peripheral nervous system, consisting of intra-epidermal sensory neurons whose axons form nerves projecting into the sensory vesicle of the brain (Takamura, 1998; Imai & Meinertzhagen, 2007b). The nervous system of cephalochordates (lancelets) is similar to the tripartite neural tube of the tunicate tadpole (Lacalli, 1996, 2004, 2006; Wicht & Lacalli, 2005). However, it possesses significantly higher numbers and diversity of neurons in particular along the spinal cord, which features dorsal sensory neurons and ventral motor neurons, in addition to glia, similar to the spinal cord of vertebrate larvae (Wicht & Lacalli, 2005).

Neural Development in Ambulacrarians

The early steps of neurogenesis have been addressed in a number of recent studies on the embryos and larvae of sea urchins and other echinoderms (Burke, 1983; Byrne & Cisternas, 2002; Nakajima et al., 2004; Yaguchi et al., 2006; Nakano et al., 2006; Bishop & Burke, 2007; Byrne et al., 2007; Dupont et al., 2009; Angerer et al., 2011; Bishop et al., 2013; Garner et al., 2016) and hemichordates (Nielsen & Hay-Schmidt, 2007; Miyamoto et al., 2010; Kaul & Stach, 2010; Cunningham & Casey, 2014; Kaul-Strehlow et al., 2015). In ambulacrarian larvae, called dipleurulae, the nervous system is comprised of an apical organ, as well as peripheral arrays of sensory neurons and neurite bundles extending along the bands of ciliated cells that surround the mouth and ventral epidermis (Fig. 9A, B), very similar to what has been described for the lophotrochozoan trochophore (see earlier). However, unlike trochophore larvae, dipleurulae also possess a large number of neurons forming a diffuse nerve net. The specification of the neurectodermal field that gives rise to the larval nervous system begins with the expression of SoxB transcription factors, which appear ubiquitously at the blastula stage (Lowe et al., 2003; Cunningham & Casey, 2014, for Saccoglossus kowalevskii; Garner et al., 2016, for Strongylocentrotus purpuratus; Fig. 9C, D). Genes associated with the early formation of an apical brain (e.g., six3/6) become focused to the anterior of the embryo as a result of the Wnt signaling pathway (Lowe et al., 2003; Angerer et al., 2011; Fig. 9E). The anterior tip of this ectodermal region gives rise to the apical organ. Further posteriorly, BMP signaling shapes the neurogenic ectoderm into a specific domain, the “pre-ciliary band ectoderm” (Angerer et al., 2011; Burke et al., 2014; Fig. 9E), that generates the neurons along and near the ciliary band. In the similarly shaped hemichordate embryo, neurogenesis is initially not patterned along the dorsoventral axis despite an existing BMP gradient (Lowe et al., 2006; Cunningham & Casey, 2014; Fig. 9C). It is not yet clear what genetic mechanism orchestrates the increased production of neurons that form the ciliary bands in hemichordate larvae, or the dorsal and ventral cords that appear at a later stage.

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Figure 9 Neural development in Ambulacraria (Echinodermata + Hemichordata). (A and B) Schematic side views (ventral to the left, anterior up) of larvae of echinoderm (A) and hemichordate (B1: early or “Heider” stage; B2: late or “Agassiz” stage). Spatial patterns of neuronal cell bodies and neurite bundles are shown in red. Typical distribution of early differentiating pioneer neurons is indicated by purple spheres. (C and D) Schematic sagittal section of a hemichordate embryo (C) and echinoderm embryo (D) at the onset of neurogenesis. (E) Schematic lateral view of echinoderm embryo and larva at sequential stages of development, explaining role of Wnt and BMP signaling in the specification of the neuroecoterm (from Angerer et al., 2011, with permission). (F–I) Schematic cross sections of developing nervous system in hemichordate embryo and metamorphosing larva; (F) illustrates neurectoderm (blue) and proliferating/ delaminating neural progenitors and precursors (purple/orange); (G) shows differentiated nerve net (red) and anlage of adult neurons (blue) in larva; (H and I) show invagination of dorsal collar ectoderm, giving rise to dorsal nerve cord.

The specification and proliferation of neural progenitors that generate the nervous system of ambulacrarian larvae and juveniles takes place within the ectoderm (Fig. 9C, D, F). In the sea urchin, SoxC is expressed in a discrete domain within the apical ectoderm and then becomes restricted to neural progenitors when they undergo their final mitosis, which gives rise to neurons marked by typical neural genes, such as the POU transcription factors Brn1/2/4 and Elav (Garner et al., 2016). The progenitor mitosis appears to be asymmetric, resulting in the expression of the Notch ligand Delta in only one of the daughter cells. In the hemichordate, markers for postmitotic neurons (Elav, Synaptotagmin) are expressed in scattered cells within the ectoderm from an early stage onward (Cunningham & Casey, 2014).

The initial phase of neurogenesis in ambulacrarians gives rise to a basiepithelial nerve net, supplemented by basiepithelial neurite bundles associated with the ciliary bands and apical organ (Fig. 9F, G). At a later stage, parts of the ectoderm with neurogenic potential invaginate, forming internalized tubular structures in whose walls additional neurons form (Fig. 9H, I). In hemichordates, the dorsal ectoderm of the collar region gives rise the dorsal nerve cord (Kaul-Strehlow et al., 2015); in some directly developing echinoderms (e.g., sea cucumber Eupentacta fraudatrix), part of the ventral ectoderm surrounding the embryonic mouth invaginates to form the rudiment of the adult nervous system (Mashanov et al., 2007). Most echinoderms are indirect developers in which the adult body, including the nervous system, forms largely independently of the larval organs. Typically, a rudiment of the adult develops on one side of the body in close association with one of the mesodermal coelomic pouches (Juliano et al., 2010; Bessodes et al., 2012). The nervous system appears de novo within this rudiment, although in some cases (e.g., sea urchin Hemicentrotus pulcherrimus), larval nerve cords are still around when adult neurons begin to appear and establish transient connections (Katow et al., 2009).

Neural differentiation and axon extension in echinoderms and hemichordates follow the pattern sketched in the previous section for lophotrochozoan larvae. Small numbers of neurons in the apical organ and at posterior positions within the preciliary band ectoderm extend axons across the midline and toward each other. These are followed by additional cells that intercalate in between the early ones, but also form at new positions (e.g., nerve ring surrounding the mouth and lateral ganglia in sea urchin; Bisgrove & Burke, 1986; Beer et al., 2001; Garner et al., 2016). Neurons establish axon tracts forming a commissure underneath the apical organ, and nerve bundles along the ciliary bands and around the mouth. Neurons also project outside the ciliated ring, forming a basiepithelial nerve net. In hemichordates, indirect developers with larvae (tornaria) similar to echinoderm larvae are distinguished from direct developers. In the former, the larvally established tracts along the ciliary bands and apical organ disappear during metamorphosis (Nielsen & Hay-Schmidt, 2007; Miyamoto et al., 2010; Kaul-Strehlow et al., 2015). In both types, the diffuse nerve net characteristic of the adult stage makes an early appearance. By contrast, the dense longitudinal bundles of the dorsal and ventral cord, as well as a number of circular tracts, appear secondarily.

Little is known about the molecular mechanisms directing axonal growth in ambulacrarians. Studies in sea urchin (Katow, 2008; Abe et al., 2013) identified Netrin, expressed at a high level in a domain straddling the dorsal (=aboral) midline, and at lower levels ventrally, as an important factor for the formation of the commissural tract underlying the apical organ. Interestingly, the bimodal action of netrin as an attractant (toward the midline) and repellant (for fibers after they crossed the midline; see section earlier) also seems to operate in the sea urchin larva; the Unc5 receptor, known to mediate the repulsive action of netrin, is expressed in neurons forming the apical commissure (Abe et al., 2013).

Neural Development in Urochordates and Cephalochordates

In urochordates and cephalochordates, as well as the vertebrates (that are not further considered in this chapter), the neurectoderm is restricted to the dorsal side of the embryo by the combined action of BMP/BMP antagonist and Wnt/Wnt antagonist signaling (cephalochordates [Branchiostoma floridae]: Holland et al., 2000, 2005; Lu et al., 2012; Onai et al., 2012; urochordates [Halocynthia roretzi]: Darras & Nishida, 2001; Miya & Nishida, 2003; vertebrates: reviewed in Niehrs, 2010; Fig. 10A). In all three clades, the dorsal neurectoderm invaginates to form the neural tube (Fig. 10B–D; H–J). All cells of the neural tube possess the potential to form neurons and glia. In the ascidian Ciona intestinalis, the proliferation of neural progenitors is determinate and produces fixed lineages, similar to what has been described earlier for leech. Following cleavage and gastrulation, the dorsal neurectoderm (neural plate) consists of 48 uniquely identifiable cells that are arranged in a regular orthogonal pattern of four columns and six rows (Fig. 10E, F). After the invagination of the neural tube, neural progenitors perform an additional two to three rounds of mitosis (Fig. 10C). The majority of cells maintain their epithelial structure and differentiate into ependymal cells (the same cell type that lines the inner lumen [ventricle] of the vertebrate CNS. Fewer than 100 cells delaminate and differentiate into the invariant set of larval neurons (Nicol & Meinertzhagen, 1988a, 1988b; Lemaire, 2009; Fig. 10D–F). The proneural cassette, in addition to other signaling pathways (FGFR, Nodal) that mediate cell–cell interactions between individual cells forming part of this minute neuroepithelium (Hudson & Yasuo, 2005; Hudson et al., 2007; Esposito et al., 2017), control the number and cell type of neurons produced. Many of the larval neurons undergo cell death during metamorphosis; only some cholinergic motor neurons and interneurons are retained. However, the ependymal cells forming the anterior part of the larval neural tube undergo more proliferation and contribute to the adult nervous system (cerebral ganglion; Horie et al., 2011; Fig. 10G). In addition, other structures of the larva that are in close spatial contact with the neural tube, including the neurohypophyseal duct, produce neurons of the adult cerebral ganglion (Manni et al., 1999; Dufour et al., 2006).

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Figure 10 Neural development in invertebrate Chordata (Urochordata + Cephalochordata).

(A) Schematic sagittal section of a chordate embryo at the onset of neurogenesis. (B–D) Schematic cross sections of developing nervous system in urochordate embryo; (B and C) illustrate infolding neurectoderm (neural plate; blue, purple) which contains invariant set of neural progenitors expressing intrinsic determinants (small colored circles). (D) Illustration of neural tube. (E) Line drawing of gastrula stage embryo of Ciona intestinalis (urochordates), vegetal (approximately posterior) view. Prospective neural progenitors gving rise to invariant neural lineages are color coded and named. (F) Left: Schematic dorsal view of neural plate of Ciona, illustrating regular pattern of neural progenitors (numbered and color coded). Right: Schematic drawing of Ciona larva, illustrating origin of cells of sensory vesicle, visceral ganglion, and tail nerve cord from invariant sets of progenitors in neural plate (E and F from Hudson et al., 2007; Hudson & Yasuo, 2005; with permission). (G) Schematic drawing of Ciona larva (left) and adult (right), illustrating contribution of neural progenitors located within larval brain to adult brain (from Horie et al., 2011, with permission). (H–J) Schematic cross sections of developing nervous system in cephalochordate embryo; (H and I) illustrate infolding neurectoderm (neural plate; blue, purple), and (J) shows neural tube with differentiated neurons located within neuroepithelium (small arrow) and around basal surface of epithelium. (K) Side views (anterior to the left; dorsal up) of cephalochordate embryo at sequential stages (in hr after fertilization). Probe against neural marker Elav labels neural plate (top), followed by scattered sensory neuron precursors in epidermal ectoderm (bottom) (from Satoh et al., 2001, with permission). (L and M) Sagittal confocal section of Ciona embryo (L) and larva (M), showing expression of Fibrinogen-like protein (green) on cytoplasmic processes of notochord cells reaching around neural tube. (N) RNAi-mediated knock-down of Fibrinogen-like protein results in abnormalities of neuronal trajectories along nerve cord (L–N from Yamada et al., 2009, with permission). (O and P) Sensory nervous system in chordates. (O) Schematic side views of embryos; (P) is a schematic cross section of dorsal nerve cord of cephalochordate, illustrating shape and location of neurons (from Wicht & Lacalli, 2005, with permission). At an early phase of embryonic development (left side of O), BMP signaling inhibits neurogenic potential, restricting neuroectoderm to dorsal side of embryo. During a later phase, progenitors of nonciliated, centrally located sensory neurons (dark purple shading in O and P) are specified in a BMP-dependent manner within the lateral domain of neural plate (in cephalochordates). In vertebrates, most of these sensory neurons arise from the neural crest, located laterally of the neural plate (O, bottom center). Intra-epithelial, ciliated sensory neurons (indicated by cyan shading in O and P) arise in a BMP-dependent manner throughout the ectoderm of urochordates and cephalochordates. In vertebrates, these cells are confined to the sensory placodes which form adjacent to the neural crest (O, bottom center).

Neural development in cephalochordates appears to be similar to that seen in urochordates, aside from the important fact that a fixed lineage pattern has not been observed. The proliferation observed within the neural plate and neural tube appears to follow a symmetric pattern of mitosis (Holland & Holland, 2006; Fig. 10H, I). Neurons become specified and initiate differentiation at an early stage, even prior to infolding of the neural plate, based on the finding that panneural markers like Elav (found in postmitotic neural precursors) are widely expressed in the neural plate (Satoh et al., 2001; Fig. 10K). Pointing toward the same conclusion, expression of the proneural cassette of genes (bHLH gene neurogenin; Notch ligand Delta) is seen in dispersed epithelial cells of the neural plate (Holland et al., 2000; Rasmussen et al., 2007).

Little is known about the sequence of axonal outgrowth or molecular mechanisms directing this process in urochordates or cephalochordates. In the larva of Ciona, a simple pattern of axon tracts interconnecting different parts of the CNS has been described, using antibodies against stabilized tubulin (Takamura, 1998; Fig. 10 L). Two main nerve tracts, projecting posteriorly from the visceral ganglion along the spinal cord, exist. The involvement of interactions between the neural tube and underlying notochord has been demonstrated by a recent study of Ci-Fbrn, a fibrinogen-like protein localized on processes of notochordal cells directly underlying the neural tube (Fig. 10M; knock-down of this factor resulted in disruptions of the spinal cord tracts (Yamada et al., 2009; Fig. 10N).

A fascinating question, both from a developmental-genetic and evolutionary standpoint, is the formation of the peripheral nervous system in chordates, compared to other animals. As noted in preceding sections (and as will be evident in the following subchapter on ecdysozoans), sensory neurons in most animals are intraepithelial, ciliated cells distributed all over the body. This also applies to sensory organ progenitors (SOPs), which are not restricted to the neurectoderm but can form at all positions within the ectoderm. The BMP signaling pathway, which inhibits the formation of CNS-forming neurectoderm during an early phase (Fig. 10O), does not negatively affect the formation of SOPs; to the contrary, in a later phase of neurogenesis that follows after the neural plate has been specified, BMP activity stimulates the formation of sensory neurons within the lateral neural plate, as well as the epidermal ectoderm (Lu et al., 2012; Schlosser et al., 2014; Fig. 10O). In the chordate lineage we encounter two different classes of sensory neurons. Thus, aside from the ciliated, intraepithelial receptors described earlier (“ciliated sensory neurons”), one observes sensory neurons whose cell bodies are located within the CNS, or in sensory ganglia located close to the CNS, and whose long “axonal” processes extend peripherally, terminating in specialized structures underneath the epidermis or in the musculoskeletal tissues (“nonciliated sensory neurons”; Fig. 10P). Most vertebrate neurons sensing touch, vibration, pain, and temperature belong to this second class; outside the vertebrates they are also found in cephalochordates (Wicht & Lacalli, 2005; Fig. 10P). In both vertebrates and cephalochordates, nonciliated sensory neurons arise in the lateral neural plate in a BMP-dependent manner (Fig. 10O). In vertebrates, a narrow fringe of neurectoderm forming the lateral boundary of the neural plate, called the neural crest, gives rise to most nonciliated sensory neurons. Neural crest-derived cells migrate widely in the body to form sensory ganglia. Ciliated sensory neurons of cephalochordates arise outside the neural plate in the entire ectoderm (Fig. 10K, O). By contrast, in vertebrates, the formation of ciliated sensory neurons (e.g., olfactory receptors; auditory receptors) is restricted to the sensory placodes, regions of the epidermal ectoderm adjacent to the neural crest (Schlosser et al., 2014; Fig. 10O).

Urochordates, now considered to be the sister group of the vertebrates, present a chimaeric case: on the one hand, progenitor cells with the molecular “fingerprint” of neural crest and sensory placodes were found adjacent to the neural plate (Abitua et al., 2012, 2015). These cells produce the (ciliated) sensory neurons of the head. On the other hand, ascidian larvae possess a second population of ciliated sensory neurons (and progenitors) in the trunk and tail, derived from the nonneural ventral ectoderm (Pasini et al., 2006; Tang et al., 2013; Waki et al., 2015). Even though at the stage of specification and differentiation, sensory neurons are guided by the same genetic mechanisms as central neurons (e.g., BMP, proneural cassette, neural differentiation genes), these genetic mechanisms involve signaling interactions that differ between central and peripheral system. Ascidians will provide an ideal system to shed more light on the problem of development and evolution of the sensory system.

Ecdysozoa

The ecdysozoa were recognized as the large clade of animals with a cuticular exoskeleton that is renewed during molts. Two superphyla are distinguished, the pan-arthropods and cycloneuralians (a diverse, probably paraphyletic grouping; Telford et al., 2008; Borner et al., 2014). Cycloneuralians, which are unsegmented, worm-like animals, include nematodes (round worms; Fig. 11A, B), nematomorphs (hair worms), kinorhynchs, loriciferans, and priapulids (Fig. 11C, D). The pan-arthropods (called “arthropods” for simplicity in the following) are characterized by segmented legs and a segmented body (Fig. 11E, F). Arthropods are by far the largest clade of animals, and this include insects and crustaceans (“tetraconata”), next to chelicerates (horseshoe crabs, scorpions, spiders, mites), myriapods (centipedes, millipedes), onychophorans (velvet worms), and tardigrades (“water bears”).

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Figure 11 Nervous system architecture in the Ecdysozoa, shown as schematic sagittal sections. (A and B) Nematoda (round worms); (C and D) Priapulida (acorn worms); (E and F) Arthropoda. (G and H) Line drawings based on Golgi stainings of nervous system of nematode Ascaris and fly Musca domestica, showing structure of representative neurons (from Hanström, 1968; Strausfeld, 1976, with permission). (I) Line drawing of cross section of segmental ganglion of grasshopper Locusta migratoria, illustrating regular orthogonal pattern of longitudinal and commissural tracts (from Tyrer and Gregory, 1982, with permission). (J–L) Schematic dorsal view of sensory organs and brain of arthropods (J: Onychophora; K: Hexapoda/Crustacea; L: Chelicerata). Sensory nerves are rendered in blue; structured neuropil compartments are shown in purple. Hatched lines show boundaries between segmental ganglia. Structural and molecular criteria were used to establish homologies between the segmental neuromeres that make up the brain of different arthropod phyla (for details, see text). The green bar is positioned at the level of the tritocerebrum (hexapod), pedipalpal ganglion (chelicerate), and ganglion innervating oral papilla (onychophoran), which are considered as homologous neuromeres.

Neural Architecture of the Ecdysozoa

Cycloneuralia

Cycloneuralians have a basiepithelial nervous system, formed by a ring-shaped brain around the pharynx (hence the name cycloneuralia). The nerve ring of nematodes and kinorynchs projects several pairs of nerve cords (ventral, lateral, and dorsal nerve cord; Hanström, 1968; White et al., 1986; Herranz et al., 2013; Schmidt-Rhaesa & Henne, 2016) innervating the trunk (Fig. 11B). Priapulids (“acorn worms”) possess a single ventral nerve cord (Rehkämper et al., 1989; Rothe & Schmidt-Rhaesa, 2010; Fig. 11D). For most cycloneuralia, neurons of the CNS are small in number and uniquely identifiable. For example, the nematode C. elegans, one of the classical invertebrate genetic models, has 302 neurons, each one with a unique fate and a fixed pedigree that was followed back to the zygote (Sulston & Horvitz, 1977; Sulston et al., 1983; White et al., 1986). Due to the small number of neurons, neuron shape is simple (typically one or two unbranched processes able to accommodate all synaptic contacts; Fig. 11G), and the expanse of the neuropil does not exceed the nerve ring around the pharynx and the longitudinal cords. A diffuse peripheral nerve net is absent.

Arthropoda: General Neuroanatomical Characteristics

Arthropods possess a subepidermal, ganglionic nervous system and lack a diffuse peripheral nerve net. Ganglia include an anterior brain and a chain of ventrally located segmental ganglia, called ventral nerve cord (Fig. 11F). In the ventral nerve cord, various subsets of ganglia may be fused. Part of the brain also consists of several fused, strongly modified segmental ganglia. Brain and ventral nerve cord of most arthropods (excepting the tardigrades) exhibit large numbers of neurons of considerable size and branching complexity (Fig. 11H), and a voluminous neuropil with several highly structured compartments (e.g., optic lobe, antennal lobe, central complex, mushroom body; Fig. 11H, J; see chapter by Strausfeld, this volume), reflecting the presence of large and complex sensory organs and limbs involved in locomotion, feeding, and reproduction. The different arthropod clades are distinguished by the pattern underlying the structure and modifications of body segments, which in turn dictates to a large extent the organization of the central nervous system.

Arthropoda: Onychophora and Tardigrada

Onychophora (velvet worms), the most basal branch of the pan-arthropods, have a series of more or less identical segments carrying short, stubby limbs. The head carries small cup eyes and large chemosensory antennae, as well as paired mandibles and oral papillae involved in feeding. The brain (supraesophageal ganglion) consists of the large anterior protocerebrum, innervating eyes and antennae, and the posterior deuterocerebrum, associated with the mouthparts (Hanström, 1986; Mayer et al., 2010; Mayer, 2016; Fig. 11J). The phylogenetic position of the tardigrades is debated; according to most current trees, they form a basal branch of the pan-arthropods. Tardigrades have few segments with short appendages. Their CNS includes a brain (supraesophageal ganglion) and a ventral nerve cord with four segmental ganglia (Hanström, 1968; Mayer et al., 2013; Schulze & Persson, 2016). Tardigrades possess only small numbers of neurons (20–30 per ganglion); complex sensory organs or structured neuropil domains are absent.

Arthropoda: Crustacea, Hexapoda, and Myriapoda

In crustaceans and insects, which together form the clade tetraconata (based on the similar structure of lenses of the compound eye), body segments have become modified and partially fused such as to form three domains, the head, thorax, and abdomen. The head comprises a large anterior component (acron) and five segments that surround the mouth. The acron (by some considered to represent a segmental unit called “ocular segment”; e.g., Schmidt-Ott et al., 1995) carries the compound eyes; the first two segments possess large chemosensory appendages (two pairs of antennae in crustaceans; one pair of antennae and a gustatory organ in insects; Fig. 11K). Appendages of the three segments located right behind the mouth opening (gnathal segments) are adapted for feeding. Eight segments (most crustaceans) or three segments (insects) make up the thorax; the number of segment number in the abdomen is highly variable among different species. Each of the segments is innervated by a segmental ganglion. The neuropil of a segmental ganglion is highly structured by longitudinal bundles (connectives) and transverse bundles (commissures) of axons which belong to specific, invariant sets of neurons. Longitudinal tracts in insects form three sets, located at a medial, intermediate, and lateral position; commissures form an anterior and posterior set in each segment (Fig. 11I). Sensory neurons form part of small sensory organs, called sensilla, that are scattered all over the body wall. Different sensilla are specialized for the modalities touch, vibration, stretch, temperature, taste, and olfaction; their axons bundle in compact peripheral nerves, but then project to separate neuropil domains, according to modality and location, as discussed for annelids (see earlier discussion). Sensory neuropil domains are mostly located at ventral positions; neuropil domains receiving dendrites of motor neurons are dorsal (Strausfeld, 1976).

The brain of insects and crustaceans consists of two main parts, the supraesophageal ganglion and subesophageal ganglion, each one derived from several fused segmental ganglia. The supraesophageal ganglion is further divided into the protocerebrum, deuterocerebrum, and tritocerebrum (Fig. 11K). The large protocerebrum is the ganglion of the so-called acron; it includes the multilayered optic lobes associated with the compound eyes, and the central brain, which receives sensory input of all modalities (Hanström, 1968; Strausfeld, 1976; Strausfeld & Nässel, 1980; see chapter by Gelperin, this volume). The deuterocerebrum, which represents the segmental ganglion of the antennal segment, receives predominantly olfactory input from the first antenna; the tritocerebrum is associated with the second antenna (crustaceans) and/or taste sensory organs located in the mouth cavity and the gut (insects; Fig. 11K). The tritocerebrum also forms sensory and motor connections with the stomatogastric ganglia, which innervate the gut. The subesophageal ganglion results from the fusion of the three segmental ganglia of the gnathal segments, which innervate the mouthparts. Longitudinal and commissural axon tracts connecting the neuropils of the ventral nerve cord continue into the (modified) ganglia of the brain, where many additional tract components are added (Hartenstein et al., 2018).

Myriapods (centipedes, millipedes) differ from tetraconata in regard to the high number and uniformity of body segments. The general structure of the brain and ventral nerve cord of myriapods is similar to that of insects (Hanström, 1968; Sombke et al., 2012; Sombke & Rosenberg, 2016; see also Gelperin, this volume).

Arthropoda: Chelicerata

The segmental organization of chelicerates (horseshoe crabs, spiders, scorpions, mites) differs significantly from that of tetraconata and myriapoda. The body includes the cephalo-thorax (fused head and thorax) and the abdomen. Within the cephalothorax one distinguishes the anterior acron, associated with the eyes, from six segments that have appendages involved in sensory reception, feeding, and locomotion. The first two segments form the cheliceres (“pincers” for grasping food) and the pedipalps (leg-like tactile organs in spiders; claws with pincers in scorpions), respectively (Fig. 11L). The posterior four segments are equipped with walking legs. Similar to the body segments, ganglia of the ventral nerve cord and brain have also fused into a supraesophageal ganglion and a ventral ganglion (Babu, 1985; Weygoldt, 1985). The supraesophageal ganglion of chelicerates that are considered to be basal (e.g., Xiphosurae, horseshoe crabs) is likely homologous to the protocerebrum of insects and crustaceans. It has large structured neuropil compartments associated with the eyes and other sensory structures (Fig. 11L). Both anatomical and molecular evidence suggests that the ganglion innervating the cheliceres corresponds to the deuterocerebrum in insects/crustaceans (Damen et al., 1998; Mittmann & Scholtz, 2003); the following ganglion, associated with the pedipalps, corresponds to the tritocerebrum (Fig. 11L). In higher chelicerates (spiders, scorpions) the cheliceral ganglion has merged with the protocerebrum to become part of the supraesophageal ganglion.

Neural Development in Cycloneuralia

Early Neurogenesis

The formation of the nervous system has not been studied for the majority of cycloneuralian clades, with the exception of the nematodes. The nematode Caenorhabditis elegans is one of the classical forward genetic model systems among invertebrates, and work carried out by many laboratories over more than five decades has supplied the reagents and tools which make it possible to conduct a detailed analysis of neurogenesis in this organism. The neuroectoderm of C. elegans occupies the ventral and anterodorsal part of the embryo shortly after gastrulation (Wadsworth & Hedgecock, 1992; Fig. 12A). Subsequently, the neuroectoderm is internalized into the embryo by epiboly: Cells of the laterally adjacent epidermal ectoderm spread out and move over the neural primordium (Chisholm & Hardin, 2005; Fig. 12B–D). Most cells of the internalized neurectoderm behave as dedicated neural progenitors (neuroblasts) that, during very few additional divisions, give rise to fixed neural lineages. Neuroblasts in the anterior part of the embryo form the neurons of the nerve ring around the pharynx; progenitors at further posterior locations produce the ventral nerve cord and the ganglia of the tail (Sulston et al., 1983). Some progenitors, called P-, T-, and V-blast cells, initially stay within the epidermal ectoderm at the surface of the embryo. They resume proliferation during the larval stage; P-and V-cell progeny become, in part, neurons and, in part, epidermal cells (Sulston, 1976; Sulston & Horvitz, 1977; Fig. 12E).

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Figure 12 Neural development in Nematoda. (A–D) Key events of early neurogenesis in nematode C. elegans. Upper panel (A) shows SEM photograph of postgastrula embryo (lateral view; anterior to the left). Mass of neural progenitors is rendered in purple; epidermal ectoderm in gray. Arrows indicate direction of later epiboly of epidermal ectoderm over neural progenitors. (B–D) Schematic cross sections of ventrally located dividing neural progenitors (purple; intrinsic determinants of neural fate shown as small colored circles), overgrown by epidermal ectoderm proliferating neural progenitors. (E) Schematic drawing of C. elegans larva (lateral view; anterior to the left), illustrating pattern of differentiated larval neurons (small elements) and progenitors of adult-specific neurons/epidermis (blast cells; large circles). (F–H) Role of proneural genes (F), Wnt signaling (G1, G2), and intrinsically expressed transcription factors (H) during the specification of neural fate in individual neural lineages. V5-blast in wild type generates neurons and epidermis; loss of proneural gene lin-32 converts neurons into epidermal cells (F). Wnt signal (lin-44) from posterior tail epidermis (G1) acts on mitotic spindle of T-blast; loss of lin-44 results in inverted spindle and positioning of T-derived daughter cells (G2). The POU transcription factor unc-86 is required in many lineages, among them the Q-lineage (H) to signal the exit from self-renewing divisions and onset of differentiation; loss of unc-86 results in multiple iterations of the Q-lineage. (I and J) Schematic cross section (I) and ventral view (J) of C. elegans larva. Axons of laterally located sensory neurons (yellow) extend ventrally; axons of ventrally located motor neurons (red) extend dorsally. Concentration gradients of signals Netrin (ventral), Slit (dorsal), and TGFβ (dorsal) are illustrated in (I) (from Chisholm et al., 2016, with permission. (K–N) Role of Wnt signaling in directing axonal trajectories. Panels show schematic drawings of C. elegans embryo (K) and larvae (L–N) in lateral view. (K) Expression of Wnt signals, encoded by genes lin-44, egl-20, cwn-1, cwn-2, and sfrp-1, in distinct domains along the antero-posterior axis. (L) Wild type. Long axon of sensory neurons ALM and PLM project anteriorly; short dendrites project posteriorly. Loss of posteriorly expressed Wnt signals (e.g., Lin-44) affects posterior neuron PLM, resulting in reversal of axon/dendrite growth direction (M); loss of anterior signals (e.g., cwn-2) has the same effect on the anterior neurons (N after Ackley, 2014, with permission).

The fate of neurons descending from C. elegans neuroblasts is specified by intrinsic determinants at an early stage, starting even before these neuroblasts become internalized (Fig. 12B). Many of these intrinsic fate determinants have been identified, and the attempt to list their respective roles would go far beyond the scope of this chapter. It is important to note that the early step of selecting a restricted number of neuroblasts from initially larger proneural clusters, as explained earlier in this chapter, appears to occur in a different manner in C. elegans, possibly as a result of the small overall number of cells. Genes of the proneural cassette, which in many other animal systems globally control the neuroblast selection process (see the second section of this chapter and following), are all present in the nematode genome but seem to operate only in a limited number of cells. For example, bHLH transcription factors of the Achaete-scute (HLH-14) and Atonal (LIN-32) family are only required in a small subset of neuroblasts; loss of function of these genes leads to a transformation of these neuroblasts into epidermal cells (Zhao & Emmons, 1995; Frank et al., 2003; Zhu et al., 2014; Fig. 12F), but it has no effect on neurogenesis in general. The activity of Notch (LIN-12/GLP-1) and its downstream target, Hairy/Enhancer of Split homolog (LIN-22) is also required only in a few lineages (e.g., vulval lineage; lineages V1-4) to promote the epidermal and inhibit the neural fate (Wrischnik & Kenyon, 1997; Sternberg, 2005).

Given the small overall number of neurons, neuroblasts undergo only few, asymmetric divisions. Many intrinsic and extrinsic factors that control the orientation of the mitotic spindle, as well as the onset and offset of mitotic activity, have been identified in the worm. Mitotic spindles are typically oriented anterior-posteriorly, parallel to the embryonic surface, forming an anterior (a) and a posterior (p) daughter cell at any given division. The worm Wnt homolog, lin-44, is required as an extrinsic cue for the correct polarization of the division of the T-neuroblast, one of the postembryonic neuroblasts located in the larval tail (Herman et al., 1995). Lin-44 is expressed by several epidermal cells located at the tip of the tail (Fig. 12E, G1, G2). The neighboring T-neuroblast produces an anterior daughter, which in turn gives rise to epidermal cells, and a posterior daughter, which forms neurons. In the absence of lin-44, the polarity of this division is reversed, with the anterior daughter of T forming neurons and the posterior one epidermis. A well-studied example for an intrinsic factor needed for controlling neuroblast mitosis is the dPOU transcription factor, UNC-86, expressed in many neuroblasts. The postembryonic neuroblast Q produces an anterior and a posterior daughter cell, Ql.a and Ql.p. Both of these cells undergo one final division in which the posterior daughter generates two sensory neurons (PVM, SDQ), and the anterior daughter one interneuron (PQR) and one cell that undergoes programmed cell death. UNC-86 is turned on only in the posterior daughter cell and is required to “flag” the mitosis of this cell as a final division. If unc-86 is absent, QI.p behaves as the neuroblast Q itself and continues to divide (Chalfie et al., 1981; Finney & Ruvkun, 1990; Baumeister et al., 1996; Fig. 12H).

Pathfinding and Connectivity

The main structures of the C. elegans CNS are the pharyngeal nerve ring and the ventral nerve cords. Branches of several motor axons distributed over the entire length of the animal leave the ventral cords and travel to the dorsal side, where they form a thin dorsal cord. The pharyngeal nerve ring and ventral cords are pioneered by a small number of identified neurons located at strategic positions within the embryo (Durbin, 1987; Chisholm et al., 2016). Two neurons located in the tail of the animal, PVPR and AVG, project anteriorly and lay down the trajectory for the ventral cords. After reaching the pharyngeal ring ganglion, PVPR is joined by the axon of the anteriorly located AVKR neuron that, after crossing the midline, projects posteriorly. Small groups of other identified neurons (AVA, AVB) pioneer the nerve ring. Two peripherally located sensory neurons, AVM and PVM, project ventrally and then anteriorly, joining the ventral cords. Four additional sensory neurons (paired ALMs and PLMs) project straight anteriorly along a lateral pathway. These isolated, peripheral neurons, in addition to several later born neurons that are also easily identifiable, such as the hermaphrodite-specific HSN neuron located in the tail and extending its axon along the ventral cords, have served as a paradigm for axonal pathfinding in C. elegans, as briefly outlined in the following.

The choice of axons into which direction and how far to grow is controlled by the attractive and repulsive cues provided by the Netrin/Netrin receptor and the Slit/Robo cassette, as well as the Wnt signaling pathway (Araújo & Tear, 2003; Killeen & Sybingco, 2008; Chisholm et al., 2016). C. elegans Netrin (UNC-6) is expressed by different cell types along the ventral midline (Wadsworth, 2002; Ogura et al., 2012; Fig. 12I, J). A Netrin receptor mediating attraction (UNC-40) is expressed on the growth cones of neurons that grow toward the midline, among them AVM, PVM, and HSN. In unc-6 or unc-40 mutants, axons of these neurons, which normally project their axons toward the ventral cord, project longitudinally instead. This demonstrates that axon elongation per se is not diminished, but axons are unable to sense the direction they are growing in. Netrin is also responsible for the repulsive activity excerted by the midline on other, noncommissural axons. Thus, a second Netrin receptor, encoded by the unc-5 gene, interacts with Netrin such that axons of the lateral ALM and PLM neurons are repelled from the midline. Ectopic expression of functional UNC-5 protein leads to the dorsal extension of many axons. This effect depends on the presence of the unc-6 (netrin) gene; in unc-6 mutants, unc-5 overexpression does not change axonal trajectories, providing genetic evidence for netrin and the unc-5 receptor interactions. The Slit/Robo system of C. elegans mediates axon repulsion, as in other systems. Interestingly, the pattern of expression is the opposite of that found in other animals. Slit is expressed along the dorsal side of the worm, rather than ventrally, as in flies or vertebrates (Chisholm et al., 2016; Fig. 12I); this demonstrates that signaling pathways with highly conserved function can nevertheless be recruited to different (“nonhomologous”) positions to help orchestrate a pattern of nerve connections that is appropriate.

The Wnt signaling pathway, already mentioned in the context of patterning early embryonic events along the anterior-posterior axis, also determines the polarity of axon outgrowth along the ap-axis. This has been studied for the peripherally located sensory neurons, AVM/AVL and PVM/PVL, all of which are bipolar cells, with long, anteriorly extending axons forming efferent synapses, and short, posteriorly directed dendrites responsible for stimulus (touch) reception (Killeen & Sybingco, 2008; Ackley, 2014; Fig. 12L). Wnt signals (e.g., LIN-44, EGL-20, CWN-1, CWN-2) are required for the anterior growth of axons; loss of these signals resulted in posterior axonal growth or symmetric (anterior plus posterior) axon growth (Fig. 12M, N). Detailed genetic studies showed that, unlike Netrin or Slit, Wnt does not act as a directional cue that attracts or repels growth cones; instead, Wnt appears to localize protein complexes (including a complex that contains its own receptor, Frizzled/LIN-17) within the neuron in such a way that the cytoskeletal machinery drives axon extension in the proper direction.

Neural Development in Arthropods

Early Neurogenesis

In arthropods, two ectodermal domains become specified as neurectoderm by the expression of early transcription factors, such as SoxB in insects (Buescher et al., 2002). An anterior (procephalic or head) neurogenic region gives rise to progenitors of the brain, and a ventral neurogenic region produces the ventral nerve cord (Fig. 13A). The cellular mechanisms of early neurogenesis have been well described for representatives of the different arthropod clades and differ considerably. In onychophorans, considered to be basal among the arthropods, neural progenitors delaminate from the neuroectoderm (Eriksson & Stollewerk, 2010; Fig. 13B). Subsequently, progenitors appear to divide symmetrically into intermediate neural progenitors, which in turn give rise to postmitotic neural cells ( Mayer & Whitington, 2009; Eriksson & Stollewerk, 2010; Fig. 13C, D). Neural progenitors of onychophorans appear to be quite variable in pattern, in contrast to the situation in other arthropods. Thus, in insects, neural progenitors arise at positions that are genetically prespecified by the expression of proneural genes (Campos-Ortega, 1995; Bertrand et al., 2002; Hartenstein & Wodarz, 2013; Fig. 13E, F). The specification is accomplished by the “prepatterning genes,” whose domains of expression form an orthogonal system of anteroposterior and dorsoventral stripes. The three genes conveying positional information along the dorsoventral axis, vnd/Nkx, ind/Gsx, and msh/Msx, are highly conserved and were found in animals from cnidarian to vertebrates (Cornell & Ohlen, 2000; Arendt et al., 2016; Fig. 13F). The procephalic neurectoderm gives rise to approximately 100 neuroblasts that form the central brain. The ventral neurectoderm is organized segmentally, with each segment giving rise to an identical segmental set, called neuromere, of approximately 60 neuroblasts (Hartenstein & Campos-Ortega, 1984; Doe, 1992; Hartenstein & Wodarz, 2013). Neuroblasts do not delaminate all at once, but form several groups (S1–S5) which move in sequentially following a tightly controlled pattern (Fig. 13G, H). In addition to the bilaterally symmetric pattern of neuroblasts, neurectodermal cells located directly along the midline of the embryo (“midline cells”) form neural and glial progenitors of a special kind.

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Figure 13 Early neurogenesis in Arthropoda. (A–R) Key events of early neurogenesis in Onychophora (A–D), Hexapoda (E–J, O–R), and Chelicerata/Myriapoda (K–N). Upper panels of left and upper middle panel sets (A, E, K) represent schematic sagittal section of an early embryo at the onset of neurogenesis. Panels below show schematic cross sections of neurectoderm (blue) at sequential stages of segregation of neural progenitors (purple), intermediate progenitors (yellow), and neural precursors (orange). Panel set (O–R) shows early development of Drosophila optic lobe. Optic placode invaginates from embryonic ectoderm (P) and forms two neuroepithalial layers (blue) at surface of larval brain (shown in dorsal view in O). Neuroepithelium grows by symmetric mitosis (P and Q) and transforms into asymmetrically dividing neuroblasts (purple; Q and R) following a linear gradient (arrow in Q). (F) shows lateral view of postgastrula Drosophila embryo; expression pattern of proneural genes (purple) and prepatterning genes is shown in different colors. (J) is a schematic cross section of Drosophila neurectoderm and neuropblasts, illustrating disposition of molecular factors controlling spindle orientation and polarity of neuroblast division. (S–U) Origin of glia in Drosophila. Schematic cross sections of embryo (S: early stage, shortly after neuroblast/neuro-glioblast delamination; T: intermediate stage during blast proliferation; U: late stage, following neuron/glia differentiation). Different types of glia are shown color-coded.

After delamination from the neuroectoderm, neuroblasts of insect embryos proliferate in what is known as a stem cell mode (Fig. 13H). Delaminated neuroblasts divide asymmetrically into one large and one small daughter cell with very different fates. The large cell (still called a neuroblast) continues dividing in the stem cell mode for a variable number of rounds divisions. The small cell, called a ganglion mother cell (GMC), typically divides only one more time, after which its two daughter cells become postmitotic and differentiate into neurons or glial cells. Recent data demonstrated that this type of proliferation that generates GMCs (“type I proliferation”) occurs only during the beginning of neuroblast proliferation; at later stages, neuroblasts switch to a “type 0” pattern that directly produces postmitotic neurons (Baumgardt et al., 2014). Since the mitotic spindle of neuroblasts is typically directed perpendicular to the plane of the neuroblast layer, ganglion mother cells and immature neurons form a stack on top of the neuroblast from which they originated (Fig. 13H, I). In this manner, the entire progeny (lineage) of a neuroblast remains spatially close to each other.

The mitosis performed by the GMC is also asymmetric, generating an “A” daughter neuron and a “B” daughter neuron (Truman et al., 2010; Fig. 13H). The group of “A” neurons and “B” neurons form what has been termed “A” hemilineage and “B” hemilineage. Within a lineage or hemilineage, neurons are further divided into smaller units of cells born successively within a given time interval. These units, called sublineages (Fig. 13H), are specified by transcription factors that are expressed in a defined temporal sequence by the dividing neuroblast (Kohwi & Doe, 2013).

Asymmetric neuroblast division is controlled by cytoplasmic protein complexes which are localized in a polarized manner in the neuroblast (Betschinger & Knoblich, 2004; Wodarz, 2005; Lu & Johnston, 2013; Fig. 14J). These complexes, which also play a central role in asymmetric divisions outside the nervous system (many of them were originally discovered as factors controlling early cleavage divisions in C. elegans; Bowerman & Shelton, 1999), include the Par3 and Insc complexes that are localized at the apical neuroblast cortex and control the vertical orientation of the mitotic spindle, as well as the basal trafficking of other proteins, including Miranda (Mir), Numb, and Prospero (Pros). As a result, following mitosis, only the basal daughter cell (i.e., the GMC) will receive these proteins, which block further proliferation and trigger neuronal differentiation.

Development of the Nervous System of InvertebratesClick to view larger

Figure 14 Axonal pathfinding and target recognition in Drosophila. (A) Schematic drawing of lineages/hemilineages derived from individual neuroblasts. Neurons of a lineage form distinct neurite bundles, where later born neurons project their axons along early axons. Axon bundles and neurite arborizations of individual lineages scaffold the compartments of the neuropil. (B) Digital 3D model of two neuromeres of embryonic ventral nerve cord (dorsal view; anterior to the top; left: stage 13; right: stage 14), showing individual identified neurons pioneering tracts and peripheral nerves (after Nassif et al., 1998). The first longitudinal axon tracts are laid down by neurons that descend from the midline (MP1) and from two neuroblasts adjacent to the midline. The first axons to grow out are from pCC and vMP2. A similar pair of fasciculating pioneer axons grows out from MP1 and dMP2 in posterior direction. Eventually, the pCC/vMP2 and the MP1/dMP2 pairs meet and fasciculate with each other over a short distance. Upon reaching their serial homologs in the posteriorly adjacent segment, MP1/dMP2 fasciculate with these axons. Axons of pCC and vMP2 do the same with their homologs, so that two longitudinal fascicles are formed. Small bundles of pioneer axons similar to pCC/vMP2 and the MP1/dMP2 lay down a simple axon scaffold in the brain as well. Later differentiating fibres grow along this scaffold, completing the formation of the embryonic nervous system. (C and D) Positioning of neurite bundles of the ventral nerve cord by the Slit/Robo system. The repellant Slit is expressed along the midline (C; schematic cross section, showing longitudinal tracts [red] and Slit gradient [gray]). The distance from the midline a given axon maintains is determined by the combination of Robo receptors it expresses, as depicted in D. Expression of only the low-affinity receptor Robo (orange) allows for closer approach to the midline; concomitant expression of Robo and additional, high-affinity receptors (Robo3, purple; Robo2, green) causes axons to join the intermediate and lateral fascicle, respectively, which are further away from the midline (from Araújo & Tear, 2003, with permission). (E) Confocal image of ventral nerve cord; neuropil shown in red. GFP-labeled neuron (green) in wild type forms projection close to midline (arrowheads in left half of panel); overexpression of repulsive Netrin receptor (right half of panel) causes this neuron to project axon peripherally, away from midline (from Keleman & Dickson, 2001, with permission). (F) Line drawings of motor neurons innervating Drosophila longitudinal flight muscles at different stages (in hours after puparium formation; dorsal view). Note exuberant branching at 18 hr and 24 hr (from Hebbar & Fernandes, 2004, with permission). (G and H) Synapse formation in the Drosophila visual system. (G) Schematic cross section of retina with photoreceptors R1–R8, and lamina. Axon terminal of R1–R6 and dendritic branches of lamina neurons L1–L5 form modular units called cartridges. Part of one cartridge is shown at bottom of G. Principal synapse is the tetradic synapse formed between one of the photoreceptor terminals and dendritic branches of lamina neurons L1, L2, and Am (boxed; shown at higher magnification in H). (H) This shows sequential steps in the formation of the R-L1-L2-Am tetrad. Initially (left in panel), many synaptic sites containing two L1 endings, or two L2 endings, are formed (arrowheads). These sites are eliminated in a Dscam-mediated mechanism (red arrows), with the result that in the end, only sites with L1-L2 combinations are retained (from Millard et al., 2010, with permission).

In crustaceans, neuroblasts appear in similar numbers and show a similar arrangement as seen in insects, and cell lineage analysis suggests that at least some lineages could be homologous (Scholtz, 1992; Harzsch, 2003; Ungerer & Scholtz, 2008). The mode of crustacean neuroblast proliferation appears to differ slightly from that in insects: Rather than delaminating, crustacean neuroblasts remain integrated in the ectodermal epithelium; dividing perpendicularly to the plane of the epithelium, neuroblasts deliver their progeny interiorly. In myriapods and chelicerates, stem cell–like neuroblasts are absent. In these clades, the neurectoderm is divided up into many metamerically arranged “precursor groups,” which invaginate and subsequently dissociate into clusters of neurons and glia (Stollewerk et al., 2001; Dove & Stollewerk, 2003; Harzsch et al., 2005; Döffinger et al., 2010; Brenneis et al., 2013; Hartenstein & Stollewerk, 2015; Fig. 13K–N). In regard to number and pattern, precursor groups resemble the neuroblast lineages in insects and crustaceans. Thus, it appears that the expression of proneural genes, in chelicerates/myriapods and insects/crustaceans alike, defines small groups of neurectodermal cells as proneural clusters (Stollewerk, 2002; Stollewerk et al., 2003). In insects/crustaceans, each cluster “elects” a neural progenitor that subsequently divides and produces a lineage of neurons/glia. By contrast, in chelicerates/myriapods, proneural clusters remain epithelial and appear to grow by symmetric division (Fig. 13L); toward the end of this proliferatory phase, clusters invaginate and then dissociate into postmitotic neurons (Fig. 13L–N). Note that the invagination mode of early neurogenesis, encountered in chelicerates/myriapods which branch more basally than insects and crustaceans, has to be evolutionarily older than the neuroblast mode of the more derived insects and crustaceans (Harzsch et al., 2005).

Some parts of the nervous system of insects and crustaceans, notably the visual system (compound eye and optic lobe), for many species by far the most cell-rich part of the CNS, develops from proliferating neuroepithelia, rather than stem cell–like neuroblasts. This mode of neurogenesis may hark back to the invagination of precursor clusters that is the common mode of early neurogenesis in basal arthropods (see earlier). In hemimetabolous insects and crustaceans, a vertically oriented epithelial growth zone remains in the surface ectoderm of the embryonic head after neuroblasts of the central brain have delaminated/proliferated (Harzsch et al., 1999). One side of the growth zone divides in a directed manner, such that rows of differentiating eye ommatidia are “budded off” toward posteriorly. The other side produces progenitors of the optic lobe. These cells, similar to neuroblasts, divide asymmetrically, pushing their progeny interiorly toward the brain, where they differentiate as visual interneurons. In holometabolous insects (e.g., Drosophila), the growth zones invaginate and form the eye imaginal disc that gives rise to the compound eye, as well as a neuroepithelial “optic anlage” that forms the large number of neurons of the optic lobe (Fig. 13O, P). The epithelial optic anlage gradually transforms into neuroblasts (Egger et al., 2011; Joly et al., 2016; Fig. 13Q, R). Both the epithelium-to-neuroblast transition, as well as the subsequent neuroblast proliferation, follow a temporal gradient that translates into the directed growth of the eye and optic lobe; a similar temporally ordered growth has been observed in the vertebrate visual system and could be evolutionarily ancient (Joly et al., 2016).

Development of Glial Cells in Insects

Glial cells of vertebrate nervous systems are structurally defined by their processes that form sheaths around neuronal cell bodies, neurites, and synapses. On the basis of this definition, glial cells are sparse or even nonexistent in many basal taxa, suggesting that the types of glia that exist in derived taxa may not be homologous to each other or to vertebrate glia (Hartline, 2011; Verkhratsky & Butt, 2013). In derived invertebrate groups, among them arthropods, annelids, and mollusks, a considerable variety of glial cell types have been identified. The three main classes that have been defined in insects include glial cells that form sheaths around the entire nervous system and peripheral nerves (surface glia), those that ensheath the neuropile or axon bundles within the neuropile (neuropil glia), and those that form sheaths around individual neurons (cell body glia; Hartenstein, 2011; Freeman, 2015; Fig. 13U).

Insect glial cells are derived from progenitors that form both neurons and glial cells (“neuroglioblasts”); only a minor fraction of glial cells come from other progenitors (“glioblasts”) that are dedicated to the production of glial cells only (Fig. 13S). A distinctive feature of glial cells in the nervous system of insects, but most likely of many other taxa, is their widespread migration (von Hilchen et al., 2008; Yuva-Aydemir & Klämbt, 2011; Omoto et al., 2016). For example, in Drosophila, most glial cells are born from a small group of neuroglioblasts and glioblasts located laterally in the neural primordium. From this position, glial cells spread out in all directions to eventually cover the entire surface of the CNS. Other glial cells migrate through the cortex and form sheaths around the neuropil and neuronal cell bodies (Fig. 13T, U). Many glial cells remain mitotically active during their phase of spreading, even after they have differentiated.

Genetic studies in Drosophila have demonstrated that glial and mixed neuronal-glial progenitors are initially specified by the same mechanism as neuroblasts. However, two transcriptional regulators, Glial cells missing (Gcm) and Repo, are expressed in the (postmitotic or still dividing) progeny that embarks on the glial pathway (Jones, 2005; Soustelle & Giangrande, 2007; Altenhein et al., 2016). These factors inhibit neural genes. Specific glial determinants have so far not been identified in other taxa, including vertebrates. Here, the repo gene does not exist, and gcm appears to play a variety of roles, including development of the placenta and neural stem cells (Cross et al., 2002; Hitoshi et al., 2011). Various other transcription factors, among them Olig2, play an early role in glia, but also various types of neurons and other cells (Ono et al., 2009). This diversity in glial determinants is another indication for the presumption that glial cells evolved independently in different animal taxa.

Axonal Pathfinding

The formation of axon tracts has been studied in several arthropod species, and genetic analyses in Drosophila uncovered many of the molecular cassettes controlling axonal pathfinding (Evans, 2016). In Drosophila and other insects, neurons forming a hemilineage typically remain clustered together throughout development and project their neurites along a coherent tract, with later born axons following earlier ones (Larsen et al., 2009; Lovick et al., 2016; Fig. 14A). In many cases, one hemilineage undergoes programmed cell death, leaving a single cluster/neurite tract (Kumar et al., 2009; Truman et al., 2010; Lovick et al., 2016). Once tracts reach their destination, they differentiate and form axonal and dendritic branches (“arborization”). It has been documented for many (hemi)lineages that the spatial pattern of arborization is also very similar for many neurons of a given lineage, in particular those born during the later phase of neuroblast proliferation; lineages and their tracts represent a scaffold of connections that define the “macrocircuitry” of the insect nervous system (Lovick et al., 2013; Fig. 14A).

The orthogonal system of connectives and commissures (see earlier discussion) is laid down at an early stage by pioneer neurons, which are the firstborn neurons from several lineages. Pioneer neurons can be labeled by a number of global markers, such as the IG-like adhesion molecule Fasciclin II, as well as specific genetic markers; as a result, a detailed map of the individual neurons pioneering tracts of the central ganglia and the brain has been established (Goodman & Doe, 1993; Nassif et al., 1998, 2003; Hartenstein et al., 2015). The first axons forming the anterior and posterior commissure grow toward the midline at the level of two glial progenitors; pioneers of the posterior commissure grow slightly posteriorly. The first longitudinal axon tracts are formed by neurons that descend from neuroblasts in or adjacent to the midline of each segment (vMP2/aCC/pCC; dMP2/MP1; Fig. 14B). Axons produced by these neurons interact with a group of neuropil glial cells, the longitudinal glia cells. The resulting pioneer tracts prefigure the medial and intermediate longitudinal tracts (Fig. 14C–E); the lateral tracts appear slightly later. Similarly, small pioneer bundles prefigure the complex system of tracts in the brain.

The pattern of pioneer tracts is controlled by the signaling systems introduced at the top of the chapter (see earlier discussion). A number of signal/receptor pathways have been identified that appear to form a “coordinate system” of positional information used by axons to adjust their orientation and targeting. Most of the involved factors, including the Semaphorins/Plexins and the Slit/Robo system, act as repellants, where receptor activation causes growth cone collapse and the turning away of the corresponding axon from the signal presenting source (Yu & Kolodkin, 1999; Araújo and Tear, 2003; Dickson & Gilestro, 2006; Chilton, 2006; Evans, 2016). The repellant Slit is expressed by cells along the ventral midline and forms a mediolateral gradient (Fig. 14C, D). Most neurons express different types of Slit receptors, which are encoded by the roundabout (robo) genes. In Drosophila, three Robo receptors (Robo, Robo2, Robo3) with different affinity to Slit have been identified (Fig. 14D). Robo represents a low-affinity receptor, found on neurons with axons that do not cross the midline. In mutant embryos lacking Robo, these axons are able to cross back and forth multiple times. The high-affinity receptors Robo 2 and 3 are high-affinity receptors expressed by neurons whose axons extend along the intermediate and/or lateral tracts (Fig. 14D). Knock-out of Robo 2 or Robo 3 results in changed pathways, where an axon normally staying far away from the midline is able to approach closer (e.g., switch from lateral to medial connective).

The role of netrins in the insect nervous system parallels its function described for C. elegans. The netrin signal is expressed by the ventral midline. Receptors are able to mediate either attraction or repulsion. The receptor Frazzled (Fra; homolog of C. elegans unc-40) causes attraction. One mechanism of action is that upon Netrin binding, the cytoplasmic domain of Fra is cleaved off and acts as a transcriptional activator of the gene commissureless (com), which in turn inhibits the expression of Robo. Thus, Netrin and Slit/Robo signaling intersects with Netrin and Fra activation, neutralizing the Robo-mediated repulsion and allowing for crossing. The Netrin receptor causing repulsion, homolog of C. elegans unc-5, is expressed on motor neurons, whose axons grow away from the midline (Keleman & Dickson, 2001); overexpression of this receptor in axons that normally extend close to the midline causes these fibers to grow laterally, thereby leaving the CNS (Fig. 14E).

The Semaphorins/Plexins represent another system of repulsive cues. They were originally found in grasshopper, where the semaphorin G-Sema I is expressed in certain epidermal domains and functions to repel sensory axons from these domains, thereby directing the pattern of peripheral axonal pathways (Kolodkin et al., 1993; Bonner & O’Connor, 2000). In Drosophila, several semaphorins are expressed both in the CNS and subsets of muscle cells. A repelling function of these molecules has been shown in case of neuromuscular connectivity: Ectopic expression of semaphorins in muscle precursors prohibits motor axons from innervating these muscles. Semaphorins/plexins also order the dorsoventral architecture of the ventral nerve cord (Zlatic et al., 2009) and control several pathway choices in the brain (see later).

Connectivity

The final phase in neurogenesis entails the selection of specific synaptic partners from among a large number of potential targets. The typical scenario, in vertebrates and invertebrates alike, is a large number of growth cones of many different neurons colliding in a small space; growth cones emitting numerous processes that all interweave, probing each other for the presence of the proper molecular cues, and engaging in transient contacts; competition for synaptic space, and the lack of proper molecular recognition, resulting in the dissolution of many contacts, and the withdrawal of the processes; consolidation and differentiation of the remaining contacts into functional synapses.

The establishment of specific synaptic connections and underlying control mechanisms has been investigated in a number of insect species. Initial studies had to fight against the preconceived notion that connections in insects (and invertebrates in general) were less plastic than those of vertebrates, a notion likely based on the smaller cell number and often invariant cell types found in the insect nervous system (Murphey, 1986). However, the importance of activity for shaping neuronal connectivity in insects is amply demonstrated by studies in flies and various social insects (bee, wasp, ant). Neuropils of the brain involved in processing of sensory stimuli, notably the antennal lobe and mushroom body, showed significant decreases in volume, as well as number of axons and synapses number in animals reared in sensory and social deprivation (Withers et al., 1993; Farris et al., 2001; Jones et al., 2009). Several studies also docucmented that in individual neurons, competition for synaptic space plays as much a role in insects as it does in vertebrates; for example, experiments done in cricket (Murphey & Chiba, 1990) showed that after removing one presynaptic sensory neuron (“S1”) that normally shares synaptic space with another neuron (“S2”) on the membrane of a postsynaptic interneuron (“I1”), S2 sprouted additional branches, forming exuberant contacts with I1. Furthermore, the detailed reconstruction of the development of connections within the insect nervous system provided many examples where neurons initially formed an excess of branches that were later pruned. An example shown in Fig. 14F are the motor neurons innervating longitudinal flight muscles in Drosophila; during an early phase, many secondary and tertiary branches are formed, which are later lost prior to the appearance of the definitive synaptic boutons (Hebbar & Fernandes, 2004). As in vertebrates, electric activity plays an important part in the pruning and the selection of definitive synapses. Mutations in ion channels that rendered the Drosophila neuromuscular junction hyperexcitable resulted in an initial increase in motorneuronal branch density, followed by a much stronger degree of pruning. A recent study of the pruning that occurs during the metamorphosis of sensory dendrites in Drosophila pupae documented that local calcium transients precede the severing of individual branches, and that blocking these transients inhibited pruning (Kanamori et al., 2013).

The molecular mechanism controlling the formation of specific synaptic connections has also been investigated in a number of Drosophila neuropils, most prominently the optic lobe (Clandinin & Zipursky, 2002; Fischbach & Hiesinger, 2008; Sanes & Zipursky, 2010; Hadjieconomou et al., 2011). Insects have large image-forming eyes formed by repeated modules, called ommatidia. Each ommatidium has eight photoreceptor cells (R1–R8); of these, the outer photoreceptors R1–R6 are mainly involved in motion detection, the inner photoreceptors R7/R8 in color vision (Fig. 14G). R1–R6 project their short axons into the first optic neuropil, called lamina. R7 and R8 bypass the lamina and target the next compartment, called medulla. The modular architecture of the eye continues in the lamina and medulla (as well as the deeper optic lobe neuropils, lobula, and lobula plate). Lamina and medulla neurons form modules, called cartridges and columns, respectively, that match the number of ommatidia (Fig. 14G). Photoreceptors R1–R6 give direct synaptic input on two (L1, L2) out of several types of interneurons that make up each lamina cartridge, and the control of this specific connection has served as a paradigm for target recognition (Millard et al., 2007, 2010; Tadros et al., 2016). The large photoreceptor output synapses all have the same composition, whereby one receptor contacts one L1, one L2, and two other lamina interneurons. The lamina synapses are representative for the typical sensory output synapses found in insect brains: The sensory axonal terminal forms a large, rounded, or cylindrical end bulb (called glomerulus in some systems); on this bulb are found many (tens to hundreds) individual presynaptic sites, each site characterized by a presynaptic density, clustered synaptic vesicles, and a so-called synaptic ribbon (“T-bar”), which is formed by proteins involved in vesicle docking (Nieratschker et al., 2009; Wichmann & Sigrist, 2010). Each presynaptic site contacts multiple postsynaptic partners (polyadic synapses); in case of the photoreceptor synapse, there are four partners (tetrad).

During development of the tetrad synapse, the axons of one L1 and L2 fasciculate along the cylindrical endbulb of a photoreceptor (R). L1 and L2 then send out multiple short branches that sweep over the surface of R, forming transient contacts, many of which involve two L1, or two L2, targeting the same presynaptic site (arrowheads in Fig. 14H, left). These combinations, which do not exist in the mature lamina, are eliminated during later metamorphosis by a process of pruning (Fig. 14H, center, right) that involves the system of Dscam membrane molecules introduced for the process of “self-avoidance” of neurons at the head of the chapter. Loss of Dscam1 and 2 results in tetrads with frequent pairings of two L1 or two L2 branches (Millard et al., 2010).

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