Sensory cilia in arthropods

Arthropod Structure & Development 41 (2012) 515e534

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Review

Sensory cilia in arthropods Thomas A. Keil* Max-Planck-Institute of Biochemistry, Department of Molecular Structural Biology, Am Klopferspitz 18, D-82152 Martinsried, Germany

a r t i c l e i n f o

a b s t r a c t

Article history: Received 8 March 2012 Accepted 3 July 2012

In arthropods, the modified primary cilium is a structure common to all peripheral sensory neurons other than photoreceptors. Since its first description in 1958, it has been investigated in great detail in numerous sense organs (sensilla) of many insect species by means of electron microscopy and electrophysiology. The perfection of molecular biological methods has led to an enormous advance in our knowledge about development and function of sensory cilia in the fruitfly since the end of the last century. The cilia show a wealth of adaptations according to their different physiological roles: chemoreception, mechanoreception, hygroreception, and thermoreception. Divergent types of receptors and channels have evolved fulfilling these tasks. The number of olfactory receptor genes can be close to 300 in ants, whereas in crickets slightest mechanical stimuli are detected by the interaction of extremely sophisticated biomechanical devices with mechanosensory cilia. Despite their enormous morphological and physiological divergence, sensilla and sensory cilia develop according to a stereotyped pattern. Intraflagellar transport genes have been found to be decisive for proper development and function. Ó 2012 Elsevier Ltd. All rights reserved.

Keywords: Cilia Primary Chemoreception Mechanoreception Sensory transduction Intraflagellar transport Sensilla

1. Introduction Cilia have gained increasing attention in cell biology approximately during the last 20 years. First, intraflagellar transport was one of the main topics (cf. Johnson and Rosenbaum, 1993; Kozminski et al., 1993, 1995; rev. Rosenbaum and Witman, 2002). Second, their role in determination of the lefteright-asymmetry during vertebrate embryogenesis made highlights (cf. Hirokawa et al., 2006). And last but not least, it became more and more clear that numerous human diseases are caused by ciliary malfunctions (first described by Afzelius et al., 1975; rev. by Afzelius, 2004; Marshall, 2008). Cilia which lacked the central microtubule pair and grew from deep inside cells have been described first by Sorokin (1962). Initially, they were called “rudimentary”, and later “primary” cilia. They were considered to be immotile. It turned out by and by that these cilia occur in many, if not most, animal cell types, disappearing only during cell division, when their basal bodies turn into centrioles and become associated with the mitotic spindle. The function of mammalian cilia as receptors and signaling centers has been investigated in great detail during the last 10 years (reviewed recently by Berbari et al., 2009; Gerdes et al., 2009). Primary cilia have a mechanoreceptive function in the kidney,

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where they sense fluid flow, but the mechanism is not clear (cf. Aznar and Billaud, 2010). Bloodgood (2010) came to the conclusion that all cilia have a sensory function. A recent theory on the phylogeny of cilia by Jékely and Arendt (2006) says that they originated from a “sensory membrane patch”, their original role being that of a “sensorium” (Quarmby and Leroux, 2010). The electron microscope (EM) showed during the 1950s that the sensory cells of the vertebrate retina have a modified cilium as their receptive structure (De Robertis, 1956). Cilia or flagella were believed not to exist in insects, with the exception of the spermatozoan tail, until the first epidermal sensory organs (“sensilla”) of insects were investigated in the EM by Slifer et al. (1957) and then by Gray and Pumphrey (1958). It was found that the sensory cells bear a modified cilium as the receptive structure, and a very detailed description was given by Gray (1960). With the exception of the photoreceptors and multidendritic receptors, a modified primary cilium (i.e., without the central microtubule pair) is the common structure of sensilla in the world of insects and all other arthropods. An exemplary insect sensillum (Fig. 1) is a complex consisting of specialized external cuticular structures, mostly shaped as hair or peg, and a number of bipolar sensory neurons surrounded by several accessory, or “enveloping” cells which perform a dual function. First, these cells build the cuticular structures during development (Fig. 1aec; Hartenstein, 2005; Keil, 1997b). The outermost “tormogen” cell forms the socket, the “trichogen” cell the hair, and the innermost “thecogen” or “scolopale” cell an extracellular, probably cuticular sheath

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Fig. 1. Showing the later developmental stages of an olfactory sensillum of the silkmoth Antheraea polyphemus, modified after Keil and Steiner (1991). Red: sensory neuron; green: thecogen (¼sheath-forming) cell; orange: trichogen (¼hair-forming) cell; purple: inner tormogen (socket-forming) cell; blue: outer tormogen cell. Note that this type of sensillum has two tormogen cells, the inner one will degenerate. (a) Arrangement of cells at the beginning of cuticle deposition. The cilium (“primary dendrite”), enclosed by the “cuticular” dendrite sheath (arrowhead), projects sideways from the hair-forming process (cf. Fig. 5b,c). (b) The inner tormogen cell has disappeared except a small rudiment, and the “primary dendrite” and sheath have been lost. Now the dendrite has retracted, the cilium is growing out a second time, and the definite sheath is formed (arrowhead). (c) Deposition of cuticle is completed, the hair-forming process retracts from the hair shaft, and the sensory cilium grows out into the hair. (d) Adult stage: Trichogen and tormogen cell have retracted from the hair and now form the sensillum lymph cavity (slc). The cilium is now the “outer dendritic segment” (od).

around the sensory cilia. Second, tormogen and trichogen cell form the physiological environment (“sensillum lymph cavity”) for the receptors during the functional stage (Fig. 1d). The receptors proper are localized in the primary cilia (called “outer dendritic segments”) which are the distal projections of the sensory dendrites (“inner dendritic segments”). The cilium has adapted to functions such as olfaction, gustation, hygro-, thermo-, and mechanoreception, and developed the most sophisticated cellular apparatus for each “application”. The same, of course, holds true for the cuticular structures which are highly specialized for their different tasks (rev. by Keil, 1997a,b, 1999; Keil and Steinbrecht, 1984; Steinbrecht, 1998, 1999). Since the turn of the century, the focus of cilia research seems to shift more and more toward the fruitfly, Drosophila melanogaster. Here, all tools of genetics are at the researcher’s service, and it is possible to investigate primary cilia (9 þ 0) in the sensory neurons of epidermal sensilla in parallel with motile (9 þ 2) cilia in sperm cells. The hot topic of intraflagellar transport has been successfully approached in sensory organs of this species (e. g., Avidor-Reiss

et al., 2004). And finally, investigations on Drosophila suggested that an essential reason for the association of centrioles with the mitotic spindle might be their proper distribution to the daughter cells as a prerequisite for the generation of cilia. This was shown by Basto et al. (2006) in Drosophila Dsas-4 mutants, which have lost all centrioles by the third larval stage. These animals develop into morphologically normal looking adults but they cannot develop (mechano) sensory cilia and consequently are uncoordinated and die. The question, therefore, arose whether the centrioles are “active players or merely passengers” during mitosis (Debec et al., 2010; Gogendeau and Basto, 2010). Richard Eakin proposed in 1965 that there exist two phylogenetic lines of photoreceptors: first, the ciliary, and second, the microvillar. But Eakin and Westfall (1965) had already found a “rudimentary cilium” in the microvillar photoreceptor cell of Peripatus, and centrioles are present in developing insect photoreceptor cells (Home, 1972; Wachmann and Hennig, 1974). Finally, it became clear that both types of photoreceptors co-exist in several phyla (e. g., annelids, mussels, chordates: Fain et al., 2010).

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As it turns out, both lines of receptor cells, adapted to different sensory modalities, exist in most animal species (e. g., in mammals the olfactory and photoreceptors are of the ciliary type, whereas the auditory and taste receptors are of the microvillar type; in insects the chemo-, thermo-, and mechanoreceptors are of the ciliary type and the photoreceptors are of the microvillar type). Ulrich Thurm demonstrated the mechanosensitivity of mussel gill cilia in 1968, and in 1969 drew the attention to the ciliary type in plathelmintes, annelids, and arthropods. Wiederhold (1976) gave the first extensive review on mechanosensory transduction in cilia. Altner and Prillinger (1980) reviewed the ultrastructure of chemo-, thermo-, and hygroreceptors, summarizing what was then known about sensory cilia of invertebrates. After 32 years which have passed since this work, it seems worthwhile to have a closer look at our recent knowledge about sensory cilia in insects to get a more complete view of this highly variable organelle. Since D. melanogaster has taken the center stage of sensillum research, a wealth of new insight, especially in the field of mechanoreception, has been gained. Therefore, chapter 7 is a little more extensive. Wherever possible, previously unpublished micrographs have been used to illustrate this review. If no other reference is given, they were taken from my own work. The specimens were chemically fixed unless noted otherwise. 2. The normal and not so normal structure of insect sensory cilia Gray and Pumphrey (1958) were the first to find a ciliary structure in the ear of Locusta migratoria. Gray (1960) did a thorough electron microscopical investigation of the chordotonal organ and gave the first detailed description of an insect sensory cilium. Slifer and Sekhon (1960) described the ciliary region of the sensory dendrite in the plate organ of Apis as follows: “each dendrite suddenly decreases in diameter and assumes the fine structure of a typical cilium”. The most detailed investigation of the sensory cilia in larval sensilla of Dysdercus intermedius (Heteroptera) and Agrion spec. (Odonata) was done by Gaffal and Bassemir (1974). It is now clear that the basic construction of the sensory cilium is similar in all insects, but shows variations with respect to the sensory modality. An extensive discussion can also be found in Field and Matheson (1998). Fig. 2a is a cartoon of a typical ciliated bipolar sensory neuron. It gives off an axon at its basal and a dendrite at its apical pole. In the distal tip of the dendrite, a centriole pair arranged in tandem is found (Figs. 2b and 3a), which sends more or less elaborate crossstriated rootlets in the proximal direction. The distal end of the inner dendritic segment very often forms marked “shoulders” after chemical fixation (e.g., Fig. 5 in Ernst, 1969, Fig. 13 in Keil and Steiner, 1991). On the other hand, the transition from inner to outer dendritic segment is always very smooth in cryofixed specimen (Fig. 4; Steinbrecht, 1980). Maybe this indicates that the relatively slow process of chemical fixation can induce a contraction of the ciliary rootlet which is avoided by the fast cryofixation. Root fibers originate from both centrioles (Fig. 2b). The first population arises from the base of the distal centriole, tightly enclosing the proximal centriole which gives off a second population of root fibers. Both populations unite beneath the proximal centriole, either remaining as parallel running individuals or mostly fusing into a massive common root bundle. It is somewhat depending on the sensory modality of the cilium how far they run into the inner dendritic segment: in chordotonal sensilla, they can even enter the cell body and reach beyond the nucleus. The rootlets always show a periodic cross striation (e. g., Schmidt, 1969). It

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seems that the rootlets had already been observed by the light microscopists (several figures in Eggers, 1928). Wolfrum (1991, 1997) tested the ciliary rootlets in chordotonal sensilla of Periplaneta for the presence of collagen. The result was negative, in contrast to control preparations definitely known to contain collagen. But he was able to detect centrin (a protein also present in the ciliary rootlets of green algae) and alpha-actinin. Rootletin is a large coiled-coil protein which has been found to be the main component of the ciliary rootlets in mammalian photoreceptor cells (Yang et al., 2002). It is needed for the stability of the cilia (Yang et al., 2005). It interacts with kinesin and therefore might be involved in intracellular transport (Yang and Li, 2005). It also serves as a physical linker between the centrioles (Yang et al., 2006). Drosophila has genes encoding a rootletin homolog (Kernan, 2007), it is reasonable to assume a central function for this protein in insect sensory cilia. The distal centriole (basal body) forms a 9  2 þ 0-cilium. The microtubule pairs may run into the very tip of this cilium. In the case of the chordotonal organ, this ciliary structure does not change. In most other sensillum types, the cilia can assume the most variable shapes depending on their sensory modality, and increase the number of microtubules up to 1000 in certain mechanoreceptors (Nicklaus et al., 1967). The number of microtubule doublets can also increase to over 40 in Acheta domesticus (Keil, unpublished observation). These variations will be described below. In most sensilla, the microtubules appear smooth and hollow, but it has been observed in chordotonal sensilla that the Atubule of each pair appears solid, and that it projects two tiny side arms similar to dynein (Fig. 9b; Corbière-Tichané, 1971b; Toh and Yokohari, 1985; rev. by Field and Matheson, 1998). Lateral projections on the microtubule pairs have been found in several sensilla at the ciliary base in the region of the ciliary necklace (Fig. 3b); they often form Y-shaped connections to the membrane (Fig. 3c; Fig. 2; Gaffal and Bassemir, 1974). The molecular structure of these connectors and their possible function as controls of the entry of soluble proteins into the cilium has recently been discussed in detail by Garcia-Gonzalo and Reiter (2012). In freeze-fractured preparations of olfactory and gustatory sensilla, Menco and van der Wolk (1982) found approximately 200 membrane particles per square micrometer in this region. The central microtubule pair is always missing, but sometimes a small vesicular structure has been observed in its place (Fig. 3a,b). The basalmost region of the cilium can be interpreted as a “transition zone” (e.g., Ma and Jarman, 2011). These authors investigated the dilatory mutant of Drosophila in which formation of chordotonal cilia is disturbed. The DILA protein normally found in this transitory region seems to be necessary for intraflagellar transport at the ciliary base, probably interacting with the UNC (uncoordinated) protein. The transition zone acts as a diffusion barrier necessary for membrane compartmentalizing, separating the ciliary membrane proper from the cell membrane (Hu and Nelson, 2011) and regulating the traffic between both compartments (Chih et al., 2012). Recently, it turned out that some proteins which belong to the nuclear pore complex are found also in the barrier region of the ciliary base (cf. Obado and Rout, 2012). There are a few exceptions to the typical appearance. For example, the tandem arrangement of the centrioles is disturbed in filiform sensilla on the cricket cercus. Another case is the so-called “granular” or “fibrillar” body found in mechanoreceptors of certain flies (the blowfly, Calliphora: Richter, 1964; Smith, 1969; Keil, 1978; but not the fruitfly, Drosophila : Chevalier, 1969; Avidor-Reiss et al., 2004). The centriole pair is formed normally, but during development the distal one as well as the cilium base increase in diameter and deposit a granular material (Keil, 1978). A similar structure has been described also by Eakin and Westfall (1964) in photoreceptor

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Fig. 2. (a) Unspecified bipolar sensory neuron of an insect with apical cilium. (b) Ciliary region of a chemosensory neuron at higher magnification. The sequence of the cross sections from proximal to distal is indicated by numbers 1e10. 1,2: Sections through the root fiber bundles. 3. Section through the proximal basal body (centriole). 4: Section between the basal bodies (centrioles). 5: Section through the distal basal body (centriole). 6,7: Sections through the transition zone. 8,9: Sections through the level of the Y-shaped connectors. 10: At this level, the cilium has increased its diameter and number of microtubules. (b) From Gaffal and Bassemir (1974), Fig. 7. With kind permission from Springer Science þ Business Media.

cells of the arrowworm Sagitta. Nothing can be said about its function. Other structures not normally found in cilia are the 10 nm-filaments which have been described in the mechanosensory cilia of Calliphora (Keil, 1978; Smith, 1969; Voelker, 1982). Variations in the number of ciliary doublets (up to 11) have been found in sensilla of the tick Amblyomma americanum (Foelix and Axtell, 1971, 1972). In monociliated sensory neurons of crustaceans, only one single centriole seems to be the rule (cf. Schmidt and Gnatzy, 1984; Crouau, 1987). 3. The way of the centrioles There is only limited knowledge about the origin and development of the ciliary basal bodies in insect sensory cells, and the following description is based entirely on electron microscopic observations in embryos of the cockroach Periplaneta americana (Seidl, 1991), and developing pupae of the blowfly Calliphora vicina (De Kramer and van der Molen, 1984) and the silkmoth Antheraea polyphemus (Keil and Steiner, 1990a,b, 1991). Centrioles can be found everywhere in the epidermis (cf. Whitten, 1973; Bassemir

and Hansen, 1980), they can even form short ciliary stumps (Seidl, 1991). Once a sensillum mother cell (or sensory organ precursor 1: SOP 1) has been singled out in the epidermis (cf. Hartenstein, 2005), a centriole pair becomes localized in its apex. In the cells which originate from the differential mitoses of this cell, the centrioles are first found in close association with the mitotic spindles, and finally with the re-constituting cell nuclei. They then migrate to the distal regions of their respective cells. Each cell destined to become a sensory neuron forms a slender distal process, which becomes the “inner dendritic segment” (in analogy to the vertebrate photoreceptor). In the tip of the inner dendritic segment, the centriole pair can be found (Fig. 5e), from where it gives rise to a primary cilium which is to become the “outer dendritic segment”. 4. How cilia grow, and how they stick to an ancient developmental principle The primary cilium grows out from inside the apical process of the prospective sensory neuron above the epidermal surface (Figs. 1

Fig. 3. Ciliary structures in different types of insect sensilla. (a) Longitudinal section of the ciliary base in an olfactory s. trichodeum of the silkmoth Antheraea polyphemus. Note the submembrane density and the central vesicle in the neck region (transition zone, arrowheads), and the very short ciliary root just reaching from the distal slightly beneath the proximal basal body (bb). Bar ¼ 0.5 mm. (b) Cross section through the ciliary base of an unspecified mechanoreceptor cell in the housecricket Acheta domesticus. A tiny central vesicle is visible, as well as small outward projections of the microtubule doublets. In this type of neuron, the Y-shaped connectors are absent. Bar ¼ 0.2 mm. (c) Cross section through the ciliary base of an olfactory neuron of a pore plate organ of the honeybee, Apis mellifera. The Y-shaped connectors between microtubule doublets and ciliary membrane are clearly visible. The doublets themselves are interconnected by a ring-shaped structure. Bar ¼ 0.1 mm. (d) Cross section through the base of an unidentified sensillum in Acheta, showing a cross-sectioned centriole in the inner dendritic segment (id) as well as the centriole tandem in the thecogen cell (th). Bar ¼ 0.5 mm.

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Fig. 4. Ciliary base after cryofixation. Independent of the method used, the transition is smooth. (a) Olfactory sensillum on the antenna of Bombyx mori. Plunge-frozen, cryosubstituted, embedded in Epon. The centrioles are not visible in this section. Bar ¼ 1 mm. From Steinbrecht (1980), by permission of Elsevier. (b) Olfactory sensillum on the antenna of Drosophila melanogaster. High-pressure frozen, cryosubstituted, refrozen and cryosectioned according to Tokuyasu (Method: Ripper et al., 2008). Note that membranes are not visible after this procedure. Arrowheads point at centrioles. Bar ¼ 1 mm.

and 5; Ernst, 1972; Keil and Steiner, 1991; Seidl, 1991). The latter authors clearly showed that ciliary outgrowth starts with a vesicle forming over the distal basal body, into which the cilium grows e exactly as described by Sorokin (1962). Additionally, numerous coated membrane vesicles are found in the tip of the inner dendritic segment (Fig. 8a in Seidl, 1991). It seems that the membrane material needed for early outgrowth of the cilium into its own cavity is simply taken from the outer membrane of the dendrite, which retracts during this time (see also De Kramer and van der Molen, 1984). Quite remarkable is the fact that Seidl also observed the initial stages of cilium development in the three enveloping cells of the sensillum, but he claims that the “basal bodyevesicle complexes” are lost later in development. However, Bassemir and Hansen (1980) found the centriole pairs to persist in all 3 enveloping cells of larval sensilla of the damselflies Agrion puella and Ischnura elegans, and at least in the thecogen cell a centriole pair arranged in tandem and forming rootlets, looking exactly as in the sensory cell, has been found to persist in Acheta (Keil, unpublished observations: Fig. 3d). The developmental time course in chordotonal organs of the Drosophila embryo has been investigated by Hartenstein (1988) and Carlson et al. (1997). From these papers, it can be concluded that the outgrowth of the sensory cilia takes place within 2 h. In the external sensilla, the outgrowing cilium is partly enclosed by a thin cuticular sheath secreted by the innermost enveloping (thecogen) cell (Fig. 5a,d). Following the outgrowth of the cilium, the second enveloping (trichogen) cell forms an apical process

(“trichogen sprout”), which encloses the cilium at its base in olfactory sensilla (Fig. 5b,c,d), whereas it encloses the cilia for almost their whole length in gustatory sensilla (Hansen and Hansen-Delkeskamp, 1983; De Kramer and van der Molen, 1984). The cilia show a 9  2 þ 0-cross section only in the basalmost region, but this is lost already inside the trichogen sprout (Fig. 5d). The primary cilia still project over the surface of the epidermal and the hair-forming cells, while these start to secrete cuticle, but are lost when the cuticle gets thicker (Fig. 1a,b). In an external mechanoreceptor, the tip of the cilium remains attached to the inner cuticle of the hair base (Schmidt and Gnatzy, 1971). In olfactory sensilla, the cilia grow out a second time after the trichogen cell has formed the cuticular hair shaft and retracts from it (Fig. 1b,c; Ernst, 1972; Keil and Steiner, 1991). In chemosensilla of flies, considered to be the highest evolved insects, the cilia grow only once into the already formed trichogen sprout (Kuhbandner, 1984). This means that the primary outgrowth is omitted. Why, then, do most sensory cilia grow out two times? Why, as an example, do the primary cilia project sideways from the base of the developing olfactory hairs in the pupae of Antheraea and Necrophorus, only to be lost soon? The answer can be found in the (phylogenetically old) hemimetabolous insects (i.e., grasshoppers, crickets and cockroaches: Slifer et al., 1959; Schmidt and Gnatzy, 1971; Gnatzy and Schmidt, 1971; Moran, 1971; Moran et al., 1976; Gnatzy, 1978), and also in the “apterygote” insects (Berg, 1994; Berg and Schmidt, 1996, 1997). In preparation for moulting, the old cuticle is detached first from

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Fig. 5. Stages of sensillum outgrowth in the pupa of Antheraea. (a) The primary cilia are projecting over the epidermis. They are partly enclosed by their cuticular sheath; the second from the left projects distally (arrowhead). The trichogen sprouts (s) are just beginning to grow (Nomarski optics). Bar ¼ 20 mm. (b) The trichogen sprouts have already reached a certain length, while the cilia are projecting sidewards from their bases. The right one can be seen projecting from its sheath (arrowheads) (Nomarski optics). From Keil and Steiner (1991), Fig. 7. Bar ¼ 20 mm. (c) Same stage, showing the base of the trichogen sprout with the cilia (arrowheads). Bar ¼ 1 mm. (d) Cross section of the cilia within the trichogen sprout. They have already lost their typical 9  2 þ 0-configuration and are tightly enclosed in the dendrite sheath (arrowhead). From Keil and Steiner (1991), Fig. 19. Bar ¼ 0.2 mm. (e) Cross section of a centriole in the early developing dendrite (before outgrowth of cilium). Some of the 9 radial spokes as well as the central hub are clearly visible (cf. Kitagawa et al., 2011). From Keil and Steiner (1990), Fig. 14. Bar ¼ 0.2 mm.

the epidermis (apolysis; described in detail by Gnatzy and Romer, 1980). However, the sensory organs continue to function normally (Gnatzy and Tautz, 1977); therefore, the sensory cilia keep their connection to the old sensilla. They grow out considerably while the distance between the old cuticle and the epidermis increases. In order to be protected from the moulting fluid, they have to be enclosed in a cuticular sheath, which is secreted by the thecogen cell and grows with the elongating cilia. The new cuticle as well as the new sensilla develop under the old cuticle. This means that openings in the walls of the new sensilla must persist, through which the cilia can project. In most sensilla, this opening is found at the base of the hair (e. g., mechano- and olfactory sensilla), or the cilia project from the apical opening in gustatory sensilla. The sensilla thus can remain functional until the old cuticle is shed (Gnatzy and Tautz, 1977). The elongated cilia are lost together with

their sheaths; the remains can be found attached to the inner side of the exuvia (Schmidt and Gnatzy, 1971; Moran et al., 1976). Obviously, this elementary mechanism has persisted into the holometabolous insects, where it is of course needed during larval development, but no longer in the pupal stage where it has been skipped only very late in evolution (that means, in the dipterans). 5. Olfaction and taste: from simple cilia to spirals and branches The structure of insect olfactory sensilla has been reviewed extensively by Keil (1999) and Steinbrecht (1999). Chemosensory cilia, such as found in olfactory and gustatory sensilla, can reach a considerable length of up to 400 mm, as on the antenna of the hawkmoth Manduca (Keil, 1989; Lee and Strausfeld, 1990). Their

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morphological differentiation is surprisingly low. Three-dimensional reconstruction from serial sections of freeze-substituted sensilla trichodea of the silkmoth Antheraea showed first, that the sensory cilia can be wound around each other like tendrils, and second, that they show numerous swellings along their length (Figs. 6 and 7b; Keil, 1984a). In sensilla trichodea, there is just an increase in diameter and microtubule number (Fig. 7a), whereas in s. basiconica and s. placodea they can be split up into numerous thin, but loosely packed, branches (Fig. 7b). The swellings have been shown very clearly in cryosubstituted chemosensilla of P. americana (Seidl, 1991). Another feature of the olfactory cilia to be mentioned is the negatively charged surface coat of their membranes, which has been demonstrated by application of cationic markers (Keil, 1984b, 1987). Leal et al. (2005) pointed out that these negative charges result in a local low pH close to the receptor membrane, necessary for interaction with the olfactory binding protein-pheromonecomplex. Taste sensilla in the fly gave the first opportunity to get direct electrophysiological access to the sensory cilia because of their apical opening (Wolbarsht and Dethier, 1958). In the extraordinary long (up to 300 mm) olfactory sensilla of the giant silkmoth Antheraea, direct access to olfactory cilia could be gained by pinching off the tips with sharpened tweezers (Kaissling et al., 1991). In the same species, it was then possible to directly extrude the cilia from the opened tips (Williams, 1988; Keil, 1993) and make patchclamp recordings from them (Zufall and Hatt, 1991).

6. Carbon dioxide and temperature: how cilia increase their surface Stange and Stowe (1999) reviewed the structural peculiarities of the CO2-receptors in detail. In these sensilla, the cilia show a highly elaborated structure. They are characterized by an extensive increase of membrane surface, which is reached in two different ways: 1. intensive branching, while the branches tightly fill the lumen of the sensillum, and 2. an intensive folding (Lee et al., 1985). The single cilium increases in thickness until it completely fills the hair lumen, and then suddenly divides either into numerous thin branches, such as in the honeybee (Fig. 8a), or tightly packed lamellae, such as in moths (Fig. 8b,c). The dense packing of the membranes is considered as advantageous, as it also would mean dense packing of receptors (Stange and Stowe, 1999). However, exceptions have been observed in CO2-receptors of caterpillars, where the lamellae are only loosely packed (Keil, 1996). In Drosophila, the CO2-receptors are not located within a separate sensillum. The large antennal basiconic sensillum LB-II 4 contains 4 sensory neurons, the cilia of which split into numerous loosely packed thin branches, and one of these cilia can assume a complicated lamellar shape (Fig. 8d; Shanbhag et al., 1999). Electrophysiological investigations by De Bruyne et al. (2001) showed that 3 of these neurons respond to olfactory stimuli, while the fourth responds to CO2. Thermoreceptors are combined with hygroreceptors within the same sensillum (rev. by Steinbrecht, 1998). The former are characterized by a highly lamellated sensory cilium ending directly under the hair base, whereas the latter are tightly enclosed in the cuticle of the hair shaft. The most elaborate lamellation of sensory cilia had been described in thermo-hygroreceptors of a cavernicole beetle (Corbière-Tichané, 1971a), combination with dense packing has also been found in this type of sensillum (Steinbrecht, 1998). The membranes of these lamellae are interconnected by very conspicuous dense surface structures, which have been found in chemically fixed (Corbière-Tichané, 1971a) as well as cryosubstituted (Haug, 1985) specimens. They have been termed “BOSSes” by Steinbrecht (1989). It must be pointed out that this differentiation has also been described in the diplopod Polyxenus lagurus (Nguyen Duy-Jacquemin, 1983). 7. Mechanoreception: from 9 3 2 D 0 to giant microtubule complexes

Fig. 6. Reconstruction of the course of two olfactory cilia in a s. trichodeum of the silkmoth Antheraea. Their normal length would be between 250 and 300 mm, it is foreshortened in this drawing by a factor of 17.

Two different types of mechanoreceptors will be discussed in the following: 1. the internal and 2. the external. The internal receptors belong mostly to the chordotonal or scolopidial type, which is characterized by a straight and slender 9  2 þ 0-cilium (Fig. 9); the multiple dendritic receptors (cf. Bodmer and Jan, 1987) are not considered here. The sensory cilium is attached to the body wall indirectly via a cuticular cap and an inner enveloping cell reinforced by thick actin bundles (Wolfrum, 1990). The earlier light microscopists described these sensilla as “stiftführende Sinnesorgane” (Eggers, 1928). The only differentiations of these cilia are first, the dynein-like arms in the proximal region (Fig. 9b) and second, a swelling (“dilation”) in the distal region, which contains an electron-dense structure. The cilium either ends within the cap, or runs through it, ending in a slight bulbous swelling. The membrane is connected to the microtubules on its inner face, and to the cuticular cap on its outer face (Fig. 9c). Some, if not most, investigators detected a marked bend in the proximal region of the chemically fixed cilium (cf. Schmidt, 1969; Moran et al., 1977; Toh and Yokohari, 1985). The dynein-like arms in the proximal region of the chordotonal cila have been observed in the EM long ago (cf. Schmidt, 1969;

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Fig. 7. Olfactory sensory cilia. (a) Cross section of a thick and a thin olfactory cilium in a s. trichodeum of the silkmoth Antheraea. Note the electron dense structures (arrowheads) squeezed between peripheral microtubules and cell membrane. From Keil (1993), Fig. 4. Bar ¼ 0.2 mm. (b) Longitudinal section of a cryosubstituted s. basiconicum of Bombyx mori, showing numerous ciliary branches with swellings (arrowheads). Bar ¼ 0.5 mm. From Steinbrecht (1980), by permission of Elsevier.

Corbière-Tichané, 1971b), but an interpretation could not be given. It is remarkable that these structures have been found only in these, but never in other insect sensory cilia. During the last few years, interest has focussed on the chordotonal cilium of Drosophila, and it is somewhat surprising how many different proteins are localized very precisely in different regions of this organelle (Fig. 9a). In the transition (necklace) zone at the base, DILA is found (Ma and Jarman, 2011). In the next region, dynein arms and nanchung/inactive (¼TRPV-channel subunits) are localized (Gong et al., 2004; Cheng et al., 2010; Lee et al., 2010). The ciliary dilation contains REMP A and DCX-EMAP (Lee et al., 2008; Bechstedt et al., 2010). And finally in the ciliary tip within the cuticular cap the channel protein NompC ¼ TRPN has been detected (Cheng et al., 2010; Liang et al., 2011). It would be interesting to know when during development these proteins

are transported to their destinations, and whether there is a turnover during lifetime. The dendrite sheath, or cap, of insect mechanoreceptors acts as the mechanical connection between the sensory cilium and the stimulus-transmitting extracellular structures. During development, it is secreted by the thecogen cell in external, or by the scolopale cell in internal mechanoreceptors. Chung et al. (2001) showed that the nompA gene is expressed in these cells, and that the protein becomes located in the sheath or cap. In mutants, cilium and cap are formed, but not connected, and the transduction chain is interrupted. The external receptors (rev. by Keil, 1997a) belong to morphologically completely different sensilla types, such as the hair-shaped s. trichodea (to be distinguished from the olfactory s. trichodea!), the filiform, the club-shaped, the dome-shaped campaniform, and

Fig. 8. Carbon-dioxide sensing sensilla. (a) Oblique section through a s. ampullaceum on the antenna of Apis mellifera. In the basal region, the enlarged cilium completely fills the hair lumen, but then forms numerous tightly packed branches. Bar ¼ 1 mm. (b) Cross section of a s. basiconicum from the labial palp of the moth Helicoverpa armigera. As in Apis, the enlarged cilium completely fills the basal lumen of the hair. Bar ¼ 0.5 mm. (c) Cross section of a s. basiconicum from the labial palp of the moth Helicoverpa armigera in the more distal region. The cilium now forms numerous thin lamellae. Bar ¼ 0.5 mm. (d) Cross section of the LB-II 4 s. basiconicum from Drosophila melanogaster, cryosubstitution. Three neurons give off branched cilia, whereas one gives off a lamellated (in this case appearing circular) cilium. It might be speculated that the latter is the CO2-receptor. Micrograph courtesy of Alexander Steinbrecht. Bar ¼ 0.5 mm.

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Fig. 9. Chordotonal cilia. (a) Schematic drawing of a chordotonal cilium. The different colors indicate the different functional regions and the localization of proteins. orange: inner dendritic segment. yellow: transition zone; DILA. blue: proximal ciliary region; dynein, nanchung/inactive. green: ciliary dilation; REMP A, DCX-EMAP. purple: distal ciliary region; NompC ¼ TRPN. black: cuticular cap; NompA. dark yellow: scolopale cell; actin. brown: attachment cell. (b) Cross section of the proximal region (indicated by thick arrowhead in a) of a chordotonal cilium on the maxillary palp of the Helicoverpa armigera caterpillar, clearly showing the dynein arms (arrowhead) and the dense lumen of the A-tubulus. Bar ¼ 0.1 mm. (c) Cross section of a chordotonal cilium on the maxillary palp of the Helicoverpa caterpillar through the cuticular cap (thick arrowhead in a). Note the regular arrangement of the microtubule doublets, as well as connectors to the cell membrane. (b) and (c) are from the same cryosubstituted specimen and shown at the same magnification. Note that the microtubule doublets are much closer together in level (c), resulting in a thinner axoneme. Bar ¼ 0.1 mm.

numerous different stout bristles, and also the gustatory sensilla. Their sensory cilia are characterized by a highly elaborated cytoskeleton in the distal tip, consisting of a short, but compact assembly of microtubules mostly embedded into an electron-dense matrix and approximately 1 mm in length, the “tubular body” (Fig. 10; “Sinnesstift” of the earlier light microscopists). The number of microtubules might be as high as 1000 (Nicklaus et al., 1967). An overview of the microtubule numbers in different tubular bodies is given by Gnatzy and Tautz (1980). The peripheral tubules are connected to the membrane via “membrane-integrated cones”, or MICs (Fig. 9b, insert; Thurm et al., 1983), and these are connected to the dendrite sheath via fine filaments. Could these filaments be identical with the nompA protein? The tubular body is typical for the primary land-dwelling insects, spiders (cf. Christian, 1971; Foelix and Chu-Wang, 1973), and myriapoda (Keil, 1976; Tichy, 1987; Nguyen Duy-Jacquemin, 1997); a similar structure has been described also in the tardigrada (Walz, 1978). It has been found only in a few crustaceans, namely, the land-dwelling isopod Hemilepistus reaumuri (Gaffal et al., 1975). Otherwise, the sensory cilia in crustacean mechanoreceptors are organized according to the chordotonal type (Ball and Cowan, 1977; Crouau, 1997). The latter author discusses the possibility that the development of a tubular body is merely a question of the environment, that means, the physical properties of the surrounding medium (water vs. air).

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Tubular bodies are most variable with respect to their shape. Their cross section is more or less circular in mechanoreceptors which answer to undirected touch. Some mechanoreceptors, such as the extremely sensitive filiform sensilla on the cricket cercus, have a very precise directional sensitivity, and here the tubular bodies show a polarized structure (Fig. 10b). The biomechanical construction of the hair base is highly sophisticated in these sensilla, and Thurm et al. (1983) calculated that a movement in the range of 0.1 nm would be sufficient to yield an electrophysiological response. Erler (1983a,b) treated identified mechanoreceptive sensilla of Acheta with vinblastine, then recorded their electrical responses to stimuli after increasing exposure time, and finally investigated these sensilla in the electron microscope. He found that the sensitivity decreased with increasing destruction of the tubular body. Thus, the necessity of an intact microtubular cytoskeleton for proper mechanoreceptor function could be shown. The question for the “electron-dense substance” remained open. A few unpublished observations with the LAMMA (Laser Microprobe Mass Analysis; W. Gnatzy, Frankfurt) and EDX-microprobes (Energy Dispersive X-ray Analysis; V. Krefting, T. Keil, Münster) showed an increased amount of Caþþ in the tubular body, but there have been no further investigations. The nature of the electron-dense substance has been approached only recently by Bechstedt et al. (2010): It could be shown to be missing in both types of mechanosensory cilia in Drosophila-DCX-EMAP-mutants. In these animals, the tubular body in campaniform sensilla is not formed properly and is completely disorganized, whereas the ciliary dilation in their chordotonal sensilla e including the curved microtubules e appears otherwise normal. The flies are uncoordinated and deaf, and no electrical response could be recorded from the antennal nerve upon stimulation of Johnston’s organ. Bechstedt et al. (2010) interpreted the dilation as the transduction zone. However, the NompC-protein (¼TRPN-channel protein) is localized only in the tip of the chordotonal cilium where it is attached to the cuticular cap, and in the distal tubular body region in bristle and campaniform sensilla (Lee et al., 2010; Liang et al., 2011), where also the DCX-EMAP protein is found (Bechstedt et al., 2010). Maybe a combination of both is needed for mechanotransduction. Anyway, it is tempting to speculate that the nompC protein forms the intracellular, and the nompA protein the extracellular link of the transduction chain. The proteins found in mechanoreceptors of Drosophila are summarized in Table 1. A most fascinating variation of the mechanoreceptor theme is the infrared receptor which has been discovered 15 years ago in buprestid beetles, but later also in some other insects (Schmitz et al., 1997, 2007). This sensillum is characterized by an extremely elaborated cuticular cupola, into which the tip of a typical mechanoreceptor cilium with a tubular body is embedded. The cuticle is composed in a way which causes it to swell if hit by infrared radiation, and the whole sensillum is nothing but an extremely modified mechanoreceptive hair, as shown by numerous intermediate types. 8. Ciliary dynamics: transport and movement Intraflagellar transport (IFT) has been intensively studied during the early 1990s (e. g., Kozminski et al., 1993, 1995; rev. Insinna and Besharse, 2008). It could be shown to play a decisive role for proper formation of the functioning sensory cilium first, in mammalian photoreceptors and then also in Drosophila mechanoreceptors. The formation of sensory cilia in Johnston’s organ is seriously disturbed in the DmKap (Kinesin associated protein)-, and also the klp64D (kinesin II)-mutant (Sarpal et al., 2003). Consistent with this, Avidor-Reiss et al. (2004) showed that in the oseg2 and the klp64D-

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Fig. 10. Cross sections through tubular bodies in different mechanoreceptor types, showing a richness of functional shapes. The arrowheads indicate the direction of the stimulus. (a) Campaniform sensillum of the cockroach Periplaneta americana, from Moran et al. (1971), Fig. 10. Ó 1971 Rockefeller University Press, originally published in Journal of Cell Biology 48, 155e173. Bar ¼ 1 mm. (b) Filiform sensillum of Acheta. Insert shows the connectors (MICs) between the peripheral microtubule layer and the cell membrane. From Keil and Steinbrecht (1984), Fig. 12 a,b. With kind permission from Springer Science þ Business Media B.V. Bar ¼ 0.5 mm, bar in insert ¼ 0.1 mm. (c) Macrochaeta of the fruitfly, Drosophila melanogaster. Bar ¼ 0.5 mm. (d) Macrochaeta of the blowfly, Calliphora vicina. This section is adjacent to the one shown in Keil (1978), Fig. 5. Bar ¼ 0.5 mm. (e) Campaniform sensillum from a cryosubstituted haltere of Calliphora. Micrograph by Wolfgang Voelker, From Keil (1998). Bar ¼ 0.5 mm.

mutants of Drosophila the transport of tubulin, or of microtubule subunits, into the cilium of mechanoreceptor bristles was disturbed. Consequently, the tubular body was not formed, and microtubule fragments assembled under the base of the cilium. A similar effect had been found in mouse intraflagellar transport mutants (Pazour et al., 2002), in which the transport of membrane material into the photoreceptor cilium was disturbed and membrane material assembled under the base of the cilium. The high number of genes involved in Drosophila ciliogenesis has been reviewed by Zur Lage et al. (2011), they emphasized 7 genes involved in transport from the Golgi apparatus to the cilium, 11

genes involved in anterograde and 7 genes involved in retrograde ciliary transport. An important element of IFT is the structure originally called “raft”, later “train”, a slender electron-dense structure squeezed between peripheral microtubules and the ciliary membrane (Kozminski et al., 1993, 1995; for a detailed ultrastructural 3Dinvestigation see Pigino et al., 2009). Such structures had also been found in the olfactory cilia of the silkmoth Antheraea (Fig. 7a; Keil, 1982, 1993), but could then not be interpreted. There are indications for turnover processes in insect olfactory cilia. Vesicles have been found in the distal region of sensory cilia (Marshall, 1973);

T.A. Keil / Arthropod Structure & Development 41 (2012) 515e534 Table 1 Proteins that have been localized in mechanosensory cilia of Drosophila. Protein TilB DILA/UNC

Location

Dendrite Transition zone Dynein Cilium base Nanchung/ Cilium inactive ¼ TRPV base DmKAP/KLP 64D Cilium Oseg2 Cilium REMP A DCX-EMAP NOMPC ¼ TRPN NOMPA

Function

References

Dynein assembly Transport regulator

(Kavlie et al., 2010) (Ma and Jarman, 2011) (Kavlie et al., 2010 and others) (Gong et al., 2004)

Motor Ion channel Transport Tubulin transport

(Sarpal et al., 2003) (Avidor-Reiss et al., 2004) Dilation Transport (Lee et al., 2008) Dilation Electron-dense (Bechstedt et al., substance, transduction 2010) Cilium tip Microtubule-membrane- (Cheng et al., 2010; connector, ion channel Liang et al., 2011) Cap Membrane(Chung et al., 2011) cap-connector?

multivesicular bodies have been shown to squeeze through the ciliary constriction in sensilla of the housefly larva (Chu and Axtell, 1971), and small vesicles have been observed frequently in the center of the ciliary base (Fig. 3a,b). In the rare cases of longitudinal sections through the tips of trichoid sensilla in the hawkmoth Manduca sexta, the distal ends of the cilia clearly disintegrate into vesicles (Keil, unpublished observations: Fig. 11b). Permanent growth of sensory cilia e especially the chemosensory ones e would need a permanent supply of membrane material, comparable to the vertebrate photoreceptor (Pazour et al., 2002). In fact, vesicles have been found in the tip of the inner dendritic segment, sometimes in very high numbers (Fig. 11a; Keil and Steinbrecht, 1987; Brandt, 1988; Keil, 1989). No decision could be made whether these have an exo- or endo-cytotic function: at least, they never show a coat (discussed in detail by Keil, 1989). “Polarized exocytosis” to the base of the mammalian photoreceptor cilium, which also shows an intense membrane turnover, has recently been discussed by Nachury et al. (2010), and the transport of membrane-bound proteins from the Golgi apparatus to the cilium by Garcia-Gonzalo and Reiter (2012). It is suggested here that a similar process is active in the insect olfactory cilia. In chemically fixed, but also cryosubstituted specimens, numerous swellings, or beads, have been found in the cilia in the EM (Keil, 1984a, 1989, 1993; Seidl, 1991; Steinbrecht, 1980; see Fig. 7b), and in the light microscope as well (Williams, 1988; Kumar and Keil, 1998b). Beading in mammalian axons had been described earlier (cf. Ochs et al., 1996). Time-lapse observations on apically opened olfactory hairs showed first, that the beads can move along the cilia in vitro and second, that the latter are able to make active movements, which means they are able to elongate (Williams, 1988; Keil, 1989) and shorten again repeatedly. Motor proteins could be demonstrated immunohistochemically by Kumar and Keil (1998a). Another indication for the ability of the cilia to make active movements comes from the chordotonal organs. Here, it had been found by several investigators that after chemical fixation, a marked bend occurs at the base of the cilium (e. g., Gray and Pumphrey, 1958; Schmidt, 1969; Moran et al., 1977; rev. by Field and Matheson, 1998). Moran et al. (1977) were the first to suggest an active role of these cilia in mechanotransduction. The best argument for active motility was presented by Göpfert and Robert (2003). They could show that the funiculus of the Drosophila antenna is vibrated by movements of the sensory neurons of Johnston’s organ in response to auditory stimulation. Spontaneous oscillations (SO) of the antennal flagellum driven by the Johnston’s organ have been described in Culex (Warren et al., 2010). The

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question is of course for the mechanism of these movements. The latter authors have suggested a microtubule-dependent mechanism, because the SOs could be suppressed by application of colchicine. The dynein-like side arms in the region proximal of the ciliary dilation might play the decisive role in the generation of this movement (Kavlie et al., 2010). These authors found that in the deaf tilB (touch insensitive larva B) mutant of Drosophila, these arms are missing. Dynein arms are assembled near the basal body in Chlamydomonas and then transported into the cilium (Hou et al., 2007), the TilB protein is thought to be involved. It is very likely that the side-arms are really identical with dynein, and are responsible for the active movements of the chordotonal cilia. This means that structures typical for motile (9 þ 2)-cilia are found at least in one type of primary cilium. On the other hand, it could be imagined that the very long ciliary rootlets in the chordotonal neurons are contractile. This has been described in Chlamydomonas by Salisbury and Floyd (1978), while Sleigh (1979) pointed out the possible function of the rootlets in vertebrate rod cells. A thorough discussion was given by Wolfrum (1991, 1997). The amplification process found in the Johnston’s organ shows parallels to the vertebrate ear, where the outer hair cells act as amplifiers (rev. by Ashmore, 2008). Contractions of these cells are driven by the protein prestin, which is localized in the basolateral membrane. It is an anion transporter acting as a “motor” by conformational changes. It has also been localized in the Johnston’s organ of mosquitoes and flies (Weber et al., 2003). This has been taken as a further argument for a common phylogenetic origin of vertebrate and insect hearing organs, a view which, however, is severely doubted by Hartenstein (2005). 9. Receptors and channels 9.1. Mechanoreception Mechanosensors answer to stimuli with only a very short latency (few milliseconds), thus it could be expected that there must be a very direct action on the receptor. Rice et al. (1973) investigated the mechanoreceptive setae on the mouthparts of the Tsetse fly and came to the conclusion that hole-shaped ion channels were opened directly by stretching of the receptor membrane. Thurm pointed out that very often connectors between the microtubular cytoskeleton and the membrane had been observed (first in the spider Tegenaria by Christian, 1971) and suggested that these structures themselves be the receptors (“membrane integrated cones”, or MICs: Thurm et al., 1983). Mutants unresponsive to mechanical stimuli (Mec-.) of Caenorhabditis elegans (Hamill and McBride, 1994, 1996) allowed a closer analysis of the molecular mechanisms. Comparison with the transduction process in mammalian stereocilia (Gillespie and Walker, 2001) showed great similarities. There were thorough investigations of Drosophila Nomp (No Mechanoreceptor Potential) mutants, and it became clear that TRP (Transient Receptor Potential)-channels are involved in insect mechanotransduction (cf. Árnadòttir and Chalfie, 2010). A molecular model was presented by Howard and Bechstedt (2004) which takes into account the earlier electron microscopical observations (e. g., Voelker, 1982; Thurm et al., 1983, Fig. 12a). In their model, the ion channel in the membrane is connected to the microtubule cytoskeleton via 29 ankyrin units arranged as a 20 nm long complex of 4 chains, which most probably corresponds with a MIC and acts as the “gating spring”. The NompC (¼TRPN) protein is localized in the tubular body region in sensory bristles (Lee et al., 2010), campaniform sensilla, and the distal region of the cilia in chordotonal organs (Cheng et al., 2010; Liang et al., 2011) of Drosophila. The presence of the NompC protein in the distal tip, but not the ciliary dilation, of

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Fig. 11. Turnover processes in olfactory s. trichodea of Manduca sexta. (a) Cross section of two inner dendritic segments in the region of the centriole pairs (bb). Note numerous smooth vesicles. Bar ¼ 0.5 mm. (b) Disintegration of the olfactory cilia in the sensillum tip. Proximally, the cilium looks intact (arrowhead). Bar ¼ 0.5 mm.

the chordotonal cilium e just where it is inserted in the cuticular cap e opens the question for the proper site of mechanotransduction. Additionally, Chung et al. (2001) detected NompA in the cap surrounding the ciliary tip (cf. x7). Cheng et al. (2010) could also show that the ankyrin repeats which had already been found

by Howard and Bechstedt (2004) are necessary for connection to the microtubules. At least, cross sections through this region show connectors between microtubules, membrane, and cap (Fig. 9c). It must be pointed out that in the proximal region of the chordotonal cilium (that is, beneath the dilation), the Nanchung/

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9.2. Chemoreception

Fig. 12. Cartoon of some sensory transduction mechanisms. This figure does not intend to represent molecular details, but only to give a rough impression. (a) Mechanotransduction, modified after Gillespie and Walker (2001) and Howard and Bechstedt (2004). The ion channel (green) is connected to the cytoskeleton via the ankyrin chains (“MIC”, orange) and to the cuticle via an as yet unidentified connector (purple). Note that the NompA protein is located somewhere on the inside of the cuticular cap, and NompC in the ciliary tip. Any mechanical deformation of the cuticle opens the channel for cations (red). CUT, cuticle; MT, microtubule. (b) Olfactory transduction, according to Sato et al. (2008): Binding of the odor molecule (blue) to the receptor (yellow) results in direct opening of the channel (green). (c) Olfactory transduction according to Wicher et al. (2008): Binding of the odor molecule (blue) to the G-protein coupled receptor (yellow) triggers a second-messenger-cascade via adenylate cyclase (AC), in which cAMP opens the channel. (d) In the olfactory CO2-receptor, the stimulating molecule acts directly on the channel (green). Modified after Scott (2011).

inactive complex could be detected (Lee et al., 2010; Cheng et al., 2010). So obviously insect mechanosensory cilia can carry different types of receptors/channels in different regions. Additionally, parts of the receptor complex are localized in extracellular structures, exactly as had been found in Caenorhabditis (Hamill and McBride, 1996). It could be shown that TRP-channels play the decisive role in insect hygroreceptors (Liu et al., 2007), and also in thermoreceptors on the antenna of Drosophila (Gallio et al., 2011) which have lamellated sensory cilia with BOSSes (Foelix et al., 1989); in any case, TRP-channels are responsive for thermoreception in mammals (Patapoutian et al., 2003). A short review of the structure and function of TRP-channels has been given by Montell (2011).

Olfactory receptors are characterized by long latency times (Kaissling and Priesner, 1970). It could therefore be imagined that a second-messenger-system, similar to that found in vertebrate olfactory neurons (cf. Lancet, 1986; Buck and Axel, 1991; Anholt, 1993), also plays a role in insect olfaction (Breer et al., 1990). However, despite all efforts by different research groups, the issue remained controversial. In homogenates of whole antennae, an increased activity of second messengers was found upon stimulation with odorants (Breer et al., 1990; Ziegelberger et al., 1990). But in contrast to this, the latter authors showed in the same paper that in isolated trichoid olfactory hairs of A. polyphemus, which contain no other cellular structures than the olfactory cilia (cf. Klein and Keil, 1984), do not have an increased level of cGMP after stimulation. Patch-clamp recordings from extruded olfactory cilia of A. polyphemus showed first, that the AC1-ion channel is activated by cGMP and protein kinase C-activators in inside-out patches, and second, that the density of these channels in the membrane is surprisingly low, ranging between 0.33 and 1.6 per square micrometer (Zufall and Hatt, 1991). By the end of the century, there was some consensus about the molecular mechanism of insect olfaction (rev. by Stengl et al., 1999): first, the odor molecule docks to a receptor in the membrane, then the receptor activates a G-protein, which in turn activates phospholipase C (PLC), which then generates diacylglycerole (DAG) and inositol-triphosphate (IP3). Both of these then open different Caþþ and Kþ channels. Compared with mammals, the number of olfactory receptor (OR) genes in insects is quite low: ca. 1200 in the mouse vs. 62 in the fruitfly, only one (rarely two) of these being expressed in each olfactory sensory neuron (ORN) (rev. by Benton, 2006). However, the number of insect olfactory genes is quite variable and obviously depends on the ecological importance of the olfactory sense for different species (rev. by Hansson and Stensmyr, 2011): there are 10 in the body louse Pediculus and at least 297 in the fireant Solenopsis (Wurm et al., 2011). These OR proteins share no sequence similarity to the G-protein coupled receptors (GPCRs) such as found in mammals and show a membrane topology very distinct from the latter (Benton, 2006). A special case is Or 83b which is expressed in probably all ORNs, where it forms a complex with each specific OR and is necessary for the transport into the sensory cilium (Benton, 2006). The controversy was revived by two papers appearing simultaneously in 2008 in Nature. However, both papers do not report direct measurements on sensory neurons, but the respective receptor proteins were expressed in mammalian cell lines or Xenopus oocytes. Sato et al. (2008) come to the conclusion that the complex of OR and Or 83b proteins forms a ligand-gated ion channel (Fig. 12b). Wicher et al. (2008) report that on the one hand, cells co-expressing the Or 22a and Or 83b proteins form ligandgated channels, whereas cells expressing only Or 83b form ion channels needing cAMP or cGMP (Fig. 12c). This issue was discussed in detail by Pellegrino and Nakagawa (2009) who suggest that the G-protein independent response is the faster one, followed with some delay by a longer-lasting G-protein coupled response (see also discussion by Kaupp, 2011). Gustatory receptors still pose more problems regarding their molecular nature, as pointed out in the recent review by Montell (2009). There are 68 known gustatory receptor (GR) proteins in Drosophila, which sense “sweet” and “bitter” tastants, as well as nonvolatile pheromones on the cuticular surface. In Drosophila, two GRs (GR21a, GR63a) are sensitive to CO2, but surprisingly in olfactory neurons on the antenna (see x6; Jones et al., 2007; Scott, 2011, Fig. 12d). Their way of action is still not well known: in mammals, the enzyme carbonic anhydrase is involved in CO2-sensing, but not in

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Drosophila (rev. by Luo et al., 2009). The sensitivity for water depends on the osmosensitive ion channel PPK28 (Cameron et al., 2010). 10. Sensory cilia in other arthropods In myriapoda, it has repeatedly been found that some sensory neurons form two cilia (e. g., Bedini and Mirolli, 1967; Haupt, 1971; Keil, 1975; A. Ernst, 1981, 1983: Chilopoda; Schönrock, 1981; Nguyen Duy-Jacquemin, 1997: Diplopoda). In these cases, the centrioles have changed their position: The tandem arrangement is disrupted, and both centrioles are lying side by side, each producing its own cilium. The same feature has been described in aesthetasc chemoreceptors on the antennules of crustacea, for example Pagurus hirsutiusculus (Ghiradella et al., 1968), Neomysis integer, Idotea baltica (Guse, 1983), and Asellus aquaticus (Heimann, 1984). The same author found multiciliated dendrites (up to 25 cilia, each of which arises from a single basal body) in the ostracod Conchoecia spinirostris (Heimann, 1979). Two cilia per neuron have been described in some Collembola (Karuhize, 1971; Altner and Thies, 1976, 1978). In pterygote insects, however, this number is an exception (Steinbrecht, 1988). An overview on the occurrence of double cilia is given by Altner and Thies (1976). MICs connecting the peripheral microtubules in sensory cilia have been found in trichobothria of the spider Tegenaria derhami (Christian, 1971), in the tardigrade Macrobiotus hufelandi (Walz, 1978), and the millipede P. lagurus (Nguyen Duy-Jacquemin, 1988). In the latter, tightly lamellated cilia looking identical to those known from thermo-hygroreceptors of insects (including the BOSSes, cf. Steinbrecht, 1998) have also been described by Nguyen Duy-Jacquemin (1983). In the funnel-canal organs of the shore crab Carcinus maenas which are presumably bimodal mechano- and chemo-receptors (Schmidt and Gnatzy, 1984), one type of sensory cilia shows the typical basal bend as well as the dynein-like side-arms, whereas the other is staight, lacks the side-arms, but contains numerous electron-dense vesicles in the proximal region. Hayes (1971) has investigated the chemoreceptors (unfortunately not the mechanoreceptors as well) in a living fossil, the horseshoe crab Limulus polyphemus. These most probably are the phylogenetically oldest sensilla of euarthropods. Their sensory cilia do not differ much from those of insects. What is most interesting about them are the dynein-like arms in their proximal region. In insects, these have only been found in the chordotonal cilia. Anyhow, it is remarkable that the (most probable) dynein arms are present on these primary, or “immotile”, cilia. We may speculate that their presence is phylogenetically the original state, and they are lost in the higher evolved arthropods with the exception of the chordotonal cilia. The tubular body of the external mechanoreceptors in land-dwelling arthropods most probably is a secondary adaptation (Crouau, 1997). 11. Concluding remarks Cilia have entered the center stage of cell research, papers on this topic are published at an increasing rate. For research on cilia, the fruitfly Drosophila has become highly popular for several reasons: first, a large number of ciliary mutants is available, and second, it is possible to investigate primary (sensory) and motile (sperm) cilia in the same species. It could hardly be imagined that “the insect sensory cilium” would show such an incredible morphological, physiological, and biochemical diversity when Edwin G. Gray published his famous paper 50 years ago. The beauty of Gray’s reconstruction was only surpassed 9 years later by KlausDieter Ernst’s basiconic sensillum of Necrophorus (Ernst, 1969). Approximately until the turn of the century, the spectrum of insect

species investigated by electron microscopical and electrophysiological methods was very broad, ranging from “Apterygota” over locusts, crickets, cockroaches to beetles, moths, and flies. Already during the late 80s, the focus shifted more and more toward the fruitfly D. melanogaster. In this species, molecular genetics, often in combination with the two previous methods, has yielded new insight into functional as well as developmental problems which could not be dreamed of in 1980. But it must be pointed out that Drosophila hardly is “the typical insect” e merely a useful model. In any case, the fundamental work done from the 60s to the 90s in numerous other insect species formed the basis for today’s findings in Drosophila e this fact must never be forgotten, as often seems to be the case in recent publications! Independent of their sensory modality, all insect sensilla are built more or less according to the same scheme and follow the same developmental pattern (cf. Keil and Steinbrecht, 1984; Keil, 1997b; Hartenstein, 2005). Their sensory neurons can hardly be distinguished morphologically, differences are confined to their cilia. A large number of different sensory transduction processes has been investigated so far. The issue of mechanoreception seems to be quite clear by now, and there obviously are many parallels to the transduction processes in mammals and nematodes. Despite many efforts by different research groups, the issue of olfaction is not as clear as it seemed to be 10 years ago: Is it a secondmessenger-dependent or a ligand-gated mechanism or a combination? Infrared reception seems to be only a modification of mechanoreception. The receptor for water is an osmosensor in taste sensilla. Receptors for temperature and humidity seem to belong to the TRP family. So there remain plenty of open questions. An issue to be addressed further is the intraflagellar transport and possible turnover in insect sensory cilia. Different proteins have been localized in very distinct regions of mechanosensory chordotonal cilia, the question remains how they can be transported so precisely to their respective destinations. Could it be that there is a turnover in the tips of the cilia, as has been found in Chlamydomonas (Marshall and Rosenbaum, 2001)? Can a turnover of olfactory cilia, which is indicated only by a few unpublished EMobservations, be verified by in-situ-observations in fluorescent living cilia? Could in vivo-fluorescence observations help to address the problem of vesicle traffic to/from the distal region of the inner dendritic segment? Could the up and down of the centrioles during morphogenesis be clarified? Furthermore, the assembly of the tubular body, together with the localization of Caþþ, would be worthwhile for detailed analysis. And finally, the by now well established method of electron tomography of freeze-substituted specimens (e.g., Hoenger and McIntosh, 2009) could be applied to membrane-associated structures such as the MICs or the BOSSes. Acknowledgments This review is partly based on work done at the Dept. of Neurophysiology, University of Münster, and the MPI for Behavioral Physiology at Seewiesen. Ulrich Thurm stimulated my interest in cilia, Maaike Thomas, Heidi Hoesch, Cornelia Bock lent me their hands during this time. Thanks to Werner Gnatzy for allowing me to cite his unpublished work, Wolfgang Voelker for his micrograph, and to Alexander Steinbrecht for helpful comments and micrographs. Dedicated to the memory of Jacobus Jan (“Koos”) de Kramer, who died on September 24, 2011. References Afzelius, B.A., 2004. Cilia-related diseases. Journal of Pathology 204, 470e477. Afzelius, B.A., Eliasson, R., Johnsen, O., Lindholmer, C., 1975. Lack of dynein arms in immotile human spermatozoa. Journal of Cell Biology 66, 225e232.

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