Cilia are multifunctional organelles that are constructed using intraflagellar transport (IFT) of cargo to and from their tip. It is widely held that the retrograde IFT motor, dynein-2, must be controlled in order to reach the ciliary tip and then unleashed to power the return journey. However, the mechanism is unknown. Here, we systematically define the mechanochemistry of human dynein-2 motors as monomers, dimers, and multimotor assemblies with kinesin-II. Combining these data with insights from single-particle EM, we discover that dynein-2 dimers are intrinsically autoinhibited. Inhibition is mediated by trapping dynein-2's mechanical 'linker' and 'stalk' domains within a novel motor–motor interface. We find that linker-mediated inhibition enables efficient transport of dynein-2 by kinesin-II in vitro. These results suggest a conserved mechanism for autoregulation among dimeric dyneins, which is exploited as a switch for dynein-2's recycling activity during IFT.
Subscribe to Journal
Get full journal access for 1 year
only $17.42 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Rosenbaum, J.L. & Witman, G.B. Intraflagellar transport. Nat. Rev. Mol. Cell Biol. 3, 813–825 (2002).
Lechtreck, K.F. IFT-Cargo interactions and protein transport in cilia. Trends Biochem. Sci. 40, 765–778 (2015).
Fliegauf, M., Benzing, T. & Omran, H. When cilia go bad: cilia defects and ciliopathies. Nat. Rev. Mol. Cell Biol. 8, 880–893 (2007).
Ishikawa, H. & Marshall, W.F. Ciliogenesis: building the cell's antenna. Nat. Rev. Mol. Cell Biol. 12, 222–234 (2011).
Kozminski, K.G., Johnson, K.A., Forscher, P. & Rosenbaum, J.L. A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523 (1993).
Johnson, K.A. & Rosenbaum, J.L. Polarity of flagellar assembly in Chlamydomonas. J. Cell Biol. 119, 1605–1611 (1992).
Hao, L. et al. Intraflagellar transport delivers tubulin isotypes to sensory cilium middle and distal segments. Nat. Cell Biol. 13, 790–798 (2011).
Bhogaraju, S. et al. Molecular basis of tubulin transport within the cilium by IFT74 and IFT81. Science 341, 1009–1012 (2013).
Kubo, T. et al. Together, the IFT81 and IFT74 N-termini form the main module for intraflagellar transport of tubulin. J. Cell Sci. 129, 2106–2119 (2016).
Scholey, J.M. Kinesin-2: a family of heterotrimeric and homodimeric motors with diverse intracellular transport functions. Annu. Rev. Cell Dev. Biol. 29, 443–469 (2013).
Andreasson, J.O., Shastry, S., Hancock, W.O. & Block, S.M. The mechanochemical cycle of mammalian kinesin-2 KIF3A/B under load. Curr. Biol. 25, 1166–1175 (2015).
Albracht, C.D., Guzik-Lendrum, S., Rayment, I. & Gilbert, S.P. Heterodimerization of kinesin-2 KIF3AB modulates entry into the processive run. J. Biol. Chem. 291, 23248–23256 (2016).
Prevo, B., Mangeol, P., Oswald, F., Scholey, J.M. & Peterman, E.J. Functional differentiation of cooperating kinesin-2 motors orchestrates cargo import and transport in C. elegans cilia. Nat. Cell Biol. 17, 1536–1545 (2015).
Gibbons, B.H., Asai, D.J., Tang, W.J., Hays, T.S. & Gibbons, I.R. Phylogeny and expression of axonemal and cytoplasmic dynein genes in sea urchins. Mol. Biol. Cell 5, 57–70 (1994).
Pazour, G.J., Wilkerson, C.G. & Witman, G.B. A dynein light chain is essential for the retrograde particle movement of intraflagellar transport (IFT). J. Cell Biol. 141, 979–992 (1998).
Pazour, G.J., Dickert, B.L. & Witman, G.B. The DHC1b (DHC2) isoform of cytoplasmic dynein is required for flagellar assembly. J. Cell Biol. 144, 473–481 (1999).
Porter, M.E., Bower, R., Knott, J.A., Byrd, P. & Dentler, W. Cytoplasmic dynein heavy chain 1b is required for flagellar assembly in Chlamydomonas. Mol. Biol. Cell 10, 693–712 (1999).
Signor, D. et al. Role of a class DHC1b dynein in retrograde transport of IFT motors and IFT raft particles along cilia, but not dendrites, in chemosensory neurons of living Caenorhabditis elegans . J. Cell Biol. 147, 519–530 (1999).
Hou, Y. & Witman, G.B. Dynein and intraflagellar transport. Exp. Cell Res. 334, 26–34 (2015).
Roberts, A.J., Kon, T., Knight, P.J., Sutoh, K. & Burgess, S.A. Functions and mechanics of dynein motor proteins. Nat. Rev. Mol. Cell Biol. 14, 713–726 (2013).
Carter, A.P., Diamant, A.G. & Urnavicius, L. How dynein and dynactin transport cargos: a structural perspective. Curr. Opin. Struct. Biol. 37, 62–70 (2016).
Cianfrocco, M.A., DeSantis, M.E., Leschziner, A.E. & Reck-Peterson, S.L. Mechanism and regulation of cytoplasmic dynein. Annu. Rev. Cell Dev. Biol. 31, 83–108 (2015).
Patel-King, R.S., Gilberti, R.M., Hom, E.F. & King, S.M. WD60/FAP163 is a dynein intermediate chain required for retrograde intraflagellar transport in cilia. Mol. Biol. Cell 24, 2668–2677 (2013).
Asante, D., Stevenson, N.L. & Stephens, D.J. Subunit composition of the human cytoplasmic dynein-2 complex. J. Cell Sci. 127, 4774–4787 (2014).
Schmidts, M. et al. TCTEX1D2 mutations underlie Jeune asphyxiating thoracic dystrophy with impaired retrograde intraflagellar transport. Nat. Commun. 6, 7074 (2015).
Mencarelli, C., Mitchell, A., Leoncini, R., Rosenbaum, J. & Lupetti, P. Isolation of intraflagellar transport trains. Cytoskeleton 70, 439–452 (2013).
Stepanek, L. & Pigino, G. Microtubule doublets are double-track railways for intraflagellar transport trains. Science 352, 721–724 (2016).
Hancock, W.O. Bidirectional cargo transport: moving beyond tug of war. Nat. Rev. Mol. Cell Biol. 15, 615–628 (2014).
Williams, C.L. et al. Direct evidence for BBSome-associated intraflagellar transport reveals distinct properties of native mammalian cilia. Nat. Commun. 5, 5813 2014).
Ichikawa, M., Watanabe, Y., Murayama, T. & Toyoshima, Y.Y. Recombinant human cytoplasmic dynein heavy chain 1 and 2: observation of dynein-2 motor activity in vitro. FEBS Lett. 585, 2419–2423 (2011).
Schmidt, H., Zalyte, R., Urnavicius, L. & Carter, A.P. Structure of human cytoplasmic dynein-2 primed for its power stroke. Nature 518, 435–438 (2015).
Burgess, S.A., Walker, M.L., Sakakibara, H., Knight, P.J. & Oiwa, K. Dynein structure and power stroke. Nature 421, 715–718 (2003).
Kon, T., Mogami, T., Ohkura, R., Nishiura, M. & Sutoh, K. ATP hydrolysis cycle-dependent tail motions in cytoplasmic dynein. Nat. Struct. Mol. Biol. 12, 513–519 (2005).
Roberts, A.J. et al. AAA+ ring and linker swing mechanism in the dynein motor. Cell 136, 485–495 (2009).
Kon, T. et al. The 2.8 Å crystal structure of the dynein motor domain. Nature 484, 345–350 (2012).
Schmidt, H., Gleave, E.S. & Carter, A.P. Insights into dynein motor domain function from a 3.3-Å crystal structure. Nat. Struct. Mol. Biol. 19, 492–497, S1 (2012).
Roberts, A.J. et al. ATP-driven remodeling of the linker domain in the dynein motor. Structure 20, 1670–1680 (2012).
Torisawa, T. et al. Autoinhibition and cooperative activation mechanisms of cytoplasmic dynein. Nat. Cell Biol. 16, 1118–1124 (2014).
Reck-Peterson, S.L. et al. Single-molecule analysis of dynein processivity and stepping behavior. Cell 126, 335–348 (2006).
Shima, T., Imamula, K., Kon, T., Ohkura, R. & Sutoh, K. Head-head coordination is required for the processive motion of cytoplasmic dynein, an AAA+ molecular motor. J. Struct. Biol. 156, 182–189 (2006).
Qiu, W. et al. Dynein achieves processive motion using both stochastic and coordinated stepping. Nat. Struct. Mol. Biol. 19, 193–200 (2012).
Trokter, M., Mücke, N. & Surrey, T. Reconstitution of the human cytoplasmic dynein complex. Proc. Natl. Acad. Sci. USA 109, 20895–20900 (2012).
Imai, H. et al. Direct observation shows superposition and large scale flexibility within cytoplasmic dynein motors moving along microtubules. Nat. Commun. 6, 8179 (2015).
Nicholas, M.P. et al. Control of cytoplasmic dynein force production and processivity by its C-terminal domain. Nat. Commun. 6, 6206 (2015).
McKenney, R.J., Huynh, W., Tanenbaum, M.E., Bhabha, G. & Vale, R.D. Activation of cytoplasmic dynein motility by dynactin-cargo adapter complexes. Science 345, 337–341 (2014).
Schlager, M.A., Hoang, H.T., Urnavicius, L., Bullock, S.L. & Carter, A.P. In vitro reconstitution of a highly processive recombinant human dynein complex. EMBO J. 33, 1855–1868 (2014).
Amos, L.A. Brain dynein crossbridges microtubules into bundles. J. Cell Sci. 93, 19–28 (1989).
Gibbons, I.R. et al. The affinity of the dynein microtubule-binding domain is modulated by the conformation of its coiled-coil stalk. J. Biol. Chem. 280, 23960–23965 (2005).
Kon, T. et al. Helix sliding in the stalk coiled coil of dynein couples ATPase and microtubule binding. Nat. Struct. Mol. Biol. 16, 325–333 (2009).
Cole, D.G. et al. Chlamydomonas kinesin-II-dependent intraflagellar transport (IFT): IFT particles contain proteins required for ciliary assembly in Caenorhabditis elegans sensory neurons. J. Cell Biol. 141, 993–1008 (1998).
Piperno, G. & Mead, K. Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella. Proc. Natl. Acad. Sci. USA 94, 4457–4462 (1997).
Taschner, M. & Lorentzen, E. The intraflagellar transport machinery. Cold Spring Harb. Perspect. Biol. 8, a028092 (2016).
Taschner, M. et al. Intraflagellar transport proteins 172, 80, 57, 54, 38, and 20 form a stable tubulin-binding IFT-B2 complex. EMBO J. 35, 773–790 (2016).
Derr, N.D. et al. Tug-of-war in motor protein ensembles revealed with a programmable DNA origami scaffold. Science 338, 662–665 (2012).
Engel, B.D., Ludington, W.B. & Marshall, W.F. Intraflagellar transport particle size scales inversely with flagellar length: revisiting the balance-point length control model. J. Cell Biol. 187, 81–89 (2009).
Shih, S.M. et al. Intraflagellar transport drives flagellar surface motility. eLife 2, e00744 (2013).
Li, W., Yi, P. & Ou, G. Somatic CRISPR-Cas9-induced mutations reveal roles of embryonically essential dynein chains in Caenorhabditis elegans cilia. J. Cell Biol. 208, 683–692 (2015).
Verhey, K.J. & Hammond, J.W. Traffic control: regulation of kinesin motors. Nat. Rev. Mol. Cell Biol. 10, 765–777 (2009).
Belyy, V. et al. The mammalian dynein-dynactin complex is a strong opponent to kinesin in a tug-of-war competition. Nat. Cell Biol. 18, 1018–1024 (2016).
Yu, I., Garnham, C.P. & Roll-Mecak, A. Writing and reading the tubulin code. J. Biol. Chem. 290, 17163–17172 (2015).
Brunnbauer, M. et al. Regulation of a heterodimeric kinesin-2 through an unprocessive motor domain that is turned processive by its partner. Proc. Natl. Acad. Sci. USA 107, 10460–10465 (2010).
Imanishi, M., Endres, N.F., Gennerich, A. & Vale, R.D. Autoinhibition regulates the motility of the C. elegans intraflagellar transport motor OSM-3. J. Cell Biol. 174, 931–937 (2006).
Liang, Y. et al. FLA8/KIF3B phosphorylation regulates kinesin-II interaction with IFT-B to control IFT entry and turnaround. Dev. Cell 30, 585–597 (2014).
Pedersen, L.B., Geimer, S. & Rosenbaum, J.L. Dissecting the molecular mechanisms of intraflagellar transport in Chlamydomonas . Curr. Biol. 16, 450–459 (2006).
Urnavicius, L. et al. The structure of the dynactin complex and its interaction with dynein. Science 347, 1441–1446 (2015).
Chowdhury, S., Ketcham, S.A., Schroer, T.A. & Lander, G.C. Structural organization of the dynein-dynactin complex bound to microtubules. Nat. Struct. Mol. Biol. 22, 345–347 (2015).
Goodman, B.S. & Reck-Peterson, S.L. Engineering defined motor ensembles with DNA origami. Methods Enzymol. 540, 169–188 (2014).
Castoldi, M. & Popov, A.V. Purification of brain tubulin through two cycles of polymerization-depolymerization in a high-molarity buffer. Protein Expr. Purif. 32, 83–88 (2003).
Hyman, A. et al. Preparation of modified tubulins. Methods Enzymol. 196, 478–485 (1991).
Kon, T., Nishiura, M., Ohkura, R., Toyoshima, Y.Y. & Sutoh, K. Distinct functions of nucleotide-binding/hydrolysis sites in the four AAA modules of cytoplasmic dynein. Biochemistry 43, 11266–11274 (2004).
Cho, C., Reck-Peterson, S.L. & Vale, R.D. Regulatory ATPase sites of cytoplasmic dynein affect processivity and force generation. J. Biol. Chem. 283, 25839–25845 (2008).
Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).
Uyeda, T.Q., Kron, S.J. & Spudich, J.A. Myosin step size: estimation from slow sliding movement of actin over low densities of heavy meromyosin. J. Mol. Biol. 214, 699–710 (1990).
Wickham, H. ggplot2: Elegant Graphics for Data Analysis (Springer Science & Business Media, 2009).
Scheres, S.H. RELION: implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol. 180, 519–530 (2012).
Mindell, J.A. & Grigorieff, N. Accurate determination of local defocus and specimen tilt in electron microscopy. J. Struct. Biol. 142, 334–347 (2003).
van Heel, M., Harauz, G., Orlova, E.V., Schmidt, R. & Schatz, M. A new generation of the IMAGIC image processing system. J. Struct. Biol. 116, 17–24 (1996).
Ludtke, S.J., Baldwin, P.R. & Chiu, W. EMAN: semiautomated software for high-resolution single-particle reconstructions. J. Struct. Biol. 128, 82–97 (1999).
Frank, J. et al. SPIDER and WEB: processing and visualization of images in 3D electron microscopy and related fields. J. Struct. Biol. 116, 190–199 (1996).
Pettersen, E.F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).
We thank C. Moores, G. Zanetti and A. Osborne for critical comments on the manuscript; M. Williams, S. Reck-Peterson and members of the Birkbeck EM group for advice; S. Nofal, S. Miah, and L. Stejskal for initial experiments; and A. Carter (MRC-LMB, Cambridge) for plasmids. This work was supported by a Sir Henry Dale Fellowship to A.J.R. from the Wellcome Trust and Royal Society [104196/Z/14/Z].
The authors declare no competing financial interests.
Integrated supplementary information
Supplementary Figure 1 Influence of microtubule length and buffer conditions on dynein-2 microtubule gliding.
(a) Kymographs of microtubule gliding at 4 nM Dyn2motor. Microtubule lengths are shown below. (b) Plot of gliding velocity as a function of microtubule length at 4 nM input concentration of Dyn2motor. N = 63 microtubules. Microtubule length and velocity are correlated with a Spearman coefficient of 0.56 (p < 0.0001). (c) Plot of mean microtubule gliding velocity (± s.d.) at different dynein-2 motor input concentrations in buffer lacking 50 mM KCl. Slower microtubule gliding velocities were seen for all three dynein-2 constructs in the absence of KCl (compare with Fig. 2e, 3d and 5e). Number of microtubules analyzed per concentration as follows: Dyn2motor 0.2 nM (32), 0.5 nM (46), 2 nM (47), 5 nM (43), 20 nM (48), 200 nM (90), GST-Dyn2motor 0.2 nM (70), 2 nM (32), 10 nM (42), 20 nM (51), 100 nM (46), 200 nM (51), GST-Dyn2(DQR)motor 0.2 nM (35), 1 nM (56), 2 nM (52), 10 nM (51), 20 nM (52), 200 nM (58). Fitted values (± standard error of the fit): Dyn2motor Vmax = 219.3 ± 3.3 nm/s, f = 0.2 ± 0.01, GST-Dyn2motor Vmax = 168.0 ± 2.1 nm/s, f = 0.1 ± 0.004, GST-Dyn2(DQR)motor Vmax = 259.0 ± 2.7 nm/s, f = 0.3 ± 0.01.
(a) EM micrographs GST-Dyn2motor dimers in different nucleotide and buffer conditions. High salt, 500 mM KCl. GST-Dyn2motor domains appear stacked in ADP.Vi and separated in other conditions. GST-Dyn2motor domains are also stacked in ATP conditions (micrograph shown in Fig. 3c). Arrow, particularly well-resolved GST-Dyn2motor dimer in the separated configuration in which stalks are visible in the raw micrograph. (b) EM micrograph GST-Dyn2(DQR)motor dimers in ATP. (c) Histograms showing the distribution of 2D motor–motor distances in EM class averages of GST-Dyn2motor and GST-Dyn2(DQR)motor. Total number of molecules analyzed: GST-Dyn2motor ATP (917), ADP.Vi (1160), no-nucleotide (890), ADP (853), ATP High salt (846), GST-Dyn2(DQR)motor ATP (891). The GST-Dyn2motor distributions in ATP and ADP.Vi conditions show a peak at low motor–motor separation indicative of stacking. Stacking is largely abolished in GST-Dyn2(DQR)motor.
Supplementary Figure 3 Linker-mediated stacking model and consistency with dynein-1 and dynein-2 data.
(a) Upper panel: Model of the autoinhibited state of dynein-2, derived from PDB 4RH7 (Schmidt, H. et al., Nature. 518, 435-8, 2015). The linker domains (magenta) are trapped in the motor–motor interface, while the C-terminal domains (CTDs) are on the periphery of the dimer and do not interact. All motor–motor interfaces are labeled. Lower panel: previous model of the cytoplasmic cytoplasmic dynein-1 phi particle (Torisawa, T. et al., Nature Cell Biology. 16, 1118-24, 2014), derived from PDB 3VKG (Kon, T. et al., Nature. 484, 345-50, 2012). The motor domains interact via their CTDs and the linker domains are free to move on the periphery of the dimer. (b) Enlarged views of dynein-2 interfaces in the new model. Amino acids involved in inter-motor domain interactions are labeled. (c) Top row, EM averages of GST-Dyn2motor in ATP and ADP.Vi conditions. Bottom row, highest scoring projections of the new model based on cross correlation. GST-Dyn2motor shows preferred orientations on the EM support, which are matched by the new model with Euler angles shown (ϕ, θ; SPIDER convention). GST/SNAPf density is labeled with an arrowhead in the EM average and is absent in projections. (d) Equivalent analysis carried out for human cytoplasmic dynein-1 holoenzyme in the phi particle configuration. The tail and associated subunits of the holoenzyme are labeled with an arrow in the EM average. Scale bar, 10 nm. The linker-mediated stacking model is consistent with class averages of both dynein-1 and dynein-2.
(a) Sequence diagrams of dimeric GST-Dyn2(DQR)motor, and the Dyn2(DQR)motor construct used to assess if the DQR mutations impact activity in the context of a dynein-2 monomer. (b) Size-exclusion chromatograms. Dyn2(DQR)motor was normalized to the peak value of GST-Dyn2(DQR)motor. (c) Velocity of microtubule gliding at 1 and 20 nM concentrations of Dyn2motor and Dyn2(DQR)motor. Black and pink lines show mean ± s.d. Number of microtubules analyzed per concentration: Dyn2motor 1 nM (39), 20 nM (56), Dyn2(DQR)motor 1 nM (29), 20 nM (49). (d) Microtubule-stimulated ATPase activity of Dyn2(DQR)motor (Dyn2motor values from Fig. 2f shown in gray for comparison). Experiments were carried out in triplicate, mean values ± s.d. are shown. Fitted values (± standard error of the fit): kcat = 4.7 ± 0.3 s-1, kbasal = 1.6 ± 0.1 s-1, Km(MT) = 8.0 ± 2.2 μM.
(a) Sequence diagrams of Kif3 and Kif3 Δtail, in which mutations that prevent autoinhibition are introduced and putatively disordered C-terminal regions are removed. Yellow box, SNAPf tag. (b) Size-exclusion chromatogram of Kif3 Δtail and schematic of the construct. (c) SDS-PAGE of Kif3 ∆tail after the final purification step. (d) Plot of mean microtubule gliding velocity (± s.d.) at different Kif3 Δtail input concentrations. Number of microtubules analyzed per concentration: 0.5 nM (50), 0.7 nM (48), 2 nM (43), 5 nM (48), 20 nM (50), 60 nM (47). Fitted values (± standard error of the fit): Vmax = 537.0 ± 2.0 nm/s, f = 0.99 ± 0.002. (e) Kymograph showing single-molecule motility of Kif3 Δtail labeled with Alexa647 via its SNAPf tag. (+) and (–) indicate microtubule polarity. (f) Velocity histogram of Kif3 Δtail single molecules (N = 311 molecules).
(a) Gel shift assay showing migration of DNA origami chassis samples with three (3x) or seven (7x) attachment sites and no dynein (‘none’), GST-Dyn2motor (‘WT’), or GST-Dyn2(DQR)motor (‘DQR’) bound. (b,c) Kymographs showing behavior of DNA origami assemblies with (b) seven GST-Dyn2motor and (c) seven GST-Dyn2(DQR)motor sites. (+) and (–) indicate microtubule polarity.
Supplementary Figures 1–6 (PDF 1380 kb)
Coordinates for the linker-stacking model of dynein-2 autoinhibition, derived from PDB 4RH7. (TXT 3609 kb)
Microtubule gliding powered by Dyn2motor at different concentrations. See also Fig. 2d. Videos are shown at 24X real time. (MOV 4884 kb)
Class averages of GST-Dyn2motor in no nucleotide and 1 mM ATP conditions. See also Fig. 4b. Videos show 38 class averages in each condition, looped 4 times. (MOV 3134 kb)
Analysis of PDB 4RH7 crystal lattice reveals pairs of monomeric dynein-2 motor domains matching the stacked architecture of Dyn2motor dimers observed by single-particle electron microscopy. (MOV 8736 kb)
About this article
Cite this article
Toropova, K., Mladenov, M. & Roberts, A. Intraflagellar transport dynein is autoinhibited by trapping of its mechanical and track-binding elements. Nat Struct Mol Biol 24, 461–468 (2017). https://doi.org/10.1038/nsmb.3391
The Generation of Dynein Networks by Multi-Layered Regulation and Their Implication in Cell Division
Frontiers in Cell and Developmental Biology (2020)
International Journal of Molecular Sciences (2020)
Journal of Cell Science (2020)
Seminars in Cell & Developmental Biology (2020)