The docking complex is a molecular complex necessary for assembly of outer dynein arms (ODAs) on the axonemal doublet microtubules (DMTs) in cilia and flagella. The docking complex is hypothesized to be a 24-nm molecular ruler because ODAs align along the DMTs with 24-nm periodicity. In this study, we rigorously tested this hypothesis using structural and genetic methods. We found that the ODAs can bind to DMTs and porcine microtubules with 24-nm periodicities even in the absence of the docking complex in vitro. Using cryo-electron tomography and structural labeling, we observed that the docking complex took an unexpectedly flexible conformation and did not lie along the length of DMTs. In the absence of docking complex, ODAs were released from the DMT at relatively low ionic strength conditions, suggesting that the docking complex strengthens the electrostatic interactions between the ODA and DMT. Based on these results, we conclude that the docking complex serves as a flexible stabilizer of the ODA rather than as a molecular ruler.
The beating motions of cilia and flagella are driven by the outer and inner dynein arms (ODAs and IDAs, respectively; Fig. 1A). The arrangements of ODAs and IDAs on the outer doublet microtubules (DMTs) of the axonemes are formed by highly ordered repeating structures with 24-nm and 96-nm periodicities, respectively (Fig. 1B). For IDAs, we have previously shown in Chlamydomonas reinhardtii that a molecular ruler complex made of two proteins FAP59 and FAP172 determines the 96-nm repeats (Oda et al., 2014a,b). The two proteins are Chlamydomonas homologues of mammalian CCDC39 and CCDC40, and form a 96-nm coiled-coil complex that governs both the repeat length and arrangements of IDAs, radial spokes and nexin–dynein regulatory complexes.
By contrast, Takada et al. (2002) have proposed that the 24-nm periodicity of ODAs is established by the docking complex of ODA through a similar mechanism. The docking complex comprises three proteins, DC1, DC2 and DC3 (encoded by ODA3, ODA1 and ODA14, respectively), and Chlamydomonas mutants lacking DC1 and DC2 fail to assemble ODAs in the flagella (Koutoulis et al., 1997; Takada et al., 2002; Wakabayashi et al., 2001). The recombinant protein complexes of the three subunits form a 24-nm-long complex, and the complex binds to the microtubules in a cooperative manner (Owa et al., 2014). Docking complexes enter the flagella separately from the ODAs and are thought to provide microtubule-binding sites for ODAs (Takada and Kamiya, 1994; Wakabayashi et al., 2001). However, these previous studies do not provide evidence that the docking complex is actually required for the 24-nm periodicity of ODAs.
In this study, we purified the ODAs devoid of the docking complex and found that the ODAs were able to bind to DMTs and porcine brain microtubules with 24-nm periodicity independently of the docking complex. We identified the precise positions of the subunits of the docking complex using cryo-electron microscopy and structural labeling, and found that the docking complex was flexible and localized mostly away from the microtubules. Our results suggest that the 24-nm periodicity of ODA is determined by ODA itself.
RESULTS AND DISCUSSION
The docking complex is not necessary for the 24-nm periodicity of ODA
In order to determine whether the docking complex is required for the 24-nm periodicity of the ODA, it is necessary to purify ODA particles that are devoid of docking complexes. We fractionated the high-salt extract of C. reinhardtii axonemes using a MonoQ anion-exchange chromatography column (Goodenough et al., 1987). ODAs, originally three-headed αβγ particles, were separated into two-headed αβ and single-headed γ heavy chain particles, and the docking complex was eluted between the αβ and γ peaks (the α, β and γ chains are encoded by ODA11, ODA4 and ODA2, respectively) (Fig. S1A). As we detected trace amounts of docking complex in the fractions containing αβ particles, we depleted the remaining docking complex by incubating the fractions with oda3-mutant axonemes (oda3 axonemes), which lack ODA and docking complex. Both ODA and the docking complex bound to the oda3 axonemes, but only docking complexes were depleted because there was an excess of ODA in the sample (Fig. S1B). After three rounds of depletion, we successfully obtained ODA particles that lacked detectable docking complexes (Fig. S1C and D). Note that we detected ODA using an antibody against ODA intermediate chain 2 (IC2) (D6168, Sigma-Aldrich).
We incubated ODAs with oda3 axonemes in the presence and absence of docking complex (Fig. S1E), and reconstructed the three-dimensional (3D) structures of the DMTs using cryo-electron tomography (Fig. 1C and D). Surprisingly, the purified ODA particles arranged with 24-nm periodicities along the DMT, irrespective of the presence of docking complex (Fig. 1D). The overall structures of the ODA with and without docking complex were not substantially different from one another, except for two small bumps on the outer surface of the ODA (Fig. 1D, arrowheads), suggesting that most of the densities of the docking complex were averaged out owing to structural flexibility.
In order to determine whether any other axonemal proteins, such as CCDC103 (King and Patel-King, 2015; Panizzi et al., 2012), were required for the 24-nm periodicity of the ODA, we also incubated ODAs with cytoplasmic microtubules that had been purified from porcine brains. The ODAs regularly bound to the porcine microtubules with 24-nm periodicity in the presence and absence of docking complex (Fig. 1E and F). These results suggest that the 24-nm periodicity of ODA does not require the docking complex and is, presumably, determined by ODA molecules themselves. Alignment of ODA along cytoplasmic microtubules with 24-nm periodicity has been reported previously (Haimo et al., 1979; Satir et al., 1981), but the extracted and purified ODAs used in those studies are likely to have contained docking complexes (Oda et al., 2007; Sakato and King, 2003).
Docking complexes stabilize the interaction between ODA and DMTs
We hypothesized that docking complexes reinforce the interaction between the ODA and DMT because the docking complex has a higher affinity for DMTs when compared with the affinity for the intermediate chains of the ODA (Ide et al., 2013; Owa et al., 2014). To test this hypothesis, we increased the ionic strength of the buffer used for resuspension of the ODA-reconstituted oda3 axonemes from 50 mM CH3CO2K to 100 mM NaCl. We prepared cryo-samples of the axonemes in the high-salt buffer and examined the binding of ODA to DMTs. We found that ODAs dissociated from the DMT in high-salt buffer in the absence of the docking complex (Fig. 2A and B, high salt). By contrast, ODAs appeared unchanged in high-salt buffer in the presence of the docking complex. As ODAs were extracted from the axonemes with 400–600 mM KCl or NaCl (Pfister et al., 1982; Dean and Mitchell, 2013), the dissociation of ODAs from DMTs in 100 mM NaCl indicates that the interaction between the ODA and DMT is substantially weaker in the absence of docking complex when compared with that in the presence of docking complex. Lack of ODA in the mutant strains that lack the docking complex is probably due to the reduced affinity of the ODA for DMTs.
Docking complexes localize away from DMTs
In order to visualize the interactions between the docking complex and ODA, and between the docking complex and DMT, we structurally labeled DC1 and DC2 at four positions each (Fig. 3A; Table S1, Fig. S2) using the biotin–streptavidin system (Oda and Kikkawa, 2013; Oda et al., 2014a,b), and identified their 3D positions in the axonemes (Fig. 3B,C). The densities of the labels on DC1 and DC2 appeared to be distant from the surface of DMTs, except for the labeled construct DC1-M216 (Table S1), which was located in close proximity to the microtubule-binding site of the ODA. Most of the labels on DC1 were located near to the distal edge of the ODA. These results suggest that the docking complex makes contact with the DMT in the middle segment of DC1 and that the rest of the complex binds to the outer surface of the ODA with heterogeneous conformations, rather than lying along the length of the DMT.
The electrophoresis of the biotin-tagged docking complexes appeared odd; the band representing the labeled construct DC2-M507 (Table S1) was doubled, and the mobilities of the tagged proteins were varied (Fig. S2A). As all the tagged docking complexes were properly biotinylated (Fig. S2A and B), and expression of the tagged docking complexes restored motility defects of the mutants (Table S1); we suppose that insertion of the tags into the middle of the sequence affected the behavior of the proteins during the electrophoresis and resulted in the irregular band patterns.
Outer-inner dynein linkers are not required for ODA localization
Although the 24-nm periodicity of the ODA appears to be determined by ODA itself, there must be a mechanism that keeps the 24-nm repeats in fixed positions relative to the 96-nm repeats. If the positions of the ODA 24-nm repeats were not fixed relative to the 96-nm repeats, we would be unable to visualize the structures of the two repeats simultaneously on an averaged subtomogram of the DMT. As the ODAs and IDAs are structurally cross-bridged by the outer–inner dynein (OID) linkers (Fig. S3A, arrowhead) (Nicastro et al., 2006; Bui et al., 2009; Oda et al., 2013), we suspected that the OID linkers coordinate the relative positions of the 24-nm and 96-nm repeats. We generated a triple mutant ida1pf2pf3, which lacks all the major OID linkers, and examined the structural relationship between the 24-nm and 96-nm repeats (Fig. S3A). In contrast to our expectations, the 24-nm repeats of the ODAs were in the correct positions relative to the 96-nm repeats of the radial spokes. The mechanism for the structural coordination between the 24-nm and 96-nm repeats remains to be elucidated.
Note that one of the OID linkers was not visible on the ODA-reconstituted oda3 axoneme structures regardless of the presence of docking complex (Fig. S3B, arrowheads). This result supports our conclusion above that OID linkers are not necessary for fixing the relative positions of ODAs and IDAs.
Role of the ATP-dependent microtubule-binding sites of ODA
As a single ODA complex has ATP-dependent microtubule-binding sites as well as ATP-independent binding regions (Haimo et al., 1979; Oda et al., 2007), we examined the effect of ATP on the alignment of ODAs along microtubules. We incubated ODA and oda3 axonemes in the presence of ATP and vanadate to fix the ATP-dependent microtubule-binding sites in a weak-binding state (Fig. S3C) (Shimizu and Johnson, 1983). We found sporadic binding of ODA, but alignment with 24-nm periodicity was not observed. We also incubated ODA with porcine microtubules in the presence of ATP and vanadate, but no decoration with ODA was observed (Fig. S3D). In previous studies, ODAs have been seen to sometimes align along a single microtubule with 24-nm periodicity only through the ATP-dependent binding sites, and they do not form cross-bridges (Haimo et al., 1979; Satir et al., 1981; Oda et al., 2007), suggesting that the 24-nm periodicity of ODA does not require microtubule-binding through the ATP-independent binding sites. Our results suggest that the ATP-dependent binding sites play a major role in the recruitment of ODAs to DMTs and that the inter-molecular interactions between adjacent ODAs determines the periodical alignment. Note that even one ATP-dependent binding site would suffice for microtubule binding of ODA because it has been shown that ODAs lacking two of the three ATP-dependent binding sites can bind to the axonemes in vivo (Sakakibara et al., 1993).
Possible meaning in the flexibility of docking complexes
Our results suggest that the docking complex is unnecessary to establish the 24-nm periodicity of ODAs along DMTs. Instead, the docking complex is thought to work as a flexible adaptor that stabilizes the interaction between the ODA and DMT (Fig. 4). The structure of the docking complex was first observed as a small projection on the DMT by performing thin-section electron microscopy analysis of oda6-mutant axonemes (oda6 axonemes), which lack ODAs but have docking complexes (Takada and Kamiya, 1994). However, we were unable to detect substantial docking-complex-like densities in the averaged subtomogram of oda6 axonemes (Fig. S4A) (Bui et al., 2009). We compared the thin-section electron micrographs of oda3 and oda6 axonemes. In agreement with our conclusion, we found that the shapes of the small projections representing the docking complexes on oda6 axonemes were highly variable and often invisible (Fig. S4B).
It is counter-intuitive that the regularly aligned ODAs are stabilized by heterogeneous structures. We suppose that the structural flexibility allows the docking complex to accommodate the large conformational changes occurring in the mechano-chemical cycles of the ODAs (Oda et al., 2007; Movassagh et al., 2010; Lin et al., 2014). The large and fast movements of the dynein head domains might require a structurally flexible support, rather than a rigid base, to exert forces for ciliary and flagellar beating.
MATERIALS AND METHODS
Strains and reagents
C. reinhardtii wild-type cells were grown in Tris-acetate-phosphate (TAP) medium. To screen transformants, cells were grown on TAP agar that had been supplemented with 10 µg/ml paromomycin. Strains used for this study are listed in Table S1.
Preparation of axonemes
Chlamydomonas cells were deflagellated with dibucaine-HCl, and axonemes were collected by centrifugation (Piperno et al., 1977). Flagella were de-membraned with 1% nonidet P-40 in HMDENa/K buffer composed of 30 mM Hepes-NaOH pH 7.2, 5 mM MgCl2, 1 mM dithiothreitol, 1 mM EGTA, 50 mM NaCl or 50 mM CH3CO2K.
Purification and reconstitution of ODAs
ODA particles were purified using a MonoQ anion-exchange column (GE Healthcare). Peak fractions containing ODA αβ and γ particles were combined and concentrated to a final concentration of 2 mg/ml. Triple-headed ODA αβγ particles containing docking complexes were purified using a UnoQ anion-exchange column (Bio-Rad), as described previously (Furuta et al., 2009). Peak fractions containing ODA αβγ particles were also concentrated to 2 mg/ml and used for reconstitution of docking complex (+) axonemes. For absorption and reconstitution of ODAs onto oda3 axonemes, the concentrated solutions of ODA particles were dialyzed against HMDEK buffer overnight and incubated with 50 µg of axonemes for 2 h. To test for the decoration of microtubules, ODA particles in HMDEK buffer were mixed with 0.1 mg/ml porcine brain microtubules in the presence of 10 µM paclitaxel for 1 h.
Construction of DC1 and DC2 expression vectors
Fragments spanning from the start codon to immediately before the stop codon for the genes encoding DC1 and DC2 were amplified with PCR using genomic DNA from the wild-type strain, and were then inserted into pIC2 plasmids (Oda et al., 2015). We inserted the tag sequence corresponding to amino acids 141–228 of Chlamydomonas acetyl-CoA carboxylase biotin carboxyl carrier protein (BCCP) into the middle of the sequences of DC1 and DC2.
Labeling of BCCP-tagged axonemes was performed as described previously (Oda et al., 2014a,b). Plunge-frozen grids were transferred to a JEM-3100FEF transmission electron microscope (JEOL) with a 914 cryo-transfer holder (Gatan Inc.). Tilt series images were recorded at −180°C using a TemCam-F416 CMOS camera and EM-TOOLs program (TVIPS). The angular range was ±60° with 2.0° increments. The total electron dose was limited to 100 e−/Å2. Images were recorded at 300 keV, with 6–9 µm defocus, at a magnification of ×25,700 and a pixel size of 6 Å. An in-column energy filter was used with a slit width of 20 eV.
Image processing for subtomogram averaging was performed as described previously (Oda and Kikkawa, 2013) using the IMOD software package (Kremer et al., 1996), custom Ruby-Helix scripts (Metlagel et al., 2007) and the PEET software suite (Nicastro et al., 2006). The effective resolutions were ∼4.9 nm (Fig. S2C). Surface renderings were generated using the UCSF Chimera program (Pettersen et al., 2004).
In order to identify statistically significant differences, we applied a Student's t-test to compare wild-type and streptavidin-labeled axonemes, as described previously (Oda and Kikkawa, 2013). The isosurface threshold values were t>7.17, with a one-tailed probability of <0.1%.
A part of this work was conducted in Research Hub for Advanced Nano Characterization, The University of Tokyo, under the support of ‘Nanotechnology Platform’ (project number 12024046) by Ministry of Education, Culture, Sports, Science and Technology, Japan.
The authors declare no competing or financial interests.
T.O. conceived the project; T.O., T.A. and H.Y. conducted experiments; and T.O. and M.K. drafted the manuscript.
This work was supported by CREST, the Japan Science and Technology Agency (to M.K.); the Kazato Research Foundation (to T.O.); the Takeda Science Foundation (to M.K. and T.O.); the Japan Society for the Promotion of Science KAKENHI [grant number 15642352 (to T.O.)]; and the Institute for Fermentation, Osaka (to T.O.).
The electron microscopy maps of averaged DMTs are available at the EM Data Bank (www.emdatabank.org) under the accession numbers EMD-6504 to EMD-6511.
Supplementary information available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.184598/-/DC1
- Received December 9, 2015.
- Accepted February 25, 2016.
- © 2016. Published by The Company of Biologists Ltd