Regulation and Expression of Metazoan Unconventional Myosins

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1 Regulation and Expression of Metazoan Unconventional Myosins Anna M. Sokac 1 and William M. Bement 1,2 Program in Cellular and Molecular Biology 1 and Department of Zoology, 2 University of Wisconsin, Madison, Wisconsin Unconventional myosins are molecular motors that convert adenosine triphosphate (ATP) hydrolysis into movement along actin filaments. On the basis of primary structure analysis, these myosins are represented by at least 15 distinct classes (classes 1 and 3 16), each of which is presumed to play a specific cellular role. However, in contrast to the conventional myosins-2, which drive muscle contraction and cytokinesis and have been studied intensively for many years in both uni- and multicellular organisms, unconventional myosins have only been subject to analysis in metazoan systems for a short time. Here we critically review what is known about unconventional myosin regulation, function, and expression. Several points emerge from this analysis. First, in spite of the high relative conservation of motor domains among the myosin classes, significant differences are found in biochemical and enzymatic properties of these motor domains. Second, the idea that characteristic distributions of unconventional myosins are solely dependent on the myosin tail domain is almost certainly an oversimplification. Third, the notion that most unconventional myosins function as transport motors for membranous organelles is challenged by recent data. Finally, we present a scheme that clarifies relationships between various modes of myosin regulation. KEY WORDS: Myosin, f-actin, Mechanochemistry, Gene expression, Phosphorylation, Myosin-binding proteins Academic Press. I. Introduction Myosins are proteins that convert adenosine triphosphate (ATP) hydrolysis into movement along actin filaments. Although first recognized as a 197 International Review of Cytology, Vol. 200 Copyright 2000 by Academic Press /00 $35.00 All rights of reproduction in any form reserved.

2 198 ANNA M. SOKAC AND WILLIAM M. BEMENT major component of skeletal muscle, where they drive sarcomere contraction, it is now clear that the myosins represent a vast and complex family of proteins. On the basis of historical convenience, members of the myosin gene family are referred to as being conventional or unconventional. Conventional myosins are the myosins-2 two-headed, filament-forming myosins that have been studied intensively for many years due to their abundance in muscle and their role in driving muscle contraction. (Myosin classes traditionally have been designated by Roman numerals, e.g., myosins-ii. However, because there are at least 16 and possibly 19 or more different classes, we prefer to use Arabic numerals, e.g., myosin-16 instead of myosin-xvi.) Myosin-2 is also found in nonmuscle cells where it performs a variety of functions in conjunction with filamentous actin (f-actin), such as facilitating cell crawling ( Jay et al., 1995; Svitkina et al., 1997) and providing contractile force for cytokinesis (Mabuchi and Okuno, 1977; Shelton et al., 1999), embryogenesis (Young et al., 1993, 1999; Shelton et al., 1999), and wound healing (Bement et al., 1993; Brock et al., 1996). Myosins from all other classes classes 1 and 3 16 are considered unconventional. The functions of the unconventional myosins are not as well-understood as those of the conventional myosins, and their biochemistry and regulation are, likewise, poorly characterized. However, there is intense interest in the unconventional myosins, which is reflected in the explosion of publications describing the characterization, expression, regulation, and possible functions of these motors. The interest stems from (1) the recognition that unconventional myosins comprise part of a large and complex gene family (see Section II.B), (2) the growing awareness of the potential roles of unconventional myosins in intracellular transport (see Section II.E), and (3) the discovery that defects in several different unconventional myosins underlie a number of human and murine genetic diseases (see Section II.E). In this review, we consolidate what is presently known about the regulation and expression of unconventional myosins in vertebrates. In so doing, we frequently discuss lessons learned from conventional myosins as well as lessons learned from protozoan unconventional myosins, because these systems provide context essential for understanding the vertebrate unconventional myosins. We also discuss putative functions of unconventional myosins because function is intimately intertwined with regulation. It is our goal to provide the reader with both a basic background in the biology of unconventional myosins as well as an understanding of how the activity of these fascinating molecules may be controlled in metazoan systems.

3 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 199 II. The Myosin Superfamily A. General Structural Features of Myosins Myosins are multimeric, composed of one or more heavy chains, each of which is associated with one or more light chains (with the predicted exception of myosins-14; see Sections II.C and III.B). The heavy chains comprise three functional parts: the head or motor domain, the neck, and the tail (Fig. 1). The amino-terminal motor domain of myosins couples ATP binding and hydrolysis to f-actin binding and translocation. This domain is highly conserved and essentially defines a myosin as a myosin. In almost all myosins, the motor domain is followed by a central neck region where light chains bind. The neck contains one or more IQ motifs, 23 amino acid repeats containing the sequence IQXXXRGXXXRK that are responsible for light-chain binding (Cheney and Mooseker, 1992). Most, but not all, of the myosin light chains are either calmodulin or members of the calmodulin superfamily. In addition to binding light chains, and perhaps as a result of it, the neck works in concert with the motor domain to produce movement. Beyond the neck is the highly variable carboxyterminal tail domain that mediates myosin interactions with target molecules and, for those myosins that dimerize or form filaments, with other myosin heavy chains. Several motifs have been identified in myosin tails that direct protein protein interactions. Myosins-2 provide the classic example in that their tails are composed almost exclusively of -helical coiled coil, which drives dimerization of the heavy chains (Fig. 2). A number of the unconventional myosin heavy chains also contain -helical coiled coil in their tails, including myosins 5 8, 10 12, and 17; consequently, they are predicted to form homodimers. To date, however, myosin-5 is the only unconventional myosin that has been directly shown to be a dimer (Cheney et al., 1993). Many unconventional myosin tails contain protein motifs first FIG. 1 Schematic diagram showing the general primary structure of most myosins: the aminoterminal head, followed by the neck, followed by the carboxy-terminal tail.

4 200 ANNA M. SOKAC AND WILLIAM M. BEMENT FIG. 2 Schematic diagram showing the primary structure of the myosin-2 heavy chain. The tail is composed almost entirely of coiled coil, which drives dimerization of the heavy chain. identified in other proteins, including talin homology domains and pleckstrin homology (PH) domains (Mermall et al., 1998). Four distinct domains that are found in several unconventional myosin tails have been described and designated as TH or MyTH (for tail homology or myosin tail homology, respectively) domains 1 4 (Hammer, 1994; Bähler et al., 1994; Hasson and Mooseker, 1996; Mermall et al., 1998). The MyTH1 domain usually is found in the amino-terminal portion of the tail, is extremely basic, and is thought to be at least partly responsible for the interaction of myosin with acidic phospholipids. The MyTH2 domain usually is downstream of the MyTH1 domain and is rich in proline and glycine residues. In the myosins-1 from Acanthamoeba, the MyTH2 domain has been shown to comprise a second actin-binding domain that, in contrast to the actin-binding site in the head, is ATP-insensitive (Lynch et al., 1986). Although MyTH2 domains are found in myosins-1 from the metazoa, efforts to demonstrate f-actin binding by metazoan MyTH2 domains so far have been unsuccessful (e.g., Stöffler and Bähler, 1998). The MyTH3 domains are the same as src homology 3 (SH3) domains that are found in many proteins involved in signal transduction and are known to bind to proline-rich regions of other proteins (Hammer, 1994). The function of the MyTH4 domain presently is unknown. In addition to protein protein interaction motifs, some myosin tails contain catalytic motifs, such as the GTPase-activating protein (GAP) motif, and, in an extreme case, what appears to be an entire enzyme (chitin synthase, see following discussion). Although these diverse structures have been identified in myosin tails, there is little information regarding their cellular significance when linked to a myosin motor domain. One popular assumption is that the tail domain largely defines myosin localization and function. However, as will be discussed, this is probably an oversimplification because, in spite of their conserved nature, the heads display considerable diversity in structure and biochemistry that is quite likely to influence function.

5 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 201 B. Molecular Phylogeny The high conservation of the myosin head domain has permitted the identification of multiple unconventional myosins by both low-stringency hybridization screening (Titus et al., 1989) and degenerate polymerase chain reaction (PCR) (Sherr et al., 1993; Bement et al., 1994a). As a result, the acquisition of sequence information for unconventional myosins has far outstripped the acquisition of biochemical and functional information for these proteins. Nevertheless, sequence information alone can be quite useful when used in conjunction with phylogenetic analyses that permit the classification of myosins on the basis of evolutionary relatedness. The notion is that members of a given class will share similar properties and, perhaps, similar functions within a cell. Class assignments are determined by a distance matrix methodology in which motor domain sequences are aligned progressively by pairwise comparisons. Distance scores are calculated for each pair and a distance matrix is created. Although this analysis does not identify a common ancestral myosin, it does demonstrate that phylogenetic classes are equally unrelated (Goodson and Spudich, 1993; Mooseker and Cheney, 1995; Mermall et al., 1998). That is, the classes grouped together as unconventional myosins (classes 1 and 3 16) are no more closely related to each other than to the conventional myosins (class 2). In those cases where phylogenetic data are bolstered by detailed biochemical analysis, it appears that, within a given class, structural, functional, and biochemical character is indeed conserved to some extent. The heads and necks of members of a particular myosin class contain common regulatory elements, including phosphorylation sites and type(s) of light chains bound. Tail sequences are class-specific in that the tails of class members contain common functional domains. Thus, phylogenetic relationships can help to predict the structure, biochemistry, and possibly function of newly discovered myosins. Phylogenetic analysis also facilitates communication within the field. This is not trivial as the myosin literature is muddled by a complex and inconsistent naming scheme. Whereas a given group of investigators may adopt a consistent, internally logical nomenclature, such as those based on order of identification, chromosomal location, or mutant phenotype, when considered within the context of the field as a whole, such names can be extremely bewildering (Table I). For example, budding yeast myosins myo2 and myo4 are class 5 myosins, myo1 is a class 2 myosin, and myo3 and myo5 are class 1 myosins. Phylogenetic classification of myosins can provide a common and descriptive nomenclature. Throughout this review, myosins originally named according to phenotype or by other naming systems are followed by the corresponding phylogenetic class in parentheses to facilitate recognition.

6 202 ANNA M. SOKAC AND WILLIAM M. BEMENT TABLE I A Myosin by Any Other Name Given name Organism Class TgM-A, 1 -B, 1 -C 1 Toxoplasma gondii 14 HMWM 2 Acanthamoeba 4 myoa, 3 -B, 3 -C, 3 -D, 4 -E, 5 -F, 6 -K 6 Dictyostelium 1 myoi 6 Dictyostelium 7 myoj 6 Dictyostelium 5/11 myoa 7 Aspergillus nidulans 1 Myo1 8 Saccharomyces cerevisiae 2 Myo2, S. cerevisiae 5 Myo3, S. cerevisiae 1 MYO22 13 Anemia phyllitidis 8 MYO15 13 A. phyllitidis 11 ATM1, Arabidopsis 8 MYA1, Arabidopsis 11 ninac 18 Drosophila 3 95F 19 Drosophila 6 35B 20 Drosophila 7 HUM-1, Caenorhabditis elegans 1 HUM-2 21 C. elegans 5 HUM-3 21 C. elegans 6 HUM-4 21 C. elegans 12 HUM-6 21 C. elegans 7 dilute 22 Mouse 5 myr1, 23-2, 24-3, Rat 1 myr5 27 Rat 9 myr6 28 Rat 5 myr7 29 Rat 9 1 Heintzelman and Schwartzman, Horowitz and Hammer, Titus et al., Jung et al., Urrutia et al., Titus et al., McGoldrick et al., Watts et al., Johnston et al., Haarer et al., Goodson and Spudich, Geli and Riezman, Moepps et al., , 15 Knight and Kendrick-Jones, Kinkema and Schiefelbein, Kinkema and Schiefelbein, Montell and Rubin, Kellerman and Miller, Chen et al., Baker and Titus, Mercer et al., Ruppert et al., Ruppert et al., Stöffler et al., Bähler et al., Reinhard et al., Zhao et al., Chieregatti et al., To date, phylogenetic analyses have defined at least 15 distinct classes of myosins (Fig. 3), and analysis of other published sequences suggests that this number can now be increased to 19. Comparison of a PCR fragment obtained in a screen for Xenopus unconventional myosins to a human myosin-like protein (MLP) found in the database revealed the existence of a class 16 (Sokac and Bement, 1998; Sokac and Bement, manuscript submitted). Two myosins from fungi that are fusions of a myosin motor domain with chitin synthase define class 17 (Baker and Titus, 1998). A

7 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 203 FIG. 3 An unrooted phylogenetic tree of the myosin superfamily. Myosin head sequences were aligned according to the default settings of Clustal X. The tree was generated using Drawtree from the Phylip software package. Sequence divergence is proportional to branch length. Myosin classes are shown in bold Arabic numbers. Myosin-1 subclasses are shown parenthetically. Asterisks indicate that only a very short stretch of head sequence ( 30 residues) is available for that myosin. These short sequences were omitted from alignment and tree generation and are instead listed with their closest myosin relative. Note that Dictyostelium myoj, previously classified as a myosin-11, did not group within class 11 in any of a number of alignment tree generation attempts. Likewise, Xenopus myosin-14, although presumed to be a member of class 14 due to its small transcript size ( 3 kb) and consistency in falling near myosins-14 in all alignment tree generation attempts, did not group directly with the myosins-14. Accession numbers for myosin sequences are as follows: Acanthamoeba HMWM ; Acetabularia myo , myo ; Anemia MYO ; Arabidopsis ATM ; MYA ; bovine M ; C. elegans HUM ; chicken BB M , SKMM , SMM ; Dictyostelium MHC , myoj ; Drosophila ninac , 95F ; human M-1C , M-1D , M-1B , NMMHCA , M , M-7A , M-9A , MLP ; Limulus M ; mouse dilute , M-7A , MYO ; Rana M ; rat myr , myr , myr , myr ; S. cerevisiae Myo ; T. gondii M-A , M-B ; Xenopus NMM2A , NMM2B Accession numbers for remaining Xenopus myosins are pending.

8 204 ANNA M. SOKAC AND WILLIAM M. BEMENT myosin identified in a PCR screen of Tetrahymena (Garces and Gavin, 1998) defines class 18. Last, a subtractive PCR screen has identified Dictyostelium myom (Schwarz et al., 1999), which defines myosin class 19. C. Overview of Unconventional Myosin Classes Each of the 16 established myosin classes significantly differs from the other classes at the primary structure level and, where studied, at the biochemical and cellular levels as well. A comprehensive discussion of each of the classes is well beyond the scope of this review (Mooseker and Cheney, 1995; Coluccio, 1997; Mermall et al., 1998). However, the basic features of each class must be discussed in order for the similarities as well as differences in regulation to be understood. 1. Myosins-1 Myosins-1 were first discovered in Acanthamoeba by preparative biochemistry (Pollard and Korn, 1973). The heavy chains are comparatively small ( kda) and bind 1 6 light chains. In vertebrates, all of the myosin- 1 light chains identified so far are calmodulin. Myosin-1 heavy chains are thought to be monomeric based on the absence of significant stretches of coiled coil in the primary structures. However, chicken brush border myosin-1 (Coluccio and Bretscher, 1987; Conzelman and Mooseker, 1987), rat liver myr-1 (Coluccio and Conaty, 1993), adrenal myosin-1 (Wagner and Molitoris, 1997), and rat myr3 (Stöffler and Bähler, 1998) are all capable of cross-linking purified f-actin. Although this bundling could be attributed to a second actin-binding site in the tail domain of these myosins, as is found in myosins-1 from Acanthamoeba (see previous discussion), there is no evidence for such a domain in any of these particular myosins, suggesting that under some conditions these myosins may be capable of forming multimers. The myosins-1 are the most diverse of any of the unconventional myosin classes and are represented by at least four, phylogenetically distinct subclasses (Figs. 3 and 4). Subclass 1 contains the amoeboid myosins-1, represented by myosins-1 from Acanthamoeba, Dictyostelium, Aspergillus, budding yeast, and several vertebrate myosins-1, including human myosin- 1c, human myosin-1d (Bement et al., 1994b), and rat myr3 (Stöffler et al., 1995). These myosins have MyTH1, MyTH2, and SH3 domains in their tails. In vertebrates, subclass 1 myosins bind to a single calmodulin light chain via the single IQ motif in their necks (Stöffler et al., 1995). The tail domains of some of these myosins have been directly demonstrated to interact with acidic phospholipids (Doberstein and Pollard, 1992), appar-

9 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 205 FIG. 4 Schematic diagram showing the primary structure of the heavy chains from the four subclasses of myosins-1. Abbreviations: MyTH1, myosin tail homology 1 domain; MyTH2, myosin tail homology 2 domain; SH3, src homology 3 domain. ently by way of the MyTH1 domain. Subclass 2 is represented by brush border myosins-1 (Hoshimaru and Nakanishi, 1987; Garcia et al., 1989), rat myr1 (Ruppert et al., 1993), and mouse myosin-1 (Sherr et al., 1993). These myosins have varying numbers of IQ motifs in their necks and, consequently, bind 3 6 calmodulin light chains (Wolenski, 1995; Coluccio, 1997). Brush border myosins-1 have been shown to associate with membranes via their tails (Hayden et al., 1990), as has myr1 (Ruppert et al., 1995), although no precise binding region has been mapped. Subclass 3 is represented by the myosin-1 isoforms that have been found in a variety of vertebrates (Barylko et al., 1992; Sherr et al., 1993; Reizes et al., 1994; Solc et al., 1994). Subclass 3 also contains Drosophila myosin-1b (Morgan et al., 1994) and rat myr2 (Ruppert et al., 1995). These myosins bind three calmodulin light chains via the three IQ motifs in their necks (Barylko et al., 1992). Subclass 4 comprises mouse myosin-1 (Sherr et al., 1993), rat

10 206 ANNA M. SOKAC AND WILLIAM M. BEMENT myr4 (Bähler et al., 1994), Drosophila myosin-1a (Morgan et al., 1994), and Caenorhabditis elegans myosin-1a (Baker and Titus, 1997). These myosins are thought to bind at least two calmodulins via the two IQ motifs in the neck and may bind a third in a Ca 2 -dependent manner via the tail (Bähler et al., 1994). 2. Myosins-3 Myosins-3 were first identified as a result of analysis of Drosophila mutants deficient in phototransduction (ninac; Montell and Rubin, 1988). Myosin- 3 heavy chains are highly unusual in that the amino terminus encodes a kinase domain (Fig. 5). The heavy chain is 174 or 135 kda, depending on the splice form. The monomeric heavy chain binds two calmodulin light chains via the IQ motifs in its neck (Porter et al., 1993). A 122-kDa myosin-3 has also been identified in the horseshoe crab, Limulus (Battelle et al., 1998). 3. Myosin-4 This myosin was identified by hybridization screening (Horowitz and Hammer, 1990). To date, myosin-4 has been found only in Acanthamoeba castellanii. The heavy chain is 180 kda and is assumed to be monomeric based on the lack of any significant coiled coil within the tail (Fig. 5). Its light chains have not been characterized, but it does have a single, putative, light-chain-binding site in the neck. The myosin-4 tail has a SH3 domain at the carboxy terminus and a MyTH4 domain amino-terminal to that (Mermall et al., 1998). 4. Myosins-5 Myosins-5 were first detected as an abundant chicken brain protein (Larson et al., 1990) and subsequently identified as the gene product of the mouse dilute locus (Mercer et al., 1991). The 212-kDa heavy chains form a dimer by way of coiled-coil regions in their tails (Fig. 5; Cheney et al., 1993). The neck region is long and contains six IQ motifs (Espreafico et al., 1992). As purified, the heavy chain is associated with four calmodulin light chains (Cheney et al., 1993) as well as a 23- and a 17-kDa light chain. In addition, the tail binds an 8-kDa light chain, which has been identified as a light chain for dynein (Benashski et al., 1997). The tail also contains a canoe/ AF-6 domain. These domains are found in the mammalian AF-6 proteins and in Drosophila canoe, both of which have been shown to interact with the small GTPase, ras, in other cell types (Kuriyama et al., 1996).

11 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 207 FIG. 5 Schematic diagram showing the primary structure of the heavy chains from myosins- 3 through myosins-15. The? indicates myosin regions for which the sequence is presently unavailable. Abbreviations: MyTH4, myosin tail homology 4 domain; rho-gap, rho GTPaseactivating protein; SH3, src homology 3 domain; PH, pleckstrin homology domain.

12 208 ANNA M. SOKAC AND WILLIAM M. BEMENT 5. Myosins-6 Myosins-6 were first identified by a screen for Drosophila actin-binding proteins (Miller et al., 1989). These subsequently were shown by sequence analysis to represent an unconventional myosin (Kellerman and Miller, 1992) and by genetic analysis to underlie the Snell s Waltzer phenotype in mice (Avraham et al., 1995). The heavy chains are kda and are thought to be dimeric on the basis of the presence of coiled coil in the tail, just past the neck (Fig. 5). Each of the heavy chains binds to a calmodulin light chain presumably via the single IQ motif in their necks (Hasson and Mooseker, 1994). 6. Myosins-7 The myosins-7 were first identified as partial sequences from Drosophila (Chen et al., 1991) and humans (Bement et al., 1994a). These subsequently were identified as underlying the Shaker-1 deafness phenotype in mice (Gibson et al., 1995) and Usher syndrome in humans (Weil et al., 1995). The myosin-7 heavy chains are large (250 kda or larger) and probably dimeric (Fig. 5). Each heavy chain has five IQ motifs in the neck, two MyTH4 domains in the tail, and two talin homology domains in the tail (Chen et al., 1996; talin is a cytoskeletal protein found at focal adhesions). 7. Myosins-8 Myosin-8 was identified in Arabidopsis by PCR (Knight and Kendrick- Jones, 1993) and has not been found in any animal systems to date. The heavy chain is predicted to be 131 kda and has three putative light-chainbinding sites. A region of coiled coil is found in the tail suggesting that the heavy chain may dimerize (Fig. 5). 8. Myosins-9 Myosins-9 were first identified by a PCR screen for unconventional myosins (Bement et al., 1994a). The heavy chains are 230 or 300 kda, depending on the isoform (Chieregatti et al., 1998), and are predicted to be monomeric (Fig. 5). The heavy chain has four light-chain-binding sites in the neck and a tail structure that is suggestive of a role in intracellular signaling (Reinhard et al., 1995; Wirth et al., 1996). That is, there is a putative zinc-binding domain and a rho-gtpase-activating (rhogap) domain in the tail. 9. Myosins-10 Myosin-10 was first identified in the bullfrog by a PCR screen for unconventional myosins (Solc et al., 1994). The complete primary structure has been

13 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 209 reported only for the cow (accession number u55042), but preliminary reports indicate that it is expressed in a variety of vertebrates (Oliver et al., 1996; Sokac and Bement, 1998). The heavy chain of myosin-10 is predicted to be 230 kda and to dimerize via a short region of coiled coil in the tail (Fig. 5). The neck has three putative light-chain-binding sites, while the tail has a MyTH4 domain, a talin homology domain, and several PH domains. 10. Myosins-11 Like myosin-8, myosins-11 were first identified by PCR in Arabidopsis (Kinkema and Schiefelbein, 1994). Unlike myosin-8, myosin-11 has been found in another system; specifically, Dictyostelium, where it is referred to as MyoJ (Peterson et al., 1996). The heavy chain is 170 kda, has six light-chain-binding sites, and is predicted to be dimeric on the basis of the presence of coiled coil in the tail (Fig. 5). 11. Myosins-12 Myosin-12 was identified in a PCR screen for C. elegans unconventional myosins (Baker and Titus, 1997). The heavy chain is predicted to be very large (307 kda) and to bind two light chains. Its tail structure is unusual in that whereas there is a small amount of coiled coil, indicating that the heavy chain may dimerize, it is well beyond the neck instead of being immediately carboxy-terminal to it, as found in most other myosins (see previous discussion). Like myosins-7, the myosin-12 heavy-chain tail has two MyTH4 domains (Fig. 5; Baker and Titus, 1997). 12. Myosin-13 Myosin-13 was identified in the plant Acetabularia and has yet to be identified in any other system. The heavy chain is relatively small (125 kda) and probably monomeric and is predicted to bind seven light chains (Fig. 5). 13. Myosins-14 Myosins-14 were identified by a PCR screen for unconventional myosins in Toxoplasma gondii (Heintzelman and Schwartzman, 1997), a protozoan parasite, although preliminary studies indicate that they may be expressed in animal systems as well (Sokac and Bement, manuscript submitted). The heavy chains of these myosins are unusually small, ranging from 90 to 110 kda. In fact, they are scarcely more than a head and an extremely

14 210 ANNA M. SOKAC AND WILLIAM M. BEMENT diminutive tail, with no neck or light-chain-binding sites (Fig. 5; Heintzelman and Schwartzman, 1997). 14. Myosins-15 Myosins-15 were discovered simultaneously in humans and mice by genetic and molecular genetic approaches to identify the genes underlying Shaker- 2 (another mouse deafness mutation) and nonsyndromic human deafness DFNB3 (Probst et al., 1998; Wang et al., 1998a). Although the complete sequence of the myosins-15 is not yet available, the heavy chain is greater than 200 kda and is predicted to bind two light chains on the basis of the presence of two IQ motifs (Fig. 5). The tail has a single MyTH4 domain and a talin-like repeat. There is no evidence of coiled coil in the tail, indicating that the heavy chains are monomeric, like the myosins-7 responsible for the Shaker-1 and Usher syndromes. Of the preceding classes we shall not consider myosin-4 further because it has yet to be found in the metazoans. Myosins-8, -10, -11, -12, -13, -16, -17, -18, and -19 will not be considered further simply because there is very little information available for them beyond their primary structure. Thus, the remainder of this review will be concerned primarily with myosins- 1, -3, -5, -6, -7, -9, -14, and -15. The number and diversity of the different myosin classes and the fact that some classes have been found in only one or two organisms should not obscure the observation that many different classes are expressed within a single organism and, indeed, within a single cell type. For example, oocytes of Xenopus laevis, the African clawed frog, express five different myosins- 1, two different myosins-2, a myosin-5, a myosin-6, two myosins-9, a myosin- 10, a myosin-14, and a member of the novel myosin class 16 (Sokac and Bement, manuscript submitted). Similar results have been obtained in cultured cell lines (Bement et al., 1994a). Thus, the structural diversity of the different myosin classes does not simply reflect the diversity of the different organisms used for the study of unconventional myosins, but instead is likely to reflect real functional and regulatory differences among the different classes. D. The Basis of Myosin Motility The large number of myosins and myosin classes suggests that, as organisms became more complex, myosins evolved to take on new functions and, accordingly, acquired increased structural diversity. As noted previously, however, the head is responsible for actin binding and ATP hydrolysis, so it is not surprising that this region is relatively well-conserved, even among

15 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 211 the different classes. The conservation implies that basic rules established for myosin-2 motor activity will be largely applicable to the other myosins. This is fortunate because there is a wealth of information on the structure and kinetics of the skeletal muscle myosin-2 motor domain as it transits through the ATPase/actin-binding cycle. The coordinated cycle of ATPase activity and actin binding often is referred to as the mechanochemical cycle because chemical energy derived from ATP hydrolysis is converted to mechanical work, which drives movement along an actin filament. Identification of the basic steps of this cycle, together with structural information now available for the myosin-2 head and neck (referred to as S1 ), has provided an extraordinarily detailed understanding of myosin mechanochemistry. That is, the tertiary structure of chicken skeletal myosin-2 motor and neck domains has been determined to a resolution of 2.8 Å by X-ray crystallography (Rayment et al., 1993b). This information has been combined with the 3D actin structure, crosslinking data, and cryoelectron microscopy images of myosin-decorated actin filaments to further define the actin myosin interface (Rayment et al., 1993a). A complete description of myosin topology can be found in Rayment et al. (1993b). Here we will only highlight those structural elements most fundamental to motility and regulation. With respect to tertiary structure, at the core of the globular motor domain is a large, seven-stranded, mostly parallel -sheet that contributes to the nucleotide-binding pocket. This central sheet is enveloped by a complex arrangement of secondary structural elements that comprise the rest of the nucleotide-binding pocket and the actin interface. A prominent crevice that can exist in either an open or closed conformation extends from the base of the nucleotide-binding pocket to the outer surface of the myosin at the actin contact site. Communication between the nucleotidebinding pocket and the actin-binding region is thought to be conveyed by the conformation of this crevice in the following way: The crevice divides the actin-binding surface into an upper and a lower region. When the crevice is closed, the actin contact site is intact and myosin tightly binds f-actin. When the crevice is open, the actin contact site is disrupted and myosin dissociates or loosely binds f-actin. The crevice is forced open when the -P i of ATP or ADP- -P i (adenosine diphosphate- -inorganic phosphate) occupies the nucleotide-binding pocket and is closed when the nucleotide pocket is unoccupied by P i. Consequently, the binding of P i at the nucleotide-binding pocket is mutually exclusive with tight binding of myosin to f-actin. This crevice thus confers the ATP-sensitive binding of myosin to actin. The neck extends as a single long -helix from the globular head. Lateral binding of light chains reinforces the -helix. The resulting arm is rigid and swings relative to the head domain via a region termed the hinge. This

16 212 ANNA M. SOKAC AND WILLIAM M. BEMENT swing occurs as a result of conformational changes in the globular motor domain. The hinge is also referred to as the converter region because it converts shape changes in the head to the swing of the neck. Thus, conformational change that takes place in the head is further amplified by the neck. This elaborate and interdependent topology sets the stage for myosin mechanochemistry. When the cycle is started in the absence of nucleotide, the nucleotide-binding pocket is open and unoccupied, the connecting crevice is closed, and myosin is very tightly bound to actin (rigor binding). As ATP enters the nucleotide-binding pocket, -P i binding forces open the crevice and so disrupts the interaction between myosin and actin (no binding). As the nucleotide-binding pocket closes around the ATP, a major conformational change occurs in the globular motor and the ATP is hydrolyzed to ADP- -P i. The conformational change is transmitted to the neck via the converter region, resulting in a swing with respect to the motor domain. This swing is termed the recovery stroke. The myosin ADP- -P i complex associates with f-actin via weak ionic interactions (loose binding). A tight association forms between the actin and myosin ADP- -P i complex as additional ionic and hydrophobic interactions are recruited and the upper and lower regions of the actin-binding surface are pulled together. Reconstitution of the actin-binding surface drives closure of the crevice, forcing -P i from the nucleotide-binding pocket. Loss of -P i reverses the conformational change induced by nucleotide pocket closure, resulting in the concomitant swing of the neck. Because the myosin ADP complex is tightly bound to the actin, this swing moves the actin and therefore is called the power stroke. The nucleotide-binding site is now open and ADP is released to restore rigor binding. One simple way to understand the myosin-2 ATPase cycle is to think of an old-fashioned baby carriage (Fig. 6). The actin-binding surface is represented by the top and bottom of the front of the carriage. The nucleotide-binding pocket is encompassed by the top and bottom of the rear of the carriage. The neck is represented by the handle. A hinge at the base of the handle (the converter region) allows the handle to swing with respect to the rest of the carriage. Because opening of the carriage front causes closing of the carriage rear and vice versa, closure of the carriage resembles the mutually exclusive relationship between the crevice at the actin-binding surface and the nucleotide-binding pocket of the myosin-2 head. That is, in the carriage analogy conformational changes of the myosin head are represented by the front of the carriage closing (actin binding) or the rear of the carriage closing (nucleotide binding; Fig. 6). In the absence of nucleotide, the carriage front is closed tightly on the actin filament (represented by an iterated array of babies in Fig. 6). The

17 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 213 FIG. 6 Schematic diagram of the myosin mechanochemical cycle. The baby carriage represents the myosin head and the line of babies represents the actin filament. The front of the carriage defines the actin-binding site, the back of the carriage defines the nucleotide-binding site, and the handle defines the neck. Tighter binding is indicated by increasing discomfort on the faces of the babies.

18 214 ANNA M. SOKAC AND WILLIAM M. BEMENT handle on the carriage is up. Binding of ATP causes partial closing toward the rear of the carriage, thereby disrupting the actin-binding site at the front of the carriage and causing the release of f-actin. Closure of the nucleotide-binding pocket and hydrolysis of ATP to ADP- -P i close the rear of the carriage further and result in movement of the handle to the down position (this corresponds to the recovery stroke). In this position the carriage is only able to bind f-actin loosely. Interactions are recruited between the carriage front and the f-actin, pulling the front closed again. As the front is closed further, the carriage achieves tighter f-actin binding until finally the actin-binding surface is restored, forcing -P i release and causing the handle to swing and resume the up position (the power stroke). The nucleotide-binding pocket is now open and ADP is released to achieve rigor binding. Many years of analyses provide kinetic context for the structural changes characteristic of the skeletal muscle myosin-2 ATPase cycle. One of the features of the skeletal muscle myosin-2 mechanochemical cycle is that release of the -P i apparently is the rate-limiting step of the cycle. Consequently, it is at the level of P i release that myosins often are regulated. As will be described later (see Section III.D and III.F), depending on the myosin in question, this regulation may be achieved by differential phosphorylation of the heavy-chain head or of the light chains or by regulation at the level of the actin filament. A second important feature of the skeletal muscle myosin-2 mechanochemical cycle is its characteristically low duty ratio. That is, in the presence of cellular concentrations of ATP and f-actin, and in the absence of any factors that limit access of the myosin to the f-actin or otherwise regulate the myosin-2, the myosin is not bound tightly to f-actin for most of the ATPase cycle (duty ratio time bound tightly to actin/total time of cycle). Indeed, addition of ATP to preparations containing both f-actin and myosin-2 is one of the standard approaches for separating the two. What is the significance of a low duty ratio? From the cellular standpoint, any motor with a low duty ratio is likely to be nonprocessive. That is, when considered in isolation, such motors are not expected to move along actin filaments continuously because they would spend most of their time unbound and therefore would tend to diffuse away. Efficient actin-based motility is expected to require that motors either have high intrinsic duty ratios or attain circumstances that increase the effective duty ratio. For skeletal muscle myosins-2, a high effective duty ratio is accomplished by the organization of the sarcomere, which has a high concentration of both myosin and f-actin that is physically restrained from diffusing. For other myosins the effective duty ratio could be increased in other manners, such as increasing the density of motors, tethering the motors to membranes to prevent diffusion, and so forth (Ostap and Pollard, 1996).

19 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 215 One study indicated that, in contrast to skeletal muscle myosin-2, myosin- 5 is inherently processive (Mehta et al., 1999). This was demonstrated by following the number of steps taken by single myosin-5 molecules on actin filaments. Each step is thought to correspond to a single ATPase cycle. Even at low ATP concentrations, single myosin-5 molecules were able to take 3 5 steps before stalling or releasing, indicating that one or both of the heads remain bound to the actin filament throughout much of the mechanochemical cycle (Mehta et al., 1999). Myosin-5 differs from myosins-2 in that it has a large step size ( 36 nm for myosin-5 versus 4 17 nm for myosin-2) and the ability to step backward (i.e., toward the minus end of the actin filament) when placed under high load. These features of myosin-5 mechanochemistry have been proposed to contribute to its higher processivity (Howard, 1997; Mehta et al., 1999). Myosin-5 is not the only unconventional myosin that exhibits important differences in mechanochemical properties relative to skeletal muscle myosin-2. Kinetic analyses of several myosins-1 have revealed both similarities and interesting differences relative to skeletal muscle myosins-2 and each other (Ostap and Pollard, 1996; Jontes et al., 1997; Coluccio and Geeves, 1999). All of the myosins-1 studied to date share similar affinities for f-actin in the absence of nucleotide, and all have relatively low duty ratios. However, there is a wide variability in rate constants for ATPinduced dissociation from f-actin among the myosins-1, with some being relatively fast (Ostap and Pollard, 1996) and others being extremely slow (Coluccio and Geeves, 1999). For those myosins-1 that are slow to release ADP, time spent binding tightly to actin is increased ( Jontes et al., 1997), suggesting that the duty ratio is increased. In fact, the slow release of ADP for both brush border myosin-1 and myr3 is correlated with a two-step power stroke (Viegel et al., 1999). Thus, as with myosin-5, differences in the mechanical properties of the unconventional myosins with respect to skeletal muscle myosin-2 could influence the kinetics and cellular functions of these myosins. E. Putative Functions of the Unconventional Myosins Much of the excitement surrounding the study of unconventional myosins stems from demonstrations that mutations in a number of different unconventional myosin genes result in genetic diseases in mice and humans (Table II). Mutations in a myosin-5 gene underlie the dilute phenotype in mice, which is characterized by diluted pigmentation, seizures, and death, depending on the allele. In humans, mutations in a myosin-5 gene are associated with Griscelli disease, which is characterized by diluted pigmentation and

20 TABLE II Unconventional Myosins Encoded by Disease Loci Myosin Class Disease Pathology Mouse myosin-5a 5 Dilute 1 Washed-out coat color; severe alleles produce neurological disorders resulting in death of the juvenile animal Human myosin-5a 5 Griscelli disease 2 Partial albinism, immunodeficiency, neurological disorders Mouse myosin-6 6 Snell s Waltzer 3 Deafness, vestibular dysfunction resulting in a waltzing gate Mouse myosin-7a 7 Shaker-1 4 Deafness, vestibular dysfunction resulting in hyperactivity, head tossing, and circling Human myosin-7a 7 Usher syndrome type 1B 5,6 Deafness, vestibular dysfunction, blindness resulting from retinitis pigmentosa Nonsyndromic recessive deafness 7 Deafness DFNB2 nonsyndromic profound Deafness, vestibular dysfunction in some cases deafness 6 Mouse MYO15 15 Shaker-2 8 Deafness, vestibular dysfunction resulting in head tossing and circling Human MYO15 15 DFNB3 nonsyndromic deafness 9 Deafness 1 Mercer et al., Pastural et al., Avraham et al., Gibson et al., Weil et al., Weil et al., Liu et al., Probst et al., Wang et al., 1998a.

21 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 217 unregulated activation of macrophages and lymphocytes (Pastural et al., 1997). Mutations in a myosin-6 gene underlie the Snell s Waltzer phenotype in mice (Avraham et al., 1995), which is characterized by circling behavior and deafness. Mutations in a myosin-7 gene result in the Shaker-1 phenotype in mice (Gibson et al., 1995), which is characterized by head tossing, hyperactivity, and deafness. Mutations in a myosin-7 gene in humans result in Usher syndrome (Weil et al., 1995), which is characterized by late onset deafness/blindness. Mutations in a myosin-15 gene underlie the Shaker-2 phenotype in mice (Probst et al., 1998), which is characterized by circling behavior and deafness. Mutations in a myosin-15 gene in humans result in a form of nonsyndromic deafness (Wang et al., 1998a). Thus, there is good reason to think that unconventional myosins are critical for normal organismal function and that a detailed understanding of the cellular roles of these proteins will have a major clinical impact. At first blush, the cellular role(s) of the unconventional myosins may appear obvious they are f-actin-based motors and so must be involved in f-actin-based transport. In particular, given the ability of many unconventional myosins to bind membranes, they presumably are involved in f-actinbased transport of membranes and/or organelles. One general model of unconventional myosin transport function that is widely accepted posits that unconventional myosins serve as motors that work in concert with microtubule-based motors to achieve the asymmetric distribution of specific organelles typically observed in eukaryotic cells. That is, microtubules are considered to serve as highways, tracks responsible for transport over long distances to and from the cell interior, whereas f-actin acts as local roads, tracks responsible for transport of membranous organelles within the cell cortex. Kinesins and dyneins then serve as motors for long-range transport, whereas unconventional myosins are thought to serve as motors for local transport. The notion is simple and very appealing because the spatial distribution of the two cytoskeletal elements is consistent with this notion microtubules typically extend from a single area in the cell interior (namely, the microtubule organizing center) to the cell periphery, whereas f-actin is most abundant in the cell cortex. Further, there are now numerous examples of kinesin-based transport of organelles to the cell periphery and dynein-based transport of organelles to the cell interior. However, as will be described later, while unconventional myosins may serve to transport organelles in some cases, at present it is not at all clear whether this role is as representative of unconventional myosin function as is widely assumed. We will argue that much of the evidence in support of unconventional myosins acting as organelle transporters can also be interpreted as supporting roles for unconventional myosins as organelle anchors or cross-linkers.

22 218 ANNA M. SOKAC AND WILLIAM M. BEMENT 1. Organelle Transporters? The unconventional myosin discovered first was Acanthamoeba castellanii myosin-1 (Pollard and Korn, 1973). Because of the strengths of this organism for preparative biochemistry, the Acanthamoeba myosins-1 are the beststudied unconventional myosins, at least at the biochemical level. Indeed, it was studies of purified Acanthamoeba myosins-1 that led to the initial proposal that unconventional myosins might serve as motors involved in membrane trafficking (Pollard et al., 1991). This idea rests on the facts that these proteins (1) bind both natural (Adams and Pollard, 1986, 1989) and artificial (Adams and Pollard, 1989; Doberstein and Pollard, 1992; Zot et al., 1992) membranes via their tail domains, (2) localize to membrane compartments in vivo (Baines et al., 1992), and (3) can support f-actindependent movement of organelles and purified lipid vesicles in vitro (Adams and Pollard, 1986). The notion of unconventional myosins as membrane transport motors is supported further by observations of f-actin-dependent organelle/particle trafficking in a number of different systems, including squid axoplasm (Kuznetsov et al., 1992; Bearer et al., 1993, 1996; Langford et al., 1994), sea urchin coelomocytes (D Andrea et al., 1994), Drosophila embryos (Mermall et al., 1994), mammalian neurons (Morris and Hollenbeck, 1995; Evans and Bridgman, 1995), and yeast (Simon et al., 1997). In addition, isolated organelles have been either directly shown to have unconventional myosins associated with them by immunoblotting of organelle fractions (Fath and Burgess, 1993; Evans et al., 1998) or inferred to have unconventional myosins associated with them by the demonstration of ATP-dependent binding of f-actin and/or translocation of f-actin by organelles (Langford et al., 1994; Bearer et al., 1996; Evans et al., 1998). Further, genetic analyses showed that disruptions of unconventional myosin genes in Aspergillus (McGoldrick et al., 1995), Dictyostelium (Novak et al., 1995), and Saccharomyces cerevisiae ( Johnston et al., 1991; Govindan et al., 1995; Geli and Reizman, 1996) impair membrane trafficking processes, such as endocytosis and exocytosis, as well as resulting in abnormal pigment granule distribution in melanocytes of dilute mice (Mercer et al., 1991). Similarly, overexpression of myosin truncates in cultured cells perturbs endocytotic pathways (Raposo et al., 1999). Thus, it is hardly surprising that the notion of unconventional myosins as organelle transporters became firmly entrenched in the literature and is now in several textbooks (Alberts et al., 1994). Nevertheless, critical analysis of the evidence indicates that the situation may not be so simple. At least four criteria should be satisfied before a given situation can be considered to be representative of unconventional myosin-dependent organelle transport. First, it must be demonstrated that the transport occurs on f-actin within intact cells. Although transport of a

23 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 219 given organelle on f-actin in vitro certainly is outstanding evidence of an association of a myosin with that organelle, it cannot be assumed that the in vivo role of the myosin is to transport the organelle. It is equally likely that the myosin is responsible for tethering the organelle in question to the actin cytoskeleton and/or that the myosin is a passive passenger en route to its final destination at, say, the plasma membrane (see discussion below). Second, it must be demonstrated that what is being transported is actually an organelle. Complexes of proteins, including myosins, undergo directed locomotion within cells (Verkhovsky et al., 1995; Kolega, 1998). Thus, it cannot be assumed that all moving particles within cells represent organelles. Third, it must be demonstrated that the organelle in question is associated with an unconventional myosin. Ideally, this would include unambiguous identification of the myosin in question, although this is not always possible. Fourth, and most importantly, it must be demonstrated that inhibition, inactivation, or removal of that myosin results in the cessation of the transport in question in vivo. To date, in no system have all four criteria been satisfied, although in some cases two or three have. These are described next. a. Yeast Mitochondria In the budding yeast, S. cerevisiae, mitochondria colocalize with f-actin cables (Drubin et al., 1993) and undergo directed motility (Simon et al., 1997). This motility is inhibited in temperaturesensitive actin mutants following shift to the restrictive temperature, as well as following depolymerization of f-actin by latrunculin treatment (Boldogh et al., 1998). Microtubule depolymerization, however, has no apparent effect on mitochondrial movement or distribution. Further, isolated yeast mitochondria display ATP-dependent binding and translocation of f-actin (Simon et al., 1995), implying the existence of a myosin-like motor on the surface of mitochondria. Thus, mitochondrial motility in budding yeast satisfies the first three criteria required of an organelle transporter. Surprisingly, however, it does not satisfy the fourth. All of the budding yeast unconventional myosins have now been identified by genomic sequencing (there are four two myosins-1 and two myosins-5), and mutations in any one of them have no effect on mitochondrial motility either in terms of absolute velocity or in terms of the final distribution of mitochondria (Simon et al., 1995; Goodson et al., 1996). Whereas double myosin-1 and double myosin-5 mutations result in modest differences in mitochondrial distribution, this effect apparently results from mutation-induced reorganization of the actin cytoskeleton rather than a transport defect (Simon et al., 1995; Goodson et al., 1996). Thus, unless the f-actin motor activity associated with yeast mitochondria is encoded by a gene that is strikingly different from all other known

24 220 ANNA M. SOKAC AND WILLIAM M. BEMENT myosin genes, it cannot be concluded that unconventional myosins serve as mitochondrial transport motors in yeast. b. Neurons Kuznetsov et al. (1992) demonstrated by high-resolution, differential-interference contrast (DIC) microscopy that organelles could, in extruded squid axoplasm, jump from microtubule tracks to f-actin tracks and continue moving. Since that time several reports have documented that organelles from squid axoplasm are associated with myosin-like motors. Much of the attention has focused on myosin-5 as a potential neuronal organelle motor because mutations in myosin-5 result in neuronal deficits in dilute mice. Consistent with this notion, myosin-5 has been shown to associate with a subset of synaptic vesicles (Prekeris and Terrian, 1997; Evans et al., 1998) and neuronal endoplasmic reticulum (Tabb et al., 1998). In both cases, the purified organelles can translocate on actin filaments in vitro. Collectively, the preceding observations suggest that unconventional myosins in general may transport neuronal organelles and that myosin-5 in particular may act as a neuronal organelle transport motor. Consistent with this notion, translocation of uncharacterized particles on f-actin has been observed in growth cones of neurons treated with low doses of nocodazole and cytochalasin (Evans and Bridgman, 1995). In addition, mitochondria translocate in axons even after complete microtubule depolymerization, although in a predominantly retrograde fashion (that is, toward the cell body) and with markedly different kinetics than are observed in control cells (Morris and Hollenbeck, 1995). At present, it is unclear how these diverse observations fit together, and none of the movements hypothesized to be myosin-dependent satisfies all four criteria for myosin-based organelle transport. For example, the nature of the particles observed by Evans and Bridgman (1995) to translocate on f-actin in the growth cone of cytochalasin and nocodazole-treated neurons is unknown, although they are too small to be mitochondria. Conversely, mitochondria distribution apparently is normal in dilute mice, and there is no evidence that myosin-5 binds to neuronal mitochondria (Evans et al., 1997). If myosin-5 is a mitochondrial motor, then this motor must be encoded by a myosin-5 gene different from the one represented by the dilute gene. Regardless of the myosin motor or motors associated with neuronal mitochondria, the fact that the direction and kinetics of mitochondrial movement are significantly different when microtubules are depolymerized suggests that actin-based motors do not normally drive mitochondrial transport, although they may make it more efficient (Morris and Hollenbeck, 1995). The direct demonstration of myosin-5 on purified organelles (Prekeris and Terrian, 1997; Evans et al., 1998; Tabb et al., 1998) and the repeated

25 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 221 observation of f-actin-dependent movement of different neuronal organelles in vitro (Langford et al., 1994; Bearer et al., 1996) indicate that many neuronal organelles have unconventional myosins associated with them. However, as noted previously, the myosins may be acting as passive passengers or as eventual tethers that link the organelles in question to the cytoskeleton. In fact, as will be described later, analyses of pigment granule movement in melanocytes of wild-type and dilute mice indicate that myosin- 5 functions primarily as an organelle tether. c. Melanocytes The results of studies designed to evaluate the role of unconventional myosin-dependent transport of pigment granules in melanocytes are complex, but several of the criteria required for unconventional myosin-dependent movement have been satisfied. In mice, pigment granule distribution is dependent on both f-actin and microtubules (Wu et al., 1998a). Moreover, in dilute mice, melanocyte pigment granule distribution is abnormal with most of the granules clustered in the perinuclear region. Thus, it naturally was suggested that myosin-5 is responsible in part for transporting pigment granules in these cells, perhaps in concert with microtubule-based motors (Wei et al., 1997; Wu et al., 1997). However, in vivo analysis of pigment granule dynamics in melanocytes derived from wild-type and dilute mice has revealed that, in fact, the myosin-5 does not transport pigment granules on f-actin, but rather anchors them to f-actin following transport to the cortex by microtubules. That is, in wild-type melanocytes, pigment granules constantly are being shuttled to and from the cortex along microtubules, but a subset of them are captured in the cortex in an f-actin-dependent manner. In contrast, in dilute melanocytes, anterograde and retrograde movement of granules is still observed, but capture does not occur and pigment cannot accumulate in the cortex (Wu et al., 1998a). Note that prior to this study, myosin-5 in mouse melanocytes was considered an example par excellence of an unconventional myosin acting as an organelle motor on the basis of the dilute phenotype, the fact that dilute encodes a myosin-5 (Mercer et al., 1991), and the localization of myosin-5 on pigment granules (Provance et al., 1996; Wu et al., 1997). In melanocytes of frogs and fish, studies have suggested that an unconventional myosin, presumably myosin-5, works with microtubule-based motors to transport pigment granules from the cell center to dispersed locations throughout the cell (Rogers and Gelfand, 1998; Rodionov et al., 1998). Put simply, the notion is that pigment granules move outward toward the cell periphery on microtubules and then jump onto and move along f-actin tracks to achieve uniform distribution throughout the cytoplasm. This model is based on the following findings: First, frog melanosomes (pigment granules) have an associated myosin-5 and undergo f-actin-dependent motility in vitro (Rogers and Gelfand, 1998). Second, when microtubules are depoly-

26 222 ANNA M. SOKAC AND WILLIAM M. BEMENT merized, melanosomes nevertheless can undergo redistribution from the cell center, albeit relatively slowly (Rogers and Gelfand, 1998). Third, when melanosome dynamics are analyzed in fragments of fish melanocytes (experimental fragmentation was employed to reduce both the number of microtubules and the number of melanosomes), f-actin-dependent melanosome translocation can be observed (Rodionov et al., 1998). Fourth, depolymerization of f-actin in intact fish melanocytes prevents the normal dispersal of melanosomes (Rodionov et al., 1998). These findings certainly are consistent with f-actin-dependent transport of melanosomes, but are not quite definitive. In light of the demonstration that myosin-5 links pigment granules to the cortical actin cytoskeleton in mouse melanocytes (Wu et al., 1998a), it is possible that, under normal conditions, the melanosomes are not normally transported on f-actin in fish and frog melanocytes, but rather are cross-linked to it via myosin-5. That is, the partial or complete removal of microtubules from melanocytes might exaggerate or even result in f-actin-dependent melanosome motility, as may also be true for mitochondrial motility in neurons (see preceding discussion). For example, f-actin-dependent melanosome movement can be observed in mouse melanocytes depleted of microtubules, but this movement apparently is unimportant for the regulated distribution of melanosomes (Wu et al., 1998a). Conversely, given the apparent abundance of both microtubules and f-actin in intact melanocytes, it could be possible to achieve dispersed melanosomes simply by tethering the melanosomes to f-actin rather than actual transport along f-actin. The preceding points having been made, we should also point out that it is probable that unconventional myosins do act as organelle transporters in some cases, including some of those described previously. The technical difficulties inherent in trying to image f-actin-dependent motility in vivo are great because f-actin does not typically run for long distances as microtubules do. Further, in order to distinguish unconventional myosins acting as organelle/f-actin cross-linkers versus organelle transporters will require tools that specifically inhibit myosin motor activity versus cross-linking activity. These difficulties notwithstanding, there are several clear-cut examples of f-actin-dependent organelle movement observable in the absence of experimental manipulations (Nothnagel et al., 1981; King-Smith et al., 1997), and unconventional myosins remain the most likely candidates as motors for these movements. 2. Protein/mRNA Transporters? There is also evidence that unconventional myosins may serve to transport proteins or RNA. In Drosophila, the ninac gene product (myosin-3) is required for proper localization of calmodulin within the eye (Porter et al.,

27 REGULATION AND EXPRESSION OF METAZOAN MYOSINS ). Because the myosins-3 bind calmodulin and because the localization of the myosins themselves is dependent on the motor domain, it follows that these proteins may serve, in part, to control the distribution of calmodulin within the fly eye by actin-based transport. Of course, it is also possible that they serve to localize calmodulin by acting as cross-linkers rather than motors per se, as discussed previously for myosins-5. Budding yeast myo4 (myosin-5) has been shown to be required for localization of ASH1 mrna (which encodes a transcription regulator) to the daughter cell (Bobola et al., 1996; Jansen et al., 1996). myo4 complexes with ASH1 mrna in a manner that is dependent on several other proteins (Chartrand et al., 1999; Münchow et al., 1999). In addition, ASH1 mrna has been directly shown to undergo directed movement (Bertrand et al., 1998; Beach et al., 1999), and this movement is inhibited in myo4 mutants (Bertrand et al., 1998). Thus, on the basis of the criteria set out previously, ASH1 mrna transport is likely to be mediated by a yeast myosin Membrane/f-Actin Cross-Linkers? If unconventional myosins are not necessarily acting as organelle transporters, but their disruption impairs processes that involve membrane trafficking (Table III), such as endocytosis and exocytosis, what might they be doing? As discussed previously, one reasonable possibility is that they function to cross-link membranous organelles to the actin cytoskeleton. In this context, the ATPase activity of the myosin would not be used for directed transport along continuous f-actin tracks but instead used as a means to provide regulation for the cross-linking. That is, myosins have an intrinsic means of regulation for f-actin binding, namely, ATP binding (see Section II.D). Although this might seem like a poor mechanism for having regulated cross-linking since the ATP level in cells is thought to be constitutively high, other factors in the cell may allow myosins to play this role. These factors might include having multiple unconventional myosins per organelle (see Section III.B.2), negative regulation of the ATPase activity (see Section III.D,G,H), or regulation at the level of the actin filament itself (see Section III.F). In the case of myosin-5, which is more processive than other myosins, high levels of cellular ATP are of less concern because myosin-5 is expected to be bound to f-actin even in the presence of high ATP. In addition to the mouse melanocyte results (Wu et al., 1998a), which provide a clear example of an unconventional myosin acting to cross-link an organelle to the actin cytoskeleton, many of the observations discussed previously also provide support for unconventional myosins acting as membrane f-actin cross-linkers. That is, association of specific unconventional myosins with organelle fractions, localization of unconventional myosins to organelles by immunoblotting and immunomicroscopy, and translocation

28 TABLE III Putative Unconventional Myosin Functions Myosin Class Putative function Organelle transport S. cerevisiae Myo2 5 Transport of vesicles 1 and the vacuole 2 to the daughter cell Drosophila 95F 6 Cell cycle-dependent transport of particles in the syncytial blastoderm 3 Mouse dilute 5 Transport of smooth endoplasmic reticulum in dendritic spines 4 Xenopus myosin-5 5 Transport of pigment granules to a uniform distribution in melanophores 5 Membrane/f-actin cross-linking Mouse dilute 5 Cross-linking between melanosomes and f-actin in melanocytes 6 Membrane trafficking Acanthamoeba myosin-1c 1 Contractile vacuole pumping 7 Dictyostelium myoa, -B, -C, -D 1 Fluid phase pinocytosis 8,9 Dictyostelium myoa, -B 1 Secretion of lysosomal enzymes 10 Dictyostelium myob, -C 1 Phagocytosis 9,11 Aspergillus nidulans myoa 1 Polarized secretion, 12 endocytosis 13 S. cerevisiae Myo3, Myo5 1 Receptor-mediated endocytosis 14 Cell motility Dictyostelium myoa, -B 1 Regulation of pseudopod Chicken myosin-5 5 Extension of filopodia in neuronal growth cones 19 Morphogenesis/cell polarity Dictyostelium myoa, -B, -C 1 Fruiting body formation (in suspension), 8 generation of cortical tension in resting cells

29 Aspergillus nidulans myoa 1 Polarized cell growth 12 S. cerevisiae Myo3, Myo5 1 Cell growth, 21 polarization of the actin cytoskeleton 21 Protein/mRNA transport S. cerevisiae Myo4 5 Transport of ASH1 mrna to daughter cell 22,23 Drosophila ninac 3 Localization of calmodulin 24 Signal transduction Rat myr5 9 Rho-mediated actin remodeling 25 Sensory functions Drosophila ninac 3 Phototransduction, 26 maintenance of photoreceptor structure, 26 photoreceptor membrane turnover 27 Limulus myosin-3 3 Coordination of circadian rhythm and photoreceptor shape changes 28 Bullfrog myosin-1 1 Auditory function via regulation of stretch-gated channels of inner-ear hair cells 29,30 Mouse myosin-6 6 Auditory and vestibular function via basal anchorage of stereocilia of inner-ear hair cells 31,32 Mouse myosin-7a 7 Auditory and vestibular function via lateral stabilization of stereocilia of inner-ear hair cells Mouse MYO15 15 Auditory and vestibular function 36,37 Human myosin-7a 7 Auditory and vestibular function via lateral stabilization of stereocilia of inner-ear hair cells 32,38,39 1 Johnston et al., Hill et al., Mermall et al., Takagishi et al., Rogers and Gelfand, Wu et al., 1998a. 7 Doberstein et al., Novak et al., Jung et al., Temesvari et al., Jung and Hammer, McGoldrick et al., Yamashita and May, Geli and Riezman, Wessels et al., Titus et al., Wessels et al., Novak and Titus, Wang et al., Dai et al., Goodson et al., Bobola et al., Jansen et al., Porter et al., Müller et al., Porter and Montell, Hicks and Williams, Battelle et al., Gillespie et al., Metcalf et al., Avraham et al., Hasson et al., 1997a. 33 Gibson et al., Weil et al., Self et al., Wang et al., 1998a. 37 Probst et al., Weil et al., Liu et al.,

30 226 ANNA M. SOKAC AND WILLIAM M. BEMENT of isolated organelles on f-actin or vice versa are just as consistent with a cross-linking role as they are with a role in translocation, in that they simply demonstrate the presence of a myosin on the organelle in question. If, as seems likely (Gottlieb et al., 1993), organelle linkage to the actin cytoskeleton is required for normal endocytosis and exocytosis, then organelle factin cross-linking could explain the observed membrane-trafficking phenotypes that result from mutations in one or more unconventional myosin genes. It is also possible that some of the observed defects in membrane trafficking result from a general disruption of the cortical f-actin cytoskeleton that can result from unconventional myosin disruption, as suggested by Temesvari et al. (1996). Some unconventional myosins are associated with the plasma membrane, as demonstrated by both immunomicroscopy and biochemical fractionation (Baines et al., 1992; Wagner et al., 1992; Heintzelman et al., 1994). Further, perturbation of unconventional myosins (in particular, myosins-1) results in disruption of the cortical f-actin cytoskeleton in Dictyostelium (Peterson and Titus, 1994; Novak et al., 1995) and yeast (Goodson et al., 1996), as well as cell locomotion defects in Dictyostelium (Titus et al., 1993; Wessels et al., 1996; Novak and Titus, 1997). The disruption is manifest as an abnormal distribution of cortical f-actin in Dictyostelium myosin-1 double mutants (Novak et al., 1995) and loss of cortical f-actin polarization in budding yeast myosin-1 double mutants (Goodson et al., 1996). These results imply that the myosins-1 may be responsible for stabilizing or controlling cortical f-actin. Consistent with this notion, brush border myosin-1 is most abundant in enterocyte microvilli, structures that are generally characterized as exceptionally stable examples of plasma membrane cytoskeleton linkage (Bement and Mooseker, 1996). This is not to say that unconventional myosins only act as static crosslinkers of the f-actin cytoskeleton and the plasma membrane. In addition to the motility deficits observed following myosin-1 disruption in Dictyostelium (see earlier discussion), immunofluorescence studies have documented the redistribution of myosins-1 (Stöffler et al., 1995) and myosin-6 (Buss et al., 1998) to the plasma membrane in response to the addition of lectins and growth factors, respectively. These results imply that unconventional myosins may act as regulatable cross-linkers that are employed by the cell in specific situations and in response to specific extracellular signals. Such a role is also consistent with the presence of signaling domains in the tails of many unconventional myosins. That is, as discussed previously, SH3 domains are present in both protozoan and metazoan myosins-1. In addition, PH domains, which have been shown to be responsible for protein protein and protein phospholipid interactions in other proteins (Shaw, 1996), are present in myosins-10. Further, rho-gtpase-activating protein (GAP) domains and putative lipid-/zinc-binding domains are present in

31 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 227 myosins-9. Although this notion has yet to be tested, such myosins might serve to act as templates for the assembly of signaling complexes at the plasma membrane. Such complexes would be ideally positioned to transmit or propagate signals from the plasma membrane to the cell interior. 4. Microtubule/f-Actin Cross-Linkers? A considerable amount of evidence suggests that the actin cytoskeleton and microtubule cytoskeletons interact in vivo in most, if not all, cell types (Waterman-Storer and Salmon, 1999). The connections between the two systems may be indirect, with microtubules regulating the actomyosin cytoskeleton via members of the rho family of small GTP-binding proteins (Waterman-Storer et al., 1999; Ren et al., 1999), or they may be more directly connected (Sider et al., 1999). Several lines of evidence suggest that the unconventional myosins may act as linkers for the two cytoskeletal systems. Indirect evidence comes from (1) the myosin-7a mutant phenotype that is often characterized by disruption of structures containing cilia (see Section III.B.2), (2) the localization pattern of myosin-7a, which frequently associates with cilia (see Section III.B.2), and (3) the localization pattern of myosin-5a, which associates with microtubules in some cell types (see Section III.B.2). Direct evidence comes from the demonstration that myosin-5a binds kinesin (Huang et al., 1999), whereas myosin-6 forms a complex with the microtubule-binding protein, CLIP-190, in Drosophila embryos (Lantz and Miller, 1998). III. Unconventional Myosin Regulation A. Myosin Structure The unconventional myosins display a large range of diversity in their primary structure, both across and within classes. Presumably, much of this diversity is responsible for imparting differences in function and/or regulation to the different myosins. As will be seen, structural diversity is evident in all three of the major regions of the myosin heavy chain the head, neck, and tail. 1. Variation in the Head Several unconventional myosins contain unique amino-terminal extensions, presently of unknown function. The 140- to 150-amino acid extensions of the human and rat members of class 9 (Reinhard et al., 1995; Wirth et al.,

32 228 ANNA M. SOKAC AND WILLIAM M. BEMENT 1996; Chieregatti et al., 1998) resemble the Ras-binding domain of Raf and Ral-GEF, but do not bind Ras (Kalhammer et al., 1997; Chieregatti et al., 1998). Similarly, Caenorhabditis elegans HUM4 (class 12) contains a 150- amino acid amino-terminal extension (Baker and Titus, 1997). Last, the Drosophila and Limulus myosin-3 proteins contain extensions that encode a kinase domain, including those residues required for nucleotide binding and phosphotransfer (Montell and Rubin, 1988; Battelle et al., 1998). The Drosophila myosin-3 extension shares identity with 15 of the 16 most highly conserved amino acids of serine threonine kinases, tyrosine kinases, and viral serine threonine kinases (Montell and Rubin, 1988). Although kinase activity has been demonstrated for Drosophila myosin-3 (Ng et al., 1996), and this domain is required for myosin function in phototransduction and the maintenance of retinal structure (Porter and Montell, 1993), no intramolecular regulatory functions have yet been assigned to it. Within the globular catalytic head domain are several flexible loops that do not appear in the myosin-2 crystal structure (Table IV; Rayment et al., 1993b). Among the myosins, these loops are poorly conserved in length and amino acid composition (Spudich, 1994). Consequently, it has been proposed that these loops impart specificity to myosin function and/or regulation. Indeed, these loops appear to adjust the kinetics of myosin enzymatics. Two of these loops correspond to preferred sites of attack by proteases and often are considered in terms of their position with respect to the major products of limited proteolysis. For example, loop 1 is the tryptic digestion site linking 25-kDa and 50-kDa fragments of skeletal muscle myosin-2. This loop is positioned near the nucleotide-binding pocket (Rayment et al., 1993b). In a series of studies, chimeric myosins, created by fusing the backbone of a given myosin to the loop 1 region of a donor myosin, have been used to determine what influence loop 1 has on myosin kinetics. A Dictyostelium myosin-2 backbone was combined with loop 1 sequences donated from rabbit skeletal myosin-2 or Acanthamoeba myosin- 2. This loop 1 substitution had no effect on actin activation of the mechanochemical cycle, but it did influence the velocity of actin translocation and ADP release in a manner that correlated with the donor myosin (Murphy and Spudich, 1998). This result is in agreement with an earlier hypothesis that loop 1 controls the rate of ADP release (Spudich, 1994). A second study, in which chick smooth muscle myosin-2 was used as the backbone of the chimera and loop 1 sequences were donated by a number of other myosins, including skeletal, cardiac, and nonmuscle myosin-2, demonstrated that, of all the kinetic steps, loop 1 most greatly influences the rate of ADP release. These workers further demonstrated that the more flexible loop 1, the more efficient the myosin at releasing ADP.

33 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 229 Although loop 1 most significantly influences the rate of ADP release, it cannot be generalized that loop 1 only influences this kinetic parameter. The chick smooth muscle myosin chimeras as well as native isoforms of smooth muscle myosins with inserted sequences in loop 1 exhibit changes not only in ADP-binding affinity and actin translocation but also in the rate of ATPase activity (Sweeney et al., 1998; Rovner et al., 1997; Kelley et al., 1993). It is not obvious whether the increase in ATPase rate is dependent on the increased rate of ADP release, as may be inferred from the dependency of sequential events in the mechanochemical cycle, or whether this effect is independent of the other cycle kinetics. Nevertheless, loop 1 is a major regulator of myosin activity. In the majority of unconventional myosins this loop is minimized relative to myosin-2, with the exception of Dictyostelium myoj (class 11), which contains a large 25- to 30- amino acid insert in loop 1 relative to myosin-2 (Hammer and Jung, 1996; Peterson et al., 1996). At present, the significance of this insertion is unknown. Loop 2 is the linker between the 50-kDa tryptic digestion fragment and a 20-kDa tryptic digestion fragment. Loop 2 mediates the initial weak electrostatic interactions between myosin ADP-P i and the negatively charged amino terminus of an actin monomer within the actin filament (Rayment et al., 1993a). By using chimeras with Dictyostelium myosin-2 backbone and loop 2 sequences donated from vertebrate skeletal, smooth, and cardiac muscle myosins-2, it was observed that the actin-activated ATPase activity (measured as P i release) of a given chimera corresponded well with the donor myosin (Uyeda et al., 1994). This result suggested that loop 2 controls ATPase activity by modulating the rate at which the myosin ADP- -P i forms the tight binding complex and releases P i.inan analysis, loops of up to 20 amino acids carrying charge variations of 1 to 20 relative to the native myosin-2 were inserted into a Dictyostelium myosin-2 backbone. It was demonstrated that the size of loop 2, when accompanied by small charge changes (not more than 2 charge units), had little influence on the rate of ATPase activity or the affinity between f- actin and myosin. In contrast, loop 2 sequences carrying four or more newly introduced positive charges stabilizes f-actin myosin binding and increases f-actin-dependent ATPase activity (Furch et al., 1998). Thus, loop 2 insertions carrying positive charges increase the affinity of myosin ADP-P i for actin, thereby making tight binding more efficient. In addition, loop 2 affects the regulatory properties of myosin. That is, certain myosins-2 that normally are activated by phosphorylation of their regulatory light chain (see Section III.D.1.a) become constitutively active with modifications of loop 2 (Rovner et al., 1995). Both the human and rat isoforms of class 9 myosins contain a very large ( 150 amino acids) insert at loop 2 (Wirth et al., 1996; Reinhard et al.,

34 TABLE IV Functional and Regulatory Elements of the Myosin Heavy Chain Residue number in chicken skeletal Myosin structure Also known as muscle M2 Description Loop to 50-kDa junction Glu204 Gly216 Located near the nucleotide-binding pocket, influences ADP loop release rate Hypertrophic cardiac loop Arg405 Lys415 Located within the actin-binding interface, contains TEDS site, myopathy (HCM) functional/regulatory significance not well-defined loop Loop to 50-kDa junction Gly627 Phe647 Located within the actin-binding interface, influences Pi release loop rate Phosphate (P) loop Not strictly defined Located within the nucleotide-binding pocket, GESGAGKT sequence highly conserved among other nucleotide-binding proteins, including ras and kinesin TEDS site Glu411 Located at the tip of the HCM loop, 16 residues upstream of the highly conserved DALAK sequence, generally encodes either a negatively charged residue (E or D) or a phosphorylatable residue (T or S) Converter region Hinge region Not strictly defined The region encompassing the junction between the globular head and the -helical neck, the head swings from this point during the power and recovery stroke 230

35 Reactive sulfhydryls SH1 and SH2 Cys707 and Cys697, Located in the converter region, the cysteines move from 19 Å respectively apart to 2 Å upon nucleotide binding, functional/regulatory significance not well-defined, not conserved among all myosins IQ domain Thr790 Arg800 Located in the neck, conserved core sequence Ile816 Val826 IQXXXRGXXXRK binds EF-hand family members, including M2 regulatory light chain, M2 essential light chain, and calmodulin Tail homology 1 TH1, polybasic region No corresponding region Located in the tail, rich in basic residues, binds nonspecifically (MyTH1) domain to acidic phospholipids Tail homology 2 TH2,GPA- or GPQ- No corresponding region Located in the tail, rich in Gly, Pro, Ala, and/or Gln residues, (MyTH2) domain rich region mediates ATP-independent actin binding Tail homology 3 TH3, Src homology 3 No corresponding region Located in the tail, encodes SH3 domain, mediates (MyTH3) domain (SH3) domain protein protein interactions Tail homology 4 No corresponding region Located in the tail, functional/regulatory significance not defined (MyTH4) domain 231

36 232 ANNA M. SOKAC AND WILLIAM M. BEMENT 1995; Chieregatti et al., 1998). In human myosin-9b, this insertion is highly basic with pi Kinetic studies of human myosin-9b show that this myosin is a very slow motor, translocating actin filaments at a fraction of the speed recorded for other myosins (Post et al., 1998). Myosin-9b is unusual in that it is capable of translocating f-actin in the in vitro motility assay for far longer periods of time than other myosins. Myosin-9b binds to f-actin in an ATP-insensitive manner that perhaps contributes to the longevity of actin filament translocation powered by this motor (Post et al., 1998). Although the necessity of the large insertion at loop 2 as yet is unknown, its localization at the actin-binding interface places it in optimal position to mediate nucleotide-independent binding to f-actin. A second loop known to contact actin is referred to as the Arg405 Thr414 loop (Cope et al., 1996) or the HCM loop on the basis of the finding that a mutation in this loop in human cardiac myosin-2 is associated with hypertrophic cardiomyopathy (HCM; Geisterfer-Lowrance et al., 1990; Rayment et al., 1993a). In addition to the phenotype of HCM (enlargement of the heart and heart failure), the importance of this loop is demonstrated by the finding that the same mutation that causes HCM impairs both cardiac myosin-2 f-actin-activated ATPase activity and rate of f-actin translocation (Sweeney et al., 1994). Further, this loop is also the locus of the so-called TEDS site, a position that is almost invariably occupied by either a phosphorylatable or negatively charged residue (Bement and Mooseker, 1995; see Section III.D.2.a). Because most of the information about this loop comes from studies of differential phosphorylation, this loop will be discussed further (see Section III.D.2). An additional region that may be critical for myosin regulation is the converter region of the head that surrounds the reactive sulfhydryl pair SH1 and SH2 (Table IV; Rayment et al., 1993b). This region is predicted to be the hinge for neck swinging (see Section II.D; Jontes and Milligan, 1997; Whittaker et al., 1995; Uyeda et al., 1996). At the primary structure level, this region is conserved within most myosin classes, but not necessarily between myosins of different classes (Cope et al., 1996). The converter undergoes a major rearrangement when the nucleotide pocket closes, bringing the two sulfhydryls from a position of 19 Å apart to only 2 Å apart (Rayment and Holden, 1994). Interestingly, significant variation is generated in this region of the Drosophila myosin-2 heavy chain (MHC) by alternative splicing events (Bernstein and Milligan, 1997). The resulting myosin isoforms have diverse physiological roles and predicted kinetics, suggesting that this region may dictate major events of the mechanochemical cycle. As the hinge between the head and neck, this region converts events at the nucleotide- and actin-binding sites into movement of the carboxy-terminal portions of the myosin. Communication may also occur in the reverse order from the light-chain-binding domain to the catalytic

37 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 233 head. It remains to be seen whether or not this region is critical to unconventional myosin activity, although this is the pivot point identified for BBM-1 upon ADP binding ( Jontes and Milligan, 1997). In addition, Cope et al. (1996) noted variation in this region between the different unconventional myosin classes. 2. Variation in the Neck and Light Chain Composition The neck domain is a long -helix that extends from the globular head domain and is stabilized by light-chain binding. The resulting rigid lever arm swings in response to conformational changes in the head. Because the neck rotates as a rigid unit, it is thought to amplify the small conformational changes in the head to produce a larger power stroke. Variation in the neck has the potential to impact myosin function by at least two distinct mechanisms. First, the neck is the site where the regulatory light chains bind. Under normal conditions myosins require binding of their light chains for maximal ATPase activity and motility (Lowey et al., 1993; Wolenski et al., 1993; Trybus et al., 1994). Several myosin light chains have now been identified, and each imparts its own mode of regulation to the mechanochemical cycle (see Section III.C and III.D). Consequently, regulation of a given myosin may be defined by its light-chain composition, which is, in turn, defined by the light-chain-binding sites of the neck. Second, the length of the lever arm is proportional to the working distance of the myosin, defined as the distance that the distal carboxy terminus moves relative to the previous actin-binding site. That is, the longer the lever arm, the greater the displacement of its carboxy terminus caused by a swing of a given angle. It has been suggested that lever arm length therefore can influence both the speed and processivity of a myosin (Spudich, 1994; Howard, 1997). In support of the former possibility, the speed at which Dictyostelium myosin-2 translocates actin filaments has been demonstrated to be proportional to the length of the lever arm (Uyeda et al., 1996). In support of the latter possibility, myosin-5, with its exceptionally long neck, takes unusually large steps and displays high processivity (Mehta et al., 1999). Thus, variation in the length of the neck has the potential to significantly impact the motor properties of a given myosin. Consistent with the notion that variation in the neck may control variation in myosin regulation and function, there is dramatic variation in neck length, not only among classes but also within classes (Table V; Wolenski, 1995). Among classes the myosins-14 have no apparent IQ motifs and therefore are considered to be neckless (Heintzelman and Schwartzman, 1997). In contrast, the myosins-13 have up to seven IQ motifs, presumably permitting binding of up to seven light chains, although this has yet to be demonstrated.

38 TABLE V Alternative Isoforms of Unconventional Myosins Proposed mechanism Myosin Transcript size Protein size of variation Description of variation T. gondii M-B, -C 1 114, 124 kda Alternative splicing Differ in the carboxy-terminal tail region Drosophila M-1A 2 3.8, 4.4 kb Unknown Unknown ninac 3 3.6, 4.8 kb 132, 174 kda Alternative splicing Differ in the carboxy-terminal tail region M-5 4 Broad 6-kb band Unknown Unknown 95F 5 Broad 4.5- to 4.7-kb Alternative splicing, Differ in the 5 -UTR, 3 -UTR, and/or the central or band alternative usage carboxy-terminal tail region of multiple polya sites Chicken BB M-1 6 Alternative splicing The slightly larger isoform contains an additional calmodulin-binding motif Mouse MM , 4.9, 6.5 kb Alternative usage of Unknown multiple polya sites dilute 8 7, 8, 12 kb Alternative splicing Differ in central tail region 234

39 Rat MM-1 7 5, 6.9 kb Unknown Unknown MM , 7 kb Unknown Unknown myr1a, -b, -c 9 Broad 5.7- to 6.2-kb Triplet at 130 kda Alternative splicing Differ in number of IQ domains (contain 6, 5, or 4) band myr5 10 7, 10 kb Unknown Unknown myr kb Alternative splicing Differ in central tail region myr , 10.5 kb Doublet at 300 kda Alternative splicing Bullfrog M , 160 kda Unknown Unknown Human myosin-1a , 130, Unknown Unknown 140 kda myosin-1c , 3 kb Unknown Unknown myosin , 8 kb Unknown Unknown myosin-7a 17,18 Alternative splicing Seven clones isolated with alternative splicing sites present in the head and tail regions myosin-9b , 240 kda Alternative splicing Differ in carboxy-terminal tail region MYO , 5.5 kb Unknown Unknown 1 Heintzelman et al., Morgan et al., Montell and Rubin, Bonafé and Sellers, Kellerman and Miller, Halsall and Hammer, Sherr et al., Huang et al., Ruppert et al., Reinhard et al., Zhao et al., Chieregatti et al., Hasson et al., Bement et al., 1994b. 15 Bement et al., 1994a. 16 Avraham et al., Weil et al., Chen et al., Post et al., Wang et al., 1998a. 235

40 236 ANNA M. SOKAC AND WILLIAM M. BEMENT Within the myosins-1, neck lengths vary from the very short of subclass 1, which have a single IQ motif (Bement et al., 1994b), to the relatively long, as in some myosins of subclass 2 that have six IQ motifs (Ruppert et al., 1993). Neck length can also vary between isoforms of a given myosin. At least two different metazoan myosins-1, both from subclass 2, undergo differential splicing that results in variation in the number of IQ motifs. Avian brush border myosin-1 is encoded by two alternative splice forms, one of which contains three IQ motifs (Garcia et al., 1989) and one of which contains four (Halsall and Hammer, 1990). The binding of calmodulin light chains to the neck regions has been demonstrated directly using both proteolytic fragments of brush border myosin-1 (Collucio and Bretscher, 1988, 1989) and synthetic peptides (Halsall and Hammer, 1990). Similarly, rat myr1 is expressed as three alternative splice forms called myr1a, -1b, and -1c, whose necks contain six, five, and four IQ motifs, respectively (Ruppert et al., 1993). That the neck domains of these myosins can indeed bind the expected number of calmodulin light chains is indicated by the fact that partially purified myr1a binds calmodulin in an overlay assay (Ruppert et al., 1993) and by the fact that a myosin-1 independently identified by Coluccio and Conaty (1993), shown by peptide sequencing to be myr1 (Williams and Coluccio, 1994), was found to have six associated calmodulin light chains (Coluccio, 1994). 3. Variation in the Tail A significant number of unconventional myosins are subject to alternative splicing to generate isoforms that are identical with the exception of variable sequences in the tail domain (Table V). Although a few of these splice variants differ in regions in the middle of the tail, including some forms of fly myosin 95f (class 6; Kellerman and Miller, 1992), mouse dilute (class 5a; Huang et al., 1998b), and rat myr6 (class 5; Zhao et al., 1996), the majority of variants are at the carboxy tip of the tail. An alternate splice form of human myosin-9b has been identified in leukocytes that replaces a 99-amino acid proline-rich region with a novel 228-amino acid region (Post et al., 1998). Toxoplasma gondii M-B and M-C (both class 14) represent two splice forms with unique carboxy-terminal coding sequences (Heintzelman and Schwartzman, 1997). Two myosins in Drosophila, myosin-3 and myosin- 6, utilize alternative splicing to generate unique tail ends (Montell and Rubin, 1988; Kellerman and Miller, 1992). Interestingly, these myosin- 3 and myosin-6 splice variants exhibit differential spatial and temporal expression patterns, respectively (Porter et al., 1992; Kellerman and Miller, 1992).

41 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 237 A dissection of mouse dilute mutant alleles further demonstrates the importance of alternative splicing in the generation of variable tail regions and shows that such variants are often cell-type specific (Huang et al., 1998b). Out of 10 dilute alleles studied thus far, 4 were splice mutants. Among these splicing mutations were Myo5a d-105h and Myo5a 2ENUR, which encode a protein missing a tail exon. This protein is expressed in tissues including spleen and skin, but not brain. The allele-specific dilute phenotype corresponds with this tissue-specific expression, as the Myo5a d-105h and Myo5a 2ENUR mutants only exhibit pale fur color but not neuronal defects. B. Differential Myosin Expression One of the simplest means to regulate protein function is by differential expression. By turning on or off the synthesis of specific unconventional myosins, a cell or tissue can precisely control the activity of these proteins. Obviously, if a protein is absent from a given cell or tissue, it is not necessary for the various processes executed by that cell or tissue. Consequently, studies of unconventional myosin expression not only yield insights into mechanisms of regulation but also provide clues as to the likely function of specific unconventional myosins. Such studies also are a useful adjunct to genetic analyses of the role of unconventional myosins in disease, because there is often a tight correlation between the spatial and temporal distribution of a given unconventional myosin and the observed phenotype that results from the loss or inactivation of that myosin in genetic disorders. For example, myosins-3 have been identified only in the eyes of Drosophila (Montell and Rubin, 1988) and Limulus, the horseshoe crab (Battelle et al., 1998). Drosophila ninac mutants that lack or have nonfunctional myosin-3 are characterized by phototransduction defects and retinal deterioration (Porter and Montell, 1993). Likewise, myosins-7 are most highly expressed in cell types with cilia, and patients with Usher syndrome (a myosin-7 deficit, see preceding discussion) typically exhibit defects associated with such ciliated cells (Wolfrum et al., 1998). Further, the myosin-5 encoded by the dilute gene is particularly abundant in neural tissues, and the dilute phenotype, which results from myosin-5 mutations, is, in severe alleles, characterized by aberrant neuronal function (Huang et al., 1998a). On the other hand, many unconventional myosins are expressed broadly, at least when considered on a class-by-class basis. For example, myosins- 1, myosins-6, and myosins-9 are expressed in many cell types. Conversely, some tissues, such as testes, abundantly express virtually all classes of unconventional myosins. Therefore, it is difficult to draw broad conclusions about unconventional myosin function on the basis of expression alone. In spite of this, within a given class striking patterns occasionally are observed.

42 238 ANNA M. SOKAC AND WILLIAM M. BEMENT For example, a member of subclass 1 of the myosins-1, myosin-1c, is expressed in virtually every tissue type analyzed, whereas brush border myosin-1, a member of subclass 2 of the myosins-1, is largely limited to a single cell type, the enterocyte (see following discussion). The logical assumption is that some isoforms play broad roles required in most or all cell types, whereas other isoforms play more specialized roles. Of course, when considering specialized cell functions for specific proteins, it is well to keep in mind the ecclesiastical dictum, There is nothing new under the sun. In general, even highly specialized cell types do not employ processes totally foreign to other cells, but instead use modified or exaggerated forms of universal cellular processes, such as endocytosis, exocytosis, ion transport, organelle transport, etc. Thus, it is probably a mistake to assume that even myosins with very restricted patterns of expression fulfill roles that are limited only to the cell type within which they are expressed. It is also important to keep in mind other more mundane considerations when evaluating expression data. First, there is not always a tight correlation between RNA expression and protein expression. For example, at the level of RNA, human myosin-1c is most abundant in the kidney (Bement et al., 1994b), but at the level of protein, it is only moderately expressed in this organ (Skowron et al., 1998). However, the situation in the thymus is exactly the opposite. Second, when very specific probes are used, it is possible that some proteins will not be detected. For example, many of the different classes of myosins are represented by multiple isoforms, and different RNA and antibody probes to a given isoform may not necessarily cross-react with another isoform. Thus, in the following cases where a given tissue is described as lacking a particular myosin class, it may be that it actually does express that myosin, but that its expression was not detected simply because it does not cross-react with the probe employed. 1. Tissue and Cell-Type Distribution a. Myosins-1 Myosins-1 are the most abundant and most diverse of the unconventional myosins (Coluccio, 1997). As noted previously, the myosins- 1 can be separated into four subclasses on the basis of their primary structure. All cell types analyzed so far express at least one representative of these different myosin-1 subclasses, and most express representatives from all four subclasses (Table VI). On this basis and the fact that unicellular organisms, such as budding yeast (Goodson et al., 1996; Geli and Reizman, 1996), Dictyostelium (Titus et al., 1989), and Acanthamoeba (Maruta et al., 1979), express two or more myosin-1 isoforms, it is generally believed that most myosins-1 play roles that are associated with eukaryotic cell housekeeping rather than cell-type- or tissue-specific functions.

43 TABLE VI Tissue Distribution of Myosins-1 16 Heart Brain Placenta Lung Liver Stomach Skeletal muscle Spinal cord Kidney Inner ear Pancreas Spleen Thymus Prostate Testis Ovary Intestine Small intestine Large intestine Leukocytes Subclass 1 myr3 (Rat) 1 Myosin-1c (human) 2 Subclass 2 myr1a (Rat) 3 myr1b (Rat) 3 myr1c (Rat) 3 MM-1 (mouse) 4 BB M-1 (chicken) 5 BB M-1 (cow) 6 BB M-1 (mouse) 7 BB M-1 (human) 8 Subclass 3 M-1B (Drosophila) 9 M-1 (bullfrog) M-1 (cow) 13 MM-1 (mouse) 4 Myr2 (rat) 14 Subclass 4 M-1A (Drosophila) 9 myr4 (rat) 15 MM-1 (mouse) 4 1 Stöffler et al., Bement et al., 1994b. 3 Ruppert et al., Sherr et al., Garcia et al., Kawakami et al., Skowron and Mooseker, Skowron et al., Morgan et al., Solc et al., Metcalf et al., Gillespie et al., Zhu et al., Ruppert et al., Bähler et al., , RNA expression detected;, no expression detected;, protein expression detected; blank, expression not determined.

44 240 ANNA M. SOKAC AND WILLIAM M. BEMENT Within the myosins-1, the different subclasses display some differential expression in adult tissues, although the information available at this time reveals no striking patterns. For example, human myosin-1c and rat myr3 (members of subclass 1) are expressed in virtually every cell type analyzed so far (Bement et al., 1994b; Stöffler et al., 1995; Skowron et al., 1998). In contrast, rat myr1a c and mouse myosin-1 (members of subclass 2) are much more restricted in expression (Ruppert et al., 1993; Sherr et al., 1993). Myosins-1 of subclasses 3 and 4 apparently are expressed more broadly than those of subclass 2 and less broadly than those of subclass 1 (Table VI). Whereas most of the myosins-1 are broadly expressed, brush border myosin-1 displays a highly restricted pattern of expression in that it is found almost exclusively in enterocytes, the absorptive epithelial cells that line the lumen of the gut (Bement and Mooseker, 1996). In the enterocytes it is localized to the interface between the f-actin and plasma membrane in microvilli (see Section III.B.3). Brush border myosin-1 is most abundant in the enterocytes of the small intestine and somewhat less abundant in the enterocytes of the large intestine (Heintzelman and Mooseker, 1990; Skowron et al., 1998; Skowron and Mooseker, 1999). This difference in expression correlates with the difference in microvillar length between the small and large intestines: the microvilli in the small intestine typically are twice as long as those in the large intestine. Whereas transcripts for brush border myosin-1 have been detected by ultrasensitive methods in other tissues, such as liver and testes (Kawakami et al., 1992; Balish and Coluccio, 1995), and a protein with some of the biochemical characteristics of brush border myosin-1 has been detected in kidney brush borders (Coluccio, 1991), quantitative comparisons of brush border myosin-1 RNA or protein levels suggest that if brush border myosin-1 is expressed in tissues besides intestine, it is only at extremely low levels (Hoshimaru and Nakanishi, 1987; Garcia et al., 1989; Skowron et al., 1998; Skowron and Mooseker, 1999). One possible explanation for this discrepancy is that these other tissues express alternative splice forms of brush border myosin-1 or closely related myosin-1 isoforms that are not recognized by the probes available (Balish and Coluccio, 1995). Just as brush border myosin-1 displays the most restricted pattern of expression in adult cells among all of the myosins-1 studied, it also has the most striking temporal pattern of expression during early development and terminal differentiation. During early development, brush border myosin- 1 is not detectable until approximately day 7 in chicken embryo enterocytes (Shibayama et al., 1987) and approximately day 18 in mouse embryo enterocytes (Skowron and Mooseker, 1999). During terminal differentiation in human cells, which takes place along the crypt villus axis in the gut (Bement and Mooseker, 1996), brush border myosin-1 and myosin-1c display a fascinating pattern of reciprocal expression. In the crypt, which houses the

45 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 241 immature enterocytes, brush border myosin-1 expression is relatively low, whereas myosin-1c expression is relatively high (Skowron et al., 1998). This situation is exactly reversed in the villus, the location of mature (fully differentiated) enterocytes, suggesting that whereas brush border myosin- 1 is required for processes dispatched by the quiescent, mature enterocyte, myosin-1c is more important in the processes characteristic of the rapidly dividing immature enterocyte. Therefore, it will be of considerable interest to determine whether tumors of the gut (and other tissues) characteristically express high levels of myosin-1c, reflecting the lack of differentiation of cancerous cells. The developmental expression of Drosophila myosins-1a (a member of subclass 4) and -1B (a member of subclass 3) has also been characterized (Morgan et al., 1994, 1995). Whereas both of these myosins are expressed in the adult fly, they are most highly expressed in the gut during the larval stages, coincident in space and time with the formation of a well-developed brush border (Morgan et al., 1995). Myosin-1B is also found in the egg chamber at more or less constant levels throughout oogenesis. Within the egg chamber, it localizes to the follicle cells, which are known to possess microvilli (Morgan et al., 1995). Both the gut and the follicle cell expression patterns are consistent with roles for these myosins-1 in maintaining microvillar structure or in carrying out microvillar function. b. Myosins-3 The myosins-3 are the most highly restricted class of myosin with respect to expression, both in terms of organism and tissue, although this may very well change as more and more organisms are studied. Known representatives of myosins-3 have only been identified in Drosophila and the horseshoe crab, Limulus (Table VII). Myosin-3 was first found in Drosophila, where it was shown to underlie the ninac phenotype (Montell and Rubin, 1988), which is characterized by abnormal phototransduction and degeneration of the rhabdomere. The Limulus myosin-3 was identified more recently (Battelle et al., 1998). In both cases myosin-3 expression is limited to the eye (Table VII), consistent with the observed phenotype of ninac mutants. Interestingly, Limulus myosin-3 undergoes differential phosphorylation in response to manipulation of the light dark cycle, suggesting that it may also be involved in phototransduction. c. Myosins-5 On the basis of a number of independent approaches, it now appears that, like the myosins-1, myosins-5 are broadly expressed. Myosins-5 have been identified in chickens (Espreafico et al., 1992) and squid (Molyneaux and Langford, 1997) by biochemical approaches, in yeast ( Johnston et al., 1991) and mice (Mercer et al., 1991) by genetic approaches, in plants by immunological approaches (Miller et al., 1995), and in humans (Bement et al., 1994a), rats (Zhao et al., 1996), and Drosophila (Bonafé

46 TABLE VII Tissue Distribution of Myosins-3 through Myosins Heart Brain/CNS Placenta Lung Liver Stomach Skeletal muscle Spinal cord Kidney Inner ear Pancreas Spleen Thymus Prostate Testis Ovary Intestine Small intestine Large intestine Leukocytes Eye Myosins-3 ninac (Drosophila) 1 M-3 (Limulus) 2 Myosins-5 M-5 (Drosophila) 3,4 M-5 (chicken) 5 dilute (mouse) 6 myr6 (rat) 7 Myosins-6 95F (Drosophila) 8 242

47 M-6 (pig) 9 M-6 (rat) 9 M-6 (mouse) 10 M-6 tx 8 (human) 10 M-6 tx 6 (human) 10 Myosins-7 M-7a (mouse) 11,12 M-7 (rat) 13 M-7a (human) 12,14,15 Myosins-9 myr5 (rat) 16,17 myr7 (rat) 16 M-9b (human) 18 Myosins-15 MYO15 (mouse) 19 MYO15 (human) 20 1 Porter et al., Battelle et al., Bonafé and Sellers, MacIver et al., Espindola et al., Mercer et al., Zhao et al., Lantz and Miller, Hasson and Mooseker, Avraham et al., Gibson et al., Liu et al., Hasson et al., Weil et al., Chen et al., Chieregatti et al., Reinhard et al., Wirth et al., Probst et al., Wang et al., 1998a. 21, RNA expression detected;, no expression detected;, protein expression detected; blank, expression not determined. 243

48 244 ANNA M. SOKAC AND WILLIAM M. BEMENT and Sellers, 1998; MacIver et al., 1998) by molecular biological approaches. Although the isoform diversity of myosins-5 does not approach that of the myosins-1, two myosins-5 are found in yeast (Haarer et al., 1994). Similarly, comparison of different partial and complete myosin-5 sequences from mice, rats, and humans suggests that mammals may have at least three myosin-5 isoforms (Zhao et al., 1996). With respect to expression in adult tissues, chicken and mouse myosins- 5 are found in several other tissues, but are most abundant in the brain. High levels of brain expression are, of course, consistent with the phenotype of severe alleles of the dilute gene. Conversely, rat myr6 (myosin-5) is not found in the brain, but is abundant in heart, lung, liver, and other tissues where the dilute myosin-5 is not expressed. Presumably, when the different myosin-5 isoforms for these organisms are all identified and their expression characterized, the different orthologues may show more consistent patterns of expression across species. In Drosophila there is no information concerning the protein distribution of myosin-5; however, Northern blot analysis indicates that myosin-5 expression is limited almost exclusively to the germ line (MacIver et al., 1998), and in situ hybridizations suggest that some myosins-5 may be expressed in the hindgut of late-stage embryos (Bonafé and Sellers, 1998). d. Myosins-6 In contrast to myosins-1 and myosins-5, myosins-6 are not present in budding yeast and have not been identified in either Dictyostelium or Acanthamoeba, suggesting that the myosins-6 may be specific to the metazoa. Like the myosins-5, the myosins-6 are less diverse than the myosins-1, although there is evidence for alternative forms (Table V). Whereas the Snell s Waltzer phenotype circling behavior as a result of inner-ear degeneration might lead one to expect the expression of myosins-6 predominantly in the ear, they are in fact rather broadly expressed (Table VII). Porcine and rat myosins-6 are expressed in virtually every tissue analyzed, although the absolute levels vary considerably within different tissues (Hasson and Mooseker, 1994). In human tissues the expression of myosin-6 mrna is only slightly more limited than its expression in pig or rat. Whereas human myosin-6 mrna is expressed within the inner ear, it does not appear to be more abundant there than it is in the brain on the basis of RT-PCR analysis (Avraham et al., 1997). Likewise, although there are high levels of myosin-6 in the ear, in mouse this message appears equally abundant in kidney and is prominently expressed in brain, liver, and testes (Avraham et al., 1995). Drosophila is the only system in which the expression of myosin-6 during development has been studied in detail. It is expressed throughout the life of the fly at more or less constant levels, although it tends to be most abundant in later stage embryos (Kellerman and Miller, 1992). Within the

49 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 245 adult, it is highly expressed in nervous tissues, whereas in the embryo it is found in a variety of cell types. e. Myosins-7 Myosins-7 have been found in several vertebrates, including humans, mice, and frogs (Bement et al., 1994a; Gibson et al., 1995; Weil et al., 1995; Solc et al., 1994). They have also been identified in nematodes (Baker and Titus, 1997), and a partial myosin sequence similar to that of myosin-7 has been isolated from Dictyostelium (Schwarz et al., 1999). Defects in the genes for myosin-7 underlie the Shaker-1 phenotype in mice and Usher syndrome, a form of progressive deafness/blindness, in humans (see Section II.E). Consistent with these phenotypes, the myosins-7 are highly expressed in the eyes and ears of mice (Weil et al., 1995; Hasson et al., 1997b) and rats (Hasson et al., 1995). It is also expressed at moderate to high levels in testes, lung, and kidney (Gibson et al., 1995; Hasson et al., 1995). Notably, all of these tissues are abundant in cell types that contain microvilli, cilia, or both, and, as will be described later, myosins-7 typically are found in such structures. Analysis of myosin-7 expression during early mouse embryogenesis is also consistent with a role for these proteins in cilia and microvilli, as it is restricted to epithelial cell types containing microvilli or cilia throughout development (Sahly et al., 1997; Self et al., 1998). f. Myosins-9 Myosins-9 have only been identified in vertebrates, including rats (Reinhard et al., 1995; Chieregatti et al., 1998), humans (Bement et al., 1994a; Wirth et al., 1996), and frogs (Sokac and Bement, manuscript submitted). Two isoforms of myosin-9, human myosins-9a and -9b, originally were identified as short sequences from the head domain in a PCR screen for human unconventional myosins (Bement et al., 1994a). Myosin- 9a corresponds to rat myr7 (Chieregatti et al., 1998), whereas myosin-9b corresponds to rat myr5 (Reinhard et al., 1995; Wirth et al., 1996). Human myosin-9b is broadly expressed, but is most abundant in peripheral blood leukocytes and the spleen (Wirth et al., 1996). Interestingly, induced differentiation of the HL-60 cell line (a leukocyte model) results in a striking ( 5-fold) increase in the level of myosin-9b expression. Rat myr5 also is widely expressed and is most abundant in spleen, lung, and testes (Chieregatti et al., 1998). myr7, in contrast, although also abundant in the spleen, is much less abundant in lung, spleen, and liver than myr5 and much more abundant in the brain. Thus, these two myosins-9 display reciprocal expression, similar to the relationship between brush border myosin-1 and myosin-1c in the human gut (Skowron et al., 1998; see preceding discussion). g. Myosins-14 Myosins-14 were discovered in Toxoplasma gondii, a protozoan parasite, where they are represented as three isoforms encoded by

50 246 ANNA M. SOKAC AND WILLIAM M. BEMENT two different genes (Heintzelman and Schwartzman, 1997). The myosins- 14 also are expressed in the amoeba, Plasmodium falciparum (Pinder et al., 1998). Although there are no published reports of metazoan orthologues of these unusually small myosins, we have found, using a PCR screen for unconventional myosins, that myosin-14 is expressed in Xenopus laevis (Sokac and Bement, manuscript submitted). The pattern of myosin-14 expression during Xenopus oogenesis mirrors that of other housekeeping cytoskeletal proteins, such as actin and myosin-2, suggesting that myosin- 14 plays a constitutive role during early amphibian development (Sokac and Bement, manuscript submitted). h. Myosins-15 Like the myosins-14, myosins-15 have been discovered relatively recently (Probst et al., 1998; Wang et al., 1998a). Consequently, expression information about these myosins is very limited. Like myosins- 6 and myosins-7, mutant forms of myosins-15 are associated with genetic deafness in mice (Probst et al., 1998) and humans (Wang et al., 1998a). Therefore, it is not surprising that myosins-15 are expressed in the inner ear. Additionally, mouse myosin-15 is expressed in brain, liver, and kidney (Probst et al., 1998). Human myosin-15 is expressed at high levels in the brain and low levels in a number of other tissues (Wang et al., 1998a). 2. Subcellular Localization Another simple means for cells to regulate protein function is to control the distribution of that protein within a cell. This gives the cell more flexibility than simply turning expression on or off, in that the protein can be shuttled about from place to place as needed. Regulation by redistribution is also much faster than regulation at the level of expression. From the experimental standpoint, if studying expression at the cellular and tissue levels is useful in identifying putative myosin functions, then looking at subcellular localization is even more useful. Of course, the attempt to infer function from subcellular localization is subject to many of the concerns already listed for extrapolation of function from mutant phenotypes, and so localization data must be interpreted cautiously. For example, punctate staining within the cytoplasm often is assumed to be vesicular staining and indicative, therefore, of a role in membrane transport. However, myosin-2 staining is often punctate (Verkhovsky et al., 1995; Kolega, 1998), yet these punctae are not associated with membranes but rather result from self-association of the myosins. Further, it has been observed that myosin-5a punctae in neuronal cells are not detergentextractable (Evans et al., 1997), implying that these structures may not be vesicular in nature. In addition, as with immunoblotting and RNA expression analysis, certain technical concerns must be kept in mind. First, differ-

51 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 247 ent antibodies directed against the same protein may yield different staining patterns. Likewise, a single antibody may give drastically different staining patterns when different fixation conditions are employed. Even if staining is observed consistently in a given pattern, one may be misled as to the site of protein function if staining was conducted on resting cells. That is, a number of proteins, including myosins, undergo rapid, reversible translocation within the cell in response to specific stimuli or cellular situations, and identification of their site of action therefore depends on catching them during activation. Finally, as noted by others, the location of a given motor protein may not necessarily predict its most important site of action (Lin et al., 1996). a. Myosins-1 Because myosins-1 are the most diverse of the unconventional myosins, perhaps it is not surprising that they display the greatest variation in intracellular distribution. In the amoeboid protozoa, Dictyostelium and Acanthamoeba, they typically are found in the cytoplasm, at the plasma membrane, in the cortex, on small cytoplasmic vesicles, in pseudopodia and lamellipodia at the leading edge, in phagocytic cups, and associated with the contractile vacuole (Table VIII). In the metazoa, an even greater diversity of localization patterns is seen. Depending on the isoform in question, the cell type, and the specific situation, metazoan myosins-1 have been localized to a variety of intracellular structures and organelles, including microvilli, golgi, the plasma membrane, endosomes, the nucleus, cell cell contacts, and focal adhesions (Table IX). Although there is significant overlap in terms of subcellular localization between the different subclasses (at least one representative from each subclass is localized to microvilli, for example), and a given isoform may be found in several different places, there are some general patterns. For example, only the members of subclass-1 have been found to be localized to cell cell contacts and the adherens junction (Stöffler et al., 1995, 1998; Skowron et al., 1998). The brush border myosins-1 and Drosophila myosins-1a and -1B are the only myosins-1 that are localized almost exclusively to the microvilli. (Although a small fraction of brush border myosin-1 is associated with golgi-derived vesicles, this may simply represent newly synthesized brush border myosin- 1 that is en route to the microvilli.) Further, ectopic expression of brush border myosin-1 in cells that normally lack both a brush border and brush border myosin-1 nevertheless results in these proteins becoming localized in microvilli (Footer and Bretscher, 1994; Collins and Matsudaira, 1995). Surprisingly, given the localization of several different protozoan myosins-1 to the plasma membrane of locomoting cells, myosin-1 and myr1 are the only metazoan myosin-1 isoforms that have been found to have significant localization in areas of dynamic plasma membrane activity (Wagner et al., 1992; Ruppert et al., 1995). That is, whereas several reports

52 248 ANNA M. SOKAC AND WILLIAM M. BEMENT TABLE VIII Subcellular Localization of Unconventional Myosins in Unicellular Organisms 12 Myosin Subcellular localization Method Acanthamoeba Myosin-1A 1,2 Cytoplasm, cell cortex, small cytoplasmic vesicles IF/IEM Myosin-1B 1,2 Dynamic regions of the PM, large vacuole IF/IEM membranes Myosin-1C 2,3 PM, large vacuole membranes, contractile vacuole IF/IEM membrane Dictyostelium myob 4,5 Pseudopodia/lamellipodia of the leading edge, IF phagocytic cups, f-actin-rich surface crowns in adherent cells myoc 6 Pseudopodia of the leading edge IF myod 7 Pseudopodia of the leading edge IF Aspergillus myoa 8 Hyphal tip, PM IF/CF S. cerevisiae Myo2 9 A cap at or near the PM of unbudded cells, in IF small buds, or at schmoo tips, the neck just prior to cytokinesis and septation Myo4 10 Bud IF Myo5 11 Actin patches IF 1 Baines et al., Baines et al., Baines and Korn, Novak et al., Fukui et al., Jung et al., Jung et al., McGoldrick et al., Lillie and Brown, Jansen et al., Goodson et al., IF, immunofluorescence; IEM, immunoelectron microscopy; CF, cell fractionation; PM, plasma membrane. have described punctae of other myosin-1 isoforms at the plasma membrane (Lewis and Bridgman, 1996), only myosin-1 and myr1 display continuous leading edge staining (Table IX) reminiscent of the myosins-1 from the amoeboid protozoa. Given the ability of myosins-1 to bind membranes, often it is assumed that the punctate cellular staining patterns represent localization to intracellular organelles. This is certainly the case for some of the myosins-1; however, in other cases punctate localization is observed even after detergent extraction. Further, the fact that punctate staining of the plasma membrane can be observed (Evans et al., 1997) implies that at least some of the punctae may represent clusters or plaques of myosin-1 rather than myosin-1 associated with vesicles. As noted previously (see Section II.D), by clustering multiple myosins together, it may be possible to achieve greater processivity (Ostap and Pollard, 1996). One of the more surprising patterns of myosin-1 localization reported is that described by Nowak et al. (1997), who provided convincing evidence

53 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 249 that a putative myosin-1 isoform is localized to the nucleus of several different cell types. Presently, the significance of this observation remains to be determined. However, given the long-standing but mysterious observation that actin is a common nuclear component (Clark and Rosenbaum, 1979), it will be of great interest to determine whether this myosin-1 acts as a nuclear motor. The preceding discussion focused on stable patterns of myosin-1 localization. However, at least two myosins-1 undergo regulated redistribution. The first is brush border myosin-1, which undergoes a developmentally regulated shift from the basolateral domain to the apical microvilli during enterocyte differentiation (Fath et al., 1990; Heintzelman and Mooseker, 1990; Peterson et al., 1993). How this occurs is unknown, although one possibility is that it associates with a binding protein that becomes localized to the microvilli. If so, the fact that ectopically expressed brush border myosin-1 localizes to microvilli in nonepithelial cells suggests that such binding proteins must be present in other cell types as well. myr3 also undergoes regulated redistribution. Specifically, following the treatment of cells with a lectin, concanavalin A (ConA), myr3 is targeted to sites of cell cell contact (Stöffler et al., 1995). Targeting of myr3 to cell cell contacts can also be achieved when cells are induced to form adherens junctions by transient transfection with a constitutively active form of Cdc42, a rho family GTPase (Stöffler et al., 1998). Surprisingly, this induced localization does not require the SH3 domain of the myr3 tail, although it does require the portion of the tail that is immediately downstream of the neck as well as a functional motor domain. These findings indicate that regulated localization is the combined result of an unidentified determinant in the myr3 tail as well as actin-dependent ATPase activity. b. Myosins-3 The ninac gene, which encodes the Drosophila myosins- 3, gives rise to two alternative proteins of 132 and 174 kda (Montell and Rubin, 1988). As noted previously, both are expressed exclusively in the retinal photoreceptor cells. In these cells, phototransduction is accomplished by the rhabdomere, a rod-shaped stack of microvilli. Within the photoreceptor cells, the 132-kDa protein is found predominantly in the cytoplasm, whereas the 174-kDa protein is found predominantly in the rhabdomere (Table X; Hicks and Williams, 1992; Porter et al., 1992). Similar to the results described earlier for myr3, mutant analysis shows that motor activity is at least partly required for localization of p174 to the rhabdomere. Part of the tail domain is also required based on the fact that p132, which lacks much of the tail of p174, does not localize to the rhabdomere (Porter and Montell, 1993). The myosin-3 kinase domain is not responsible for localization to the rhabdomere, because both p132 and p174 have identical

54 TABLE IX Subcellular Localization of Metazoan Myosins-1 19 Myosin Cell type(s) Subcellular localization Method Subclass 1 myr3 (rat) 1,2 Rat intestinal epithelium, retinal Zonula adherens IF sensory epithelium NRK Elongate structures at sites of cell cell contact, cytoplasmic IF punctae Myosin-1c (human) 3 Caco-2BBe Cell cell contacts, cytoplasmic punctae, microvilli Subclass 2 myr1 (rat) 4 NRK Ruffling edges, cytokinetic furrow, microvilli IF MM1- (mouse) 5,6 Primary fetal rat cervical ganglion Cytoplasmic punctae, tubulovesicular structures in the cell IF/IEM neurons body, nonvesicle clusters on PM and/or f-actin of the growth cone BB M-1 (chicken) 7 10 Chicken intestinal epithelium Cross-link between f-actin and PM in microvilli, a IF/IEM/CF subpopulation of golgi-derived vesicles BB M-1 (mouse) 11 Mouse intestinal epithelium Microvilli IF BB M-1 (human) 3 Human intestinal epithelium, Microvilli IF/CF Caco-2BBe 250

55 Subclass 3 M-1B (Drosophila) 12 Drosophila intestinal epithelium Microvilli IF/IEM/CF M-1 (bullfrog) Frog inner-ear hair cells Cytoplasmic punctae within the hair cell body, pericuticular IF/IEM necklace, stereocilia tips and rootlets M-1 (cow) 16, 17 MDCK, CHO, NRK Cytoplasmic punctae often enriched in the perinuclear region, IF dynamic regions of the cell periphery, including filopodia and ruffles PC12, primary rat brain cells Cytoplasmic punctae, growth cones IF Swiss 3T3 Cytoplasmic punctae often enriched in the perinuclear region, IF/IEM dynamic regions of the cell periphery, including filopodia and ruffles, inside the nucleus myr2 (rat) 4 NRK Cytoplasmic punctae IF Subclass 4 M-1A (Drosophila) 12 Drosophila intestinal epithelium Microvilli IF/CF myr4 (rat) 18 Rat brain neurons Cell body and apical dendrite punctae IF 1 Stöffler et al., Stöffler et al., Skowron et al., Ruppert et al., Lewis and Bridgman, Raposo et al., Matsudaira and Burgess, Drenckhahn, Heintzelman and Mooseker, Fath and Burgess, Skowron and Mooseker, Morgan et al., Hasson et al., 1997a. 14 Gillespie et al., Metcalf, Wagner et al., Nowak et al., Bähler et al., IF, immunofluorescence; IEM, immunoelectron microscopy; CF, cell fractionation; PM, plasma membrane. 251

56 TABLE X Subcellular Localization of Myosins-3 through Myosins Myosin Cell type(s) Subcellular localization Method Myosins-3 ninac p174 (Drosophila) 1,2 Drosophila photoreceptor Rhabdomeres IEM ninac p132 (Drosophila) 1,2 Drosophila photoreceptor Extrarhabdomeral cytoplasm IEM Myosin-3 (Limulus) 3 Limulus lateral eye photoreceptor Cell body including the rhabdom IF Myosins-5 Myosin-5 (chicken) 4 10 Chick brain Small cytoplasmic (synaptic?) vesicles CF/IEM Chicken intestinal epithelium Distal tips of microvilli, f-actin-rich region at the base of the microvilli IF Rat Purkinje cells Dendrites; perinuclear region of cell body IF Rat cerebral neurons Small synaptic vesicles CF Primary rat hippocampal neuron Punctae in axons, dendrites, and perinuclear region of the cell body IF Guinea pig inner-ear neurons Synaptic terminals contacting hair cells, afferent processes IF MDCK, OC-k3, B16-F10, primary rat Centriole pericentriolar material in interphase, punctae on spindle in IF hippocampal glial cells M-phase dilute (mouse) Primary mouse fibroblasts and melanocytes, Cytoplasmic punctae enriched at MTOC in interphase, punctae on IF Swiss 3T3, B16 F10 spindle in M-phase Primary rodent cervical ganglion neurons Punctae in growth cone, small microtubule-bound organelles, actin IF/IEM filaments, PM B16 F10 Cytoplasmic punctae, PM, ER, melanosomes IF/IEM Myosins-6 95F (Drosophila) Drosophila embryonic CNS neurons Punctae in axons IF Drosophila syncytial embryo Punctae in cortical cytoplasm and pseudocleavage furrow, posterior pole IF Myosin-6 (chicken) 18 NRK Dynamic regions of PM, golgi, perinuclear region, macropinosomes IF/IEF/CF A431 following EGF stimulation Ruffles, contraction fibers IF Myosin-6 (pig) 4,10,19 Mouse kidney proximal tubule f-actin-rich region at the base of the microvilli IF Frog inner-ear hair cells Stereocilia rootlets, cytoplasmic punctae, cuticular plate, pericuticular IF/IEM necklace Guinea pig inner-ear hair cells Cell body cytoplasm, cuticular plate IF Chicken intestinal epithelium f-actin-rich region at the base of the microvilli IF Myosin-6 (human) 20 Mouse inner-ear hair cell Cuticular plate IF 252

57 Myosins-7 Myosin-7a (mouse) Human, mouse, rat photoreceptors Near the PM of the connecting cilium IEM Mouse, rat retinal pigmented epithelium Microvilli, cell body cytoplasm IEM Guinea pig inner-ear hair cells Stereocilia, cuticular plate, cell body cytoplasm IF Mouse olfactory supporting cells Apical surface, microvilli IF/IEM Mouse olfactory sensory cells Cilia IEM Mouse kidney proximal tubule epithelium Microvilli IEM Mouse kidney distal tubule epithelium Proximal region of cilia IEM Mouse lung bronchial epithelium Proximal region of the cilia and microvilli IEM Mouse testis Sertoli cells Ectoplasmic specializations surrounding the developing spermatozoan IEM head Mouse intestinal epithelium Distal tips of the microvilli IEM Bovine photoreceptors Axonemes CF Myosin-7a (human) 4,24 Frog inner-ear hair cells Pericuticular necklace, stereocilia, base of the stereocilia at sites IF/IEM coincident with extracellular links between adjacent stereocilia Rat, mouse, guinea pig inner-ear hair cells Stereocilia, cuticular plate, cell body IF Mouse testis Sertoli cells Ectoplasmic specializations surrounding developing spermatozoan head IF Myosins-9 myr5 (rat) 25,26 Rat brain neurons Cell body, dendrites NRK Cytoplasmic punctae enriched in the perinuclear region, stress fibers, IF dynamic regions of the PM myr7 (rat) 25 Rat brain neurons Cell body, dendrites IF myosin-9b (human) 27 Differentiated HL-60 Cytoplasmic punctae enriched in perinuclear region IF 1 Hicks and Williams, Porter et al., Battelle et al., Hasson et al., 1997a. 5 Espindola et al., Espreafico et al., Espreafico et al., Prekeris and Terrian, Evans et al., Heintzelman et al., Wu et al., 1998b. 12 Evans et al., Nascimento et al., Lantz and Miller, Kellerman and Miller, Mermall and Miller, Mermall et al., Buss et al., Hasson and Mooseker, Avraham et al., Liu et al., Wolfrum et al., Hasson, Hasson et al., 1997b. 25 Chieregatti et al., Müller et al., Wirth et al., Antibodies used for subcellular localization were raised against the named myosin from the organism listed parenthetically. IF, immunofluorescence; IEM, immunoelectron microscopy; CF, cell fractionation; PM, plasma membrane; ER, endoplasmic reticulum; MTOC, microtubule-organizing center. 253

58 254 ANNA M. SOKAC AND WILLIAM M. BEMENT kinase domains and deletion of the kinase domain does not prevent localization of p174 at the rhabdomere (Porter and Montell, 1993). c. Myosins-5 Because of the interest in the dilute phenotype, the intracellular distribution of myosins-5 has been the subject of considerable study. In general, the subcellular distribution of myosin-5 has been investigated in three general cell types or tissues: (1) neuronal/brain tissue, based on the neuronal deficit phenotype of severe dilute alleles, (2) melanocytes and other pigment-containing cells, based on the pale pigment phenotype of weaker dilute alleles, and (3) cultured cell lines, based on their wellcharacterized cytoskeletal and organelle systems. i. Neuronal/Brain Distribution The most consistent subcellular localization pattern observed for myosins-5 in neurons is in small punctae. These punctae are located in the dendrites and perinuclear region of the Purkinje cells (Espreafico et al., 1992) and in the cell body, axons, dendrites, and growth cones of primary cultures of mammalian neurons (Table X). Although the identity of these punctae has not been established, punctate staining is commonly thought to represent association with membranous organelles (see preceding discussion). Consistent with this notion, a detailed biochemical analysis of myosin-5 distribution in rat brain synaptosomes demonstrated that myosin-5 associates with a subpopulation of synaptic vesicles via a Ca 2 -dependent interaction with the synaptobrevin synaptophysin complex (Prekeris and Terrian, 1997). Similarly, Evans et al. (1998) used immunoelectron microscopy to demonstrate that myosin-5 in rat brain is associated with vesicles, most of which ( 80%) are also immunoreactive for SV2, a synaptic vesicle protein. This should not be taken to mean that most of the brain myosin-5 is associated with synaptic vesicles, however, because half or more is associated with other membrane fractions (Evans et al., 1998). The nature of these other fractions has yet to be defined, although some of the vesicles presumably are derived from the plasma membrane where myosin-5 has been shown to localize in growth cones (Evans et al., 1997). It is possible that some of it may be endoplasmic reticulum because myosin-5 has been found in association with endoplasmic reticulum from extruded squid axoplasm (Tabb et al., 1998). ii. Melanocyte Distribution In the melanocytes of a variety of species, myosin-5 is associated with melanosomes, as demonstrated by immunofluorescence (Wu et al., 1997), immunoelectron microscopy (Wu et al., 1997), and biochemical fractionation (Rogers and Gelfand, 1998). Curiously, Nascimento et al. (1997) noted that much of the myosin-5 associated with melanosomes appeared to be localized there indirectly. That is, it seems as if the myosin-5 is associated with small vesicles or filaments that are associated with melanosomes. The significance of this observation remains to be addressed, although inspection of the micrographs published in an-

59 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 255 other study support this finding (Wu et al., 1997). As in the brain, myosin- 5 not only is restricted to melanosomes in melanocytes but also is localized at the perinuclear region and at the tips of melanocyte processes (Nascimento et al., 1997). Consistent with the observations of myosin-5 distribution in extruded squid axoplasm (Tabb et al., 1998), much of the myosin-5 in melanocytes appears to be associated with the melanocyte endoplasmic reticulum. iii. Cultured Cell Line Distribution Investigation of the distribution of myosin-5a in a variety of different cultured cell lines by two different labs led to the surprising finding that it is consistently associated with microtubule-related structures (Table X; Espreafico et al. 1998; Wu et al., 1998b). By using several different fixation protocols and two different myosin-5 antibodies, Espreafico et al. (1998) found that myosin-5 in several different cell lines is concentrated in the microtubule-organizing center (MTOC). This result was confirmed by using a GFP myosin-5a tail construct, indicating that the localization is not an immunofluorescence artifact and that the tail domain is responsible for the MTOC localization. Wu et al. (1998b), also by employing several different myosin antibodies and fixation protocols, found that myosin-5 is localized both to the centrosome and to punctae that colocalize with microtubules in interphase cultured cells. During M-phase, myosin-5 associates with the spindle. It was further demonstrated that this staining pattern was insensitive to extraction with nonionic detergent, implying that the association was not indirect by way of a vesicle or other membranous structure. This finding is consistent with results indicating that myosin-5 may interact directly with a kinesin-like, microtubule-based motor (see Section III.G; Huang et al., 1999). In a study of unconventional myosin distribution in A431 cells, regulated redistribution of myosin-5 was observed (Buss et al., 1998). Specifically, it was demonstrated that myosin-5 is predominantly cytosolic in resting cells, with some localization at areas of the plasma membrane undergoing ruffling. However, following stimulation of the cells with epidermal growth factor (EGF), which triggers increased ruffling as well as filapodia formation at the plasma membrane, myosin-5 is recruited rapidly and dramatically from the cytosol to the newly formed ruffles and filapodia. In summary, myosins-5 display a variety of different distributions, depending on the cell type. In many cell types, they colocalize with membranous organelles, including melanosomes, synaptic vesicles, and endoplasmic reticulum. In other cell types, they display detergent-insensitive association with microtubules and/or microtubule-based structures. In yet other cell types, myosins-5 localize to actively ruffling areas of the plasma membrane in what appears to be a regulated fashion. It remains to be seen how the differences in intracellular distribution from cell type to cell type are controlled. In many cases, it is not clear which of the three putative myosin-

60 256 ANNA M. SOKAC AND WILLIAM M. BEMENT 5 isoforms is being studied, possibly contributing to the differences in subcellular localization found in different studies. d. Myosins-6 The intracellular distribution of myosins-6 has garnered far less attention than the myosins-5, in spite of the fact that they have been linked to genetic disease in the mouse (see Section II.E). In Drosophila nurse cells, myosin-6 is localized to small particles as well as to the ring canals (Bohrmann, 1997). In the syncytial embryo of Drosophila, myosin- 6 is localized to small particles of uncharacterized composition (Mermall et al., 1994). These particles undergo cell-cycle-regulated movement that apparently is powered by myosin-6. During interphase, myosin-6 is found throughout the cytoplasmic regions surrounding the syncytial nuclei but is concentrated in areas where f-actin and microtubules are associated with the nuclei (Mermall and Miller, 1995). During prophase, myosin-6 localizes to the presumptive metaphase furrows prior to the localization of f-actin at these sites. Later, in metaphase, myosin-6 apparently is lost from these furrows, whereas f-actin is still localized there. In other words, the appearance of myosin-6 at the metaphase furrows that separate the nuclei of the syncytium precedes the localization of f-actin there, whereas the disappearance of myosin-6 at the furrow precedes the disappearance of f-actin (Mermall and Miller, 1995). Myosin-6 also is concentrated at the posterior pole of the syncytial embryo (Lantz and Miller, 1998) and is found in association with small particles and putative organelles in neurons and hemocytes. The finding that myosin-6 interacts with and colocalizes with CLIP-190, a microtubule-binding protein, implies that at least some aspect of the localization of myosin-6 is microtubule-dependent. On the other hand, microtubule depolymerization has much less effect than actin depolymerization on myosin-6/clip-190 localization at the posterior pole, suggesting that f-actin is more important than microtubules for the maintenance of pole localization (Lantz and Miller, 1998). In vertebrates, the intracellular distribution of myosin-6 has only been described in a handful of cell types. In either undifferentiated or differentiated LLCPK cells (cultured kidney epithelial cells), myosin-6 is largely soluble (cytosolic) and does not associate to any great degree with f-actincontaining structures, as assessed by both immunofluorescence and cell fractionation (Hasson and Mooseker, 1994). In contrast, myosin-6 is associated with the apical microvilli of proximal tubule cells in the adult mouse kidney (Hasson and Mooseker, 1994) and with the microvilli of the avian intestinal brush border (Heintzelman et al., 1994), implying that the LLCPK cell line may lack key localization determinants for myosin-6 in situ. Like myosin-5, myosin-6 has also been shown to undergo EGF-dependent redistribution. In resting A431 cells, myosin-6 was found to localize predominantly to the golgi apparatus as well as being present in the cytosol and

61 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 257 having a small pool localized to dynamic regions of the plasma membrane (Buss et al., 1998). Immunoelectron microscopy showed that plasmamembrane-associated myosin-6 was most highly concentrated in microvilli, ruffles, and other surface protrusions. In response to EGF treatment, the pool of plasma-membrane-associated myosin-6 increased dramatically, as described previously for myosin-5 (Buss et al., 1998). Importantly, EGF treatment also resulted in phosphorylation of myosin-6 at the TEDS site, a region of the heavy chain implicated in the regulation of myosins-1 from Acanthamoeba, yeast, and Dictyostelium (see Section III.D.2). e. Myosins-7 The initial demonstration that defects in the gene encoding myosins-7 result in genetic deafness in mice and deaf blindness in humans prompted a number of studies designed to characterize the intracellular distribution of myosins-7 in these sites of presumptive function (i.e., ear and eye tissue). In mammalian retina, myosin-7a is concentrated in the connecting cilium of the photoreceptor cells (Liu et al., 1997). Within the ear, it is found in the stereocilia of the hair cells as well as in the pericuticular necklace (Hasson et al., 1995, 1997a; see following discussion). Patients with Usher syndrome 1B display a broad variety of pathologies associated with ciliated tissues. In addition to degeneration of the cochlear and retinal tissues, Usher 1B patients have been reported to have decreased motility of sperm and defective nasal cilia (Wolfrum et al., 1998). This observation led to the proposal that myosin-7a is required for normal ciliary function (Gibson et al., 1995). Wolfrum et al. (1998) tested this notion by immunoelectron microscopy analysis of a broad variety of cilia from many different sources. They found that myosin-7a is indeed a common component of cilia and typically is concentrated toward the base of the cilia. Wolfrum et al. (1998) also found that myosin-7 was a common component of microvilli. In addition, myosin-7 is localized in the f-actin-rich ectoplasmic specializations of the Sertoli cells of the testes (Hasson et al., 1997b). Thus, myosin-7 typically is found in association with areas of the plasma membrane that are underlaid by highly ordered arrays of f-actin or microtubules. f. Myosins-9 There is relatively little information available on the subcellular distribution of myosins-9. In resting HL-60 cells, a model for leukocyte differentiation, most of the human myosin-9b is localized to the f-actinrich cortex (Wirth et al., 1996). Following differentiation, myosin-9b both redistributes to a more diffuse cytoplasmic localization and concentrates in a perinuclear spot that may correspond to the golgi. Similarly, in NRK cells, rat myr5 localizes both to the cell cortex and to punctae within the cytoplasm, including perinuclear staining in the region of the golgi (Müller et al., 1997).

62 258 ANNA M. SOKAC AND WILLIAM M. BEMENT 3. Localization of Multiple Myosins in a Single Cell Type Studies wherein several unconventional myosins are localized within a single cell type are particularly informative. Such analyses have been performed on both the intestinal epithelium (Heintzelman et al., 1994) and the inner-ear hair cell (Hasson et al., 1997a). These cell types are excellent subjects in this capacity because the cell contains many functional domains that have been fairly well-characterized. Four myosins have been localized within the hair cell by combined immunofluorescence and immunoelectron microscopy. In stereocilia, which are fingerlike projections that extend from the apical plasma membrane with actin bundles at their core, myosin-1 colocalizes in the region where gated ion channels concentrate (Hasson et al., 1996; Metcalf, 1998). Myosin-7a colocalizes with extracellular cross-links at the base of the stereocilia that are assumed to lend stability to the structure by linking each stereocilium to the adjacent stereocilium. The proximal end of the actin-bundle core of the stereocilia is embedded in an actin filament meshwork near the apex of the cell known as the cuticular plate. Myosin-6 is found within the cuticular plate in position to cross-link actin filaments of the stereocilia core and actin filaments of the meshwork, firmly anchoring the stereocilia (Hasson et al., 1997a). In the intestinal brush border cell, the distribution of four myosins has been determined by immunofluorescence, electron microscopy, and biochemical fractionation (Fig. 7). Brush border myosin-1 localizes to the apical projections, referred to as microvilli, where it tethers the actin-bundle core to the plasma membrane. Vesicle-associated brush border myosin-1 also is located within the terminal web, a meshwork of actin filaments that surrounds and anchors the roots of the microvilli (Drenckhahn, 1988). Myosin-2 cross-links the roots of adjacent microvilli within the apex of the cell body to lend structural rigidity to the microvilli (Mooseker, 1985). Myosin-2 is also localized within a circumferential bundle of actin filaments that encircles the cell apex and contacts the adherens junctions. Myosin-6 is concentrated at the base of the microvilli, perhaps anchoring the microvilli or mediating some aspect of vesicle traffic. Last, myosin-5 is positioned both at the base of the microvilli and at the distal microvillar ends, where it may link the central actin bundle and the plasma membrane (Heintzelman et al., 1994). 4. Mechanisms of Subcellular Localization How are the characteristic subcellular distributions of unconventional myosins achieved? Simplistically, it might be supposed that the simple presence of an f-actin-binding domain in the head and a lipid-binding domain in the

63 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 259 FIG. 7 Schematic diagram of the distribution of myosins and f-actin in the enterocyte brush border. The brush border is divided into major domains: (1) the microvillar domain extends into the lumen of the gut, and (2) the terminal web sits immediately beneath the microvillar domain. At least four different myosin classes are found in the brush border, as indicated. tail would be enough to ensure localization in structures like microvilli, stereocilia, ruffles, and filapodia because such domains are characterized by relatively high concentrations of f-actin in close proximity to membranes. However, a number of observations argue that the situation probably is more complex. First, as described previously, members of classes 1, 5, 6, and 9 undergo regulated redistribution in response to specific stimuli and cellular conditions. Second, whereas many unconventional myosins do indeed localize to structures that are rich in both f-actin and membrane, many more do not but instead show localization to microtubule-rich domains, membranous organelles, or uncharacterized punctate structures. Third, the same myosin may show a strikingly different pattern of localization within different cell types. Fourth, as described previously for both hair cells and enterocytes, even within a specific f-actin- and membrane-rich structure, such as a stereocilium or microvillus, different unconventional myosins are not distributed uniformly but are localized to specific subdomains of that structure. These observations argue that other mechanisms are likely to contribute to the characteristic pattern of unconventional myosin localization observed within different cell types. We next consider several different

64 260 ANNA M. SOKAC AND WILLIAM M. BEMENT mechanisms. Some of these have only been identified for conventional myosins, but nevertheless may be relevant for unconventional myosins. Other mechanisms have received experimental support for sorting of unconventional myosins. One possibility is that the myosin localization determinant resides in the myosin transcript. During rapid growth of adult muscle, the myosin-2 messenger RNA (mrna) gathers at sites where there is nascent construction of functional bipolar filaments (Russell and Dix, 1992). Likewise, it has been reported that myosin-2 protein is polymerized into filaments as it emerges from the translating ribosome in cultured skeletal muscle, a process known as cotranslational assembly (Isaacs and Fulton, 1987). In neonatal cardiac myocytes, the subcellular localization of cardiac myosin- 2 to sites of filament assembly is dependent on the 3 -untranslated region (3 -UTR) of the myosin transcript (Goldspink et al., 1997). Likewise, the 3 -UTR of the rabbit skeletal muscle myosin transcript directs a reporter sequence to the perinuclear region, the site of initial sarcomere assembly in cultured myotubes (Wiseman et al., 1997). Because the subcellular distribution of their mrnas has not yet been characterized, it is unknown whether sorting of unconventional myosins relies on a similar mechanism. If it occurs posttranslationally, subcellular sorting of unconventional myosins could be controlled via activities specified by any combination of the motor domain (or by the neck domain insofar as it influences the motor domain) or the tail domain. Differential motor activity could influence localization patterns of unconventional myosins in one of several different ways. One of the simplest modes of motor-dependent control of subcellular myosin sorting was proposed by Kolega (1998) in his study of myosin-2a and myosin-2b distribution in cultured bovine aortic endothelial cells. In these cells, myosins-2a and -2B preferentially localize to the leading and trailing edges of the cell, respectively, although neither isoform is excluded from any structure. When exogenous, fluorescently labeled myosin-2a and myosin-2b were coinjected and observed by time-lapse fluorescence imaging with each exogenous isoform differentially distributed in the same pattern as for the endogenous isoform, indicating that the myosin protein (rather than the mrna) contains all of the information necessary for subcellular sorting (Kolega, 1998). In this study it was also observed that myosin-2a associated with newly forming structures and dissociated from disassembling structures more rapidly than myosin-2b. Kolega proposed that differential motor activity could account for the disparate isoform localization and rates of dynamics. That is, because myosin 2A translocates actin filaments 3 times faster than myosin 2B in vitro and has higher actinactivated ATPase activity (Kelley et al., 1996), the observed distributions of isoforms could reflect differences in motor properties when in the context of the cytoplasm. No information is available as to whether unconventional

65 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 261 myosins employ a similar mechanism, although some do differ dramatically in terms of their motor properties. For example, myosins-9 are comparatively slow motors, driving f-actin movement at a rate of 15 nm/s (Post et al., 1998), whereas myosins-5 are comparatively fast, driving movement at a rate of 400 nm/s (Cheney et al., 1993). Further, Baines et al. (1995) showed that the subcellular distributions of Acanthamoeba myosins-1a, -1B, and -1C vary as a function of motor activation by heavy-chain phosphorylation, as would be expected if the ability to translocate f-actin or to rapidly pass through the mechanochemical cycle were important for subcellular localization. Differential motor activity may influence unconventional myosin localization by more subtle means as well. That is, myosins that display reduced kinetics for any given step of the ATPase cycle would be expected to spend more time in association with f-actin. For example, Ostap and Pollard (1996) showed that Acanthamoeba myosins-1a and -1B differ in their rate constants for ATP-induced dissociation from f-actin, with myosin-1a being similar to smooth muscle myosin-2 and myosin-1b being similar to skeletal muscle myosin-2. They suggested that this difference could contribute to the differences in subcellular localization and function of these two myosin- 1 isoforms. Jontes et al. (1997) found that, in brush border myosin-1, the rate constant for ATP-induced dissociation from f-actin is roughly equal to those of Acanthamoeba myosin-1a and smooth muscle myosin-2. Coluccio and Geeves (1999) found that ATP-induced dissociation of rat myr1 from f-actin is even slower, roughly 10% that of brush border myosin-1. This myosin is also relatively slow with respect to other steps of the ATPase cycle, which presumably would increase the amount of time it spends in association with f-actin and, consequently, the cortical f-actin cytoskeleton (Coluccio and Geeves, 1999). The role of the motor domain in determining subcellular localization has been tested directly for several unconventional myosins. For example, subcellular localization of myr1, myr2, and myr3 is altered when these proteins are expressed as headless truncates, but to different degrees. myr3, which normally localizes to actin-rich structures at cell cell contacts, is unable to localize at such structures in the absence of the head (Stöffler et al., 1998). On the other hand, myr1 and myr2, both of which are found at least in part at intracellular membranes, still localize when expressed as headless truncates, albeit to a lesser degree than the endogenous myr1 and myr2 (Ruppert et al., 1995). In these myosins, then, it appears that both the head and the tail regions are required for normal localization, but that the relative contributions of each varies among the different isoforms. A similar situation was observed within the myosins-3. The two different splice forms of the ninac protein (p134 and p174) differ only in their tail regions, yet the 174-kDa form is found in the f-actin rich rhabdomeres,

66 262 ANNA M. SOKAC AND WILLIAM M. BEMENT whereas the 134-kDa form is found in the cytoplasm. Presumably, then, it is the tail that contains the sorting information. Nevertheless, ninac head mutants fail to localize to the rhabdomere, indicating that this protein requires contributions from both the head and the tail for proper subcellular sorting (Porter and Montell, 1993). Of course when headless truncates are used, it cannot be determined whether subcellular localization is influenced by myosin motor activity, by the ability of myosin to bind f-actin, or simply by myosin head sequences. Several studies suggest that myosin motor activity is at least partially dispensable for proper subcellular localization. For example, Dictyostelium myosin-2 mutants that cannot hydrolyze ATP still concentrate in the cleavage furrow of the dividing cell at a rate equal to that for wild-type myosin- 2 (Yumura and Uyeda, 1997), as do those lacking the entire head domain (Zang and Spudich, 1998). Similarly, Novak and Titus (1997) demonstrated that Dictyostelium myob (a myosin-1) displays a normal subcellular distribution when the heavy-chain phosphorylation site required for motor activation is mutated to a nonphosphorylatable residue, making the motor nonactivatable. Finally, a detailed study of Shaker-1 mutant alleles, the gene that encodes mouse myosin-7a, showed that regardless of the predicted severity of a mutation on myosin motor activity, myosin-7a subcellular localization was not disrupted, at least at the resolution of fluorescence microscopy (Hasson et al., 1997b). Likewise, f-actin binding, an activity attributed to the head and considered diagnostic for myosin function, is, in some cases, entirely dispensable for subcellular myosin localization. For example, Xenopus nonmuscle myosin-2a is recruited to the borders of puncture wounds in the absence of f-actin (Bement et al., 1999). Further, in budding yeast, myosins-2 localize to the incipient cytokinetic furrow in an f-actin-independent manner (Lippincott and Li, 1998; Bi et al., 1998). Thus, the contribution made by the myosin head to subcellular localization remains poorly defined. In metazoan myosins, it is widely thought that the tail is responsible for proper targeting, although this point has been tested directly in only a handful of cases. For example, as noted previously, myr1 and myr2 localize to their normal intracellular positions when expressed as headless truncates, although not as completely because some of the truncates partition into the soluble fraction (Ruppert et al., 1995). Tail-dependent subcellular targeting of metazoan myosins-1 is also suggested by the finding that the tail of brush border myosin-1 can partially displace myosin-1 from membrane fractions of hepatoma cells (Raposo et al., 1999). Overall, the evidence for tail-directed subcellular targeting is strongest for the myosins-5. In budding yeast, a point mutation in the tail disrupts normal targeting of myo2 to sites of polarized growth and to vacuoles (Catlett and Weisman, 1998). Further, expression of tail constructs displaces

67 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 263 the endogenous myo2p from sites of polarized growth (Reck-Peterson et al., 1999). Similarly, when expressed in mouse melanocytes, fusions of GFP with the tail of myosin-5a are targeted to melanosomes, a site where myosin- 5 normally localizes (Wu et al., 1998a). In addition, several dilute alleles have been shown to map to the tail of myosin-5a (Huang et al., 1998b). The preceding findings highlight the importance of the unconventional myosin tails as determinants of subcellular distribution. How, exactly, do the tails confer specific patterns of localization upon unconventional myosins? Although there is no definitive answer to this question, evidence suggests that at least two mechanisms may be operative binding of acidic phospholipids and binding of myosin docking proteins. Acidic phospholipid binding by myosin tail domains has been reported for vertebrate conventional myosins (Murakami et al., 1994), as well as for protozoan unconventional myosins. In the latter, phospholipid binding has been mapped to the TH1 region of the tail, which is rich in basic amino acids (Adams and Pollard, 1989; Doberstein and Pollard, 1992). Binding of acidic phospholipids by metazoan unconventional myosins has been documented for brush border myosin-1. Consistent with the results from the protozoan myosins-1, the tail appears to mediate binding, although the binding region has yet to be mapped precisely (Hayden et al., 1990). The fact that unconventional myosin tails can bind to pure phospholipids via regions in their tails suggests that such interactions may contribute to the observed tendency of many unconventional myosins to localize to membranous structures in vivo. However, as noted previously, it is unlikely that the mere tendency to bind acidic phospholipids can explain the diversity of unconventional myosin localization patterns. In particular, it seems unlikely to account for the finding of differential unconventional myosin localization within subdomains of structures like the brush border or hair cell stereocilia. Rather, it seems more likely that this binding may promote initial association of unconventional myosins with membranes, with the final distribution being dictated by interaction of the myosin in question with other cytoplasmic cues. Evidence that such cytoplasmic cues are provided by myosin docking proteins comes from several different sources. Xu et al. (1995) identified a 125-kDa protein that binds to the SH3 domain of Acanthamoeba myosin- 1C and colocalizes with this myosin on intracellular organelles. This protein subsequently was shown to bind to myosin-1c by virtue of tandemly arrayed proliné-rich regions (Xu et al., 1997). This is consistent with the general observation that SH3 domains mediate protein protein interactions via proline-rich regions in their binding partners (Ren et al., 1993). Similarly, the SH3 domain of S. cerevisiae myo5p (class 1) has been shown to interact with the proline-rich protein, verprolin, as demonstrated by yeast two hybrid analysis and immunoprecipitation (Anderson et al., 1998). Further, myo5p

68 264 ANNA M. SOKAC AND WILLIAM M. BEMENT is mislocalized in a verprolin deletion mutant, suggesting that verprolin is the cytoplasmic receptor for myo5p. In support of this hypothesis, verprolin is distributed asymmetrically in budding yeast and colocalizes with myo5p (Anderson et al., 1998). The importance of the SH3 domains in myosin-1 tails for normal subcellular targeting apparently is not universal. For example, a Dictyostelium myob mutant lacking its SH3 domain nevertheless properly localizes when expressed in vivo (Novak and Titus, 1997). Similarly, removal of the SH3 and the TH1 region from the myr3 tail did not prevent this myosin from localizing normally in cultured mammalian cells (Stöffler et al., 1998). Although no docking proteins for metazoan myosins-1 have been identified, potential candidates as docking proteins for myosin-3 and myosin-5 have been identified. A PDZ-domain-containing protein, INAD, has been shown to bind to the ninac p174, and the portion of p174 that binds to INAD is sufficient to target heterologous proteins to the rhabdomere. On the other hand, the situation is complicated by the fact that p174 distribution apparently is normal in INAD mutants, suggesting that INAD, while sufficient for docking, is not absolutely necessary. With respect to the myosins-5, as noted previously (see Section III.B.2), myosin-5 in rat brain binds to the synaptophysin synaptobrevin complex, and this binding may be responsible for its localization to synaptic vesicles because conditions that disrupt the interaction of myosin-5 with the complex result in release of the myosin- 5 from synaptic vesicles (Prekeris and Terrian, 1997). C. Ca 2 /Calmodulin Studies of Ca 2 -mediated regulation have been conducted for three unconventional metazoan myosin classes: 1, 5, and 9. From these studies, three general effects of Ca 2 have been identified. First, in contrast to most calmodulin-binding proteins, the unconventional myosins bind calmodulin in the absence of Ca 2. Myosins purified in the presence of EGTA retain their associated calmodulin light chains (Wolenski, 1995), whereas Ca 2 causes the dissociation of some, but not all, calmodulin light chains from the myosin heavy chain (Coluccio and Bretscher, 1987; Collins et al., 1990; Swanljung-Collins and Collins, 1991; Wolenski et al., 1993; Nascimento et al., 1996; Post et al., 1998). Second, disruption of the myosin calmodulin complex results in the reduced velocity of translocation of actin filaments in an in vitro motility assay (Collins et al., 1990; Wolenski et al., 1993; Cheney et al., 1993; Post et al., 1998), and some motility is restored by the addition of exogenous calmodulin (Collins et al., 1990; Wolenski et al., 1993; Cheney et al., 1993). Third, Ca 2 influences actin-activated ATPase activity

69 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 265 (Conzelman and Mooseker, 1987; Collins et al., 1990; Wolenski et al., 1993; Cheney et al., 1993; Nascimento et al., 1996). The effects of Ca 2 on chicken brush border myosin-1 have been investigated by several different labs. Whereas there are some qualitative differences between studies, there is general agreement on several different points. First, one of the major effects of Ca 2 on brush border myosin-1 is to cause the release of one or more calmodulin light chains when the free Ca 2 concentration is 10 M or higher, although the precise concentration required for dissociation varies from study to study (Collins et al., 1990; Swanljung-Collins and Collins, 1991; Wolenski et al., 1993). Second, at concentrations higher than 1 M, Ca 2 stimulates the actin-activated ATPase of brush border myosin-1 (Collins et al., 1990; Swanljung-Collins and Collins, 1991; Wolenski et al., 1993). This increase plateaus at 10 M, although again, the absolute numbers vary in the different studies. Third, and somewhat surprisingly, the Ca 2 -induced increase in actin-activated ATPase is accompanied by a decrease in the rate at which brush border myosin-1 translocates actin filaments (Collins et al., 1990; Wolenski et al., 1993). In fact, at extremely high Ca 2 concentrations (greater than 10 M), f-actin motility ceases altogether. These effects of Ca 2 are mediated via Ca 2 interaction with the light chains rather than the heavy chain, as inclusion of free calmodulin prevents the irreversible loss of motility at high Ca 2 concentrations and removal of the light-chain-binding region by limited proteolysis of the heavy chain eliminates the effects of Ca 2 on the actinactivated ATPase (Wolenski et al., 1993). Results obtained for rat myr1 were similar in some respects to those obtained for chicken brush border myosin-1, although interesting differences were also observed (Williams and Collucio, 1994). As with brush border myosin-1, Ca 2 stimulates myr1 actin-activated ATPase while inhibiting translocation of f-actin. However, the inhibition of motility is manifest at much lower free Ca 2 levels 1 M in myr1 versus 10 M or greater for brush border myosin-1. In addition, inclusion of exogenous calmodulin completely suppresses the decrease in motility triggered by up to 10 M Ca 2, rather than just rescuing it. Similar results were obtained in studies of myosin-1. As with myr1 and brush border myosin-1, Ca 2 elevation increases the actin-activated ATPase (Barylko et al., 1992), inhibits f-actin translocation (Zhu et al., 1996, 1998) and causes the dissociation of one of the calmodulin light chains (Zhu et al., 1996). However, it was found that the concentration of Ca 2 required to increase the actin-activated ATPase activity of myosin-1 was much higher than that required to inhibit transport of f-actin (10 versus 1 M, respectively). Myosin-5 has also been studied with respect to its Ca 2 sensitivity (Cheney et al., 1993). Again, as with the myosins-1 discussed previously, Ca 2 stimu-

70 266 ANNA M. SOKAC AND WILLIAM M. BEMENT lates the actin-activated ATPase (Nascimento et al., 1996) while slowing translocation of f-actin (Cheney et al., 1993). Similar to brush border myosin- 1, inclusion of exogenous calmodulin in the assay permits the rescue of motility when Ca 2 levels are lowered following exposure of myosin-5 to high Ca 2 (Cheney et al., 1993). Myosin-9 also shows Ca 2 -dependent slowing of f-actin translocation (Post et al., 1998), although it is not yet known what effects Ca 2 has on the myosin-9 actin-activated ATPase. In contrast to the unconventional myosins described previously, Ca 2 apparently inhibits the actin-activated ATPase of myr3 (Stöffler and Bähler, 1998). Further, the single calmodulin light chain of this subclass 1 myosin- 1 remains bound to the heavy chain in the presence of elevated Ca 2. The reason for these differences is unknown. However, because Ca 2 elevation only causes the release of some of the calmodulins from brush border myosin-1 (one or two out of four) and myosin-1 (one out of three), it seems reasonable to speculate that the IQ motifs of the neck are not identical in terms of their ability to bind calmodulin at different Ca 2 concentrations. Indeed, it has been shown that the two IQ motifs in the neck of myr4 differ in their ability to bind calmodulin at different Ca 2 concentrations (Bähler et al., 1994). In any case, further characterization of the effects of Ca 2 on actin filament translocation by myr3 is required to determine whether Ca 2 can be considered as a general inhibitor of the actin-translocating activity of myosins-1. How Ca 2 binding to calmodulin light chains modulates the mechanochemistry of myosin remains poorly understood. Two proposals have been offered to explain the available data: (1) Ca 2 defines light-chain composition by inducing the reversible dissociation of the calmodulin myosin complex (Wolenski, 1995) and/or (2) Ca 2 binding to the light chains causes a conformational change that is transmitted to the heavy chain (Houdusse et al., 1996). These proposed mechanisms are not mutually exclusive, and an in vitro study suggests that both contribute to the Ca 2 /calmodulinmediated regulation of myosins. A significant increase in the rate of actinactivated and actin-independent ATPase activity of bovine myosin-1 corresponds with Ca 2 -induced dissociation of one of three calmodulins from the heavy chain. With regard to the velocity of actin filament translocation, however, the concentration of Ca 2 required for motility inhibition is less than that required for dissociation and, instead, corresponds to a major Ca 2 -induced conformational change of calmodulin (Zhu et al., 1998). In other words, the increase in actin-activated ATPase may result from Ca 2 - induced loss of one of the light chains, whereas the observed inhibition of f-actin translocation may result from Ca 2 -induced conformational changes in the remaining calmodulins. The in vivo significance of the interaction between Ca 2, calmodulin, and unconventional myosins remains to be assessed. However, work in fungi

71 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 267 suggests that this mode of myosin regulation is more subtle than predicted by the in vitro data. Yeast myo2 (class 5) associates with mutated calmodulin that lacks Ca 2 -binding capacity, yielding a phenotype indistinguishable from that of cells with wild-type calmodulin (Brockerhoff et al., 1994). Furthermore, yeast myo5 (class 1) and calmodulin (Cmd1p) are essential in the process of receptor-mediated endocytosis, and Cmd1p associates with myo5 via the myo5 IQ motifs. Interestingly, deletion of the myo5 IQ motifs (myo5 IQ1d2dp) does not significantly inhibit receptor-mediated endocytosis (Geli et al., 1998). Thus, it seems that Ca 2 regulation of the unconventional myosins is dispensable for some cellular processes. These data do not exclude the possibility that the completely naked neck domain (i.e., completely devoid of associated light chains) of myosin is autoinhibitory and must be masked by a light chain to attain full myosin activity, as has been suggested previously (Nascimento et al., 1996; Geli et al., 1998), nor do they exclude the possibility that phenotypes resulting from elimination of light-chain binding are too subtle to be easily detected. D. Phosphorylation Regulation of myosins by phosphorylation has been studied intensively in a variety of model organisms. Whereas most of the work has been conducted on myosins-2, regulation of protozoan myosins-1 has been studied in some detail, and there are hints that the findings from such studies will be relevant to metazoan unconventional myosins. 1. Conventional Myosins a. Light-Chain Phosphorylation A wealth of studies have documented the effects of light-chain phosphorylation on the activity of smooth muscle and nonmuscle myosins-2 and, indeed, such phosphorylation events are thought to be the major mechanism by which these myosins are regulated (Table XI). Briefly, each myosin-2 heavy chain binds one essential light chain (ELC) and one regulatory light chain (RLC). Thus, the whole myosin- 2 molecule comprises two pairs of heavy chains, two RLCs, and two ELCs. In smooth muscle and nonmuscle myosins-2, the RLCs are subject to phosphorylation of the activating residue (serine19 in most vertebrates), which has two effects increased actin-activated ATPase activity and increased filament formation (Bresnick, 1999). The RLCs may also be subject to phosphorylation on inhibitory residues (serine1,2 or threonine9 in most vertebrates), which may reduce actin-activated ATPase activity or inhibit subsequent phosphorylation of the activating residues (Bresnick, 1999).

72 TABLE XI Effects of Phosphorylation on Conventional Myosin Activity 33 Actin-activated Myosin Pi site Kinase Filament assembly Mg 2 -ATPase In vitro motility Dictyostelium M2 1 3 HC-T1823 MHCK Inhibits filament assembly HC-T1833 HC-T2029 Dictyostelium M2 4,5 HC MHC-specific PKC Inhibits filament assembly Dictyostelium M2 6,7 RLC-S13 MLCK No effect Acanthamoeba M HC-S1489 No effect HC-S1494 HC-S1499 Vertebrate SMM RLC-S19 MLCK Promotes filament assembly 34 RLC-T18 Vertebrate SMM RLC-S1 PKC (following 34 No effect MLCK Pi) RLC-S2 RLC-T9 Vertebrate SMM2 17,19 RLC-S1 PKC (only) No effect 34 No motility RLC-S2 RLC-T9 Vertebrate SMM2 20 RLC-S19 Protease-activated kinase I Vertebrate SMM2 21 HC-S1954 Casein kinase II Vertebrate SMM2 22 RLC-S19 rho-kinase Vertebrate NMM2 14,17,23 25 RLC-S19 MLCK Promotes filament assembly Vertebrate NMM2 17,26,27 RLC-S1 PKC (following No effect 35 No effect MLCK Pi) RLC-S2 RLC-T9 Vertebrate NMM2 17,27 RLC-S1 PKC (only) No effect No motility RLC-S2 RLC-T9 268

73 Vertebrate NMM RLC-S1 Cyclin-p34 cdc2 RLC-S2 RLC-T9 Vertebrate NMM2A 30 HC-S1944 Casein kinase II No effect 36 Vertebrate NMM2A HC-S1917 PKC No effect 36 Vertebrate NMM2B 30 HC-S1975 Casein kinase II Inhibits filament assembly 36 HC-S1965 HC-T1960 HC-S1956 HC-S1952 Vertebrate NMM2B 30 HC-S1922 PKC Inhibits filament assembly 36 HC-S1935 HC-T1937 HC-S1938 HC-S1939 HC-S1941 Vertebrate NMM2B 30 HC-S1975 Casein kinase II Promotes filament assembly 36 HC-S1965 HC-T1960 HC-S1956 Vertebrate NMM2B 30 HC-S1922 PKC Inhibits filament assembly 36 HC-S1933 HC-S1938 HC-S1939 HC-S Vaillancourt et al., Lück-Vielmetter et al., Kuczmarski and Spudich, Ravid and Spudich, Dembinsky et al., Ostrow et al., Griffith et al., Côté et al., Hammer et al., Collins et al., Kuznicki et al., Ganguly et al., Ikebe and Hartshorne, Craig et al., Sellers, Sellers et al., Umemoto et al., Bengur et al., Nishikawa et al., Tuazon and Traugh, Kelley and Adelstein, Amano et al., Bresnick, Scholey et al., Adelstein and Conti, Kawamoto et al., Ikebe and Reardon, Yamakita et al., Satterwhite et al., Murakami et al., Conti et al., Murakami et al., HC, heavy chain; RLC, regulatory light chain; MHCK, myosin heavy chain kinase; MLCK, myosin light-chain kinase; PKC, protein kinase C. 34 Assays used heavy meromyosin. 35 Effect based on K a not V max. 36 Assay used a tail fragment of the respective myosin. 269

74 270 ANNA M. SOKAC AND WILLIAM M. BEMENT The in vivo requirements for light-chain phosphorylation have been assessed in several organisms. In Dictyostelium, RLC nulls are defective in cytokinesis and other processes thought to be dependent on myosin- 2 (Chen et al., 1994). Surprisingly, however, RLC nulls can be rescued by expression of RLC that cannot be phosphorylated at the activating residue (Ostrow et al., 1994). Likewise, replacement of the endogenous fission yeast myosin-2 RLC with a nonphosphorylatable mutant has no apparent effect on the timing or execution of cytokinesis, nor does replacement with RLC containing substitutions of acidic amino acids (to mimic phosphorylation) at the RLC phosphorylation sites (McCollum et al., 1999). In Drosophila, on the other hand, expression of nonmuscle myosin-2 RLC that cannot be phosphorylated on the activating residue results in slight defects in cytokinesis and major defects in other actomyosin-dependent developmental events ( Jordan and Karess, 1997). Thus, the relative importance of light-chain phosphorylation in vivo varies not only from organism to organism but also for different cellular events within a single organism. b. Heavy-Chain Phosphorylation Rather less is known about heavychain phosphorylation of myosins-2, although it has been the subject of intensive study in Dictyostelium and Acanthamoeba (Brzeska and Korn, 1996). In Dictyostelium, regulated assembly of myosin-2 into bipolar filaments is dependent on both the central regions and the carboxy-terminal regions of the tail. More specifically, the central region of the tail is necessary and sufficient to drive assembly (O Halloran et al., 1990; Lee et al., 1994), but the assembly process is controlled by a domain at the end of the tail. Phosphorylation of three threonines in the regulatory region near the carboxy terminus inhibits filament assembly by stabilizing an intramolecular interaction between tail regions (Vaillancourt et al., 1988; Egelhoff et al., 1993). This yields a bent tail conformation that is assembly-incompetent (Pasternak et al., 1989). Whereas failure to polymerize does not directly alter the motor activity of these myosins, it does prevent these myosins from acting as contractile units. Moreover, filament formation is directly correlated with the ability of these myosins to be recruited to the cytokinetic apparatus (Sabry et al., 1997). Acanthamoeba myosin-2 is also subject to heavy-chain-tail phosphorylation, and as with Dictyostelium myosin-2, the phosphorylation has an inhibitory effect (Table XI; Collins and Korn, 1980). However, this effect appears to be at the level of the actin-activated ATPase more directly than is the case for Dictyostelium myosins-2. That is, in Acanthamoeba myosin-2, three serine residues near the carboxy terminus, in a region called the tailpiece, are phosphorylated (Collins et al., 1982) and the resulting phosphorylation suppresses motor activity (Ganguly et al., 1992). Suppression of the motor

75 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 271 activity apparently is due to conformational changes in the head that are thought to result from interaction of the head with the tail (Kuznicki et al., 1983). This interaction apparently is favored within the context of an intact filament, rather than within isolated myosin-2 molecules, because the effects of tail phosphorylation on actin-activated ATPase are much more pronounced in the myosin filament (Ganguly et al., 1992). As in the myosins-2 from the protozoa, the heavy-chain phosphorylation sites for vertebrate nonmuscle myosins are near the carboxy-terminal portion of the tail (Kelley and Adelstein, 1990; Murakami et al., 1990, 1998; Conti et al., 1991). The effects of the phosphorylation depend upon both the isoform of myosin in question and the residues phosphorylated. Phosphorylation of the tail of myosin-2a is not known to have any effects on filament assembly, whereas phosphorylation of isoforms of the myosin-2b tail can, under certain circumstances, inhibit filament assembly. Specifically, when a single serine in the tail is phosphorylated by protein kinase C (PKC), myosin-2b and myosin-2b filament formation is inhibited. Likewise, when multiple serines are phosphorylated by casein kinase II, myosin- 2B filament formation is inhibited (Murakami et al., 1998). Interestingly, the region phosphorylated by PKC and casein kinase II is also the site of interaction with the myosin-2-regulating protein, Mts1 (see Section III.G; Kriajevska et al., 1998). 2. Unconventional Myosins a. Heavy-Chain Phosphorylation Virtually all of the information available for unconventional myosin phosphorylation indicates that, when regulation by phosphorylation does occur, it occurs on the heavy chain rather than the light chains (Table XII). In fact, whereas calmodulin has been reported to be the target of phosphorylation under some circumstances (Corti et al., 1999), there are no reports of phosphorylation of unconventional myosin calmodulin light chains. The regulatory effects of heavy-chain phosphorylation for the unconventional myosins that are best understood are those that follow phosphorylation of the head rather than the tail. Specifically, actin-activated ATPase activity of all of the myosins-1 from Acanthamoeba and Dictyostelium studied to date absolutely requires phosphorylation of the heavy-chain head (Brzeska and Korn, 1996). The site of phosphorylation is a serine or threonine residue residues from the amino terminus, depending on the myosin-1 in question. This residue is contained within the HCM loop, one of the two loops of the myosin head known to contact actin. The position occupied by this residue has been referred to as the TEDS site on the basis of the observation that in other myosins, the corresponding

76 TABLE XII Effects of Phosphorylation on Unconventional Myosin Activity 13 Actin-activated Myosin Pi site Kinase(s) Mg 2 -ATPase In vitro motility Dictyostelium myob 1 HC Acanthamoeba M1-HCK Dictyostelium myod 2 HC Dictyostelium M1-HCK Dictyostelium myod 3 HC STE20, CLA4, PAK-3 Acanthamoeba M-1B 4,5 HC-S315 Acanthamoeba M1-HCK Acanthamoeba M-1C 6 HC-S329 Acanthamoeba M1-HCK Acanthamoeba M-1C 7 HC Human PAK-1 S. cerevisiae Myo3 8 HC-S STE20, CLA4 Aspergillus myoa 9 HC-S Drosophila ninac 10 ninac Vertebrate BB M-1 11 PKC Vertebrate myosin-6 12 PAK-3 1 Côté et al., Lee and Côté, Wu et al., Albenesi et al., Brzeska et al., Wang et al., 1998b. 7 Brzeska et al., Wu et al., Yamashita and May, Ng et al., Swanljung-Collins and Collins Buss et al., HC, heavy chain; HCK, heavy chain kinase; PKC, protein kinase C; PAK, p21-activated kinase. 14 Assay used a head fragment. 15 Based on site-directed mutagenesis that resulted in a phenotype.

77 REGULATION AND EXPRESSION OF METAZOAN MYOSINS 273 residue almost invariably is phosphorylatable (S or T) or acidic (D or E; Fig. 8). In the absence of heavy-chain phosphorylation, Acanthamoeba and Dictyostelium are completely motor-inactive they lack both actin-activated ATPase and the ability to translocate f-actin (Maruta and Korn, 1977; Albanesi et al., 1985). Following heavy-chain phosphorylation, this situation is reversed and the myosins express full motor activity. It is not clear exactly how phosphorylation at the TEDS site activates myosins-1. A negative charge at the TEDS site has no obvious effect on the static structure of Acanthamoeba myosin-1c, relative to myosins-1c with an uncharged amino acid in this position (Carragher et al., 1998). On the basis of kinetic studies, Ostap and Pollard (1996) concluded that the most likely step to be affected by TEDS site phosphorylation is P i release, because there were no significant differences in the rate constants of the steps preceding phosphate release in phosphorylated versus unphosphorylated myosins-1. As noted in Section II.D, the P i release step of the ATPase cycle is the target of regulation in other myosins, making this suggestion plausible. The importance of this phosphorylation has been confirmed by studies in which the TEDS site residue is made acidic (to mimic phosphorylation) or nonphosphorylatable. Myosins-1 from unicellular organisms that cannot be phosphorylated at the TEDS site are inactive both in vitro (Wang et al., 1998b) and in vivo (Novak and Titus, 1998). Myosins-1 with an acidic substitution at the TEDS site display constitutive actin-activated ATPase in vitro (Wang et al., 1998b) and, when expressed in Aspergillus, drive constitutive endocytosis (Yamashita and May, 1998). Moreover, in budding yeast, a mutant myo3 with the TEDS site serine substituted with aspartate can rescue a myo3 myo5 double mutant, whereas myo3 with alanine at this site cannot (Wu et al., 1997). b. TEDS Rule and Its Implications As discussed previously, the site of myosin-1 heavy-chain phosphorylation in myosins-1 from unicellular organisms is contained within the HCM loop, one of the two flexible surface loops known to contact f-actin. Multiple sequence alignment of the HCM loop from myosins-1 of amoeboid organisms with the HCM loop in other myosins revealed a TEDS rule. The TEDS rule is simply the observation that in nearly all myosins, the residue that corresponds to the residue phosphorylated in amoeboid myosins-1 is either phosphorylatable (S or T) or acidic (E or D) (Fig. 8; Bement and Mooseker, 1995). There are two immediate implications of the TEDS rule: (1) a negative charge at the TEDS site is critical for actin-activated ATPase activity, and (2) any myosin having a phosphorylatable residue at the TEDS site is predicted to be reversibly activated by phosphorylation (Bement and Mooseker, 1995). The position of the TEDS site at the tip of the HCM loop, which is known

78 274 ANNA M. SOKAC AND WILLIAM M. BEMENT