XB-ART-10619J Cell Biol. July 24, 2000; 150 (2): 361-76.
Microtubules remodel actomyosin networks in Xenopus egg extracts via two mechanisms of F-actin transport.
Interactions between microtubules and filamentous actin (F-actin) are crucial for many cellular processes, including cell locomotion and cytokinesis, but are poorly understood. To define the basic principles governing microtubule/F-actin interactions, we used dual-wavelength digital fluorescence and fluorescent speckle microscopy to analyze microtubules and F-actin labeled with spectrally distinct fluorophores in interphase Xenopus egg extracts. In the absence of microtubules, networks of F-actin bundles zippered together or exhibited serpentine gliding along the coverslip. When microtubules were nucleated from Xenopus sperm centrosomes, they were released and translocated away from the aster center. In the presence of microtubules, F-actin exhibited two distinct, microtubule-dependent motilities: rapid ( approximately 250-300 nm/s) jerking and slow ( approximately 50 nm/s), straight gliding. Microtubules remodeled the F-actin network, as F-actin jerking caused centrifugal clearing of F-actin from around aster centers. F-actin jerking occurred when F-actin bound to motile microtubules powered by cytoplasmic dynein. F-actin straight gliding occurred when F-actin bundles translocated along the microtubule lattice. These interactions required Xenopus cytosolic factors. Localization of myosin-II to F-actin suggested it may power F-actin zippering, while localization of myosin-V on microtubules suggested it could mediate interactions between microtubules and F-actin. We examine current models for cytokinesis and cell motility in light of these findings.
PubMed ID: 10908578
PMC ID: PMC2180232
Article link: J Cell Biol.
Grant support: GM24364 NIGMS NIH HHS , GM52932-01A2 NIGMS NIH HHS , R29 DC003299 NIDCD NIH HHS
Genes referenced: actn1 dync1li1
Article Images: [+] show captions
|Figure 1. Microtubule-independent F-actin motility in Xenopus egg extracts. (A) Low magnification, time-lapse view of an Alexa 488 phalloidin-stained F-actin network formed in the presence of nocodazole (10 μM). The number of F-actin bundles decreases and their thickness increases over time as bundle zippering results in annealing of adjacent bundles. (B) Trajectory plot of the position of F-actin bundles in a time-lapse series of 600 × 600 pixel images. Time points are separated by 15 s; arrowheads indicate the direction of movement in final frame. Most of the bundles move only short distances and display no consistent direction of movement. (C) High magnification, time-lapse view of F-actin bundle zippering. Adjacent bundles (arrows) zipper together at intersections, resulting in the formation of larger bundles and a decrease in the number of intersections. (D) Depletion of ATP by apyrase treatment (10 μg/ml) of extracts blocks F-actin zippering. The number of intersections between adjacent F-actin bundles was quantified in extracts at various times after the initiation of imaging in the absence (control) and presence (apyrase) of apyrase. In the absence of apyrase, the number of intersections decreases over time; in the presence of apyrase the number of intersections fails to decrease and the networks did not change over time. Data shown are mean ± SEM; n = 2. (E) High magnification, time-lapse view of F-actin bundle serpentine gliding. The F-actin bundle moves sinuously across the substrate. Images in successive panels were acquired at 15-s intervals. (F) Plot of the X-Y coordinates of the point of peak fluorescence intensity of the F-actin bundle shown in E. This demonstrates the serpentine path followed by the F-actin bundle. (G) Serpentine gliding is prevented by depletion of ATP with apyrase and is stimulated by BDM. The number of gliding events per field of view in samples subjected to the indicated treatment was compared with the number of events in control samples from the same experiment. Apyrase almost completely eliminated serpentine gliding. BDM at 20 mM had a slight stimulatory effect, while BDM at 50 mM more than tripled the number of bundles moving by serpentine gliding. Data shown are mean ± SEM; n = 2. Time is in min/s. Also see supplemental videos 1–3 at http://www.jcb.org/cgi/content/full/150/2/361/DC1.|
|Figure 2. Microtubule gliding and aster expansion in Xenopus egg extracts. (A) FSM of astral microtubule release and gliding along the substrate. The microtubule nucleated at the aster center (arrow) grows several microns and is then released at its minus end from the aster. The dark gap (arrowhead) in the speckled microtubule moves relative to the substrate demonstrating that the microtubule is translocating away from the aster via the action of a minus end–directed motor on the coverslip. (B) Low magnification, time-lapse view showing expansion of a sperm aster. Immediately after the initiation of imaging, the aster is well organized into a radial array containing many microtubules. Over time, microtubules are released and translocate away from the aster, and small radial arrays of microtubules (arrowheads) wander away from the original aster. (C) Low magnification, images of asters that were formed by adding demembranated sperm to extracts in solution and fixed at the time point shown and then placed in a slide-coverslip chamber. At the earliest time point, asters are well focussed, similar to those observed in thin preparations (see B). At the 5-min time point, asters have expanded, and the microtubules do not extend from a single, well-defined point. At 10- and 20-min time points, expansion and disorganization is even more pronounced. Time is in min/s. Also see supplemental videos 4 and 5 at http://www.jcb.org/cgi/content/full/150/2/361/DC1.|
|Figure 3. Microtubules remodel F-actin networks by rapid jerking of F-actin bundles. (A) Low magnification, time-lapse view of an F-actin network formed in the presence of randomly arrayed microtubules. An F-actin bundle network forms and undergoes zippering, but is also subject to rapid jerking movements that grossly distort the organization of the network. (B) Trajectory plot of the position of F-actin bundles in a time-lapse series of 600 × 600 pixel images in the presence of randomly arrayed microtubules. Individual bundles move long distances, but do not move in any consistent direction, and change direction rapidly. Time points are separated by 15 s; arrowheads indicate the direction of movement in final frame. (C) Low magnification, time-lapse view of an F-actin network formed in the presence of microtubules organized by a demembranated sperm MTOC (position marked by an asterisk). The F-actin network undergoes zippering, but is progressively cleared centrifugally from the area occupied by the MTOC. Over time, thick actin bundles (arrows) accumulate around the periphery of the MTOC. (D) Trajectory plot of F-actin bundle motility in the presence of aster-organized microtubules. Individual bundles move long distances, in a consistently centrifugal direction out from the position of the MTOC center (asterisk). Time points are separated by 15 s; arrowheads indicate the direction of movement in final frame. (E) High magnification, time-lapse view of part of an F-actin network subjected to jerking motility in the presence of randomly arrayed microtubules. An F-actin bundle (arrowhead) making up one branch of the network undergoes directed movement (0:00–00:30) and collides with an adjacent bundle (at 00:30) forming a V-shaped junction between the bundles. The junction then undergoes a change in trajectory (time 00:45) and rapidly moves in unison with a neighboring bundle (arrow; 02:15–03:15). The movement brings the moving bundles into collisions with other bundles. (F) Microtubules accelerate the bundling of F-actin networks. The number of intersections between F-actin bundles within a network was quantified at various time after the initiation of imaging in four control experiments in the presence of microtubules (with MTs) and three experiments using extract containing 10 μM nocodazole (without MTs). The presence of microtubules accelerates the rate of bundling relative to the rate of bundling in their absence, as judged by the more rapid decrease in the number of intersections. To control for variation in the number of starting intersections, results are expressed as the % of the starting number of intersections. Results shown are mean ± SEM. Time is in min/s. Also see supplemental videos 6–8 at http://www.jcb.org/cgi/content/full/150/2/361/DC1.|
|Figure 4. F-actin bundles form lengthwise associations with microtubules. Double label images of microtubules obtained from time-lapse movies were digitally processed to remove the background of unpolymerized fluorescent label in the extract, color encoded, and combined into an RGB overlay to reveal the codistribution of the two polymers. Microtubules are randomly arrayed on the coverslip surface. Some F-actin is not associated with microtubules, but a substantial amount is aligned lengthwise along microtubules (arrowheads). Occasionally, ends of F-actin bundles terminate on the side of microtubules (large arrow) and microtubule and F-actin bundles can associate end-to-end (small arrow).|
|Figure 5. F-actin jerking results from F-actin binding to translocating microtubules. (A) Time-lapse views of an F-actin bundle (arrow) that undergoes jerking motility. (B) Time-lapse, FSM views of the microtubule associated with the F-actin bundle in A. That this microtubule is gliding is demonstrated by movement of the boundary between a dark region and a fluorescent spot on the lattice of the speckled microtubule (arrowhead). (C) Dual-wavelength fluorescence imaging of actin (green) and tubulin (red) showing the combined images depicted in A and B. The images were digitally processed to reduce the background, color-coded, and combined into an RGB overlay to reveal the codistribution of the two polymers. The movement of the F-actin bundle mirrors the movement of the speckled microtubule, indicating that the F-actin is attached to the lattice of the gliding microtubule. Images in A–C were acquired at 15-s intervals. (D) Plot of the distance of the tip (arrow in A) of the F-actin bundle and the speckle mark (arrowhead in B) on the microtubule from the origin (the position of the F-actin bundle tip at time 0:00) vs. time. The plot shows the exact correspondence in the movement of the two polymers. (E) Centrifugal clearing of F-actin around a sperm aster. Demonstrated by dual-wavelength fluorescence imaging of X-rhodamine tubulin (red) and Alexa 488 phalloidin labeling of F-actin (green) in Xenopus egg extracts. Time in min/s. F-actin is extensively aligned along astral microtubules (times 00:00–6:15). Over time, F-actin bundles are translocated away from the sperm aster center (arrowheads, times 00:00–2:15) while remaining bound to microtubules are released and translocate away from the aster as the aster fragments (arrow) and expand (time 10:01, arrow). Also see supplemental videos 9 and 10 at http://www.jcb.org/cgi/content/full/150/2/361/DC1.|
|Figure 6. Cytoplasmic dynein powers microtubule gliding, aster expansion, and F-actin bundle jerking. (A) Low magnification, time-lapse views of microtubules (left) seeded from a demembranated Xenopus sperm and F-actin (right) in the presence of 2 mg/ml anti-cytoplasmic dynein intermediate chain antibody 70.1. The antibody inhibits the translocation of microtubules away from the MTOC, thus inhibiting aster expansion but does not inhibit microtubule nucleation and growth, thus inducing a striking accumulation of microtubules (left; compare to Fig. 2 B). The antibody also inhibits F-actin jerking and prevents centrifugal clearing of F-actin from the aster (right). Time is in min/s. (B) Low magnification views showing asters that were fixed at various times after the addition of demembranated sperm to Xenopus extracts in solution in the presence of 2 mg/ml anti-cytoplasmic dynein intermediate chain antibody 70.1. In the presence of the antibody, even at late time points, the aster remains tightly focussed and has many microtubules emanating from it (compare to Fig. 2 C). (C) Cytoplasmic dynein activity is required for microtubule gliding. Quantification of the rate of microtubule gliding across the substrate was calculated in control extracts (n = 11), extracts containing 2 mg/ml 70.1 (n = 9), and extracts containing 250 μM sodium orthovanadate (n = 5). Inhibition of cytoplasmic dynein drastically slowed the rate of microtubule gliding. (D) Cytoplasmic dynein activity is required for F-actin jerking. Quantification of the rate of F-actin jerking was determined from the same experiments used for C. Inhibition of cytoplasmic dynein with 70.1 antibodies or orthovanadate completely eliminated F-actin jerking. (E) Cytoplasmic dynein is not required for F-actin zippering. Quantification of the rate of F-actin zippering was determined from the same experiments used for C. Cytoplasmic dynein inhibition with 70.1 antibodies or orthovanadate does not affect F-actin zippering. Also see supplemental video 11 at http://www.jcb.org/cgi/content/full/150/2/361/DC1.|
|Figure 7. F-actin can translocate along the lattice of stationary microtubules. (A) Time-lapse fluorescence imaging shows F-actin moving in a straight path. (B) Time-lapse FSM view of a stationary microtubule in the same field of view shown in A. The speckle mark indicated by the arrowhead does not move with respect to the substrate, showing that this microtubule is stationary throughout the period of imaging. (C) Color overlay of F-actin from A and the microtubule from B shows that the F-actin moves with respect to the stationary speckle marks on the microtubule lattice. Images in A–C were acquired at 15-s intervals (also see supplemental video 12 at http://www.jcb.org/cgi/content/full/150/2/361/DC1). (D) Plot of the distance of the tip (arrow in A) of the F-actin bundle and the speckle mark on the microtubule (arrowhead in B) from the origin (the position of the F-actin bundle tip at time 0:00) vs. time. This demonstrates that the F-actin moves relative to the microtubule lattice.|
|Figure 8. Translocating microtubules fail to transport F-actin in the absence of extract factors. (A) Time-lapse, dual-label fluorescence views of Alexa 488 phalloidin–labeled skeletal muscle F-actin/alpha actinin bundles (green) and microtubules (red) assembled from 99.5% porcine brain tubulin and 0.5% X-rhodamine–labeled tubulin that are gliding across a coverslip coated with rat brain kinesin. A microtubule translocates across the field of view, as judged by following the translocation of its speckle mark (arrow). En route, it makes contact with multiple F-actin bundles, three of which are indicated (#1–3). In spite of these multiple contacts, the moving microtubule does not transport any F-actin bundles. (B) Plot of the distance of the microtubule speckle mark (arrow in A) and of the three different F-actin bundles indicated in A (#1–3) from the origin (position of the speckle at time 00:00) vs. time. This demonstrates that the contact with the microtubule doesn't change the position of the actin bundles. Also see supplemental video 13 at http://www.jcb.org/cgi/content/full/150/2/361/DC1.|
|Figure 9. Localization of myosin-IIA and -V in Xenopus egg extracts. (A) Triple label fluorescence analysis of myosin-IIA (blue), microtubules (red), and F-actin (green) in rapidly frozen extracts. Myosin-IIA punctae (arrowheads) are found predominantly on F-actin bundles. In some regions, myosin-2, microtubules, and F-actin are colocalized (triple arrowheads); in other regions, F-actin and microtubules are colocalized in the absence of myosin-2 (arrows). (B) Triple label fluorescence analysis of myosin-V (blue), microtubules (red), and F-actin (green) in rapidly frozen extracts. Most of the myosin-V (arrowheads) is found on microtubules or the substrate. In both A and B colocalization of red, blue and green result in white.|