Functional specificity of actin isoforms

CYTOLOGY V202 - AP - 5155 / C2-35 / 09-02-00 06:38:04

Functional Specificity of Actin Isoforms Sofia Yu. Khaitlina Institute of Cytology, Russian Academy of Sciences, St. Petersburg 194064, Russia

Actin, one of the main proteins of muscle and cytoskeleton, exists as a vaiety of highly conserved isoforms whose distribution in vertebrates is tissue-specific. Synthesis of specific actin isoforms is accompanied by their subcellular compartmentalization, with both processes being regulated by factors of cell proliferation and differentiation. Actin isoforms cannot substitute for each other, and the high-level synthesis of exogenous actins leads to alterations in cell organization and morphology. This indicates that the highly conserved actins are functionally specialized for the tissues in which they predominate. The first goal of this review is to analyze the data on the polymerizability of actin isoforms to show that cytoskeleton isoactins form less stable polymers than skeletal muscle actin. This difference correlates with the dynamics of actin microfilaments versus the stability of myofibrillar systems. The three-dimensional actin structure as well as progress in the analysis of conformational changes in both the actin monomer and the filament allows us to view the data on the structure and polymerization of isoactins in terms of structure–function relationships within the actin molecule. Most of the amino acid substitutions that distinguish actin isoforms are located apart from actin–actin contact sites in the polymer. We suggest that these substitutions can modulate the ability of actin monomers to form more or less stable polymers by long-range (allosteric) regulation of the contact sites. KEY WORDS: Muscle actin, Cytoplasmic actin, Intracellular sorting, Structure–function relationship, Actin conformation. 䉷 2001 Academic Press.

I. Introduction Actin is one of the major structural proteins of eukaryotic cells that plays a key role in muscle contraction and cell motility, as well as in many other cell processes and functions. Actin microfilaments are involved in International Review of Cytology, Vol. 202 0074-7696/01 $35.00

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Copyright 䉷 2001 by Academic Press All rights of reproduction in any form reserved.

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maintaining cell shape, cell division, endocytosis, exocytosis, secretion, signal transduction, and the regulation of enzyme activities (Pollard and Cooper, 1976; Welch et al., 1998). They have been shown to regulate the activity of membrane channels (Neguljaev et al., 1996; Maximov et al., 1997) and participate in transcription (Scheer et al., 1984), mRNA transport and translation (Bassell and Singer, 1997), and synaptic transmission (Rosenmund and Westbrook, 1993; Prekesis et al., 1996). A number of functions, including cytoplasmic streaming, movement of chloroplasts, cytokinesis, karyokinesis, maintenance of cell shape and architecture, and organelle transport, are proposed for microfilaments in plants (Meagher, 1995; Kropf et al., 1998). Alterations in the organization of microfilaments result in disorganized cell arrangement and orientation, uncontrolled cell growth, and abnormal responses to the environment, all of which are characteristic of neoplastic cell transformation (Pollack et al., 1975; Kakunaga et al., 1984; Janmey and Chaponnier, 1995). In contrast to thin filaments in myofibrils of skeletal and cardiac muscles that assemble during myogenesis and exist as a stable microfilament system, the most characteristic property of actin-based cytoskeleton is its dynamics, i.e., its ability to assemble and disassemble depending on cell requirements. Assembly of actin filaments is observed when a cell is spreading on a substrate (Moreau and Way, 1999; Small et al., 1999; Watterman-Storer and Salmon, 1999), during acrosome reaction of Thyone sperm (Tilney et al., 1973, 1983), and during mating of Chlamidomonas gametes (Detmers, 1985). A striking example of intracellular assembly of actin filaments is the formation of a comet-like tail on the surface of pathogenic bacteria invading nonphagocytic mammalian cells. The rate of bacterial movement within the cytoplasm of the host cell is shown to be equal to the rate of actin polymerization (Tilney et al., 1992a,b; Theriot et al., 1992; Sechi et al., 1997; Welch et al., 1998; Gossart, 1995; Gossart and Leguit, 1998; Lasa et al., 1998). The dynamics of actin cytoskeleton is maintained by two factors: the ability of actin to undergo reversible transformation from the monomeric state (G-actin) to the polymeric state (F-actin) and the interaction of actin with actin-binding proteins, which can inhibit or stimulate actin polymerization, sever the polymers, cross-link actin filaments into bundles, and bind them to cell membrane (Sheterline et al., 1995; Jokusch and Hinssen, 1996). Whereas the interaction of actin with actin-binding proteins is discussed in numerous experimental and review papers (Way and Weeds, 1990; Bretcher, 1993; Sun et al., 1995; Puius et al., 1998; Carlier 1998; McGough, 1998), the present article will concentrate on the properties of actin itself that may underlie a higher or lower stability of the polymer and, thus, provide an actin-based mechanism for the microfilament dynamics. Multiple isoforms of actin are synthesized in different muscle and nonmuscle cells. The primary structure of actin is highly conserved: amino acid

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sequences of the nonmuscle isoactins are more than 90% identical to those of the muscle isoactins (Vandekerckhove and Weber, 1978,a,b). Nevertheless, tissue-specific and developmentally determined synthesis of actin isoforms (Garrels and Gibson, 1976; Storti et al., 1976; Wahlen et al., 1976; Rubenstein and Spudich, 1977; Wiens and Spooner, 1983; Owens and Thompson, 1986), as well as compartmentalization of actin isoforms within the cell (Pardo et al., 1983; Otey et al., 1988; Herman, 1993; Gunning et al., 1998), was observed. It stimulated a search for differences in the properties of isoactins that could correlate with their utilization in different contractile systems. Some of these results have been reviewed previously (Rubenstein, 1990; Herman, 1993; Hennessey et al., 1993). The three-dimensional structure of actin has now been solved (Kabsch et al., 1990; McLaughlin et al., 1993; Schutt et al., 1993). This information, as well as the progress in the analysis of conformational changes in the actin monomer (Strzelecka-Golaszewska et al., 1993, 1995; Crosbie et al., 1994; Muhlrad et al., 1994; Kim et al., 1995; Moraczewska et al., 1996, 1999; Chik et al., 1996; Mounier and Sparrow, 1997) and the filament (Orlova and Egelman, 1992, 1993, 1997; Orlova et al., 1995, 1997; Muhlrad et al., 1994; Strzelecka-Golaszewska et al., 1996), allows us to view the data on the structure and polymerization of isoactins in terms of structure–function relationships within the actin molecule. The aim of this article is to demonstrate how variations in the structure and conformation of isoactins could result in differences between their functional properties.

II. General Characteristics of Actin Isoforms A. Temporal and Spatial Segregation 1. Classification of Actin Isoforms Multiple forms of actin isolated from different tissues and cells form a family of eukaryotic proteins encoded by individual genes. Actin sequences are conserved to a much greater degree than those in almost any other proteins. Although amino acid substitutions are distributed throughout the actin sequences, there are several regions that are much more variable than others. One of these regions is the N-terminal segment of the polypeptide chain, which contains a cluster of acidic residues. This variability contributes to a net charge on the actin molecules, which can be detected by isoelectric focusing. According to the differences in mobilities of different actins at isoelectric focusing, actins have been classified as 움-, 웁-, and 웂-isoforms in order of increasing basicity (Garrels and Gibson, 1976; Storti et al., 1976;

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Wahlen et al., 1976; Rubenstein and Spudich, 1977). The N-terminal segments of 움-actins contain four charged amino acid residues; in skeletal muscle actin the sequence is Asp-Glu-Asp-Glu. The corresponding segments of 웁- and 웂-actin contain three acidic residues, which are AspAsp-Asp and Glu-Glu-Glu for 웁- and 웂-actin, respectively (Vandekerckhove and Weber, 1978a). These isoforms can also be distinguished by two-dimensional high-voltage paper electrophoresis of their N-terminal tryptic peptide (Vandekerckhove and Weber, 1981). Another classification of actin isoforms is based on the specificity of amino-terminal processing. The acetylated amino-terminal residue of actin (N-acetyl-Asp or N-acetyl-Glu) appears as a result of posttranslational stepwise acetylation-dependent processing reactions (Rubenstein and Martin, 1983; Solomon and Rubenstein, 1985; Rubenstein, 1990). The isoactins, called the class I molecules, are encoded by genes that specify a Met-Asp/ Glu N-terminus acetylated early in translation and then removed. The new N-terminal residue, Asp or Glu, is acetylated to yield the mature actin form. This class combines cytoplasmic 웁- and 웂-actins and smooth muscle 웂-actin. The class II molecules are encoded as polypeptides with a MetCys-Asp/Glu N-terminus. In these actins, final acetylation of the N-terminal Asp or Glu follows stepwise removal of Met and acetyl-Cys. Class II includes skeletal, cardiac, and smooth muscle 움-actins. Actins from most lower eukaryotes and invertebrates also are class II molecules (Rubenstein 1990). By taking into account the facts that the nucleotide- and cationbinding domains of class I and II isoactins are almost identical to the corresponding domains of actin-related proteins and that, in general, the actin-related proteins share 74% similarity with the mammalian isoactins, the latter are combined in the class III molecules (Herman, 1993; Mullins et al., 1996, 1998). 2. Tissue Specificity of Actin Isoforms Sequence analysis has shown that the main classes of isoactins found in mammals and birds are heterogeneous (Vandekerckhove and Weber, 1978, a,b). Although the differences between the actin species are small, there are distinct 움-actin isoforms in skeletal and cardiac muscles. One more 움isoform, smooth muscle 움-actin, is the predominant isoactin in vascular smooth muscle. In intestinal smooth muscle, the predominant actin isoform is 웂-actin, which differs from cytoplasmic 웂-actin (Table I). Nonmuscle cells contain cytoplasmic 웁- and 웂-actin isoforms (Storti et al., 1976; Rubenstein and Spudich, 1977). Table I shows that cardiac and smooth muscle 움-actins differ from their skeletal muscle counterpart by four and eight substitutions, respectively. Smooth muscle 웂-actin differs from skeletal muscle 움-actin by six substitu-

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ACTIN ISOFORMS TABLE I Amino Acid Substitutions between Mammalian Actin Isoformsa

Residue

Skeletal muscle 움-actin

1 2 3 4 5 6 10 16 17 76 89 103 129 153 162 176 201 225 260 267 272 279 287 297 299 358 360 365

Asp Glu Asp Glu Thr Thr Cys Lue Val Ile Thr Thr Val Leu Asn Met Val Asn Thr Ile Ala Tyr Ile Asn Met Thr Glu Ala

Cardiac 움-actin Asp Glu

Leu Ser

Smooth muscle 움-actin

Smooth muscle 웂-actin

Glu

–

Glu Asp Ser

Glu

Cys

Cys

Ser

Ser

Leu Ser

Leu Ser Pro

Cytoplasmic 웁-actin

Cytoplasmic 웂-actin

– Asp

– Glu

Asp Ile Ala Val Met Cys Val

Ile Ala lle Met Cys Val

Val Thr Met Thr Leu Thr Gln Ala Leu Cys Phe Val Thr Leu Ser

Val Thr Met Thr Leu Thr Gln Ala Leu Cys Phe Val Thr Leu Ser

Ser

Ser

a Empty boxes indicate that there is no substitution of the corresponding residue in comparison with that in skeletal muscle actin. A (–) at position 1 indicates that the corresponding actins do not contain any residue at this position because the amino acid sequence of 움-actins consists of 375 residues whereas those of 웁- and 웂-actins consist of 374 residues. Actin sequences were taken from Vandekerckhove and Weber (1978a,b).

tions, and cytoplasmic 웁- and 웂-actin are almost identical. The sequences of the cytoplasmic isoactins are about 10% different from those of any of the muscle isoactins. The most different form of cytoplasmic actin is yeast actin (Ng and Abelson, 1980; Gallwitz and Sures, 1981). In addition, two acidic actin isoforms, along with two common 웁- and 웂-actins, were found in the nuclei of mouse leukemia cells (Nakayasu and Ueda, 1986).

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In amphyoxus Branchiostoma lanceolatum, the closest living relative of vertebrates, at least two muscle isoforms, skeletal and smooth muscle 움actins, as well as cytoplasmic 웁-actin were found (Fagotti et al., 1998). However, most, if not all, invertebrate muscle actins are homologous to each other and to vertebrate cytoplasmic actins (Vandekerckhove and Weber, 1984; Schoenenberger et al., 1995). Specifically, mollusk muscles contain 웁-like isoactin, which, being immunologically non-cross-reactive to mammalian actins (DeCouet, 1983), differs from mammalian 웁-actin by only four substitutions (Table II) (Vandekerckhove and Weber, 1984). Several cytoplasmic actin genes are also expressed in invertebrates (Fang and Brandthorst, 1996). Specific actin isoforms are found in protozoa (Gor-

TABLE II Amino Acid Substitutions in 웁-like Scallop Adductor Muscle Actin as Compared with Skeletal Muscle and Nonmuscle Isoactins Amino acid residue

Skeletal muscle 움-actin

1 2 3 4 5 6 10 16 17 76 103 129 159 162 176 201 225 260 267 272 279 287 297 299 358 365

Asp Glu Asp Glu Thr Thr Cys Leu Val Ile Thr Val Leu Asn Met Val Asn Thr Ile Ala Tyr Ile Asn Met Thr Ala

Scallop adductor muscle 웁-like actin

Cytoplasmic 웁-actin

Asp

Asp

Asp Val Ala Val Met Cys Val Val Ala

Asp Ilea Ala Val Met Cys Val Val Thr a Meta Thr Leu Thr Glna Alaa Leu Cysa Phea Val Thr Leu Ser Ser

Thr Leu Thr Ser Leu

Val Thr Leu Ser Ser

a Amino acid substitutions that distinguish nonmuscle 웁-actin from 웁-like scallop actin. Actin sequences were taken from Vandekerckhove and Weber (1984).

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don et al., 1977; Uyemura et al., 1978; Vandekerckhove and Weber, 1978c, 1981; Vandekerckhove et al., 1984; Sussman et al., 1984; Pahlic, 1985; Hirono et al., 1987, 1990) and in plants (Meagher, 1995). An unconventional form of actin was also found in motile cyanobacteria (Labbe et al., 1996; Usmanova et al., 1998). Thus, synthesis of actin isoforms is not random but tissue-specific. Different muscle and nonmuscle motile systems contain different actin isoforms. It implies that, in differentiating muscle cells that undergo transformation from a nonmuscle precursor into the mature myotubes, cytoplasmic actin isoforms should be replaced by the muscle-specific counterparts and at the intermediate stages of differentiation different actin isoforms may coexist. 3. Transitions of Actin Isoforms during Myogenesis During both the development of skeletal muscles in vivo and the differentiation of muscle cells in culture, myoblasts line up, elongate, and fuse to form multinuclear myotubes. This process is accompanied by a change in the isoactin pattern. In early muscle development and in prefused cultured myoblasts, only cytoplasmic 웁- and 웂-actins are detected by direct analysis of the total cell extracts or in the actin samples purified by the polymerization–depolymerization procedure (Storti et al., 1976; Wahlen et al., 1976). As the development progresses, the relative amount of 움-actin increases until it becomes a predominant actin species. In chicken thigh muscle it happens by day 20 of embryonic development (Storti et al., 1976). In agreement with these data, mRNA from the 13- and 16-day embryonic thigh muscles directed cell-free synthesis of all three isoactins (Storti and Rich, 1976). Following the fusion of cultured chicken embryonic myoblasts, synthesis of 움-actin begins to increase at about 44 hr after plating. At 96 hr, when the fusion is complete and the myotubes can spontaneously contract, 움-actin becomes the major component in the culture (Rubenstein and Spudich, 1977). In addition, analysis of protein and mRNA populations provided evidence for the synthesis of cardiac 움-actin, along with skeletal muscle 움actin, in embryonic skeletal muscles (Minty et al., 1982; Paterson and Eldridge, 1982; Vandekerckhove et al., 1986; Hayward and Schwartz, 1986; Hayward et al., 1988; Lyons et al., 1991). In embryonic chicken breast muscle, both in vivo and in vitro, the cardiac 움-actin mRNA accounted for more than 90% of the sarcomeric actin transcripts, whereas 5 weeks after hatching the skeletal 움-actin mRNA was the only detectable sarcomeric actin transcript (Paterson and Eldridge, 1982). In embryonic mouse muscle, only 30–40% of mRNA was identified to be the cardiac 움-actin mRNA, 60–70% of mRNA was the skeletal 움-actin species (Minty et al., 1982), and nearly the same proportion of cardiac to skeletal muscle actins was

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determined at the protein level (Vandekerckhove et al., 1986). In cultured embryonic mouse muscles cells, accumulation of the cardiac (fetal) actin mRNA was shown to begin at the same time as the accumulation of adult skeletal muscle 움-actin mRNA, at the onset of cell fusion, and coordinately with the decline in the level of the cytoplasmic 웁- and 웂-actin mRNAs. It is followed by a stepwise rise in the levels of skeletal muscle 움-actin mRNA and protein (Minty et al., 1982; Vandekerckhove et al., 1986; Garner et al., 1989). Comparison of the quantitative data shows different proportions of cardiac 움-actin in different animals, as well as in similar muscle cells depending on whether experiments were performed at the RNA or protein level and whether cultured cells or muscles were studied. This difference can reflect different rates of mammalian and avian myogenesis, the different rates of protein and RNA turnover, and the specificity or cells in culture compared with the same cells in muscle. It is important, however, that in the adult skeletal muscles of all of the animals, cardiac 움-actin is not synthesized. On the other hand, synthesis of cardiac 움-actin was shown to be strongly activated in skeletal muscle fibers and satellite cells (undifferentiated myoblasts) of injured muscle, in correlation with muscle regeneration (Franke et al., 1996). Cardiac 움-actin is also synthesized in other muscle pathologies and in muscle-derived tumors (Franke et al., 1996). Furthermore, before the skeletal muscle 움-actin appears, transient expression of a smooth muscle 움-actin gene was observed in the mouse L6 cell line, as well as in primary mouse myogenic cells in culture (Caplan et al., 1983). Thus, both cardiac and smooth muscle 움-actins can be regarded as fetal forms during skeletal muscle development. The transition of actin isoforms during development of the heart is very similar to that occurring in skeletal muscle myogenesis. During development, five out of six mammalian genes are expressed in cardiomyocytes (Wiens and Spooner, 1983; Vandekerckhove et al., 1986). In chicken embryo, cytoplasmic 웁- and 웂-actins were shown to be synthesized in the precardiac mesoderm and adjacent ectoderm, as well as in the primitive heart. Synthesis of 움-actin was detected in the precardiac mesoderm preceding myofibrillar organization and its proportion increased with development, whereas the synthesis of 웁- and 웂-actins was reduced (Wiens and Spooner, 1983). In addition, during fetal cardiac development, skeletal muscle 움-actin and smooth muscle 움-actin are synthesized (Vandekerckhove et al., 1986; Ruzicka and Schwartz, 1988; Garner et al., 1989; van Bilsen and Chien, 1993). In mature cardiomyocytes, cardiac 움-actin becomes a strongly predominating isoform (Vandekerckhove et al., 1986). However, partial reversion occurs from the adult to the fetal phenotype, with resynthesis of smooth muscle and skeletal muscle 움-actins, during cardiac hypertrophy in vivo and in cultured cardiomyocytes from adult animals

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(Eppenberger-Eberhardt et al., 1990; van Bilsen and Chien, 1993). This resynthesis is regulated by hormones and growth factors (Harden et al., 1996, 1998; Schaub et al., 1997). Smooth muscle cells contain smooth muscle 움- and 웂-actins as well as cytoplasmic 웁- and 웂-actins in different proportions in different animals and tissues. All of these isoforms can be detected in vascular smooth muscle cells during fetal life or in culture with cytoplasmic 웁-actin being the predominant species (Franke et al., 1980; Owens et al., 1986; Skalli et al., 1987) The corresponding differentiated cells are characterized by a significant accumulation of smooth muscle 움-actin, which becomes a predominant isoform and a marker of smooth muscle cell populations and tissues (Owens and Thompson, 1986; Skalli et al., 1987). In embryonic and adult chicken gizzard muscles, one muscle isoform, smooth muscle 웂-actin, and cytoplasmic 웁- and 웂-actins are synthesized (Saborio et al., 1979; Hirai and Hirabayashi, 1983; Kuroda, 1985). Nonmuscle 웂-actin exists only at early stages of development, before 15 days of embryogenesis (Kuroda, 1985). With development of the embryo, the relative amount of muscle 웂-actin gradually rises (Saborio et al., 1979; Hirai and Hirabayashi, 1983; Kuroda, 1985). The ratio of 웂- to 웁-actin accumulated in chicken gizzard by hatching was determined to be 3.6 at the protein level (Hirai and Hirabayashi, 1983) and 2.2 when mRNAs were analyzed (Saborio et al., 1979). After hatching, the content of muscle 웂-actin slowly increased (Hirai and Hirabayashi, 1983; Kuroda, 1985). Similar to the changes observed in skeletal and cardiac muscles, synthesis of actin isoforms characteristic of the differentiated smooth muscle cells can be altered under pathological conditions or when smooth muscles undergo adaptation to some physiological processes. For instance, during pregnancy, in both the human and rat myometrium considerable elevation of the relative content of smooth muscle 웂-actin occurs with a corresponding reduction in the level of smooth muscle 움-actin (Skalli et al., 1987). A marked decrease in the content of smooth muscle 움-actin occurred in smooth muscle cells from atheromatous lesions in humans and experimental animal models (Gabbiani et al., 1984) or upon the stimulation of smooth muscle cell growth in culture (Owens et al., 1986) and in vivo (Clowes et al., 1988). In contrast, a growth arrest was associated with increased smooth muscle 움-actin synthesis (Owens et al., 1986; Strauch and Rubenstein, 1984). These data suggest an inverse relationship between the synthesis of smooth muscle 움-actin and cell proliferation or, in other words, a correlation between specific muscle cell differentiation and accumulation of the muscletype isoform (Owens et al., 1986; Owens and Thompson, 1986). In addition, the presence of the skeletal muscle 움-actin isoform was revealed at a late stage of differentiation of BC3H1 smooth-muscle-like cells (Strauch and Reeser, 1989).

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Thus, muscle cells maintain a specific distribution of coexisting actin isoforms in proportions regulated by factors of cell proliferation and differentiation (Table III). At the intermediate stages of differentiation, simultaneous synthesis of cytoplasmic and muscle-specific actin isoforms takes place. Moreover, in muscle cells, several muscle isoforms can be synthesized simultaneously. In the myogenic cells in culture, a higher level of embryonic and fetal actin isoforms is preserved, probably due to the lack of tissuespecific control and physiological stimuli like load-bearing. The two cytoplasmic isoactins also coexist in every cell. Questions arise concerning whether there is intracellular sorting of isoactins within muscle and nonmuscle cells and whether actin isoforms can substitute for each other in various cell processes. 4. Compartmentalization of Isoactins within Muscle Cells Staining of skeletal muscle cells with specific antibodies directed against different actin species revealed localization of 움-actin in the center of the cell in the contracting myofibrils, whereas cytoplasmic isoactins were found at the cell periphery in the cortical array of actin filaments (Lubit and Schwartz, 1980; Pardo et al., 1983; Otey et al., 1988). Cytoplasmic isoactins also were detected in sarcolemmal costamers where myofibrils are attached to the cell membrane and in the Z-disk lattice. (Craig and Pardo, 1983). Antibodies specific to 웂-actin revealed its colocalization with mitochondria (Pardo et al., 1983). 웁-Actin was found to associate with the motor end plate in the neuromuscular junction (Hall et al., 1981) and with acetylcholine receptor (Lubit, 1984). Ventricular adult rat cardiomyocytes cultured in the presence of growth factors or hormones synthesize, along with cardiac 움-actin, a high proportion of smooth muscle 움-actin (Eppenberger-Eberhardt et al., 1990). The cardiac actin is restricted to a central perinuclear region, whereas smooth muscle 움-actin is accumulated in nonstriated stress fibers outside the sharp boundary of the myofibrillar area (Harden et al., 1996, 1998). Similarly, cultured vascular smooth muscle cells and pericytes restrict vascular 움-actin within the stress fibers, which are counterparts of myofibrils in these cells, away from nonmuscle actins localized in regions of advancing, cortical cytoplasm (DeNofrio et al., 1989; Herman, 1993). In chicken gizzard in vivo, antibodies that cross-reacted with smooth muscle 움- and 웂-isoactins but did not react with 웁-isoform stained the myosin-containing contractile regions, whereas antibodies to cytoplasmic 웁-actin specifically stained the cytoskeletal compartments, including dense bodies where 웁-actin was colocalized with desmin and filamin (North et al., 1994). These data are consistent with the results of biochemical experiments. Myofilaments immunoprecipitated with antibodies against muscle-specific

a

In cell culture

In vivo

In cell culture

In vivo

In cell culture

In vivo

See text for more references.

Smooth

Cardiac

Skeletal

Muscle

웁- and 웂-cytoplasmic, 움-skeletal, 움-cardiac, 움-smooth 웁- and 웂-cytoplasmic, 움-skeletal, 움-cardiac, 움-smooth 웁- and 웂-cytoplasmic, 움-smooth

움-Skeletal, 웁- and 웂-cytoplasmic

웁- and 웂-cytoplasmic, 움-cardiac, 움-skeletal 웁- and 웂-cytoplasmic, 움-cardiac, 움-skeletal, 움-smooth

웁- and 웂-cytoplasmic, 웂-smooth, 움-smooth

움-Smooth, 웁- and 웂-cytoplasmic

움-cardiac, 웁- and 웂-cytoplasmic

움-Cardiac, 웁- and 웂-cytoplasmic

움-Skeletal, 웁- and 웂-cytoplasmic

Adult (differentiated)

Fetal (nondifferentiated)

TABLE III Actin Isoform Patterns at Different Stages of Myogenesis

Storti et al., 1976; Wahlen et al., 1976; Vandekerckhove et al., 1986a Rubenstein and Spudich, 1977; Vandekerckhove et al., 1986; Caplan et al., 1983a Wiens and Spooner, 1983; Vandekerckhove et al., 1986a Eppenberger-Eberhardt et al., 1990; van Bilsen and Chien, 1993a Gabbiani et al., 1984; Owens et al., 1986; Skalli et al., 1987a Franke et al., 1980; Gabbiani et al., 1984a

References

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tropomyosin were enriched in muscle-specific 움-actin (Lin and Lin, 1986). On the other hand, gentle extraction of actin from BC3H1 muscle-like cells, human rhabdomyosarcoma cells, and embryonic chicken muscle cells released abnormally large amounts of nonmuscle isoactins compared with their relative content in the cell. This suggests that nonpolymerized or easily depolymerized actin as well as cortical actin filaments preferentially consist of the cytoplasmic isoactins (Rubenstein, 1990). Thus, when multiple actin isoforms are synthesized simultaneously in muscle cells, compartmentalization takes place. Muscle isoactins are restricted to the myofibrillar systems, whereas nonmuscle isoforms are utilized in the cytoplasmic structures. 5. Intracellular Sorting of Cytoplasmic ␤- and ␥-Actins The proportion of the two cytoplasmic actin isoforms, 웁- and 웂-actins, is different in various cells (Table IV). Mammalian erythrocytes contain only 웁-actin (Pinder and Gratzer, 1983), whereas 웂-actin markedly predominates in the brush borders and microvilli from chicken intestinal epithelium (Bretscher and Weber, 1978; Vandekerckhove and Weber, 1981) and in chicken auditory hair cells (Hofer et al., 1997). However, in most cells, despite different origins and morphological differences, 웁- and 웂-actin coexist at a constant ratio of approximately 2 : 1 (Table IV). Cells appear to maintain the constant 웁- to 웂-actin ratio even if synthesis of these isoforms is reduced due to, for instance, the expression of an introduced mutant 웁-actin gene (Leavitt et al., 1987). It indicates that the mechanisms responsible for regulation of the steady-state actin level do not discriminate between the actin isoforms, and the specific ratio between the two cytoplasmic isoforms seems to be maintained by mechanisms different from the autoregulatory feedback actin concentration (Leavitt et al., 1987). Specific localization of 웁-actin versus the entire F-actin pool was revealed in microvascular pericytes (Hoock et al., 1991). Pericyte 웁-actin was found beneath the plasma membrane as well as within fan lamellae and in association with the ends of stress fibers terminating in the plasma membrane, whereas fluorescent phalloidin staining was observed preferentially along stress fibers. In endothelial and 3T3 cell monolayers migrating after injury, 웁-actin was localized in association with ruffling membranes, pseudopods, and the veil of cytoplasm present within fan lamellae. Once the cells began to contact one another and stopped moving, the anti-웁-actin staining at the leading edge disappeared. Simultaneous staining of filamentous actin with fluorescent phalloidin revealed an abundance of 웂-actin in quiescent regions that were not stained with anti-웁-actin (Hoock et al., 1991). Results of these experiments indicate that 웁-actin accumulates at the membrane–

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ACTIN ISOFORMS TABLE IV Ratio of 웁-Actin to 웂-Actin in Various Tissues and Cells Tissues and cells

웁:웂

References

Human erythrocytes Chicken auditory hair cells Chicken intestinal epithelium brush border Rabbit gastric parietal cells Human platelets Embryonic chicken brain Rat liver Bovine thymus Endothelial cells from bovine aorta Rat brain Rat heart Rat kidney Rat lung Rat spleen Rat osteoblasts Cell lines Human carcinoma (HeLa) Baby hamster kidney (BHK) Rat kangaroo kidney (PtK2) Chicken embryo fibroblasts Human fibroblasts KD and HuT12 Mouse C2 myoblasts Human smooth muscles Myometrium Myometrial leiomyoma Gastric Tracheal Tanea coli

웁 0.5 0.6

Pinder and Gratzer, 1983 Hofer et al., 1997 Vandekerckhove and Weber, 1981

1.4 2.0 1.0 2.5 2.6 2.3 1.9 6.1 2.2 1.8 1.8 2.9

Yao et al., 1995 Gordon et al., 1977

2.1 1.7 1.9 3.5–4.2 1.7 2.0 3.4 4.7 2.0 3.0 3.6

Vandekerckhove and Weber, 1981 Gabbiani et al., 1984 Otey et al., 1987

Watanabe et al., 1998 Vandekerckhove and Weber, 1981

Leavitt et al., 1987 Schevzov et al., 1992 Skalli et al., 1987

cytoskeleton interface in regions of moving cytoplasm, whereas 웂-actin seems to be restricted to stress fibers. Spatial and temporal segregation of 웁- and 웂-actins also was found in gastric parietal cells (Yao et al., 1995), auditory hair cells (Hofer et al., 1997), osteoblasts (Watanabe et al., 1998), and neurons (Weinberger et al., 1996; Ulloe and Avila, 1996: Hannan et al., 1998; Micheva et al., 1998). A feature common to all of these cells is a more ubiquitous distribution of 웂actin. The location of 웁-actin, on the contrary, depends on the physiological function of the cell and the stage of its differentiation, with clear preferences for cell processes and newly formed compartments. In chicken auditory hair cells, 웁-actin is not a predominant isoform, and its localization is specifically restricted to the actin filament core bundle of stereocilia that

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is extensively cross-linked by fimbrin (Hofer et al., 1997). Similarly, in rabbit gastric parietal cells, 웁-actin predominates in the apical plasma membrane, where its colocalization with the actin-binding protein ezrin was revealed (Yao et al., 1995). Differentiating neurons in culture show accumulation of 웁-actin in the growth cone (Ulloe and Avila, 1996; Hannan et al., 1998; Bassell et al., 1998; Micheva et al., 1998). In adult neurons, 웁-actin was found to be located in dendrites and, to a lesser extent, in the cell body, but was not detected in the axon (Weinberger et al., 1996). In the developing cerebellar cortex, the accumulation of 웁-actin was consistently observed in actively growing structures, e.g., growth cone filopodia, cell bodies, and axonal tracts. In the adult cerebellar cortex, 웁-actin was found preferentially in dendritic spines—structures known to retain their capacity for morphological modifications in the adult brain (Micheva et al., 1998). Thus, both the ratio and localization of cytoplasmic 웁- and 웂-actins are specific for distinct cells and subcellular compartments. As will be discussed later, high-level expression of incorporated 웁- and 웂-actin genes results in dramatic changes in cell phenotype emphasizing the role of cytoplasmic actins, in contrast to the muscle isoforms, in maintaining cell morphology. 6. Can Actin Isoforms Substitute for Each Other? Subcellular sorting of actin isoforms does not necessarily imply that the isoforms are functionally different, as they may be located separately but perform similar functions. On the other hand, isoforms located in the same cell compartment may be responsible for different functions. To address the question of whether actin isoforms can substitute for each other functionally in myofibrils and cytoskeletal structures, several approaches were applied that allow the alteration of ratios of various actin isoforms by the introduction of exogenous protein, incorporation of a gene, or transformation of gene expression. When fluorescently labeled skeletal muscle and cytoplasmic actins were coinjected into cultured muscle and nonmuscle cells (McKenna et al., 1985), the injected skeletal muscle and brain actins were not differentiated by either embryonic chicken fibroblasts or cardiac myocytes. Fibroblasts that synthesize only trace amounts, if any, of skeletal muscle 움-actin (Rubenstein and Spudich, 1977) utilized injected skeletal muscle and cytoplasmic actins to the same extent in stress fibers and other actin-containing structures. Similarly, injected muscle and brain actins were localized equally in cardiac myofibrils. Moreover, all of the F-actin-containing structures of these cells were associated with fluorescently labeled actin from either source (McKenna et al., 1985). These data suggested that preexisting actin-containing structures do not discriminate between different isoforms of exogenous

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actin. In agreement with these results, fluorescently labeled muscle and cytoplasmic actins were incorporated into developing skeletal muscle myofibrils (Hayakawa et al., 1996). Incorporation of sarcomeric (cardiac) 움-actin into cytoskeleton was also observed in mouse L-cells transfected by human cardiac actin gene. The cardiac actin was synthesized in the transfected cells and partitioned between the Triton-X100-insoluble and -soluble cytoskeleton fractions to the same extent as the endogenous 웁-actin (Gunning et al., 1984). In neonatal and adult rat cardiomyocytes transfected by muscle-specific actin genes encoding two sarcomeric (움-skeletal muscle and 움-cardiac) and two smooth muscle (움-vascular and 웂-enteric) actins synthesis of these actins did not interfere with cell contractility and did not disturb the localization of endogenous sarcomeric proteins, indicating that they participate in the formation of sarcomeres (von Arx et al., 1995). In fibroblasts transfected by each of the six vertebrate isoactin genes, all of the actin isoforms associated with the endogenous microfilaments (von Arx et al., 1995), as well as transfected striated and smooth muscle actins were recruited by stress fibers in smooth muscle cells (Mounier et al., 1997). However, in endothelial and epithelial cells, exogenous skeletal muscle and cardiac actins rarely were incorporated into stress fibers, instead remaining scattered within the cytoplasm and frequently forming long crystallike inclusions (Mounier et al., 1997). Cardiac 움-actin synthesized in transfected neurons also formed cylindrical deposits and was present in dendritic spines only rarely (Kaech et al., 1997). Moreover, the quantitative characteristics of the incorporation of muscle and cytoplasmic actins into heterologous microfilament structures seem to be different. Although both 움- and 웁-actin isoforms were incorporated into myofibrils in a cell-free system, 3fold greater incorporation of the 움-isoform was observed (Peng and Fischman, 1991). Fluorescently labeled muscle actin became detectable in striated structures of developing muscle cells sooner than cytoskeletal actin (Hayakawa et al., 1996). In cultured ventricular adult rat cardiomyocytes, upregulation of smooth muscle 움-actin with basic fibroblast growth factor or triiodothyronine interfered with sarcomere assembly and stopped myofibrillar growth, whereas down regulation of smooth muscle 움-actin by insulin-like growth factor promoted myofibrillar growth. These results indicate that the high level of smooth muscle 움-actin synthesis seems to be incompatible with the formation of cross-striated contractile structures (Harden et al., 1998). It was also shown that induction of smooth muscle 움-actin synthesis in fibroblasts specifically led to decreased motility of these cells, whereas resting, nonactivated smooth muscle 움-actin-negative cell strains migrated at a higher speed. The increased motility of fibroblasts was due to a reduction in filamentous smooth muscle 움-actin by electroinjected

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specific antibodies or antisense oligodeoxynucleotides (Ronnov-Jessen and Petersen, 1996). Fluorescence photobleaching recovery following the incorporation of labeled muscle and nonmuscle actins into cultured embryonic chicken cardiac myocytes and fibroblasts showed that the exchangeability of labeled muscle actin in striated myofibrils was faster than that of cytoplasmic actin in the immature nonstriated portion of myofibrils. These data indicate that, although cardiomyocytes were unable to discriminate between the incorporation of muscle and cytoskeletal actins into myofibrils, actin molecules in cardiac myofibrils cannot be readily exchanged by heterotypic cytoplasmic actin (Shimada et al., 1997; Suzuki et al., 1998). Functional nonequivalence of muscle and cytoplasmic isoactins was revealed in experiments with transgenic animals. When the murine cardiac 움-actin gene was disrupted by homologous recombination, the majority of the mice lacking cardiac 움-actin did not survive to birth, and the rest generally died within 2 weeks of birth. Although an increased level of skeletal and vascular smooth muscle 움-actins was observed in the hearts of newborn animals, it was insufficient to maintain the myofibrillar integrity in homozygous mutants. Ectopic expression of the intestinal smooth muscle 웂-actin gene in the cardiac 움-actin-deficient mice resulted in the development of hypodynamic, considerably enlarged and hypertrophied hearts. The transgenically expressed intestinal smooth muscle 웂-actin gene reduced cardiac contractility in wild-type and heterozygous mice (Kumar et al., 1997). The Drosophila melanogaster Act88F gene encodes a specific actin isoform in the indirect flight muscle of the fly. This actin is similar to vertebrate cytoplasmic 웁-actin and differs from vertebrate skeletal muscle actin by 28 substitutions (Schoenenberger et al., 1995). Inactivation of the Act88F gene causes the fly to be unable to fly, which can be reversed when the wildtype Act88F gene or vertebrate cardiac 움-actin gene is introduced into the fly (Karlik et al., 1984; Fyrberg, 1989; Schoenenberger et al., 1995). However, transgenic Drosophilas that are null mutations for endogenous Act88F were unable to fly if they were transfected by the cytoplasmic 웁-actin gene (Schoenenberger et al., 1995). To test the significance of the amino acid replacements that distinguish muscle and nonmuscle Drosophila isoactins, directed mutagenesis was used to introduce these substitutions into the Act88F flight-muscle-specific actin gene. Transformation of files with this construct led to the disruption of flight muscle structure and function (Fyrberg et al., 1998), indicating that cytoplasmic actin cannot substitute the muscle isoform in the muscle structure and contraction. Similarly, in fusioncompetent myogenic cells derived from the quail fibrosarcoma, which do not synthesize skeletal muscle 움-actin, no sarcomeres are formed during

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differentiation, although nonmuscle actin genes were expressed (Antin and Ordahl, 1991). Tranfection of neonatal and adult rat cardiomyocytes with two cytoplasmic actin genes also resulted in the rapid cessation of cell contraction and dramatically changed cell morphology, including the outgrowth of filopodia and cell flattening. The newly synthesized cytoplasmic actins were localized diffusely in nonstriated filamentous structures and submembrane sites, whereas myofibrillar staining normally revealed by phalloidin disappeared, indicating that sarcomeres were depleted of F-actin (von Arx et al., 1995). Similar alterations of cardiomyocyte morphology and function were observed in cardiomyocytes transfected with a construct encoding the chimeric protein containing the first 83 amino acid residues of cardiac 움-actin and the C-terminal portion of cytoplasmic 웂-actin. The protein containing the N-terminal portion of cytoplasmic 웂-actin and residues 84– 375 of cardiac 움-actin was distributed in a manner similar to that of muscle actin (von Arx et al., 1995). Isoform-specific distribution of these chimeric actins corresponding to their C-terminal sequences also was observed in dendritic spines (Kaech et al., 1997). The distinct roles of the two cytoplasmic actin isoforms were revealed in experiments with overexpression of cytoplasmic 웁- and 웂-actin genes in mouse C2 myoblasts (Schevzov et al., 1992). Elevated synthesis of 웁-actin promoted cell spreading. In contrast, C2 clones with a high level of 웂-actin showed a strong decrease in cell surface area. In addition, in the 웂-actin transfectants, a diffuse organization of cytoskeleton rather than well-defined stress fibers was observed (Schevzov et al., 1992). The specificity of the effects was supported by the fact that high-level expression of a mutant 웁actin gene, giving rise to accumulation of actin with lower polymerizability, produced a decrease in the cell surface area and disruption of the actin microfilament network similar to that seen with transfection of the 웂-actin gene (Schevzov et al., 1992). An increase in the 웁- to 웂-actin ratio due to expression of the two different mutant 웁-actin genes in normal human fibroblasts and the cell line Hut12 produced a dramatic effect on cell morphology (Leavitt et al., 1987). However, at variance with these results is that transfection by the 웁- or 웂-actin genes of cardiomyocytes (von Arx et al., 1995), smooth muscle cells, and endothelial and epithelial PtK2 cells (Mounier et al., 1997) revealed no differential effect of the two incorporated cytoplasmic actin isoforms. Thus, the experimental evidence of the inability of different actin isoforms to substitute for each other is not quite consistent, a discrepancy that can probably be accounted for by the cell type, stage of cell differentiation, and extent to which the incorporated actin isoform is in excess over the endogenous ones. In this context, it is noteworthy that in most studies performed so far the exogenous actin is synthesized on a background of

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endogenous actin and, as a rule, represents only a fraction of total actin. In addition, epitope tagging used as a method to monitor the fate of the exogenous protein can interfere with actin function (Brault et al., 1999). In spite of these limitations, the main body of the results indicates that the muscle actin isoforms are not able to replace cytoplasmic actin in stress fibers and other cytoskeleton structures if the muscle actin genes normally are not expressed in these cells. On the other hand, cytoplasmic actin isoforms cannot either replace muscle actins in striated myofibrils or maintain muscle contraction. The high-level synthesis of exogenous actins leads to alterations in cell organization and morphology. These data indicate that the highly conserved actins functionally are specialized for the tissues in which they predominate. 7. Cellular Mechanisms of Actin Isoform Sorting The results of biochemical, immunocytochemical, and molecular biology experiments reviewed previously demonstrate that appearance, amount, and localization of actin isoforms are strongly controlled by cell machinery. Although at the early stages of cell differentiation the expression of any actin gene is possible, under normal physiological conditions, while differentiation proceeds the synthesis of specific actin isoforms is regulated temporally and the proteins produced are segregated spatially. Pathological situations of tissue injury or mammalian disease correlate either with up- and downregulation of distinct actin genes returning to a fetal gene program or with a failure to sort actin isoforms. On the other hand, changes in expression of specific actin genes are accompanied by alterations in cell structure and function, suggesting that specific actin isoforms perform unique cellular functions. However, the physiological significance and mechanisms of isoform sorting are poorly understood. In other words, so far it is not clear why different actin isoforms do not substitute for each other and what mechanisms regulate their segregation in the cell. To explain the sorting of actin isoforms, its correlation with the sorting of actin mRNAs is considered (Herman, 1993; Gunning et al., 1998). Developmental changes in the actin isoform patterns qualitatively are similar at both the mRNA and protein levels. Actin isoforms and their corresponding mRNAs often are colocalized in cell compartments, as is documented for the distribution of 웁-actin and its mRNA in the motile leading edge of fibroblasts (Lawrence and Singer, 1986; Sundell and Singer, 1991) and microvascular pericytes (Hoock et al., 1991; Herman, 1993), as well as in dendrites of developing and mature neurons (Bassell et al., 1998; Hannan et al., 1998). It is possible, therefore, that the sorting of actin isoform mRNAs prior to translation contributes to protein segregation. This suggestion is consistent with two lines of evi-

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dence. The peripheral location of 웁-actin mRNA within cultured myoblasts and fibroblasts is maintained by serum, and relocalization of diffusely distributed preexisting 웁-actin mRNA into the leading lammellae is regulated by growth-factor-induced signal transduction mechanisms within minutes (Hill et al., 1994; Latham et al., 1994). The distribution of 웁-actin mRNA correlates with the magnitude and direction of cell movement (Kislauskis et al., 1997). It is also shown that isoform-specific 3⬘-untranslated sequences of actin mRNAs play a key role in directed localization of specific mRNAs in distinct cell compartments (Kislauskis et al., 1993). Restraint of 웁-actin mRNA localization with a specific antisense oligodeoxynucleotide resulted in the loss of cell polarity and reduced cell motility (Kislauskis et al., 1994, 1997). These observations indicate that sequence-specific targeting of actin mRNAs to distinct cellular regions followed by synthesis of their corresponding proteins may be a way of actin isoform sorting. Studies addressing the question of actin mRNA targeting mechanisms showed that exposure of spread chicken embryo fibroblasts to cytochalasin D or plating of these cells in the presence of cytochalasin D disturbed the peripheral localization of actin mRNA (Sundell and Singer, 1991). These results imply that transport and subsequent anchorage of mRNA in the cell periphery is microfilament-dependent. It suggests that actin-bound proteins of microfilaments may participate in mRNA sorting. Alternatively, the required interaction between microfilaments and mRNA can occur via RNA-containing structures. It is interesting that prosomes, mRNA-associated ribonucleoprotein-containing particles, form a pseudo-sarcomeric structure concomitantly with or even prior to the integration of sarcomeric actin into a future sarcomeric structure (De Conto et al., 1998). On the other hand, actin isoforms frequently are distributed over a wider area than their respective mRNAs. In myoblasts, 웂-actin mRNA is localized in perinuclear and nearby cytoplasm, whereas the protein was found in stress fibers and at the cell plasma membrane and did not correspond to its mRNA distribution (Hill and Gunning, 1993). Still more strikingly, in developing neurons, the majority of 웁-actin is concentrated in the growth cone, whereas only a small portion of the corresponding mRNA is located there (Hannan et al., 1998). It is plausible therefore, that isoform-specific patterns of mRNA targeting determine the sites of synthesis and assembly rather than localization of actin isoforms in the cell (Hannan et al., 1998; Gunning et al., 1998). The other factor that can influence the specific distribution of actin isoforms is the different localization of actin-binding proteins. The appearance of muscle-specific actin isoforms during myogenesis is coordinated with changes in isoform patterns of myosin, tropomyosin, troponin, and other contractile proteins (Caplan et al., 1983; Bandman, 1992). In nonmuscle cells, spatial and temporal segregation of isoactins is accompanied by

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the segregation of isoforms of tropomyosin and myosin (Gunning et al., 1998), as well as by specific locations for actin-binding proteins ezrin (Shuster and Herman, 1995; Yao et al., 1995) and fimbrin (Hofer et al., 1997). The detailed comparison of intracellular distributions of actin and tropomyosin isoforms in neurons indicated that each neuronal compartment has a complex combination of isoforms in both early-differentiating and adult neurons (Gunning et al., 1998). Thereby localization of some tropomyosin isoforms coincides with that of 웁- or 웂-actin, suggesting that tropomyosin isoforms may control the level of actin assembly and, hence, actin targeting (Hannan et al., 1998). In chicken auditory hair cells, 웁-actin filaments are cross-linked selectively by fimbrin (Hofer et al., 1997)—a protein from the plastin–fimbrin family whose members are shown to differ in their interactions with actin isoforms (Prassler et al., 1997). In rabbit gastric parietal cells, not only is 웁-actin colocalized with ezrin but, in addition, a possible preference of ezrin for 웁-actin over 웂-actin has been documented (Yao et al., 1995, 1996). Colocalization of 웁-actin with ezrin was also revealed in the motile cytoplasm of cultured bovine vascular cells (Shuster and Herman, 1995). Purified pericyte ezrin was shown to be bound to 웁but not 움-actin; this interaction was indirect and most likely mediated through another accessory protein(s) (Shuster and Herman, 1995). Thus, regardless of the mechanism of actin isoform sorting, it finally is associated with actin filament structures. Specific anchorage of mRNA as well as the specific interaction of actin isoforms with actin-binding proteins indicates that these components recognize the actin isoforms’ specificity. In other words, it implies that the properties of actin isoforms themselves contribute to their specific localization and function. In summary, results of the in vivo experiments show that the distribution of actin isoforms in vertebrates is tissue-specific rather than species-specific. On the other hand, the unique actin isoforms can be found in invertebrates, particularly in lower eukaryotes. The synthesis of specific actin isoforms is accompanied by their subcellular compartmentalization, with both processes being regulated by factors of cell proliferation and differentiation. The mechanisms of actin isoform sorting seem to involve targeting of mRNAs that contain isoform-specific 3⬘-untranslated sequences as well as the specific interaction of actin isoforms with actin-binding proteins. It is plausible, however, that the specificity of the actin isoforms themselves contributes to the specificity of this recognition. B. Functional Specificity in Vitro 1. Actin Polymerization Both in vivo and in vitro, actin can exist as a monomer (globular, or Gactin) and as a polymer (fibrillar, or F-actin), with the G 씮 F transformation

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being a reversible process. However, the functional form of actin in cells appears to be the polymer. Therefore, the ability of actin to form stable polymers that, in turn, can be easily depolymerized may be regarded as the main functional property of actin. The main steps and mechanisms of actin polymerization in vitro are wellestablished. Globular actin spontaneously polymerizes into filaments at physiological concentrations of neutral salts, with the optimum being 50– 150 mM KCl and/or 1–2 mM MgCl2. The presence of Mg2⫹ accelerates polymerization induced by monovalent cations, whereas millimolar concentrations of Ca2⫹ inhibit the reaction (Selden et al., 1983, 1986). Actin polymerization can also be induced at low salt concentrations by polycations (Oriol-Audit, 1978; Gawlitta et al., 1981; Grant and Oriol-Audit, 1982) and by myosin and its proteolytic fragments (Miller et al., 1988). In the cell, spontaneous actin polymerization is inhibited by the monomer-sequestering proteins, profilin and thymosin (Sun et al., 1995). The polymerizing conditions induce conformational changes in actin monomers that promote their association and, therefore, are regarded as the monomer activation (Rich and Estes, 1976; Rouayrenc and Travers, 1981; Pardee and Spudich, 1982; Khaitlina et al., 1996; Strzelecka-Golaszewska, 2000). However, filament assembly can occur only above a threshold (critical) actin concentration, which varies with polymerizing conditions. The monomers associate rather slowly to form short oligomers (nuclei), which consist of at least three monomers. In the time course of actin polymerization, the nucleation step usually is observed as a lag phase (Carlier, 1989, 1991, 1998; Estes et al., 1992; Sheterline et al., 1995; Steinmetz et al., 1997a,b). Once the nuclei are formed, they are elongated rapidly until the monomer concentration falls to the critical level and the system reaches a steady state. The two ends of the polymer are not equivalent, with the polymerization rate at one end (the barbed, fast, or ⫹ end) being much higher than that at the other (the pointed, slow, or ⫺ end) (Pollard and Mooseker, 1981; Pollard, 1986). Thereby the addition of actin monomers to the barbed end is accompanied by dissociation of monomers from the pointed end, resulting in a flux of monomers along the filament. This process is called head-to-tail assembly or treadmilling (Wegner, 1976, 1982; Wegner and Isenberg, 1983). The actin polymerization does not involve a length-limiting step; therefore, the length of an actin filament is limited only by its mechanical stability. Actin polymerization is accompanied by stoichiometric hydrolysis of the tightly bound ATP, which occurs not during the assembly step but subsequent to incorporation of the monomer into the filament to yield an ADPcontaining filament. An intermediate in this process is a subunit carrying ADP-Pi. As ATP hydrolysis lags behind assembly, the hydrolysis step thus is uncoupled from the assembly step (Korn et al., 1987). Under conditions

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where the rate of assembly exceeds the rate of hydrolysis and phosphate release, an individual filament will contain segments carrying ATP and ADP-Pi. The affinities of the ATP- and ADP-containing filament ends for monomeric actin are different; therefore, the ATP-containing cap at the filament end stabilizes the filament against disassembly. When the concentration of free monomers and, hence, the rate of assembly decreases below the rate of ATP hydrolysis, the ATP cap can disappear (Carlier, 1989, 1991, 1998). Thus, the ability of actin to polymerize can be characterized by two parameters: the critical concentration for polymerization and the rate of reaction. The latter is different at different stages of polymerization, as well as at the different ends of the polymer. Another relevant parameter is filament length, which depends on the mechanical properties of the filaments. These parameters seem to correlate with the thermodynamic stability of actin filaments and characterize the monomer–monomer interactions in the filament. Furthermore, the rate of ATP hydrolysis is significant for characterizing the association–dissociation of monomers at the filament ends. 2. Lower Polymerizability of ␤- and ␥-Actins Relative to Skeletal Muscle ␣-Actin Comparison of actins isolated from developing and adult rabbit skeletal muscles provided the first evidence for lower polymerizability of actin from the low-differentiated muscles relative to mature skeletal muscle actin (Khaitlina and Pinaev, 1976). In this early work, the isoform composition of the actin preparations was not examined. However, the transition of actin isoforms during myogenesis (Wahlen et al., 1976; Storti et al., 1976) suggested that the lower polymerizability of neonatal actin is characteristic of 웁- and 웂-actin isoforms, whose level is high at this stage of muscle differentiation. This suggestion was confirmed in detailed studies on the purified cytoplasmic and muscle actin isoforms. a. Critical Concentration for Polymerization The polymerization properties of nonmuscle cytoplasmic 웁- and 웂-isoforms were shown to be qualitatively similar but quantitatively different from those of skeletal muscle 움actin (Gordon et al., 1976, 1977; Koffer and Dickens, 1987). The critical concentrations of actins from human platelets, embryonic chicken brain, and rat liver were compared with the critical concentration of rabbit skeletal muscle actin under two sets of ionic conditions, 2 mM MgCl2 and 0.1 M KCl, and at two temperatures, 25 and 5⬚C (Gordon et al., 1976). In 2 mM MgCl2 at 25⬚C, the critical concentrations were similar for all of the actin

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species (Table V). At 5⬚C, the nonmuscle actins showed a small, but reproducible decrease in the slope of the plot of specific viscosity versus actin concentration, although no measurable difference in critical concentration was detected. However, in 0.1 M KCl at 25⬚C, the critical concentrations for nonmuscle actins were 2–3-fold higher than those for skeletal muscle actin, and in 0.1 M KCl at 5⬚C, a 5-fold difference between critical concentrations for polymerization of 웁/웂- and 움-actins was observed (Table V). The stronger dependence of critical concentrations on temperature and the higher values of the critical concentration at low temperature than those for skeletal muscle 움-actin were also reported for the ␦-actin isoform from Acanthamoeba castellanii (Gordon et al., 1976), actin from Dictiostellium discoideum (Uyemura et al., 1978), intestinal smooth muscle 웂-actin from chicken gizzard (Prochniewicz and Yanagida, 1981), vascular smooth muscle 움-actin from bovine aorta (Strzelecka-Golaszewska et al., 1985b), and scallop adductor muscle 웁-like actin (Khaitlina, 1986) (Table V). Figure 1 demonstrates the temperature- and salt-dependent changes of critical concentrations for the polymerization of scallop adductor muscle 웁-like actin and rabbit skeletal muscle 움-actin (Khaitlina, 1986). Whereas no difference between the two actin species was detected in 0.1 M KCl at 20⬚C, the critical concentration of scallop actin became much higher than that of skeletal muscle actin upon a decrease in the temperature and KCl concentration (Fig. 1A). The critical concentration for polymerization is connected with the binding constant for addition of the monomer to the polymer ends. Therefore, determination of critical concentrations at various temperatures makes possible calculation of the thermodynamic constants, the free energy, enthalpy, and entropy, for polymerization. These calculations showed that a more pronounced increase in the extent of polymerization of 웁- and 웂actins with temperature could be attributed to higher positive changes in enthalpy and entropy for polymerization of these actins compared to skeletal muscle 움-actin (Gordon et al., 1976, 1977; Prochniewicz and Yanagida, 1981). These results indicate quantitative differences in intermonomer interactions involved in the polymerization of different actin species, as well as a possible difference in cooperativity between the monomers of the corresponding polymers (Prochniewicz and Yanagida, 1981) or in the polymer stabilities (Khaitlina, 1986). It is important that no difference in critical concentrations of skeletal muscle and cardiac 움-actins was found (Strzelecka-Golaszewska et al., 1985b). Thus, under certain conditions, cytoplasmic 웁- and 웂-actins, as well as smooth muscle 움- and 웂-actins, polymerize less readily than sarcomeric 움actin isoforms. This difference seems to reflect the specificity of intermonomer contacts within the corresponding filaments.

0.7 2.1 1.7 1.9 2.1 1.43 11.4 0.7

0.7 0.7 0.7 0.5 0.5

움-Skeletal 웁,웂-Cytoplasmic

␦ 움-Skeletal 웁,웂 움-Skeletal 웂-Smooth 움-Smooth 움-Cardiac 움-Skeletal 웁-like

Rabbit skeletal muscle Human platelets Embryonic chicken brain Rat liver Acanthamoeba Rabbit skeletal muscle Saccharomyces cerevisiae Rabbit skeletal muscle

Chicken gizzard Bovine aorta Bovine heart Rabbit skeletal muscle Scallop adductor muscle

35.7b 21.4b 4.8b 2.4 7.2

4.8

2.4 12.1 11.9 11.4 10.7

5⬚C

15.1d 14.7d 5.9d 18 18

135c about 15d 5.9d

7.1 9.6c 9.6c 9.6c 21.7c 14 about 7d

b

a

c

Activation of myosin Mg2⫹ATPase, Kapp(웂M )

Every set of data includes properties of rabbit skeletal muscle 움-actin as a reference. The values were determined at 0⬚C. c Activation of skeletal muscle heavy meromyosin Mg2⫹-ATPase was measured. d Activation of skeletal muscle myosin or myosin subfragment 1 Mg2⫹-ATPase was measured.

25⬚C

Isoform

Critical concentration for polymerization with KCl (애M ) Actin

TABLE V Properties of Different Actin Isoformsa

Khaitlina, 1987; Hue et al., 1988

Prochniewicz and Yanagida, 1981; Mossakowska and StrzeleckaGolaszewska, 1985

Gordon et al., 1976 Greer and Schekman, 1982; Miller et al., 1996

Gordon et al., 1977

References

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FIG. 1 Critical concentrations for polymerization of scallop adductor muscle and rabbit skeletal muscle actins under different conditions. (A) Rabbit (䊊) and scallop (䊉) intact Ca-actins, from left to right; 0.1 M KCl, 20⬚C; 0.1 M KCl, 4⬚C; and 0.02 M KCl, 4⬚C. (B) Rabbit (䊉, 䊊) and scallop (䊏, ⵧ) ECP-cleaved Mg-actins at 0.1 M KCl, 20⬚C. Open and filled symbols show results from two independent experiments.

b. Rate of Polymerization To illustrate the kinetics of polymerization of different actin isoforms, Fig. 2 compares the time courses of polymerization of scallop adductor muscle 웁-like actin and rabbit skeletal muscle 움-actin in Ca and Mg forms (Khaitlina et al., 1999). In the presence of 0.1 M KCl, the steady-state extent of polymerization of both actin species was the same. However, the rate of polymerization of scallop Ca-actin was lower

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FIG. 2 Time course of polymerization of scallop adductor muscle and rabbit skeletal muscle Ca- and Mg-actins. Rabbit skeletal muscle (12 애M ) or scallop Ca-actins (curves 1 and 2) or Mg-actins (curves 3 and 4) were polymerized by the addition of 0.1 M KCl. Light-scattering intensities were recorded at 334 nm. Curves 1 and 3, scallop adductor muscle actin; curves 2 and 4, rabbit skeletal muscle actin. Reproduced with permission from Khaitlina et al. (1999).

than that of skeletal muscle actin (Fig. 2). The difference in the rate of polymerization was abolished upon replacement of tightly bound Ca2⫹ by Mg2⫹ (Fig. 2), indicating that it was due neither to different inactivations of scallop and rabbit actins nor to the presence of contaminants in these preparations. A striking difference in the polymerization properties of yeast and skeletal muscle actins was observed (Greer and Schekman, 1982b; Kim et al., 1996). In the presence of 0.1 M KCl, no polymerization of 25 애M yeast Ca-actin was detected even after 18 hr of incubation at 4⬚C. At the same time, Mg-induced polymerization of yeast Ca-actins was faster than that of skeletal muscle actin due to a higher rate of replacement of tightly bound Ca2⫹ with Mg2⫹ in yeast actin (Kim et al., 1996) and/or more intensive fragmentation of yeast actin filaments, which increases the number of filament ends serving as nuclei for filament elongation and, thus, promotes the nucleation step of yeast actin polymerization (Buzan and Frieden, 1996). The rates of polymerization of Acanthamoeba (Gordon et al., 1976) and BHK (Koffer and Dickens, 1987) Ca-actins with KCl at 25⬚C also were shown to be lower than that of skeletal muscle actin. At 5⬚C, the difference

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between the rates of polymerization of Acanthamoeba and skeletal muscle actins was observed not only in 0.1 M KCl but also in 5 mM MgCl2 (Gordon et al., 1976). On the other hand, the difference was partially abolished in the presence of skeletal muscle F-actin added as the seeds, suggesting that the rate of nucleation of different actins can be different (Gordon et al., 1976; Koffer and Dickens, 1987). The presence of seeds allows one to circumvent the nucleation step of polymerization and, thus, to evaluate the elongation rates. These experiments showed that in KCl elongation of BHK actin was slower than that of skeletal muscle actin (Koffer and Dickens, 1987). According to preliminary data, the rate of the elongation of scallop Ca-actin in KCl is slower than that of skeletal muscle actin in the presence of both F-actin and gelsolin used as the seeds (Khaitlina and Hinssen, 1999). Thus, under conditions that are not optimal for actin polymerization (low temperature, low salt concentration, absence of Mg2⫹ ), assembly of skeletal muscle actin is faster than that of other actins studied, and this difference can be due to the higher nucleation and elongation rates. At the same time, the rate of depolymerization of skeletal muscle actin can be slower than that of other actins, as was shown for skeletal muscle and scallop actins depolymerized in the presence of DNaseI (Khaitlina, 1986). These data again suggest that different actins can form polymers of different stability. c. Stability of Polymers Formed by Different Actin Isoforms The stability of actin polymer is shown to depend dramatically on the integrity of the polypeptide chain within the DNase-binding loop at the top of the monomer (Khaitlina et al., 1988, 1991, 1993; Schwyter et al., 1989). Actin cleaved between Gly42 and Val43 within the DNase loop by the protease ECP 32 from Escherichia coli A2 strain retains the principal properties of intact actin, but loses its ability to polymerize and interact with DNaseI (Khaitlina et al., 1988, 1991). Polymerizability of the proteolytically modified actin was partially restored when the tightly bound Ca2⫹ was replaced by Mg2⫹ and became similar to that of intact actin in the presence of phalloidin stabilizing the filament (Khaitlina et al., 1993). Experiments (Khaitlina et al., 1999) have shown that cleavage of scallop actin with the E. coli protease impaired the polymerization of scallop Mg-actin more intensively than that of skeletal muscle actin (Fig. 3). The critical concentration for polymerization of ECP-cleaved scallop Mg-actin was 6.8 애M, compared to 2.8 애M for ECP-cleaved skeletal muscle actin (Fig. 1B). These values are similar to the critical concentrations for polymerization of scallop and skeletal muscle actins in 0.1 M KCl at 4⬚C (Fig. 1A, Table VI), suggesting that the destabilizing effects of low temperature and damage to the DNase loop might be of a similar nature.

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FIG. 3 Time course of polymerization of scallop adductor muscle and rabbit skeletal muscle Mg-actins upon cleavage with the E. coli protease ECP32. Rabbit skeletal muscle (24 애M ) or scallop ECP-cleaved Mg-actins were polymerized with 0.1 M KCl. Light-scattering intensities were recorded at 334 nm. Curves 1 and 3, scallop adductor muscle actin; curves 2 and 4, rabbit skeletal muscle actin. Reproduced with permission from Khaitlina et al. (1999).

Other evidence for different stabilities of polymers formed by actin isoforms was obtained in rheological experiments (Allen et al., 1996). The viscoelastic properties of a polymer solution strongly depend on filament flexibility and length. When rheological parameters of skeletal muscle 움actin, smooth muscle 웁- and 웂-actins, and erythrocyte 웁-actin were compared, the skeletal muscle actin gel was the most elastic, the smooth muscle 웁- and 웂-actin gels were less elastic, and cytoplasmic 웁-actin did not form an elastic gel under the experimental conditions used. The lower elasticity of smooth muscle actin isoforms relative to skeletal muscle actin correlated with the lower average lengths of these actins polymerized in the presence of labeled phalloidin (Allen et al., 1996). The length of erythrocyte 웁-actin filaments was similar to that of skeletal muscle 움-actin, indicating that these filaments might be more flexible than the other filaments studied (Allen et al., 1996). Lower stability of yeast actin filaments was observed during the in vitro motility measurements. Compared to the behavior of skeletal muscle actin, much stronger fragmentation of yeast actin filaments took place (Kim et

ECP-cleaved

Intact

Actin 100 mM KCl, 20⬚C 100 mM KCl, 4⬚C 20 mM KCl, 4⬚C 100 mM KCl, 25⬚C 50 mM KCl, 20⬚C

Conditions

Sc (웁) 0.5 애M 7.2 애M 18 애M ⬎76 애M

Rb (움) 0.5 애M 2.4 애M 6 애M

Ca-actin

TABLE VI Critical Concentrations for Polymerization of Scallop and Skeletal Muscle Actins under Different Conditions

2.7 애M 2.8 애M

R (움)

Mg-actin

6.8 애M

Sc (웁)

Khaitlina et al., 1993 Khaitlina et al., 1999

Khaitlina, 1986

Reference

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al., 1996). The more extensive fragmentation of yeast actin filaments also was suggested to explain the faster nucleation of yeast Mg-actin polymerization (Buzan and Frieden, 1996; Kim et al., 1996). In summary, the results of the in vitro studies show that actins isolated from different contractile systems differ in their polymerizability, 웁- and 웂-isoactins differ from skeletal muscle 움-actin by a lower rate of polymerization if they contain Ca2⫹ as a tightly bound cation, by a higher critical concentration for polymerization at low temperature and/or low concentration of polymerizing salt or upon cleavage of the actin polypeptide chain within subdomain 2, and by faster depolymerization in the presence of DNaseI. These data point to lower stability of the polymers formed by 웁and 웂-isoactins compared with those of sarcomeric 움-actins. The lower stability of yeast F-actin manifests itself in strong filament fragmentation. These differences might be explained by the difference in structure of actin isoforms, which affects the efficiency of the actin–actin contacts in the polymer.

C. Structural Specificity 1. Three-Dimensional Structure of Actins The three-dimensional structure of the actin molecule was resolved for skeletal muscle actin complexed with DNaseI (Kabsch et al., 1990) or gelsolin S1 (McLaughlin et al., 1993) and for nonmuscle 웁-actin complexed with profilin (Schutt et al., 1993). The overall structures are similar though not identical. The actin molecule consists of one polypeptide chain of 42 kDa folded in such a fashion that it fits into a square of side length 5.5 nm; the flat side is of width 3.5 nm (Fig. 4). The molecule is clearly divided into two domains of roughly equivalent size (which, for historical reasons, are called the large and small domains) with a cleft containing the bound nucleotide (usually ATP) and cation (Ca2⫹ or Mg2⫹ ). These two domains are connected covalently with only two strands of the polypeptide chain that are close together at the base of the molecule, allowing relative movement of the domains. Therefore, this region is regarded as a hinge (Lorenz et al., 1993; Tirion and ben Abraham, 1993) or a shear (Page et al., 1998) region of the molecule. Each domain is divided into two subdomains, so that the whole monomer consists of four subdomains. By definition, subdomains 1 (residues 1–32, 70–144, and 338–375) and 2 (residues 33–69) constitute the small domain, whereas subdomains 3 (residues 145–180 and 270–337) and 4 (residues 181–269) represent the large domain of actin (Kabsch et al., 1990). The bound nucleotide contacts residues from all four subdomains and functions as the coordinating center of the molecule. The similarity of

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FIG. 4 Schematic presentation of the three-dimensional structure of actin monomer (Kabsch et al., 1990). The gray balls map the monomer–monomer contact sites along the long-pitch helix of actin polymer predicted by the atomic model of F-actin (Holmes et al., 1990; Lorenz et al., 1993). For better resolution of these sites as well as the sites involved in interstrand contacts, see Fig. 7.

this structure to known structures of other multidomain proteins (Flaberty et al., 1991; Bork et al., 1992) allows the prediction that actin can undergo conformational changes during polymerization or related processes (Lorenz et al., 1993; Tirion et al., 1995; Chik et al., 1996; Page et al., 1998). Indeed, in the models of actin filament based on the structure of skeletal muscle G-actin derived from the actin–DNase complex, a small shift between the large and small domains and a few mobile surface loops were suggested to optimize the fit of the structure to the X-ray fiber diffraction data (Holmes et al., 1990; Lorenz et al., 1993; Tirion et al., 1995). The resultant changes in the 움-actin monomer are assumed to reflect conformational changes associated with polymerization. As compared to G-actin, in monomers of the actin filament subdomain 2 comes to a lower radius and subdomain 4 undergoes shifts, thus causing the cleft between these domains to narrow (Lorenz et al., 1995; Tirion et al., 1995). In terms of these models, actin polymerization also is accompanied by reorientation of the DNasebinding loop and shift of the C-terminus. Together with loop 262–274, these two regions may participate in the intermonomer hydrophobic interactions that stabilize the filament (Tirion et al., 1995).

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Much greater conformational transitions have been revealed by structural analysis of the 웁-actin–profilin crystals obtained under different conditions (Chik et al., 1996; Page et al., 1998). A most striking transition is rotation of the large domain relative to the small domain by nearly 10⬚, which results in sufficient opening of the nucleotide-containing cleft. The nucleotide in the open state is 25% more exposed to solvent than in the closed (tight) one (Chik et al., 1996). The closure of the cleft principally is due to 14.7⬚ rotation of subdomain 2. The N-terminus (residues 2–7) is very different between the two states, adopting a helical structure in the tight state and a turn in the open state. In addition, local conformational changes in the hydrophobic pocket surrounding Cys374 as well as within the DNase loop were observed (Chik et al., 1996). Transition from the tight to the open state seems to involve several interdomain (hinge and shear) regions and the loops of subdomains 2 and 4, whereas the secondary and tertiary structure of the rigid domain cores is largely maintained (Page et al., 1998). Thereby, subdomains 3 and 4 rotate as a single rigid body, whereas subdomains 1 and 2 rotate independently and separately from subdomains 3 and 4. Similar conformational changes are expected to be associated with actin polymerization (Page et al., 1998). It is not clear to what extent the difference between these structures is due to inherent properties of actins and/or due to the presence of either DNaseI or profilin in the crystals, which are bound at the top and bottom of the monomer, respectively, and can affect actin conformation differently. It would be interesting, therefore, to extend the analysis to the solved structure of the yeast actin–gelsolin segment 1 complex (Vorobiev et al., 1998) and compare the actin’s open and tight states in the available actin– gelsolin segment 1 complexes. On the other hand, comparison of the four determined crystal structures showed that three of them—움-actin–DNaseI (Kabsch et al., 1990), 움-actin–gelsolin segment 1 (McLaughlin et al., 1993), and 웁-actin–profilin structure (Schutt et al., 1993)—are similar in conformation and seem to correspond to the tight state. This may imply that during modeling of the actin filament, when the structure of these tight-state monomers was used as a staring point, not all but only additional conformational changes were revealed, which are on a smaller scale. At the same time, all of the structural data clearly show that the multidomain structure of the actin monomer provides a good basis for the regulation of actin functions via large-scale and local conformational changes. 2. Structure of ␤-Actin in the Actin–Profilin Complex Although so far the large conformational transitions from the open to the tight state are found only in 웁-actin, they cannot be regarded as an inherent property of this actin isoform because the three-dimensional structure of

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bovine cytoplasmic 웁-actin generally is similar to the structure of skeletal muscle 움-actin (Schutt et al., 1993). However, several differences between the two actin species are significant (Schutt et al., 1993). The main one is a 5⬚ rotation of the small domain of the tight-state 웁-actin relative to the position of this domain in skeletal muscle 움-actin. This rotation does not affect the conformation of the nucleotide-containing cleft, suggesting that 움-actin complexed with DNaseI is also in the tight state (Schutt et al., 1993). Differences in the amino acid conformations between the two actin isoforms fall into four categories (Schutt et al., 1993) (Table VII). The first and second groups include residues whose main-chain atoms can be superimposed and whose resulting side-chain conformations are very similar, with only subtle differences in interaction with neighboring residues. Residues of the first group are buried away from solvent, whereas residues of the second group are partially or fully exposed to solvent. The third group consists of residues with significant changes in side-chain conformations. Residues of this group may project freely into solvent without interaction with other residues, or they accommodate different intermolecular contacts in the two crystals. The fourth group includes residues whose main-chain conformations are not superimposable, leading to significantly changed side-chain orientations. Residues that bind ATP belong to the first and second groups. As a result, the calcium–nucleotide complex adopts an almost identical conformation in the two isoforms. Structural differences between the two actins involve the N-terminal region of actin: stretch 1–6 containing five amino acid substitutions is helical in 움-actin. These substitutions also induce some changes in the interactions among the neighboring residues. In addition, differences in conformations

TABLE VII Amino Acid Replacements in Cytoskeletal 웁-Actin versus Skeletal Muscle 움-Actin Classified According to Their Effect on Actin Conformation (Schutt et al., 1993) Effecta

Residues

1 group

Subtle effect on side-chain conformations

2 group

Subtle effect on side-chain conformations

3 group

Sufficient changes in sidechain conformations Changes in main-chain conformations

Cys/Val10, Val/Cys17, Thr/Val103, Val/Thr129, Leu/Mat153, Asn/Thr163, Thr/Ala260, Asp/ Ala297, Met/Leu299 Leu/Met16, Ile/Val76, Met/Leu176, Asn/Thr201, Ile/leu267, Ala/Cys272, Tyr/Phe279, Ile/ Val287, Thr/Ser258, Ala/Ser365 Asn/Glu225

4 group a

Glu/Asp2, Glu/Asp4, Thr/Ile5, Thr/Ala6

See text for explanations of the effects.

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of the DNase-binding loop and the C-terminal segment are observed (Schutt et al., 1993). The residues within the DNase-binding loop of 웁-actin show significant differences in orientation, which may be explained by their different environment in crystals that do not contain DNaseI. In both actins, the DNase loop is a region of high mobility (Kabsch et al., 1990; McLaughlin et al., 1993; Schutt et al., 1993; Tirion et al., 1995). Another region of high mobility, the C-terminus, was removed from 움-actin before crystallization (Kabsch et al., 1990). Thus, as can be expected from the conservation of the amino acid sequences, the three-dimensional structures of skeletal muscle 움-actin and cytoplasmic 웁-actin are very similar. However, they differ in the overall position of the small domain relative to the large domain, as well as in local conformations at the N-terminus and, probably, at the C-terminus and within the DNase-binding loop. These regions are the regions of high mobility in all actins, which suggests their adaptive role in both general actin functions and specific functions of actin isoforms. 3. Conformational Mobility of G-Actin Experimental evidence for the conformational mobility of the actin molecule was obtained in spectroscopic and biochemical experiments, which showed that the conformation of G-actin in solution can be modulated at several sites by the type of divalent cation and nucleotide as well as by cation binding at the multiple moderate-affinity sites (Frieden et al., 1980; Carlier et al., 1986; Strzelecka-Golaszewska et al., 1993, 1996; Crosbie et al., 1994; Muhlrad et al., 1994; Kim et al., 1995; Khaitlina et al., 1996; Moraczewska et al., 1996, 1999; Strzelecka-Golaszewska, 2000). The nucleotide bound in G-actin usually is ATP, whereas actin subunits in the polymer contain ADP. The cation bound in isolated G-actin usually is Ca2⫹. However, in vivo actin most likely contains Mg2⫹ as a tightly bound cation. Limited proteolysis and fluorescent probes conjugated to Gln41 within the DNase-binding loop or at the penultimate Cys374 showed that exchange of tightly bound Ca2⫹ for Mg2⫹ results in a decrease in susceptibility to proteolysis at Lys62 and Lys68 within the interdomain nucleotidecontaining cleft as well as at Lys373 in the C-terminus of the monomer (Strzelecka-Golaszewska et al., 1993). In contrast, replacement of tightly bound ATP for ADP enhances exposure of the C-terminal amino acids and, in addition, diminishes susceptibility of the DNase-binding loop (Strzelecka-Golaszewska et al., 1993; Kim et al., 1995; Moraczewska et al., 1996, 1999). Similar intramolecular effects are observed in the subunits of F-actin. Release of phosphate following ATP hydrolysis results in reorientation of subdomain 2 (Orlova and Egelman, 1992; Muhlrad et al., 1994). The subti-

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lisin and trypsin cleavage sites in segments 61–69 and 227–235 and between Lys373 and Cys374 are less accessible in Mg-F-actin than in Ca-F-actin (Strzelecka-Golaszewska et al., 1996). These data suggest that the presence of Mg2⫹ at the high-affinity cation-binding site brings the monomer from an open state to a closed (tight) conformation resembling that of the Factin subunit and, thus, favorable for helical polymer formation. On the other hand, conformational changes induced by ATP–ADP replacement following ATP hydrolysis destabilize the filaments, providing a basis for their disassembly (Egelman and Orlova, 1995; Strzelecka-Golaszewska, 2000). It seems likely that transition from the open to the tight state can also account for decreased proteolytic susceptibility of the monomeric actin in the presence of salt, which is regarded as a manifestation of monomer activation (Rich and Estes, 1976; Rouayrenc and Travers, 1981; Pardee and Spudich, 1982; Khaitlina et al., 1996; Strzelecka-Golaszewska, 2000). Similar to the effect of tightly bound Mg2⫹, the effect of salt can be explained as stabilization of one of the different conformations that the monomer can adopt upon thermal motion of its domains or subdomains (Steinmetz et al., 1997a; Strzelecka-Golaszewska, 2000). Monomer activation also involves conformational changes at the C-terminus (Carlier et al., 1986). Thus, the experimental data revealed conformational changes in actin monomer that can be relevant for actin polymerization and polymer stability. In skeletal muscle 움-actin, these changes are consistent with intramonomer transitions suggested by structural analysis to occur during polymerization. Similar approaches can be applied for comparison of conformational changes in different actin isoforms. 4. Conformational Specificity of Different G-Actins To analyze the conformation of actin isoforms, biochemical as well as spectral approaches developed for studying skeletal muscle actin were used. Specifically, most proteolytic enzymes cleave globular actin within subdomain 2 (Mornet and Ue, 1981), providing a good tool for probing conformational changes within this part of the monomer (Hue et al., 1988; Hozumi, 1988; Strzelecka-Golaszewska et al., 1993). Accordingly, to compare conformations of different actin isoforms, limited proteolysis with trypsin (between residues 62–63 and 68–69; Jacobson and Rosenbusch, 1976), chymotrypsin (between residues 44–45 and 67–68; Konno, 1988), subtilisin (between residues 47–48; Schwyter et al., 1989), and E. coli protease ECP 32 (between residues 42–43; Khaitlina et al., 1991) was used. Kinetics of the cleavage of bovine aorta Ca-G-actin with trypsin, chymotrypsin, and thermolysin showed that this actin is more susceptible to proteolysis than skeletal muscle actin (Cavadore et al., 1985). In contrast to these

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data, the rates of limited proteolysis of scallop Ca-G-actin with trypsin, subtilisin, and ECP 32 were nearly the same as those of skeletal muscle actin (Hue et al., 1988; Khaitlina et al., 1999). The further C-terminal cleavage of the core 33-kDa fragment with trypsin was faster in scallop actin than in skeletal muscle actin (Fig. 5), suggesting that the site within subdomain 1, between Lys359 and Gln360, may be more exposed in scallop actin due to substitutions Thr/Ser358 and Ala/Ser365 (Khaitlina et al., 1999).

FIG. 5 Proteolysis by trypsin of segment 61–69 in scallop adductor muscle and rabbit skeletal muscle G-actins. Ca- or Mg-G-actins (12 애M ) were digested with trypsin at an enzyme:actin mass ratio of 1 : 5 at 20⬚C (A) and 0⬚C (B). Time of digestion, MW markers, and molecular masses of the fragments are indicated. SBTI ⫽ soybean trypsin inhibitor. Reproduced with permission from Khaitlina et al. (1999).

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The inhibition of tryptic cleavages within segment 61–69 upon substitution of tightly bound Mg2⫹ for Ca2⫹ (Strzelecka-Golaszewska et al., 1993) also was observed with scallop actin. Replacement of tightly bound Ca2⫹ by Mg2⫹ did not, however inhibit trypsinolysis of scallop G-actin as efficiently as with skeletal muscle actin, indicating that in scallop Mg-G-actin the nucleotide-containing cleft is more open than in skeletal muscle actin (Fig. 5). The difference was more pronounced at 0 than at 20⬚C. Conformational differences between actin isoforms also were revealed in experiments comparing the binding of cation and ATP at the high-affinity site in different actin isoforms (Strzelecka-Golaszewska et al., 1985a,b; Chen et al., 1995; Kim et al., 1996). No significant difference was found in the equilibrium constants for the tight binding of Ca2⫹ in chicken gizzard smooth muscle 웂-actin and skeletal muscle 움-actin (Strzelecka-Golaszewska et al., 1985a). On the other hand, polymerization behavior of yeast actin indicates that the exchange of tightly bound Ca2⫹ for Mg2⫹ occurs in yeast actin faster than in skeletal muscle actin (Kim et al., 1996). A 3-fold lower value of the equilibrium constant for ATP binding to chicken gizzard smooth muscle 웂-actin than to skeletal muscle 움-actin was documented (Strzelecka-Golaszewska et al., 1985a). This difference correlated with a higher rate of spontaneous or EDTA-induced inactivation of chicken gizzard smooth muscle 웂-actin and bovine aorta smooth muscle 움-actin relative to cardiac and skeletal muscle 움-actins (StrzeleckaGolaszewska et al., 1980, 1985b). Skeletal muscle 움-actin also has a higher affinity for ATP than does yeast cytoplasmic actin in the presence of Ca2⫹ (Chen et al., 1995), and nucleotide exchange was found to proceed in yeast actin at a 5-fold higher rate than in skeletal muscle 움-actin (Miller et al., 1995). These properties resemble properties of the mutant yeast actin in which conversion of Ser14 to Ala resulted in a 40- to 60-fold decrease in affinity for ATP, alterations in the susceptibility of subdomain 2 to proteases, and decreased thermostability (Chen et al., 1995). The hydroxyl group of Ser14 is assumed to form a hydrogen bond with the amide nitrogen of Gly74, thus stabilizing the monomer conformation (Chen et al., 1995). Substitutions Cys/Val10 and/or Val/Cys17 in the proximity of this site in nonsarcomeric actins may influence the actin conformation in a similar fashion, as does mutation-induced elimination of the Ser14 hydroxyl group. Amino acid differences within subdomain 1 of skeletal muscle and scallop actins appeared to result in a small but reproducible difference in the intrinsic fluorescence. The ratio of fluorescence intensities at 320 and 360 nm, which is a sensitive indicator of spectral shifts (Turoverov et al., 1976), was determined to be 2.60 ⫾ 0.03 and 2.70 ⫾ 0.03 for monomeric skeletal muscle and scallop Ca-actins, respectively, pointing to a blue shift of intrinsic fluorescence of scallop versus skeletal muscle actin. This difference may be explained by substitution Thr/Val103 in the vicinity of Trp356, which

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makes the microenvironment of this tryptophan more hydrophobic. Differences in circular dichroism spectra of platelet and sarcomeric actins also were observed (Coue et al., 1982). Thus, the data of limited proteolysis and kinetics of ATP binding show that, in different nonsarcomeric Ca-actins, the nucleotide-containing cleft either has the same conformation or can be more accessible than in skeletal muscle actin. In the former case, the difference between the G-actin species can appear upon replacement of tightly bound Ca2⫹ with Mg2⫹, suggesting lower efficiency of the Mg-induced conformational transitions. In other words, the experimental data allow us to assume that—either initially or upon replacement of tightly bound Ca2⫹ with Mg2⫹ —skeletal muscle 움actin monomer is in a more closed conformation than other actin isoforms. In addition, small spectral differences between the actin isoforms point to local conformational variations within subdomain 1 of the monomer. 5. Dynamics and Cooperativity of F-Actins Actin filament is a helical polymer several micrometers in length. The filament structure can be described in two ways: either as a one-start lefthanded genetic helix with a pitch of 5.9 nm or as a two-start right-handed helix with a pitch of 72 nm (Holmes and Kabsch, 1991). According to current models (Holmes et al., 1990; Lorenz et al., 1993), the contacts between the monomers along the long-pitch helical strands are stronger than those between the strands, making the filament sufficiently rigid. The system, however, is not completely stiff but can be regarded as a semiflexible polymer. This polymer can exist in a number of different structural states produced by changes in intermonomer contacts and/or in intramonomer conformation (Oosawa et al., 1972, 1977; Oosawa, 1980; Yanagida et al., 1984, Orlova and Egelman, 1992, 1993). Because the conformation of actin monomer depends on a type of bound nucleotide and cation (StrzeleckaGolaszewska et al., 1993; Kim et al., 1995; Moraczewska et al., 1996, 1999), these factors also affect the filament structure (Orlova and Egelman, 1992, 1993; Muhlrad et al., 1994; Isambert et al., 1995; Strzelecka-Golaszewska et al., 1996; Strzelecka-Golaszewska, 2000). Image analysis of negatively stained actin filaments followed by their three-dimensional reconstruction revealed the difference between the Factin-ADP state and the ADP-BeF3⫺ state, the latter being an analogue of the F-actin-ADP-Pi state or the F-actin-ATP state (Orlova and Egelman, 1992). As compared with the control F-ADP-actin in which subdomain 2 is not visualized, subdomain 2 is visualized in F-ADP-BeF3-actin, suggesting that the release of inorganic phosphate following ATP hydrolysis is accompanied by disordering of the DNase-binding loop, probably due to a break in one of the longitudinal contacts in the filament (Orlova and Egelman,

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1992). In F-ADP-BeF3-actin, subdomain 2 is more strongly protected against proteolysis than in F-ADP-actin (Muhlrad et al., 1994). These conformational changes correlate with destabilization of actin filaments accompanying the release of phosphate after ATP hydrolysis (Korn et al., 1987; Carlier, 1989; Isambert et al., 1995), suggesting that a more open conformation of actin monomer can contribute to lower polymer stability. This suggestion was directly confirmed on yeast actin in which Val159 (the residue making a hydrogen bond with the 웂-phosphate of the tightly bound ATP) was mutated to Asn (Belmont et al., 1999; Belmont and Drubin, 1998). The mutated F-actin depolymerized slowly and had a lower critical concentration for polymerization, indicating that the more stable filaments were formed. The three-dimensional structure of these filaments resembled that of F-ADP-BeF3-actin, with a more closed nucleotide cleft than in the wild-type F-ADP-actin (Belmont et al., 1999). These results imply that the newly formed ADP-Pi-actin filaments may be more stable than the old FADP-actin (Isambert et al., 1995). The changes induced in G-actin by replacement of tightly bound Ca2⫹ with Mg2⫹ are preserved in F-actins. However, similar changes are produced in G-Ca-actin during salt-induced polymerization. Therefore, the differences between Ca- and Mg-F-actins revealed by limited proteolysis are not so large as those between the corresponding G-actins (StrzeleckaGolaszewska et al., 1996: Strzelecka-Golaszewska, 2000). No difference in flexibility and persistence filament lengths of fluorescently labeled F-Caactin and F-Mg-actin was detected by means of fluorescence optical videomicroscopy (Isambert et al., 1995). On the other hand, a striking Ca-induced decrease in the flexibility of Mg-actin filaments was observed in negatively stained electron microscopic samples (Orlova and Egelman, 1993), probably due to binding of Ca2⫹ at the sites of low and moderate affinity. The flexible structure of actin filaments obtained in the absence of Ca2⫹ suggested an apparent rotation of actin subdomain 2 or the DNase-binding loop, decreasing steric hindrance to bending along the long-pitch helixes (Orlova and Egelman, 1993). Similar rotation of subdomain 2 or the DNase-binding loop may be induced by interaction with myosin (Egelman and Orlova, 1995). The other intrinsic property of actin filaments is their cooperativity, i.e., the capability to propagate conformational changes that occur in actin monomer throughout the filament (Egelman and Orlova, 1995). Therefore, the structure of the filaments can be modulated by low amounts of ligands and actin-binding proteins. A near-maximal change in the fluorescence of a probe attached to F-actin was found at substoichiometric amounts of heavy meromyosin (Oosawa et al., 1972; Miki et al., 1982). Propagation of the structural changes also was observed in gelsolin-nucleated filaments (Orlova et al., 1995; Prochniewicz et al., 1996; Khaitlina and Hinssen, 1997)

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and in the presence of substoichiometric concentrations of phalloidin or beryllium fluoride (Drewes and Faulstich, 1990, 1993; Muhlrad et al., 1994; Orlova and Egelman, 1995). Chemically modified actin protomers cooperatively affected the interaction of myosin with unmodified protomers of the same actin filament (Prochniewicz et al., 1993). Interestingly, the cooperativity in the interaction with heavy meromyosin was observed for Mg- but not for Ca-actin (Orlova and Egelman, 1997). This shows that cooperative properties of actin polymer can be used to regulate actin functions, and ligand-induced structural variations within the filament are important for this regulation. Thus, actin filaments are flexible, are highly cooperative, and can undergo different structural transitions. Thereby, the conformational transitions between the open and the closed states of the monomer involving reorientation of subdomain 2 seem to regulate filament stability. 6. Structural Specificity of Different F-Actins F-Actin is fairly resistant to proteolysis (Rich and Estes, 1983). However, when subtilisin is used at a high enzyme to protein ratio, it cleaves F-actin within subdomains 2 and 4 as well as at the C-terminus (Muhlrad et al., 1994; Vahdat et al., 1995; Strzelecka-Golaszewska et al., 1996). This was used as a tool to compare conformations of yeast and skeletal muscle Factin (Kim et al., 1996) as well as those of scallop and skeletal muscle Factins (Khaitlina et al., 1999). When scallop and rabbit skeletal muscle F-Ca-actins were incubated with subtilisin at an enzyme:actin mass ratio of 1 : 50, the yields of the 19- and 21-kDa fragments, corresponding to the cleavage of actin polypeptide chain between Leu67 and Lys68 (Strzelecka-Golaszewska et al., 1996), were higher in scallop actin than in skeletal muscle actin (Fig. 6), indicating that segment 61–69 in scallop F-Ca-actin was more accessible to proteolysis with subtilisin (Khaitlina et al., 1999). Thus, in the polymeric form, the nucleotide-containing cleft in 웁-like scallop actin remains in a more open conformation than in skeletal muscle 움-actin. Three-dimensional reconstruction of negatively stained yeast actin filaments revealed that yeast F-actin displays the cleft more openly than skeletal muscle actin: it is seen as a ‘‘hole’’ between subdomains 2 and 4 of the actin subunit and appears to result in weaker connectivity between subdomain 2 of one subunit and subdomain 1 of the subunit above it on the same strand of actin filament (Orlova et al., 1997). This structural difference correlates with a lower stability of yeast filaments (Buzan and Frieden, 1996; Kim et al., 1996) and is consistent with higher susceptibility of yeast F-actin to limited proteolysis by subtilisin within the DNase loop (Kim et al., 1996) and with weaker binding of rhodamine phalloidin (De

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FIG. 6 Proteolysis by subtilisin of scallop and rabbit skeletal muscle F-actin. Ca- or Mg-Factins (12 애M ) taken at the steady state of polymerization monitored by light scattering were digested with subtilisin at an enzyme:actin mass ratio of 1 : 50 at 20⬚C. Time of digestion, MW markers, and molecular masses of the fragments are indicated. Reproduced with permission from Khaitlina et al. (1999).

La Cruz and Pollard, 1996) than has been observed for skeletal muscle F-actin. Because nucleotide exchange was found to proceed faster in yeast Gactin than in skeletal muscle G-actin (Chen et al., 1995; Miller et al., 1995), the open cleft seems to be characteristic of yeast actin monomer, and this feature is preserved and may be responsible for the difference in the filament structures. In contrast, as can be judged from the limited proteolysis data, the conformation of subdomain 2 in scallop adductor muscle G-Ca-actin is similar to that in rabbit skeletal muscle G-Ca-actin, with the top of the cleft exposed to nearly the same extent (Khaitlina et al., 1999). It was only in the polymers that the nucleotide-containing cleft in 웁-like scallop actin exhibited a more open conformation than in skeletal muscle 움-actin, suggesting the involvement of polymerization-related structural changes in regulation of the different state of the cleft in the two actin species. Thus, the differences in stability of isoactin filaments discussed previously correlate with a more open position of the cleft in yeast and scallop F-Caactins than that in skeletal muscle F-Ca-actin. In yeast actin, this difference can be due to variations in the monomer structure. The conformations of scallop and skeletal muscle G-Ca-actins are similar, but the cleft remains more open after Mg-induced transitions in subdomain 2 as well as in the

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polymer. It can be assumed, therefore, that in scallop 웁-like actin the shift of subdomain 2 to the position in has in the polymer occurs less efficiently than in skeletal muscle 움-actin. This raises the question of how amino acid substitutions in actin isoforms can produce this difference.

III. Correlation between Amino Acid Substitutions and Actin’s Functional Properties A. Effect of Point Alterations in the Actin–Actin Contact Sites on Actin Polymerizability According to the model of actin polymer (Holmes et al., 1990; Lorenz et al., 1993), the monomer–monomer contacts along the strands of the twostart helix are formed by interaction between residues 322 and 325 in subdomain 1 with residues 243–245 in subdomain 4, between residues 286–289 in subdomain 3 with residues 202–204 in subdomain 4, and between residues 166–169 and 375 in subdomains 3/1 with residues 40–45 in subdomain 2. In contrast to these potentially extensive interactions along the filament, the density in the center of the model is low, possibly involving interactions between hydrophobic loop 264–273 on one strand and a hydrophobic pocket formed by residues 166, 169, 171, 173, 285, 289, 63, 64, and 40–45 between the pair of monomers on the opposite strand of the polymer (Fig. 4). This hydrophobic interaction is considered to be the main ‘‘internal’’ factor of filament stability. Modifications in each of these sites may affect the actin–actin contacts and, hence, the efficiency of actin polymerization as well as filament properties (Sheterline et al., 1995). Indeed, polymerization of skeletal muscle actin was completely inhibited upon chemical modification of His40 (Muhlrad et al., 1969; Hegyi et al., 1974) and Lys61 (Burtnick, 1984), which are involved in the extensive contacts between neighboring molecules along the long two-start helix. Polymerization also is stopped by labeling of Tyr53 (Bender et al., 1976) and impaired by labeling of Tyr59 (Lehrer and Elzinga, 1972), which are adjacent to the actin–actin interface. The strong inhibition of actin polymerization upon proteolytic modifications within the DNasebinding loop also was shown (Khaitlina et al., 1988, 1991, 1993; Schwyter et al., 1989). It was demonstrated that the 웁-actin mutant naturally occuring in the human cell line HUT14, which contains Asp245 instead of Gly (Leavitt and Kakunaga, 1980; Vandekerckhove et al., 1980; Kakunaga et al., 1984), exhibits strongly impaired filament assembly (Leavitt et al., 1982; Taniguchi et al., 1988; Aspenstrom et al., 1992). A somewhat smaller effect was ob-

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served when Gly245 was changed to Lys (Aspernstrom et al., 1992). The mutated residue is involved in interactions along the two-start actin helix (Holmes et al., 1990). Consistent with the proposal that the hydrophobic interactions of loop 266–269 stabilize the actin filament (Holmes et al., 1990; Tirion et al., 1995), the decreasing hydrophobicity of this loop due to point mutation Leu266Asp resulted in cold-sensitive polymerization defects (Chen et al., 1993; Kuang and Rubenstein, 1997a,b). The stability of actin filaments also is decreased upon cutting off the two or three C-terminal residues (O’Donohgue et al., 1992; Drewes and Faulstich, 1993; Mossakovska et al., 1993). Mutations of the C-terminal Cys374 for the negatively charged residues followed by synthesis of the mutant actins in the transfected mammalian cell resulted in disorganization of microfilaments, increasing sensitivity to destabilizing effects of cytochalasin B, and decreased incorporation of the mutant actins into the Tritoninsoluble microfilament fraction (Tsapara et al., 1999). The C-terminal modification also was shown to affect incorporation of the modified actin into myofibrils of Drosophila indirect flight muscles (Brault et al., 1999). Thus, even local modifications within the actin–actin contact sites, and specifically within subdomain 2 of the actin molecule (for instance, cleavage of one polypeptide bond), are crucial for maintaining polymer stability and can result in the loss of polymerizability. Modifications at the C-terminus strongly affect the filament dynamics, which can shift an equilibrium between monomeric and polymeric forms of actin in the cell.

B. Localization of Amino Acid Substitutions in the ThreeDimensional Structure and Effect of These Substitutions on Actin Properties Amino acid substitutions that distinguish actin isoforms are distributed primarily in subdomains 1 and 3 of the actin monomer (Table I; Fig. 7; see color insert). Only a few of the substitutions coincide with the monomer– monomer contact sites. Figure 7 also shows that, with the exception of yeast actin (Ng and Abelson, 1980; Gallwitz and Sures, 1981), there are no substitutions in subdomain 2, consistent with the strong effect of modifications within subdomain 2 on actin polymerization, which may lead to the elimination of actin functions. The large cluster of amino acid substitutions that determines the specificity of actin isoforms involves the N-terminus of the polypeptide chain, including the N-terminal acetyl moiety. This region of the actin molecule participates in interactions with many actin-binding proteins (Sheterline et al., 1995). It is rather mobile in F-Mg-actin (Heintz et al., 1996). According

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to computer analysis of hydrogen bonds related to the N-terminal substitutions, the N-terminus appears to be less compact and more flexible in nonmuscle G-actins than in muscle actin isoforms (Mounier and Sparrow, 1997). Modifications of the N-terminal residues by directed mutagenesis did not cause significant changes in actin polymerizability, but did affect the acto–myosin interaction (Hennessey et al., 1993; Sheterline et al., 1995). It has been shown, however, that specific monoclonal antibodies against the N-terminal decapeptide of smooth muscle 움-actin promoted the polymerization of this actin, but the effect was not observed in skeletal muscle 움-actin (Chapponier et al., 1995). Moreover, microinjection of the smooth-muscle-specific decapeptide (but not the skeletal-muscle-specific Nterminal decapeptide) in cultured smooth muscle cells induced the selective disappearance of smooth muscle 움-actin from stress fibers, indicating that the N-terminal sequence unique to actin isoforms plays a role in the regulation of actin assembly in vitro and in vivo (Chapponier et al., 1995). A drastic reduction of actin polymerization is induced by ADP-ribosylation of Arg177 in cytoskeletal actin with botulinum C2 toxin (Aktories et al., 1986, 1992; Vandekerckhove et al., 1988). In intact cells, this effect leads to disorganization of the microfilament network and accumulation of monomeric actin (Reuner et al., 1987). Skeletal muscle actin is not ADPribosylated by botulinum C2 toxin (Aktories et al., 1986), but it can be modified similarly at Arg177 by other Clostridium toxins (Vandekerckhove et al., 1987; Mauss et al., 1984). It is also shown that the ADP-ribosylation of actin accelerates ATP exchange and inhibits ATP hydrolysis (Geipel et al., 1989). These data indicate that the environments of Arg177 in actin isoforms are different, probably due to the substitution at position 176 (Met/Leu, Table I), which may be part of a structural motif related to the ATP-binding domain (Mounier and Sparrow, 1997). Interestingly, the substitution of several internal residues is accompanied by another change that is symmetrical to the first: Cys10/Val17 and Thr103/ Val129 in muscle actins become, respectively, Val10/Cys17 and Val103/ Thr129 in cytoplasmic actins; Leu153, Met176, and Met299 become Met153, Leu176, and Leu299. The axact symmetries of the compensatory pairs of residues suggest that there are very specific requirements for the internal packing of amino acids in actin, probably to preserve and maintain the overall structure (Mounier and Sparrow, 1997). In this context, it is noteworthy that this symmetry is altered in smooth muscle actins, which contain both Cys10 and Cys17. The distance between these residues is too large to form the S–S bridge. However, this structural specificity might account for the similarity in polymerization properties of smooth muscle 웁- and 웂actins (Strzelecka-Golaszewska et al., 1985). Thus, most of the amino acid substitutions existing in different actin isoforms are located far away from the actin–actin contact sites. Some of

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these substitutions can modulate actin structure and/or nucleotide binding. Variability of the surface N-terminal segment in different actin isoforms is, in this respect, of special significance. These indirect effects may be enhanced by allosteric regulatory mechanisms, which are known to be very intensive in actin monomer.

C. Long-Distance Conformational Transitions and Conformational Coupling within Actin Monomer A large number of studies provide evidence for intensive, long-range conformational coupling within the actin monomer, which may be essential for actin function. The types of divalent cation and tightly bound nucleotide determine the structural transitions at the C-terminus of action (Frieden and Patane, 1985; Valentin-Ranc and Carlier, 1989; Strzelecka-Golazsewska et al., 1993; Orlova and Egelman, 1993). In the reciprocal direction, modifications at the C-terminus of actin were shown to affect the binding of nucleotide (Drummond et al., 1992; Crosbie et al., 1994; StrzeleckaGolazsewska et al., 1995) and to slow down the rate of ATP hydrolysis during polymerization (Mossakovska et al., 1993; Strzelecka-Golazsewska et al., 1995). Conformational communication of the N-terminus, namely the 18–29 loop, with the nucleotide cleft and other regions of the actin monomer was demonstrated. Interaction with antibodies directed against the 18–29 peptide slowed down the rate of nucleotide exchange and increased the fluorescence intensity of actin-bound ␧-ATP and a pyrene probe attached to Cys374 (Adams and Reisler, 1994). Communication between the DNase-binding loop at the top of the monomer and the C-terminus was also shown. Removal of the three C-terminal amino acids caused inhibition of subtilisin cleavage within the DNaseIbinding loop (Strzelecka-Golazsewska et al., 1995). On the other hand, modifications within the DNaseI loop upon binding of DNaseI (Crosbie et al., 1994; Usmanova et al., 1998) or by limited proteolysis (Kuznetsova et al., 1996) resulted in structural alterations within actin subdomain 1. In addition, gelsolin, which is known to interact with subdomains 1 and 3 of the actin molecule, affected the structure of the DNaseI-binding loop (Khaitlina and Hinssen, 1997), whereas modifications within the DNaseI loop changed the interaction of actin with gelsolin (Khaitlina and Hinssen, 1997) and Cap Z (Usmanova et al., 1998). A prolonged delay in polymerization resulting from modification of Arg95 and Arg372 within subdomain 1 also is documented ( Just et al., 1995). Thus, subdomain 1 is significantly involved in long-range conformational coupling between distant regions of the actin monomer. Therefore, it may be assumed that amino acid substitutions within subdomain 1 might be

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responsible, at least in part, for less efficient structural transitions in subdomain 2 during polymerization of scallop actin and, hence, responsible for the more open position of the cleft (Khaitlina et al., 1999). In line with this assumption, computer analysis of hydrogen bonds in muscle and nonmuscle actins revealed a higher number of internal hydrogen bonds in cytoplasmic actins than in muscle actins. These extra internal hydrogen bonds may affect the internal dynamics of cytoplasmic actins, probably making them less flexible and more compact (Mounier and Sparrow, 1997). This implies that substitutions in the primary structure of isoactins that are distant from actin–actin contact sites can modulate the ability of the monomers to form more or less stable polymers by long-range (allosteric) effects on the contact sites.

D. Effect of Actin-Binding Proteins on the Conformation of Actin Isoforms The relatively small size of the actin monomer, the subdomain structure, and the intensive cooperative coupling within the monomer allow the expectation of conformational changes in response to interaction with any actinbinding protein. Some of these effects have been shown. Binding of myosin subfragment 1 caused structural transitions within actin subdomain 2, similar to the changes induced by the tightly bound Mg2⫹ (Chen et al., 1992; Fievez and Carlier, 1993, Wawro et al., 1996). In addition, myosin subfragment 1 restored the polymerizability of actin cleaved within the DNasebinding loop (Wawro et al., 1996). Gelsolin bound to subdomains 1 and 3 of G-actin induced conformational changes within the DNase-binding loop and restored the nucleating activity of proteolytically modified Ca-actin (Khaitlina and Hinssen, 1997). Profilin accelerates the exchange of the actin-bound nucleotide (Goldschmidt-Clermont et al., 1992), and this effect correlates with an open state of the actin monomer in the 웁-actin–profilin complexes (Chik et al., 1996). These data indicate that actin-binding proteins can promote actin polymerization via the long-range effects on actin conformation that stabilize the polymer. The structure of actin polymer can be changed upon interaction with cofilin, which decreases the filament twist and thus produces filaments with approximately 25% shorter ‘‘actin crossover’’ than normal actin filaments (McGough et al., 1997). Rotation of subdomain 2 or the DNaseI-binding loop by about 25⬚ was established in the actin–scruin complexes (Owen and DeRosier, 1993). As discussed previously, gelsolin also was shown to change the structure of actin filament (Orlova et al., 1995; Prochniewicz et al., 1996; Khaitlina and Hinssen, 1997). If the interaction of actin isoforms

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with actin-binding proteins is different, these effects provide additional mechanisms for the regulation of actin isoform specificity. However, the interaction of actin isoforms with actin-binding proteins has not been investigated much. Most of the actin-binding proteins are bound to subdomain 1 (Sheterline et al., 1995), which contains many amino acid substitutions (Fig. 7). Therefore, the interaction of actin-binding proteins with different actin isoforms can be different. Indeed, different affinities of actin isoforms to myosin (Prochniewicz and Strzelecka-Golaszewska, 1980; Strzelecka-Golaszewska and Sobieszek, 1981; Mossakowska and Strzelecka-Golaszewska, 1985) were shown. Differences in conformational dynamics of the C-terminal region of yeast and skeletal muscle F-actins may account for the less efficient interaction of yeast actin with myosin (Prochniewicz and Thomas, 1999). Tropomyosins also bind actin with different affinities (Pitterger et al., 1994). Preferential interaction of cytoplasmic actins with profilin (Larsson and Lindberg, 1988; Ohshima et al., 1989) as well as that of 웁-actin with proteins of the plasmin-fimbrin family (Hofer et al., 1997; Prassler et al., 1997) and with ezrin (Shuster and Herman, 1995; Yao et al., 1995, 1996) has been reported. Distrophin and utrophin constructs also bind to nonmuscle (platelet) actin with an affinity fourfold higher than to skeletal muscle actin. The different binding of these proteins to actin isoforms can modulate their effects on the dynamics of actin polymerization and filament properties. Furthermore, together with specific localization of actin-binding proteins within the cell, the different affinity can be sufficient for proper targeting of isoactins into the cell compartments.

IV. Concluding Remarks Actin is involved in various cell processes. Therefore, it is tempting to correlate the appearance of different actin isoforms with specific intracellular localization and/or cell functions. Indeed, the expression of actin isoforms is under strong tissue-specific and developmental control. Isoactins are sorted specifically within the cell and cannot be substituted for each other without changes in cell morphology and function. Changes in the tissue-specific synthesis of actin isoforms induced by the abolishment or alteration of control mechanisms usually coincide with alterations in cell morphology and function. These data suggest a functional specificity for actin isoforms. The question is whether this specificity may be accounted for by isoactin’s properties. The main property of actin is its ability to polymerize reversibly. On the other hand, the most striking difference between muscle contractile systems and the cytoskeleton is the difference in their dynamics: whereas myofibrils

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are fairly stable structures, microfilament structures of cytoskeleton are assembled and disassembled depending on cell requirements. Therefore, an attempt was made to correlate polymerizability and dynamics of actin isoforms with stability and dynamics of the contractile systems that utilize these isoactins. Actin isoforms are segregated within the cell in such a way that 움-actin is involved in contractile activity, whereas 웁- and 웂-actins are associated with cytoskeleton. In accord with their specific localization in the dynamic structures, 웁- and 웂-isoactins polymerize not as efficiently as and depolymerize faster than skeletal muscle 움-actin when polymerization conditions deviate from the optimal. This implies that microfilaments of cytoskeleton consist of actin isoforms that in vitro form less stable polymers than sarcomeric actin isoforms. It is also possible that the lower rate and higher critical concentration for the polymerization of 웁- and 웂-actins may play a direct role in the targeting of actin isoforms, if concentrations of isoactins in different cell compartments are different. Intracellular sorting of the two cytoplasmic actin isoforms seems to result in 웁-actin being associated with actively moving cytoplasm, whereas 웂-actin forms a more constant part of the cytoskeleton. The polymerizability of 웁like scallop actin is, however higher than that of smooth muscle 웂-actin (Prochniewicz and Yanagida, 1981; Khaitlina, 1986). On the other hand, in the only work that compared the polymerizability of purified cytoskeletal 웁- and 웂-isoactins (Toyama and Toyama, 1988), the rate of polymerization of 웁-actin was slower than that of 웂-actin, consistent with the involvement of 웂-actin in the more stable part of cytoskeleton. This shows that scallop and gizzard actins, although good analogues of cytoplasmic 웁- and 웂-actins, are only analogues, and more experiments should be done to reveal the differences between the two cytoplasmic actin species. Higher stability of skeletal muscle 움-actin polymers correlates with a more closed nucleotide-containing cleft than in 웁-actin. It is known that a closed position of the cleft induced by the replacement of tightly bound Ca2⫹ with Mg2⫹ correlates with more efficient polymerization. In contrast, under conditions that reduce actin polymerizability (when ADP rather than ATP is a tightly bound nucleotide, at low temperature, or upon cleavage of the peptide bond within the DNase loop), the nucleotide cleft seems to be more open. Hence, the more open position of the cleft in 웁- and 웂-Factins may contribute to their lower stability. A structural basis for this difference seems to be the different conformations of subdomains 2 and 4 in the F-actin subunits, which affect the state of the cleft. The cleft can be more open in 웁/웂-actin monomers, as is shown for yeast actin. On the other hand, in scallop adductor muscle 웁-like actin, the difference appears during polymerization and seems to be due to less efficient transition of subdomain 2 to the position it has in the polymer.

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The three-dimensional structures of muscle and nonmuscle actins are similar, and most of the substitutions are located far away from the actin-actin contact sites where their appearance could result in a loss of polymerizability and interaction with actin-binding proteins. However, substitutions located in other regions of the molecule may affect both the structure and stability of actin polymers. The structures formed by the two sets of variable residues in subdomain 1 (Cys10, Ala18, Ala29, and Leu105) and subdomain 3 (Leu153, Asn162, Met176, and Met299) seem to be involved in ATP-related stabilization of the molecule (Mounier and Sparrow, 1997). Interaction between Ser14 and Gly74 is suggested to play a role in controlling the conformation of subdomain 2 relative to subdomain 1 (Chen et al., 1995), and variable residues at positions 10, 16, 17, and 76 may influence this interaction. Furthermore, subdomain 1 is involved significantly in long-range conformational coupling between distant regions of the actin monomer. Because lower internal dynamics of nonmuscle actins has been predicted (Mounier and Sparrow, 1997) it is plausible that amino acid sequence differences within subdomain 1 may be responsible, at least in part, for the less efficient structural transitions in subdomain 2 during polymerization and, hence, responsible for the more open state of the cleft. It may be assumed, therefore, that differences in amino acids that are distant from actin–actin contact sites can modulate the ability of the actin monomers to form more or less stable polymers by long-range (allosteric) regulation of the contact sites. The number of amino acid substitutions between different actin isoforms is small, and many of these substitutions are conservative. This raises a debate about faint effects of substitutions on actin properties. In this respect, the experiments on the Drosophila flight-muscle-specific Act88F gene (Fyrberg et al., 1998) are worth noting. When 10 substitutions characteristic of different Drosophila actins were introduced independently into the Act88F gene and the chimeric proteins were synthesized in flies, only 1 of the 10 transformations perturbed myofibrillar function, demonstrating that most of these isoform-specific amino acid replacements are of minor significance. Similar results were obtained when portions of the Act88F gene were substituted. However, transformation of flies with the muscle-specific actin gene, which was completely converted to a nonmuscle isoform by the introduction of 18 amino acid replacements, resulted in the disruption of flight muscle structure and function. This implies that the effects of amino acid substitutions, while minor individually, collectively confer unique properties (Fyrberg et al., 1998). Further modulation of actin assembly by actin-binding proteins can contribute to the regulation of actin cytoskeleton dynamics. Transient enhancement of the microfilament stability required for various cell functions may be achieved by recruiting actin-binding proteins that also can regulate the

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stability and properties of actin filaments allostericaly. In addition, specific interaction of actin-binding proteins with actin subdomain 1 may be significant for proper targeting of actin isoforms in the cell, which is determined by their C-terminal sequences (von Arx et al., 1995; Kaech et al., 1997). It is interesting that the highly variable C-terminal cysteine motifs are required for specific targeting and membrane association of Rab proteins (Chavnier et al., 1991). Thus, subdomain 1 of the actin molecule plays a role of the regulatory part of the monomer, and the substitutions within its primary structure that specify actin isoforms are important to modulate actin functions directly or via allosteric regulatory mechanisms.

Acknowledgment I am grateful to Prof. Marcus Schaub and Prof. Hanna Strzelecka-Golaszewska for helpful discussions of the manuscript. This work was supported in part by Grant 99-04-49482 from the Russian Foundation for Basic Research.

References Adams, S. B., and Reisler, E. (1994). Sequence 18–29 on actin: Antibody and spectroscopic probing of conformational changes. Biochemistry 33, 14426–14433. Aktories, K., Wille, M., and Just, I. (1992). Clostridial actin-ADP-ribosylating toxins. Curr. Top. Microbiol. Immunol. 175, 97–113. Aktories, K., Barmann, M., Ohishi, I., Tsuyama, S., Jacobs, K. H., and Habermann, E. (1986). Botulinum C2 toxin ADP-ribosylates actin. Nature 322, 390–392. Allen, P. G., Shuster, C. B., Kas, J., Chaponnier, C., Jamney, P. A., Herman, L. M. (1996). Phalloidin binding and rheological difference among actin isoforms. Biochemistry 35, 14062– 14069. Antin, P. B., and Ordahl, C. P. (1991). Isolation and characterization of an avian myogenic cell line. Dev. Biol. 143, 111–121. Aspenstrom, P., Engkvist, H., Lindberg, U., and Karlsson, R. (1992). Characterization of yeast-expressed 웁-actin, site-specifically mutated at the tumor-related residue Gly245. Eur. J. Biochem. 207, 315–320. Bandman, E. (1992). Contractile protein isoforms in muscle development. Dev. Biol. 154, 273–283. Bassell, C. G., and Singer, R. H. (1997). mRNAs and cytoskeletal filaments. Curr. Opin. Cell Biol. 9, 109–115. Bassell, C. J., Zhang, H., Byrd, A. L., Femino, A. M., Singer, R. H. Taneja, K. L., Lifshitz, L. M., herman, I. M., and Kosik, K. S. (1998). Sorting of 웁-actin mRNA and protein in neurites and growth cones in culture. J. Neurosci. 18, 251–265. Belmont, L. D., and Drubin, D. G. (1998). The yeast V159N actin mutant reveals roles for actin dynamics in vitro. J. Cell Biol. 142, 1289–1299. Belmont, L. D., Orlova, A., Drubin, D. G., and Egelman, E. H. (1999). A change in conformation associated with filament instability after Pi release. Proc. Natl. Acad. Sci. USA 96, 29–34.

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