The interior of a cell is a hive of activity, filled with proteins and other items moving from one location to another. A network of filaments called microtubules forms tracks along which so-called motor proteins carry these items. Kinesins are one group of motor proteins, and a typical kinesin protein has one end (called the ‘motor domain’) that can attach itself to the microtubules. The other end links to the cargo being carried, and a ‘neck’ connects the two. When two of these proteins work together, flexible regions of the neck allow the two motor domains to move past one another, which enable the kinesin to essentially walk along a microtubule in a stepwise manner.

To take these steps along microtubules, each kinesin motor domain in the pair must undergo alternating cycles of tight association and release from their tracks. This cycle is coordinated by binding and breaking down a molecule called ATP, which also provides the energy needed to take the next step. How the cycle of loose and tight microtubule attachment is coordinated with the release of the breakdown products of ATP, and how the energy from the ATP molecule is converted into the force that moves the motor along the microtubule, has been unclear.

Atherton et al. use a technique called cryo-electron microscopy to study—in more detail than previously seen—the structure of the motor domains of two types of kinesin called kinesin-1 and kinesin-3. Images were taken at different stages of the cycle used by the motor domains to extract the energy from ATP molecules. Although the two kinesins have been thought to move along the microtubule tracks in different ways, Atherton et al. find that the core mechanism used by their motor domains is the same.

When a motor domain binds to the microtubule, its shape changes, first stimulating release of the breakdown products of ATP from the previous cycle. This release makes room for a new ATP molecule to bind. The structural changes caused by ATP binding are relatively small but produce larger changes in the flexible neck region that enable individual motor domains within a kinesin pair to co-ordinate their movement and move in a consistent direction. This mechanism involves tight coupling between track binding and fuel usage and makes kinesins highly efficient motors.

The structures uncovered by Atherton et al. reveal a mechanism that links microtubule binding, the energy supplied to the motor domain and the force that moves the kinesin along a microtubule. Future work will clarify whether the key features observed in the motor domains of kinesin-1 and kinesin-3 are also found in other types of kinesin motors.

Kinesins are a large family of microtubule (MT)-based motors that play important roles in many cellular activities including mitosis, motility, and intracellular transport (Vale, 2003; Hirokawa and Noda, 2008; Hirokawa et al., 2010). Their involvement in a range of pathological processes also highlights their significance as therapeutic targets and the importance of understanding the molecular basis of their function (Mandelkow and Mandelkow, 2002; Greber and Way, 2006; Henry et al., 2006; Stokin and Goldstein, 2006; Liu et al., 2012b). Kinesins are defined by their motor domains that contain both the MT and ATP binding sites. Three ATP binding motifs—the P-loop, switch I, switch II–are highly conserved among kinesins (Sablin et al., 1996), myosin motors, and small GTPases (Vale, 1996). Kinesins also share a conserved mode of MT binding (Woehlke et al., 1997; Alonso et al., 1998) such that MT binding, ATP binding, and hydrolysis are functionally coupled for efficient MT-based work.

A number of kinesins drive long distance transport of cellular cargo (Hirokawa et al., 2010; Soppina et al., 2014) with dimerisation allowing them to take multiple 8 nm ATP-driven steps toward MT plus ends (Svoboda et al., 1993). Their processivity depends on communication between the two motor domains, which is achieved via the neck linker that connects each motor domain to the dimer-forming coiled-coil (Hackney, 1994; Rice et al., 1999; Tomishige and Vale, 2000; Clancy et al., 2011). In the presence of MTs, ATP binding stimulates neck linker association (docking) with the motor domain towards the MT plus end, while ATP hydrolysis and MT release cause neck linker undocking (Rice et al., 1999; Vale and Milligan, 2000; Skiniotis et al., 2003; Asenjo et al., 2006); thus, the neck linker is required for both intra-dimer communication and directionality. However, even when the role of the motor N-terminus in reinforcing neck linker movement via cover neck bundle (CNB) formation is considered (Hwang et al., 2008; Khalil et al., 2008), the contribution of neck linker docking to the force generating mechanism(s) of these kinesins remains uncertain (Rice et al., 1999; Vale and Milligan, 2000; Rice et al., 2003). New insights into the conformational rearrangements of these motors when bound to MTs are essential to reveal how they produce force.

The high resolution X-ray structures of a range of kinesin motor domains have established a major communication route from the nucleotide binding site via helix-α4 (the so-called relay helix) to the neck linker, such that alternate conformations of helix-α4 either block or enable neck linker docking (Vale and Milligan, 2000; Kikkawa et al., 2001). However, the neck linker conformation seen in these MT-free structures is not always correlated to the nucleotide bound (Vale and Milligan, 2000; Grant et al., 2007). Cryo-electron microscopy (cryo-EM) has played a major role in elucidating several aspects of MT-bound kinesin mechanochemistry (Sosa and Milligan, 1996; Sosa et al., 1997; Rice et al., 1999; Skiniotis et al., 2003; Hirose et al., 2006; Kikkawa and Hirokawa, 2006; Sindelar and Downing, 2007, 2010; Goulet et al., 2012, 2014). Despite these contributions, and despite recent advances in the study of kinesin–tubulin complexes using X-ray crystallography (Gigant et al., 2013), several outstanding questions concerning kinesin mechanochemistry remain. Specifically, the mechanism by which MT binding stimulates the kinesin ATPase and in particular enhances Mg-ADP release by several orders of magnitude is not clear (Hackney, 1988; Ma and Taylor, 1997; Sindelar, 2011). Although several speculative models have been proposed, an unambiguously interpretable structure of nucleotide-free MT-bound kinesin is currently lacking and is clearly critical in establishing how such transitions are achieved. Such a structure would also provide key insights into how ATP binding is coupled to both neck linker docking and force generation.

To address these major questions, we describe the MT-bound mechanochemical cycles of two plus-end directed human kinesin motor domains, a kinesin-1 (Kif5A) and a kinesin-3 (Kif1A), using cryo-EM structure determination at subnanometer resolution. Kinesin-1s (Kin1) and kinesin-3s (Kin3) are both important neuronal plus-end directed transport motors (Hirokawa et al., 2009b), but recent data suggest that Kin3 rather than Kin1 motors specifically are involved in long distance transport (Soppina et al., 2014). Their motor domains share 41% sequence identity, but profoundly different mechanochemistries—in which Kin1 dimers take processive steps and Kin3 monomers diffuse along MT tracks—have been proposed for these motors (Hirokawa et al., 2009a; Sindelar, 2011). Thus, we wanted to investigate these differences and compare the motors side by side. The high quality of our reconstructions, coupled to flexible fitting, enables new insights into the kinesin mechanism. In particular, nucleotide-free reconstructions for both motor domains reveal a conserved mechanism, whereby MT binding stimulates changes at the nucleotide-binding site favouring Mg-ADP release and conformationally primes the motor to receive Mg-ATP. We also show that relatively small structural transitions occur at the nucleotide-binding site on Mg-ATP binding, but that these lead to larger scale conformational changes and neck linker docking. Structural analysis of two different transport kinesins allows a direct comparison of their conserved mechanochemical features and identification of attributes that confer distinctive properties on each motor.

Kinesin mechanochemistry and the extent of mechanistic conservation within the motor superfamily are open questions, critical to explain how MT binding, and ATP binding and hydrolysis drive motor activity. Our structural characterisation of two transport motors now allows us to propose a model that describes the roles of mechanochemical elements that together drive conserved MT-based motor function (Figure 7).

Figure 7

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In the Mg-ADP-bound kinesin, association with the MT surface is experienced directly by loop11 and the N-terminus of helix-α4 biasing their conformations towards more structured states. Full stabilisation of these elements is not achieved until Mg-ADP is released, and the additional contacts with the MT surface may in particular serve to nucleate the single turn helix in loop11. This is consistent with the well-documented role of loop11 in sensing MT attachment and triggering Mg-ADP release via interactions with α-tubulin (Woehlke et al., 1997; Yun et al., 2001; Ebbing et al., 2008; Uchimura et al., 2010). Loop9 does not directly contact the MT before or after Mg-ADP release but dramatically changes conformation, unfurling, and extending around the nucleotide-binding site. The structured conformations of loop11 and the N-terminus of helix-α4 are sterically compatible with the conformations of loop9 before and after Mg-ADP release—that is, no clashes are seen in either case. However, the extended conformation of loop9 and the ordered conformations of helix-α4/loop11 are likely to be mutually stabilising due to formation of additional contacts, and thereby mediate communication between the nucleotide and MT-binding sites (Woehlke et al., 1997; Yun et al., 2001; Farrell et al., 2002; Ebbing et al., 2008; Nitta et al., 2008). Critically, however, the water network coordinating Mg-ADP is stabilized exclusively by the retracted helical conformation of loop9 (Figure 2—figure supplement 2). The transition towards the extended conformation of loop9 promotes Mg-ADP release by destabilisation of Mg coordination (Nitta et al., 2008). These structural rearrangements therefore indicate that sequential conformational changes of the switch loops in the presence of MTs stimulate Mg-ADP release, the rate-limiting step of motors in solution (Hackney, 1988). These rearrangements allow formation of a nucleotide-free motor that is strongly bound to its MT track (Nakata and Hirokawa, 1995), at least in part due to additional contacts formed between loop11 and the MT.

Conformational changes at the nucleotide-binding site that lead to Mg-ADP release also appear to prime the kinesin motor domain for Mg-ATP binding. However, the primed conformation clearly does not lead to neck linker docking in the absence of Mg-ATP, contrary to previous predictions (Nitta et al., 2008). Multiple strands of evidence suggest that the neck linkers of transport kinesins in solution explore both docked and undocked conformations independent of the nucleotide state (Rice et al., 1999; Nitta et al., 2008; Scarabelli and Grant, 2013). Thus, tight MT binding is critical in strongly biasing neck linker conformation in the absence of nucleotide such that it will be undocked and, in our reconstructions, directed albeit flexibly towards the MT minus end. Interaction of helix-α6 with α-tubulin's H12 (Uchimura et al., 2010) may therefore help to prevent neck linker docking in the absence of nucleotide, despite changes in the conformations of the switch loops at the active site.

Mg-ATP binding does not cause large rearrangements of the nucleotide-binding site of MT-bound motor domains. However, the presence of the pre-hydrolysis γ-phosphate of Mg-ATP is critical for the pincer-like movement of loop11 and loop9 towards each other. Along with formation of strong additional contacts between these loops, the helix-α4 N-terminus and the P-loop (see Figure 2—figure supplement 4 and Parke et al., 2010; Chang et al., 2013; Gigant et al., 2013), this new local connectivity induces the larger rearrangements that cause neck linker docking. The resulting conformational changes cannot be described only as a tilt of the motor domain relative to static contacts with the MT including helix-α4; in addition to β-sheet tilting, multiple changes across the domain reinforce mechanical amplification and neck linker docking when Mg-ATP binds. The resolution of our reconstructions also allows us to detect subtle distortion of the central β-sheet edges on Mg-ATP binding. However, arguably the most important consequences of Mg-ATP binding are the changes—extension, tilting, and translation—in helix-α6 that allow neck linker docking. This conformation is stabilised by contacts between its N-terminus and elements in the nucleotide-binding pocket (see Figure 2—figure supplement 4 and Parke et al., 2010; Chang et al., 2013; Gigant et al., 2013).

Neck linker docking is essential for both defining the directionality of kinesin motility and mediating head–head tension to ensure processive dimer stepping (Rice et al., 1999; Tomishige and Vale, 2000; Vale and Milligan, 2000; Skiniotis et al., 2003; Clancy et al., 2011; Sindelar, 2011), but whether docking itself can generate the force required for kinesin stepping has been questioned (Rice et al., 2003). Thus, the structural basis of ATP-dependent force generation remains a matter of debate in the field (Visscher et al., 1999; Cross and McAinsh, 2014). The conformational changes associated with helix-α6 during the ATPase cycle—in which contacts with the MT formed in the ADP/NN state are broken as Mg-ATP-dependent rotation pulls it away from the MT surface—reinforce neck linker movements and may also contribute to mechanical amplification and force generation. The translation/extension of helix-α6 into the hydrophobic cavity that is created by β-sheet tilting when Mg-ATP binds may ensure that this tilting is not reversed. Intriguingly, mutagenesis of residues at the helix-α6/neck linker junction has a profound effect on the activity of kinesin monomers (Case et al., 2000), pointing to the importance and likely conservation of structural transitions in this region (Case et al., 1997). Importantly, movement of helix-α6 also relieves steric blocking of neck linker docking and presumably biases the mobile neck linker trajectory. In collaboration with the motor N-terminus, formation of the CNB reinforces the plus end directionality of this bias. Thus, we propose that the helix-α6 is a key mechanical element within the kinesin motor domain, and that its Mg-ATP-dependent movement is essential to plus-end directed stepping.

Once the neck linker has docked, ATP hydrolysis occurs, ensuring efficient coupling between kinesin stepping, Mg-ATP binding and hydrolysis (Schnitzer et al., 2000; Hahlen et al., 2006). A detailed reaction mechanism for hydrolysis has been proposed based on the conformations of loop9 and loop11 (a so-called ‘phosphate tube’) with Mg-ATP-analogue bound (Parke et al., 2010). Consistent with MT binding being important in the catalytic enhancement of kinesins (Ma and Taylor, 1997), this hydrolysis competent configuration of the switch loops is rarely seen in Mg-ATP-analogue kinesin structures in the absence of MTs (e.g., Kikkawa et al., 2001; Nitta et al., 2004; Cochran et al., 2009, with Parke et al., 2010; Chang et al., 2013 being the notable exceptions); those in complex with tubulin always adopt this configuration (Sindelar and Downing, 2010; Goulet et al., 2012; Gigant et al., 2013). On Mg-ADP release, loop9 and loop11 are stabilized into conformations quite close to catalytically competent ones. This suggests that the conformational changes triggered by MT binding that lead to MT-stimulated ADP release also contribute to setting up the catalytic site for ATP hydrolysis. Thus, a subset of mutations in MT-sensing residues in loop11 or which decouple MT affinity and ADP-release also affect MT-stimulated ATP-hydrolysis (Woehlke et al., 1997; Song and Endow, 1998; Yun et al., 2001; Ebbing et al., 2008; Uchimura et al., 2010). Following hydrolysis and phosphate release, we would predict that the Mg-ADP remaining in the catalytic site causes retraction of loop9, subsequent destabilization of loop11 and the helix-α4 N-terminus, leading to track detachment.

This model allows several previously proposed hypotheses, in particular concerning MT-stimulated Mg-ADP release, to be excluded. Mechanisms that involve MT-induced ‘opening’ of the nucleotide pocket, disordering of the switch loops around the nucleotide pocket to destabilise Mg-ADP coordination, or in which loop9 extends into the nucleotide pocket to perturb the P-loop and eject Mg-ADP (Yun et al., 2001; Kikkawa and Hirokawa, 2006; Sindelar and Downing, 2007; Nitta et al., 2008; Sindelar, 2011) are not supported by our observations that: (1) both loop9 and loop11 move towards the nucleotide-binding pocket on Mg-ADP release, (2) these loops adopt well-defined and conserved conformations that are clearly visualised after Mg-ADP release, and (3) the conformation of these loops does not sterically interfere with nucleotide binding or disrupt the P-loop. Another prominent idea is that a significant twist of the core β-sheet caused by MT attachment would promote Mg-ADP release analogous to the equivalent release step in myosin (Coureux et al., 2003; Hirose et al., 2006; Kull and Endow, 2013). However, comparison of our Kin3-Mg-ADP and Kin3-NN reconstructions (Figure 5—figure supplement 1A) does not support β-sheet twist as a mechanism for Mg-ADP release in kinesins.

The structural elements involved in these mechanochemical transitions are extremely well conserved amongst kinesins, and it is likely that the mechanisms we describe are utilised by all superfamily members. We previously characterised the MT-bound ATPase cycle of human kinesin-5 (Kin5, Goulet et al., 2012, 2014). Although the resolutions of those cryo-EM reconstructions (∼10 Å) do not provide the level of detail of the current work, many of our current hypotheses are consistent with a conserved mechanochemistry, specifically conformational coupling of loops9 and 11 to bring about MT-induced Mg-ADP release and Mg-ATP induced neck linker docking. Superimposed on this conserved mechanochemistry, family-specific modifications were also detected; most strikingly for Kin5, these include the proposed role of the Kin5-extended loop5 in controlling nucleotide binding and the stiffer properties of the Kin5 neck linker that undergoes an order-to-order transition on Mg-ATP binding. Family-specific insertions elsewhere in the motor domain are likely to have other modifying roles, such as Kin3's loop12, which enhances the initial interaction between these highly processive motors and their tracks (Soppina and Verhey, 2014). A tantalising hint of how insertions in loop2 may be coupled to MT depolymerisation in for example kinesin-13s (Desai et al., 1999; Moores et al., 2002; Asenjo et al., 2013) and kinesin-8s (Varga et al., 2006; Peters et al., 2010) is provided by its proximity to the MT surface and the mechanical amplifier helix-α6, and by its large displacement on Mg-ATP binding. Future studies at high resolution will provide further insights into the ways this conserved mechanochemistry is modified in diverse functional contexts within the kinesin superfamily.

1. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-3 motor domain in absence of nucleotides

   Publicly available at The Electron Microscopy Data Bank.

   http://www.ebi.ac.uk/pdbe/entry/EMD-2765
2. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-3 motor domain in absence of nucleotides

   Publicly available at RCSB Protein Data Bank.

   http://www.pdb.org/pdb/explore/explore.do?structureId=4uxo
3. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 3-protofilament microtubule-bound human kinesin-3 motor domain with AMPPNP

   Publicly available at The Electron Microscopy Data Bank.

   http://www.ebi.ac.uk/pdbe/entry/EMD-2766
4. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 3-protofilament microtubule-bound human kinesin-3 motor domain with AMPPNP

   Publicly available at RCSB Protein Data Bank.

   http://www.pdb.org/pdb/explore/explore.do?structureId=4uxp
5. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-3 motor domain with ADPAlFx

   Publicly available at The Electron Microscopy Data Bank.

   http://www.ebi.ac.uk/pdbe/entry/EMD-2767
6. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-3 motor domain with ADPAlFx

   Publicly available at RCSB Protein Data Bank.

   http://www.pdb.org/pdb/explore/explore.do?structureId=4uxr
7. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-3 motor domain with ADP

   Publicly available at The Electron Microscopy Data Bank.

   http://www.ebi.ac.uk/pdbe/entry/EMD-2768
8. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-3 motor domain with ADP

   Publicly available at RCSB Protein Data Bank.

   http://www.pdb.org/pdb/explore/explore.do?structureId=4uxs
9. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) 13-protofilament microtubule-bound human kinesin-1 motor domain in absence of nucleotides

   Publicly available at The Electron Microscopy Data Bank.

   http://www.ebi.ac.uk/pdbe/entry/EMD-2769
10. 1. Atherton J
    2. Farabella I
    3. Yu IM
    4. Rosenfeld SS
    5. Houdusse A
    6. Topf M
    7. Moores CA

    (2014) 13-protofilament microtubule-bound human kinesin-1 motor domain in absence of nucleotides

    Publicly available at RCSB Protein Data Bank.

    http://www.pdb.org/pdb/explore/explore.do?structureId=4uxt
11. 1. Atherton J
    2. Farabella I
    3. Yu IM
    4. Rosenfeld SS
    5. Houdusse A
    6. Topf M
    7. Moores CA

    (2014) 13-protofilament microtubule-bound human kinesin-1 motor domain with AMPPNP

    Publicly available at The Electron Microscopy Data Bank.

    http://www.ebi.ac.uk/pdbe/entry/EMD-2770
12. 1. Atherton J
    2. Farabella I
    3. Yu IM
    4. Rosenfeld SS
    5. Houdusse A
    6. Topf M
    7. Moores CA

    (2014) 13-protofilament microtubule-bound human kinesin-1 motor domain with AMPPNP

    Publicly available at RCSB Protein Data Bank.

    http://www.pdb.org/pdb/explore/explore.do?structureId=4uxy
13. 1. Atherton J
    2. Farabella I
    3. Yu IM
    4. Rosenfeld SS
    5. Houdusse A
    6. Topf M
    7. Moores CA

    (2014) 13-protofilament microtubule-bound human kinesin-1 motor domain with ADPAlFx

    Publicly available at The Electron Microscopy Data Bank.

    http://www.ebi.ac.uk/pdbe/entry/EMD-2771
14. 1. Atherton J
    2. Farabella I
    3. Yu IM
    4. Rosenfeld SS
    5. Houdusse A
    6. Topf M
    7. Moores CA

    (2014) 13-protofilament microtubule-bound human kinesin-1 motor domain with ADPAlFx

    Publicly available at RCSB Protein Data Bank.

    http://www.pdb.org/pdb/explore/explore.do?structureId=4uy0

1. 1. Alonso MC
   2. van Damme J
   3. Vandekerckhove J
   4. Cross RA

   (1998) Proteolytic mapping of kinesin/ncd-microtubule interface: nucleotide-dependent conformational changes in the loops L8 and L12

   The EMBO Journal 17:945–951.

   https://doi.org/10.1093/emboj/17.4.945

   • Google Scholar
2. 1. Asenjo AB
   2. Chatterjee C
   3. Tan D
   4. DePaoli V
   5. Rice WJ
   6. Diaz-Avalos R
   7. Silvestry M
   8. Sosa H

   (2013) Structural model for tubulin recognition and deformation by kinesin-13 microtubule depolymerases

   Cell Reports 3:759–768.

   https://doi.org/10.1016/j.celrep.2013.01.030

   • Google Scholar
3. 1. Asenjo AB
   2. Weinberg Y
   3. Sosa H

   (2006) Nucleotide binding and hydrolysis induces a disorder-order transition in the kinesin neck-linker region

   Nature Structural & Molecular Biology 13:648–654.

   https://doi.org/10.1038/nsmb1109

   • Google Scholar
4. 1. Case RB
   2. Pierce DW
   3. Hom-Booher N
   4. Hart CL
   5. Vale RD

   (1997) The directional preference of kinesin motors is specified by an element outside of the motor catalytic domain

   Cell 90:959–966.

   https://doi.org/10.1016/S0092-8674(00)80360-8

   • Google Scholar
5. 1. Case RB
   2. Rice S
   3. Hart CL
   4. Ly B
   5. Vale RD

   (2000) Role of the kinesin neck linker and catalytic core in microtubule-based motility

   Current Biology 10:157–160.

   https://doi.org/10.1016/S0960-9822(00)00316-X

   • Google Scholar
6. 1. Chang Q
   2. Nitta R
   3. Inoue S
   4. Hirokawa N

   (2013) Structural basis for the ATP-induced isomerization of kinesin

   Journal of Molecular Biology 425:1869–1880.

   https://doi.org/10.1016/j.jmb.2013.03.004

   • Google Scholar
7. 1. Chen S
   2. McMullan G
   3. Faruqi AR
   4. Murshudov GN
   5. Short JM
   6. Scheres SH
   7. Henderson R

   (2013) High-resolution noise substitution to measure overfitting and validate resolution in 3D structure determination by single particle electron cryomicroscopy

   Ultramicroscopy 135:24–35.

   https://doi.org/10.1016/j.ultramic.2013.06.004

   • Google Scholar
8. 1. Clancy BE
   2. Behnke-Parks WM
   3. Andreasson JO
   4. Rosenfeld SS
   5. Block SM

   (2011) A universal pathway for kinesin stepping

   Nature Structural & Molecular Biology 18:1020–1027.

   https://doi.org/10.1038/nsmb.2104

   • Google Scholar
9. 1. Cochran JC
   2. Sindelar CV
   3. Mulko NK
   4. Collins KA
   5. Kong SE
   6. Hawley RS
   7. Kull FJ

   (2009) ATPase cycle of the nonmotile kinesin NOD allows microtubule end tracking and drives chromosome movement

   Cell 136:110–122.

   https://doi.org/10.1016/j.cell.2008.11.048

   • Google Scholar
10. 1. Coureux PD
    2. Wells AL
    3. Menetrey J
    4. Yengo CM
    5. Morris CA
    6. Sweeney HL
    7. Houdusse A

    (2003) A structural state of the myosin V motor without bound nucleotide

    Nature 425:419–423.

    https://doi.org/10.1038/nature01927

    • Google Scholar
11. 1. Cross RA
    2. McAinsh A

    (2014) Prime movers: the mechanochemistry of mitotic kinesins

    Nature Reviews Molecular Cell Biology 15:257–271.

    https://doi.org/10.1038/nrm3768

    • Google Scholar
12. 1. Desai A
    2. Verma S
    3. Mitchison TJ
    4. Walczak CE

    (1999) Kin I kinesins are microtubule-destabilizing enzymes

    Cell 96:69–78.

    https://doi.org/10.1016/S0092-8674(00)80960-5

    • Google Scholar
13. 1. Ebbing B
    2. Mann K
    3. Starosta A
    4. Jaud J
    5. Schols L
    6. Schule R
    7. Woehlke G

    (2008) Effect of spastic paraplegia mutations in KIF5A kinesin on transport activity

    Human Molecular Genetics 17:1245–1252.

    https://doi.org/10.1093/hmg/ddn014

    • Google Scholar
14. 1. Farrell CM
    2. Mackey AT
    3. Klumpp LM
    4. Gilbert SP

    (2002) The role of ATP hydrolysis for kinesin processivity

    The Journal of Biological Chemistry 277:17079–17087.

    https://doi.org/10.1074/jbc.M108793200

    • Google Scholar
15. 1. Fourniol FJ
    2. Moores CA

    (2011) Snapshots of kinesin motors on microtubule tracks

    Methods in Molecular Biology 778:57–70.

    https://doi.org/10.1007/978-1-61779-261-8_5

    • Google Scholar
16. 1. Frank J
    2. Radermacher M
    3. Penczek P
    4. Zhu J
    5. Li Y
    6. Ladjadj M
    7. Leith A

    (1996) SPIDER and WEB: processing and visualization of images in 3D electron microscopy and related fields

    Journal of Structural Biology 116:190–199.

    https://doi.org/10.1006/jsbi.1996.0030

    • Google Scholar
17. 1. Gigant B
    2. Wang W
    3. Dreier B
    4. Jiang Q
    5. Pecqueur L
    6. Pluckthun A
    7. Wang C
    8. Knossow M

    (2013) Structure of a kinesin-tubulin complex and implications for kinesin motility

    Nature Structural & Molecular Biology 20:1001–1007.

    https://doi.org/10.1038/nsmb.2624

    • Google Scholar
18. 1. Goddard TD
    2. Huang CC
    3. Ferrin TE

    (2007) Visualizing density maps with UCSF Chimera

    Journal of Structural Biology 157:281–287.

    https://doi.org/10.1016/j.jsb.2006.06.010

    • Google Scholar
19. 1. Goulet A
    2. Behnke-Parks WM
    3. Sindelar CV
    4. Major J
    5. Rosenfeld SS
    6. Moores CA

    (2012) The structural basis of force generation by the mitotic motor kinesin-5

    The Journal of Biological Chemistry 287:44654–44666.

    https://doi.org/10.1074/jbc.M112.404228

    • Google Scholar
20. 1. Goulet A
    2. Major J
    3. Jun Y
    4. Gross SP
    5. Rosenfeld SS
    6. Moores CA

    (2014) Comprehensive structural model of the mechanochemical cycle of a mitotic motor highlights molecular adaptations in the kinesin family

    Proceedings of the National Academy of Sciences of USA 111:1837–1842.

    https://doi.org/10.1073/pnas.1319848111

    • Google Scholar
21. 1. Grant BJ
    2. McCammon JA
    3. Caves LS
    4. Cross RA

    (2007) Multivariate analysis of conserved sequence-structure relationships in kinesins: coupling of the active site and a tubulin-binding sub-domain

    Journal of Molecular Biology 368:1231–1248.

    https://doi.org/10.1016/j.jmb.2007.02.049

    • Google Scholar
22. 1. Greber UF
    2. Way M

    (2006) A superhighway to virus infection

    Cell 124:741–754.

    https://doi.org/10.1016/j.cell.2006.02.018

    • Google Scholar
23. 1. Grigorieff N

    (2007) FREALIGN: high-resolution refinement of single particle structures

    Journal of Structural Biology 157:117–125.

    https://doi.org/10.1016/j.jsb.2006.05.004

    • Google Scholar
24. 1. Hackney DD

    (1994) Evidence for alternating head catalysis by kinesin during microtubule-stimulated ATP hydrolysis

    Proceedings of the National Academy of Sciences of USA 91:6865–6869.

    https://doi.org/10.1073/pnas.91.15.6865

    • Google Scholar
25. 1. Hackney DD

    (1988) Kinesin ATPase: rate-limiting ADP release

    Proceedings of the National Academy of Sciences of USA 85:6314–6318.

    https://doi.org/10.1073/pnas.85.17.6314

    • Google Scholar
26. 1. Hahlen K
    2. Ebbing B
    3. Reinders J
    4. Mergler J
    5. Sickmann A
    6. Woehlke G

    (2006) Feedback of the kinesin-1 neck-linker position on the catalytic site

    The Journal of Biological Chemistry 281:18868–18877.

    https://doi.org/10.1074/jbc.M508019200

    • Google Scholar
27. 1. Henry T
    2. Gorvel JP
    3. Meresse S

    (2006) Molecular motors hijacking by intracellular pathogens

    Cellular Microbiology 8:23–32.

    https://doi.org/10.1111/j.1462-5822.2005.00649.x

    • Google Scholar
28. 1. Hirokawa N
    2. Nitta R
    3. Okada Y

    (2009a) The mechanisms of kinesin motor motility: lessons from the monomeric motor KIF1A

    Nature Reviews Molecular Cell Biology 10:877–884.

    https://doi.org/10.1038/nrm2807

    • Google Scholar
29. 1. Hirokawa N
    2. Niwa S
    3. Tanaka Y

    (2010) Molecular motors in neurons: transport mechanisms and roles in brain function, development, and disease

    Neuron 68:610–638.

    https://doi.org/10.1016/j.neuron.2010.09.039

    • Google Scholar
30. 1. Hirokawa N
    2. Noda Y

    (2008) Intracellular transport and kinesin superfamily proteins, KIFs: structure, function, and dynamics

    Physiological Reviews 88:1089–1118.

    https://doi.org/10.1152/physrev.00023.2007

    • Google Scholar
31. 1. Hirokawa N
    2. Noda Y
    3. Tanaka Y
    4. Niwa S

    (2009b) Kinesin superfamily motor proteins and intracellular transport

    Nature Reviews Molecular Cell Biology 10:682–696.

    https://doi.org/10.1038/nrm2774

    • Google Scholar
32. 1. Hirose K
    2. Akimaru E
    3. Akiba T
    4. Endow SA
    5. Amos LA

    (2006) Large conformational changes in a kinesin motor catalyzed by interaction with microtubules

    Molecular Cell 23:913–923.

    https://doi.org/10.1016/j.molcel.2006.07.020

    • Google Scholar
33. 1. Hirose K
    2. Amos LA

    (2007) High-resolution structural analysis of the kinesin-microtubule complex by electron cryo-microscopy

    Methods in Molecular Biology 392:213–230.

    https://doi.org/10.1007/978-1-59745-490-2_15

    • Google Scholar
34. 1. Hwang W
    2. Lang MJ
    3. Karplus M

    (2008) Force generation in kinesin hinges on cover-neck bundle formation

    Structure 16:62–71.

    https://doi.org/10.1016/j.str.2007.11.008

    • Google Scholar
35. 1. Khalil AS
    2. Appleyard DC
    3. Labno AK
    4. Georges A
    5. Karplus M
    6. Belcher AM
    7. Hwang W
    8. Lang MJ

    (2008) Kinesin's cover-neck bundle folds forward to generate force

    Proceedings of the National Academy of Sciences of USA 105:19247–19252.

    https://doi.org/10.1073/pnas.0805147105

    • Google Scholar
36. 1. Kikkawa M
    2. Hirokawa N

    (2006) High-resolution cryo-EM maps show the nucleotide binding pocket of KIF1A in open and closed conformations

    The EMBO Journal 25:4187–4194.

    https://doi.org/10.1038/sj.emboj.7601299

    • Google Scholar
37. 1. Kikkawa M
    2. Okada Y
    3. Hirokawa N

    (2000) 15 A resolution model of the monomeric kinesin motor, KIF1A

    Cell 100:241–252.

    https://doi.org/10.1016/S0092-8674(00)81562-7

    • Google Scholar
38. 1. Kikkawa M
    2. Sablin EP
    3. Okada Y
    4. Yajima H
    5. Fletterick RJ
    6. Hirokawa N

    (2001) Switch-based mechanism of kinesin motors

    Nature 411:439–445.

    https://doi.org/10.1038/35078000

    • Google Scholar
39. 1. Konishi Y
    2. Setou M

    (2009) Tubulin tyrosination navigates the kinesin-1 motor domain to axons

    Nature Neuroscience 12:559–567.

    https://doi.org/10.1038/nn.2314

    • Google Scholar
40. 1. Kull FJ
    2. Endow SA

    (2013) Force generation by kinesin and myosin cytoskeletal motor proteins

    Journal of Cell Science 126:9–19.

    https://doi.org/10.1242/jcs.103911

    • Google Scholar
41. 1. Kull FJ
    2. Sablin EP
    3. Lau R
    4. Fletterick RJ
    5. Vale RD

    (1996) Crystal structure of the kinesin motor domain reveals a structural similarity to myosin

    Nature 380:550–555.

    https://doi.org/10.1038/380550a0

    • Google Scholar
42. 1. Liu HL
    2. Hallen MA
    3. Endow SA

    (2012a) Altered nucleotide-microtubule coupling and increased mechanical output by a kinesin mutant

    PLOS ONE 7:e47148.

    https://doi.org/10.1371/journal.pone.0047148

    • Google Scholar
43. 1. Liu JS
    2. Schubert CR
    3. Fu X
    4. Fourniol FJ
    5. Jaiswal JK
    6. Houdusse A
    7. Stultz CM
    8. Moores CA
    9. Walsh CA

    (2012b) Molecular basis for specific regulation of neuronal kinesin-3 motors by doublecortin family proteins

    Molecular Cell 47:707–721.

    https://doi.org/10.1016/j.molcel.2012.06.025

    • Google Scholar
44. 1. Ludtke SJ
    2. Baldwin PR
    3. Chiu W

    (1999) EMAN: semiautomated software for high-resolution single-particle reconstructions

    Journal of Structural Biology 128:82–97.

    https://doi.org/10.1006/jsbi.1999.4174

    • Google Scholar
45. 1. Ma YZ
    2. Taylor EW

    (1997) Kinetic mechanism of a monomeric kinesin construct

    The Journal of Biological Chemistry 272:717–723.

    https://doi.org/10.1074/jbc.272.2.717

    • Google Scholar
46. 1. Mandelkow E
    2. Mandelkow EM

    (2002) Kinesin motors and disease

    Trends in Cell Biology 12:585–591.

    https://doi.org/10.1016/S0962-8924(02)02400-5

    • Google Scholar
47. 1. Moores CA
    2. Yu M
    3. Guo J
    4. Beraud C
    5. Sakowicz R
    6. Milligan RA

    (2002) A mechanism for microtubule depolymerization by KinI kinesins

    Molecular Cell 9:903–909.

    https://doi.org/10.1016/S1097-2765(02)00503-8

    • Google Scholar
48. 1. Nakata T
    2. Hirokawa N

    (1995) Point mutation of adenosine triphosphate-binding motif generated rigor kinesin that selectively blocks anterograde lysosome membrane transport

    The Journal of Cell Biology 131:1039–1053.

    https://doi.org/10.1083/jcb.131.4.1039

    • Google Scholar
49. 1. Nitta R
    2. Kikkawa M
    3. Okada Y
    4. Hirokawa N

    (2004) KIF1A alternately uses two loops to bind microtubules

    Science 305:678–683.

    https://doi.org/10.1126/science.1096621

    • Google Scholar
50. 1. Nitta R
    2. Okada Y
    3. Hirokawa N

    (2008) Structural model for strain-dependent microtubule activation of Mg-ADP release from kinesin

    Nature Structural & Molecular Biology 15:1067–1075.

    https://doi.org/10.1038/nsmb.1487

    • Google Scholar
51. 1. Okada Y
    2. Hirokawa N

    (1999) A processive single-headed motor: kinesin superfamily protein KIF1A

    Science 283:1152–1157.

    https://doi.org/10.1126/science.283.5405.1152

    • Google Scholar
52. 1. Okada Y
    2. Hirokawa N

    (2000) Mechanism of the single-headed processivity: diffusional anchoring between the K-loop of kinesin and the C terminus of tubulin

    Proceedings of the National Academy of Sciences of USA 97:640–645.

    https://doi.org/10.1073/pnas.97.2.640

    • Google Scholar
53. 1. Pandurangan AP
    2. Shakeel S
    3. Butcher SJ
    4. Topf M

    (2014) Combined approaches to flexible fitting and assessment in virus capsids undergoing conformational change

    Journal of Structural Biology 185:427–439.

    https://doi.org/10.1016/j.jsb.2013.12.003

    • Google Scholar
54. 1. Pandurangan AP
    2. Topf M

    (2012) Finding rigid bodies in protein structures: Application to flexible fitting into cryoEM maps

    Journal of Structural Biology 177:520–531.

    https://doi.org/10.1016/j.jsb.2011.10.011

    • Google Scholar
55. 1. Parke CL
    2. Wojcik EJ
    3. Kim S
    4. Worthylake DK

    (2010) ATP hydrolysis in Eg5 kinesin involves a catalytic two-water mechanism

    The Journal of Biological Chemistry 285:5859–5867.

    https://doi.org/10.1074/jbc.M109.071233

    • Google Scholar
56. 1. Peters C
    2. Brejc K
    3. Belmont L
    4. Bodey AJ
    5. Lee Y
    6. Yu M
    7. Guo J
    8. Sakowicz R
    9. Hartman J
    10. Moores CA

    (2010) Insight into the molecular mechanism of the multitasking kinesin-8 motor

    The EMBO Journal 29:3437–3447.

    https://doi.org/10.1038/emboj.2010.220

    • Google Scholar
57. 1. Reubold TF
    2. Eschenburg S
    3. Becker A
    4. Kull FJ
    5. Manstein DJ

    (2003) A structural model for actin-induced nucleotide release in myosin

    Nature Structural Biology 10:826–830.

    https://doi.org/10.1038/nsb987

    • Google Scholar
58. 1. Rice S
    2. Cui Y
    3. Sindelar C
    4. Naber N
    5. Matuska M
    6. Vale R
    7. Cooke R

    (2003) Thermodynamic properties of the kinesin neck-region docking to the catalytic core

    Biophysical Journal 84:1844–1854.

    https://doi.org/10.1016/S0006-3495(03)74992-3

    • Google Scholar
59. 1. Rice S
    2. Lin AW
    3. Safer D
    4. Hart CL
    5. Naber N
    6. Carragher BO
    7. Cain SM
    8. Pechatnikova E
    9. Wilson-Kubalek EM
    10. Whittaker M
    11. Pate E
    12. Cooke R
    13. Taylor EW
    14. Milligan RA
    15. Vale RD

    (1999) A structural change in the kinesin motor protein that drives motility

    Nature 402:778–784.

    https://doi.org/10.1038/45483

    • Google Scholar
60. 1. Sablin EP
    2. Kull FJ
    3. Cooke R
    4. Vale RD
    5. Fletterick RJ

    (1996) Crystal structure of the motor domain of the kinesin-related motor ncd

    Nature 380:555–559.

    https://doi.org/10.1038/380555a0

    • Google Scholar
61. 1. Sali A
    2. Blundell TL

    (1993) Comparative protein modelling by satisfaction of spatial restraints

    Journal of Molecular Biology 234:779–815.

    https://doi.org/10.1006/jmbi.1993.1626

    • Google Scholar
62. 1. Scarabelli G
    2. Grant BJ

    (2013) Mapping the structural and dynamical features of kinesin motor domains

    PLOS Computational Biology 9:e1003329.

    https://doi.org/10.1371/journal.pcbi.1003329

    • Google Scholar
63. 1. Schnitzer MJ
    2. Visscher K
    3. Block SM

    (2000) Force production by single kinesin motors

    Nature Cell Biology 2:718–723.

    https://doi.org/10.1038/35036345

    • Google Scholar
64. 1. Shen MY
    2. Sali A

    (2006) Statistical potential for assessment and prediction of protein structures

    Protein Science 15:2507–2524.

    https://doi.org/10.1110/ps.062416606

    • Google Scholar
65. 1. Sindelar CV

    (2011) A seesaw model for intermolecular gating in the kinesin motor protein

    Biophysical Reviews 3:85–100.

    https://doi.org/10.1007/s12551-011-0049-4

    • Google Scholar
66. 1. Sindelar CV
    2. Downing KH

    (2010) An atomic-level mechanism for activation of the kinesin molecular motors

    Proceedings of the National Academy of Sciences of USA 107:4111–4116.

    https://doi.org/10.1073/pnas.0911208107

    • Google Scholar
67. 1. Sindelar CV
    2. Downing KH

    (2007) The beginning of kinesin's force-generating cycle visualized at 9-A resolution

    The Journal of Cell Biology 177:377–385.

    https://doi.org/10.1083/jcb.200612090

    • Google Scholar
68. 1. Skiniotis G
    2. Surrey T
    3. Altmann S
    4. Gross H
    5. Song YH
    6. Mandelkow E
    7. Hoenger A

    (2003) Nucleotide-induced conformations in the neck region of dimeric kinesin

    The EMBO Journal 22:1518–1528.

    https://doi.org/10.1093/emboj/cdg164

    • Google Scholar
69. 1. Song H
    2. Endow SA

    (1998) Decoupling of nucleotide- and microtubule-binding sites in a kinesin mutant

    Nature 396:587–590.

    https://doi.org/10.1038/25153

    • Google Scholar
70. 1. Soppina V
    2. Norris SR
    3. Dizaji AS
    4. Kortus M
    5. Veatch S
    6. Peckham M
    7. Verhey KJ

    (2014) Dimerization of mammalian kinesin-3 motors results in superprocessive motion

    Proceedings of the National Academy of Sciences of USA 111:5562–5567.

    https://doi.org/10.1073/pnas.1400759111

    • Google Scholar
71. 1. Soppina V
    2. Verhey KJ

    (2014) The family-specific K-loop influences the microtubule on-rate but not the superprocessivity of kinesin-3 motors

    Molecular Biology of the Cell 25:2161–2170.

    https://doi.org/10.1091/mbc.E14-01-0696

    • Google Scholar
72. 1. Sosa H
    2. Dias DP
    3. Hoenger A
    4. Whittaker M
    5. Wilson-Kubalek E
    6. Sablin E
    7. Fletterick RJ
    8. Vale RD
    9. Milligan RA

    (1997) A model for the microtubule-Ncd motor protein complex obtained by cryo-electron microscopy and image analysis

    Cell 90:217–224.

    https://doi.org/10.1016/S0092-8674(00)80330-X

    • Google Scholar
73. 1. Sosa H
    2. Milligan RA

    (1996) Three-dimensional structure of ncd-decorated microtubules obtained by a back-projection method

    Journal of Molecular Biology 260:743–755.

    https://doi.org/10.1006/jmbi.1996.0434

    • Google Scholar
74. 1. Sousa D
    2. Grigorieff N

    (2007) Ab initio resolution measurement for single particle structures

    Journal of Structural Biology 157:201–210.

    https://doi.org/10.1016/j.jsb.2006.08.003

    • Google Scholar
75. 1. Stokin GB
    2. Goldstein LS

    (2006) Linking molecular motors to Alzheimer's disease

    Journal of Physiology 99:193–200.

    https://doi.org/10.1016/j.jphysparis.2005.12.085

    • Google Scholar
76. 1. Svoboda K
    2. Schmidt CF
    3. Schnapp BJ
    4. Block SM

    (1993) Direct observation of kinesin stepping by optical trapping interferometry

    Nature 365:721–727.

    https://doi.org/10.1038/365721a0

    • Google Scholar
77. 1. Tomishige M
    2. Vale RD

    (2000) Controlling kinesin by reversible disulfide cross-linking. Identifying the motility-producing conformational change

    The Journal of Cell Biology 151:1081–1092.

    https://doi.org/10.1083/jcb.151.5.1081

    • Google Scholar
78. 1. Topf M
    2. Lasker K
    3. Webb B
    4. Wolfson H
    5. Chiu W
    6. Sali A

    (2008) Protein structure fitting and refinement guided by cryo-EM density

    Structure 16:295–307.

    https://doi.org/10.1016/j.str.2007.11.016

    • Google Scholar
79. 1. Uchimura S
    2. Oguchi Y
    3. Hachikubo Y
    4. Ishiwata S
    5. Muto E

    (2010) Key residues on microtubule responsible for activation of kinesin ATPase

    The EMBO Journal 29:1167–1175.

    https://doi.org/10.1038/emboj.2010.25

    • Google Scholar
80. 1. Vale RD

    (1996) Switches, latches, and amplifiers: common themes of G proteins and molecular motors

    The Journal of Cell Biology 135:291–302.

    https://doi.org/10.1083/jcb.135.2.291

    • Google Scholar
81. 1. Vale RD

    (2003) The molecular motor toolbox for intracellular transport

    Cell 112:467–480.

    https://doi.org/10.1016/S0092-8674(03)00111-9

    • Google Scholar
82. 1. Vale RD
    2. Milligan RA

    (2000) The way things move: looking under the hood of molecular motor proteins

    Science 288:88–95.

    https://doi.org/10.1126/science.288.5463.88

    • Google Scholar
83. 1. Varga V
    2. Helenius J
    3. Tanaka K
    4. Hyman AA
    5. Tanaka TU
    6. Howard J

    (2006) Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner

    Nature Cell Biology 8:957–962.

    https://doi.org/10.1038/ncb1462

    • Google Scholar
84. 1. Visscher K
    2. Schnitzer MJ
    3. Block SM

    (1999) Single kinesin molecules studied with a molecular force clamp

    Nature 400:184–189.

    https://doi.org/10.1038/22146

    • Google Scholar
85. 1. Woehlke G
    2. Ruby AK
    3. Hart CL
    4. Ly B
    5. Hom-Booher N
    6. Vale RD

    (1997) Microtubule interaction site of the kinesin motor

    Cell 90:207–216.

    https://doi.org/10.1016/S0092-8674(00)80329-3

    • Google Scholar
86. 1. Yun M
    2. Zhang X
    3. Park CG
    4. Park HW
    5. Endow SA

    (2001) A structural pathway for activation of the kinesin motor ATPase

    The EMBO Journal 20:2611–2618.

    https://doi.org/10.1093/emboj/20.11.2611

    • Google Scholar

Author details

1. Joseph Atherton

   Institute of Structural and Molecular Biology, Department of Biological Sciences, Birkbeck College, University of London, London, United Kingdom

   Contribution

   JA, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Irene Farabella

   Institute of Structural and Molecular Biology, Department of Biological Sciences, Birkbeck College, University of London, London, United Kingdom

   Contribution

   IF, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

3. I-Mei Yu

   Structural Motility, Institut Curie, Centre National de la Recherche Scientifique, Paris, France

   Contribution

   I-MY, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

4. Steven S Rosenfeld

   Department of Cancer Biology, Lerner Research Institute, Cleveland Clinic, Cleveland, United States

   Contribution

   SSR, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

5. Anne Houdusse

   Structural Motility, Institut Curie, Centre National de la Recherche Scientifique, Paris, France

   Contribution

   AH, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

6. Maya Topf

   Institute of Structural and Molecular Biology, Department of Biological Sciences, Birkbeck College, University of London, London, United Kingdom

   Contribution

   MT, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

7. Carolyn A Moores

   Institute of Structural and Molecular Biology, Department of Biological Sciences, Birkbeck College, University of London, London, United Kingdom

   Contribution

   CAM, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   c.moores@mail.cryst.bbk.ac.uk

   Competing interests

   The authors declare that no competing interests exist.

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