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date: 29 February 2020

Morphology of Invertebrate Neurons and Synapses

Abstract and Keywords

Despite their often small numbers, the neurons in invertebrate nervous systems can nevertheless constitute many classes, and the nervous systems of little studied or entirely new species still offer significant opportunities for discovery. Circuit analyses and connectomic data are of particular significance, as are the relationships of these to behavior, and the organization of simple larval brains. Functional analyses of synaptic circuits still require knowledge of the neurotransmitter and neurotransmitter receptor for each identified neuron. Synapse complexity ranges widely; undifferentiated pathways in basal species may have unpolarized synapses with presynaptic sites opposite each other, and specialized pathways may have polyadic synapses.

Keywords: neuron, synapse, dendrite, axon, connectome, neuron size, neuron number, circuit, evolution

The title of this chapter incorporates a taxonomic paradox, one that distinguishes vertebrates, a rather coherent and relatively closely related set of chordate orders, from invertebrates, a diverse assemblage of all other animal groups, ~30 identified phyla, as well as invertebrate chordates. This paradox is only somewhat ameliorated by the recent upgrade of vertebrates to the status of a phylum (Satoh et al., 2014). Consequently, in tackling the topic of the morphology of invertebrate neurons, we have to anticipate the vast diversity of invertebrate forms, most with a central nervous system (CNS) or brain of some sort or other. The intensity of work on invertebrate nervous systems peaked half a century ago, driven by three major features of their neurons: that many have simple numerical proportions; that they are diverse in organization and composition and thus offer many different scientific opportunities; and that many have large neurons, favorable for impalement and landmark electrophysiological recordings. These advantages led to the development of a number of canonical invertebrate preparations, from the squid giant axon to the compound eye of Limulus, which have revealed many principles in neurobiology (Wiersma, 1967; Sattelle & Buckingham, 2006; Clarac & Pearlstein, 2007), while model genomic invertebrate species, especially Drosophila (Miklos, 1993; Venken et al., 2011) and Caenorhabditis elegans (White et al., 1986; Zhen & Samuel, 2015), have since revealed the molecular bases of many others. The structure of invertebrate neurons was the subject of an early summary (Hanström, 1928) and together with their function, the topic has been famously immortalized in a compendious two-volume treatise by Bullock and Horridge (1965), which has been summarized more recently (e.g., Meinertzhagen, 2010) but otherwise remains a pinnacle, unchanged for the last half century and in bad need of a major update. Neuron structure in invertebrates continues to offer enormous opportunities for discovery, especially aided now by greatly enhanced imaging opportunities.

The range of invertebrate species is vast: Many are marine, and most are abundant, readily available, cheap and expendable, and still relatively (p. 248) unstudied. Often valued for their simplicity, most invertebrate species are smaller and have fewer cells than vertebrates. Some are not necessarily simple in size or behavior, however, and the overall range of invertebrate nervous systems is enormous. To choose two examples at random, the complex brains of higher cephalochordates (for Octopus: Young, 1971) rival those of many fishes or more advanced vertebrates (Packard, 1972), while the rhopalial eyes of box jellyfish (Nilsson et al., 2005) are far more complex than their basal grade would suggest. At the other extreme lie examples like the numerically simple and determinate nervous systems of forms such as the rotifer Asplanchna (Ware, 1971), and the unspecialized basal nervous systems of cnidarians (Grimmelikhuijzen & Westfall, 1996) and ctenophores (Moroz, 2015). Nevertheless, many invertebrate brains are indeed simple and offer tractable alternatives to solve the complex problems of larger brains. Some exhibit constancy of cell number, as shown long ago in the nematode Ascaris (Goldschmidt, 1908, 1909) and the larvaceans Oikopleura and Fritillaria (e.g., Martini, 1909a, 1909b) and named eutely (Martini, 1909c). Some modern reviews provide morphological data on other less studied groups, such as tunicates (Mackie & Burighel, 2005), and marine invertebrate larvae are also an excellent source of simple brains in tiny bodies, well suited to connectomic approaches using electron microscopy (EM; Lacalli, 2009). Besides these, a search of the old German literature armed with some time and a dictionary can still be rewarded by the discovery of new nervous systems with yet more favorable features. Two recent examples for such new introductions are the tiny brain of the model pygmy squid, the first cephalopod brain to be viewed in whole-mount (Koizumi et al., 2016), and the 71-neuron photoreceptor to motor neuron circuit of the larval marine polychaete Platynereis (Randel et al., 2014, 2015). The latter study provides a library of cell types for the neurons of the visuomotor pathway, each with a rather simple morphology, reconstructed from EM series.


Let us consider first, what is a neuron? There are very many different manifestations of excitability in metazoan animals, and these must have arisen from different elements present even in bacteria when these converged on protists, sponges, cnidarians, and ctenophores (Meech & Mackie, 2007). Later, during the course of neural evolution, the condensation of neural cells into ganglia was accompanied by the segregation of different classes of neurons and glia. These features seem so obvious as not to require acknowledgment, but they raise the question of what it is that we mean when we refer to a neuron. Anctil (2015) gives an historical treatment for the quest to find the evolutionary origin of neurons among cnidarians and ctenophores. Certain as most neurobiologists are on this topic, the features that distinguish a neuron from cells of all other classes are in fact quite indistinct at the borders, for example in basal groups such as coelenterates and sponges, or between neurons and neuroendocrine cells. Thus, two cardinal features of neurons, electrical excitability and the secretion of neurotransmitters, exist in many other cell types. These include muscles of most forms in all species (for review, Hoyle, 1969); endocrine cells (for review, Ozawa & Sand, 1986); and epithelial cells in many species (e.g., those of coelenterates that produce action potentials: Mackie & Passano, 1968). Propagated action potentials generated by a sodium conductance mechanism occur even in sponges, which lack any morphological form of neurons (Leys et al., 1999). There is likewise overlap between the calcium-dependent mechanisms of transmitter release in neurons and those of neuroendocrine cells, such as chromaffin cells (Neher & Zucker, 1993). A third criterion for most neurons is the possession of long processes or neurites. Possession of a unitary axon, or other extended neurite, is possibly the least universal of all criteria, and various examples of anaxonal neurons exist, for example vertebrate hair cells and coronet cells in the larval CNS of Ciona (Ryan et al., 2016), while some neurons have bifid axons (see later). Even though these structural features may be distributed far more widely among different cell populations than in nervous systems, all three together—electrical excitability, the secretion of neurotransmitters, and the possession of long neurites—cluster within most neurons and provide the clearest triad of features to discriminate neurons from all other cell types.

During the course of early evolution, increase in neuron number was accompanied by the increased centralization of all neurons in ganglia and brains. Eventually these condensations became augmented by the addition of glia (see later), which are poorly reported in basal groups such as echinoderms and turbellarians, no less than are their morphology and interactions with neurons in more developed nervous systems. Thus, especially the glia of polychaete annelids and molluscs are rather poorly (p. 249) reported, whereas leeches, by contrast, have well-studied glia (Kuffler & Potter, 1964; Macagno, 1980; Kai-Kai & Pentreath, 1981; Dahl & Muller, 2014), with somata in ganglionic packets.

Structural Features of All Neurons

Two cardinal differences between invertebrate and vertebrate brains are, first, that the somata of invertebrate neurons dwell in a cortex or rind surrounding the neuropile, to which they contribute neurites; and, second, that the vast majority of invertebrate neurons lack myelin. These differences have far-reaching consequences for the organization of different brains. A correlate of soma position, the axosomatic placement of synapses in vertebrates, is often assumed to be a canonical vertebrate feature, but it is not invariably so. Conversely, some invertebrate neurons do have axosomatic synapses that are heterodox, as in the procerebrum (Zs-Nagy & Sakharov, 1969, 1970), or the peripheral intestinal nerve (Elekes et al., 1985) of pulmonate gastropods, or transiently in the developing abdominal ganglion of Aplysia (Schacher et al., 1979). All neuron types will be considered with the exception of photoreceptors, which are reviewed elsewhere (e.g., Land & Nilsson, 2002).

Insofar as the soma of an invertebrate neuron lies outside the neuropile it lacks the somatic dendrites that are the major site of synaptic integration in vertebrate neurons. Instead, integration occurs within the neuropile itself where neurites form both pre- and postsynaptic contacts, mixed promiscuously. As first promulgated by van Gehuchten (1891), Cajal formulated the rule that impulses propagate from dendrite to axon according to his law of dynamic polarization (Cajal, 1891). This rule holds for most neurons in the vertebrate brain, except notably the retina, but is widely violated in invertebrate brains (Bullock & Horridge, 1965; Meinertzhagen & Lee, 2012). Its presumed correlate is to generate synaptic currents close to the spike initiation zone and conduct impulses down the axon, a requirement that is diminished for neurons with short dimensions that do not spike, as for many in invertebrates. Cajal’s law of dynamic polarization required neurons with dendrites that are exclusively postsynaptic and axon terminals that are presynaptic, as indeed are most vertebrate neurons; synapses were inferred and circuits then constructed exclusively on this basis. And this perspective dominated neuroanatomy, the simple dichotomy between the synaptic roles of axons and dendrites challenged only by few neuron types, most notably dorsal root ganglion cells, or brain regions such as the inner retina (Dowling, 2012). For the better part of a century, conclusions were made largely by matching the depth relations between the terminals of input neurons and the dendrites of their presumed targets; this practice became transferred to arthropods and cephalopods. The practice may be true for particular terminals or dendrites, and in vertebrates, but it has been long gainsaid by many examples in invertebrates. In the particular case of Drosophila, now known from dense reconstructions, EM denies any such simple dichotomy in important details, with many terminals having postsynaptic sites and many dendrites that are presynaptic as well as postsynaptic (Takemura et al., 2008).

The second special case is myelin, which in vertebrates originated in chondrichthian ancestors (Bullock et al., 1984). Together with jaws, increased size, speed, and a more complex nervous system, myelin made the seas a whole lot less safe for invertebrate prey. Early chordates have naked axons: Myelin is absent in ascidian larvae (Katz, 1983), the lancelet (Lacalli, 2002), and in the hagfish and lampreys (Bullock et al., 1984). Amphioxus can dart with extreme rapidity and may therefore also be suspected to have fast conduction pathways, but nevertheless they lack oligodendrocytes that generate myelin (Lacalli & Kelly, 2002) and the gene for myelin protein P0 (Holland et al., 2008). Many small vertebrate axons retain an unmyelinated state, but many have an insulating sheath of myelin, identified in classical reports by its osmiophilia, negative birefringence, and saltatory conduction (Hartline, 2008).

For many years myelin was thought to be the special child of vertebrates, a major source of their success (Zalc & Colman, 2000) that increases the conduction velocity of impulses; contributes to a reduction in, and increased cellular density of, vertebrate nervous systems (Castelfranco & Hartline, 2015); and reduces the energetic costs of their neural activity (Sengupta et al., 2010). The nerve membranes of protostomes express glucose-containing glycosphingolipids, whereas glycosphingolipids from deuterostomes contain predominantly galactose (Okamura et al., 1985). Glial investments that show some similarities to vertebrate myelin also exist in at least four invertebrate groups: oligochaete annelids, penaeid and caridean decapod Crustacea, and calanoid copepods (Hartline & Colman, 2007). Like their vertebrate counterpart, these also have a multilamellar membrane wrapping and long myelinated segments (p. 250) interspersed with “nodal” loci where the myelin terminates and the nerve impulse propagates along the axon by “salutatory” conduction, peaking in velocity in penaeid shrimps (Xu & Terakawa, 1999). Although the cytoplasmic spaces of these glial wrappings differ from those in vertebrate myelin and vary considerably among themselves (Hartline, 2008), their structure indicates that myelin has arisen convergently on at least three occasions (Roots, 2008). In the first three groups the myelin arises as in vertebrates, from a wrapping by glia, whereas in the last group its origin is neuronal and not glial (Wilson & Hartline, 2011). In other invertebrate groups, the morphology of invertebrate glia and their interactions with axons are not well defined and may reward patient search, particularly in polychaete annelids and molluscs.

Given the lack of myelin in all other invertebrate axons, the requirement for rapid conduction along long nerves is supposed to have driven the increased axon diameters seen in certain escape pathways of invertebrate brains (Eaton, 1984). As a result, conduction velocity increased—although not necessarily in proportion to the square root of axon diameter, as proposed from purely theoretical considerations (Rushton, 1951). Examples include the much-studied giant axons that innervate the mantle of squid, up to 1 mm in diameter (Young, 1936), and medial giant fibers in the crayfish Procambarus, with a diameter of 200–250 µm that tapers to 100–150 µm (Johnson, 1924).

Neuron Numbers and Sizes Among Invertebrate Groups

The number of constituent neurons that populate invertebrate brains ranges over at least six orders of magnitude (Table 9.1), and it is usually not recorded with any accuracy. Indeed, for technical reasons it is rarely possible to count with any great accuracy cells in excess of a hundred or so, nor indeed particularly instructive to do so. As cell number increases, so does the number of interneurons, and usually their degeneracy. Each group has species that reveal a considerable range in cell numbers; for example, molluscs have probably the largest range with, of these, gastropods typically having the lower numbers, and cephalopod brains the upper, especially in Octopus (Young, 1971). Cell numbers vary considerably, especially with growth, increasing a thousandfold from hatchling to adult in Octopus (Young, 1963, 1971; Packard & Albergoni, 1970; Giuditta et al., 1971). Behavioral complexity is a poor correlate of these numbers in different species, and in basal groups it may be augmented by various forms of neuroid or epithelial conduction (Anderson, 1980), as for example in tunicates (Bone & Mackie, 1975; Mackie & Bone, 1976). Larvaceans are possible champions for doing most with least, as remarkably effective adult filter feeders that work with a CNS of only ~130 neurons (Søviknes & Glover, 2007), while among invertebrate larvae the trochophore of the polychaete Spirobranchus has only ~36 neurons (Lacalli, 1984).

This list of examples comes from studies that are often simple and descriptive, and while most are neither modern nor comprehensive, the numbers they report, if accurate, are lasting and remain valid forever, serving as groundwork to studies at greater depth. There is still room for studies that adopt new model species, for which a good example is the recent description of the brain of the pygmy squid from wholemounts (Koizumi et al., 2016), which offers one prospect to rejuvenate work on this important group.

The sizes of neuronal somata encompass a range of 104, from a diameter of ~1 µm in the smallest insects to ~1 mm for the largest cells in Aplysia. On the small side, miniaturization is associated with reduced numbers and small sizes of neurons (Polilov, 2015). In that case cell size generally decreases with increasing miniaturization until a lower limit is reached, with somata down to 2 µm in diameter, most of it occupied by the nucleus (Beutel & Haas, 1998; Polilov, 2015). In an extreme case, the smallest insects such as the tiny parasitic wasp Megaphragma have evolved anucleate neurons, in which over 95% of the neurons are reported to undergo lysis of their nuclei and cell bodies after the formation of the adult nervous system, before the adult emerges from the pupa (Polilov, 2012). This lysis greatly decreases the absolute and relative volume of the nervous system. Even so, the relative size of the CNS in the smallest insects is greater than that in larger ones. In extreme cases, the relative volume of the CNS can reach 12% in adults in Trichogramma evanescens (Polilov, 2012). In the mushroom body the density of synaptic complexes in highly polymorphic leaf-cutting worker ants limits brain miniaturization (Groh et al., 2014). The cells of the compound eye provide a model system for the study of brain miniaturization (Fischer et al., 2014). On the large size, gastropods such as Aplysia have 1 mm neurons easily visible to the naked eye, with growth cones that are claimed to be the largest in the animal kingdom (Lovell & Moroz, 2006). These contrast among molluscs with the procerebral (p. 251) (p. 252) neurons of pulmonates, which are very small and numerous (Chase, 2000).

Table 9.1 Approximate Numbers of Cells in Representative Invertebrate Brains



Neuron/Cell Number






196 brain cells


Ware (1971)




Chitwood and Chitwood (1950)

C. elegans

302 neurons


White et al. (1986)



6-~20 × 103 cells hatchling to adult


Ogawa (1939)


0.6–1 × 103 ganglion XXX hatchling to adult


Ogawa (1939)



~400 cells/ganglion


Macagno (1980)


389–398 cells, segment 10


Macagno (1980)


700 in two sex segments


Macagno (1980)







500 cells, 3rd abdominal ganglion


Kendig (1967)


~0.31 × 106 cells/hemi deuterocerebrum


Schmidt and Ache (1996)






0.42 × 106 cells


Guthrie (1961)


0.85 × 106 cells


Witthöft (1967)


0.34 × 106 cells


Strausfeld (1976)


1.0 × 105 cells (est)


Chiang et al. (2011)

Acheta terminal ganglion

2,100 cells


Gymer and Edwards (1967)

Carausius 2nd abdominal ganglion

663 neurons


Becker (1965)





Aplysia californica large abdominal ganglion



Coggeshall (1967)

Pulmonate CNS

250 × 103


Chase (1986)

Helix procerebrum

20 × 103


Ratté and Chase (2000)

Limax procerebrum

50 × 103


Gelperin and Tank (1990)

4.5 kg octopus

4 × 108


Young (1971)



Oikopleura dioica

120–130 neurons


Søviknes et al. (2005), Søviknes and Glover (2007)

Ascidian larva


177 neurons


Ryan et al. (2016)

~340 cells

serial LM

Nicol and Meinertzhagen (1991)

Ascidian adult



Mackie and Burighel (2005)


Amphioxus adult spinal cord

20 × 103 (estimate)

Nicol and Meinertzhagen (1991)

The Numbers of Neurons


The most sparsely populated examples range from numerically finite examples, such as the CNS in rotifers: 196 brain cells for Asplanchna (Ware, 1971); nematodes: 102–103 (Chitwood & Chitwood, 1950), with 302 identified in the C. elegans hermaphrodite (White et al., 1986), but see Varshney et al. (2011); to intermediate numbers in the CNS of annelids: 103–104 (Ogawa, 1939, who gives 6–~20,000 for the oligochaete Pheretima). Segmentation multiplies these numbers but less so the numbers of cell types, many of which are repeated metamerically. Definitive counts exist for the leech Hirudo with about 400 cells per ganglion, and in particular 389–398 for segment 10 and ca. 700 in the two sex segments; numbers vary by ± ~2% for the same ganglia in different individuals and species (Macagno, 1980). The larger numbers of cells documented in arthropods include those in crustaceans: about 105–106, with the closest recorded numbers of ~310,000 per hemibrain in the deuterocerebrum of the spiny lobster Panulirus (Schmidt & Ache, 1996); insects: 105–106, with an estimate of 0.42 × 106 for the hemipteran Gerris (Guthrie, 1961), counts of 0.85 × 106 from the honeybee Apis (Witthöft, 1967), and 0.34 × 106 in the housefly Musca (Strausfeld, 1976), or variously reported as about 100,000 (Chiang et al., 2011) or 135,000 (Kaiser, 2015) in Drosophila. Of the latter, the mushroom body contains roughly 2,200 neurons, including seven types of Kenyon cells and 21 types of output cells, as well as 20 types of neurons that are genetically qualified to use the neurotransmitter dopamine (Aso et al., 2014). Also in Drosophila each compound eye contains about 6,200 (776 ommatidia, each with eight) photoreceptors (Ready et al., 1976), whereas cell bodies, neurons plus glia, in the underlying optic lobe at the mid-pupal stage total about 27,000 for the second neuropile, or medulla, and 10,500 for the third neuropile, or lobula plate, of each side (Meinertzhagen & Sorra, 2001), in aggregate a clear majority for the entire CNS.

For individual insect ganglia, Gymer and Edwards (1967) give 2,100 cells for the terminal ganglion of Acheta, whereas Becker (1965) gives a numerically more modest adjusted total of 663 neurons for the second abdominal ganglion of the stick insect Carausius. Among mollusc brains neuron numbers range widely, between 103 and 108. Gastropods typically having the lower order of magnitude within this range (e.g., for the large abdominal ganglion of Aplysia californica: Coggeshall, 1967), and the upper range is set by cephalopod brains, with 106–108–4 × 108 given for the brain of a 4.5 kg octopus (Young, 1971). This list of examples comes from studies that are often simple and descriptive, and while neither modern nor comprehensive, the numbers they report remain valid and are backed by many other similar cases.


Cell numbers in protochordate deuterostomes are also of imaginable and therefore manageable (p. 253) proportions. Among urochordates the ascidian larva has for example ~340 counted accurately in larval Ciona intestinalis (Nicol & Meinertzhagen, 1991), of which about 177 are neurons (Ryan et al., 2016), counted in a different larva; adult nervous systems have far more, in the range 102–103 (numbers given in Mackie & Burighel, 2005). Larvaceans are simpler still, with only a hundred or so cells. Martini (1909b) gives 87 for Fritillaria, and Søviknes and Glover (2007) give further details for Oikopleura dioica. Among cephalochordates, the spinal cord of adult amphioxus has about 20,000 cells (Nicol & Meinertzhagen, 1991), although this number is an estimate and the larva has far fewer. Simple minded as it is, there are a host of qualifications to append to even such a cursory list. Some cases show, for example, large developmental changes, and counts that exceed about a hundred become problematic for methodological reasons.

Neuron Numbers Versus Neuron Types

Many nervous systems in invertebrates have few neurons, but these can often be identified from animal to animal, and in some cases are known in great detail. In groups having a more complex CNS, the morphological classes of neurons can be very precise and discrete, but characterizing this precision first required compilation of entire libraries of cell morphologies. These are well illustrated by traditional studies using Golgi impregnation on insects, especially flies (e.g., Strausfeld, 1970, 1971, 1976; Fischbach & Dittrich, 1989; Hanesch et al., 1989), and have been augmented by alternative methods, especially cobalt backfills and injections (Pitman et al., 1972), variously applied and modified (see Altman & Tyrer, and other chapters in Strausfeld & Miller, 1980). These methods reveal the morphological forms of selected neurons in their entirety, opening a window on the analysis of these and other tiny brains of invertebrates. Golgi impregnations in particular require skill to interpret, and all methods that reveal new classes of neurons of course fail to identify their unlabeled neighbors, which by definition are in a majority.

Cell size is not uniform throughout the brain. In insects most neurons have somata of rather uniform size, most typically 2–10 µm in diameter, but many nervous systems contain neurons of widely disparate sizes. For example, relative to the neurons of the mesocerebrum in the brains of pulmonate molluscs, the procerebrum has many small neurons, some 20,000 per lobe in Helix (Chase, 1986; Ratté & Chase, 2000) and Achatina each 5–8 µm in diameter (Chase & Tolloczko, 1993) and about 50,000 in Limax (Gelperin & Tank, 1990), their number being at least equal to the number of all other central neurons, whereas the mesocerebrum has fewer larger neurons (Ratté & Chase, 1997), the largest between 60 and 80 µm in diameter (Chase, 2000).

In the free-living polyclad flatworm Notoplana, neurons have a range of cell morphologies that form several groups, as reported from Golgi impregnation (Keenan et al., 1981), but these classes are very broad and loose. Historically, impregnation by the Golgi method has been the most widely used method, but when they work, the technically more demanding cell injection methods are equally powerful. For example, the morphology of leech neurons was first clearly seen only after injecting horseradish peroxidase, HRP (Muller & McMahan, 1976).

Earlier light microscopy reports, in Drosophila (e.g., Fischbach & Dittrich, 1989; Hanesch et al., 1989 and as cited earlier for other insect species, are now mostly supplanted by extensive libraries of reporter lines for cell types generated using especially the GAL4/UAS system (Brand & Perrimon, 1993). These drive the expression of markers such as green fluorescent protein (GFP), either in all cells or in small or single-cell clones to provide comprehensive libraries of cell types (e.g., Wolff et al., 2015 or Takemura et al., 2013, 2015; as expatiated later), with more comprehensive documentation in Shinomiya et al. (2011).

A significant problem is to know when all cell types have been identified. The chief strategem is to proceed until no new cell type appears, suggesting that the number of classes has thus become fully saturated. At least some insect neuropiles examined reveal that cells form discrete, discontinuous groups, but it is still harder to be sure that all cell (types) have been identified (see the Discussion by Fischbach & Dittrich, 1989). Comparing Drosophila medulla cells, for example, reveals 47 forms impregnated by the Golgi method (Fischbach & Dittrich, 1989), yet this falls far short of a total of at least 77 separate types so far identified by genetic single-cell lines, some (especially Dm and Mi cell types) with minor but consistent distinctions in their arbor; tangential Mt cells are further additions not included here and not easily recognized as Golgi impregnates. To these totals are added two cells seen by EM but by neither Golgi impregnation nor from a genetic reporter line (Takemura et al., 2013, 2015; Takemura, personal communication). To some extent these numbers can be seen as mere technical details, but (p. 254) the magnitudes of encountered variations in the detection of cell types, as well as the overall number of types (maybe >100 for all optic neuropiles) now covered in three saturation screens, could possibly represent what is to be found in other brain regions when their turn comes to be intensively sampled for cells with determinate morphologies. By comparison, only 17 morphologically distinct cell types so far identified using GAL4 lines in combination with a multicolor flip-out technique arborize in the protocerebral bridge of the central complex in Drosophila (Wolff et al., 2015).

Unlike studies based on light microscopy, the dense EM reconstruction of an entire neuropile, ganglion, or CNS reveals all cells within a reconstructed volume. Because they reveal all cells so that no cell can hide out undetected in the neuropile, such reconstructions may reveal some neurons not reported by other means, and thus incorporate various surprises. It may still require the imprimatur of genetic methods then to identify and endorse all cell classes definitively. CT1 is one such surprise: a novel wide-field complex tangential cell in the Drosophila optic lobe that arborizes over the facing strata of both medulla and lobula neuropiles (Takemura et al., 2016).

Some of the most clearly distinguishable classes of neurons are found in the most sparsely populated nervous systems, many of which exhibit eutely. These include the CNS of the ascidian larva, rotifers such as Asplanchna, certain nematodes such as C. elegans, and the larvae of various marine invertebrate groups. So far these have been comprehensively characterized in few species. For example, the 302 CNS neurons of a hermaphrodite C. elegans incorporate about 118 classes (White et al., 1986), albeit their cell total is only about 90% complete (Varshney et al., 2011). Based on their connections, considerable differences, however, are reported between the neuron classes of the bacteria-feeding C. elegans and the predatory nematode Pristionchus pacificus (Bumbarger et al., 2013). Fewer than half of the synapse classes identified in P. pacificus are found in C. elegans, and those in P. pacificus are more highly connected. In the C. elegans pharynx, 6 out of 14 neuron classes are considered interneurons because they lack motor synapses, whereas in P. pacificus 13 of 14 neuron classes are presynaptic to muscle cells. Only a single neuron class, I4, is anatomically an interneuron by this criterion in P. pacificus. These results suggest fundamental changes in the number and types of connectivity classes of neurons in P. pacificus compared with those of C. elegans (Bumbarger et al., 2013). There is, in fact, a surprising degree of neuroanatomical variation both within and among different nematode groups, variation for example in the numbers of neurons in the ventral nerve cord (Han et al., 2016) that would not have been predicted from the constancy of cell number in C. elegans and the close similarity between C. elegans and Ascaris.

Along with the growing number of identified features of the Drosophila nervous system, the CNS of C. elegans is frequently compared to neurons in vertebrate brains, as if these constitute equal arms of neural evolution. In fact, C. elegans and Drosophila are both ecdysozoan protostomes, and to compare them directly with vertebrates overlooks many examples in the second entire superphylum of Lophotrochozoa, which includes important annelid and mollusc models. Likewise vertebrates are a single phylum of deuterostomes (Satoh et al., 2014) with many specialized features, especially in having brains with huge numbers of neurons. Tunicates are a widely canvassed sister phylum to vertebrates, and their CNS is now known in some cellular detail in the larvae of ascidians and offers a far more balanced deuterostome counterpart to that of C. elegans (Nicol & Meinertzhagen, 1991; Ryan et al., 2016). In particular, the larva of C. intestinalis and the hermaphrodite of C. elegans have nervous systems with comparable cell numbers (Nicol & Meinertzhagen, 1991). Thus, in addition to the case of the nematodes quoted earlier, three tadpole larvae of Ciona intestinalis, the sibling progeny of a single mating, have been reported to exhibit a CNS complement of 331–339 cells (Nicol & Meinertzhagen, 1991). Of these, a fourth unrelated larva had similar cell numbers, including 177 neurons, each with an axon, that comprise at least 25 different types (Imai & Meinertzhagen, 2006; Ryan et al., 2016) and 52 subtypes (Ryan et al., 2016), although some of the latter are not discrete, clearly distinguishable by all morphological features. In the 36 cells of the larval nervous system of the Spirobranchus trochophore larva, including the tuft cells and the pigment cell of the eye, there are between 16 and 18 demonstrably different cell types (Lacalli, 1984). Thus, in all cases there are 2–3 cells per morphological class of neuron.

Among larger nervous systems, no comprehensive attempt has been made to identify morphological correlates of the large number of types, about 100, for which an electrophysiological signature is reported from the circumoesophageal connective of the crayfish Procambarus (Wiersma, 1958). These (p. 255) include receptor neurons, and compared with a total of 84 types in the connective tissue between the third and fourth abdominal ganglia, each is electrophysiologically distinct (Wiersma & Hughes, 1961), for which by comparison the third ganglion is reported to contain only ~500 cells (Kendig, 1967) or the anterior abdominal ganglia to contain 600–700 neurons (Kondoh & Hisada, 1986). Segmentation increases the numbers of cells more than the numbers of cell types, because many cells are serially homologous, repeated metamerically.

The crustacean stomatogastric ganglion is an extensively characterized nervous system in miniature. An early account in the decapods Callinectes sapidus, Homarus americanus, and Panulirus argus reports the relative constancy of cell number and type. Thus, the stomatogastric ganglion of the crab Cancer borealis has 25–26 neurons (Kilman & Marder, 1996), 4–5 fewer than in 13 serially sectioned Panulirus ganglia, in which two had 30 cells, seven 29, and four 28 (Maynard & Dando, 1974). In a 500 g lobster, the wholemounted ganglion was about 1.1 mm long, with neurobiotin coupled neurons numbering an average of 30.9 ± 0.7 (SD) (Bucher et al., 2007). Most neurons are members of a single class (types AB, LP, VD, IC, Int1, DG, LG, MG, and AM) or with just two copies (types PD, LPG), and 10 to 13 PY and GM neurons, with varying numbers of each type, plus 6 other neurons. All cells but one are monopolar. Despite the electrophysiological constancy of each member, the morphology of a specific neuron type can be quite variable from animal to animal (Bucher et al., 2007).

In addition, there are many examples of specific classes of neurons, either single neurons or those with multiple members having a morphological signature that has been particularly well documented. They include but are not limited to the locust descending contralateral movement detector, or DCMD (O’Shea & Williams, 1974; Rind, 1984); various neurons of the locust thoracic flight motor (e.g., Watson & Burrows, 1982; Watson et al., 1985); the many modular representatives of optic lobe neurons, for example the 12 interneuron classes of the first optic neuropile, the lamina, especially L1 and L2 (Tuthill et al., 2013); or medulla cells such as Tm2 (Meinertzhagen et al., 2009); and the lobula plate motion-sensing horizontal (HS) and vertical (VS) system tangential cells of the blowfly (Cuntz et al., 2008). Cluster analyses of the latter indicate how tightly such neurons constitute a class, different from the members of each other related class.

The Concept of an Identified Neuron

The incorporation in nervous systems of relatively large numbers of cell types each with relatively few individual cells lead, early on, to the recognition that in many invertebrate species each type is sufficiently discrete to be recognized from animal to animal, as an identified neuron (Kandel, 1976; Bullock, 2000; Comer & Robertson, 2001). Many of these are known in great detail, each by its size, shape, and network connections, like members of a European family. But it should also be acknowledged that in many invertebrate nervous systems having few neurons the neurons nevertheless lack morphological determinacy. Yet others may not be known in sufficient detail from multiple samples to make valid comparisons.

This cornerstone concept of the identified neuron underlies much that simple brains have contributed to studies in invertebrate neurobiology, but it is not restricted to invertebrates. Thus, vertebrate brains have, for example, Mauthner and other reticulospinal neurons that are also individually identifiable (Bullock, 1978). Lampreys, in particular, also have various identified Müller cells that are well characterized (Rovainen, 1979).

Traditional criteria by which an identified neuron is recognized include its electrophysiological signature (the chief criterion defining crayfish interneurons; see earlier), its neurotransmitter phenotype, and its morphological form (cell body position, arborization pattern, and other criteria by which, for example, fly optic lobe cells have been highly differentiated). Morphological features provide a subtle phenotype, often with high stereotypy in many studied invertebrate systems. Golgi impregnation (e.g., Cajal & Sánchez, 1915; Cajal, 1917) and vital stains such as methylene blue (e.g., Alexandrowicz, 1951; reviewed in Plotnikova & Nevmyvaka, 1980) were historical standards for such studies and are now augmented by dye fills and injections using Lucifer Yellow, HRP, fluorescent dextrans, diI, and so on (examples in Strausfeld & Miller, 1980; & Strausfeld, 1983). In Drosophila such methods are largely superseded by the use of genetic markers, especially by the use of the GAL4/UAS system of transgenes to drive cell-specific expression of reporter genes, such as that for GFP. One library of such markers now numbers more than 10,000 different lines (Shih et al., 2015), as part of the The FlyCircuit database of ~16,000 individual neurons (Chiang et al., 2011), and these confirm that the cell types recognized empirically by different morphological methods can be mimicked (p. 256) by genetic reagents, suggesting that morphological cell types are actually genetic cell types as well (Meinertzhagen et al., 2009). Genetic markers offer many additional advantages, providing a means to label specific neurons directly and repeatedly, either with fluorescent markers such as GFP, or—at the EM level—membrane-targeted HRP, as well as a means to construct comprehensive libraries of cell types that reveal the numerical and morphological range of cell types in the Drosophila brain (Meinertzhagen et al., 2009).

Evolutionary Origins of Neurons

The evolutionary origins of neurons are often revealed by their morphological features. Nervous systems provide structurally the most complex examples of biological organization, and networks of neurons have arisen through the concatenation of many ancestral networks along with remodeling of old circuits that supported now-extinct behaviors (Arbas et al., 1991; Katz & Harris-Warrick, 1999; Shinomiya et al., 2015).

The brains of invertebrate and vertebrate groups are said by some to have had a common origin in the ancestral bilaterian (e.g., Arendt & Nübler-Jung, 1999; Holland et al., 2013) and by others to have independent origins (e.g., Holland, 2003), while yet others invoke many origins. Thus, in the extreme case, complex brains are claimed to have evolved many times (Moroz, 2012) and to have acquired differentiated neurons along independent paths (Moroz, 2009). Each of these diverse invertebrate examples has, through its ancestors, successfully sustained life on Earth for a vast period of evolutionary history, and so can be considered a functional success no less than the more complex brains of vertebrates.

Until now most attention has focussed on seeking a transition from basal forms to those in which the brain’s structure attains a canonical stage (e.g., Arendt et al., 2008), with generally diffuse stages in neural organization progressively becoming more centralized to form a brain. The CNS takes precedence over the peripheral nervous system (PNS) in most accounts. Synaptic circuits for identified behaviors have typically been considered from either a developmental or functional perspective without reference to how the circuits might have been inherited from ancestral forms.

Neuronal isomorphs have been recognized in different members of the same invertebrate group. Examples among insects include lamina L-cells in flies (Shaw & Meinertzhagen, 1986; Fischbach & Dittrich, 1989). L1 and L2’s partnership has been highly conserved during the evolution of ancestral fly groups, but their input at photoreceptor synapses arose first at a dyad which became augmented by the incorporation of two additional postsynaptic elements to form the tetrad typical of more recent forms (Shaw & Meinertzhagen, 1986; see later). More extensive synaptic changes have occurred at other lamina cells (Shaw & Moore, 1989). Further afield, T1 in Drosophila and a decapod crustacean are remarkably isomorphic (Fischbach & Dittrich, 1989), and even T4 and T5, similar motion-sensing neurons in two neuropils that might have arisen by the duplication of a common ancestral cell type (Shinomiya et al., 2015). At one site, the photoreceptor tetrad synapse, the cluster of postsynaptic elements actually increased during the course of evolution of the Diptera, by the single-step recruitment of two amacrine cell processes to the ancestral dyad form of the synapse (Shaw & Meinertzhagen, 1986).

Identified neurons in gastropod molluscs provide quantifiable examples of neurons identified from both their number and the size and relative positions of their somata. Immunoreactivity to 5-HT (e.g., Newcomb et al., 2006) or to GABA (e.g., Gunaratne et al., 2014) has been widely exploited as a probe. Such accounts from various nudibranch species provide examples for this fertile area of descriptive studies and cite many others. Most initial studies emphasize the numbers and positions of somata, revealing the constant number and positions of many neurons. These are taken to support the common evolutionary origins of cells and cell clusters, while interspecies differences in the number of cells per cluster and even intraspecies variation in the clusters themselves indicate homoplasies. Although most accounts now document cells by means of confocal image stacks, most fail to provide additional morphological information on the arbors, or the relational data that link an immunolabeled neuron morphologically to its partners. Additional examples of the evolution of neurons are given by Katz and Harris-Warrick (1999).

Despite such examples, in general the same cell type has not been identified between different phyla unless by using very relaxed criteria. A promising place to look for such counterparts would be among the paired lobed or dome-shaped neuropils of the protocerebrum in mandibulate and chelicerate arthropods and the nonganglionic brains of polychaete annelids, polyclad planarians, and (p. 257) nemerteans, brain regions that share a structural ground plan with insect mushroom bodies (Wolff & Strausfeld, 2015).


Not strictly belonging to a consideration of the morphology of neurons and synapses but nevertheless warranting brief mention, invertebrate glia have their own parallel story. Basal groups lack clear glia, or at any rate lack reported forms. To this group, the acoelomorphs, interstitial flatworms with very simple cellular organization and currently at the base of the bilaterian phylogeny, do however possess glia-like cells, suggesting that glia had an early, evolution parallel to the neurons they surround, but basal taxa lacking convincing glia are found in urochordates, hemichordates, bryozoans, rotifers, and basal platyhelminths (Hartline, 2011). However, the larval ascidian Ciona has various CNS cell types that are not clearly neuronal or ependymal (Ryan et al., 2016). Historically, glia have mostly been identified by exclusion from the criteria used to recognize neurons. They were first tabulated by Roots (1978, 1981) and by Radojcic and Pentreath (1979), and recently updated by additional molecular criteria (Hartline, 2011). New glia are probably destined to come to light. Thus, Mashanov et al. (2010) identify a major cell type in the holothurian CNS that resembles vertebrate radial glia, and that might be more widespread in echinoderm nervous systems. Confirmation of the latter possibility awaits the development of new antibody markers. The role of glia in myelination is important in distinguishing invertebrate nervous systems from their vertebrate counterparts. This may well have begun in early deuterostome chordates. In the amphioxus larva, for example, a floor plate cell type is reported that might be a homologue of oligodendrocytes, but with a cell guidance rather than myelinating function (Lacalli & Kelly, 2002).

Glia are extensively reported in the medicinal leech Hirudo and other leeches. The abdominal ganglia each have 10 giant glial cells about 80 µm in radius; the geometrical arrangement of glial packets is generally invariant, and morphological features are reported in Kuffler and Potter (1964) and later studies. In both Haemopis and Hirudo six of the giant glia compartmentalize the neurons of the cortex into “packets”: two anteriolateral, two posteriolateral, and two ventromedial; two more giant glia each populate the neuropile core and connectives. Each packet contains 50–60 neurons with somata arranged several layers deep. These are grouped into large (>30 µm in diameter), more numerous medium (18–30 µm), and small (10–18 µm), the latter located in the inner layers. Neurons in a packet invaginate a single giant glial cell, processes from which ramify between the neurons (Kai-Kai & Pentreath, 1981). In the horse leech Haemopsis each abdominal ganglion is unusually large, with 12,000–14,000 small microglial cells in addition to the giant glial cells, and approximately 300 neurons (Kai-Kai & Pentreath, 1981). Four leech species show relative constancy in cell numbers and invariance in the geometry of their glial packets (Macagno, 1980). A system of extracellular channels continuous with the basal lamina penetrates the giant glial cells and partially extends around the neurons. These channels contain a matrix, and hemidesmosomes join the matrix-filled channels to both the neurons and neighboring giant glial cells; neuropile areas rarely contain glial cell processes (Kai-Kai & Pentreath, 1981).

Glia have received special treatment in Drosophila both from the perspective of their developmental origins and functions (Hartenstein, 2011). Based on the position and morphology of glial cell bodies in the larva, glia in the Drosophila ventral ganglion are classified as surface associated, cortex associated, or neuropil associated (Ito et al., 1995). In adult Drosophila, Repo-expressing glial cells also reveal the presence of three classes and five types in the central brain, the same as in the larval CNS. Comprehensive definition of glial cell types is available in the Drosophila lamina, where three main types are again surface, cortex, and neuropile glia (Edwards & Meinertzhagen, 2010); and glial ultrastructure has been reported (Trujillo-Cenóz, 1965). A specific organelle, the capitate projection (Trujillo-Cenóz, 1965), at which photoreceptor terminals receive invaginations from surrounding epithelial glia, is a proposed integrated site of membrane and transmitter recycling (Fabian-Fine et al., 2003). The same three classes of glia as in the optic lobe are also resolved in the antennal lobe in Drosophila (Awasaki et al., 2008) and other neuropiles contain similar glial subtypes, but unlike the antennal lobes, they have not been described in great detail. There are about 80 glial cells in each adult Drosophila antennal lobe, and other numbers are listed in Edwards and Meinertzhagen (2010). Related glial types are also reported in the antennal glomeruli of Manduca (Oland et al., 1999), but these differ from those in Drosophila, in which glial cells in the developing antennal lobe are unlikely to play a strong role in either axon sorting or glomerulus stabilization and, (p. 258) in the adult, do not electrically isolate neighboring glomeruli (Oland et al., 2008). Of the three cell types: (a) surface glia form the blood–brain barrier, regulating the flow of substances into and out of the nervous system, both for the brain as a whole and the optic neuropiles in particular; (b) cortex glia provide a second level of barrier, wrapping axon fascicles and isolating neuronal cell bodies both from neighboring brain regions and from their underlying neuropiles; and (c) neuropile glia can be generated in the adult and a subtype, ensheathing glia, are responsible for cleaning up cellular debris during Wallerian degeneration. Both the neuropile ensheathing and astrocyte-like glia may be involved in clearing neurotransmitters from the extracellular space (Edwards & Meinertzhagen, 2010). In the locust metathoracic ganglion seven major types of glial cell, with subdivisions, form invaginations into neurons of four different kinds: regular, chunky, filigree, and ridge (only at axon hillocks). Such types (Hoyle, 1986) reveal the limitations of purely morphological studies and are not easily related to those reported for other insect neuropiles. Glial cell morphologies have occasionally been reported from Golgi impregnation (Sánchez y Sánchez, 1935; Strausfeld, 1976; Cantera & Trujillo-Cenoz, 1996), and glial cell types are summarized for other insect groups in Cantera and Trujillo-Cenoz (1996). It is unlikely that different types of glial cell have been fully distinguished, even in arthropods in which they are known best. Two types of glia with different ultrastructures are identified in the buccal ganglion of the snail Helisoma (Berdan et al., 1987). Gap junctions formed by both glia and neurons do not differ discriminably. Improbably well-developed glia of several morphological types are reported in highly derived and parasitic cestode Platyhelminthes. Four types have been identified: multilamellar light-cytoplasm, cells of the main trunks, fibroblast-like cells secreting extracellular matrix, “sandwich” cells wrapping neuropile with alternating cellular and extracellular matrix layers, and even cells forming myelin-like structures (Biserova et al., 2010).

Immunocytochemical Detection of Neurons

Although not technically a morphological feature, neurotransmitters have been widely used to identify neuronal morphologies by means of immunocytochemistry. Wide-field neurons immunoreactive for candidate neuromodulators offer the most obvious examples, for example for 5-HT and various neuropeptides. Essentially all nervous systems examined express FMRF amide-related peptides (e.g., Elphick & Mirabeau, 2014; Zatylny-Gaudin & Favrel, 2014), which together with 5-HT provide excellent markers for early differentiation and neuron morphologies in wholemounted invertebrate larvae. Examples include gastropod veliger molluscs (e.g., Dickinson & Croll, 2003), with a relatively well-developed nervous system having 26–28 apical cells, 80–100 neurons in the CNS, and 200–300 peripherally located neurons by the late larval stage, the early phoronid actinotroch larva (Hay-Schmidt, 1990), and echinoderm dipleurula and pluteus (Byrne et al., 2007) larvae. A full listing is beyond the scope of this chapter, but FMRF amide, catecholamines, acetylated α-tubulin, and serotonin, either alone or in different combinations, are the bases of numerous larval studies, especially with respect to the apical organ. For the expression of immunoreactivity to serotonin in insects, see the following review (Nässel, 1987), while for neuropeptide expression in specific neurons of larval and adult insects, see the following (Nässel, 2002; Nässel & Homberg, 2006; Nässel & Winther, 2010).

Immunoreactivity of presynaptic elements to neurotransmitters has been exploited at favorable synaptic sites and for favorable antibodies. For example, the synaptic engagements of GABA-positive profiles are documented in the lip of the honeybee calyx (Ganeshina & Menzel, 2001). Antibodies against GABA and 5-HT are technically reliable, densely expressed, and as a result bestow particular advantages. Those against other neurotransmitters, such as the neuromodulators in insects, octopamine and tyramine (Kononenko et al., 2009), provide alternatives with sparser patterns of expression.

From a survey of mollusc species, Sakharov (1974) viewed transmitter specificity as one of the most evolutionarily conserved characteristics of neurons (Moroz, 2009). In general, invertebrate brains utilize the same neurotransmitters as those found in the brains of vertebrates, albeit with some exceptions. The latter includes expression of octopamine and tyramine, which are exclusive to invertebrates (Roeder, 1999), while acetylcholine is the most widely expressed neurotransmitter in the C. elegans CNS (Pereira et al., 2015) and also more widely expressed in arthropod brains than in the brains of vertebrates, for example among most sensory systems in insects and some other arthropods (Osborne, 1996). The documentation of expression types is far from complete, however, and would require an atlas of atlases.

The main features for insect neurons, their types, morphologies, and fine structure, have been (p. 259) selectively reported by Strausfeld and Meinertzhagen (1998), to which the interested reader is referred.

The Morphological Components of Neurons

The morphological features of a neuron—its soma, axon, and neurites; their relative sizes, placement, and branching—offer for some neurons a morphological signature which can be recognized uniquely, from individual to individual. In many cases, detailed knowledge of a neuron’s lineage, transmitter, morphology, and synaptic network has been recorded in intimate detail, like a member of one of Europe’s finest families. Organelles include not only synapses but also cilia.


In many invertebrate groups, neurons are monopolar, having cell bodies in a rind or cortex surrounding a central neuropile within which neurites arborize or are lateral to the neuropile in pulmonates. The cell bodies lack neurites and synaptic contacts, a major difference from the somata of vertebrate neurons. This pattern is typical of the arthropod, mollusc, and annelid groups from which most invertebrate preparations are drawn. In other groups, the segregation between somata and neuropile is less strict. In the CNS of larval Ciona, for example, individual neurons are mostly monopolar, but up to 42% have a soma with a single or few dendrites, and an axon that usually has a clear terminal (Ryan et al., 2016).


Typically this tends toward a unitary cylindrical form, but some neurons that mediate local circuits may lack an axon, as for anaxonal amacrine neurons in the insect optic lobe (Strausfeld, 1976). Axons may also have collaterals, while others may have a bifid axon. Examples of the latter occur in the larval CNS of Ciona (Ryan et al., 2016), in the adult larvacean Oikopleura (Olsson et al., 1990), and in the rotifer Asplanchna (Ware, 1971), all associated with small cell numbers; it is not clear whether such cases reflect the condensation of two ancestral cells or the division during evolution of the axon from a single cell to yield one with two axons.

Axon diameters range from large, up to about 1 mm in the case of squid giant axons, to small, down to 50 nm as recorded after chemical fixation for EM in the case of amacrine cell axons in the lamina in Drosophila (Meinertzhagen & O’Neil, 1991) or 100–150 nm in the larval polychaete Platynereis (Randel et al., 2014). Examples of large axons saw service in early conduction studies and include motor axons of Crustacea, especially decapod crustaceans, gastropods, and annelids, especially earthworms. Examples are listed in Bullock and Horridge (1965; their Table 3.4). Within arthropods, crustaceans have some large-diameter, mainly motor, axons. Axon size is more modest in insects, but even so insect ocellar second-order neuron axons are up to 25 µm in diameter in large dragonflies and some Hemiptera (Chappell et al., 1978; Meinertzhagen, unpublished). By comparison with these the smallest axon sizes in C. elegans are 0.2–0.5 µm (White et al., 1986) and in larval Ciona intestinalis axons are 0.3–1.0 µm in diameter (Ryan et al., 2016), while axons in Drosophila are typically at least 0.5 µm, but with the finest as small as 70 nm (Meinertzhagen & O’Neil, 1991), close to the limit that can support conduction (Llinás, 2003). For the most slender axons, a lower limit to neurite size is placed by ion-channel noise. At a diameter below 0.1 µm axons become inoperable because single, spontaneously opening Na+ channels generate rates of spontaneous action potentials that disrupt communication (Faisal et al., 2005).


Given the variable and often coextensive placement of pre- and postsynaptic sites over invertebrate neurons, many examples of invertebrate neuron violate the law of dynamic polarization proposed for vertebrates. In view of the resulting ambiguous status of dendrites, which as part of a vertebrate consensus should be postsynaptic, it seems safest to refer to all extensions of invertebrate neurons, even those arising from the soma, simply as neurites. These have variable branching patterns with a complexity of the arbor that differs considerably among the brains of different animals. It is generally simple in sparsely populated brains such as those of the rotifer Asplanchna (Ware, 1971), larval Ciona intestinalis (Imai & Meinertzhagen, 2006; Ryan et al., 2016), a larvacean Oikopleura dioica (Olsson et al., 1990), or C. elegans (White et al., 1986; Varshney et al., 2011), all with less than a few hundred cells. We may suspect that branching simplicity results from each neuron’s few input or output partners, but this correlation lacks clear analysis.


Looked at in sufficient detail, many neurons—possibly most—have cilia. These are best seen with tubulin antibodies in wholemounts or brain (p. 260) slices, and for sampling reasons are seen only rarely from EM in vertebrate brains (Fuchs & Schwark, 2004). As a result, most cases formerly were either not identified or were dismissed as idiosyncratic instances. Cilia contain a number of microtubules surrounded by a cell membrane, which together extend from a basal body within the soma. The microtubules (axonemes) exist in one of two common arrangements in cross section, 9 + 2, which possess dynein arms and are motile, or 9 + 0, which are primary cilia (Dutcher, 2003; Davis et al., 2006). Nearly all cells in vertebrates possess primary cilia (Marshall & Nonaka, 2006; Singla & Reiter, 2006), although the timing and nature of their existence may vary within and between cell types. Are cilia also universal for the neurons of invertebrates? As invertebrate deuterostomes, urochordates such as Ciona intestinalis help arbitrate this question, at least for chordates. Historically, it was believed that cilia within the Ciona larval CNS were limited to peripheral or specialized sensory neurons or to nonneuronal ependymal cells, which constitute a large population of CNS cells (Dilly, 1969; Meinertzhagen et al., 2004; Konno et al., 2010). A complete connectome derived from serial-section EM now reveals that of the ~177 neurons in the larval CNS, 120 are ciliated, and all remaining cells appear to contain basal bodies, even if they may lack canonical ciliary extensions from the soma. Of the 120, 14 project to juxtaposed membranes, 14 with synaptic-like structures, 11 to vacuoles near their membranes, and 18 to internalized vacuoles that may contain extracellular material. Thus, no neuron completely lacks a ciliary component, even though the extension of a ciliary shaft into the extracellular space is lacking in 80 neurons and instead most ciliated neurons extend cilia into the cerebrospinal fluid of the neural canal. This universality is consistent with the features reported for various vertebrate nervous systems, and it suggests that primary ciliated neurons within the CNS of other invertebrate deuterostomes, such as echinoderms, may simply have been overlooked.

Arendt and Nübler-Jung (1999) argue that the CNS in both protostomes and deuterostomes arose from a single ancestral centralization of ectoderm. Regardless of one’s position on this point, in chordates this centralization occurs during neurulation, in which the ectoderm invaginates as a whole. Garstang (1928) first proposed that the dorsal tubular nervous system evolved from two halves of a ciliated band in an auricularia-like larva of the kind found in echinoderms and hemichordates. Initially overstated, his argument is increasingly supported by recent evidence. According to his interpretation, the presence of cilia in the chordate CNS was probably inherited as an ancestral feature. In protostomes, a ventral nerve cord arises when ganglionic masses detach from the ventral neuroectoderm to form a ladder of connectives and commissures (Arendt & Nübler-Jung, 1999). Was this neuroectoderm ciliated? No clear answer can be given, even though it would be relatively simple to label cilia in embryonic or larval wholemounts of selected protostomes. However, for two canonical protostomes the lack of universal cilia in cells of the CNS in both Drosophila (Basto et al., 2006) and C. elegans (Hall, 2014: may instead prove a specialty of ecdysozoans.


Synaptic Ultrastructure

Studies on synaptic ultrastructure are legion. Most accounts have used thin-section transmission EM, although freeze-fracture studies are available for a number of synapses—for example, those in the cockroach metathoracic ganglion (Wood et al., 1977) and the antennal lobe of the moth Manduca sexta (Tolbert & Hildebrand, 1981); fly photoreceptor tetrad synapses (Fröhlich, 1985; and squid giant synapses (Pumplin & Reese, 1978). Synaptic ultrastructure is particularly clear in arthropods: not only insects (Armett-Kibel et al., 1977; Leitinger et al., 2012) but also in other arthropods, crustaceans (Hafner, 1974), and spiders (Trujillo-Cenóz & Melamed, 1967; Fabian-Fine et al., 1999, 2002). Early standards for studies on insects in particular were set by early EM descriptions from the group of Akert (Steiger, 1967; Lamparter et al., 1969) and for the fly’s lamina by Trujillo-Cenóz (1965).

Synaptic ultrastructure has been reported in many descriptive studies conducted especially during the 1970s. It is mostly no longer necessary to wade through these detailed studies, which have been extensively reviewed elsewhere, for coelenterates (Westfall, 1996; Kass-Simon & Pierobon, 2007) and many other invertebrates (Cobb & Pentreath, 1978), and notably for locusts and other insects (Watson & Schürmann, 2002), especially Drosophila (Prokop & Meinertzhagen, 2006). The review by Cobb and Pentreath (1978) highlights the great difficulties in documenting most synapses in many, chiefly marine, invertebrate species. In most such reports, synaptic ultrastructure is lacking, indistinguishable, or unspecialized, and diffusion fixation is insufficient to capture structure (p. 261) that is clear or distinctive. Molluscs, in particular, represent something of an anomaly. Thus, although famous for the size of its neurons and the range and diversity of their synaptic mechanisms, the appearance of synapses in the opistobranch sea hare Aplysia is nevertheless disappointingly nondescript. For all Aplysia has told us about synaptic physiology, it has done so with synapses that are structurally very poorly differentiated monads (Coggeshall, 1967) and for which synaptic morphology has been largely reported in the context of various forms of memory (e.g., Bailey & Kandel, 2008). In a sample of ~100 synapses in Aplysia, only four serial synapses were identified, all small, perhaps indicating a relative impoverishment in network richness; in the same sample, 40% of the postsynaptic profiles also contained vesicles similar to presynaptic vesicles (Graubard, 1978). An exception that occurs at the neurites of mechanoreceptor sensory neurons is provided by the indented synapses of Aplysia, which have postsynaptic invaginations of the presynaptic terminal (Bailey et al., 1979; Bailey & Thompson, 1979). In annelids, the best accounts of synapses are those of Schürmann (Gunther & Schürmann, 1973; Schürmann & Günther, 1973) on earthworms, Muller and McMahan (1976) for the leech, and Schürmann (1978) for Onycophora. More recent studies quantify synaptic ultrastructure, for example in the larval polychaete Platynereis (Randel et al., 2014) and the leech Hirudo (Pipkin et al., 2016), in which larger vesicles containing presynaptic profiles are juxtaposed by two or more smaller postsynaptic processes. Some leech neurons segregate their arbors into exclusively input and mixed input/output regions, while others have exclusively mixed input/output neurites; yet others have exclusively input neurites.

Unlike many terrestrial species, especially arthropods, synaptic ultrastructure is far less clear and less highly differentiated in marine invertebrates. The nervous systems of many marine invertebrate larvae, in particular, are diffusely organized and lack clear synapses (e.g., Lacalli, 1984, 1990). However, it is not clear whether this absence of evidence can fully be taken as evidence of their absence, given the rather flimsy appearance typical of synapses in basal nervous systems, and that the conditions for EM fixation may be critical in preserving synaptic vesicles. Indeed, the primary literature on synaptic ultrastructure needs careful interpretation, especially with respect to the effects of EM fixation and the tilt angle of sections; cellular fine structure in general, and synaptic ultrastructure in particular, is generally far less well preserved in marine species than terrestrial (Cobb & Pentreath, 1977). In general, a correlation exists between the speed of a species’ behavioral repertoire and the structural differentiation of its synapses, but this correspondence lacks serious analysis and may reward further examination. One might suspect a correlation between slowness of movement, especially in most gastropods, and lack of synaptic differentiation, but this is apparently denied by fast-moving cephalopods such as octopus, which also has simple synapses lacking ultrastructural specializations, especially in having only a single postsynaptic element rather than dyadic synapses (Gray and Young, 1964; Woodhams, 1977). An exception are the vesicle-filled bag endings of photoreceptors (Dilly et al., 1963), which make synaptic contacts with small invaginating spines from postsynaptic granule cells, and with tunnel fibers from the outer granule cells running through channels in the synaptic bags.

Synapse complexity ranges widely; undifferentiated pathways in basal species may have unpolarized synapses with presynaptic vesicles on either side of a synaptic cleft, as is the case among ascidians, for larval Ciona (Ryan et al., 2016). Among other deuterostomes, the report of Cobb and Pentreath on starfish, as well as on gastropod molluscs (Cobb & Pentreath, 1977), documented in a joint review by these authors, councils care in interpreting the rather unspecialized structure of chemical synapses compared with synapses in vertebrates (Cobb & Pentreath, 1978). Collectively these accounts may lack more recent molecular perspectives, and indeed these are treated elsewhere (e.g., Zhai & Bellen, 2004; Chia et al., 2013; Ackermann et al., 2015).

Presynaptic Organelles

Presynaptic Active Zones

Although most invertebrate synapses lack clear presynaptic specializations, a range of organelles signifies release sites used by synaptic vesicles. At some these incorporate features of vertebrate ribbon synapses (Sterling & Matthews, 2005), for which photoreceptor synapses in insects were named (Fröhlich & Meinertzhagen, 1982; Nicol & Meinertzhagen, 1982) that have been renamed T-bars in Drosophila (Zhai & Bellen, 2004; Prokop & Meinertzhagen, 2006; Takemura et al., 2013). Not all synapses have such organelles, however. In the mushroom body calyx of Drosophila, for example, the large, ChAT-positive (cholinergic) boutons of antennal lobe projection neuron have three forms—unilobed, clustered, or elongated—and (p. 262) have clear- or dense-cored vesicles and those with a dark cytoplasm having both (Butcher et al., 2012). The boutons are 2–7 µm in diameter, the largest identified cholinergic synapses in the Drosophila brain (Yasuyama et al., 2002), and are enwrapped by the actin-enriched, claw-shaped endings of Kenyon cells (Leiss et al., 2009. Each forms synapses, membrane densities with or without a T-bar ribbon; all bouton types have more synapses with T-bar ribbons than without, especially the dark boutons, and ribbon synapses are larger and with more postsynaptic elements (2–14) than non-ribbon (1–10) synapses (Butcher et al., 2012).

Synaptic Differentiation

In the snail Helix aspersa, procerebral neurons with different sites of arborization have patterns of synapse distribution that are distinct (Ratté & Chase, 2000), so that although the synapses themselves are poorly differentiated they are obviously highly regulated. By contrast, synaptic differentiation is particularly clear in flies, especially synapses that have T-bar presynaptic ribbons, but at most invertebrate synapses such organelles are lacking, and the presynaptic site marked by a simple cumulus of vesicles. Vesicle diameters range from 30 to 80 nm for clear vesicles and much larger, typically up to 180 nm, for those with dense cores. Their definition was a major preoccupation of early synaptic ultrastuctural studies, one that provides an entire chapter in itself, but one that is frustratingly inconclusive. Examples are listed for different invertebrates in Cobb and Pentreath (1978). Adequate fixation is one problem in vesicle preservation, as shown at the stretch receptor organ of the crayfish (Tisdale & Nakajima, 1976).

Postsynaptic Composition

Divergent Polyads

Even though many synapses are monads, having a single postsynaptic element opposite the presynaptic site, many others are polyads and thus divergent. Dyad and triad synapses, which in vertebrates are heterodox and known mostly only from the retina (Dowling & Boycot, 1966; Sterling & Matthews, 2005), are widely distributed in invertebrate nervous systems, and higher order polyads are also recorded. In the antennal lobe of the moth Manduca, Tolbert and Hildebrand (1981) report two to seven postsynaptic elements juxtaposed to the presynaptic site in pairs, so that in cross section the synapse appears a dyad. Multiple postsynaptic elements appose the presynaptic element in pairs, so that in cross section the synapse looks like a dyad. The report of Tolbert and Hildebrand (1981) summarizes the literature at that time, with most details still current.

Synaptic polyads are probably a basal condition for some synapses in many nervous systems. Even the neurons of nervous systems, such as those of the peripheral parts of the plexus of the marine polyclad flatworm Notoplana (Koopowitz & Chien, 1975), or the dense network of unmyelinated fibers in the neuropile of the brain in the turbellarian Microstomum (Reuter, 1981), have many dyad (“shared”) synapses, indicating that ancestral dyads were already present at this grade of neural evolution. Leech synapses are often dyads (Muller & McMahan, 1976) and C. elegans has many dyads, especially at neuromuscular junctions, and some triad synapses (White et al., 1986). A detailed record from a single larva in Ciona reveals that of the 8,601 synapses between neurons in the larval Ciona CNS, 921 were polyadic, with multiple postsynaptic elements. Considering only those between CNS cells, so excluding peripheral synapses and synapses onto muscle or the basal lamina, reveals that ~14% of the >6,600 synaptic contacts are divergent polyads, having multiple postsynaptic elements opposite a single presynaptic site (Ryan et al., 2016). Of those cases, 93% were dyads. On the other hand, the synapses of molluscs lack dyads (Dr. Craig Bailey, personal communication).

Elsewhere in invertebrate brains, synaptic dyads are widespread. In insects such synapses are probably a majority, with few synapses having a single postsynaptic element. In the CNS of adult Drosophila, most synapses are divergent polyads with multiple postsynaptic elements, mostly two or three in the mushroom body output lobe (Takemura, personal communication), but more than 10 at ribbon synapses in its input calyx neuropile (Butcher et al., 2012). A detailed record is reported for ~900 reconstructed cells in seven columns of the optic lobe’s medulla neuropile in Drosophila, with ~53,500 pre- and ~315,500 postsynaptic sites (Takemura et al., 2015), the average thus having ~6 elements per synapse, many with the profiles of two or more elements captured in a single cross section (Takemura et al., 2015). Flies have strictly four postsynaptic elements at their photoreceptor tetrads. In the Drosophila lamina each photoreceptor terminal forms about 50 such synapses (Meinertzhagen & Sorra, 2001), while Musca photoreceptors each have 200 (Nicol & Meinertzhagen, 1982); each is a fixed tetrad (p. 263) (Nicol & Meinertzhagen, 1982; Fröhlich, 1985; Meinertzhagen & O’Neil, 1991). Salticid spider photoreceptor synapses are also tetrads (Oberdorfer, 1977), while in the dragonfly Sympetrum, by contrast, the eqivalent synapses are triads (Armett-Kibel et al., 1977). In flies, Sterling and Laughlin (2015) interpret this organization as a means to increase the efficiency of presynaptic vesicle release, which multiple elements share at each site, albeit at the morphogenetic cost of assembling those elements in fixed ratios.

In other selected examples, dyad synapses have been observed in locusts: in output synapses, some of those at motor neurons (Watson & Burrows, 1982) and most at nonspiking interneurons (Watson & Burrows, 1988) of the metathoracic ganglion; and in the antennal lobe and mushroom body (Leitch & Laurent, 1996). The output synapses of crustacean nonspiking interneurons are also dyads, similar to those in the locust (Kondoh & Hisada, 1986). Dyads have also been seen elsewhere in crustaceans. In the neuropile of the stomatogastric ganglion (King, 1976a), at the median photoreceptor of the barnacle ocellus (Schnapp & Stuart, 1983), and the photoreceptors of the crustacean compound eye in Artemia salina (Nässel et al., 1978), synapses are also dyads. By contrast dyad synapses are not observed in the lamina of Daphnia magna (Nässel et al., 1978), which seems therefore to be secondarily simplified. Postsynaptic sites at many of these synapses often appear structurally closely matched, suggesting that divergent transmission especially to two postsynaptic elements at dyads is equally balanced.

Synaptic Convergence

In addition to divergence at synaptic sites, clear cases of synaptic convergence are also reported. Thus, the lobula giant movement detector (LGMD) neuron of the locust visual system receives retinotopic inputs from medulla units that are presynaptic to fine dendrites of the LGMD in the distal lobula. These make dyad synapses not only upon the LGMD itself but also onto neighboring input terminals at immediately adjacent sites. These complexes assemble in large numbers that completely cover LGMD neurites, and when the latter are cut in cross section, they form so-called synaptic rosettes (Rind & Simmons, 1998). Rosette motifs of convergent dyads are also seen in the mushroom body of locusts (Leitch & Laurent, 1996), and as well in Drosophila (Rivlin, personal communication), and synaptic convergence was reported from an early study on the antennal lobe of the migratory locust (Schürmann & Wechsler, 1970). Their functional significance is not known, but in the LGMD the retinotopic units are thought to excite the LGMD, but inhibit each other, the synapses forming a substrate for a critical race between excitation caused by edges moving out over successive photoreceptors and inhibition spreading laterally (Rind & Simmons, 1998).

Synaptic Vesicles

Synapses almost invariably incorporate a cluster of presynaptic vesicles, although the number of these may often not be well preserved, probably because they discharge during fixation for electron microscopy. Vesicles differ by number, shape (round, elliptical, pleiomorphic), content (clear or dense cored), and size (~30 nm for small clear vesicles up to about 180 nm for dense cored vesicles), which are reported and as were discussed previously in this review. Good examples come from the work of Ratté and Chase (2000) on pulmonate molluscs. Differing shapes and sizes of synaptic vesicles have been correlated with specific transmitters (Atwood et al., 1972; Tisdale & Nakajima, 1976). In the stomatogastric ganglion of the crab Cancer borealis, five synaptic profile classes contain a presynaptic apparatus (of dense bars and small clear vesicles), two also containing dense core vesicles that comprise multiple immunocytochemical classes, proposed neuromodulators, not readily distinguished by structural criteria (Kilman & Marder, 1996). The somata of all neurons are reported to contain dense-core vesicles in the stomatogastric ganglion of the lobster (Friend, 1976).

Vesicles behave as miniature osmometers with a shape and size that shrinks and swells with fixative molarity. Relative depletion studies by differential stimulation of excitor and inhibitor innervation to crustacean muscle have been used to identify the vesicles in either synapse type. Very little consideration in most studies is given to the time taken to fix synaptic vesicles, which is not instantaneous with aldehyde fixation and can be several times the duration of the vesicle cycle (Macintosh & Meinertzhagen, 1995) so that vesicles are shed and recycled in uncontrolled ways even while in aldehyde fixatives. Studies using high-pressure freezing and freeze substitution offset these problems and have been employed in vertebrate systems (e.g., Rostaing et al., 2006; Korogod et al., 2015) but with the exception of methods for C. elegans (e.g., (p. 264) Manning & Richmond, 2015) have yet to be widely adopted for invertebrate nervous systems.

The reported cleft widths of synapse types in Drosophila all fall within 10–20 nm, in contrast to vertebrates, in which the synaptic cleft at neuromuscular junctions is significantly wider than at central synapses (Peters et al., 1991). The features of synaptic organelles have been conserved, not only at different sites in the same species (as at neuromuscular and photoreceptor synapses in Drosophila: Prokop & Meinertzhagen, 2006) but also in different species in insects (Watson & Schürmann, 2002) or even, to a surprising extent, at equivalent sites of different groups (as in the convergence in photoreceptor synapses of insects and vertebrates: Meinertzhagen, 1993). In fact, the acquisition of synaptic transmission and the recruitment of a full suite of synaptic organelles had already occurred in ancestral Cnidaria, long preceding the origins of organized nervous systems, where their function was possibly first refined at neuromuscular junctions (Spencer, 1989).

Synaptic Partnerships

Most early studies fail to reveal the exact identity of cells participating at synaptic sites, except the few that report findings from labeled neurons, for which HRP fills of identified neurons (e.g., Muller & McMahan, 1976; Tolbert & Hildebrand, 1981; Watson & Burrows, 1981, 1982) have been particularly valuable. More recent dense EM reconstruction approaches use serial-section EM (ssEM) or focused ion beam milling, in which all cells are reconstructed and identified morphologically, and therefore in which none goes undetected. Specific methods for staining synaptic organelles are reported in Schürmann (1980). In Drosophila, combined GAL4 and UAS reagents have been used to target HRP to the plasma membrane of specific neurons, providing a valuable means to identify EM profiles of expressing neurons after the brain is incubated in peroxidase and diaminobenzidene (DAB; Larsen et al., 2003; Edwards & Meinertzhagen, 2009). Specificity and strength of expression is conferred by the particular GAL4 driver. In a recent upgrade of this approach, a recently reported “two-tag” double labeling system has been used to highlight both pre- and postsynaptic neurons in the same preparation, combining two orthogonal expression systems and two different peroxidases, either membrane-targeted HRP to target a postsynaptic neuron, as earlier, in combination with mitochondria-targeted APX (ascorbate peroxidase; Martell et al., 2012) to label its presynaptic partner, with both labels appearing at synaptic sites after their common incubation in DAB (Lin et al., 2016). These methods circumvent more laborious serial sectioning methods, but they are not for the faint-hearted; they combine the need for meticulous ultramicrotomy to sample the correct contact region between the two neurons, with the need to develop clever genetic reagents. The latter at least are now available for future studies.

Numbers of Synapses

Numerous biases in identifying synaptic profiles in single sections give notoriously false impressions of the relative abundance of organelles or between which parts neurons connect, which stereological methods alone can help offset (Weibel, 1979). Serial-section EM (Shibata et al., 2015) and 3D reconstruction methods have advanced rapidly in recent years (e.g., Chklovskii et al., 2010; Xu et al., 2013) that can, however, also help offset these problems. Evaluations of synapse numbers need careful ultrastructural and statistical attention. The dissector method is preferred for volumetric counts of synaptic numbers (Coggeshall & Lekan, 1996). Uncorrected profile counts give no reliable evidence for volumetric densities of synaptic sites and often are best disregarded. Recent studies on more highly differentiated synapses incorporate laborious EM tomographic methods to provide images of individual synaptic organelles with resolution increased in the z-axis (e.g., Leitinger et al., 2012).

Complete numbers of synapses are reported in only a few cases from dense reconstructions using EM. Thus, the 302 CNS cells in C. elegans have 6,393 chemical synapses, 890 gap junctions, and 1,410 neuromuscular junctions (White et al., 1986), as updated by Varshney et al. (2011), for an average of 21.2 synapses + 2.95 gap junctions per cell. In the L4 larva of C. elegans, 3,462 synapses and 754 gap junctions were distributed among 183 neurons, or 18.9 synapses and 4.12 gap junctions per cell (Durbin, 1987). Of these, 9% of neurons form partnerships connected by synapses and 7% by gap junctions (Durbin, 1987). Numerically comparable, the 177 CNS neurons in a larval Ciona, all with a synapse and most with an axon, had 6,618 synapses each occupying >1 60 nm section, including 1,772 neuromuscular junctions; there were also 1,206 gap junctions of >1 section (Ryan et al., 2016). These formed 2,835 synaptic partnerships. Each neuron thus formed 37 synapses or neuromuscular junctions. By comparison, in the lamina of Drosophila (Meinertzhagen & (p. 265) O’Neil, 1991), each module, or cartridge, has 37 synapses per cell (Meinertzhagen & Sorra, 2001), while in the medulla each neuron has on average 132 synapses per cell, with a total of 2,634 synaptic partnerships for the 20 cells of the so-called core connectome (Takemura et al., 2015). In a less intensively documented example, the Daphnia lamina cartridge has 51 synapses per neuron (Macagno et al., 1973). The ~30 neurons and ~150 input axons of the crustacean stomatogastric ganglion form an estimated million synapses (King, 1976a).

Synaptic Puncta Viewed by Light Microscopy

The small sizes of many invertebrate brains and ganglia and their transparency in many cases, combined with the improved design of modern microscope objectives, have meant that many studies have pioneered the use of entire brains in wholemounts. This is especially true of Drosophila, which spearheaded huge advances in neurobiology by allowing the visualization of neurons and synaptic proteins in many studies using confocal imaging of neurons. But many workers have been reluctant to acknowledge fully the fact that sites of synaptic contact cannot be resolved by light microscopy alone and that because of its z-axis resolution, light microscopy cannot reliably assign neurons to pre- and postsynaptic sites at the same synapse. Some molecular labels do, it is true, help to diagnose the polarity of synaptic contacts; the synaptic protein Bruchpilot (BRP) from anti-Brp (nc82) or genetic reagents based on this protein help identify sites which at photoreceptor synapses have been shown to be presynaptic in both Drosophila (Wagh et al., 2006; Hamanaka & Meinertzhagen, 2010) and the locust (Leitinger et al., 2012), while Denmark is reported as a dendritic marker in Drosophila (Nicolaï et al., 2010) and thus predominantly postsynaptic.

Notwithstanding, synapses have been visualized by light microscopy using genetic constructs to facilitate synaptic counting methods in Drosophila, especially using the split GFP so-called GRASP system, as applied in C. elegans (Feinberg et al., 2008), or the mushroom body of Drosophila (Pech et al., 2013) or the so-called synaptic tagging with recombination (STaR) method for marking synapses, as applied in the Drosophila medulla (Chen et al., 2014). The latter targets expression of Bruchpilot, which localizes to the platform of the T-bar ribbon (Wagh et al., 2006; Hamanaka and Meinertzhagen, 2010). UAS-brp-shortcherry, a fluorescently tagged fragment of BRP which depends on endogenous BRP for its correct localization (Schmid et al., 2008), is claimed to represent a reliable marker for active zones (e.g., Kremer et al., 2010) and thus to offer a molecular proxy for the presynaptic site, one that does still require validation by electron microscopy. It might be, for example, that not all synapses have a T-bar ribbon or may have a T-bar that lacks a presynaptic platform.

Neuromuscular Junctions

Neuromuscular junctions have provided a special opportunity for morphological studies of invertebrate synapses. Foremost in detail are those of arthropod muscle, both the neuromuscular junctions of decapod crustacea documented in studies especially from the Atwood group (see, e.g., Cooper et al., 1996 and studies cited therein) and phyletic upgrade studies on the larval neuromuscular junctions of Drosophila (Atwood et al., 1993; as reviewed, Prokop & Meinertzhagen, 2006). In these cases the muscle provides a simplified postsynaptic partner to those at the frequent dyad synapses of the CNS, in being a single cell with a clear postsynaptic density. The latter often has a highly developed subsynaptic reticulum.

The Locations of Synapses

Axo-axonic synapses are commonplace, for example in the procerebrum of pulmonate gastropods Helix and Limax (Zs-Nagy & Sakharov, 1970), the intestinal nerve of the peripheral nervous system in Helix (Elekes et al., 1985), or the larval CNS of Ciona (Ryan et al., 2016). In the latter, detailed plots of synaptic loci reveal that most synapses are in fact axo-axonic and axoterminal, especially among relay interneurons, rather than synapses forming upon dendrites or somata. Thus, presynaptic sites are most commonly located over axons or their terminals, while dendrites and somata are heterodoxically presynaptic. In contrast, terminals of sensory neurons bear most of the neurons’ presynaptic sites. The distributions of synapses along the length of the axon, and the dimensions of neurons, most with axons less than 100 µm and some not half this value, suggest that signaling occurs by local potentials (so as to prevent antidromic conduction or the collision of impulses) and, even if by impulses, that the distributions of synapses does not readily support a clear polarity of conduction, which must await appropriate imaging studies.

In the crustacean stomatogastric ganglion, neuropile is differentiated into two regions: a core of coarse neuropile and a surrounding region of (p. 266) fine-textured synaptic neuropile. Synapses are found only in the latter and not in the core (King, 1976a). Nearly every secondary neurite of all neurons forms both pre- and postsynaptic contacts and thus lacks a distinct pre- and postsynaptic region (King, 1976b). The connections between a specific pair of neurons are also distributed and do not cluster. In antennal lobe glomeruli of adult Drosophila, the input olfactory receptor neurons form a hollow crown within which fit the neurites of projection neurons and local interneurons (Tanaka et al., 2012).

Few reports for the exact placement and pattern of synapses over the surfaces of reconstructed neurons have been made, for the obvious reason that for neurons of any branching complexity this task is no simple matter. But Ratté and Chase (2000) report input and output synaptic sites along reconstructed stretches of the neurites of procerebral neurons in the pulmonate snail Helix, while in insects the synaptic distribution is reported for a reconstructed DCMD neuron and the inputs it receives from the LGMD, and unidentified neurons (Killmann et al., 1999). The DCMD dendrites emerging from the integrative segment vary in form and number between individuals and sexes but always form a flattened dendritic domain. The arborizations and the integrative segment are exclusively postsynaptic. Input to the DCMD is received at two types of synapse, having either round (Type 1) or pleiomorphic synaptic vesicles (Type 2). LGMD’s synapses are smaller Type 1 synapses. Killman et al. (1999) calculate for an entire DCMD dendritic arbor, a total of 8,500 active zones that includes at least 2,250 LGMD-synapses and about 1,000 Type 2 synapses. Reconstructions of identified HRP-injected locust thoracic motor neurons indicate that input and output synapses may occur within 1 µm of each other (Watson & Burrows, 1981, 1982). In three glomeruli of the Drosophila antennal lobe, each of the three major classes of neurons (receptor, projection, and intrinsic neurons) is both pre- and postsynaptic to the other two, projection neuron neurites have an approximate ratio of three presynaptic sites for each postsynaptic one, and input and output sites are partially segregated along the proximo-distal gradient of neurites, with the thinnest being solely presynaptic (Rybak et al., 2016).

Synaptic Organelles

The most obviously universal synaptic organelles are presynaptic vesicles. These offer few clues, or none at all, to the identification of their contents. Functional analyses of synaptic circuits still require knowledge of the neurotransmitter and neurotransmitter receptor for each identified neuron, a next phase for the morphological connectome studies that have been reported to date (e.g., Takemura et al., 2013, 2015).

The polarity of transmission is sometimes unclear. From structural criteria, the direction of transmission is assumed from the presence of a clear cumulus of presynaptic vesicles, although nervous systems can also reveal symmetrical neuronal partners in which a clear presynaptic site lies on both sides of a synaptic cleft, and transmission is therefore assumed to be nonpolarized. First reported in coelenterates (Horridge & Mackay, 1962) later instances have been reported in the tentacular neuropile of pulmonates (McCarrager & Chase, 1985) and in the larval CNS of Ciona (Ryan et al., 2016), in which neurons so connected form both reciprocal and serial synapses.

Gap Junctions

Gap junctions, sites of electrical transmission, are characterized by membrane apposition between neuron neighbors (Bennett & Goodenough, 1978). They incorporate one of two major channel proteins, connexins in deuterostomes and innexins in protostomes (Goodenough, 1974; Phelan & Starich, 2001). A third family of channel proteins, the pannexins (Panchin et al., 2000) are homologues of innexins and coexist with connexin junctions in vertebrates (Baranova et al., 2004). Genes for innexins (pannexins) are reported in all Metazoa except echinoderms, whereas connexins are exclusive to chordates (Abascal & Zardoya, 2013) and invertebrate deuterostomes (White et al., 2004). No fewer than 12 gap junction proteins (pannexins and or innexins but not chordate-specific connexins) are reported in the ctenophore Pleurobrachia (Moroz et al., 2014).

In thin-section EM, gap junctions appear as a narrowing of the intercellular cleft, with—in insects—a close membrane apposition of 2–4 nm, sometimes with various membrane densities or organelles, or with an increased linearity of the sectioned membranes, but with ultimate if rare validation by freeze-fracture images. Most studied using both thin-section EM (Ribi, 1978; Shaw & Stowe, 1982; Shaw et al., 1989) and freeze-fracture (Shaw & Stowe, 1982) techniques are those between photoreceptor terminals in flies, and those in the motor system onto giant fibers in flies (Strausfeld & Bassemir, 1983; Blagburn et al., 1999). Contact areas between giant fibers and tergotrochanteral (p. 267) motor neurons or interneurons in Drosophila contain chemical synapses intermixed with regions of close membrane apposition (having a separation of 3.25 nm ± 0.12), with faint cross striations and an array of 41 nm vesicles on the giant fiber side (Blagburn et al., 1999). Gap junctions with a separation between membranes of 1.41 ± 0.08 nm are abundant between peripheral perineurial glial processes (Blagburn et al., 1999). Individual haltere afferents in Drosophila may also make mixed synapses with the B1 motor neuron (Trimarchi & Murphey, 1997), and parallel axons from Johnston’s organ display extensive contacts, including putative gap junctions. Their synaptic boutons have both chemical synapses as well as putative gap junctions, indicating mixed synapses (Sivan-Loukianova & Eberl, 2005).

The cleft at a gap junction depends critically on how it is measured, and whether membrane thickness is included or not. In C. elegans, gap junctions have a cleft of about 8 nm including the apposed membranes, and the region of close apposition is usually in the form of a plaque about 350 nm diameter (White et al., 1986). In thin-section EM many gap junctional profiles are sectioned obliquely, compromising the requirement to image the cleft clearly. As a result, most estimates of gap junction numbers and distributions are almost certain to be underestimates, even in EM series in which comprehensive documentation of gap junctions is reported, as for example in the complete connectome of C. elegans (White et al., 1986) and larval Ciona intestinalis (Ryan et al., 2016). Despite minor difference in the intercellular cleft between innexin GJs in Drosophila and connexin GJs in a vertebrate, morphological differences are minor. Gap junctions often copopulate synapses between neurons with chemical synapses, and mixed electrical and chemical synaptic transmission is reported in both vertebrates (e.g., Lin & Faber, 1988) and gap junctions and chemical synapses in invertebrates (e.g., Strausfeld & Bassemir, 1983). In the larval CNS of Ciona, 8% of all partners connected by chemical synapses also have putative gap junctions, while only 1% are connected only by gap junctions (Ryan et al., 2016).

Some neurons, such as T1 in the fly’s medulla, lack chemical presynaptic contacts (Takemura et al., 2013), and for such cases we might suspect the existence of electrical transmission via gap junctions as an alternative. The ultrastructure of candidate gap junctions in the medulla has not been validated, however, and these are less clear than chemical synapses, dependent in particular on the plane of section.

Beyond Synapses to Synaptic Networks

The most obvious significance of the synaptic bookkeeping just presented is that at least in the case of the three preparations for which we have comprehensive data from dense reconstructions, the number of chemical synapses per neuron is not large, on average about 50 (Ciona, Drosophila) or 20 (C. elegans) but with a wide range of ~1–430, albeit divergence is commonplace, with many cases of each synapse providing input to multiple postsynaptic targets. Correlated with the paucity of their number, most neurons are sparsely connected.

Many synaptic features of all nervous systems have been recorded with particular precision in the nervous systems of invertebrates, but they are unlikely to be special to them. For example, the metathoracic fast extensor tibiae and mesothoracic tergosternal flight motor neurons have both output and input synapses on their neuropilar neurites. These synapses are involved in serial, reciprocal, and recurrent relationships (Watson & Burrows, 1982), revealing that the structural equivalent of a physiological synapse may be complex. Reciprocal and serial synapses are in fact commonplace in all synaptic networks. In the C. elegans connectome, for example, the probability of a connection from neuron B to neuron A given that there is also one from A to B is high (14% compared with 9% for the respective probabilities) (Durbin, 1987). In the larval CNS of Ciona, reciprocal synapses constitute approximately 15% of the synapses between neurons, excluding those onto muscle cells (Ryan et al., 2016).

Synaptic network complexity may be enhanced further not only by programming connections between increased numbers of neurons but also by the additional complexity of individual synaptic sites. A recent study proposes that the complementary expression of paired members of two immunoglobulin families Dip and Dpr interacting proteins act during circuit assembly in the Drosophila visual system (Tan et al., 2015). Such programming may ensure that a presynaptic neuron forms a synapse with a single postsynaptic neuron it contacts, associated especially when both costratify in the neuropiles, as in the optic lobe (Fischbach & Dittrich, 1989) or central complex (Wolff et al., 2015). But there is enriching network complexity for a fixed number of synapses, many synapses are divergent (see earlier under “Polyadic Synapses”), and serial synapses are (p. 268) also commonplace. The latter serve to spread activity between adjacent pathways, as in the amacrine cell networks of the vertebrate inner retina (Dowling, 2012), and they also abound in invertebrate networks. For example, they are clearly seen at peripheral locations in the neuromuscular innervation of the crab and crayfish (Pearce & Govind, 1993). The slit sensilla neurons of the spider leg (Fabian-Fine et al., 1999, 2000) and crustacean muscle receptor organ (Elekes & Florey, 1987) both provide rich sources of accessible peripheral synaptic organization generated between the peripheral efferent innervation of sensory neurons (Fabian-Fine et al., 2002). Basal Cnidaria and Hydrozoa also appear to exhibit presynaptic inhibition mediated by serial synapses, which confirms that this type of synaptic arrangement existed even in the first nervous systems (Westfall, 1996; Westfall et al., 2002).

Denying any simple dichotomy between the roles of pre- and postsynaptic sites, autapses are heterodox synapses at which a neuron is presynaptic to itself. Such sites are occasionally seen in the Drosophila medulla and provide a special case of abnormal synapse, one seen at elevated frequency in two particular cells, and thus that is probably programmed (Takemura et al., 2015). The polarities of transmission at autapses are generally assumed to be opposite for the forward and feedback directions, and feedback at most autapses is thought to be inhibitory and therefore self-limiting, for example maintaining the precision with which a neuron fires action potentials (Tamás et al., 1997; Ikeda & Bekkers, 2009).

How is such synaptic business regulated? We obviously have much to learn, but several studies now confirm that the volumetric density of synapses approximates 1–2 synapses per µm3, at least in Drosophila (Meinertzhagen, 2010; Rybak et al., 2016), and similar densities can be extracted from reports on other systems, including those of vertebrates, as summarized in Meinertzhagen (2010). So, despite many differences, some overall packing limits, possibly energetic in origin, guide the construction of neural circuits.

The Centralization of Neuron Populations

With progressive centralization, nervous systems became arranged in characteristic neuropiles and axon tracts. As a result, the numbers of processing centers in invertebrate brains increase with cell number and complexity. In Drosophila, the total neuropile volume in a brain, excluding the optic laminas, is 4.27 × 106 µm3 for 16 identified brain regions (Rein et al., 2002), but many more may exist. Some so-called glomerular neuropiles have many repeating processing elements, compartments, modules, or columns; others are diffuse (Hanström, 1928), in which a substructure cannot be, or has not so far been, distinguished by light microscopy, has not been sought by requisite methods, or, more probably, simply does not exist. Subdivisions can be orthogonal, as in columns in the arthropod compound eyes’ optic neuropiles (one per ommatidium) and their stratification, as seen in a range of arthropods (Sinakevitch et al., 2003) with 10 strata reported in detail for Drosophila (Fischbach & Dittrich, 1989; Takemura et al., 2013). These subdivisions serve to reduce the combinatorial complexity of synaptic connections, but similar arrangements exist in other brain regions, notably the central body (Hanesch et al., 1989; Wolff et al., 2015), where they may serve the needs of rapid timing. Other neuropiles are less morphologically determinate but nevertheless glomerular, as in the antennal lobe: with 54 glomeruli in Drosophila visible in both dissected and in vivo brains (Grabe et al., 2015), but many more (~160) in honeybees (Galizia et al., 1999). Borders between glomeruli are in some cases hard to discern and partly ambiguous. Yet other neuropiles, of which the lateral horn and suboesophageal ganglion are volumetrically large examples, are referred to as diffuse neuropils and may occupy as much as 90% of the central brain; they lack a clear substructure and have a circuit organization that is mostly not known. A hierarchical nomenclature system, using the brain of Drosophila as its reference establishes 47 brain regions comprising the entire insect brain (Ito et al., 2014); this system builds on previous reports for specific species, such as that for the housefly Musca domestica (Strausfeld, 1976). Similar conventions will be needed for other invertebrate groups, especially the brains of more divergent molluscs. Cellular organization in the latter is less well reported than in arthropods, and in cephalopods is treated histologically as an honoraray vertebrate, without identified neurons.

Fasciculation and Pathways

Neurons project to different brain regions by means of axons arranged in tracts, and the study of these, historically known by their hodological (i.e., connectionist) anatomical pathways (Fortuyn, 1920), underlies the generation of connectome data. Typical fiber tracts, such as the first projection of photoreceptor axon bundles to the lamina, (p. 269) or the second projection via the external chiasma, show scrupulous retinotopy of fiber bundles (Meinertzhagen, 1976). Most comprehensively detailed in Drosophila, axons in insect neuropiles fasciculate strictly, typically in bundles of up to ~15 (Takemura et al., 2008). Fiber tracts comprise multiple such fascicles, as in the chiasmata of the optic lobe, sensory pathways, or cervical connectives. A total of 58 tracts between 41 neuropiles and 6 hubs are documented comprehensively in Drosophila and a total of 16,000 neuron classes fasciculate consistently within these (Chiang et al., 2011). Similar compositional fixity is seen in C. elegans with many minor tracts contributing to the circumpharyngeal nerve ring (White et al., 1986). The chief tracts are the ventral cord, the dorsal cord, the excretory canal–associated processes, and the posteriorly directed sublateral processes. There is a close association between the neurites of specific neurons, or between a neurite and the basal lamina (White et al., 1986). Axons in the larger tracts of the CNS of the first nauplius stage of a harpacticoid copepod Dactylopusia tisboides, 80 µm long, contain roughly a dozen fibers each (Lacalli, 2009).

A striking feature is the precision with which axons position themselves relative to neighbors within bundles (White et al., 1986). The same is true in the insect compound eye, most noticeably among axon bundles of ommatidia, which likewise fail to braid their relative positions even when the bundle rotates as a whole (Meinertzhagen, 1976). In more basal groups, by contrast, axons may often lack strict fasciculation. So, for example, axons in the larval brain of Ciona do not strictly bundle together and may braid their positions (Ryan et al., 2016). In the larvacean Oikopleura, a brain nerve comprising a single axon terminates on two types of touch receptors in the oral region (Olsson et al., 1990). The two nerves are the dendrites of two perikarya in the forebrain and are the master neurons for ciliary reversal in the stigmata, which is a two-neuron reflex. They form axoaxonal synapses with one motor neuron in the midbrain, the command neuron for ciliary reversal. This and other details exhibit a remarkable sidedness, including the left and right branches of a bifid neuron in nerve 3 (Olsson et al., 1990).

The neural canal of the tubular nerve cord in various protochordate groups examined (tunicates, larvaceans, and amphioxus) harbors a still-enigmatic Reissner’s fiber (Olsson, 1972).

Circuits, Connectomes, and Network Motifs

It is a source of some frustration for the timing of this review that morphological studies have recently entered a new phase of circuit analysis to which invertebrate brains have much to contribute. Given their small size and numerical simplicity, invertebrate brains lend themselves to the complete documentation of all their synaptic circuits by electron microscopy, leading to what has become known as a connectome (Lichtman & Sanes, 2008). This is no new concept, merely one that is newly resurrected, given life by emerging digital and molecular technologies. Until recently a complete connectome was available only for a single species, the nematode C. elegans (White et al., 1986; Varshney et al., 2011). Elsewhere, comprehensive morphological circuits are known only for specific brain regions, such as the fly’s optic lamina (Boschek, 1971; Meinertzhagen & O’Neil, 1991; Meinertzhagen & Sorra, 2001) or medulla (Takemura et al., 2008, 2013), and now the motor circuits of the larva (Fushiki et al., 2016). These and the methods used to obtain them seem destined to open up the brain of Drosophila to complete connectomic analysis, given that no neuropile exceeds the depth (50–75 µm) accessible to EM reconstruction, and given the range and power of genetic methods available in Drosophila (Pfeiffer et al., 2010; Jenett et al., 2012). The larval CNS is the likely next Drosophila target of a future complete connectome (see Ohyama et al., 2015), as indeed recently reported (Fushiki et al., 2016), while reconstruction of an entire adult brain is a singular and ambitious objective of the Janelia campus of the Howard Hughes Medical Institute (Plaza et al., 2014). In Drosophila, objectives at the EM level are aided by new tools for neuroanatomy and neurogenetics (Pfeiffer et al., 2008), 3D registration (Lam et al., 2010; Peng et al., 2011), and wiring networks (Chiang et al., 2011; Shi et al., 2015). Once constructed, brain regions in Drosophila must be identified and named for maximum uniformity with the brains of other insects (Ito et al., 2014), and their patterns of gene expression identified, especially using methods for automated whole-brain in situ expression patterns of multiple genes (Lin et al., 2015). Finally, extensive libraries of reporter lines for cell types are now available for specific brain regions, for which the following are at best only a partial listing for the wealth of opportunities these now provide: the antennal lobe (Tanaka et al., 2012; and corresponding neurons in the larva: Masuda-Nakagawa et al., 2010); the mushroom body (p. 270) (Tanaka et al., 2008); the optic lobe (Otsuna et al., 2006, 2014); central body protocerebral bridge (Lin et al., 2013; Wolff et al., 2015); the gustatory center (Miyazaki & Ito, 2010); auditory projecting neurons (Kamikouchi et al., 2006); neurons of gravity and hearing (Kamikouchi et al., 2009); paired giant descending neurons (GDNs, or giant fibers) and paired giant antennal mechanosensory descending neurons (GAMDNs) (Mu et al., 2014); neurons controlling motor programs (Flood et al., 2013a,b); octopaminergic neurons (Busch et al., 2009); PDF neurons of the circadian system (Shafer et al., 2006; Hermann et al., 2013); and the neurons that constitute the fly’s motion-detection pathways, the T4 and T5 neurons (Fischbach & Dittrich, 1989; Maisak et al., 2013; Takemura et al., 2013), HS and VS neurons (Scott et al., 2002). Neurons in the visual system, from the lamina (Tuthill et al., 2013), medulla (Takemura et al., 2013, 2015), and lobula (Otsuna & Ito, 2006; Otsuna et al., 2014), are also comprehensively covered by lines selected from major screens generated at the Janelia Campus of Howard Hughes Medical Institute and that constitute part of the IMP Vienna stock collection ( These reports augment earlier light microscopy reports from Golgi impregnation in Drosophila (Fischbach & Dittrich, 1989; Hanesch et al., 1989, and as cited earlier for other insect species).

In deuterostome species, the connectome for the 177 CNS neurons of the ascidian tadpole larva, Ciona intestinalis, has now also recently been completed using ssEM (Ryan et al., 2016), based on a simple library of neurons viewed with synaptotagmin-driven GFP expression (Imai & Meinertzhagen, 2006). Details from Ryan et al. (2016) include an anterior brain vesicle with 10 types of neurons and 19 subtypes, a posterior brain vesicle with 7 types of neurons and 14 subtypes, and a motor ganglion with 4 types (including two types of motor neurons totally five pairs in all) and 6 subtypes. Antenna relay neurons project mainly to right-side motor interneurons, and photoreceptor relay neurons to the left side. Many of the motor ganglion’s synaptic connections occur on only one side or are more numerous on one side than the other. The CNS is essentially an epithelial tube, which lacks stratification and highly differentiated tracts, and circuits of mostly axo-axonal connections. Overall, the network is highly bidirectional, with more feedback loops in two-node motifs than would be predicted from a random network. Where neurons form such loops synaptic polarity presumably differs between the two directions. Such motifs have been proposed to provide robust dynamical stability (Prill et al., 2005). This trend of bidirectionality holds true in both three- and four-node motifs as well, with the number of fully connected motifs (those in which all components are reciprocally connected) much higher than would be expected by chance. These types of motif generally indicate that a network contains well-connected groups (“cliques”) and exhibits a great deal of pathway redundancy. This is a common trait of biological networks that are small and highly connected, with few neurons coordinating complex behaviors.

Some details are also reported for the larvacean Oikopleura (Olsson et al., 1990), but these are hard to reconcile with the Ciona larval CNS.

Accuracy and Asymmetry in Invertebrate Neurons

Many invertebrate brains have a high level of numerical and morphological determinacy. It is therefore perhaps surprising that few workers appear to have thought it worthwhile to record variations in the number and distribution of neurons and their synapses. The first of these, Hertweck (1931), provides examples of variation in the branching patterns of peripheral nerves and in the numbers of neurons in chordotonal organs of the Drosophila larva. In the polychaete Platynereis there is a high level of stereotypy between individual cells of the same type, in the number of cells, their neurite projections, and synaptic connections, including left-right asymmetries in the connections of several neurons (Randel et al., 2015). Few other examples provide similar or quantified data, for which studies on the visual pathways of the insect compound eyes have proved exemplary. The projection accuracy of photoreceptor axons in the fly Calliphora offered a precedent (Horridge & Meinertzhagen, 1970), with a projection error rate of about 1.5% in an error-prone region of the fly’s eye. A more recent report provides similar data for Drosophila by means of genetic reporters, revealing error rates, measured directly for the differentiation and axon targeting of large numbers of photoreceptors, of approximately 0.08% for missing photoreceptors and 0.04% for axons that innervate the wrong cartridge (Schwabe et al., 2013). These rates are at least an order of magnitude less inaccurate than the synaptic errors reported among 20 neuron classes in the Drosophila medulla, as part of a so-called core connectome, those neurons present in seven neighboring columns. These connections identify (p. 271) circuits for motion, and their detailed documentation reveals that <1% of contacts overall are not part of a consensus circuit but incorporate errors either of omission or commission (Takemura et al., 2015). Autapses are also occasionally seen, and the elevated frequency of these in two cells, Dm9 and Mi1, which form ≥20-fold more autapses than do other neurons (Takemura et al., 2015), suggests that they are programmed and do not differ by chance.

An aspect of neuron accuracy is not only morphological stereotypy between animals but how closely the neurons of either side of the brain mirror the connections of their contralateral partners. Various instances of asymmetry in invertebrate nervous systems are usefully reviewed by Frasnelli et al. (2012), and several obvious examples serve as illustrations:

  1. a) In the simple brain of the rotifer Asplanchna, an unpaired cell far from the midline on the posterior dorsal exterior of the brain surface differs in outline from all others, being amoeboid with numerous highly infolded processes (Ware, 1971). One process penetrates the neuropile, and none projects very far beyond the surface. Another, monopolar medial cell 3, is shifted slightly to the opposite side of the midline from the unpaired cell.

  2. b) In larval neurons of the ascidian Ciona, the number, placement, projections, and connections exhibit various forms of sidedness (Ryan et al., 2016). Thus, photoreceptor input to the brain vesicle is mostly on the left, antennal (gravity sense) input mostly on the right, and the inputs of relay neurons to the motor ganglion show considerable left-right differences, with overall more on the left than right side, even though connections between the motor neurons are more numerous on the right-hand side (Ryan et al., 2016).

  3. c) Left-right asymmetries are reviewed for the nervous system of C. elegans by Hobert et al. (2002). Two thirds of the neurons of C. elegans (198 out of 302) are paired bilaterally. Most of the remainder, including 75 ventral nerve cord neurons, are located on or very close to the midline and lack a contralateral partner. Four unilateral neurons in particular (AVL, RIS, RIH, RID) occur in the head ganglia. Insofar as the mitotic ancestry of all cells has been recorded, it is clear that similar lineage histories are neither sufficient nor necessary for two cells to adopt bilaterally symmetrical cell fates. Other details are given in Hobert et al. (2002).

  4. d) On the right side of the Drosophila brain, the asymmetrical body, a 10 µm diameter FasII-positive region next to the ellipsoid, provides a fascinating brain asymmetry required for long-term memory formation (Pascual et al., 2004).

  5. e) Among various decapod Crustacea, the claws exhibit bilateral specializations with corresponding differences between the motor innervation to major (crusher) and minor (cutter) claws (Govind, 1992).

  6. f) Molluscs exhibit sidedness, most obviously gastropod molluscs as the result of torsion. In pulmonates such as Helix, the right mesocerebrum is large and has many neurons with large somata, while the left is small (as reviewed in Chase, 2000). A clear example comes from an early differentiating neuron in the vetigastropod Haliotis kamtschatkana that expresses 5-HT and anticipates the trajectory of the pleurovisceral nerve cords prior to torsion (Page, 2006). At that time, the soma lies at the ventral midline and extends two neurites anteriorly toward the apical sensory organ, but these do not cross over during ontogenetic torsion. Full crossing of the cords occurs gradually during later development as the mantle cavity deepens and expands leftward. Unlike gastropods, the slug Limax has a CNS that is bilaterally symmetrical, but even so the procerebrum on one side only, randomly determined, is used to store odor-aversion memory (Matsuo et al., 2010).

Examples of other brain asymmetries for which an underlying morphological or synaptic basis is still wanting include the asymmetry in gastropod mating behavior that results from anatomical asymmetry controlled by a maternal effect locus (i.e., che sinistral or dextral shell coil, of the snail’s chirality) in pond snails, Lymnaea stagnalis (Asami et al., 2008; Davison et al., 2009).

Neuronal Plasticity

Like those of vertebrates, the brains of many invertebrate species are not fixed, even if they may be highly predictable, those of different individuals normally showing close similarities but nevertheless capable of change under a variety of conditions. The many recorded instances of neuronal plasticity in invertebrates have been reviewed, at least for insects (Meinertzhagen, 2001), especially Diptera and Hymenoptera (Groh & Meinertzhagen, 2010).

Despite their normal invariance, synaptic networks may be susceptible to experience, and in Drosophila and other cultured species we may (p. 272) suspect that some of the stereotypy of these circuits in different individuals can be the product of stereotypy in the conditions under which individuals are reared. Clear cases of structural synaptic plasticity in response to different conditions of rearing have been reported (e.g., Kral & Meinertzhagen, 1989), which accompany and doubtless may underlie volumetric changes both for specific neuropiles (Barth et al., 1997) and the entire brain (Heisenberg et al., 1995) in Drosophila. Projection neuron boutons in the calyces of the honeybee mushroom body transform between 1-day-old bees and foragers, with the proportion of ribbon synapses, as well as the numbers of the Kenyon cell dendrites postsynaptic at each, higher in the latter; ribbon and nonribbon synapses form mainly dyads in the 1-day-old bee, and triads later on in the forager (Groh et al., 2012). These changes are associated with age and/or experience and are mainly caused by the outgrowth and connectivity of Kenyon cell dendrites.

An obvious case of morphological plasticity is provided by the growth of brain regions in hemimetabolous insects. A subset of Kenyon cells with sprouting neurites can be distinguished from neighoring Kenyon cells by avid labeling for f-actin (Mashaly et al., 2008). Short-term plasticity of synaptic sites in response to light adaptation is also reported for photoreceptor synapses (Rybak & Meinertzhagen, 1997), for which the presynaptic T-bar ribbon may be an assembly point and plasticity organelle (Wichmann & Sigrist, 2010).

A specific form of synaptic plasticity, which is reversible, is circadian in origin (Pyza & Meinertzhagen, 1993) and accompanies volumetric changes in lamina L-cells (Pyza & Meinertzhagen, 1995) and their dendrites (Weber et al., 2009). L2-feedback synapses increase at night, and in the subjective night, relative to their numbers in the day and subjective day (Pyza & Meinertzhagen, 1993).

Finally, in an extreme and remarkable form of plasticity, deganglionated ascidians Chelyosoma and Corella survive without regenerating a CNS ganglion. Apparently the sea squirts residual innervation that survives deganglionation, comprising either interconnected motor nerve terminals, interconnected sensory neurites, or some combination of both, is sufficient to maintain a deganglionated animal (Mackie & Wyeth, 2000). The response repertoire of deganglionated animals is largely intact, albeit slower, but it is not recorded whether deganglionation significantly diminishes the animal’s personality.

Postscript: Drawing the Line

At the conclusion of this review, the author apologizes to the many workers and some colleagues whose studies have not been cited and regrets any impression of partiality in the inevitable restriction to topics with which he is most familiar, especially in the nervous systems of flies and ascidian larvae. The final review is, as a result, heavily circumscribed by morphological findings, the topic of this chapter, with the particular omission of many recent studies having a more molecular or functional emphasis and on groups, such as molluscs, in which morphological analyses especially of synapses are less intensively reported. Also omitted are most of the numerous studies on invertebrate sensory receptor neurons and neuromuscular systems. No real justification can be offered for topping and tailing the literature in this way, except that to have included these topics would have doubled the length of this review, and that many reviews already exist on these topics, especially for eyes and other ocular organs on the one hand, and neuromuscular innervation on the other.


The author acknowledges with thanks Dr. Kerrianne Ryan (Dalhousie University) for permission to quote from her unpublished PhD thesis on the larval nervous system of Ciona intestinalis, and Dr. Shinya Takemura (Janelia Research Campus, HHMI) for permission to cite unpublished data on the synaptic organization in Drosophila, and to both of these colleagues for reading an earlier draft of this chapter. The author’s original research on invertebrate brains has been supported for many years by Discovery grant A-0000065 from the Natural Sciences and Engineering Council of Canada.


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