Research Highlights

1) Recombinant expression of dynein

A large body of structural, biochemical and biophysical experiments on other motor proteins, such as myosin V and kinesin, has led to their mechanism of movement being understood in considerable detail. Underpinning much of this research has been the ability to recombinantly express, tag and manipulate their genes as well as the ability to produce them as dimers that move processively for multiple steps. The large size of the dynein gene has, until recently [1, 2], made recombinant expression of its motor domain particularly challenging.

Using homologous recombination to modify the cytoplasmic dynein gene in S. cerevisiae. In the first round a URA3 gene is inserted into the dynein gene. In the second step this URA3 marker is replaced with the desired modification.

Saccharomyces cerevisae (bakerís yeast) offers two significant advantages for the study of dynein. The first is that dynein is not essential for growth, which allows the genomic copy of the gene to be deleted or modified at will. The second is that homologous recombination can be used to alter only the desired region. This does away with the need for dealing with large plasmids, difficult ligation reactions and the use of rare restriction enzyme sites and makes it relatively simple to manipulate the gene [1]. A typical scheme for truncating dynein, adding some tags and a strong promoter is shown in the figure to your right.

Top:An engineered dynein construct for motility assays: the dynein motor is dimerised by fusing it to GST. The head is tagged with the genetic HaloTag (Promega) that can be labelled with different fluorescent dyes such as rhodamine.

Bottom:Press the play button for a single molecule movie of this construct moving along axonemes Click here to download movie

Using this system we artificially dimerised the dynein motor domain using the constitutive dimer glutathione-S-transferase (GST) [3]. We also added a HaloTag from Promega, which is a genetically encoded tag that can be labelled very specifically with a range of different ligands. This allowed us to tag the protein with stable fluorescent dyes, such as rhodamine, or even quantum dots (inorganic crystals that fluoresce very brightly under UV light). The movie (press the play button) shows single molecules of these fluorescently labelled GST-dynein constructs moving along axonemes. These constructs have now helped open the door to the study of dynein.

2) High precision tracking of single molecules.

The movie in the above section shows single, fluorescently labelled dynein molecules moving along microtubules. It is possible to track the position of these fluorescent spots with a precision of a few nanometers as long as they are bright enough [4, 5]. This means that it is possible to follow the individual steps that a motor protein makes as it walks along a microtubule. The two traces in the figure below show the movement of a kinesin and a dynein molecule tagged with a very bright fluorescent molecule called a quantum dot. The kinesin molecule takes regular 16nm steps, which are defined by the spacing of tubulin molecules [6]. In contrast and not unexpectedly given its size and long stalk, the movement of dynein is much less regular and exhibits a large number of forward and backward movements.

Traces of kinesin and dynein tagged with quantum dots stepping along microtubules. Note the traces are oriented for ease of display. Dynein and kinesin in fact move in opposite directions.

Cartoon of dynein construct during movement.

By attaching quantum dots to different parts of the molecule, we were able to propose an initial model for how dynein moves [3]. In the future, advances in this high precision tracking, such as the ability to monitor two different colour fluorophores simultaneously [7], will allow us to build up a complete picture of how dynein moves along microtubules. Some questions that need to be answered include what angle the stalks make with respect to the microtubule and whether its orientation changes as dynein moves.

3) Structure of the dynein stalk

One of the remarkable features of dynein is that its microtubule binding domain (MTBD) is found at the tip of a long stalk [8] and is thus separated by over 10 nm from the rest of the motor. This is in stark contrast to the kinesin and myosin motors where the polymer binding site is an intrinsic part of the motor domain and raises the question of how the two parts of dynein communicate with each other. Such communication is a key part of how motor proteins couple the cycle of ATP hydrolysis to movement. Nucleotide hydrolysis and conformational changes in the AAA+ ring that drive motility occur only after dynein has bound to microtubules. Reciprocally, the MTBD is released from microtubules when ATP binds to the AAA+ ring [9]. An explanation for the communication mechanism has now begun to emerge from recent biochemical experiments and the determination of the crystal structure of dyneinís MTBD, all carried out in collaboration with Ian Gibbons (UC Berkeley) [10, 11].

A) Fusion of seryl-tRNA synthetase (cyan) and the tip of the dynein stalk (red). B) Close up of the dynein stalk (helices CC1 and CC2) and the rest of the microtubule binding domain colored in a rainbow of colors from purple (CC1) to red (CC2). Helices H1, H3 and H6 are proposed to form the surface that interacts with microtubules. The conserved proline residues are shown in red spacefilling representation.

In order to produce a stable protein construct for crystallization, the dynein MTBD and the top part of the stalk, which forms an antiparallel coiled coil of two alpha helices (CC1 and CC2), was fused into the coiled coil of a protein called seryl-tRNA synthetase (SRS). The 2.3Å crystal structure we solved confirms the coiled-coil nature of the dynein stalk and shows that it is kinked by the presence of two highly conserved proline residues. The rest of the MTBD consists of a novel fold of alpha helices that pack against the top of the stalk. The top face (including helices H1, H3, and H6) was identified as the site of interaction with microtubules, based on mutations that interfere with binding. Confirmation was provided by docking the crystal structure into an electron-density map of a dynein stalk bound to microtubules that was obtained by cryoelectron microscopy.

A) Cryo electron microscopy (EM) reconstruction of dynein stalk (blue) bound to microtubules (green). B) Close up showing a tubulin protofilament (made up of α and β tubulins) and a dynein microtuble binding domain (MTBD) docked into the EM map (blue mesh).

We propose that the mechanism of communication along the stalk involves a sliding movement of helix CC1 with respect to the helix CC2 [10, 11]. This model is based on 3 lines of evidence.

  1. Changing the fusion site between the stalk and the SRS base by removing exactly four amino acids from CC1 leads to an almost ten fold increase in the affinity of the MTBD for microtubules. This suggests that conformational changes in the AAA ring could alter the affinity of the MTBD by driving a sliding of CC1 relative to CC2 by the equivalent of four amino acids (see the movie below for an animation).

    Movie showing a model for stalk sliding. The first frame is a close up of the stalk region of the dynein MTBD crystal structure showing the amino acids that pack together in the coiled coil's core (dark blue). The residues from seryl-tRNA synthetase are shown in red. Removal of 4 amino acids (cyan) from CC1 increases the affinity of the MTBD for microtubules.

    This movie will simulate removal of these amino acids and show how the left hand helix (CC1) can slide. Some residues slide from one side of the core to the other, while others (marked with arrows) will move out of the core all together. When you press play you will see residues appear (marked in yellow) that will move into the core in their place. Then the cyan region of CC1 will be removed and the remainder will slide downwards. Click here to download movie

  2. In a standard coiled coil, such a sliding motion would be highly entropically unfavourable as it would expose some of the hydrophobic residues in the core of the coiled coil (see the movie on the website). The dynein stalk, however, has a conserved feature which may lower the barrier to such a conformational change. The residues in the core of a standard coiled-coil are arranged as two offset stripes of hydrophobic amino acids running up each of the two helices. In the dynein stalk the residues in CC2 mostly follow this regular pattern (see figure below). By contrast, CC1 has one stripe of hydrophobic residues and one of hydrophilic residues. The consequence of this arrangement is that if CC1 slides relative to CC2 then the only residues to be exposed would be the hydrophilic ones whereas the hydrophobic residues simply move from one side of the coiled-coil core to the other. As no hydrophobic residues are exposed to the solvent, the entropic cost of sliding should be considerably reduced in comparison to a regular coiled coil that has predominantly hydrophobic amino acids in its core.

    A conserved feature of the stalk may aid sliding of the two helices in the stalk coiled coil. Left panel: spacefilling model of the stalk showing the two helices in stalk (CC1 in blue & CC2 in red) and the microtubule binding domain in white. Right panel: the two helices have been open up to show the amino acids that are packed together to form their core. These residues are arranged in a zig-zag pattern, marked with arrows. In CC2 most of the residues are hydrophobic (green) whereas in CC1, those on the left are hydrophobic, whereas those on the right are hydrophilic (orange).

  3. Other features of the stalk structure also hint that helix sliding may provide the communication mechanism. CC2 makes multiple contacts with other helices in the MTBD (See figure below), whereas CC1 only makes a small number of interactions with a single helix (H4). This arrangement is consistent with CC1 being relatively free to move, while CC2 is held in place by multiple interactions. Furthermore CC1 is followed by one of the main helices at the microtubule binding interface (H1). This suggests that microtubule binding would cause a rearrangement in the position of H1, which would be propagated back up to the AAA ring by a sliding of CC1 against the relatively rigid CC2. Reciprocally nucleotide driven conformational changes in the AAA ring would, via the movement of CC1, alter the conformation of H1 and thus change the affinity of dynein for the microtubule.

    Cartoon showing the sequence of amino acids in the dynein stalk. Blue lines show packing of residues in the coiled-coil core. Residues that contact other parts of the microtubule binding domain are marked with a red dot. In comparison to the extensive contacts made by CC2, the first helix (CC1) makes only a small number of contacts with H4. This is consistent with CC2 being relatively rigid, while CC1 can move in response to changes at the microtubule binding interface.


  1. Reck-Peterson, S.L. and R.D. Vale, Molecular dissection of the roles of nucleotide binding and hydrolysis in dynein's AAA domains in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A, 2004. 101(6): p. 1491-5.(pdf)
  2. Nishiura, M., et al., A single-headed recombinant fragment of Dictyostelium cytoplasmic dynein can drive the robust sliding of microtubules. J Biol Chem, 2004. 279(22): p. 22799-802.
  3. Reck-Peterson, S.L., et al., Single-molecule analysis of dynein processivity and stepping behavior. Cell., 2006. 126(2): p. 335-48.
  4. Yildiz, A., et al., Myosin V walks hand-over-hand: single fluorophore imaging with 1.5-nm localization. Science, 2003. 300(5628): p. 2061-5.
  5. Thompson, R.E., D.R. Larson, and W.W. Webb, Precise nanometer localization analysis for individual fluorescent probes. Biophys J, 2002. 82(5): p. 2775-83.
  6. Yildiz, A., et al., Kinesin walks hand-over-hand. Science, 2004. 303(5658): p. 676-8.
  7. Churchman, L.S., et al., Single molecule high-resolution colocalization of Cy3 and Cy5 attached to macromolecules measures intramolecular distances through time. Proc Natl Acad Sci U S A, 2005. 102(5): p. 1419-23.
  8. Gee, M.A., J.E. Heuser, and R.B. Vallee, An extended microtubule-binding structure within the dynein motor domain. Nature, 1997. 390(6660): p. 636-9.
  9. Numata, N., et al., Molecular mechanism of force generation by dynein, a molecular motor belonging to the AAA+ family. Biochem Soc Trans, 2008. 36(Pt 1): p. 131-5.
  10. Carter, A.P., et al., Structure and functional role of dynein's microtubule-binding domain. Science, 2008. 322(5908): p. 1691-5.(pdf)
  11. 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, 2005. 280(25): p. 23960-5.(pdf)