Abstract
Motile cilia are microtubule-based organelles that play important roles in most eukaryotes. Although it is known that microtubules in cilia are sufficiently stable to withstand their beating motion, it remains unknown how they are stabilized while serving as tracks for axonemal dynein and intraflagellar transport. To address this question, we identified a new class of microtubule-associated proteins, named FAP45 and FAP52, in Chlamydomonas. These proteins are conserved among eukaryotes with motile cilia. Using cryo-electron tomography (cryo-ET) and high-speed atomic force microscopy (HS-AFM), we established that lack of these proteins leads to a loss of inner protrusions in B-tubules and less stable microtubules. These inner protrusions are located near the inner junctions of doublet microtubules and lack of FAP45, FAP52, and FAP20 results in detachment of the B-tubule from the A-tubule, as well as flagellar shortening. These results demonstrated that FAP45 and FAP52 bind to the inside of microtubules and stabilize ciliary axonemes.
Introduction
Cilia and flagella are microtubule-based organelles that operate as both antennae and propellers in eukaryotic cells. The structures and related genes of cilia and flagella are well conserved among eukaryotes. Cilia are classified as either non-motile or motile. Non-motile cilia, or “primary cilia”, function as antennae and are involved in signal transduction through the hedgehog, Wnt, and Ca2+ signaling1. Motile cilia and flagella beat at 20-60 Hz2, 3 and drive cellular motility and fluid flow. Since the cells in most tissues are ciliated, ciliary dysfunction leads to various types of human diseases termed ciliopathies, which include hydrocephalus, situs inversus, retinal degeneration, and nephronophthisis4. Furthermore, recent studies revealed that the assembly of cilia is significantly decreased in several types of tumors5, 6, suggesting a close relationship between cilia and tumorigenesis.
The axoneme is the core structure of cilia and flagella and is highly conserved from protists to vertebrates. The axoneme is composed of a central pair of microtubules cylindrically surrounded by nine doublet microtubules (DMTs). This arrangement is often referred to as the 9+2 structure (Fig. 1a). In DMTs, 10 protofilaments of the B-tubule are attached to 13 protofilaments of the A-tubule at the inner and outer junction (Fig. 1a). In motile cilia and flagella, structures essential for motility, such as axonemal dyneins, radial spokes, and the nexin-dynein regulatory complex (N-DRC), are arranged on DMTs with a 96-nm repeating unit7-9. Axonemal dyneins bound on A-tubules slide on the neighboring B-tubules and this sliding propagates along the axoneme, generating a bending force. The activity of dyneins is regulated by the interaction between the radial spokes and the central pair of singlet MTs10-12.
Recent advances in cryo-ET techniques have dramatically revealed structural details of DMTs, yet a significant question remains—What stabilizes DMTs? Cytoplasmic MTs frequently bend with various degrees of curvature13, 14 and this bending occasionally induces MT breakage15. Furthermore, a recent study demonstrated that cytoplasmic MTs were damaged even after bending only several times16. Therefore, to ensure a high beat frequency, the DMTs of motile cilia and flagella should be structurally robust compared with cytoplasmic MTs. However, the mechanism that provides DMTs with such robustness remains to be clarified.
Nicastro and colleagues have provided insights into the structural basis of DMTs using cryo-ET of Chlamydomonas flagella. They reported periodic high densities on the inner surfaces of A-tubules and B-tubules, which they named microtubule inner proteins (MIPs, Fig.1a) 7, 17. MIPs have also been observed in the axonemes of higher organisms 7, 17-20, implying that these inner structures are essential for the integrity of DMTs in motile cilia and flagella.
In this paper, we explored the mechanisms stabilizing DMTs using Chlamydomonas mutants and various techniques, including cryo-ET and HS-AFM. We found that two uncharacterized flagellar-associated proteins (FAP), FAP45 and FAP52, are essential for the stability of B-tubules. Lack of both FAP45 and FAP52 leads to the loss of MIPs in B-tubules, resulting in disruption of the MT walls. Furthermore, the B-tubule wall detached from the A-tubule at the inner junction, and flagellar shortening was observed when both FAP52 and FAP20 were absent. These results indicate that the B-tubule is reinforced by microtubule inner proteins and its stability is essential for maintaining the structure of the axoneme.
Results
Identification of proteins essential for DMT stabilization in flagella
To identify proteins that stabilize DMTs, we searched the Chlamydomonas flagellar proteome database21 using the assumption that these proteins are (1) abundant in the axoneme fraction, (2) tightly bound on the axoneme even in the presence of high salt, and (3) highly conserved among ciliated organisms. The proteins that satisfy these criteria are listed in Supplementary Table S1. We focused on FAP45, an uncharacterized protein whose peptides were most frequently found in the proteomic analysis (Table S1, “total unique peptide” and “Axo” columns). FAP45 is a coiled-coil protein composed of 501 amino acids, has a predicted molecular mass of a ~59 kDa, and is conserved among organisms with motile cilia. The human ortholog of FAP45 is coiled-coil domain-containing protein 19 (CCDC19), also known as NESG1. The transcript of this protein is enriched in the nasopharyngeal epithelium and trachea22. However, the functions of FAP45/CCDC19 are totally unclear and thus we focused on FAP45.
We first investigated the partner that interacts with FAP45 on the axoneme using chemical crosslinking. We treated isolated wild type axonemes with 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC, zero-length crosslinker). The crosslinked products were solubilized and immunoprecipitated with a polyclonal anti-FAP45 antibody. Tubulin was identified as the major crosslinked partner of FAP45 using western blot (Fig. 1c, open arrowhead; data not shown), suggesting a direct interaction between FAP45 and tubulin. In addition, a mass spectrometric analysis revealed that the ~130 kDa product is composed of FAP45 and FAP52 proteins, probably in a 1:1 ratio (Fig. 1c and d, filled arrowhead; Supplementary Table S2).
FAP52 is the Chlamydomonas ortholog of human WD40 repeat domain 16 (WDR16) and is listed in Supplementary Table S1. It was previously shown that WDR16 knockdown in zebrafish led to severe hydrocephalus23. Furthermore, a recent study reported that homozygous deletion of the WDR16 gene in human leads to situs anomalies, which are typical phenotypes of ciliopathy24. However, the function of the FAP52/WDR16 protein in flagellar motility and axonemal structure remains unclear. Therefore, we hypothesized that FAP45 and FAP52 proteins play important roles in stabilizing DMTs.
Isolation and characterization of Chlamydomonas FAP45 and FAP52 mutants
We used Chlamydomonas mutants lacking FAP45 and FAP52 to investigate their function. The mutants were isolated from ~10,000 clones in an insertionally-mutagenized Chlamydomonas library25. In these mutant strains, the ~1.8 kbp aphVII fragment was inserted into the 5’UTR of the FAP45 gene or the first exon of the FAP52 gene (Supplementary Fig. S1b). These strains did not express detectable FAP45 or FAP52 protein as analyzed by a western blot of axonemal fractions (Fig. 1b, Supplementary Fig. S1c). On the other hand, fap45 and fap52 axonemes retained the known major axonemal components, such as outer arm dyneins, inner arm dyneins, radial spokes, and the dynein regulatory complex (N-DRC) (Fig. 1b). Despite FAP45 and FAP52 being crosslinked by a zero-length crosslinker, they localized on the axoneme independent of each other (Fig. 1b). Consistent with the mass spectroscopic analysis, the ~130 kDa crosslinked product of FAP45 and FAP52 was not detected in the crosslinked fap45 or fap52 axoneme (Fig. 1c and d, filled arrowheads), suggesting that FAP45 and FAP52 are neighbors on the axoneme.
Next, we examined the motility phenotype of fap45 and fap52. The swimming velocity and beat frequency of fap45 cells were slightly reduced (Fig. 1e, Movie 1), whereas the phenotype of fap52 was quite similar to that of wild type (Fig. 1f, Movie 1). Intriguingly, the double mutant fap45 and fap52 (fap45fap52) swam significantly slower than wild type, with a low beat frequency (Fig. 1g, Movie 1), even though there was no decrease in the major known axonemal components essential for flagellar motility (Fig. 1b). Furthermore, there was an accumulation of non-motile fap45fap52 cells under confluent culture conditions (Movie 2). Since the medium under confluent culture conditions is more viscous than that of log phase conditions, a viscous load may affect the motility of fap45fap52. To explore this possibility, we observed the swimming behavior of wild type and fap45fap52 log phase cells in this viscous medium. Most of the wild type cells swam slowly but smoothly (Movie 3) whereas many of the fap45fap52 cells stopped swimming or struggled to swim against the viscous load (Movie 3), indicating that fap45fap52 flagella cannot beat properly under these conditions. Since the fap45fap52 axonemes had normal levels of axonemal dyneins, radial spokes, and N-DRC, these results suggest that the lack of both FAP45 and FAP52 causes structural defects in the DMT.
FAP45 and FAP52 are luminal proteins in B-tubules
We biochemically localized FAP45 and FAP52 in DMTs by fractionating the axonemal proteins using increasing concentrations of sarkosyl26 (Supplementary Fig. S2a). FAP45 began to be extracted from the pellet at 0.2% sarkosyl, half of the FAP45 was extracted at 0.3%, and the protein was completely extracted from the pellet at 0.7% (Supplementary Fig. S2b). FAP52 started to be solubilized at 0.3% sarkosyl and almost all the protein was solubilized at 0.7%, although a small amount of protein still remained in the pellet. These behaviors correlate well with solubilization of the B-tubule.
To distinguish whether FAP45 and FAP52 are located outside or inside the B-tubule, we prepared axonemes containing biotinylated FAPs and tested whether or not the proteins are accessible to streptavidin. These axonemes were purified from rescue strains expressing FAP proteins whose N- or C-terminus was fused to biotin carboxyl carrier protein tag (BCCP tag, Fig. S2b and c) 27, 28. However, essentially no signal of biotinylation was detected on those axonemes (Supplementary Fig. S2d), suggesting that streptavidin access to BCCP tag was prevented, presumably by the B-tubule wall. Therefore, we treated axonemes with 0.15% sarkosyl, which partially broke the B-tubule wall and, as expected, resulted in detection of the tagged proteins (Supplementary Fig. S2e). Taken together with the results that the loss of FAP45 and FAP52 did not affect the incorporation of other axonemal proteins bound on the outer surface of the DMT (Fig. 1b), these data indicate that FAP45 and FAP52 are luminal proteins enclosed by the B-tubule.
Lack of FAP45 and/or FAP52 causes structural defects in B-tubules
To reveal the defect caused by the loss of FAP45 and FAP52, we observed the mutant axonemes by cryo-ET. Interestingly, the mutants had structural defects inside of the B-tubule (Fig. 2). For further visualization of missing structures in fap45 and fap52, we also applied Student’s t-tests to compare wild type and mutant density maps27 (Fig. 2e and f, right).
A comparison between wild type and fap45 DMTs shows that the luminal surface of wild type B-tubule is smoother than that of fap45 (Fig. 2d and e). In the fap45 DMTs, the grooves between B-tubule protofilaments B7-B9 are prominent (Fig. 2b and e; Supplementary Fig. S3e), which are similar to bare microtubules. The grooves seem to be partially filled with filamentous densities as shown in the t-map (Fig. 2e right, highlighted in green). These densities are arranged in a 48-nm repeat, and reminiscent of filamentous densities extended from MIP3 in Tetrahymena DMTs29. In both our Chlamydomonas DMT and Tetrahymena DMT28, these lateral filaments appear to connect MIP3 to protofilaments B7-B9. Therefore, we named these densities that are missing in fap45 mutant as “MIP3c” (Fig 2e right and Fig. 6a).
A comparison between the wild type and fap52 DMT revealed an arch-like density is missing in the B-tubule of the fap52 mutant. In the wild type DMT, there are a large arch-like density and a small spike-shaped density in a 16-nm repeat inside of the B-tubule on the protofilament B9 and B10, called MIP3a and MIP3b, respectively (Fig. 2a and d, highlighted in yellow and orange; Supplementary Fig. S3d)17. The DMTs of fap52 completely lacked the densities of MIP3a, whereas MIP3b appear to be unchanged (Fig. 2c and f; Supplementary Fig. S3d).
Next, we tested whether the predicted atomic structures of FAP45 and FAP52 can fit into these densities. Chlamydomonas FAP52 contains eleven WD40 domains (Supplementary Fig. S1a). Since WD40 repeat domains typically form a seven-bladed β-propeller, FAP52 most likely contains two β-propellers. Indeed, a model comprising two-propellers fit well into the density map, suggesting that MIP3a corresponds to most of FAP52 protein (Supplementary Fig. S3g). For MIP3c, predicted coiled-coils of FAP45 protein were fit into the density (Supplementary Fig. S3f). FAP45 is mainly composed of helices and coiled-coils that account for 85% of the total sequence (Supplementary Fig. S1a). The total length of helices and coiled coils in FAP45 is ~60 nm. Therefore, the circled densities in Supplementary Fig. S3f are probable one unit of MIP3c. Thus, MIP3a and MIP3c most likely consist of FAP52 and FAP45, respectively.
To investigate the reasons why the double mutant fap45fap52 cells swim significantly slower than single mutants, we observed the fap45fap52 axoneme using cryo-ET. In this case, we could not apply 3D sub-tomographic averaging because many of the axonemes were frayed and the B-tubules were partially missing (Fig. 3a and b). Since cryo-ET is not suitable for observing the shapes of individual DMTs due to the missing wedge effects, thin sections of the fap45fap52 axoneme were observed by conventional EM and revealed that the B-tubules were missing in ~33.8% of the outer DMTs (Fig. 3c). All of the protofilaments in the B-tubules were completely missing in some DMTs, whereas DMTs remaining several protofilaments in the B-tubules were also observed. These data suggest that the loss of both FAP45 and FAP52 decreases structural stability between the B-tubule protofilaments. Of note, we also found that the density of dynein e, a species of inner arm dyneins was decreased in fap45 and fap52 (Supplementary Fig. S3a-c). The density of dynein e is smaller than that of other inner arm dyneins even in wild type (Supplementary Fig. S3a), implying that binding of dynein e to the B-tubule is flexible compared to that of other dyneins. Therefore, it is possible that the lack of FAP45 or FAP52 slightly changes the shape of B-tubules and affects the interaction between dynein e and B-tubules.
Direct observation of B-tubule depolymerization by HS-AFM
The above “static” DMT structures suggested that the B-tubules of fap45fap52 are less stable than those of wild type. We directly observed the stability of the B-tubule using HS-AFM30, in which a small cantilever tip (tip diameter ~ 1-2 nm) intermittently taps the DMT surface fixed on a mica stage. This intermittent contact can eliminate frictional force during lateral scanning and damage to the DMT caused by scanning because the feedback operation maintains a constant tip-sample interaction force and the force is applied for a short time (< 100 ns).
We first observed in vitro polymerized MTs in the absence of a stabilizing agent such as taxol. It was previously reported25 that MTs depolymerize spontaneously within two or three frames (1.0 to 1.5 s), suggesting that depolymerization requires less than 50 nm/s. Actual depolymerization could be faster26 but our HS-AFM has insufficient temporal resolution to verify this.
Compared with in vitro polymerized MTs, DMTs are far more stable and do not depolymerize spontaneously under normal HS-AFM conditions. To determine how a defect influences the stability of DMTs, we made a small hole by thrusting the tip of the cantilever into the B-tubule (Time=0, Fig. 4a) and observing whether or not the damage propagates. In wild type, the hole rarely grew during the observation period (Fig. 4b, Movie 4). On the other hand, in the mutants, the hole gradually expanded in both the proximal and distal directions (Fig. 4c, Movie 5-7). Time-course analysis of the depolymerized B-tubule length demonstrated that the tendency for depolymerization followed the order as fap45fap52 > fap45 > fap52 ≥ wild type (Fig. 4d). These data strongly suggest that FAP45 and FAP52 stabilize the doublet B-tubule and prevent depolymerization induced by damage to the MT wall.
MIP3a is important for B-tubule anchoring to the A-tubule at the inner junction of DMT
Since MIP3a is located near the inner junction between the A- and B-tubules and appeared to attach to the A-tubule28, we investigated whether FAP52 is involved in stabilizing the junction. We previously reported that FAP20 is a component of the inner junction and a null mutant of FAP20 (fap20) showed an abnormal swimming phenotype, although the structure of the DMTs appeared to be normal31. Based on the hypothesis that MIP3a anchors the B-tubule to the A-tubule together with FAP20, we constructed the double and triple mutants fap20fap45, fap20fap52, and fap20fap45fap52. Most fap20fap45 cells were trembling and their motility appeared slightly worse than that of fap20 cells (Movie 8). In contrast, fap20fap52 and fap20fap45fap52 cells were essentially paralyzed (Movie 8). Furthermore, the flagellar lengths of fap20fap52 and fap20fap45fap52 deviated greatly from wild type, fap20, and fap20fap45 (Fig. 5c), with a far higher ratio of short flagella. To identify the cause of these defects, we observed the axonemes of the mutants by thin section TEM. The axonemes and DMTs of fap20fap45 appeared similar to those of fap20 (Fig. 5c and d). Interestingly, several doublet B-tubules in fap20fap52 and fap20fap45fap52 were detached from the A-tubules (Fig. 5c arrowheads; Fig. 5d) whereas no detachment of B-tubules from A-tubules was observed in fap20fap45 axonemes. Given that such defects were rarely observed in fap20 axonemes31, these results demonstrate that MIP3a is required for anchoring the B-tubule to the A-tubule at the inner junction together with FAP20, and the junction is important for stabilizing the axonemal structure.
Discussion
Ciliary/flagellar MTs are remarkably stable compared with cytoplasmic MTs. Of the “9+2” MTs, DMTs are directly stressed by the axonemal dynein-generated force and constantly bent and straightened in Chlamydomonas flagella or trachea cilia, yet their structures remain intact. The mechanism that stabilizes axonemal MTs has not been clarified. Here, we identified FAP45 and FAP52 as proteins that stabilize DMTs by binding to the inner lumen of the B-tubule. This is the first report to identify factors essential for the stabilization of ciliary/flagellar MTs.
Based on our studies and existing data, we propose a schematic model of FAP45 and FAP52 (Fig. 6b). Our cryo-ET observations show that MIP3a and MIP3c are composed of FAP52 and FAP45, respectively (Fig. 6a). FAP52/MIP3a anchors the protofilaments A13 and B10, whereas FAP45/MIP3c is bound inside of B7, B8, and B9 protofilaments. Besides, biochemical crosslinking shows that FAP45 and FAP52 directly interact with each other. Thus, these two proteins bundle B7-B10 protofilaments to reinforce the B-tubule. Consistent with this model, the B-tubules of fap45 were more vulnerable to physical stress than those of fap52 in the HS-AFM observation (Fig. 4d, Movie 6). The MIP3a structure was observed also in vertebrates7, 18, but MIP3c was not clearly described probably due to the limitation of resolution. However, the amino acid sequence of FAP45 is highly conserved from protists to mammals (Supplementary Fig. S4), suggesting that the functions of FAP45 are conserved also in higher organisms as in Chlamydomonas.
Our data also suggest that DMTs are stabilized by “fail-safe” mechanisms. The single mutants of FAP45 and FAP52 did not show significant decreases in swimming speed and only the double mutant fap45fap52 showed slow swimming and defects of the B-tubules. Although the B-tubules of fap45fap52 were more easily depolymerized than wild type B-tubules, the speed of depolymerization was slower than that of in vitro polymerized pure MTs. This suggests that B-tubules are also stabilized by other mechanisms, such as tubulin acetylation32 (Supplementary Fig. S5) or fMIPs that bind inside the B-tubule along its length29.
The inner junctions of DMTs are also stabilized by “fail-safe” mechanisms involving multiple proteins. We previously demonstrated that FAP20 is a constituent of the inner juction31. In addition to the “true” inner junction, our data clarified that MIP3a also connects B-tubules to A-tubules (Fig. 5c and d). Furthermore, some inner junctions appeared to be attached in fap20fap52 and fap20fap45fap52 mutants (Fig. 5c). In the triple mutant, additional proteins, such as tektin and PACRG, may contribute to stabilizing the inner junction, given our previous observation that these two proteins were partially retained in fap2031.
Ciliopathy caused by FAP45/CCDC19 mutation has not been reported to date. On the other hand, patients lacking FAP52/WDR16 were reported to have abdominal situs inversus and situs inversus totalis24. Despite abnormal laterality, those patients did not have typical symptoms of primary ciliary dyskinesia, such as recurrent bronchiolitis, hydrocephalus, or low nasal nitric oxide levels. These findings are consistent with the fap52 phenotype in Chlamydomonas, where we could not detect abnormal phenotypes other than the lack of MIP3a structure. Given that the phenotype of fap45 was more severe than that of fap52 in Chlamydomonas, we predict that the lack of FAP45/CCDC19 in human causes ciliopathy.
Compared to B-tubules, the functions of MIPs in the A-tubules are unclear. A recent paper clearly showed that hyper-stable “ribbons” were composed of four protofilaments to which MIP4 is bound33, suggesting MIP4 stabilizes the ribbon structure. However, no mutant lacking MIPs in A-tubules has been isolated based on the swimming phenotypes of Chlamydomonas, implying that MIPs in A-tubules might also have fail-safe mechanisms, similar to MIP3a and c.
Materials and Methods
Strains, culture conditions, and isolation of fap45 and fap52 mutants
The Chlamydomonas strains used in this study are listed in Supplementary Table S3. The double and triple mutants were constructed by standard methods3. All cells were cultured on Tris-acetate-phosphate (TAP) plates with 1.5% agar or in TAP medium. fap45 and fap52 mutants were isolated from a library of mutants generated by aphVIII gene insertion25. The mutants were backcrossed with wild type several times before use.
Motility assay using the CLONA system
To assay the motilities of Chlamydomonas cells in TAP medium or TAP with ficoll, videos were recorded using a high-speed camera (EX-F1; Casio) attached on a dark-field light microscope (BX51; Olympus) at 600 fps. The videos were analyzed using the CLONA system34 and the results were plotted using R Project software.
Antibodies
The antibodies used in this study are described in Supplementary Table S4. Anti-FAP45 and anti-FAP52 antibodies were raised against full-length FAP45 and FAP52 proteins, respectively. The full-length cDNA of FAP45 or FAP52 was cloned between the EcoRI and BamHI sites of pMAL-c2x (New England Biolabs). In both constructs, a 6×His tag was inserted into the HindIII site of pMAL-c2x to enable purification in two steps. For affinity purification, each cDNA was also cloned between the NdeI and EcoRI sites of pColdI (Takara). Expression of the recombinant proteins was induced in Escherichia coli BL21(DE3) with 0.3 mM IPTG, and almost all of the expressed protein was solubilized from each construct. MBP-FAP45-6×His and MBP-FAP52-6×His were purified with amylose resin (New England Biolabs) and then further purified with Ni-NTA agarose (Qiagen). These purified proteins were used as antigens. Each antibody was affinity purified using polyvinylidene difluoride membranes blotted with FAP45-6×His or FAP52-6×His.
Chemical crosslinking of axonemes
Isolated axonemes of wild type, fap45, and fap52 were treated with 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC, Thermo Scientific) in HMEK (30 mM Hepes-KOH, pH 7.4, 5 mM MgSO4, 1 mM EGTA, 50 mM K-acetate) for 60 min at room temperature. After quenching the reaction, crosslinked axonemes were analyzed by SDS-PAGE and western blotting with anti-FAP45 and anti-FAP52 antibodies. The crosslinked products were immunoprecipitated with anti-FAP45 antibody following a previously described method35. Mass spectrometry analysis (LC tandem MS) of the precipitates was performed at the Proteomics and Mass Spectrometry Facility (University of Massachusetts Medical School).
Immunofluorescence microscopy
Nucleoflagellar apparatus (NFA) was prepared as previously described36. After fixation with 2% formaldehyde for 10 min at room temperature, NFAs were treated with cold methanol (−20°C). The samples were immunostained as previously described37. Images were taken with a CCD camera (ORCA-R2; Hamamatsu Photonics) linked to a fluorescence microscope (IX70; Olympus).
Generation of BCCP-tagged strains
Fragments from the start codon to immediately before the stop codon in the FAP45 and FAP52 genes were amplified by genomic PCR and inserted into pIC2 plasmids38. In the FAP52 construct, a 3×HA tag was inserted into the C terminus of FAP52. Each construct was linearized and transformed into fap45 or fap52 cells by electroporation.
Streptavidin-Alexa546 staining of the axonemes was performed as previously described.31
Thin-section TEM
Samples for thin-section TEM were prepared as previously described39 except that we used 50 mM Na-phosphate, pH 7.0 instead of cacodylate buffer. The samples were observed using a transmission electron microscope (JEM-3100FEF; JEOL) equipped with a 4,096×4,096-pixel CMOS camera (TemCam-F416; TVIPS). All images were taken at 300 keV, with ~3 μm defocus, at a magnification of 40,000 and a pixel size of 2.5 Å.
Cryo-sample preparation
Purified axonemes in HMDEK buffer (30 mM Hepes-KOH (pH7.4), 5 mM MgSO4, 1 mM dithiothreitol, 1 mM EGTA, 50 mM potassium acetate, protease inhibitor cocktail (Nacalai Tesque)) were incubated with anti-beta-tubulin antibody (T0198; SIGMA) for 15 min on ice, followed by incubation with goat anti-mouse IgG (H+L) 15 nm gold (BB International) and 15 nm colloidal gold conjugated with BSA. The mixtures were loaded onto home-made holey carbon grids and plunged into liquid ethane at −180°C using an automated plunge-freezing device (EM GP; Leica).
Image acquisition
Grids were transferred into the JEM-3100FEF using a high-tilt liquid nitrogen cryo-transfer holder (914; Gatan Inc.). Tilt series images were recorded at −180°C using a K2 summit direct detector (Gatan) and the serialEM40. The angular range was ±60° with 2.0° increments. The total electron dose was limited to 100 e-/Å2 and the nominal magnification was 6,000x. An in-column energy filter was used with a slit width of 20 eV and a pixel size of 7.1 Å.
Image processing
Image processing for subtomogram averaging of DMT was carried out as previously described27, 28. Tilt series images were aligned and back-projected to reconstruct 3D tomograms using IMOD41. Alignment and averaging of the subtomograms were performed using custom Ruby-Helix scripts42 and PEET software7 to average the 96-nm repeats of DMTs. UCSF Chimera was used for isosurface renderings43.
HS-AFM observation
Flagella were demembranated with 0.5% Nonidet P-40 in HMDEK (30 mM Hepes-KOH, 30 mM Hepes-KOH, pH 7.4, 5 mM MgSO4, 1 mM DTT, 1 mM EGTA, 50 mM K-acetate, protease inhibitor cocktail (Nacalai Tesque)), followed by centrifugation for 2 min. The axoneme pellets were suspended with ATP buffer (HMDEK containing 0.1 mM ATP, 1 mM ADP and 0.5% polyethylene glycol (20,000 mol wt)) and incubated for 1 min at room temperature. Frayed axonemes were then diluted with HMDEK and placed on a mica stage. The AFM images were recorded using a home-built high-speed atomic force microscope44-46 at frame rates of 1-2 fps. All observations were performed at room temperature.
Author contribution
MO and MK designed the experiments. MO performed most of the experiments and analyzed the data. MO and MK wrote the paper. MO and TU performed AFM analyses. HY produced the BCCP-tagged constructs and helped with several EM experiments. TY, HI, and HF provided the Chlamydomonas mutant library. KW advised on chemical crosslinking experiments. TA advised on AFM analyses.
Movies 1
Swimming of wild type, fap45, fap52, and fap45fap52 in TAP medium.
Movie 2
Swimming of confluent cultured fap45fap52 in TAP medium.
Movie 3
Swimming of wild type and fap45fap52 in TAP with 7.5% ficoll.
Movie 4
HS-AFM movie of wild type DMT on a mica surface.
Movie 5
HS-AFM movie of fap45fap52 DMT on a mica surface.
Movie 6
HS-AFM movie of fap45 DMT on a mica surface.
Movie 7
HS-AFM movie of fap52 DMT on a mica surface.
Movie 8
fap20, fap20fap45, fap20fap52, and fap20fap45fap52 in TAP.
Acknowledgements
We thank Dr. John Leszyk (UMMS) for mass spec analysis and Dr. Brian Dynlacht (NYU) for helpful discussion. MO is a recipient of the fellowship from Japan Society for Promotion of Sciences for young scientists. This work was supported by CREST, the Japan Science and Technology Agency to M.K. and T.A., the Japan Society for the Promotion of Science (JSPS) KAKENHI (Grant number 25120714) and JST ALCA to H. F. The authors have no competing financial interests to declare.
References
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