Glial Cells: Invertebrate☆

Glial Cells: Invertebrate☆ JA Coles, University of Glasgow, Glasgow, UK ã 2015 Elsevier Inc. All rights reserved.

Introduction The Anatomical Distribution of Glial Cells in Invertebrates Cnidaria: Nervous Systems without Glia Bilateria (Triploblastica) Nematoda Annelida Insecta Urochordata Ascidiacea (sessile tunicates) Thaliacea (pelagic tunicates and salps) Cephalochordata Functions of Glia in Adult Invertebrates Glia as a Blood–Brain Barrier Glia and Homeostasis of Extracellular Ion Concentrations Ion homeostasis in the retina of the bee (Apis) Possible role of glial cells in homeostasis of extracellular K+ around the giant axon of the squid Glia and the Removal of Neurotransmitter Glia as a Site of Glycogen Metabolism Glia as Suppliers of Fuel to Neurons Glia and the Maintenance of Axon Function Local synthesis of axon proteins Rapid signaling from neurons to glia Glia as Actors in the Development of the Nervous System Early Development of the Nervous System Development of Sensory Systems The olfactory system of the moth, Manduca sexta A glial function, myelination, is responsible for major differences in the sizes and life styles of vertebrates and invertebrates Lessons from Invertebrate Glial Cells

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Introduction Glia are cells that contact and invest neurons, and interact with them, but play at most an ancillary role in rapid electrical signaling. They generally share a common ectodermal origin with neurons. In general, glia surround neurons; sometimes the area of contact is increased by glial projections that fill neuronal invaginations. Present in annelids and insects, these processes are usually called trophospongia (Figure 1(b)) or, at the synapses of insect photoreceptors, capitate processes. Like vertebrate glia, invertebrate glia often contain glycogen granules and show electrical coupling. They also have a prominent cytoskeleton, usually of microtubules in arthropods and of filaments in mollusks and annelids. Large lysosomes called gliosomes (diameter, 1–4 mm) are frequent, and gliograna (diameter, 0.3 mm) are also found. Research on glial cells has overwhelmingly concentrated on the rat (with some interest in the mouse), and the amount of research done on the several million other animal species, almost all of which are invertebrates, is tiny in comparison. However, in terms of what has been learned about major principles of glial function, studies on invertebrates have made great contributions. Invertebrate glia are most often mentioned in work on the early development of the fruit fly, Drosophila. This is because Drosophila lends itself to studies based on analysis of mutants and because precursor cells give rise to glial cells and neurons in similar numbers. The way in which genes are switched on and off to determine whether a precursor cell becomes a neuron or a glial cell and also the study of the gene products involved in neuron–glia interactions are very active fields of research. This inextricable interplay of neurons and glia in development is only briefly outlined here. Instead, the emphasis is on the physiology of neuron–glia interactions in adult invertebrates. From the 1960s, until more powerful techniques became available for studying mammalian glia in the 1980s, a small number of invertebrate preparations with experimentally accessible glia were intensively investigated with the expectation that the results would reveal principles underlying the physiology of glia in general. The giant glial cell of the segmental ganglion of several species ☆

Change History: February 2015. JA Coles updated Section text, Figures and further readings.

Reference Module in Biomedical Sciences

http://dx.doi.org/10.1016/B978-0-12-801238-3.04613-4

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Figure 1 (a) Transverse section through a ganglion of the ventral nerve cord of the leech Hirudo medicinalis. Between the inner and outer capsules are six packet glial cells (five shown), each enclosing a packet of neuronal cell bodies. Two giant neuropil glial cells (one shown) lie on the axis of the cord and send processes among the neuropil within the inner capsule. (b) More detailed scheme showing the cell bodies of two packet neurons lying within a packet glial cell. From Coggeshall RE and Fawcett DW (1964) The fine structure of the central nervous system of the leech, Hirudo medicinalis. Journal of Neurophysiology 27: 229–289.

of leech (an Annelid), the Schwann cells of giant axons of squid and crayfish, and the ‘outer pigment cell’ of the retina of the honey bee drone have probably given rise to the majority of publications with this approach. Analysis of the signaling between neurons and glia necessary to the development of a nervous system is under way, particularly in insect sensory systems. However, altogether, the physiology of probably fewer than ten types of invertebrate glial cell has been studied. Work on these model preparations has been at the origin of several general and important concepts that have been subsequently applied to mammals, and these concepts can usefully be approached through the original invertebrate work.

The Anatomical Distribution of Glial Cells in Invertebrates Cnidaria: Nervous Systems without Glia The phylum Cnidaria includes sea anemones, hydra, and jellyfish, all of which have nervous systems without glial cells. One or two ring nerves are found in cubozoans (box jellyfish) and hydrozoan jellyfish. In addition, the small ( 1 cm diameter) cubozoan jellyfish, notably Tripedalia cystophora, have elaborate visual systems with up to more than 24 eyes, each with advanced neural circuitry composed of many hundreds of neurons. Despite the absence of glia, these circuits allow rapid and complex hunting behavior.

Bilateria (Triploblastica) Glial cells are reported in all Bilateria species that have been examined.

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Nematoda The hermaphrodite form of the nematode Caenorhabditis elegans (Figure 2(a)) has 56 glial cells. Each of the 24 sensilla has a sheath glial cell and a socket glial cell (or two socket cells for the phasmids). In addition, six glial cells, labeled GLR cells, extend sheetlike projections that contact muscle arms in the head. These associate with 302 neurons. The glia create a large part of the architecture of each sensillum. In the case of the amphid sensillum, the socket glial cell has a long process, which follows the dendrites of 12 neurons (only one of which is shown in (Figure 1(b)). At the cuticle, it takes a doughnut shape to create a pocket open to the exterior. At the inner end of the pocket, the sheath cell takes the form of a sieve with eight channels through which pass the sensory cilia of eight of the neurons. Laser ablation of single cells is contributing to understanding how the cells act together to form the sensillum.

Annelida In leeches such as Hirudo medicinalis, the ventral nerve cord has 23 ganglia, each containing the cell bodies of monopolar neurons grouped in six packets. The neuronal somata and initial axon segments of each packet lie within a packet glial cell (Figure 1). Two further giant neuropil glial cells invest the neuropil in the center of the ganglion. The axons connecting neighboring ganglia are enclosed in a glial cell sheath. There is no myelin (here or in any invertebrate); the larger axons are separated one from another by glial processes, but smaller axons are in bundles with no intervening glia. The neuropil glial cell has a cell body more than 100 mm across and extensive processes; it is probably the largest of all glial cells. For this reason, it was chosen in the 1960s by Stephen Kuffler and colleagues for the first systematic study of glial cell physiology. They found that the ionic basis of the membrane potential depended more strongly on potassium ions than is the case for neurons. They showed that glucose (and other compounds) can diffuse freely from outside the ganglion to the surfaces of the neurons, and they concluded that these leech glial cells do not normally supply metabolic substrate to neurons. This is certainly not universally the case, and it may be exceptional. Over subsequent decades, it has been shown that these glial cells do, in fact, do a lot of interesting things (see Figure 3(a)). (a)

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Figure 2 Glia in the nematode Caenorhabditis elegans. (a) Schematic of an adult hermaphrodite, approximately 1 mm long, showing the outline of the pharynx. Some neuronal tracts are shown in green, and one amphid sheath glial cell is shown in red. (b) Enlarged view of the anterior region. The nerve ring (green) is a neuropil where most synaptic interactions between neurons occur. Elements of one amphid sensillum are shown: one of the 12 neuronal dendrites (green), the sheath glial cell (red), and the socket glial cell (blue). Adapted from figures in www.wormatlas.org and from Procko, C, Lu Y, Shaham S, (2011) Glia delimit shape changes of sensory neuron receptor endings in C. elegans. Development 138: 1371–1381.

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Figure 3 Invertebrate neuron–glia interactions are divers and usually have parallels in vertebrates. (a) Some of the receptors, channels, and transporters found on the giant neuropil glial cell of the leech. iGlu, ionotropic (AMPA/kainate) glutamate receptor; mGlu, metabotropic glutamate receptor; nACh, nicotinic acetylcholine receptor; 5-HTR, receptor for 5-hydroxytryptamine; MMR, receptor for myomodulin, a neurotransmitter in leech. (b) Signaling from giant axon to Schwann cell in the crayfish. Propagation of action potentials along the axon causes release of N-acetylaspartylglutamate (NAAG). NAAG is split into N-acetylaspartate (NAA) and glutamate (glu) by the extracellular enzyme glutamate carboxypeptidase II (GCPII). Glutamate acts on metabotropic glutamate receptors, which activate parallel signaling pathways, including signaling through extracellular acetylcholine. (c) On glia of Drosophila brain, glutamate transporters of the EAAT1 class reduce oxidative stress. In addition, the glia contain a cysteine protease, cathepsin, which is normally inhibited by Cer. Approximately 3 h after conditioned learning, the establishment of long-term memory involves a decrease in Cer concentration leading to increased cathepsin activity. (a) Modified from a scheme kindly supplied by JW Deitmer. (b) Based on results in Urazaev AK, Grossfeld RM, and Lieberman EM (2005) Regulation of glutamate carboxypeptidase II hydrolysis of N-acetylaspartylglutamate (NAAG) in crayfish nervous tissue is mediated by glial glutamate and acetylcholine receptors. Journal of Neurochemistry 93: 605–610 and papers cited therein. (c) Based on results in Rival T, Soustelle L, Strambi C, Besson MT, Iche M, and Birman S (2004) Decreasing glutamate buffering capacity triggers oxidative stress and neuropil degeneration in the Drosophila brain. Current Biology 14: 599–605, and Comas D, Petit F, and Preat T (2004) Drosophila long-term memory involves regulation of cathepsin activity. Nature 430: 460–463.

Insecta In the central nervous system of insects (ventral nerve cord and brain) there are three main classes of glia: surface (or subperineurial) glia form a sheath around the entire central nervous system (Figure 4), cortex glia provide a matrix in which are embedded neuronal cell bodies, and neuropil-associated glia send processes into the neuropil. Unlike the neuropil glia of the leech (Figure 1), the cell bodies of insect neuropil-associated glia are numerous (25–30 in each neuromere segment of Drosophila) and are located outside the neuropil.

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Figure 4 A glial blood–brain barrier. The ventral nerve chord (central nervous system) of the cockroach Periplaneta americana. (a) The connectives running between the ganglia are avascular and contain giant axons and small axons (all unmyelinated). (b) The neural lamella provides a connective tissue sheath. Some work suggests that the main barrier to extracellular diffusion of molecules to the axons is formed by tight junctions between the perineurial sheath cells, which are sometimes classed as glial cells. However, work on Drosophila suggests that these cells are of mesodermal origin and also that the barrier is constituted by the underlying glial cells. Adapted from Treherne JE and Scholfield PK (1981) Mechanisms of homeostasis in the central nervous system of an insect. Journal of Experimental Biology 95: 61–73.

In addition, several types of glia in the periphery can be defined, such as the antennal nerve glial cells of Manduca and the outer pigment cells of Apis retina (Figure 6).

Urochordata Ascidiacea (sessile tunicates) The nervous systems of the pelagic larval forms of sea squirts (Urochordata and Ascidia) are laid out much like those of vertebrates. Nonneural cells, usually called ependymal cells, outnumber neurons. When the larvae become sessile, the brain is remodeled, becoming an order of magnitude larger. In this adult form, glial cells may be absent: the axons are unsheathed, but a few nonneural cells are present within the brain.

Thaliacea (pelagic tunicates and salps) There are no glia in the peripheral nervous system. They are not in evidence in the brain: this is covered by a two-layered epithelium, but the cells of this differ markedly from most glia in that they conduct action potentials.

Cephalochordata The amphioxus or lancelet, Branchiostoma lanceolatum, has glial cells. Thin glial lamellae containing gliofilaments enclose the rhabdom of the Joseph photoreceptor cells.

Functions of Glia in Adult Invertebrates Glia as a Blood–Brain Barrier The nervous systems of invertebrates are not vascularized; thus, unlike vertebrates, if they have a blood–brain barrier, it is not between capillaries and the brain parenchyma but, rather, around the various elements of the more or less distributed nervous system. The structures usually responsible for occluding extracellular clefts are tight junctions. They are found between glia in the sheaths around the central nervous system of insects and at least some mollusks (e.g., cuttlefish). Insects have a remarkable blood– brain barrier that supports large differences in the concentrations of ions and molecules. In the ventral nerve cord of the cockroach,

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Figure 5 The retina of the drone (male) honey bee: a model for studying glia–neuron exchanges. (a) Drawing of a slice of the head, caudal upwards. Much of the chitin that normally covers the caudal surface of the head has been removed to expose the retina and brain. The facetted cornea covers much of the surface of the head (here, the facets are shown too large; there are approximately 10 000 in total). Behind each facet, the cell bodies of six large photoreceptor neurons extend for 400–500 mm to the basal membrane (brown line). (b) In section, the large photoreceptor cells appear clustered in sixes, like the petals of a rosette. The outlines the glial cells can be seen clearly in the lower left. The rosettes are approximately 25 mm in diameter. (c) Stimulation of the neurons with their physiological stimulus (light) causes large movements of ions. The initial and final values are the measured mean values; the time courses are schematic. Green, K+; red, Na+; blue, Cl . The volumes of the fluid in the three compartments have been estimated as follows: photoreceptors, 0.29 l l 1 of tissue; glial cells, 0.44 l; and extracellular space, 0.05 l. K+ is removed from the extracellular space both by net uptake into the glia and by spatial buffering. (d) Metabolic exchanges. Glucose is taken up only by the glia. Pyruvate formed from glucose or glycogen is converted to alanine, which is transferred to the neurons. Ammonium, liberated when alanine is reconverted to pyruvate, returns to the glia being taken up on a specific Cl dependent cotransporter. (a, b) Adapted from Coles JA (1989) Functions of glial cells in the retina of the honeybee drone. Glia 2: 1–9. (c) Data from Coles JA, Orkand RK, Yamate CL, and Tsacopoulos M (1986) Free concentrations of Na, K, and Cl in the retina of the honeybee drone: Stimulus-induced redistribution and homeostasis. Annals of the New York Academy of Sciences 481: 303–317, and Coles JA, Orkand RK, and Yamate CL (1989) Chloride enters glial cells and photoreceptors in response to light stimulation in the retina of the honey bee drone. Glia 2: 287–297. (d) Based on Tsacopoulos M, Poitry-Yamate CL, and Poitry S (1997) Ammonium and glutamate released by neurons are signals regulating the nutritive function of a glial cell. Journal of Neuroscience 17: 2383–2390, Marcaggi P, Thwaites DT, Deitmer JW, and Coles JA (1999) Chloride-dependent transport of NH4+ into bee retinal glial cells. European Journal of Neuroscience 11: 167–177 and Coles JA, Martiel JL, and Laskowska K (2008) A glia-neuron alanine-ammonium shuttle is central to energy metabolism in bee retina. Journal of Physiology 586: 2077–2091.

the barrier is ascribed to tight junctions between the outer perineural sheath cells (Figure 4(b)). Some authors call these cells glia, but examination of Drosophila mutants suggests that they are not of ectodermal origin. The retinas of insect compound eyes also have a glial blood-retina barrier, and in Apis, at least, this barrier separates fluids with different concentrations of amino acids and carbohydrates (Figure 5(a)). In contrast, in ganglia of the leech (Annelida) and the snail (Gastropoda), ions and molecules with a molecular weight of up to approximately 10 kDa can diffuse readily through the extracellular clefts of the glial sheath to reach the neurons.

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Glia and Homeostasis of Extracellular Ion Concentrations Although groups of short axons not separated by any glial sheath are found in animals from jellyfish to man, activity more subtle than the transmission of action potentials benefits from a regulated ionic environment. Potassium is the ion of most concern since extracellular [K+] affects the membrane potential of neurons, and active neurons release K+, thus tending to increase extracellular [K+]. Extracellular [K+] is normally quite low, with the highest reported value being 10 mM (in bee retina), so addition of modest amounts of K+ produces considerable fractional increases in [K+]. Homeostasis of extracellular [K+] has been studied particularly in the retina of the honey bee drone and in the giant nerve fiber of the squid. Regulation of extracellular [H+] (pH) by transport of H+ or HCO3 has been studied in ganglia of the leech, in which the presence of a glial Na+-HCO3 cotransporter was discovered.

Ion homeostasis in the retina of the bee (Apis) The drone (male) honey bee has a highly performing eye, which it uses in pursuing queen bees. The functional units of the retina, the ommatidia, are arranged with an almost crystalline regularity imposed by the facets of the cornea (Figure 5(a) and 5(b)). The retina is composed essentially of two populations of cells – the photoreceptor neurons and the major glial cells, which were originally called ‘outer pigment cells.’ The drone retina is the only nervous tissue in which measurements have been made (using ion-selective microelectrodes) of the free concentrations of the major ions, K+, Na+, and Cl , in the three compartments – neurons, glial cells, and extracellular space. Stimulation of the photoreceptor neurons with light causes considerable shifts of ions between the neurons and glia, whereas the concentrations in the extracellular clefts show only modest changes (Figure 5(c)). It was found that the glial cells clear excess extracellular K+ in two ways. The glial cells are connected to each other by gap junctions. When the activation of the neurons is nonuniform, extracellular [K+] rises locally, and K+ enters the glial cells and somewhat depolarizes the functional syncytium of glial cells. This depolarization drives K+ out of the glial cells in places where extracellular [K+] is only slightly raised. The process is called ‘spatial buffering.’ In addition, glial cells of drone retina also clear extracellular K+ by taking it up together with Cl and water, which leads to swelling of the glial cell. This behavior is not shared by inactive neurons, which take up very little K+. The situation appears to be similar in mammalian brain, in which raised extracellular [K+] causes astrocytes to swell more than do neurons.

Possible role of glial cells in homeostasis of extracellular K+ around the giant axon of the squid The giant axon of the squid (a cephalopod) is unmyelinated but achieves a high conduction velocity by having a large diameter, which can exceed 500 mm. It is surrounded by a thin continuous sheath composed of Schwann glial cells, each approximately 2 mm thick, 20 mm wide, and 100–600 mm long. This sheath is further surrounded by connective tissue. The ion channels of both the axon and the Schwann cells are well characterized. Two Schwann cell conductances have been described – an L-type Ca2+ conductance and an outwardly rectifying K+ conductance activated by entry of Ca2+. These currents are functionally almost identical to conductances found on mammalian Schwann cells. Although propagation of action potentials along the axon causes release of K+ through the axon membrane, little increase in [K+] in the periaxonal space is observed in healthy squid nerves. The K+ conductances of the Schwann cells are not suited to clearing K+. Possibly, [K+] does not rise excessively simply because the extracellular space is large.

Glia and the Removal of Neurotransmitter Astrocytes in vertebrates take up neurotransmitter glutamate. The neuropil glial cells of the leech (Figures 1 and 3), and glial cells of Drosophila (Figure 3(c)), are also equipped to do this. In the retina of the honey bee drone, the neurons release glutamate and the glial cells take it up in the absence of synapses – a situation with parallels to vertebrate nerve tracts, which also release glutamate. In Drosophila, the absence of the transporter dEEAT1 increases oxidative stress. Photoreceptors in insects and barnacle (Balanus, Crustacea) use histamine as their principal neurotransmitter. In barnacle, the glial cells make little or no contribution to recycling this histamine. In contrast, in Drosophila, glial cells are involved in at least two ways. Formation of neurotransmitter vesicles occurs on the neuronal membrane close to glial capitate processes that invaginate the neurons. Also, the enzymes coded by the genes ebony and tan, which are involved in histamine metabolism in the glial cells, are essential to normal neurotransmission.

Glia as a Site of Glycogen Metabolism In many insects, reserves of glycogen and lipids (principally in the glia) are depleted during starvation, and the glial trophospongia which invaginate neurons are well placed to mediate transfer of substrate to the neurons. In isolated slices of bee head, not supplied with glucose or other exogenous metabolic substrate, glial glycogen is broken down to provide metabolic substrate (alanine) which is transferred to the photoreceptor neurons (Figure 5(d)). However, the metabolism of glycogen is active and regulated in complex ways, suggesting that it is not a passive store but has physiological roles in normally fed animals. In isolated ganglia of the leech, electrical stimulation in the absence of exogenous glucose first causes a decrease, then an increase, and then a decrease in glycogen content. Light stimulation in vivo of Apis eye increases, rather than decreases, incorporation of glucose into glycogen in the retinal glia. In isolated leech ganglia, glycogen content is decreased by 5-hydroxytryptamine, dopamine, histamine, and octopamine and increased by GABA and glycine. In the retinas of bee drones, inhibition of glycogen phosphorylase prevents use of glucose suggesting that, as in mammals, glycogen metabolism is essential to normal energy metabolism.

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Glia as Suppliers of Fuel to Neurons Do glial cells, in addition to providing fuel from glycogen during starvation, have a role in the normal supply of metabolic fuel to the neurons? A clear answer is available only for the retina of the drone honey bee. In slices of drone retina, glucose appears to be taken up exclusively by the glia. Pyruvate, formed by glycolysis, is converted to alanine, which leaves the glia and is taken up by the photoreceptors. In the photoreceptors, alanine is reconverted to pyruvate, liberating ammonium, which passes back to the glia, being taken up on a Cl cotransporter specific for NH+4 over K+ (Figure 4(d)).

Glia and the Maintenance of Axon Function Local synthesis of axon proteins It is quite simple to squeeze out most of the axoplasm from a squid giant axon, or to perfuse it, and the ability to do this has contributed greatly to understanding how proteins are maintained in axons. Proteins move down axons at a rate of 1–4 mm day 1 and most have half-lives of less than 1 or 2 weeks. Axons can be many meters long (as in giant squid); thus, at least in axons more than a few centimeters long, proteins have to be produced locally. It is well established that when squid axons, severed from their cell bodies but still ensheathed by glial cells, are bathed in radioactive amino acids, labeled proteins appear in the axoplasm. Furthermore, when the axon is bathed in a labeled RNA precursor ([3H]uridine), labeled RNA appears in the cytoplasm of the axon. Transcription machinery appears to be present within the squid axon. The only known available DNA templates are in the glial cells; thus, presumably RNA is transferred from the glial cells to the axon. The mode of this transfer is unknown and the idea is controversial. Similar conclusions have been reached for the terminals of squid photoreceptors and for the giant Mauthner neuron of goldfish.

Rapid signaling from neurons to glia Neurotransmitter-mediated effects of neuronal activity on glia have been studied particularly in the neuropil glia of the leech (Figures 1 and 3), in the glia surrounding the giant axons of squid and crayfish (Figure 3(b)), and in the glia ensheathing neurons of the snail. In all these cases, bursts of neuronal activity activate a Ca2+-dependent K+ conductance, usually leading to a hyperpolarization. This may improve the clearance of K+ from the extracellular space and the efficacy of Na+-dependent membrane transport, such as Na+–HCO3 cotransport, and glutamate uptake. A striking feature of these glial cells is the presence of receptors for a wide variety of transmitters. Leech neuropil cells, for example, respond to glutamate, 5-hydroxytryptamine, acetylcholine, adenosine phosphates, and myomodulin (a transmitter best known in gastropods). The glial response to an agonist can be extremely complex. In the ventral nerve cord of the crayfish, the passage of a train of impulses along the giant axons causes the axons to release N-acetylaspartylglutamate (NAAG), which leads, by a cascade of events, to hyperpolarization of the periaxonal glia. The main pathway requires the presence of the extracellular enzyme glutamate carboxypeptidase, which splits off glutamate from NAAG, and metabotropic glutamate receptors. Acetylcholine plays a role in the cascade: it is released by the glial cells and acts back on glial acetylcholine receptors leading to activation of K+ channels (Figure 3(b)). All the details of the overlapping pathways have not been worked out, but the analysis has been carried further than in vertebrates, in which elements of this signaling system have been found at the rat neuromuscular junction. At least some elements of this signaling are also found between the giant axon and its Schwann cells in the squid. Signaling between glia and neurons in Drosophila In the mushroom bodies of Drosophila brain, cer encodes an inhibitor of a subfamily of cysteine proteinases named cathepsins. Three hours after conditioned learning, expression of cer in glial cells is transiently reduced (Figure 3(c)). Interference with this pattern of expression impairs long-term memory. In mammals, prolonged activation of cysteine proteinases is associated with neuronal degeneration in Alzheimer’s disease. This is an example of how the genetic manipulations that can be done relatively quickly in Drosophila can give clues to the functioning of mammals. No evidence of multiple neuron–glia signals in bee retina In marked contrast to leech giant glial cells, snail glial cells, and the periaxonal glia of squid and crayfish, no neurotransmitter signaling to the glial cells of drone retina has been reported.

Glia as Actors in the Development of the Nervous System For analyzing the genetic and molecular mechanisms involved in the development of a nervous system, the nematode Caenorhabditis elegans and a few insects have proved useful models. The development is rapid, the number of cells involved is not enormous, and genetic tools are available. At various stages, glial cells can regulate the proliferation, migration, and apoptosis of neurons.

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Early Development of the Nervous System For both Caenorhabditis and Drosophila, neurons and glia are roughly equal players in early development and are derived from common precursors. A landmark discovery was the role in Drosophila of the gene glial cells missing (gcm), whose expression determines the glial fate of all the glia except the midline glia, a small class of neuropil-associated glia. The gene product, Gcm, is a transcription factor that directly or indirectly activates transcription of approximately 50 genes, most of which have mammalian orthologs. Some Gcm-activated genes (e.g., repo, loco, and pnt) determine that a precursor becomes a glial cell rather than a neuron, whereas tramtrack represses neuronal fate. Other Gcm-activated genes are expressed only during a certain time or only in a subset of glial cells. This means that cofactors are required that determine the maturation and diversity of the glia.

Development of Sensory Systems Most insects have sensory systems that are highly perfected yet relatively small, and some have been selected as models for a detailed analysis of their development. This can be illustrated by work on the olfactory system of the moth Manduca sexta.

The olfactory system of the moth, Manduca sexta The main olfactory receptor neurons in insects are in the antennae, and their axons extend to the olfactory lobe of the brain. In Manduca sexta, the receptor neurons for different molecular attributes of odorants are spatially intermixed in the antennal epithelium, but their axons, on entering the olfactory lobe, are sorted into fascicles according to their odorant specificity and terminate in glomeruli whose number increases during maturation to reach 64  1 in the adult. This system develops over approximately 18 days during the metamorphosis from larva to adult. The axons of the olfactory receptor neurons initially grow down the antenna without glia (Figure 6(a)). On arriving outside the antennal lobe, the axons encounter sorting zone glial cells and reciprocal neuron–glia interactions occur. The axons send signals to the glia, which then increase in number. Also, the glia contribute to the sorting of the axons: On contacting glia, the growth cones enlarge, explore, turn, and find like axons (Figure 6(c), AN

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Figure 6 Glial cells contribute to the development of the olfactory system in the moth Manduca sexta. (a, b) The antennal lobe with the base of the antennal olfactory nerve (top left). During metamorphosis, axons from the olfactory epithelium (not shown) are sorted to appropriate odor-specific glomeruli. Probable neuron–glia interactions are numbered in (c). (1) The first axons to reach the lobe encounter central glial cells and induce them to proliferate. (2) Glia migrate into the nerve to become sorting zone glia. Axons encountering sorting zone glia form large growth cones. (3) In vitro, the growth cones of axons contacting neuropil glia cells become larger and more complex. (4) Glia arising in the olfactory receptor epithelium migrate along the olfactory nerve (not shown in b). (5) Receptor axons induce neuropil glial cells to extend stabilizing processes around protoglomeruli. (6) Antennal nerve glial cells enwrap bundles of receptor axons. (7) Neuropil glia define the borders of the glomeruli. (d) The signaling mechanism involved in step (5). Acetylcholine released by the axon terminal depolarizes the glial cell, and the depolarization opens voltage-gated Ca2+ channels, leading to an increase in intracellular [Ca2+]. (a, b, d) Adapted from Lohr C and Deitmer JW (2006) Calcium signalling in invertebrate glial cells. Glia 54: 642–649. (c) Adapted from Tolbert LP, Oland LA, Tucker ES, et al. (2004) Bidirectional influences between neurons and glial cells in the developing olfactory system. Progress in Neurobiology 73: 73–105.

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stages 4–6). Glial cells of a second kind, the olfactory ensheathing cells, migrate along the axons from the receptors and, later, enwrap bundles of axons. In the olfactory lobe, the neuronal cell bodies are clustered in its periphery, outside an initially spherical zone of neuropil (Figure 6(a)). The cell bodies of the neuropil glial cells surround this neuropil. The outer parts of the neuropil progressively separate into glomeruli, and the layer of neuropil glial cells is deformed to follow this change of shape (Figure 6(b)). The neuropil glia show reciprocal interactions with the receptor cell axons. The axons induce properties of the neuropil glia, including expression of Ca2+ channels. Also, the neuropil glia induce synapse formation. Some elements of the neuron–glia signaling are known. In the case of the neuropil glia, acetylcholine released by the ingrowing receptor axon terminals depolarizes the glia. This activates voltage-dependent Ca2+ channels, and the resulting increase in intracellular free [Ca2+] in some way initiates glial cell migration (Figure 6(d)).

A glial function, myelination, is responsible for major differences in the sizes and life styles of vertebrates and invertebrates Electrical signaling by changing the electrical potential across a neuronal plasma membrane consisting of a lipid bilayer is slow and energetically expensive. This is because the capacitance of the lipid bilayer is very high (1 mF cm 2), which means that many ions have to cross it to change the potential. In many vertebrate nerves, the effective membrane capacity of the axon membrane is reduced by a myelin sheath, produced by glial cells. This makes possible the high nerve impulse conduction velocities necessary for large animals capable of rapid and complex movements, such as dinosaurs and humans. The structure most resembling myelin in invertebrates may be the several loosely wrapped layers of membrane that are sometimes found around axons of Drosophila, although Drosophila possesses homologs of only a few of the genes required for myelin. This wrapping is probably not useful for conducting active nerve impulses, but it would speed up passive, electrotonic conduction. The invertebrate solution to the problem of nerve conduction velocity is either to stay small (as are insects) or to make axons of very large diameter. These giant axons take up much space and their use is mainly limited to stereotyped escape reactions, as is the case in annelids and squid.

Lessons from Invertebrate Glial Cells The conclusion from the small number of invertebrate glial cells whose functions have been studied is that in adult animals the proteins that serve as ion channels, transporters, receptors, and elements of intracellular signaling pathways have properties very similar to those of vertebrate glia. It is true that some invertebrate glia show behavior that has not been clearly described in vertebrates, such as a possible involvement of cathepsin in memory formation in Drosophila, or the choice of alanine, rather than lactate, as a substrate transferred from glia to neurons, as in bee retina. However, it would be premature to state that similar phenomena do not occur in vertebrates. The involvement of glial cells in the development of insect nervous systems also shows many close parallels with mammals, although many of the genes involved appear to have different functions in vertebrates. It should be borne in mind that an invertebrate nervous system has evolved through many more life cycles than has that of Homo sapiens, even though the body shape of the animal may have changed little over several hundred million years.

Further Reading Carlson SD and SaintMarie RL (1990) Structure and function of insect glia. Annual Review of Entomology 35: 597–621. Coggeshall RE and Fawcett DW (1964) The fine structure of the central nervous system of the leech, Hirudo medicinalis. Journal of Neurophysiology 27: 229–289. Coles JA, Orkand RK, Yamate CL, and Tsacopoulos M (1986) Free concentrations of Na, K, and Cl in the retina of the honeybee drone: Stimulus-induced redistribution and homeostasis. Annals of the New York Academy of Sciences 481: 303–317. Coles JA and Deitmer JW (2005) Extracellular potassium and pH: Homeostasis and signalling. In: Kettenmann H and Ransom BR (eds.) Neuroglia, 2nd edn., pp. 334–345. New York: Oxford University Press. Coles JA (1989) Functions of glial cells in the retina of the honeybee drone. Glia 2: 1–9. Comas D, Petit F, and Preat T (2004) Drosophila long-term memory involves regulation of cathepsin activity. Nature 430: 460–463. Coutinho-Budd J and Freeman MR (2013) Probing the enigma: Unraveling glial cell biology in invertebrates. Current Opinion in Neurobiology 23: 1073–1079. Freeman MR, Delrow J, Kim J, Johnson E, and Doe CQ (2003) Unwrapping glial biology: Gcm target genes regulating glial development, diversification, and function. Neuron 38: 567–580. Garm A, Ekstro¨m P, Boudes M, and Nilsson DE (2006) Rhopalia are integrated parts of the central nervous system in box jellyfish. Cell and Tissue Research 325: 333–343. Gommerat I and Gola M (1996) Glial potassium channels activated by neuronal firing or intracellular cyclic AMP in Helix. Journal of Physiology 495: 649–664. Inoue I, Tsutsui I, Abbott NJ, and Brown ER (2002) Ionic currents in isolated and in situ squid Schwann cells. Journal of Physiology 541: 769–778. Ito K, Urban J, and Technau GM (1995) Distribution, classification, and development of Drosophila glial cells in the late embryonic and early larval ventral nerve cord. Roux’s Archives of Developmental Biology 204: 284–307. Kuffler SW and Potter DD (1964) Glia in the leech central nervous system: Physiological properties and neuron-glia relationshhip. Journal of Neurophysiology 27: 290–320. Lohr C and Deitmer JW (2006) Calcium signalling in invertebrate glial cells. Glia 54: 642–649. Marcaggi P, Thwaites DT, Deitmer JW, and Coles JA (1999) Chloride-dependent transport of NH4+ into bee retinal glial cells. European Journal of Neuroscience 11: 167–177. Parker RJ and Auld VJ (2006) Roles of glia in the Drosophila nervous system. Seminars in Cell & Developmental Biology 17: 66–77. Pentreath VW (1995) Metabolic interactions between neurons and glial cells in leech and snail ganglia. In: Vernadakis A and Roots BI (eds.) Neuron–glia interactions during phylogeny, pp. 161–196. Totowa, NJ: Humana Press. Pereneanu W, Shy D, and Hartenstein V (2005) Morphogenesis and proliferation of the larval brain glia in Drosophila. Developmental Biology 283: 191–203. Rival T, Soustelle L, Strambi C, Besson MT, Iche M, and Birman S (2004) Decreasing glutamate buffering capacity triggers oxidative stress and neuropil degeneration in the Drosophila brain. Current Biology 14: 599–605.

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Shaham S (2005) Glia–neuron interactions in nervous system function and development. Current Topics in Developmental Biology 69: 39–66. Tolbert LP, Oland LA, Tucker ES, et al. (2004) Bidirectional influences between neurons and glial cells in the developing olfactory system. Progress in Neurobiology 73: 73–105. Tsacopoulos M, Poitry-Yamate CL, and Poitry S (1997) Ammonium and glutamate released by neurons are signals regulating the nutritive function of a glial cell. Journal of Neuroscience 17: 2383–2390. Urazaev AK, Grossfeld RM, and Lieberman EM (2005) Regulation of glutamate carboxypeptidase II hydrolysis of N-acetylaspartylglutamate (NAAG) in crayfish nervous tissue is mediated by glial glutamate and acetylcholine receptors. Journal of Neurochemistry 93: 605–610.

Relevant Websites http://flybase.bio.indiana.edu FlyBase. http://www.sdbonline.org Society for Developmental Biology. http://www.wormatlas.org Wormatlas. Altun ZF and Hall DH (eds.) 2002–2006.