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. Author manuscript; available in PMC: 2013 Dec 15.
Published in final edited form as: Vision Res. 2012 Aug 16;75:11–18. doi: 10.1016/j.visres.2012.08.006

Transport and localization of signaling proteins in ciliated cells

Mehdi Najafi 1, Peter D Calvert 1,2
PMCID: PMC3514659  NIHMSID: NIHMS401881  PMID: 22922002

Abstract

Most cells in the human body elaborate cilia which serve a wide variety of functions, including cell and tissue differentiation during development, sensing physical and chemical properties of the extracellular milieu and mechanical force generation. Common among cilia is the transduction of external stimuli into signals that regulate the activities of the cilia and the cells that possess them. These functions require the transport and localization of specialized proteins to the cilium, a process that many recent studies have shown to be vital for normal cell function and, ultimately, the health of the organism. Here we discuss several mechanisms proposed for the transport and localization of soluble and peripheral membrane proteins to, or their exclusion from the ciliary compartment with a focus on how the structure of the cytoplasm and the size and shape of proteins influence these processes. Additionally, we examine the impact of cell and protein structure on our ability to accurately measure the relative concentrations of fluorescently tagged proteins amongst various cellular domains, which is integral to our understanding of the molecular mechanisms underlying protein localization and transport.

Keywords: Cilia, photoreceptor, diffusion, signal transduction

INTRODUCTION

Cilia are microtubule-based organelles that extend from the surface of most eukaryotic cells where they play important roles in numerous signaling pathways (Wheatley, 1995). For example during development cilia are involved in establishing the left-right organ asymmetry (Supp et al., 1997; McGrath et al., 2003) and in mediating hedgehog signaling (Corbit et al., 2005; Huangfu and Anderson, 2005; Rohatgi et al., 2007). Later in life they house signal transduction modules involved in several sensory systems including vision, chemosensation (McEwen et al., 2008), and mechanosensation (Praetorius and Spring, 2003). Mutations that impact the elaboration, maintenance and function of cilia result in a heterogeneous group of pathologies collectively termed ciliopathies (Fliegauf et al., 2007). Thus a clear view of the processes that govern cilium maintenance and function is essential for understanding ciliary disease mechanisms.

To serve their various signaling roles, cilia maintain protein and membrane compositions that are distinct from the rest of the cell. Specific sets of soluble and membrane proteins are localized to the cilium while other cellular proteins are excluded from this compartment. Two distinct, but not mutually exclusive mechanisms proposed to regulate protein access to the ciliary compartment are gating, in which diffusion barriers regulate protein transit through the base of cilia (reviewed in Rosenbaum and Witman, 2002; Baldari and Rosenbaum, 2010; Nachury et al., 2010; Hu and Nelson, 2011), and partitioning, whereby segregation of proteins between the cilium and the cell body is governed by the recently described steric interactions between proteins and cell structures (Najafi et al., 2012) or binding affinities between proteins and relatively immobile cytoplasmic elements (reviewed in Calvert et al., 2006).

Evaluation of the contributions of these different mechanisms to the regulation of protein complement in primary cilia has proven difficult due in large part to uncertainties in the geometry of the cytoplasmic aqueous spaces and the structural elements in cells as well as the finite resolution of conventional fluorescence microscopy (Peet et al., 2004; Calvert et al., 2010; Najafi et al., 2012). Moreover, a complete picture of the mechanisms underlying protein localization to cilia requires quantification of molecular dynamics within and between various compartments of live cells, which has recently begun to be applied but suffers from similar pitfalls. Here we examine the mechanisms of ciliary protein localization with a particular focus on the challenges of quantifying the relative distribution patterns and dynamics of fluorescently tagged proteins in live cells. We highlight a solution to these problems of imaging resolution and uncertainties in cell structure offered by the ciliary photoreceptors of the vertebrate retina.

A DIFFUSION BARRIER AT THE CILIUM BASE

Evidence for a membrane diffusion barrier

A diffusion barrier at the base of flagella was originally inferred from experiments that examined the distribution patterns of glycoproteins, including the sexual agglutinins, in the flagellar membrane and the cell bodies of Chlamydomonas reinhardtii and Chlamydomonas eugametos (Adair et al., 1983; Musgrave et al., 1986). Agglutinins are involved in adhesion of mating gametes during Chlamydomonas fusion and are dispersed on both ciliary and flagellar membranes. However, only the flagellar agglutinins can participate in mating and have adhesive activity, suggesting that two populations of agglutinins are present on the membranes of Chlamydomonas that are segregated to different cellular membrane compartments. Further evidence for an agglutinin diffusion barrier came from a study where agglutinin levels were spatially perturbed. Exposure of Chlamydomonas to agglutinin antibodies resulted in depletion of agglutinin specifically from the flagella (Hunnicutt et al., 1990). Since the agglutinin molecules on the plasma membrane were intact after this treatment, but did not readily replace the flagellar agglutinins, it was concluded that an agglutinin diffusion barrier prevented their diffusion from the cell body into the flagellar membrane. Similar evidence for diffusion barriers at the bases of cilia and flagellar in other cell types have been reported as well, including sperm cells (Gold Myles et al., 1984; Nehme et al., 1993), epithelia cells (Vieira et al., 2006; Hu et al., 2010), and photoreceptors (Spencer et al., 1988), where intrinsic membrane proteins including G protein coupled receptors (GPCRs) are shown to be confined to the ciliary membranes.

Additional evidence for a molecular diffusion barrier at the bases of cilia and flagella comes from the analysis of the lipid compositions of the cilium and plasma membranes. The lipid composition of ciliary and flagellar membranes were found to be significantly different from the membrane surrounding the cell body in many species, with the cilia/flagella membranes containing higher spatial densities of sterols, glycolipids and sphingolipids (Kaya et al., 1984; Chailley and Boisvieux-Ulrich, 1985; Kaneshiro, 1990) which may form lipid rafts into which specific ciliary membrane proteins partition (Tyler et al., 2009).These results suggested that, in addition to membrane proteins, the lipid components of the plasma membrane and the ciliary membrane do not readily exchange.

Molecular identity of the membrane diffusion barrier

On a structural level, freeze-fracture electron microscopy studies show an orderly arrangement of membrane particles extending approximately 100 nm distally from the transition fibers, called the ciliary necklace, that is found at the base of many cilia (Gilula and Satir, 1972; Weiss et al., 1977). In photoreceptor cells the necklace covers a larger region of membrane, extending over most of the approximately 1 µm “connecting cilium” membrane (Rohlich, 1975). While the ciliary necklace has been suggested to be the physical manifestation of the diffusion barrier (Christensen et al., 2007; Rohatgi and Snell, 2010), it should be noted that it is not present in all ciliated or flagellated cells, including sperm from some species (Gilula and Satir, 1972). Moreover, at the spatial frequency of ~20 nm, the particles do not appear to be at high enough density to significantly impede diffusion of membrane proteins the size of GPCRs, which typically have a diameter in the membrane of ~ 4–5 nm.

Although there appear to be many proteins localized to the bases and transition zones of cilia, direct experimental tests for a role in a diffusion barrier have been reported for only one so far, a member of a class of GTPase proteins called septins (Hu et al., 2010). Septins, of which there are currently 13 family members (Hall et al., 2005), were originally shown to be involved in forming a diffusion barrier for membrane proteins at the mother-bud neck of budding yeast (Barral et al., 2000; Takizawa et al., 2000). Recently they were also shown to be located at the base of primary cilia in epithelial cells (Hu et al., 2010; S K Kim et al., 2010) and at the annulus of sperm cell flagella (Ihara et al., 2005; Kissel et al., 2005). Knockout of spetin 4 in mouse resulted in mislocalization of an integral membrane protein, basigin, in sperm cells (Kwitny et al., 2010) suggesting a role as a diffusion barrier in flagella.

In an elegant and insightful set of experiments Hu et al. (2010) used partial siRNA knockdowns of septin2 in IMCD3 cells to show that replenishment of ciliary targeted membrane proteins tagged with GFP, including the somatostatin receptor 3, serotonin receptor 6, and Smoothened, after photobleaching in the cilium was hastened and that the ratio of fluorescence in the cilium versus the apical membrane, which they term the “barrier index”, declined when septin 2 expression was reduced. Together these results suggested that septin 2 was integral to the formation of the diffusion barrier in primary cilia.

Several lines of evidence, however, suggest that septins may not be the sole molecular players in the diffusion barrier. While the recovery of the membrane proteins to the ciliary membrane compartment in the Hu et al. (2010) study was accelerated in septin2 knock down cells, relative to untreated cells, it remained significantly slower than the equilibration of the proteins within the cilium and in the apical membrane after local bleaches, suggesting a significant barrier remained.

Moreover, that the most severe knockdown of septin2 resulted in complete loss of cilia suggests a more general role of septins as structural proteins that are required to maintain the structural integrity of cilia. Thus it can’t be ruled out that the reduction in the barrier to diffusion in the septin2 knockdown cells is an indirect effect of septins on the maintenance of ciliary structures that ultimately constitute the diffusion barrier. For example, it has been proposed that the sharp hairpin turn that the ciliary membrane makes where it adjoins the plasma membrane may itself impede diffusion of membrane proteins and lipids between apical and ciliary membranes due to torsion or a similar physical phenomenon that septins help to maintain (Nachury et al., 2010). Finding an experimental avenue to test the membrane torsion hypothesis is thus important to gaining a full mechanistic understanding of the membrane protein diffusion barrier.

Finally, while it is clear that photoreceptor membrane proteins are retained within the plasma membranes of the outer segments of cone and rod cells, recent studies examining the distribution of various septin family members in retina so far have not demonstrated their presence in the photoreceptor ciliary transition zone (Pache et al., 2005; Hara et al., 2007). Thus it is possible that the involvement of septins in ciliary diffusion barrier is cell-type specific.

It is important to note that although it is often stated in the literature that diffusion barriers slow molecular diffusion between subcellular compartments, in order to be effective in maintaining separation of molecules a diffusion barrier, if acting alone, has to completely prevent molecular flux. Without additional mechanisms, such as active transport, that would counter the back flux through a leaky barrier, i.e. a structure that retards but does not completely block diffusion, it is clear that the molecules would eventually equilibrate amongst the compartments to which they have access, albeit with a slower rate. Therefore, complete understanding of the impact of membrane regions that impede macromolecule diffusion (diffusion barriers) on the localization of molecules within cilia requires quantification of molecular flux rates into and out of the cilium and models of molecular dynamics that take the membrane geometries and local diffusivities into account, for example using an approach similar to that recently developed for photoreceptors (Calvert et al., 2010). Such analyses will provide crucial information as to whether or not a barrier is present, the magnitude of its permeability, and the degree to which a barrier is compromised in experiments like the septin 2 knockdowns.

ACCESS OF SOLUBLE AND PERIPHERAL MEMBRANE PROTEINS TO THE CILIARY COMPARTMENT

Unlike the seven membrane spanning G protein-coupled receptors intrinsic to sensory cilia membranes discussed above, many of the proteins involved in signal transduction in cilia are soluble or peripherally membrane bound via post-translationally added lipid moieties. The mechanisms regulating the access, retention or exclusion of these crucial signaling proteins to the ciliary compartment are not well understood.

Nuclear pore-like activity at the cilium base

One exciting possibility that has gained recent experimental support is that the cilium base may operate in a manner similar to nuclear pore complexes in regulating soluble protein access to the ciliary compartment. Dishinger et al. (2010) recently demonstrated that access of a cytosolic motor protein, KIF17, to cilia of cultured epithelial cells depends on a cilium-to-cytosol gradient of the GTP bound form of the nuclear import molecule, RanGTP, and that importin-β2 was an important cofactor in transport. This work was recently followed up with another interesting report from the same group demonstrating that several nucleorporins (NUPs), components of the nuclear pore complex, NUP37, 35, 93, and 62, are found at the base of primary cilia (Kee et al., 2012). Based on these results the Verhey group proposed a model for trafficking of cytoplasmic proteins into the ciliary compartment whereby entry of proteins larger than 30–40 kDa is mediated by nuclear import-like events (Kee et al., 2012).

Is there a diffusion barrier for soluble proteins at the cilium base?

For the above mechanism to be viable, a molecular size-dependent diffusion barrier at the cilium base is expected to prevent molecules from drifting into or out of the ciliary compartment. Based on measurements of the ratio of fluorescence elicited from fluorescently labeled dextran polymers of different sizes in cilia and the cytoplasm, also referred to as the “barrier index”, Kee et al. (2012) have proposed that a molecular size-dependent barrier to soluble molecules exists at the cilium base. In experiments reminiscent of those carried out by the Luby-Phelps group (Janson et al., 1996), Kee et al. found that of the 3, 10, 40 and 70 kDa dextrans they injected into live, ciliated hTERT-RPE cells, 3 and 10 kDa sizes enter the cilia with similar barrier indices that were approximately six-fold larger than the barrier indices for 40 and 70 kDa dextrans. Interestingly this study also showed that fluorescently labeled globular proteins of various sizes through 41 kDa that were expressed or injected into cells all had similar barrier indices which were nearly two fold higher than the 3 and 10 kDa dextrans. 67 kDa BSA had a lower index than the other globular proteins, but was intermediate between the 10 and 40 kDa dextrans. The authors concluded that the cilium base possess a size-dependent barrier that limits access to the cilium via diffusion to soluble molecules below ~ 40 kDa. However, owing to uncertainties in quantifying fluorescence ratios between different subcellular regions, and because fluorescence above background was in fact detectable in the cilia for all probes used, the Kee et al. result may better support an alternative mechanism for size-dependent soluble macromolecule access to the cilium based on steric interactions with ciliary and cellular structures (Najafi et al., 2012; see below).

CONTROL OF SOLUBLE AND PERIPHERAL MEMBRANE PROTEIN LEVELS IN PHOTORECEPTOR SENSORY CILIA REGULATES LIGHT SENSITIVITY

The distribution of soluble and peripheral membrane proteins in photoreceptor sensory cilia play key roles in setting photoreceptor light sensitivity (reviewed in Calvert et al., 2006; Slepak and Hurley, 2008; Burns and Pugh, 2010; Arshavsky and Burns, 2012). Most notably transducin, the G protein involved in phototransduction cascade activation, and arrestin-1, the GPCR binding protein involved in photoresponse termination, undergo a stimulus-dependent redistribution between the ciliary rod outer segment, where light signaling is initiated, and the rod cell body. Transducin moves out of the outer segment and arrestin moves into this compartment in response to light stimulation. While these phenomena were originally described more than 25 years ago (Broekhuyse et al., 1985; Brann and Cohen, 1987; Philp et al., 1987), the mechanistic underpinnings have only recently begun to emerge.

Until recently the prevailing hypotheses for the mechanisms underlying the light dependent redistribution of arrestin and transducin were that the proteins localize to either the cell body or to the outer segment by binding to immobile structures or intrinsic membrane proteins within these compartments. An important demonstration that the distribution patterns of arrestin and transducin in dark- and light-adapted rods was due to localization, rather than simple cytoplasmic space filling, came from a ground breaking study by the laboratory of Ed Pugh that demonstrated that the distribution of arrestin in dark and light adapted rods differed significantly from the distribution of transgenically expressed EGFP, a “teflon” protein that exhibits little binding affinity for cell structures and membranes (Peet et al., 2004). Thus, in the presence of steady illumination, arrestin was proposed to abandon weak binding sites that localized it to the cell body in favor of higher affinity sites in the outer segment, including phospho-rhodopsin (Nair et al., 2005) and phosphoinositides (Lee et al., 2003), with diffusion being the mode of transport. A series of in vitro biochemical experiments have led to the suggestion that α-tubulin (Nair et al., 2004), enolase 1 (Smith et al., 2011) and NSF (Huang et al., 2010), each of which is present primarily throughout the cell body and into the synaptic spherule of rods, may be biding partners that hold arrestin out of the outer segment of dark-adapted rods. However, high resolution imaging shows that the distribution patterns of α-tubulin (Peet et al., 2004) and NSF (Huang et al., 2010) do not match that of arrestin in the dark adapted rod cell body. Moreover, the estimated total masses of α-tubulin (Hiller and Weber, 1978) and enolase 1 (Smith et al., 2011), appear to be 10–100-fold below that required to localize the > 109 copies of arrestins in each rod.

Transducin, on the other hand, was proposed to localize to the outer segment of dark-adapted rods owing to its association with disc membranes via post-translationally added lipid moieties, Tα being acylated (Neubert et al., 1992) and Tγ being farnesylated (Fukada et al., 1990). The importance of the specific lipid moieties placed on the transducin subunits, in terms of the light-induced translocation was demonstrated by Kassai et al. (2005) who showed that replacing the farnesyl with geranylgeranyl on Tγ inhibited light-dependent transport of transducin in mice, possibly by altering association of Tγ with a protein called phosducin which is thought to induce folding of the Tγ farnesyl moiety into a cleft in the Tβ propeller domain (Gaudet et al., 1996), thus shielding it from membrane association. The importance for transducin subunit dissociation from disc membranes was demonstrated by adding a palmitoylation site to Tα, which again inhibited light-dependent transport to the cell body (Kerov and Artemyev, 2011). It was proposed that the heterotrimeric Tαβγ complex has higher membrane affinity than the Tα and Tβγ subunits alone so that dissociation of the complex upon light activation and exchange of GDP for GTP on Tα would lead to their dissociation from outer segment disc membranes and association with UNC119 (Zhang et al., 2011), a soluble prenyl binding protein, and phosducin, which would increase their solubility thus allowing them to diffuse into the cell body where they would presumably associate with membranes or other structures to maintain reduced outer segment levels (Calvert et al., 2006). However, to date cell body localized binding partners for transducin subunits have not been identified.

A diffusion barrier in the ciliary transition zone of photoreceptors was proposed as an alternative explanation for the cell body localization of arrestin and transducin (Pulvermuller et al., 2002). However, direct assessment of the permeability of the connecting cilium in rods for proteins in the size range of transducin and arrestin showed their transport through this structure was not substantially impeded (Calvert et al., 2010; Najafi et al., 2012). Thus while outer segment localization of arrestin and transducin appeared to be explainable by binding to disc membranes or disc-associated proteins, localization to the cell body remained paradoxical owing to the lack of evidence for diffusion barriers or appropriate binding partners in sufficient capacity to account for their inner segment localization.

THE INTERPLAY BETWEEN STERIC VOLUME EXCLUSION AND LOCAL BINDING IN REGUALTING SOLUBLE PROTEIN ACCESS TO PHOTORECEPTOR SENSORY CILIA

Our recent work offers a resolution to this paradox. We showed that the distribution patterns of soluble proteins in Xenopus ciliated rod photoreceptors were steeply molecular size-dependent with larger proteins being present in the outer segment in considerably lower abundance, relative to the cell body, than smaller molecules. Moreover, we showed that this effect was not due to a size-dependent diffusion barrier at the base of the rod cilium, but rather that it was mediated by steric interactions between molecules and cell structures that produced variable reductions in the aqueous cytoplasmic volumes available to the molecules (Najafi et al., 2012). This effect, often termed steric volume exclusion (SVE), has been known to physical chemists and ion channel biophysicists for decades (Giddings et al., 1968; Hille, 2001) but has largely been overlooked by cell biologists, with a few notable exceptions (Minton, 1992; Janson et al., 1996).

The physical underpinnings of the steric volume exclusion effect is that, at steady state, the concentration of a given soluble molecule that does not have significant binding interactions with cytoplasmic elements will be the same in all contiguous subcellular aqueous compartments to which it has access. Importantly, the volume on which the concentration depends is not the geometric volume of the cytoplasmic aqueous spaces subtended by the cell structures, but rather the volume accessible to the center of mass of the molecule (Fig. 1). An important conclusion to be gleaned from this insight is that the volume available to a given molecule will depend on the size and shape of the molecule as well as the shape of the aqueous space (Giddings et al., 1968; Minton, 1992; Janson et al., 1996; Najafi et al., 2012) (see below). Thus, the distribution pattern of soluble proteins in cells will depend on the size of the proteins and the spatially variable patterns and densities of cell structures.

Figure 1. The Steric Volume Exclusion effect.

Figure 1

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Cell structures (A) and macromolecules (B) reduce the volume available to a solute molecule by an amount proportional to the radius of gyration (rs) and the geometry of the space, represented in (A) as the distance between two parallel membranes (L). The result is that the total mass of a given protein found in a more structurally dense region of a cell will be lower than in a less dense region, and the relative magnitude of the difference will depend on the size of the molecule (C).

Spatial variations in the cytoplasmic structures found in photoreceptors are well characterized. The two major cytoplasmic compartments in rod photoreceptors possess significantly different densities of structures. The ciliary outer-segment compartment contains thousands of disc-shaped membranes that are organized into parallel plates with a uniform, ~ 12–15 nm spacing, that approaches the size of the transduction cascade molecules themselves (Rosenkranz, 1977; Nickell et al., 2007; Najafi et al., 2012). In contrast, the cell body appears to be much less densely populated with structures that are less organized (Townes-Anderson et al., 1985). This arrangement of neighboring cytoplasmic compartments is expected to result in significant partitioning of molecules out of the outer segment and into the cell body due simply to steric volume exclusion.

From the dependence of the steric volume exclusion effect on molecular size and shape, and on the geometry of the cytoplasmic aqueous spaces, directly follows an important and novel mechanism for regulating the mass of soluble proteins within the rod outer segment, or any other cellular microcompartment for that matter. Changes in molecular size, or the geometry of the aqueous spaces, will lead to changes in the mass of soluble proteins within the microcompartments and thus possibly to changes in the efficiency of reactions in which they are involved. In the case of photoreceptors, changes in the sizes of arrestin and transducin have been demonstrated through binding interactions. Arrestin forms dimers and tetramers at physiological concentrations when not bound to rhodopsin (Schubert et al., 1999; Imamoto et al., 2003; Hanson et al., 2007; Hanson et al., 2008; M Kim et al., 2011). Transducin subunits associate with proteins that shield their lipid moieties and increase their effective size (Gaudet et al., 1996; Zhang et al., 2011). These interactions coupled with steric volume exclusion may thus be the mechanism driving arrestin and transducin out of the outer segment under appropriate illumination conditions (Najafi et al., 2012).

CELL STRUCTURES AND THE RESOLUTION LIMITS OF FLUORESCENCE MICROSCOPY

Several studies have used the ratio of fluorescence intensities of the ciliary compartment versus the cell body as a metric for the relative concentrations or spatial densities of fluorescently tagged proteins, and ultimately to support assertions that the proteins are localized, excluded or neither from a particular subcellular region. An important consideration when interpreting fluorescence intensities from various regions within a cell is the geometry of the cell structures being imaged and the spatial resolving power of the microscope optics. While the resolving power of an optical system is fairly straightforward to evaluate, in most cells the specific variation in subcellular structural densities is more difficult to define since, despite having a wealth of high-resolution EM studies that have revealed cytoplasmic ultrastructure, in general no two cells possess precisely the same spatial distribution of structures in the cytoplasm. Notable exceptions to this rule are the ciliary photoreceptors of the vertebrate retina (Fig. 2), which have emerged as unparalleled models for understanding the crucial interplay between cytoplasmic architecture and protein structures in regulating cell function and signaling (Peet et al., 2004; Calvert et al., 2010; Najafi et al., 2012).

Figure 2. Cell structures and the resolution limit of fluorescence microscopy.

Figure 2

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Most cell structures are below the resolution limit of fluorescence microscopy, making quantitative comparisons of fluorescent protein concentrations amongst different cell regions challenging. This is particularly true of ciliated cells where the ~0.35 µm diameter of the cilium itself places it well below the resolution limit the excitation beam of confocal microscopy (the point spread function, psf, red ellipse) (A). Amphibian rod photoreceptors offer the advantage that both the ciliary compartment and the cell body are large enough to accommodate the full psf (B). In addition to the envelope geometry, sub-resolution structures within the cytoplasm, and the regions near them that are excluded to the radius of gyration of a given fluorescent molecule (shown in grey), occlude the sampling volume of the psf (C). Together these factors lead to variation in fluorescence levels measured between different cell regions despite uniform concentrations of fluorescent molecules within the contiguous aqueous cytoplasm. (See text for details)

The study by Peet et al. (2004) was the first to quantify the impact of subcellular structures on the spatial fluorescence pattern of a soluble fluorescent molecule, enhanced green fluorescent protein (EGFP) in a ciliated cell. In this study, live rod photoreceptors from retinas of the African clawed frog, Xenopus lavis, that expressed EGFP under the opsin promoter were oriented in the imaging chamber such that the axial dimensions of the photoreceptors were orthogonal to the direction of light propagation in the microscope, and imaged confocally. The authors showed that the fluorescence intensities in the ciliary outer segment were about half that of the brightest regions of the cells, which was predicted based in the well characterized cytoplasmic architecture of the ciliary outer segment that contains a highly ordered and uniformly spaced stacks of membranous discs. The key insight was that at any given position within the cell, the resolution limit of confocal imaging resulted in the averaging of fluorescence within the cytoplasmic space sampled by the psf which itself consisted of two fractions: (1) the aqueous spaces to which the soluble EGFP had access and, (2) the space occupied by sub-resolution cytoplasmic structures to which it did not. Thus, the fluorescence intensity recorded at any position within a cell is a fraction of the maximum intensity that would have been recorded from a psf sampling a hypothetical unobstructed volume of the cytoplasm that contains no cytoplasmic structures (Fig. 2).

Recently we have extended and modified the hypothesis posited by Peet et al. (2004) to include the impact of accessible cytoplasmic aqueous spaces that vary depending on the size of fluorescent solutes (Najafi et al., 2012; see below). The complete expression for the fluorescence, F, elicited by excitation with a diffraction limited beam in a confocal imaging system is

F(x,y,z)=I(x,y,z)·c(x,y,z)·αγk·faq(x,y,z)·fac(x,y,z), (Eq. 1)

where I(x,y,z) is the Gaussian intensity profile of the excitation beam in 3D (i.e. the point spread function, psf), c(x,y,z) is the concentration of fluorescent molecules, αγ is the product of the absorbance cross section and quantum yield of the fluorophore, k is the efficiency of the imaging system in capturing emitted photons, and faq(x,y,z) and fac(x,y,z) are the fraction of the geometric volume encompassed by the psf that is occupied by aqueous cytoplasm and the fraction of the psf volume that is accessible to the center of mass of a particular solute, respectively (Fig. 2). Importantly, this expression shows that, because I is not varying and with the reasonable assumption that c is uniform for soluble proteins within the accessible cytoplasmic spaces, the ratio of fluorescence recorded from any two positions within a cell reflects the ratio of the local fractional psf sampling volumes,

F(x,y,z)F(x,y,z)'=faq(x,y,z)·fac(x,y,z)faq(x,y,z)'·fac(x,y,z)'. (Eq. 2)

The key insight from Eq. 2 is that, in the case of soluble molecules in the contiguous aqueous cytoplasm, the variation in fluorescence measured in different cell regions does not necessarily reflect differences in the concentrations of the molecules relative to the accessible cytoplasm. Moreover, because the fraction of accessible volumes depends on the size of a macromolecule, the ratio may significantly change with molecules of different sizes. Thus without some a priori knowledge of the geometry of the cytoplasmic spaces being probed, it is difficult to make arguments about the relative access of proteins to different cell regions at either the qualitative or quantitative levels.

IS THERE A ROLE OF STERIC VOLUME EXCLUSION IN REGULATING PROTEIN ACCESS TO PRIMARY CILIA?

With these considerations, it is clear that caution should be used when interpreting fluorescence ratios from different cell regions. Photoreceptors offer the unprecedented advantage of detailed knowledge of the geometry of the cytoplasmic spaces, as well as accurate estimations of the numbers, shapes and volumes of proteins in the outer segment. Although many detailed EM studies have been carried out on primary cilia and flagella that show that the axoneme and associated structures appear to be less spatially constraining than the discs of rod outer segments, we know less about the numbers, sizes and shapes and the volumes occupied by ciliary proteins, which are generally not detected by EM. Thus, the fluorescence ratios between cilium and cell body used to characterize putative diffusion barriers should be interpreted with some caution. For instance, the presence of fluorescence within the cilia reported in Kee et al. (2012) for even the largest molecules examined, 70kDa dextran and 67 kDa BSA, may better support a steric volume exclusion effect similar to that described by Najafi et al. (2012), perhaps caused by ciliary protein crowding, over the proposed diffusion barrier. Further analysis will be required to resolve this question.

CONCLUSIONS AND PERSPECTIVES

Transport and retention of signaling proteins within cilia is crucial for ciliary signaling and its regulation. A variety of mechanisms appear to be at play. While integral membrane proteins such as GPCRs are likely delivered into the ciliary compartment via IFT or similar transport systems and retained there by a membrane diffusion barrier, soluble and peripheral membrane proteins appear to segregate between the cilium and the cell body via partitioning mechanisms, including local binding and steric volume exclusion. Importantly, changes in signaling protein size coupled with steric volume exclusion may be a key mechanism for regulating transduction protein levels in photoreceptor sensory cilia as well as other cilia and cell signaling microcompartments. Finally, a nuclear pore-like diffusion barrier may control access and egress to the ciliary compartment for soluble and peripheral membrane proteins, although the molecular weight cutoff for this potential mechanism remains unclear.

Highlights.

  • Brief review of signaling protein transport and localization within ciliated cells.

  • Focus on the mechanisms of protein retention within, or exclusion from cilia.

  • Impact of steric volume exclusion on cell signaling protein distributions.

  • Resolution limit of fluorescent imaging and interpretation of protein distribution.

ACKNOWLEDGEMENTS

The work in the author’s lab is supported by National Eye Institute grant R01 EY018421 (PDC). PDC is the recipient of a Career Development Award from Research to Prevent Blindness (RPB). The SUNY Upstate Department of Ophthalmology is the recipient of an unrestricted grant from RPB and receives support from Lions district 20-Y1.

Footnotes

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Submitted by invitation for a special issue of Vision Research devoted to the 14th Vision Research Conference

REFERENCES

  1. Adair WS, Hwang C, Goodenough UW. Identification and visualization of the sexual agglutinin from the mating-type plus flagellar membrane of Chlamydomonas. Cell. 1983;33(1):183–193. doi: 10.1016/0092-8674(83)90347-1. [DOI] [PubMed] [Google Scholar]
  2. Arshavsky VY, Burns ME. Photoreceptor Signaling: Supporting Vision across a Wide Range of Light Intensities. Journal of Biological Chemistry. 2012;287(3):1620–1626. doi: 10.1074/jbc.R111.305243. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Baldari CT, Rosenbaum J. Intraflagellar transport: it's not just for cilia anymore. Curr Opin Cell Biol. 2010;22(1):75–80. doi: 10.1016/j.ceb.2009.10.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Barral Y, Mermall V, Mooseker MS, Snyder M. Compartmentalization of the cell cortex by septins is required for maintenance of cell polarity in yeast. Molecular Cell. 2000;5(5):841–851. doi: 10.1016/s1097-2765(00)80324-x. [DOI] [PubMed] [Google Scholar]
  5. Brann MR, Cohen LV. Diurnal expression of transducin mRNA and translocation of transducin in rods of rat retina. Science. 1987;235(4788):585–587. doi: 10.1126/science.3101175. [DOI] [PubMed] [Google Scholar]
  6. Broekhuyse RM, Tolhuizen EF, Janssen AP, Winkens HJ. Light induced shift and binding of S-antigen in retinal rods. Current eye research. 1985;4(5):613–618. doi: 10.3109/02713688508999993. [DOI] [PubMed] [Google Scholar]
  7. Burns ME, Pugh EN., Jr. Lessons from photoreceptors: turning off g-protein signaling in living cells. Physiology (Bethesda) 2010;25(2):72–84. doi: 10.1152/physiol.00001.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Calvert PD, Schiesser WE, Pugh EN., Jr Diffusion of a soluble protein, photoactivatable GFP, through a sensory cilium. J Gen Physiol. 2010;135(3):173–196. doi: 10.1085/jgp.200910322. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Calvert PD, Strissel KJ, Schiesser WE, Pugh EN, Jr, Arshavsky VY. Light-driven translocation of signaling proteins in vertebrate photoreceptors. Trends Cell Biol. 2006;16(11):560–568. doi: 10.1016/j.tcb.2006.09.001. [DOI] [PubMed] [Google Scholar]
  10. Chailley B, Boisvieux-Ulrich E. Detection of plasma membrane cholesterol by filipin during microvillogenesis and ciliogenesis in quail oviduct. Journal of Histochemistry and Cytochemistry. 1985;33(1):1–10. doi: 10.1177/33.1.3965567. [DOI] [PubMed] [Google Scholar]
  11. Christensen ST, Pedersen LB, Schneider L, Satir P. Sensory cilia and integration of signal transduction in human health and disease. Traffic. 2007;8(2):97–109. doi: 10.1111/j.1600-0854.2006.00516.x. [DOI] [PubMed] [Google Scholar]
  12. Corbit KC, Aanstad P, Singla V, Norman AR, Stainier DYR, Reiter JF. Vertebrate Smoothened functions at the primary cilium. Nature. 2005;437(7061):1018–1021. doi: 10.1038/nature04117. [DOI] [PubMed] [Google Scholar]
  13. Dishinger JF, Kee HL, Jenkins PM, Fan S, Hurd TW, Hammond JW, Truong YNT, Margolis B, Martens JR, Verhey KJ. Ciliary entry of the kinesin-2 motor KIF17 is regulated by importin-B2 and RanGTP. Nature Cell Biology. 2010;12(7):703–710. doi: 10.1038/ncb2073. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Fliegauf M, Benzing T, Omran H. When cilia go bad: Cilia defects and ciliopathies. Nature Reviews Molecular Cell Biology. 2007;8(11):880–893. doi: 10.1038/nrm2278. [DOI] [PubMed] [Google Scholar]
  15. Fukada Y, Takao T, Ohguro H, Yoshizawa T, Akino T, Shimonishi Y. Farnesylated gamma-subunit of photoreceptor G protein indispensable for GTP-binding. Nature. 1990;346(6285):658–660. doi: 10.1038/346658a0. [DOI] [PubMed] [Google Scholar]
  16. Gaudet R, Bohm A, Sigler PB. Crystal structure at 2.4 angstroms resolution of the complex of transducin betagamma and its regulator, phosducin. Cell. 1996;87(3):577–588. doi: 10.1016/s0092-8674(00)81376-8. [DOI] [PubMed] [Google Scholar]
  17. Giddings JC, Kucera E, Russell CP, Myers MN. Statistical theory for the equilibrium distribution of rigid molecules. J Phys Chem. 1968;72(13):4397–4408. [Google Scholar]
  18. Gilula NB, Satir P. The ciliary necklace. A ciliary membrane specialization. Journal of Cell Biology. 1972;53(2):494–509. doi: 10.1083/jcb.53.2.494. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Gold Myles D, Primakoff P, Koppel DE. A localized surface protein of guinea pig sperm exhibits free diffusion in its domain. Journal of Cell Biology. 1984;98(5):1905–1909. doi: 10.1083/jcb.98.5.1905. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Hall PA, Jung K, Hillan KJ, Russell SE. Expression profiling the human septin gene family. The Journal of pathology. 2005;206(3):269–278. doi: 10.1002/path.1789. [DOI] [PubMed] [Google Scholar]
  21. Hanson SM, Dawson ES, Francis DJ, Van Eps N, Klug CS, Hubbell WL, Meiler J, Gurevich VV. A model for the solution structure of the rod arrestin tetramer. Structure. 2008;16(6):924–934. doi: 10.1016/j.str.2008.03.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Hanson SM, Van Eps N, Francis DJ, Altenbach C, Vishnivetskiy SA, Arshavsky VY, Klug CS, Hubbell WL, Gurevich VV. Structure and function of the visual arrestin oligomer. Embo J. 2007;26(6):1726–1736. doi: 10.1038/sj.emboj.7601614. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Hara A, Taguchi A, Niwa M, Aoki H, Yamada Y, Ito H, Nagata K-i, Kunisada T, Mori H. Localization of septin 8 in murine retina, and spatiotemporal expression of septin 8 in a murine model of photoreceptor cell degeneration. Neuroscience Letters. 2007;423(3):205–210. doi: 10.1016/j.neulet.2007.07.030. [DOI] [PubMed] [Google Scholar]
  24. Hille B. Ionic Channels of Excitable Membranes. Third ed. Sunderland, MA, USA.: Sinauer; 2001. [Google Scholar]
  25. Hiller G, Weber K. Radioimmunoassay for tubulin: a quantitative comparison of the tubulin content of different established tissue culture cells and tissues. Cell. 1978;14(4):795–804. doi: 10.1016/0092-8674(78)90335-5. [DOI] [PubMed] [Google Scholar]
  26. Hu Q, Nelson WJ. Ciliary diffusion barrier: the gatekeeper for the primary cilium compartment. Cytoskeleton. 2011;68(6):313–324. doi: 10.1002/cm.20514. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Hu Q, Milenkovic L, Jin H, Scott MP, Nachury MV, Spiliotis ET, Nelson WJ. A septin diffusion barrier at the base of the primary cilium maintains ciliary membrane protein distribution. Science. 2010;329(5990):436–439. doi: 10.1126/science.1191054. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Huang SP, Brown BM, Craft CM. Visual Arrestin 1 acts as a modulator for N-ethylmaleimide-sensitive factor in the photoreceptor synapse. J Neurosci. 2010;30(28):9381–9391. doi: 10.1523/JNEUROSCI.1207-10.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Huangfu D, Anderson KV. Cilia and Hedgehog responsiveness in the mouse. Proceedings of the National Academy of Sciences of the United States of America. 2005;102(32):11325–11330. doi: 10.1073/pnas.0505328102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Hunnicutt GR, Kosfiszer MG, Snell WJ. Cell body and flagellar agglutinins in Chlamydomonas reinhardtii: The cell body plasma membrane is a reservoir for agglutinins whose migration to the flagella is regulated by a functional barrier. Journal of Cell Biology. 1990;111(4):1605–1616. doi: 10.1083/jcb.111.4.1605. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Ihara M, et al. Cortical organization by the septin cytoskeleton is essential for structural and mechanical integrity of mammalian spermatozoa. Developmental Cell. 2005;8(3):343–352. doi: 10.1016/j.devcel.2004.12.005. [DOI] [PubMed] [Google Scholar]
  32. Imamoto Y, Tamura C, Kamikubo H, Kataoka M. Concentration-dependent tetramerization of bovine visual arrestin. Biophys J. 2003;85(2):1186–1195. doi: 10.1016/S0006-3495(03)74554-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Janson LW, Ragsdale K, Luby-Phelps K. Mechanism and size cutoff for steric exclusion from actin-rich cytoplasmic domains. Biophysical Journal. 1996;71(3):1228–1234. doi: 10.1016/S0006-3495(96)79367-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Kaneshiro ES. Lipids of ciliary and flagellar membranes. In: Bloodgood, editor. Ciliary and Flagellar Membranes. New York: Plenum Press; 1990. pp. 241–265. [Google Scholar]
  35. Kassai H, et al. Farnesylation of retinal transducin underlies its translocation during light adaptation. Neuron. 2005;47(4):529–539. doi: 10.1016/j.neuron.2005.07.025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Kaya K, Ramesha CS, Thompson GA., Jr Temperature-induced changes in the hydroxy and non-hydroxy fatty acid-containing sphingolipids abundant in the surface membrane of Tetrahymena pyriformis NT-1. Journal of lipid research. 1984;25(1):68–74. [PubMed] [Google Scholar]
  37. Kee HL, Dishinger JF, Lynne Blasius T, Liu CJ, Margolis B, Verhey KJ. A size-exclusion permeability barrier and nucleoporins characterize a ciliary pore complex that regulates transport into cilia. Nature Cell Biology. 2012;14(4):431–437. doi: 10.1038/ncb2450. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Kerov V, Artemyev NO. Diffusion and light-dependent compartmentalization of transducin. Molecular and cellular neurosciences. 2011;46(1):340–346. doi: 10.1016/j.mcn.2010.10.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Kim M, Hanson SM, Vishnivetskiy SA, Song X, Cleghorn WM, Hubbell WL, Gurevich VV. Robust self-association is a common feature of mammalian visual arrestin-1. Biochemistry. 2011;50(12):2235–2242. doi: 10.1021/bi1018607. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Kim SK, et al. Planar cell polarity acts through septins to control collective cell movement and ciliogenesis. Science. 2010;329(5997):1337–1340. doi: 10.1126/science.1191184. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Kissel H, Georgescu MM, Larisch S, Manova K, Hunnicutt GR, Steller H. The Sept4 septin locus is required for sperm terminal differentiation in mice. Developmental Cell. 2005;8(3):353–364. doi: 10.1016/j.devcel.2005.01.021. [DOI] [PubMed] [Google Scholar]
  42. Kwitny S, Klaus AV, Hunnicutt GR. The annulus of the mouse sperm tail is required to establish a membrane diffusion barrier that is engaged during the late steps of spermiogenesis. Biology of Reproduction. 2010;82(4):669–678. doi: 10.1095/biolreprod.109.079566. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Lee SJ, Xu H, Kang LW, Amzel LM, Montell C. Light adaptation through phosphoinositide-regulated translocation of Drosophila visual arrestin. Neuron. 2003;39(1):121–132. doi: 10.1016/s0896-6273(03)00390-8. [DOI] [PubMed] [Google Scholar]
  44. McEwen DP, Jenkins PM, Martens JR. Chapter 12 Olfactory Cilia: Our Direct Neuronal Connection to the External World, edited. 2008. pp. 333–370. [DOI] [PubMed] [Google Scholar]
  45. McGrath J, Somlo S, Makova S, Tian X, Brueckner M. Two populations of node monocilia initiate left-right asymmetry in the mouse. Cell. 2003;114(1):61–73. doi: 10.1016/s0092-8674(03)00511-7. [DOI] [PubMed] [Google Scholar]
  46. Minton AP. Confinement as a determinant of macromolecular structure and reactivity. Biophys J. 1992;63(4):1090–1100. doi: 10.1016/S0006-3495(92)81663-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Musgrave A, de Wildt P, van Etten I, Pijst H, Scholma C, Kooyman R, Homan W, van den Ende H. Evidence for a functional membrane barrier in the transition zone between the flagellum and cell body of Chlamydomonas eugametos gametes. Planta. 1986;167(4):544–553. doi: 10.1007/BF00391231. [DOI] [PubMed] [Google Scholar]
  48. Nachury MV, Seeley ES, Jin H. Trafficking to the ciliary membrane: how to get across the periciliary diffusion barrier? Annu Rev Cell Dev Biol. 2010;26:59–87. doi: 10.1146/annurev.cellbio.042308.113337. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Nair KS, Hanson SM, Kennedy MJ, Hurley JB, Gurevich VV, Slepak VZ. Direct binding of visual arrestin to microtubules determines the differential subcellular localization of its splice variants in rod photoreceptors. J Biol Chem. 2004;279(39):41240–41248. doi: 10.1074/jbc.M406768200. [DOI] [PubMed] [Google Scholar]
  50. Nair KS, et al. Light-dependent redistribution of arrestin in vertebrate rods is an energy-independent process governed by protein-protein interactions. Neuron. 2005;46(4):555–567. doi: 10.1016/j.neuron.2005.03.023. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Najafi M, Maza NA, Calvert PD. Steric volume exclusion sets soluble protein concentrations in photoreceptor sensory cilia. Proceedings of the National Academy of Sciences of the United States of America. 2012;109(1):203–208. doi: 10.1073/pnas.1115109109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Nehme CL, Cesario MM, Myles DG, Koppel DE, Bartles JR. Breaching the diffusion barrier that compartmentalizes the transmembrane glycoprotein CE9 to the posterior-tail plasma membrane domain of the rat spermatozoon. Journal of Cell Biology. 1993;120(3):687–694. doi: 10.1083/jcb.120.3.687. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Neubert TA, Johnson RS, Hurley JB, Walsh KA. The rod transducin alpha subunit amino terminus is heterogeneously fatty acylated. J Biol Chem. 1992;267(26):18274–18277. [PubMed] [Google Scholar]
  54. Nickell SP, Park SH, Baumeister W, Palczewski K. Three-dimensional architecture of murine rod outer segments determined by cryoelectron tomography. Journal of Cell Biology. 2007;177(5):917–925. doi: 10.1083/jcb.200612010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Pache M, Zieger B, Bläser S, Meyer P. Immunoreactivity of the Septins SEPT4, SEPT5, and SEPT8 in the Human Eye. Journal of Histochemistry & Cytochemistry. 2005;53(9):1139–1147. doi: 10.1369/jhc.4A6588.2005. [DOI] [PubMed] [Google Scholar]
  56. Peet JA, Bragin A, Calvert PD, Nikonov SS, Mani S, Zhao X, Besharse JC, Pierce EA, Knox BE, Pugh EN., Jr Quantification of the cytoplasmic spaces of living cells with EGFP reveals arrestin-EGFP to be in disequilibrium in dark adapted rod photoreceptors. J Cell Sci. 2004;117Pt 14:3049–3059. doi: 10.1242/jcs.01167. [DOI] [PubMed] [Google Scholar]
  57. Philp NJ, Chang W, Long K. Light-stimulated protein movement in rod photoreceptor cells of the rat retina. FEBS letters. 1987;225(1–2):127–132. doi: 10.1016/0014-5793(87)81144-4. [DOI] [PubMed] [Google Scholar]
  58. Praetorius HA, Spring KR. The renal cell primary cilium functions as a flow sensor. Current Opinion in Nephrology and Hypertension. 2003;12(5):517–520. doi: 10.1097/00041552-200309000-00006. [DOI] [PubMed] [Google Scholar]
  59. Pulvermuller A, Giessl A, Heck M, Wottrich R, Schmitt A, Ernst OP, Choe HW, Hofmann KP, Wolfrum U. Calcium-dependent assembly of centrin-G-protein complex in photoreceptor cells. Molecular and cellular biology. 2002;22(7):2194–2203. doi: 10.1128/MCB.22.7.2194-2203.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Rohatgi R, Snell WJ. The ciliary membrane. Curr Opin Cell Biol. 2010;22(4):541–546. doi: 10.1016/j.ceb.2010.03.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Rohatgi R, Milenkovic L, Scott MP. Patched1 regulates hedgehog signaling at the primary cilium. Science. 2007;317(5836):372–376. doi: 10.1126/science.1139740. [DOI] [PubMed] [Google Scholar]
  62. Rohlich P. The sensory cilium of retinal rods is analogous to the transitional zone of motile cilia. Cell and tissue research. 1975;161(3):421–430. doi: 10.1007/BF00220009. [DOI] [PubMed] [Google Scholar]
  63. Rosenbaum JL, Witman GB. Intraflagellar transport. Nature Reviews Molecular Cell Biology. 2002;3(11):813–825. doi: 10.1038/nrm952. [DOI] [PubMed] [Google Scholar]
  64. Rosenkranz J. New aspects of the ultrastructure of frog rod outer segments. International Review of Cytology. 1977;Vol. 50:25–158. doi: 10.1016/s0074-7696(08)60098-4. [DOI] [PubMed] [Google Scholar]
  65. Schubert C, Hirsch JA, Gurevich VV, Engelman DM, Sigler PB, Fleming KG. Visual Arrestin Activity May Be Regulated by Self-association. Journal of Biological Chemistry. 1999;274(30):21186–21190. doi: 10.1074/jbc.274.30.21186. [DOI] [PubMed] [Google Scholar]
  66. Slepak VZ, Hurley JB. Mechanism of light-induced translocation of arrestin and transducin in photoreceptors: interaction-restricted diffusion. IUBMB life. 2008;60(1):2–9. doi: 10.1002/iub.7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Smith WC, Bolch S, Dugger DR, Li J, Esquenazi I, Arendt A, Benzenhafer D, McDowell JH. Interaction of arrestin with enolase1 in photoreceptors. Invest Ophthalmol Vis Sci. 2011;52(3):1832–1840. doi: 10.1167/iovs.10-5724. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Spencer M, Detwiler PB, Bunt-Milam AH. Distribution of membrane proteins in mechanically dissociated retinal rods. Investigative Ophthalmology and Visual Science. 1988;29(7):1012–1020. [PubMed] [Google Scholar]
  69. Supp DM, Witte DP, Steven Potter S, Brueckner M. Mutation of an axonemal dynein affects left-right asymmetry in inversus viscerum mice. Nature. 1997;389(6654):963–966. doi: 10.1038/40140. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Takizawa PA, DeRisi JL, Wilhelm JE, Vale RD. Plasma membrane compartmentalization in yeast by messenger RNA transport and a septin diffusion barrier. Science. 2000;290(5490):341–344. doi: 10.1126/science.290.5490.341. [DOI] [PubMed] [Google Scholar]
  71. Townes-Anderson E, MacLeish PR, Raviola E. Rod cells dissociated from mature salamander retina: ultrastructure and uptake of horseradish peroxidase. J Cell Biol. 1985;100(1):175–188. doi: 10.1083/jcb.100.1.175. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Tyler KM, Fridberg A, Toriello KM, Olson CL, Cieslak JA, Hazlett TL, Engman DM. Flagellar membrane localization via association with lipid rafts. Journal of Cell Science. 2009;122(6):859–866. doi: 10.1242/jcs.037721. [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Vieira OV, Gaus K, Verkade P, Fullekrug J, Vaz WLC, Simons K. FAPP2, cilium formation, and compartmentalization of the apical membrane in polarized Madin-Darby canine kidney (MDCK) cells. Proceedings of the National Academy of Sciences of the United States of America. 2006;103(49):18556–18561. doi: 10.1073/pnas.0608291103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Weiss RL, Goodenough DA, Goodenough UW. Membrane particle arrays associated with the basal body and with contractile vacuole secretion in Chlamydomonas. Journal of Cell Biology. 1977;72(1):133–143. doi: 10.1083/jcb.72.1.133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Wheatley DN. Primary cilia in normal and pathological tissues. Pathobiology : journal of immunopathology, molecular and cellular biology. 1995;63(4):222–238. doi: 10.1159/000163955. [DOI] [PubMed] [Google Scholar]
  76. Zhang H, et al. UNC119 is required for G protein trafficking in sensory neurons. Nature neuroscience. 2011;14(7):874–880. doi: 10.1038/nn.2835. [DOI] [PMC free article] [PubMed] [Google Scholar]

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