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Table of Contents
Abstract
The non-motile cilium, once believed to be a vestigial cellular structure, is now increasingly associated with the ability of a wide variety of cells and organisms to sense their chemical and physical environments. With its limited number of sensory cilia and diverse behavioral repertoire, C. elegans has emerged as a powerful experimental system for studying how cilia are formed, function, and ultimately modulate complex behaviors. Here, we discuss the biogenesis, distribution, structures, composition and general functions of C. elegans cilia. We also briefly highlight how C. elegans is being used to provide molecular insights into various human ciliopathies, including Polycystic Kidney Disease and Bardet-Biedl Syndrome.
Cilia are slender microtubule-based subcellular organelles that emanate from the cell surfaces of virtually all eukaryotic organisms. Two types of cilia exist: motile cilia (alternatively termed flagella), which are used for locomotion or for the generation of fluid flow, and non-motile (primary) cilia, which are implicated in sensing the chemical and/or physical extracellular environments. Eukaryotic cilia are evolutionarily distinct from the similarly-shaped microvilli or stereocilia that are built from an actin cytoskeleton, and from the bacterial flagellum that drives motility in some prokaryotes.
In a letter to Max Perutz dated June 5, 1963, Sydney Brenner wrote that a key unresolved question in biology was how the nervous system developed, and proposed that another important area of research could be how multicellular organisms controlled flagellation and ciliation (http://elegans.swmed.edu/Sydney.html). Fittingly, many of the early studies on C. elegans were centered on its chemotactic behaviors, which we now understand depend on the functions of cilia present in sensory neurons, and some of the earliest mutants to be isolated were defective in their abilities to sense environmental conditions (Ward, 1973; Dusenbery, 1974; Dusenbery et al., 1975). Although at that time the link between chemosensation and cilia was not firmly established, electron microscopic reconstruction of the environmentally-exposed, cilia-based sensory system at the anterior end of the animal helped to make the link more evident (Ward et al., 1975; Ware et al., 1975).
Unlike many organisms, including humans, the only ciliated cell type in C. elegans is the sensory neuron, and none of the cilia in the nematode are motile. Of the 302 neurons found in the adult hermaphrodite, a substantial number (60) possess cilia at the ends of their dendritic processes.
Cilia from all studied organisms are known to nucleate from a modified centriolar structure termed a ‘basal body’. Most often, the basal body is positioned in proximity to the cellular membrane from where the cilium emanates. C. elegans basal bodies have been described as more ‘degenerate’ and termed ‘transition zones’ by Perkins et al. (1986). Ultrastructurally, the C. elegans transition zone (also termed ‘proximal segment’) typically possesses a circular array of doublet microtubules (as opposed to the triplet microtubule arrangement most often associated with basal bodies in other organisms). In amphid and phasmid cilia, the transition zone is followed by a so-called ‘middle segment’ characterized by a canonical arrangement of 9 doublet microtubules, and this middle segment transforms into a ‘distal segment’ built of singlet microtubules (Figure 1); notably, in some ciliated neurons, these ultrastructural features may be somewhat divergent (Ward et al., 1975; Ware et al., 1975; Perkins et al., 1986). Additional singlet microtubules are often present in the central region of the C. elegans cilia (Figure 1), but these microtubules are likely distinct from the central pairs observed in motile cilia. On the whole, this organization of doublets transitioning to singlets at the distal end is very similar to that seen in the flagella of mating Chlamydomonas cells (Mesland et al., 1980) and may be a general property of sensory cilia, as it has also been observed in several vertebrate cell types (e.g., pancreatic, renal and olfactory cells; Reese 1965; Webber and Lee, 1975; Hidaka et al., 1995). The nature and positions of the C. elegans ciliated cell bodies and of representative dendritic ciliated endings are shown schematically in Figure 1.
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Figure 1. Ultrastructures of cilia and relative positions of all known ciliated neurons (cell bodies and associated dendrites) in the C. elegans hermaphrodite. The top two panels show electron micrograph cross-sections of amphid cilia in the middle segment (microtubule doublets; left panel) and distal segment (microtubule singlets; right panel; adapted from Evans et al., 2006). The worm figures illustrate the positions of the all ciliated cell bodies and their dendritic extensions. The four insets show, schematically, electron micrograph reconstructions of known ciliated endings (adapted from Perkins et al. (1986) for amphids and phasmids and Ward et al. (1975) for labial and cephalic neurons). Cu, cuticle; CR, ciliary rootlet; SCu, subcuticle; So, socket cell; Sh, sheath cell.
The primary chemosensory organ of C. elegans is built from a collection of amphid neurons whose cell bodies are located in the anterior region of the pharyngeal bulb and possess axons that associate with the nerve ring. The dendrites of these neurons extend to the anterior end of the animal and terminate with diverse ciliated structures (Figure 1). The proximal regions of amphid cilia are typically protected by a sheath cell and extend through a channel created by socket cells to become partially exposed to the external environment. The majority of amphid neurons possess cilia shaped as single rods (ASE, ASG, ASH, ASI, ASJ, ASK) or pairs of rods (ADF, ADL). Other amphids boast cilia that have membrane elaborations and possess unusual shapes; these are the wing neurons (AWA, AWB, AWC) and the amphid finger neuron (AFD), in which a small cilium is surrounded by approximately 50 villi. Both the wing and AFD neuron cilia terminate within a sheath cell, and thus are not exposed to the external environment. The lengths of amphid cilia range from ∼7.5 μm (in the ASE, ASG, ASH, ASI, ASJ and ASK neurons) to ∼1.5 μm for the AFD cilium (Ward et al., 1975; Ware et al., 1975; Perkins et al., 1986). Similar in structure to the single rod-like cilia found in amphids are the PHA and PHB phasmid cilia. These are located slightly posterior to the anus of the worm and are exposed to the external environment (Hall and Russell, 1991).
The inner labial neuron types (IL1, IL2) are both arranged symmetrically in sets of 6 cells, ultimately terminating in the 6 “lips” that surround the mouth of the worm. Originating from a position anterior to the amphids, the dendrites of these neurons terminate in shorter cilia, and possess a seemingly more degenerate basal body. While the IL1 cilia consistently originate from basal bodies consisting of 7 doublet microtubules, those of IL2 neurons are more variable (ranging from 5–7 doublets). These neurons are further distinguished by the fact that, while the IL1 cilia ultimately terminate, or embed, in the subcuticle, the IL2 cilia are exposed to the external environment via openings in the cuticle (Ward et al., 1975; Ware et al., 1975).
The outer labial (2 lateral outer labial, or OLL neurons, and 4 quadrant outer labial, or OLQ neurons) and cephalic (CEP; 4 neurons) neurons similarly terminate, albeit in a more restricted fashion, in the cuticle near the sub-dorsal, sub-ventral, and lateral lips of C. elegans. The cilia found at the dendritic termini of CEP neurons possess a degenerate transition zone (6–8 doublet microtubules), while those found in the OLL/OLQ neurons have a canonical, 9 microtubule doublet arrangement. The cilia of CEP neurons are unusual in that, ∼1μm from the basal body (within the subcuticle), the axonemal microtubules associate with additional microtubules, generating an electron-dense structure difficult to reconstruct via EM (Ward et al., 1975; Ware et al., 1975).
Interestingly, the IL1, OLL and OLQ neurons are unique in the fact that they have striated rootlet structures descending from their transition zones (WormAtlas; Ward et al., 1975; Ware et al., 1975). Ciliary rootlets are prominent fibrous polymers of the protein rootletin that emanate from the proximal end of the basal body (Yang et al., 2002). Rootlets have been implicated in the maintenance and longevity of vertebrate sensory cilia (Yang et al., 2005), as well as in providing scaffolding for kinesin-1-based intracellular transport (Yang and Li, 2005). It should be noted that very little is known about the rootlets of C. elegans; even a rootletin homolog has yet to be clearly identified.
Two unusual ciliated cell types, AQR (located near the pharynx) and PQR (found posterior to the phasmids in the tail), are found, along with their cilia, to be directly exposed to the pseudocoelomic cavity of the worm. Extremely little is known about the ultrastructure of the cilia of these neurons, although they can be identified under a compound microscope using, for example, the GCY-36 protein fused to GFP (Cheung et al. 2004).
The 4 lateral, cervical deirid neurons are found in pairs, at the posterior end of the pharyngeal bulb (ADE) and slightly anterior to the anus (PDE). Like many of the other neurons discussed in this review, their ciliated dendritic endings are in a channel formed by a socket cell and an invaginated sheath cell. The cilia of both ADE and PDE terminate in the subcuticle, and thus are not exposed to the external environment. These ADE/PDE cilia are remarkably similar to those found in the 4 CEP neurons, and, interestingly, these 8 neurons constitute the complete dopaminergic neuron set for the hermaphrodite worm (Sulston and Brenner, 1975; Ward et al., 1975; Ware et al., 1975).
BAG and FLP are two relatively uncharacterized ciliated neurons whose cilia both terminate in or near the lateral lips of the worm. Unlike many of the other neurons described in this review, their cilia are not surrounded by support cells. Furthermore, their ultrastructures are quite complex, appearing via EM reconstruction as “bags” (BAG) or “flaps” (FLP) (Ward et al., 1975; Ware et al., 1975; Perkins et al., 1986).
C. elegans males have 52 additional ciliated sensory neurons, the majority of which are found in the male tail rays/hooks, where the cilia perform sensory functions (Peden and Barr 2005). It should be noted, however, that only 48 of these 52 neurons are confirmed by EM to have cilia (Sulston et al., 1980). General descriptions of the structure and function of male-specific cilia are described in Table 1. While in many organisms spermatozoa possess motile cilia (flagella), those of C. elegans are aflagellar, relying on amoeboid locomotion to reach and fertilize oocytes (Nelson et al., 1982).
Table 1. Description of individual ciliated neuron types and their reported functions
| Ciliated neuron | Cilium structure | Exposed? | Embedded? | Dye fills? | General role | Reference(s) |
|---|---|---|---|---|---|---|
| ASE (L/R) | Single rod | Y | Chemoattraction | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | ||
| ADF (L/R) | Pair of rods | Y | FITC, DiI | Dauer entry | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| ASG (L/R) | Single rod | Y | Chemoattraction | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | ||
| ASH (L/R) | Single rod | Y | FITC, DiI | Mechanosensory (Nose touch), chemorepulsion, osmo-avoidance | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986; Kaplan and Horvitz, 1993 | |
| ASI (L/R) | Single rod | Y | FITC, DiI | Chemoattraction | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| ASJ (L/R) | Single rod | Y | FITC, DiI | Dauer exit/recovery | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| ASK (L/R) | Single rod | Y | FITC, DiI | Chemoattraction | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| ADL (L/R) | Pair of rods | Y | FITC, DiI | Chemorepulsion | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| AWA (L/R) | Winged | N | Sheath cell | Chemoattraction | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| AWB (L/R) | Winged | N | Sheath cell | Chemorepulsion | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| AWC (L/R) | Winged | N | Sheath cell | Chemoattraction | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| AFD (L/R) | Small, surrounded by dendritic villi | N | Sheath cell | Thermosensation | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | |
| IL1 (DL/DR/ L/R/VL/V) | N | Subcuticle | Mechanosensation (Nose touch) | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986; Hart et al., 1995 | ||
| IL2 (DL/DR/ L/R/VL/V) | Y | DiI, DiO | Unknown (presumably chemosensory) | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | ||
| CEP (DL/DR/ VL/VR) | N | Cuticle | FITC (occasionally) | Mechanosensation (Basal slowing response) | Ward et al. 1975; Ware et al. 1975; | |
| OLQ (DL/DR/ VL/VR) | N | Cuticle | Mechanosensation (Nose touch and basal slowing response) | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986; Kaplan and Horvitz, 1993; | ||
| OLL (L/R) | N | Cuticle | Mechanosensation (putative) | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986 | ||
| BAG (L/R) | N | Behind cuticle | Unknown | Perkins et al. 1986 | ||
| FLP (L/R) | N | Behind cuticle | Mechanosensation (Nose touch) | Perkins et al. 1986; Kaplan and Horvitz, 1993 | ||
| ADE (L/R) | Single rod | N | Subcuticle | FITC (occasionally) | Mechanosensation (Basal slowing response) | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986; Sulston and Brenner, 1975 |
| PDE (L/R) | Single rod | N | Subcuticle | FITC (occasionally) | Mechanosensation (Basal slowing response) | Ward et al. 1975; Ware et al. 1975; Perkins et al. 1986; Sulston and Brenner, 1975 |
| PHA (L/R) | Single rod | Y | FITC, DiI | Chemorepulsion | Hall and Russell, 1991 | |
| PHB (L/R) | Single rod | Y | FITC, DiI | Chemorepulsion | Hall and Russell, 1991 | |
| AQR | Y (pseudocoelom) | Oxygen-sensation, social feeding | Cheung et al., 2005 | |||
| PQR | Y (pseudocoelom) | Oxygen-sensation, social feeding | Hall and Russell, 1991; Cheung et al., 2005 | |||
| Male-specific ciliated neurons | ||||||
| CEM (DL/DR/ VL/VR) | Y | Male chemotaxis (putative) | Sulston et al. 1980 | |||
| RnA (L/R) (n=1–9) | N | structural cell | rarely | Male mating behavior | Sulston et al. 1980 | |
| RnB (L/R) (n=1–9) | Y (not R6B) | rarely | Male mating behavior | Sulston et al. 1980 | ||
| HOA | N | subcuticle | Sensing vulva in male-mating behavior | Sulston et al. 1980 | ||
| HOB | Y | Sensing vulva in male-mating behavior | Sulston et al. 1980 | |||
| PCA (L/R) | N | Sensing vulva, inducing spicule prodding behavior | Sulston et al. 1980 | |||
| SPD (L/R) | Y | Sperm transfer | Sulston et al. 1980 | |||
| SPV (L/R) | Y | Sperm transfer | Sulston et al. 1980 | |||
Ciliogenesis depends on the intraflagellar transport (IFT) of ciliary precursors from the transition zone, which sits at the junction between the dendrite of the sensory neuron and the cilium, to the growing ciliary structure (Figure 2). The many known components of the IFT machinery, some of which were first identified in C. elegans (Scholey et al. 2004 and see below) are listed in Table 2. Using time-lapse microscopy it has been shown that in C. elegans, two IFT motors of the kinesin-2 family, heterotrimeric kinesin-II and homodimeric OSM-3, move IFT-particles (consisting of two multi-protein subcomplexes, A and B; Cole et al., 1998) and presumably ciliary precursor proteins from the base of cilium to their sites of incorporation; this anterograde IFT-machinery, and probably also turnover products, are then transported back to the base of the cilium using the IFT-dynein motor (Movie 1; Figure 2; Orozco et al., 1999; Signor et al., 1999b; Snow et al., 2004). These two anterograde motors cooperate to build the middle and distal segments of cilia. In the middle segment, kinesin-II and OSM-3-kinesin function redundantly to move the same IFT-particles and to assemble the middle segment of the axoneme. In this segment, the slower-moving kinesin-II (0.5 μm s-1) reduces the speed of the faster-moving OSM-3 (∼1.3 μm s-1) to give rise to the intermediate rate of motor-IFT-particle transport observed (∼0.7 μm s-1). Subsequently, at the middle-distal segment boundary, kinesin-II returns to the base of the cilium, liberating OSM-3, which now moves IFT-particles and bound cargo to the distal tip at its own faster velocity to extend the distal singlets of the axoneme (Snow et al., 2004; Figure 2). Thus, animals lacking functional kinesin-II (e.g., kap-11 mutants) build a full-length cilium due to the redundant function of OSM-3, osm-3 mutants specifically lack the distal segment, and osm-3; kap-1 double mutants fail to make cilia because of the absence of functional Kinesin-II or OSM-3 (Snow et al., 2004). It should be noted that OSM-3 alone specifically extends distal singlets on some axonemes, but not others (Evans et al., 2006). An additional kinesin, KLP-6, has been implicated in male mating behavior and is required for proper localization of the human polycystin-2 homolog, PKD-2, to the cilium (Peden and Barr, 2005). This finding indicates that additional kinesin motors might be involved in ciliary transport, although intriguingly, IFT-like movement of the KLP-6 kinesin was not observed.
Table 2. Components and available mutants of the intraflagellar transport machinery
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Figure 2. Intraflagellar transport in C. elegans. Intraflagellar transport in C. elegans. Components of the IFT machinery and ciliary cargo assemble at or near the transition zone (basal body). Two kinesins, heterotrimeric kinesin-II and homodimeric OSM-3-kinesin, separately bind IFT particle subcomplexes A and B, respectively, and transport these together with IFT-dynein and cargo along the middle segment in the anterograde (+) direction. In the distal segment, OSM-3-kinesin alone transports the IFT particles and dynein/cargo. BBS proteins act to stabilize the association between the motors and IFT particle subcomplexes A and B. Components of the IFT machinery and presumably other ciliary molecules are recycled back to the base of the cilium using the IFT-dynein molecular motor. The lengths of the transition zone (1 μm), middle segment (4 μm) and distal segment (2.5 μm) regions are shown (for amphid cilia) along with transverse view schematics of the microtubule arrangements (on top).
The two sequential anterograde IFT-pathways are coordinated by at least two types of regulator proteins. Two C. elegans homologs of human Bardet-Biedl Syndrome (BBS) proteins (BBS-7 and BBS-8) have been shown to stabilize the IFT-particle subcomplexes A and B which are bound to the Kinesin-II and OSM-3 IFT-motors, respectively (Blacque et al., 2004; Ou et al., 2005a; Snow et al., 2004). Abbrogation of BBS protein function results in slightly truncated cilia and chemosensory or lipid accumulation defects (Blacque et al., 2004; Mak et al., 2006). The implications for this observation are of interest given that BBS, which is characterized by a diverse array of ailments, including obesity, cystic kidneys, and retinal degeneration, is one of a growing number of known ciliopathies (Beales, 2005; Blacque and Leroux, 2006). At least eight genes encoding BBS proteins are present in C. elegans (Table 2). The second modulator of the sequential IFT pathway, a conserved ciliary protein also first characterized in C. elegans, DYF-1, specifically docks the OSM-3 kinesin onto IFT-particles and simultaneously activates its motor activity; a dyf-1 mutant therefore specifically lacks the distal segment singlet microtubules (Ou et al., 2005a).
Movie 1. cilia of C. elegans as seen by time-lapse microscopy of GFP-labelled OSM-1.
The ability to analyze strains bearing GFP-tagged IFT proteins by time-lapse microscopy in C. elegans has provided researchers with a powerful means to dissect IFT function and study ciliary mutants (Orozco et al., 1999). Until now, this technique to study cilia function has distinguished C. elegans from the other prominent ciliary model organism, Chlamydomonas. In addition to providing crucial information about BBS and various IFT-associated proteins such as DYF-1, such in vivo studies are complemented by the fact that in many C. elegans ciliary mutants, abnormal IFT causes defects in sensory cilia structures and sensory behavior. For example, osm-3 and che-3 mutants possess defects in the functions of the anterograde IFT-kinesin and retrograde IFT-dynein, respectively, and display structural defects in the sensory cilia and corresponding deficiencies in osmotic avoidance and chemotaxis (Signor et al., 1999a; Wicks et al., 2000). Notably, the first evidence that biochemically-fractionated IFT-particle subunits identified in Chlamydomonas are essential for ciliary assembly was based on the phenotypes of the corresponding C. elegans mutants, such as osm-1/IFT172, osm-6/IFT52, osm-5, che-2, che-11 and che-13 and daf-10 (Brazelton et al., 2001; Cole et al., 1998; Qin et al. 2001; Scholey et al., 2004; Table 3). In addition, other components of the IFT machinery present in the Chlamydomonas flagellar proteome but not specifically identified in biochemical fractionations of IFT-particles (Pazour et al., 2005) have first been described in C. elegans, including DYF-2, a protein that may help bridge the IFT subcomplexes A and B (Efimenko et al., 2006), DYF-3, a protein associated with polycystic kidney disease that is likely part of IFT subcomplex B (Ou et al., 2005b), and IFTA-1 (IFT-Associated protein 1), a likely subcomplex A protein (Blacque et al., 2006). Each of these mutants are characterized phenotypically as having cilia structure and chemosensory defects.
Table 3. Cilia-related genes with corresponding genetic map positions and phenotypes
| Name | Other name | Gene model | Genetic position (cM) | Ref. allele | che | daf | dyf | osm | Annotation | Reference |
|---|---|---|---|---|---|---|---|---|---|---|
| che-1 | tax-1, tax-5 | C55B7.12 | I:1.20 +/− 0.015 | e1034 | + | / | / | +/− | C2H2-type transcription fact. | Uchida et al., 2003 |
| che-2 | F38G1.1 | X:-19.76 +/− 0.06 | e1033 | + | + | + | + | IFT-particle B | Fujiwara et al., 1999 | |
| che-3 | osm-2, che-8, avr-1, caf-2 | F18C12.1 | I:2.47 +/− 0.023 | e1124 | + | + | + | + | IFT-dynein heavy chain | Wicks et al., 2000 |
| che-6 | IV:0.00 +/− 0.000 | e1126 | + | / | / | / | Abnormal IL2 basal bodies | |||
| che-10 | II:−2.80 +/− 0.244 | e1809 | + | / | + | + | ||||
| che-11 | C27A7.4 | V:3.67 +/− 0.031 | e1810 | + | + | + | + | IFT-particle A | Qin et al., 2001 | |
| che-12 | V:2.28 +/− 0.105 | e1812 | + | / | +/− | + | Sheath cell secretion | |||
| che-13 | che-9 | F59C6.7 | I:5.05 +/− 0.029 | e1805 | + | + | + | + | IFT-particle B | Haycraft et al., 2003 |
| che-14 | ptd-1 | F56H1.1 | I:0.45 +/− 0.015 | e1960 | + | / | +/- | / | Transmembrane receptor | Michaux et al., 2000 |
| daf-6 | ptr-7 | F31F6.5 | X:21.50 | e1377 | + | + | + | + | Sheath cell function | Perens and Shaham, 2005 |
| daf-10 | osm-4 | IV:4.05 +/− 0.002 | e1387 | + | + | + | + | IFT-particle A | Bell et al., 2006 | |
| daf-19 | daf-24 | F33H1.1a | II:2.11 +/− 0.012 | m86 | + | + | + | + | RFX family transription fact. | Swoboda et al., 2000 |
| dyf-1 | F54C1.5a | I:-0.53 +/− 0.177 | mn335 | + | / | + | + | OSM-3-kinesin activator | Ou et al., 2005 | |
| dyf-2 | ZK520.1/.3 | III:21.40 +/− 0.10 | m160 | + | / | + | / | IFT protein | Efimenko et al., 2006 | |
| dyf-3 | C04C3.5a | IV:-6.09 +/− 1.34 | mn331 | + | / | + | / | IFT protein associated with PKD | Murayama et al., 2005 | |
| dyf-4 | V:4.31 +/− 0.29 | m158 | + | / | + | / | ||||
| dyf-5 | I:3.62 +/− 0.036 | mn400 | + | / | + | / | ||||
| dyf-6 | X:2.19 +/− 0.050 | m175 | + | / | + | / | IFT protein | Bell et al., 2006 | ||
| dyf-7 | X:2.18 +/− 0.046 | m537 | + | / | + | / | ||||
| dyf-8 | C43C3.3 | X:1.44 +/− 0.005 | m539 | + | + | + | + | Transmembrane receptor (endoglin family) | Wicks and Plasterk, pers. comm. | |
| dyf-9 | V:24.22 +/− 0.35 | n1513 | + | + | + | / | ||||
| dyf-10 | I:1.53 +/− 0.040 | e1383 | + | / | + | / | ||||
| dyf-11 | X:-18.26 +/− 0.25 | ad1303 | + | / | + | / | ||||
| dyf-12 | X:2.18 +/− 0.074 | nr2344 | + | + | + | / | ||||
| dyf-13 | C27H5.7a | II:0.25 +/− 0.017 | mn396 | + | + | + | / | Distal segment assembly | Blacque et al., 2005 | |
| osm-1 | T27B1.1 | X:24.06 +/− 0.029 | p808 | + | + | + | + | IFT-particle B | Bell et al., 2006 | |
| osm-3 | caf-1, klp-2 | M02B7.3a | IV:-2.27 +/− 0.087 | p802 | + | + | + | + | IFT-kinesin | Shakir et al., 1993 |
| osm-5 | Y41G9A.1 | X:-12.68 +/− 0.02 | p813 | + | + | + | + | IFT-particle B | Haycraft et al., 2001 | |
| osm-6 | R31.3 | V:3.52+/− 0.025 | p811 | + | + | + | + | IFT-particle B | Collet et al., 1998 | |
| osm-12 | bbs-7 | Y75B8A.12 | III:16.09+/−0.297 | n1606 | + | / | + | + | Distal segment assembly, IFT particle stability | Blacque et al., 2004 |
| bbs-1 | Y105E8A.5 | I:24.52 +/− 0.030 | ok1111 | + | / | + | + | Distal segment assembly, IFT particle stability | May et al., 2006 | |
| bbs-8 | T25F10.5 | V:0.13 +/− 0.001 | nx77 | + | / | + | + | Distal segment assembly, IFT particle stability | Blacque et al., 2004 |
Whereas our understanding of the IFT transport process has matured significantly in the last few years, very little is known about the nature of the proteins that require IFT-mediated transport to reach their ciliary destination. Indeed, only radial spoke proteins had been found to be bona fide IFT cargo proteins in Chlamydomonas (Qin et al., 2004); now, several have surfaced in C. elegans. One class of IFT-cargo are the cilia-localized TRP-type channels OSM-9 and OCR-2, which are implicated in various chemosensory responses (Tobin et al., 2002). Both have been shown to undergo IFT (Qin et al., 2005), marking the first account of a non-axonemal component being visualized to move along a cilium. Interestingly, OSM-9 and OCR-2 depend on each other for their ciliary localization, and ectopic expression of OCR-2 in AWC neurons is sufficient to drive OSM-9 to the cilia in this neuron (Tobin et al., 2002). Another apparent IFT cargo is
