RESEARCH ARTICLE

The route of the visual receptor rhodopsin along the ciliumAbhishek Chadha1, Stefanie Volland1, Natella V. Baliaouri1, Elaine M. Tran1 and David S. Williams1,2,3,*

ABSTRACTThe photoreceptor outer segment is the most elaborate primary cilium,containing large amounts of rhodopsin (RHO) in disk membranes thatgrow from a connecting cilium. The movement of RHO along theconnecting cilium precedes formation of the disk membranes.However, the route that RHO takes has not been clearly determined;some reports suggest that it follows an intracellular, vesicular routealong the axoneme, possibly as an adaptation for the high load ofdelivery or the morphogenesis of the disk endomembranes. Weaddressed this question by studying RHO in cilia of IMCD3 cells andmouse rod photoreceptors. In IMCD3 cilia, fluorescence recovery afterphotobleaching (FRAP) experiments with fluorescently tagged RHOsupported the idea of RHOmotility in the ciliary plasmamembrane andwas inconsistent with the hypothesis of RHO motility within the lumenof the cilium. In rod photoreceptors, FRAP of RHO–EGFP was alteredby externally applied lectin, supporting the idea of plasmalemmal RHOdynamics. Quantitative immunoelectron microscopy corroborated ourlive-cell conclusions, as RHO was found to be distributed along theplasma membrane of the connecting cilium, with negligible labelingwithin the axoneme. Taken together, the present findings demonstrateRHO trafficking entirely via the ciliary plasma membrane.

This article has an associated First Person interview with the firstauthor of the paper.

KEY WORDS: Photoreceptor, Rhodopsin, Trafficking, Cilium

INTRODUCTIONPrimary cilia have important sensory functions inmany types of cells.Receptors or channels in the cilium function in the detection ofextracellular signals, and their movement into and along the cilium isessential for ciliary function. Cilium dysfunction causes ciliopathies,many of which are syndromic, including disease of the brain, kidneyand retina (Badano et al., 2005; Hildebrandt et al., 2011).The most elaborate primary cilium is the outer segment (OS) of a

photoreceptor cell, consisting of a connecting cilium, which iscomparable to a transition zone (Horst et al., 1990), and a stack ofdisk membranes, containing the light receptor rod photoreceptoropsin (rhodopsin; RHO). The delivery of RHO to the OS is requiredfor the continuous renewal of the disk membranes, which is criticalfor photoreceptor cell function and viability, and requires anextraordinary amount of trafficking along the connecting cilium tothe site of disk membrane morphogenesis (Young, 1967, 1968;

Papermaster et al., 1985; Deretic and Papermaster, 1991). RHOis added to a mouse rod OS at an average of 72 molecules/s(Williams, 2002), with the larger frog rod OS requiring nearly1000 molecules/s (Papermaster et al., 1985). Dislocalization of RHOis a hallmark of many retinal degenerations (Nemet et al., 2015).

Early reports on rod photoreceptor cells proposed that RHO travelsalong the membrane of the connecting cilium after entering from theadjacent periciliary membrane (Papermaster et al., 1985, 1986). Thismodel is consistent with simple primary cilia, whose membranereceptors must reside in the ciliary plasma membrane in order tointeract with extracellular ligands. Indeed, studies on the hedgehogsignaling receptor smoothened (SMO) and the somatostatin receptor3 (SSTR3) (Ye et al., 2013), as well as an SSTR3–RHO chimera (Leeet al., 2018) have demonstrated the movement of these proteins alongthe ciliary plasma membrane of IMCD3 cells.

However, most of the RHO in an OS resides in disk membranesthat are discrete endomembranes, separate from the plasmamembrane, and it has been proposed, based on electron microscopy(EM) images of mouse photoreceptors, that RHO may travel invesicles within the connecting cilium, and these intracellular vesiclesthen directly coalesce into the disk endomembranes (Chuang et al.,2007, 2015; Sung and Chuang, 2010). CryoEM studies identifiedsmall vesicles, which were especially abundant in Bbs4-knockoutmice, a ciliopathy model, between the axoneme and the plasmamembrane of the connecting cilium (Gilliam et al., 2012). Morerecent work from different labs (Burgoyne et al., 2015; Ding et al.,2015; Volland et al., 2015) support an earlier model of diskmorphogenesis (Steinberg et al., 1980) in which the diskendomembranes form as a result of outgrowths of the ciliaryplasma membrane and subsequent closure of the spaces betweenadjacent outgrowths (Arikawa et al., 1992). However, the notion ofRHO vesicles traveling within the connecting cilium to the site ofmembrane outgrowth remains a possibility. Use of an intracellularroute, perhaps in addition to a plasmalemmal one, could be anadaptation in response to the far greater flow of RHO along theconnecting cilium compared with other receptor proteins alongsimple primary cilia. Investigation of such a possibility is importantfor understanding the breadth of sensory cilium adaptations.

In the present study, we have used live mouse IMCD3 andphotoreceptor cells to investigate the localization of RHO as itmoves along the cilium. In particular, we developed an N-terminallytagged RHO that is localized in IMCD3 cell cilia at a comparablelevel to that of untagged RHO and can be used to test forextracellular exposure. We provide evidence that all of the mobileciliary RHO is located in the plasma membrane. Results of the live-cell studies were supported by a quantitative immunoEM study ofRHO localization.

RESULTSRHO in cilia of IMCD3 cellsTo characterize the ciliary mobility of RHO, we first used IMCD3cell cultures. Monolayer cultures of IMCD3 cells were transientlytransfected with a RHO construct, and induced to undergoReceived 3 January 2019; Accepted 2 April 2019

1Departments of Ophthalmology and Neurobiology, Jules Stein Eye Institute, DavidGeffen School of Medicine at UCLA, University of California, Los Angeles, CA90095, USA. 2Molecular Biology Institute, University of California, Los Angeles,CA 90095, USA. 3Brain Research Institute, University of California, Los Angeles,CA 90095, USA.

*Author for correspondence (dswilliams@ucla.edu)

D.S.W., 0000-0002-7758-3932

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© 2019. Published by The Company of Biologists Ltd | Journal of Cell Science (2019) 132, jcs229526. doi:10.1242/jcs.229526

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http://dx.doi.org/10.1242/jcs.233577http://dx.doi.org/10.1242/jcs.233577mailto:dswilliams@ucla.eduhttp://orcid.org/0000-0002-7758-3932

ciliogenesis by serum deprivation with reduced (0.5%) fetal bovineserum (FBS).For the live-cell experiments (below), we generated a RHO

construct that contained a small 15-amino-acid avidity (Avi) tag,added to the extracellular N-terminus (Fig. 1A). Avi-labeledproteins are biotinylated in the ER by co-transfected biotin ligase,and subsequent addition of fluorescently labeled streptavidin to theextracellular space enables labeling of Avi tags on the cell surface(Ye et al., 2013). To compare the ciliary localization of Avi-RHOand non-tagged RHO, we immunolabeled permeabilized IMCD3cells, expressing either Avi-RHO or RHO, with RHO monoclonalAb (mAb) 4D2. Through imaging cilia that were horizontal andessentially lying on the apical plasma membrane, we found that theconcentration of Avi-RHO in the IMCD3 cell cilium wascomparable to that of untagged RHO, as detected by RHO

immunofluorescence, although the apparent concentration of bothRHO and Avi-RHO varied among different cilia, presumablyaccording to differences in expression level (Fig. 1B).

To determine the relative amounts of RHO and Avi-RHO in thecilium versus the adjacent plasma membrane, we calculated acilium-to-plasma membrane fluorescence intensity ratio (CPIR)(Madugula and Lu, 2016). Because the cilium lies on the plasmamembrane, the fluorescence in the apical membrane was subtractedfrom the cilium fluorescence. The resulting net cilium fluorescencewas then divided by the fluorescence in the apical membrane(Fig. 1C). Fig. 1D shows the CPIR for individual cilia expressingAvi-RHO or RHO, with the CPIR values normalized to the medianCPIR for RHO. The CPIR values for the Avi-RHO and RHOcovered a comparable range (Fig. 1D), indicating that the N-terminal Avi tag does not cause major changes in RHO trafficking orlocalization.

The CPIR for RHO appeared lower than that for SSTR3–mKate2(Fig. 1B); this difference could be due to there being less RHO in thecilium or more RHO in the plasma membrane. It has been calculatedthat a cilium, representing a cylindrical shape lying on an apicalmembrane, should have 2.3 times as much signal from a givenmembrane protein as the adjacent membrane if the two regionscontain the same concentration of the membrane protein (Genevaet al., 2017). However, this calculation assumes the apical membraneto be a planar surface, whereas, in polarized epithelial cells, the apicalmembrane consists of an array of microvilli. The apical membraneunderlying the IMCD3 cilia in our images is likely to contain morethan one membrane surface, so that we have not attempted to quantifyan absolute CPIR value for RHO. Since the experiments below arefocused onRHOmovement within the cilium, and the contribution ofthe plasma membrane to fluorescence signals is always subtractedout, the results should not be affected by the CPIR.

Live-cell tests for RHO in the ciliary plasma membrane inIMCD3 cellsWe first determined that the Avi-RHO in the cilium could bedetected by adding extracellular streptavidin–Alexa Fluor 647. Thisindicated that at least some of the Avi-RHOwas in the ciliary plasmamembrane (with its N-terminal Avi tag exposed on the outside of thecell) (Fig. 2A, left panel).

To study the mobility of RHO along the cilium, we performedfluorescence recovery after photobleaching (FRAP) experimentswith Avi-RHO in the distal part of the cilium of IMCD3 cells. Wesubtracted any FRAP contribution of the apical plasma membranefrom the distal cilium fluorescence signal, by subtracting the meanfluorescence of a region adjacent to the cilium, within thephotobleached zone. Bleaching the distal cilium was followed byrecovery of Avi-RHO fluorescence and a simultaneous decline inthe fluorescence in the proximal cilium, indicating movement of theAvi-RHO along the cilium (Fig. 2A,B). The time for half of thefluorescence recovery, t½, was 14±3.3 s (mean±s.e.m.). A comparisonof t½ measurements from cilia with different normalized Avi-RHOCPIR values indicated that CPIR had no significant effect on the t½(R2=8.8%).

The mobility of Avi-RHO in the ciliary plasma membrane shouldbe inhibited by extracellular cross-linking agents. To test whetherAvi-RHO FRAP could be inhibited by crosslinking cell surfaceglycoproteins, we added lectins to the medium prior to FRAPexperimentation. Treatment with 75 µg/ml wheat germ agglutinin(WGA) abrogated ciliary recovery of Avi-RHO (Fig. 2C,D).Concavalin A (ConA) had a similar effect. Each molecule of thishomotetrameric lectin has four binding sites for the glycosylated

Fig. 1. Presence of Avi-RHO andRHO in cilia of IMCD3 cells. (A) Diagram ofAvi-RHO. (B) Cilia of IMCD3 cells containing Avi-RHO or native RHO. RHOwas detected by labeling with RHO mAb4D2. Cilia were identified bycolocalization with SSTR3–mKate2, a robust ciliary marker. In separateexperiments (not shown), we found comparable results when using acetylatedtubulin labeling to identify cilia, indicating that SSTR3–mKate2 does notadversely affect RHO localization. Scale bar: 3 µm. (C) Method for defining thecilium-to-plasma membrane fluorescence intensity ratio (CPIR), adopted fromMadugula and Lu (2016), on a sample image of untagged bovine RHO,immunolabeled with RHO mAb4D2. The CPIR is a ratio of fluorescencespecific to the cilium (Fc) relative to fluorescence from the adjacent membrane(FAM). Fc was defined as the fluorescence measured from the cilium, less thebackground fluorescence of the adjacent plasma membrane (AM), which,because of its proximity (the cilia imaged were essentially lying on the apicalplasma membrane), is included in the measured fluorescence of the cilium.The ends of the depicted line profile could also be used to define AM; however,a circular ROI was selected to obtain a larger sampling for a more accuratemeasurement of AM fluorescence. (D) Comparison of the CPIR for bovine Avi-RHO and unmodified bovine RHO in IMCD3 cell cilia, quantified by using thefluorescence signal from the RHO mAb4D2 plus secondary antibody labelingof fixed cultures. To facilitate comparison of CPIR values between Avi-RHOand RHO, all CPIR values presented in Fig. 1D are normalized to the medianCPIRof RHO. n=33 and 35, respectively, from three separate cultures;P=0.47,Mann–Whitney test. The CPIRs for individual cilia are plotted, and the medianwith interquartile range is indicated for each construct.

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N-terminus of membrane receptors (Edelman and Wang, 1978),such as RHO (Xie et al., 2011). Treatment with 120 µg/ml ConAresulted in near complete inhibition of the recovery of Avi-RHO,following partial cilium photobleaching (Fig. 2E,F), whereastreatment with 60 µg/ml enabled fluorescence recovery, but with asignificantly increased recovery time (Fig. 2G,H). This reductionin recovery is consistent with what has previously beendemonstrated for ciliary membrane receptors, such as SSTR3(Breslow et al., 2013; Ye et al., 2013).Although the Avi-RHO construct enabled us to study RHO in the

plasma membrane, we sought to test whether some additional RHOmight be trafficked within the cilium in intraciliary vesicles. We

generated an Avi-RHO–EGFP-C8 construct (Fig. 3A), which, whenexpressed in IMCD3 cells, allowed for the simultaneouscomparison of the mobility of cell surface RHO (labeled withstreptavidin–Alexa Fluor 647) with that of total RHO (labeled withEGFP) (Fig. 3B, traces in lighter green and red). If RHO traffickedin intraciliary vesicles, its mobility is likely to be distinguishablefrom the mobility of RHO in the ciliary plasma membrane, so thatwewould expect to observe differences in the rate or extent of FRAPfor total RHO and cell surface RHO. Instead, we observed similarFRAP curves for the Alexa Fluor 647 and the EGFP fluorescence(Fig. 3B–D), consistent with the hypothesis that all RHO is movingalong the ciliary plasma membrane.

Fig. 2. FRAP of Avi-RHO in the ciliary plasma membrane in IMCD3 cells. (A) Distal cilium FRAP of Avi-RHO labeled with streptavidin–Alexa Fluor647 in IMCD3 cells. Cells were transfected with Avi-RHO and biotin ligase. Biotinylated Avi tag on the cell surface was then labeled with extracellular streptavidin-Alexa Fluor 647. Pre indicates prior to the photobleach; the time after the bleach is shown in seconds for the other panels. (B) Recovery of distal ciliumfluorescence after photobleaching coincides with a decrease in fluorescence from the proximal cilium after photobleaching. (C,D) FRAP recovery images (C) andproximal and distal fluorescence (D) after treatment with 75 µg/ml WGA. (E,F) FRAP recovery images (E) and proximal and distal fluorescence (F) after treatmentwith 120 µg/ml ConA. (G) FRAP recovery curves from IMCD3 cells, untreated or treated with 60 µg/ml ConA. Data were fitted using a monoexponentialassociation function. (H) FRAP half recovery times of streptavidin–Alexa Fluor 647-labeled Avi-RHO in untreated and ConA (60 µg/ml)-treated IMCD3 cells.n=11 for each, pooled from two WT and three ConA-treated experiments. *P=0.04, Mann–Whitney test. Error bars indicate s.e.m. AU, arbitrary units.

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We repeated this test with Avi-RHO-EGFP-C8, but in thepresence of 75 µg/ml WGA, which blocks the FRAP of Avi-RHO(Fig. 2C,D). If a portion of RHO trafficked within the cilium, thenwe would expect, in this test, to detect some recovery of EGFPfluorescence, but not streptavidin–Alexa Fluor 647 fluorescence,following crosslinking of cell surface proteins. Instead, both surfaceand total pools of RHO were completely blocked by thismanipulation (Fig. 3B,E), indicating that all RHO traffics via thecilium plasma membrane.

Live-cell tests for RHO in the ciliary plasmamembrane of rodphotoreceptor cellsIn a previous study (Trivedi et al., 2012), we developed a method forFRAP imaging of the connecting cilium region of a photoreceptor inRho-EGFP+/−; Rpe65−/− mouse retinal explants, thus measuringthe in situ mobility of RHO in this region. The Rho-EGFP+/− micewere placed on an Rpe65-null background so that the rodphotoreceptors would not be photoexcited, and potentiallyphotodamaged, upon exposure to the EGFP excitation laser;Rpe65 encodes for the isomerase that is required for theregeneration of RHO photopigment (Redmond et al., 1998). Weused RHO–EGFP heterozygotes, since homozygotes undergoretinal degeneration (Chan et al., 2004), most likely because theEGFP was added directly to the C-terminus, resulting ininterference with the C-terminal ciliary-targeting signal of RHO(Wang et al., 2012; Wang and Deretic, 2015). Here, we used FRAPanalysis with Rho-EGFP+/−; Rpe65−/− mouse retinal explants totest for the presence of RHO mobility in the plasma membrane ofthe connecting cilium.As illustrated in Fig. 4A, the bleached region was a spot of 2 µm

in diameter, and the fluorescence recovery was measured from arectangular area that corresponded to the connecting cilium withinthe bleached region. Because the periciliary plasma membrane isclosely adjacent to the ciliary plasma membrane, it was alsoincluded in the FRAP measurements. Extending the bleached

region so that it included different amounts of the outer segment(including even the entire outer segment) had no effect on thefluorescence recovery of the connecting cilium region, indicatingthat recovery came from RHO–EGFP in the inner segment, and notretrograde flow from the outer segment.

Based on the preceding experiments with IMCD3 cilia, wedetermined whether extracellular ConA impeded FRAP ofRHO–EGFP. Treatment with 120 µg/ml ConA appeared tocompletely curtail fluorescence recovery (Fig. 4B), as wasobserved in IMCD3 cilia (Fig. 2E). To test whether thisobservation reflected reduced RHO mobility, rather thantoxicity of the lectin, given the sensitivity of retinal explants toperturbations, we treated retinas with a lower concentration ofConA (60 µg/ml; Fig. 4D,E). As with IMCD3 cells (Fig. 2G,H),the lower concentration of ConA did not completely blockFRAP, but it did increase the half-time of recovery [10.1±1.9 s untreated versus 19.7±3.9 s ConA (mean±s.e.m.); n=11and 13 cells from seven and eight animals for untreated andConA treated groups, respectively; P=0.0154, Mann–WhitneyU-test].

A limitation of this method is that instead of obtaining FRAPmeasurements on just one half of the cilium, as with the IMCD3cells, we can only bleach and measure recovery from a region thatincludes the entire connecting cilium and also the periciliarymembrane. The contribution of the periciliary membrane to theFRAP signal is likely to be minor, given that we measured itsconcentration of RHO to be only one-third of that in the ciliaryplasma membrane (see below). The inhibition of RHO–EGFPFRAP by extracellular ConA is therefore consistent with movementof RHO along the ciliary plasma membrane. However, since FRAPof the entire cilium is dependent upon the entry of RHO into theconnecting cilium, as well as movement along it, the impaired effectof the ConA on the photoreceptor connecting cilium could havebeen due to interactions of the lectin with the adjacent plasmamembrane as well as the ciliary plasma membrane. Nevertheless,

Fig. 3. FRAP of Avi-RHO-EGFP-C8 in the ciliary plasmamembrane in IMCD3 cells. (A) Schematic of the Avi-RHO-EGFP-C8 construct. (B) Example of a FRAP recovery curve fortotal (EGFP) or cell surface (streptavidin–Alexa Fluor 647labeled) RHO in a single cilium. Example traces for are alsoshown for a cell treatedwith 75 µg/mlWGA (darker shaded greenand red). (C,D) The fractional extent of recovery (C) and halfrecovery time (D) were not significantly different (ns) for thesurface and total RHO; n=8 and 9, respectively, from sevenseparate cultures; P=0.49 for (C) and 0.38 for (D), Wilcoxonmatched-pairs signed rank test. Error bars indicate s.e.m.(E) FRAP recovery images after treatment with 75 µg/mlWGA fortotal (EGFP) and cell surface (streptavidin–Alexa Fluor 647labeled) pools of Avi-RHO-EGFP-C8.

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the observation that externally applied ConA did have an effect onRHO–EGFP recovery indicates that RHO is routed via the cellsurface at some stage, refuting the hypothesis that it remains entirelyintracellular, as proposed by Chuang and Sung and coworkers(Chuang et al., 2007, 2015; Sung and Chuang, 2010).

ImmunoEMofRHO in theciliaryandpericiliarymembranesofmouse photoreceptor cellsThe above live-cell experiments have greater physiologicalrelevance than studies of fixed cells, but, as noted, they do notallow sufficient resolution to distinguish between the ciliary plasmamembrane and the periciliary membrane, which are ∼100 nm apartin mouse photoreceptor cells. In order to measure the relative RHOconcentrations in the two membranes, we performed quantitativeimmunoEM.As reported by others (Besharse et al., 1985; Hicks andBarnstable, 1986; Besharse and Wetzel, 1995; Liu et al., 1999;Wolfrum and Schmitt, 2000), RHO is labeled quite poorly in theconnecting cilium by standard immunoEM (Fig. 5A). Therefore, weaggregated immunogold label from many similarly aligned,longitudinal sections that passed through the center of theconnecting cilium of mouse photoreceptors and showed the ciliaryand periciliary plasma membranes in profile. The compilation isshown in Fig. 5B. First, our data support our live-cell analyses,indicating that all of the RHO in the connecting cilium is present in theplasma membrane, with label in the central cilium not significantlyabove background. Second, it shows that the concentration of RHO inthe ciliary plasma membrane is 3-fold greater than that in thepericiliary membrane, which is not significantly different fromthat measured in the other apical inner segment plasmamembrane (Fig. 5C).

DISCUSSIONRouting of RHO to the outer segment of photoreceptors is criticalfor visual function, and dislocalization of RHO to the cilium is ahallmark of many retinal degenerations (Nemet et al., 2015). Here,we sought to determine the route taken by RHO as it traverses theconnecting cilium to the site of disk membrane morphogenesis.From a combination of different approaches, our results indicate thatRHO is present and mobile in the ciliary plasma membrane, and notwithin the lumen of the cilium.

First, we generated an N-terminally tagged RHO for live-cellimaging of RHO in vitro. We attempted to add a SNAP tag to theN-terminal of RHO, as described for SNAP–SMO (Milenkovicet al., 2009), but SNAP–RHO did not reach the plasmamembrane, probably due to misfolding. The structure of theN-terminal of RHO appears to be critical for correct RHO foldingand is sensitive to alterations. A P23H missense mutation in theN-terminal domain of RHO results in misfolded RHO that fails toreach the plasma membrane when heterologously expressed (Liuet al., 1996), and is a relatively common cause of retinitispigmentosa in North America (Dryja et al., 1990). A successfulconstruct was achieved by adding a small 15-amino-acid avidity(Avi) tag to the N-terminus (Fig. 1A). As shown, Avi–RHO canbe labeled by extracellular fluorescent streptavidin (Fig. 2), andit localizes to IMCD3 cilia to a similar extent to untagged RHO(Fig. 1B,D). This construct should be a useful tool for otherlive-cell studies of RHO.

Historically, the paucity of RHO immunolabeling in theconnecting cilium has been a puzzle, given the calculated flowof RHO to the outer segment disk membranes. It has led tosuggestions that RHO might circumvent the connecting cilium en

Fig. 4. RHO in the connecting cilium of rod photoreceptor cells. (A) Schematic of the inner segment, connecting cilium and proximal disks of a photoreceptorcell. FRAP was performed on the region surrounding the connecting cilium. The dashed circle represents photobleached spot, and the dashed rectanglerepresents the region quantified for recovery. (B) Treatment of photoreceptor cells with 120 µg/ml ConA curtailed recovery in the cilium. (C,D) Connecting cilia ofrod photoreceptors in retinal explants fromRho-EGFP+/−;Rpe65−/−mice, imaged by FRAP in the presence (D) or absence (C) of 60 μg/ml ConA. Representativephotoreceptors before photobleach (left), upon photobleach (middle) and after recovery (right) are shown. Note that although the fluorescence signal in theouter segment shows signs of saturation, we confirmed that pre-bleach fluorescence in the region of the connecting cilium, which was monitored for recovery,did not exceed saturation. Scale bar: 1 µm. (E) Representative FRAP traces for RHO–EGFP in control (blue) and 60 µg/ml ConA-treated (red) cells. Data werefitted using monoexponential association functions (black lines).

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route to the disk membranes (Besharse and Wetzel, 1995). Thefirst demonstration of significant RHO immunolabel in the mouseconnecting cilium was by immunoEM of Myo7a-mutant mice.This study showed abnormally high levels of RHO in theconnecting cilium compared with wild-type (WT) mice.Notably, RHO was localized along the plasma membrane (Liuet al., 1999). Subsequently, two quantitative studies (Wolfrum andSchmitt, 2000; Burgoyne et al., 2015), the first using silverenhancement of nanogold immunolabeling, found RHO to beprimarily localized on the plasma membrane and not in the lumenof the WT mouse rod connecting cilium. The quantitative

immunoEM results of the present study also support localizationto the ciliary plasma membrane, with negligible RHO in the lumenof the cilium.

Only ∼11% of IMCD3 cell cilia have a basal part that is closelyassociated with a periciliary membrane that forms a pocket(Molla-Herman et al., 2010). This represents a small fraction ofIMCD3 cell cilia; it contrasts with the cilia of hTERT-RPE1 cells,which all arise from a pocket (Molla-Herman et al., 2010).Nevertheless, in order to avoid any involvement of RHO in thispocket membrane in the FRAP signal, we focused our FRAPstudies (Figs 2 and 3) on the distal cilium of IMCD3 cells. The

Fig. 5. ImmunoEMof RHO in and around the connecting cilium ofmouse photoreceptor cells. (A) Examples of immunoEM images of longitudinal sections,passing through the center of the connecting cilium of mouse photoreceptors, showing gold particle labeling of RHO [e.g. blue arrows in the left panel, indicating agold particle on the ciliary plasma membrane (left) and another on the periciliary plasma membrane (right)]. Scale bar: 200 nm. (B) Compilation of RHOimmunogold particle localization from 22 longitudinal sections from three separate mice. Shaded yellow indicates the different regions that were compared.Blue dots correspond to blue arrows in the top-left panel of A. (C) Bar graph of immunogold particles located within 30 nm of each of the following: the outerand inner plasma membranes of the connecting cilium (outer membrane is outside of the pocket, inner membrane is inside the pocket and thus adjacent to thepericiliary membrane); the center of the connecting cilium (CC center), shown as a dotted line in B; the periciliary plasmamembrane; and the apical inner segment(IS) plasma membrane, on the opposite side from the cilium. *P=0.013, CC center versus IS membrane; *P=0.04, CC center versus periciliary membrane;***P

connecting cilium of a mouse rod photoreceptor cell possesses apartial pocket (Liu et al., 2007), so that the FRAP signal measuredin our experiments with rod connecting cilia would have includedfluorescence from the periciliary membrane (Fig. 4). However,since we found a relative scarcity of RHO in the periciliarymembrane compared with the ciliary plasma membrane, thispericiliary fluorescence is likely to have been minimal. Thepericiliary membrane has been proposed as a docking site forRHO-containing vesicles, prior to entry into the cilium. In frogrod photoreceptors, this membrane region is highly amplified intoa ‘periciliary ridge complex’, most likely to accommodate thehigher delivery rate of OS membrane proteins (Peters et al., 1983);thus it could be a site of RHO accumulation. Our data indicatethat, at least in mouse, as RHO passes through this region, it doesnot accumulate beyond the concentration detected in other parts ofthe distal inner segment plasma membrane (Fig. 5).The mobility of some receptor proteins in cilia has been studied

directly. By using extracellular tags, such as a SNAP tag(Milenkovic et al., 2015), fluorescent streptavidin (Ye et al.,2013) and antibody-linked quantum dots (Lee et al., 2018), SMO,SSTR3 and an SSTR3-RHO chimera have been shown to movealong the plasmamembrane of simple primary cilia. However, RHOhas not been similarly tested previously, and, as mentioned in theIntroduction, RHO could be a special case, given the specializationsof the photoreceptor cilium, which include a high flux of RHOalong the cilium, and the formation of a stack of endomembranedisks that contain the rod visual receptor RHO (Papermaster et al.,1985, 1986). It has been argued that the delivery of RHO to the siteof disk membrane morphogenesis occurs through vesiculartransport directly from the trans-Golgi to the ciliary lumen andalong the center of the connecting cilium (Chuang et al., 2007,2015; Sung and Chuang, 2010). Although physical constraintsimposed by the basal body transition zone would appear to requirean energy-intensive deformation to accommodate the movement ofvesicles into the ciliary lumen (Nachury et al., 2010), this routecould be necessitated by the formation of large amounts ofendomembrane disks. However, the present study shows that theciliary trafficking route of RHO is like that of other ciliary receptors,that is, along the ciliary plasma membrane, thus supportingconservation of a fundamental cilium mechanism.Such conservation is likely to be present in other aspects of RHO

trafficking. Although precise molecular mechanisms have not beenestablished, the specialized photoreceptor cell has been shown to bedependent upon intraflagellar transport (IFT), the fundamentalprocess that is required for all motile flagella/cilia and sensoryprimary cilia, with components including IFT motors, IFT particlecomplexes (Rosenbaum and Witman, 2002; Prevo et al., 2017) andBBS protein complexes (Jin et al., 2010; Liew et al., 2014; Nachury,2018; Ye et al., 2018). Mutations in genes encoding the anterogrademotor, kinesin-2 (e.g. Marszalek et al., 2000; Lopes et al., 2010),IFT20 (Keady et al., 2011), IFT88 (Pazour et al., 2002), IFT140(Crouse et al., 2014), IFT122 (Boubakri et al., 2016), IFT57 (Krockand Perkins, 2008), and the BBS proteins, BBS2 (Nishimura et al.,2004), BBS3 (Zhang et al., 2011), BBS4 (Abd-El-Barr et al., 2007),BBS6/MKKS (Ross et al., 2005) and BBS8 (Hsu et al., 2017; Dilanet al., 2018) have all been shown to affect RHO delivery, outersegment formation and maintenance, and typically result inphotoreceptor degeneration.Taken together, the results of the present study represent the first

rigorous test of the trafficking route for RHO along the cilium. Theydemonstrate that even in the most specialized case of ciliary receptortrafficking, the route conforms to conserved principles.

MATERIALS AND METHODSConstructsThe RHO-EGFP-C8 construct was described previously; it consists of full-length bovine RHO cDNA followed by an EGFP tag, plus a repeat of theterminal eight amino acids of RHO, encoding the sequence ETSQVAPA(Trivedi et al., 2012). The plasmid is available from Addgene, catalog#45399. Native RHO was generated from the RHO-EGFP-C8 construct byremoving the EGFP and C8. For cell surface experiments, we added anAvidity (Avi) tag, MGLNDIFEAQKIEWHE, to the N-terminus of bothRHO and RHO-EGFP-C8. pDisplay-BirA-ER was from Addgene (catalog#20856, deposited by Alice Ting; Howarth et al., 2005). Vector schematicsare displayed using Protter (ETH Zurich).

Tissue cultureIMCD3 cells were obtained from the ATCC (Mannassas, VA; CRL-2123),and were validated by visual inspection of their epithelial morphology inbright-field microscopy. IMCD3 cells were cultured in DMEM/F12medium with 1% penicillin/streptomycin and 10% FBS under standardconditions (all tissue culture reagents were obtained from Thermo FisherScientific, Waltham, MA). Cilia were induced by 24–48 h of treatment with0.5% FBS medium, and visualized by using fluorescently tagged ciliummarkers. Transfections were performed with Lipofectamine 3000 (ThermoFisher Scientific). For cell-surface labeling experiments, cells were co-transfected with an Avitag-containing construct, pDisplay-BirA-ER (biotinligase) and SSTR3-mKate2, and treated with 10 μM biotin (Avidity, Aurora,CO) for 24 h. pDisplay-BirA-ER is expressed in the endoplasmic reticulum(Howarth et al., 2005) and biotinylates Avi-RHO during its synthesis,enabling subsequent labeling with extracellular streptavidin–Alexa Fluor 647(Thermo Fisher Scientific). Prior to imaging, cells were washed three timeswith PBS containing 0.9 mM Ca2+ and 0.5 mMMg2+, treated with 20 μg/mlstreptavidin–Alexa Fluor 647 for 10 min at room temperature, washed threetimes again with PBS containing Ca2+ and Mg2+, and then imaged.

Cilium-to-plasma membrane intensity ratio of RHO fluorescenceThe cilium-to-plasma membrane intensity ratio (CPIR) of the fluorescencefrom immunolabeled RHO was defined as described previously for ciliaryproteins (Madugula and Lu, 2016). CPIR values for both RHO andAvi-RHO (Fig. 1D) were obtained using RHO mAb4D2 labeling ofpermeabilized tissue. First, the fluorescence intensity of the midpoint of aline profile intersecting a representative region of the cilium was measured.Second, we subtracted the fluorescence in a representative patch of plasmamembrane, which, because the analyzed cilia were lying on this membrane,is included in the measured fluorescence of the cilium. The CPIR wasdetermined by dividing the result by the adjacent membrane fluorescence.We normalized CPIR values to the median CPIR of non-tagged RHO.

Fluorescence recovery after photobleachingFRAP experiments were performed at 37°C with IMCD3 cells in 0.5% FBSwith 20 mMHEPES, or with retinal explants from Rho-EGFP+/−; Rpe65−/−

mice of either sex at approximately postnatal day (P)30 in Ames mediumwith 20 mM HEPES. To photobleach an IMCD3 cilium, we targeted acircular spot with a diameter of 2 μm. For photoreceptor cells, we dissectedretinas into approximately six pieces, and typically found photoreceptorsoriented horizontally near the edges of the explant. These horizontallyaligned photoreceptors, which remained as part of the retinal piece (i.e. theywere not isolated cells), were selected for FRAP imaging. Typically, we alsophotobleached a circular spot of 2 μm in diameter (Fig. 4A). All bleachingwas performed using five cycles of bleaching with an Ultraview spinningdisk confocal microscope (PerkinElmer, Waltham, MA). For all FRAPimaging experiments, we confirmed that the pre-bleach fluorescence wassub-saturating. Recovery was measured using custom regions of interest(ROIs) (with ImageJ) including an IMCD3 cilium or the connecting ciliumof a photoreceptor cell (rectangle in Fig. 4A). The FRAP measurementswere corrected for acquisition photobleaching and background signal fromother parts of the cell, and fitted to monoexponential association curves todetermine half recovery times and extents of recovery using GraphPad Prism(GraphPad Software, San Diego, CA).

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MiceFor animal use, we followed NIH guidelines with an approved protocol. Wekept mice on a 12-h-light–12-h-dark cycle with 10–50 lux of fluorescentlighting during the light cycle. For live-cell imaging of RHO–EGFP, weused the RHO–EGFP knock-in line generated by the laboratory of Dr TedWensel (Chan et al., 2004); we placed these mice on an Rpe65-knockoutbackground (Redmond et al., 1998), so that the photoreceptors would notrespond to the excitation laser. RPE65 is a component of the visual cycle andenables regeneration of 11-cis-retinal, which is essential for light activationof RHO. Note that we used heterozygous RHO–EGFP mice, sincehomozygotes for RHO-EGFP undergo retinal degeneration (Chan et al.,2004). For immunoEM studies, we used C57BL/6J mice. In all cases, themicewere∼30 days old (P30), and both genders were used indiscriminately.

AntibodiesMouse RHO mAb 4D2 was generated and validated in previous work(Millipore, Temecula, CA) (Hicks andMolday, 1986). It was used it at 1:100for immunoEM and 1:200 for immunofluorescence.

ImmunoEMThe method for immunoEM studies of the connecting cilium followed thoseused previously (Arikawa and Williams, 1989; Liu et al., 1997). Mice werekilled by cervical dislocation. Eyes were enucleated and fixed overnight at4°C in 4% formaldehyde and 0.2% glutaraldehyde in 0.1 M sodiumcacodylate buffer. Following removal of the anterior segment, the resultingeyecups were dissected along the dorsal ventral axis, dehydrated with anethanol series (30–90%), infiltrated with LR-white resin (fixatives andembedding reagents from ElectronMicroscopic Sciences, Hatfield, PA) andembedded in gelatin capsules. Samples were polymerized at 55°C for 15 h,and ultrathin sections were collected on formvar- and carbon-coated, nickelmesh grids. The grids were treated with 0.1% glycine in 0.1 M phosphatebuffer for 15 min, blocked in 2% BSA in 0.1 M phosphate buffer for20 min, and incubated at 4°C with RHO mAb 4D2 (1:100) overnight. Afterwashing, grids were incubated with secondary antibody conjugated to12-nm colloidal gold (Jackson ImmunoResearch Labs,West Grove, PA) at aconcentration of 1:20. After washing, sections were stained with 5% uranylacetate in ethanol for 5 min, and imaged with a JEM 1200-EX (JEOL USA,Inc., Peabody, MA) transmission electric microscope at 80 kV, usingmagnifications of 30,000 to 60,000. Images of longitudinally alignedphotoreceptor connecting cilia were acquired from sections taken from threedifferent animals.

The RHO labeling density was determined by counting the number ofgold particles within a 60-nm wide ROI, centered on the plasma membranesof the connecting cilia, the periciliary membrane, the inner segment plasmamembrane on the opposite side of the cell and the center of the connectingcilia. The counts were used to determine the number of gold particles perμm, and thus relative density, using Fiji (ImageJ Version 2.0.0; imageprocessing software package, available at https://fiji.sc/) and MicrosoftExcel. Background labeling was based on labeling along random 60-nmwide strips in the adjacent retinal pigment epithelium, and subtracted fromthe raw counts. Additionally, the location of each gold particle was recordedon a schematic of the inner–outer segment.

Experimental design and statistical analysisExperiments were performed on multiple cultures or animals (see figurelegends), and, for all statistical analyses, were repeated until an n value of∼10was achieved with the exception of the experiments in Figs 1 and 4 in which alarger sample size was obtained to better define the distribution of points.After multiple data sets were obtained, statistical comparisons were performedto test the probability of no significant difference between groups. For eachcomparison, data were assessed for normality using the Kolmogorov–Smirnov test. For single comparisons, data were then compared using eitherthe Student’s t-test or the Mann–Whitney U-test, depending on whether thedata were normally distributed. For multiple comparisons, data were analyzedusing either a one-way ANOVA or Kruskal–Wallis test, followed by post-hoccomparisons that were corrected for type II errors, and were chosen based onwhether the data were normally distributed.

AcknowledgementsWe thank Barry Burgess for technical support, Vanda Lopes and Emilie Lemesre fortheir involvement in early stages of the project, and other members of the Williamslab for comments on the manuscript.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsConceptualization: A.C., S.V., D.S.W.; Methodology: A.C., S.V.; Validation: A.C.;Formal analysis: A.C., S.V.; Investigation: A.C., S.V., N.V.B., E.M.T., D.S.W.;Resources: D.S.W.; Writing - original draft: A.C., D.S.W.; Writing - review & editing:A.C., S.V., E.M.T., D.S.W.; Visualization: A.C., D.S.W.; Supervision: A.C., D.S.W.;Project administration: D.S.W.; Funding acquisition: D.S.W.

FundingThe study was supported by National Institutes of Health grants F32EY026318 (toA.C.), R01EY013408 and EY027442 (to D.S.W.) and P30EY0331. Deposited inPMC for release after 12 months.

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