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Flagellar microtubule doublet assembly in vitro reveals a regulatory role of tubulin C-terminal tails

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Science  18 Jan 2019:
Vol. 363, Issue 6424, pp. 285-288
DOI: 10.1126/science.aav2567

Assembly of the ciliary microtubule doublet

The cilium is a conserved organelle that is crucial for motility as well as for sensing the extracellular environment. Its core structure is characterized by nine microtubule doublets (MTDs). The mechanisms of MTD assembly are unclear. Schmidt-Cernohorska et al. developed an assay to reconstitute MTD assembly in vitro. Tubulin carboxyl-terminal tails played a critical inhibitory role in MTD formation. Molecular dynamics revealed that carboxyl-terminal tails of the A11 microtubule protofilament regulated MTD initiation. Furthermore, live-cell imaging showed an unexpected bidirectional isotropic elongation of the MTD.

Science, this issue p. 285

Abstract

Microtubule doublets (MTDs), consisting of an incomplete B-microtubule at the surface of a complete A-microtubule, provide a structural scaffold mediating intraflagellar transport and ciliary beating. Despite the fundamental role of MTDs, the molecular mechanism governing their formation is unknown. We used a cell-free assay to demonstrate a crucial inhibitory role of the carboxyl-terminal (C-terminal) tail of tubulin in MTD assembly. Removal of the C-terminal tail of an assembled A-microtubule allowed for the nucleation of a B-microtubule on its surface. C-terminal tails of only one A-microtubule protofilament inhibited this side-to-surface tubulin interaction, which would be overcome in vivo with binding protein partners. The dynamics of B-microtubule nucleation and its distinctive isotropic elongation was elucidated by using live imaging. Thus, inherent interaction properties of tubulin provide a structural basis driving flagellar MTD assembly.

The cilium is an organelle crucial for motility, as well as for sensing environmental cues such as signaling molecules, light, and mechanical stimuli (1). The core structure of the cilium is characterized by nine microtubule doublets (MTDs) (2). In Chlamydomonas, MTDs form a double-track railway for intraflagellar transport trains (3), which carry ciliary building blocks along microtubules during the assembly and disassembly of the cilium (4). These MTDs display distinctive structural features and are formed by a complete A-microtubule composed of 13 protofilaments and an incomplete B-microtubule of 10 protofilaments, which starts from the outer junction (OJ) between protofilaments A10 and A11 of the A-microtubule (Fig. 1A and fig. S1A) (58). Cryo–electron microscopy (cryo-EM) analysis of the Tetrahymena ciliary MTD reveals that this OJ involves noncanonical, surface-to-side tubulin-tubulin contacts (9). It also reveals an inner sheath composed of microtubule inner proteins (MIPs) inside the MTD.

Fig. 1 MTD assembly in vitro.

(A) Schematic of a procentriole with A- and B-microtubules (green) and the procentriole cartwheel structure (gray). B-microtubule branching occurs at the OJ, highlighting the protofilaments A10, A11, and B1. The dashed box corresponds to the closer view to the right. Black arrows, seam of the A-microtubule and inner junction. (B to E) Representative cryo-EM images of microtubules (MT) (B), microtubules supplemented with tubulin pretreated with subtilisin (MT+Tub_S) (C), subtilisin-treated microtubules (MT_S) (D), and MT_S incubated with tubulin (MTD) (E) with their corresponding schematics. Arrowhead, MTD. Tub, tubulin. Scale bars, 25 nm. (F) Percentage of MTD formation (three independent experiments): 0% for MT (n = 825 microtubules), 3 ± 2% for MT+Tub_S (n = 729), 8 ± 6% for MT_S (n = 1515), and 72 ± 8% for MTDs (n = 2341). Errors bars represent SD.

MTD assembly occurs at the centriolar level, with the B-microtubule nucleating and elongating bidirectionally onto the surface of the A-microtubule, as assessed by cryo–electron tomography (cryo-ET) in human centrosomes (10). However, the molecular mechanism enabling B-microtubule nucleation at the surface of the A-microtubule is unclear. C-terminal tails of tubulin may play a role (11) because their limited proteolytic digestion by subtilisin induces the nucleation of hooked microtubules and protofilament bundles (fig. S1, B and C).

We hypothesized that B-microtubule assembly could be mediated solely through tubulin-tubulin interactions. We first set out to address whether tubulin devoid of the C termini alone could assemble MTDs. We developed an in vitro assay to mimic the sequential assembly of MTDs by A-microtubule formation followed by B-microtubule nucleation. First, stable microtubules (12) were assembled (Fig. 1B) and subsequently incubated with subtilisin-treated tubulin without C-terminal tails (Tub_S) (11, 13) (materials and methods in the supplementary materials). This did not result in MTD formation (Fig. 1C). We next assessed whether the removal of C-terminal tails of the A-microtubule would promote MTD formation. Microtubules treated with subtilisin (MT_S) (13, 14) (fig. S1D) looked identical to untreated microtubules (Fig. 1, B and D). When we added free tubulin to subtilisin-treated microtubules, 72% of these microtubules formed assemblies that resembled MTDs (Fig. 1, E and F), reaching a median length of 0.66 ± 0.5 μm after 15 min (fig. S1E). By contrast, only ~7.5% of MTD-like structures were observed among microtubules treated with subtilisin alone. This possibly reflects some depolymerization of the tubulin lacking C-terminal tails; this tubulin would reattach at the surface of the A-microtubule and nucleate efficiently because of having a lower critical concentration than untreated tubulin (15) (Fig. 1F). Thus, C-terminal tails of the A-microtubule negatively regulate a noncanonical, surface-to-side tubulin interaction, allowing microtubule branching.

By using cryo-EM, we next investigated whether branching occurs at the tip or on the main body of the A-microtubule and found that B-microtubules assembled mainly on the body of the A-microtubule (fig. S2A), corroborating previous in vivo findings (10). Cryo-ET of these reconstituted MTDs showed structural similarity to the ciliary MTDs (Fig. 2, A and B; movie S1; and fig. S2B), with 23% of multiple MTDs surrounding a single A-microtubule (Fig. 2C and fig. S2C). This indicates that B-microtubule nucleation in vitro was not restricted to one protofilament of the A-microtubule and that, in vivo, additional proteins may be needed to provide positional information. By using subtomogram averaging to improve the resolution of the OJ to 17.2 Å (fig. S2D), we confirmed the typical triangular junction formed among protofilaments A10 and A11 of the A-microtubule and protofilament B1 of the B-microtubule in the reconstituted MTDs (Fig. 2, D to G, and fig. S2E) (9). Additionally, inspecting the curvature of the B-microtubule junction from individual MTDs in cryo-ET (fig. S3A) revealed a curvature similar to that observed in in vivo MTDs (Fig. 2F). This suggested that the surface-to-side tubulin-tubulin interaction at the OJ is sufficient to drive the correct angle for MTD assembly. We noticed an important mobility of the B-microtubule at the MTD inner junction, possibly because of the lack of MIPs or because of a protein such as FAP20, which closes the MTDs in cilia (Figs. 1A and 2, I to J) (16).

Fig. 2 Cryo-EM reconstruction of in vitro MTDs.

(A) Representative image of a cryo-ET section. Scale bar, 25 nm. (B) zx view of a cryo-ET section. Scale bar, 25 nm. (C) zx view of a cryo-ET section showing an MTD flower. Scale bar, 25 nm. Arrowheads in (A) to (C) indicate B-microtubules. (D and F) Subtomogram averaging of in vitro MTDs at 17-Å resolution (D) and of Tetrahymena ciliary MTDs at 5.7 Å (EMD-8528 map from the Electron Microscopy Data Bank) (F). Scale bars, 25 nm. (E and G) Closer view of the OJ for the in vitro MTD (E) and the OJ of the ciliary MTD (G). Arrowheads indicate the triangular shape of the A10, A11, and B1 protofilaments. (H) Traces of the B-microtubules starting at the OJ and highlighting the curvatures of the B-microtubules in vitro compared with those of the in vivo ciliary MTDs (n = 44 microtubules). (I and J) Plot profiles at the positions indicated by the arrows in (H) showing that the curvature of the OJ is stable (I) whereas the end of the B-microtubule is more flexible (J). The black line indicates the position of the B-microtubule in vivo. A.U., arbitrary units. (K) Side and top views of an MTD model with the A-microtubule in green and gray (α-tubulin, gray; β-tubulin, green), the tubulin C-terminal tails of the A-microtubule in red, and the protofilament B1 in blue and beige (α-tubulin, blue; β-tubulin, beige). Atoms are represented as spheres. (L) Closer view of the OJ highlighting the interactions of the tubulin C-terminal tails of the protofilaments A10 and A11 with B1. The arrowhead indicates the conflict between the C-terminal tails of A11 and B1. (M and N) Plots of the interaction energy between tubulin C-terminal tails of A10 and the protofilament B1 (M) or A11 and the protofilament B1 (N).

By using an in silico approach, we explored how the C-terminal tails of the A-microtubule hinder MTD assembly. In our simulations, the A-microtubule was composed of 13 protofilaments of three αβ-tubulin dimers each, where all atoms were taken into account. Because tubulin tails are unstructured, they may adopt random conformations. To obtain a representative sample of these conformations, we used molecular dynamics simulations (see materials and methods). The first protofilament from the B-microtubule, B1, was added to the A-microtubule between A10 and A11 according to the method of (9) (fig. S3, B to D). To capitalize on all the A-microtubule sampled tails, every two successive protofilaments of the A-microtubule (A1-A2, A2-A3, A3-A4, ... A12-A13) were superimposed on A10-A11 of the same microtubule in order to obtain a variety of tail positions at the OJ (Fig. 2K and fig. S3C). Then, for each of these couples of protofilaments, the tails were relaxed and their interaction energy with the entire protofilament B1 was calculated. For the A10 tails, this energy was distributed around 0 kcal/mol, indicating that these tails did not play a role in the insertion of B1 (Fig. 2M). By contrast, for A11 tails, the interaction energy was highly repulsive in 11% of the cases, with an energy value of several thousands of kilocalories per mole, which is sufficient to strongly hinder the insertion of B1 (Fig. 2N). Visual inspection of the constructed junctions confirmed that A11 tails did interpenetrate the core of B1, whereas A10 tails did not (Fig. 2L). This provides an explanation for the results of the in vitro experiments but not for the formation of MTDs in vivo despite the presence of these tails. Observation of the in vivo MTD structure isolated from flagella (9) showed the presence of an unidentified MIP (MIP7) at the junction between A11 and B1, at the same location as the tails of A11 in our model. This MIP7 has been proposed to stabilize the interaction between B1 and A10-A11 (9). We hypothesized that MIP7 action is not in stabilizing the interaction but rather in binding A11 protofilament tubulin tails to enable the B1 insertion.

Next, we monitored the assembly dynamics of MTDs. We immobilized subtilisin-treated Alexa 488–labeled A-microtubules on a glass slide. Rhodamine-labeled free tubulin was added to the reaction mixture to trigger B-microtubule assembly (Fig. 3A), and we monitored the reaction by using total internal reflection fluorescence (TIRF) microscopy. With either guanosine triphosphate or guanosine-5′-[(α,β)-methyleno]triphosphate (GMPCPP) in solution, we observed the usual elongation of template A-microtubules at both tips. However, we observed nucleation and elongation of patches of fluorescent rhodamine signal on the A-microtubule only in the presence of GMPCPP (movies S2 and S3), the same condition used in our cryo-EM experiments. We thus interpreted these patches as B-microtubules (Fig. 3B and fig. S4A). This result suggests that MTD formation requires a certain level of stabilization, mediated in our experiments by GMPCPP and in vivo possibly by the presence of MIPs. Investigating the growth rates of A- and B-microtubule tips showed that, unlike the plus and minus tips of the A-microtubules, which are known to grow at different rates (17), B-microtubules grow at the same rate in both directions (Fig. 3, C and D, and fig. S4, A to D). The isotropic B-microtubule growth rate was faster than the growth rate of the plus tip of the A-microtubule (Fig. 3D), and this rate correlated with increasing tubulin concentration (Fig. 3D and fig. S4D). Thus, B-microtubules are dynamic structures nucleating on the lattice of the A-microtubule and elongating in both directions without apparent anisotropy. To estimate the protofilament number in B-microtubules, we compared the rhodamine fluorescent signal in the center of the B-microtubules with the steady-state rhodamine fluorescent signal at the tips of the template A-microtubules, which are formed by 14 protofilaments because of the presence of GMPCPP (18). This quantification suggested that after 40 min, B-microtubules were assembled, with on average (±SD) 5.7 ± 2.6 and 13.8 ± 3.8 protofilaments at the free tubulin concentrations of 1 and 2 μM, respectively (fig. S4E and movies S2 and S4). Finally, we repeated the experiment with A-microtubules treated with decreasing subtilisin:tubulin ratios (1:1, 1:50; 1:100, and 1:1000), leading to predominant β-tubulin C-terminal tail removal (fig. S5) (13). We found that MTD nucleation decreased markedly (fig. S5 and movie S5), suggesting that the removal of C-terminal tails from both α- and β-tubulin is necessary for the MTD nucleation.

Fig. 3 Dynamics of MTD assembly.

(A) Protocol to visualize MTD assembly by using TIRF microscopy. (B) Montage showing MTDs. Subtilisin-treated A-microtubules are in green; A-microtubule tips and B-microtubules formed by rhodamine-labeled tubulin are depicted in magenta. White arrowheads indicate the tip elongation of the A-microtubule. Yellow arrowheads point to the B-microtubule assembling on the surface of the A-microtubule. Scale bar, 1 μm. (C) Montage showing MTDs and the corresponding multichannel kymograph. Scale bars: horizontal, 5 μm; vertical, 15 min. (D) Polymerization rate of B-microtubules at 2 μM free tubulin. ns, not significant, *P < 0.0001, determined by the Mann-Whitney test. Plus and minus tips of A-microtubules polymerize at 53.93 ± 11.56 nm/min (n = 25 tips) and 37.13 ± 13.13 nm/min (n = 22 tips), respectively. The polymerization rates of B-microtubules toward plus and minus tips of the A-microtubules are 83.51 ± 30.44 nm/min (n = 53 tips) and 96.52 ± 45.87 nm/min (n = 52 tips) [values are averages (represented by red lines) ± SD]. (E) Model of MTD formation. In vitro, MTD assembly initiates on the surface of an A-microtubule (green) deprived of tubulin C-termini (red tails) by the addition of a protofilament owing to a noncanonical surface-to-side tubulin interaction. Protofilaments in the B-microtubule (purple) continue to assemble to ultimately lead to a near-complete MTD.

In vivo, MTDs are composed of heterodimers of α- and β-tubulin and dozens different MIPs. Our work establishes that the C-terminal tail of tubulin exhibits an inhibitory effect that, in vivo, may prevent uncontrolled MTD formation. Molecular simulations suggested that the C-terminal tails of one specific protofilament hinder the attachment of protofilament B1 at the internal side of the OJ. We propose that in vivo, specific MIPs bind and displace the C-terminal tails of A11 and allow for the formation of a B-microtubule that elongates bidirectionally (Fig. 3E). Moreover, such proteins may be needed to precisely position the MTD branching to a specific protofilament on the A-microtubule, as well as to stabilize the entire MTD. The requirement for such protein is alleviated in our in vitro minimal system by providing GMPCPP.

In summary, our work highlights the crucial role of tubulin C-terminal tails in regulating MTDs, which are key to the assembly and function of centrioles, cilia, and flagella.

Supplementary Materials

www.sciencemag.org/content/363/6424/285/suppl/DC1

Materials and Methods

Figs. S1 to S5

References (1932)

Movies S1 to S5

References and Notes

Acknowledgments: We thank N. Klena for critical reading of the manuscript and M. Braun for helpful discussions. We thank N. Olieric for initially providing the subtilisin enzyme. We thank the BioImaging Center of the University of Geneva. Funding: This work was supported by ERC StG 715289 (ACCENT), granted to P.G., and V.H., P.G., and M.L.G. are supported by Swiss National Science Foundation (SNSF) PP00P3_157517. E.S. and S.B. are supported by the University of Geneva. Z.L. is supported by the Czech Science Foundation (18-08304S), the project BIOCEV (CZ.1.05/1.1.00/02.0109) from the ERDF, and CAS (RVO: 86652036). I.Z. is supported by GAUK (1372218). We acknowledge the Centre of Imaging Methods core facility, Faculty of Science, Charles University, supported by the MEYS CR (LM2015062 Czech-BioImaging). Author contributions: M.S.-C. prepared the samples, analyzed the cryo-EM data, and set up the immunofluorescence, Coomassie, and Western blot experiments. M.L.G. performed subtomogram averaging. I.Z. and Z.L. performed and analyzed the TIRF experiments. R.A. and L.M. performed the molecular modeling analysis. D.D. provided access and assistance with the cryo-EM F20 microscope. E.S. prepared samples for the cryo-EM session and performed immunofluorescence experiments. S.B. prepared the SDS–polyacrylamide gel electrophoresis gels and Western blots. P.G. performed the cryo-EM. P.G., Z.L., L.M., and V.H. designed, analyzed, and supervised the work, and P.G., V.H., L.M., M.S.-C., and Z.L. wrote the manuscript. Competing interests: The authors declare no competing interests. Data and materials availability: All data are available in the main text or the supplementary materials.

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