Structure of the Microtubule-Binding Domain of Flagellar ... - Cell Press

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Article Structure of the Microtubule-Binding Domain of Flagellar Dynein Yusuke S. Kato,1,2,3,* Toshiki Yagi,4,5,6 Sarah A. Harris,7 Shin-ya Ohki,8 Kei Yura,9,10 Youske´ Shimizu,2 Shinya Honda,3 Ritsu Kamiya,4,11 Stan A. Burgess,1 and Masaru Tanokura2,* 1Astbury

Centre for Structural Molecular Biology, Institute of Molecular and Cellular Biology, University of Leeds, Leeds LS2 9JT, UK of Applied Biological Chemistry, University of Tokyo, Bunkyo-ku, Tokyo 113-8657, Japan 3Biomedical Research Institute, National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba 305-8566, Japan 4Department of Biological Sciences, University of Tokyo, Bunkyo-ku, Tokyo 113-0033, Japan 5Structural Biology, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan 6Department of Cell Biology and Anatomy, University of Tokyo, Bunkyo-ku, Tokyo 113-0033, Japan 7School of Physics and Astronomy, University of Leeds, Leeds LS2 9JT, UK 8Center for Nano Materials and Technology (CNMT), Japan Advanced Institute of Science and Technology (JAIST), Nomi 923-1292, Japan 9Graduate School of Humanities and Sciences, Ochanomizu University, Bunkyo-ku, Tokyo 112-8610, Japan 10National Institute of Genetics, 1111 Yata, Mishima, Shizuoka 411-8540, Japan 11CREST, Japan Science and Technology Agency, Kawaguchi 332-0012, Japan *Correspondence: [email protected] (Y.S.K.), [email protected] (M.T.) http://dx.doi.org/10.1016/j.str.2014.08.021 2Department

SUMMARY

Flagellar dyneins are essential microtubule motors in eukaryotes, as they drive the beating motions of cilia and flagella. Unlike myosin and kinesin motors, the track binding mechanism of dyneins and the regulation between the strong and weak binding states remain obscure. Here we report the solution structure of the microtubule-binding domain of flagellar dynein-c/DHC9 (dynein-c MTBD). The structure reveals a similar overall helix-rich fold to that of the MTBD of cytoplasmic dynein (cytoplasmic MTBD), but dynein-c MTBD has an additional flap, consisting of an antiparallel b sheet. The flap is positively charged and highly flexible. Despite the structural similarity to cytoplasmic MTBD, dynein-c MTBD shows only a small change in the microtubule-binding affinity depending on the registry change of coiled coil-sliding, whereby lacks the apparent strong binding state. The surface charge distribution of dynein-c MTBD also differs from that of cytoplasmic MTBD, which suggests a difference in the microtubule-binding mechanism.

INTRODUCTION Dyneins are minus-end directed microtubule motors whose activities govern diverse biological events in eukaryotes. In cilia and flagella, ensembles of distinct dynein motors drive the generation and propagation of bends along the entire length of the axoneme (Kamiya, 2002). In the cytoplasm, single dynein dimers, stepping processively along microtubules, traffic various types of cargoes, including vesicles, proteins, RNA, and viruses (Hirokawa, 1998; Levy and Holzbaur, 2006). The widespread functions of dynein mean that its dysfunction is associated with diverse medical conditions, such as the brain disorders lissence-

phaly and hydrocephaly, neurodegenerative diseases including motor neuron disease, and ciliopathies causing mirrored internal organ localization and male infertility (Fliegauf et al., 2007; Levy and Holzbaur, 2006). Cytoskeletal motors produce directed movement by converting chemical energy from ATP hydrolysis into mechanical force (Vale and Milligan, 2000). Cycles of ATP hydrolysis and track attachment and detachment must be coordinated to produce this movement. Communication between the sites of ATP hydrolysis and track binding is known to be bidirectional: the nucleotide content of the active site alters the affinity of track binding, while track binding in turn alters the rate of hydrolysis within the active site. The actin-based motor myosin and the microtubule-based motor kinesin share a common evolutionary ancestry within the G protein superfamily (Kull et al., 1998). A structural comparison between myosin and kinesin shows that they share a common core fold, and that the sites of track binding and ATP hydrolysis are close together within a single protein domain, 30 A˚ apart. By contrast, dynein is a member of the AAA+ superfamily (Neuwald et al., 1999), a protein family with common ATPase activity and hexameric ring structure. In addition, dynein’s track binding and ATP hydrolysis sites have been shown to be separated by as much as 250 A˚, with the microtubule-binding domain (MTBD) at the distal end of a 100 A˚ long, antiparallel coiled coil structure called the stalk (Burgess et al., 2003; Kon et al., 2012; Roberts et al., 2009). An antiparallel coiled coil is composed of two a helices that go in opposite directions. These differences between dynein and the other cytoskeletal motors suggest that the mechanism underlying dynein’s activity is fundamentally different and pose interesting questions about the mechanism of bidirectional control along the antiparallel coiled coil. How communication is achieved between ATP hydrolysis within the AAA+ domains at one end of the stalk and microtubule-binding at the other end remains obscure, although a model is emerging. The mechanism of communication along the stalk involves sliding of the two helices of the coiled coil relative to one another, changing their registry by half a heptad along their entire length (Gibbons et al., 2005). This communication model is

1628 Structure 22, 1628–1638, November 4, 2014 ª2014 Elsevier Ltd All rights reserved

Structure Structure of MTBD of Flagellar Dynein

Because of these functional differences between axonemal and cytoplasmic dyneins, microtubule-binding properties and underlying mechanism of their respective MTBDs may differ. To investigate this, we have solved the solution structure of the MTBD of flagellar dynein-c/DHC9 (dynein-c MTBD), the inner arm flagellar dynein from C. reinhardtii. We have also characterized some aspects of its microtubule-binding using a variety of techniques, and we compare these findings to those from the MTBD of cytoplasmic dynein1 (cytoplasmic MTBD) (Carter et al., 2008; Gibbons et al., 2005). RESULTS

Figure 1. Structure of the Microtubule-Binding Domain of Dynein-c (A) Superposition of the 20 best structures of dynein-c MTBD and the nomenclatures of secondary structure elements with rainbow colors from the N- to C-termini (blue to red). This figure is depicted with Pymol (Schro¨dinger, LLC., New York, NY). (B) Mean structure viewed from the opposite side.

supported by recent electron microscopy (EM) structure of the MTBD of cytoplasmic dynein (cytoplasmic MTBD) bound to the microtubule, in which an imposed change in the coiled coil registry induces a structural change within the MTBD from the weak binding to the strong binding conformation (Redwine et al., 2012). In this conformation, coiled coil helix 1 (CC1) and helix 1 (H1) of cytoplasmic MTBD is rearranged compared with the previously reported MTBD structure in the weak binding state. These EM studies have shown the first atomic model for the binding geometry of any dynein MTBD to a microtubule. Helixhelix crosslinking studies have also demonstrated that signaling by helix sliding occurs within the context of an otherwise fully functional cytoplasmic dynein motor (Kon et al., 2009) However, whether this is a general mechanism that applies to all dyneins, including the various isoforms found in the axonemes of cilia and flagella, remains unknown. Functional differences between cytoplasmic and axonemal dyneins imply that their respective stalks operate in very different contexts. Cytoplasmic dynein performs long-range processive stepping using two identical motor domains. To prevent complete dissociation of the complex, one of the motors must remain tightly bound to the track while the other moves forward. Axonemal dyneins, by contrast, operate in large ensembles (comprised of more than ten distinct heavy chain isoforms) within the confines of the microtubular array of the axoneme (Kamiya, 2002; Yagi et al., 2009). Since multiple distinct dynein isoforms anchored on the same cargo microtubule via their tails operate on the same track, it is expected that they must be responsive to the activity of one another within the ensemble. During oscillatory beating of cilia and flagella, dynein ensembles undergo switching between phases of actively driven microtubule translocation and passive reverse-direction sliding of the microtubule (Lindemann, 2007).

Structure Determination Cloning dynein-c MTBD was carried out using a Chlamydomonas cDNA library (Yagi et al., 2005). Dynein-c MTBD was expressed in soluble fractions using an E. coli expression system. The length of the dynein-c MTBD construct was designed referring to those of previous studies for cytoplasmic MTBD (Mizuno et al., 2004; Shimizu et al., 2008) (Figure S1 available online). Our dynein-c MTBD construct, A2791–A2942, has an intermediate coiled coil sequence between the long sequence of cytoplasmic MTBD used for crystal structure analysis (Carter et al., 2008) and the short sequence of cytoplasmic MTBD used by Shimizu et al. (2008). The MTBD protein by Shimizu et al. (2008) binds microtubules tightly and was sufficiently soluble to provide relatively good nuclear magnetic resonance (NMR) spectra, but the protein stability was not ideal for long time NMR measurements. The dynein-c MTBD protein in the present studies is highly stable in aqueous solution and showed excellent quality of 2D NMR spectra, enabling us to determine its structure using 3D tripleresonance NMR (Figure 1 and Movie S1). The structure shows that the fold of dynein-c MTBD is globular and includes a short region of the stalk’s distal coiled coil helices (CC1 and CC2). Structural statistics are summarized in Table 1. Dynein-c MTBD consists of eight a helices and has similar overall structure to that of cytoplasmic MTBD in the weak binding state (Figures 1 and 2). The N- and C-terminal helices, CC1 and CC2, together form an antiparallel coiled coil. CC2 is located in the interior of the molecule, directly interacting with all of the helices except H3. By contrast, CC1 is much more solventexposed and interacts only with H4 and CC2, suggesting possible mobility of this helix. A continuous region of H4-L7H5-L8-H6 fringes CC2. A region of L7-H5-L8 shows an irregular structure: H5 is more extended and curved than typical a helices. Notable in the new structure of dynein-c MTBD is the existence of a protrusion called the flap. The flap is located between H2 and H3, consists of two b strands (S1 and S2) and an intervening loop (L4), and corresponds to the 13 amino acid insert (V2846–Y2858). Several other axonemal dynein isoforms possess an insert of similar size at the same location in their amino acid sequences, but this insert is absent in cytoplasmic dyneins (Figures 3 and S1), suggesting that the flap has a unique functional role in flagellar beating (discussed later). The flap presumably has high mobility because it does not contact other structural elements of the MTBD fold and may play a crucial role when dynein-c binds to the microtubule. 1H-15N heteronuclear NOE experiments (Figures 4A and 4C) and molecular dynamics (MD) simulations (Figures 4B and 4D) support the idea that this region is flexible.

Structure 22, 1628–1638, November 4, 2014 ª2014 Elsevier Ltd All rights reserved 1629


Table 1. Structural Statistics of Dynein-c MTBD NMR Distance and Dihedral Constraints Total restraint used

2,646

Total NOE

2,421

Intraresidue

663

Interresidue

1,758

Sequential (ji jj = 1)

687

Medium-range (1 < ji jj < 6)

703

Long-range (ji jj R 6)

368

Total dihedral angle restraints (F,J)

225

Structure statistics Distance restraint violations Rms (A˚)

0.0093 ± 0.0012

Maximum (A˚)

0.37 ± 0.10

Dihedral angle restraint violations Rms ( )

0.6372 ± 0.0562

Maximum ( )

5.05 ± 0.83

van der Waals distance restraints violations Sum (A˚)

11.7 ± 0.6

Maximum (A˚)

0.35 ± 0.01

CYANA-2.0 target function (A˚2) Average Pairwise Rmsda (A˚)

3.17 ± 0.20

Heavy atoms

1.45 ± 0.11

Backbone

0.75 ± 0.15

Ramachandran Plot (20 Ensembles) Most favored

88.1% (2,414)

Additionally allowed

11.7% (321)

Generally allowed

0.2% (5)

Disallowed

0.0% (0)

a

Pairwise rmsd was calculated among 20 refined structures (Asp2792Gly2840 and Ser2862-Val2932).

Principal-component analysis (PCA), performed on two independent MD simulations of dynein-c MTBD, shows that motion of the flap dominates the dynamics of the system (Movie S2). This is also supported by calculations of the root-mean-square deviation (rmsd) of all residues in the structure (Figures 4B and 4D). The most dominant motion mode of the flap (shown in Movie S2) is almost independent of the rest of the MTBD fold. This mode shows a swinging motion of the flap relative to the MTBD fold. The relative arrangement of H1 and H3, proposed to form the interface with the microtubule (Carter et al., 2008), differs between the dynein-c and cytoplasmic MTBD structures (Figure 2C). This difference may be a consequence of the mobility of H1 within the MTBD, as shown by the recent EM studies (Redwine et al., 2012). The starting pair of the heptad repeat in CC1 is L2807 and in CC2 is G2914 (Figure 2D). The coiled coil registry of dynein-c MTBD is therefore the same as that of the crystal structure of cytoplasmic MTBD, whose coiled coil has the socalled +b-registry (Figure 2C). A slight kink in the helix of CC1 of dynein-c MTBD coincides with a completely conserved proline (P2798, Figures 2A, 2B, and S1). Cytoplasmic MTBD has a similar kink at this position and a second one in CC2 correspond-

Figure 2. The Microtubule-Binding Domains of Dynein-c and Cytoplasmic Dynein (A) Dynein-c MTBD from C. reinhardtii. Conserved prolines in the distal coiled coil are illustrated as spheres (red and blue). (B) Cytoplasmic MTBD from mouse (Carter et al., 2008). (C) Superposition of dynein-c MTBD (pink) and mouse cytoplasmic MTBD (blue). (D) Details of the distal coiled coil of dynein-c MTBD. G2914 and L2807 are the distal end pair of the stalk coiled coil. L2800 and L2807 in CC1 (blue) are at the ‘‘d’’ positions in the heptad repeat. A2806 and L2799 are at the ‘‘e’’ positions. A2921 and G2914 in CC2 (red) are at the ‘‘a’’ positions.

ing to another completely conserved proline (P2934 in the dynein-c sequence) that lies outside the converged region in our structure of dynein-c MTBD. These prolines in cytoplasmic MTBD occur close together and bend the path of the whole coiled coil near its junction with the helix-rich terminal domain. This bend in the coiled coil has also been reported in EM images of intact molecules of both cytoplasmic dynein (Roberts et al., 2009) and flagellar dynein-c (Burgess et al., 2003) and is therefore likely to be a general feature of all dynein stalks.



Figure 3. Alignment of Microtubule-Binding Domains of various Dyneins Alignment of MTBD sequences from flagellar inner arm dynein-c from C. reinhardtii, flagellar outer arm dynein-b from sea urchin (Anthocidaris crassispina) and cytoplasmic dynein from mouse. Residue numbers of dynein-c are shown. Heptad repeats (‘‘d’’ positions in CC1 and ‘‘a’’ positions in CC2) are indicated in pink (located in the dynein-c MTBD structure) and blue bands (hypothetical). Blue and red bars define respectively, the CC1 and CC2 numbers that represent the dynein-c MTBD boundaries of each construct incorporated into an SRS-MTBD chimera. Proline residues (green asterisks) are the starting points for the nomenclatures of these numbers. For example, SRS-MTBD19:9 has 19 residues in CC1 and nine in CC2 counted from these prolines.

The structure of the coiled coil region (CC1 and CC2) is different from those of parallel coiled coils, in which the positions ‘‘a’’ and ‘‘d’’ in the heptad repeat face the ‘‘a’’ and ‘‘d’’ residues in the counterpart helix, respectively. A parallel coiled coil is often composed of a homo-dimer, in which the coiled coil structure is symmetrical. In contrast, the coiled coil of dynein-c MTBD is asymmetric, in which the ‘‘a’’ and ‘‘d’’ positions face respectively the ‘‘d’’ and ‘‘a’’ positions in the counterpart helix (Hadley and Gellman, 2006). The residues in ‘‘a’’ and ‘‘d’’ positions are important in providing the knobs-into-holes interaction between the two helices and are usually occupied by large aliphatic residues to stabilize the coiled coil structure (Hadley et al., 2008). The ‘‘d’’ positions on CC2 of dynein-c MTBD are, however, occupied by small residues (G2914 and A2921; Figure 2D). A2921 is conserved among most flagellar and cytoplasmic dyneins, whereas G2914 is conserved only in some flagellar dyneins. These suggest possible mobility of CC1 relative to the rest of the MTBD fold, as shown in cytoplasmic MTBD (Redwine et al., 2012). Helix Sliding and Microtubule-Binding Affinity While our NMR structural studies have shown that there is a high overall similarity between the structures of cytoplasmic and dynein-c MTBDs, the flap of dynein-c is a novel feature. This raises the question of the influence of the flap on microtubule-binding. In cytoplasmic dynein, a change in registry between the two helices of the coiled coil has been shown to alter the microtubule-binding affinity of the MTBD and reciprocally the ATPase activity within the AAA+ ring (Kon et al., 2009). Thus, helix sliding has been proposed as the mechanism that controls microtubule

attachment and detachment during stepping of this processive motor. Earlier work had demonstrated the influence of helix registration on microtubule-binding affinity using a series of chimera proteins in which the distal stalk of cytoplasmic dynein, including regions both proximal and distal to the proline pair described above, was fused to the antiparallel coiled coil region of seryl tRNA synthetase (SRS) (Gibbons et al., 2005). In that work, various chimeras were created by differential splicing of dynein’s antiparallel coiled coil helices onto those of SRS, thereby imposing different registrations between the two helices of dynein’s coiled coil, and the proteins were assayed for microtubule-binding affinity. Two characteristic binding states were found: a high-affinity state in the so-called a-registry, and a weaker affinity state in the +b-registry (Gibbons et al., 2005). To test whether a similar helix sliding mechanism occurs in the stalk of our flagellar dynein-c, we generated a similar series of chimera proteins of the dynein-c stalk region and SRS (SRSMTBD) to impose different coiled coil registries on the stalk (Figures S2A–S2E) and estimated their apparent binding affinities. The highest microtubule affinity (apparent KD = 30 mM) in SRS-MTBD chimeras of flagellar dynein-c was observed with helices in the a-registry (Figures 5A, 5B, S2F, and S2G). A similar apparent microtubule-binding affinity was also found in the a+1registry, in which CC1 is shifted relative to the a-registry by one whole heptad (i.e., seven amino acid residues) into the MTBD. This is a 20-fold lower apparent affinity compared with that of SRS-MTBD of cytoplasmic dynein in the same registry (KD = 1 3 mM) and in similar buffer conditions (Gibbons et al., 2005). This is a remarkable difference between dynein-c MTBD and the cytoplasmic counterpart. All other registries tested, including b-registries, gave much lower apparent binding affinities to microtubules. The difference in apparent KD values between the a- and b-registries was 2-fold in the case of dynein-c MTBD. This is much smaller than that found in cytoplasmic MTBD, whose a- and b-registries have at least 10-fold difference in KD values. Free dynein-c MTBD, without the SRS tag, also shows weak binding (apparent KD value = 40 mM), similar to those of SRSMTBD chimeras, including in buffers used for NMR measurements for structure determination (Figures S2H and S2I). This suggests that free dynein-c MTBD also binds to microtubules much more weakly than the corresponding free MTBD of cytoplasmic dyneins (KD = 110 mM) (Mizuno et al., 2004; Shimizu et al., 2008), and that the microtubule is incapable of inducing dynein-c MTBD to adopt tight binding (KD = 13 mM), even when the helices of the stalk are presumably free to shift their registry. Finally, we tested the binding properties of full-length dyneinc/DHC9 molecules from C. reinhardtii. Tight binding can be demonstrated by copelleting of dynein molecules with microtubules upon centrifugation. The dynein-c/DHC9 fraction that we used also contained dynein-b/DHC5, and only a very small amount of both dynein isoforms copelleted with microtubules in all the experimental conditions tested (Figure 5C). This indicates that both dynein-c and dynein-b isoforms showed weak microtubule-binding, even in the absence of nucleotide, which typically produces tight binding by the other cytoskeletal motors including cytoplasmic dynein. Both dynein-c and dynein-b are flagellar inner arm dyneins (IADs) categorized into the IAD-3



Figure 4. Dynamic Analysis of the Microtubule-Binding Domain of Dynein-c (A) Plot showing the result of 1H-15N heteronuclear NOE experiments. Vertical axis shows 1H-15N heteronuclear NOE values, which approximately correspond to the structural stability of backbone amides in dynein-c MTBD. Together with (B), the horizontal axes show the residue numbers in the MTBD. Residues with high flexibility show low NOE values. Error bars indicate the SD. (B) Plot of rmsd values from all trajectories of MD simulations. We carried out two sets of MD simulations, 45 ns each, using two structures out of the ensemble of 20 NMR structures. Red and blue lines plot rmsd values of these two sets of trajectories. Both of the plots show the similarity of the dynamic behaviors of the two individual simulations. High rmsd regions correspond to flexible portions in the simulation. The region of the flap (P2843–Y2858) shows high rmsd values corresponding to the low NOE region in (A). (C) Mapping the result of 1H-15N heteronuclear NOE experiments shown in (A). Residues shaded orange corresponds to NOE < 0.7 and yellow to 0.7 < NOE < 0.75. (D) Mapping of one of the rmsd plots shown in (B) on the structure of dynein-c MTBD.

Taken together, our results show that the binding of dynein-c to microtubules at its highest affinity is weaker than that of cytoplasmic dynein under the conventional conditions used for testing microtubule-binding. However, there is still the possibility that a strong binding state of dynein-c exists in vitro. First, not all possible registries of the SRS-MTBD chimeras have been assessed, as this requires a systematic examination of the effect of changing the lengths of the coiled coil helices by one amino acid residue at a time. Moreover, we did not check the binding of full-length dynein-c in all possible nucleotide states. Nevertheless, our results suggest that there are marked differences in the microtubule-binding properties of dynein-c and cytoplasmic dynein.

subfamily (Wickstead and Gull, 2007). Both possess the sequences corresponding to the flap in the MTBD (Figure S1). By contrast, dynein-fa/DHC1 molecules, an inner arm isoform categorized into the IAD-1A subfamily that lacks the flap (see Cr_IAD1A in Figure S1 for sequence information), shows strong binding to microtubules in the absence of nucleotide.

Microtubule-Binding Mode Our observation that the MTBD of dyneinc/DHC9 only exhibits the weak binding mode under conventional experimental conditions, in spite of its structural similarity to the cytoplasmic MTBD, remains puzzling. A binding geometry for cytoplasmic MTBD on the microtubule lattice has been proposed in the recent EM studies (Redwine et al., 2012), in which H1 and H3 of cytoplasmic MTBD are placed on the surface region of the microtubule, similar to that for kinesin-binding. To examine the microtubule-binding surface of dynein-c MTBD, we made use of a bioinformatics



Figure 6. Prediction of Binding Surface of the Microtubule-Binding Domain of Dynein-c to Microtubule from KYG Analysis Hot colors indicate a high propensity for microtubule-binding. Buried residues are colored deep blue.

Figure 5. Microtubule-Binding Assays of SRS-MTBD Chimeras and Full-Length Dyneins (A) Microtubule-binding assays of SRS-MTBD chimeras of dynein-c with imposed registries of the coiled coil. There are four different chimeras with the same CC1 length, but different registries that were centrifuged with and without microtubules (upper and lower panels, respectively). The paired number nomenclature for constructs (e.g., 19:9) is the same as that used in previous studies (Carter et al., 2008; Gibbons et al., 2005) and denotes the number of residues between the SRS splice site and the proline marking the stalk-MTBD boundary for CC1 and CC2, respectively. (B) Summary of apparent microtubule affinity of the SRS-MTBD chimeras with different lengths of CC1 and CC2. Dashed diagonal lines show the ‘‘+4’’ (a-registry), ‘‘+8’’ (+b-registry), and ‘‘+11’’ (a+1-registry) constructs. (C) Axonemal dyneins from the oda1 mutant of C. reinhardtii (i.e., lacking outer arm dyneins) were copelleted in the presence and absence of microtubules and ATP. The bands of inner arm dynein species -b and -c overlap as expected (Kagami and Kamiya, 1992), and both show little copelleting under any condition tested here, whereas most of the dynein-fa copellets under the so-called rigor condition (+ microtubule and ATP).

approach called KYG method (Kim et al., 2006; Yura, 2008), which predicts protein-protein interfaces based on amino acid sequence and protein structure. This method predicted clusters of residues on helices H2, H5, and at the tip of the flap should contact the microtubule (Figure 6). A comparison of the surface charge distributions of cytoplasmic and dynein-c MTBDs also

reveals striking differences (Figure 7). The surface area composed of H3, L2, L3, S1, and S2 is positively charged in dynein-c MTBD, whereas that composed of H1, H3, and the loop corresponding to L2 of dynein-c is positively charged in cytoplasmic MTBD. This difference is partly due to the substitutions on H1, H3, and H6 to acidic residues in dynein-c (D2814, E2860, and E2906) (Figure S1) and may cause the weaker affinity of dynein-c MTBD in the a-registry by reducing the electrostatic attraction to microtubules. K3298, K3299, R3306, R3337, R3342, and R3382 of cytoplasmic MTBD are all involved in salt bridges to microtubules in the a-registry (black pentagrams in Figure S1) (Redwine et al., 2012). Some of the corresponding residues in dynein-c, K2811, K2819, K2863, and K2907, are conserved, but the substitutions in others, such as P2812 and E2868, are unfavorable for salt bridge formation. Therefore, it is not clear whether the dynein-c MTBD should adopt a similar binding geometry to that of cytoplasmic MTBD when in the a-registry. Moreover, there is no experimental information concerning the binding geometry of MTBD of any dynein in the weak-binding state including the +b-registry and other nonspecific binding modes. An additional consideration is the potential for steric clashes between the flap and the microtubule surface. When the solution structure of dynein-c MTBD is docked on a microtubule in the binding geometry of cytoplasmic MTBD proposed by Redwine et al. (2012), the average solution NMR structure of dynein-c MTBD exhibits serious overlap with the electron



Figure 7. Surface Charge Distribution Ribbon diagram and surface representation with calculated electrostatic charges of the MTBDs of dynein-c/DHC9 (left) and cytoplasmic dynein1 (right). Blue and red in surface representation correspond to positive and negative charges. This figure is depicted with Swiss PDB Viewer (Guex and Peitsch, 1997).



density of the protofilament adjacent to that with the docked dynein-c MTBD (Figure S3A). MD and PCA analyses have both shown, however, that the flap is highly mobile, which may well enable these clashes to be avoided through bending (Figure S3C). In contrast, an extended flap conformation results in clashes with H9 and H10 of b-tubulin in the adjacent protofilament (Figure S3B). DISCUSSION Here we have solved the structure of the dynein-c MTBD, an inner arm subspecies from Chlamydomonas reinhardtii flagella. The structure of dynein-c MTBD comprises the most distal part of dynein’s MT-binding stalk domain and includes portions of the a helices that form the antiparallel coiled coil of the stalk. Dynein-c MTBD shares the same fold as cytoplasmic MTBD from mice (Carter et al., 2008), but with a notable difference: the flap composed of 13 amino acid residues. The insert corresponding to the flap is seen only in the amino acid sequence of some axonemal dynein isoforms, but is absent in those of cytoplasmic dyneins (Figure S1). NMR data and MD simulations both suggest that the flap is highly mobile with respect to the rest of the helix-rich MTBD. The results of PCA performed on the MD trajectory show the most dominant motion of the flap (Movie S2). In spite of the high similarity in the overall structure between dynein-c MTBD and cytoplasmic MTBD, the microtubule-binding properties are remarkably different. Dynein-c MTBD binds to microtubules weakly in the a-registry state, in which cytoplasmic MTBD binds to microtubules strongly. The registry change in the coiled coil of dynein-c, therefore, has a smaller impact on the change in the microtubule-binding affinity of its MTBD than that in cytoplasmic dynein. Such characteristics of the coiled coil and MTBD of dynein-c could technically be attributed to three factors; namely, the presence of the flap, surface charge distribution of the MTBD, and the communication mechanism between the coiled coil and the MTBD. If it is assumed that the binding geometry of dynein-c MTBD to a microtubule is similar to that of cytoplasmic MTBD, then steric clashes with the flap of dynein-c MTBD could well have a negative impact on microtubule-binding. However, the binding geometry of dynein-c MTBD remains unknown. Several surface residues on dynein-c MTBD, including those important for microtubule-binding, are replaced with negative charges compared with the cytoplasmic counterpart (Figure S1). These substitutions cumulatively result in striking differences between the surface charge distribution in dynein-c and cytoplasmic MTBDs, which would be expected to reduce the binding affinity to microtubules, since microtubules contain negatively charged surface patches. In addition, the registry change at the coiled coil may have a smaller impact on the conformational change of dynein-c MTBD than that of cytoplasmic MTBD. Thus, these three factors taken together would be sufficient to produce the different microtubule-binding characteristics of dynein-c MTBD in the a-registry compared with the cytoplasmic MTBD. Some of the microtubule-binding residues for the tight binding, however, remain conserved between dynein-c and cytoplasmic MTBDs; so further scrutiny is necessary to establish whether dynein-c really lacks the strong binding state.

Flagellar dynein can diffuse up to 170 nm along the longitudinal axis of a microtubule (Vale et al., 1989), whereas no such diffusive properties have been reported for cytoplasmic dyneins. Microtubule-binding proteins have often had positively charged flexible regions, which have been proposed to enhance their 1D diffusion on the microtubule lattice (Cooper and Wordeman, 2009). The flap of dynein-c is also flexible and abundant in positive charges, so the flap may contribute to the 1D diffusion of flagellar dyneins on the longitudinal axis of a microtubule through the interaction with the acidic residues on the surface of tubulins. Thus, the characteristic surface charge distribution and the existence of the flap may together contribute to the difference in the nonspecific binding mode of flagellar dynein compared with that of cytoplasmic dynein. Oscillatory bending within cilia and flagella is driven by the activity of ensembles of multiple distinct dynein motors switching between phases of locally driven active sliding of microtubules and passive reverse sliding driven by dyneins elsewhere in the axoneme. Temporal and spatial coordination of this activity likely involves feedback from the bending state of the axoneme (Morita and Shingyoji, 2004), which may involve a tension-sensing mechanism via the dynein motors’ slender stalk domains (Lindemann, 2007). The various dynein isoforms within these ensembles display differences in their motor properties. For example, microtubule translocation velocities of inner arm species in vitro vary between 2 and 11 mm/s, and some species (including dynein-c) rotate microtubules as they translocate them in vitro (Kagami and Kamiya, 1992). Such diverse properties of various flagellar dyneins might be relevant to the presence or absence of the flap. Many flagellar isoforms including dynein-c/DHC9, dynein-b/DHC5, and dynein-d/DHC2 possess amino acid sequences corresponding to the flap, but others, including dynein-fa/DHC1, do not. IADs such as dynein-c, -d, and -fa, are categorized into the IAD-3, -4, and -1a subfamilies, respectively, according to the classification by Wickstead and Gull (2007) (Figure S1). As shown in Figure 5C, dynein-fa binds to microtubules tightly, so the lack of the flap might provide an advantage for microtubule-binding. We have made homology models of MTBDs of dynein-d and dynein-fa to compare the surface charges distribution among dyneins categorized into different subfamilies (Figure S4) (Guex and Peitsch, 1997). The surface charge distribution of the homology model of the MTBD of dynein-fa is similar to that of cytoplasmic MTBD, even though dynein-fa and cytoplasmic dynein are categorized into different subfamilies. The MTBDs of dynein-fa and cytoplasmic dynein contain numerous positive charges around H1, H3, and the loop corresponding to L2 of dynein-c (Figures 7 and S4). H1 and H3 are the microtubule-binding interface of cytoplasmic MTBD (Redwine et al., 2012). The surface of a microtubule is rich in negative charges, so it is likely that the positive charges on H1 and H3 of dynein contribute toward strong microtubule-binding in these isoforms. By contrast, the positive charges around H1 and H3 of dynein-c are fewer than those of cytoplasmic dynein and dynein-fa (Figure 7), and there is an observed lack of strong microtubule-binding in this dynein. The homology model of the MTBD of dynein-d/DHC2, belonging to a different IAD subfamily from both dynein-c and dynein-fa, shows even fewer positive charges around H1 and H3 than those of the rest of the dyneins, indicating diversity in the surface charge



distribution of MTBDs. Taken together, these findings suggest that the presence or absence of the flap, and differences in the surface charge distribution of MTBDs, might contribute to the observed diversity in their motor properties. How multiple distinct dynein motors cooperate within the context of an individual axonemal ensemble remains mysterious (Kamiya, 2002). Inner arm dynein-c has recently been shown to accelerate the microtubule translocation velocity in response to an assisting external force, but not to a force resisting the direction of translocation, implying that it would not retard the activity of faster dyneins (Kikushima and Kamiya, 2009). The lack of the strong binding state is presumably advantageous for such cooperative property of dynein-c, if there is too strong an affinity between the motor and the track then this would provide too much resistance to external forces, which would impede function. In addition, it is expected that the diffusible property of flagellar dyneins on microtubules may also contribute to their responses to external forces, and consequently their cooperation with other dynein molecules. The present studies have determined the structure of dynein-c MTBD using solution NMR. We have shown that there is a strong similarity in the overall structure of MTBDs of dynein-c and cytoplasmic dynein, but that there are important variations in the microtubule-binding properties between those MTBDs, which are presumably related to their contrasting functions. It is proposed that these differences may be due to three factors: the presence of the flap, differences in the surface charge distribution between those MTBDs, and differences in the consequences of changes in registry on the structures of the MTBDs. The impact of these factors on the binding affinities of many other dyneins should be further studied in the future. EXPERIMENTAL PROCEDURES Plasmid Construction Dynein-c MTBD was cloned from Chlamydomonas reinhardtii (Yagi et al., 2005). Dynein-c MTBD from Ala2791 to Ala2942 was cloned into pGEX-4T1 (GE Healthcare) with a tobacco etch virus (TEV) protease cleavage site instead of a thrombin site. The SRS-MTBD chimera constructs were produced by cloning various lengths of the stalk region of dynein-c (including the MTBD region) into the SalI-HindIII sites of the monomer SRS expression system (Gibbons et al., 2005). The sequences of the SRS-MTBD chimeras are shown in Figures S2C–S2E. Sample Preparations All protein expression was carried out with BL21(DE3) (Merck Millipore). Dynein-c MTBD fused with glutathione S-transferase expressed in a soluble fraction was purified with Glutathione Sepharose (GE Healthcare) following manufacturer’s instructions. After cleavage by AcTEV protease (Life Technologies), denatured dynein-c MTBD was refolded in a gel-filtration column by exchanging solvents. The denaturing/refolding procedure was necessary to improve the quality of NMR spectra. The sample without this refolding treatment showed numerous minor signals that made further analysis difficult. Dynein-c MTBD was unfolded by adding 6 7 M guanidine hydrochloride, 0.5 M Tris-HCl pH 8.5, 0.5 M dithiothreitol (DTT), and 10 mM EDTA to the MTBD solution eluted from glutathione Sepharose. The unfolded sample was loaded onto a Superdex75 column (GE Healthcare) equilibrated with 10 mM Na-phosphate (pH 6.4) and 100 mM NaCl. 15N- and 13C15N-labeled dynein-c MTBD were produced following the M9 trace elements protocol with some modification (Cai et al., 1998). The SRS-MTBD chimera proteins were also produced using methods similar to those in a previous report (Gibbons et al., 2005) without refolding procedures. Tubulins were prepared from porcine brain as described previously (Borisy et al., 1975; Williams and Lee, 1982).

Nuclear Magnetic Resonance Measurements and Structure Determination The NMR spectra required for structure determination were acquired at 25 C (298 K) on either a Unity INOVA 600 or 750 spectrometer equipped with a Narolac z axis gradient probe or a Varian Cold Probe (Agilent Technologies). There was 0.5 mM dynein-c MTBD that was dissolved in 10 mM Na-phosphate buffer (pH 6.4), 50 mM arginine, 50 mM glutamate (Golovanov et al., 2004), 100 mM NaCl, 0.02% NaN3, 1 mM fully deuterated DTT, and 5100% 2H2O. Data acquisition, processing, analysis, and structural calculation were carried out using a method similar to that previously described (Kato et al., 2006; Herrmann et al., 2002). The mixing time of NOESY-related measurements was 100 ms. The torsion angle restraint was produced using the TALOS module in the NMRPipe suite of programs (Delaglio et al., 1995; Shen et al., 2009). The 1H-15N heteronuclear NOE experiment was carried out with a 4.0 s relaxation delay (Tate and Kainosho, 1994). Molecular Dynamics Simulations Simulations used the Duan 2003 forcefield (Duan et al., 2003) in conjunction with the AMBER9 suite of programs (Case et al., 2005). Restraint of bonds to hydrogen with SHAKE enabled a 2 femtosecond timestep to be used in the numerical integration of the equations of motion of the atoms in the biomolecule. Simulations were run at constant temperature (300 K) and pressure (1 atm). All protein structures were surrounded by 10 A˚ of TIP3P water and used periodic boundary conditions in conjunction with the fast particle-mesh Ewald summation method for accurate treatment of long-range electrostatic interactions. Simulations were run for 50 ns using 32 processors of the Leeds 404 processor Opteron-Myrinet Supercomputer, the last 45 ns were selected for data analysis. The MD trajectories were manipulated using the PTRAJ module available within AMBER9. To extract the major modes of motion, PCA was performed by diagonalization of the Cartesian coordinate covariance matrix on the MD trajectories (Meyer et al., 2006). The nature of the structural change associated with each eigenvector was determined by producing animations of the components (see Movie S2). Prediction of Microtubule-Binding Interface The protein interface prediction using the KYG method was performed by extending the method originally developed by Kim et al. (2006) for the prediction of RNA interfaces for RNA-binding proteins (Yura, 2008). The singlet propensity is the preference for a certain type of residue at the interface, and the doublet propensity is the preference for a residue pair at the interface. Singlet and doublet residue propensities in the protein-protein interface of approximately 1,000 biological complexes from the Protein Data Bank (Berman et al., 2007) were calculated to predict the binding surface of dynein-c MTBD. The prediction of protein interface residues using these propensities was evaluated by a Jackknife test and achieved approximately 60% specificity. Binding Assay by Copelleting The SRS-MTBD chimera proteins were dialyzed against 30 mM HEPES (pH 7.4), 5 mM MgSO4, 1 mM DTT, 1 mM EGTA, and 50 mM potassium acetate (HMDEK). Tubulins were polymerized with 10% DMSO, 1 mM Mg-GTP, 1 mM EGTA, and 20 mM taxol (Sigma-Aldrich) at 37 C. The buffer was replaced with HMDEK with 20 mM taxol before copelleting. Copelleting assays were carried out with 9 mM SRS-MTBD and 10.7 mM microtubules centrifuged at 100,000 3 g at 20 C for 30 min in HMDEK with 20 mM taxol. Density of bands in SDS-PAGE gels was analyzed with ImageJ to evaluate amounts of SRSMTBD in pellet and supernatant fractions. KD values were calculated from the following equation: KD = [free SRS-MTBD][free microtubule]/[complex]. The estimate of the KD value was determined based on the results from three time independent experiments for each construct. For each SRS-MTBD construct tested, determination of the KD value was based on the results from the copelleting experiments with a single SRS-MTBD concentration rather than fitting of the data from the copelleting experiments with multiple SRS-MTBD concentrations to a hyperbolic function. Thus, our estimated KD values may contain more errors than those from the hyperbolic fitting, which is why we refer to them as apparent KD values. Axonemal dyneins from the oda1 mutant (Kamiya, 1988) of C. reinhardtii, which lacks outer arm dyniens (dynein-a, -b, and -g), were purified following the method described previously



(Witman et al., 1978). The microtubule-binding assay was carried out under the same conditions as in the cases of SRS-MTBD with or without 4.0 mM ATP. Assignment of the band of dynein-c in SDS-PAGE was carried out previously (Kagami and Kamiya, 1992). ACCESSION NUMBERS The NMR chemical shifts for dynein-c MTBD have been deposited in the Biological Magnetic Resonance Bank, http://www.bmrb.wisc.edu (accession no. 11057). The atomic coordinates have been deposited in the Protein Data Bank (PDB), http://www.pdb.org (PDB ID code 2RR7). SUPPLEMENTAL INFORMATION Supplemental Information includes four figures and two movies and can be found with this article online at http://dx.doi.org/10.1016/j.str.2014.08.021. AUTHOR CONTRIBUTIONS Y.S.K., T.Y., S.A.H., R.K., S.A.B., and M.T. designed the study and wrote the manuscript. Y.S.K., T.Y., S.A.H., S.O., K.Y., and S.A.B. carried out the experiments, analyses and/or simulations. T.Y., S.A.H., K.Y., Y.S., S.H., R.K., S.A.B., and M.T. provided advices and supports for the experiments and strategy. ACKNOWLEDGMENTS We thank Drs. Peter J. Knight, Thomas A. Edwards, Anthony J. Roberts (University of Leeds), Keiko Hirose, and Taro Q. P. Uyeda (National Institute of Advanced Industrial Science and Technology [AIST] Japan) for their intense discussions and/or critical readings of the manuscript, Ian R. Gibbons and Joan E. Garbarino (University of California, Berkeley) for kindly providing the SRS-chimera expression systems, Masahide Kikkawa (University of Tokyo) and Ronald D. Vale (University of California, San Francisco) for kindly providing the 3D EM map and suggestions, Ken Downing (Lawrence Berkeley Laboratory) for kindly providing microtubule coordinates, and Jun-ichi Kurita (Agilent Technologies Japan, Ltd.), Takashi Shimizu, and Kenji Kanazawa (AIST Japan) for their dedicated technical support. Maintenance of the 750 MHz-NMR spectrometer used in this work has been supported by the Center for Nano Materials and Technology and the Japan Advanced Institute of Science and Technology. Y.S.K. was supported in part by the Biotechnology and Biological Sciences Research Council, United Kingdom (to S.A.B.). K.Y. was supported by Platform for Drug Discovery, Informatics, and Structural Life Science from the Ministry of Education, Culture, Sports, Science and Technology (MEXT) of Japan. This work was supported in part by the National Project on Protein Structural and Functional Analysis, Targeted Proteins Research Program, and Grant-in-Aid for Scientific Research, of MEXT of Japan. Received: June 1, 2014 Revised: August 27, 2014 Accepted: August 27, 2014 Published: October 23, 2014

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