Cytoskeleton PDF

A cell in culture has been fixed and labeled to show its cytoplasmic arrays of microtubules (green) and actin filaments (red). This dividing cell has been labeled to show its spindle microtubules (green) and surrounding cage of intermediate cell cortex MICROTUBULES...

A cell in culture has been fixed and labeled to show its cytoplasmic arrays of microtubules (green) and actin filaments (red). This dividing cell has been labeled to show its spindle microtubules (green) and surrounding cage of intermediate cell cortex MICROTUBULES 100 n m ili 25 n m Microtubules are long, hollow cylinders made of the protein tubulin. With an outer diameter of 25 nm, they are much more rigid than actin filaments. Microtubules are long and straight and frequently have one end attached to a microtubule-organizing center (MTOC) called a centrosome. (i) Single microtubule; (ii) cross section at the base of three cilia showing triplet microtubules; (iti) interphase microtubule array (green) and organelles (red); (iv) Micrographs courtesy of R. Wade (i); DT.. Woodrow and RW.. Linck (ii);.J Seemann (iii); ciliated protozoan.. Burnette D i (v)  INTERMEDIATE FILAMENTS 100 mm 25 nm Intermediate filaments are ropelike fibers with a diameter of about 10 nm; they are made of intermediate filament proteins, which constitute a large and heterogeneous family. One type of intermediate filament forms a meshwork called the nuclear lamina just beneath the inner nuclear membrane. Other types extend across the cytoplasm, giving cells mechanical strength. In an epithelial tissue, they span the cytoplasm from one cell-cell junction to another, thereby strengthening the entire epithelium. (i) Individual intermediate filaments; (ii) Intermediate filaments (blue) in neurons and (ili) epithelial cell; (iv) nuclear lamina. Large-scale cytoskeletal structures can change or persist, according to need, lasting for lengths of time ranging from +e\\ea2a12jeejUe:eece++\+ee1eBjeee2d}dja+ macromolecular components that make up these structures are in a constant state of flux. As a result, a structural rearrangement in a cell requires little extra energy when conditions change. For example, actin filaments form many types of cell- surface projections. Some of these are dynamic structures, such as the filopodia, lamellipodia, and pseudopodia that cells use to explore territory and move around. Motility is initiated by an actin-dependent protrusion of the leading edge, which is composed of lamellipodia and filopodia. These protrusive structures contain actin filaments, with elongating barbed ends orientated towards the plasma membrane time 0 min 1 min 2 min 3 min From a video recorded by David Rogers 15 um A neutrophil in pursuit of bacteria. In this preparation of human blood, a small clump of bacteria (white arrow) is about to be captured by a neutrophil. As the bacteria move, the neutrophil quickly reassembles the dense actin network within a pseudopod ("false foot") at its leading edge (highlighted in red) to push toward the location of the bacteria (Movie 16.1). Rapid disassembly and reassembly of the actin cytoskeleton ni this cell enable ti to change its orientation and direction of movement within a few minutes. (From a video recorded by David Rogers.) The regulation of actin filament formation is an important mechanism by which cells control their shape and movement. Actin subunits can spontaneously bind one another, but the association is unstable until subunits assemble into an initial oligomer, or nucleus, that is stabilized by multiple subunit subunit contacts and can then elongate rapidly by addition of more subunits. This process is called filament nucleation. N U C L E AT I O N A helical polymer is stabilized by multiple contacts between adjacent subunits. In the case of actin, two actin molecules bind relatively weakly to each other, but addition of a third actin monomer to form a trimer makes the entire group m o r e stable. Further monomer addition can take place onto this trimer, which therefore acts as a nucleus for polymerization. For tubulin, the nucleus is larger and has a more complicated structure (possibly a ring of 13 or more tubulin molecules) —but the principle is t h e same. The assembly of a nucleus is relatively slow, which explains the lag phase seen during polymerization. The lag phase can be reduced or abolished entirely by adding premade nuclei, such as fragments of already polymerized micro tubule s or actin filame nts. TIME COURSE OF POLYMERIZATION The ni vitro assembly of a protein into a long polymer such as a cytoskeletal filament typically shows the following time course: EQUILIBRIUM PHASE amo unt of polymer GROWTH LAG PHASE PHASE time — The lag phase corresponds ot time taken for nucleation. The growth phase occurs as monomers add to the exposed ends of the growing filament, causing filament elongation. The equilibrium phase, or steady state, si reached when the growth of the polymer due to monomer addition precisely balances the shrinkage of the polymer due to disassembly back to monomers. PLUS A N D M I N U S E N D S The two ends of an actin filament or microtubule polymerize at different rates. The fast-growing end is called the plus end, minus plus whereas the slow-growing end si called the minus end. The end end difference in the rates of growth at the two ends is made possible by changes in the conformation of each subunit as SLOW FAST it enters the polymer. free subu nit in subunit polymer This conformational change affects the rates at which subunits add to the two ends. loss, which determines the equilibrium constant for its association Even though kon and kof wil have different values for the plus and with the end, is identical at both ends: if the plus end grows four minus ends of the polymer, their ratio Koffkon-and hence C-must be times faster than the minus end, it must also shrink four times the same at both ends for a simple polymerization reaction (no ATP or faster. Thus, for C> Ca both ends grow; for C< C, both ends GTP hydrolysis). This si because exactly the same subunit interactions shrink are broken when a subunit is lost at either end, and the final state of The nucleoside triphosphate hydrolysis that accompanies the subunit after dissociation si identical. Therefore, the AG for subunit actin and tubulin polymerization removes this constraint. NUCLEOTIDE HYDROLYSIS Each actin molecule carries a tightly bound ATP molecule that is hydrolyzed to a tightly bound ADP molecule soon after its assembly into the polymer. Similarly, each tubulin molecule carries a tightly bound GTP molecule that si converted to a tightly bound GDP molecule soon after the molecule assembles into the polymer. T( = monomer carrying ATP or G T P T T D (D = m o n o m e r carrying ADP free m o n o m e r subunit in polymer or G D P Hydrolysis of the bound nucleotide reduces the binding affinity of the subunit for neighboring subunits and makes it more likely to dissociate from each end of the filament. It is usually the T form that adds to the filament and the D form that leaves. Considering events at the plus end only: K T on D D D D D T D D D D D D off As before, the polymer will grow until C= C. For illustrative purposes, we can ignore T on and k off since they are usually very small, so that polymer growth ceases when K D off KoonnC= kf、 o =ko r =C kkonon This is a steady state and not a true equilibrium, because the ATP or GTP that is hydrolyzed must be replenished by a nucleotide exchange reaction of the free subunit ( D → T ).  DYNAMIC INSTABILITY a n d TREADMILLING a r e t w o b e h a v i o r s observed in cytoskeletal polymers. Both are associated with nucleoside triphosphate hydrolysis. Dynamic instability is believed to predominate in microtubules, whereas treadmilling may predominate in actin filaments. TREADMILLING One consequence of the nucleotide hydrolysis that accompanies polymer formation is to change the critical concentration at the two ends of the polymer. Because KDoff and KTon refer to different reactions, their ratio on need not be the same at both ends of the polymer, so that: C (minus end) >cC (plus end) Thus, fi both ends of a polymer are exposed, polymerization proceeds until the concentration of free monomer reaches a value that si above C, for the plus end but below C, for the minus end. At this steady state, subunits undergo a net assembly at the plus end and a net disassembly at the minus end at an identical rate. The polymer maintains a constant length, even though there is a net flux of subunits through the polymer, known as treadmilling. Subunits with bound ATP (T-form subunits) polymerize at both ends of a growing filament and then undergo ATP hydrolysis within the filament. As the filament grows, elongation is faster than hydrolysis at the plus end in this example, and the terminal subunits at this end are therefore always in the T form. However, hydrolysis is faster than elongation at the minus end, and so terminal subunits at this end are in the D form. Treadmilling occurs at intermediate concentrations of free subunits. The critical concentration for polymerization on a filament end in the T form, Cc(T), is lower than that for a filament end in the D form, Cc(D). If the actual subunit concentration is somewhere between these two values, the plus end grows while the minus end shrinks, resulting in treadmilling.  The Functions of Actin Filaments Are Inhibited by Both Polymer-stabilizing and Polymer-destabilizing Chemicals TA B L E 1 6 - Chemical Inhibitors of Actin and Microtubules Effe ct o n Chemical fi l a m e n t s Mechanism Original source Actin Latrunc ulin Depolymerizes Binds actin subunits Spo nge s Cytochalasin B Depolymerizes Caps filament plus ends Fungi Phalloidin Stabilizes Binds along filaments Amanita mushroom Microtubules Taxol (paclitaxel) Stabilizes Binds along filaments Yew tree Nocodazole Depolymerizes Binds tubulin subunits Synthetic Colchicine Depolymerizes Caps both filament ends Autumn crocus ACTIN FILAMENTS + formin Arp2/3 complex nucleates assembly and remains nucleates assembly to form associated with the growing a branched network and remains plus end associated with the minus end thymosin binds subunits, profilin prevents assembly bin ds mo nom ers , + concentrates them at sites of filament assembly tropomodulin actin s u b u n i t s prevents assembly and disassembly at minus end ACTIN FILAMENTS + a c t i n fi l a m e n t + tropomyosin stabilizes filament, modulates cofilin binding of other accessory proteins binds ADP-actin filaments, accelerates disassembly gelsolin capping protein severs filaments and binds to plus end prevents assembly and disassembly at plus end filament bundling, cross-linking, and attachment to membranes plasma fi m b r i n mem bran e a-actinin filamin spectrin ERM Some of the major accessory proteins of the actin cytoskeleton. Except for the myosin motor proteins, an example of each major type is shown. Each of these is discussed in the text. However, most cells contain more than a hundred different actin-binding proteins, and ti is likely that there are important types of actin-associated proteins that are not yet recognized.  plus end plus end p l us e n d D, from.I Rouiller et al., J. Cell Biol. 180:887-895, 2008. © 2008A. Rouiller et al. Originally published ni J. Cel Biol. 180:887-895. https://doi.org/10.1083/jcb.200709092. With permission from Rockefeller (A) actin [*] Arp2 nucleation-promoting factor (NPF) other minus plus proteins end end + Arp3 Arp2 inactive actin n u c l e a t e d actin fi l a m e n t Arp2/3 complex active Arp2/3 complex monomers (B) (D) 25 n m daught er actin fi l a m e n t 70° accessory 1-7- proteins University Press. preexisting actin fi l a m e n t (E) (C) Once nucleated, rapid polymerization of actin depends on the further addition of actin monomers at the plus end of each filament. A key factor is the monomer-binding protein profilin. Profilin binds to the face of the actin monomer opposite the ATP-binding cleft, blocking the side of the monomer that would normally associate with the filament minus end, while leaving exposed the site on the monomer that binds to the plus end. When the profilin actin complex binds a free plus end, a conformational change in actin reduces its affinity for profilin and the profilin falls off, leaving the actin filament one subunit longer.  The ActA protein on the bacterial surface activates the Arp2/3 complex to nucleate new filament assembly along the sides of existing filaments. Filaments grow at their plus end until capped by capping protein. Actin is recycled through the action of cofilin, which enhances depolymerization at the minus ends of the filaments. By this mechanism, polymerization is focused at the rear surface of the bacterium, propelling it forward bacteria in blue and actin filaments in red (A) Electron micrograph of a myosin II thick filament isolated from frog muscle. Note the central bare zone, which is free of head domains. (B) Schematic diagram, not drawn to scale. The myosin II molecules aggregate by means of their tail regions, with their heads projecting to the outside of the filament. The bare zone in the center of the filament consists entirely of myosin II tails. (C) A small section of a myosin II filament as reconstructed from electron micrographs. In the relaxed (noncontracting) state, the two heads of a myosin molecule are bent backward and sterically interfere with each other to switch off their activity. An individual myosin molecule in the inactive state is highlighted in green. The cytoplasmic myosin II filaments in non-muscle cells are much smaller, although similarly organized Figure 16 23 Direct evidence for the motor activity of the myosin head. In this experiment, purified myosin heads were attached to a glass slide, and then actin filaments labeled with fluorescent phalloidin were added and allowed to bind to the myosin heads. (A) When ATP was added, the actin filaments began to glide along the surface, owing to the many individual steps taken by each of the dozens of myosin heads bound to each filament. The video frames shown in this sequence were recorded about 0.6 second apart; the two actin filaments shown (red) were moving in opposite directions at a rate of about 4 1 /sec. (B) Diagram of the experiment. The large red arrows indicate the direction of actin filament movement (Movie 16.4). (A, courtesy of James Spudich.) Figure 16 25 The force of a single myosin molecule moving along an actin filament measured using an optical trap. (A) Schematic of the experiment, showing an actin filament with beads attached at both ends and held in place by focused beams of light called optical tweezers (Movie 16.6). The tweezers trap and move the bead and can also be used to measure the force exerted on the bead through the filament. In this experiment, the filament was positioned over another bead to which myosin II motors were attached, and the optical tweezers were used to determine the effects of myosin binding on movement of the actin filament. (B) These traces show filament movement in two separate experiments. Initially, when the actin filament is unattached to myosin, thermal motion of the filament produces noisy fluctuations in filament position. When a single myosin binds to the actin filament, thermal motion decreases abruptly, and a roughly 10-nm displacement results from movement of the filament by the motor. The motor then releases the filament. Because the ATP concentration is very low in this experiment, the myosin remains attached to the actin filament for much longer than it would in a muscle cell. (Adapted from C. Rne et al., News Physiol. Sci. 17:213 218, 2002. With permission from the American Figure 16 26 Skeletal muscle cells (also called muscle fibers). (A) These huge multinucleated cells form by the fusion of many muscle cell precursors, called myoblasts. Here, a single muscle cell is depicted. In an adult human, a muscle cell is typically 50 1 in diameter and can be up to several centimeters long. (B) Fluorescence micrograph of rat muscle, showing the peripherally located nuclei (blue) in these giant cells. Myofibrils are stained red. (B, courtesy of Nancy L. Kedersha.) Figure 16 27 Skeletal muscle myofibrils. (A) Low-magnification electron micrograph of a longitudinal section through a skeletal muscle cell of a rabbit, showing the regular pattern of cross- striations. The cell contains many myofibrils aligned in parallel (see Figure 16 26). (B) Detail of the skeletal muscle shown in A, showing portions of two adjacent myofibrils and the definition of a sarcomere (black arrow). (C) Schematic diagram of a single sarcomere, showing the origin of the dark and light bands seen in the electron micrographs. The Z discs, at each end of the sarcomere, are attachment sites for the plus ends of actin filaments (thin filaments); the M line, or midline, is the location of proteins that link adjacent myosin II filaments (thick filaments) to one another. (D) When the sarcomere contracts, the actin and myosin filaments slide past one another without shortening. (A and B, courtesy of Roger Craig.) Figure 16 28 Electron micrographs of an insect flight muscle viewed in cross section. The myosin and actin filaments are packed together with almost crystalline regularity. Unlike their vertebrate counterparts, these myosin filaments have a hollow center, as seen in the enlargement on the right. The geometry of the hexagonal lattice is slightly different in vertebrate muscle. (From J. Auber and R. Couteaux, J. Microsc. 2:309 324, 1963. With UeX1\\:2X:1S:céeéFXa2ça\ede Microscopie É+eceX:2Wje.) Figure 16 30 T tubules and the sarcoplasmic reticulum. (A) Drawing of the two membrane systems that relay the signal to contract from the muscle cell plasma membrane to all of the myofibrils in the cell. (B) Electron micrograph showing a cross section of a T tubule. Note the position of the large Ca2+-release channels in the sarcoplasmic reticulum membrane that connect to the adjacent T-tubule membrane. (C) Schematic diagram showing how a Ca2+-release channel in the sarcoplasmic reticulum membrane is thought to be opened by the activation of a voltage-gated Ca2+ channel in the membrane of the T tubule (Movie 16.7). (B, courtesy of Clara Franzini-Armstrong.) Figure 16 31A The control of skeletal muscle contraction by troponin. (A) A thin filament of a skeletal muscle cell, showing the positions of tropomyosin and troponin along the actin filament. Each tropomyosin molecule has seven evenly spaced regions with similar amino acid sequences, each of which is thought to bind to an actin subunit in the filament. (A, adapted from G.N. Phillips et al., J. Mol. Biol. 192:111 131, 1986.) Figure 16 32 Smooth muscle contraction. (A) Upon muscle stimulation by activation of cell-surface receptors, Ca2+ released into the cytoplasm from the sarcoplasmic reticulum (SR) binds to calmodulin (see Figure 15 34). Ca2+- bound calmodulin then binds myosin light-chain kinase (MLCK), which phosphorylates myosin light chain, stimulating myosin activity. Non-muscle myosin is regulated by the same mechanism (see Figure 16 34). (B) Smooth muscle cells in a cross section of cat intestinal wall. The outer layer of smooth muscle is oriented with the long axis of its cells extending parallel along the length of the intestine, and upon contraction will shorten the intestine. The inner layer is oriented circularly around the intestine and when contracted will cause the intestine to become narrower. Contraction of both layers squeezes material through the intestine, much like squeezing toothpaste out of a tube. (C) A model for the contractile apparatus in a smooth muscle cell, with bundles of contractile filaments containing actin and myosin (red) oriented obliquely to the long axis of the cell. Their contraction greatly shortens the cell. In this diagram, the bundle angles are exaggerated to schematically illustrate the effect of contraction. In addition, only a few of the many bundles are shown. (B, Figure 16 36 Myosin V walks along actin filaments. (A) The lever arm of myosin V is long, allowing it to take a bigger step along an actin filament than can myosin II (see Figure 16 24). (B) Atomic force microscopy images showing myosin V (green) walking along an actin filament. Myosin V functions to carry cargo in cells. (B, adapted from N. Kodera and T. Ando, Biophys. Rev. 6:237 260, 2014. Reproduced with permission of SNCSC.) Figure 16 37 The structure of a microtubule and its subunit. (A) The subunit of each UX:e:+a1e2e\aejbj+2eeeX:d1eX:X1edX:1aee++2’eUaX: d - a2d -tubulin 1:2:1eX\TeGTP1:+ecj+e2ee -tubulin monomer is so tightly bound that it can be c:2\deXeda22eeXa+UaXe:eeUX:ee2TeGTP1:+ecj+e2ee -tubulin monomer, however, is less tightly bound and has an important role in filament dynamics. GTP is shown in red. (BO2eejbj+2\jbj2e -heterodimer) and one protofilament are shown schematically. Each protofilament consists of many adjacent subunits with the same Figure 16 37 The structure of a microtubule and its subunit. (C) (D) A short segment of a microtubule viewed in an electron microscope. (E) Electron micrograph of a cross section of a microtubule showing a ring of 13 protofilaments. (A, PDB code: 1JFF; D, courtesy of Richard Wade; E, courtesy of Richard Linck.) Figure 16 38 The preferential growth of microtubules at the plus end. Microtubules grow faster at one end than at the other. A stable bundle of microtubules obtained from the core of a cilium (called an axoneme) was incubated for a short time with tubulin subunits under polymerizing conditions. Microtubules grew fastest from the plus end of the microtubule bundle, the end on the right in this micrograph. (Courtesy of Gary Borisy.). Salmon and Clare Waterman Courtesy of Wendy C 3 1 2 3 1 2 4 4 time 0 sec 125 sec 307 sec 669 sec 10 um DYNAMIC INSTABILITY Microtubules depolymerize about 100 times faster from an end containing GDP-tubulin than from one containing GTP-tubulin. AGTP cap favors growth, but fi ti is lost, then depolymerization ensues. GTP cap GROWING SHRINKING Individual microtubules can therefore alternate between a period of growth and a period of rapid disassembly, a phenomenon called dynamic instability. Figure 16 40A Dynamic instability due to the structural differences between a growing and a shrinking microtubule end. (A) If the free tubulin concentration in solution is near the critical concentration, a single microtubule end may undergo transitions between a growing state and a shrinking state. A growing microtubule has GTP-containing subunits at its end, forming a GTP cap. If GTP hydrolysis proceeds more rapidly than subunit addition, the cap is lost and the microtubule begins to shrink, an event called a catastrophe. But GTP-containing subunits may still add to the shrinking end, and if enough add to form a new cap, then microtubule growth resumes, an event called a rescue. More recent studies indicate that free tubulin subunits possess a similar bent conformation in both the T form and the D form. Growing microtubules with curved protofilaments at their tips have now been observed both in vitro and in vivo, and a straightening of the T form containing protofilaments may occur subsequent to subunit incorporation as favorable lateral interactions zip them into the microtubule lattice. Regardless of the mechanism of microtubule assembly, the loss of a GTP cap and subsequent catastrophe causes protofilaments containing D-form subunits to spring apart and depolymerize. Figure 16 41C Microtubule nucleation by the g-tubulin ring complex. (CI21a2ce++eUe\ee -TuSC spiral a\\:caee\eadde:2a+acce\\:XUX:ee2\e::X1ee -ejbj+2X2c:1U+eP -TuRC), which is likely to nucleate the minus end of a microtubule as shown here. Note the longitudinal discontinuity between two protofilaments, which results X:1ee\UXa+:Xe2eae:2:ee -ejbj+2\jbj2e\McX:ejbj+e\:ee2a}e:2e\jc\ea1bXea’2ee:eeX\e uniform helical packing of the protofilaments. A centriole consists of a cylindrical array of short, modified microtubules arranged into a barrel shape with striking ninefold symmetry. Together with a large number of accessory proteins, the centrioles recruit the pericentriolar material, where microtubule nucleation takes place. The pericentriolar material consists of a dense spherical matrix that is thought to form through a process of biomolecular condensation. The centrosome duplicates before mitosis, forming a pair of centrosomes that each contain a centriole pair. When mitosis begins, the two centrosomes move apart to form the poles of the mitotic spindle. Charge distribution on the surface of microtubule (red indicates negative charges, blue positive charges) Localization of MAPs in the axon and dendrites of a neuron. This immunofluorescence micrograph shows the distribution of the proteins tau (green) and MAP2 (orange) in a hippocampal nMAP2 staining is confined to the cell body Organization of microtubule bundles by MAPs. (A) One end of MAP2 binds along the microtubule lattice and extends a long projecting arm. (B) Tau possesses a shorter projection arm. (C) Electron micrograph showing a cross section through a microtubule bundle in a cell overexpressing MAP2. The regular spacing of the microtubules (MTs) in this bundle results from the constant length of the projecting arms of MAP2. (D) Similar cross section through a microtubule bundle in a cell overexpressing tau. Here the microtubules are spaced more closely together than they aXe2Cbecaj\e:eaj\Xe+ae}e+\:XeUX:$ece2aX1 Microtubule branching by augmin. (A) Augmin b2d\a+:2ee\d:a2e e P\e21cX:ejbj+ea2dXecXje\a - tubulin ring complex that nucleates a new microtubule with a low branching angle. (B) Fluorescence micrographs showing augmin (orange) nucleating a microtubule branch in the cortex of an epidermal cell in the plant Arabidopsis thaliana. (C) Depletion of augmin severely stunts plant growth The effects of proteins that bind to microtubule ends. The transition between microtubule growth and shrinkage is controlled in cells by a variety of proteins. Catastrophe factors such as kinesin-13, a member of the kinesin motor protein superfamily, bind to microtubule ends and pry them apart, thereby promoting depolymerization. On the other hand, a MAP such as XMAP215 promotes rapid microtubule polymerization. XMAP215 binds tubulin dimers and delivers them to the microtubule plus end, thereby increasing the microtubule growth rate. Frames from a fluorescence time-lapse movie of the edge of a cell expressing fluorescently labeled tubulin that incorporates into microtubules (green) as well as the +TIP protein EB1 tagged with a different color (red). The same microtubule is marked (asterisk) in successive movie frames. When the microtubule is growing (frames 1, 2), EB1 is associated with the tip. When the microtubule undergoes a catastrophe and begins shrinking, EB1 is lost (frames 3, 4). The labeled EB1 is regained when growth of the microtubule is rescued (frame 5) Sequestration of tubulin by stathmin. Structural studies with electron microscopy and crystallography suggest that the elongated stathmin protein binds along the side of two tubulin heterodimers Microtubule severing by katanin can destabilize or amplify microtubules. (A) Taxol-stabilized, fluorescently labeled microtubules were adsorbed on the surface of a glass slide, to which purified katanin was added along with ATP. There are a few breaks in the microtubules 30 seconds after the addition of katanin. (B) Three minutes after the addition of katanin, the filaments have been severed in many places, leaving a series of small fragments at the previous locations of the long microtubules. (C) Incorporation of GTP-tubulin subunits from the soluble pool into sites of katanin-induced damage in the microtubule lattice stabilizes the severed end or generates an island of GTP-tubulin that promotes rescue. MICROTUBULES + Y-TuRC augmin nucleates assembly and centrosome nucleates microtubule remains associated with branching t h e minus end +¥ stathmin +TIPs binds subunits, aß-tubulin dimers associate with growing plus ends prevents assembly a n d can link t h e m to other structu res, such as m e m b r a n e s MICROTUBULES - microtubule - + ХМА Р215 stabilizes plus ends, kinesin-13 promotes rapid microtubule + growth induces catastrophe and disassembly + katanin MAPS severs microtubules stabilize microtubules by binding along sides filament bundling and cross-linking 170 tau MAP2 plectin links to i n t e r m e d i a t e fi l a m e n t s There are two major classes of microtubule-based motors, kinesins and dyneins, which perform three major functions: 1) they move cargo such as organelles and macromolecules within the cell. Unlike actin-based transport, however, microtubule-based motors are used to transport cargo over long distances. 2) motors can slide microtubules relative to one another, thereby generating specific arrangements of microtubules, as in neurons and epithelial cells and in the mitotic spindle. 3) a subset of microtubule-based motors regulates microtubule dynamics, as illustrated by kinesin-13  Kinesins As in the myosin superfamily, only the motor domains are conserved. Kinesin-1 has the motor domain at the N-terminus of the heavy chain and moves toward the microtubule plus end. The middle domain forms a long coiled-coil, mediating dimerization. The C-terminal domain forms a tail that attaches to cargo, such as a membrane-enclosed organelle. Kinesin-5 forms tetramers in which two dimers associate by their tails. The bipolar kinesin-5 tetramer is able to slide two microtubules past each other, analogous to the activity of the bipolar thick filaments formed by myosin II. Kinesin-13 has its motor domain located in the middle of the heavy chain. It is a member of a family of kinesins that have lost typical motor activity and instead bind to microtubule ends to promote depolymerization. Kinesin-14 is a C-terminal kinesin. Unlike most kinesins, Structures of four kinesin superfamily members members of the kinesin-14 family travel toward the microtubule minus end.  Dyneins Cytoplasmic dynein. (A) Cryo-electron microscopy (cryoEM) reconstruction of a molecule of cytoplasmic dynein. Like myosin II and kinesin-1, cytoplasmic dynein is a two-headed molecule. The dynein head is very large compared with the head of either myosin or kinesin. (B) Schematic depiction of cytoplasmic dynein showing the two heavy chains that contain a motor head with domains for microtubule binding and ATP hydrolysis, connected by a long stalk. The tail domain consists of a linker that connects the motor heads to a dimerization domain. Bound to the linker domain are multiple intermediate chains and light chains (blue) eaee+Ue:1edaee1a2:d2e2\j2ce:2\ C) The organization of domains in a dynein heavy chain. This is a huge polypeptide, containing more than 4000 amino acids. The conserved dynein motor head domain contains six AAA domains, four of which retain ATP-binding sequences, but only one of which has the major ATPase activity (brown). The tail domain is not as highly conserved as the head domain The power stroke of dynein. Illustration of the movement of a monomeric axonemal dynein found in the flagellum of the unicellular green alga Chlamydomonas reinhardtii. As in cytoplasmic dynein, the motor-containing head domain of axonemal dynein connects to a long, coiled-coil stalk with the microtubule-binding site at the tip. The tail aeeace\e:a2ad$ace2e1cX:ejbj+e2eeaP:2e1eM:}e1e2e\e:jee::ccjXeX:ja+2’eX -swing, dynein- 2c1eca2\1 ATP binding and hydrolysis cause the linker to throw the head domain toward the microtubule minus end like a fishing hook. The microtubule-binding domain reattaches 8 nm along the microtubule. Release of ATP and phosphate then leads to a large conformational power stroke in the linker domain, pulling the tail and its attached microtubule toward the minus end. Each cycle generates a step of about 8 nm, thereby contributing to flagellar beating (see Figure 16 60). In the case of cytoplasmic dynein, the tail is attached to a cargo such as a vesicle, and a single power stroke transports the cargo about 8 nm along the microtubule toward its minus end (see Figure 16 56). The arrangement of microtubules in a flagellum or cilium. (A) Electron micrograph of the flagellum of a green- alga cell (Chlamydomonas\:22cX:\\\ece:2++j\eXae2eed\e2ce}eXmQaXXa2e1e2e: microtubules. (B) Diagram of the parts of a flagellum or cilium. The various projections from the microtubules link the microtubules together and occur at regular intervals along the length of the axoneme. (C) High-resolution electron tomography image of an outer microtubule doublet showing structural details and features inside the microtubules called microtubule inner proteins (MIPs). Axonemal dynein. CryoEM reconstruction of a sea urchin sperm flagellum showing dynein arms connecting the A microtubule of one doublet with the B microtubule of an adjacent doublet at regular intervals. Sperm axonemal dynein is dimeric. The tail of the molecule binds tightly to an A microtubule, while the two globular heads each have a stalk that connects to an ATP-dependent binding site on a B microtubule (see Figure 16 58). When the heads hydrolyze their bound ATP, they move toward the minus end of the B microtubule, thereby producing a sliding force between the adjacent microtubule doublets in a cilium or flagellum The bending of an axoneme. (A) When axonemes are exposed to the proteolytic enzyme trypsin, the flexible protein links holding adjacent microtubule doublets together are broken. In this case, the addition of ATP allows the motor action of the dynein heads to slide one microtubule doublet against the adjacent doublet. (B) In an intact axoneme (such as in a spermatozoon), the flexible protein links prevent the sliding of the doublet. The motor action therefore causes a bending motion, creating waves or beating motions. Primary cilia. (A) Electron micrograph and diagram of the basal body of a mouse neuron primary cilium. The axoneme of the primary cilium (black arrow) is nucleated by the mother centriole at the basal body, which localizes at the plasma membrane near the cell surface. (B) Centrioles function alternately as basal bodies and as the core of centrosomes. Before a cell enters the cell-division cycle, the primary cilium is shed or resorbed. The centrioles recruit pericentriolar material and duplicate during S phase, generating two centrosomes, each of which contains a pair of centrioles. The centrosomes nucleate microtubules and localize to the poles of the mitotic spindle. Upon exit from mitosis, a primary cilium again grows from the mother centriole. An electron micrograph of intermediate filaments A model of intermediate filament construction The monomer shown in (B) pairs with another monomer to form a dimer, in which the conserved central rod domains are aligned in parallel and wound together into a coiled- coil. (C) Two dimers then line up side by side to form an antiparallel tetramer of four polypeptide chains. Dimers and tetramers are the soluble subunits of intermediate filaments. (D) Within each tetramer, the two dimers are offset with respect to one another, thereby allowing it to associate with another tetramer. (E) In the final 10-nm-diameter filament, tetramers are packed together in a rope-like array, which has 16 dimers (32 coiled-coils) in cross section. Half of these dimers are pointing in each direction. An electron micrograph of intermediate filaments is shown on the upper left (Movie 16.16).  basal cell of epidermis basal lamina defective keratin hemidesmosomes filament network (A) (B) (C) 40 um Figure 16-64 Blistering of the skin caused by a mutant keratin gene. A mutant gene encoding a truncated keratin protein (lacking both the N - and C - terminal domains) was expressed in a transgenic mouse. The defective protein assembles with the normal keratins and thereby disrupts the keratin filament network ni the basal cells of the skin. Light micrographs of cross sections of (A) normal and (B) mutant skin show that the blistering results from the rupturing of cells ni the basal layer of the mutant epidermis (short red arrows). (C) Asketch of three cells ni the basal layer of the mutant epidermis, as observed by electron microscopy. As indicated by the red arrow, the cells rupture between the nucleus and the hemidesmosomes (discussed ni Chapter 19), which connect the keratin filaments to the underlying basal lamina. A ( and B, © 1991 P.A. Coulombe et al. Originally published ni J. Cell Bio/. https://doi.org/10.1083/jcb.115.6.1661. With permission from Rockefeller University Press.) Two types of intermediate filaments in cells of the nervous system. (A) Freeze-etch electron microscopy image of neurofilaments in a nerve cell axon, showing the extensive cross-linking through protein cross-bridgese an arrangement believed to give this long cell process great tensile strength. The cross-bridges are formed by the long, nonhelical extensions at the C-terminus of the largest neurofilament protein (NF-H). (B) Freeze-etch image of glial filaments in glial cells, showing that these intermediate filaments are smooth and have few cross- bridges. (C) Conventional transmission electron micrograph of a cross section of an axon showing the regular side-to-side spacing of the neurofilaments, which greatly outnumber the microtubules. Plectin cross-linking of diverse cytoskeletal elements. Plectin (green) is seen here making cross- links from intermediate filaments (blue) to microtubules (red). In this electron micrograph, the dots (yellow) are gold particles linked to anti-plectin antibodies. The entire actin filament network was Xe1:}ee:Xe d }ea+ee\eUX:ee2\0PXXUTM Svitkina et al. Originally published in J. Cell Biol. http://doi.org/10.1083/jcb.135.4.991. With permission from Rockefeller University Press.) GTP-binding proteins called septins serve as an additional filament system in all eukaryotes except terrestrial plants. Septins assemble into nonpolar filaments that form rings and cage-like structures, which act as scaffolds to compartmentalize membranes into distinct domains or to recruit and organize the actin and microtubule cytoskeletons. In primary cilia, a ring of septin filaments assembles at the base of Cell compartmentalization by septins. (A) Septins form filaments in the the cilium and serves as a diffusion neck region between a mother yeast cell and bud. (B) In this barrier at the plasma membrane, photomicrograph of human cultured cells, the DNA is stained blue and restricting the movement of septins are labeled in green. The microtubules of primary cilia are labeled membrane proteins and with an antibody that recognizes a modified (acetylated) form of tubulin establishing a specific composition (red) that is enriched in the axoneme. (C) A magnified image reveals a in the ciliary membrane collar of septin at the base of the cilium. The dramatic effects of Cdc42, Rac, and Rho on actin organization in fibroblasts. In each case, the actin filaments have been labeled with fluorescent phalloidin. (A) Serum-starved fibroblasts have actin filaments primarily in the cortex and relatively few stress fibers. (B) Microinjection of a constitutively activated form of Cdc42 results in many long filopodia at the cell periphery. (C) Microinjection of a constitutively activated form of Rac causes the formation of an enormous lamellipodium that extends from the entire circumference of the cell. (D) Microinjection of a constitutively activated form of Rho causes the rapid assembly of many prominent stress fibers. Cdc42 establishes yeast-cell polarity. (A) A positive feedback loop by which Cdc42-GTP recruits its own GEF to the plasma membrane to generate a focal site of Cdc42 activity. (B) Local activation of a formin protein by Cdc42-GTP nucleates actin filament assembly. Transport of vesicles along these actin filaments toward their plus ends by myosin V delivers cargoes necessary for growth of the bud. PAR proteins establish two distinct cortical domains in C. elegans. (A) Symmetry breaking and polarity establishment occur in the fertilized egg before the maternal and paternal nuclei meet. After sperm entry, its centrosome duplicates and nucleates microtubules that decrease Rho GEF activity at what will become the posterior end of the embryo. This leads to accumulation of the anterior PAR proteins (including Par-3), allowing posterior PAR proteins (including Par-2) to bind the cortex at the posterior end. (B) Prior to fertilization, bundles of fluorescently labeled myosin II (white) are distributed throughout the cortex of the unpolarized egg because of the uniform distribution of activated Rho all along the plasma membrane, resulting in uniform cortical acto- myosin contractility. (C) After fertilization, local depletion of the Rho GEF near the sperm entry site (at right in this image) reduces myosin levels and contractility in the posterior cortex of the cell. (D) Par complex localization after polarization with Par-3 (red) at the anterior and Par-2 (green) at the posterior of the zygote. Multiple mechanisms operate to maintain this asymmetry through mutual antagonism between anterior and posterior PAR Cell-polarity protein modules identified in Drosophila. (A) PAR and Crumbs proteins cooperate to assemble the apical domain and junctional complexes, whereas Scribble defines the basolateral domain. Scribble and PAR are mutually antagonistic, whereas PAR and Crumbs reinforce each other. Cdc42 helps to recruit PAR proteins. The contrasting effects of Rac and Rho activation on actin organization. (A) Activation of the small GTPase Rac leads to alterations in actin accessory proteins that promote the formation of protrusive actin networks in lamellipodia and pseudopodia. (B) Activation of the related GTPase Rho leads to nucleation of actin filaments by formins and increases contraction by myosin II, promoting the formation of contractile actin bundles at the rear of the cell and assembly of stress fibers. myosin heavy chain (MHC) myosin light-chain kinase (MLCK) Neutrophil polarization and chemotaxis. (B) Binding of bacterial molecules to G-protein- coupled receptors on the neutrophil stimulates directed motility. These receptors are found all over the surface of the cell, but are more likely to be bound to the bacterial ligand at the front. Two d\e2ce\2a+2Uaea\c:2eXbjeee:eece++\ polarization. At the front of the cell, stimulation of the Rac pathway leads, via the trimeric G protein Gi, to growth of protrusive actin networks. Second messengers within this pathway are short-lived, so protrusion is limited to the region of the cell closest to the stimulant. The same receptor also stimulates a second signaling pathway, via the trimeric G proteins G12 and G13, that triggers the activation of Rho. The two pathways are mutually antagonistic. Because Rac-based protrusion is active at the front of the cell, Rho is activated only at the rear of the cell, stimulating contraction of the cell rear and assisting directed movement.