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Originally published In Press as doi:10.1074/jbc.M103184200 on May 30, 2001

J. Biol. Chem., Vol. 276, Issue 32, 29754-29763, August 10, 2001
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Flagellar Protein Dynamics in Chlamydomonas*

Lin Song and William L. DentlerDagger

From the Department of Molecular Biosciences, University of Kansas, Lawrence, Kansas 66045

Received for publication, April 10, 2001

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Cilia and flagella appear to be stable, terminal, microtubule-containing organelles, but they also elongate and shorten in response to a variety of signals. To understand mechanisms that regulate flagellar dynamics, Chlamydomonas cells with nongrowing flagella were labeled with 35S, and flagella and basal body components were examined for labeled polypeptides. Maximal incorporation of label into the flagella occurred within 3 h. Twenty percent of the flagellar polypeptides were exchanged. These included tubulins, dyneins, and 80 other axonemal and membrane plus matrix polypeptides. The most stable flagellar structure is the PF-ribbon, which comprises part of the wall of each doublet microtubule and is composed of tubulin and three other polypeptides. Most 35S was incorporated into the high molecular weight ribbon polypeptide, rib240, and little, if any, 35S is incorporated into PF-ribbon-associated tubulin. Both wild-type (9 + 2) and 9 + 0 flagella, which lack central microtubules, exhibited nearly identical exchange patterns, so labeling is not due to turnover of relatively labile central microtubules. To determine if flagellar length is balanced by protein exchange, 35S incorporation into disassembling flagella was examined, as was exchange in flagella in which microtubule assembly was blocked by colchicine. Incorporation of 35S-labeled polypeptides was found to occur into flagellar axonemes during wavelength-dependent shortening in pf18 and in fla10 cells induced to shorten flagella by incubation at 33 °C. Colchicine blocked tubulin addition but did not affect the exchange of the other exchangeable polypeptides; nor did it induce any change in flagellar length. Basal bodies also incorporated newly synthesized proteins. These data reveal that Chlamydomonas flagella are dynamic structures that incorporate new protein both during steady state and as flagella shorten and that protein exchange does not, alone, explain length regulation.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Cilia and eucaryotic flagella are ubiquitous microtubule-containing organelles that mediate a variety of functions and are prominently displayed on cells in the heart, kidney, ovary, uterus, liver, central nervous system, spleen, inner ear, and olfactory epithelium and in adrenal, thyroid, pancreatic, thymus, and pituitary glands. Cilia propel vertebrate sperm, display visual pigments in vertebrate photoreceptors and sensory molecules on olfactory and chemosensory neurons. During development, they are prominent on embryonic surfaces.

Pathological conditions associated with defects in ciliogenesis include liver disorders, drug-induced cirrhosis, gastritis, hyperparathyroidism, retinal dysplasia, Mulvihill-Smith progeria-like syndrome, Oak Ridge polycystic kidney disease (1), Usher's syndrome and retinitis pigmentosa. Patients with primary ciliary dyskinesia have abnormally aligned cilia or cilia with motility defects, which leads to infertility and decreased resistance to lower respiratory tract infections, chronic rhinitis, sinusitis, and otitis media. Ciliary movement is essential for the development of asymmetry in developing mammals and is particularly important for development and positioning of the heart (2, 3). There are likely to be many unknown links between cilia and embryonic development. Cilia line embryonic surfaces of developing neural tubes and ventral and notochordal plate cells in developing mouse embryos (4, 5), and they appear during neural tube closure (6). Cell- or tissue-specific defects in cilia formation may cause currently unidentified defects during early embryonic development.

The growth, maintenance, and disassembly of cilia and flagella is a highly orchestrated process in which the length of each of the nine doublets and the two central microtubules is precisely regulated as is the addition of more than 250 different polypeptides that attach to the microtubules or are associated with the membrane and soluble matrix. Isolated cilia and flagella are extremely stable microtubular structures that can only be dissociated by ionic detergents, heat, or chaotropic agents (7).1 Despite their stability in vitro, fully grown flagella in many cells can be completely disassembled (8). In sea urchin embryos, cilia can be stimulated to grow by the application of several different agents, including concanavalin A (9) and theophylline (10). Some of the most dramatic examples of flagellar dynamics occur in Chlamydomonas, in which flagella normally disassemble prior to cell division and new flagella grow after division is completed. If one flagellum is amputated, the remaining flagellum shortens and then elongates once a new flagellum has started to grow on the "amputated" basal body (11). Chlamydomonas flagellar growth can be stimulated with lithium (12, 13), induced to disassemble by IBMX and other agents known to be involved with signal transduction pathways (13), and destabilized by cytochalasin D, which induces periods of rapid shortening and elongation (14). A temperature-sensitive mutant, fla10, shortens its flagella at 33 °C but not at 22 °C (15, 16). Chlamydomonas pf18 mutants shorten flagella under reduced red light (13), and long flagellar (lf) mutants produce up to 3 times longer than normal flagella (17). When the cytoplasm of cells with abnormally long flagella is mixed with that of cells with normal length flagella, the long flagella rapidly shorten (17), indicating that signaling mechanisms that regulate flagellar length are dominant in cells with normal length flagella. Although isolated flagellar microtubules are the most stable microtubules known, these microtubules clearly can be dynamic in vivo. Little is understood about the mechanisms that regulate flagellar dynamics, although recent studies point to a role of signal transduction pathways (13, 18).

Flagellar assembly, or elongation, requires the transport of subunits from the cytoplasm, where they are synthesized, to the tips of the flagellar microtubules, where they are added to the microtubule ends (19). Some of the transport to the distal tips requires the heterotrimeric kinesin, kinesin-II, which is associated with intraflagellar transport (IFT) (20). Early studies by Morris and Scholey (21) revealed that sea urchin embryonic cilia could grow nearly half-length if IFT is blocked by kinesin-II antibodies but that further growth could not occur. Subsequent studies have confirmed that kinesin-II is required for flagellar assembly in Chlamydomonas (16, 22), C. elegans (23-26), Tetrahymena (27), tracheal epithelia (28), and mouse embryonic nodal cilia (29, 30). Kinesin-II appears to deliver flagellar protein to the distal tips of the flagellum, the plus-ends of the microtubules, where they are added to the microtubules.

The components transported by kinesin-II require recycling to flagellar bases for flagellar growth or maintenance to occur, and this movement requires cytoplasmic dyneins (31-33). Direct movement of cytoplasmic dynein has been observed in sensory ciliated neurons in C. elegans (26). Additionally, cytoplasmic dynein is required for the delivery of rhodopsin to vertebrate retinal cilia, and the failure to do so may lead to retinitis pigmentosa (34, 35).

The dependence of flagellar length on kinesin-II and cytoplasmic dynein indicates that flagellar proteins and/or signal transduction pathway components are shuttled back and forth along the flagellum and that some flagellar components are likely to be exchanged in the fully assembled flagellum. One indication of the importance of this shuttling is provided by examination of Chlamydomonas cytoplasmic dynein mutants. These cells can assemble short flagella, but shortly after they are formed, the flagella shorten (31-33) and, in many cases, the microtubules start to fray into disorganized filamentous structures (31). Marshall and Rosenbaum (36) showed that epitope-tagged tubulin could be added to steady state flagella and revealed the presence of retrograde tubulin flux. Moreover, the retrograde flux was balanced by IFT (37). Taken together, these studies clearly indicate that the balance of anterograde and retrograde IFT and, possibly, some flagellar protein turnover is essential for the maintenance of steady state length. Imbalances in anterograde transport (as in kinesin-II mutants) or in retrograde transport (as in cytoplasmic dynein mutants) may lead to flagellar disassembly or, possibly, elongation.

Several early studies of pulse-labeled flagella indicated that a low amount of protein exchange, or turnover, occurred in steady state flagella. Low levels of protein turnover were detected in whole flagella and in the membrane plus matrix and axonemal fractions (11, 38-41), and some tubulin turnover was detected in Chlamydomonas flagella (40) and Tetrahymena cilia (42). The only prominent axonemal protein detected to turnover in Chlamydomonas flagella was a nontubulin 55-kDa "protein X" (43). However, none of these studies rigorously ruled out the possibility that the incorporation of label was not due to protein exchange but, rather, was due to the assembly of new flagella due to flagellar amputation, which up-regulates flagellar protein synthesis. In contrast to protozoans, embryonic cilia on marine organisms are considerably more dynamic. Steady state cilia on sea urchin blastulae exhibit exchange of numerous membrane plus matrix and axonemal polypeptides, including tubulin and tektin A (44, 45).

The recent studies of sea urchin ciliary turnover and the importance of IFT motors in maintaining flagellar length prompted us to critically reexamine flagellar protein turnover in steady state Chlamydomonas flagella. Moreover, with the availability of mutants that shorten flagella due to light or temperature shifts and long flagellar mutants, we could compare the dynamics of protein turnover in shortening or abnormally long flagella. The results reported here show that steady state flagella exhibit a considerable amount of protein exchange, with a minimum 20% of flagellar proteins being exchanged within a 6-h period. Although tubulin turnover occurs, tubulin turnover could be blocked without affecting the turnover of nontubulin components. Turnover of most flagellar proteins continues to occur as flagella shorten, thus demonstrating that flagellar length is not strictly coupled to the turnover of major flagellar proteins. The single peptide exhibiting the greatest exchange is rib240, a component of the axonemal PF-ribbons. Unlike the ribbon fractions in sea urchin cilia, neither significant tubulin nor tektin-like proteins are exchanged in the PF-ribbons, but a new high molecular weight component, rib240, is exchanged. Since the ribbons have been suggested to be important stabilizers or length determinants in flagella, the role of rib240 in flagellar assembly deserves serious consideration.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Cell Culture and 35S Labeling-- For all experiments, Chlamydomonas reinhardtii wild-type (CC-2929), pf18 (CC-1036), fla10 (CC-1919), and lf4-6 (P. Lefebvre, University of Minnesota) were grown in minimal medium (M medium)2 bubbled with air and were synchronized by growing cells on a 12-h light/12-h dark cycle. For each experiment, t = 0 was the start of the light period.

Vegetative cells were grown to a density of 5 × 106 cells/ml, harvested in sterile bottles, and incubated in low sulfur M medium (MgSO4 was reduced from 1.2 to 0.012 mM, and MgCl2 was added to 1.18 mM) for 24 h. For each labeling experiment, 4 µCi/ml 35S (as sulfuric acid; PerkinElmer Life Sciences) was added to starved cells from a 1 mCi/ml stock solution in sterile distilled water. For colchicine experiments, freshly prepared colchicine was added to cells at a final concentration of 2 mg/ml. Each batch of colchicine was tested to ensure that it inhibited flagellar growth after deflagellation of cells by pH shock.

For chase experiments, 1 liter of starved cells were labeled with 4 µCi/ml 35S for up to 6 h and then pelleted at 1100 × g for 5 min at 20 °C in a JA-10 (Beckman) rotor and washed twice with 400 ml of low sulfur M medium, and cells were gently resuspended in 1 liter of low sulfur M medium supplemented with 30 ml of 10% MgSO4. The final cell suspension was examined by phase microscopy to ensure that all cells were uniformly flagellated. After the chase period, cells were pelleted, and flagella were isolated and fractionated as described below.

To induce shortening in light-synchronized pf18 cells, cells were harvested and incubated under low red lights with gentle shaking at 1000 rpm, described by Tuxhorn et al. (13).

To measure flagellar length, cells were fixed in an equal volume of 2% glutaraldehyde for 20 min. Flagellar length was determined by phase-contrast microscopy using an ocular micrometer. For each sample, flagellar lengths were averaged from 50 biflagellated cells.

Flagellar Isolation and Fractionation-- Flagella were amputated using dibucaine and were isolated following procedures described by King (46). Cells were collected by centrifugation (1100 × g, 5 min) and were suspended in 50 ml of ice-cold HMDS (10 mM HEPES, 5 mM MgSO4, 1 mM dithiothreitol, 4% sucrose, pH 7.5). Dibucaine was added to a final concentration of 1 mM, and cells were swirled for 1-2 min and examined by phase-contrast microscopy to ensure that all cells were deflagellated. All subsequent steps were carried out at 4 °C. Cell bodies were pelleted (1100 × g, 5 min), and flagella were recovered from the supernatant by centrifugation at 7500 × g for 10 min. Flagellar pellets were suspended and centrifuged over HMD (10 mM HEPES, 5 mM MgSO4, 1 mM dithiothreitol) with 25% sucrose at 1100 × g for 20 min. Purified flagella were harvested at the interface. The membrane plus matrix fractions were isolated by incubating flagella with 0.1% Nonidet P-40 (Sigma) for 20 min on ice and then centrifuging the suspension at 12,000 × g for 10 min. The pelleted axonemes were then twice suspended in HMDS plus 0.6 M KCl, incubated for 20 min on ice, and centrifuged at 12,000 × g for 10 min.

PF-ribbons were purified as described by Norrander et al. (47). Isolated axonemes were incubated in 0.7% Sarkosyl in TED (10 mM Tris, pH 8, 0.1 mM EDTA, 1 mM dithiothreitol) overnight with gentle shaking. Ribbons were pelleted 100,000 × g for 1 h, washed with 1 ml of TED, and pelleted a final time.

Basal bodies were isolated using the method of Snell (48) with the following modifications. Sulfur-starved cells were incubated with H235SO4 for 2 or 6 h as described above. Cells were collected by centrifugation (1100 × g, 5 min, 20 °C), suspended in 100 ml of TE (10 mM Tris, 1 mM EDTA, pH 7.5), and deflagellated with 1 mM dibucaine (Sigma). After centrifugation (1500 × g, 3 min, 20 °C), the supernatant was collected to purify flagella, and the pellet (cell bodies) was suspended in autolysin (49) for 40 min to remove cell walls. Cells were diluted with an equal volume of 1% Nonidet P-40 in TE, stirred for 20 min, and homogenized using a 25-ml glass-Teflon homogenizer with 20 up and down strokes. Cell lysates were layered over 20 ml of 25% sucrose-TE in 50 ml of conical polycarbonate tubes and centrifuged in a swinging bucket rotor (1500 × g, 10 min, 4 °C) to sediment cells. The supernatant was then layered over 25% sucrose-TE and recentrifuged. This supernatant was layered over a 40-50% sucrose-TE gradient and centrifuged in a swinging bucket rotor (14,000 × g, 60 min, 4 °C). Basal bodies were collected from the 40-50% interface, diluted with equal volume of TE, and pelleted by centrifugation (35,000 × g, 30 min, 4 °C). The basal bodies were suspended in 3 ml of 0.5% Nonidet P-40 in TE, homogenized in a 5-ml glass-Teflon homogenizer, layered over a 50-55-60% sucrose-TE gradient, and centrifuged in a swinging bucket rotor (14,000 × g, 45 min, 4 °C). Basal bodies were collected from the 55-60% interface and treated with 0.6 M NaCl on ice for 20 min. After centrifugation (35,000 × g, 30 min, 4 °C), basal bodies were collected from the pellet.

Flagellar Regeneration-- Synchronized wild-type cells were starved in low sulfur M medium for 24 h and deflagellated by pH shock. Cells were concentrated in 200 ml of low sulfur M medium, and 0.5 N acetic acid was added dropwise until pH 4.5 was reached. After 30 s, the pH was raised to 7.0 by adding 0.5 N KOH. Cells were then pelleted and resuspended in low sulfur M medium. 35S-Labeled sulfuric acid was added to the cell culture (4 µCi/ml) and incubated for at least 2 h. Cells were checked periodically with a phase-contrast microscope to ensure that flagella regenerated to full-length before flagella were isolated.

Electrophoresis and Fluorography-- Flagellar and cell body polypeptides were separated by SDS-PAGE, using the urea-SDS method of Lefebvre et al. (50). To compare labeled polypeptides in each fraction, each gel lane contained the same amount of protein. Gels were stained with Coomassie Blue, treated with Enhance (PerkinElmer Life Sciences), air-dried between cellophane, and were exposed on a preflashed Kodak X-Omat XR-5 x-ray film for 2-20 days. For some experiments, dried gels were exposed and read using a Molecular Imager FX System (Bio-Rad).

Electron Microscopy-- Ten-µl aliquots of purified basal bodies, microtubules, or PF-ribbons were applied to carbon-over-formvar grids and stained with 2% uranyl acetate. Samples were stained and photographed using a JEOL 1200EXII microscope operating at 80 kV.

Quantitation of 35S-Labeled Proteins in Whole Cells and Flagella-- To assay 35S incorporation in whole cells during the light cycle, 2 ml of sulfur-starved cells were taken from the culture every hour during the light period and incubated with 4 µCi/ml 35S for 30 min. Duplicate 50-µl aliquots of cells were spotted onto filter disks (2.3 cm; Whatman 3MM), dropped immediately into cold 20% perchloric acid, incubated for 1 h, washed twice with 50% ethanol, rinsed with distilled water, and air-dried. Each disc was placed in a standard 20-ml plastic scintillation vial with 5 ml of scintillation liquid and counted using a liquid scintillation counter (Packard Instrument Co. 1600 TR Tri-Carb).

To measure the incorporation of labeled proteins into each flagellar compartment, duplicate 10-µg samples from intact flagella, membrane plus matrix fraction, KCl-soluble proteins, KCl-insoluble axonemes, and PF-ribbons were spotted onto filter disks, treated with perchloric acid, and counted as described above. To compare the incorporation in different time periods or in different cell strains, radioactivity was calculated and converted as either total cpm in each fraction or cpm/µg for each fraction.

For some experiments, polypeptides were sliced from the SDS-polyacrylamide gels containing equal amount proteins for each component after silver staining and were placed into a 20-ml scintillation vial. Ten ml of 5% Soluene-350 in Insta-Fluor (Packard Instrument Co.) were added to dissolve the gel slices. After incubation at room temperature overnight, samples were counted as described above.

Incorporation of 35S in selected polypeptides was also measured from the fluorograms using NIH Image 1.61. Equal amounts of protein for each flagellar compartment were separated by SDS-PAGE, dried, and exposed to a preflashed x-ray film. Fluorograms were scanned into the computer with an Eastman Kodak Co. density gradient step, and the full-range digital image was analyzed in NIH Image 1.61 calibrated with the density standard. Optical densities for individual polypeptides and the total flagellar fraction were determined. The percentage of labeled individual polypeptides and the total labeled proteins in each compartment were obtained by comparing densities of individual polypeptides with the optical density of the fraction. At least two separate gels were measured for each polypeptide. The S.D. among experiments was statistically insignificant.

Polyclonal Antibody Generation-- A polyclonal antibody was raised against rib240 (51). Briefly, PF-ribbons isolated from 4 liters of wild type cells were separated in SDS-PAGE and stained with Coomassie Blue. The band containing rib240 was excised, homogenized in PBS, and emulsified with Freund's complete adjuvant (Sigma). Three mice were immunized intraperitoneally. Serum was obtained after second boost for antibody detection.

Immunoblot Analysis-- Purified PF-ribbons were separated in SDS-PAGE. The gel was briefly soaked in 49.6 mM Tris, 384 mM glycine, 20% (v/v) methanol, 0.1% (w/v) SDS and blotted onto a nitrocellulose sheet for 60 min at 24 V in a blotting apparatus (Idea Scientific).

For antigen detection, nitrocellulose was blocked with blocking buffer (5% Carnation dry milk, 0.05% Tween 20, 0.15 M NaCl, 10 mM Tris-HCl, pH 7.25) for 1 h. The blot was incubated in either preimmune or postimmune serum diluted 1:1000 in blocking buffer for 1 h at room temperature. The blot was washed three times with TBS (150 mM NaCl, 10 mM Tris, pH 7.5) and incubated in alkaline phosphatase-goat anti-mouse IgG (Zymed Laboratories Inc.) for 1 h. The blot was then washed with Tris-buffered saline and developed with nitro blue tetrazolium and 5-bromo-4-chloro-3-indolyl phosphate in alkaline phosphatase buffer (100 mM Tris, 100 mM NaCl, 5 mM MgCl, pH 9.5). Blot was scanned into a computer with a flatbed scanner and saved as a digital image.

Immunofluorescence Microscopy-- Coverslips were treated with 5 mM EDTA for 5-10 min, rinsed in deionized water, and air-dried. Before samples were applied, coverslips were coated with 0.1% polyethyleneimine, rinsed with deionized water, and air-dried. A drop of isolated axonemes was placed on a glass coverslip for 10 min and fixed in 3% formaldehyde in PBSA (10.6 mM Na2HPO4, 1.47 mM KH2PO4, 137 mM NaCl, 2.68 KCl, 3% bovine serum albumin, pH 7.4) for 20 min at room temperature. Coverslips were incubated in postimmune serum (1:250 diluted in PBSA), anti-beta -tubulin antibody (R. H. Himes, University of Kansas) (1:500 diluted in PBSA), or PBSA alone for 4 h at 37 °C, washed in PBS (PBSA without albumin), incubated in Texas Red goat anti-mouse IgG (Molecular Probes, Inc., Eugene, OR) for 1 h at 37 °C, washed in PBS, and mounted on glass slides in anti-bleach solution (1 mg/ml p-phenylenediamine in PBS and glycerol, pH 8.0). Axonemes were viewed with a Zeiss standard microscope equipped with an epifluorescence unit containing a 450-490-nm excitation filter and 520-560-nm barrier filter. Samples were photographed using a × 63, 1.4 NA objective lens, × 10 projection eyepiece, and a Dage SIT66 camera. The image was frame-averaged with an Avio digital image processor (Nippon Avionics Co.) and captured as a TIFF file with NIH Image 1.61.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Newly Synthesized Flagellar Polypeptides Are Incorporated into Fully Grown Flagella-- Chlamydomonas cells were grown on a light/dark cycle in minimal medium to ensure that all cells were at the same point in the cell cycle. Under these conditions, the lengths of wild-type flagella did not change throughout the light period, so labeling of flagellar polypeptides occurred without flagellar growth. The optimal time to label flagella was determined by adding 35S to sulfur-starved cells at 60-min intervals following the start of the light period and measuring 35S incorporation into perchloric acid-precipitable polypeptides (Fig. 1). Labeling was greatest during the first 4-5 h of the light period, so most experiments were carried out during this period.


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  		Fig. 1.   Incorporation of 35S into perchloric acid-precipitable polypeptides. Light-synchronized wild-type (diamond ) and pf18 (black-diamond ) cells were sampled at 60-min intervals and labeled with 35S for 30 min, and protein was precipitated. Radioactivity was measured and calculated as the percentage of maximal incorporation for each time point. These data were averaged from three separate experiments. In this and all subsequent figures, t = 0 is the start of the light period.

To determine if full-length flagella incorporate newly synthesized polypeptides, 35S was added to cells for 4 h, and the flagella were amputated and fractionated into detergent-soluble membrane plus matrix, KCl-soluble polypeptides, and KCl-insoluble microtubular axonemes. Polypeptides were separated by SDS-PAGE, and stained gels were analyzed by fluorography to identify labeled flagellar polypeptides. In each experiment, cells were fixed, and flagellar lengths were measured to ensure that there was no change in flagellar length during the labeling period (Fig. 2A).


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  		Fig. 2.   Incorporation of 35S in wild-type cells. A, samples of cells incubated with (open circles) or without (filled circles) colchicine were fixed, and flagellar lengths were measured on at least 50 cells at each time point. B, SDS-PAGE and fluorography. Cells were incubated with or without colchicine and harvested after a 4-h incubation with 35S. Flagella were isolated and fractionated, and polypeptides were separated by SDS-PAGE. Stained gels and fluorographs were examined to identify radioactive polypeptides in cell bodies and flagella. Newly synthesized polypeptides were prominent in detergent-solubilized extracts (M), including the major 350-kDa membrane glycoprotein (arrow), mastigonemes (M'), a major anomalously migrating protein (bracket), and several IFT proteins (I). Newly incorporated dynein heavy chains (D) were prominent in the KCl extracts (K) as high molecular weight polypeptides, radial spoke components (R), rib240 (asterisk), and tubulins (<) in detergent- and KCl-extracted axonemes (A). Tubulins were not incorporated in flagella isolated from colchicine-treated cells. C, cell bodies; M, membrane plus matrix; K, KCl-soluble protein; A, KCl-insoluble axonemes.

More than 80 35S-labeled polypeptides were resolved by SDS-PAGE (Fig. 2B). Prominent polypeptides in the membrane plus matrix fractions included a 350-kDa membrane protein, 230-kDa mastigonemes, a major anomalously migrating membrane protein (~150 kDa), several peptides comigrating with IFT proteins (83, 57/55, 46, and 20 kDa), and a 90-kDa polypeptide that comigrated with FLA10 kinesin. Prominently labeled polypeptides released from axonemes with 0.6 M KCl included high molecular weight polypeptides that comigrated with flagellar dynein heavy chains. The axonemes contained numerous labeled polypeptides, including tubulins and radial spoke polypeptides (Fig. 2B).

To ensure that flagellar 35S incorporation was due to protein exchange and not to flagellar regeneration by cells deflagellated during handling, 35S was added to cells incubated in colchicine, under conditions that prevent flagellar regeneration after deflagellation. Fully grown flagella did not shorten during colchicine treatment (Fig. 2A). Flagella isolated from 35S-labeled and colchicine-treated cells contained the same labeled polypeptides that were found in untreated cells (Fig. 2B), with the exception that no labeled tubulin was incorporated into flagella on colchicine-treated cells. These results confirm that newly synthesized protein is incorporated without flagellar growth and demonstrate that tubulin exchange is not necessary for the exchange of other flagellar polypeptides.

After a 3-h incubation with 35S, flagella were isolated and fractionated, and the amount of radioactivity in each fraction was measured as a percentage of the total labeled flagella. The membrane plus matrix fraction contained 45 ± 17% (196 ± 48 cpm/µg; four experiments) of the total labeled protein (478 ± 190 cpm/µg), while 55 ± 17% (281 ± 186 cpm/µg) of the labeled protein was in the axoneme. Of the axonemal proteins, 60% (147 ± 26 cpm/µg) were solubilized by KCl, and 40% (134 ± 41 cpm/µg) were in the KCl-insoluble, axonemal fraction.

Flagellar Protein Exchange in a High Molecular Weight Component of PF-ribbons-- In sea urchin embryonic cilia, an integral outer doublet microtubule component, tektin-A, was shown to undergo rapid turnover in steady state cilia (44, 45, 52). To examine the possibility that stable axonemal structures in Chlamydomonas also may exhibit high levels of turnover, we isolated PF-ribbons (47) from cells that were labeled for 6 h with 35S and examined their labeling pattern. As shown in Fig. 3, PF-ribbons were composed of three filaments and five major polypeptides. Neither alpha  or beta  tubulins nor the rib43a polypeptide was significantly labeled. By contrast, two high molecular weight polypeptides (>240 kDa) were replaced by labeled polypeptides during the labeling period. Most of the label (42%) was incorporated into a single 240-kDa polypeptide, rib240 (Fig. 3A). To confirm that the labeled protein was an axonemal component, a polyclonal antibody was generated against rib240 (Fig. 4A). The antiserum stained the entire length of demembranated axonemes (Fig. 4B), confirming that it is a linear axonemal component.


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  		Fig. 3.   Incorporation of labeled polypeptides into PF-ribbons. Cells were incubated with 35S for 6 h, and PF-ribbons were isolated from the outer doublets. A, SDS-PAGE of PF-ribbons, stained with Coomassie Blue (S) and 35S-labeled ribbon polypeptides (P) analyzed with a PhosphorImager (Molecular Dynamics, Inc., Sunnyvale, CA). B and C, negatively stained fractions of PF-ribbons in low (B) and high (C) magnification. Bar, 100 nm.


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  		Fig. 4.   Identification of rib240 in flagella. A, PF-ribbons were separated in SDS-PAGE and transferred onto a nitrocellulose membrane. The membrane was probed with a preimmune serum (a) and a postimmune serum containing a polyclonal antibody against rib240 (b). B, immunofluoresence staining of isolated axonemes using anti-beta -tubulin antibody (a), buffer alone (b), and a polyclonal antibody against rib240 (c).

Turnover Rate of Flagellar Polypeptides-- To examine the incorporation rate of polypeptides into steady state flagella, light-synchronized wild-type cells were incubated with 35S with or without colchicine, and flagella were isolated from equal numbers of cells at 1- or 3-h intervals. In both cases, 35S incorporation into flagella reached a maximum by 3 h (Fig. 5A). Incorporation of polypeptides into the membrane plus matrix fraction reached a maximum within 3 h, while labeling in the KCl-soluble and insoluble axoneme fractions was not saturated until 6 h (Fig. 5B), suggesting that axonemal proteins exchange with a pool of soluble proteins transported into the flagellum.


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  		Fig. 5.   Incorporation of labeled flagellar polypeptides. A, wild-type cells were incubated with 35S for a continuous 12 h in the absence (open circles) or presence (filled circle) of colchicine. Flagella were isolated from 1.25 × 109 cells/h, flagellar polypeptides were precipitated by perchloric acid, and radioactivity was quantified. Radioactivity was calculated as a percentage of the maximal incorporation and averaged from three separate experiments. Error bars show the S.D. from three separate experiments. B, SDS-PAGE and fluorograms of flagella isolated at 3-h intervals during a 12-h incubation and fractionated into membrane plus matrix, KCl-soluble fraction, and the KCl-insoluble axonemes.

The exchange rate of individual polypeptides did not vary during the time of labeling. Synchronized cells with fully grown flagella were labeled for 3, 6, and 9 h, and flagella were isolated, fractionated on SDS-PAGE, and fluorographed. Each gel lane contained identical amounts of protein. Fluorographs were scanned, using a calibrated scanner with a 3.4 dynamic range, and the density of each peak was determined using NIH Image. Of the KCl-soluble protein, two high molecular weight dynein heavy chains contained 60% of the 35S. Of the KCl-insoluble axoneme fraction, alpha - and beta -tubulins contained 6% of the 35S, and three radial spoke proteins contained 3, 3, and 2% of the 35S (Fig. 5). Each of the polypeptides contained the same proportion of 35S when quantified after 3-, 6-, and 9-h labeling periods.

If exchange with newly synthesized protein is continuous, then labeled flagellar polypeptides should decrease after 35S removal. Cells were labeled for 3 h to achieve maximal incorporation and were then transferred to medium containing unlabeled sulfur. After 35S removal, labeled flagellar polypeptides decreased over 3 h (Fig. 6A). It was estimated that all labeled polypeptides would be chased from the flagella within 6 h. In cells labeled for 6 h and chased for 6 h, unlabeled polypeptides replaced most of the labeled polypeptides (Fig. 6B).


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  		Fig. 6.   Chase of incorporated polypeptides in wild-type cells. A, cells were labeled for 3 h and chased for another 3 h in unlabeled sulfur medium. Flagella were isolated and precipitated by perchloric acid. Radioactivity at each time point was calculated as the percentage of radioactivity after a 3-h incubation with 35S. The arrow indicates the time point when label was replaced by cold sulfate. B, cells were transferred to regular M medium after a 6-h incubation with 35S (-6). Flagella were isolated and fractionated after 1 (1) or 6 h (6).

To What Extent Are Flagellar Polypeptides Replaced?-- To determine the extent of flagellar protein exchange, we compared the amount of 35S incorporated into fully assembled flagella over a 6-h period (during which time the maximum amount of exchange occurred) with the amount of 35S incorporated into flagella after cells were deflagellated and incubated in 35S for 2 h, as flagella grew to the length of steady state labeled flagella. Labeling of regenerating cells was carried out for 2 h to minimize any incorporation due to protein turnover once the flagella reached full length.

Axonemes isolated from fully grown flagella contained 19% of the 35S incorporated in axonemes in regenerating flagella (Table I). The membrane plus matrix in fully grown flagella contained 20% of the label incorporated into regenerating flagella. Thus, during a 6-h labeling period, ~40% of the flagellar polypeptides exchanged with newly assembled polypeptides.

                              
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Table I
Comparison of labeled proteins in steady state flagella with total flagellar proteins

The incorporation of 35S into 18 individual polypeptides also was examined. The exchange rate of individual polypeptides varied significantly (Fig. 7 and Table II). Some polypeptides, including the membrane polypeptides M1 and M4, were completely exchanged with newly synthesized proteins, while another polypeptide (M2) exhibited little significant exchange. In the axonemes, some polypeptides, such as A2, A3, A4, and A5, exchanged completely, while others, including A1 and A8, showed no significant exchange. Polypeptides A6 and A7 are likely alpha  and beta  tubulins, because the incorporation of these labeled polypeptides is abolished when cells are incubated with colchicine. Thus, exchange of axonemal polypeptides is not uniform.


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  		Fig. 7.   Coomassie Blue-stained SDS-PAGE showing selected polypeptides that were sliced from the gel for radioactivity analysis, shown in Table II. Membrane plus matrix (M), axonemes (A), and PF-ribbons (R) are shown. A6 and A7 comigrated with tubulins. R2, rib240.

                              
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Table II
Comparison of labeled protein in steady state and regenerating flagella

One of the major polypeptides that was exchanged in nongrowing flagella is rib240 (R2 in Table II), a component of the PF-ribbon complex, the most insoluble component of microtubule walls. During a 6-h labeling period, 3% of the 35S incorporated into axonemes was found in the PF-ribbons, which comprise less than 2% of the total axonemal protein. Most 35S in PF-ribbons was found in a single polypeptide, rib240 (Fig. 3). rib240 composed 0.1% of total flagellar protein and 9% of the PF-ribbon complex, but this minor flagellar protein also incorporated 42% of the 35S incorporated into PF-ribbons and 1% of the total flagellar 35S. Incorporation of 35S into rib240 is easily visualized in fluorograms of Coomassie Blue-stained gels in which rib240 cannot be detected (Fig. 2, asterisk).

Flagellar Proteins Exchange as Flagella Disassemble-- Is the exchange we observe solely located in the relatively labile central microtubules (54)? To examine this, pf18, a Chlamydomonas "9 + 0" mutant that lacks central microtubules, was labeled with 35S and compared with 35S incorporation into 9 + 2 wild type flagella. As shown in Fig. 8B, the same major labeled polypeptides were found in the axonemes and in the detergent- and salt-soluble fractions of pf18 flagella and in wild-type flagella. Thus, turnover does not occur in the central microtubules alone but, rather, in all of the axonemal microtubules and associated proteins. More important, these data revealed that subunit exchange occurred as flagella shortened. During the labeling period, pf18 cells were incubated under low red light, which induces flagellar shortening (13) (Fig. 8A). Despite shortening by ~14%, labeled polypeptides were found in each of the flagellar compartments.


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  		Fig. 8.   35S labeling in pf18 cells. A, cells were incubated with (filled circles) and without (open circles) colchicine during the labeling period, cells were sampled and fixed, and flagellar lengths were measured as described in Fig. 2. B, cells were labeled as described in Fig. 2, and cell bodies (C) were compared with isolated flagella fractionated into detergent-soluble membrane plus matrix (M) and detergent-insoluble axonemes (A). A 350-kDa protein (arrow), mastigoneme (M'), IFT proteins (I), radial spokes (R), dyneins (D), and tubulins (<) were labeled.

Since flagellar subunit exchange occurred in shortening pf18 flagella, we examined exchange in more rapidly shortening flagella, using the temperature-sensitive flagellar assembly mutant fla10. Synchronously cultured fla10 cells were labeled for 2 h at 20 °C, at which flagellar length was constant, or for 2 h at 33 °C, at which flagella shortened to half-length during the labeling period (Fig. 9A). Flagella isolated from 20 °C cultures contained the same labeled polypeptides found in wild type cells (Fig. 9B). As flagella shortened, new polypeptides continued to be added to the flagellar membrane plus matrix, salt-soluble components, and the salt-insoluble axonemal microtubules. The rate of incorporation of newly synthesized proteins into shortening flagella was reduced to ~25% that of control cells at 20 °C (Table III). However, incorporation of 35S into total cell body protein also decreased to ~24% of the control level (605 cpm/µg versus 2529 cpm/µg). Since the reduction of newly incorporated flagellar protein matched the decrease in 35S incorporation into total cell body proteins, it appears that flagellar polypeptide turnover continues to occur as flagella disassemble at nearly the same rate as it occurs in nonshortening flagella.


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  		Fig. 9.   Incorporation of 35S in fla10 at 20 °C (filled circles) or 33 °C (diamond ). A, flagellar lengths during a 2-h labeling period. B, analysis of labeled polypeptides in cell bodies (C), membrane plus matrix (M), KCl-solubilized protein (K), and KCl-insoluble axonemes (A) when labeling was carried out at the two temperatures.

                              
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Table III
Radioactivity of labeled proteins in each flagellar compartment after fla10 cells were incubated with 35S for 2 h
Equal numbers of cells were used at each temperature.

Flagellar disassembly in fla10 is due to a temperature-sensitive mutation in a kinesin-II microtubule motor required for IFT (15, 22). Since IFT continues for up to 90 min after the temperature shift (22), the labeling study described above was carried out while IFT continued. To determine if protein turnover continues after IFT cessation, fla10 cells were incubated at 33 °C for 90 min, 35S was added, and cells were incubated for an additional 2 h. During this time, flagella shortened from 12 to 4.6 µm. When the short flagella were isolated, no significant labeling was detected (data not shown). However, incubation at 33 °C also reduced 35S incorporation into cell body protein by 90% (from 2529 to 251 cpm/µg). Whether incubation at 33 °C resulted in an inhibition of 35S uptake into cysteine or inhibition of protein synthesis was not determined. The lack of incorporation of 35S into protein made it impossible to determine if turnover occurred after IFT had completely stopped.

Does Exchange Occur in Long Flagella?-- Chlamydomonas lf mutants have flagella up to 3 times longer than do those on wild type cells (17). Since assembly requires transport of material to the flagellar tips, we considered the possibility that the extra long flagella may exhibit slower turnover than that observed in shorter flagella. Synchronously cultured lf4-6 cells were labeled with 35S for 4 h. Flagella were isolated and fractionated, and SDS-polyacrylamide gels were examined by fluorography (Fig. 10A). In these experiments, lf4 flagella were nearly twice as long as wild-type flagella, but the incorporation of labeled polypeptides in the membrane plus matrix, KCl-soluble, and KCl-insoluble axonemes occurred at essentially the same rate in wild type and lf4 flagella (Table IV). On average, long flagella incorporated label at up to 4 times that of wild type cells, but the lf4 cell bodies incorporated 35S at a rate 5-fold higher than wild type cells. When compared with the amount of 35S uptake into cell bodies, lf4 axonemes and membrane plus matrix exchanged the same polypeptides at the same rate as did wild type cells (Fig. 10B).


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  		Fig. 10.   35S incorporation in lf4-6. A, cells were labeled for 4 h, and flagella were isolated from the cell bodies (C) and fractionated into the membrane plus matrix fraction (M), KCl-soluble dyneins (K), and the KCl-insoluble axonemes (A). Flagella on these cells averaged 21 µm. B, percentage of labeled flagellar proteins in cell bodies.

                              
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Table IV
Comparison of labeled proteins in lf4 and wild type cells after a 4-h incubation with 35S

Protein Exchange in Basal Bodies-- Flagellar outer doublet microtubules are continuous with basal bodies, so it was of interest to determine if basal bodies underwent protein turnover during the light period, when no new basal body assembly occurs (55). Cells were labeled with 35S for 2 and 6 h, and basal bodies were isolated and fractionated by SDS-PAGE. Numerous polypeptides were labeled, some of which comigrated with flagellar polypeptides and may be components of the axoneme (Fig. 11A, dots). Some polypeptides were not labeled during a 2-h period but were labeled after a 6-h labeling period (Fig. 11A, asterisks). Labeled polypeptides were also found in basal bodies if cells were labeled for 1-3 h and then chased with cold sulfate for 3-4 h (data not shown). Thus, the basal bodies, like flagella, exchange subunits with newly synthesized polypeptides in the absence of new basal body assembly. Whether these polypeptides exchange directly from a soluble cytoplasmic pool or enter from the flagellar compartment remains to be discovered.


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  		Fig. 11.   Incorporation of labeled polypeptides in basal bodies. A, wild-type cells were labeled for 2 and 6 h. Flagella were fractionated into the membrane and matrix fraction (M) and the detergent-insoluble axonemes (A). Basal bodies (BB) were isolated to examine if they incorporated labeled polypeptides. Several polypeptides were labeled after a 2-h incubation (dot), while a few bands were found labeled by 6 h (asterisk). B, transmission electron micrographs show basal bodies after labeling. Bar, 200 nm.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Previous studies of sea urchin embryos provided important insight into the dynamics of protein turnover in cilia (44, 45). Early studies of Chlamydomonas flagella (11, 38-41) and Tetrahymena (42) cilia indicated the possibility of protein turnover. However, the reported turnover was low, and it was possible that labeled polypeptides found in isolated flagella and cilia could have been due to new protein synthesis induced during flagellar assembly in a small proportion of the cells and not protein turnover in steady state flagella. Remillard and Witman (43) reported the labeling of a nontubulin 55-kDa flagellar polypeptide in pulse-labeled flagella, and Marshall and Rosenbaum (36) reported the incorporation of epitope-tagged tubulin in nongrowing flagella, but the exchange of other flagellar components was not examined. Recent discoveries of the importance of the IFT motors kinesin-II and cytoplasmic dynein in flagellar assembly and maintenance in a variety of organisms (16, 21-30) combined with earlier studies showing that flagellar growth and/or shortening can be induced by light (13), various drugs and ions (12-14), and temperature (15, 24), prompted a careful examination of protein turnover in steady state Chlamydomonas flagella.

Our experiments confirm that more than 80 different polypeptides are exchanged with a cytoplasmic protein pool in steady state flagella. However, many polypeptides are not exchanged in these flagella, and the turnover of some components is not necessarily linked to turnover of other components. For example, tubulin incorporation was blocked by colchicine without effecting the exchange of most of the other axonemal and membrane plus matrix polypeptides. It is possible that the lack of labeled tubulin incorporation in the presence of colchicine was due to the lack of new tubulin synthesis, due to tubulin autoregulation (56). However, we also tested our colchicine to ensure that the batch and concentration of colchicine used in each experiment completely blocked flagellar regrowth after deflagellation. Even if colchicine blocked new tubulin synthesis, there would be a sufficient pool of tubulin to assemble half-length flagella (11), so we are confident that the turnover observed in the presence of colchicine occurred in the absence of microtubule assembly.

Polypeptides associated with the most insoluble component of the axoneme, the PF ribbons (44, 47, 52, 53), were labeled, but rib43a and a 70-kDa ribbon polypeptide were lightly labeled. Tubulins were not labeled in the ribbon fractions, in contrast to the significant labeling of tubulin found in sea urchin embryonic ciliary ribbons (44, 52). The lack of labeling of individual polypeptides might reflect low numbers of cysteine or methionine residues, but tubulin is prominently labeled in the axonemes and is rich in each of these amino acids. Rib43a lacks cysteines but contains 10 methionine residues (47), so the lack of rib43a labeling in our experiments is unlikely to be due to a lack of 35S incorporation due to its amino acid composition. rib240 exhibits a high level of exchange in Chlamydomonas, and, after a 6-h labeling period, it contained 1% of the total flagellar 35S while comprising less than 0.1% of the axonemal protein. It will be interesting to further characterize this protein and to determine why it exchanges so rapidly and how it moves in and out of the insoluble and, presumably, stable PF-ribbons. The lack of tubulin exchange in PF-ribbons in Chlamydomonas but the presence of high levels of tubulin exchange in sea urchin junctional protofilaments may reveal differences in length regulation or in the positions of these insoluble flagellar components in the doublet microtubules.

Based on 35S incorporation, 20% of the exchangeable flagellar protein is replaced in steady state flagella within a 6-h period. On synchronized cells, flagella remain at constant length for greater than 12 h, so nearly 50% of the flagellar proteins would be expected to undergo replacement during the normal 12-h light period for Chlamydomonas. This is a low estimate of the turnover that occurs in vivo, because flagellar protein synthesis is relatively low in fully flagellated cells (57), and significant turnover would be expected to occur with unlabeled flagellar polypeptides synthesized before the addition of 35S in our experiments. New synthesis is not required to maintain flagella, because cells can be maintained in the presence cycloheximide or LiCl, both of which block protein synthesis required for flagellar regeneration, without any detectable changes in flagellar length or motility (13). Protein exchange also exceeds the rate at which flagella shorten, since fla10, at restrictive temperatures, and pf18, with reduced red light, incorporate newly synthesized flagellar proteins, including tubulin, at essentially the same rates as control cells even as the flagella shorten at rates up to 3.5 µm/h.

We expected that flagellar turnover would occur more slowly or not at all in mutants with exceptionally long flagella, but we discovered that proteins turn over at the same rate or, in some experiments, at a slightly faster rate than wild-type flagella. Flagellar protein exchange might occur throughout the flagella, in which case flagellar length would not be relevant to the rate of exchange. Alternatively, signals that regulate flagellar growth or maintenance might be more active in long flagellar mutants, and these might increase the rate at which new proteins are transported to the flagellar tips. The latter explanation is consistent with the rapid shortening of long flagella in dikaryons formed during mating of lf mutants with cells with normal length flagella (17), which suggests that one or more of the mutations responsible for long flagella may be in a signaling system.

Signals that regulate the addition or removal of tubulin and, therefore, flagellar microtubule length are probably independent of those that regulate flagellar protein turnover. Although colchicine blocked flagellar growth and did not stimulate shortening, it did not affect the exchange of other axonemal or membrane plus matrix components. Colchicine induces flagellar shortening in pf18 cells treated with low levels of red light and in pf18 and wild-type cells treated with cytochalasin D (14), but these appear to reflect unique properties of pf18 (13, 14) and a yet unexplained effect of cytochalasin D on flagellar length. Axonemal protein exchange may make the axonemes more responsive to signals that regulate length, but turnover alone does not appear to drive flagellar length changes.

Where does flagellar protein exchange occur? It is well established that flagellar growth occurs by the addition of tubulin at the distal tips of flagella (11, 19), in association with the microtubule-capping structures that link the microtubules to the membrane (58-61). In fully grown flagella, new tubulin also is added to the distal tips of each microtubule (36). Radial spokes are added to the distal tips, and the addition of spokes to the axonemes proceeds proximally (19). Each of these results supports the existence of tubulin treadmilling (62) in the axoneme, and Marshall and Rosenbaum (37) have presented evidence that retrograde flux of epitope-tagged tubulin is balanced by IFT.

Our data are consistent with a treadmilling model with several important exceptions. First, tubulin exchange or treadmilling in the flagellum must occur independently of the turnover of other axonemal components, because most exchangeable axonemal components continue to turnover when tubulin addition is blocked by colchicine under conditions in which colchicine blocks flagellar regeneration after amputation. Moreover, if tubulin exchange occurs at the distal microtubule tips, one would expect that colchicine would lead to flagellar shortening; tubulin would be removed from one end of a microtubule, but no new tubulin could be added at the distal tip. We reported that colchicine increases flagellar shortening in a Chlamydomonas mutant, pf18, incubated under reduced red light (13, 14) and that colchicine induces flagellar shortening in cytochalasin D-treated pf18 and wild type Chlamydomonas cells (14). In each of these experiments, colchicine had no effect on wild type Chlamydomonas cells incubated under normal light. Based on our cytochalasin D results, we first proposed that flagellar microtubules are dynamic and undergo substantial tubulin exchange at the tips (14). However, since colchicine does not lead to significant flagellar disassembly, except in an unusual example (13, 14), it is unlikely that the maintenance of flagellar length is solely due to tubulin exchange being balanced by IFT.

IFT (and the kinesin-II and cytoplasmic dynein motors that drive it) is essential for the growth and maintenance of flagellar length. Disruption or inhibition of kinesin II blocks ciliary assembly in sea urchin blastulae (21), Tetrahymena (27), Caenorhabditis elegans (23-26), and mammalian embryos (29, 30). In Chlamydomonas, mutations of kinesin-II or of cytoplasmic dynein and associated proteins prevent flagellar assembly or make it impossible to maintain flagella (16, 31-33). The role of IFT in flagellar protein turnover is unclear, since data reported here revealed that Chlamydomonas cells that contain normal level of kinesin-II still resorb their flagella at restrictive temperature (15). Data reported by Piperno et al. (63) and our data clearly show that flagellar protein turnover continues to occur in fla10 kinesin-II mutants at restrictive temperatures.

In our experiments, 35S incorporation into flagellar proteins continued to occur as fla10 cells were maintained at restrictive temperature. Flagella shortened during this period, although IFT continues for 60-90 min after shifting cells to restrictive temperature (22). We attempted to label cells by adding 35S to cells after 60 or 90 min at restrictive temperature, but these cells failed to incorporate 35S into protein. These experiments do, however, clearly show that IFT alone is not responsible for flagellar protein exchange. This increases the importance of identifying the specific components carried by IFT, because IFT is essential for flagellar length control, and IFT probably carries signals that regulate length, not simply mediate polypeptide exchange. IFT components are beginning to be identified (23, 33, 63), and their characterization will probably reveal the functions of IFT. The recent discovery that cytoplasmic kinesins can carry membrane-associated signal transduction components to the tips of neurons (64) suggests that similar signaling components may be carried to the flagellar tips by the kinesin motors and that these components will regulate ciliary assembly. Roles for signaling systems in flagellar growth control have been proposed (13, 18), and it will be interesting to determine the roles of these or other signaling components in flagellar assembly regulation.

Taken together, these results confirm that protein exchange or turnover occurs in flagella and refocuses the topic of flagellar length regulation from one involved with simple assembly or growth to one of flagellar microtubule length maintenance by a cell. Future studies will reveal more about the motors recently discovered to move components within flagella and on mechanisms that regulate the addition, removal, or exchange of specific microtubule components. The targets of regulation probably will be associated with the microtubule cap structures at the flagellar assembly sites and on the newly discovered rib240 component of the insoluble axonemal PF-ribbons.

    ACKNOWLEDGEMENTS

We thank Dr. Ray Stephens for critically reviewing and providing valuable suggestions about the manuscript as well as providing the intellectual stimulus for examining the dynamics of steady state flagella. We also thank Dr. Richard Linck for providing rib43 antibodies and Dr. Richard Himes for tubulin antibodies.

    FOOTNOTES

* This work was supported by National Institutes of Health Grant GM32556.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed. Tel.: 785-864-3490; Fax: 785-864-5321; E-mail: wdent@ukans.edu.

Published, JBC Papers in Press, May 30, 2001, DOI 10.1074/jbc.M103184200

1 W. L. Dentler, unpublished observations.

    ABBREVIATIONS

The abbreviations used are: M medium, minimal medium; PAGE, polyacrylamide gel electrophoresis; PBS, phosphate-buffered saline.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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