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Centrosomal protein CEP104 (ChlamydomonasFAP256) moves to the ciliary tip during ciliary assembly

Trinadh V. Satish Tammana, Damayanti Tammana, Dennis R. Diener and Joel Rosenbaum*Department of Molecular Cellular and Developmental Biology, Yale University, New Haven, CT 06520, USA

*Author for correspondence (joel.rosenbaum@yale.edu)

Accepted 31 July 2013Journal of Cell Science 126, 5018–5029� 2013. Published by The Company of Biologists Ltddoi: 10.1242/jcs.133439

SummaryThe ciliary tip has been implicated in ciliary assembly and disassembly, and signaling, yet information on its protein composition islimited. Using comparative, quantitative proteomics based on the fact that tip proteins will be approximately twice as concentrated in

half-length compared with full-length flagella, we have identified FAP256 as a tip protein in Chlamydomonas. FAP256 localizes to thetips of both central pair and outer doublet microtubules (MTs) and it remains at the tip during flagellar assembly and disassembly.Similarly, its vertebrate counterpart, CEP104, localizes on the distal ends of both centrioles of nondividing cells until the mother

centriole forms a cilium and then localizes at the tip of the elongating cilium. A null mutant of FAP256 in Chlamydomonas and RNAi invertebrate cells showed that FAP256/CEP104 is required for ciliogenesis in a high percentage of cells. In those cells that could formcilia, there were structural deformities at the ciliary tips.

Key words: CEP104, Centrosomal proteins, Ciliary tip, Ciliogenesis, FAP256, Flagellar assembly/disassembly

IntroductionThe cilia tip is a specialized region in the ciliary apparatus that isinvolved in cilia assembly/disassembly, signaling and sensoryfunction (Johnson and Rosenbaum, 1992; Marshall and Rosenbaum,

2001; Pedersen et al., 2006; Goetz and Anderson, 2010; Gluenzet al., 2010). In comparison to the basal body and axonemal shaft ofthe cilium, little is known about the ultrastructure or composition of

the tip. However, it is an important hub where tubulin subunits areadded or removed at the plus ends of the axonemal MTs duringciliary assembly and disassembly (Rosenbaum and Child, 1967;Johnson and Rosenbaum, 1992; Marshall and Rosenbaum, 2001).

At the tip, tubulin and other axonemal precursors from thecytoplasm are delivered for assembly by intraflagellar transport(IFT), axonemal turnover products are removed for return to the

cytoplasm, IFT is remodeled from long anterograde to shortretrograde trains and the anterograde kinesin motor is inactivatedwith the simultaneous activation of the dynein retrograde motor

(Kozminski et al., 1993; Cole et al., 1998; Pedersen et al., 2006;Pigino et al., 2009). Moreover, recent findings showing recruitmentof the Gli family transcription activators Gli2 and Gli3, and a

regulator of Sonic hedgehog pathway, Sufu, to the tip of theprimary cilium upon hedgehog pathway activation, suggest that theciliary tip acts as an important signaling center during mammalianembryonic development (Haycraft et al., 2005; Liem et al., 2009;

Tukachinsky et al., 2010; Goetz and Anderson, 2010).

In addition to the activities primarily involving IFT, specificproteins probably involved in the assembly and disassembly of

the ciliary MTs, are localized at the tips. Among these are the MTplus-end-binding proteins EB1 and EB3 and the nonprocessivekinesin motor kinesin-13. The latter has been shown to be

involved in MT depolymerization at the plus and minus ends ofMTs (Hunter et al., 2003; Wordeman, 2005) and homologs ofKinesin-13 have been linked to axonemal assembly and

disassembly through overexpression and knockout studies in a

variety of protistans (Blaineau et al., 2007; Chan and Ersfeld,

2010; Dawson et al., 2007). Another member of the kinesin

superfamily, Kif19A, which depolymerizes MTs from the plusends, also localizes at the tips of motile cilia. Mice with

mutations of the Kif19a gene showed elongated cilia in brain,

oviduct and tracheal epithelial cells, suggesting an important role

for this protein in ciliary length regulation (Niwa et al., 2012).Although the Chlamydomonas homolog of kinesin-13 family

proteins, CrKinesin-13 was not seen at the flagellar tip, RNAi-

mediated depletion of this protein resulted in cells with shorter

flagella that failed to regenerate after deflagellation (Piao et al.,2009), suggesting a role for CrKinesin-13 in flagellar assembly.

Similarly, depletion of Kif24, a kinesin-13 family protein that

specifically depolymerizes centriolar MTs, resulted in aberrant

cilia (Kobayashi et al., 2011). Another class of MT bindingproteins, the EB family of MT plus-end-binding proteins, which

help in recruiting other proteins to the plus ends of MTs, were

reported to be localized to the flagellar tips in Chlamydomonas,Giardia and mammalian motile cilia (Pedersen et al., 2003;

Dawson et al., 2007; Schrøder et al., 2011 and Brooks and

Wallingford, 2012). The Chlamydomonas homolog of an EB

family protein, CrEB1, localizes to the tip of the flagellaraxoneme and interacts with IFT protein IFT172, indicating its

possible role in regulating IFT events at the flagellar tip

(Pedersen et al., 2003; Pedersen et al., 2005). EB3 is localized

to the tips of motile cilia in bronchial epithelial cells, anddepletion studies showed that this protein is required for the

formation of primary cilia in cultured human retinal pigment

epithelial (RPE1) cells (Schrøder et al., 2011). Moreover, GFP-

EB3 was recently reported to be localized to the tip in the motilecilia of Xenopus and the primary cilia of RPE1 cells (Brooks and

Wallingford, 2012 and Larsen et al., 2013). Finally, a 97 kDa

5018 Research Article

mailto:joel.rosenbaum@yale.edu

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Tetrahymena protein antigenically related to mammalian

kinetochore proteins was also reported to be present at the

flagellar tip, but the biochemical composition and function of this

protein were not known (Miller et al., 1990).

The fine structure of the ciliary tip was described decades ago

through ultrastructural studies in various organisms including

Chlamydomonas, Tetrahymena and vertebrates (Dentler and

Rosenbaum, 1977; Dentler, 1980; Foliguet and Puchelle, 1986).

In Chlamydomonas and Tetrahymena at the tip, plug-like

structures with thin filaments were shown linking the A-tubule

of each outer doublet to the ciliary membrane and the central pair

MTs terminate in a cap-like structure connected to the ciliary

membrane. These structures were reported to remain intact

during flagellar regeneration and resorption (Dentler and

Rosenbaum, 1977; Dentler, 1980). In mammalian tracheal and

oviduct cilia, several filamentous structures called lateral spokes

connect the A-tubule of the outer doublet to the ciliary

membrane: the central pair and A-tubules terminate in an

electron-dense capping structure, which, in turn, is connected to

the ciliary membrane and claw-like structures called ciliary

crowns project from the surface of the ciliary tip (Foliguet and

Puchelle, 1986). Similarly, electron-dense structures were also

observed at the tip of the flagellar axoneme in other protistans,

such as Trypanosoma brucei, Leishmania major and Crithidia

daenei (Woolley et al., 2006). Among the few known proteins

that localize at the tip, sentan is perhaps the only protein related

to a known structure at the ciliary tip. It localizes to the bridging

structures between the ciliary membrane and peripheral singlet

MTs at the distal tip in tracheal and oviduct motile cilia (Kubo

et al., 2008). Although various large-scale comparative genomics

and proteomics studies have identified many cilia- and

ciliogenesis-related genes (Ostrowski et al., 2002; Li et al.,

2004; Pazour et al., 2005; Ishikawa et al., 2012), the proteins that

make up these apical tip structures remain unknown.

In this report, we describe a method to identify ciliary tip proteins

based on the hypothesis that proteins specifically localized at the

tip will be approximately twice as concentrated in half-length

in comparison to full-length cilia. The relative abundance of

individual proteins in half-length compared with the full-length

flagella was quantified by a mass-spectrometry-based proteomic

analysis, iTRAQ (Ross et al., 2004). Using this method, we

identified a new ciliary tip protein, CEP104/FAP256, which is a

conserved centrosomal protein (Jakobsen et al., 2011) that is

required for cilia assembly in both RPE1 cells and Chlamydomonas.

ResultsIdentification of flagellar tip proteins using iTRAQ

Identification of C. reinhardtii flagellar tip proteins was based on

the hypothesis that proteins present exclusively at the flagellar tip

will be enriched around twofold to threefold when equal

Fig. 1. Flagellar tip and basal body localization of

FAP256 in Chlamydomonas cells with full-length,

assembling and resorbing flagella. (A) Wild-type

Chlamydomonas cells were fixed and stained with antibodies

against acetylated a-tubulin (green, ciliary axonemal

marker) and FAP256 (red). Insets show the tip region of one

of the flagella stained with FAP256 at the apical end.

(B,C) Chlamydomonas cells were fixed at different stages of

flagellar growth and stained for the axoneme and FAP256.

(D,E) Wild-type cells with full-length flagella were induced

to resorb flagella and processed for IFM. Arrowheads

indicate flagellar tips and arrows show basal body staining of

FAP256. Scale bar: 5 mm.

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quantities of short flagella are compared with the full-length

flagella by quantitative mass spectrometry analysis, iTRAQ.

Chlamydomonas cells grown on agar plates lose their flagella in a

few days (Lewin, 1953) and will synchronously and rapidly

regenerate them when transferred to liquid medium. Flagella

were isolated from cells that were allowed to regenerate for 10 or

120 minutes. The isolated short (3.4561.56 mm, n5103) andfull-length (9.861.1 mm n5108) flagella were labeled separatelywith different iTRAQ tags (see the Materials and Methods) and

the relative abundance of individual proteins in the two samples

was obtained from the ratios of the tags as determined by

quantitative mass spectrometry analysis. The ratios for a

particular protein were averaged and proteins were categorized

into groups based on their relative abundance in either short or

full-length flagella. Proteins with an average fold increase of

$1.4 were considered to be enriched in the short flagella. Out ofthe 348 proteins identified at $95% confidence cut-off in theproteomic analysis, 74 proteins were found to be enriched $1.4-fold in short flagella and 59 proteins were found to be enriched

$1.4-fold in full-length flagella (supplementary materialTable S1). Chlamydomonas end binding protein (CrEB1), a

Chlamydomonas MT-plus-end-binding protein that is known to

localize at the tip of the flagellum (Pedersen et al., 2003), was

found to be enriched twofold in the short flagella and served as a

proof of principle that tip proteins should be increased ,twofoldin the half-length flagella. In short flagella, several known

Chlamydomonas proteins involved in flagellar assembly, such as

IFT polypeptides (Marshall et al., 2005) and motor proteins were

enriched. Also enriched in the short flagella were 18

uncharacterized flagellar-associated proteins (FAPs), which

would be of particular interest as candidates for new tip proteins.

Eleven Chlamydomonas flagellar proteins that were increased

twofold to fourfold and have close human homologs were

selected for localization studies in the primary cilia of RPE1

cells. Antibodies against many mammalian proteins are

commercially available, and screening could be done quickly by

immunofluorescence microscopy (IFM) of the RPE1 cells, rather

than making specific antibodies against Chlamydomonas proteins

(see supplementary material Fig. S2). One of the proteins that was

increased ,twofold in half-length flagella, FAP256 (supplementarymaterial Table S2), was a conserved uncharacterized protein

identified in the flagellar proteome (Pazour et al., 2005). Using

FAP256 as query, an NCBI-Blastp search identified only one

human protein of low E-value (2E-76) with 85% coverage. Both

FAP256 and mammalian CEP104 contain a GlyBP domain

(glycine-glutamate-trienylcyclohexylpiperidine binding protein).

Using antibodies against the vertebrate homolog of FAP256

(CEP104) on RPE1 cells, CEP104/FAP256 was localized to the

tip of the primary cilium and at the centrioles (see results below).

FAP256 localizes to the flagellar tip in full-length, growing

and resorbing flagella

To determine the localization of FAP256 in Chlamydomonas

flagella, rabbit polyclonal antibodies were raised against a

recombinant 150 amino acid internal fragment of this protein.

Immunofluorescence analysis using affinity-purified anti-

FAP256 antibodies revealed that FAP256 localizes to the tips

of the flagella in Chlamydomonas cells (Fig. 1A). FAP256 was

also seen at the basal bodies (Fig. 1A). To determine whether

FAP256 localizes at the axonemal tip during flagellar growth and

resorption, IFM was carried out using antibodies against FAP256

and acetylated a-tubulin (axonemal marker). Flagellar tip andbasal body staining of FAP256 was clearly seen in the short

flagellar stubs that appear at the early stages of flagellar assembly

and during later stages of flagellar elongation (Fig. 1B,C).

FAP256 was also localized at the tips in the flagella

Fig. 2. Loss of FAP256 in the Chlamydomonas insertional

mutant Roc22. (A) Chlamydomonas wild-type (CBR34mt+)

and Roc22 mutant cells were fixed and stained with

antibodies against acetylated a-tubulin (green, ciliary

axonemal marker) and FAP256 (red). Loss of FAP256

staining at the flagellar tip (arrowheads) and the basal body

region (arrows) can be seen in Roc22 mutants. Insets show

the tip region of one the flagella stained with FAP256 at the

apical end. Scale bar: 5 mm. (B) Western blots showing theabsence of FAP256 in Roc22 insertional mutant. Equal

amounts of lysates from whole cell and isolated flagella of

Chlamydomonas wild-type (CBR34mt+) and Roc22 mutants

were probed with antibodies against FAP256 and

intermediate chain IC69 (loading control). (C) Roc22

mutants fail to regenerate flagella upon deflagellation.

Roc22 mutants (gray) as well as wild-type cells (CBR34mt+,

black) were deflagellated by pH shock and allowed to

regenerate flagella. Percentage of nonflagellated cells is

shown at various time points (minutes). ,200 cells werecounted for each experiment. Values shown are mean 6 s.d.

from three independent experiments. (D) Flagellar lengths

were measured in Roc22 cells that regenerated flagella at

various time points and mean flagellar lengths are plotted.

Flagella of Roc22 cells were shorter than those of wild-type

cells. ,100 flagella were counted at each time point andP-values at various time points were calculated by unpaired

t-test (30 minutes, P50.0006; 60, 90, 120 and 180 minutes,

P50.0001).

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undergoing resorption (Fig. 1D,E). These results indicate that

FAP256 localizes to the flagellar tips in full-grown flagella and

remains at the tip of the flagellum during flagellar assembly and

disassembly.

FAP256 has a role in flagellar assembly

Ciliary growth occurs by the addition of tubulin subunits at the

plus ends of the axonemal MTs at the ciliary tip (Johnson and

Rosenbaum, 1992; Marshall and Rosenbaum, 2001). The

localization of a conserved centrosomal protein, FAP256, in

this position suggests that it has a role in flagellar assembly. To

test this possibility, a Chlamydomonas insertional mutant for the

FAP256 gene, Roc22, was obtained (Matsuo et al., 2008).

Immunofluorescence showed the absence of FAP256 from the

flagellar tips and the basal bodies of the mutant cells (Fig. 2A).

Western blot analysis also revealed its absence in the null

mutants (Fig. 2B). The ability of cells to form flagella after pH-induced deflagellation was analyzed. About 70% of the mutant

cells (69.468.2%) failed to regenerate their flagella afterdeflagellation (Fig. 2C). Among the Roc22 cells that

regenerated their flagella, the flagella were slightly shorter thanthose of wild-type cells (CBR34 mt+) (Fig. 2D). The flagella ofRoc22 cells were 10.161.40 mm (n5126) compared with12.861.51 mm (n5121) in the wild-type cells. Together, theseresults indicate that FAP256 plays a role in flagellar assembly but

that its absence is not completely inhibitory to flagellar assembly.

FAP256 localizes to the tips of outer-doublet and central-pair MTs

To determine the precise localization of FAP256 at the flagellartip, immunofluorescence and immunogold antibody labeling was

performed on whole mounts of isolated Chlamydomonas flagellaraxonemes splayed apart at their distal tips (see the Materials and

Methods). IFM revealed that FAP256 localizes at the distal endsof the splayed out axonemal MTs (Fig. 3A), indicating thatFAP256 is present at the tips of both outer-doublet and central-

pair MTs (Fig. 3A). To determine this more precisely,immunogold labeling with EM was performed on negatively

stained whole mounts of flagellar axonemes. Clear labeling bythe gold particles was seen at the distal ends of individual outer-doublet and central-pair MTs (Fig. 3B).

Lack of FAP256 results in abnormalities of the flagellartip region

Immunofluorescence and immunogold EM results indicated thatFAP256 is present at the distal ends of the axonemal MTs at theflagellar tip. Mid-sagittal sections of flagella from control and

Roc22 cells were examined with TEM to determine whether theloss of FAP256 altered the ultrastructure of the flagellar tip. As

Fig. 3. FAP256 localizes at the tip of the central-pair and outer-doublet

MTs in splayed axonemes. (A) Chlamydomonas flagellar axonemes were

splayed and processed for IFM using antibodies against acetylated a-tubulin

(green) and FAP256 (red). Arrows indicate the tip localization of FAP256 in

splayed axonemes. Scale bar: 5 mm. (B) Immunogold labeling of FAP256 onsplayed axonemes stained with 2% uranyl acetate. Representative

micrographs showing gold particles at the tip of the splayed axoneme

(arrows). Inset shows enlarged flagellar tip region. Arrows show the gold

particles labeling individual outer-doublet MTs and arrowheads indicate

central-pair MTs of the splayed axoneme.

Fig. 4. The tips of flagella from Roc22 differ dramatically from those of

wild-type cells. (A,B) In the tip of wild-type flagella the central-pair MTs are

capped by an end plate (arrows). (C–E) In Roc22 flagella, the endplate is less

distinct or absent (an endplate might be present in E, arrowheads).

Furthermore, in Roc22, the central pair MTs can be of unequal length

(D, arrowheads), nearly reaching the overlying membrane (C–E, arrowheads).

In wild-type flagella, the outer-doublet MTs end proximal to the central-pair

endplate (* in A,B); in the tip, they are devoid of radial spokes so that the

matrix appears empty. In Roc22, the outer-doublet microtubules can be nearly

as long or longer (* in C,D) than the central pair, and radial spokes connect

them to the central pair almost to the end of the flagellum (.,, in C).

The narrowing of the flagellum seen in the tips of the wild type is not apparent

in Roc22 (compare B and C). The flagellar tips of Roc22 are more blunt than

in the wild type, which often end with a cone-shaped bulge (most distinct

in B).

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previously described (Ringo, 1967; Dentler and Rosenbaum,

1977; Dentler, 1980; Rogowski et al., 2013), the flagellar tip of

wild-type cells appears different from the rest of the axoneme

(Fig. 4A,B). At the tip, the matrix of the axoneme becomes much

less dense, which is due in part to the lack of radial spokes. The

central-pair MTs end together, proximal to the flagellar

membrane and are covered with a bipartite endplate. The outer-

doublet MTs are reduced to the single A-tubules, which terminate

short of the central pair. Because of this early termination of the

doublets, the diameter of the flagellum decreases near the distal

end (Ringo, 1967). Often a small cone of membrane is seen at the

very tip of the flagellum (Fig. 4A,B). These tip specializations

were missing or less apparent in Roc22 flagella (Fig. 4C–E). The

central-pair MTs often appeared to extend all the way to the

membrane and were sometimes of slightly different lengths. The

central pair generally did not appear to be capped with an end

plate. The outer doublets often extended nearly to the overlying

membrane of the tip and radial spokes extending from the A-

tubule of the doublet could be seen almost to the very tip. The

diameter of the distal end did not appear to be normally reduced,

and the entire tip appeared blunt and more rounded than in the

wild type, lacking the membranous cone above the central pair.

These diverse alterations in the structure of the flagellar tip of

Roc22 cells suggest that FAP256 is important in the formation of

the overall configuration of the flagellar tip.

CEP104, the mammalian homolog of FAP256, moves to the

tip during ciliary growth

The mammalian homolog of FAP256, CEP104, is a conserved

protein recently reported in a proteomic analysis of isolated

human centrioles and hence named centrosomal protein of

104 kDa or CEP104 (Jakobsen et al., 2011). Localization studies

using mouse polyclonal antibodies against this protein revealed

that in ciliated RPE1 cells, CEP104 localizes to the tip of the

primary cilium (Fig. 5A). The tip localization of CEP104 was

also seen in the motile cilia of frozen human tracheal sections

(Fig. 5B). In addition to its localization at the ciliary tip in RPE1

cells, CEP104 is present at the distal end of the daughter

centriole, but not on the mother centriole (Fig. 5A). This is

different from the localization seen in interphase non-ciliated

cells, where CEP104 localized to the distal ends of both the

mother and daughter centrioles (Fig. 6A) (see also Jiang et al.,

2012). In non-ciliated cells, the intensity of the CEP104 signal

appeared to be similar at both the centrioles. To determine the

fate of CEP104 on the mother centriole during the formation of

the primary cilium, CEP104 localization was studied in cells

fixed at different time intervals during cilia assembly induced by

serum removal. During cilia assembly, even in the very early

stages of ciliary growth (i.e. ,1 mm long), CEP104 becamerelocalized to the tip of the elongating cilium and remained at the

ciliary tip throughout the growth of the cilium (Fig. 6B–D); the

daughter centriole retained the distal localization of CEP104

throughout ciliary growth.

CEP104 remains at the ciliary tip during ciliary resorption

To determine whether CEP104 is retained at the tip during ciliary

disassembly, cells with fully-grown cilia were induced to

disassemble their cilia by serum addition. Ciliary resorption was

clearly seen after 3 hours of serum addition with most of the cells

having short cilia of ,3 mm. After 24 hours of serum addition,most of the RPE1 cells in the culture had lost their cilia. CEP104

was localized to the tip of the full-length cilium at 0 hours of serum

addition (Fig. 7A) and at various time points during cilia shortening

(Fig. 7B). Finally by 24 hours of serum addition, most of the non-

ciliated interphase cells again contained CEP104 on both the

mother and daughter centrioles (Fig. 7C), as seen at the beginning

of ciliogenesis. These results show that throughout the ciliary

disassembly process, CEP104 remained at the tip of the cilium.

CEP104 localizes to the centrioles during cell division

To determine whether the centriolar localization of CEP104 is

maintained during cell division, actively growing RPE1 cells

were observed during growth by staining with CEP104 and c-tubulin antibodies (marker for centrosomes). As described earlier,

CEP104 is present at the distal end of both mother and daughter

centrioles in the interphase non-ciliated cells (supplementary

material Fig. S1A). In S phase, the centrosome splits and

separates and assembly of new daughter centrioles is initiated.

Throughout S phase, CEP104 appears as two distinct spots in

each centrosome, one spot for each mother and daughter centriole

(supplementary material Fig. S1B,C). CEP104 was present at

both the spindle poles during metaphase and telophase stages of

cell division (supplementary material Fig. S1D,E). These results

indicate that in actively dividing RPE1 cells, CEP104 is recruited

Fig. 5. Tip localization of CEP104 in the primary cilia of

RPE1 cells and motile cilia of human tracheal epithelial

cells. (A) Confluent RPE1 cells were serum starved to

induce cilia formation and stained with antibodies against

Arl13B (ciliary marker, red, bracket), c-tubulin (centrosomal

marker, also in red) and CEP104 (green). Arrows indicate

ciliary tips and arrowheads show daughter centriole

localization of CEP104. (B) Frozen sections of human

trachea were processed for IFM using antibodies against b-

tubulin, to label tracheal cilia (red, bracket) and CEP104

(green). A ridge of cilia on the tracheal epithelium is shown

at higher magnification in the inset. Arrows indicate the

staining of ciliary tips with CEP104. Faint cytosolic staining

of CEP104 can also be seen in these cells. Nuclei are stained

with DAPI (cyan). Scale bars: 5 mm.

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to the daughter centrioles at very early stages of their assembly

and remains associated with the centrosome at all stages of

mitosis prior to interphase and ciliogenesis.

CEP104 is indispensible for ciliogenesis in RPE1 cells

The presence of CEP104 at the distal tip on the growing RPE1

cilia raises the possibility that it plays an important role in

regulating ciliary assembly similar to that observed with its

Chlamydomonas homolog, FAP256. To assess the role of

CEP104 in the formation of primary cilia in mammalian cells,

RNAi using siRNA oligos specific to CEP104 was used to

deplete this protein in RPE1 cells (see the Materials and

Methods). Nontargeting siRNAs were used for control ‘mock’

transfections. Western blots of the whole-cell lysates showed that

the level of CEP104 protein in the siRNA-treated samples was

significantly reduced when compared with the mock-transfected

cells, confirming the specificity of anti-CEP104 antibodies

(Fig. 8D). Immunofluorescence analysis showed that the

amount of CEP104 was highly reduced at both mother and

daughter centrioles in siRNA-treated non-ciliated RPE1 cells

compared with centriolar localization seen in mock-transfectedcells (Fig. 8A). Staining of CEP104 at the tip of the primary

cilium and the daughter centriole was completely lost in thedepleted cells compared with the staining observed at the tip anddaughter centriole in mock-transfected cells (Fig. 8B). The

ability of cells to grow cilia was significantly reduced inCEP104-depleted cultures: only 23.760.4% of the depleted cellsformed cilia, compared with 85.663.2% in mock-transfectedcells (Fig. 8C). Finally, the length of the cilia that did form in the

CEP104-knockdown culture was significantly reduced whencompared with mock-transfected cells. The average length of thecilia in CEP104-depleted cells was ,3.0 mm (3.0460.69 mm;n540) compared with ,5.4 mm (5.3960.8 mm; n553) in mock-transfected cells. Together, these results indicate that ciliogenesisin mammalian cells, is severely compromised by the depletion of

CEP104, similar to Chlamydomonas flagellogenesis seen with theFAP256-null mutant.

DiscussionComparative proteomics of short and full-length flagellareveals tip proteins

In this report, we show that proteins stationed at the ciliary orflagellar tip can be identified by quantitative comparison of the

proteomes of isolated short and full-length flagella. Thereasoning behind the strategy is that proteins present at the tipof short flagella should be about twice as concentrated on a per-protein basis as in full-length flagella. The results presented here

show that one of the proteins enriched in short flagella, FAP256,localizes at the tip during flagellar assembly and disassembly andis involved in flagellar biogenesis in Chlamydomonas. The

mammalian homolog of FAP256, CEP104, also localizes at theciliary tip in primary, as well as motile, cilia and is indispensiblefor ciliogenesis in RPE1 cells.

The present study was carried out using Chlamydomonasbecause it is a well-studied model genetic system where intactflagella of defined lengths can be isolated. The current approach

for identification of proteins enriched in short flagella is amodification of a previous attempt for the identification offlagellar tip proteins by Schneider and colleagues (Schneider

et al., 2008) wherein regenerating flagella were isolated by twodeflagellation steps and protein identification was done bydifference gel electrophoresis. In our case, short and full-lengthflagella from Chlamydomonas cells grown on plates were

isolated by a single deflagellation step; this allowed theisolation of highly purified flagella uncontaminated by othercell organelles (Lewin, 1953; Marshall et al., 2005). The

identification of proteins that were twice as concentrated inshort versus full-length flagella relied on iTRAQ analysis, whichis a sensitive, mass-spectrometry-based procedure used to

compare quantitatively, the amount of individual proteins intwo different samples, such as full-length and short flagella.CrEB1, a known flagellar tip protein in Chlamydomonas

(Pedersen et al., 2003) was enriched twofold in short flagella,indicating that the strategy was successful.

Through iTRAQ analysis, a list of putative new tip proteins

was generated for Chlamydomonas flagella, some of which havehomologs in vertebrates. Mammalian homologs of six of theChlamydomonas putative tip proteins identified in the iTRAQ

procedure (RABGDI, SBP-1, CDPK-1, MSD-3, Serpin and FBB-11), localized to the basal bodies or centrioles, but not to theprimary cilium in RPE1 cells (supplementary material Fig. S2).

Fig. 6. CEP104 from the mother centriole moves to the ciliary tip during

ciliary assembly. Confluent RPE1 cells were serum starved and fixed at

various time intervals during cilia assembly. Cells were processed for IFM

using antibodies against Arl13B (ciliary marker, red), c-tubulin (centrosomal

marker, also in red) and CEP104 (green). (A) CEP104 (arrows) localizes to

both mother and daughter centrioles in non-ciliated RPE1 cells. (B–D) Ciliary

tip and daughter centriole staining of CEP104 (green) during various stages of

ciliary growth. Loss of CEP104 at the mother centriole and tip localization

can be seen from a very early stage of ciliary growth (B). Arrowheads indicate

ciliary tip and arrows show daughter centriole localization of CEP104. Nuclei

are stained with DAPI (cyan). Approximately 200 cells were analyzed at each

time point. Scale bars: 5 mm.

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All of these proteins, however, are present in the Chlamydomonas

flagellar proteome (Pazour et al., 2005). The absence of these

flagellar proteins in primary cilia of RPE1 cells might be due tothe differences in the tip structures between nonmotile primary

cilia and the motile Chlamydomonas flagellum. Indeed, thecentral-pair MTs, which form a part of the distal cap in

Chlamydomonas, are not present in primary cilia. Reflecting

the difference in tip structure, the composition of the ciliary tipsalso differs. For example, Sentan is present at the tip of the

tracheal and oviduct motile cilia but not in the primary cilia ofRPE1 cells (Kubo et al., 2008). Alternatively the inability to

detect these proteins at the tip could be due to (1) the very low

abundance of these proteins in the primary cilia of RPE1 cells;(2) the inaccessibility of certain antigens in the ciliary

compartment as previously documented for IFT20 (Follit et al.,2006); or (3) the methodology used, which enriched proteins that

are concentrated during early stages of ciliary growth.

FAP256 is involved in flagellar assembly and disassembly

One of the putative tip proteins identified in the comparative

iTRAQ analysis of Chlamydomonas flagella and followed indetail in this study, FAP256, is a centriolar protein that can also

be found at the tip of the flagellum. The inability of a majority ofFAP256-null mutants to grow flagella after deflagellation

suggests that this protein is involved in flagellar assembly, butbecause some cells do grow flagella, it seems that FAP256 is not

essential for the assembly process, or that some other protein can

partially assume its function. The occurrence of structuralabnormalities at the tips of null mutants that do form flagella

suggest that FAP256 is involved in flagellar assembly byregulating the stability of the apical structures at the flagellar

tip (described later). In addition to this function at the tip,

FAP256 could also have a role at the basal bodies during initialstages of Chlamydomonas flagella formation as seen with its

vertebrate homolog, CEP104 (described later).

Flagellar assembly occurs by the addition of axonemal

precursors to the distal tips (Witman, 1975; Johnson andRosenbaum, 1992; Marshall and Rosenbaum, 2001; Rosenbaum

and Child, 1967), and resorption likewise appears to occur by MT

disassembly at the tips (Johnson and Porter, 1968; Rosenbaumet al., 1969; Marshall and Rosenbaum, 2001). Generally, cells

resorb their flagella before cell division (Bloodgood, 1974;

Cavalier-Smith, 1974) and flagellar resorption can also be

triggered chemically with isobutyl methylxanthine (IBMX) or

sodium pyrophosphate (Lefebvre et al., 1980; Lefebvre et al.,

1978), yet very little is known about the molecular events thatoccur in the flagellum upon induction of flagellar disassembly.

Recent studies in Chlamydomonas showing increased

methylation and ubiquitylation of flagellar proteins during

flagellar disassembly suggest an active role of these proteinmodifications in the disassembly process (Schneider et al., 2008;

Huang et al., 2009). Furthermore, various genetic and

biochemical data suggest that protein phosphorylation has a

key role in flagellar disassembly (Cao et al., 2009). Comparativephosphoproteome analysis of shortening versus steady-state

flagella revealed that ,89 flagellar proteins were specificallyphosphorylated in the resorbing flagella compared with the

control (Pan et al., 2011). Interestingly, FAP256 was detected in

the phosphoproteome of disassembling flagella but not in thecontrol (Pan et al., 2011), indicating that this protein is

specifically phosphorylated during flagellar shortening. The

presence of FAP256 at the flagellar tip at steady state and

during resorption, shown in this study, together with itsphosphorylation during flagellar resorption, suggests that this

protein plays an important role in maintaining flagellar length

and its role might be altered or regulated by phosphorylation

during flagellar resorption.

Loss of FAP256 results in structural abnormalities at theflagellar tip

At the flagellar tip, the central pair normally end just short ofthe flagellar membrane, and are attached to it by a capping

structure, whereas the outer doublets become singlets and end

,100–200 nm from the flagellar tip (Ringo, 1967; Dentler andRosenbaum, 1977). The A-tubule of each outer doublet hasa carrot-shaped plug in it, and each plug has a thin filament

connecting it to the flagellar membrane. These structures

are present in the A-tubule throughout both assembly

and disassembly of the flagellum (Dentler and Rosenbaum,

1977; Dentler, 1980). Although these MT-end-binding structureshave been known for over 35 years, nothing is known about

their composition or function. Our microscopic analysis with

Fig. 7. Localization of CEP104 at the tip during ciliary

disassembly. Cells with fully grown cilia were induced for cilia

disassembly by adding the serum and fixing at various time intervals

after serum addition. Cells were stained with Arl13B (ciliary marker,

red), c-tubulin (centrosomal marker, also in red) and CEP104

(green). (A,B) Tip localization of CEP104 (arrowheads) can be seen

in full-length and disassembling cilia. (C) By 24 hours of serum

addition, most cells in the culture had lost their cilia and CEP104

staining was seen on both mother and daughter centrioles. Arrows

indicate centriole localization of CEP104. Nuclei are stained with

DAPI (cyan). Approximately 200 cells were analyzed at each time

point. Scale bars: 5 mm.

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anti-FAP256 antibodies and EM localizations with gold-labeled

antibodies showed that in addition to its presence at the central

pair, FAP256 also localizes to the tips of individual outer

doublets. Because the outer doublets at the tip are reduced to just

the A-tubule of the doublet, it is with this part of the doublet that

FAP256 is associated.

About 70% of the FAP256-null cells do not grow flagella, and

the flagella that are assembled are blunt at their tips rather than

tapered as seen with the wild-type cells. The bluntness of the tips

in FAP256 mutant flagella is due to the fact that the outer

doublets are of the same length as the central pair MTs. This

structural abnormality could arise because the outer doublet MTs

grew to the tip of the flagellum instead of ending proximal to the

central pair or because the central pair MTs were unstable and

depolymerized at the tip region. The overall length of the

flagellum in the mutant cells is shorter than in wild-type cells,

thereby ruling against the possibility of the overgrowth of the

outer doublet MTs. A third explanation is that in the absence of

FAP256, the flagellum fails to initiate tip formation, never

forming the specialized trademarks of the wild-type tip. In

support of this third mechanism, the loss of FAP256 had

pleotropic effects on the overall structure of the flagellar tip,

including loss of the end plate, extension of outer-doublet and

central-pair microtubules to the flagellar membrane, attachment

of radial spokes to the outer doublet MTs in the tip and loss of the

membranous cone at the tip of the flagellum. These diverse

effects of the FAP256-null mutants suggest that FAP256 might

be required for the structural integrity of the specialized

structures seen at the flagellar tip and the loss of this protein

affected the overall formation of these apical structures.

CEP104 regulates ciliary assembly by interacting with

proteins that suppress cilia formation

The vertebrate homolog of the newly identified tip protein,

CEP104 was previously reported as a SXIP-motif-containing

EB1-interacting protein. CEP104 localizes on both mother and

daughter centrioles in non-ciliated RPE1 cells, but only on the

daughter centriole in ciliated cells (Jakobsen et al., 2011; Jiang

Fig. 8. Depletion of CEP104 in RPE1 cells impairs cilia

formation. (A,B) Both non-targeting (mock) and CEP104-

siRNA-treated samples were processed for IFM using antibodies

against Arl13B (ciliary marker, red) and CEP104 (green).

Depletion of CEP104 at the centrosomal region (arrows) in non-

ciliated cells (A) and ciliary tip (arrowheads) in ciliated cells

(B) was seen in CEP104-siRNA-treated cells when compared

with mock-transfected cells. Nuclei were visualized by DAPI

staining (cyan). Scale bars: 5 mm. (C) Percentage of full-lengthcilia, no cilia and short cilia bearing cells in mock (black) and

CEP104-siRNA-treated (gray) populations were plotted from

immunofluorescence images. ,200 cells were counted for eachexperiment. Values shown are mean 6 s.d. from three

independent experiments. (D) Western blots showing the

depletion of CEP104 in RPE1 cells treated with CEP104 siRNA.

Equal amount of cell lysates from mock treated, siRNA-treated

and untreated RPE1 cells serum starved for 24 hours were

probed with mouse anti-CEP104 antibodies. Antibodies against

a-tubulin were used as a loading control.

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et al., 2012). In addition, we report here that CEP104 moves fromthe mother centriole to the tip of the primary cilium during

ciliogenesis. Consistent with previous results, our data showedthat siRNA-mediated depletion of this protein significantlyreduced the ability of RPE1 cells to form primary cilia (Jianget al., 2012).

During ciliogenesis, the mother centriole functions as a basalbody by nucleating cilia when the cells are in the G0 stage of thecell cycle, and, upon re-entry to the division phase, cells resorb

their cilia allowing the centrioles to form spindle poles (Sorokin,1968, Rieder et al., 1979, Wheatley et al., 1996). The molecularbasis for the conversion of the mother centriole into the ciliary

basal body is not understood. Studies focused on the transitionbetween the mother centriole and the ciliary basal body identifiedan important centriolar protein complex CP110–CEP97 thatinhibits ciliary assembly. Loss of these proteins promotes cilia

formation (Spektor et al., 2007). In ciliated cells, these proteinswere seen only on the daughter centriole, not on the mothercentriole or in the cilium proper (Spektor et al., 2007; Kuhns

et al., 2013). The mechanism by which CP110 and CEP97 areremoved from the mother centriole during ciliogenesis remainsunknown (Bettencourt-Dias and Carvalho-Santos, 2008). Based

on previous reports and the results presented in this study, weenvisage that CEP104 might be involved in the removal of theCP110–CEP97 inhibitory complex from the mother centriole.Recent studies from Jiang and co-workers, showed that CEP104

interacts with CP110 and CEP97 and recruits CEP97 to the plusends of cytosolic MTs in HEK293T cells, perhaps through EB1binding (Jiang et al., 2012). Likewise, the interaction of CEP104

with CP110 and CEP97 might be holding these proteins at themother centriole before the onset of ciliogenesis. At the veryinitial stages of ciliogenesis, as shown in this report, CEP104

moves from the mother centriole to the tip of the cilium. UnlikeCEP104, neither CP110 nor CEP97 was found in the cilium(Spektor et al., 2007) or at the ciliary tip (our data, not shown).

Dissolution of a CEP104–CP110–CEP97 complex might releaseCP110 and CEP97 from the mother centriole, thereby allowingthe centriole to participate in ciliogenesis. Once the complex isbroken, CEP104 would be free to assume its function at the tip of

the nascent cilium. Furthermore, recent reports showing the roleof centriolar kinases MARK4 and Ttbk2 in the removal of CP110and CEP97 from the mother centriole, and in turn in ciliogenesis

(Kuhns et al., 2013; Goetz et al., 2012), raise the possibility that aphosphorylation event might be involved in the removal ofCP110 and CEP97 from the mother centriole. Neither MARK4

nor Ttbk2 were shown to directly interact with the proteins of thecilia suppression pathway (CP110–CEP97) or CEP104. MARK4interacts and phosphorylates mother centriolar protein ODF2

(Kuhns et al., 2013) and Ttbk2 was recently identified as an SXIPcontaining EB1 binding protein (Jiang et al., 2012) suggesting theoccurrence of a complex cascade of events involving MT bindingproteins and other centriolar proteins at the mother centriole

during initial stages of ciliogenesis. Hence, further studies areneeded to elucidate how CEP104, CP110 and CEP97 interact atthe onset of ciliogenesis.

Through proteomic analysis of Chlamydomonas flagella,FAP256 was identified as a protein required for ciliogenesis inChlamydomonas and vertebrate cells. It is essential for the

formation of specialized structures at the flagellar tip inChlamydomonas. The mammalian homolog of this proteinCEP104 interacts with centriolar proteins that inhibit

ciliogenesis, detaches from these proteins at the onset of ciliaformation and moves to the tip of the nascent cilium. These

results suggest that CEP104/FAP256 has dual functions: one atthe centriole to initiate ciliogenesis and one at the ciliary tip,where it is required for the formation or stability of thespecializations characteristic of the tip. Because of its integral

role in cilia formation, CEP104 can be viewed as a new candidateciliopathy protein.

Materials and MethodsCultures

Immortalized human retinal pigment epithelial cells (RPE1-hTERT) cells werecultured in DMEM and F12 nutrient mix 1:1 (Gibco) supplemented with 10% heat-inactivated FBS (Gibco) and 1% penicillin-streptomycin (Penstrep, Sigma).Cultures were grown at 37 C̊, with 5% CO2 and passaged upon confluency (4–5days) by trypsinization (0.25% Trypsin-EDTA, Gibco). Cultures up to 20 passageswere used for the experiments.

Wild-type C. reinhardtii cells (cc125mt+) were obtained from ChlamydomonasGenetics Center (Durham, NC). Roc22 insertional mutants and CBR34mt+cultures were obtained from Masahiro Ishiura, Nagoya University, Japan. All thestrains except Roc22 insertional mutants were maintained in solid mediasupplemented with 1.5% agar. Roc22 cells were grown in similar cultureconditions as described above supplemented with 10 mg/ml hygromycin. Forexperimentation, cultures were either grown in liquid minimal medium M1 or Trisacetate phosphate (TAP) medium (Harris, 1989) at 18 C̊ with 12 hour light anddark cycles and constant aeration.

Cilia assembly and disassembly

For the induction of primary cilia in RPE1 cultures, 36105 cells/well were seededon sterile coverslips in six-well culture plates (BD Falcon) and allowed to grow inDMEM+F12 nutrient mix supplemented with 10% FBS to ,80% confluency. Thecells were then subjected to serum starvation in DMEM+F12 nutrient mixcontaining 0.5% FBS for 24 hours. For ciliary assembly studies, cells were fixed atvarious time intervals to 24 hours after serum removal and processed for IFM. Forciliary disassembly, RPE1 cells growing on coverslips at ,50% confluency weresubjected to serum starvation for 24 hours for cilia formation. Cells with fullygrown cilia were then induced for ciliary disassembly by adding the serum back,fixed at various time intervals after serum addition and processed for IFM. Thegrowth of full-length cilia after 24 hours of serum starvation was checked bystaining the cells on duplicate coverslips for the ciliary membrane marker Arl13B.

siRNA transfection

For the transfection of siRNAs in RPE1 cells, ,36105 cells were plated on sterilecoverslips on six-well plates 1 day before the transfection in DMEM+F12 nutrientmix containing 10% FBS without antibiotics. Next day, the growth medium wasreplaced with fresh medium without antibiotics 1 hour before transfection. Thecells were then transfected using 50 nM CEP104 siRNA (ON-TARGET plusSMART pool, Dharmacon) using 5 ml per well Lipofectamine2000 (Invitrogen)or Dharmafectduo (Dharmacon) transfection reagents as described in themanufacturer’s protocol. For control transfections, 50 nM of nontargetingsiRNA pool were transfected in RPE1 cells in similar conditions. The next day,the culture medium was replaced with fresh medium containing 10% FBS and thecells were allowed to grow at 37 C̊ till confluency. Upon reaching confluency, thecells were subjected to serum starvation in DMEM+F12 nutrient mix containing0.5% FBS for 24–48 hours. Cilia formation and CEP104 depletion was assessed byIFM and western blotting using mouse anti-CEP104 antibodies.

Antibodies and affinity purification

For the generation of rabbit anti-FAP256 antiserum, a synthetic gene fragmentof 480 bp from the internal region of C. reinhardtii FAP256 gene(CTGGGCAAG………AGCGGCGGT) without a stop codon (synthesized atGenscript, NJ) was cloned into NdeI and HindIII sites of E. coli expression vectorpET21A (Novagen). The cloned fragment was expressed with a C-terminal Hexa-Histag in pET21A in E.coli BL21 (DE3) Gold strain (Agilent Technologies). Purificationof recombinant protein was carried out as described (Tammana et al., 2008) withminor modifications. FAP256-transformed E.coli cells were grown at 37 C̊ up to 0.5OD and induced with 0.5 mM IPTG. Cells were then harvested after 5 hours ofgrowth and resuspended in lysis buffer (50 mM sodium phosphate, 300 mM sodiumchloride, 1 mg/ml lysozyme and protease inhibitors) for 30 minutes at 4 C̊ and lysedby sonication. Lysates were further clarified by centrifugation at 14,000 rpm at 4 C̊for 30 minutes. The pellet was resuspended in solubilization and wash buffer (8 Murea, 0.5 M sodium chloride, 20 mM sodium phosphate, 20 mM imidazole) andapplied to Ni-NTA agarose affinity columns. The columns were washed with washbuffer thoroughly and eluted by increasing imidazole concentration to 500 mM. Finalelutes were then concentrated, subjected to buffer exchange using centrifugal filters

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of 10,000 MW (Amicon Ultra, Millipore) and analyzed by SDS PAGE (12%resolving) for purity. Purified recombinant FAP256 protein fragment was used forraising antiserum in rabbits (Pocono Rabbit Farm and Laboratory). Finally anti-FAP256 antibodies were purified from the test bleeds by the blot-strip method usingthe purified recombinant FAP256 protein fragment. The specificity of the antibodieswas tested by western blotting and immunofluorescence analysis. The sources anddilutions of all other primary antibodies used in mammalian and Chlamydomonasimmunofluorescence analysis are listed in supplementary material Table S3.Secondary antibodies for western blotting analysis, alkaline-phosphatase-conjugated goat anti-rabbit IgG antibodies and alkaline-phosphatase-conjugatedgoat anti-mouse IgG antibodies (Invitrogen) were used at 1:10,000 dilution.

Western blotting

For western blotting, flagella were isolated from Chlamydomonas cc125mt+,Roc22mt+ and CBR34mt+ cells growing in 8L TAP liquid medium according to(Cole et al., 1998). For preparing RPE1 cell lysates, cells growing on tissue cultureflasks (BD Labware) were scraped using a cell scraper and washed twice in coldPBS containing protease inhibitors. Lysates from the cell pellets or isolatedflagella were prepared by resuspending and boiling them for 5 minutes in 16SDSsample buffer. Samples were resolved on SDS-PAGE (12% resolving) and electro-blotted onto nitrocellulose membrane. Rabbit anti-FAP256 antibodies and mouseanti-CEP104 antiserum were used for detecting Chlamydomonas FAP256 andmammalian CEP104, respectively. Blots were developed by using colorimetric orchemiluminescent methods.

Immunofluorescence microscopy

For immunofluorescence analysis, RPE1 cells growing on coverslips were washedthree times in 16PBS and fixed using cold methanol and acetone 1:1 mix for5 minutes at 220 C̊. The coverslips were air dried and rehydrated in PBS and thenblocked for 1 hour at room temperature in blocking solution (0.5% BSA,10% normal goat serum and cold water fish skin gelatin in 16PBS).Immunofluorescence analysis on Chlamydomonas strains was done essentially asdescribed previously (Pedersen et al., 2003). Primary antibodies against specificproteins and markers for cilia (mouse anti-acetylated a-tubulin and rabbit Arl13bantibodies) and basal bodies (rabbit c-tubulin antibodies) were used at dilutionsgiven in supplementary material Table S3. Unless otherwise noted, secondaryantibodies for IFM, Alexa-Fluor-488-conjugated goat anti-mouse antibodies andAlexa-Fluor-594-conjugated goat anti-rabbit antibodies (Molecular Probes) wereused at 1:1000 dilution. Finally, the coverslips were mounted on clean glass slidesusing Prolong gold anti-fade mounting medium containing DAPI (Invitrogen) andthe images were captured on a Zeiss LSM510 META confocal microscope using a1006 1.4 NA (oil) plan apochromate lens.

Chlamydomonas flagellar growth and resorption

For flagellar growth, Chlamydomonas cells grown on TAP-agar plates werescraped into 10 mM HEPES and allowed to regenerate flagella. Cells were fixed atvarious time intervals and processed for IFM. Flagellar resorption was induced incells with full-length flagella by treating cells with 20 mM sodium pyrophosphate(Lefebvre et al., 1978); cells were fixed at various time intervals for IFM.

Chlamydomonas flagellar axoneme splaying

Axoneme splaying for IFM was carried out as described (Johnson, 1998) withminor modifications. Briefly, Chlamydomonas cells adhered on the coverslipswere treated with HMDEK buffer (10 mM HEPES, pH 7.2, 5 mM MgSO4, 1 mMDTT, 0.5 mM EDTA and 25 mM KCl) containing 0.01% NP40 and 1 mM ATPfor 3 minutes, fixed and permeabilized with ice-cold methanol. The slides werethen processed for IFM using antibodies against acetylated a-tubulin and FAP256as described above.

Flagellar and ciliary length measurements

For flagellar length measurements from different Chlamydomonas strains,cells were fixed with 1% glutaraldehyde in M1 medium and were attachedto coverslips coated with 0.1% poly-L-lysine (Sigma-Aldrich). Imageswere captured with a Nikon microscope (Eclipse TE2000; Nikon) equipped withplain Apo 406 objective lens and a camera (Cascade 512B; Photometrics). Thelengths of at least 100 flagella were measured for each time point usingthe MetaMorph software package (MDS Analytical Technologies). For measuringthe lengths of primary cilia in siRNA transfection experiments, control and siRNA-treated cells (after 24 hour serum starvation) were subjected to staining with mouseanti-acetylated a-tubulin antibodies and rabbit anti-c-tubulin antibodies for labelingof ciliary axoneme and basal bodies, respectively. Images were captured using theabove microscope at 1006magnification and length measurements were made usingMetaMorph software as described above.

Immunogold labeling of whole mounts

Whole mounts of axonemes were prepared as described by Johnson andRosenbaum (Johnson and Rosenbaum, 1992). Concentrated drops of cells were

allowed to settle on formvar-coated nickel grids for 10 minutes. Flagellaraxonemes were splayed by inverting the grids onto HMDEK buffer containing0.01% NP40 and 1 mM ATP for 3–5 minutes. The grids were then treatedsequentially with drops of fixative containing 2% PFA and 0.5% glutaraldehyde(15 minutes), PBS (365 minutes), permeabilizing solution (PBS and 0.5%Triton X-100, 265 minutes), wash buffer (PBS and 0.05% Tween20,365 minutes) and blocking buffer (wash buffer with 0.5% BSA, 10% normalgoat serum and 0.02% NaN3, 30 minutes). After blocking, the grids wereinverted onto drops of affinity purified anti-FAP256 antibodies (1:50) inblocking buffer for 3–4 hours at room temperature, followed by washing withwash buffer (465 minutes). The grids were then incubated with 12 nm goldconjugated to goat anti-rabbit IgG (Jackson Immunologicals, PA) diluted to1:100 in blocking buffer for 2 hours. Grids were then rinsed in wash buffer(465 minutes), PBS (265 minutes), water (265 minutes) and then negativelystained with 2% aqueous uranyl acetate for 1 minute. Finally, uranyl acetate wasrinsed from the grids with drops of water.

Transmission EM of flagellar tips

EM of flagellar tips was carried out as described (Pigino et al., 2009) with minormodifications. CBR34 (wild-type) and Roc22 cells were allowed to adhere onpoly-L-lysine-coated coverslips for 5 minutes. The cells were fixed on thecoverslips with 2.5% glutaraldehyde in MI medium for 30 minutes at roomtemperature. Tannic acid was added to 0.1% and fixation continued for another30 minutes. The coverslips were then washed in 10 mM HEPES pH 7.4 and fixedfor 30 minutes at 4 C̊ in 1% osmium tetroxide in 50 mM HEPES. After washing inwater, the cells were stained with 2% uranyl acetate for 1 hour at roomtemperature. The samples were dehydrated through ethanol and propylene oxideand embedded in Epon 812. Following polymerization of the resin, the coverslipswere dissolved from the Epon with hydrofluoric acid. Desired areas of the resinwere cut out and glued to blocks of Epon with 5 minute two-part epoxy (Devcon).Sections were stained with lead citrate (Reynolds, 1963) and uranyl acetate, andwere viewed with a JEOL 1230 microscope equipped with a Hamamatsu Orca HRdigital camera.

Chlamydomonas flagella isolation for iTRAQ analysis

For iTRAQ analysis, we used a method to isolate short and full-length flagellafrom C. reinhardtii (cc125mt+) cells growing on air-agar interphase (Lewin, 1953;Marshall et al., 2005) to avoid multiple steps of deflagellation at acidic pH.Chlamydomonas cells grown on agar plates lose their flagella in a few days andwill synchronously and rapidly regenerate them when transferred to liquidmedium. Briefly, cells growing on solid medium containing TAP with 1.5% agarfor 5–6 days were gently scraped into an ice cold solution of 10 mM HEPES andallowed to regenerate flagella at 18 C̊ with constant aeration. Following this,flagella were isolated from aliquots of cells by pH shock at 10 minutes and120 minutes when the flagella were ,3.4561.56 mm (n5103) and 9.861.1 mm(n5108) long, respectively. Isolation of flagella was carried out essentially asdescribed previously (Cole et al., 1998; Witman et al., 1972).

iTRAQ analysis

iTRAQ (Isobaric tag for relative and absolute quantification, Applied Biosystems)is a mass-spectrometry-based protein-profiling method that allows simultaneousidentification and comparative quantification of peptides by covalent labeling withreporter ions (Ross et al., 2004). iTRAQ analysis for comparing the relativeabundance of individual proteins in short and full-length flagella isolated fromC. reinhardtii (CC125mt+) cells was done at the Center for Mass Spectrometry andProteomics, University of Minnesota. Briefly, the total protein content in theflagellar preparations was estimated, and equal quantities (,100 mg) of eachsample were subjected to protein reduction and cysteine blocking, followed bytrypsin digestion resulting in the generation of large number of peptides. Thesepeptides were then labeled with iTRAQ reagents containing signature ions 114,115, 116 and 117 separately: 114 and 115 for peptides generated from shortflagella; 116 and 117 for peptides generated from long flagella. This labelingstrategy ensures that the samples will be replicated and internal comparison can bedone using 114:115 and 116:117 ratios. The samples were then pooled, followedby the removal of trypsin and hydrolyzed iTRAQ reagents on a mixed bed resin(MCX) cartridge. The samples were then subjected to strong cation exchangechromatography for fractionation followed by LC/MS/MS analysis. Massspectrometry data was collected on 4800 TOF/TOF MALDI analyzer. Proteinidentification was performed using the C. reinhardtii NCBI Refseq gene databaseand Augustus version 9.0 database searches. The relative abundance of signatureions among the two samples determines the abundance of a particular proteinpresent in the sample with respect to the other sample. Data analysis was carriedout using Protein Pilot software (Applied Biosystems) and peptides were filtered at$95% confidence cut-off. To calculate the relative abundance of individualproteins, ratios from different tags were averaged. Proteins were categorized intogroups based on their relative abundance in short flagella or full-length flagella.Proteins with average fold abundance of $1.4 were considered to be enriched ineach category.

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AcknowledgementsWe are grateful to Dr Masahiro Ishiura, Nagoya University, Japan forsharing Chlamydomonas mutant strains and Dr Caspary Tamara,Emory University, USA for providing anti-Arl13b antibodies.

Author contributionsThe project was conceived and designed by J.L.R.; experiments weredesigned by T.V.S.T and J.L.R.; experiments were carried out andanalyzed by T.V.S.T., D.T. and D.R.D.; manuscript was prepared byT.V.S.T., D.R.D. and J.L.R.

FundingThis work was supported by the National Institutes of Health [grantnumber GM014642 to J.L.R.]. Deposited in PMC for release after 12months.

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.133439/-/DC1

ReferencesBettencourt-Dias, M. and Carvalho-Santos, Z. (2008). Double life of centrioles:

CP110 in the spotlight. Trends Cell Biol. 18, 8-11.

Blaineau, C., Tessier, M., Dubessay, P., Tasse, L., Crobu, L., Pagès, M. and Bastien,P. (2007). A novel microtubule-depolymerizing kinesin involved in length control ofa eukaryotic flagellum. Curr. Biol. 17, 778-782.

Bloodgood, R. A. (1974). Resorption of organelles containing microtubules. Cytobios 9,142-161.

Brooks, E. R. and Wallingford, J. B. (2012). Control of vertebrate intraflagellartransport by the planar cell polarity effector Fuz. J. Cell Biol. 198, 37-45.

Cao, M., Li, G. and Pan, J. (2009). Regulation of cilia assembly, disassembly, andlength by protein phosphorylation. Methods Cell Biol. 94, 333-346.

Cavalier-Smith, T. (1974). Basal body and flagellar development during the vegetativecell cycle and the sexual cycle of Chlamydomonas reinhardii. J. Cell Sci. 16, 529-556.

Chan, K. Y. and Ersfeld, K. (2010). The role of the Kinesin-13 family protein TbKif13-2 in flagellar length control of Trypanosoma brucei. Mol. Biochem. Parasitol. 174,137-140.

Cole, D. G., Diener, D. R., Himelblau, A. L., Beech, P. L., Fuster, J. C. and

Rosenbaum, J. L. (1998). Chlamydomonas kinesin-II-dependent intraflagellartransport (IFT): IFT particles contain proteins required for ciliary assembly inCaenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993-1008.

Dawson, S. C., Sagolla, M. S., Mancuso, J. J., Woessner, D. J., House, S. A., Fritz-Laylin, L. and Cande, W. Z. (2007). Kinesin-13 regulates flagellar, interphase, andmitotic microtubule dynamics in Giardia intestinalis. Eukaryot. Cell 6, 2354-2364.

Dentler, W. L. (1980). Structures linking the tips of ciliary and flagellar microtubules tothe membrane. J. Cell Sci. 42, 207-220.

Dentler, W. L. and Rosenbaum, J. L. (1977). Flagellar elongation and shortening inChlamydomonas. III. structures attached to the tips of flagellar microtubules and theirrelationship to the directionality of flagellar microtubule assembly. J. Cell Biol. 74,747-759.

Foliguet, B. and Puchelle, E. (1986). Apical structure of human respiratory cilia. Bull.Eur. Physiopathol. Respir. 22, 43-47.

Follit, J. A., Tuft, R. A., Fogarty, K. E. and Pazour, G. J. (2006). The intraflagellartransport protein IFT20 is associated with the Golgi complex and is required for ciliaassembly. Mol. Biol. Cell 17, 3781-3792.

Gluenz, E., Höög, J. L., Smith, A. E., Dawe, H. R., Shaw, M. K. and Gull, K. (2010).Beyond 9+0: noncanonical axoneme structures characterize sensory cilia from protiststo humans. FASEB J. 24, 3117-3121.

Goetz, S. C. and Anderson, K. V. (2010). The primary cilium: a signalling centreduring vertebrate development. Nat. Rev. Genet. 11, 331-344.

Goetz, S. C., Liem, K. F., Jr and Anderson, K. V. (2012). The spinocerebellar ataxia-associated gene Tau tubulin kinase 2 controls the initiation of ciliogenesis. Cell 151,847-858.

Harris, E. H. (1989). The Chlamydomonas Sourcebook. A Comprehensive Guide toBiology and Laboratory Use, xiv, 780 pp. San Diego, CA: Academic Press.

Haycraft, C. J., Banizs, B., Aydin-Son, Y., Zhang, Q., Michaud, E. J. and Yoder,B. K. (2005). Gli2 and Gli3 localize to cilia and require the intraflagellar transportprotein polaris for processing and function. PLoS Genet. 1, e53.

Huang, K., Diener, D. R. and Rosenbaum, J. L. (2009). The ubiquitin conjugationsystem is involved in the disassembly of cilia and flagella. J. Cell Biol. 186, 601-613.

Hunter, A. W., Caplow, M., Coy, D. L., Hancock, W. O., Diez, S., Wordeman,

L. and Howard, J. (2003). The kinesin-related protein MCAK is a microtubuledepolymerase that forms an ATP-hydrolyzing complex at microtubule ends. Mol. Cell11, 445-457.

Ishikawa, H., Thompson, J., Yates, J. R., III and Marshall, W. F. (2012). Proteomicanalysis of mammalian primary cilia. Curr. Biol. 22, 414-419.

Jakobsen, L., Vanselow, K., Skogs, M., Toyoda, Y., Lundberg, E., Poser, I.,

Falkenby, L. G., Bennetzen, M., Westendorf, J., Nigg, E. A. et al. (2011). Novel

asymmetrically localizing components of human centrosomes identified bycomplementary proteomics methods. EMBO J. 30, 1520-1535.

Jiang, K., Toedt, G., Montenegro Gouveia, S., Davey, N. E., Hua, S., van der Vaart,

B., Grigoriev, I., Larsen, J., Pedersen, L. B., Bezstarosti, K. et al. (2012). AProteome-wide screen for mammalian SxIP motif-containing microtubule plus-end

tracking proteins. Curr. Biol. 22, 1800-1807.

Johnson, K. A. (1998). The axonemal microtubules of the Chlamydomonas flagellumdiffer in tubulin isoform content. J. Cell Sci. 111, 313-320.

Johnson, U. G. and Porter, K. R. (1968). Fine structure of cell division inChlamydomonas reinhardi. Basal bodies and microtubules. J. Cell Biol. 38, 403-425.

Johnson, K. A. and Rosenbaum, J. L. (1992). Polarity of flagellar assembly inChlamydomonas. J. Cell Biol. 119, 1605-1611.

Kobayashi, T., Tsang, W. Y., Li, J., Lane, W. and Dynlacht, B. D. (2011). Centriolarkinesin Kif24 interacts with CP110 to remodel microtubules and regulate ciliogenesis.Cell 145, 914-925.

Kozminski, K. G., Johnson, K. A., Forscher, P. and Rosenbaum, J. L. (1993). Amotility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad.Sci. USA 90, 5519-5523.

Kubo, A., Yuba-Kubo, A., Tsukita, S., Tsukita, S. and Amagai, M. (2008). Sentan: anovel specific component of the apical structure of vertebrate motile cilia. Mol. Biol.Cell 19, 5338-5346.

Kuhns, S., Schmidt, K. N., Reymann, J., Gilbert, D. F., Neuner, A., Hub, B.,

Carvalho, R., Wiedemann, P., Zentgraf, H., Erfle, H. et al. (2013). Themicrotubule affinity regulating kinase MARK4 promotes axoneme extension duringearly ciliogenesis. J. Cell Biol. 200, 505-522.

Larsen, J., Grigoriev, I., Akhmanova, A. and Pedersen, L. B. (2013). Analysis ofmicrotubule plus-end-tracking proteins in cilia. Methods Enzymol. 524, 105-122.

Lefebvre, P. A., Nordstrom, S. A., Moulder, J. E. and Rosenbaum, J. L. (1978).Flagellar elongation and shortening in Chlamydomonas. IV. Effects of flagellardetachment, regeneration, and resorption on the induction of flagellar protein

synthesis. J. Cell Biol. 78, 8-27.

Lefebvre, P. A., Silflow, C. D., Wieben, E. D. and Rosenbaum, J. L. (1980). Increasedlevels of mRNAs for tubulin and other flagellar proteins after amputation or

shortening of Chlamydomonas flagella. Cell 20, 469-477.

Lewin, R. A. (1953). Studies on the flagella of algae. Il. Formation of flagella byChlamydomonas in light and darkness. Ann. N. Y. Acad. Sci. 56, 1091-1093.

Li, J. B., Gerdes, J. M., Haycraft, C. J., Fan, Y., Teslovich, T. M., May-Simera, H.,

Li, H., Blacque, O. E., Li, L., Leitch, C. C. et al. (2004). Comparative genomicsidentifies a flagellar and basal body proteome that includes the BBS5 human disease

gene. Cell 117, 541-552.

Liem, K. F., Jr, He, M., Ocbina, P. J. and Anderson, K. V. (2009). Mouse Kif7/Costal2 is a cilia-associated protein that regulates Sonic hedgehog signaling. Proc.

Natl. Acad. Sci. USA 106, 13377-13382.

Marshall, W. F. and Rosenbaum, J. L. (2001). Intraflagellar transport balances

continuous turnover of outer doublet microtubules: implications for flagellar lengthcontrol. J. Cell Biol. 155, 405-414.

Marshall, W. F., Qin, H., Rodrigo Brenni, M. and Rosenbaum, J. L. (2005). Flagellar

length control system: testing a simple model based on intraflagellar transport andturnover. Mol. Biol. Cell 16, 270-278.

Matsuo, T., Okamoto, K., Onai, K., Niwa, Y., Shimogawara, K. and Ishiura, M.

(2008). A systematic forward genetic analysis identified components of theChlamydomonas circadian system. Genes Dev. 22, 918-930.

Miller, J. M., Wang, W., Balczon, R. and Dentler, W. L. (1990). Ciliary microtubule

capping structures contain a mammalian kinetochore antigen. J. Cell Biol. 110, 703-714.

Niwa, S., Nakajima, K., Miki, H., Minato, Y., Wang, D. and Hirokawa, N. (2012).

KIF19A is a microtubule-depolymerizing kinesin for ciliary length control. Dev. Cell23, 1167-1175.

Ostrowski, L. E., Blackburn, K., Radde, K. M., Moyer, M. B., Schlatzer, D. M.,

Moseley, A. and Boucher, R. C. (2002). A proteomic analysis of human cilia:identification of novel components. Mol. Cell. Proteomics 1, 451-465.

Pan, J., Naumann-Busch, B., Wang, L., Specht, M., Scholz, M., Trompelt, K. and

Hippler, M. (2011). Protein phosphorylation is a key event of flagellar disassemblyrevealed by analysis of flagellar phosphoproteins during flagellar shortening inChlamydomonas. J. Proteome Res. 10, 3830-3839.

Pazour, G. J., Agrin, N., Leszyk, J. and Witman, G. B. (2005). Proteomic analysis ofa eukaryotic cilium. J. Cell Biol. 170, 103-113.

Pedersen, L. B., Geimer, S., Sloboda, R. D. and Rosenbaum, J. L. (2003). TheMicrotubule plus end-tracking protein EB1 is localized to the flagellar tip and basalbodies in Chlamydomonas reinhardtii. Curr. Biol. 13, 1969-1974.

Pedersen, L. B., Miller, M. S., Geimer, S., Leitch, J. M., Rosenbaum, J. L. and Cole,

D. G. (2005). Chlamydomonas IFT172 is encoded by FLA11, interacts with CrEB1,and regulates IFT at the flagellar tip. Curr. Biol. 15, 262-266.

Pedersen, L. B., Geimer, S. and Rosenbaum, J. L. (2006). Dissecting the molecularmechanisms of intraflagellar transport in chlamydomonas. Curr. Biol. 16, 450-459.

Piao, T., Luo, M., Wang, L., Guo, Y., Li, D., Li, P., Snell, W. J. and Pan, J. (2009). A

microtubule depolymerizing kinesin functions during both flagellar disassemblyand flagellar assembly in Chlamydomonas. Proc. Natl. Acad. Sci. USA 106, 4713-4718.

Pigino, G., Geimer, S., Lanzavecchia, S., Paccagnini, E., Cantele, F., Diener, D. R.,

Rosenbaum, J. L. and Lupetti, P. (2009). Electron-tomographic analysis ofintraflagellar transport particle trains in situ. J. Cell Biol. 187, 135-148.

Journal of Cell Science 126 (21)5028

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.133439/-/DC1http://dx.doi.org/10.1016/j.tcb.2007.11.002http://dx.doi.org/10.1016/j.tcb.2007.11.002http://dx.doi.org/10.1016/j.cub.2007.03.048http://dx.doi.org/10.1016/j.cub.2007.03.048http://dx.doi.org/10.1016/j.cub.2007.03.048http://dx.doi.org/10.1083/jcb.201204072http://dx.doi.org/10.1083/jcb.201204072http://dx.doi.org/10.1016/S0091-679X(08)94017-6http://dx.doi.org/10.1016/S0091-679X(08)94017-6http://dx.doi.org/10.1016/j.molbiopara.2010.08.001http://dx.doi.org/10.1016/j.molbiopara.2010.08.001http://dx.doi.org/10.1016/j.molbiopara.2010.08.001http://dx.doi.org/10.1083/jcb.141.4.993http://dx.doi.org/10.1083/jcb.141.4.993http://dx.doi.org/10.1083/jcb.141.4.993http://dx.doi.org/10.1083/jcb.141.4.993http://dx.doi.org/10.1128/EC.00128-07http://dx.doi.org/10.1128/EC.00128-07http://dx.doi.org/10.1128/EC.00128-07http://dx.doi.org/10.1083/jcb.74.3.747http://dx.doi.org/10.1083/jcb.74.3.747http://dx.doi.org/10.1083/jcb.74.3.747http://dx.doi.org/10.1083/jcb.74.3.747http://dx.doi.org/10.1091/mbc.E06-02-0133http://dx.doi.org/10.1091/mbc.E06-02-0133http://dx.doi.org/10.1091/mbc.E06-02-0133http://dx.doi.org/10.1096/fj.09-151381http://dx.doi.org/10.1096/fj.09-151381http://dx.doi.org/10.1096/fj.09-151381http://dx.doi.org/10.1038/nrg2774http://dx.doi.org/10.1038/nrg2774http://dx.doi.org/10.1016/j.cell.2012.10.010http://dx.doi.org/10.1016/j.cell.2012.10.010http://dx.doi.org/10.1016/j.cell.2012.10.010http://dx.doi.org/10.1371/journal.pgen.0010053http://dx.doi.org/10.1371/journal.pgen.0010053http://dx.doi.org/10.1371/journal.pgen.0010053http://dx.doi.org/10.1083/jcb.200903066http://dx.doi.org/10.1083/jcb.200903066http://dx.doi.org/10.1016/S1097-2765(03)00049-2http://dx.doi.org/10.1016/S1097-2765(03)00049-2http://dx.doi.org/10.1016/S1097-2765(03)00049-2http://dx.doi.org/10.1016/S1097-2765(03)00049-2http://dx.doi.org/10.1016/j.cub.2012.01.031http://dx.doi.org/10.1016/j.cub.2012.01.031http://dx.doi.org/10.1038/emboj.2011.63http://dx.doi.org/10.1038/emboj.2011.63http://dx.doi.org/10.1038/emboj.2011.63http://dx.doi.org/10.1038/emboj.2011.63http://dx.doi.org/10.1016/j.cub.2012.07.047http://dx.doi.org/10.1016/j.cub.2012.07.047http://dx.doi.org/10.1016/j.cub.2012.07.047http://dx.doi.org/10.1016/j.cub.2012.07.047http://dx.doi.org/10.1083/jcb.38.2.403http://dx.doi.org/10.1083/jcb.38.2.403http://dx.doi.org/10.1083/jcb.119.6.1605http://dx.doi.org/10.1083/jcb.119.6.1605http://dx.doi.org/10.1016/j.cell.2011.04.028http://dx.doi.org/10.1016/j.cell.2011.04.028http://dx.doi.org/10.1016/j.cell.2011.04.028http://dx.doi.org/10.1073/pnas.90.12.5519http://dx.doi.org/10.1073/pnas.90.12.5519http://dx.doi.org/10.1073/pnas.90.12.5519http://dx.doi.org/10.1091/mbc.E08-07-0691http://dx.doi.org/10.1091/mbc.E08-07-0691http://dx.doi.org/10.1091/mbc.E08-07-0691http://dx.doi.org/10.1083/jcb.201206013http://dx.doi.org/10.1083/jcb.201206013http://dx.doi.org/10.1083/jcb.201206013http://dx.doi.org/10.1083/jcb.201206013http://dx.doi.org/10.1016/B978-0-12-397945-2.00007-Xhttp://dx.doi.org/10.1016/B978-0-12-397945-2.00007-Xhttp://dx.doi.org/10.1083/jcb.78.1.8http://dx.doi.org/10.1083/jcb.78.1.8http://dx.doi.org/10.1083/jcb.78.1.8http://dx.doi.org/10.1083/jcb.78.1.8http://dx.doi.org/10.1016/0092-8674(80)90633-9http://dx.doi.org/10.1016/0092-8674(80)90633-9http://dx.doi.org/10.1016/0092-8674(80)90633-9http://dx.doi.org/10.1111/j.1749-6632.1953.tb30293.xhttp://dx.doi.org/10.1111/j.1749-6632.1953.tb30293.xhttp://dx.doi.org/10.1016/S0092-8674(04)00450-7http://dx.doi.org/10.1016/S0092-8674(04)00450-7http://dx.doi.org/10.1016/S0092-8674(04)00450-7http://dx.doi.org/10.1016/S0092-8674(04)00450-7http://dx.doi.org/10.1083/jcb.200106141http://dx.doi.org/10.1083/jcb.200106141http://dx.doi.org/10.1083/jcb.200106141http://dx.doi.org/10.1091/mbc.E04-07-0586http://dx.doi.org/10.1091/mbc.E04-07-0586http://dx.doi.org/10.1091/mbc.E04-07-0586http://dx.doi.org/10.1101/gad.1650408http://dx.doi.org/10.1101/gad.1650408http://dx.doi.org/10.1101/gad.1650408http://dx.doi.org/10.1083/jcb.110.3.703http://dx.doi.org/10.1083/jcb.110.3.703http://dx.doi.org/10.1083/jcb.110.3.703http://dx.doi.org/10.1016/j.devcel.2012.10.016http://dx.doi.org/10.1016/j.devcel.2012.10.016http://dx.doi.org/10.1016/j.devcel.2012.10.016http://dx.doi.org/10.1074/mcp.M200037-MCP200http://dx.doi.org/10.1074/mcp.M200037-MCP200http://dx.doi.org/10.1074/mcp.M200037-MCP200http://dx.doi.org/10.1021/pr200428nhttp://dx.doi.org/10.1021/pr200428nhttp://dx.doi.org/10.1021/pr200428nhttp://dx.doi.org/10.1021/pr200428nhttp://dx.doi.org/10.1083/jcb.200504008http://dx.doi.org/10.1083/jcb.200504008http://dx.doi.org/10.1016/j.cub.2003.10.058http://dx.doi.org/10.1016/j.cub.2003.10.058http://dx.doi.org/10.1016/j.cub.2003.10.058http://dx.doi.org/10.1016/j.cub.2005.01.037http://dx.doi.org/10.1016/j.cub.2005.01.037http://dx.doi.org/10.1016/j.cub.2005.01.037http://dx.doi.org/10.1016/j.cub.2006.02.020http://dx.doi.org/10.1016/j.cub.2006.02.020http://dx.doi.org/10.1073/pnas.0808671106http://dx.doi.org/10.1073/pnas.0808671106http://dx.doi.org/10.1073/pnas.0808671106http://dx.doi.org/10.1073/pnas.0808671106http://dx.doi.org/10.1083/jcb.200905103http://dx.doi.org/10.1083/jcb.200905103http://dx.doi.org/10.1083/jcb.200905103

Journ

alof

Cell

Scie

nce

Reynolds, E. S. (1963). The use of lead citrate at high pH as an electron-opaque stain inelectron microscopy. J. Cell Biol. 17, 208-212.

Rieder, C. L., Jensen, C. G. and Jensen, L. C. (1979). The resorption of primary ciliaduring mitosis in a vertebrate (PtK1) cell line. J. Ultrastruct. Res. 68, 173-185.

Ringo, D. L. (1967). Flagellar motion and fine structure of the flagellar apparatus inChlamydomonas. J. Cell Biol. 33, 543-571.

Rogowski, M., Scholz, D. and Geimer, S. (2013). Electron microscopy of flagella,primary cilia, and intraflagellar transport in flat-embedded cells. Methods Enzymol.524, 243-263.

Rosenbaum, J. L. and Child, F. M. (1967). Flagellar regeneration in protozoanflagellates. J. Cell Biol. 34, 345-364.

Rosenbaum, J. L., Moulder, J. E. and Ringo, D. L. (1969). Flagellar elongation andshortening in Chlamydomonas. The use of cycloheximide and colchicine to study thesynthesis and assembly of flagellar proteins. J. Cell Biol. 41, 600-619.

Ross, P. L., Huang, Y. N., Marchese, J. N., Williamson, B., Parker, K., Hattan, S.,Khainovski, N., Pillai, S., Dey, S., Daniels, S. et al. (2004). Multiplexed proteinquantitation in Saccharomyces cerevisiae using amine-reactive isobaric taggingreagents. Mol. Cell. Proteomics 3, 1154-1169.

Schneider, M. J., Ulland, M. and Sloboda, R. D. (2008). A protein methylationpathway in Chlamydomonas flagella is active during flagellar resorption. Mol. Biol.Cell 19, 4319-4327.

Schrøder, J. M., Larsen, J., Komarova, Y., Akhmanova, A., Thorsteinsson, R. I.,

Grigoriev, I., Manguso, R., Christensen, S. T., Pedersen, S. F., Geimer, S. et al.

(2011). EB1 and EB3 promote cilia biogenesis by several centrosome-relatedmechanisms. J. Cell Sci. 124, 2539-2551.

Sorokin, S. P. (1968). Reconstructions of centriole formation and ciliogenesis inmammalian lungs. J. Cell Sci. 3, 207-230.

Spektor, A., Tsang, W. Y., Khoo, D. and Dynlacht, B. D. (2007). Cep97 and CP110suppress a cilia assembly program. Cell 130, 678-690.

Tammana, T. V., Sahasrabuddhe, A. A., Mitra, K., Bajpai, V. K. and Gupta, C. M.

(2008). Actin-depolymerizing factor, ADF/cofilin, is essentially required in assemblyof Leishmania flagellum. Mol. Microbiol. 70, 837-852.

Tukachinsky, H., Lopez, L. V. and Salic, A. (2010). A mechanism for vertebrateHedgehog signaling: recruitment to cilia and dissociation of SuFu-Gli proteincomplexes. J. Cell Biol. 191, 415-428.

Wheatley, D. N., Wang, A. M. and Strugnell, G. E. (1996). Expression of primary ciliain mammalian cells. Cell Biol. Int. 20, 73-81.

Witman, G. B. (1975). The site of in vivo assembly of flagellar microtubules. Ann. N. Y.Acad. Sci. 253, 178-191.

Witman, G. B., Carlson, K., Berliner, J. and Rosenbaum, J. L. (1972).Chlamydomonas flagella. I. Isolation and electrophoretic analysis of microtubules,matrix, membranes, and mastigonemes. J. Cell Biol. 54, 507-539.

Woolley, D., Gadelha, C. and Gull, K. (2006). Evidence for a sliding-resistance at thetip of the trypanosome flagellum. Cell Motil. Cytoskeleton 63, 741-746.

Wordeman, L. (2005). Microtubule-depolymerizing kinesins. Curr. Opin. Cell Biol. 17,82-88.

Role of FAP256/CEP104 in ciliogenesis 5029

http://dx.doi.org/10.1083/jcb.17.1.208http://dx.doi.org/10.1083/jcb.17.1.208http://dx.doi.org/10.1016/S0022-5320(79)90152-7http://dx.doi.org/10.1016/S0022-5320(79)90152-7http://dx.doi.org/10.1083/jcb.33.3.543http://dx.doi.org/10.1083/jcb.33.3.543http://dx.doi.org/10.1016/B978-0-12-397945-2.00014-7http://dx.doi.org/10.1016/B978-0-12-397945-2.00014-7http://dx.doi.org/10.1016/B978-0-12-397945-2.00014-7http://dx.doi.org/10.1083/jcb.34.1.345http://dx.doi.org/10.1083/jcb.34.1.345http://dx.doi.org/10.1083/jcb.41.2.600http://dx.doi.org/10.1083/jcb.41.2.600http://dx.doi.org/10.1083/jcb.41.2.600http://dx.doi.org/10.1074/mcp.M400129-MCP200http://dx.doi.org/10.1074/mcp.M400129-MCP200http://dx.doi.org/10.1074/mcp.M400129-MCP200http://dx.doi.org/10.1074/mcp.M400129-MCP200http://dx.doi.org/10.1091/mbc.E08-05-0470http://dx.doi.org/10.1091/mbc.E08-05-0470http://dx.doi.org/10.1091/mbc.E08-05-0470http://dx.doi.org/10.1242/jcs.085852http://dx.doi.org/10.1242/jcs.085852http://dx.doi.org/10.1242/jcs.085852http://dx.doi.org/10.1242/jcs.085852http://dx.doi.org/10.1016/j.cell.2007.06.027http://dx.doi.org/10.1016/j.cell.2007.06.027http://dx.doi.org/10.1083/jcb.201004108http://dx.doi.org/10.1083/jcb.201004108http://dx.doi.org/10.1083/jcb.201004108http://dx.doi.org/10.1006/cbir.1996.0011http://dx.doi.org/10.1006/cbir.1996.0011http://dx.doi.org/10.1111/j.1749-6632.1975.tb19199.xhttp://dx.doi.org/10.1111/j.1749-6632.1975.tb19199.xhttp://dx.doi.org/10.1083/jcb.54.3.507http://dx.doi.org/10.1083/jcb.54.3.507http://dx.doi.org/10.1083/jcb.54.3.507http://dx.doi.org/10.1002/cm.20159http://dx.doi.org/10.1002/cm.20159http://dx.doi.org/10.1016/j.ceb.2004.12.003http://dx.doi.org/10.1016/j.ceb.2004.12.003

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