A section of mouse intestine stained for actin (red). the extracellular matrix protein laminin (green), and DNA (blue). Each blue dot of DNA

indicates the presence of a cell. Actin in the microvilli on the apical end

of the epithelial cells can be seen lining the surface facing the lumen (top). Actin can also be seen prominently in the smooth muscle that

surrounds the intestine (bottom). [Micrograph courtesy ofThomas Deerinck

and Mark Ellisman.]

W hen we look through a microscope at the wonder­ful diversity of cells in nature, the variety of cell shapes and movements we can discern is astonish­

ing. At first we may notice that some cells, such as vertebrate sperm, ciliates such as Tetrahymena, or flagellates such as Chlamydomonas, swim rapidly, propelled by cilia and fla­gella. Other cells, such as amebas and human macrophages, move more sedately, propelled not by external appendages but by coordinated movement of the cell itself. We also might notice that some cells in tissues attach to one another, forming a pavementlike sheet, whereas other cells-neurons, for example-have long processes, up to 3 ft in length, and make selective contacts between cells. Looking more closely at the internal organization of cells, we see that organelles have characteristic locations; for example, the Golgi appara­tus is generally near the central nucleus. How is this diversity of shape, cellular organization, and motility achieved? Why is it important for cells tq have a distinct shape and clear internal organization? Let us first consider two examples of cells with very different functions and organizations.

The epithelial cells that line the intestine form a tight, pave­mentlike layer of brick-shaped cells, known as an epithelium

OUTLINE

17.1 Microfilaments and Act in Structures 776

17.2 Dynamics of Actin Filaments 779

17.3 Mechanisms of Actin Filament Assembly 784

17.4 Organization of Actin-Based Cellular Structures 790

Cell Organization and Movement 1: Microfilaments

CHAPTER

(Figure 17-1 a, b). Their function is to import nutrients (such as glucose) from the intestinal lumen across the apical (top) plasma membrane and export them across the basolateral (bottom-side) plasma membrane toward the bloodstream. To perform this directional transport, the apical and basolateral plasma membranes of epithelial cells must have different pro­tein compositions. Epithelial cells are attached and sealed to­gether by cellular junctions (discussed in Chapter 20), which create a physical barrier between the apical and basolateral domains of the membrane. This separation allows the cell to place the correct transport proteins in the plasma membranes of the two surfaces. In addition, the apical membrane has a unique morphology, with numerous fingerlike projections called microvilli that increase the area of the plasma membrane available for nutrient absorption. To achieve this organization, epithelial celb must have an internal structure to give them shape and to deliver the appropriate proteins to the correct membrane surface.

Now consider macrophages, a type of white blood cell that seeks our infectious agents and destroys them by an en­gulfing process called phagocytosis. Bacteria release chemi­cals that attract the macrophage and guide it to the infection.

17.5 Myosins: Actin-Based Motor Proteins 793

17.6 Myosin-Powered Movement s 801

17.7 Cell Migration: Mechanism, Signaling, and Chemotaxis 808

DEMO : Purchase from www.A-PDF.com to remove the watermark

(a) (b) Microvilli Cell junctions

Extracellular matrix

(c) (d)

-- Microfilaments

- Microtubules

Basolateral domain

-- lntermed{ate filaments

Filopodium

Direction of migration

FIGURE 17-1 Overview of the cytoskeletons of an epithelial cell and a migrating cell. (a) Transmission electron micrograph of a thin

section of an epithelial cell from the small intestine, showing the cytoskeletal components of the microvilli. (b) Epithelial cells are highly

polarized, with distinct apical and basolateral domains. An intestinal epithelial cell transports nutrients into the cell through the apical domain

and out of the cell across the basolateral domain. (c) Transmission

electron micrograph of part of the leading edge of a migrating cell. The cell was treated with a mild detergent to dissolve the membranes, which

As the macrophage follows the chemical gradient, twisting and turning to get to the bacteria and phagocytose them, it has to constantly reorganize its cell locomotion machinery. As we will see, the internal motile machinery of macrophages and other crawling cells is always oriented in the direction that they crawl (Figure 17-lc, d).

These are just two examples of cell polarity-the ability of cells to generate functionally distinct regions. In fact, as you think about all types of cells, you will realize that most of them have some form of cell polarity. An additional and fundamental example of cell polarity is the ability of cells to divide: they must first select an axis for cell division and

also allows solubilization of most cytoplasmic components. The

remaining cytoskeleton was shadowed with platinum and visualized in the electron microscope. Note the meshwork of actin filaments visible in this micrograph. (d) A migrating cell, such as a fibroblast or a macro­

phage, has morphologically distinct domains, with a leading edge at the front. Microfilaments are indicated in red, microtubules in green, and

intermediate filaments in dark blue. The position of the nucleus (light blue oval) is also shown. [Part (a) Courtesy of Mark Mooseker; Part (c) from

T. M. Svitkina et al., 1999,1. Cell 81ol. 145:1 009, courtesy ofTatyana Svitkina.)

then set up the machinery to segregate their organelles a long that axis.

A cell's shape, internal organization, and functional po­larity are provided by a three-dimensional filamentous pro­tein network called the cytoskeleton. The cytoskeleton can be isolated and visualized after treating cells with gentle de­tergents that solubilize the plasma membrane and internal organelles, releasing most of the cytoplasm (Figure 17-lc). The cytoskeleton extends throughout the cell and is attached to the plasma membrane and internal organelles, thus pro­viding a framework for cellular organization. The term cyto­skeleton may imply a fixed structure like a bone skeleton. In

774 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

.·

... Microfilaments Microtubules Intermediate filaments ·c: :I

..Q :I

(/)

Actin aP-Tubulin dimer Various

Q)

:; g 7-9 nm t 4111fiJ·Iilllll·IIII·B•• .. u;

FIGURE 17-2 The components of the cytoskeleton. Each filament

type is assembled from specific subunits in a reversible p rocess so that

cells can assemble and disassemble filaments as needed. Bottom

panels show localization of the three filament systems in cultured cells

fact, the cytoskeleton can be very dynamic, with components capable of reorganization in less than a minute, or it can be quite stable for hours at a time. As a result, the lengths and dynamics of filaments can vary greatly, filaments can be as­sembled into diverse types of structures, and they can be regulated loca lly in the cell.

The cytoskeleton is composed of three major filament systems, shown in Figure 17-1 b, d as well as in Figure 17-2, all of which are organized and regulated in time and space. Each filament system is composed of a polymer of assembled subunits. The subunits that make up the filaments undergo regulated assembly and disassembly, giving the cell the flexi­bility to assemble or disassemble different types of structures as needed.

• Microfilaments are polymers of the protein actin orga­nized into functional bundles and networks by actin-binding proteins. Microfilaments are especially important in the or­ganization of the plasma membrane, giving shape to surface structures such as microvilli. Microfilaments can function on their own or serve as tracks for A TP-powered myosin motor proteins, which provide a contractile function (as in muscle) or ferry cargo along microfilaments.

• Microtubules are long tubes formed by the protein tubulin and organized by microtubule-associated proteins. They of­ten extend throughout the cell, providing an organizational framework for associated organelles and structural support to cilia and flagella. They also make up the structure of the mitotic spindle, the machine for separating duplicated chro­mosomes at mitosis. Molecular motors called kinesins and

10nmt~

as seen by immunofluorescence microscopy of actin, tubulin, and an

intermediate filament protein, respect ively. (Actin and tubulin courtesy

of D. Garbett and A. Bretscher; intermediate filaments Copynght Molecular

Expressions, Nikon & FSU.]

dyneins transport cargo along microtubules and, like myo­sin, are also powered by A TP hydrolysis.

• Intermediate fi laments are t issue-specific filamentous structures that serve a number of different functions, includ­ing lending structural support to the nuclear membrane, pro­viding structura l integrity to cells in tissues, and serving structural and barrier functions in skin, hair, and nails. Un­like the situation for microfilaments and microtubules, there are no motors that use intermediate filaments as tracks.

As we can see in Figure 17- 1, cells can construct very dif­ferent arrangements of their cytoskeletons. To establish these arrangements, cells must sense signals-either from soluble factors bathing the cell, from adjacent cells, or from the extra cellular matrix-and interpret them (Figure 17-3). These sig­nals are detected by cell-surface receptors that activate signal-transduction pathways that ultimately converge on fac­tors that regulate cytoskeletal organization.

The importance of the cytoskeleton for normal cell func­tion and motility is evident when a defect in a cytoskeletal component- or in cytoskeletal regulation--causes a disease. For example, about 1 in 500 people has a defect that affects the contractile apparatus of the heart, which results in car­diomyopathies varying in degree of severity. Many diseases of the red blood cell affect the cytoskeletal components that support these cells' plasma membranes. Metastatic cancer cells exhi bit unregulated motility due to misregulation of the cytoskeleton, breaking away from their tissue of origin and migrating to new locations to form new colonies of uncon­trolled growth.

CHAPTER 17 • Cell Organization and Movement 1: Microfilaments 775

Signals from soluble factors, other cells, the extracellular matrix

I 1 \

1 I Signal transduct ion

pathways

1 Cytoskeleton

I \ Organization Cell shape,

and movement movement,

/

Plasma membrane with receptors

of organelles and contraction

FIGURE 17-3 Regulation of cytoskeleton function by cell signaling. Cells use cell-surface receptors to sense external signals from the extracellular matrix, other cells, or soluble factors. These signals are transmitted across the plasma membrane and activate specific cytosolic signaling pathways. Signals-often integrated from more than one receptor-lead to the organization of the cytoskeleton to provide cells with their shape, as well as to determine organelle distribution and movement. In the absence of external signals, cells still organize their internal structure, but not in a polarized manner.

In this and the following chapter, we discuss the struc­ture, function, and regulation of the cytoskeleton. We will see how a cell arranges its cytoskeleton to determine cell shape and polarity, to provide organization and motility to its or­ganelles, and to be the structural framework for such pro­cesses as cell swimming and cell crawling. We will discuss how cells assemble the three different filament systems and how signal-transduction pathways regulate these structures both locally and globally. How the cytoskeleton is regulated during the cell cycle is discussed in Chapter 19, and how it participates in the functional organization of tissue is covered in Chapter 20. Our focus in this chapter is on microfilaments and actin-based structures. Although we initially examine the cytoskeletal systems separately, in the next chapter we will see that microfilaments cooperate with microtubules and in­termediate filaments in the normal functioning of cells.

17.1 Microfilaments and Actin Structures Microfilaments can assemble into a wide variety of different types of structures within a cell (Figure 17-4a). Each of thr~e diverse structures underlies particular cellular functions. Mi­crofilaments can exist as a tight bundle of filaments making up the core of the slender, fingerlike microvilli, but they can also be found in a less ordered network beneath the plasma membrane, known as the cell cortex, where they pro\'ide support and organization. In epithelial cells, microfilaments

form a contractile band around the cell, the adherens belt, that is intimately associated with adherens junctions (Chapter 20) to provide strength to the epithelium. In migrating cells, a network of microfilaments is found at the front of the cell in the leading edge, or lamellipodium, which can also have pro­truding bundles of filaments called filopodia. Many cells have contractile microfilaments called stress fibers, which at­tach to the external substratum through specialized regions c.:allcd focal adhesions or focal contacts (discussed in Chap­ter 20). Specialized cells such as macrophages use contractile microfilaments in a process called phagocytosis to engulf and internalize pathogens (such as bacteria), which are then destroyed internally. Highly dynamic, short bursts of actin filament assembly can power the movement of endocytic vesicles away from the plasma membrane. At a late stage of cell division in animals, after all the organelles have been duplicated and segregated, a conbractile ring forms and con­stricts to generate two daughter cells in a process known as cytokinests. Thus cells use actin filaments in many ways: in a structural role, by harnessing the power of actin polymeriza­tion to do work, or as tracks for myosin motors. The elec­tron micrograph in Figure 17-4b shows microfilaments in microvilli. Different arrangements of microfilaments often coexist within a single cell, as shown in Figure 17-4c, in this case a migrating fibroblast.

The basic building block of microfilaments is actin, a protein that has the remarkable property of reversibly as­sembling into a polarized filament with functionally distinct ends. These filaments are then molded into the various struc­tures described in the previous paragraph by actin-binding proteins. The name microfilament refers to actin in its po­lymerized form, with its associated proteins. In this section, we look at the actin protein itself and the filaments into which it assembles.

Actin Is Ancient, Abundant, and Highly Conserved

Actin is an abundant intracellular protein in eukaryotic cells. In muscle cells, for example, actin comprises 10 percent by weight of the total cell protein; even in nonmuscle cells, actin makes up 1-5 percent of the cellular protein. The cytosolic concentration of actin in nonmuscle cells ranges from 0.1 to 0.4 mM; in special structures such as microvilli, however, the local actin concentration can be as high as 5 mM. To grasp how much actin is present in cells, consider a typical liver cell, which has 2 X 104 insulin receptor molecules but approximately 5 X 108

, or half a billion, actin molecules. Because they form structures that extend across large parts of the cell interior, cytoskeletal proteins are among the most abllndant proteins in a cell.

Actin is encoded by a large family of genes that gives rise to some of the most conserved proteins within and across species. The protein sequences of actins from amebas and from animals are identical at 80 percent of the amino acid positions, despite about a billion years of evolution. The multiple actin genes found in modern eukaryotes are related

776 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

• 0 •

(a)

Microvilli

Filopodia

Cell cortex

Lamellipodium/ leading edge

(b) (c)

Phagocytosis Moving endocytic vesicles Contractile ring

FIGURE 17-4 Examples of microfilament-based structures. (a) In (c) A cell moving toward the top of the page, stained for actin with flue-each panel, microfilaments are depicted in red. (b) Scanning electron rescent phalloidin, a drug that specifically binds F-actin. Note how micrograph of the apical region of a polarized epithelial cell, showing many different organizations can exist in one cell. [Part (b) courtesy of the bundles of actin filaments that make up the cores of the microvilli. N. Hirakawa; Part (c) courtesy of J. V. Small.]

to a bacterial gene that has evolved to have a role in bacterial cell-wall synthesis. Some single-celled euka ryotes, such as yeasts and amebas, have one or two ancestral actin genes, whereas multicellular organisms often contain multiple actin genes. For instance, humans have six actin genes, and some plants have more than 60 actin genes (although most are pseudogenes, which do not encode functional actin pro­teins). Each func~ional actin gene encodes a different isoform of the protein. Actin isoforms can be classified into three groups: the a-actins, [3-actins, and ')'-actins. In vertebrates, four actin isoforms are present in specific types of muscle cells, and two isoforms are found in nonmuscle cells. These six isoforms differ at only about 25 of the 375 residues in the complete protein, or show about 93 percent identity. Al­though these differences may seem minor, the three types of isoforms have different functions: a-actin is associated with contractile structures, )'-actin accounts for filaments in stress fibers, and [3-actin is enriched in the cell cortex and leading edge of motile cells.

G-Actin Monomers Assemble into Long, Helical F-Actin Polymers

Actin exists as a globular monomer called G-actin and as a filamentous polymer called F-actin, which is a linear chain of G-actin subunits. Each actin molecule contains a Mg2

+ ion complexed with either ATP or ADP. The importance of the interconversion between the ATP and the ADP forms of actin is discussed later.

X-ray crystallographic analysis reveals that the G-actin monomer is separated into two lobes by a deep cleft (Fig­ure 17-5a). At the base of the cleft is the ATPase fold, the site where ATP and Mg2 are bound. The floor of the cleft acts as a hinge that allows the lobes to flex relative to each other. When ATP or ADP is bound to G-actin, the nucleo­tide affects the conformation of the molecule; in fact, with­out a bound nucleotide, G-actin denatures very quickly. The addition of cations-Mg2 , K.,., or NaT -to a solution of G-actin will induce the polymerization of G-actin into F-actin filaments. The process is reversible: F-actin depolymerizes into G-actin when the ionic strength of the solution is low­ered. The F-actin filaments that form in vitro are indistin­guishable from microfilaments seen in cells, indicating that F-actin is the major component of microfilaments.

From the results of x-ray diffraction studies of actin fila­ments and the actin monomer structure shown in Figure 17-5a, scientists have determined that the subunits in an actin fila­ment arc arranged in a helical structure (Figure 17-5b) . In this arrangement, the filament can be considered as two heli­cal strands wound around each other. Each subunit in the strucmre contacts one subunit above, one below in one strand, and two in the other ~trand. The subunits in a single strand wind around the back of the other strand and repeat after 72 nm or 14 actin subunits. Since there are two strands, the actin filament appears to repeat every 36 nm (see Figure 17-5b). When F-actin is negatively stained with uranyl acetate for electron microscopy, it appears as a twisted string whose diameter varies between 7 and 9 nm (Figure l7-5c).

17.1 Microfilaments and Actin Structures 777

(a)

FIGURE 17-5 Structures of monomeric G-actin and F-actin filaments. (a) Model of actin monomer (measuring 5.5 X 5.5 X 3.5 nm)

shows that it is divided by a central cleft into two approximately

equal-sized lobes and four subdomains, numbered I-IV. ATP (red) binds

at the bottom of the cleft and contacts both lobes (the yellow ball

represents Mg2 ). TheN- and ( -termini lie in subdomain I. (b) An actin

filament appears as two strands of subunits. One repeating unit consists

of 28 subunits (14 in each strand, indicated by* for one strand),

covering a distance of 72 nm. The ATP-binding cleft of every actin

F-Actin Has Structural and Functional Polarity All subunits in ·an actin filament are oriented the same way. Consequently, a filament exhibits polarity; that is, one end differs from the other. As we will see, one end of the filament is favored for the addition of actin subunits and is designated the ( +) end, whereas the other end is favored for subunit dissociation, designated the (-) end. At the ( +) end, the ATP-binding cleft of the terminal actin subunit contacts the neighboring subunit, whereas on the (-) end, the cleft is ex­posed to the surrounding solution (see Figure 17-Sb).

Without the atomic resolution afforded by x-ray crystal­lography, the cleft in an actin subunit and therefore the po­larity of a filament is not detectable. However, the polarity

EXPERIMENTAL FIGURE 17-6 Myosin 51 decoration demonstrates the polarity of an actin filament. Myosin 51 head domains bind to actin subunits in a particular orienta­

tion. When bound to all the subunits in a filament, 51 appears

to spiral around the filament. This coating of myosin heads

produces a series of arrowheadlike decorations (arrows),

most easily seen at the wide views of the filament. The

polarity in decoration defines a pointed (- ) end and a

barbed (+ )end. [Courtesy of R. Craig.)

( ) end

(b ) (c)

(-)end

r 36 nm

1

I 36nm

1 (+)end

subunit is oriented toward the same end of the filament. The end of a

filament with an exposed binding cleft is the (-)end; the opposite end

is the ( +) end. (c) In the electron microscope, negatively stained actin

filaments appear as long, flexible, and twisted strands of beaded

subunits. Because of the twist, the filament appears alternately thinner

(7-nm diameter) and thicker (9-nm diameter) (arrows). (The microfila­

ments visualized in a cell by electron microscopy are F-act in filaments

plus any bound proteins.) [Part (a) adapted from C. E. Schutt et al., 1993,

Nature 365:810, courtesy of M. Rozycki. Part (c) courtesy of R. Craig.]

of actin filaments can be dcmomtrated by electron microscopy in "decoration" experiments, which exploit the ability of the motor protein myosin to bind specifically to actin filaments. In this type of experiment, an excess of myosin Sl, the actin­binding head domain of myosin, is mixed with actin fila­ments a nd binding is permitted to take place. Myosin attaches to the sides of a filament with a slight tilt. When all the actin subunits are bound by myosin, the filament appears "decorated" with arrowheads that all point toward one end of the fi lament (Figure 17-6).

The ability of the myosin S 1 head to bind and coat F­actin is very useful experimentally-it has allowed research­ers to identify the polarity of actin filaments, both in vitro

778 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

·.

·.·

and in cells. The arrowhead points to the (-) end, and so the (-)end is often called the "pointed" end of an actin filament; the ( +) end is known as the "barbed" end. Because myosin binds to actin filaments and not to microtubules or interme­diate filaments, arrowhead decoration is one criterion by which actin filaments can be definitively identified among the other cytoskeletal fibers in electron micrographs of cells.

KEY CONCEPTS of Section 17.1

Microfilaments and Actin Structures

Microfilaments can be assembled into diverse structures, many associated with the plasma membrane (see Figure 17-4a).

Actin, the basic building block of microfilaments, is a ma­jor protein of eukaryotic cells and is highly conserved.

• Actin can reversibly assemble into filaments that consist of two helices of actin subunits.

• The actin subunits in a filament are all oriented in the same direction, with the nucleotide-binding site exposed on the(-) end (see Figure 17-5).

17.2 Dynamics of Actin Filaments

The actin cytoskeleton is not a static, unchanging structure consisting of bundles and networks of filaments. Although microfilaments may be relatively static in some structures, in others they are highly dynamic, growing or shrinking in length. These changes in the organization of actin filaments can generate forces that cause large changes in the shape of a cell or drive i1,1tracellular movements. In this section, we consider the mechanism and regulation of actin polymeriza­tion, which is largely responsible for the dynamic nature of cells. We will see that several actin-binding proteins make important contributions to these processes.

Actin Polymerization in Vitro Proceeds in Three Steps The in vitro polymerization of G-actin monomers to form F-actin filaments can be monitored by viscometry, sedimen­tation, fluorescence spectroscopy, or fluorescence micros­copy (Chapter 9). When actin filaments grow long enough to become entangled, the viscosity of the solution increases, which is measured as a decrease in its flow rate in a viscom­eter. The basis of the sedimentation assay is the ability of ultracentrifugation ( 1 OO,OOOg for 30 minutes) to sediment F-actin but not G-actin. The third assay makes use of G-actin covalently labeled with a fluorescent dye; the fluorescence spectrum of the labeled G-actin monomer changes when it is polymerized into F-actin. Finally, growth of the fluorescently

labeled filaments can be imaged with fluorescence video mi­croscopy. These assays are useful for kinetic studies of actin polymerization and for characterization of actin-binding proteins to determine how they affect actin dynamics or how they cross-link actin filaments.

The mechanism of actin assembly has been studied ex­tensively. Remarkably, one can purify G-actin at a high pro­tein concentration without it forming filaments-provided it is maintained in a buffer with ATP and low levels of cat­ions. However, as we saw earlier, if the cation level is in­creased (e.g., to l 00 mM K and 2 mN1 Mg1

), G-actin will polymerize, with the kinetics of the reaction depending on the starting concentration of G-actin. The polymenzation of pure G-actin in vitro proceeds in three sequential phases (Figure 17-7a):

1. The nucleation phase is marked by a lag period 111 which G-actin subunits combine into two or three subunits. When the oligomer reaches three subunits in length, it can act as a seed, or nucleus, for the next phase.

2. During the elongation phase, the short oligomer rapidly increases in length by the addition of actin monomers to both of its ends. As F-actin filaments grow, the concentra­tion of G-actin monomers decreases until equilibrium is reached between the filament ends and mot:omers, and a steady state is reached.

3. In the steady-state phase, G-actin monomers exchange with subunits at the filament ends, but there is no net change in the total length of filaments.

The kinetic curves in Figure 17-7b, c show the state of fila­ment mass during each phase of polymerization. In Figure 17-7c we see that the lag period is due to nucleation, because it can be eliminated by the addition of a small number ofF­actin nuclei to the solution of G-actin.

How much G-actin is required for spontaneous filament assembly? Scientists have placed various concentrations of ATP-G-actin under polymerizing conditions and found that, below a certain concentration, filaments cannot assemble (Figure 17-8). Above this concentration, filaments begin to form, and when steady state is reached, the incorporation of more free subunits is balanced by the dissociation of sub­units from filament ends to yield a mixture of filaments and monomers. The concentration at which filaments are formed is known as the overall critical concentration, Cc. Below C, , filaments will not form; above C0 filaments form. At steady state, the concentration of monomeric actin remains at the critical concentration (see Figure 17-8).

Actin Filaments Grow Faster at ( +) Ends Than at (-) Ends We saw earlier that myosin Sl head decoration experiments reveal an inherent structural polarity ofF-actin (see Fig­ure 17-6). If free A TP-G-actin is added to a preexisting myosin­decorated filament, the two ends grow at very different rates

17.2 Dynamics of Actin Filaments 779

0 FOCUS ANIMATION: Actin Polymerization

(a) ~

a ~

Nucleus""- )

---+ a,...--.lo.W a

• Nucleation

(b)

!+- Nucleation ~I+- Elongation ~~~Steady state -?j

rn c: Q)

E ~ ;;:::

0 rn rn

"' ~

Time

FIGURE 17-7 The three phases of in vitro G-actin polymerization. (a) In the initial nucleation phase, ATP-G-actin monomers (red) slowly form stable complexes of actin (purple). These nuclei are rapidly elongated in the second phase by the addition of subunits to both ends of the filament. In the third phase, the ends of actin filaments are

(Figure 17-9). In fact, the rate of addition of ATP-G-actin is nearly 10 times faster at the ( +) end than at the (-) end. The rate of addition is, of course, determined by the concen­tration of free ATP-G-actin. Kinetic experiments have shown that the rate of addition at the ( +) end is about 12 J.LM 1 s 1

and about 1.3 J.LM 1 s 1 at the (-) end (Figure 17-lOa). This means that if 1 J.LM free ATP-G-actin is added to preformed filaments, on average 12 subunits will be added to the (+)end every second, wherea~ only 1.3 will be added at the (-) end every second. What about the rate of subunit loss from each end? By contrast, the rates of dissociation of ATP-G-actin subunits from the two ends are quite similar, about 1.4 s 1

FIGURE 17-8 Determination of filament formation by actin concent ration. The critical concentration (C,) is the concentration of G-actin monomers in equilibrium with actin filaments. At monomer

concentrations below the C" no polymerization takes place. When polymerization is induced at monomer concentrations above the C,, filaments assemble until steady state is reached and the monomer concentration falls to C,.

Elongation Steady state

(c)

I+- Elongation ~~---- Steady state----~

rn c: Q)

E ~ ;;::: -0 rn rn

"' ~

Time

in a steady state with monomeric G-actin. (b) Time course of the in vitro polymerization reaction reveals the initial lag period associated with nucleation, the elongation phase, and steady state. (c) If some short, stable actin filament fragments are added at the start of the reaction to act as nuclei, elongation proceeds immediately, without any lag period.

from the ( +) end and 0.8 s 1 from the (-) end. Since this dissociation is simply the rate at which subunits leave ends, it does not depend on the concentration of free A TP-G-actin.

What implications do these association and dissociation rates have for actin dynamics? first let's consider just one end, the ( +) end. As we noted above, the rate of addition

rn rn

"' ~

Tota l actin concentration (monomer and filaments)

780 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

EXPERIMENTAL FIGURE 7 .. 9 The two ends of a myosin­decorated actin filament grow unequally. When short actin filaments are decorated with a myosin S 1 head and then used to nucleate actin polymerization, the resulting actin subunits add much more efficiently to the (+ )end than the (- )end of the nucleating filament. This result indicates that G-actin monomers are added much faster at the (+)end than at the (- )end. [Courtesy ofT. Pollard.]

depends on the free A TP-G-actin concentration, whereas the rate of loss of subunits does not. Thus subunits will be added at high free ATP-G-actin concentrations, but as the concen­tration is lowered, a point will be reached at which the rate of addition is balanced by the rate of loss and no net growth occurs at that end. This is called the critica l concentration C ... , for the ( +) end, and we can calculate it by setting the rate of assembly equal to the rate of disassembly. Thus at the

(a) (-) end ADP-actin

c-c = o.so J.!M

(b) ,H

~-

ADP- G-actin ~~-

FIGURE 17-10 Act in treadmilling. ATP-actin subunits add faster at the ( ... )end than at the(-) end of an actin filament, resulting in a lower critical concentration and tread mill ing at steady state. (a) The rate of addition of ATP-G-actin is much faster at the(+ ) end than at the(-) end, whereas the rate of dissociation of ADP-G-actin is similar at the two ends. This difference results in a lower critical concentration at the

critical concentration, the rate of assembly is C ... , times the measured rate of addition of 12 1-1-M - 1 s 1 (C c 12 s 1

),

whereas the rate of disassembly is independent of the free actin concentration, namely, 1.4 s 1

• Setting these equal to each other yields C\ = 1.4 s 1/12 1-1-M- 1 s 1 or 0.12 11-M for the ( +) end. Above this free ATP-G-actin concentration, subunits add to the ( + ) end and net growth occurs, whereas below it, there is a net loss of subunits and shrinkage occurs.

Now let's consider just the (- ) end. Because the rate of addition is much lower, 1.3 11-M -I s 1

, yet the rate of disso­ciation is about the same, 0.8 s 1

, we expect the critical con­centration C c at the (-) end ro be higher than C\ . Indeed, as we just did for the ( + ) end, we can calculate C , to be about 0.8 s- 1/1.3 11-M 1 s- \ or 0.6 11-M. Thus at less than 0.6 11-M free ATP-G-actin, say, 0.31-1-M, the( -) end will lose subunits. But notice that at this concentration the ( +) end will grow, since 0.3 11-M is above C c· Because the critical concentrations are different, at steady state the free ATP-G­actin will be intermediate between c+ c and c-C) so the ( +) end will grow and the (-) end will lose subunits. This phe­nomenon is known as treadmilling, because particular sub­units, such those shown in blue in Figure 17-lOb, appear ro move through the fi lament.

The ability of actin filaments to treadmill is powered by hydrolysis of ATP. When ATP-G-actin binds to a (+) end, ATP is hydrolyzed to ADP and P,. The P, is slowly released from the subunits in the filament, so that the filament be­comes asymmetric, with A TP-actin subunits at the ( + ) end of the filament followed by a region with ADP-P,-actin and then, after P, release, ADP-actin subunits toward the (- )

ADP-P;-actin

(+)

ATP-actin ~

(+)end

12 J.!M 1 s 1

1.4 s 1

., C\ = 0.12 J.!M

(+)end. At steady state, ATP-actin is added preferentially at the ( + ) end, giving rise to a short region of the filament containing ATP-actin and regions containing ADP-P,-actin and ADP-actin toward the (-) end. (b) At steady state, ATP-G-actin subunits add preferentially to the (+ )end, while ADP-G-actin subunits disassemble from the (- )end, giving rise to tread milling of subunits.

17.2 Dynamics of Actin Fi laments 781

end (see Figure 17-IOa). During hydrolysis of ATP and subsequent release of P, from subunits in a filament, actin undergoes a conformational change that is responsible for the different association and dissociation rates at the two ends. Here we have considered only the kinetics of ATP-G­actin, but in reality it is ADP-G-actin that dissociates from the (-)end. Our analysis also relies on a plentiful supply of A TP-G-actin, which, as we will see, turns out to be the case in vivo. Thus actin can use the power generated by hydroly sis of ATP to treadmill, and treadmilling filaments can do work in vivo, as we will see later.

Actin Filament Treadmilling Is Accelerated by Profilin and Cofilin Measurements of the rate of actin trcadmilling in vivo show that it can be several times higher than can be achieved with pure actin in vitro under physiological conditions. Consis­tent with a treadmilling model, growth of actin filaments in vivo only ever occurs at the (+)end. How is enhanced tread­milling achieved, and how does the cell recharge the ADP­actin dissociating from the (-)end to ATP-actin for assembly at the ( +) end? Two different actin-binding proteins make important contributions to these processes.

The first is profilin, a small protein that binds G-actin on the side opposite to the nucleotide-binding cleft. When pro­filin binds ADP-actin, it opens the cleft and greatly enhances the loss of ADP, which is replaced by the more abundant cellular ATP, yielding a profilin-ATP-actin complex. This complex cannot bind to the (-) end because profilin blocks the sites on G-actin for (-) end assembly. However, the pro­fit in-A TP-actin complex can bind efficiently to the ( +) end, and profilin dissociates after the new actin subunit is bound (Figure 17-11 ). This function of profilin on its own does not enhance treadmilling rate, but it does provide a supply of ATP-actin from released ADP-actin; as a consequence, es­sentially all the free G-actin in a cell has bound ATP.

Profilin has another important property: it can bind other proteins with sequences rich in proline residues at the same time as binding actin. We will see later how this prop­erty is important in actin filament assembly.

Cofilin is also a small protein involved in actin treadmill­ing, bur it binds specifically to F-actin in which the subunits contain ADP, which are the older subunits in the filament toward the(-) end (see Figure 17-lOa). Cofilin binds by bridgtng two actin monomers and inducing a small change in the twist of the filament. This small twist destabilizes the filament, breaking it into short pieces. By breaking the fila­ment in this way, cofilin generates many more free (-) ends and therefore greatly enhances the disassembly of the (-) end of the filament (see Figure 17-11 ). The released ADP­actin subunits are then recharged by profilin and added to the(+) end as described above. In this way, profilin and cofilin can enhance treadmilling in vitro more than tenfold, up to levels seen in vivo. As might be anticipated, the cell

1-l end (+}end

Actin-ADP Actin-ADP-P;

;r: \ thymosin-134

\.. \ cycle

~~

ATP

~ ADP-actin

(I ATP-actin \:::1 Profilin 0 Cofilin J Thymosin-~4 FIGURE 1 7-1 1 Regulation of filament formation by actin-bi nding proteins. Actin-binding proteins regulate the rate of assembly and

disassembly as well as the availability of G-actin for polymerization. In the profilin cycle D. profilin binds ADP-G-actin and catalyzes the exchange of ADP for ATP. The ATP-G-actin-profilin complex can deliver actin to the ( +) end of a filament with dissociation and recycling of profilin. In the cofilin cycle f) , cofilin binds preferentially to filaments containing ADP-actin, inducing them to fragment and thus enhancing depolymerization by making more filament ends. In the thymosin-134 cycle 11. G-actin available from the actin-profilin equilib­rium is bound by thymosin-13"' sequestering it from polymerization. As the free G-actin concentration is lowered by polymerization, G-actin-thymosin-134 dissociates to make free G-actin available for association with profilin and further polymerization.

uses signal-transduction pathways to regulate both profilin and cofilin, and thereby the turnover of actin filaments.

Thymosin-(34 Provides a Reservoir of Actin for Polymerization It has long been known that cells often have a very large pool of unpolymerized acrin, sometimes as much as half the actin in the cell. Since cellular actin levels can be as high as 100-400 j.t.M, this means that there can be 50-200 j.t.M un­polymerized actin in cells. Since the critical concentration in vitro is about 0.2 j.t.M, why doesn't all this actin polymerize? The answer lies, at least in part, in the presence of actin

782 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

.·.

monomer sequestering proteins. One of these is thymosin­[34, a small protein that binds to ATP-G-actin in such a way that it inhibits addition of the actin subunit to either end of the filament. Thymosin-[34 can be very plentiful, for exam­ple, in human blood platelets. These discoid-shaped cell fragments are very abundant in the blood, and when they are activated during blood clotting, they undergo a burst of actin assembly. Platelets are rich in actin: they are estimated to have a total concentration of about 550 fLM ::~crin, of which about 220 fLM is in the unpolymerized form. They also contain about 500 fLM thymosin-[34 , which sequesters much of the free actin. However, as in any protein-protein interaction, free actin and free thymosin-[34 are in a dynamic equilibrium with the actin-rhymosin-[34. If some of the free actin is used up for polymerization, more actin-thymosin-[34 will dissociate, providing more free actin for polymerization (see Figure 17-11 ). Thus thymosin-[34 behaves as a buffer of unpolymerized actin for when it is needed.

Capping Proteins Block Assembly and Disassembly at Actin Filament Ends The treadmilling and dynamics of actin filaments are further regulated in cells by capping proteins that specifically bind to the ends of the filaments. If this were not the case, actin filaments would continue to grow and disassemble in an un­controlled manner. As one might expect, two classes of pro­teins have been discovered: ones that bind the ( +) end and ones that bind the (-)end (figure 17-12).

A protein known as CapZ, consisting of two closely re­lated subunits, binds with a very high affinity ( =0. 1 nM) to the (+)end of actin filaments, thereby inhibiting subunit ad­dition or loss. The concentration of CapZ in cells is generally sufficient to rapidly cap any newly formed ( +) ends. So how can filaments grow at their( +) ends? At least two mecha-

(-}end

Tropomodulin

FIGURE 17-12 Capping proteins. Capping protPins block assembly and disassembly at filament ends. CapZ blocks the (+}end, which is where filaments normally grow, so its function is to limit actin dynamics to the(- } end. The capping protein tropomodulin blocks(- }

ends, where filament disassembly normally occurs; thus the major function of tropomodulin is to stabilize filaments.

nisms regulate the activity of CapZ. First, the capping activ­ity of CapZ is inhibited by the regulatory lipid P1(4,5)P1,

found in the plasma membrane (Chapter 16). Second, recent work has shown that certain regulatory proteins are able to bind the ( +) end and simultaneously protect it from CapZ while still allowing assembly there. Thus cells have evolved an elaborate mechanism to block assembly of actin filaments at their (+)ends except when and where assembly is needed.

Another protein called tropomodulin binds to the ( ) end of actin filaments, also inhibiting assembly and disassembl y. This protein is found predominantly in cells in which actin filaments need to be highly stabilized. Two examples we will encounter later in this chapter are the short actin filaments m the cortex of the red blood cell and the actin filaments in muscle. As we will see, in both cases tropomodulin works with another protein, tropomyosin, which lies along the fila­ment to stabilize it. Tropomodulin binds to both tropomyo­sin and actin at the (-) end to greatly stabilize the filament.

In addition to CapZ, another class of proteins can cap the ( +) ends of actin filaments. These proteins also can sever actin filaments. One member of this family, gelsolin, is regu­lated by increased levels of Ca1

+ ions. On binding Ca1+, gel­

solin undergoes a conformational change that allows it to bind to the side of an actin filament and then insert itself between subunits of the helix, thereby breaking the filament. It then remains bound to and caps the ( +) end, generating a new (-) end that can disassemble. As we discuss in a later section, actin cross-linking proteins can provide linkages be­tween individual actin filaments to turn a solution ofF-actin into a gel. If gelsolin is added to such a gel, and the level of Ca2+ is elevated, gelsolin will sever the actin filaments and turn it back into a liquid solution. This ability to turn a gel into a sol is why the protein was named "gclsolin."

KEY CONCEPTS of Section 17.2

Dynamics of Actin Filaments

The rate-limiting step in actin assembly is the formation of a short actin oligomer (nucleus) that can then be elongated into filaments.

The critical concentration ( Cc) is the concentration of free G-actin at which the assembly onto a filament end is bal­anced by loss from that end.

When the concentration of G-actin is above the C0 the filament end will grow; when it is less than the C0 the fila­ment will shrink (see Figure 17-8).

• ATP-G-actin adds much faster at the ( +) end than at the (-)end, resulting in a lower critical concentration at the ( +) end than at the (-) end.

At steady state, actin subunits treadmill through a fila­ment. ATP-actin is added at the (+)end, ATP is then hydro­lyzed to ADP and P., P, is lost, and ADP-actin dissociates from the (-) end.

17.2 Dynamics of Actin Filaments 783

• The length and rate of turnover of actin filaments is regu­lated by specialized actin-binding proteins (see Figure 17-11 ). Profilin enhances the exchange of ADP for ATP on G-actin; cofilin enhances the rate of loss of ADP-actin from the fila­ment (-) end, and thymosin-P4 binds G-actin to provide re­serve actin when it is needed. Capping proteins bind to fila­ment ends, blocking assembly and disassembly.

17.3 Mechanisms of Actin Filament Assembly

The rate-limiting step of actin polymerization is the forma­tion of an initial actin nucleus from which a filament can grow (see Figure 17-7a) . In cells, this inherent property of actin is used as a control point to determine where actin fila­ments are assembled-this is how the different actin assem­blies within a single cell are generated (see Figures 17-1 and 17-4). Two major classes of actin nucleating proteins, the formin protein family and the Arp213 complex, nucleate actin assembly under the control of signal-transduction pathways. Moreover, they nucleate the assembly of different actin organizations: formins lead to the assembly of long actin filaments, whereas the Arp2/3 complex leads to branched networks. We will discuss each separately and see

how the power of actin polymerization can drive motile pro­cesses in a cell. We will then touch on the recent discoveries of new, specialized actin nucleating factors.

Form ins Assemble Unbranched Filaments Formins are found in essentially all eukaryotic cells as quite a diverse family of proteins: seven differenr classes are present in vertebrates. Although they are diverse, all formin family mem­bers have two adjacent domains in common, the so-called FHl and FH2 domains (formin-homology domains 1 and 2). Two FH2 domains from two individual monomers associate to form a doughnut-shaped complex (Figure 17-13a). This complex has the ability to nucleate actin assembly by binding two actin subunits, holding them so that the (+)end is toward the FH2 domains. The nascent filament can now grow at the ( +) end while the FH2 domain di~er remains attached. How is this possible? As we saw earlier, an actin filament can be thought of as two intertwined strands of subunits. The FH2 dimer can bind to the two terminal subunits. It then probably rocks between the two end subunits, letting go of one to allow addition of a new subunit and then binding the newly added subunit and freeing up space for the addit ion of another sub­unit to the other strand. In this way, rocking between the two subunits on the end, it can remain attached while simultane­ously allowing growth at the (+)end (see Figure 17-13a).

0 FOCUS ANIMATION: Elongation of Actin Filament by Formin FH2 Dimer

(a)

~ \

II -----+

Dimer of formin FH2 domain

(b)

FIGURE 17-13 Actin nucleation by the form in FH2 domain. (a) Formins have a domain called FH2 that can form a dimer and nucleate filament assembly. The dimer binds two actin subunits (step 0 ), and, by rocking back and forth (steps lfJ- EJ), can allow insertion of additional subunits between the FH2 domain and the (+)end of the growing filament. The FH2 domain protects the ( +)

(+) end

(- ) end

end from being capped by capping proteins. (b) The FH2 domain of a form in was labeled with colloidal gold (black dot) and used to nucleate assembly of an actin filament. The resulting filament was visualized by electron microscopy after staining with uranyl acetate. Formins assemble long unbranched filaments. [Part (b) from D. Pruyne et

al., 2002, Science 297:612.]

784 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

Exterior

Plasma membrane

Cytosol

Formin protein /

~A,;.,;., Profilin­ATP-actin

FIGURE 17-14 Regulation of form ins by an intramolecular interaction. Some of the form in classes found in vertebrates are

regulated by an intramolecular interaction. The inactive form in is activated by binding its Rho-binding domain (RBD) to membrane-bound

active Rho-GTP, resulting in exposure of the formin's FH2 domain, which

can then nucleate the assembly of a new filament. All form ins have an FH1 domain adjacent to the FH2 domain; the proline-rich FH1 domain is

a site for recruitment of profilin-ATP-G-actin complexes that can then be "fed" into the growing (+)end. For simplicity of representation, a single

form in protein is shown, but as shown in Figure 17-13, the FH2 domain functions as a dimer to nucleate actin assembly. Regulation of the Rho

family o f small GTPases is detailed in Figure 17-42.

The FHl domain adjacent to the FH2 domain a lso makes an important contribution to actin filament growth (Figure 17-14). This domain is rich in proline residues that are sites for the binding of several profilin molecules. We dis­cussed earl ier how profil in can exchange the ADP nucleotide on G-actin to generate profilin- ATP-actin. The FHl domain behaves as a landing site to increase the local concentration of profilin-A TP-G-actin complexes. The actin from the lo­calized profilin-actin complexes is fed into the FH2 domain to add actin to the ( +) end of the filament with the concomi­tant release of profit in, thereby allowing rapid FH2-mediated filament assembly (see Figure 17-14 ). Since the form in allows addit ion of actin subunits to the ( +) end, long fi laments with a formin at their (+)end are generated (figure 17-13b). In this manner, formin s nucleate actin assembly and have the remarkable ability to remain bound to the ( +) end while also a llowing rapid assembly there. To ensure the continued growth of the filament, formins bind to the ( +) end in such a way that precl udes binding of a (+)end capping protein such as CapZ, which would normally terminate assembly.

To be useful to a cell, formin activity has to be regulated. Many formins exist in a folded inactive conformation as a result of an interaction between the fi rst half of the protein and the C-terminal tail. These formins are activated by mem­brane-bound Rho -GTP, a Ras-rela ted small GTPase (d is­cussed in Section 17.7). When Rho is switched from the inactive Rho-GDP form into its activated Rho-GTP state, it can bind and activate the formin (see Figure 17-14).

Recent studies have shown that formins are responsible for the assembly of long actin filaments such as those found in stress fibers, filopodia, and in the contractile ring during cytoki­nesis (see Figure 17-4 ). The actin-nucleating role of formins was only discovered recently, so the roles performed by this diverse protein family are only now being uncovered. Since there are many different formin classes in animals, it is likely that formins will be found to assemble additional actin-based structures.

The Arp2/3 Complex Nucleates Branched Filament Assembly

The Arp2/3 complex is a protein machine consisting of se\en subunits, two of which are actin-related proteins (" Arp" ), explaining its name (Figure 17-15a). It is found in essentially all eukaryotes, including plants, yeasts, and animal cells. The Arp2/3 complex alone is a very poor nucleator. To nucleate the assembly of branched actin, Arp2/3 needs to be activated by interacting with a nucleation promoting factor (NPF), 10

addition to associating with the side of a preexisting actin filament. Although there are many different NPFs, the major family is characterized by the presence of a region called WCA (WH2, connector, acidic). Experiments have shown that if you add the WCA domain into an actin assembly assay together with preformed actin fi laments, Arp2/3 be­comes a potent nucleator of further actin assembly.

How do the Arp2/3 complex and NPF nucleate filaments? The NPF binds an actin subunit through its WH2 domain and activates the Arp2/3 complex through interaction with its acidic domain. Jn the inactive Arp2/3 complex, the two actin­rela ted polypeptides-Arp2 and Arp3-are in the wrong con­figuration to nucleate filament assembly (see Figure 17-15a ). When activated by the NPF, Arp2 and Arp3 move into the correct configuration, and the complex binds the side of pre­existing actin filament. The actin subunit brought in by the WH2 domain of the NPF binds to the Arp2/3 template to nucleate filament assembly at the(+) end (Figure 17-lSb). This new ( +) end then grows as long as A TP-G-actin is a va il­able or until it is capped by a ( +) end capping protein such as CapZ. The angle between the old filament and the new one is 70° (figure 17-lSc). This is also the angle observed experi­mentally in branched filaments at the leading edge of monic cells, which is believed to be formed by the action of the acti­vated Arp2/3 complex (Figure 17-15d). As we discuss in sub­sequent sections, the Arp2/3 complex can be used to drive actin polymerization to power intracellular motility.

Actin nucleation by the Arp2/3 complex is exquisitel y controlled, and the NPFs are part of that regulatory process. One NPF is called WASp, as it is defective in patients with Wiskott-Aidrich syndrome, an X-linkcd disease characterized by eczema, low platelet count, ::~nd immune deficiency. WASp exists in a fo lded inactive conformation, so that the WCA domain is not avai lable (Figure 17-16). One mechanism to activate the protein involves the small Ras-relatcd GTP-binding protein Cdc42 (discussed in Section 17. 7), which in the GTP­bound state binds to and opens WASp, making the WH2 actin-binding and acidic activation domains accessible.

17.3 Mechanisms of Actin Filament Assembly 785

~ VIDEO: Direct Observation of Actin Filament Branching Mediated by Activated Arp2/3

(b)

~ Actin

llh)- s-ub_\ __ n-it-~ ~ Part of nucleation promoting factor (NPF)

D ~ A rp 2/3 complex

FIGURE 17-15 Actin nucleation by the Arp2/3 complex. (a) X-ray structure of the Arp2/3 complex, with five of the subunits in gray, and the

Arp2 and Arp3 subunits in green and blue. (b) To nucleate actin assembly

efficiently, the act ivating part of an NPF is shown with its W (WH2),

C (connector), and A (acidic) domains. An actin subunit b inds to the

W domain (step 0 ); and then the A domain binds the Arp2/3 complex (step fJ). This interaction induces a conformational change in the Arp2/3

complex, and after binding to the side of an actin filament, the actin subunit bound to theW domain binds to the Arp2/3 complex (step i}),

Although form ins and the Arp2/3 complex are found in fungi, plants, and animals, additional actin nucleators have re­cently been discovered in animal cells. One of these, called Spire, has four tandem WH2 domains, so it can bind four actin mono­mers. It does this in such a manner to allow the actins to as­semble into a filament, although the exact mechanism remains to be understood. As actin fi laments perform so many functions in cells, it is likely that additional nucleators will be discovered.

FIGURE 17-16 Regulation of the Arp2/3 complex by WASp. WASp is inactive due to an Intramolecular interaction that masks the WCA domain. On binding t he membrane-bound active small G protein Cdc42-GTP (a member of the Rho family) through its

Rho-binding domain (RBD), the intramolecular interaction in WASp

is relieved, exposing theW domain to bind actin and the acidic A domain for activation of the Arp2/3 complex. Regulation of the Rho family of small GTPases is detai led in Figure 17-42.

(c)

!+lend

l-l end

(d )

w hich then initiates the assembly of an actin fi lament at the available ( +)

end (step [)). The Arp2/3 branch makes a characteristic 70 angle

between the filaments. (c) Averaged image compiled from several electron micrographs of Arp2/3 at an actin branch. (d) Image of actin

filaments in the leading edge, with a magnification and coloring of individual branched filaments. [Part (a) PDB ID 2P9L; part (c) from C. Egile

et al., 2006, PLoS Btol. 3:e383; part (d) from T. M. Svitkina and G. G. Borisy, 1999, J. Cell Bioi. 145:1 009.)

Exterior

Plasma membrane

Cytosol

WASp

~ation (+)end

(- )end

786 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

',

.·

Intracellular Movements Can Be Powered by Actin Polymerization How can actin polymerization be harnessed to do work? As we have seen, actin polymerization involves the hydrolysis of actin-ATP to actin-ADP, which allows actin to grow prefer­entially at the ( + ) end and disassemble at the (- ) end. If an actin filament were to become fixed in the meshwork of the cytoskeleton and you could bind and ride on the assembling ( +) end, you would be transported across the cell. This is just what the intracellular bacterial parasite Listeria monocyto­genes does to get around the cell. The study of Listeria motility

was, in fact, the way the nucleating activity of the Arp2/3 pro­tein was discovered. As we shall see shortly, Listeria has hi­jacked a normal cell motility process for its own purposes; we discuss Listeria first, as it is currently much better understood than the normal processes that employ similar mechanisms.

Listeria is a food-borne pathogen that causes mild gas­trointestinal symptoms in most adults but can be fatal to elderly or immunocompromised individuals. It enters animal cells and divides in the cytoplasm. To move from one host cell to an­other, it moves around the cell by polymerizing actin into a comet taillike the plume behind a rocket (Figure 17-17a, b), and when it runs into the plasma membrane, it pushes its way

Q) VIDEO: In Vivo Assembly of Actin Tails in Listeria-infected Bacteria

(a) (c)

(b)

~ • ~ • • • • •

& •

• • ~

Actin assembly (+ )end

In vitro reaction, • • Actin • ~ • or within host cell disassembly cytoplasm

EXPERIMENTAL FIGURE 17-17 Listeria utilizes the power of actin polymerization for intracellular movement. (a) Fluorescence

microscopy of a cu ltured cell stained with an antibody to a bacterial

surface protein (red) and fluorescent phalloidin to localize F-actin (green). Behind each Listeria bacterium is an actin "comet tail" that

propels the bacterium forward by actin polymerization. When the bacterium runs into the plasma membrane, it pushes the membrane out into a structure like a filopodium, which protrudes into a neighbor­

ing cell. (b) Listeria motility can be reconstituted in vitro with bacteria

and just four proteins: ATP-G-actin, Arp2/ 3 complex, CapZ, and cofilin.

(-)end • • • This phase micrograph shows bacteria (black), behind which are the

phase-dense actin tails. (c) A model of how Listeria moves using just

four proteins. The ActA protein on the cell surface activates the Arp2/3

complex to nucleate new filament assembly from preexisting filaments. Filaments grow at their (+)end until capped by CapZ. Actin is recycled

through the action of cofilin, which enhances depolymerization at the

(-) end of the filaments. In this way, polymerization is confined to the back of the bacterium, which propels it forward. [Part (a) courtesy of

J. Theriot and T. Michison; part (b) from T. P. Loisel et al., 1999, Nature 401 :613.]

17.3 Mechanisms of Actin Filament Assembly 787

into the adjacent cell to infect it. How does it recruit the host cell actin to propel itself? Listeria has on its surface a protein called ActA that mimics an NPF by having an actin-binding site and an acidic region to activate the Arp2/3 complex (Fig­ure 17-17c). The ActA protein also binds a protein known as VASP, which has three important properties. First, VASP has a proline-rich region that can bind profilin-ATP-actin for enhancing ATP-actin assembly into the newly formed barbed ends generated by the Arp2/3 complex. Second, it can hold on to the end of the newly formed filament. Third, it can protect the ( +) end of the growing filament from cap­ping by CapZ. These properties allow VASP to enhance actin assembly and confine it to the rear of the bacterium. The assembling filaments then push on the bacterium. Since the filaments are embedded in the stationary cytoskeletal matrix of the cell, the Listeria cell is pushed forward, ahead of the polymerizing actin. Researchers have reconstituted Listeria motility in the test tube using purified proteins to see what the minimal requirements for Listeria motility are. Re­markably, the bacteria will move when just four proteins are added: ATP-G-actin, the Arp2/3 complex, CapZ, and cofilin (see Figure 17-17b, c). We have discussed the role of actin and Arp2/3, but why are CapZ and cofilin needed? As we have seen earlier, CapZ rapidly caps the free (+)end of actin filaments, so when a growing filament no longer contributes to bacterial movement, it is rapidly capped and inhibited from further elongation. In this way, assembly occurs only adjacent to the bacterium where ActA is stimulating the Arp2/3 complex. Cofilin is necessary to accelerate the disas­sembly of the (-) end of the actin filament, regenerating

(a) Endocytic site

initiation Invagination

Release and movement

into cell

~~------+-------,_-------r-------+~Time 0 15 20 25 30 (seconds)

End~s~ assembly factors

Nucleation promoting factors (e.g. WASp)

Arp2/3· dependent ~._,,. assembly of actin filaments

cargo

\ Plasma

membrane

FIGURE 17-18 Arp2/3-dependent actin assembly during endocytosis. (a) Clathrin-mediated endocytosis is a rapid and ordered

process. It has been best studied in yeast, where the temporal order of specific steps has been delineated. In vivo imaging has shown that endocytosis assembly factors recruit nucleation-promoting factors

that activate the Arp2/3 complex. A burst of Arp2/3-dependent actin assembly drives internalized endocytic vesicles away from the

free actin to keep the polymerization cycle going (see Figure 17-11). This minimal rate of motility can be increased by the presence of other proteins, such as VASP and profilin, as mentioned above.

To move inside cells, the Listeria bacterium, as well as other opportunistic pathogens such as the Shigella species that cause dysentery, hijack s a normal, regulated cellular process involved in cell locomotion. As we discuss in more detail later (Section 17.7), moving cells have a thm sheet of cytoplasm that protrudes from the front of the cell called the leading edge (see Figures 17-lc, 17-4, and 17-15d). This thin sheet of cytoplasm consists of a dense meshwork of actin filaments that are continually elongating at the front of the cell to push the membrane forward. Factors in the leading­edge membrane activate the Arp2/3 complex to nucleate these filaments. Thus the power of actin assembly pushes the membrane forward to contribute to cell locomotion.

Microfilaments Function in Endocytosis

As we saw in Chapter 14, endocytosis describes the processes that cells use to take up particles, molecules, or fluid from the external medium by enclosing them in plasma membrane and then internalizing them. The uptake of molecules or liquid is called receptor-mediated or fluid-phase endocytosis, and the uptake of large particles is called phagocytosis ("cell eating"). Microfilaments participate in both of these processes.

Fluid-phase endocytosis is a very highly organized process, and recent studies have shown that the power of actin assembly contributes to this mechanism. Endocytosis assembly factors

(b)

plasma membrane, just like the movement of Listeria. (b) Endosome movement can be reconstituted in vitro. Endosomes isolated from cells

that had taken up fluorescently labeled transferrin (red) were added to a cell extract containing fluorescently labeled actin (green). The

endosomes bind WASp, which then activates the Arp2/3 complex to assemble actin tails that propel them through the cytoplasm.

[Part (b) from Taunton et al., 2000, J. Cell Bioi. 148:5 19].

788 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

Antibody to component on bacterial surface

aj Phagosome

Surface protein

Opsonized bacterium

Fe receptors

Leukocyte

~O Lysosomes

"0

FIGURE 17-19 Phagocytosis and actin dynamics. Actin assembly and contraction drives the internalization of phagocytic particles. Shown here is the phagocytosis and degradation of a bacterium by a leukocyte. An invading bacterium is coated by specific antibodies to a cell-surface protein in a process known as opsonization (step 0 ).

The Fe region ofthe bound antibodies is displayed on the bacterial surface and recognized by a specific receptor, the Fe receptor, on the leukocyte surface (step f)). This interaction signals the cell to assemble a contractile actin structure that results in the internalization and engulfment of the bacterium (step IJ). Once it has been internalized into a phagosome, the bacterium is killed and degraded by enzymes delivered from lysosomes (step 19).

recruit NPfs so that as the endocytic vesicles invaginate and pinch off from the membrane, they are then driven into the cytoplasm, powered by a rapid and very short-lived burst (a few seconds in duration) of actin polymerization driven by the Arp2/3 complex (Figure 17-lSa). This actin-based movement of endocytic vesicles involving the Arp2/3 complex can be re­constituted in vitro (Figure 17-lSb) and is mechanistically very similar to leading-edge formation and Listeria motility.

Phagocytosis is a viral process in the recognition and re­moval of pathogens, such as bacteria, by white blood cells. The immune system identifies the bacterium as foreign mate­rial and makes antibodies that recognize components on its surface. As we discussed in Chapter 3, each antibody has a region called the Fab domain that binds specifically to its an­tigen, in this case a component on the bacterial cell surface. As antibodies coat the bacterium through interaction be­tween their tab domains and the cell-surface antigen, a sec­ond antibody domain known as the Fe domain is exposed. This process is known as opsonization (Figure 17-19, step 0 ). The white blood cells have a receptor on their cell surface, the Fe receptor, that recognizes the antibodies on the bacterium; this interaction signals the cells to bind and engulf the patho­gen (steps 6 and 10). The signal also tells the cell to assemble microfilaments at the interaction site with the bacterium, and the assembled microfilaments, together with myosin motor proteins, provide the force necessary to draw the bacterium into the cell, ultimately fully enclosing the pathogen m plasma membrane (step 19). Once internalized, the newly formed phagosome fuses with lysosomes, where the pathogen is killed and degraded by lysosomal enzymes.

Toxins That Perturb the Pool of Actin Monomers Are Useful for Studying Actin Dynamics Certain fungi and sponges have developed toxins that target the polymerization cycle of actin and are therefore roxie to animal cells. Two types of toxins have been characterized. The first class is represented by two unrelated roxins, cyrocha­lasin D and larrunculin, which promote the depolymerizarion of filaments, though by different mechanisms. Cytochalasin D, a fungal alkaloid, depolymerizes actin filaments by bind­ing to the ( +) end ofF-actin, where it blocks further addition of subunits. Latrunculin, a toxin secreted by sponges, binds and sequesters G-actin, inhibiting it from adding to a filament end. Exposure ro either toxin thus increases the monomer

17.3 Mechanisms of Actin Filament Assembly 789

pool. When cytochalasin D or latrunculin is added to live cells, the actin cytoskeleton disassembles and cell movements such as locomotion and cytokinesis are inhibited. These ob­servations were among the first to implicate actin filaments in cell motility. Latrunculin is especially useful because it binds actin monomers and prevents any new actin assembly. Thus if you add latrunculin to a cell, the rate at which actin-based structures disappear reflects their normal rate of turnover. This has revealed that some structures have half-lives of lc~~ than a minute, whereas others are much more stable. For ex­ample, experiments with latrunculin show that the leading edge of motile cells turns over every 30-180 seconds, and stress fibers turn over every 5-10 minutes.

In contrast, the monomer-polymer equilibrium is shifted in the direction of filaments by jasplakinolide, another sponge toxin, and by phalloidin, which is isolated from Amanita phal­loides (the "angel of death" mushroom). jasplakinolide en­hances nucleation by binding and stabilizing actin dimers and thereby lowering the critical concentration. Phalloidin binds at the interface between subunits in F-actin, locking adjacent sub­units together and preventing actin filaments from depolymer­izing. Even when actin is diluted below its critical concentration, phalloidin-stabilized filaments will not depolymerize. Because many actin-based processes depend on actin filament turnover, the introduction of phalloidin into a cell paralyzes all these systems and the cell dies. However, phalloidin has been very useful to researchers, as fluorescent-labeled phalloidin, which binds only to F-actin, is commonly used to stain actin filaments for light microscopy (see Figure 17-4).

KEY CONCEPTS of Section 17.3

Mechanisms of Actin Filament Assembly

Actin assembly is nucleated by two classes of proteins: formins nucleate the assembly of unbranched filaments (see figure 17-13), whereas the Arp2/3 complex nucleates the as­sembly of branched actin networks (see Figure 17-15). The activities of formins and Arp2/3 are regulated by signal­transduction pathways.

Functionally different actin-based structures are assem­bled by formins and Arp2/3 nucleators. Formins drive the assembly of stress fibers and the contractile ring, whereas Arp2/3 nucleates the assembly of branched actin filaments found in the leading edge of motile cells.

The power of actin polymerization can be harnessed to do work, as is seen in the Arp2/3-dependent intracellular move­ment of pathogenic bacteria (see Figure 17-17) and inward movement of endocytic vesicles (see Figures 17-18 and 17-19).

Several toxins affect the dynamics of actin polymerization; some, such as latrunculin, bind and sequester actin mono­mers, whereas others, such as phalloidin, stabilize filamen­tous actin. Fluorescently labeled phalloidin is useful for staining actin filaments.

17.4 Organization of Actin-Based Cellular Structures

We have seen that actin filaments are assembled into a wide variety of different arrangements and how many associated proteins nucleate actin assembly and regulate filament turn­over. Dozens of proteins in a vertebrate cell organize these filaments into diverse functional structures. Here we discuss just a few of these proteins, giving examples of typical types of actin cross-linking proteins found in cells, and also discuss the proteins involved in making functional links between actin and membrane proteins. One fascinating problem, about which very little is known, is how cells assemble dif­ferent actin-based structures within the same cytoplasm of a cell. Some of this organization must be due to local regula­tion, a topic we come to at the end of the chapter.

I

Cross-Linking Proteins Organize Actin Filaments into Bundles or Networks

When one assembles actin filaments in a test tube, they form a tangled network. In cells, however, actin filaments are found in a variety of distinct structures, such as the highly ordered filament bundles in microvilli or the meshwork char­acteristic of the leading edge (sec Figure 17-4a). These differ­ent organizations are determined by the presence of actin cross-linking proteins. To be able to organize actin, an actin cross-linking protein must have two F-actin-binding sites (Figure 17-20a).

Cross-linking ofF-actin can be achieved by having two actin-binding sites within a single polypeptide, as with fim­hrin, a protein found in microvilli that builds bundles of filaments all having the same polarity (Figure 17-20b). Other actin cross-linking proteins have a single actin-bind­ing site in a polypeptide chain and then two chains associ­ate to form dimers that bring together two actin-binding sites. These dimeric cross-linking proteins can assemble to

generate a rigid rod connecting the two binding sites, as happens with o.-actinin. Like fimbrin, o.-actinin also bun­dles parallel actin filaments, but farther apart than fimbrin. Another protein, called spectrin, is a tetramer with two actin-binding sites; spectrin spans an even greater distance between actin filaments and makes networks under the plasma membrane (shown in Figure 17-21 and discussed in the next section). Other types of cross-linking proteins, such as filamin, have a highly flexible region between the two binding sites, functioning like a molecular leaf spring, so they can make stabilizing cross-links between filaments in a meshwork (Figure 17-20c), as is found in the leading edge of motile cells. The Arp2/3 complex, which we dis­cussed in terms of its ability to nucleate actin filament as­sembly, is also an important cross-linking protein, attaching the (-)end of one filament to the side of another filament (see Figure 17-15).

790 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

(a)

Fimbrin

a-actinin

Spectrin

0 : I ~ill -!! !!!~ -

Filamin

Dystrophin Plasma membrane

~ FIGURE 17-20 Actin cross-)inking prote ins. Actin cross-linking

proteins mold F-actin filaments into diverse structures. (a) Examples of four F-actin cross-linking proteins, all of which have two domains (blue) that bind F-actin. Some have a Ca2+ -binding site (purple) that inhibits

their activity at high levels of free Ca2+. Also shown is dystrophin, which

has an actin-binding site on its N-terminal end and a (-terminal domain

that binds the membrane protein dystroglycan. (b) Transmission electron

Adaptor Proteins Link Actin Filaments to Membranes

To contribute to the structure of cells and also to harness the power of actin polymerization, actin filaments are very often attached to membranes or are associated with intracellular structures. Actin filaments are especially abundant in the cell

Location:

Microvilli, filopodia, focal adhesions

Stress fibers. filopodia, muscle Z line

Cell cortex

Leading edge, stress fibers, filopodia

Linking membrane proteins to actin cortex in muscle

(b)

(c)

micrograph of a thin section of a stereocilium (an unfortunate name, since it is really a giant microvillus) on a sensory hair cell in the inner ear. This

structure contains a bundle of actin filaments cross-linked by fimbrin, a small cross-linking protein that allows for close and regular interaction of

actin filaments. (c) Long cross-linking proteins such as fila min are flexible

and can thus cross-link actin filaments into a loose network. (Part (b) from L. G. Tilney, 1983, J. Cell Bioi. 96:822; part (c) courtesy of J. Hartwig.]

cortex underlying the plasma membrane, to which they give support. Actin filaments can interact with membranes either laterally or at their ends.

Our first example of actin filaments attached to mem­branes is the human erythrocyte-the red blood cell. The erythrocyte consists essentially of plasma membrane enclosing

17.4 Organization of Actin-Based Cellular Structures 791

(a) (b ) (c)

(d)

: ~ (-) ends (+ ) ends

; ) Actin filaments

FIGURE 17-21 Lateral attachment of microfilaments to membranes. (a) Electron micrograph of t he erythrocyte membrane showing the spoke-and-hub organization of the cortical cytoskeleton

supporting the plas·ma membrane in human erythrocytes. The long

spokes are composed mainly of spectrin and can be seen to intersect at the hubs, or membrane-attachment sites. The darker spots along

the spokes are ankyrin molecules, which cross-link spect rin to integral membrane proteins. (b) Diagram of the erythrocyte cytoskeleton,

showing the two main types of membrane attachment: D ankyrin

and f) band 4.1. (c) Actin is incorporated into the tip of stereocilia (giant microvilli). Cells w ith stereocilia were transfected to express

GFP-actin for a short period of time and then counterstained with

a high concentration of the protein hemoglobin to transport oxygen from the lungs to tissues and carbon dioxide from tis­sues back to the lung-all powered by the magnificent muscle known as the heart. Erythrocytes must be able to survive the raging torrents of blood flow in the heart, then flow down arteries and survive squeezing through narrow capillaries be­fore being cycled through the lungs via the heart. To survive this grueling process for thousands of cycles, erythrocytes have a microfilament-based network underlying their plasma mem­brane that gives them both the tensile st rength and the flexibil­ity necessary for their journey. This network is based on short actin filaments of about 14 subunits in length, stabilized on

rhodamine-phalloidin to stain all the F-actin. The experiment shows

that new actin is incorporated at the tips of the stereocilia. (d) Ezrin, a

member of the ezrin-radixin-moesin (ERM) family, links actin filaments laterally to the plasma membrane in surface structures such as

microvi lli; attachment can be direct or indirect. Ezrin, activated by phosphorylation (P), links directly to the cytoplasmic region of

transmembrane proteins (right) or indirectly through a scaffolding protein such as EBPSO (left). [Part (a) from T. J. Byers and D. Branton, 1985,

Proc. Nat'/ Acad. Sci. USA 82:6153, courtesy of D. Branton; part (b) adapted from

S. E. Lux, 1979, Nature 281:426, and E. J. Luna and A. L. Hitt, 1992, Science

258:955; part (c) from A. K. Rzadzinska et al., 2004, J. Cell Bioi. 1 64:887;

(d) adapted from R. G. Fehon et al., 2010, Nature Rev. Mol. Cei/Bio/. 1 1 :276.)

their sides by tropomyosin (discussed in more detail in Section 17.6) and by the capping protein tropomodulin on the (-) end. These short filaments serve as hubs for binding about six flexible spectrin molecules, generating a fishing-net type of structure (Figure 1 7-21a). This network gives the erythrocyte both its strength and flexibility. Spectrin is attached to mem­brane proteins through two mechanisms: through a protein called ankyrin to the bicarbonate transporter (a transmem­brane protein also known as band 3) and t hrough a spectrin and F-actin-binding protein called band 4.1 to another trans­membrane protein called glycophorin C (Figure 17-21b). Al­though this spectrin-based network is highly developed in the

792 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

·.

.•

erythrocyte, similar types of linkages occur in many cell types. For example, a related type of ankyrin-spectrin attachment links the Na + IK-'- ATPase to the actin cytoskeleton on the basolateral membrane of epithelial cells.

1t1f.1 Genetic defects in proteins of the red blood cell cyto­H skeleton can result in cells that rupture easily, giving rise to diseases known as hereditary spherocytic anemias (spheroC)'tic been usc the cells arc rounder, anemias bet:au:-e there is a shortage of red blood cells) and hence a shorter life span. In human patients, mutations in spectrin, band 4.1, and ankyrin can cause this disease. •

In addition to the spectrin-based type of support in the cell cortex, microfilaments provide the support for cell-surface structures such as microvilli and membrane ruffles. If we look at a microvillus, it is clear that it must have an end-on attach­ment at the tip and lateral attachments down its length. What is the orientation of actin filaments in microvilli? Decoration of microvillar filaments by the Sl fragment of myosin show that it is the ( +) end at the tip. Moreover, when fluorescent actin is added to a cell, it is incorporated at the tip of a micro­villus, showing that not only is the ( +) end there, actin fila­ment assembly occurs there (Figure 17-21c). At present it is not known how actin filaments are attached at the microvil­lus tip, but a likely candidate is a fonnin protein. This ( +) end orientation of actin filaments with respect to the plasma membrane is found almost universally-not just in microvilli but also, for example, in the leading edge of motile cells. The lateral attachments to the plasma membrane are provided, at least in part, by the ERM (ezrin-radixin-moesin) family of proteins. These are regulated proteins that exist in a folded, inactive form. When activated by phosphorylation in re­sponse to an external signal, F-actin and membrane-protein­binding sites of the ERM protein are exposed to provide a lateral linkage t.o actin filaments (Figure 17-2ld). At the plasma membrane, ERM proteins can link the actin filaments directly or indirectly through scaffolding proteins to the cyto­plasmic domain of membrane proteins.

The types of actin membrane linkages we have discussed so far do not involve areas of the plasma membrane attached directly to other cells in a tissue or to the extracellular ma­trix. Contact between epithelial cells is mediated by highly specialized regions of the plasma membrane called adherens ;unctions (see Figure 17-1b). Other specialized regions of as­sociation called focal adhesions mediate attachment of cells to the extracellular matrix. fn turn, these specialized types of attachments connect to the cytoskeleton, as will be described in more detail when we discuss cell migration (Section 17.7) and cells in the context of tissues (Chapter 20).

.. Muscular dystrophies are genetic diseases that are ' often characterized by the progressive weakening of

skeleta l muscle. One of these genetic diseases, Duchenne muscular dystrophy, affects the protein dystrophin, whose gene is located on the X chromosome, and so the disease is much more prevalent in males. Dystrophin is a modular protein

whose function is to link the cortical actin network of mus­cle cells to a complex of membrane proteins that link to the extrace llular matrix. Thus dystrophin has an N-terminal actin-binding domain, followed by a series of spectrinlike repeats and terminating in a domain that binds the trans­membrane dystroglycan complex to the extracellular matrix protein laminin (see Figure 17-20a). In the absence of dys­trophin, the plasma membrane of muscle cells becomes weakened by cycles of muscle contraction and eventually ruptures, resulting in death of the muscle myofibril. •

KEY CONCEPTS of Section 17.4

Organization of Actin-Based Cellular Structures

• Actin filaments are organized by cross-linking proteins that have two F-actin-binding sites. Actin cross-linking pro­teins can be long or short, rigid or flexible, depending on the type of structure involved (see Figure 17-20).

• Actin filaments are attached laterally to the plasma mem­brane by specific classes of proteins, as seen in the red blood cell or in cell-surface structures such as microvilli (see Figure 17-21).

• The ( +) end of actin filaments can also· be attached to

membranes, with assembly mediated between the filament end and the membrane.

• Several diseases have been traced to defects in the microfilament-based cortical cytoskeleton that underlies the plasma membrane.

17.5 Myosins: Actin-Based Motor Proteins

In Section 17.3 we discussed how actin polymerization nu­cleated by the Arp2/3 complex can be harnessed to do work, such as in the movement of vesicles during endocytosis, at the leading edge of motile cells, and the propulsion of Liste­ria bacterium across the eukaryotic cell. In addition to actin­polymerization-based motility, cells have a large family of motor proteins called myosins that can move along actin filaments. The first myosin discovered, myosin II, was isolated from skeletal muscle. For a long time, biologists thought that this was the only type of myosin found in nature. However, they then discovered other types of myosins and began to ask how many different functional classes might exist. Today we know that there are several different classes of myosins, in addition to the myosin II of skeletal muscle, that move along actin. Indeed, with the discovery anc.l analysis of all these actin-based motors and the corresponding microtu­bule-based motors described in the next chapter, the former relatively static view of a cell has been replaced wtth the re­alization that the cytoplasm is incredibly dynamic-more like an organized but busy freeway system with motors busily ferrying components around.

17.5 Myosins: Actin-Based Motor Proteins 793

Myosins have the amazing ability to convert the energy released by A TP hydrolysis into mechanical work (move­ment along actin). All myosins convert ATP hydrolysis into work, yet different myosins can perform very different types of functions. For example, many molecules of myosin II pull together on actin filaments to bring about muscle contrac­tion, whereas myosin V binds to vesicular cargo to transport it along actin filaments. The other classes of myosin provide a myriad of functions, from moving organelles around cells to contributing to cell migration.

To begin to understand myosins, we first discuss their general domain organization. Armed with this information, we explore the diversity of myosins in different organisms and describe in more detail some of those that are common in eukaryotes. To understand how such diverse functions

(a)

can be accommodated by one type of motor mechanism, we will investigate the basic mechanism of how the energy re­leased by ATP hydrolysis is converted into work and then see how this mechanism is modified to tailor the properties of specific myosin classes for their specific functions.

Myosins Have Head, Neck, and Tail Domains with Distinct Functions

Much of what we know about myosins comes from studies of myosin II isolated from skeletal muscle. In skeletal mus­cle, hundreds of individual myosin II molecules are assem­bled into bundles called bipolar thick filaments (Figure 17-22a). In a later section we will discuss how these myosin filaments interdigitate wi th actin filaments to bring about

«--------- 160 nm ------~ +-----------------325nm--------------~

(b) Myosin II (c) Head and neck domain

Head Neck Tail

~Chymotrypsin cleavage

Heavy chain

LMM

! Papain cleavage

S2

FIGURE 17-22 Structure of myosin 11. (a) Organization of myosin II in filaments isolated from skeletal muscle. Myosin II assembles into bipolar filaments in which the tails form the shaft of the filament with heads exposed at the ends. Extraction of bipolar filaments with high salt and ATP disassembles the filament into individual myosin II molecules. (b) Myosin II molecules consist of two identical heavy chains (light blue) and four light chains (green and dark blue). The tail of the heavy chains forms a coiled coil to dimerize; the neck region of each heavy chain has two light chains associated with it. Limited proteolytic

cleavage of myosin II generates tai l fragments- LMM and 52-and the 51 motor domain. (c) Three-dimensional model of a single 51 head domain shows that it has a curved, elongated shape and is bisected by a cleft. The nucleotide-binding pocket lies on one side of this cleft, and the actin-binding site lies on the other side near the tip of the head. Wrapped around the shaft of the a-helical neck are two light chains. These chains stiffen the neck so that it can act as a lever arm for the head. Shown here is the ADP-bound conformation.

794 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

muscle contraction. Here, we first investigate the properties of the individual myosin molecule itself.

It is possible to dissolve the myosin thick filament in a solution of ATP and high salt, generating a pool of individ­ual myosin II molecules. The soluble myosin II molecule is actually a protein complex consisting of six polypeptides. Two of the subunits are identical high-molecular-weight polypeptides known as myosin heavy chains . Each consists of a globular head domain and a long tail domain, connected by a flexible neck domain. The tails of the two myosin heavy chains intertwine, so that the head regions are in close prox­imity. The remaining four subunits of the myosin complex are smaller in size and are known as the light chains. There are two types of light chain, the essential light chain and the regulatory light chain. One light chain of each type associates with the neck region of each heavy chain (Figure 17 -22b, top) . The myosin heavy chain and the two types of light chains are encoded by three different genes.

The soluble myosin II molecule has A TPase activit), re- · fleeting its ability to power movements by hydrolysis of A TP. But which part of the myosin complex is responsible for this activity? To identify functional domains in a protein, a stan­dard approach is to cleave the protein into fragments with specific proteases and then ask which fragments have the ac­tivity. Soluble myosin II can be cleaved by gentle treatment with the protease chymotrypsin to yield two fragments, one called heavy meromyosin (HMM; mero means "part of") and the other, light meromyosin (LMM) (Figure 17-22b, middle). The heavy meromyosin can be further cleaved by the protease papain to yield subfragment 1 (51) and subfragment 2 (52) (Figure 17-22b, bottom ). By analyzing the properties of the various fragments-51, 52, and LMM-it was found that the intrinsic ATPase activity of myosin resides in the Sl fragment, as does its F-actin-binding site. Moreover, it was

found that the ATPase activity of the Sl fragment was greatly enhanced by the presence of filamentous actin, so it is said to have an actin-activated A TPase activity, which is a hallmark of all myosins. The Sl fragment of myosin II consists of the head and neck domains with associated light chains, whereas the 52 and LMM regions make up the tail domain.

X-ray crystallographic analysis of the head and neck do­mains revealed its shape, the positions of the light chains, and the locations uf the ATP-binding and actin-binding sttes (hg­ure 17-22c). At the base of the myosin head is the a-helical neck, where two light-chain molecules wrap around the neck like C-clamps. In this position, the light chains stiffen the neck region. The actin-binding site is an exposed region at the tip of the head domain; the ATP-binding site is also in the head domain, within a cleft opposite the actin-binding sire.

How much of myosin II is necessary and sufficient for "motor" activity? To answer this question, one needs a simple in vitro motilitT assay. In one such assay, the sliding-filament assay, myosin molecules are tethered to a coverslip to which is added stabilized, fluorescently labeled actin filaments. Be­cause the myosin molecules are tethered, they cannot slide; thus any force generated by interaction of myosin heads with actin filaments forces the filaments to move relative to the myosin (Figure 17-23a). If ATP is present, added actin fila­ments can be seen to glide along the surface of the coverslip; if A TP is absent, no filament movement is observed. Using this assay, one can show that the Sl head of myosin II is suf­ficient to bring about movement of actin filaments. This movement is caused by the tethered myosin Sl fragments (bound to the coverslip) trying to "move" toward the (+) end of a filament; thus the filaments move with the (-) end leading. The rate at which myosin moves an actin filament can be determined from video recordings of sliding-filament assays (Figure 17-23b).

9 TECHNIQUE ANIMATION: In Vitro Motility Myosin Assay

EXPERIMENTAL FIGURE 17 ·23 Sliding-filament assay is used to detect myosin-powered movement. (a) After myosin molecules are adsorbed onto the surface of a glass coverslip, excess unbound

myosin is removed; the coverslip then is placed myosin-side down on a glass slide to form a chamber tl:lrough which solutions can flow. A

solution of actin filaments, made visible and stable by staining with rhodamine-labeled phalloidin, is allowed to flow into the chamber. In

the presence of ATP, the myosin heads walk toward the (+)end of filaments by the mechanism illustrated in Figure 17-26. Because

myosin tails are immobilized, walking of the heads toward the (+)ends

causes sliding of the filaments, which appear to be moving with their (-)ends leading the way. Movement of individual filaments can be

observed in a fluorescence light microscope. (b) These photographs show the positions of three actin filaments (numbered 1, 2, 3) at 30-second intervals recorded by video microscopy. The rate of filament

movement can be determined from such recordings. [Part (b) courtesy of M. Footer and S. Kron.]

(a)

~(-)-p

(b)

Sl head of Actin myosin (+)

" ·pp ~ (+)

17.5 Myosins: Actin-Based Motor Proteins 795

All myosins have a domain related to the S1 domain of myosin II, comprising the head and neck domains, which is responsible for their motor activity. However, as we will see in a later section, the length of the neck domain and the num­ber and type of light chains associated with it varies in differ­ent myosin classes. The tail domain does not contribute to motility but rather defines what is moved by the S !-related domain. Thus, as might be expected, the tail domains can be very different and are tailored to bind specific cargoes.

Myosins Make Up a large Family of Mechanochemical Motor Proteins

Since all myosins have related 51-motor domains with con­siderable similarity in primary amino acid sequence, it is pos­sible to determine how many myosin genes, and how many different classes of myosins, exist in a sequenced genome. There are about 40 myosin genes in the human genome (Fig­ure 17-24), nine in Drosophila, and five in budding yeast. Computer analysis of the sequence relationships between the myosin head domains suggests that about 20 distinct classes

/Deafness/blindness

Myosin II

Myosin I

Myosin V

FIGURE 17-24 The myosin superfamily in humans. Computer analysis of the relatedness of S 1 head domains of all of the approxi­

mately 40 myosins encoded by the human genome. Each myosin is

indicated by a blue dot, with the length of the black lines indicating phylogenetic distance relationship~. Thus myosins connected by short

lines are closely related, whereas those separated by longer lines are more distantly related. Among these myosins are three classes­myosins I, II, and V-widely represented among eukaryotes, with

others having more specialized functions. Indicated are examples in

which loss of a specific myosin causes a disease. [Redrawn and modified from R. E. Cheney, 2001, Mol. Bioi. Ce// 12:780.]

of myosins have evolved in eukaryotes, with greater sequence similarity within a class than between. As indicated in Figure 17-24, the genetic basis for some diseases has been traced to

genes encoding myosins. All myosin head domains convert ATP hydrolysis into mechanical work using the same general mechanism. However, as we will see, subtle differences in this mechanism can have profound effects on the functional prop­erties of different myosin classes. How do these different classes relate with respect to their tail domams? Amazingly, if one takes just the protein sequences of the tail domains of the myosins and uses this information to place them in classes, they fall into the same groupings as the motor domains. This implies that head domains with specific properties have co­evolved with specific classes of tail domains, which makes a lot of sense, suggesting that each class of myosin has evolved to carry out a specific function.

Among all these different classes of myosins are three es­pecially well-studied ones, which are commonly found in ani­mals and fungi: the so-called myosin I, myosm II, and myosin V families (Figure 17-25). In humans, eight genes encode heavy chains for the myosin I family, 14 for the myosin II fam­ily, and three for the myosin V family (see Figure 17-24).

The myosin II class assembles into bipolar filaments, with opposite orientations in each half of the bipolar filament so that there is a cluster of head domains at each end of the fila­ment. This organization is important for its involvement in contraction; indeed, this is the only class of myosins involved in contractile functions. The large number of members in this class reflects the need for myosin II filaments with the slightly different contractile properties seen in different muscles (e.g., skeletal, cardiac, and various types of smooth muscle) as well as in nonmuscle cells.

The myosin II class is the only one that assembles into bipolar filaments. All myosin II members have a relatively short neck domain, with two light chains per heavy chain. The myosin I class is quite large, has a variable number of light chains associated with the neck region, and is the only one in which two heavy chains are not associated through their tail domains and so arc single-headed. The large size and diversity of the myosin I class suggests that these myo­sins perform many functions, most of which remain to be determined, but some members of this family connect actin filaments to membranes, and others are implicated in endo­cytosis. Members of the myosin V class have two heavy chains, giving a motor with two heads, long neck regions with six light chains each, and tail regions that dimerize and terminate in domains that bind to specific organelles to be transported. As we will see shortly, the length of the neck region affects the rate of myosin movement.

ln every case that has been tested so far, myosins move toward the (+)end of an actin filament- with one exception, the myosin VI found in animals. This remarkable myosin has an insert in its head domain to make it work in the opposite direction, and so motility is toward the (- ) end of an actin filament. Myosin V1 is believed to contribute to endocytosis by moving the endocytic vesicles along actin filaments away from the plasma membrane. Recall that membrane-associated

796 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

.·

. '

Class Function Step size

10-14 nm 1-llz l+l Membrane association, endocytosis

II 8nm

36 nm

FIGURE 17-25 Three common classes of myosin. Myosin I consists

of a head domain with a variable number of light chains associated with

the neck domain. Members of the myosin I class are the only myosins to have a single head domain. Some of these myosins are believed to

associate directly with membranes through lipid interactions. Myosin lis

actin filaments have their ( +) ends toward the membrane, so a motor directed toward the (-)end would take them away from the membrane toward the center of the cell.

Conformational Changes in the Myosin Head Couple ATP Hydrolysis to Movement

Studies of muscle .contraction provided the first evidence that myosin heads slide or walk along actin filaments. Unraveling the mechanism of muscle contraction was greatly aided by the development of in vitro motility assays and single-molecule force measurements. On the basis of information obtained with these techniques and the three-dimensional strucwre of the myosin head (see Figure 17-22c), researchers developed a general model for how my6sin harnesses the energy released by A TP hydrolysis to move along an actin filament (Figure 17-26). Because all myosins are thought to use the same basic mechanism to generate movement, we will ignore whether the myosin tail is bound to a vesicle or is part of a thick filament, as it is in muscle. The most important aspect of this model is that the hydrolysis of a single ATP molecule is coupled to each step taken by a myosin molecule along an actin filament.

How can myosin convert the chemical energy released by ATP hydrolysis into mechanical work? This question has long intrigued biologists. It has been known for a long time that the Sl head of myosin is an ATPase, having the ability to hydrolyze ATP into ADP and P,. Biochemical analysis re­vealed the mechanism of myosin movement (Figure 17-26a).

Vesicle _/

Contraction

Organelle transport

have two head domains and two light chains per neck and are the only class that can assemble into bipolar filaments. Myosin Vs have two head

domains and six light chains per neck. They bind specific receptors

(brown box) on organelles, which they transport. All myosins in these three classes move toward the (+ )end of actin filaments.

In the absence of A TP, the head of myosin binds very tightly to F-actin. When ATP binds, the affinity of the head for F-actin is greatly reduced and releases from actin. The myosin head then hydrolyzes the ATP, and the hydrolysis products, ADP and P., remain bound. The energy provided by the hydrolysis of ATP induces a conformational change in the head that results in the head domain rotating with respect to the neck. This is known as the "cocked" position of the head (Figure 17-26b, top) . In the absence ofF-actin, release of P, is excep­tionally slow-the slowest part of the A TPase cycle. How­ever, in the presence of actin, the head binds F-actin tightly, inducing both release of Pi and rotation of the head back to its original position, thus moving the actin filament relative to the neck domain (Figure 17-26b, bottom ). In this way, binding to F-actin induces the movement of the head and release of P., thereby coupling the two processes. This step is known as the power stroke. The head remains bound until the ADP leaves and a fresh ATP binds the head, releasing it from the filament. The cycle then repeats, and the myosin can move again against the filament.

How is hydrolysis of ATP in the nucleotide-binding pocket converted into force? The results of structural studies of myosin in the presence of nucleotide~, and nucleotide ana­logs that mimic the various steps in the cycle, indicate that the binding and hydrolysis of a nucleotide cause a small con­formational change in the head domain. This small move­ment is amplified by a "converter" region at the base of the head, acting like a fulcrum and causing the leverlike neck to rotate . This rotation is amplified by the rodlike lever arm,

17.5 Myosins: Actin-Based Motor Proteins 797

0 FOCUS ANIMATION: Myosin-Actin Cross-Bridge Cycle PODCAST: Myosin Movement Against Actin Filaments

(a) Thick Filament

D "Power strok~e~"~: ........ _.IIC!~ Release of P and elastic energy straightens myosin; moves actin filament left

(b)

FIGURE 17-26 ATP-driven myosin movement along actin filaments. (a) In the absence of ATP, the myosin head is firmly attached to the actin f ilament. Although this state is very short-lived in living

muscle, it is the state responsible for muscle stiffness in death (rigor

mortis). Step 0 : On binding ATP, the myosin head releases from the actin filament. Step If) : The head hydrolyzes the ATP to ADP and P1,

which induces a rotation in the head with respect to the neck. This

"cocked state" stores the energy released by ATP hydrolysis as elastic energy, like a stretched spring. Step D: Myosin in the "cocked" state

binds actin. Step ~: When it is bound to actin, the myosin head

couples release of P with release of the elastic energy to move the actin filament. This is known as the "power stroke," as it involves moving the actin filament with respect to the end of the myosin neck

domain. Step Ill: The head remains tightly bound to the filament as ADP IS released and before fresh ATP is bound by the head.

(b) Molecular models of the conformational changes in the myosin head involved in "cocking" the head (upper pane() and during the

power stroke (lower pane(). The myosin light chains are shown in dark blue and green; the rest of the myosin head and neck are colored in

light blue, and actin is red [Part (a) adapted from R. D. Vale and R. A. Milligan, 2002, Sc1ence 288:88; part (b) courtesy of Mike Geeves.]

798 CHAPTER 17 • Cell Organization and Movement L Microfilaments

4

0 3 Q)

~ E 2- 2 c '() 0

Q) >

E: ERI1 E 'T FIG RE 27 The length of the myosin II neck domain determines the rate of movement. To test the lever-arm model of myosin movement, investigators used recombinant

DNA techniques to make myosin heads attached to different-length

neck domains. The rate at which they moved on actin filaments was determined. The longer the lever arm, the faster the myosin moved,

supporting the proposed mechanism. [Redrawn from K. A. Ruppel and

J. A. Spudich, 1996, Annu. Rev. Cell Mol. Bioi. 12:543-573.]

which constitutes the neck domain, so the actin filament moves by a few nanometers (see figure 17-26b).

This model makes a strong prediction: the distance a myosin head moves along actin during hydrolysis of one ATP-the myosin ste{J size-should be proportional to the length of the neck domain. To test this, mutant myosin mol­ecules were constructed with different-length neck domains and the rate at which they moved down an actin filament was determined. Remarkably, there is an excellent corre­spondence between the length of the neck domain and the rate of movement (Figure 17-27).

Myosin Heads Take Discrete Steps Along Actin Filaments The most critical feature uf myosin is its ability to generate a force that powers movements. Researchers have used optical traps to measure the forces generated by single myosin mole­cules (Figure 17-28 ). In this approach, myosin is immobilized on beads at a low density. An actin filament, held between two optical traps, is lowered toward the bead until it contacts a myosin molecule on the bead. When ATP is added, the myo­sin pulls on the actin filament. u~ing a mechanical feedback mechanism controlled by a computer, one can measure the distance pulled and the forces and duration of the movement. The results of optical trap studies show that myosin ll does not interact with the actin filament continuously but rather binds, moves, and releases it. In fact, myosin U spends on av­erage only about 10 percent of each ATPase cycle in contact with F-actin-it is said to have a duty ratio of 10 percent. This

FIGURE 17-28 Optical trapping of actin. Optical trap techniques

can be used to determine the step size and force generated by a single myosin molecule. In an optical trap, the beam of an infrared laser is

focused by a light microscope on a latex bead (or any other object that

does not absorb infrared light), which captures and holds the bead in the center of the beam. The strength of the force holding the bead is adjusted by increasing or decreasing the intensity of the laser beam. In

this experiment, an actin filament is held between two optical traps. The actin filament is then lowered onto a third bead coated with a

dilute concentration of myosin molecules. If the actin filament

encounters a myosin molecule in the presence of ATP, the myosin will pull on the actin filament, which allows the investigators to measure both the force generated and the step size the myosin takes.

will be important later when we consider that in contractmg muscle, hundreds of myosin heads pull on actin filaments, so that at any one time, 10 percent of the heads are engaged to

provide a smooth contraction. When myosin II does contact F-actin, it takes discrete steps,

which average out to about 8 nm (figure 17-29, top), and gen­erates 3-5 piconewtons (pN) of force, approximately the same force as that exerted by gravity on a single bacterium.

If we now look at a similar optical trap experiment with myosin V, the curves look completely different (Figure 17-29, bottom). Now we can easily discern clear steps of about 36 nm in length. This larger ~rep size reflects the longer neck domain­the lever arm-of myosin V. Moreover, we see that the motor takes many sequential steps without releasing from the actin­it is said to move processively. This is because its ATPase cycle is modified to have a much higher duty ratio( > 70 per­cent) by slowing the rate of ADP release; thus the head re­mains in contact with the actin filament for a much larger percentage of the cycle. Since a single myosin V molecule has two heads, a duty ratio of> 50 percent ensures that one head is in contact at all times as it moves down an actin filament, so that it does not fall off.

Myosin V Walks Hand over Hand down an Actin Filament

The next question is, how do the two heads of myosin V work together to move down a filament? One model pro­poses that the two heads walk down a filament hand over hand with alternately leading heads (Figure 17-30a). An

17.5 Myosins: Actin-Based Motor Protems 799

40

E 20 c Q) 0 c ~ (/)

i:5

E E Q) 0 c ~ (/)

i:5

0

252

216

180

i44

108

72

36

0 0.2

Myosin II

0.5 Time (s)

Myosin V

0.4 Time (s)

0.6 0.8

EXPERIMENTAL FIGURE 17 ·29 Optical trap experiments measure the step size and processivity of myosins. Using an optical

trap setup similar to the one described in Figure 17-28, investigators

have analyzed the behavior of myosin II (top trace) and myosin V

(bottom trace). As shown by the peaks in the trace, myosin II takes erratic small steps (S-15 nm), which means it binds the actin filament,

moves, and then lets go. It is therefore a nonprocessive motor. By contrast, single-headed myosin V takes clear 36-nm steps one after the

other, so it has a step size of 36 nm and is highly processive-that is, it does not let go of the actin filament. [Part (a) from Finer et al., 1994, Nature

368:113; part (b) from M. Rief et al., 2000, Proc. Nat'/ Acad. Sci. USA 97:9482.]

alternative possibility is an inchworm model, in which the leading head takes a step, the second head is pulled up behind it, and then the leading head takes another step (Figure 17-JOb). How can one distinguish between these models? In the inchworm model, each individual head takes 36-nm steps, whereas in the walking model, each takes 72-nm steps. Scien­tists have managed to attach a fluorescent probe to just one neck rt-eion of myosin V and watch it walk down an actin filament: it takes 72-nm steps (Figure 17-30c), and so it walks hand over hand down a filament. Why is the step size of my­osin V so large? If we compare its step size of 36 nm to the structure of the actin filament, we see that it is the same as the length between helical repeats in the actin filament (see Fig­ures 17-5b and 17-30a), so myosin V steps between equiva-

(a) Hand over hand

T

(b) Inchworm

Label on neck

(+)

(c)

E 500 E c 0

·.;::; 'iii 0 c...

0 10 20 30 40 50 60

Time (s)

EXPERIMENTAL FIGURE 17-30 Myosin V has a step size of 36 nm, yet each head moves in 72-nm steps, so it moves hand over

hand. Two models for myosin V movement down a filament have been suggested. (a) In the hand-over-hand model, one head binds an actin fi lament, and the other then swings around and binds a site 72 nm

ahead. (b) In the inchworm model, the leading head moves 36 nm,

then the lagging head moves up behind it, allowing the leading head to take another 36-nm step. (c) Two-headed myosin V labeled with a

fluorescent tag on just one head appears to have a step size of 72 nm.

Thus myosin V walks hand over hand. [Adapted from A. Yildiz et al., 2003, Science 300:2061.]

lent binding sites as it walks down one side of an actin filament. Myosin V has presumably evolved to take large steps the size of the helical repeat of actin and to do this very processively so that it rarely dissociates from an actin fila­ment. These arc exactly the properties one would expect for a motor designed to transport cargo along an actin filament.

800 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

·.

·.

' ·

.·

KEY CONCEPTS for Section 17.5

Myosins: Actin-Based Motor Proteins

• Myosins are actin-based motors powered by ATP hydrolysis.

• Myosins have a motor head domain, a lever-arm neck do­main, and a cargo-binding tail domain (see Figure 17-22).

• There arc many classes of myosin, with three classes pres­ent in many eukaryoll:l>: myosin I has a single head domain, myosin II has two heads and assembles into bipolar fila­ments, and myosin V has two heads but does not assemble into filaments (see Figure 17-25).

• Myosins convert ATP hydrolysis to mechanical work by am­plifying a small conformational change in their head through their neck domain when the head is bound to F-actin (see Fig­ure 17-26).

• Myosin heads take discrete steps along an actin filament, which can be small (8 nm) and nonprocessive in the case of myosin II, or large (36 om) and processive for myosin V.

17.6 Myosin-Powered Movements

We have already discussed how myosins have head and neck domains responsible for their motor properties. We now come to the tail regions, which define the cargoes that myosins move. The function of many of the newly discovered classes of myosins found in metazoans is not yet known. In this section, we give just two examples where we have a good idea of spe­cific myosin functions. Our first example is skeletal muscle, which is where myosin II was discovered. In muscle, many myosin I! heads bundled into bipolar filaments, each with a short duty cycle, work together to bring about contraction. Similarly organiied contractile machineries function in the contraction of smooth muscle and in stress fibers, as well as in the contractile ring during cytokinesis. We then turn to the myosin V class, which has a long duty cycle that allows these myosins tO transport cargoes over relatively long distances without dissociating from actin filaments.

Myosin Thick Filaments and Actin Thin Filaments in Skeletal Muscle Slide Past One Another During Contraction Muscle cells have evolved to carry out one highly specialized function: contraction. Muscle contractions must occur quickly and repetitively, and they must occur through long distances and with enough force to move large loads. A typi­cal skeletal muscle cell is cylindrical, large (1-40 J.Lm in length and 10-50 J.Lm in width), and multinucleated (containing as many as 100 nuclei) (Figure 17-31a). Within each muscle celt are many myofibrils consisting of a regular repeating array of a specialized structure called a sarcomere (Figure 17-31 b). A sar­comere, which is about 2 J.Lm long in resting muscle, shortens

by about 70 percent of its length during contraction. Electron microscopy and biochemical analysis have shown that each sarcomere contains two major types of filaments: thick fila­ments, composed of myosin II, and thin filaments, containing actin and associated proteins (Figure 17-31c).

The thick filaments are composed of myosin II bipolar filaments, in which the heads on each half of the filament have opposite orientations (see Figure 17-22a). The thin actin filaments arc assembled with their ( +) ends embedded in a densely staining structure known as the Z disk, so that the two sets of actin filaments in a sarcomere have opposite orientations (Figure 17-32). To understand how a muscle contracts, consider the interactions between one myosin head (among the hundreds in a thick filament) and a thin (actin) filament, as diagrammed in Figure 17-26. Dunng these cyclical interactions, also called the cross-bridge cycle, the hydrolysis of ATP is coupled to the movement of a myo­sin head toward the Z disk, which corresponds to the ( +) end of the actin thin filament. Because the thick filament is bipolar, the action of the myosin heads at opposite ends of the thick filament draws the thin filaments toward the center of the thick filament and therefore toward the center of the sarcomere (see Figure 17-32). This movement shortens the sarcomere until the ends of the thick filaments abut the Z disk. Contraction of an intact muscle results from the activ­ity of hundreds of myosin heads on a single. thick filament, amplified by the hundreds of thick and thin filaments in a sarcomere and thousands of sarcomeres in a muscle fiber. We can now see why myosin II is both nonprocessive and needs to have a short duty cycle: each head pulls a short distance on the actin filament and then lets go to allow other heads to pull, and so many heads working together allow the smooth contraction of the sarcomere.

The heart is an amazing contractile organ-it contracts without interruption about 3 million times a year, or a

fifth of a billion times in a lifetime. The muscle cells of the heart contain contractile machinery very similar to that of skel­etal muscle except that they are mono- and hi-nucleated cells. In each cell, the end sarcomeres insert into structures at the plasma membrane called intercalated disks, which link the cells into a contractile chain. Since heart muscle cells are only generated early in human life, they cannot be replaced in re­sponse to damage, such as occurs during a heart attack. Many different mutations in proteins of the heart contractile machin­ery give rise to hytJertrophic cardiomyopathies-thickening of the heart wall muscle, which compromises its function. For example, many mutations have been documented in the car­diac myosin heavy-chain gene that compromise the protein's contractile function even in heterozygous individuals. ln such individuals, the heart tries to compensate by hypertrophy (en­largement), often resulting in fatal heart arrhythmia (irregular beating). In addition to myosin heavy-chain defects, defects that result in cardiomyopathies have been traced to mutations in other components of the contractile machinery, including actin, myosin light chains, tropomyosin and troponin, and structural components such as titin (discussed below). •

17.6 Myosin-Powered Movements 801

FIGURE 17-31 Structure of the skeletal muscle sarcomere. (a) Skeletal muscles consist of muscle fibers made of bundles of multinucleated cells. Each cell contains a bundle of myofibrils, which consist of thousands of repeating contractile structures called sarcomeres. (b) Electron micrograph of mouse striated muscle in longitudinal section, showing one sarcomere. On either side of the Z disks are the lightly stained I bands, composed entirely of actin thin filaments. These thin filaments extend from both sides of the Z disk to interdigitate with the dark-stained myosin thick filaments in the A band. (c) Diagram of the arrangement of myosin and actin filaments in a sarcomere. [Part (b) courtesy of S. P. Dadoune.]

Skeletal Muscle Is Structured by Stabilizing and Scaffolding Proteins The structure of the sarcomere is maintained by a number of accessory proteins (Figure 17-33 ). The actin filaments are stabilized on their ( +) ends by CapZ and on their (-) ends by tropomodulin. A giant protein known as nebulin extends along the thin actin filament all the way from the Z disk to tropomodulin, to which it binds. Ncbulin consists of repeat­mg domains that bind to the actin in the filament, and it is believed that the number of actin-binding repeats, and there­fore the length of nebulin, determines the length of the thin filaments. Another giant protein, called titin (because it is so large), has its head associated with the Z disk and extends to the middle of the thick filament, where another titin mole­cule extends to the subsequent Z disk. Titin is believed to be an elastic molecule that holds the thick filaments in the mid­dle of the sarcomere and also prevents overstretching to en­sure that the thick filaments remain interdigitated between the thin filaments.

Contraction of Skeletal Muscle Is Regulated by Ca2+ and Actin-Binding Proteins Like many cellular processes, skeletal muscle contraction is initiated by an increase in the cytosolic Ca2

c concentration. As described in Chapter 11, the Ca2

+ concentration of the cytosol is normally kept low, below 0.1 J..LM. In skeletal mus­cle cells, a low cytosolic CaH level is maintained primarily by a unique Ca2

+ A TPase that continually pumps Ca2+ ions

from the cytosol containing the myofibrils into the sarco­plasmic reticulum (SR), a specialized endoplasmic reticulum of the muscle cells (Figure 17-34). This activity establishes a reservoir of Ca2 in the SR.

The arrival of a nerve impulse (or action potential; see Chapter 22) at a neuromuscular junction triggers an action potential in the muscle-cell plasma membrane (also known as the sarcolemma). The action potential travels down in­vaginations of the plasma membrane known as transverse tubules, which penetrate the cell to lie around each myofi­bril. The arrival of the action potential in the transverse tu­bules stimulates the opening of voltage-gated Ca2

+ channels in the SR membrane, and the ensuing release of CaH from the SR raises the cytosolic Ca2 concentration in the myofi-

(a)

Bundle of muscle fibers

:....._ ____ Multinucleated muscle cell

Myofibril

"------- Nuclei >(..-.----y ~~ ~:~ ~~ ~

- "J Sarcomere (b)~

Z disk Z disk -- I band ~+----- A band ____ .,._+--- I band -(c)

~

,.;;;:::. --:-. ·~· ··::

''-----~.r------/ y

Myosin filaments

Actin filaments Actin fi laments

brils. This elevated Ca1 concentration induces a change in two accessory proteins, tropomyosin and troponin, which are bound to the actin thin filaments and normally block myosin binding. The change in position of these proteins on the actin thin filaments in turn permits the myosin-actin

802 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

Relaxed Actin Myosin Actin

(+) (+) l (-) J (-) I (+) (+)

"W; ----i ... .J"9 ~ ""'".,...

~~ - •Ae ~ ~

~ ' ~'~~

z di~k Z disk

Contracted l +ATP, Ca2+

FIGURE 17-32 The sliding-filament model of contraction in striated muscle. The arrangement of thick myosin and thin actin

filaments in the relaxed state is shown in the top diagram. In the presence of ATP and Ca2

, the myosin heads extending from the thick

filaments walk toward the (+)ends of the thin fi laments. Because the

thin filaments are anchored at the Z disks (purple), movement of myosin pulls the actin filaments toward the center of the sarcomere,

shortening its length in the contracted state, as shown in the

bottom diagram.

interactions and hence contraction. This type of regulation is very rapid and is known as thin-filament regulation.

Tropomyosin (TM) is a ropelike molecule, about 40 nm in length, that binds to seven actin subunits in an actin fi la­ment. TM molecules are strung together head to tai l, forming a continuous chain along each side of the actin thin filament (Figure 17-35a, b). Associated with each tropomyosin is tro­ponin (TN), a cqmplex of three subunits, TN-T, TN-I, and TN-C. Troponin-C is the calcium-binding subunit of troponin .

(a)

Sarcolemma

FIGURE 17-34 The sarcoplasmic reticulum regulates the level of free Ca2+ In myofibrils. (a) When a nerve impulse stimulates a muscle cell, the action potential is transmitted down a transverse tubule

(yellow), which is continuous with the plasma membrane (sarcolemma),

leading to release of Ca2 from the adjacent sarcoplasmic reticulum

CapZ Tropomodulin (-)

Z disk (+) (+)

FIGURE 17-33 Accessory proteins found in skeletal muscle. To stabilize the actin filaments, CapZ caps the (+)end of the thin filaments at the Z disk, whereas tropomodulin caps the(-) end. The

giant protein titin extends through the thick filaments and attaches to the Z disk. Nebulin binds actin subunits and determines the length

of the thin fi lament.

TN-C controls the position of TM on the su~face of an actin filament through the TN-I and TN-T subunits.

Under the control of Ca2+ and TN, TM can occupy two

positions on a thin filament-switching from a state of mus­cle relaxation to contraction. In the absence of Ca2

+ (the relaxed state), TM blocks myosin's interaction with F-actin and the muscle is relaxed . Binding of Ca2 ions to TN-C triggers movement of TM to a new site on the filament, thereby exposing the myosin-binding sires on actin (see Fig­ure 17-35b). Thus, at Ca2

+ concentrations greater than lj..lM, the inhibition exerted by the TM-TN complex is relieved and contraction occurs. The Ca2 ' -dependent cycling between

(b)

(blue) into the myofibrils. (b) Thin-section electron micrograph of

skeletal muscle, showing the intimate relationsh ip of the sarcoplasmic reticulum to the muscle fibers. [Part (b) from K. R. Porter and C. Franzini­

Armstrong, ASCB Image & Video Library, August 2006:FND-14. Available at: httpJ /cellimages.ascb.org/u? /p4041 co Ill, 83.)

17.6 Myosin-Powered Movements 803

(a)

Tropomyosin (TM)

(b)

FIGURE 17-35 Ca2+ -dependent thin-filament regulation of skeletal muscle contraction. (a) Model of the tropomyosin-troponin regulatory complex on a thin filament. Troponin is a protein complex that is bound to the long a.-helical tropomyosin molecule. (b) Three­dimensional electron-microscopic reconstructions of the tropomyosin helix (yellow) on a muscle thin filament. Tropomyosin in the relaxed state (top) shifts to a new position (arrow) in the state inducing

relaxation and contraction states in skeletal muscle is sum­marized in Figure 17-35c.

Actin and Myosin II Form Contractile Bundles in Nonmuscle Cells In skeletal muscle, actin thin filaments and myosin II thick filaments assemble into contractile structures. Nonmuscle cells contain several types of related contractile bundles com­posed of actin and myosin II filaments, which are similar to skeletal muscle fibers but much less organized. Moreover, they lack the troponin regulatory system and arc instead reg­ulated by myosin phosphorylation, as we will discuss later.

In epithelial cells, contractile bundles are most commonly found as an adherens belt, also known as the circumferential belt, which encircles the inner surface of the cell at the level of the adherens junction (see Figure 17-4a) and are important in maintaining the integrity of the epithelium (discussed in Chap­ter 20). Stress fibers, which are seen along the lower surfaces of cells cultured on artificial (glass or plastic) surfaces or in extracellular matrices, are a second type of contractile bundle (see Figure 17-4a, c) important in cell adhesion, especially on deformable substrates. The ends of stress fibers terminate at imegrin-containing focal adhesions, special structures that at­tach a cell to the underlying substratum (see Figure 17-41 and Chapter 20). Circumferential belts and stress fibers contain several proteins found in the contractile apparatus of smooth muscle and exhibit some organizational features resem-

Myosin-binding site

(c)

Relaxation Myosin-binding site masked

Actin • TM • TN

-Ca2

Actin • TM • TN- Ca2+

Myosin-binding site exposed Contraction

contraction (bottom) when the Ca2+ concentration increases. This

movement exposes myosin-binding sites (red) on actin. (Troponin is not shown in this representation, but it remains bound to tropomyosin in both states.) (c) Summary of the regulation of skeletal muscle contraction by Ca2 binding to troponin. [Part (b) ad'lpted from W. Lehman, R. Craig, and P. Vi bert, 1993, Nature 123:313, courtesy of P. Vibert.]

bling those of muscle sarcomeres. A third type of contrac­tile bundle, referred to as a contractile ring, is a transient structure that assembles at the equator of a dividing cell, encircling the cell midway between the poles of the mitotic spindle (Figure 17-36a). As the ring contracts, pu lling the plasma membrane in, the cytoplasm is divided and eventu­ally pinched into two parts in a process known as cytokine­sis, giving rise to two daughter cells. Dividing cells stained with antibodies against myosin I and myosin II show that myosin II is localized to the contractile ring, whereas myo­sin I is at the distal regions, where it links the actin cortex to the plasma membrane (Figure 17-36b). Cells deleted for the gene encoding the heavy chain of myosin II are unable to undergo cytokinesis, thereby establishing a role for myo­sin II in cell division. Instead, these cells form a multinucle­ated syncytium because cytokinesis, but not nuclear division, is inhibited.

Myosin-Dependent Mechanisms Regulate Contraction in Smooth Muscle and Nonmuscle Cells Smooth muscle is a specialized tissue composed of contractile cells that is found in many internal organs. For example, smooth muscle surrounds blood vessels to regulate blood pressure, surrounds the intestine to move food through the gut, and restricts airway passages in the lung. Smooth muscle cells contain large, loosely aligned contractile bundles that

804 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

(a)

Microtubule Contractile Chromosome ring

EXPERIMENTAL F GURE 1 ·36 Fluorescent antibodies reveal the localization of myosin I and myosin II during cytokinesis. (a) Diagram of a cell going through cytokinesis, showing the mitotic spindle (microtubules green, chromosomes blue) and the contractile ring with actin filaments (red). (b) Fluorescence micrograph of a Dictyostelium ameba during cytokinesis reveals that myosin II (orange)

resemble the contractile bundles in epithelial cells. The con­tractile apparatus of smooth muscle and its regulation consti­tute a valuable model for understanding how myosin activity is regulated in a nonmuscle cell. As we have just seen, skeletal muscle contraction is regulated by the tropomyosin-troponin complex bound to the actin thin filament switching between the contraction-inducing state in the presence of Ca2 and the relaxed state in its absence. In contrast, smooth muscle con­traction is regulated by the cycling of myosin II between on and off states. Myosin II cycling, and thus contraction of smooth muscle and nonmuscle cells, is regulated in response to many extracellular signaling molecules.

Contraction ,of vertebrate smooth muscle is regulated primarily by a pathway in which the myosin regulatory light chain (LC) associated with the myosin II neck domain (see Figure 17-22b) undergoes phosphorylation and dephosphor­ylation. When the regulatory light chain is not phosphory­lated, the smooth muscle myosin II adopts a folded conformation and its A TPase cycle is inactive. When the regulatory LC is phosphorylated by the enzyme myosin LC kinase, whose ac­tivity is regulated by the level of cytosolic free Ca2

, the myosin II unfolds and assembles into active bipolar filaments and becomes active to induce contraction (Figure 17-37). The Ca2 -dependent regulation of myosin LC kinase activity is mediated through the Cal+ -binding protein calmodulin (see Figure 3-31 ). Calcium first binds to calmodulin, which induces a conformational change in the protein, and the Ca2+ /calmodulin complex then binds to myosin LC kinase and activates it. When the Ca2+ returns to its resting level, myosin LC kinase becomes inactive and myosin light-chain phosphatase removes the phosphates to allow the system to return to its relaxed state. This mode of regulation relies on the diffusion of Cah over greater distances than in sarco­meres and on the action of protein kinases, so contraction

(b)

is concentrated in the contractile ring, also known as the cleavage furrow, whereas myosin I (green) is localized at the poles of the cell. The cell was stained with antibodies specific for myosin I and myosin II, with each antibody preparation linked to a different fluorescent dye. [Courtesy of Y. Fukui.]

is much slower in smooth muscle than in skeletal muscle. Because this regulation involves myosin,, it is known as thick-filament regulation.

The role of activated myosin LC kinase can be demon­strated by microinjecting a kinase inhibitor into smooth muscle cells. Even though the inhibitor does not block the rise in the cyrosolic Ca2+ level that occurs following stimula­tion of the cell, injected cells cannot contract.

Unlike skeletal muscle, which is stimulated to contract solely by nerve impulses, smooth muscle cells and nonmuscle cells are regulated by many types of external signals. For ex­ample, norepinephrine, angiotensin, endothelin, histamine, and other signaling molecules can modulate or induce the contraction of smooth muscle, or elicit changes in the shape and adhesion of nonmuscle cells by triggering various signal­transduction pathways. Some of these pathways lead to an increase in the cytosolic Ca2

+ level; as previously described, this increase can stimulate myosin activity by activating myo­sin LC kinase (see Figure 17-37). As we will discuss below, other pathways activate Rho kinase, which is also able to activate myosin activity by phosphorylating the regulatory light chain, although in a Ca2

+ -independent manner.

Myosin-V-Bound Vesicles Are Carried Along Actin Filaments In contrast to the contractile functions of myosin II filaments, the myosin V family of proteins are the most processive myo­sin motors known and transport cargo down actin filaments. In the next chapter we discuss how they can work together with microtubule motors to bring about transport of organelles. Al­though not a lot is known about their functions in mammalian cells, myosin V motors are not unimportant: defects in a

17.6 Myosin-Powered Movements 805

FIGURE 17-37 Myosin phosphorylation mechanism for regulating smoot h muscle contraction. In vertebrate smooth muscle, phosphor­ylation of the myosin regulatory light chain (LC) activates contraction. At Ca2

· concentrations< 10 6 M, the regulatory light chain is not phosphorylated, and the myosin adopts a folded conformation. When the Ca2

• level rises, it binds calmodulin (CaM), which undergoes a conformational change (CaM*). The CaM*-Ca2

- complex binds and activates myosin light-chain kinase (M LC kinase), which then phos­phorylates the myosin LC. This phosphorylation event unfolds the myo­sin, which is now active and can assemble into bipolar filaments to participate in contraction. When the Ca2 levels drop, the myosin LC is dephosphorylated by myosin light-chain (MLC) phosphatase, which is not dependent on Ca2 for activity, causing muscle relaxation.

specific myosin V protem can cause severe diseases, such as seizures (see Figure 17-24).

Much more is known about myosin V motors in more experimentally accessible and simpler systems such as the budding yeast. This well-studied organism grows by bud­ding, which requires its secretory machinery to target newly synthesized material to the growing bud (Figure 17-38a). Myosin V transports secretory vesicles along actin filamems at 3 ~J..m/s into the bud. However, this is not the only func­tion of myosin V proteins in yeast. At a later stage of the cell cycle, all the organelles have to be distributed between the mother and daughter cells. Remarkably, myosin Vs in yeast

Relaxation Myosin folded and inactive

Myosin LC

CaM+ + c az• CaM·-Ca2'- MLC MLC kinase ~ MLC kinase phosphatase

(inactive) _ caz• (active)

Myosin unfolded and active

Regulatory light chain Myosin LC-®

I

~:1:::=== ~ l·-=::":"'~""""'i,S!SiS!*iS!SSa.'SS\S!SS!!'iS!"!S"a.'S'"S!'ii:"'

Heavy chains (tail)

Head

ll Contraction

Myosin filament assembly

~ OVERVIEW ANIMATION: Movement of Multiple Cargoes by Myosin V in Yeast

FIGURE 17-38 Cargo movement by myosin Vs in budding yeast. (a) The yeast Saccharomyces cerevisiae (used in making bread, beer, and wine) grows by budding. Secretory vesicles are transported into the bud, which swells to about the size of the mother cell. The cells then go through cytokinesis to form two daughter cells, and each divides again. (b) Diagram of a medium-sized bud showing how myosin Vs transport secretory vesicles (SV) down actin cables nucleated by form ins (purple) located at the bud tip and bud neck. Myosins Vs are also used to segregate organelles, such as the vacuole (the yeast equivalent of a lysosome), peroxisomes, endoplasmic reticulum (ER), trans-Golgi

network (TGN), and even selected mRNAs into the bud. Myosin V also binds the end of cytoplasmic microtubules (green) to orient the nucleus in preparation for mitosis. [Adapted from D. Pruyne et al., 2004,

Ann. Rev. Cell Bioi. 20 :559.]

(a)

c

(b)

(-)

Var:uole

(-)

806 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

~ Bud

(+) ~~ mRNA ER

(-) ~ Yj+JP (+')~

·.

'·

provide the transport system for segregation of many organ­elles, including peroxisomes, lysosomes (also known as vac­uoles), endoplasmic reticulum, the trans-Golgi network, and even transport the ends of microtubules and some specific messenger RNAs into the bud (Figure 17-38b). Whereas budding yeast uses myosin V and polarized actin filaments in the transport of many organelles, animal cells, which are much larger, employ microtubules and their motors to trans­port many of these organelles over relatively long distances. We discuss these transport mechanisms in the next chapter.

(a)

Vacuole

Nitella cell

:~~:~,m { ER

(+) <\ Nonmoving { cortical cytoplasm

Plasma membrane ~ Cell wall/

(-)

FIGURE 17-39 Cytoplasmic streaming in cylindrical giant algae. (a) Cells of Nitella, a freshwater alga commonly found in ponds in the

summer. The cytoplasmic movement, described below, is amazing and can readily be observed with a simple microscope, so go find some Nitel/a (or related alga) and watch this wonderful phenomenon! (b) The

center of a Nitella cell is fil led with a single large water-fil led vacuole,

which is surrounded by a layer of moving cytoplasm (blue arrows). A nonmoving layer of cortical cytoplasm filled with chloroplasts lies just under the plasma membrane (enlarged in bottom figure). On the inner

Perhaps the most dramatic use of myosin Vs is seen in the giant green algae, such as Nitella and Chara. ln these large cells, which can be 2 em in length, cytosol flows rapidly, at a rate approaching 4.5 mm/min, in an endless loop around the inner circumference of the cell (Figure 17-39). This cytoplasrmc streammg is a principal mechanism for distributing cellular me­tabolites, especially in large cells such as plant cells and amebas.

Close inspection of objects caught in the flowing cytosol, such as the endoplasmic reticulum (ER) and other membrane­bounded vesicles, shows that the velocity of streaming increases

(c)

side of this layer are bundles of stationary actin filaments (red), all

oriented with the same polarity. A motor protein (blue), a plant myosin V, carries parts of the endoplasmic reticulum (ER) along the actin filaments. The movement of the ER network propels the entire viscous

cytoplasm, including organelles that are enmeshed in the ER network.

(c) Electron micrograph of the cortical cytoplasm showing a large vesicle connected to an underlying bundle of actin filaments. [Part (a)

from James C. French; part (c) from B. Kachar.]

17.6 Myosin-Powered Movements 807

from the cell center (zero velocity) to the cell periphery. This gradient in the rate of flow is most easily explained if the motor generating the flow lies at the membrane. In electron micro­graphs, bundles of actin filaments can be seen aligned along the length of the cell, lying above stationary chloroplasts located adjacent to the membrane. The bulk cytosol is propelled by myosin V (also known as myosin XI in plants) attached to parts of the ER adjacent to the actin filaments. The flow rate of rhe cytosol in Nitella is ahout 15 times as fast as the movement produced by any other known myosin.

KEY CONCEPTS of Section 17.6

Myosin-Powered Movements

• In skeletal muscle, contractile myofibrils are composed of thousands of repeating units called sarcomeres. Each sarco­mere consists of two interdigitating filament types: myosin thick filaments and actin thin filaments (see Figure 17-31 ).

Skeletal muscle contraction involves the ATP-dependent sliding of myosin thick filaments along actin thin filaments to shorten the sarcomere and hence the myofibril (see Fig­ure 17-32).

The ends of the actin thin filaments in skeletal muscle are stabilized by CapZ at the ( +) end and by tropomodulin at the (--)end. Two large proteins, nebulin associated with the thin filaments and titin with the thick filaments, also con­tribute to skeletal muscle organization.

Skeletal muscle contraction is subject to thin-filament reg­ulation. At low levels of free Cal+, the muscle is relaxed and tropomyosin blocks the interaction of myosin and F-actin. At elevated levels of free Cal·, the troponin complex associ­ated with tropomyosin binds Ca2+ and moves the tropomyo­sin to uncover the myosin-binding sites on actin, allowing contraction (sec Figure 17-35).

Smooth and nonmuscle cells have contractile bundles of actin and myosin filaments, with a similar organization as skeletal muscle but less well ordered.

Contractile bundles are subject to thick-filament regula­tion. A myosin light chain is phosphorylated by myosin light-chain kinase, which activates myosin and hence in­duces contraction. The myosin light-chain kinase is activated by binding Ca2 -calmodulin when the free Cal concentra­tion rises (see Figure 17-3 7).

• Myosin V transports cargo by walking processively along actin filaments.

17.7 Cell Migration: Mechanism, Signaling, and Chemotaxis

We have now examined the different mechanisms used by cells to create movement-from the assembly of actin filaments and the formation of actin-filament bundles and networks to the contraction of bundles of actin and myosin and the transport of

organelles by myosin molecules along actin filaments. Some of these same mechanisms constitute the major processes whereby cells generate the forces needed to migrate. Cell migration re­sults from the coordination of motions generated in different parts of a cell, integrated with a directed endocytic cycle.

The study of cell migration is important to many fields of biology and medicine. For example, an essential feature of animal development is the migration of specific cells along pre­determined paths. [pitheliall:ells in an adult animal migrate to

heal a wound, and white blood cells migrate to sites of infec­tion. Less obvious are the continual slow migration of intesti­nal epithelial cells along the villi in the intestine and the slow but constant migration of endothelial cells that line the blood vessels. The inappropriate migration of cancer cells after breaking away from their normal tissue results in metastasis.

Cell migration is initiated by the formation of a large, broad membrane protrusion at the leading edge of a cell. Video microscopy reveals that a major feature of this movement is the polymerization of actin at the membrane. Actin filaments at the leading edge are rapidly cross-linked into bundles and networks in a protruding region, called a lamellipodium in vertebrate cells. In some cases, slender, fingerlike membrane projections, called filopodia, also extend from the leading edge. These structures form stable contacts \vith the underly­ing surface (such as the extracellular matrix) that the cell moves across. In this section, we take a closer look at how cells coordinate various microfilament-based processes with endocytosis to move across a surface. We also consider the role of signaling pathways in coordinating and integrating the actions of the cytoskeleton, a major focus of current research.

Cell Migration Coordinates Force Generation with Cell Adhesion and Membrane Recycling

A moving fibroblast (connective tissue cell) displays a charac­teristic sequence of events-initial extension of a membrane protrusion, attachment to the substratum, forward flow of cytosol, and retraction of the rear of the cell (Figure 17-40). These events occur in an ordered pattern in a slowly moving cell such as a fibroblast, but in rapidly moving cells, such as macrophages, all of them are occurring simultaneously in a coordinated manner. We first consider the role of the actin cytoskeleton, and then the involvement of the endocytic cycle.

Membrane Extension The network of actin filaments at the leading edge is a type of cellular engine that pushes the mem­brane forward in a manner very similar to the propulsion of Listeria by actin polymerization (Figure 17 -41d; for Listeria, see Figure 17-17c). Thus at the membrane of the leading edge, actin is nucleated by the activated Arp2/3 complex and filaments are elongated hy ac;sembly onto ( +) ends adjacent to the plasma membrane. As the actin meshwork is fixed with respect to the substratum, the front membrane is pushed our as the filaments elongate. This is very similar to Listeria, which "rides" on the polymerizing actin tail, which is also fixed within the cytoplasm. Actin turnover, and thus treadmilling, is mediated, as it is in the comet tails of Liste­rta, by the action of profilin and cofilin (see Figure 17-41d).

808 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

(;) VIDEO: Mechanism of Fish Keratinocyte Migration VIDEO: Actin Filaments in the Lamellipodium of a Fish Keratinocyte

FIGURE 17-40 Steps in cell movement. Movement begins with the

extension of one or more lamellipodia from the leading edge of the cell 0 ; some lamellipodia adhere to the substratum by focal adhesions f) .

Then the bulk of the cytoplasm in the cell body flows forward due to contraction at the rear of the cell !D. The trailing edge of the cell

remains attached to the substratum until the tail eventually detaches and retracts into the cell body. During this cytoskeleton-based cycle,

the endocytic cycle internalizes membrane and integrins at the rear of the cell and transports them to the front of the cell (arrows) for reuse in making new adhesions m.

Ce ll-Substrate Adhesions When the membrane has been ex­tended and the cytoskeleton has been assembled, the plasma membrane becomes firmly attached to the substratum. Time­lapse microscopy shows that actin bundles in the leading edge become anchored to structures known as focal adhesions (Fig­ure 17-41c). The attachment serves two purposes: it prevents the leading lamella from retracting, and it attaches the cell to the substratum, allowing the cell to move forward . Given the im­portance of focal adhesions and their regulation during cell lo­comotion, it is not surprising that they have been found to be very rich in molecules involved in signal-transduction pathways. Focal adhesions are discussed in more detail in Chapter 20, when we discuss cell-matrix interactions.

The cell-adhesion molecules that mediate most cell-matrix interactions are membrane proteins called integrins. These proteins have an external domain that binds to specific com­ponents of the extracellular matrix, such as fibronectin and collagen, and a cytoplasmic domain that links them to the actin cytoskeleton (see Figure 17-4lc and Chapter 20). The cell makes attachments at the front, and as the cell migrates forward, the adh.esions eventually assume positions toward the back.

Cell-Body Translocation After the forward attachments have been made, the bulk contents of the cell body are translo­cated forward (see Figure 17-40, step IJ). It is believed that the nucleus and the other, organelles embedded in the cyto­skeleton are moved forward by myosin [[-dependent cortical contraction in the rear part of the cell, like squeezing the lower half of a tube of toothpaste. Consistent with this model, myosin II is localized to the rear cell cortex.

Breaking Cell Attachments Finally, in the last step of move­ment (de-adhesion), the focal adhesions at the rear of the cell are broken, the imegrins recycled, and the freed tail brought forward. In the light mi~.:ruscope, the tail is seen to "snap" loose from its connections-perhaps by the contraction of stress fibers in the tai l or by elastic tension-and it some­times leaves a little bit of its membrane behind, still firmly attached to the substratum.

The abi lity of a cell to move corresponds to a balance between the mechanical forces generated by the cytoskeleton

Direction of movement ---+

D Extension

lamellipodium

/

fJ Adhesion

IJ Translocation Cell body movement ---+

IJ De-adhesion and endocytic recycling

and the resisting forces generated by cell adhesions. Cells cannot move if they are ei ther too strongly attached or not attached to a surface. This relationship can be demonstrated by measuring the rate of movement in cells that express varying levels of integrins. Such measurements show that the fastest migration occurs at an intermediate level of adhesion, with the rate of movement falling off at high and low levels of adhesion. Cell locomotion thus results from traction forces exerted by the cell on the underlying substratum.

Recycling Membrane and lntegrins by Endocytosis The dy­namic changes in the actin cytoskeleton alone are not suffi­cient to drive cell migration; it is also dependent on endocytic recycling of membranes. T he membrane needed during la­mellipodium extension is provided from internal endosomes fo llowing their exocytosis. Adhesion molecules in focal adhesions at the rear of the cell are internalized from disas­sembling foca l contacts and transported by an endocytic cycle to the front to make new substrate attachments (Figure 17-40, step 19). This cycle of adhesion molecules in a migrat­mg cell resembles the way a tank uses its treads to move for­ward. This movement of membrane internally through the cell also generates a rearwards membrane flow across the sur­face of the cell. Indeed, this type of flow may contribute to cell locomotion, as it has recently been found that white blood cells can move in a liquid ("swim") in the absence of attach­ment to a substrate.

17.7 Cell Migration: Mechanism, Signaling, and Chemotaxis 809

0 VIDEO: Actin Dynamics in a Migrating Fibroblast VIDEO: Microtubule and Actin Movements in Migrating Cells

(a)

(c)

Focal adhesion~/ -

FIGURE 17-41 Actin-based structures involved in cell locomotion. (a) Localization of actin in a fibroblast expressing

GFP-actin. (b) Diagram of the classes of microfi laments involved in cell migration. The meshwork of actin fi laments in the leading edge

advances the cell forward. Contracti le fibers in the cell cortex squeeze

the cell body forward, and stress fibers terminating in focal adhesions also pull the bulk of the cell body up as the rear adhesions are released.

The Small GTP-Binding Proteins Cdc42, Rae, and Rho Control Actin Organization

A striking feature of a moving cell is irs polarity: a cell has a front and a back. When a cell makes a turn, a new leading edge forms in the new direction. If these extensions formed in all directions at once, the cell would be unable to pick a new direction of movement. To sustain movement in a particu lar direction, a cell requires signa ls to coordinate events at the front of the cell with events at the back and, indeed, signals to tell the cell where its front is. Understanding how this coordi­nation occurs emerged from studies with growth factors.

Growth factors, such as epidermal growth factor (EGF) and platelet-derived growth factor (PDGF), bind to specific

(b)

Stress f ibers

ATP-G-actin -1 Profilin - ... -{

A DP-G-actin __. "-

Leading edge

(c) The st ructure of focal adhesions involves the attachment of the ends

of stress fibers through integrins to the underlying extracellular matrix. Focal adhesions also conta in many signaling molecules important for cell locomotion. (d) The dynamic actin meshwork in the leading edge is

nucleated by the Arp2/3 complex and employs the same set of factors that control assembly and disassembly of actin filaments in the Listeria ta il (see Figure 17-17). [Part (a) Courtesy of J. Vic Small.)

cell-surface receptors (Chapter 16) and stimulate cells to move and then to divide. For example, in a wound, blood platelets become activated by being exposed to collagen in the extracellular matrix at the wound edge, which helps the blood to clot. Activated platelets also secrete PDGF to at­tract fibroblasts and epithelial celb to enter the wound and repair it. It is possible to watch part of this process in vitro. If you grow cells in a culture dish and, after starving them of growth factors, you add some fresh growth factor, within a minute or two the cells respond by forming membrane ruf­fles. Membrane ruffles are very similar to the lamellipodia of migrating cells: they are a result of activation of the machin­ery that controls exocytosis of endosomes coupled with actin assembly.

810 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

.·

·.

Scientists knew that growth factors bind to very specific receptors on the cell surface and induce a signal-transduction pathway on the inner surface of the plasma membrane (Chap­ter 15), but how that linked up to the actin machinery was mysterious. Research then revealed that the signal-transduction pathway activates Rae, a member of the small GTPase super­family of Ras-relared proteins (Chapter 15). Rae is one member of a family of proteins that regulate microfilament organiza­tion; two others are Cdc42 and Rho. Unfortunately, due to the history of their discovery, this family of proteins also has been collectively named "Rho proteins," of which Cdc42, Rae, and Rho are members. To understand how these pro­teins work, we first have to recall the way small GTP-binding proteins function.

Like all small GTPases of the Ras superfamily, Cdc42, Rae, and Rho act as molecular switches, inactive in the GOP­bound state and active in the GTP-bound state (Figure 17-42). In their GOP-bound state, they exist free in the cytoplasm in an inactive form bound to a protein known as guanine nucle­otide dissociation inhibitor (GDI). Growth factors can bind and activate their receptors to turn on specific membrane­bound regulatory proteins, guanine nucleotide exchange fac­tors (GEFs), which activate Rho proteins at the membrane by releasing them from GOI and catalyzing the exchange of GOP for GTP. The GTP-bound active Rho protein associates with the plasma membrane, where it binds effector proteins to ini­tiate the biological response. The small GTPase remains active until the GTP is hydrolyzed to GDP, which is stimulated by specific GTPase-activating proteins (GAPs). An important ap­proach to unraveling the functions of Rho proteins has been

Extracellular signal

.""' Exterior '>I

Cytosol TP

~2] Effect~~\ proteins 2.:)

FIGURE 17-42 Regulation of the Rho family of small GTPases. The Rho family of small GTPases are molecular switches regulated by

accessory proteins. Rho proteins exist in the Rho-GOP bound form complexed with a protein known as GDI (guanine nucleotide dissocia­tion inhibitor), which retains them in an inactive state in the cytosol.

Membrane-bound signaling pathwily~ bring Rho proteins to the membrane and, through the action of a GEF (guanine nucleotide

exchange factor), exchange the GOP for GTP, thus activating them. Membrane-bound activated Rho-GTP can then bind effector proteins

that cause changes in the actin cytoskeleton. The Rho protein remains

in the active Rho-GTP state until acted on by a GAP (GTPase-activating protein), which returns it to the cytoplasm. [Adapted from s. Etienne­Manneville and A. Hall, 2002, Nature 420:629.]

to introduce into cells mutant proteins that are locked either in the active-Rho-GTP-state or in the inactive-Rho­GDP-state. A mutant small GTPase that is locked in the active state is said to be a dominant-active protein. Such a dominant-active protein binds the effector molecules constitu­tively, and one can then assess the biological outcomes. Alter­natively, one can introduce a different mutant that is dominant negative, which binds and thereby inhibits the relevant Gil­protein. Thus introduction of a dominant-negative protein in­terferes with the signal-transduction pathway, so one can now assess what processes are blocked.

Cdc42, Rae, and Rho were implicated in regulation of mi­crofilament organizations because introduction of dominant­active mutants had dramatic effects on the actin cytoskeleton, even in the absence of growth factors. It was discovered that dominant-active Cdc42 resulted in the appearance of filopodia, dominant-active Rae resulted in the appearance of membrane ruffles, and dominant-active Rho resulted in the formation of stress fibers that then contracted (Figure 17-43). How can one tell if dominant-active Rae and growth-factor stimulation, both of which stimulate membrane ruffle formation, operate in the

Control

Dominant active Rae

Dominant active Rho

Dominant active Cdc42

.XPERIMENTA FIGURE 17-43 Dominant-active Rae, Rho, and Cdc42 induce different act in-containing structures. To look at the effects of constitutively active Rae, Rho, dnd Cdc42, growth-factor­

starved fibroblasts were microinjected with plasmids to express

dominant-active versions of the three proteins. The cells were then treated with fluorescent phalloidin. which stains filamentous actin.

Dominant-active Rae induces the formation of peripheral membrane ruffles, whereas dominant-active Rho induces abundant contractile stress fibers and dominant active Cdc42 induces filopodia. [From A. Hall,

1998, Setence 279:509.]

17.7 Cell Migration: Mechanism, Signaling, and Chemotaxis 811

FIGURE 17-44 Summary of signal-induced changes in the actin cytoskeleton. Specific signals, such as growth factors and lysophospha­tidic acid (LPA), are detected by cell-surface receptors. Detection leads to the activation of the small GTP-binding proteins, which then interact with effectors to bring about cytoskeletal changes as indicated.

Actin polymerizat ion

Actin polymerization

Myosin activity

Act in polymerization

~ ~ { ~ Filopodia Lamellipodia Stress fiber formation formation formation and contraction

c_=J C) ~ ~ > ~

same signal-transduction pathway? If growth-factor stimula­tion leads to Rae activation, introduction of a dominant-nega­tive Rae protein into a cell should block the ability of a growth factor to induce membrane ruffli ng. This is precisely what is found. Using this and many other biochemical strategies, scien­tists have identified signaling pathways involving Cdc42, Rae, and Rho (Figure 17-44).

Some of the pathways that these proteins regulate con­tain proteins we are familiar with. Activation of Cdc42 stim­ulates actin asse~bly by Arp2/3 through activation of WASp, a nucleation-promoting-factor (NPF) protein (see Figure 17-16), resulting in the formation of filopodia. Activation of Rae also induces Arp2/3, mediated by theW AVE complex, lead­ing to the assembly of branched actin filaments in the lead­ing edge. Activation of Rho has at least two effects. First, it can activate a formin for unbranched actin filament assem­bly. Second, through activation of Rho kinase, it can phos­phorylate the myosin light chain to activate nonmuscle myosin II and also inhibit light-chain dephosphorylation by phosphorylating myosin light-chain phosphatase to inhibit its activity. Both actions of Rho kinase lead to a higher level of myosin light-chain phosphorylation and therefore higher myosin activity and contraction. The three Rho proteins, Cdc42, Rae, and Rho, are also linked by activation and inhi­bition pathways, as shown in Figure 17-44.

Cell Migration Involves the Coordinate Regulation of Cdc42, Rae, and Rho

How does each of these small GTP-binding proteins con­tribute to the regulation of cell migration? To answer this question, researchers developed an in vitro wound-healing

assay (Figure 17-45a). Cells in culture are grown in a petri dish with growth factors, allowing them to grow unti l they are confluent and form a tight monolayer, at which point they stop d ividing. The cell monolayer is then scratched with a needle to remove a swath of cells to generate a "wound" containing a free edge of cells. The cells on the edge sense the loss of their neighbors and, in response to components of the extracellular matrix now exposed on the dish surface, move to fi ll up the empty wound area (figure 17-45b). To do this, they orient themselves toward the free area, first putting out a lamellipodium and then moving in that direc­tion. In this way one can study the induction of directed cell migration in vitro.

Using this system, researchers have introduced dominant­negative Rae into cells on the wound edge to see how it affects the ability of the cells to migrate and fill the wound. Since Rae is needed for activation of the Arp2/3 complex to form the lamellipodium, it is not surprising that the cells fail to form this structure and do not migrate, and so the wound does not close (Figure 17-45c). A very interesting result is obtained when dominant-negative Cdc42 is introduced into the cells at the wound edge: they can form a leading edge but do not ori­ent in the correct direction-in fact, they try to migrate in random directions. This suggests that Cdc42 is critical for regulating the overall polarity of the cell. Studies from yeast (where C.dc42 was first described), wounded-cell monolaycrs, epithelial cells, and neurons reveal that Cdc42 is a master regulator of polarity in many different systems. Part of this regulation in animals involves Cdc42 binding to its effector, Par6, a polarity protein that functions in nematodes (where it was first discovered), neurons, and epithelial cells. We explore these polarity pathways in more detail in Chapter 21.

812 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

Confluent cells Scratch Move into wound

EXPERIMENTAL FIGURE 17-45 The wounded-cell

monolayer assay can be used to dissect signaling pathways in d irected cell movement . (a) A confluent layer of cells is scratched to remove a swath of about three cells wide to generate a free cell border. The cells detect the free space and newly exposed extracellular matrix and, over a period of hours, fill the area. (b) Localization of actin in a wounded monolayer 5 minutes and 3 hours after scratching; the cells have migrated into the wounded area. (c) Effect of introducing domindnt-negatlve Cdc42, Rae, and Hho into cells at the wound edge; all affect wound closure.

(b) (c)

(/) (I)

; c E lD

Dominant negative Rae

Dominant negative Cdc42

Dominant negative Rho

[Parts (b) and (c) from C. D. Nobes and A. Hall, 1999, J. Cell Bioi.

144:1235.]

25 50 75 100 Percent wound closure

Studies such as these suggest a general model of how cell migration is controlled (Figure 17-46). Signals from the environment are transmitted to Cdc42, which orients the cell. The oriented cell has high Rae activity at the front, to induce the formation of the leading edge; Rho activity is high in the rear, to assemble contractile structures and activate the myosin-Il-based contractile machinery. It is important to notice that different regions of the cell can have different levels of active Cdc42, Rae, or Rho, so these regulators are controlled locally within the cell. Part of this spatial regula-

Back: Rho activation ~

Leading to myosin II ----f activation

Contraction of myosin II f ilaments in both stress fibers and cell cortex

Cdc42 Front: Rae activation ~ act ivation

leading to Arp2/3 at the front activation

Actin filament assembly and treadmilling in the leading edge

FIGURE 17-46 Contribution of Cdc42, Rae, and Rho to cell movement. The overall polarity of a migrating cell is controlled by Cdc42, which is activated at the front of a cell. Cdc42 activation leads to active Rae in the front of the cell, which generates the leading edge, and active Rho at the back of the cell, which causes myosin II activation and contraction. Active Rho inhibits Rae activation, ensuring the asymmetry of the two active G-proteins.

tion occurs because some small G proteins can work antago­nistically. For example, active Rho can stimulate pathways that lead to the inactivation of Rae. This might help ensure that no leading-edge structures form at the rear of the cell.

Migrating Cells Are Steered by Chemotactic Molecules

Under certain conditions, extracellular chemical cues guide the locomotion of a cell in a particular direction. In some cases, the movement is guided by insoluble molecules in the underlying substratum, as in the wound-healing assay described above. In other cases, the cell senses soluble molecules and follows them, along a concentration gradient, to their source-a process known as chemotaxis. For example, leukocytes (white blood cells) are guided toward an infection by a tripeptide secreted by many bacterial cells (Figure 17-47a). In another example, dur­ing the development of skeletal muscle, a secreted protein signal called scatter factor guides the migration of myoblasts to the proper locations in limb buds. One of the best-studied exam­ples of chemotaxis is the migration of Dictyostelium amebas during their starvation response. When these soil amebas are stressed, they begin to secrete cAMP, which is an extracellular chemotactic agent in this organism. Other Dictyostelium cells move up the cAMP concentration gradient toward its source (see Figure 17-4 7a). Thus the amebas move toward each other, aggregate into a migratory slug, and then differentiate into a fruiting body in which starvation-resistant spores are formed.

Despite the variety of different chemotactic molecules­sugars, peptides, cell meta bolitcs, cell-wall or membrane lipids-they all work through a common and familiar mecha­nism: binding to cell-surface receptors, activation of intracel­lular signaling pathways, and remodeling of the cytoskeleton through the activation or inhibition of various actin-binding

17.7 Cell Migration: Mechanism, Signaling, and Chemotaxis 813

G) VIDEO: Chemotaxis of a Single Dictyostefium Cell to cAMP

(a) (b) (c)

Receptors c::) cAMP~ G-protein c::)

PI-3K C) PTEN c:)

PI(3,4,5)P3 C)

KPE E A

Dictyostelium amebae

migrating to cAMP

G RE

Human neutrophils migrating to fMLP

Chemotaxis involves elevated levels of signaling phosphoinositides, which signal to the actin cytoskeleton. (a) Dictyostelium cells migrate toward a pipet of cAMP (left), and human neutrophils (a type of leukocyte) migrate toward a pipet of fMLP (formylated Met-Leu-Phe), a chemotactic peptide produced by bacteria (right). In the lower two panels are individual chemotaxing Dictyoste/ium and neutrophil cells that look remarkably similar, despite about 800 million years of evolution separating them. (b) Summary of results of studies exploring the localization of

proteins. What is quite amazing is that just a 2 percent differ­ence in the concentration of chemotactic molecules between the front and back of the cell is sufficient to induce directed cell migration. Equally amazing is the finding that the internal signal-transduction pathways used in chemotaxis have been conserved between Dictyostelium amebas and human leuko­cytes despite almost a billion years of evolution.

Chemotactic Gradients Induce Altered Phosphoinositide levels Between the Front and Back of a Cell To investigate how Dzctyostelium amebas sense a chemotactic gradient, investigators have studied the cell-surface receptors for extracellular cAMP and downstream signaling pathways in the expectation that these must somehow sense the concen­tration gradient. Before we discuss the details, let's consider how such a system might work. If a cell can sense a 2 percent difference in concentration across its length, it is unlikely that simply activating actin assembly 2 percent more at the front than at the back could lead to directed movement. Rather, there must be some mechanism that amplifies this small exter­nal signal difference mto a large internal biochemical differ­ence. One way to do this would be for the cell to subtract the average signal from the front and back and only respond to a difference m signal. It is believed that this is how the system works. To try to understand this mechanism, investigators have looked at the concentration of active components of the signaling pathway to see where the amplification occurs.

Micrographs of cAMP receptors tagged with green fluo­rescent protein (GFP) show that the receptors are distributed

Actin C) Myosin II c:)

: =• Chemotactic gradient

Lipid phosphatase

(PTEN)

Lipid kinase (PI-3K)

components of signaling pathways (green) in Dictyostelium cells undergoing chemotaxis toward cAMP. Also shown are the localization of actin and myosin (red). (c) The enzyme Pl-3 kinase, which generates PI(3.4,S)P3, is enriched at the front of chemotaxing cells, whereas PTEN, the phosphatase that hydrolyses PI(3.4,5)P3, is enriched at the back. These distributions result in elevated PI(3,4,5)P3 at the front of the cells, which signals the polarity for movement. [Part (a) from C. Parent, 2004, Curr. Opin. Cell Bioi. 16:4; part (c) from M. lijima et al., 2002, Dev. Cell 3:469.)

uniformly on the surface of an ameba cell (Figure 17-47b); therefore an internal gradient must be established by another component of the signaling pathway. Because cAMP recep­tors signal through trimeric G proteins (Chapter 16), a sub­unit of the trimeric G protein and other downstream signaling proteins were tagged with GFP to look at their distributions. Fluorescence micrographs show that the concentration of rrimeric G proteins is also rather uniform. Downstream of the rrimeric G proteins is Pl-3 kinase, an enzyme that phos­phorylares membrane-bound inositol phospholipids (phos­phoinositides), such as PI4,5-biphosphare [PI(4,5)P2],

creating the signaling lipid PI3,4,5-rriphosphare [Pl(3,4,5)Pd (see Figure 16-25). Remarkably, the enzyme PI-3 kinase is highly enriched at the front of a migrating cell, as are its products. PTEN, the phosphatase that dephosphorylates the signaling lipid PI(3,4,5)P3 back to PI(4,5)P2 , is enriched in the rail of the migrating cell (Figure 17-4 7b, c). This asym­metry is believed to be established in the following way. Prior to the cell's exposure to a cAMP gradient, the phos­phatase PTEN is associated uniformly with the plasma mem­brane. When a cell "sees" a gradient, PI 3-kinase is activated a bit more at the front than the back. This results in slightly higher levels of the signaling phospholipid at the front. The association of the phosphatase PTFN with the memhrane is very sensitive to the level of PI(3,4,5)P3, so it is preferentially depleted from the front. Since it is less effective at dephos­phorylating the PI(3,4,5)P3 at the front and more effective at dephosphorylating PI(3,4,5)P3 at the rear, a strong asymme­try of Pl(3,4,5)P3 results. Thus the phosphatase PTEN con­tributes to the background subtraction necessary for a cell to

sense a shallow gradient of chemoattractant.

814 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

·.

.·

The difference in local PI(3,4,5)P3 concentration now sig­nals to the actin cytoskeleton to assemble a leading edge at the front and contraction at the rear (Figure 17-4 7b), and the cell is on its way to the source of chemoattractant. A very similar mechanism has been implicated in the chemotaxis of leuko­cytes. This cell polarization is not stable in the absence of the chemotactic gradient, so if the gradient changes, as might hap­pen with a leukocyte chasing a moving bacterium, the cell will also change its direction and follow the gradient to its source.

l(fV CONCEPTS of SP('tion 17.7

Cell Migration: Mechanism, Signaling, and Chemotaxis

Cell migration involves the extension of an actin-rich lead­ing edge at the front of the cell, the formation of adhesive contacts that move backward with respect to the cell, and their subsequent release, combined with rear contraction to push the cell forward (see Figure 17-40).

Cell migration also involves a directed endocytic cycle, taking membrane and adhesion molecules from the rear of the cell and inserting them at the front.

The assembly and function of actin filaments is controlled by signaling pathways through small GTP-binding proteins of the Rho family. Cdc42 regulates overall polarity and the formation of filopodia, Rae regulates actin meshwork for­mation through the Arp2/3 complex, and Rho regulates both actin filament formation by formins as well as contrac­tion through regulation of myosin II (see Figure 17-44).

Chemotaxis, the directed movement toward an attractant, involves signaling pathways that establish differences in phosphoinositides between the front and rear of the cell, which in turn regulate the actin cytoskeleton and direction of cell migration (see Figure 17-47).

Perspectives for the Future

In this chapter, we have seen that cells have intricate mecha­nisms for the regulated spatial and temporal assembly and turnover of microfilameqts to perform their many functions. Biochemical analyses of actin-binding proteins coupled with protein inventories provided by sequences of whole gcnomes have allowed the catalogjng of many different classes of actin­binding proteins. To understand how this large group of pro­teins can assemble specific structures in a cell, it will be important to know the concentration of all the components, how they interact, and their regulation by signaling pathways. Although this might seem like a daunting task, new microscopic methods to detect the location of specific protein-protein inter­actions and the location of many of the key signaling pathways suggest that rapid progress will be made in this area.

The protein inventories provided by genomic sequences have also documented the large number of myosin families, yet the biochemical properties of many of these motors, or their biological functions, remain to be elucidated. Again, recent

technical developments, including the ability to tag motors with fluorescent tracers such as GFP, or knocking down their expression with RNAi technologies are providing very power­ful avenues to help reveal motor functions. However, some important aspects of motors remain largely unexplored. For example, a motor that transports an organelle down a filament first has to bind the organelle, then transport it, and then re­lease it at the destination. However, little is known about how these different events are coordinated or how these types of myosin-based motors are returned to pick up new cargo.

Twenty years ago it was believed that all actin assembly was driven by activation of the Arp2/3 complex. Then the nucleating and capping activities of form ins were discovered, and more recently, additional actin nucleators, with colorful names such as Spire, Cordon-bleu, WASH, and WHAMM have been discovered. It is likely that additional nucleators will continue to be discovered.

Finally, although we have generally discussed microfila­ments without regard to tissue type-except for the special­izations found in skeletal and smooth muscles-many actin-binding proteins show cell-type-specific expression, and so the array and relative levels of these protein!. are tai­lored to specific functions of different cell types. This is clearly the case, as revealed by proteomic analysis of cell-specific pro­tein expression and by the fact that many diseases arc a con­sequence of tissue-specific expression of actin-binding proteins or myosins.

Key Terms actin 775

actin cross-linking proteins 790

Arp2/3 complex 784

CapZ protein 783

Cdc42 protein 811

cell migration 808

cell polarity 774

chemotaxis 813

cofilin 782

contractile bundles 804

critical concentration, Cc 779

cytoskeleton 774

duty ratio 799 F-actin 777

filopodia 776 formin 784

G-actin 777

intermediate filaments 775

lamellipodium 776

leading edge 776

microfilaments 775

microtubules 775

microvilli 773

motor protein 775

myosin 793

myosin light-chain kinase 805

nucleation promoting factor (NPF) 785

power stroke 797

profilin 782

Rae protein 811

Rho protein 811

skeletal muscle 793

smooth muscle 804

step size 799

stress fibers 776

thick filaments 801

thin filaments 801

thymosin ~4 783

treadmilling 781

tropomodulin 783

tropomyosin 803

WASp protein 785

Key Terms 815

Review the Concepts

1. Three systems of cytoskeletal filaments exist in most eu­karyotic cells. Compare them in terms of composition, func­tion, and structure.

2. Actin filaments have a defined polarity. What is filament polarity? How is it generated at the subunit level? How is filament polarity detectable?

3. In cells, actin filaments form bundles and/or networks. Huw do cells form such structures, and what specifically determines whether actin filaments will form a bundle or a network?

4. Much of our understanding of actin assembly in the cell is derived from experiments using purified actin in vitro. What techniques can be used to study actin assembly in vitro? Explain how each of these techniques works. Which of these techniques would tell you whether the mass of actin filaments is comprised of many short actin filaments or fewer longer filaments?

5. The predominant forms of actin inside a cell are ATP-G­actin and ADP-F-actin. Explain how the interconversion of the nucleotide state is coupled to the assembly and disassem­bly of actin subunits. What would be the consequence for actin filament assembly/disassembly if a mutation prevented actin's ability to bind ATP? What would be the consequence if a mutation prevented actin's ability to hydrolyze ATP?

6. Actin filaments at the leading edge of a crawling cell are believed to undergo treadmilling. What is treadmilling, and what accounts for this assembly behavior?

7. Although purified actin can assemble reversibly in vitro, various actin-binding proteins regulate the assembly of actin filaments in the cell. Predict the effect on a cell's actin cyto­skeleton if function-blocking antibodies against each of the following are independently microinjected into cells: pro­filin, thymosin-~4, CapZ, and the Arp2/3 complex.

8. Predict how ·actin would polymerize on an arrowhead decorated seed (as shown in Figure 17 -9) in the presence of CapZ, tropomodulin, or profilin-actin.

9. Compare and contrast the ways in which formin and WASp arc activated, and how each stimulates actin filament formation.

10. There are at least 20 different types of myosin. What properties do all types share, and what makes them differ­ent? Why is myosin II the only myosin capable of producing contractile force?

11. The ability of myosin to walk along an actin filament may be observed with the aid of an appropriately equipped microscope. Describe how such assays are typically per­formed. Why is ATP required in these assays? How can such assays be used to determine the direction of myosin move­ment or the force produced by myosin?

12. Contractile bunJh:~ occur in nonmuscle cells; these structures are less organized than the sarcomeres of muscle cells. What is the purpose of nonmuscle contractile bundles? Which type of myosin is found in contractile bundles?

13. How does myosin convert the chemical energy released by ATP hydrolysis into mechanical work?

14. Myosin II has a duty ratio of 10 percent, and its step size is 8 nm. In contrast, myosin V has a much higher duty ratio (about 70 percent) and takes 36-nm steps as it walks down an actin filament. What differences between myosin II and myosin V account for their different properties? How do the different structures and properties of myosin ll and myosin V reflect their different functions in cells?

15. Contraction of both skeletal and smooth muscle is trig­gered by an increase in cytosolic Cah. Compare:: rhe mecha­nisms by which each type of muscle converts a rise in Cal+ into contraction.

16. Phosphorylation of myosin light-chain kinase (MLCK) hy protein kinase A (PKA) inhibits MLCK activation by Ca2 -calmodulin. Drugs such as albuterol bind to ~-adrenergic receptor, which cause~ a rise in cAMP in cells and activation of PKA activity. Explain why albuterol is useful for treating the severe contraction of the smooth muscle cells surrounding airway passages involved in an asthma attack.

17. Several types of cells utilize the actin cytoskeleton to power locomotion across surfaces. How are different assem­blies of actin filaments involved in locomotion?

18. To move in a specific direction, migrating cells must uti­lize extracellular cues to establish which portLOn of the cell will act as the front and which will act as the back. Describe how G proteins appear to be involved in the signaling pathways used by migrating cells to determine direction of movement.

19. Cell motility has been described as being like the motion of tank treads. At the leading edge, actin filaments form rapidly into bundles and networks that make protrusions and move the cell forward. At the rear, cell attachments are broken and the tail end of the cell is brought forward. What provides the trac­tion for moving cells? How does cell-body translocation hap­pen? How are cell attachments released as cells move forward?

Analyze the Data

Myosin Vis an abundant nonmuscle myosin that is responsi­ble for the transport of cargo such as organelles in many cell types. Structurally, it consists of two identical polypeptide chains that dimerize to form a homodimer. The motor do­mains reside at theN-terminus of each chain and contain both ATP- and actin-binding sites. The motor domain is followed by a neck region containing six "IQ" motifs, each of which hinds calmodulin, a Ca2+ -binding protein. The neck domain is followed by a region capable of forming coiled coils, via which the two chains dimerize. The final 400 amino acid residues form a globular tail domain (GTD), to which cargo binds. Myosin V would consume large amounts of ATP if its motor domain were always active, and a number of studies have been conducted to understand how this motor is regulated.

a. The rate of ATP hydrolysis (i.e., ATP molecules hydro­lyzed per second per myosin V) was measured in the presence of increasing amounts of free Ca2

. The concentration of cyto­solic free Ca2

+ is normally less than 10 6 M but can be ele­vated in localized areas of the cell and is often elevated in

816 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

response to a signaling event. What do these data s uggest

about myosin V regulation?

12 I [

• •• • , a..(/) Sf- -1- ·- • <t~ --o e • <1>"0 ... > f2.r. 4f- • -

• • • .. ·- .. , I I 0

10 8 10 6 10-4

!Ca2+J. M

b. In additional studies, the ATPase activity of myosin V was measured in the presence of increasing amounts ofF

actin in the presence or absence of 10 6 M free Ca2 • What

additional information about myosin V regulation do these

data provide?

a.. (/) 1- ·­<t~ --oe <1>"0 ... > f2.r.

16.----------------------------.

, 0

+Ca2+ ----

- Ca2+ ------

,---------------------5 10 15

[Actinl.).JM 20

c. Next, the behavior of truncate d m yosin V, which

lacks just its C-terminal globular tai l, was examined and

compared to the behavior of intact m yosin V. From this ex­periment, what ca n you d e duce about the mechanism by which myosin V is regulqted?

16.----------------------------.

12

4

, , , , , ,'

,~

0 5

-------------­~~-

+Ca2~ ---

- Ca2+ ------

10 15 20 [Acti n], ).1M

References General References

Bra), D. 200 I . Cell MoL'ements. Garland. Howard, J. 2001. The Mechamcs of Motor Proteins and the

Cytoskeleton. Sinauer. Kreis, T., and R. Vale. 1999. Guidebook to the Cytoskeletal and

Motor Proteins. Oxford Umversiry Pre\s.

Web Sites

The myosin home page: hrrp://www .mrc-lm b.cam.ac. u k/myosin/myosin.html

The kinesin home page: hrrp://www.cellbio.duke.edu/kinesin/

Microfilaments and Actin Structures

Holmes, K. C., et al. 1990. Atomic model of the actin filament. Nature 347:44-49.

Kabsch, W., et al. 1990. Atomic structure of the actin:DNase l complex. Nature 347:37-44.

Pollard, T. D., and J. A. Cooper. 2009. Aeon, a central player 111

cell shape and movement. Sctence 326:1208- 1212. Pollard, T. D., L Blanchoin, and R. D. Mullins. 2000. \llolecu­

lar mechamsms controlling actin filament dynamtcs 111 nonmuscle cells. Ann. Rev. Bwphys. Btomol. Struc. 29:545-576.

Dynamics of Actin Filaments

Paavilainen, V. 0 ., et al. 2004. Regulation of cytoskeletal dynamics b) actin-monomer-binding proteins. Trends Cell B10/. 14:386-394.

Thenot, J. A. !997. Accelera ting on a treadm1ll: ADF/cofihn promotes rapid actin fi lament turnover in the dynamic cytoskeleton. ]. Cell Bml. 136: 1165-1168.

Mechanisms of Actin Filament Assembly

Campellone, K. G., and M. D. Welch. 2010. A nucleator arms race: cellular control of actin assembly. Nat. Reu. Mol. Cell Bml. 11:237-251.

Chesarone, M. A., et al. 2010. Unleashing formins to remodel the actm and microtubule cyroskelerons. Nat. Reu. Mol. Cell Bioi. 11:62-74.

Goode, B. L., and M. J. Eck. 2007. Mechanism and function of fo rmins in the control of actin assembly. Ann. Ret•. Biochem. 76:593-627.

Gouin, E., M. D. Welch, and P. Cossart. 2005. Actin-based motility of intracellular pathogens. Curr. Opin. Mtcrobiol. 8:35-45.

Higgs, H . N. 2005. Formm proteins: a domain-based approach. Trends Biochem. Sci. 30:342-353.

Pruyne, D., et al. 2002. Role of formins in actin assembly: nucleation and barbed end association. Sctence 297:612-615.

Rouiller, I., et al. 2008. The structural basis of actin filament branching by rhe Arp2/3 complex.]. Cell Bioi. 180:887-895.

Organization of Actin-Based Cellular Structures

Bennett, V., and A.]. Baines. 2001. Spectrin and ankynn-based pathways: metazoan inventions for integrating cells into tissues. Pbysiol. Rev. 81:1353-1 392.

Fehon, R.G., A. I. McClatchey, and A. Bretscher. 2010. Organizing the cell cortex: the role of ERM proteins. Nat. Rev. Mol. Cell Bioi. 11:276-287.

References 817

McGough, A. 1998. f-actin-bindmg proteins. Curr. Opin. Struc. Bwl. 8:166-176.

Srossel, T. P., et al. 200 l. Filamms as mtegrators of cell mechanics and signalling. Nat. Rev. Mol. Cell Bioi. 2:138-145.

Myosins: Actin-Based Motor Proteins

Berg, J. S., B. C. Powell, and R. E. Cheney. 200 I. A millennia! myosin census. Mol. Bioi. Cel/12:780-7 94.

,\lermall, V., P. L. Post, and M.S. Mooseker. 1998. Unconven­tional myosin in cell movement, membrane traffic, and s1gnal transduction. SCience 279:527-533.

Rayment, I. 1996. The structural bas1s of the myosin ATPase activity.]. Brol. Chern. 271:15850-15853.

Vale, R. D. 2003. The molecular motor toolbox for mtracellular transport. Cel/112:467-480.

Vale, R. D., and R. A . .Milligan. 2000. The way things move: looking under the hood of molecular motor proteins. Scrence 288:88-95.

Myosin-Powered Movements

Bretscher A. 2003 . Polarized growth and organelle segregation in yeast-the tracks, motors, and receptors.]. Cell Bwl. 160:811-816.

Clark, K. A., et al. 2002. Stnated muscle cytoarchitecture: an intricate web of form and function. Ann. Rev. Cell De[/. Bwl. 18:637-706.

Graz1er, H. L., and S. Labeit. 2004. The giant protein min: a major player in myocardial mechanics, signaling, and disease. Circ. Res. 94:284-295.

Cell Migration: Signaling and Chemotaxis

Bonsy, G. G., and T. ~1. Svitkina. 2000. Acnn machinery: pushing the envelope. C:urr. Opin. Cell Bioi. 12: I 04 I 12.

Bumdge, K., and K. Wennerberg. 2004. Rho and Rae take cenrer stage. Cel/116:167-179.

Etienne-Jvlanneville, S. 2004. Cdc42-the centre of polarity. ]. Cell Sci. 117:1291-1300.

Etienne-~1annevdle, S., and A. Hall. 2002. Rho GTPascs in cell biology. Nature 420:629-63'5.

Manahan, C. L., et al. 2004. Chemoattractant signaling in Drctyostelrum discoidewn. Ann. Rev. Cell Dev. Brol. 20:223-253 .

Pollard, T. D., and G. G. Borisy. 2903 . Cellular motility driven by assembly and disassembly of actin filaments. Cell 112:453-465.

Ridley, A. J., et al. 2003. Cell migration: integrating signals from the front to back. Science 302:1704-1709.

Small, j. V., T. Strada, E. Vigna!, and K. Rottner. 2002. The larnellipodium: where motility begins. Trends Cell Bioi. 12:112-120.

818 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

·.

Looking at Muscle Contraction H. Huxley and J. Hanson, 1954, Nature 173:973-976

The contraction and relaxation of stria ted muscles allow us ro per­

form all of our daily tasks. How does this happen? Scientists have long looked to see how fused muscle cells, called myofibrils, differ from other cells that cannot perform powerful movement. In 1954, Jean Hanson and Hugh Huxley published their micros­copy studies on muscle contraction, which demonstrated the mechanism by which it occurs.

Background

The ability of muscles to perform work has long been a fascinating process. Voluntary muscle contraction is per­formed by striated muscles, which are named for their appearance when viewed under the microscope. By the 1950s, biologists studying myofibrils had named many of the structures they observed under the microscope. One contracting unit, called a sarcomere, is made up of two main regions called the A band and the I band. The A band contains two darkly colored thick stria­tions and one thin striation. The I band is made up pri~arily of light-colored striations, which are divided by a darkly colored line known as the Z disk . Although these structures had been characterized, their role in muscle contraction remained unclear. At the same time, biochemists ,also tried to tackle this problem by looking for pro­teins that are more abundant in myofi­brils than in nonmuscle cells. They found muscles ro contain large amounts of the structural proteins actin and my­osin in a complex with each other. Actin and myosin form polymers that can shorten when treated with adeno­sine triphosphate (ATP).

With these observations in mind, Hanson and Huxley began their study of cross striations in muscle. In a few short years, they united the biochemical

data with the microscopy observations and developed a model for muscle con traction that still holds true today.

The Experiment

Hanson and Huxley primarily used phase-contrast microscopy in their studies of striated muscles that they isolated from rabbits. The technique allowed them to obtain clear pictures of the sarcomere and to take careful measurements of the A and the I bands. By treating the muscles with a variety of chemicals, then studying them under the phase-contrast microscope, they were able to successfully combine bio­chemistry with microscopy to describe muscle structure as well as the mecha­nism of contraction.

In their first set of studies, Hanson and Huxley employed chemicals that are known to specifically extract either myosin or actin from myofibrils. First, they treated myofibrils with a chemical that specifically removes myosin from muscle. They used phase-contrast mi­croscopy to compare untreated myofi­brils to myosin-extracted myofibrils. In the untreated muscle, they observed the previously identified sarcomeric struc­ture, including the darkly colored A band. When they looked at the myosin­extracted cells, however, the darkly col­ored A band was not observed. Next, they extracted actin from the myosin­extracted muscle cells. When they ex­tracted both myosin and actin from the myofibril, they could see no identifiable structure to the cell under phase-contrast microscopy. From these experiments, they concluded that myosin was located primarily in the A band, whereas actin is found throughout the myofibril.

With a better understanding of the biochemical nature of muscle struc­tures, Huxley and Hanson went on to study the mechanism of muscle con­traction. They isolated individual myo-

CLASSIC EXPERIMENT 17.1

fibrils from muscle tissue and treated them with ATP, causing them to con­tract at a slow rate. Using this tech­nique, they could take pictures of various stages of muscle contraction observed using phase-contrast micros­copy. They could also mechanically in­duce stretching by manipulating the coverslip, which allowed them to also observe the relaxation process. With these techniques in hand, they exam­ined how the structure of the myofibril changes during contraction and stretch.

First, Huxley and Hanson treated myofibrils with A TP, then photo­graphed the images they observed under phase-contrast microscopy. These pictures allowed them to mea­sure the lengths of both the A band and the I band at various stages of con­traction. When they looked at myofi­brils freely contracting, they noticed a consistent shortening of the lightly col­ored I band, whereas the length of the A band remained constant (F1gure 1). Within the A band, they observed the formation of an increasingly dense area throughout the contraction.

Next, the two scientists examined how the myofibril structure changes during a simulated muscle stretch. They stretched isolated myofibrils mounted on glass slides by manipulating the coverslip. They again photographed phase-contrast microscopy images and measured the lengths of the A and the I bands. During stretch the length of the I band increased, rather than shortened, as it had in contraction. Once again, the length of the A band remained un­changed. The dense zone that formed in the A band during contraction became less dense during stretch.

From their observations, Hanson and Huxley developed a model for muscle contraction and stretch (Fig­ure 1 ). In their model, the actin fila­ments in the I band are drawn up into the A band during contraction, and thus

Looking at Muscle Contraction 819

--I band~~ A band ----7+---- I band -FIGURE 1 Schematic diagram of muscle contraction and stretch observed by Hanson and Huxley. The lengths ofthe sarcomere (5), the A band (A), and the I band (I) were

measured in muscle samples contracted 60 percent in length relative to the relaxed muscle (bottom) or stretched to

Zd;,k~ St;~~;0ed : : i Relaxed :1: ~ i 100%

Contracted :1: a; 90%

~ E=l: ii¢

s 2.8J1 A 1.5J1 I 1.3J1

s 2.3J1 A 1.5J1 I 0.8J1

s 2.0J1 A 1.5J1 I 0.5p

120 percent (top). The lengths ofthe sarcomere, the I band, and the A band are noted on the right. Notice that from

120 percent stretch to 60 percent contraction the A band

does not change in length. However, the length of the I band can stretch to 1.3 J..Lm, and at 60 percent contraction, it disappears as the sarcomere shortens to the overall length

of the A band. [Adapted from J. Hanson and H. E. Huxley, 1955,

Symp. Soc. Exp. Bioi. Fibrous Proteins and Their Biological Significance

9:249.]

Contracted :13!. =·iii: I: s 1.8J1 80% A 1.5p

I 0.3J1

Contracted :IM5ii~ s 1.5J1

60% A 1.5J1 I O.OJ1

the I band becomes shorter. This al­lows for increased interaction between the myosin located in the A band and the actin filaments. As the muscle stretches, the actin filaments withdraw from the A band. From these data, Hanson and Huxley proposed that muscle contraction is driven by actin filaments moving in and out of a mass of stationary myosin filaments.

Discussion

By combining microscopic observa­tions with known biochemica l treat­ments of muscle fibers, Hanson and Huxley were able to describe the bio­chemical nature of muscle structures and outline a mechanism for muscle contraction. A large body of research continues to focus on understanding

820 CHAPTER 17 • Cell Organization and Movement 1: Microfilaments

the process of muscle contraction. Scientists now know that muscles contract by A TP hydrolysis, driving a conformational change in myosin that allows it to pull on actin. Re­searchers are continuing to uncover the molecular details of this process, whereas the mechanism of contrac­tion proposed by Hanson and Huxley remains in place.

·.