# Myosin VI Must Dimerize and Deploy Its Unusual Lever Arm in Order to Perform Its Cellular Roles

Monalisa Mukherjea<fnref n="1" />,*, M. Yusuf Ali<fnref n="2" />,*, Carlos Kikuti<fnref n="3" />,<fnref n="4" />, Daniel Safer<fnref n="1" />, Zhaohui Yang<fnref n="1" />, Helena Sirkia<fnref n="3" />,<fnref n="4" />, Virginie Ropars<fnref n="3" />,<fnref n="4" />, Anne Houdusse<fnref n="3" />,<fnref n="4" />, David M. Warshaw<fnref n="2" />, and H. Lee Sweeney<fnref n="1" />,†

## Abstract

It is unclear if the reverse-direction myosin, myosin VI, functions as a monomer or dimer in cells and how it generates large movements on actin. We deleted a stable, single α-helix (SAH) domain that has been proposed to function as part of a lever arm to amplify movements, without impact on in vitro movement or in vivo functions. A myosin VI construct that used this SAH domain as part of its lever arm was able to take large steps *in vitro*, but did not rescue *in vivo* functions. It was necessary for myosin VI to internally dimerize, triggering unfolding of a three-helix bundle and calmodulin binding in order to step normally *in vitro* and rescue endocytosis and Golgi morphology in myosin VI-null fibroblasts. A model for myosin VI emerges in which cargo binding triggers dimerization and unfolds the three-helix bundle to create a lever arm essential for *in vivo* functions.

## Graphical Abstract

© 2014 Elsevier Inc. Published by Elsevier Inc. All rights reserved.

†To whom correspondence should be addressed: lsweeney@mail.med.upenn.edu.

*Authors contributed equally.

## SUPPLEMENTAL INFORMATION

Supplemental information includes Supplemental Experimental Procedures, two figures, and two tables and can be found with this article online at http//......

## AUTHOR CONTRIBUTIONS

M.M and M.Y.A. designed and performed experiments and contributed to writing the manuscript, C.K., D.S., Z.Y., H.S., and V.R. designed and performed experiments, and A.H., D.M.W., and H.L.S. designed experiments and wrote the manuscript.

The authors have no conflicts of interest to declare.

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<footnote n="1">¹Department of Physiology, University of Pennsylvania Perelman School of Medicine, 415 Curie Blvd, 700 CRB, Philadelphia, PA 19104-6085, USA</footnote>

<footnote n="2">²Department of Molecular Physiology and Biophysics, University of Vermont, Burlington, VT 05405, USA</footnote>

<footnote n="3">³Structural Motility, Institut Curie, centre de recherche, Paris, F-75248 France</footnote>

<footnote n="4">⁴CNRS, UMR144, 26 rue d'Ulm, 75248 Paris cedex 05, France</footnote>
<figure><img src="image_1.png" /></figure>

Keywords

unconventional myosin; processivity; calmodulin; single stable α-helix

INTRODUCTION

Myosin VI is the only actin-based motor among the known classes of molecular motors (Odronitz and Kollmar, 2007) that has been shown to move toward the minus-end of actin filaments (Wells et al., 1999). Myosin VI is ubiquitous and has been implicated in a variety of cellular functions (Buss et al., 2004; Frank et al., 2004; Sweeney and Houdusse, 2007; Chibalina et al., 2009). Biochemical and biophysical studies have demonstrated that a myosin VI dimer is capable of processive movement along an actin filament (Park et al., 2006) and load-dependent anchoring (Altman et al., 2004), and thus can fulfill a number of specialized cell biological functions, including endocytosis (Buss et al., 2001; Hasson, 2003; Dance et al., 2004; Inoue et al., 2008) and maintenance of Golgi morphology and function (Sahlender et al., 2005; Buss and Kendrick-Jones, 2008). A fundamental question and subject of considerable controversy is whether or not myosin VI functions as a monomer or dimer in any or all of its cellular functions (Rock et al., 2005; Lister et al., 2004; Park et al., 2006; Sweeney and Houdusse, 2010; Bond et al., 2012).

Myosin motors move along actin filaments driven by conformational changes in the motor domain associated with the sequential release of MgATP hydrolysis products, Pᵢ and MgADP. These conformational changes in the motor domain are amplified by the myosin lever arm, which is comprised of the C-terminal subdomain of the motor domain (known as the converter) and a helix of variable length, depending on the myosin class, to which calmodulin and/or calmodulin-like light chains bind to a series of IQ motifs. Though similar in organization as compared to other members of the myosin superfamily, myosin VI has a number of class-specific structural elements (Figure 1A). Most importantly, an insert
(known as insert 2) between the converter subdomain of the motor domain and the single IQ motif, binds an additional calmodulin and repositions the lever arm to reverse the directionality of myosin VI (Ménétrey et al., 2005, 2007). However, a lever arm comprised of the myosin VI converter, the insert 2 region with associated CaM, and the IQ motif with bound CaM would not allow a myosin VI dimer to take 30–36nm steps on actin, as has been observed (Rock et al., 2001, 2005; Yildiz et al., 2004; Park et al., 2006). Thus some region distal to the IQ motif must function as an extension of the myosin VI lever arm (Figure 1B, C, & D).

Two models have emerged that attribute the lever arm extension to totally different structures and propose very different regions of internal dimerization (Figure 1C & D), which must occur distal to the lever arm extension. The region following the IQ motif of approximately 80 residues forms a three-helix bundle in monomeric myosin VI (Figure 1B). Following the three-helix bundle of myosin VI, there is an additional domain that is predicted to form an extended, stable single α-helix (SAH) (Knight et al., 2005) (Figure 1). Other classes of myosins, including class VIIa, X, and *Dictyostelium MyoM* (Knight et al., 2005; Spink et al., 2008, Baboolal et al 2009) also contain a SAH domain distal to their respective IQ motifs. SAH domains have also been reported in smooth muscle caldesmon (Wang et al., 1991) and have been identified in more than 123 different proteins (Sivaramakrishnan et al., 2008). The predicted SAH domains of these myosins typically contain alternating clusters of negatively charged acidic and positively charged basic amino acids typically in *i*, *i+4* arrangements (Knight et al., 2005, Spink et al., 2008). It has been suggested that SAH domains are sufficiently stiff to function as mechanical extensions of the myosin lever arms and can thus provide for the bulk of the lever arm extension necessary for the large step sizes of myosin VI (Sivaramakrishnan et al 2009). In the model of Spink et al. (2008), depicted in Figure 1C, the myosin VI lever arm extension consists of the combination of the folded three-helix bundle and the SAH that follows it. Dimerization was proposed to occur within the cargo-binding domain, distal to the SAH (Spink et al., 2008).

In an alternative model proposed by Mukherjea et al. (2009), the lever arm extension for myosin VI is created by unfolding of the three-helix bundle upon myosin VI dimerization, as depicted in Figure 1D. In this model, dimerization between two myosin VI monomers occurs immediately distal to the three-helix bundle. Thus besides the nature of the lever arm extension itself, a major difference in the two models is the location of the internal dimerization region in myosin VI. In the unfolded three-helix bundle model, dimerization occurs prior to the SAH and in the model where the SAH is the major component of lever arm extension, dimerization occurs after the SAH between the two cargo-binding domains.

Note that in both models dimerization is weak, so that full-length myosin VI would normally exist as a monomer (Lister et al., 2004). It is only via interaction with its adapter proteins, many of which are themselves dimers (Tumbarello et al., 2013), that dimerization between the myosin VI monomers would occur (Spudich et al., 2007; Phichith et al., 2009). While there are two models of how myosin VI dimerizes, there also is ongoing controversy as to whether or not myosin VI actually functions as a dimer in cells (Lister et al., 2004; Park et al., 2006; Sweeney and Houdusse, 2010; Bond et al., 2012). Thus the questions we are trying to address are whether or not myosin VI functions primarily as a monomer or dimer
in cells, and in either case, whether or not it uses the SAH region as its lever arm extension or unfolds the three-helix bundle to create the lever arm extension.

# RESULTS

## Deletion of the SAH domain does not affect head gating and dimerization

To probe into the role of the SAH domain in myosin VI motor function, we designed a full-length construct lacking 37 residues (Glu942-Glu978) from the C-terminal end of the predicted SAH domain (MVI<sub>FL</sub>-ΔSAH) (Figure 1A). We preserved residues Lys915-Arg940 that may be necessary for dimerization of myosin VI constructs (Mukherjea et al, 2009). As shown in Table 1, following actin clustering of the monomers, MVI<sub>FL</sub>-ΔSAH exhibits gating (decreased ATPase activity per head), which is indicative of a dimeric myosin in which the lead head cannot complete its ATPase cycle until the rear head is released from actin. Since this dimerization is weak, clustering of myosin VI monomers on actin is required to increase the effective concentration of monomers above the low micro-molar concentrations of the ATPase assays in order to initiate this proximity-induced dimer formation (Park et al 2006). The average V<sub>max</sub> values of the actin-activated ATPase assays obtained for the single- and double-headed MVI<sub>FL</sub>-ΔSAH mutant are similar to the values obtained for the wild-type full-length protein under identical conditions (Table 1). The V<sub>max</sub> values of the dimeric MVI<sub>FL</sub>-ΔSAH construct are also in agreement with the steady-state values of the artificially zippered MVI<sub>991+zipper</sub> construct (Mukherjea et al, 2009). These results indicate that removal of the SAH after Arg940 does not affect gating, suggesting that the distal part of the predicted SAH domain is not involved in transmitting the intramolecular strain required for efficient head gating.

Park et al. (2006) were able to use rotary shadowing electron microscopy (EM) to visualize dimers of full-length myosin VI following actin clustering. However, only about 20% of the total population appeared to be dimers. Based on the ATPase values, one would expect a larger percentage of dimers. However, since the dimers are unstable, and there is a delay between when the molecules are released from the actin and when they are spayed on the mica surface, the EM is likely to be an underestimate of the number of dimers formed on actin. Nonetheless, we examined dimer formation of MVI<sub>FL</sub>-ΔSAH mutant following actin clustering, since EM can confirm the ability of a construct to dimerize (Figure S1). As summarized in Table 1, approximately 40% of the molecules appeared to be dimers. Only monomer and dimers were seen, with no larger aggregates observed.

## Myosin VI lacking the SAH can dimerize and step processively

Single molecule FIONA (Fluorescence Imaging with One Nanometer Accuracy; Yildiz and Selvin, 2005) assays provided further evidence that the MVI<sub>FL</sub>-ΔSAH dimer was sufficiently stable to function as a processive motor similar to its full-length wild-type counterpart. For these assays, the myosin VI proteins were labeled with biotin-CaM, as previously described (Park et al., 2006). To observe the stepping dynamics, streptavidin conjugated quantum dots (Q<sub>dots</sub>; λ=655 nm) were attached to the biotin-labeled monomers (Ali et al. 2011) and the monomers were bound to an actin filament in the absence of ATP to induce dimerization. Following addition of ATP, these actin-induced mutant dimers were
capable of processive movement along actin filaments with step sizes that were indistinguishable from the wild-type MVI<sub>FL</sub> actin-associated dimer (Figure 2A–C Table S1,) or the zippered MVI<sub>991+zipper</sub> construct (Mukherjea et al, 2009; Park et al., 2006). For the MVI<sub>FL</sub>-ΔSAH dimer, 31±15.5 nm (N=122) steps were observed (Figures 2B and C; Table S1) compared to the 28±18.5 nm (N=80) step sizes for the MVI<sub>FL</sub> dimer. We are confident that this behavior arises from dimers, rather than non-specific oligomerization. We have previously observed that when multiple motors labeled with the same color Qdot are linked together and observed moving, it is then impossible to discern individual steps (Ali et al., 2011). Additionally, the run length distributions of the wild type and mutant dimers were almost identical, 0.68±0.19 µm (N=75) for MVI<sub>FL</sub> and 0.72±0.16 µm (N=71) for MVI<sub>FL-ΔSAH</sub> (Figure 2D; Table S1).

## The unfolded three-helix bundle of myosin VI binds an additional CaM

Based on structures that reveal sources of compliance within the myosin VI converter and insert 2, we have been able to provide a model for the stepping behavior of the myosin VI dimer that can account for published measurements of step size distributions (Ménétrey et al., 2012). In this modeling, we noted that the published stepping data are best modeled if the unfolded three-helix bundle forms a relatively inextensible lever arm extension. A major shortcoming of the model of unfolding of the three-helix bundle is that the helices are amphipathic, which might favor a refolding of the bundle.

We have long noted that the inclusion of µM levels of free CaM in the buffer is required in order for myosin VI to show its normal processive movement in single molecule assays (Rock et al., 2001, 2005; Park et al., 2006). This has been puzzling since neither the insert-2-CaM nor the IQ-CaM readily dissociate. Furthermore, Liu et al. (2011) recently proposed the possibility of additional CaM binding within an unfolded three-helix bundle. Thus we investigated the possibility that once the three-helix bundle unfolds, additional, weaker CaM binding site(s) may be uncovered. This could lead to stabilization of the unfolded helices and create a more effective lever arm. We first examined this possibility by determining the number of CaMs that bind to our zippered myosin VI dimer at either 1 µM or 5µM free [CaM], bracketing the range required for optimal single molecule movement and within the physiological range of free CaM in a cell (Black et al., 2004). As summarized in Table S2, three CaMs bind to each heavy chain of the zippered dimer in the presence of either 1µM or 5µM CaM, consistent with a sub-micromolar affinity. Note that the monomeric myosin VI construct that contained the three-helix bundle (truncated at Lys917), but presumably in a folded form (Mukherjea et al., 2009), bound only two CaMs per heavy chain.

We next substituted glycines for hydrophobic residues found in the loop (in the folded bundle) between the first and second helix (D851-P865) of the bundle (F853G, V856G, V857G, L860G), which would not impact the helices or folding of the bundle, but could remove potential interactions for CaM. This mutation abolished the binding of the third CaM (Table S2), thus revealing that these residues are critical for the recruitment of this additional CaM upon unfolding of the three-helix bundle.

To determine the affinity and to delineate the binding site for the extra CaM, we performed microscale thermophoresis (MST) experiments with the zippered myosin VI dimer as well
as with peptides derived from the three-helix bundle and the adjacent proximal portion of the putative SAH. We measured the K<sub>d</sub> using Oregon Green-labeled CaM in the presence of 2 mM EGTA. As shown in Figure 3, the measured K<sub>d</sub> for the zippered myosin VI dimer is 0.67 µM. Also shown in Figure 3 is that the peptides revealed that CaM binding requires the proximal region of the bundle (K834-E875). The measured K<sub>d</sub> was 2.3 µM for peptide R849-R955 and 4.71 µM for peptide K834-E875. In contrast, CaM interacted weakly with the region distal to residue E875, with a K<sub>d</sub> of 93.1 µM for the peptide E875-R955 and a K<sub>d</sub> of 1502 µM for the peptide S887-R955.

We next demonstrated that removal of the hydrophobic residues from the loop connecting helix 1 and helix 2 (denoted by red ovals in the expanded three-helix bundle in Figure 1A), which eliminates binding of the extra CaM to the zippered dimer (above), also removes potential interactions for CaM in the peptides. Note that this loop was ordered in an NMR structure of the bundle (RCSB accession code 2LD3), as shown in Figure 1B. MST experiments performed with the mutated peptide (849–955; L850A, F853A, V856A, V857A, L860A) detected no CaM binding. As a control and to further delineate residues involved in CaM binding, we performed the same experiment with a construct with comparable mutations in the first helix of the bundle (834–875; I837A, L840A, V843A, L846A). These residues are denoted by green ovals in the expanded three-helix bundle sequence of Figure 1A and shown in red (space fill) in Figure 1B. The latter construct bound CaM with a K<sub>d</sub> of 4.5 µM, similar to the 8.5 µM obtained with the wild type peptide, indicating that these residues are not necessary for strong CaM binding (Figure 3). These results support a model in which an additional calmodulin binds to the proximal region of the unfolded bundle that includes residues 850–860.

A perfect SAH destroys myosin VI dimerization and prevents bundle unfolding

To delineate whether the proximal part of the putative SAH domain in myosin VI is part of a
SAH or is part of a dimerization domain in the dimer, we replaced the residues from Glu922
to Lys935 with alternate acidic and basic residues in a full-length construct (MVIFL-
SAHmimic) to match the (i, i+4) phasing observed in the distal part of the predicted SAH
domain (Figure 1A), creating a consensus SAH domain devoid of hydrophobic residues. In
order to maintain the correct registry with the legitimate SAH and to extend it, we added
three additional amino acids (one additional turn of the helix). The Vmax values of the actin-
activated ATPase activity were similar to monomeric wild type myosin VI, whether or not
actin clustering was employed, consistent with either loss of dimerization or loss of gating.
No dimers were observed with rotary shadowing EM (Table 1, Figure S1), consistent with
loss of dimerization. Furthermore, gating was observed when a leucine zipper was
introduced into the zippered MVI991-SAHmimic after amino acid 991 (wild type sequence
numbering) to give MVI991+zipper-SAHmimic. The leucine zipper will tether two monomers
in close proximity, as would occur when bound to cargoes such as Dab2 and optineurin
(Phichith et al., 2009). This zippered MVI991+zipper-SAHmimic displayed steady-state
ATPase values that were virtually identical to a dimeric, zippered wild-type construct
(MVI991+zipper), indicative of gating (Table 1). Altogether these data suggest that the
MVIFL-SAHmimic construct fails to dimerize in the steady-state ATPase assays, but if the
monomers are tethered together, the lever arms are sufficiently stiff to create gating between the heads.

Previously we were able to demonstrate unfolding of the three-helix bundle in the zippered wild type construct by introducing solvent-exposed cysteine residues into the bundle (at positions T845 and A880) and then labeling both residues with TMR (tetramethylrhodamine) to measure the fluorescence (Mukherjea et al., 2009). This places the two rhodamines close enough to stack with each other and quench the fluorescence via exciton coupling (Okoh et al., 2006). In a monomeric construct with the bundle folded, the close proximity of the two rhodamines led to quenching of the fluorescence. Upon dimerization of the construct by adding a leucine zipper, the fluorescence increased to that of two unquenched rhodamines, consistent with unfolding of the bundle (Mukherjea et al., 2009). When the same cysteines were engineered into the zippered MVI<sub>991</sub>-SAH<sub>mimic</sub> construct and labeled with TMR, the fluorescence remained quenched (Table 2), consistent with the bundle remaining folded.

We next analyzed the processive behavior of the MVI<sub>FL</sub>-SAH<sub>mimic</sub> mutant in single molecule FIONA assays. In the presence of actin clustering, no molecules exhibited processive movement. This provides further evidence that disruption of the putative dimerization domain immediately distal to the 3-helix bundle (residues 922–936), leads to loss of dimerization and thus processive movement upon actin clustering. This demonstrates that there is not sufficient interaction between the cargo binding domains to dimerize two monomers in the absence of binding to an adapter protein, and further demonstrates the involvement of the region from residues 922–935 in internal dimerization.

To determine the ability of the SAH<sub>mimic</sub> mutant to move processively if two monomers are held in close proximity as they would be by cargo binding in a cell, we compared the zippered MVI<sub>991+zipper</sub>-SAH<sub>mimic</sub> construct to the zippered wild type construct. When the single molecule motility was evaluated, the MVI<sub>991+zipper</sub>-SAH<sub>mimic</sub> was indeed processive, with a characteristic run length and average step size that were not significantly different from those of the zippered wild type (MVI<sub>991+zipper</sub>), as shown in Figure 4 and Table S1. The average velocities were also not different (0.30±0.13µm/sec vs. 0.31±0.13µm/sec for the dimeric MVI<sub>991+zipper</sub> and the dimeric MVI<sub>991+zipper</sub>-SAH<sub>mimic</sub>, respectively). Together, these results indicate that while there is no internal dimerization of the MVI<sub>FL</sub>-SAH<sub>mimic</sub> mutant, forced dimerization, and likely binding to cargo adapter proteins such as Dab2 or optineurin, that place monomers in close proximity, should allow processive movement.

## Impact of loss of either the SAH or dimerization on endocytosis

It has been shown that fibroblasts derived from the myosin VI-null *Snell's waltzer* (*sv*/*sv*) mice exhibit a significant defect in clathrin-mediated endocytosis, which is rescued by expression of the isoform of full-length myosin VI that is missing the large insert (NI) that follows residue 1036 (Puri, 2009). Since removal of a large portion of the SAH (MVI<sub>FL</sub>-ΔSAH construct) did not impact gating and processive movement under *in vitro* conditions, we investigated whether endocytosis is rescued in *sv*/*sv* fibroblasts expressing this mutant construct (NI isoform). We assessed the uptake of horseradish peroxidase (HRP) conjugated transferrin (Tf-HRP) in *sv*/*sv* fibroblasts transfected with either the wild-type MVI<sub>FL</sub> or
mutant MVI<sub>FL</sub>-ΔSAH (Figure 5A). Density scans of Western blots from transfected cells, such as shown in Figure 5A, revealed average expression levels 57.8 ± 10.6% and 63.0 ± 9.8% of myosin VI in wild type cells for the MVI<sub>FL</sub> and MVI<sub>FL</sub>-ΔSAH, respectively. As shown in Figure 5B, no significant differences were observed between the transferrin endocytosis in *sv/sv* fibroblasts expressing the wild type or the ΔSAH mutant proteins. However, in our experiments, transferrin uptake in the untransfected *sv/sv* fibroblasts is only slightly reduced compared to the wild-type transfected cells (Figure 5B).

It has been reported that in *sv/sv* fibroblasts in the absence of myosin VI, transferrin uptake occurs through a clathrin-independent, caveolae-mediated pathway (Puri, 2009). To determine whether partial inhibition of the caveolae-mediated pathway would allow us to better visualize rescue of clathrin-mediated endocytosis in the *sv/sv* fibroblasts, transferrin uptake was analyzed after the transfected cells were treated with the cholesterol uptake-blocking drug, filipin, which specifically inhibits caveolar endocytosis (Sigismund et al., 2008; Puri, 2009). The transferrin uptake in untransfected *sv/sv* fibroblasts was significantly reduced upon pre-treatment with filipin (Figure 5C), indicating that a caveolin mediated pathway is active in the absence of myosin VI as has been previously shown (Puri 2009). However, filipin treatment did not lower the transferrin uptake in either the wild type or the ΔSAH mutant myosin VI transfected *sv/sv* fibroblasts (Figure 5C), indicating that the SAH deleted mutant myosin VI can rescue the function of myosin VI in clathrin-mediated endocytosis.

Since replacement of residues between Glu922 and Glu936 with alternate acidic and basic residues in a full-length construct to create a consensus SAH (MVI<sub>FL</sub>-SAH<sub>mimic</sub>) resulted in loss of dimerization while preserving motor function, this afforded us a possibility to directly assess whether or not dimerization proximal to the SAH of myosin VI is necessary for its cellular functions. We thus repeated the endocytosis assays in *sv/sv* fibroblasts in the presence and absence of filipin with cells transfected with the MVI<sub>FL</sub>-SAH<sub>mimic</sub>. Surprisingly, given the *in vitro* performance of the artificially dimerized construct (zippered MVI<sub>991+zipper</sub>-SAH<sub>mimic</sub>), there was no discernable rescue of endocytosis in the absence of filipin and less than a fourth of the increase in uptake obtained with the wild type transfection (13% increase as opposed to a 58% increase) in the presence of filipin, despite achieving protein levels (61.9 ± 11.6% of wild type cells) that were similar to the other transfected constructs.

## Impact of removal of the SAH domain or preventing dimerization on Golgi structure

Lack of myosin VI in *sv/sv* fibroblasts has been reported to result in a significantly smaller and fragmented Golgi structure compared to the more extensive and reticulate appearance seen in wild-type fibroblasts (Warner et al, 2003). It has also been shown that expression of the full-length wild-type myosin VI in the *sv/sv* fibroblasts can rescue the Golgi morphology (Warner et al, 2003). To examine the effect of the SAH deletion on the Golgi structure, *sv/sv* fibroblasts were transfected with the wild-type MVI<sub>FL</sub>, the MVI<sub>FL</sub>-ΔSAH mutant, or the MVI<sub>FL</sub>-SAH<sub>mimic</sub> construct. The transfected fibroblasts were immunostained with an antibody against Coatomer subunit beta (β-Cop), which is primarily a cis-Golgi marker, although it is present to a lesser extent in the trans-Golgi (Griffiths et al., 1995). In the
untransfected *sv/sv* fibroblasts, the Golgi complex appeared as dispersed short tubular or vesicular structures (Figure 5D, S2), with markedly reduced Golgi area as compared to the MVI<sub>FL</sub>-transfected cells (Figure S2). The Golgi area was measured as a percentage of the total cell area (see Supplemental Experimental Procedures). Cells expressing either the wild-type MVI<sub>FL</sub> or the mutant MVI<sub>FL</sub>-ΔSAH exhibited more extensive Golgi structure (Figures 5D and S2), with the overall Golgi area increasing 2-fold and 1.6-fold, respectively. Thus compared to the wild-type transfected fibroblasts, the cells expressing the MVI<sub>FL</sub>-ΔSAH protein display only a slightly reduced Golgi size (Figures 5D and S2). In sharp contrast, the MVI<sub>FL</sub>-SAH<sub>mimic</sub> construct was not able to rescue Golgi morphology (5D and S2) and the overall Golgi area was indistinguishable from that of the untransfected cells (Figure 5D). In a few cells, as shown in Figure S2, there was less Golgi area than in untransfected cells.

# DISCUSSION

## New model for the Myosin VI dimer

New insights have emerged from this study that allow us to alter the model proposed by Mukherjea et al. (2009). Based on all of the data presented herein, we propose the model in Figure 1E for the functional dimer of myosin VI. In the model, two monomers dimerize and the three-helix bundle unfolds, recruiting an additional CaM, as suggested by the modeling of Liu et al. (2011). We determined that this additional CaM binds with sub-micromolar affinity (0.67 µM) in the region that comprises hydrophobic residues (L850, F853, V856, V857, L860) found at the end of the first helix of the folded bundle (Figure 1B, Figure 3, Table S2). This will likely stabilize the unfolded bundle to create the lever arm extension. Modifying the putative SAH to ensure that its most proximal region conforms to a consensus SAH (MVI<sub>FL</sub>-SAH<sub>mimic</sub>), allows us to rule out a model in which the nature of dimerization is purely electrostatic (Kim et al, 2010), and as discussed below, leads us to propose that the dimerization region involves some combination of residues from the distal portion of the unfolded three-helix bundle and the proximal region of what has been defined as the SAH, as illustrated in Figure 1E. Furthermore, deletion of most of the SAH, preserving the region necessary for dimerization, has no impact on *in vitro* or *in vivo* function, consistent with it being distal to the dimerization region and thus not contributing to the lever arm extension (Figures 2, 5B, 5C, and 5D; Table S1). It now remains to be determined where the lever arm extension ends and the dimerization region begins. Since it is clear that the additional CaM binding site involves the junction of the first and second helices, it is possible that all or part of the third helix of the bundle could contribute to a coiled coil for dimerization. This may explain why we see a small number of dimers in EM with a construct that is truncated at the end of the three-helix bundle (Table 1).

## Function of the SAH domain in Myosin VI

Even if the SAH never functions as a lever arm extension for myosin VI *in vivo*, there are two types of roles that it could play. Analogous to the long coiled coil of the myosin V isoforms (Trybus, 2008), the SAH could act both as a spacer between the motors and cargo, as well as create the proper geometry and interactions for folding in the monomeric state (Spink et al., 2008). In smooth muscle caldesmon, the ~65 residue SAH domain acts as a spacer and maintains the required distance between the N-terminal myosin and CaM binding
domains and the C-terminal actin and tropomyosin binding domain (Wang et al 1991; Mabuchi et al 1991). The SAH by definition acts as a spacer in the dimer since it follows the internal dimerization region and precedes the cargo-binding domain. However, the lack of major *in vivo* impact of our SAH deletion argues that the additional spacing created between the motor and cargo domains by the SAH is not necessary to allow cargo binding, at least for the functions we probed. While myosin VI does not have a dimeric, folded “off” state, as observed for myosin V (Trybus, 2008), the SAH may be necessary for proper folding of monomeric myosin VI, as was previously suggested (Spink et al., 2008). Partial deletion of the SAH in our MVI<sub>FL</sub>-ΔSAH construct could perturb this folding and may be the reason we see more dimers in EM with MVI<sub>FL</sub>-ΔSAH construct as compared to the MVI<sub>FL</sub> wild type (Table 1). This folding could prevent dimerization even if two monomers are in close proximity until the monomers are unfolded by binding to cargo adapter proteins. Once the dimer disengages from these adapter proteins, the weak dimerization likely allows the dimer to dissociate into monomers, which then refold.

We are not suggesting that SAH domains cannot play a role in extending lever arms of other classes of myosin. In myosin X, the SAH domain is part of the effective lever arm (Knight et al., 2005; Tokuo et al, 2007), as may also be the case in *Dictyostelium MyoM* (Baboolal et al 2009).

## A SAH lever arm extension can support in vitro movement but not in vivo function

Our *in vitro* experiments clearly demonstrate that the SAH does not function as a lever arm extension once myosin VI dimerizes and the three-helix bundle unfolds, and our *in vivo* experiments did not demonstrate any loss of function when the SAH was deleted. We next probed whether or not the SAH domain could serve as a lever arm extension if we prevented internal dimerization. To do so, we created the MVI-SAHi mimic constructs, in which internal dimerization and unfolding of the three-helix bundle did not occur (Tables 1, 2, S1). Since the cargo binding domains in the absence of cargo adapter proteins did not trigger dimerization, we replaced the cargo binding domain with a leucine zipper after residue 991, to force dimerization and essentially create the molecule illustrated in Figure 1C. This molecule (MVI<sub>991+zipper</sub>-SAHi mimic) was processive with run lengths nearly identical to that of the zippered wild type myosin VI (MVI<sub>991+zipper</sub>), although with increased run length variability (Figure 4 and Table S1). The molecule displayed gated ATPase activity equivalent to the zippered wild type construct (Table 1), implying that the SAH-containing lever arm is stiff enough to prevent completion of the power stroke of the lead head until the rear head has detached.

Given that the SAH domain can serve as a functional lever arm extension *in vitro*, we next addressed whether or not myosin VI must internally dimerize and unfold the three-helix bundle in order to carry out its cellular functions, or if closely spaced myosin VI monomers with folded three-helix bundles and SAH lever arm extensions can function *in vivo*. To answer this highly debated question, we examined the ability of the MVI<sub>FL</sub>-SAHi mimic construct that cannot internally dimerize and unfold the bundle (due to replacement of the putative dimerization region with a consensus SAH) to rescue endocytosis and Golgi morphology in fibroblasts lacking myosin VI. Given its *in vitro* performance, we anticipated
that the MVI<sub>FL</sub>-SAH<sub>mimic</sub> construct would demonstrate some functionality in cells.

However, it provided no rescue of either endocytosis or Golgi morphology in myosin VI null (*sv/sv*) fibroblasts (Figure 5). These results likely explain why a myosin VI mutation within this internal dimerization region, L926Q (Figure 1A), causes congenital deafness in humans (Brownstein et al., 2014).

The functions we chose to examine encompass the two types of roles myosin VI is thought to play in cells, transport and anchoring. To support endocytosis, myosin VI must act as a vesicle transporter, while to support the Golgi structural linkage to the surrounding actin network, myosin VI is likely functioning primarily as load-dependent anchor (Altman et al., 2004; Sweeney and Houdusse, 2010). Based on the outcome of these experiments, we conclude that myosin VI must function as a dimer, and not as a monomer for its two known types of cellular functions. We would anticipate that the myosin VI adapter proteins, Dab2 and optineurin, that are involved in endocytosis and Golgi localization, respectively, would place the cargo binding domains of two MVI<sub>FL</sub>-SAH<sub>mimic</sub> constructs in close proximity (as in Figure 1C), creating a similar geometry as found in the zippered MVI<sub>991+zipper</sub>-SAH<sub>mimic</sub> construct. Thus we conclude that the lever arm extension formed by unfolding the three-helix bundle is required for myosin VI to traffic in cellular actin networks, even though a lever arm extension consisting of the folded bundle and SAH is sufficient for normal movement on a single actin filament. The critical differences that account for this are unclear, but one can speculate that the unfolded three-helix bundle with one weakly associated CaM may have more flexibility than the SAH-containing lever arm. Furthermore, based on our earlier modeling (Ménétrey et al., 2012), the lever arm using the unfolded three-helix bundle would keep the unbound head closer to the same actin filament as the bound head, as compared to a more conventional lever arm. This geometry may allow the myosin to function under higher loads than can be borne by the SAH lever arm extension.

## A need for lever arm diversity

A fundamental question for myosin VI and other members of the myosin superfamily is what constraints does the lever arm design create for cellular functions, given the complexity of cellular actin networks (Svitkina et al., 1995). The unusual lever arm design (including the unusual converter rearrangements; Ménétrey et al., 2007) that has evolved for myosin VI may be specialized for different functional properties as compared to a more conventional lever arm design, such as found in myosin Va. The myosin VI lever arm design could favor myosin VI remaining on the same actin filament when trafficking in a dense cellular actin network (Sweeney and Houdusse, 2010), allowing more efficient transport as well as load-dependent anchoring. There is *in vitro* evidence suggesting that myosin VI does not switch actin filaments as easily as myosin Va (Brawley and Rock, 2009; Ali et al., 2013). Myosin Va and the closely related myosin Vb, have extended and relatively stiff lever arms, which may be necessary to stall the motors and tether vesicles within actin networks, rather than smoothly transporting (Wu et al., 1998; Provance et al., 2008). Such tethering would prevent myosin VI from being an efficient vesicular transporter in endocytosis. Our present study demonstrates that the unusual myosin VI lever arm design is necessary for its *in vivo* function. Given that this lever arm extension has helical regions that are likely compliant, but not extensible, we postulate that compliance may be a necessary
feature of the lever arms of myosin motors designed to traffic within dense cellular actin networks. Since the myosin VI lever arm extension is deployed only upon dimerization-induced unfolding of the three-helix bundle and calmodulin recruitment, dimerization of myosin VI is necessary to support its cellular functions.

# EXPERIMENTAL PROCEDURES

## Protein Constructs, Expression, EM, and ATPase assays

The myosin VI constructs were generated using a porcine myosin VI cDNA (amino acid residues 1–991) and a mouse myosin VI cDNA (spliceform without the large insert) as templates. Details of construct design, protein expression, and ATPase assays are presented in the Supplementary Experimental Procedures. The rotary shadowing electron microscopy (EM) was performed as previously described (Park et al., 2006). See Supplemental Experimental Procedures.

## Single-molecule assays

Biotin labeled CaM was used to selectively exchange only one of the two myosin VI CaMs in the motor domains of the wild-type or the mutant proteins for the single molecule studies as described before (Yildiz et al., 2004). The processive movements of MVI<sub>FL</sub> and MVI<sub>FL-ΔSAH</sub> were observed on 20 µl flow cells using total internal reflection fluorescence microscope (TIRFM) as described (Baker et al. 2005; Ali et al. 2011). Details of single molecule assays and data analysis are presented in Supplemental Experimental Procedures.

## MicroScale Thermophoresis (MST) assays

The method of Wienken et al., 2010 was adapted for probing myosin VI interactions with CaM. Details are presented in Supplemental Experimental Procedures.

## Cell culture and Transfection

Immortalized *Snell's waltzer* (*sv*/*sv*) tail fibroblasts were grown on coverslips and transient transfections were carried out with the wild type and mutant myosin VI constructs using Lipofectamine 2000 (Invitrogen, CA) following the manufacturer's protocol.

## Immunofluorescence studies

Details of immunofluorescent staining of the Golgi complex in the *sv*/*sv* fibroblasts and determination of Golgi area are presented in Supplemental Experimental Procedures.

## Transferrin uptake assays

The *sv*/*sv* fibroblasts were cultured as described above on 35mm dishes and transfected with the wild-type or mutant myosin VI constructs. Untransfected cells were used as a control. Transferrin uptake was carried out as described previously (Buss et al., 2001; Damke et al., 1995). Twenty-four hours post-transfection, the fibroblasts were serum starved by incubating in serum-free DMEM for 2 hours. Cells were washed twice with PBS and incubated with 25µg/ml horseradish peroxidase conjugated transferrin (Tf-HRP) (Jackson ImmunoResearch, PA) for various time-points at 37°C. After the incubation, culture dishes
were placed on ice and washed twice with 10mM HCl, 150mM NaCl to remove the cell surface bound transferrin. Cells were then lysed with 0.5% Triton X-100 and the lysates were incubated with 3,3',5,5'-tetramethylbenzidine (Sigma, MO) in the dark for 20 minutes. The reaction was stopped by the addition of 1N H₂SO₄ and the absorbances for the unbound, surface bound and internalized fractions were quantified at 450 nm using a microplate reader. The data was plotted as a percentage of the total Tf-HRP internalized. Each experiment was performed in triplicate.

For the transferrin uptake studies in the presence of filipin, the transfected *sv/sv* fibroblasts were serum starved for 2 hours and incubated with 1.2µg/ml filipin (Sigma, MO) for 1 hour followed by Tf-HRP uptake for 5 minutes. Control cells were not treated with filipin. The cells were then analyzed as described above and the results were plotted with the corresponding untreated controls. Data is represented in graphs as mean ± standard deviation (SD) for 6 independent sets of experiments. Student's t-test calculation was used to analyze the statistical significance of the differences among the plotted values. A value of p ≤0.05 was considered statistically significant.

**Supplementary Material**

Refer to Web version on PubMed Central for supplementary material.

**Acknowledgments**

We are grateful to Xiaoyan Liu for protein purification and to Allan B. Zong for the performance of ATPase assays. We thank Mirko Travaglia for rotary shadowing and Clara Franzini-Armstrong for access to EM equipment. This work was supported by NIH grants DC009100 (HLS) and GM094229 (DMW). Additional support was provided by the CNRS (AH), the ARC subvention fixe SFI20121205398, a FRM equipe 2010 grant and the ANR blanche BLAN 2010 1504 01 (AH). Part of this work was performed by a team (AH) belonging to the labex CelTisPhyBio 11-LBX-0038. MM is supported by a grant from the ALS Foundation for Childhood Cancer and MYA from the American Heart Association (12SDG11930002).

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## Highlights

* Unfolding of a 3-helix bundle recruits calmodulin to the myosin VI lever arm

* Deletion of the SAH domain does not impact *in vitro* movement, endocytosis, or Golgi

* Internal dimerization is required to rescue endocytosis and Golgi organization

* Internal dimerization creates a lever arm specialized for movement in cells
<figure><img src="image_2.png" /><figcaption>**Figure 1. Schematic of full-length myosin VI**</figcaption></figure>

A. In the full-length myosin VI (MVI~FL~), the motor domain at the N-terminus is followed by a unique insert (Insert 2) that binds the structural calcium-bound calmodulin (Ca²⁺-CaM), followed by an IQ motif that binds an apo-CaM. The IQ-motif is followed by a three-helix bundle, that has been proposed to unfold and form an extension of the CaM-containing lever arm when myosin VI dimerizes (Mukherjea et al., 2009). The three-helix bundle is followed by a region (shown in light and dark green) that has been predicted to be a stable single alpha helix (SAH) domain (Knight et al., 2005; Spink et al., 2008), and can potentially form an extension of the myosin VI lever arm. However, it is been proposed that the proximal end of the putative SAH (light green) contributes to dimerization (Mukherjea et al., 2009). At the C-terminal end of the molecule is the cargo-binding domain. Underneath the schematic is the amino acid sequence of the three-helix bundle and the regions flanking it (N-terminal: IQ motif; C-terminal: proximal portion of the SAH). The diagram beneath the sequence, shows the peptides that were investigated in this study. A mutant construct was made for this study with residues Glu942-Glu978 (dark green region) of the SAH domain deleted (MVI~FL~-ΔSAH). In a second mutant construct, MVI~FL~-SAH~mimic, residues from Glu922 to Arg940 (underlined in sequence) were replaced with alternate acidic and basic residues (shown in light green bar under sequence) to create a consensus SAH. The red asterisk denotes the position of leucine 926, which when mutated to a glutamine, causes congenital deafness in humans (Brownstein et al., 2014). **B. Structure of the myosin VI three-helix bundle.** The residues that when mutated destroyed CaM binding are shown in red. The
region containing these residues was disordered in the crystallographic structure of three-helix bundle (RCSB accession code 3GN4). They are shown as part of the helix in the figure according to the NMR structure of three-helix bundle (RCSB accession code 2LD3). C.

**Model of myosin VI dimer with folded three-helix bundle.** The model for the myosin VI dimer proposed by Spink et al. (2008), which employs the SAH domain as the lever arm extension. Note that in this model, dimerization would only occur between the cargo-binding domains and/or within the adapter protein bound to the cargo-binding domain. D.

**Model of myosin VI dimer with unfolded three-helix bundle.** The model for the myosin VI dimer proposed by Mukherjea et al. (2009), which uses the unfolded three-helix bundle as the lever arm extension. Note that in this model, dimerization must occur immediately following the unfolded three-helix bundle. E. Modified model of the myosin VI dimer.

Based on the experiments described in this study, we propose modifications to our previous model of the myosin VI dimer (illustrated in Figure 1D). On the left, two folded monomers are depicted with the folding stabilized by interactions with the SAH (Spink et al., 2008). We propose that dimerization is initiated when two folded monomers encounter a cargo that places their cargo binding domains in close proximity (Phichith et al., 2009). This close proximity allows dimerization to begin at the region immediately proximal to the SAH (light green in dimer cartoon) and then propagate into the third helix of the bundle (light blue), triggering the bundle's unfolding. The unfolded bundle exposes a sub-micromolar affinity CaM binding site between the first and second helices of the bundle, which recruits a free CaM. This CaM binding, coupled with participation of the third helix in dimerization, leads to stabilization of the unfolded bundle. Since there will be helical segments that are not covered by CaM or participating in dimerization, these likely contribute flexibility to the lever arm, although it will be relatively inextensible.
<figure><img src="image_3.png" /><figcaption>**Figure 2. Single molecule motility**</figcaption></figure>

Step sizes and run lengths of wild type (MVI<sub>FL</sub>) and mutant (MVI<sub>FL</sub>-ΔSAH) constructs at 2 mM [ATP]. Step size traces of (A) MVI<sub>FL</sub> and (B) MVI<sub>FL</sub>-ΔSAH. (C) Distribution of step sizes of MVI<sub>FL</sub> (black) and MVI<sub>FL</sub>-ΔSAH (red). The histograms were fit to single Gaussians, centered at 56±37 nm (N=80) for MVI<sub>FL</sub> and 62±31 nm (N=122) for MVI<sub>FL</sub>-ΔSAH. (D) Characteristic run lengths of MVI<sub>FL</sub> and MVI<sub>FL</sub>-ΔSAH are 0.68±0.19 µm (N=75) and 0.72±0.16 µm (N=71) respectively, obtained from a single exponential fit. The results are summarized in Table S1.
<figure><img src="image_4.png" /><figcaption>Figure 3. CaM binding affinity to the lever arm extension</figcaption></figure>

Binding of Oregon green-labeled calmodulin to myosin VI HMM and to myosin VI peptides as quantified using Microscale Thermophoresis (MST) experiments. See also Table S2.
<figure><img src="image_5.png" /><figcaption>Figure 4. Run lengths and step sizes of leucine zipper-dimerized wild type (MVI991+zipper) and MVI991+zipper-SAHmimic constructs</figcaption></figure>

Characteristic run lengths of (A) MVI991+zipper and (B) MVI991+zipper-SAHmimic obtained from single exponential fits to the data (as shown), were: 0.56 ± 0.23 µm (N=1493) and 0.62 ± 0.76 µm (N=537), respectively. Step size traces of (C) MVI991+zipper and (D) MVI991+zipper-SAHmimic plotted as displacement over time. (E) Histograms of step sizes of MVI991+zipper and (F) MVI991+zipper-SAHmimic were fit to single Gaussians, centered at 64±19 nm (N=124) for MVI991+zipper and 54±22 nm (N=108) for MVI991+zipper-SAHmimic. The results are summarized in Table S1.
<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /><figcaption>**Figure 5.** Endocytic uptake of transferrin and rescue of Golgi morphology in transfected *sv/sv* fibroblasts</figcaption></figure>

(A) Immunoblotting was performed on lysates from *sv/sv* fibroblasts transfected with either a full-length wild-type myosin VI (MVIFL) or a mutant construct (MVIFL-ΔSAH) with an anti-myosin VI antibody. Myosin VI was not detected in untransfected (un) cells. Fibroblasts derived from a heterozygous (*sv/+*) mouse expressing myosin VI served as a positive control. (B) Uptake of HRP conjugated transferrin in *sv/sv* fibroblasts expressing the MVIFL, MVIFL-ΔSAH, or MVIFL-SA>Hmimic proteins was measured at different timepoints. Mean data ± SD (from 6 experiments) are shown. The amount of Tf-HRP internalized is expressed as a percentage of the total transferrin. Control transferrin uptake was measured in untransfected cells. (C) Transfected *sv/sv* fibroblasts were treated with filipin and Tf-HRP internalization was measured. Mean data ± SD (n=6) were plotted as values normalized against the untreated controls. (D) To analyze the differences in Golgi area among the different constructs, the amount of area covered by the Golgi in the transfected and untransfected *sv/sv* fibroblasts were quantified with respect to the total cell area in each case. Each column and error bar represents the mean ± S.D for 120 cells (20 cells from each of 6 independent preparations of cells). \*, P < 0.0001 denotes statistical differences between untransfected and transfected *sv/sv* cells. See also images in Figure S2.
<caption>Table 1</caption>

Actin activated ATPase activity and dimerization of Myosin VI constructs

<table><thead><tr><th rowspan="2">CONSTRUCT</th><th rowspan="2">V<sub>max</sub> (head<sup>-1</sup> second<sup>-1</sup>)</th><th rowspan="2">K<sub>ATPase</sub> (μm)</th><th colspan="3">After Actin saturation</th></tr><tr><th>V<sub>max</sub> (head<sup>-1</sup> s<sup>-1</sup>)</th><th>K<sub>ATPase</sub> (μm)</th><th>% Dimers (EM)</th></tr></thead><tbody><tr><td>MVI<sub>FL</sub></td><td>5.4 ± 0.28</td><td>9.5 ± 1.3</td><td>3.5 ± 0.14</td><td>5.9 ± 1.9</td><td>17 ± 2<sup>a,b</sup></td></tr><tr><td>MVI<sub>FL</sub>-ΔSAH</td><td>4.6 ± 0.12</td><td>8.8 ± 0.6</td><td>2.5 ± 0.2</td><td>4.7 ± 1.2</td><td>38 ± 9<sup>a</sup></td></tr><tr><td>MVI<sub>FL</sub>-SAH<sub>mimic</sub></td><td>5.9 ± 0.13</td><td>8.3 ± 0.5</td><td>4.8 ± 0.2</td><td>10.7 ± 1.1</td><td>0 ± 0 <sup>a</sup></td></tr><tr><td>MVI<sub>917</sub> (truncated monomer or S1)</td><td>5.8 ± 0.22</td><td>12.1 ± 3.6</td><td>NA</td><td>NA</td><td>9 ± 6<sup>a,b</sup></td></tr><tr><td>MVI<sub>991+zipper</sub></td><td>3.0 ± 0.04</td><td>3.2 ± 1.5</td><td>NA</td><td>NA</td><td>ND</td></tr><tr><td>MVI<sub>991+zipper</sub>-SAH<sub>mimic</sub></td><td>3.2 ± 0.10</td><td>2.6 ± 1.2</td><td>NA</td><td>NA</td><td>ND</td></tr></tbody></table>

ATPase values are expressed as mean values (±S.D.) of 3–5 independent protein preparations are shown for each construct and condition.

<sup>a</sup> Rotary Shadowing EM results are mean values (±S.D.) for each construct and condition. Data were obtained from a single spray experiment; two to five electron micrographs and 100–500 molecules were inspected. See also Figure S1.

<sup>b</sup> Values taken from Park et al. (2006).
<caption>Table 2</caption>

Fluorescence changes observed by 5-TMR labeling of single or double cysteine residues inserted in MVI<sub>991+GCN4</sub>-SAH<sub>mimic</sub>

<table><thead><tr><th rowspan="2">CONSTRUCT</th><th colspan="3">Fluorescence Ratio<sup>a</sup></th><th rowspan="2">Molar Ratio of labeling per myosin head<sup>b</sup></th></tr><tr><th>without Actin or ATP</th><th>with Actin</th><th>with Actin +ATP</th></tr></thead><tbody><tr><td>MVI<sub>991+GCN4</sub>-SAH<sub>mimic</sub> T845C</td><td>362.7 ± 24.1</td><td>331.6 ± 33.2</td><td>342.3 ± 40.6</td><td>1.06 ± 0.02</td></tr><tr><td>MVI<sub>991+GCN4</sub>-SAH<sub>mimic</sub> T845C, A880C</td><td>47.7 ± 5.6</td><td>53.3 ± 9.4</td><td>57.1 ± 6.2</td><td>2.22 ± 0.37</td></tr></tbody></table>

<footnote><sup>a</sup>Fluorescence changes were analyzed by a ratio of the emission values to that of the absorption values for each construct. Absorption was measured at 552nm and emission was measured at 575nm.</footnote>