Lis1 regulates dynein by sterically blocking its mechanochemical cycle (original) (raw)

Abstract

Regulation of cytoplasmic dynein's motor activity is essential for diverse eukaryotic functions, including cell division, intracellular transport, and brain development. The dynein regulator Lis1 is known to keep dynein bound to microtubules; however, how this is accomplished mechanistically remains unknown. We have used three-dimensional electron microscopy, single-molecule imaging, biochemistry, and in vivo assays to help establish this mechanism. The three-dimensional structure of the dynein–Lis1 complex shows that binding of Lis1 to dynein's AAA+ ring sterically prevents dynein's main mechanical element, the ‘linker’, from completing its normal conformational cycle. Single-molecule experiments show that eliminating this block by shortening the linker to a point where it can physically bypass Lis1 renders single dynein motors insensitive to regulation by Lis1. Our data reveal that Lis1 keeps dynein in a persistent microtubule-bound state by directly blocking the progression of its mechanochemical cycle.

DOI: http://dx.doi.org/10.7554/eLife.03372.001

Research organism: S. cerevisiae

eLife digest

Cells use motor proteins to move ‘cargo’ from one location to another inside the cell. This cargo can range in size from a single macromolecule to something as large as the nucleus of the cell. A motor protein called dynein is the largest and least understood of the motor proteins found in cells.

Dynein molecules work in pairs to take ‘steps’ along tracks called microtubules. Dynein contains two domains: a motor domain, which is responsible for generating movement, and a ‘tail’ domain to which the cargo is attached. The motor domain is composed of a ring-like shape and two appendages—the stalk and the linker. The linker undergoes large-scale movements relative to the ring that transmits force to the tail domain.

Dynein also interacts with various accessory proteins to do its job inside the cell. One of these is a protein called Lis1 that is found across a wide range of species from yeast to humans. Defects in the gene for Lis1 result in brain developmental disorders in humans. However, it is not clear how the Lis1 protein influences the activity of dynein.

Now Toropova, Zou et al. have visualized the structure of dynein bound to Lis1 and compared it with the structure of dynein on its own in order to work out if dynein changes its shape as a result of binding to Lis1. These experiments show that when Lis1 binds to dynein, it physically blocks the linker, preventing it from making contacts with the ring-like shape that are important for the normal function of the motor.

To test the idea that this physical block is responsible for dynein molecules spending a relatively long time attached to their microtubules, Toropova, Zou et al. shortened the linker to a point where the Lis1 protein could no longer block it: this resulted in a dynein motor that was no longer sensitive to Lis1. A challenge for the future is to understand, at a molecular level, how the Lis1-mediated slowing down of dynein affects the multiple functions the motor carries out in a cell.

DOI: http://dx.doi.org/10.7554/eLife.03372.002

Introduction

Cytoplasmic dynein (‘dynein’ here), the largest and least understood of the cytoskeletal motors, uses the energy from ATP hydrolysis to move towards the minus ends of microtubules (Vale, 2003; Carter, 2013). As the major minus-end-directed motor in most eukaryotic cells, dynein's many roles include transporting a range of macromolecular cargo (Blocker et al., 1997; Jordens et al., 2001; Kural et al., 2005; Pilling et al., 2006; Driskell et al., 2007), constructing and positioning the mitotic spindle (Heald et al., 1996; Merdes et al., 1996; Kiyomitsu and Cheeseman, 2013), and polarizing and anchoring mRNAs during development (Wilkie and Davis, 2001). To perform its diverse biological functions, dynein partners with a range of regulatory co-factors, an important subset of which can alter dynein motility directly. Despite progress in understanding the architecture and mechanism of dynein's large motor domain, how this structure is acted upon by regulatory factors is not yet known.

Dynein is a homodimer of force generating units (∼500 kDa each) (Figure 1A,B). The N-terminal region of each monomer forms the ‘tail’ domain, which mediates dimerization and cargo attachment via adaptor proteins. Removal of the tail yields the ‘motor’, the minimal portion of dynein that can exert force. At the core of the motor are six AAA+ modules (AAA1–6) that fold into a ring. AAA1 is the main site of ATP hydrolysis for motility but AAA2, 3, and 4 can also bind ATP, and AAA3 and 4 can hydrolyze it (Gibbons et al., 1991; Kon et al., 2004, 2012; Cho et al., 2008; Schmidt et al., 2012). AAA5 and AAA6 have lost the ability to bind nucleotide (Kon et al., 2012; Schmidt et al., 2012). Two appendages to the ring are essential for dynein function; the ‘stalk’, an intramolecular anti-parallel coiled-coil at the end of which lies the microtubule-binding domain (Gee et al., 1997; Carter et al., 2008) and the ‘linker’, which is dynein's key mechanical element and is an elongated structure N-terminal to AAA1. The linker spans the ring and moves in a nucleotide dependent manner that is thought to transmit force to dynein's cargo (Burgess et al., 2003; Kon et al., 2005; Shima et al., 2006; Roberts et al., 2009, 2012). In order for dynein to move along microtubules, ATP binding/hydrolysis at AAA1 must be coupled with linker motion and microtubule binding and release at the tip of the stalk, located 250 Å away (Gibbons et al., 2005; Imamula et al., 2007; Kon et al., 2009; Redwine et al., 2012).

Figure 1. The binding of Lis1 to dynein changes the position of dynein's linker domain.

(A) Domain organization of dynein and Lis1 constructs used in this study. Dynein's AAA+ domains are labeled AAA1–6. MTBD: microtubule binding domain; CC: coiled coil; LisH: Lis-homology (dimerization) motif. (B) Schematic representation of dynein and Lis1, color-coded as in (A) and throughout the paper. NT: N terminus; CT: C terminus. (C) Cryo-NS EM reconstruction of the dynein motor domain in complex with Lis1 and (D) of the motor domain alone. AAA4 and AAA5 are labeled. A density present only in the dynein–Lis1 map is highlighted in (C) (brown arrowhead). The linker occupies different positions in the two maps (compare labeled densities and gray arrows), and its position in the dynein alone map is sterically incompatible with Lis1, as indicated by a semi-transparent Lis1 density. (E) Structural model of dynein's motor domain docked into the EM maps of dynein–Lis1 and (F) dynein alone. The model was built from crystal structures of the S. cerevisiae dynein ring (PDB ID: 4AKG [Schmidt et al., 2012]) and D. discoideum linker aligned to the yeast linker position (PDB ID: 3VKG [Kon et al., 2012]), the D. discoideum linker being closer in length to that in our EM construct. In (E), a homology model of the S. cerevisiae Lis1 β-propeller (brown) has been docked into the new density highlighted in (C). The linker domain in the EM map (gray arrow) is shifted away from its position in the crystal structure (purple arrow), which protrudes from the EM density and clashes with the Lis1 density. In contrast, the linker is within the EM density in the dynein alone map (F). Green circle: location of known interactions between the linker and AAA5 module in dynein (Schmidt et al., 2012). (G) Close-up view of the Lis1 density with homology model docked in, viewed along the axis indicated by the arrowhead in (C). (H) A rotated, smaller view of (E), showing the interface between Lis1 (brown arrowhead) and dynein.

DOI: http://dx.doi.org/10.7554/eLife.03372.003

Figure 1—figure supplement 1. Three-dimensional (3D) classification and refinement of the dynein and dynein–Lis1 reconstructions.

(A) SDS-PAGE of dynein and Lis1, affinity purified from S. cerevisiae. (B) Comparison between re-projections of the dynein and dynein–Lis1 reconstructions and the best-matching reference-free class averages (no nucleotide conditions). (C) Fourier Shell Correlation plots for all EM maps presented. The plots are shown as a function of resolution (1/frequency). The thresholds for the 0.5 FSC and 0.143 FSC criteria are shown. (D and E) Class averages of monomeric dynein in complex with dimeric (D) or monomeric (E) Lis1, with a schematic representation alongside each. (F–H) 3D classification of linker positions in the dynein–Lis1 (no nucleotide) data set. (F) A reconstruction using the entire data set without sorting out linker conformations. The N-terminus of the linker is mostly averaged out in this map. (G) 3D class average of a subset of dynein motors whose linkers are located on the AAA5 proximal side of Lis1. The purple arrow points to the N-terminus of the linker. (H) The linker density from (G) was overlaid on the dynein–Lis1 structure (Figure 1C) to highlight the different positions adopted by the linker in the presence of Lis1. The purple and gray arrows point to the N-terminus of the linker domain in the two conformations. The Lis1 density is indicated in all three maps. (I) A side view of the dynein motor domain in surface representation with the linker domain colored by conservation (100%, purple; 11%, white). The alignment was carried out using cytoplasmic dynein from the following species: M. musculus, H. sapiens, S. cerevisiae, C. albicans, A. fumigatus, A. nidulans, D. discoideum, D. melanogaster, and C. elegans using Muscle (Edgar, 2004). For orientation, a cartoon representation of the view is shown bottom right. The truncation site for the short linker dynein is marked with a dashed line and scissors (see Figure 5).

Figure 1—figure supplement 2. The linker's displaced position in the presence of Lis1 does not appear to involve a specific interaction with AAA4.

(A) Zoomed out view of dynein–Lis1; only the portion of the crystal structure corresponding to AAA4 is displayed, in yellow (PDB ID: 4AKG [Schmidt et al., 2012]). (B) Close-up of the N-terminal portion of the linker (left) and a view rotated by 60° (right). The latter shows a density connecting the linker and AAA4. Residues in an AAA4 helix that are located in the density connecting AAA4 to the linker are shown in atomic representation, colored by element, and labeled. (C) Kymographs of in vitro motility experiments with TMR-labeled wild-type GST-dynein331kDa or mutant GST-dynein331kDa with the five residues labeled in (C) changed to alanine (AAA4 mut). Assays were performed with dynein alone or in the presence of 200 nM Lis1. Horizontal scale bar = 2 μm, vertical = 30 s. (D) Histogram of mean velocities for each experiment ± S.D., N = 228–612, ***p < 0.0001.

Across a wide range of species, dynein interacts with a conserved regulator called Lis1 (also known as Pac1 in Saccharomyces cerevisiae) that is necessary for many dynein driven processes. Mutations in the Lis1 gene cause the neurodevelopmental disorder lissencephaly (Reiner et al., 1993). Lis1 is the only dynein regulator known to interact directly with its motor domain (McKenney et al., 2010; Huang et al., 2012). Like dynein, Lis1 acts as a dimer, with each monomer comprising an N-terminal dimerization domain (LisH) followed by a coiled-coil, a flexible loop and a C-terminal β-propeller domain of 7 WD motifs (Kim et al., 2004; Tarricone et al., 2004) (Figure 1A,B). We previously showed that the propeller domain alone can regulate dynein in vitro and used negative stain electron microscopy (EM) and two-dimensional (2D) image processing to show that Lis1 binds to dynein's motor domain near AAA3/4 (Huang et al., 2012). We and others have shown that Lis1 induces a slow-moving microtubule-attached state in dynein (Yamada et al., 2008; McKenney et al., 2010; Torisawa et al., 2011; Huang et al., 2012). Interestingly, Lis1 can accomplish this without substantially affecting dynein's overall ATP hydrolysis rate (Yamada et al., 2008; McKenney et al., 2010; Huang et al., 2012). This led us to propose that Lis1 acts as a ‘clutch’, uncoupling dynein's engine (AAA+ ring) from its track-binding region.

Our previous 2D EM data, which indicated that Lis1 binds in the vicinity of AAA3/4, raised at least two possibilities for how Lis1 can affect dynein's mechanochemistry. On the one hand, Lis1 may regulate dynein allosterically, influencing the structure or motions of AAA3 or AAA4 in the ring, and thus preventing the propagation of a signal for microtubule detachment to the stalk. Alternatively, because the linker domain lies close to AAA4 in certain nucleotide-states, Lis1 may regulate dynein sterically, affecting the linker's movement directly, and thus dynein's mechanochemistry. Establishing Lis1's mode of regulation is not possible without three-dimensional (3D) data, as it is not known if Lis1 binds on the same face of the AAA+ ring as the linker, as would be required for direct Lis1–linker interactions to occur. Moreover, it is not clear to what extent the Lis1 binding site encompasses AAA3, AAA4, or both of these modules. Thus, 3D structural information is critical to understanding the mechanistic basis of Lis1's regulation of dynein.

We set out to establish how Lis1 induces a persistent microtubule-bound state in dynein. We obtained the 3D structure of S. cerevisiae dynein bound to Lis1 to determine which elements of the motor Lis1 directly affects. Our structure revealed that Lis1 sterically prevents the linker from reaching its normal post-powerstroke locations on the ATP hydrolyzing ring that are involved in its conformational cycle. Structure-based mutagenesis also allowed us to identify residues in Lis1 responsible for binding to dynein. Single molecule analysis of a dynein motor with a shortened linker that can physically bypass Lis1 indicated that removing the steric block renders dynein insensitive to Lis1. Our combined data show that Lis1 directly blocks dynein's mechanochemical cycle, inducing a persistent microtubule-bound state, by acting on its linker domain.

Results

Structure of the dynein–Lis1 complex

In order to visualize the spatial relationship between Lis1 and dynein's multiple structural elements and to better understand the mechanism by which Lis1 regulates dynein, we used cryo-negative stain (cryo-NS) EM and single-particle image processing to obtain the 3D structure of the dynein–Lis1 complex (Figure 1C). Cryo-NS combines the structural preservation of vitrification with the high contrast provided by the negative stain (De Carlo and Stark, 2010). We found this increased contrast to be instrumental to our ability to computationally sort the different conformations that co-existed in most of our samples. We also determined a 3D map of dynein alone, as a reference, to establish whether Lis1 alters dynein's structure (Figure 1D). We used a well-characterized monomeric dynein construct (Reck-Peterson et al., 2006) but chose to use dimeric rather than monomeric Lis1 for our reconstructions. We previously showed that while a Lis1 monomer is sufficient to slow down dynein, a much higher concentration of it is required (Huang et al., 2012), presumably due to the high local concentration of the β-propeller in the context of a Lis1 dimer. Using a Lis1 dimer at much lower concentrations allowed us to minimize the background in our images.

Both dynein and Lis1 were expressed from S. cerevisiae at their genomic loci (Table 1). We imaged dynein–Lis1 and dynein alone in the absence of nucleotide and obtained structures at resolutions of 21 Å for the complex (Figure 1C,E, Figure 1—figure supplement 1C and Video 1) and of 15 Å for dynein alone (Figure 1D,F, Figure 1—figure supplement 1C and Video 2). The dynein alone map accommodates the crystal structures of the dynein motor domain well (Kon et al., 2012; Schmidt et al., 2012) (Figure 1F), with a Fourier Shell Correlation between the EM map and yeast motor domain structure (Schmidt et al., 2012) of 0.143 at a resolution of 18.8 Å.

Video 1. The three-dimensional structure of dynein–Lis1.

The movie shows the 3D reconstruction of dynein in complex with Lis1 with 360° rotation about the Y-axis. After this rotation, the EM density is made transparent to display the docked dynein crystal structure model and Lis1 homology model and is again rotated by 360° about the Y-axis.

DOI: http://dx.doi.org/10.7554/eLife.03372.007

Video 2. The three-dimensional structure of dynein.

The movie shows the 3D reconstruction of dynein alone with 360° rotation about the Y-axis. After this rotation, the EM density is made transparent to display the docked dynein crystal structure model and is again rotated by 360° about the Y-axis.

DOI: http://dx.doi.org/10.7554/eLife.03372.008

Table 1.

The dynein–Lis1 map shows two major differences relative to the dynein alone reconstruction. First, a prominent donut-shaped density is resolved in contact with the dynein ring, adjacent to the stalk (Figure 1C, brown arrowhead). This extra density matches the dimensions of a β-propeller, including the hole at its center (Figure 1E,G,H). We thus conclude that the density corresponds to Lis1. The second, and striking difference between the two maps is in dynein itself: Lis1 binds on the same face of the ring where dynein's linker domain is located and the linker is displaced by ∼44 Å in the dynein–Lis1 map relative to the dynein alone reconstruction (Figure 1C–F).

One dynein ring binds one Lis1 β-propeller

Our previous 2D image analysis of the dynein–Lis1 complex did not allow us to determine whether their interaction involved one or both of Lis1's β-propellers or whether Lis1's N-terminal LisH dimerization domain was part of the interaction as well. The extra density in the dynein–Lis1 3D map fits well a single homology model of the S. cerevisiae Lis1 β-propeller built from the crystal structure of the mouse protein (Tarricone et al., 2004) (Figure 1E,G,H). Because our map resolved the hole at the center of the propeller, the homology model could be unambiguously docked within the density in terms of its translation (Figure 1G).

In Lis1, the β-propeller is connected to the N-terminal LisH dimerization domain by a loop, predicted to be flexible, and a coiled coil (Kim et al., 2004; Tarricone et al., 2004). Consequently, the rest of Lis1 would be expected to adopt a wide range of positions relative to the dynein-bound propeller domain. In agreement with this, we did not resolve density beyond that of the single β-propeller in our dynein–Lis map. 2D image analysis of dynein–Lis1 complexes with either monomeric or dimeric Lis1 showed the same density and location for Lis1 (Figure 1—figure supplement 1D,E), further supporting a stoichiometry of one Lis1 propeller to one dynein motor domain.

Dissection of the dynein–Lis1 interface

Our structure of the dynein–Lis1 complex shows that the Lis1 β-propeller contacts dynein primarily at a surface-exposed helix at the junction of AAA3 and AAA4 (Figure 2A and Video 3), explaining why mutagenesis of four conserved, charged residues (KDEE) on this helix virtually abolished Lis1 binding and dynein regulation (Huang et al., 2012). Since the resolution of the reconstruction does not allow us to determine unambiguously the rotational orientation of the Lis1 homology model within the corresponding density, we used mutagenesis to probe the dynein–Lis1 interface and further constrain our model of the complex.

Video 3. The dynein–Lis1 interface.

The movie shows the 3D reconstruction of dynein–Lis1, with the crystal structure of the dynein motor domain and the Lis1 homology model docked in. After a few frames, the EM density disappears to show only the atomic structures and the view changes to show the interaction between dynein and Lis1 in closer detail, finishing with an open-book view of Lis1. The conserved residues that were mutated in Lis1 are annotated as well as the conserved residues in the AAA4 helix in dynein that have been shown to be necessary for Lis1 binding. Note: the rotational fit of the Lis1 propeller within the Lis1 EM density is uncertain at the current resolution of the dynein–Lis1 map.

DOI: http://dx.doi.org/10.7554/eLife.03372.012

Figure 2. Disrupting the putative dynein–Lis1 interface impairs Lis1's ability to bind to and regulate dynein.

(A) The Lis1 β-propeller engages dynein primarily at a surface helix connecting AAA3 and AAA4 (yellow arrowhead, see Video 3). Inset: a zoomed out view. (B) (Left) View along the axis highlighted in (A) by the eye/arrow; (right) rotated view. Except for the helix (yellow), the dynein density was removed for clarity. Five conserved residues on Lis1 that were mutated to alanine, either in combination (Lis15A) or individually, are labeled and shown in atomic representation. Also displayed are residues (KDEE) in dynein known to be involved in the interaction with Lis1 (Huang et al., 2012). Basic and acidic residues are labeled in blue and red, respectively. (C) No co-migration of dynein and Lis1 was detected by size-exclusion chromatography with the Lis15A and Lis1R378A mutants. Traces show elution profiles of GST-dynein331kDa (‘dynein’) with wild-type Lis1 (black), Lis15A (purple) and Lis1R378A (green). SDS-PAGE for collected fractions are shown below. (D) Kymographs of in vitro motility experiments with TMR-labeled GST-dynein331kDa alone or in the presence of 200 nM wild-type or mutant Lis1. Horizontal scale bar = 2 μm, vertical = 20 s, N = 274–542. (E) In vivo spindle oscillation assays comparing S. cerevisiae strains carrying either wild-type or mutant Lis1 or full deletions of dynein or Lis1. Inset is a Z-projection of a dividing cell with markers for the membrane (purple) and spindle pole bodies (SPBs) (cyan). BN = bud neck. Bud neck crossings by the SPBs were counted over 20 min. WT N = 53, DyneinΔ N = 32, Lis1Δ N = 55, Lis1R378A N = 58, Lis15A N = 47. For each strain the mean and SE are shown.

DOI: http://dx.doi.org/10.7554/eLife.03372.009

Figure 2—figure supplement 1. Probing of the proposed dynein–Lis1 interface by mutagenesis.

(A) Sequence identity (100%, purple; 0%, white) mapped onto the Lis1 homology model. The alignment was carried out with the following species: M. musculus, H. sapiens, S. cerevisiae, A. nidulans, D. discoideum, D. melanogaster, X. laevis, C. elegans, and D. rerio using Muscle (Edgar, 2004). (B) The Lis1 homology model was docked into the dynein–Lis1 map in two possible orientations, with either the top (middle panel) or bottom (right panel) face interacting with dynein. The rotations relating the two orientations are indicated. A cross-correlation coefficient calculated for each fit is shown below the structures. These coefficients were calculated between the Lis1 EM density and the homology model filtered to the same resolution (21 Å) (as implemented in UCSF Chimera [Pettersen et al., 2004]). (C) Size-exclusion chromatography traces for wild-type and Lis1 mutants. (D) SDS-PAGE of size-exclusion chromatography fractions for GST-dynein331kDa (shortened to ‘dynein’ in the figure) mixed with wild-type of mutant Lis1.

Figure 2—figure supplement 2. Velocity distributions for dynein alone or in the presence of wild-type or mutant Lis1.

Histogram showing the velocity distribution of single TMR-labeled GST-dynein331kDa molecules in the absence of Lis1 (black) and with 200 nM wild-type Lis1 (light gray), Lis1R378A (medium gray) and Lis15A (dark gray). Velocity distributions were unimodal and could be well fit by a single Gaussian (R 2 values between 0.8221 and 0.9937).

Within Lis1, sequence conservation is much greater on one face of the β-propeller (‘top’) compared to the other (Figure 2—figure supplement 1A), suggesting that this top face may interact with dynein. Consistent with this idea, the docked β-propeller showed a better qualitative fit and a slightly higher cross-correlation coefficient with our density map when the top face is placed at the dynein interface (Figure 2—figure supplement 1B). To test this docking orientation, we mutated highly conserved residues on the top face of the propeller (Figure 2B). Our previous finding that the KDEE residues in dynein are critical for Lis1 binding (Huang et al., 2012) suggested that interactions between Lis1 and dynein have an electrostatic component. We therefore targeted four positively charged residues on the top propeller face, as well as a surface tryptophan, all of which are conserved (Figure 2B). We mutated these residues to alanine, both singly and in combination.

We first used size-exclusion chromatography to test the ability of the Lis1 mutants to interact with a functional dimerized dynein construct (GST-dynein331kDa) (Reck-Peterson et al., 2006). When all five residues are mutated to alanine (Lis15A), no binding could be detected by size-exclusion chromatography (Figure 2C and Table 2). We also did not detect an interaction with two of the single point mutants, Lis1R378A and Lis1W419A (Figure 2C and Figure 2—figure supplement 1D). The remaining single point mutants showed decreased but detectable binding to dynein (Table 2 and Figure 2—figure supplement 1D). Thus, we conclude that highly conserved amino acids on the top face of Lis1's β-propeller are critical for dynein binding, in support of our structural model for the dynein–Lis1 complex (Figure 2A).

Table 2.

We next examined if the binding-deficient Lis1 mutants Lis15A and Lis1R378A were able to regulate dynein in vitro_._ Wild-type Lis1 decreases dynein velocity in vitro in a concentration-dependent manner (Huang et al., 2012). These assays, where the motion of single, fluorescently labeled dynein molecules along microtubules is monitored over time, are more sensitive than size-exclusion chromatography for detecting dynein–Lis1 interactions. Therefore, we expected that some of the Lis1 mutants that did not co-migrate with dynein might still exhibit weak but measurable regulation of the motor. The Lis15A mutant showed no reduction in dynein velocity, consistent with an inability to bind dynein (Figure 2D and Figure 2—figure supplement 2). The Lis1R378A mutant, on the other hand, showed a slight reduction in dynein velocity compared to dynein alone, suggesting that its binding to the motor is compromised (Figure 2D and Figure 2—figure supplement 2). Thus, the effect of the Lis1 mutations on dynein binding correlates with the ability of the Lis1 mutants to regulate dynein at the single-molecule level.

Lastly, we tested our model for the dynein–Lis1 complex by measuring the effect of disrupting the dynein–Lis1 interface in vivo. In yeast, spindle pole bodies (SPB) span the nuclear envelope and coordinate microtubule minus ends that emanate from its nuclear and cytoplasmic faces (Jaspersen and Winey, 2004). Lis1 assists in concentrating dynein at the plus ends of cytoplasmic microtubules, from where dynein is offloaded to the cell cortex (Lee et al., 2003; Sheeman et al., 2003; Roberts et al., 2014). Cortically anchored dynein exerts a pulling force that results in displacements of the entire mitotic spindle (Moore et al., 2009), giving rise to a brief series of oscillations across the bud neck. Deletion of dynein eliminates these oscillations (Yeh et al., 1995). We quantified the effect of the Lis15A and Lis1R378A mutants on spindle movement in cells treated with hydroxyurea, which prolongs the period of oscillations, by tracking fluorescently labeled SPBs over the course of 20 min. We found that disruption of the dynein–Lis1 interface resulted in a decrease in the number of bud neck crossings to a level similar to that caused by the deletion of Lis1 (Figure 2E). These results indicate that the dynein–Lis1 interface identified in our structural model is crucial for dynein's biological function.

Lis1 sterically blocks the position adopted by the linker under ADP and no nucleotide conditions but does not prevent it from reaching the pre-powerstroke position at AAA2

The end of the linker domain is displaced ∼44 Å in the dynein–Lis1 structure relative to the dynein alone map, mainly along the plane of the ring (Figure 1C,D). The structure suggests that this displacement may be a direct result of Lis1's binding to dynein: the linker position in the dynein alone structure is sterically incompatible with the presence of Lis1 (Figure 1D). This is consistent with a model where Lis1 regulates dynein motility through a steric mechanism, by physically blocking the linker's normal position in the no nucleotide state.

We next sought to test if Lis1 sterically blocks the linker in other nucleotide states. As the main mechanical element of dynein, the linker is thought to adopt at least two additional conformations during the ATPase cycle. In the presence of ATP (or ATP plus Vi, which leads to the formation of the transition state analog ADP.Vi), the linker is displaced across the ring to a position near AAA2 (Kon et al., 2005; Roberts et al., 2009, 2012) (the ‘pre-powerstroke’ position). In the presence of ADP the linker lies over AAA4 in the crystal structure of the Dictyostelium discoideum dynein (Kon et al., 2012), a ‘post-powerstroke’ position slightly different from that seen in the S. cerevisiae dynein crystal structure in the absence of nucleotide, where the linker is docked onto AAA5 (Schmidt et al., 2012). However, since dynein from the same species had not been visualized in both the no nucleotide and ADP states, and because different constructs were used in the studies cited above, it was uncertain whether the AAA4 and AAA5 linker positions corresponded to distinct mechanochemical states or were due to differences between dynein species and/or constructs.

To address this, we first obtained the structure of S. cerevisiae dynein alone in the presence of ADP. Conformational sorting revealed that the linker adopts two positions in this condition (Figure 3A). One is over AAA4, coinciding with that observed in the D. discoideum crystal structure. The other is the AAA5-docked position seen in S. cerevisiae dynein with no nucleotide. The AAA4 position was seen only in the presence of ADP and was not detectable in no nucleotide conditions. These results suggest that the linker docks at AAA5 in the absence of nucleotide but can coexist in the AAA4- and AAA5-interacting states in the presence of ADP. Importantly, both the AAA4 linker position in the ADP state and the AAA5 position in the no nucleotide state are sterically incompatible with the presence of Lis1 (Figure 3B). Thus, we conclude that binding of Lis1 to the dynein ring results in a displaced linker, away from its normal docking sites under both no nucleotide and ADP conditions.

Figure 3. Lis1 sterically blocks the linker domain's normal position on dynein's ring in ADP and no nucleotide conditions but does not prevent it from reaching the pre-powerstroke position at AAA2.

(A) Cryo-NS maps of S. cerevisiae dynein in 100 μM ADP displaying the linker next to either AAA5 (left) or AAA4 (right). The S. cerevisiae linker domain (lacking nucleotide at AAA1, PDB ID: 4AKG [Schmidt et al., 2012]) and the D. discoideum linker domain (with ADP at AAA1, PDB ID: 3VKG [Kon et al., 2012]) are displayed in purple ribbon representation and have been docked into the linker-AAA5 and linker-AAA4 maps, respectively. To enable unambiguous comparison of linker positions between the EM density and crystal structure, we aligned each EM map to the corresponding dynein motor domain crystal structure after computationally removing the linker. (B) The dynein maps in no nucleotide (blue) and ADP (purple) conditions (the latter with the linker at the AAA4 location) are overlaid to compare linker positions. The location of Lis1 in the dynein–Lis1 map is shown as a transparent brown density. Both linker positions are sterically incompatible with the presence of Lis1. Note: since the ADP AAA5 linker position is the same as that seen under no nucleotide conditions, we only show the ADP map with the linker at AAA4. (C) Schematic representation of the dynein FRET construct used to test dynein's linker swing in the presence of Lis1. eGFP (green sphere–donor) was fused to the N-terminus of the linker domain, and TMR (red sphere–acceptor) was inserted into the AAA2 domain in the ring. A pre-powerstroke linker position, where the linker moves close to AAA2 in ATP plus vanadate (Vi) conditions, would display an increased FRET efficiency between the two fluorophores (bottom) relative to the no nucleotide state, where the linker is docked at AAA5 (top). (D) FRET efficiency between the eGFP and TMR fluorophores in the absence or presence of 200 μM ATP + Vi and 840 nM Lis1, ***p < 0.001. The order of addition for the reactions containing ATP + Vi and Lis1 is indicated by arrows. Averages of three experiments ± SD are shown. (E) Cryo-NS reconstruction of dynein–Lis1 in ATP + Vi conditions with the crystal structure of the motor domain docked in (PDB ID: 4AKG [Schmidt et al., 2012]). The Lis1 density is indicated. (F) At lower contour levels, the N-terminal portion of the linker can be resolved (purple arrow).

DOI: http://dx.doi.org/10.7554/eLife.03372.014

Figure 3—figure supplement 1. FRET analysis of linker movement towards the pre-powerstroke position in the presence of Lis1.

(A) Diagram of a microtubule-gliding assay. Monomeric GFP-dynein molecules are immobilized on the coverslip via anti-GFP antibodies (Y shape). Dynein-driven gliding of fluorescently labeled (purple asterisks) microtubules is visualized using TIRF microscopy. (B) A dynein FRET construct with CoA-TMR inserted into its AAA2 domain (GFP-dyneinFRET/A2) has a microtubule gliding activity similar to that of a control construct lacking it (GFP-dynein331kDa). (C) Fitted FRET emission spectra for dynein in different nucleotide conditions (no nucleotide or 200 μM ATP + Vi) and with 0 nM or 840 nM Lis1. Arrows indicate the order of addition for the last two plots. Emission spectra for eGFP and TMR used for the fit are shown as green and red traces, respectively.

We then used two approaches to determine if Lis1 also influences the position of the linker at its pre-powerstroke position, near AAA2. First, we designed a monomeric dynein construct to use fluorescence resonance energy transfer (FRET) to measure linker movement to AAA2. We based our design, which used S. cerevisiae dynein, on a linker sensor developed for D. discoideum dynein (Kon et al., 2005). In our construct, we fused an eGFP donor to the N-terminus of the linker and coupled a tetramethylrhodamine (TMR) acceptor to AAA2 via a small acetyl-CoA-binding tag (ybbR [Yin et al., 2005]) (Figure 3C). This dynein construct (GFP-dyneinFRET/A2) slides microtubules robustly, with gliding rates ∼90% of wild-type dynein (GFP-dynein331kDa) (Figure 3—figure supplement 1A,B), showing that the tags are compatible with motor function. Under no nucleotide conditions, and in the absence of Lis1, this construct showed a low FRET efficiency (∼2%), as expected when the linker is docked at AAA5 (post-powerstroke position) and the FRET donor and acceptor are far apart (Figure 3D). In the presence of ATP and vanadate (Vi), which trap dynein as an ADP.Vi-bound complex after hydrolysis, FRET increased to ∼26% (Figure 3D). Under these conditions, the linker is biased towards the pre-powerstroke position at AAA2 and the fluorophores lie closer together. When Lis1 was added to ADP.Vi–dynein, there was no significant change in the FRET efficiency relative to ADP.Vi–dynein alone (Figure 3D). When dynein was incubated with Lis1 before adding ATP and Vi, the FRET efficiency decreased somewhat but remained close to that observed for ADP.Vi–dynein alone (Figure 3D). These results suggest that Lis1 does not affect the linker pre-powerstroke AAA2 position (ATP + Vi added before Lis1) and has only a minor effect on linkers undergoing the AAA5 to AAA2 transition (Lis1 added before ATP + Vi).

As a second method to determine if Lis1 affects the linker's ability to reach the AAA2 position under ADP.Vi conditions, we determined the EM structure of the ADP.Vi–dynein–Lis1 complex (Lis1 added before ATP + Vi). We could resolve the linker in the expected pre-powerstroke position towards AAA2 and the density for the Lis1 β-propeller at the AAA3/4 junction (Figure 3E,F). In summary, these results indicate that the presence of Lis1 does not interfere with linker movement towards its pre-powerstroke position at AAA2. In contrast, Lis1 occludes the linker binding sites at its two post-powerstroke positions (AAA4 and AAA5).

ATP turnover in the presence of Lis1 requires a hydrolysis-competent AAA1 and a functional AAA5 linker-docking site

We and others have shown that Lis1 reduces dynein's velocity without significantly affecting the motor's overall ATPase rate (Yamada et al., 2008; McKenney et al., 2010; Huang et al., 2012). However, which of dynein's AAA+ modules is responsible for this continued ATP hydrolysis was not previously addressed. To determine this, we measured microtubule-stimulated ATPase rates, with and without Lis1, in different monomeric dynein constructs.

As expected, dynein monomers continued to hydrolyze ATP in the presence of Lis1 at levels similar to those of dynein alone (Figure 4A). This hydrolysis, however, was virtually abolished, both in the presence and absence of Lis1, in a construct where AAA1, the main site of ATP hydrolysis in dynein (Gibbons et al., 1987), was rendered hydrolysis-deficient with an E to Q mutation in its Walker B motif (Kon et al., 2004) (Figure 4B). This result suggests that an intact ATP hydrolysis site at AAA1 is required for ATPase activity in the presence of Lis1.

Figure 4. ATP turnover in the presence of Lis1 requires a hydrolysis-competent AAA1 and a functional AAA5 linker-docking site.

Microtubule-stimulated ATPase activity of dynein monomers carrying (A) wild-type AAA+ modules, (B) a hydrolysis deficient E1849Q mutation in AAA1 (Kon et al., 2004), (C) a hydrolysis deficient E2819Q mutation in AAA4 (Cho et al., 2008), (D) AAA5 mutations (K3438E, R3445E, F3446D) that prevent linker docking (Schmidt et al., 2012). ATPase traces are of dynein alone (light green) or in the presence of 140 nM Lis1 (brown). Measurements were done in triplicate (A and C) or duplicate (B and D) from one preparation. Diagrams of the dynein constructs used to generate the plots are shown next to them. See Table 3 for fit equation and rate quantifications.

DOI: http://dx.doi.org/10.7554/eLife.03372.016

Figure 4—figure supplement 1. Lis1 binds to dynein ATPase mutants.

(A–D) SDS-PAGE of fractions eluted from size-exclusion chromatography runs of Lis1 mixed with each of the dynein constructs used in the ATPase assays. Lis1 co-elutes with all of the constructs.

Given that Lis1 binds at AAA4, one of the hydrolysis-competent AAA+ modules in dynein, it was possible that Lis1 might be stimulating ATP hydrolysis at that site, with AAA1 playing only an indirect role. However, dynein carrying an E to Q mutation in the Walker B motif of AAA4 (Cho et al., 2008) showed a near wild-type ATPase rate with or without Lis1 (Figure 4C). Therefore, a hydrolysis-competent AAA4 is not required for the ATPase activity observed in the presence of Lis1.

Mutations in AAA5 (an AAA+ module that cannot bind ATP) that prevent linker docking have been shown to severely reduce dynein's ATPase activity (Schmidt et al., 2012). We wondered whether Lis1 binding might rescue this mutation and restore ATPase activity to dynein. This was not the case; dynein constructs carrying the AAA5 mutation did not hydrolyze ATP even in the presence of Lis1 (Figure 4D).

Taken together, these results indicate that sustained ATP hydrolysis in a Lis1-regulated dynein requires a hydrolysis-competent AAA1 and a functional linker-docking site at AAA5.

Removing Lis1's steric block by shortening dynein's linker makes the motor Lis1-insensitive

The experiments discussed above showed that Lis1 does not regulate dynein by affecting the linker's ability to reach its pre-powerstroke position at AAA2. Our dynein–Lis1 structure shows that Lis1, however, does affect post-powerstroke linker positions as Lis1 and the linker are sterically incompatible in no nucleotide and ADP conditions (Figure 1D,E, Figure 3B). We wondered whether motility regulation was a result of this steric blocking by Lis1. Specifically, we wanted to test the hypothesis that steric blocking of the linker is necessary for inducing dynein's Lis1-dependent state of persistent microtubule attachment. To test this hypothesis, we used a dynein construct with a truncated linker that is long enough to form a functional motor but is too short to be sterically blocked by Lis1. This construct is generated by deleting 145 amino acids at the N-terminus of the dynein motor (Figure 5A).

Figure 5. A shortened linker that can physically bypass Lis1 renders dynein Lis1 insensitive.

(A) A short linker construct was designed by docking the crystal structure of the D. discoideum linker (purple ribbon) (PDB ID: 3VKG [Kon et al., 2012]) into our EM map of dynein alone and overlaying the position of Lis1 (brown mesh). Truncating the linker at residue 1365 (dashed line) yields a linker that is functional (see Figure 5—figure supplement 1) but that can no longer contact Lis1. (B) Cryo-NS reconstruction of the short linker dynein–Lis1 complex; the linker assumes the same conformation with Lis1 bound as in the absence of Lis1. (C) Diagram of the single-molecule microtubule release assay we used to test Lis1 regulation of dynein. Release from microtubules of TMR-labeled (red asterisk) dynein monomers on addition of ATP is monitored by TIRF microscopy. (D) Diagrams of predicted outcomes. Dynein's linker domain in purple, microtubule in gray, Lis1 in brown. (i) Dynein monomers release from microtubules in ATP conditions in the absence of Lis1. (ii) Our model proposes that Lis1 sterically blocks a full-length linker from assuming the normal conformation on dynein's ring, keeping dynein bound to the microtubule. (iii) In the absence of Lis1, shortening the linker would have no effect on dynein's mechanochemical cycle. (iv) Our model predicts that a shortened linker that can bypass the Lis1 steric block should render dynein insensitive to Lis1. (E) Kymographs of TMR-labeled full-length (left) or short linker (right) dynein molecules. After pre-binding to microtubules, release of dynein molecules is monitored after addition of 5 mM ATP, with and without 300 nM Lis1. Kymographs correspond to the dynein constructs shown in (D). Scale bar = 5 s. (F) Quantification of the kymographs in (D), showing the duration of microtubule attachment after addition of ATP, in the absence (gray) or presence (brown) of Lis1. Data were binned into 1 s intervals and the histograms show alternating no Lis1 and +Lis1 bars. Rare attachments longer than 10 s were excluded from the analysis and plot, N = 179–183.

DOI: http://dx.doi.org/10.7554/eLife.03372.019

Figure 5—figure supplement 1. The short linker dynein construct shows robust motility, hydrolyzes ATP, and binds Lis1.

(A) Single-molecule motility assays. Kymographs of GST-dimerized full-length and short linker dyneins. Horizontal scale bar = 2 μm, vertical = 30 s. (B) Velocity and run length for short linker and full-length linker dyneins. Similar values are seen for the two constructs, N = 265–333. (C) SDS-PAGE of elution fractions from size-exclusion chromatography of monomeric short linker dynein mixed with Lis1. Lis1 co-migrates with short linker dynein. (D) Microtubule-stimulated ATPase activity of short linker dynein with either wild-type AAA+ modules (left) or with mutations (K3438E, R3445E, F3446D) that prevent linker docking at AAA5 (Schmidt et al., 2012) (right). ATPase traces are of dynein alone (light green) or in the presence of 140 nM Lis1 (brown). Measurements were done in triplicate (wild type) or duplicate (AAA5 mutant) from one preparation. Diagrams of the dynein constructs used to generate the plots are shown next to them. (E) Addition of buffer lacking ATP to flow chambers containing TMR-labeled full-length or short linker dynein does not cause microtubule detachment. Scale bar = 5 s. (F) The short linker construct used in Figure 5 contains an N-terminal GFP connected to the short linker via 3 HA tags. We expected the GFP to be flexible in its location relative to dynein based on previous EM studies (Roberts et al., 2009) and the fact that it is averaged out in our 3D reconstruction of dynein (Figure 1D). However, to rule out interference from GFP, we also carried out the microtubule release assays with a GFP-less construct. We observed the same results: addition of ATP to GFP-less dyneins in the presence of 300 nM Lis1 caused short-linker dyneins to release from the microtubule while full-length dyneins remain attached. Scale bar = 5 s. (G) Quantification of the kymographs in (F), showing the duration of microtubule attachment of full-length (light brown) or short linker (dark brown) dynein molecules after addition of ATP, in the presence of 300 nM Lis1. Data were binned into 1 s intervals and the histograms show alternating full-length and short linker dynein bars. Rare attachments longer than 10 s were excluded from the analysis and plot, N = 141–197.

We first verified this construct functionally and structurally. A dimeric dynein motor containing this shortened linker shows robust motility properties in in vitro motility assays (Figure 5—figure supplement 1A,B) (Reck-Peterson et al., 2006). We also tested whether shortening the linker affects the microtubule-stimulated ATPase activity of dynein monomers. Monomers with a short linker showed ATPase levels comparable to those seen with a full-length linker, both in the context of a wild-type set of AAA+ modules and in the linker docking-deficient AAA5 mutant (Figure 5—figure supplement 1D). As a monomer, the short linker construct can bind Lis1 as shown both by their co-migration in size-exclusion chromatography (Figure 5—figure supplement 1C) and by our ability to obtain a 3D reconstruction of the short linker dynein–Lis1 complex (Figure 5B). Central to our testing the steric block hypothesis, our 3D structure of the short linker dynein–Lis1 complex shows the same conformation for the linker in the presence of Lis1 as we had observed for the full-length linker in the absence of Lis1 (Figure 1, Figure 5A,B). Therefore, the short linker is functional and able to physically bypass Lis1.

To directly test whether Lis1 was capable of regulating dynein with a short linker, we used a single-molecule microtubule release assay (Figure 5C). In this study, the duration of single monomeric dynein's attachments to microtubules can be measured by kymograph analysis in a flow chamber by TIRF microscopy (Huang et al., 2012). Addition of ATP triggers a low-affinity state in dynein (Kon et al., 2005; Imamula et al., 2007; Huang et al., 2012) and the dynein monomers release from microtubules, resulting in a loss of fluorescence signal. Microtubule rebinding events are short lived, likely corresponding to single turnovers of ATP. In the presence of a full-length linker, Lis1 converted dynein to a state of persistent microtubule attachment and dynein monomers stayed bound in the presence of ATP for extended periods as previously shown (Figure 5D–F) (Huang et al., 2012). Strikingly, shortening of dynein's linker eliminated Lis1's ability to induce this persistent microtubule-bound state. We quantified the durations of microtubule attachments after the addition of ATP and found the same short-lived attachments seen with dynein in the absence of Lis1 (Figure 5D–F). Thus, Lis1 is not capable of regulating microtubule attachment in the short linker construct. These data support a steric mode of dynein regulation where Lis1 physically blocks the linker.

Discussion

We previously described Lis1 as a ‘clutch’ for dynein, based on its ability to uncouple the cycles of ATP hydrolysis, which take place in the motor domain, from the cycles of microtubule binding and release at the microtubule binding domain (Huang et al., 2012). One of the functional consequences of the dynein–Lis1 interaction is that Lis1 keeps dynein in a persistent microtubule-bound state. In this study, we have determined six 3D EM structures of dynein and dynein–Lis1 in different nucleotide states. By combining these structures with single molecule motility experiments, we have established that Lis1 regulates dynein's microtubule attachment by sterically blocking its linker domain.

Together, our data suggest the following model of dynein regulation by Lis1 (Figure 6). In the current view of dynein's mechanochemical cycle, the motor domain encounters the microtubule with ADP.Pi bound at AAA1, with the linker in a pre-powerstroke position at AAA2 (Kon et al., 2005; Roberts et al., 2009, 2012). Strong microtubule binding stimulates Pi release, inducing the linker to swing to AAA4 (Kon et al., 2012). Finally, linker docking at AAA5 is thought to promote the release of ADP from AAA1, resetting the mechanochemical cycle (Schmidt et al., 2012). Our data suggest that when Lis1 is present, the linker retains its ability to adopt the pre-powerstroke AAA2 position but is prevented from reaching its normal post-powerstroke positions at AAA4 and AAA5 on dynein's ring (Figure 6). This blocking of the linker by Lis1 is critical for motility regulation; its removal by shortening dynein's linker renders the motor Lis1 insensitive.

Figure 6. Model for the regulation of dynein by Lis1.

(A–G) Current view of dynein's mechanochemical cycle. (A) ATP binding to AAA1 induces the low-affinity conformation in dynein's microtubule-binding domain and (B) release from the microtubule. (C) The linker domain changes its position from AAA5 towards AAA2, the ‘pre-powerstroke’ and ATP is hydrolyzed. (D) Binding of dynein to a new site on the microtubule triggers a change in the microtubule-binding domain to its high affinity conformation (E). (F) Release of Pi results in the ‘powerstroke’, a movement of the linker back towards AAA5. (G) Docking of the linker at AAA5 is thought to promote nucleotide exchange at AAA1, resetting the motor for a new cycle. (H–J) Model for the Lis1-regulated cycle. Lis1 prevents the linker from completing its normal conformational cycle, keeping dynein in a persistent microtubule-attached state, despite continuing ATP hydrolysis. (H) Binding of Lis1 to dynein blocks the linker from docking onto the ring at AAA5, preventing the conformational changes in the stalk and microtubule binding domain that ultimately result in dynein's release from the microtubule. (I) The linker is still capable of moving to the pre-powerstroke position at AAA2 in the presence of Lis1, and ATP is hydrolyzed. (J) Presumably, by analogy to the dynein alone cycle, Pi release triggers the power-stroke, but Lis1 sterically blocks the linker's normal position on dynein's ring in the ADP state. Our current understanding of Lis1 regulation does not yet explain the mechanism of nucleotide exchange at AAA1.

DOI: http://dx.doi.org/10.7554/eLife.03372.021

Why does Lis1's blocking the linker from adopting its normal post-powerstroke positions prevent dynein's microtubule detachment? One possibility suggested by our structures is that Lis1 disrupts the interaction between the linker and AAA5, preventing normal progression through the mechanochemical cycle. Consistent with this notion, when linker docking at AAA5 is abolished by mutagenesis, dynein displays reduced velocity and prolonged microtubule attachments (Schmidt et al., 2012), reminiscent of Lis1's effects. However, while Lis1 has little effect on dynein's ATPase, AAA5 linker docking mutants display severely reduced ATPase rates, both in the absence (Schmidt et al., 2012) and the presence of Lis1 (Figure 4D, Figure 5—figure supplement 1D). Given these results, it is not clear at this point what the mechanistic basis is for dynein's continuing ATPase in the presence of Lis1. On the one hand, it is possible that the AAA5 mutations may, in addition to preventing linker docking, disrupt dynein's mechanochemical cycle and thus also prevent ATP hydrolysis. A method to reversibly block linker docking at AAA5 (e.g., via a small molecule) would be required to determine if AAA5 docking is truly required for dynein ATPase activity. On the other hand, Lis1 may uncouple ATP hydrolysis from linker docking at AAA5 through an allosteric effect on the ring. In this scenario, the linker–AAA5 interaction, which is blocked by Lis1, would be required for the conformational changes that ultimately shift dynein's microtubule-binding domain to its low-affinity state, but not for dynein's continuing ATPase activity. Higher resolution structures of the dynein–Lis1 complex will be required to establish whether Lis1 has an effect on the structure of dynein's ring.

It is conceivable that blocking of the normal linker-docking sites by Lis1 might induce a new interaction between the linker and the AAA+ ring. Similarly, Lis1 may interact specifically with the linker itself. Either (or both) of these scenarios could in turn be responsible for preventing microtubule release. However, current evidence does not favor these possibilities. Low sequence conservation in the portion of the linker facing Lis1 argues against a specific Lis1–linker interaction (Figure 1—figure supplement 1I). Likewise, a specific interaction between a Lis1-displaced linker and dynein's ring is not supported by the apparent conformational heterogeneity of the N-terminus of the linker in the presence of Lis1, where 3D sorting is required to resolve linker positions (Figure 1—figure supplement 1F–H). Also mutating five amino acids on AAA4, proximal to the linker's displaced position (the most likely candidates to interact with the displaced linker), had minimal effect on Lis1-mediated motility regulation (Figure 1—figure supplement 2). A direct test of whether specific interactions exist among these different elements will also require a higher resolution structure, where the rotational orientation of the Lis1 homology model within its density in the EM map is unequivocal and specific interactions between the linker and Lis1 as well as the linker and the dynein ring can be distinguished from physical proximity.

In conclusion, our data show that Lis1, a conserved dynein regulator, directly disrupts dynein's mechanochemical cycle by physically blocking conformations that are required to couple the cycles of ATP hydrolysis taking place in the motor domain from those of track binding and release happening at the microtubule binding domain. This allows Lis1 to keep dynein in a persistent microtubule-bound state. This modulation of dynein's interaction with its microtubule track likely contributes to dynein's ability to carry out the variety of cellular functions it performs in different organisms, given the conservation of the amino acids at the dynein–Lis1 interface. For example, Lis1 is involved in initiation of cargo transport (Lenz et al., 2006; Egan et al., 2012; Moughamian et al., 2013), in transport of high load cargo (McKenney et al., 2010), and in targeting dynein molecules to the cell cortex via the microtubule plus end (Lee et al., 2003; Sheeman et al., 2003; Roberts et al., 2014). The displaced linker observed in the presence of Lis1 in our 3D dynein–Lis1 reconstruction may contribute to this latter task, generating an ‘unmasked’ tail domain that has been shown necessary for cortical dynein localization (Markus and Lee, 2011). In the case of the mammalian proteins, dynein and Lis1 were previously shown to form a stable complex only in ATP and Vi conditions (McKenney et al., 2010). Our 3D reconstruction of dynein–Lis1 under those conditions suggests that this might be a consequence of the linker's moving to its pre-powerstroke site at AAA2, where the linker and Lis1 are no longer sterically incompatible.

The work presented here has helped dissect the molecular mechanism by which Lis1 regulates a single dynein motor domain. The next challenge will be to understand the interactions between Lis1 and dynein dimers and of those with other regulatory factors. Future structural studies with full-length dimeric dynein–Lis1–Nudel complexes, free and bound to microtubules, will be required to answer these exciting questions.

Materials and methods

Yeast strain construction

The S. cerevisiae strains used in this study are listed in Table 1. Deletions or modifications of endogenous genomic copies of the dynein heavy chain (DYN1) and Lis1 (PAC1) were done using PCR-based methods as previously described (Longtine et al., 1998), using the URA3/5FOA ‘pop-in/pop-out’ method (Guthrie and Fink, 1991). Transformations were performed using the standard lithium acetate method (Gietz and Woods, 2002). Point mutants were generated using the PCR stitching method and verified by DNA sequencing.

Protein expression and purification

Cultures of S. cerevisiae for protein purification were grown, harvested, and frozen as described previously (Reck-Peterson et al., 2006). Dynein and Lis1 constructs were purified and labeled as described previously (Reck-Peterson et al., 2006; Huang et al., 2012), except that a modified TEV buffer for Lis1 purification was used; 50 mM Tris–HCl (pH 8.0), 150 mM potassium acetate, 2 mM magnesium acetate, 1 mM EGTA, 5% glycerol, 1 mM DTT, and 1 mM PMSF.

EM sample preparation

We chose to use cryo-NS EM, where a carbon support, combined with a heavy metal stain, resulted in highly reproducible grids with high contrast. Prior attempts at getting dynein reproducibly in open holes for standard cryo-EM were unsuccessful. Furthermore, cryo-EM on continuous carbon gave micrographs where individual dynein particles were difficult to see above the noise. The high reproducibility we were able to achieve with cryo-NS allowed us to sample a much greater range of constructs/nucleotide conditions in the same time frame than we would otherwise have been able to do in unstained, unsupported conditions. Most importantly, the improved contrast was instrumental in allowing us to sort the different dynein conformations that were present in most of our data sets. 4 μl of monomeric dynein (80–120 nM), or monomeric dynein pre-incubated for 10 min with Lis1 dimer at a 1.5-fold excess (120–180 nM), was applied to a glow discharged, continuous carbon coated, C-flat EM grids (Protochips, Raleigh, NC). Dynein samples stated to be prepared in no nucleotide conditions were treated with apyrase (0.14 U/ml) for 15 min prior to grid application to hydrolyze residual ADP left over from the dynein purification procedure. Dynein samples stated to be prepared in ADP and ATP + Vi conditions contained 100 μM ADP and 500 μM Mg-ATP/NaVO4, respectively. For the latter, nucleotide was added after the dynein–Lis1 pre-incubation step. Once applied to the grid, the samples were stained with 2% uranyl formate by floating the grid sample face down on a pool of stain. Samples were then sandwiched with a thin layer of freshly evaporated carbon, and grids were lightly blotted from the non-sample containing side and plunged into liquid nitrogen. Grids were then stored at liquid nitrogen temperatures.

EM data collection

Samples were imaged at liquid nitrogen temperatures using a Gatan 626 cryo holder (Gatan, Inc., Pleasanton, CA) on a Tecnai F20 TEM microscope (FEI, Hillsboro, OR), operating at 120 kV, equipped with a US4000 4k × 4k CCD camera (Gatan). Data were collected either manually or automatically using Leginon (Carragher et al., 2000). Dynein alone samples (no nucleotide [strain RPY844] and ADP conditions [strain RPY844]) and dynein–Lis1 (ATP + Vi conditions [strains RPY1302 and RPY816]) were imaged at 62,000× nominal magnification (1.73 Å/pixel). Dynein–Lis1 (no nucleotide [strains RPY1302 and RPY816]) was imaged at 50,000× nominal magnification (2.14 Å/pixel). Short linker dynein–Lis1 (no nucleotide [strains RPY1436 and RPY816]) was imaged at 80,000× nominal magnification (1.34 Å/pixel). Low-dose conditions during imaging (dose ∼25 e−/Å2) were used for all data sets, and micrographs were collected using a defocus range of −0.6 to −1.5 μm.

EM image pre-processing

For all data sets, ∼1,000 particles were initially selected manually in Boxer (EMAN1) (Ludtke et al., 1999) and reference-free 2D classified in IMAGIC (van Heel et al., 1996) to give class averages that were then used as templates for automated particle picking in Appion (Lander et al., 2009). Reference-free 2D classification in IMAGIC was subsequently used on the data sets to remove averages with blurred appearance or incorrect size. CTF determination and correction of image phases were carried out in Appion using Ace2 (NRAMM). Particles were band-pass filtered (high-pass = 250 Å, low-pass = 3 × sampling) in Imagic and normalized in Xmipp (Sorzano et al., 2004). For 3D classification and initial 3D refinement particles were binned by two; final 3D refinements were carried out using unbinned data.

EM image processing

Dynein (no nucleotide)

An initial model was generated using the S. cerevisiae dynein motor domain crystal structure (PDB ID: 4AKG [Schmidt et al., 2012]), low-pass Fourier filtered to 80 Å. Initial 3D refinement was carried out in EMAN2 (Tang et al., 2007). The resulting map was filtered to 40 Å and used as an initial model for 3D classification in RELION (Scheres, 2012). Five classes were generated. Particles from four of the classes were combined and refined in RELION against the class 5 map, filtered to 40 Å. 3D refinement converged after 18 iterations. The final map contained 31,839 particles (from 38,463 total) and the ‘gold-standard’ resolution using an FSC cut-off of 0.143 was 14.8 Å. The final map was filtered according to local resolutions (Cardone et al., 2013).

Dynein (ADP)

The linker domain has been shown to have a different position relative to the dynein ring in ADP conditions in D. discoideum dynein compared with that of S. cerevisiae dynein in no nucleotide conditions (Kon et al., 2012; Schmidt et al., 2012). To avoid initial model bias of linker position, the domain was computationally removed from residue 1,620 of PDB file 4AKG (Schmidt et al., 2012). The resulting map was filtered to 50 Å and used as a starting model for an initial refinement in SPIDER (Frank et al., 1996) to regain linker density at a position derived solely from the data. This map was then filtered to 40 Å and used as an initial model for 3D classification in RELION. Five classes were generated. Four of the classes showed the linker at the no nucleotide position (position 1) and 1 class at the shifted position seen with D. discoideum dynein in ADP conditions (position 2). The particles in classes corresponding to each conformation were further refined in RELION and the refinements converged after 13 and 10 iterations for linker position 1 and 2 data sets, respectively. The final maps contained 7,630 and 3,983 particles (from 17,256 total) and the ‘gold-standard’ resolutions using an FSC cut-off of 0.143 were 18.3 and 19.5 Å for linker position 1 and 2 maps, respectively. The final maps were filtered according to local resolutions (Cardone et al., 2013).

Dynein–Lis1 (no nucleotide)

The same initial model as described for dynein (no nucleotide) above was used to 3D refine an initial data set of dynein–Lis1 (RPY844, RPY816) in EMAN2. This map was then filtered to 60 Å, and EMAN2 was used to refine a larger data set of dynein–Lis1 using a dynein lacking any tags on the end of the linker (RPY1302). The resulting map was filtered to 40 Å and used for 3D classification in RELION. This process was repeated with different requested class numbers. Linker position was observed to vary across classes (Figure 1—figure supplement 1F–H). In a run with seven generated classes, classes with most density for the linker (class 1 and 6) were combined and refined against the class 1 map filtered to 40 Å in RELION. Three-dimensional refinement converged after 16 iterations. The final map contained 10,129 particles (from 35,472 total) and the ‘gold-standard’ resolution using an FSC cut-off of 0.143 was 21.4 Å. The final map was filtered according to local resolutions (Cardone et al., 2013).

Short linker dynein–Lis1 (no nucleotide)

The same initial model as described for dynein (no nucleotide) was filtered to 50 Å and used for initial 3D classification of the data set in RELION. Five classes were generated. Particles from 1 class were further refined against the class map in RELION. The refinement converged after 14 iteractions and contained 11,818 particles (from 34,805 total). The gold-standard resolution using an FSC cut-off of 0.143 was 15.4 Å. The final map was filtered according to local resolutions (Cardone et al., 2013).

Dynein–Lis1 (ATP + Vi)

The map of dynein–Lis1 (no nucleotide) filtered to 60 Å was used as a starting model for initial refinement in EMAN2. The resulting map was filtered to 40 Å and used for 3D classification in RELION. Most classes showed the linker position unresolved, indicative of variability in location as previously observed (Roberts et al., 2009, 2012), but one class resolved the linker near AAA2 when viewed at lower contour levels. The particles in this class were refined against the class map in RELION and refinement converged after 16 iterations. The final map contained 1,072 particles (from 6,600 total), and the ‘gold-standard’ resolution using an FSC cut-off of 0.143 was 23.1 Å.

Accession numbers

EM maps have been deposited with the EMDataBank. Accession codes as follows; dynein–Lis1 (no nucleotide conditions) EMDB-6008; dynein alone (no nucleotide conditions) EMDB-6013; dynein alone (ADP conditions) with position 1 and 2 linker domains, EMDB-6015 and EMDB-6014 respectively; dynein–Lis1 (ATP + Vi conditions) EMDB-6016; short linker dynein–Lis1 (no nucleotide conditions) EMDB-6017. For each entry, in addition to the final masked and filtered maps, raw half maps for each reconstruction have been deposited. We also deposited an XML file of the FSC plot between the dynein alone map and the fitted crystal structure of the motor domain (PDB ID: 4AKG [Schmidt et al., 2012]), as a supplementary file to the dynein alone submission (EMDB-6013).

Size-exclusion chromatography

Dynein and Lis1 were tested for complex formation, and Lis1 mutants were tested for structural integrity (Figure 2—figure supplement 1C) by size-exclusion chromatography. 400–800 nM dynein and 475–800 nM Lis1 were loaded separately or after being mixed for 10 min at 4°C. Samples were fractionated on a Superose 6 PC 3.2/30 column using an ÄKTAmicro system (GE Healthcare) that had been equilibrated with degassed gel filtration buffer (50 mM Tris–HCl pH 8.0, 150 mM potassium acetate, 2 mM magnesium acetate, 1 mM EGTA, 1 mM DTT). Fractions (50 μl or 90 μl) were analyzed by SDS-PAGE on 4–12% Tris-Bis gels (Invitrogen, Grand Island, NY) with SYPRO Red staining (Invitrogen) and imaged using an ImageQuant 300 (BioRad, Hercules, CA) or Typhoon (Amersham, UK) gel imaging system.

Single-molecule microscopy

Single-molecule motility assays were performed using flow chambers as previously described (Case et al., 1997). Dynein was labeled with TMR (Promega, Madison, WI), and microtubules contained ∼10% biotin-tubulin for surface attachment and ∼10% HyLite488-tubulin (Cytoskeleton Inc., Denver, CO) for visualization. For assays that included Lis1, dynein was incubated with 200 nM Lis1 for 10 min at 4°C prior to addition to the flow chamber. The imaging buffer consisted of 30 mM HEPES (pH 7.2), 50 mM potassium acetate, 2 mM magnesium acetate, 1 mM EGTA, 10% glycerol, 1 mM DTT, 20 mM taxol, 1.25 mg/ml casein, 1 mM Mg-ATP, and an oxygen scavenger system. Images were recorded every 2 s for 5 or 10 min, and dynein velocities and run-lengths were calculated from kymographs generated in ImageJ (National Institutes of Health).

In vitro motility assays were visualized on either a Zeiss Elyra PS.1 microscope with a 100× 1.46 N.A. oil immersion TIRF objective (Carl Zeiss GmbH, Germany) with an Andor EM-CCD camera or an Olympus IX-81 TIRF microscope with a 100× 1.45 N.A. oil immersion TIRF objective (Olympus, Japan) with a Hamamatsu EM-CCD camera. TMR-labeled dynein and HyLite488-microtubules were excited with 561 nm and 488 nm solid state laser lines, respectively. Images were recorded with a 100 ms exposure using Zen Black (Zeiss) or Metamorph software. Microtubule gliding assays and microtubule binding and release assays were performed as described (Huang et al., 2012). Control experiments for the microtubule release assays examined dynein release in buffer lacking ATP (Figure 5—figure supplement 1E), where dynein remained bound to microtubules as expected (Huang et al., 2012) and with dynein lacking N-terminal tags (Figure 5—figure supplement 1F,G), where untagged dynein behaved similar to tagged dynein (Figure 5E,F).

Spindle oscillation assay

To track the dynein-dependent movement of spindle pole bodies (SPBs), we used a strain containing a GFP-labeled SPB marker, SPC110, and a tdTomato-labeled cell membrane marker, HXT1 (kindly provided by Jeff Moore, University of Colorado). Mutations were introduced into the PAC1 (Lis1) locus in this strain. For control experiments, strains containing deletions of the dynein heavy chain (DYN1) and PAC1 loci were constructed. All strains were PCR verified, and mutations were additionally verified by DNA sequencing.

For image analysis, saturated overnight cultures for each strain were diluted to an OD600 of 0.1 in a total volume of 5 ml YPD media. The dilution of cultures was staggered such that the data could be collected for all strains during a single imaging session. Following dilution, each culture was incubated with rotation at 30°C for 3 hr. Hydroxyurea (HU) was then added to a final concentration of 200 mM, and the culture was incubated for an additional 2 hr with rotation at 30°C. The cells were collected by centrifugation, the media was discarded, and the cells were resuspended in 250 μl of fresh YPD + 200 mM HU. The cells were loaded into an Y04C microfluidic yeast plate (CellASIC, EMD-Millipore, Germany) and introduced into the viewing chamber with the ONIX controller (CellASIC). Imaging was performed at the Nikon Imaging Center at Harvard Medical School. All images were collected with a Yokagawa CSU-X1 spinning disk confocal with Borealis modification, on a Nikon Ti inverted microscope equipped with a Plan 60 × 1.4 N.A. objective and the Perfect Focus System (Nikon Corp., Japan). GFP-labeled SPB and tdTomato-labeled cell membrane fluorescence were excited with the 488 nm and 561 nm lines, respectively, from a LMM-5 solid state laser merge module controlled with an ATOF (Spectral Applied Research Inc., Canada). Images were acquired with a Hamamatsu ORCA-AG CCD controlled with MetaMorph 7.0 software. Images were collected for SPBs as 100 ms exposures, spanning 9 × 500 nm Z-sections (4.5 μm total Z stack) every 30 s for a total of 20 min. Cell membranes were imaged as single central Z-sections at the first and last time point. Membrane image pairs were digitally merged to allow for drift analysis; those cells with visible drift were excluded from analysis.

Image processing and particle tracking

Maximum intensity projections were calculated for Z-series at each time point for GFP-labeled SPBs. At each time point, SPBs were independently detected in the Z-projection using a wavelet detection algorithm (Aguet et al., 2013), and the two spindles were tracked throughout the course of the movie using a nearest neighbor tracking method (unpublished Matlab [Mathworks, Natick, MA] scripts). The location of the bud neck and the mother–daughter orientation were determined using the first tdTomato-labeled cell membrane exposure. The locations of tracked SPBs were used to calculate the number of bud neck crossings.

FRET

To generate the dynein FRET construct, eGFP (the FRET donor) was inserted at the dynein N-terminus and the acceptor site was inserted after L2241 in AAA2. We used the ybbR tag (GGGTVLDSLEFIASKLAGGG [Yin et al., 2005]) labeled with TMR-CoA (NEB, Ipswich, MA) as the FRET acceptor. Dynein was incubated with or without Lis1 for one hour on ice, followed by apyrase (6.6 U/ml) or 200 μM ATP.Vi for 2 min at room temperature (RT). For some experiments, dynein was first incubated with 200 μM ATP.Vi for 2 min at RT, followed by Lis1 for 1 hr on ice. Assays were performed in 30 mM HEPES (pH 7.2), 50 mM potassium acetate, 2 mM magnesium acetate, 1 mM EGTA, 1 mM DTT and the final concentrations of dynein and Lis1 were 84 nM and 840 nM, respectively. The sample was excited with 485 nm (eGFP) light, and the emitted light was detected from 505 nm to 650 nm in a SpectraMax M5 fluorimeter (Molecular Devices, Sunnyvale, CA) at RT. In order to normalize across experiments, the samples were also excited with 535 nm (TMR) light and the emitted light was detected from 555 nm to 700 nm. To analyze the FRET data, we first subtracted the fluorescence background from the buffer alone. We then used the emission spectra of dynein-labeled with eGFP and free TMR dye alone to decompose each channel in the experimental spectra. FRET efficiencies (E) were calculated using the method of Clegg (Clegg, 1995): E = {_F_aFRET/_F_aDIR − εa(485)/εa(535)}εa(535)/εd(485), where the superscripts ‘d’ and ‘a’ refer to the donor (eGFP) and the acceptor (TMR), respectively. _F_aFRET is the fluorescence intensity of the acceptor excited at 485 nm and _F_aDIR is the fluorescence intensity of the acceptor excited at 535 nm. εd(485) εa(485) and εa(535) are the molar extinction coefficients at the designated wavelengths. In our experiments εa(535)/εd(485) = 37,900 M−1 cm−1/40,000 M−1 cm−1 and εa(485)/εa(535) = 0.2.

ATPase assays

Dynein constructs used in ATPase assays were tested for complex formation with Lis1 by size-exclusion chromatography (Figure 4—figure supplement 1). ATPase assays were performed using an EnzChek phosphatase kit (Molecular Probes, Thermo Fisher Scientific Inc., Cambridge, MA) as previously described (Reck-Peterson et al., 2006; Cho et al., 2008). The final reaction consisted of 10–20 nM dynein (monomeric constructs, see Figure 4 and Figure 5—figure supplement 1D), 0 or 140 nM Lis1, 0–7.5 μM taxol-stabilized microtubules, 2 mM Mg-ATP, 200 mM MESG (2-amino-6-mercapto-7-methyl-purine riboside), 1 U/ml purine nucleoside phosphorylase, and assay buffer (30 mM HEPES (pH 7.2), 50 mM potassium acetate, 2 mM magnesium acetate, 1 mM EGTA, 1 mM DTT, and 10 mM taxol). A SpectraMax384 plate reader (Molecular Devices) was used to monitor the coupled reaction at OD360 every 12 s for 10 min. Data were fit according to Nishiura et al. (2004).

Acknowledgements

We would like to thank Jeff Moore (University of Colorado School of Medicine) for yeast strains, Vu Nguyen, Rogelio Hernandez-Lopez, Michael Cianfrocco, John Srouji, and Julie Huang for their contributions during this work, Morgan deSantis for comments on the manuscript, and Liza Sholl for administrative support. EM data were collected at Harvard's Center for Nanoscale Systems, a member of the National Nanotechnology Infrastructure Network and supported by the National Science Foundation (NSF) (ECS-0335765). Computation was performed in part on the Odyssey cluster supported by Harvard's FAS Science Division Research Computing Group. Single molecule data were collected in part at Harvard's Center for Biological Imaging, supported by an SIG award (RR1S1027990) from the NIH. Spinning disc confocal microscopy data were collected at the Nikon Imaging Center at Harvard Medical School. KT was supported by a Charles King postdoctoral fellowship. AJR is a Sir Henry Fellow supported by the Wellcome Trust (092436/Z/10/Z), sponsored by Peter J Knight, and Stan A Burgess (University of Leeds). SLR-P was supported by the Rita Allen Foundation. SLR-P and AEL were supported by the NIH (R01 GM 107214).

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Funding Information

This paper was supported by the following grants:

• Charles A. King Trust Postdoctoral Research Fellowship to Katerina Toropova.
• Wellcome Trust 092436/Z/10/Z to Anthony J Roberts.
• Rita Allen Foundation to Samara L Reck-Peterson.
• National Institutes of Health R01 GM 107214 to Samara L Reck-Peterson, Andres E Leschziner.

Additional information

Competing interests

The authors declare that no competing interests exist.

Author contributions

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

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

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

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

BSG, Analysis and interpretation of data, Drafting or revising the article.

SLR-P, Conception and design, Analysis and interpretation of data, Drafting or revising the article.

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

Additional files

Major dataset

The following datasets were generated:

Toropova K, Zou S, Roberts AJ, Redwine WB, Goodman BS, Reck-Peterson SL, Leschziner AE, 2014, Dynein-Lis1 (no nucleotide conditions) EM structure, EMD-6008; http://www.ebi.ac.uk/pdbe/entry/EMD-6008, Available at the EMDataBank repository (http://www.emdatabank.org/).

Toropova K, Zou S, Roberts AJ, Redwine WB, Goodman BS, Reck-Peterson SL, Leschziner AE, 2014, Dynein (no nucleotide conditions) EM structure, EMD-6013; http://www.ebi.ac.uk/pdbe/entry/EMD-6013, Available at the EMDataBank repository (http://www.emdatabank.org/).

Toropova K, Zou S, Roberts AJ, Redwine WB, Goodman BS, Reck-Peterson SL, Leschziner AE, 2014, Dynein (ADP conditions), linker position 1 EM structure, EMD-6015; http://www.ebi.ac.uk/pdbe/entry/EMD-6015, Available at the EMDataBank repository (http://www.emdatabank.org/).

Toropova K, Zou S, Roberts AJ, Redwine WB, Goodman BS, Reck-Peterson SL, Leschziner AE, 2014, Dynein (ADP conditions), linker position 2 EM structure, EMD-6014; http://www.ebi.ac.uk/pdbe/entry/EMD-6014, Available at the EMDataBank repository (http://www.emdatabank.org/).

Toropova K, Zou S, Roberts AJ, Redwine WB, Goodman BS, Reck-Peterson SL, Leschziner AE, 2014, Dynein-Lis1 (ATP + Vi conditions) EM structure, EMD-6016; http://www.ebi.ac.uk/pdbe/entry/EMD-6016, Available at the EMDataBank repository (http://www.emdatabank.org/).

Toropova K, Zou S, Roberts AJ, Redwine WB, Goodman BS, Reck-Peterson SL, Leschziner AE, 2014, Short linker dynein-Lis1 (no nucleotide conditions) EM structure, EMD-6017; http://www.ebi.ac.uk/pdbe/entry/EMD-6017, Available at the EMDataBank repository (http://www.emdatabank.org/).

References

1. Aguet F, Antonescu CN, Mettlen M, Schmid SL, Danuser G. Advances in analysis of low signal-to-noise images link dynamin and AP2 to the functions of an endocytic checkpoint. Developmental Cell. 2013;26:279–291. doi: 10.1016/j.devcel.2013.06.019. [DOI] [PMC free article] [PubMed] [Google Scholar]
2. Blocker A, Severin FF, Burkhardt JK, Bingham JB, Yu H, Olivo JC, Schroer TA, Hyman AA, Griffiths G. Molecular requirements for bi-directional movement of phagosomes along microtubules. The Journal of Cell Biology. 1997;137:113–129. doi: 10.1083/jcb.137.1.113. [DOI] [PMC free article] [PubMed] [Google Scholar]
3. Burgess SA, Walker ML, Sakakibara H, Knight PJ, Oiwa K. Dynein structure and power stroke. Nature. 2003;421:715–718. doi: 10.1038/nature01377. [DOI] [PubMed] [Google Scholar]
4. Cardone G, Heymann JB, Steven AC. One number does not fit all: mapping local variations in resolution in cryo-EM reconstructions. Journal of Structural Biology. 2013;184:226–236. doi: 10.1016/j.jsb.2013.08.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
5. Carragher B, Kisseberth N, Kriegman D, Milligan RA, Potter CS, Pulokas J, Reilein A. Leginon: an automated system for acquisition of images from vitreous ice specimens. Journal of Structural Biology. 2000;132:33–45. doi: 10.1006/jsbi.2000.4314. [DOI] [PubMed] [Google Scholar]
6. Carter AP. Crystal clear insights into how the dynein motor moves. Journal of Cell Science. 2013;126:705–713. doi: 10.1242/jcs.120725. [DOI] [PubMed] [Google Scholar]
7. Carter AP, Garbarino JE, Wilson-Kubalek EM, Shipley WE, Cho C, Milligan RA, Vale RD, Gibbons IR. Structure and functional role of dynein's microtubule-binding domain. Science. 2008;322:1691–1695. doi: 10.1126/science.1164424. [DOI] [PMC free article] [PubMed] [Google Scholar]
8. Case RB, Pierce DW, Hom-Booher N, Hart CL, Vale RD. The directional preference of kinesin motors is specified by an element outside of the motor catalytic domain. Cell. 1997;90:959–966. doi: 10.1016/S0092-8674(00)80360-8. [DOI] [PubMed] [Google Scholar]
9. Cho C, Reck-Peterson S, Vale R. Cytoplasmic dynein's regulatory ATPase sites affect processivity and force generation. The Journal of Biological Chemistry. 2008;283:25839–25845. doi: 10.1074/jbc.M802951200. [DOI] [PMC free article] [PubMed] [Google Scholar]
10. Clegg RM. Fluorescence resonance energy transfer. Current Opinion in Biotechnology. 1995;6:103–110. doi: 10.1016/0958-1669(95)80016-6. [DOI] [PubMed] [Google Scholar]
11. De Carlo S, Stark H. Cryonegative staining of macromolecular assemblies. Methods in Enzymology. 2010;481:127–145. doi: 10.1016/S0076-6879(10)81006-7. [DOI] [PubMed] [Google Scholar]
12. Driskell OJ, Mironov A, Allan VJ, Woodman PG. Dynein is required for receptor sorting and the morphogenesis of early endosomes. Nature Cell Biology. 2007;9:113–120. doi: 10.1038/ncb1525. [DOI] [PubMed] [Google Scholar]
13. Edgar RC. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research. 2004;32:1792–1797. doi: 10.1093/nar/gkh340. [DOI] [PMC free article] [PubMed] [Google Scholar]
14. Egan MJ, Tan K, Reck-Peterson SL. Lis1 is an initiation factor for dynein-driven organelle transport. The Journal of Cell Biology. 2012;197:971–982. doi: 10.1083/jcb.201112101. [DOI] [PMC free article] [PubMed] [Google Scholar]
15. Frank J, Radermacher M, Penczek P, Zhu J, Li Y, Ladjadj M, Leith A. SPIDER and WEB: processing and visualization of images in 3D electron microscopy and related fields. Journal of Structural Biology. 1996;116:190–199. doi: 10.1006/jsbi.1996.0030. [DOI] [PubMed] [Google Scholar]
16. Gee MA, Heuser JE, Vallee RB. An extended microtubule-binding structure within the dynein motor domain. Nature. 1997;390:636–639. doi: 10.1038/37663. [DOI] [PubMed] [Google Scholar]
17. Gibbons IR, Garbarino JE, Tan CE, Reck-Peterson SL, Vale RD, Carter AP. The affinity of the dynein microtubule-binding domain is modulated by the conformation of its coiled-coil stalk. The Journal of Biological Chemistry. 2005;280:23960–23965. doi: 10.1074/jbc.M501636200. [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Gibbons IR, Gibbons BH, Mocz G, Asai DJ. Multiple nucleotide-binding sites in the sequence of dynein beta heavy chain. Nature. 1991;352:640–643. doi: 10.1038/352640a0. [DOI] [PubMed] [Google Scholar]
19. Gibbons IR, Lee-Eiford A, Mocz G, Phillipson CA, Tang WJ, Gibbons BH. Photosensitized cleavage of dynein heavy chains. Cleavage at the “V1 site” by irradiation at 365 nm in the presence of ATP and vanadate. The Journal of Biological Chemistry. 1987;262:2780–2786. [PubMed] [Google Scholar]
20. Gietz RD, Woods RA. Transformation of yeast by lithium acetate/single-stranded carrier DNA/polyethylene glycol method. Methods in Enzymology. 2002;350:87–96. doi: 10.1016/S0076-6879(02)50957-5. [DOI] [PubMed] [Google Scholar]
21. Guthrie C, Fink GR. Guide to yeast genetics and molecular biology. Methods in Enzymology. 1991;194:1–863. [PubMed] [Google Scholar]
22. Heald RR, Tournebize RR, Blank TT, Sandaltzopoulos RR, Becker PP, Hyman AA, Karsenti EE. Self-organization of microtubules into bipolar spindles around artificial chromosomes in Xenopus egg extracts. Nature. 1996;382:420–425. doi: 10.1038/382420a0. [DOI] [PubMed] [Google Scholar]
23. Huang J, Roberts AJ, Leschziner AE, Reck-Peterson SL. Lis1 acts as a “clutch” between the ATPase and microtubule-binding domains of the dynein motor. Cell. 2012;150:975–986. doi: 10.1016/j.cell.2012.07.022. [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Imamula K, Kon T, Ohkura R, Sutoh K. The coordination of cyclic microtubule association/dissociation and tail swing of cytoplasmic dynein. Proceedings of the National Academy of Sciences of USA. 2007;104:16134–16139. doi: 10.1073/pnas.0702370104. [DOI] [PMC free article] [PubMed] [Google Scholar]
25. Jaspersen SL, Winey M. The budding yeast spindle pole body: structure, duplication, and function. Annual Review of Cell and Developmental Biology. 2004;20:1–28. doi: 10.1146/annurev.cellbio.20.022003.114106. [DOI] [PubMed] [Google Scholar]
26. Jordens II, Fernandez-Borja MM, Marsman MM, Dusseljee SS, Janssen LL, Calafat JJ, Janssen HH, Wubbolts RR, Neefjes JJ. The Rab7 effector protein RILP controls lysosomal transport by inducing the recruitment of dynein-dynactin motors. Current Biology. 2001;11:1680–1685. doi: 10.1016/S0960-9822(01)00531-0. [DOI] [PubMed] [Google Scholar]
27. Kim MH, Cooper DR, Oleksy A, Devedjiev Y, Derewenda U, Reiner O, Otlewski J, Derewenda ZS. The structure of the N-terminal domain of the product of the lissencephaly gene Lis1 and its functional implications. Structure. 2004;12:987–998. doi: 10.1016/j.str.2004.03.024. [DOI] [PubMed] [Google Scholar]
28. Kiyomitsu T, Cheeseman IM. Cortical dynein and asymmetric membrane elongation coordinately position the spindle in anaphase. Cell. 2013;154:391–402. doi: 10.1016/j.cell.2013.06.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Kon T, Imamula K, Roberts AJ, Ohkura R, Knight PJ, Gibbons IR, Burgess SA, Sutoh K. Helix sliding in the stalk coiled coil of dynein couples ATPase and microtubule binding. Nature Structural & Molecular Biology. 2009;16:325–333. doi: 10.1038/nsmb.1555. [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Kon T, Mogami T, Ohkura R, Nishiura M, Sutoh K. ATP hydrolysis cycle-dependent tail motions in cytoplasmic dynein. Nature Structural & Molecular Biology. 2005;12:513–519. doi: 10.1038/nsmb930. [DOI] [PubMed] [Google Scholar]
31. Kon T, Nishiura M, Ohkura R, Toyoshima YY, Sutoh K. Distinct functions of nucleotide-binding/hydrolysis sites in the four AAA modules of cytoplasmic dynein. Biochemistry. 2004;43:11266–11274. doi: 10.1021/bi048985a. [DOI] [PubMed] [Google Scholar]
32. Kon T, Oyama T, Shimo-Kon R, Imamula K, Shima T, Sutoh K, Kurisu G. The 2.8 Å crystal structure of the dynein motor domain. Nature. 2012;484:345–350. doi: 10.1038/nature10955. [DOI] [PubMed] [Google Scholar]
33. Kural C, Kim H, Syed S, Goshima G, Gelfand VI, Selvin PR. Kinesin and dynein move a peroxisome in vivo: a tug-of-war or coordinated movement? Science. 2005;308:1469–1472. doi: 10.1126/science.1108408. [DOI] [PubMed] [Google Scholar]
34. Lander GC, Stagg SM, Voss NR, Cheng A, Fellmann D, Pulokas J, Yoshioka C, Irving C, Mulder A, Lau PW, Lyumkis D, Potter CS, Carragher B. Appion: an integrated, database-driven pipeline to facilitate EM image processing. Journal of Structural Biology. 2009;166:95–102. doi: 10.1016/j.jsb.2009.01.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
35. Lee WL, Oberle JR, Cooper JA. The role of the lissencephaly protein Pac1 during nuclear migration in budding yeast. The Journal of Cell Biology. 2003;160:355–364. doi: 10.1083/jcb.200209022. [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Lenz JH, Schuchardt I, Straube A, Steinberg G. A dynein loading zone for retrograde endosome motility at microtubule plus-ends. The EMBO Journal. 2006;25:2275–2286. doi: 10.1038/sj.emboj.7601119. [DOI] [PMC free article] [PubMed] [Google Scholar]
37. Longtine MS, McKenzie A, Demarini DJ, Shah NG, Wach A, Brachat A, Philippsen P, Pringle JR. Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast. 1998;14:953–961. doi: 10.1002/(SICI)1097-0061(199807)14:103.0.CO;2-U. [DOI] [PubMed] [Google Scholar]
38. Ludtke SJ, Baldwin PR, Chiu W. EMAN: semiautomated software for high-resolution single-particle reconstructions. Journal of Structural Biology. 1999;128:82–97. doi: 10.1006/jsbi.1999.4174. [DOI] [PubMed] [Google Scholar]
39. Markus SM, Lee WL. Regulated offloading of cytoplasmic dynein from microtubule plus ends to the cortex. Developmental Cell. 2011;20:639–651. doi: 10.1016/j.devcel.2011.04.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
40. McKenney RJ, Vershinin M, Kunwar A, Vallee RB, Gross SP. LIS1 and NudE induce a persistent dynein force-producing state. Cell. 2010;141:304–314. doi: 10.1016/j.cell.2010.02.035. [DOI] [PMC free article] [PubMed] [Google Scholar]
41. Merdes A, Ramyar K, Vechio JD, Cleveland DW. A complex of NuMA and cytoplasmic dynein is essential for mitotic spindle assembly. Cell. 1996;87:447–458. doi: 10.1016/S0092-8674(00)81365-3. [DOI] [PubMed] [Google Scholar]
42. Moore J, Stuchell-Brereton M, Cooper J. Function of dynein in budding yeast: mitotic spindle positioning in a polarized cell. Cell Motility and the Cytoskeleton. 2009;66:546–555. doi: 10.1002/cm.20364. [DOI] [PMC free article] [PubMed] [Google Scholar]
43. Moughamian AJ, Osborn GE, Lazarus JE, Maday S, Holzbaur EL. Ordered recruitment of dynactin to the microtubule plus-end is required for efficient initiation of retrograde axonal transport. The Journal of Neuroscience. 2013;33:13190–13203. doi: 10.1523/JNEUROSCI.0935-13.2013. [DOI] [PMC free article] [PubMed] [Google Scholar]
44. Nishiura M, Kon T, Shiroguchi K, Ohkura R, Shima T, Toyoshima YY, Sutoh K. A single-headed recombinant fragment of Dictyostelium cytoplasmic dynein can drive the robust sliding of microtubules. The Journal of Biological Chemistry. 2004;279:22799–22802. doi: 10.1074/jbc.M313362200. [DOI] [PubMed] [Google Scholar]
45. Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, Ferrin TE. UCSF Chimera–a visualization system for exploratory research and analysis. Journal of Computational Chemistry. 2004;25:1605–1612. doi: 10.1002/jcc.20084. [DOI] [PubMed] [Google Scholar]
46. Pilling AD, Horiuchi D, Lively CM, Saxton WM. Kinesin-1 and Dynein are the primary motors for fast transport of mitochondria in Drosophila motor axons. Molecular Biology of the Cell. 2006;17:2057–2068. doi: 10.1091/mbc.E05-06-0526. [DOI] [PMC free article] [PubMed] [Google Scholar]
47. Reck-Peterson SL, Yildiz A, Carter AP, Gennerich A, Zhang N, Vale RD. Single-molecule analysis of dynein processivity and stepping behavior. Cell. 2006;126:335–348. doi: 10.1016/j.cell.2006.05.046. [DOI] [PMC free article] [PubMed] [Google Scholar]
48. Redwine WB, Hernandez-Lopez R, Zou S, Huang J, Reck-Peterson SL, Leschziner AE. Structural basis for microtubule binding and release by dynein. Science. 2012;337:1532–1536. doi: 10.1126/science.1224151. [DOI] [PMC free article] [PubMed] [Google Scholar]
49. Reiner O, Carrozzo R, Shen Y, Wehnert M, Faustinella F, Dobyns WB, Caskey CT, Ledbetter DH. Isolation of a Miller-Dieker lissencephaly gene containing G protein beta-subunit-like repeats. Nature. 1993;364:717–721. doi: 10.1038/364717a0. [DOI] [PubMed] [Google Scholar]
50. Roberts AJ, Goodman BS, Reck-Peterson SL. Reconstitution of dynein transport to the microtubule plus end by kinesin. Elife. 2014;3:e02641. doi: 10.7554/eLife.02641. [DOI] [PMC free article] [PubMed] [Google Scholar]
51. Roberts AJ, Malkova B, Walker ML, Sakakibara H, Numata N, Kon T, Ohkura R, Edwards TA, Knight PJ, Sutoh K, Oiwa K, Burgess SA. ATP-driven remodeling of the linker domain in the dynein motor. Structure. 2012;20:1670–1680. doi: 10.1016/j.str.2012.07.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
52. Roberts AJ, Numata N, Walker ML, Kato YS, Malkova B, Kon T, Ohkura R, Arisaka F, Knight PJ, Sutoh K, Burgess SA. AAA+ Ring and linker swing mechanism in the dynein motor. Cell. 2009;136:485–495. doi: 10.1016/j.cell.2008.11.049. [DOI] [PMC free article] [PubMed] [Google Scholar]
53. Scheres SH. RELION: implementation of a Bayesian approach to cryo-EM structure determination. Journal of Structural Biology. 2012;180:519–530. doi: 10.1016/j.jsb.2012.09.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
54. Schmidt H, Gleave ES, Carter AP. Insights into dynein motor domain function from a 3.3-Å crystal structure. Nature Structural & Molecular Biology. 2012;19:492–497. doi: 10.1038/nsmb.2272. [DOI] [PMC free article] [PubMed] [Google Scholar]
55. Sheeman B, Carvalho P, Sagot I, Geiser J, Kho D, Hoyt MA, Pellman D. Determinants of S. cerevisiae dynein localization and activation: implications for the mechanism of spindle positioning. Current Biology. 2003;13:364–372. doi: 10.1016/S0960-9822(03)00013-7. [DOI] [PubMed] [Google Scholar]
56. Shima T, Kon T, Imamula K, Ohkura R, Sutoh K. Two modes of microtubule sliding driven by cytoplasmic dynein. Proceedings of the National Academy of Sciences of USA. 2006;103:17736–17740. doi: 10.1073/pnas.0606794103. [DOI] [PMC free article] [PubMed] [Google Scholar]
57. Sorzano COS, Marabini R, Velázquez-Muriel J, Bilbao-Castro JR, Scheres SHW, Carazo JM, Pascual-Montano A. XMIPP: a new generation of an open-source image processing package for electron microscopy. Journal of Structural Biology. 2004;148:194–204. doi: 10.1016/j.jsb.2004.06.006. [DOI] [PubMed] [Google Scholar]
58. Tang G, Peng L, Baldwin PR, Mann DS, Jiang W, Rees I, Ludtke SJ. EMAN2: an extensible image processing suite for electron microscopy. Journal of Structural Biology. 2007;157:38–46. doi: 10.1016/j.jsb.2006.05.009. [DOI] [PubMed] [Google Scholar]
59. Tarricone C, Perrina F, Monzani S, Massimiliano L, Kim M-H, Derewenda ZS, Knapp S, Tsai L-H, Musacchio A. Coupling PAF signaling to dynein regulation: structure of LIS1 in complex with PAF-acetylhydrolase. Neuron. 2004;44:809–821. doi: 10.1016/j.neuron.2004.11.019. [DOI] [PubMed] [Google Scholar]
60. Torisawa T, Nakayama A, Furuta K, Yamada M, Hirotsune S, Toyoshima YY. Functional dissection of LIS1 and NDEL1 towards understanding the molecular mechanisms of cytoplasmic dynein regulation. The Journal of Biological Chemistry. 2011;286:1959–1965. doi: 10.1074/jbc.M110.169847. [DOI] [PMC free article] [PubMed] [Google Scholar]
61. Vale RD. The molecular motor toolbox for intracellular transport. Cell. 2003;112:467–480. doi: 10.1016/S0092-8674(03)00111-9. [DOI] [PubMed] [Google Scholar]
62. van Heel M, Harauz G, Orlova EV, Schmidt R, Schatz M. A new generation of the IMAGIC image processing system. Journal of Structural Biology. 1996;116:17–24. doi: 10.1006/jsbi.1996.0004. [DOI] [PubMed] [Google Scholar]
63. Wilkie GS, Davis I. Drosophila wingless and pair-rule transcripts localize apically by dynein-mediated transport of RNA particles. Cell. 2001;105:209–219. doi: 10.1016/S0092-8674(01)00312-9. [DOI] [PubMed] [Google Scholar]
64. Yamada M, Toba S, Yoshida Y, Haratani K, Mori D, Yano Y, Mimori-Kiyosue Y, Nakamura T, Itoh K, Fushiki S, Setou M, Wynshaw-Boris A, Torisawa T, Toyoshima YY, Hirotsune S. LIS1 and NDEL1 coordinate the plus-end-directed transport of cytoplasmic dynein. The EMBO Journal. 2008;27:2471–2483. doi: 10.1038/emboj.2008.182. [DOI] [PMC free article] [PubMed] [Google Scholar]
65. Yeh E, Skibbens RV, Cheng JW, Salmon ED, Bloom K. Spindle dynamics and cell cycle regulation of dynein in the budding yeast, Saccharomyces cerevisiae. The Journal of Cell Biology. 1995;130:687–700. doi: 10.1083/jcb.130.3.687. [DOI] [PMC free article] [PubMed] [Google Scholar]
66. Yin J, Straight PD, McLoughlin SM, Zhou Z, Lin AJ, Golan DE, Kelleher NL, Kolter R, Walsh CT. Genetically encoded short peptide tag for versatile protein labeling by Sfp phosphopantetheinyl transferase. Proceedings of the National Academy of Sciences of USA. 2005;102:15815–15820. doi: 10.1073/pnas.0507705102. [DOI] [PMC free article] [PubMed] [Google Scholar]

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for sending your work entitled “Lis1 regulates dynein by sterically blocking its mechanochemical cycle” for consideration at eLife. Your article has been favorably evaluated by John Kuriyan (Senior editor) and 3 reviewers, one of whom, Gabriel Lander, has agreed to reveal his identity.

The Senior editor and the three reviewers discussed their comments before we reached this decision, and the Senior editor has assembled the following comments to help you prepare a revised submission.

In the present work, the authors use electron microscopy to determine the three-dimensional structure of dynein bound to Lis1, enabling unambiguous identification of this regulator's binding site near the stalk of the motor. Although the resolutions of the reconstructions are moderate (in the 15-20Å range), the resolvable structural details were more than adequate to identify the key interactions that were involved in Lis1 regulation of dynein.

This group had a breakthrough paper in 2012, determining the Lis1-binding domain on the dynein motor domain, and using elegant single molecule studies to develop a clutch model for Lis1-mediated regulation of dynein motility. This work is a logical follow-up, with 3-D structures that support their previous binding data, and a clever short-linker mutant to test their more refined steric-blocking model.

Comparing the structure of the dynein head in the absence and presence of Lis1, the authors noticed that Lis1 binding induces a significant displacement of the dynein linker arm, a critical component of the mechanochemical cycle. Based on this finding, the authors carried out a series of well-designed biochemical and biophysical experiments to probe the mechanical relationship between bound Lis1 and the linker arm. By docking in previously determined atomic coordinates, the resulting pseudo-atomic models served as a framework for the mutagenesis and FRET experiments, which provided insight into the relationship between Lis1, nucleotide state, and the linker arm position. Truncation of the linker arm, single molecule motility experiments, and in vivo assays were performed to confirm the author's model for Lis1 regulation. Ultimately, the authors conclude that Lis1 serves as a molecular barricade, sterically preventing ATP hydrolysis from being converted into the conformational rearrangements that weaken microtubule binding.

Some of the important insights into how Lis1 works that emerge from this work are as follows:

1. That Lis1 occludes to normal position of the dynein linker in the ADP and nucleotide free (APO) conformations of dynein. In the presence of Lis1, the linker is deflected from its normal position. This EM work also provides a nice confirmation that the two conformations observed in yeast(APO) vs Dictyostellium(ADP) crystal structures can both exist in one (yeast) dynein.

2. That the motor can still enter the pre-powerstroke conformation when Lis-1 is bound. This was demonstrated by both an EM structure in the presence of ATP.vanadate and a FRET assay.

3. That the ability of Lis-1 to block release of dynein from microtubules is abolished when the linker is shortened (from the 331kD to 314kD construct). This shows that the N-terminal part of the linker is required for Lis1 function.

The EM work is expertly done, particularly in exploring the conformational variability that exists within a set of particles. The dynein motor is relatively small for detailed structural analyses by EM, and the authors' ability to distinguish between conformational subtleties in the motor arm is an impressive example of their expertise.

In general, the consensus among the reviewers is that this work is an important step forward in understanding dynein motor regulation, and provides a direct view of the conformational rearrangements that are necessary for dynein movement along the microtubule surface. The reviewers have, however, raised a number of points that you should address in preparing a revised manuscript.

1. There is some concern that much of this work follows directly from the outstanding Huang et al. 2012 paper published by this group. In this paper, the authors further refine the model developed there, and while the three-dimensional structure adds knowledge, it is limited in detail because the docking of Lis1 is not fully resolved in terms of rotational alignment. The couple of new mutants examined here could also have been predicted based on the previous structure.

The most novel idea here is that the binding of Lis1 provides a steric block to the linker swing. This is tested with a single construct with a truncated linker, and the results support their model. But there is concern that this is a single approach; the paper would have more impact if the authors could demonstrate this key point in a complementary way. The reviewers recognize that this might be a tall order, but it would be good if the authors could respond to this issue.

1. The authors interpret their data with a model in which the linker needs to get to AAA5 in order for the cycle of ATP hydrolysis to continue and so blocking access to AAA5 prevents ATP binding and therefore release from microtubules (which is thought to occur on ATP hydrolysis). The problem with this model, as the authors discuss, is that it would predict that Lis1 would inhibit the dynein ATPase. The data, however, are clear that this is not the case. It is suggested that the authors reduce the emphasis on the model that Lis1 blocks the linker from getting to AAA5. Instead the data could be presented and then this discussed as an option.

More specific points to consider:

Specific issues raised by the reviewers are given below. The authors should use their judgement in how to respond to these points, but if they choose to not address any particular issue they should explain their reasons in the cover letter accompanying the resubmitted manuscript.

1. The authors employed cryo-negative staining for their EM analysis. This involved and tedious technique is not commonly used by EM labs, as it is difficult to master. Surely the authors had a reason to employ this technique; a sentence or two explaining their rationale would be of interest to other structural labs interested in studying similarly sized macromolecular complexes.

2. A final summarizing cartoon outlining the relationship of Lis1 to the mechanochemical cycle of the dynein motor, and how it translates to MT affinity would strengthen the impact of this manuscript.

3. No mention is made of deposition of the 5 EM densities to the EMDB. It is important that these are made available to the community.

4. The authors state in their Abstract that how Lis1 regulates dynein is unknown, but in light of their previous work as well as that of the Vallee lab, this is a clear over-statement. This continues on through the Introduction, where they underplay their previous work determining the binding interface of Lis1 on dynein. This should be revised.

5. The authors set up an either/or model that seems overly simplistic, and the terms ring-centric and linker-centric are clunky and not very clear.

6. The authors report that their dynein alone map “accommodates the crystal structure of the dynein motor domain well”. While there is an overall match, there are also clearly discrepancies between the structures. How was the fit assessed quantitatively? Can anything be learned from the discrepancies?

7. The authors report that the 3D structure allows them “to hone in on the region of dynein interacting with Lis1”, but other than the displacement of the linker, they do not go further with this approach. Are there any other changes? It would be good to discuss this, and the Lis1-dynein interface in more detail.

8. It is unclear what new is learned from the section on “one dynein ring binds one Lis1 beta-propeller” Isn't this what they studied in their previous paper?

9. Did they intend to reference Carragher et al?

10. Vanadate is not so much a phosphate analog as a transition state analog. Similarly, calling the ADP-V complex a “dead-end” is awkward and imprecise terminology.

11. The authors suggest that the differences in AAA4 and AAA5 linker positions previously noted might correspond to either distinct mechanochemical states or different dynein species. But isn't it also possible that they may result from different constructs used in the different studies, some of which were truncated or chimeric?

12. Figure 2 would be stronger if the authors included parallel studies with the W419A mutant and provided more complete analysis of the motility data. Based on what is shown here, the R378A mutant effectively blocks formation of the dynein-Lis1 complex as judged by gel filtration, but only partially blocks the effects of Lis1 on dynein motility. Shoring up this point with an additional single point mutation would strengthen the point.

13. The authors report that two structures are seen in the presence of ADP and one in the presence of no nucleotide, so they conclude that the AAA4 linker position is the ADP state. This may be true, but kinetic analysis has indicated that there is isomerization between 2 ADP-bound states, so it is also possible that the two observed structures correspond to different ADP-bound states. Both should be shown in Figure 3a. Didn't the authors perform FRET analysis of the dynein-Lis1 complex alone? How come this is not included in Figure 3?

14. The authors report the ADP-V-dynein-Lis1 complex. Wouldn't it be useful to directly compare this to the ADP-V-dynein complex?

15. The experiments described in the text and in Figure 4 yield results that are predictable. Three papers, including work from this same group, have already shown that Lis1 does not significantly affect dynein's ATPase rate, so it is unclear what new is learned from this work. It could all be removed from the text, or at least moved to supplemental information. If it does remain, experiments need to be performed in triplicate, and error bars should be reported for all points.

16. In Figure 5, for the short linker construct, why are there fewer events, including fewer rebinding events? Does this construct have a weaker affinity for microtubules? If it does, how might this affect the interpretation of the results?

17. Use of dimeric Lis1. It is not clear how an avidity effect would explain how dimeric Lis1 would bind more tightly to monomeric dynein than monomeric Lis1. The avidity effect should require two binding sites for Lis1. In Huang et al there does not seem to be a direct demonstration that dimeric Lis1 binds monomeric dynein better than monomeric Lis1. Please clarify.

18. If the Lis1 purely displaces the linker by a steric mechanism, would the authors not expect to see the linker closely connected to Lis1? Instead Figure 1e suggests that the linker is making a tight connection to the AAA+ ring. Should this connection be discussed? Might it suggest that Lis1 has some allosteric effect on the AAA+ ring which, together with sterically blocking its normal position, allows the linker to dock onto the ring? Alternatively is the resolution too low to say this?

19. The authors try to draw the parallel between mutations in AAA5 and the effect of Lis1. The reviewer's reading of the data is that these two effects are different as the first abolishes ATPase activity, while the second has no effect on it.

20. One section states that the lack of conservation in the very N-terminus of the linker suggests it does not interact directly with Lis1 (which seems reasonable) and therefore that the Lis1 is acting as a steric block. The contact between the linker and the AAA+ ring in the Lis1:APO-Dynein structure appears to involve a more conserved part of the ring. This comes back to the question about whether Lis1 is acting on the AAA+ ring to allow the linker to bind to a place it normally would not.

21. Figure 2c. The blue color is difficult to distinguish from the black in the gel filtration trace.

22. Figure 3f/g. This EM map looks as though the contacts between the AAA+ ring and the Lis1 are different from in the Apo form. This may just be the different resolution of the structure, but it might be useful to discuss this somewhere.

23. Figure 4. It would be helpful if the type of mutation made was specified on this figure. b & c are E-Q mutations (residue number?). d) AAA5 mutant is not in the ATP binding site, but on the linker binding site. Specifying this on the figure (& res numbers?) would be helpful.

24. Figure 5 and Figure 5–figure supplement 1. These figures show release from microtubules is blocked by Lis1, but not if the linker is shortened. The shortening does not affect the motors activity or ability to bind Lis1. Did the authors test if the velocity of the shorter linker was affected by Lis1?

25. Figure 5 d ii: The cartoon uses the stop sign, which suggests it has a mutation in AAA5 (as in Figure 4). Is this true? If not then another symbol might be more appropriate.

26. Table 3:

Table 3.

The units of Km, kbasal and kcat should be given.

One of the reviewers has the following thought which they would like to share with you:

Another mechanism that could be mentioned in the Discussion is that Lis1 has an allosteric effect on the AAA+ ring that allows the linker to move and the AAA1 site to exchange nucleotide without triggering the conformational change in the stalk that releases dynein from microtubules. This model is also not completely satisfactory as it is harder to explain the requirement for the N-terminus of the linker.

Perhaps one (explanation would be that the transition from high to low affinity is most likely to occur before the linker moves from a post to pre-powerstroke position. That is, ATP binds to a high affinity, post-powerstroke dynein. A rearrangement occurs that leads to a low affinity, post-powerstroke dynein (that releases from MTs). This is followed by a transition to the ADP.Vi like state with a low affinity, pre-powerstroke conformation. The ATP bound, post-powerstroke, low affinity conformation could be altered so that it no longer releases from microtubules by a combination of Lis1 AND linker binding to the AAA+ ring.

1) There is some concern that much of this work follows directly from the outstanding Huang et al. 2012 paper published by this group. In this paper, the authors further refine the model developed there, and while the three-dimensional structure adds knowledge, it is limited in detail because the docking of Lis1 is not fully resolved in terms of rotational alignment. The couple of new mutants examined here could also have been predicted based on the previous structure.

The most novel idea here is that the binding of Lis1 provides a steric block to the linker swing. This is tested with a single construct with a truncated linker, and the results support their model. But there is concern that this is a single approach; the paper would have more impact if the authors could demonstrate this key point in a complementary way. The reviewers recognize that this might be a tall order, but it would be good if the authors could respond to this issue.

To further test the steric block model of Lis1 regulation of dynein we chose to explore if the specific sequence of the N-terminal portion of the linker (the part that is removed in the short linker dynein) is important for Lis1 regulation of dynein (as opposed to its simply being a linker-like protein mass that can be blocked). We chose two different sequences to add back to the short linker dynein to restore the wild type linker length: (1) The sequence from S. pombe, which appears to lack a Lis1 homolog; and (2) The sequence from an axonemal dynein from C. reinhardtii (dynein-c), which does not bind axonemal Lis1. Our goal was to then use single-molecule microtubule binding/release assays on these chimeric dyneins, alone and in the presence of Lis1, to test if Lis1 regulation was restored. Unfortunately, the chimeric axonemal dynein failed to express. The chimeric S. pombe dynein did not express well, either, but we were able to purify amounts that were sufficient for single-molecule studies. However, this chimeric dynein showed a disrupted mechanochemical cycle: it was unable to release from microtubules in the presence of ATP even in the absence of Lis1. We agree that additional evidence for the steric block hypothesis would have strengthened our model; unfortunately, given the results described above, this does seem to be too tall an order to be accomplished in a reasonable time frame.

2) The authors interpret their data with a model in which the linker needs to get to AAA5 in order for the cycle of ATP hydrolysis to continue and so blocking access to AAA5 prevents ATP binding and therefore release from microtubules (which is thought to occur on ATP hydrolysis). The problem with this model, as the authors discuss, is that it would predict that Lis1 would inhibit the dynein ATPase. The data, however, are clear that this is not the case. It is suggested that the authors reduce the emphasis on the model that Lis1 blocks the linker from getting to AAA5. Instead the data could be presented and then this discussed as an option.

We agree with this suggestion and have made the appropriate changes in the last Results section and the Discussion.

More specific points to consider:

1) The authors employed cryo-negative staining for their EM analysis. This involved and tedious technique is not commonly used by EM labs, as it is difficult to master. Surely the authors had a reason to employ this technique; a sentence or two explaining their rationale would be of interest to other structural labs interested in studying similarly sized macromolecular complexes.

Thank you for this suggestion; we have added text to both the Results and Methods sections to explain our choice.

2) A final summarizing cartoon outlining the relationship of Lis1 to the mechanochemical cycle of the dynein motor, and how it translates to MT affinity would strengthen the impact of this manuscript.

We have added a model figure (Figure 6).

3) No mention is made of deposition of the 5 EM densities to the EMDB. It is important that these are made available to the community.

We apologize for accidentally not making our intent regarding EMDB submission clear. Final maps as well as raw half maps for each reconstruction have been deposited with the EMDB. This is now also stated in the manuscript.

4) The authors state in their Abstract that how Lis1 regulates dynein is unknown, but in light of their previous work as well as that of the Vallee lab, this is a clear over-statement. This continues on through the Introduction, where they underplay their previous work determining the binding interface of Lis1 on dynein. This should be revised.

We’ve discussed this point extensively among us and feel we disagree with this assessment. What we state in our Abstract is that “The dynein regulator Lis1 is known to keep dynein bound to microtubules; however, how this is accomplished mechanistically remains unknown”. We feel this statement is correct; the key distinction here is between “what” Lis1 does (which we established in our 2012 paper), and “how” it does it (which we did not). Our Introduction summarizes how much is known, from both our and the Vallee lab’s work, about what Lis1 does to dynein. However, we feel there is a clear difference between what are ultimately descriptions of biochemical effects from a more detailed understanding of the mechanism behind them. While we have not yet answered all the questions in this regard, we think the work presented here has gone a long ways towards explaining the “how”.

Similarly, our previous work did not address the binding site on Lis1 for the dynein-Lis1 interaction but rather only residues in dynein. Some other important aspects of the dynein-Lis1 interface that were not addressed by our previous work include the face of the ring to which Lis1 binds (which would determine whether the linker might be directly affected), and which side of the Lis1 propeller interacted with dynein (even if sequence conservation had already made a strong argument for what we saw in our structure).

5) The authors set up an either/or model that seems overly simplistic, and the terms ring-centric and linker-centric are clunky and not very clear.

Thank you for the comment. We now use “steric” and “allosteric” models, and do not present them as being mutually exclusive.

1. The authors report that their dynein alone map “accommodates the crystal structure of the dynein motor domain well”. While there is an overall match, there are also clearly discrepancies between the structures. How was the fit assessed quantitatively? Can anything be learned from the discrepancies?

We have carried out a Fourier Shell Correlation (FSC) between the dynein alone map and the crystal structure of the dynein motor domain (PDB ID: 4AKG). We see an FSC of 0.143 at a resolution of 18.8 Å. We deposited an XML file of the FSC plot in the EMDB as a supplementary file to the dynein alone submission (EMDB-6013). This is briefly mentioned in Results and the deposition is noted in the Accession Numbers section in Materials and methods.

With regards to discrepancies between the EM map and the crystal structure in some regions, we felt a rigid body fit of the entire motor domain into the density was most appropriate at the resolution of our map. Although we are very interested in whether these differences exist and what they might be, we feel we do not yet have the resolution that would be required to fit individual AAA+ motifs in the density (and comment on that fit).

7) The authors report that the 3D structure allows them “to hone in on the region of dynein interacting with Lis1”, but other than the displacement of the linker, they do not go further with this approach. Are there any other changes? It would be good to discuss this, and the Lis1-dynein interface in more detail.

The displacement of the linker is the most striking difference in the dynein-Lis1 map compared to that of dynein alone. We are also very curious to see whether there are further changes in dynein’s ring upon Lis1 binding and will pursue this in future work. However, we feel that at the current ∼21 Å resolution we cannot trust, and comment on, any changes less apparent than the displaced linker domain.

With respect to the Lis1-dynein interface, we were able to directly visualize that dynein contacts Lis1 via the AAA3-AAA4 connector helix. Unfortunately, as we state in the paper, we did not have the resolution to determine Lis1’s rotational fit in the density. We therefore feel that we cannot currently go further than the view of the interface we present in Figure 2A. We worry that any additional details might lead readers into assuming the interface is fully established.

1. It is unclear what new is learned from the section on “one dynein ring binds one Lis1 beta-propeller” Isn't this what they studied in their previous paper?

We thank the reviewers for bringing this up, as we did not clearly highlight what was new in our current findings.

In our previous study, we used 2D classification of dimeric Lis1 in complex with a dimeric dynein. We observed extra density in a difference map that we could assign to Lis1 (thus establishing where on dynein Lis1 bound) but we couldn’t determine whether the density accommodated one or two propeller domains, or indeed any of the N-terminal domain of Lis1. It was not until we had the 3D map reported here that we could resolve the channel at the center of the propeller and thus were able to unambiguously show that a single propeller (and that domain alone) contacts the dynein motor domain.

We have modified this section of the Results to make the new findings stand out more clearly.

1. Did they intend to reference Carragher et al?

Thank you for spotting this error, we have replaced it with the correct reference.

10) Vanadate is not so much a phosphate analog as a transition state analog. Similarly, calling the ADP-V complex a “dead-end” is awkward and imprecise terminology.

We now refer to ADP.Vi as a transition state analog and have removed the term “dead-end”. Thank you for these suggestions.

1. The authors suggest that the differences in AAA4 and AAA5 linker positions previously noted might correspond to either distinct mechanochemical states or different dynein species. But isn't it also possible that they may result from different constructs used in the different studies, some of which were truncated or chimeric?

Thank you for pointing this out; we have added this possibility to the text.

12) Figure 2 would be stronger if the authors included parallel studies with the W419A mutant and provided more complete analysis of the motility data. Based on what is shown here, the R378A mutant effectively blocks formation of the dynein-Lis1 complex as judged by gel filtration, but only partially blocks the effects of Lis1 on dynein motility. Shoring up this point with an additional single point mutation would strengthen the point.

To give a more detailed account of the effects of the Lis1 mutants on dynein velocity we have added a histogram of velocity distributions for the motility experiments (new figure; Figure 2–figure supplement 2). We have not added additional analysis of run length data because we have previously found that velocity analysis is the most sensitive measure of Lis1’s effect on dynein. Run length analysis provides little information; since Lis1 both slows dynein down and prolongs its microtubule encounters, which have opposing effects on run length, its addition results in little change in the average travel distance [Huang et al., 2012].

With regards to the interpretation of the R378A mutant and analysis of mutant W419A, gel filtration is not a very sensitive measure of the dynein-Lis1 interaction; we would not expect a weak interaction to be detected in this assay. This is why we stated in the paper that “binding is severely weakened” for the R378A mutant rather than that this interaction does not exist (particularly given that the motility data still show that it does, albeit in a limited way). While doing the experiments with the W419A mutant suggested was feasible, these experiments would have been fairly time-consuming as the yeast strains needed for this analysis were not already available. Since we were not confident that the time investment would yield new information (given that this mutation is already encompassed in the Lis15A quintuple mutant), we chose to invest our time in experiments that would test the major mechanistic hypothesis of the paper, as suggested in general point #1 (even if those experiments were unsuccessful in the end). We have reworded the manuscript to address the apparent discrepancy between the gel filtration and single-molecule assays for R378A, making it clear that the single-molecule assays are more sensitive in terms of detecting an interaction and that not seeing co-migration by gel filtration is not an indication of a complete abolishment of that interaction.

13) The authors report that two structures are seen in the presence of ADP and one in the presence of no nucleotide, so they conclude that the AAA4 linker position is the ADP state. This may be true, but kinetic analysis has indicated that there is isomerization between 2 ADP-bound states, so it is also possible that the two observed structures correspond to different ADP-bound states. Both should be shown in Figure 3a. Didn't the authors perform FRET analysis of the dynein-Lis1 complex alone? How come this is not included in Figure 3?

Thank you for pointing this out, we have added the AAA4 linker position structure and FRET analysis of dynein-Lis1 (no nucleotide conditions) to Figure 3.

14) The authors report the ADP-V-dynein-Lis1 complex. Wouldn't it be useful to directly compare this to the ADP-V-dynein complex?

We are indeed very curious to see the structure of ADP.Vi-dynein (in the absence of Lis1) and compare it to the one we reported for the complex (ADP.Vi-dynein-Lis1) but feel this is beyond the scope of the current paper. The goal of the ADP.Vi-dynein-Lis1 structure was simply to establish (visually, in this case) whether the linker domain could reach AAA2 in the presence of Lis1.

15) The experiments described in the text and in Figure 4 yield results that are predictable. Three papers, including work from this same group, have already shown that Lis1 does not significantly affect dynein's ATPase rate, so it is unclear what new is learned from this work. It could all be removed from the text, or at least moved to supplemental information. If it does remain, experiments need to be performed in triplicate, and error bars should be reported for all points.

While it has indeed been shown before that Lis1 does not significantly affect dynein’s ATPase rate, no one had addressed the question of which AAA+ domain was responsible for the continued ATP turnover. In the work we report here we measured, for the first time, what happens to ATP hydrolysis in the presence of Lis1 when AAA1 or AAA4 are rendered hydrolysis deficient or AAA5 is linker-docking deficient. The experiments we present are therefore the first to probe which of dynein’s AAA+ domains are responsible for the observed nucleotide hydrolysis in the presence of Lis1 and, for that reason, we feel they belong in the main text.

Regarding the error bars, as the number of experiments was ≤ 3 (triplicate for experiments showing an activated ATPase rate, and duplicate for those that are ATPase deficient), we plotted each of these points separately. Error bars are not robust and can be misleading for such sample sizes [Krzywinski & Altman (2013) Nature Methods 10, 921–922]. Plotting individual data points is recommended practice in this situation, as it explicitly shows the spread in the data.

16) In Figure 5, for the short linker construct, why are there fewer events, including fewer rebinding events? Does this construct have a weaker affinity for microtubules? If it does, how might this affect the interpretation of the results?

The full-length and short linker constructs appear to have a similar affinity for microtubules based on the similar concentration of microtubules required for half-maximal activation of their ATPase rates [Km(MT)] (Table 3). The density of the full-length and short linker constructs on the microtubule in the release assays is also similar (0.289 ± 0.080 and 0.226 ± 0.081 dynein molecules per micrometer of microtubule, respectively). Small differences in density likely arise from different dilutions used. Regarding rebinding events, apart from cases where Lis1 triggers a clear persistent microtubule attachment state in dynein (i.e. full-length linker dynein with Lis1), comparative measurements are not very meaningful as rebinding events are in part dependent on the buffer flow rate through each flow chamber, which guides the rate of clearance of dynein molecules released from microtubules. This flow rate is manually driven via a syringe; so, it too can vary slightly between assays. Therefore, we do not base any of our conclusions on rebinding frequency. In summary, the data suggest that the short linker dynein has a similar microtubule affinity as the full-length linker dynein, and so our interpretations are not affected.

17) Use of dimeric Lis1. It is not clear how an avidity effect would explain how dimeric Lis1 would bind more tightly to monomeric dynein than monomeric Lis1. The avidity effect should require two binding sites for Lis1. In Huang et al there does not seem to be a direct demonstration that dimeric Lis1 binds monomeric dynein better than monomeric Lis1. Please clarify.

We apologize for the lack of clarity on this point. Previously we showed that a much greater concentration of monomeric Lis1 is needed to achieve the same effect on dynein motility observed with dimeric Lis1 [Huang et al., 2012]. This is the reason why we chose to use dimeric Lis1 in our EM work: it allowed us to use a much lower concentration of it, keeping the background in the images low. We have changed the wording for this section in the Results to discuss this in terms of local Lis1 β-propeller concentration rather than avidity.

18) If the Lis1 purely displaces the linker by a steric mechanism, would the authors not expect to see the linker closely connected to Lis1? Instead Figure 1e suggests that the linker is making a tight connection to the AAA+ ring. Should this connection be discussed? Might it suggest that Lis1 has some allosteric effect on the AAA+ ring which, together with sterically blocking its normal position, allows the linker to dock onto the ring? Alternatively is the resolution too low to say this?

It is indeed difficult at the resolution of the dynein-Lis1 map to discriminate between physical proximity to the AAA+ ring and a specific interaction. However, the fact that the linker is flexible in its position relative to the ring (Figure 1–figure supplement 1) we think argues against a tight interaction; in that case, we would have expected to be able to resolve the linker at AAA4 without having to use 3D sorting.

Despite this circumstantial evidence, we wanted to test a possible linker-AAA4 interaction directly. We mutated 5 residues on AAA4 that appear, based on our map, to be good candidates for a contact with the linker (new figure added; Figure 1–figure supplement 2). While this mutant (where all 5 residues were changed to alanine) showed a statistically significant difference in the extent to which Lis1 reduced its velocity relative to wild type, the difference was small and the AAA4 mutant was still regulated by Lis1. Taken together, the EM analysis and mutagenesis data argue strongly against a specific interaction between the linker and AAA4.

19) The authors try to draw the parallel between mutations in AAA5 and the effect of Lis1. The reviewer's reading of the data is that these two effects are different as the first abolishes ATPase activity, while the second has no effect on it.

While it is true that there is a difference in the ATPase activity between the two effects, there are also two similarities; a decrease in dynein velocity and a persistent attachment to the microtubule. We therefore feel that the comparison is useful in terms of trying to understand how Lis1 brings about a slow-moving, microtubule-attached dynein. However, we do agree that Lis1’s effect on the ATPase rate is still very much an open question, perhaps involving a secondary mechanism of allostery through the AAA+ ring. We have added new text to the Discussion to address this unresolved question.

20) One section states that the lack of conservation in the very N-terminus of the linker suggests it does not interact directly with Lis1 (which seems reasonable) and therefore that the Lis1 is acting as a steric block. The contact between the linker and the AAA+ ring in the Lis1:APO-Dynein structure appears to involve a more conserved part of the ring. This comes back to the question about whether Lis1 is acting on the AAA+ ring to allow the linker to bind to a place it normally would not.

We have addressed this in point 18 above.

21) Figure 2c. The blue color is difficult to distinguish from the black in the gel filtration trace.

We thank the reviewers for pointing this out. We have made the purple and green traces lighter to set them apart from the black more clearly.

22) Figure 3f/g. This EM map looks as though the contacts between the AAA+ ring and the Lis1 are different from in the Apo form. This may just be the different resolution of the structure, but it might be useful to discuss this somewhere.

It is likely that the AAA+ ring undergoes structural rearrangements in the ATP.Vi state that could change the position of Lis1’s binding site. However, as we pointed out, the resolutions of the two structures (apo vs. ATP.Vi) are different; since the ATP.Vi map is the lowest resolution structure we present we do not feel confident about commenting on possible differences in the AAA+ ring and/or contacts with Lis1.

23) Figure 4. It would be helpful if the type of mutation made was specified on this figure. b & c are E-Q mutations (residue number?). d) AAA5 mutant is not in the ATP binding site, but on the linker binding site. Specifying this on the figure (& res numbers?) would be helpful.

We thank the reviewers for the suggestion. All mutations and residue numbers were originally given in the figure legend because we felt that adding them directly to the figure itself would clutter it, particularly since the AAA5 linker-docking mutant is a triple mutation. In the current version we have modified the headings for each plot to make the mutation type clear without making reading the figure legend necessary.

24) Figure 5 and Figure 5–figure supplement 1. These figures show release from microtubules is blocked by Lis1, but not if the linker is shortened. The shortening does not affect the motors activity or ability to bind Lis1. Did the authors test if the velocity of the shorter linker was affected by Lis1?

It is not possible to test the effect of Lis1 on velocity in the context of a monomer gliding assay because it is known that the short linker dynein monomer moves slowly in gliding assays ([Reck-Peterson et al., 2006], Figure 3c), but moves with wild type speeds as a dimer in single-molecule assays ([Reck-Peterson et al., 2006], Figure 3b). The slow gliding by the short linker monomer is likely due to motor-glass interactions. Examining how Lis1 affects the velocity of dimeric short linker dynein is also problematic; once the short linker dynein is dimerized with GST its linker is no longer “short” (as it is extended by GST) and its ability to bypass Lis1 is now blocked by the dimerization.

25) Figure 5 d ii: The cartoon uses the stop sign, which suggests it has a mutation in AAA5 (as in Figure 4). Is this true? If not then another symbol might be more appropriate.

Thank you for pointing out the potential confusion. We have now moved all references to the linker-AAA5 interaction to the Discussion and new model figure (Figure 6). Therefore the stop sign in this figure has been removed.

26) Table 3:

The units of Km, kbasal and kcat should be given.

The units have been added.

One of the reviewers has the following thought which they would like to share with you:

Another mechanism that could be mentioned in the Discussion is that Lis1 has an allosteric effect on the AAA+ ring that allows the linker to move and the AAA1 site to exchange nucleotide without triggering the conformational change in the stalk that releases dynein from microtubules. This model is also not completely satisfactory as it is harder to explain the requirement for the N-terminus of the linker.

Perhaps one (explanation would be that the transition from high to low affinity is most likely to occur before the linker moves from a post to pre-powerstroke position. That is, ATP binds to a high affinity, post-powerstroke dynein. A rearrangement occurs that leads to a low affinity, post-powerstroke dynein (that releases from MTs). This is followed by a transition to the ADP.Vi like state with a low affinity, pre-powerstroke conformation. The ATP bound, post-powerstroke, low affinity conformation could be altered so that it no longer releases from microtubules by a combination of Lis1 AND linker binding to the AAA+ ring.

We agree that dynein is normally likely to detach from the microtubule before the linker moves to AAA2 (there is kinetic evidence that this is the case in Dictyostelium dynein [Imamula et al., 2007]), and that the transition from the high-to-low microtubule affinity (before the linker has moved) is the most-likely transition to be regulated by Lis1. However, our short linker experiments do suggest that steric regulation via the linker is critical for this regulation (i.e. it cannot be purely allosteric).