Humans have 13 class II myosin heavy chain genes. Most of these encode for the heavy chains of skeletal, cardiac and smooth muscle myosins, but three encode for the heavy chains of paralogs that are widely expressed in nonmuscle tissue. These are commonly called NM2-A, NM2-B and NM2-C, each with different heavy chains encoded by the MYH9, MYH10 and MYH14 genes, respectively (Berg et al., 2001; Vicente-Manzanares et al., 2009). Nonmuscle class II myosins (NM2) are molecular motors involved in cytokinesis, cell migration, adhesion and tissue morphogenesis (Heissler and Manstein, 2013; Vicente-Manzanares et al., 2009). The coiled-coil tail region of the heavy chain of each of these myosins homodimerizes and the neck region associates with an essential light chain (ELC) and a regulatory light chain (RLC), creating a hexameric molecule. These individual myosin molecules further self-associate via their tails to form bipolar filaments that are approximately 300 nm in length and contain, on average, either 30 myosin molecules for NM2-A and NM2-B or 16 myosin molecules for NM2-C (Billington et al., 2013; Niederman and Pollard, 1975). In addition, it has been demonstrated that NM2-A and NM2-B can co-polymerize to form heterotypic filaments in cells (Beach et al., 2014; Shutova et al., 2014). It is unknown why mammalian cells express three different NM2 paralogs, or what their individual or shared functions are (Conti et al., 2008; Vicente-Manzanares et al., 2009). The enzymatic activity and the filament assembly of each of the NM2 paralogs are regulated by phosphorylation of the RLC by myosin light chain kinase (MLCK), or other cellular kinases (Heissler and Sellers, 2016).

It has long been known that the different skeletal muscle myosins-2 have distinct enzymatic and mechanical properties from studies in muscle fibers and of isolated proteins (Bottinelli and Reggiani, 2000). Similarly, numerous enzymatic studies conducted on the soluble, single-headed subfragment-one (S1) or double-headed heavy meromyosin (HMM) fragments have revealed differences in the steady state and transient state kinetics of the three NM2 paralogs that suggests there is differentiation of function amongst them (Heissler and Manstein, 2011; Kovács et al., 2003; Rosenfeld et al., 2003; Wang et al., 2003). The enzymatic activity of all three NM2 paralogs is low compared to other myosin 2 family members. Nevertheless, of the three NM2 paralogs, NM2-A has the highest actin-activated ATPase activity and translocates actin filaments the fastest (Kim et al., 2005; Wang et al., 2003). During the course of binding and hydrolysis of ATP and subsequently product dissociation, myosin cycles through conformations that bind weakly and strongly to actin. The duty ratio (r) of a myosin is defined as the fraction of a kinetic cycle the myosin spends in a conformational state which binds strongly to actin. NM2-B has a four-fold higher duty ratio than NM2-A, meaning that it will spend a greater proportion of its ATPase cycle strongly bound to actin filaments (Kovács et al., 2003; Wang et al., 2003).

The role of myosin bipolar filaments as force generating structures in cells has been clearly demonstrated (Vicente-Manzanares et al., 2009). Myosin filaments can interact with actin filaments of opposite polarity to create a sarcomeric type contractile unit, similar to those found in striated muscles (Ebrahim et al., 2013). They may also move processively along actin filaments in cells which could aid in the organization of the actin cytoskeleton (Beach et al., 2017; Ebrahim et al., 2013; Fenix et al., 2016; Hu et al., 2017) or may serve as protein scaffolds (Joo et al., 2007; Maddox et al., 2007; Piekny and Glotzer, 2008). Recent studies using super-resolution fluorescence microscopy for the visualization of single, or small clusters of myosin filaments in cells, allow for the possibility of dissecting their detailed function (Beach et al., 2017; Beach and Hammer, 2015; Beach et al., 2014; Fenix et al., 2016; Hu et al., 2017). However, despite their fundamental roles in generating force in cells, the mechanical properties of individual NM2 filaments are unknown.

To understand their individual mechanical properties, we used purified full-length human NM2-A and NM2-B molecules expressed in Sf9 cells to reconstitute and study the ability of single filaments of NM2 myosin paralogs to move along actin filaments bound to the coverslip surface. We find that NM2-B filaments are processive even at low-solution viscosity and that 5 – 10 myosin motor domains per half filament are required to ensure processivity. In contrast, NM2-A filaments are only processive when the viscosity of the solution is raised to mimic that of the intracellular millieu using methylcellulose or when NM2-A is co-polymerized with NM2-B. These data suggest that NM2-B is more adapted to a role of maintaining a high static tension level in cells, whereas NM2-A is geared to more rapid motile activity. Our data further show that formation of heterotypic NM2 filaments is an effective mechanism to fine tune filament mechanical properties between these two modes. As cell biologists start to decipher the cellular roles of NM2 filaments using fluorescence microscopy, our study demonstrates that not all NM2 filaments are equivalent and that subunit composition must be considered in order to fully understand their function.

Our study has important implications for understanding how NM2 myosin filaments interact with actin filaments and function within the cell. Despite their almost identical structure, NM2-A and NM2-B filaments differ significantly in their motile and kinetic properties. NM2-B filaments have a higher composite duty ratio and move slower than NM2-A filaments under the unloaded conditions of the single filament in vitro motility assay. The effect of force on the kinetics of NM2 paralogs has not been measured using optical trapping, but a previous transient kinetic study showed that the rate of dissociation of ADP from both acto-NM2-A and acto-NM2-B motor domains is affected by intramolecular strain that occurs when the two motors of a myosin molecule are bound to adjacent actin monomers in the presence of ADP (Kovács et al., 2007). However, this study revealed that NM2-B is significantly more strain dependent than NM2-A. Since the dissociation of ADP from acto-NM2 limits the rate of dissociation of NM2 from actin, this suggests that under strained conditions, NM2-B will have a significantly longer lifetime on actin filaments than NM2-A. This implies that NM2-B filaments might be better suited for maintaining tension on actin filaments in cells, whereas NM2-A would be more useful as a cargo motor for rapid but unsustained contractions or in the remodeling of the actin cytoskeleton, given its higher motility rate. In this regard, it is interesting that NM2-A is more prominent in the leading edge of cells where there are active actin dynamics, whereas NM2-B is typically more concentrated with stress fibers which are linked through adhesion complexes to generate forces on the substrate and where actin is highly dynamic (Beach et al., 2014). The longer attachment lifetimes and greater strain dependence of these attachments would allow NM2-B to maintain force more effectively and efficiently than NM2-A.

We also show in the present study that NM2-A and NM2-B molecules co-polymerize randomly in vitro consistent with reports that they readily form heterotypic filaments in cells (Beach et al., 2014; Shutova et al., 2017; Shutova et al., 2014). These co-filaments move processively over a range of NM2-A to NM2-B ratios. Our data show that co-filaments containing as few as six to eight molecules of NM2-B in the presence of a majority of NM2-A molecules continue to move processively, albeit with shorter run lengths than filaments containing higher amounts of NM2-B. Interestingly, the velocity is strongly dictated by the slower NM2-B which is consistent with experiments using the sliding actin in vitro motility assay when fast and slow myosins are mixed on the surface (Cuda et al., 1997; Harris et al., 1994). These results imply that a cell can tune the mechanical output of a filament by varying its composite ratio of NM2 paralogs. Other sources of strain in a cell might affect the mechanical properties of NM2 paralogs. Strain could arise from two myosin filaments interacting with the same actin filament or from actin-binding proteins that might tether the filament to the cytoskeletal component or a membrane which might provide resistance to the NM2-directed motion.

By copolymerizing NM2-B with a tail fragment, we were able to determine that more than four motors need to be present at one end of a bipolar filament to ensure its processivity. This agrees remarkably well with calculations based on the duty ratio of the individual NM2-B motor domain (Wang et al., 2003). We also found that the rate of movement of the hybrid myosin filaments on actin were not significantly affected by the number of myosin motors available. These results suggest that the rate of movement of the myosin filaments are governed by the rate of detachment of myosin from actin. This is in contrast to a recent study with smooth muscle myosin filaments suggesting that the rate of filament movement is limited by the myosin attachment rate (Brizendine et al., 2015). There are several differences in the myosins and experimental setups that could have contributed to the different conclusions. First, smooth muscle myosin forms side-polar filaments in contrast to the bipolar filaments formed by NM2-B. Second, the duty ratio of smooth muscle myosin is low and it was necessary to include methylcellulose in the assay medium. In the presence of methycellulose, it might be possible for all the motors in a myosin filament to transiently be dissociated from actin without the myosin filament diffusing away before a new motor can bind and tether the myosin filament to actin. A second study from this same lab has recently been published containing a more elaborate model (Brizendine et al., 2017). This model allows that myosin filaments may move with attachment or detachment limited kinetics depending on the myosin, myosin motor density and ionic conditions. The higher duty ratio of NM2-B would inherently bias the system toward a detachment limited mode. The observation that the rate of movement of the filaments did not vary with the number of contributing motors was consistent with a recent study which made synthetic myosin filaments mimics using DNA nanotubes where the spacing and density of myosin motor domains could be varied (Hariadi et al., 2015).

It is significant that tail fragments that were N-terminally fused with either the Halo-tag or a fluorescent fusion protein (data not shown) form bipolar filaments that, by negatively stained transmission electron microscopy, are similar to those formed by full-length molecules. In contrast, numerous studies have shown that myosin tail fragments that are not N-terminally capped by GFP or a HaloTag form much longer and thicker aggregates or paracrystals when polymerized either in vitro or when overexpressed in cells (Cohen et al., 1970; Franke et al., 2005). One potential use of capped tail fragments would be as a dominant-negative construct for myosin in cells where they would co-polymerize with full-length NM2 and thereby reduce the average number of myosin motors in a filament. However, the polymerization of these tail fragments lacking the light chain binding sites and therefore no RLC, would not be under the control of myosin light chain kinases and could result in inappropriate assembly and mislocalization of the myosin filaments in cells (Beach et al., 2017).

In view of the experimentally determined single molecule duty ratio of 0.05 for NM2-A, it is not surprising that these filaments failed to move processively in the absence of methycellulose. Calculations using Equation (1) show that more than 50 motors are required at one end of a NM2-A bipolar filament to ensure that at least one motor would be attached to an actin filament and ensure continuation of a processive run. There are only about 30 motors available at the end of a bipolar filament and not all of these are likely to be in a geometrically favorable position for actin filament interaction at any given moment. Under these low viscosity conditions, the myosin filaments rapidly diffuse away from an actin filament when no motors were attached.

The viscosity experienced by objects the size of NM2 filaments in cells is expected to be between one and two orders of magnitude larger than that in a simple aqueous buffer. Therefore, we mimicked elements of the physiological viscosity by adding 0.5% methylcellulose to increase the viscosity by about 25-fold (Kalwarczyk et al., 2011). We show that under these higher viscosity conditions NM2-A filaments behave as processive units. A recent study of the motility of single filaments of smooth muscle myosin also found they did not move processively under low-viscosity buffer conditions, but that inclusion of methylcellulose into the assay chamber allowed for robust motility (Haldeman et al., 2014). Methylcellulose was also used in in vitro motility experiments with skeletal muscle myosin-2 (Murrell and Gardel, 2012). Methylcellulose may act in two ways in this assay. By increasing solution viscosity, the diffusion rate of the NM2-A filaments is reduced sufficiently such that upon an instantaneous dissociation from actin, it would not diffuse away before another myosin motor could attach and restore processive movement. Methylcellulose is also a crowding agent which may influence the kinetics, and therefore the duty ratio of NM2-A (Highsmith et al., 1996).

Interestingly, inclusion of methylcellulose in assays of NM2-B filament produced a ‘parking’ behavior, where filaments were unable to detach upon reaching the end of the actin filament and where multiple myosin filaments accumulated at actin intersections. This behavior precluded a quantitative comparison of NM2-B filament run lengths under viscous and nonviscous conditions. The highly processive nature of NM2-B filaments is interesting to consider in that this myosin partitions to the rear of the cell and results in the production of an extended cell rear or tail, via the production of stable and long lived actomyosin structures (Beach et al., 2014; Vicente-Manzanares et al., 2009). The parking behavior of NM2-B filaments at high viscosity has also been observed for smooth muscle myosin filaments under similar assay conditions (Haldeman et al., 2014). The spatial self-sorting behavior of NM2-A and NM2-B filaments has recently been demonstrated to rely on a gradual enrichment of NM2-B in stress fibers during retrograde flow (Beach et al., 2014; Shutova et al., 2017). Our results offer an explanation for how NM2-B can become preferentially enriched towards the cell posterior in polarized cells, since NM2-B rich filaments have an enhanced activity to remain bound to actin and will thus tend to track rearwards with retrograde flow. Conversely, NM2-A rich filaments have a greater probability of detaching from actin and will thus undergo relatively high rates of turnover and recycling with respect to actin (Shutova et al., 2017).

Another significant difference in our in vitro assays and the cellular environment is that in our system each moving myosin filament typically has contact with only one actin filament. A single cluster of motor domains emanating from one end of a bipolar filament could potentially have interactions with about a 100 nm stretch along an actin filament (Billington et al., 2013). This length of actin encompasses around 40 actin monomers, many of which would be sterically unavailable to the myosin motor domains. Inside a cell, myosin filaments may interact with a bundle of actin filaments or with multiple individual actin filaments. Both of these cases would tend to increase the ability of myosin to processively interact with actin within its cellular environment. The ability of NM2 filaments to interact with actin in an ‘end on’ manner where the cluster of myosin motors on the opposite end of the bipolar filament project outward would also enhance the ability of NM2 filaments to cross-link and bundle actin filaments or to slide actin filaments relative to one another. The single filament in vitro motility assay will be very useful in dissecting this mode of actin-myosin interaction.

Recently, studies using super-resolution light microscopy found that individual myosin filaments align to form vertical stacks. The formation of these stacks appeared to require both actin and myosin dynamics (Fenix et al., 2016; Hu et al., 2017). We observed multi-filament structures that resemble these structures being formed when two NM2 filaments meet whilst moving on an actin filament. These structures are dynamic and can shed or add individual myosin filaments. Since dynamically treadmilling actin was not present in our in vitro assays, this suggests that merging and splitting of such structures is an intrinsic property of myosin filaments when they are brought into close proximity with each other. The physical basis for forming stacks is not known. The attraction is likely to be electrostatic since fewer filament stacks are observed by electron microscopy as the ionic strength is raised. To form the folded ‘off’ state, the two motor domains of a single myosin make an asymmetric interaction in which the loop 2 region of one motor interacts with the converter domain of another (Jung et al., 2008; Wendt et al., 2001). This interaction appears to be broken upon phosphorylation of the regulatory light chains. It is possible that this weak interaction comes into significance when two filaments, each bearing 30 motors per filament end, come into close proximity. The fact that the stacks can disassemble suggests that the forces holding them together are not large.

The question remains as to why mammalian cells have three genes to encode nonmuscle myosins. Studies in mutant mouse models reveal that different nonmuscle myosin paralogs have different patterns of expression during development (for a review, see (Ma and Adelstein, 2014). For example, in the early embryo, only NM2-A is expressed in the developing visceral endoderm (Conti et al., 2004). Ablation of NM2-A is lethal at embryonic day 6 since the cells cannot form appropriate adhesion complexes between cells. The DNA encoding the NM2-B heavy chain can be inserted into the NM2-A genetic loci resulting in expression of NM2-B during this early embryonic period. The NM2-B/NM2-A switch can rescue the early embryonic lethality, but the mouse experiences other defects later in development (Wang et al., 2010). Other studies reveal cases where paralog switching is not successful. NM2-A is specifically required for placental blood vessel formation and NM2-B is required for neuronal cell migration during brain development (Ma et al., 2007; Wang et al., 2011). In these cases, the opposite paralog cannot functionally substitute, suggesting that each of these myosins have features that are not redundant for some cellular functions.

Our results suggest that relating light microscopic observations of myosin filaments in cells to their mechanical output is not trivial. With the advent of super-resolution light microscopy, it is now possible to image single myosin filaments in cells, but most experimental methods used to date are not sufficient to truly understand the composition of the observed filaments (Beach et al., 2017; Fenix et al., 2016; Hu et al., 2017). If the investigator uses overexpression of a GFP-tagged RLC, then the heavy chain composition of the filament will be unknown. Similarly, visualization of myosin filaments by overexpression of a GFP-tagged myosin heavy chain does not rule out the possibility that an observed filament could be a co-polymer with another NM2 paralog, or with myosin 18A (MYO18), which is an enzymatically inactive pseudo-myosin.

In summary, a cell has several potential mechanisms to mechanically fine tune or regulate the activity of a given myosin filament. These include forming heteromeric filaments composed of two or more NM2 paralogs (or myosin 18A) with differing mechanical properties and controlling the number of myosin molecules in a filament or the phosphorylation status of the regulatory light chain of the myosins in a filament (Beach et al., 2017; Billington et al., 2015). This means that for a given NM2 filament observed in a cell, there can be a wide range of mechanical properties which cannot be estimated by fluorescence intensity alone. It follows that that the behavior of individual NM2 filaments within a cell are highly unlikely to be uniform. Thus, increases in myosin filament density following a cellular perturbation or movement do not necessarily imply that the mechanical output will scale proportionally. These factors highlight the complexity of dissecting NM2 filament function in vivo. In this respect, the in vitro single filament motility assay is a sensitive way to provide information on the regulation of filament mechanical output and to inform future cell biological studies.

1. 1. Beach JR
   2. Shao L
   3. Remmert K
   4. Li D
   5. Betzig E
   6. Hammer JA

   (2014) Nonmuscle myosin II isoforms coassemble in living cells

   Current Biology 24:1160–1166.

   https://doi.org/10.1016/j.cub.2014.03.071

   • PubMed
   • Google Scholar
2. 1. Beach JR
   2. Hammer JA

   (2015) Myosin II isoform co-assembly and differential regulation in mammalian systems

   Experimental Cell Research 334:2–9.

   https://doi.org/10.1016/j.yexcr.2015.01.012

   • PubMed
   • Google Scholar
3. 1. Beach JR
   2. Bruun KS
   3. Shao L
   4. Li D
   5. Swider Z
   6. Remmert K
   7. Zhang Y
   8. Conti MA
   9. Adelstein RS
   10. Rusan NM
   11. Betzig E
   12. Hammer JA

   (2017) Actin dynamics and competition for myosin monomer govern the sequential amplification of myosin filaments

   Nature Cell Biology 19:85–93.

   https://doi.org/10.1038/ncb3463

   • PubMed
   • Google Scholar
4. 1. Berg JS
   2. Powell BC
   3. Cheney RE

   (2001) A millennial myosin census

   Molecular Biology of the Cell 12:780–794.

   https://doi.org/10.1091/mbc.12.4.780

   • PubMed
   • Google Scholar
5. 1. Billington N
   2. Wang A
   3. Mao J
   4. Adelstein RS
   5. Sellers JR

   (2013) Characterization of three full-length human nonmuscle myosin II paralogs

   Journal of Biological Chemistry 288:33398–33410.

   https://doi.org/10.1074/jbc.M113.499848

   • PubMed
   • Google Scholar
6. 1. Billington N
   2. Beach JR
   3. Heissler SM
   4. Remmert K
   5. Guzik-Lendrum S
   6. Nagy A
   7. Takagi Y
   8. Shao L
   9. Li D
   10. Yang Y
   11. Zhang Y
   12. Barzik M
   13. Betzig E
   14. Hammer JA
   15. Sellers JR

   (2015) Myosin 18A coassembles with nonmuscle myosin 2 to form mixed bipolar filaments

   Current Biology 25:942–948.

   https://doi.org/10.1016/j.cub.2015.02.012

   • PubMed
   • Google Scholar
7. 1. Bottinelli R
   2. Reggiani C

   (2000) Human skeletal muscle fibres: molecular and functional diversity

   Progress in Biophysics and Molecular Biology 73:195–262.

   https://doi.org/10.1016/S0079-6107(00)00006-7

   • PubMed
   • Google Scholar
8. 1. Breitsprecher D
   2. Jaiswal R
   3. Bombardier JP
   4. Gould CJ
   5. Gelles J
   6. Goode BL

   (2012) Rocket launcher mechanism of collaborative actin assembly defined by single-molecule imaging

   Science 336:1164–1168.

   https://doi.org/10.1126/science.1218062

   • PubMed
   • Google Scholar
9. 1. Brizendine RK
   2. Alcala DB
   3. Carter MS
   4. Haldeman BD
   5. Facemyer KC
   6. Baker JE
   7. Cremo CR

   (2015) Velocities of unloaded muscle filaments are not limited by drag forces imposed by myosin cross-bridges

   PNAS 112:11235–11240.

   https://doi.org/10.1073/pnas.1510241112

   • PubMed
   • Google Scholar
10. 1. Brizendine RK
    2. Sheehy GG
    3. Alcala DB
    4. Novenschi SI
    5. Baker JE
    6. Cremo CR

    (2017) A mixed-kinetic model describes unloaded velocities of smooth, skeletal, and cardiac muscle myosin filaments in vitro

    Science Advances 3:eaao2267.

    https://doi.org/10.1126/sciadv.aao2267

    • PubMed
    • Google Scholar
11. 1. Bruun K
    2. Beach JR
    3. Heissler SM
    4. Remmert K
    5. Sellers JR
    6. Hammer JA

    (2017) Re-evaluating the roles of myosin 18Aα and F-actin in determining Golgi morphology

    Cytoskeleton 74:205–218.

    https://doi.org/10.1002/cm.21364

    • PubMed
    • Google Scholar
12. 1. Cohen C
    2. Lowey S
    3. Harrison RG
    4. Kendrick-Jones J
    5. Szent-Gyorgyi AG

    (1970) Segments from myosin rods

    Journal of Molecular Biology 47:605–609.

    https://doi.org/10.1016/0022-2836(70)90329-3

    • PubMed
    • Google Scholar
13. 1. Conti MA
    2. Even-Ram S
    3. Liu C
    4. Yamada KM
    5. Adelstein RS

    (2004) Defects in cell adhesion and the visceral endoderm following ablation of nonmuscle myosin heavy chain II-A in mice

    Journal of Biological Chemistry 279:41263–41266.

    https://doi.org/10.1074/jbc.C400352200

    • PubMed
    • Google Scholar
14. Book

    1. Conti MA
    2. Kawamoto S
    3. Adelstein RS

    (2008)

    Non-muscle myosin II

    In: Coluccio C, editors. Myosins: A Superfamily of Molecular Motors. Dordrecht: Springer. pp. 223–264.

    • Google Scholar
15. 1. Cuda G
    2. Pate E
    3. Cooke R
    4. Sellers JR

    (1997) In vitro actin filament sliding velocities produced by mixtures of different types of myosin

    Biophysical Journal 72:1767–1779.

    https://doi.org/10.1016/S0006-3495(97)78823-4

    • Google Scholar
16. 1. De La Cruz EM
    2. Wells AL
    3. Rosenfeld SS
    4. Ostap EM
    5. Sweeney HL

    (1999) The kinetic mechanism of myosin V

    PNAS 96:13726–13731.

    https://doi.org/10.1073/pnas.96.24.13726

    • PubMed
    • Google Scholar
17. 1. Diensthuber RP
    2. Müller M
    3. Heissler SM
    4. Taft MH
    5. Chizhov I
    6. Manstein DJ

    (2011) Phalloidin perturbs the interaction of human non-muscle myosin isoforms 2A and 2C1 with F-actin

    FEBS Letters 585:767–771.

    https://doi.org/10.1016/j.febslet.2011.01.042

    • PubMed
    • Google Scholar
18. 1. Ebrahim S
    2. Fujita T
    3. Millis BA
    4. Kozin E
    5. Ma X
    6. Kawamoto S
    7. Baird MA
    8. Davidson M
    9. Yonemura S
    10. Hisa Y
    11. Conti MA
    12. Adelstein RS
    13. Sakaguchi H
    14. Kachar B

    (2013) NMII forms a contractile transcellular sarcomeric network to regulate apical cell junctions and tissue geometry

    Current Biology 23:731–736.

    https://doi.org/10.1016/j.cub.2013.03.039

    • PubMed
    • Google Scholar
19. 1. Fenix AM
    2. Taneja N
    3. Buttler CA
    4. Lewis J
    5. Van Engelenburg SB
    6. Ohi R
    7. Burnette DT

    (2016) Expansion and concatenation of non-muscle myosin IIA filaments drive cellular contractile system formation during interphase and mitosis

    Molecular Biology of the Cell pp. 1465–1478.

    https://doi.org/10.1091/mbc.E15-10-0725

    • PubMed
    • Google Scholar
20. 1. Franke JD
    2. Dong F
    3. Rickoll WL
    4. Kelley MJ
    5. Kiehart DP

    (2005) Rod mutations associated with MYH9-related disorders disrupt nonmuscle myosin-IIA assembly

    Blood 105:161–169.

    https://doi.org/10.1182/blood-2004-06-2067

    • PubMed
    • Google Scholar
21. 1. Haldeman BD
    2. Brizendine RK
    3. Facemyer KC
    4. Baker JE
    5. Cremo CR

    (2014) The kinetics underlying the velocity of smooth muscle myosin filament sliding on actin filaments in vitro

    Journal of Biological Chemistry 289:21055–21070.

    https://doi.org/10.1074/jbc.M114.564740

    • PubMed
    • Google Scholar
22. 1. Hariadi RF
    2. Sommese RF
    3. Adhikari AS
    4. Taylor RE
    5. Sutton S
    6. Spudich JA
    7. Sivaramakrishnan S

    (2015) Mechanical coordination in motor ensembles revealed using engineered artificial myosin filaments

    Nature Nanotechnology 10:696–700.

    https://doi.org/10.1038/nnano.2015.132

    • PubMed
    • Google Scholar
23. 1. Harris DE
    2. Work SS
    3. Wright RK
    4. Alpert NR
    5. Warshaw DM

    (1994) Smooth, cardiac and skeletal muscle myosin force and motion generation assessed by cross-bridge mechanical interactions in vitro

    Journal of Muscle Research and Cell Motility 15:11–19.

    https://doi.org/10.1007/BF00123828

    • PubMed
    • Google Scholar
24. 1. Heissler SM
    2. Manstein DJ

    (2011) Comparative kinetic and functional characterization of the motor domains of human nonmuscle myosin-2C isoforms

    Journal of Biological Chemistry 286:21191–21202.

    https://doi.org/10.1074/jbc.M110.212290

    • PubMed
    • Google Scholar
25. 1. Heissler SM
    2. Manstein DJ

    (2013) Nonmuscle myosin-2: mix and match

    Cellular and Molecular Life Sciences 70:1–21.

    https://doi.org/10.1007/s00018-012-1002-9

    • PubMed
    • Google Scholar
26. 1. Heissler SM
    2. Sellers JR

    (2016) Various themes of myosin regulation

    Journal of Molecular Biology 428:1927–1946.

    https://doi.org/10.1016/j.jmb.2016.01.022

    • PubMed
    • Google Scholar
27. 1. Highsmith S
    2. Duignan K
    3. Cooke R
    4. Cohen J

    (1996) Osmotic pressure probe of actin-myosin hydration changes during ATP hydrolysis

    Biophysical Journal 70:2830–2837.

    https://doi.org/10.1016/S0006-3495(96)79852-1

    • PubMed
    • Google Scholar
28. 1. Hu S
    2. Dasbiswas K
    3. Guo Z
    4. Tee YH
    5. Thiagarajan V
    6. Hersen P
    7. Chew TL
    8. Safran SA
    9. Zaidel-Bar R
    10. Bershadsky AD

    (2017) Long-range self-organization of cytoskeletal myosin II filament stacks

    Nature Cell Biology 19:133–141.

    https://doi.org/10.1038/ncb3466

    • PubMed
    • Google Scholar
29. 1. Joo E
    2. Surka MC
    3. Trimble WS

    (2007) Mammalian SEPT2 is required for scaffolding nonmuscle myosin II and its kinases

    Developmental Cell 13:677–690.

    https://doi.org/10.1016/j.devcel.2007.09.001

    • PubMed
    • Google Scholar
30. 1. Jung HS
    2. Burgess SA
    3. Billington N
    4. Colegrave M
    5. Patel H
    6. Chalovich JM
    7. Chantler PD
    8. Knight PJ

    (2008) Conservation of the regulated structure of folded myosin 2 in species separated by at least 600 million years of independent evolution

    PNAS 105:6022–6026.

    https://doi.org/10.1073/pnas.0707846105

    • PubMed
    • Google Scholar
31. 1. Kalwarczyk T
    2. Ziebacz N
    3. Bielejewska A
    4. Zaboklicka E
    5. Koynov K
    6. Szymański J
    7. Wilk A
    8. Patkowski A
    9. Gapiński J
    10. Butt HJ
    11. Hołyst R

    (2011) Comparative analysis of viscosity of complex liquids and cytoplasm of mammalian cells at the nanoscale

    Nano Letters 11:2157–2163.

    https://doi.org/10.1021/nl2008218

    • PubMed
    • Google Scholar
32. 1. Kengyel A
    2. Wolf WA
    3. Chisholm RL
    4. Sellers JR

    (2010) Nonmuscle myosin IIA with a GFP fused to the N-terminus of the regulatory light chain is regulated normally

    Journal of Muscle Research and Cell Motility 31:163–170.

    https://doi.org/10.1007/s10974-010-9220-y

    • PubMed
    • Google Scholar
33. 1. Kim KY
    2. Kovács M
    3. Kawamoto S
    4. Sellers JR
    5. Adelstein RS

    (2005) Disease-associated mutations and alternative splicing alter the enzymatic and motile activity of nonmuscle myosins II-B and II-C

    Journal of Biological Chemistry 280:22769–22775.

    https://doi.org/10.1074/jbc.M503488200

    • PubMed
    • Google Scholar
34. 1. Kovács M
    2. Wang F
    3. Hu A
    4. Zhang Y
    5. Sellers JR

    (2003) Functional divergence of human cytoplasmic myosin II: kinetic characterization of the non-muscle IIA isoform

    The Journal of Biological Chemistry 278:38132–38140.

    https://doi.org/10.1074/jbc.M305453200

    • PubMed
    • Google Scholar
35. 1. Kovács M
    2. Thirumurugan K
    3. Knight PJ
    4. Sellers JR

    (2007) Load-dependent mechanism of nonmuscle myosin 2

    PNAS 104:9994–9999.

    https://doi.org/10.1073/pnas.0701181104

    • PubMed
    • Google Scholar
36. 1. Kron SJ
    2. Spudich JA

    (1986) Fluorescent actin filaments move on myosin fixed to a glass surface

    PNAS 83:6272–6276.

    https://doi.org/10.1073/pnas.83.17.6272

    • PubMed
    • Google Scholar
37. 1. Ma X
    2. Bao J
    3. Adelstein RS

    (2007) Loss of cell adhesion causes hydrocephalus in nonmuscle myosin II-B-ablated and mutated mice

    Molecular Biology of the Cell 18:2305–2312.

    https://doi.org/10.1091/mbc.E07-01-0073

    • PubMed
    • Google Scholar
38. 1. Ma X
    2. Adelstein RS

    (2014) The role of vertebrate nonmuscle Myosin II in development and human disease

    BioArchitecture 4:88–102.

    https://doi.org/10.4161/bioa.29766

    • PubMed
    • Google Scholar
39. 1. Maddox AS
    2. Lewellyn L
    3. Desai A
    4. Oegema K

    (2007) Anillin and the septins promote asymmetric ingression of the cytokinetic furrow

    Developmental Cell 12:827–835.

    https://doi.org/10.1016/j.devcel.2007.02.018

    • PubMed
    • Google Scholar
40. 1. Murrell MP
    2. Gardel ML

    (2012) F-actin buckling coordinates contractility and severing in a biomimetic actomyosin cortex

    PNAS 109:20820–20825.

    https://doi.org/10.1073/pnas.1214753109

    • PubMed
    • Google Scholar
41. 1. Nagy A
    2. Takagi Y
    3. Billington N
    4. Sun SA
    5. Hong DK
    6. Homsher E
    7. Wang A
    8. Sellers JR

    (2013) Kinetic characterization of nonmuscle myosin IIb at the single molecule level

    Journal of Biological Chemistry 288:709–722.

    https://doi.org/10.1074/jbc.M112.424671

    • PubMed
    • Google Scholar
42. 1. Niederman R
    2. Pollard TD

    (1975) Human platelet myosin. II. In vitro assembly and structure of myosin filaments

    The Journal of Cell Biology 67:72–92.

    https://doi.org/10.1083/jcb.67.1.72

    • PubMed
    • Google Scholar
43. 1. Pardee JD
    2. Spudich JA

    (1982) Purification of muscle actin

    Methods in Cell Biology 24:271–289.

    https://doi.org/10.1016/S0091-679X(08)60661-5

    • PubMed
    • Google Scholar
44. 1. Piekny AJ
    2. Glotzer M

    (2008) Anillin is a scaffold protein that links RhoA, actin, and myosin during cytokinesis

    Current Biology 18:30–36.

    https://doi.org/10.1016/j.cub.2007.11.068

    • PubMed
    • Google Scholar
45. 1. Rosenfeld SS
    2. Xing J
    3. Chen LQ
    4. Sweeney HL

    (2003) Myosin IIb is unconventionally conventional

    Journal of Biological Chemistry 278:27449–27455.

    https://doi.org/10.1074/jbc.M302555200

    • PubMed
    • Google Scholar
46. Book

    1. Sellers JR

    (2006) In vitro motility assay with actin

    In: Celis J. E, editors. Cell Biology. A Laboratory Handbook, Vol. 2. Burligton: Academic Press. pp. 387–392.

    https://doi.org/10.1016/B978-012164730-8/50121-0

    • Google Scholar
47. 1. Shutova MS
    2. Spessott WA
    3. Giraudo CG
    4. Svitkina T

    (2014) Endogenous species of mammalian nonmuscle myosin IIA and IIB include activated monomers and heteropolymers

    Current Biology 24:1958–1968.

    https://doi.org/10.1016/j.cub.2014.07.070

    • PubMed
    • Google Scholar
48. 1. Shutova MS
    2. Asokan SB
    3. Talwar S
    4. Assoian RK
    5. Bear JE
    6. Svitkina TM

    (2017) Self-sorting of nonmuscle myosins IIA and IIB polarizes the cytoskeleton and modulates cell motility

    The Journal of Cell Biology 216:jcb.201705167.

    https://doi.org/10.1083/jcb.201705167

    • PubMed
    • Google Scholar
49. 1. Verkhovsky AB
    2. Svitkina TM
    3. Borisy GG

    (1995) Myosin II filament assemblies in the active lamella of fibroblasts: their morphogenesis and role in the formation of actin filament bundles

    The Journal of Cell Biology 131:989–1002.

    https://doi.org/10.1083/jcb.131.4.989

    • PubMed
    • Google Scholar
50. 1. Vicente-Manzanares M
    2. Ma X
    3. Adelstein RS
    4. Horwitz AR

    (2009) Non-muscle myosin II takes centre stage in cell adhesion and migration

    Nature Reviews Molecular Cell Biology 10:778–790.

    https://doi.org/10.1038/nrm2786

    • PubMed
    • Google Scholar
51. 1. Wang F
    2. Harvey EV
    3. Conti MA
    4. Wei D
    5. Sellers JR

    (2000) A conserved negatively charged amino acid modulates function in human nonmuscle myosin IIA

    Biochemistry 39:5555–5560.

    https://doi.org/10.1021/bi000133x

    • Google Scholar
52. 1. Wang F
    2. Kovacs M
    3. Hu A
    4. Limouze J
    5. Harvey EV
    6. Sellers JR

    (2003) Kinetic mechanism of non-muscle myosin IIB: functional adaptations for tension generation and maintenance

    The Journal of Biological Chemistry 278:27439–27448.

    https://doi.org/10.1074/jbc.M302510200

    • PubMed
    • Google Scholar
53. 1. Wang A
    2. Ma X
    3. Conti MA
    4. Liu C
    5. Kawamoto S
    6. Adelstein RS

    (2010) Nonmuscle myosin II isoform and domain specificity during early mouse development

    PNAS 107:14645–14650.

    https://doi.org/10.1073/pnas.1004023107

    • PubMed
    • Google Scholar
54. 1. Wang A
    2. Ma X
    3. Conti MA
    4. Adelstein RS

    (2011) Distinct and redundant roles of the non-muscle myosin II isoforms and functional domains

    Biochemical Society Transactions 39:1131–1135.

    https://doi.org/10.1042/BST0391131

    • PubMed
    • Google Scholar
55. 1. Wendt T
    2. Taylor D
    3. Trybus KM
    4. Taylor K

    (2001) Three-dimensional image reconstruction of dephosphorylated smooth muscle heavy meromyosin reveals asymmetry in the interaction between myosin heads and placement of subfragment 2

    PNAS 98:4361–4366.

    https://doi.org/10.1073/pnas.071051098

    • PubMed
    • Google Scholar

Author details

1. Luca Melli

   Cell Biology and Physiology Center, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, United States

   Contribution

   Conceptualization, Resources, Software, Formal analysis, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing

   For correspondence

   luca.melli@nih.gov

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-8733-047X

2. Neil Billington

   Cell Biology and Physiology Center, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, United States

   Contribution

   Conceptualization, Software, Formal analysis, Investigation, Visualization, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2306-0228

3. Sara A Sun

   Cell Biology and Physiology Center, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, United States

   Contribution

   Methodology

   Competing interests

   No competing interests declared

4. Jonathan E Bird

   Laboratory of Molecular Genetics, National Institute on Deafness and Other Communication Disorders, National Institutes of Health, Bethesda, United States

   Present address

   Department of Pharmacology and Therapeutics, University of Florida College of Medicine, Gainesville, United States

   Contribution

   Resources, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5531-8794

5. Attila Nagy

   Vaccine Production Program Laboratory, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Gaithersburg, United States

   Contribution

   Resources, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

6. Thomas B Friedman

   Laboratory of Molecular Genetics, National Institute on Deafness and Other Communication Disorders, National Institutes of Health, Bethesda, United States

   Contribution

   Resources, Supervision, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-4614-6630

7. Yasuharu Takagi

   Cell Biology and Physiology Center, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, United States

   Contribution

   Resources, Formal analysis, Validation, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared

8. James R Sellers

   Cell Biology and Physiology Center, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, United States

   Contribution

   Conceptualization, Data curation, Supervision, Funding acquisition, Investigation, Writing—original draft, Project administration, Writing—review and editing

   For correspondence

   sellersj@nhlbi.nih.gov

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-6296-564X

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