CELL MOTILITY

Historical Development of Cell Motility

In the first half of the 20^th century, it was generally accepted that myosin and actin are proteins specific for muscle, and muscle contractility is a specific interaction of actin, myosin and ATP. It came as a surprise when in 1954 Hoffmann-Berling reported that water-glycerol extracted amnion fibroblasts contract upon addition of ATP. Figures CM1a and 1b show that the elongated fibroblast cells shrink to a small globule after addition of ATP.

Fig. CM1a, shows the fibroblast without ATP (Reprinted from Biochim. Biophys. Acta, vol. 14, Hoffmann-Berling, H., Adenosintriphosphat als Betriebsstoff von Zellbewegungen, pp. 182-194, Copyright 1954, with permission from Elsevier Science).

Fig. CM1b, shows the fibroblast after addition of ATP .(Reprinted from Biochim. Biophys. Acta, vol. 14, Hoffmann-Berling, H., Adenosintriphosphat als Betriebsstoff von Zellbewegungen, pp. 182-194, Copyright 1954, with permission from Elsevier Science).

Subsequently Hoffmann-Berling showed that glycerinated epithelial cells, hen embryos, or Jensen tumor tissues could be contracted with ATP, similarly to the contraction of glycerinated muscle fibers. The idea that actin and myosin are present in mammalian cells, other than muscle, became the working hypothesis of several laboratories:

Isolated actin-like proteins were identified by SDS gel electrophoresis, polymerization-depolymerization properties, nucleotide content, binding to myosin, activation of myosin ATPase, and tryptic peptide mapping. Actin was identified in cells by electron microscopy of fibers decorated with heavy meromyosin or subfragment-1 of myosin and immunofluorescence.

Isolated myosin-like proteins were identified by ATPase activities, SDS gel electrophoresis, subunit composition, binding to actin and by electron microscopy. Labeled antibodies against myosin provided further proof for the existence of myosin in non-muscle tissues. Thus, by the middle of 1970s actin and myosin were acknowledged as regular components of non-muscle cells.

Contractile proteins were prepared, for example, from brain, oviduct, kidney, blood platelets, liver cells, sympathetic neurons, cultures of fibroblast, plasmodial slime mold, or Dyctyostelium amoebae. From all these data it appeared that the mechanism of motion follows the same principle in biology.

Actin-Binding Proteins

In situ the polymerization-depolymerization of actin is controlled by the actin-binding proteins, which combine with actin monomers, cap the ends of the actin filaments, cross-link actin filaments, or attach actin to membranes. Fundamental cellular processes such as, cytokinesis, lamellipodial and growth cone extension, chemotaxis, endocytosis, or exocytosis are regulated by actin-binding proteins:

Profilin

The classical actin-binding protein, profilin, was discovered in the middle of 1970s. It is a small (12-15 kDa), soluble protein that is present in a high concentration (20-80 µM) throughout the cytoplasm and has a high affinity to cytoplasmic actin (K_d =10^-6 M). Profilin inhibits polymerization of actin by sequestering the monomeric actin (Fig. CM2)

Fig. CM2. Schematic overview of actin polymerization and its regulation by profilin and related proteins affecting the competence of monomers to assemble into filaments (From Stossel, 1989).

The X-ray structure of profilin (Fig. CM3) reveals that the protein is bisected by an antiparallel beta-pleated sheet. Both termini are alpha-helical and pack against the same side of the central sheet, connecting to it by short loops.The X-ray structure of profilin-beta-actin is presented in Fig. CM4. Profilin forms two major contacts with actin in the crystal. The primary contact comprises a region of profilin defined by helix 3, the amino-terminal portion of helix 4, and strands 4, 5, and 6. This region makes contact with a site on actin spanning the large and the small domains of the molecule at the bases of subdomains 1 and 3.

Fig. CM3. Polypeptide fold of profilin. The four helices (H) and the seven strands (S) are shown. (From Schutt et al., reproduced with permission from Nature 365,810-816,1993, http://www.nature.com).

Fig. CM4. The profilin-actin ribbon. Profilin red, actin black. For explanation see the text. (From Schutt et al., reproduced with permission from Nature 365,810-816,1993, http://www.nature.com).

Thymosin beta4 is another actin monomer-binding protein. It is a small peptide, 43 residues, and it competes with profilin for binding to actin (Pollard et al., 2000).

Gelsolin

Of the many proteins isolated from cell extracts, gelsolin is the most potent to solubilize gelatinous (fibrous) actin (this ability is reflected in the name of the protein). Severing of the actin-gel occurs through the weakening of sufficient bonds between actin molecules within a filament to break the filament. Severing includes binding of gelsolin to actin, structural rearrangement within gelsolin, and changing the conformation of actin. After severing, gelsolin remains attached to the barbed end of the actin filament that cannot reanneal or elongate and, thus, the actin network is disassembled. Ca²⁺ is required to the severing process (Sun et al.,1999).

Structure and function: Gelsolin has two tandem homologous halves, each of which contains a 3-fold segmental repeat, segments S1-S3 and S4-S6, respectively (Fig. CM5).

Fig. CM5. Gelsolin structure-function domain. Amino acid residues are numbered and the 6 segments are indicated. Actin, PIP₂, and Ca²⁺-binding segments, and the caspase-3 protease site are shown. (From Sun et al.,1999).

The crystal structure of gelsolin shows that in the absence of Ca²⁺ gelsolin has a compact quaternary structure. Its two halves are held together by a C-terminal S6 tail which latches onto S2 (Fig. CM6). It was predicted from this structure that "Ca²⁺ must induce major conformational changes in each half and in the relation between the halves to accommodate actin binding". Indeed new X-ray diffraction studies revealed domain movement in gelsolin (Robinson et al., 1999). Upon Ca²⁺-binding the S6 domain moved by about 40 angstroms resulting in a major structural reorganization in gelsolin, i.e. the actin-binding site on S4 became exposed enabling the severing and capping of actin filaments to proceed. The various steps of gelsolin action are illustrated in Fig. CM7.

Fig. CM6. Structural model of gelsolin in the absence of Ca2+. (From Sun et al.,1999).

Fig. CM7. Schematic model for gelsolin binding, severing, and capping of an actin filament (blue). The (+) barbed end and the (-) pointed end of the filaments are indicated. Left, gelsolin in the process of Ca2+ activation. This requires tail unlatching, domain separation within each half, and separation of the halves. Middlle, S2-S3 binds to the side of an actin filament via S2. S1 wedges between two actins in the longitudinal axis. S4-S6 reaches across the filament to bind actin at the other strand. Severing occurs when sufficient actin-actin bonds are broken. Right, the terminal actin in each strand is capped by S1 and S4. PIP2 induces gelsolin to dissociate from the ends of the actin filaments; this process is called uncapping. (From Sun et al.,1999).

Phosphoinositides, particularly PIP₂ ,dissociate gelsolin from actin (Fig. CM7), thus reverse the capping of the filaments. Severing of the long actin filaments by gelsolin increases the number of the filaments. Uncapping of gelsolin from these filaments generates many polymerization-capable ends from which actin can grow to rebuild the cytoskeleton to a new specification.

The barbed ends of the actin filaments are also interacting with the capping protein, called CapZ in muscle. The protein exists at micromolar concentration in the cytoplasm, and it has a high affinity to the barbed ends, Kd = 0.1 nM (Pollard et al., 2000).

ADF/cofilin

The actin-depolymerizing factor (ADF), also called cofilin, has been recognized early as a widespread, small (15 -18 kDa) actin binding protein that plays important role in cytokinesis, endocytosis, and in the development of all embryonic tissues (Carlier et al., 1999). ADF/cofilins from different organisms present a high degree of sequence homology and the general mechanism of action of the different ADF/cofilins is conserved. The three-dimensional structure determined by X-ray crystallography is shown on Fig. CM8A. Five central beta-sheets are flanked by three to four alpha-helices. Image reconstruction (Fig. CM8B) shows that ADF interacts with two actin subunits along the long pitch helix, bridging subdomain 1 of the actins with subdomain 2 of the second subunit. As a result of the ADF binding to F-actin, there is a twist of 5^o per subunit and, therefore, the long pitch helices crossover every 27 nm on average instead of 36 nm for standard F-actin filaments.

Fig. CM8. Structure of ADF/cofilin. A, ribbon structure of cofilin showing the G- and F-actin binding domains. B, image reconstruction of a standard actin filament (a), and of a cofilin-decorated actin filament (b). (From Carlier et al., 1999)

Importantly, the stimulus-responsive function of ADF/cofilin is regulated by phosphorylation of a single serine residue. In response to stimuli, ADF is dephosphorylated. The stimuli, such as growth factors, chemotactic peptides, or agents increasing the levels of [Ca²⁺]_i and cAMP, promote the reorganization of the actin cytoskeleton. In quiescent cells, ADF/cofilin appears diffusely distributed in the cytoplasm, the activated (dephosphorylated) protein translocates to regions of the cells where actin filaments are highly dynamic like the leading edge of ruffled membranes, the cleavage furrow of dividing cells, or the neuronal growth clone. Dephosphorylation correlates with increased motility and extension of cellular processes (Carlier et al., 1999). ADF/cofilin increases the turnover of actin filaments which powers actin motility (Fig. CM9).

Fig. CM9. Enhancement of actin turnover by ADF. The F-actin filaments, containing ADF, depolymerize 30-fold faster from their pointed ends than the bare filaments, resulting in accumulation of ADF-ADP-actin pool. After dissociation of ADF, the ADP-actin assembles on the barbed end of the bare filaments. The ADF molecules are red colored. (From Carlier et al., 1999).

Arp2/3 complex and WASp/Scar proteins

Actin related proteins (Arps) participate in a diverse array of cellular processes (Schafer and Schroer, 1999). Wiskott-Aldrich syndrome proteins (WASp/Scar proteins) stimulate the formation of new actin filaments by Arp2/3 complex (Higgs and Pollard, 1999). Fig. CM10 shows that the Arp2/3 complex contains seven protein subunits, Arp2 and Arp3 are actin related proteins, the other five subunits are novel. The Arp2/3 complex cross-links filaments in an end to side manner, with the slow growing end of one filament attached to the side of another at an angle of 70 degrees. Fig CM11 shows the binding of WASp/Scar to an actin filament and to Arp2/3, creating new filaments and cross-linking them into a branching meshwork.

Fig. CM10. Arp2/3 complex structure. Based on nearest neighbor relationship of the subunits from chemical cross linking. Molecular masses in kDa are indicated. (From Higgs and Pollard, 1999).

Fig. CM11. Dendritic nucleation model. Arp2/3 complex (green) binds to the side of a preexisting actin filament (yellow), and WASp/Scar (red) bound to an actin monomer binds to Arp2/3, forming a nucleus for barbed end growth from the side of the filament (From Higgs and Pollard, 1999).

Major advances have been made in research of Arp2/3 complex recently. The crystal structure of the Arp2/3 complex has been determined at 2.0 angstrom resolution (Robinson et al., 2001). Arp2 and Arp3 are folded like actin with distinctive surface features. The structure of the five subunits has also been elucidated. Subsequently, the structure of the Arp2/3 complex in its activated state and in actin filament branch junctions has been described (Volkmann et al, 2001). Internal reflection fluorescence microscopy was used for direct real-time observation of actin filament branching mediated by Arp2/3 complex (Aman and Pollard 2001). Furthermore, it was shown that the binding of ATP to Arp2 is required for filament branching (Le Clainche et al., 2001; Dayel et al., 2001).

External stimuli drive the assembly of the actin filament network, acting through receptors and multiple signal transduction pathways, several of which converge on WSPp/Scar proteins and Arp2/3 complex. Diverse signals including those carried by the Rho family GTPases, Rac, and Cdc42 are involved (Fig. CM11A).

Fig. CM11A. Signaling pathways through WASp/Scar to Arp2/3 complex. (From Pollard et al.,with permission from the Annual Review of Biphysics and Biomolecular Sructure. vol. 29, 2000, by Annual Reviews, http://www.AnnualReviews.org).

Dendritic nucleation model

This is a model for molecular mechanisms contolling actin filament dynamics in nonmuscle cells (Fig. CM11B).

Fig. CM11B. The dendritic nucleation model (From Pollard et al., with permission from the Annual Review of Biphysics and Biomolecular Sructure. vol. 29, 2000, by Annual Reviews, http://www.AnnualReviews.org).

There are 10 steps in the figure. In the first step all the ATP-actin monomers are bound to profilin, there are no free barbed ends; the actin cytoskeletal components are held in a metastable state, poised for assembly. Activation of WASp family proteins (Step 2) leads to activation of the Arp2/3 complex and this creates new barbed ends and subsequently new filaments (Step 3). These filaments grow rapidly (Step 4) and push the membrane forward (Step 5). After a short while, growth of the barbed ends is terminated by capping (Step 6). ATP hydrolysis and Pi dissociation (Step 7) triggers severing and depolymerization of actin filaments by ADF/cofilins (Step 8). LIM kinase inhibits ADF/cofilins (Step 9). Nucleotide exchange catalyzed by profilin recycles ADP-actin to ATP-actin monomer in the pool (Step 10).

Accordingly, the molecular mechanism of motion in nonmuscle cells is a very complex cycle that converts the energy of the hydrolysis of actin-bound ATP into mechanical force through the polymerization and depolymerization of actin filaments ( treadmilling of actin filaments). In a continuously moving cell, assembly and dysassembly of actin filaments are balanced.

Reviews of Pollard et al. (2000), Higgs and Pollard (2001), and Pantaloni et al. (2001) are recommended for a further understanding of the molecular mechanism of motion in nonmuscle cells.

Actin in the cytoskeleton

Living cells have the ability to change their shape and move upon stimuli in the environment. There is a network, called the cytoskeleton, a complex of filamentous proteins, which is the engine of biological adaptation. Actin is the key component of the cytoskeleton, because it exists in two forms, globular and fibrous, and it can also form a gel; these properties are ideal for restructuring the cytoskeleton in response to a variety of signals. For instance, growth factor stimulation promotes actin assembly at the plasma membrane to generate movement, whereas apoptotic signals cause cytoskeletal destruction to elicit characteristic membrane blebbing and morphological changes.

The dynamic role of actin in modifying the cytoskeleton is regulated by the actin binding proteins. Fig. CM 12 illustrates the multiple pathways involved in the regulation. Cellular movements are initiated by extracellular stimuli and the signals transduced through the cell membrane couple the messenger response to actin assembly in the cell. This is illustrated in Fig. CM13.

Fig. CM12. Regulation of actin polymerization by the actin-binding proteins. Symbols used: "C" for monomeric binding proteins; bracket for capping and severing proteins; squiggle for cross-linking proteins. (From Pollard and Cooper, with permission from the Annual Review of Biochemistry,. vol. 35, 1986, by Annual Reviews, http://www.AnnualReviews.org).

Fig. CM13. Schematic overview of how actin-binding proteins might link stimulus-induced signal generation to actin assembly in the cell. The sequence of postulated events is indicated by the numbers in the circles. PI, phosphatidylinositol; DAG, diacylglycerol. (From Stossel, 1989).

The understanding of the mechanism of actin-based motility in cells has been greatly advanced by the work of Loisel et al.(1999), who were able to reconstitute bacterial motility from pure protein components. In addition to actin, only three other components were absolutely needed: the Arp2/3 complex, ADF, and capping protein, all which are actin-binding proteins. These results demonstrate that actin-based propulsion is driven by the free energy released by ATP hydrolysis coupled to actin polymerization, and does not require myosin.

Work on nonmuscle cells has been recently applied to muscle as well. Thus, it was reported that actin dynamics at pointed ends regulates thin filament length in striated muscle (Littlefied et al., 2001).

It was thought until recently that bacteria lacked the actin network that organize eukaryotic cytoplasm. The mreB gene is involved in determining cell shape in rod-like bacteria and recently van den Ent et al. (2001) showed that bacterial MreB protein assembles into filaments with a subunit repeat similar to that of F-actin in eukaryotic cells. In addition, the MreB crystal structure reveals a shape similar to that of actin. Accordingly, prokaryotes possess homologues of both tubulin and actin, suggesting that the building blocks for the actin cytoskeleton originated in prokaryotes before becoming the mainstay of eukaryotic cells.

Non-Muscle Myosins

Beside the conventional myosin-2, at least six more unconventional myosins, myosins-I, -V, -VI, -VII, -IX, and -X, exist (Hasson and Mooseker, 1996). A schematic comparison of conventional and unconventional myosins is shown in Fig. CM14. At the N terminus, each vertebrate unconventional myosin contains a Head, a conserved motor domain, that includes both the ATP- and actin-binding sites. Following the Head is the Neck domain, a light chain binding regulatory part of the structure, that contains a 24-30-amino acid repeat termed the IQ motif. The neck domain also serves as a binding site for calmodulin. Each class of myosin has a distinct Tail domain, that ends at the C terminus, which provides sequences that can serve to target the myosin to its particular subcellular location.

Fig. CM14. Schematic showing the Head, Neck, and Tail domains of vertebrate myosins. Molecular masses and specific binding sites are indicated. (From Hasson and Mooseker, 1996).

Thus myosin tails may have a membrane-binding site and/or a site that binds to a second actin filament independently of the head domain. Depending on its tail, the myosin molecule may move a vesicle along an actin filament, or attach an actin filament to the plasma membrane.

Functional activities of many nonmuscle myosin isoforms are regulated by heavy chain phosphorylation (Redowicz, 2001). For instance, organelle transport by myosin-V was down-regulated by this phosphorylation (Karcher et al., 2001). Mass spectrometry phosphopeptide mapping showed that the tail of myosin-V was phosphorylated in mitotic Xenopus egg extract on a single serine residue localized in the carboxyl-terminal organelle-binding domain. Phosphorylation resulted in the release of the myosin motor from the organelle. The phosphorylated site matched the consensus sequence of calcium/calmodulin-dependent protein kinase II (CaMKII), and inhibitors of CaMKII prevented myosin-V release.

A myosin I isoform was found in the nucleus (Pestic-Dragovich et al., 2000). A unique 16-amino acid amino-- terminal extension characterizes this myosin. This isoform of myosin I appears to be in complex with RNA-polymerase II and may affect transcription.

Microtubules

Microtubules based motility is somewhat different from the actin-myosin contractile system. Microtubules are hollow tubes formed from tubulin subunits; there are two kinds of tubulins, alpha and beta, each with a molecular mass of 50-kDa. The structure of the microtubules is built from 13 protofilaments, each is composed of alternating alpha and beta tubulin subunits and bundled in parallel to form a cylinder (Fig. CM16). Since the 13 protofilaments are aligned in parallel with the same polarity, the microtubule itself is a polar structure, and it is possible to distinguish a plus, fast growing, and a minus, slow growing, end (similarly to those in actin filaments). Brain tissue is very rich in tubulin (10 -20% of the total soluble proteins) but virtually every eucaryotic cell contains tubulin.

Fig. CM16. Illustration of the structure of a microtubule. (From Alberts et al, Copyright 1994, from Molecular Biology of The Cell 3rd ed., by Alberts et al., reproduced .by permission of Routledge, Inc., part of The Taylor and Francis group).

The assembly of tubulin subunits into microtubules is associated with the hydrolysis of the tubulin-bound GTP into GDP (similarly to the hydrolysis of the bound ATP during polymerization of actin). Microtubules depolymerize and repolymerize continually in animal cells (Alberts et al.,1994), and this is central to cell morphogenesis. Thus, microtubules play a major role in separating daughter chromosomes in mitosis. Microtubules are major components of eucaryotic cilia and flagella. These hairlike organelles protrude from the surface of many cells and are involved in motility. The long microtubules extend throughout the cytoplasm and govern the location of organelles and other cell components.

Kinesin and dynein: There are two classes of motor proteins, kinesin and dynein that direct organelle and particle movement along microtubules. Kinesins are a family of proteins, that are involved in organelle transport, in mitosis, in meiosis, and in the transport of synaptic vesicles along axons. Cytoplasmic dyneins are involved in organelle transport and mitosis. Kinesins and dyneins are myosin-like proteins composed of two heavy chains plus several light chains. Each heavy chain contains a conserved, globular, ATP-binding head and a tail composed of a string of rodlike domains. The two head domains are ATPase motors that bind to microtubules, while the tails generally bind to specific cell components and thereby specify the type of cargo that the protein transports. Organelles and vesicles containing kinesin move from the minus end of a microtubule (at a microtubule organizing center, such as centrosome) to the plus end (Fig. CM.17). Hence, kinesin produces movement from the center of a cell to its periphery, called anterograd transport. In contrast, cytoplasmic dynein moves the particles from the plus end to the minus end of the microtubules (Fig. CM17), called retrograd transport.

A structural view of the kinesin movement was revealed recently (Kikkawa et al., 2001). The monomeric form of the kinesin motor was crystallized and the X-ray structure obtained was combined with cryo-electron microscopy. The motor was revealed in its two functionally critical states - complexed with ADP and with a non-hydrolysable analogue of ATP. Analysis of the force generating conformational changes could be done at atomic resolution. The authors suggest that a roughly 20-degree rotation of the motor region of kinesin helps to tighten its grip on the surface of the microtubular tracks it moves along.

Fig. CM17. Movement of kinesin and cytoplasmic dynein along a microtubule. (From Alberts et al, Copyright 1994, from Molecular Biology of The Cell 3rd ed., by Alberts et al., reproduced .by permission of Routledge, Inc., part of The Taylor and Francis group).

Ciliary dynein is a much larger protein than the cytoplasmic dynein: its mass is about 2 million daltons, composed of either 2 or 3 heavy chains and 10 or more smaller polypeptides. The dynein motor protein in cilia and flagella has a head which hydrolyzes ATP and interacts with the adjacent microtubules to generate a sliding force between the microtubules. However, the microtubules are linked together, therefore, they can not slide but must bend. This local bending of the microtubules is the mechanism of ciliary movement, the beating.

Reading articles from the following authors will enhance knowledge in microtubules based motility: Hackney, 1996, Vallee and Sheetz, 1996; Samsó et al., 1998, and Rice et al., 1999.

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