STUDIES ON THE MECHANISM OF MITOSIS J . Richard McIntosh, Zacheus Cande, Judith Snyder, and Kenneth Vanderslice Department of Molecular, Cellular, and Developmental Biology University of Colorado Boitlder. Colorado 80302
A n understanding of any machine is based upon detailed knowledge of the motions of its moving parts and of the way in which it uses fuel. Studies on the mechanism of muscle contraction are a clear example of how morphological and biochemical data can fit together to provide this information about a biological machine. T h e mechanism of chromosome motion can certainly be understood by a similar analysis, and over the past years important information about the mitotic apparatus ( M A ) has appeared in the literature. The M A has proved t o be difficult experimental material, because it is small, easy to kill, and less well organized than a striated muscle cell. Nonetheless, we know something about the chemistry of the isolated M A l-’, and quite a bit about its structure.9-12 Experimental studies such as those of InouC and Sato l 3 have contributed insights into the kinds of forces that hold the spindle together, and have suggested a model for how the machinery of the spindle might function to move the chromosomes. Alternative models f o r generation of spindle force have been proposed, such as Bajer’s “zipper hypothesis” and the several “sliding filament models,” but no one model stands out as uniquely suitable to account for all the available information. Part of the problem in determining the mechanisms of mitosis has been the insufficiency of detailed information concerning the motile events that occur within the spindle, although spindle structure at different times during mitosis has been studied in many different cells (reviews appear in References 17-20). The motion of the chromosomes during their congression to the metaphase plate and their subsequent parting and segregation at anaphase has been carefully described in numerous species,” but as Mazia’“ has pointed out, the role of the chromosome at mitosis o r meiosis is a passive one. For our purposes, a chromosome may be regarded as a marker for one end of the spindle fibers that attach to its kinetochores. Almost all students of mitosis agree that the chromosomes move because they are pushed o r pulled by the spindle fiber to which they are attached. The mechanism of chromosome motion, then, is really a question of the mechanism by which the cell exerts a force o n the chromosome fiber. This fiber is a bundle of microtubules, that range in number from 1 in a fungus 2 1 to over 100 in certain plants.” Mitotic forces, therefore, are basically forces that push on microtubules, although detailed observation of mitotic cells implicates the spindle in a variety of other motions that involve W e interpret the chromosome fragments and particles of diverse job of understanding mitotic mechanism as one of describing the motions of the spindle microtubules, and of analysing the molecular mechanisms that transduce chemical energy into the mechanical work necessary to cause these motions. One would like to watch spindle fiber motions in the light microscope, but
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individual tubules are of course too small to resolve optically. Polarization microscopy can yield a quantitative measure of optical anisometry and hence a measure related to the number of aligned fibers, but our efforts to use this method for determining spindle fiber motions during anaphase have failed (for what we believe to be solid and inescapable physical reasons, which will be described in detail elsewhere). Electron microscopy therefore seems to be the method of choice for the study, in spite of the obvious problems of the preparation artifact, and the fact that different time points in this ongoing process must be viewed in different cells. The most straightforward way to view the spindle fibers is to cut sections parallel to the spindle axis and to ask where the microtubules begin and end at different times during anaphase. Unfortunately, most spindle tubules leave the section before they truly end, so the longitudinal image is not useful for our purpose. We have circumvented this difficulty by cutting thick sections for viewing in a high-voltage electron microscope, by isolating whole spindles and viewing them with the same tool, and by cutting serial cross-sections that show the microtubules in transverse orientation. These sections can be aligned to allow tracking of individual tubules from section to section to find the tubule ends. I n comparing different stages of anaphase, we can then use the tubule ends as markers for tubule position. In a parallel investigation, we have been working to discover conditions that will allow chromosomes to continue their motion after the cell membrane has been lysed with detergent. These studies are directed toward elucidation of the enzyme system that exerts force upon the spindle tubules to make them move. Neither of these investigations is complete. We can maintain chromosome motion after lysis, but we do not yet know which enzymes are important for motility. We have determined certain facts about spindle fiber motion, but we do not yet know exactly how the fibers move. This paper must therefore be regarded as a progress report, in which we will describe our findings to date and define the directions of our plans for the immediate future.
MATERIALS A N D METHODS Mammalian cells (HeLa, WI-38, CHO, and PtK,) were cultured by standard techniques. For light microscopy, the cells were subcultured on glass cover slips which can be inverted onto a glass slide (using slivers of parafilm as spacers) and waxed down to form a simple chamber that will support the initiation and completion of cell division for about 4 hours. For electron microscopy, the cells were fixed after the method of Brinkley et aI.,?O or in a mixture of 4% glutaraldehyde, 4% 20 M Carbowax@ (Cx) in 0.1 M piperazine-N-N’-(2-ethane sulfonic acid) adjusted to pH 6.9 with NaOH (PIPES buffer). This fixative must be made up fresh before use. Fixed cells were rinsed in buffer, treated with OsO,, dehydrated, and embedded by standard techniques. Single cells were selected from wafers of plastic with a phase microscope, excised, and serially sectioned in a chosen orientation for electron microscopy. For microscopy in a high-voltage electron microscope, the cells were fixed and embedded as described and sectioned at a nominal thickness of 1 pm. About 10 sections were picked up on a carbon and Formvafl-coated slot grid, stained with uranyl acetate and lead citrate for about 30 min each at room
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temperature, and observed in a J E M 1000 (operating at an accelerating voltage of 1,000 kV) . Polarization optical measurements were made with a Brace-Kohler compensator and Zeiss polarization optics. Photographs were taken o n Kodak tri-X film developed in Diafinem. Anaphase motions were recorded o n Plus-X reversal film with a Bolex movie camera and Zeiss differential interference contrast optics. W e prepared repolymerization-competent microtubule protein, using a modification of the method of Olmsted and Borisy." The brains of I-3-day-old chicks were homogenized at 0" C in 0.1 M PIPES buffer at p H 6.9 that contained 1 m M E G T A and 2.5 m M G T P . T h e homogenate was spun at 35,000 X g for 30 min at 0" C; the supernatant was collected and made 30% in glycerol, then warmed to 37" C for 25 min, and spun at 35,000 X g for 1 hour at 35" C. T h e pellet was resuspended in 0.1 M PIPES at p H 6.9, 1 m M EDTA, 1 m M G T P at 37"C, and immediately pelleted again at the same temperature. This material was frozen in liquid nitrogen and stored at -70" C for u p to 1 week. A frozen pellet was homogenized in glycerol-free buffer with 1 m M G T P at 0" C, extracted for 1 hour and spun at 35,000 X g f o r 30 min. This supernatant was our standard tubulin preparation. Its purity was assessed by electrophoresis in 0.1% sodium dodecyl sulfate, and its capacity to polymerize was measured viscometrically and by direct view of negatively stained specimens in the electron microscope. To study the motion of chromosomes in lysed cells, we made coverslip preparations as described above and selected a cell with a suitable spindle. T h e wax seal was then broken, and a preparation of tubulin a n d / o r C x with 0.02-0.2% Triton@ X-100 ( T x ) added, was drawn under the coverslip with filter paper. RESULTS Stability
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When mammalian cells at metaphase are lysed with 0.01 %-0.1% T x in a buffer that supports brain tubulin polymerization in vitro, the spindle as seen in polarization optics disappears.Z5 As the tubulin concentration is raised, the rate of disappearance decreases (FIGURE 1 ). At high concentration, the spindle birefringence ( B R ) will increase. If the preparation of tubulin is warmed t o 37" C for 25 min prior to its use for lysis, then the spindle BR stays approximately constant for all concentrations of tubulin that we have tried above 1 mg/ml. Changes in spindle BR at lysis are apparently dependent upon the concentration of microtubule subunits, not upon the total concentration of tubulin present in the lysing solution. W e have determined that the increase in spindle BR is accompanied by a significant increase in the number of tubules per spindle (FIGURE 2). While trying to correlate BR with microtubule number, we observed, as others have before,l32 O' that spindle BR can drop significantly during fixation f o r electron microscopy. Fixation in the glutaraldehyde-Carbowax mixture described above will generally preserve spindle BR in mammalian cells for the first 20 min after the addition of fixative. Higher concentrations of Cx will sometimes promote an increase in spindle BR during fixation. Multiple experiments show,
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FIGURE1. (a) A graph of BR versus time after lysis for PtK, cells treated with 0.1 M PIPES buffer (pH 6.9), 1 mM EGTA, 2 mM GTP, and 0.1% Tx,c onta hhg difl'erent concentrations of tubulin: (m)~ 0 . 4 4mg/ml; ( 0 )=0.88 mg/ml; ( A ) = 1.75 mg/ml; (0) ~ 3 . mg/ml; 5 and (0) =7.0 mg/ml. All preparations were placed at 37" C for 5 min prior t o their use as a lysing solution. FIGUREl b shows BR versus time for cells lysed in different concentrations of tubulin in the above buffer and warmed to 37" C for 25 min prior to use in lysis: ( 0 ) = 0 . 7 5 mg/ml; ( O ) = 1.5 mg/ml; and ( A ) = 3 . 0 mg/ml. The error margin is the same for both graphs, and is a statement of the range of the reproducibility of multiple measurement of the same point.
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FIGURE 2. Qualitative correlations between spindle BR and spindle tubule number. FIGURES 2a, b, and c are polarization optical and electron-microscopic pictures of a cell lysed in an equilibrium mixture of tubules and tubulin. Quantitative data are 3 and 4 (cell 71). FIGURES 2d, e, and f are similar pictures of presented in FIGURES a cell lysed in recently warmed tubulin. (This is cell 76 in FIGURES 3 and 4.) Light micrographs, x 800; electron micrographs, x 48,000.
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FIGURE 3. A graph of BR versus time, which shows the variation of BR with fixation. Three lysed cells are monitored with different concentrations of tubulin in the lysing solution, to show BR changes with time both before and during fixation. The =Cell 76 lysed in about 10 mg/ following cells are fixed in glutaraldehyde-Cx. (0) ml recently warmed tubulin; ( 0 )=cell 77 lysed in about 8 mg/ml recently warmed tubulin; ( A ) =cell 71 lysed in about 8 mg/ml tubulin, prewarmed for 15 min; 70, lysed in about 8 mg/ml tubulin warmed for 15 min and fixed in phos(.)=cell phate-buffered glutaraldehyde; ( 0 )=cell 87, an unlysed cell fixed in glutaraldehyde-
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however, that occasional cells will retain their BR in phosphate-buffered glutaraldehyde, and occasional cells will lose BR during fixation in the glutaraldehyde-Cx mixture. These results are independent of whether or not the cell has been lysed prior to fixation (FIGURE 3 ) . FIGURE 4 shows the microtubule distribution profiles for some of the cells whose BR records are presented in FIGURE 3. Our results may be summarized as follows. When BR increases, the number of tubules in the spindle increases. The relation between the two is not linear, even when BR is kept constant during glutaraldehyde fixation. If BR is allowed to vary during fixation, the relation between BR before fixation and the number of tubules as seen in the electron microscope is not significantly altered, which indicates that loss of BR during glutaraldehyde fixation is not accompanied by significant loss in the number of microtubules. These statements apply both to lysed and unlysed cells (FIGURES 3 and 4). When cells are lysed in high concentrations of microtubule subunits (5-15 mg/ml, warmed from 0" C to 37" C immediately before use in lysis), the average tubule length as well as the average tubule number increases. FIGURE 5a is a metaphase cell lysed in a mixture of tubules and tubule subunits that is approximately at equilibrium (4 mg/ml warmed for 25 min, viscosity5b is a cell lysed in recently warmed tubulin. In FIGURE plateaued). FIGURE 5b there are tubules that appear to skewer the chromosomes and continue in
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straight lines for distances that exceed the spindle length, while in FIGURE Sa, the spindle shape is well preserved. This hypertrophy of number and length is not found in all spindle tubules. We have counted the number of tubules attached to individual kinetochores in normal, fixed cells and in augmented spindles. At metaphase, the chromosomes of one PtK, cell showed a n average of 31 10 tubules per kinetochore ( N = 8 ) ; that cell had a maximum of 1,575 tubules. A spindle from the same experiment as seen in FIGURE 5b showed a maximum of 6,970 tubules, but only 30 2 8 tubules per kinetochore ( N = 4). Thus the increase in tubule number is exclusively of nonchromosomal tubules in our cells. We have previously observed that the chromosome tubules are the most stable in the spindle with respect to extraction at low tubulin concentrations,2s which indicates that both their polymerization and depolymerization rates in v i m are substantially lower than those of the nonchromosome tubules.
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The motile capacity of the spindle is considerably more labile than the structure of the spindle as seen in the light and electron microscopes. The BR of the spindle will vary u p or down with tubulin concentration after lysis in
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Section Number FIGURE 4. Microtubule number versus position along the spindle axis measured in numbers of sections. Each section is about 900 A thick. These data correspond to 3. (.)=Cell 76; ( O ) = some of the cells whose BR records are shown in FIGURE cell 77; (A)=cell 71; (.)=cell 70. The BR increase observed with lysis in high concentrations of tubule subunits is reflected in the increase in microtubule number.
FIGURE 5. (a) A cell lysed in an equilibrium mixture of tubules and tubulin. The 5b is a cell from the sections for both 5a and 5b are about 1 gm thick. FIGURE experiment that produced cell 76. The quadrupling of the normal tubule number is accompanied by an increase in average tubule length and an extension of tubules out beyond the normal domain of the spindle. Presumably the length increase accounts for the image in FIGURE, 2 in which the chromosomes are riddled with tubules, for the peak rather than a trough in the middle of the top microtubule distribution curve of FIGURE 4, and for the tubules that radiate from the poles out of the spindle 5b. x 18,700. in FIGURE
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0.1% Tx, but chromosome motion will generally cease in detergent solutions stronger than 0.03%. In 0.03% Tx, 1 mM MgCI,, 1 mM each ATP and GTP, 1 mM dithiothreitol, and about 4 mglml tubulin brought to approximate equilibrium, the chromosomes will continue to move for some time after addition of the detergent (FIGURE 6 ) . Their rate of motion is less than the physiological rate by a factor of 3-5, but movement occurs, and the cell appears to be well lysed. Addition of 2-5% Cx to the lysing solution makes the chromosome motion considerably more rapid, but it also interferes with lysis and introduces the possibility that the cell membrane is not truly disrupted. Since Cx in PIPES buffer will stabilize a ~pindle,’~ we have tried to develop a Tx-Cx lysing solution without tubulin which would give reproducible lysis and nucleotide-dependent motion of the chromosomes, in order to facilitate experiments on the nature of the enzyme system of anaphase. These effort have been only partially successful. Treatment of the cells with 0.125% Tx and 5% Cx in 0.1 M PIPES buffer will induce extensive surface blebbing of the cell. The cell processes withdraw, the mitochondria spherulate, the spindle poles become distinct, and the cytoplasm pales in about 45 sec. If the solution is left on the cell for a few min, the spindle fibers become visible in the light microscope, and the chromosomes will no longer move. If, on the other hand, the lysing solution is replaced by about 2% Cx and 0.1-2.5 mM ATP and MgCI,, the the chromosomes will continue to move (FIGURE 7 ) . Cells that are fixed while the chromosomes are moving in this solution and viewed in the electron microscope do not look extracted. The chromosomes have an unusual texture, the cell surface is pitted, and the mitochondria are peculiar, but clearly the cell is not extensively lysed (FIGURE 7b). Nonetheless, if NaF is added with the lysing solution, the chromosomes stop rapidly, while N a F added to the medium without detergent has no effect upon chromosome motion (FIGURE 7a). These results indicate that chromosome motion can occur in cells lysed in different ways, but the data are too sparse to allow us to conclude anything about the nature of the mitotic motors. Structural Studies on Fixed Cells
Several recent studies of spindle structure have employed the method of counting the numbers of microtubules per spindle cross-section as a function of position on the spindle axis at different times during anaphase, to see how the spindle tubules are redistributed.Sn-?3 The available studies on mammalian cells are in disagreement on one important point: Brinkley and Cartwright 3n found that the distribution of tubules in the anaphase interzone did not change with time, while in our earlier work, we found that it did.3l We have reinvestigated this point using CHO and PtK, cells, and have confirmed our findings with HeLa and WI-38 (FIGURES 8 and 9 ) . In early anaphase, the number of tubules in the interzone does not vary with position along the spindle axis, while in late anaphase and telophase, we find, with Brinkley et C Z ~ .that , ~ ~there is a larger number of tubules at the middle of the interzone than on either side.31 As before, however, the number of tubules in the middle of the interzone is not twice the number on either side, as might be expected from the interdigitation of two families of tubules, nor is it uniform; the ratios are about 1:1.5: 1. We are attempting to clarify the structure of the middle of the interzone by reconstructing it in 3 dimensions, using serial cross-sections to track the
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FIGURE 6. Chromosome separation versus time during anaphase in PtKl cells. In 6a ( A ) = a r e a normal, unlysed cell; (A)=& cell lysed as described in the text; A = a n observation made after lysis had begun. The bar on the ordinate shows 6b is the cell recorded in FIGURE 6a t e metaphase length of both spindles. FIGURE while at metaphase; FIGURE 6c is after the onset of anaphase, but before lysis; FIGURE 6d is just after lysis, and corresponds to the boxed triangle in FIGURE 6a. FIGURE 6c is at 2224 sec. x 1,360.
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FIGURE 7. (a) A graph of chromosome separation versus time. ( A ) = A PtKl cell in tissue culture medium (5 m M NaF was added prior to the boxed triangle); (.)=a cell treated with lysing solution prior to the first boxed square, and treated with 2.5% Cx, 2.5 mM ATP and MgCL in PIPES prior to the second boxed square; ( A ) = a cell treated with lysing solution prior to the first boxed triangle and with 2.5% Cx, 2.5 mM ATP and MgCL, and 5 mM NaF prior to the second boxed triangle. FIGURE 7b is a cell treated exactly as the cell represented by (B) in FIGURE 7a. but fixed during chromosome motion. x 13,500.
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FIGURE 8. ( a ) The microtubule distribution profile of a CHO cell fixed during early anaphase. The diagram shows the approximate distribution of the chromosomes. The C’s show where the centrioles lay. (The dotted C indicates that our sectioning started too late; this position is an estimate, based upon the convergence of the spindle tubules.) FIGURE 8b is a CHO cell later in anaphase. FIGURE 8c shows data from the midregion of the interzone of a CHO telophase.
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Section Number FIGURE 9. ( a ) The microtubule distribution profile from an early anaphase PtKl 9b shows similar data from a telophase PtK, cell. ( 0 )=data; (- - - -) = cell. FIGURE a rectangle whose width equals the length of any single dark-staining portion of the midbody microtubules. This is thought to correspond to the length of the zone of overlap. The height of the rectangle was calculated in such a way that the area under the rectangle is equal to the area under the curve defined by the data. This graphic ruse pulls the zones of persumptive overlap into register and shows that the data are consistent with a sloppy region of interdigitation.
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tubules. This method will be presented in detail elsewhere, but our preliminary results resolve the question of the structure of the late anaphase interzone. FIGURE 10 shows the output of our computer-facilitated tubule-tracking study. These data demonstrate that the bulk of the tubules in the middle of the interzone are interdigitating, and that the reason why no 1:2:1 ratios have been observed by direct counts prior to late telophase31 is that the zone of overlap is sloppy. Because tubule tracking is tedious, and because it is inherently suspect due to the large number of correspondences that must be assigned to link up successive images of the same tubule, we have sought an alternate approach to the problem of anaphase spindle structure. Longitudinal thick sections of fixed cells are not useful, because there is so much matrix material in the mammalian spindle that the microtubules are essentially invisible in the high-voltage electron microscope. We have studied the structure of isolated mammalian spindles in collaboration with Sisken,3* but while these images are graphic, they are difficult to interpret in detail, both because the structure contains so many tubules (FIGURE 11) , and because anaphase spindles generally fall apart during preparation. If, however, cells are grown on coverslips, lysed with Tx into an equilibrium mixture of tubules and tubule subunits, then fixed, embedded, and thick-sectioned parallel to their spindle axes, one reaches a satisfactory combination of extracting nontubule material from the spindle and viewing only enough tubules to be comprehensible. The resulting micrographs are most impressive by far when presented in stereo (which will not be attempted here). Even without that perspective, however, we can detect several features of the spindle tubule redistribution during anaphase. Metaphase
The metaphase plate is usually crowded with chromosomes that obscure the spindle tubules. Occasional cells will by chance have few chromosomes showing in the section that contains the spindle poles (FIGURE 12). These images show many tubules near the poles, which end as they approach the metaphase plate. Stereo viewing shows that the majority of these ends are at the surface of the section, but some tubules end within the section before reaching the metaphase plate, some end at the metaphase plate, and some extend a considerable distance into the opposite half-spindle. The images show that the lengths of the tubules that do not end on a chromosome are heterodisperse. The lengths of chromosome-connected tubules are also varied even within a single bundle. Many do not reach from the kinetochore to the pole. These observations are consistent with the distribution profiles at metaphase, which show as few as 150 tubules immediately inside the spindle pole, but the number rises to about 1500 near the metaphase plate. The ending of nonchromosomal tubules short of the metaphase plate is also corroborated by the tubule counts. PtK, cells have about 11 chromosomes, with about 31 tubules per kinetochore. One of our normal metaphase distributions shows 1540 tubules maximum and about 900 minimum at the metaphase plate. If this drop were due entirely to chromosome tubules, there would have to be more than 50 tubules per kinetochore, rather than 31. Therefore some nonchromosomal tubules must end before they reach the spindle equator. FIGURE 13 is a diagram of our current understanding of the arrangement of microtubules in the metaphase spindle.
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FIGURE 10. ( a ) A computer-constructed drawing of the digitized positions of a bundle of microtubules from the interzone of a PtK, telophase. The two polygons linked by a line served as references for the preliminary alignment of this section with the next. ( b ) A set of correspondences between digitized tubule positions on adjacent sections. (c) A projection of tubule positions. The width of the bundle has been increased by a factor of 4, to improve the visibility of the tubules. (d) A drawing of tubule positions; the abcissa is distance along the spindle axis, the ordinate has no significance. These are data from 40 sections. There are many tubules that begin and end within this length along the spindle; these fragments have been set below as 10e and f are histograms of microtubule number versus posiseparate data. FIGURES tion, as labeled beside the figures.
FIGURE 11. A HeLa metaphase spindle isolated in hexylene glycol by J. Siskin. After isolation, it was fixed in glutaraldehyde and OsO,, dehydrated in acetone, and critical-point-dried in CO,, then micrographed in the IEM 1000. x 13,600. FIGURE 12. A high-voltage electron micrograph of a metaphase PtK, cell lysed in an equilibrium mixture of tubules and tubulin and fixed in glutaraldehyde Cx, then thicksectioned. The section is about 1 p m thick. The lack of chromosomes at the metaphase plate is fortuitous; the chromosomes lay in the adjacent sections on either side. x 10,400.
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Early anaphase in a fixed P t K , cell is shown in FIGURE 14. The chromosomes have moved nearer to the poles; a larger fraction of the chromosomeconnected tubules now appear to run all the way to the poles, as if the chromosome fiber had been pushed poleward. The interzone tubules can clearly be seen. Midanaphase is shown in FIGURE 15. T h e chromosomes have moved a little closer to the poles, and the spindle has elongated as the sister chromatids have moved apart. The interzone is now well defined, and substantial interzone tubule bundles are evident. These appear to be of uniform diameter and uniform tubule number, consistent with the tubule distribution profiles at midanaphase. By late anaphase, the chromosome tubules have shortened, and the chromosomes have drawn up around the poles (FIGURE 16). Tubule bundles still run the extent of the now much elongated interzone. A special region can be detected at the midregion of the interzone: the dark-staining tubule clusters are the stem 3fi The stem bodies form when the spindle is just about twice its metaphase length. FIGURES 17, 18, and 19 are higher-magnification pictures of interzone fibers from mid- and late anaphase and from telophase, showing 20 shows our understanding the changes in their appearance with time. FIGURE of the microtubule arrangement in a mammalian cell at late anaphase.
FIGURE 13. A diagram that shows our understanding of the arrangement of spindle tubules at metaphase in PtKI. Only half the spindle is shown, for the sake of simplicity.
DISCUSSION At the onset of anaphase, sister chromatids part, and sister chromosome fibers are transported in opposite directions. Subsequently, the chromosome fibers shorten. This decrease in length could be due to a loss of subunits at either the polar or the chromosomal end of the tubules that make up the bundle. Since anaphase chromosomes have been seen to stay at rest with respect to a variety of markers thought to be identified with the chromosome fibers,". :ji it is probable that the chromosome fiber is losing subunits from its polar end.2o The motions of the nonchromosomal tubules are not so easy to determine (for want of useful markers such as the chromosome, which display the position of tubule ends). Our observations with electron microscopy are completely consistent with the idea that microtubules of essentially fixed lengths slide over one another during the initial chromosome separation. They d o not, however, prove that the nonchromosomal tubules slide. Since we have used the ends of tubules as markers for tubule positions, one can imagine that a tubule might appear to slide because it adds subunits at one end and loses them at the other (FIGURE 21). O u r data d o not exclude this possibility, but the hypothesis does
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FIGURE14. Early anaphase in PtK,. PtKi. X 10,400.
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FIGURE 16. Late anaphase in PtK,. x 5,600. FIGURE 17. An interzone fiber from the cell shown in FIGURE 15. x 32,000. FIGURE 18. An interzone fiber from the cell shown in FIGURE 16. x 32,000. FIGURE 19. A thin section of a midbody isolated from HeLa cells. Thick sections of the midbody are hard to visualize because of the tight packing of the tubules, but this thin section indicates overlapping tubules. x 32,000.
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Annals New York Academy of Sciences FIGURE 20. A diagram that shows our understanding of the arrangement of spindle tubules :it late anaphase in mammalian cells.
introduce several complications into mitotic mechanisms: (1 ) A given nonchromosomal tubule must gain subunits at one end and lose them at the other, while the chromosome tubules are only losing subunits. ( 2 ) The loss of subunits must stop just as that particular tubule end has come into the right register with others, so that an approximately constant segment of tubule overlap is achieved at telophase. ( 3 ) Since spindle elongation will continue in some cells after the zone of overlap is established,3O the net addition of subunits at the growing end of a given tubule must continue after the net loss of subunits has stopped. The observations of the motions of nonchromosomal tubules during anaphase are more easily explained by sliding: a tubule that is in equilibrium with its environment is pushed along its own axis by hypothetical sliding forces. The cessation of sliding with a limited remaining overlap is easy to understand if the sliding forces are due to tubule-tubule interactions. Subsequent growth of the spindle is presumably due to the addition of tubule subunits onto these interzone fibers from the material released by disassembly of chromosome fibers. We therefore take these data as evidence that tubules slide. Due to the lack of an apparent polarity in spindle tubules as seen in the electron microscope, it is not possible to say in which direction the spindle tubules slide, if indeed they do. Our structural investigations can contribute nothing to an understanding of what might make the tubules move. The studies by Salmon and InouC support the idea that tubule disassembly is rate-limiting during anaphase in marine e g g ~ , ~398 ,but there are still many candidates for the role of mitotic motor. In our opinion, the two enzyme systems most likely to drive the chromosomes are the dynein of cilia and flagella or the actomyosin of muscle. There is at present only limited evidence that dynein occurs in spindles, and other spindle ATPases have been found.8 There is some evidence that actinlike fibers occur in spindles,40 but that point is still in question. Moreover, one needs to have more evidence than simply the presence or absence of a given component in the
FIGURE21. Two ways to move a microtuble, both of which will appear as sliding.
McIntosh et al.: Mechanism of Mitosis
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spindle in order t o assign it a functional role; one would like to know the enzymology of the machinery that actually does the work of moving the chromosomes. We are hopeful that our in vitro system for chromosome movement will provide the experimental handle to answer this question in the not too f a r distant future.
REFERENCES 1. 2. 3. 4. 5.
6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40.
BORISY,G. G. & E. W. TAYLOR. 1967. J. Cell Biol. 34: 535. BIBRJNG, T. & J. BAXANDALL. 1969. J. Cell Biol. 41: 577. KANE,R. E. 1965. J. Cell Biol. 25(2): 137. COHEN,W. D. & L. I. REBHUN.1970. J. Cell Sci. 6: 159. MIKI,T. 1963. Exp. Cell Res. 29: 92. HOFFMAN-BERLING, H. 1954. Biochim. Biophys. Acta 14: 182. R. & E. W. TAYLOR. 1968. Exp. Cell Res. 53: 372. WEISENBERG, MAZIA,D., C. PETZELT,R. 0. WJLLIAMS & I. MEZA. 1972. Exp. Cell Res. 70: 325. KANE,R. E. 1962. J. Cell Biol. 15: 279. REBHUN, L. I. & G. SANDER.1967. J. Cell Biol. 34: 859. GOLDMAN, R. D. & L. I. REBHUN.1969. J. Cell Sci. 4: 179. COHEN,W. D. & T. GOITLIEB. 1971. J. Cell. Sci. 9: 603. INOUB,S. & H. SATO. 1967. J. Gen. Physiol. 50: 259. BAJER,A. S. 1973. Cytobios 8: 139. J. A. 1968. J. Theoret. Biol. 20: 117. SUBIRANA, J. R., P. K. HEPLER& D. G. VANWIE. 1969. Nature 224: 659. MCINTOSH, NICKLAS,R. B. 1971. Mitosis. In Advances in Cell Biology. Vol. 2. D. M. Prescott, L. Goldstein & E. H. McConkey, Eds. : 225. Appleton-CenturyCrofts. New York, N.Y. SCHRADER, F. 1953. Mitosis. (2nd Ed.) Columbia University Press. New York, N.Y. MAZJA,D. 1961. The Cell Vol. 3. J . Bracket & A . E. Minsky, Eds. : 77. Academic Press Inc. New York, N.Y. BAJER,A. & J. MOLE-BAJER.1972. Int. Rev. Cytol. 34: Suppl. 1. HEATH,I. B. 1974. J. Cell Biol. 60: 204. JENSEN,C. & A. BAJER. 1973. J. Cell Biol. 59: 156a. BAJER,A. 1958. Chromosoma 9: 319. ALLEN,R. D., A. BAJER& J. LAFOUNTAIN. 1969. J. Cell Biol. 43: 4a. NICKLAS, R. B. & C. A. KOCH. 1972. Chromosoma 39: 1. 1967. J. Cell Biol. 35: 279. & L. C. RICHARDSON. BRINKLEY, B. R., P. MURPHY OLMSTED,J. B. & G. G. BORISY.1973. Biochemistry 12: 4282. & J. R. MCINTOSH.1974. CANDE,W. Z., J. A. SNYDER,D. SMITH,K. SUMMERS Proc. Nat. Acad. Sci. 71: 1559. LAFOUNTAIN, J. R. 1974. J. Ultrastruct. Res. 46: 268. BRINKLEY, B. R. & J. CARTWRIGHT. 1971. J. Cell Biol. 50: 416. J. R. & S. C. LANDIS.1971. J. Cell Biol. 49: 468. MCINTOSH, FUGE,H. 1973. Chromosoma 43: 109. JENSEN,C. & A. BAJER. 1973. Chromosoma 44: 73. J. R., J. SISKEN & L. CHU. In preparation. MCINTOSH, BELAR,K. 1929. Wilhelm Roux Arch. Entwicklungsmech. 118: 359. PAWELETZ, N. 1967. Naturwissenschaften 2 0 533. FORER,A. 1966. Chromosoma 19: 44. SALMON, E. D. 1973. J. Cell Biol. 59: 300a. SALMON, E. D. This monograph. FORER, A. & 0.BEHNKE.1972. Chromosoma 39: 145.
A n understanding of any machine is based upon detailed knowledge of the motions of its moving parts and of the way in which it uses fuel. Studies on the mechanism of muscle contraction are a clear example of how morphological and biochemical data can fit together to provide this information about a biological machine. T h e mechanism of chromosome motion can certainly be understood by a similar analysis, and over the past years important information about the mitotic apparatus ( M A ) has appeared in the literature. The M A has proved t o be difficult experimental material, because it is small, easy to kill, and less well organized than a striated muscle cell. Nonetheless, we know something about the chemistry of the isolated M A l-’, and quite a bit about its structure.9-12 Experimental studies such as those of InouC and Sato l 3 have contributed insights into the kinds of forces that hold the spindle together, and have suggested a model for how the machinery of the spindle might function to move the chromosomes. Alternative models f o r generation of spindle force have been proposed, such as Bajer’s “zipper hypothesis” and the several “sliding filament models,” but no one model stands out as uniquely suitable to account for all the available information. Part of the problem in determining the mechanisms of mitosis has been the insufficiency of detailed information concerning the motile events that occur within the spindle, although spindle structure at different times during mitosis has been studied in many different cells (reviews appear in References 17-20). The motion of the chromosomes during their congression to the metaphase plate and their subsequent parting and segregation at anaphase has been carefully described in numerous species,” but as Mazia’“ has pointed out, the role of the chromosome at mitosis o r meiosis is a passive one. For our purposes, a chromosome may be regarded as a marker for one end of the spindle fibers that attach to its kinetochores. Almost all students of mitosis agree that the chromosomes move because they are pushed o r pulled by the spindle fiber to which they are attached. The mechanism of chromosome motion, then, is really a question of the mechanism by which the cell exerts a force o n the chromosome fiber. This fiber is a bundle of microtubules, that range in number from 1 in a fungus 2 1 to over 100 in certain plants.” Mitotic forces, therefore, are basically forces that push on microtubules, although detailed observation of mitotic cells implicates the spindle in a variety of other motions that involve W e interpret the chromosome fragments and particles of diverse job of understanding mitotic mechanism as one of describing the motions of the spindle microtubules, and of analysing the molecular mechanisms that transduce chemical energy into the mechanical work necessary to cause these motions. One would like to watch spindle fiber motions in the light microscope, but
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individual tubules are of course too small to resolve optically. Polarization microscopy can yield a quantitative measure of optical anisometry and hence a measure related to the number of aligned fibers, but our efforts to use this method for determining spindle fiber motions during anaphase have failed (for what we believe to be solid and inescapable physical reasons, which will be described in detail elsewhere). Electron microscopy therefore seems to be the method of choice for the study, in spite of the obvious problems of the preparation artifact, and the fact that different time points in this ongoing process must be viewed in different cells. The most straightforward way to view the spindle fibers is to cut sections parallel to the spindle axis and to ask where the microtubules begin and end at different times during anaphase. Unfortunately, most spindle tubules leave the section before they truly end, so the longitudinal image is not useful for our purpose. We have circumvented this difficulty by cutting thick sections for viewing in a high-voltage electron microscope, by isolating whole spindles and viewing them with the same tool, and by cutting serial cross-sections that show the microtubules in transverse orientation. These sections can be aligned to allow tracking of individual tubules from section to section to find the tubule ends. I n comparing different stages of anaphase, we can then use the tubule ends as markers for tubule position. In a parallel investigation, we have been working to discover conditions that will allow chromosomes to continue their motion after the cell membrane has been lysed with detergent. These studies are directed toward elucidation of the enzyme system that exerts force upon the spindle tubules to make them move. Neither of these investigations is complete. We can maintain chromosome motion after lysis, but we do not yet know which enzymes are important for motility. We have determined certain facts about spindle fiber motion, but we do not yet know exactly how the fibers move. This paper must therefore be regarded as a progress report, in which we will describe our findings to date and define the directions of our plans for the immediate future.
MATERIALS A N D METHODS Mammalian cells (HeLa, WI-38, CHO, and PtK,) were cultured by standard techniques. For light microscopy, the cells were subcultured on glass cover slips which can be inverted onto a glass slide (using slivers of parafilm as spacers) and waxed down to form a simple chamber that will support the initiation and completion of cell division for about 4 hours. For electron microscopy, the cells were fixed after the method of Brinkley et aI.,?O or in a mixture of 4% glutaraldehyde, 4% 20 M Carbowax@ (Cx) in 0.1 M piperazine-N-N’-(2-ethane sulfonic acid) adjusted to pH 6.9 with NaOH (PIPES buffer). This fixative must be made up fresh before use. Fixed cells were rinsed in buffer, treated with OsO,, dehydrated, and embedded by standard techniques. Single cells were selected from wafers of plastic with a phase microscope, excised, and serially sectioned in a chosen orientation for electron microscopy. For microscopy in a high-voltage electron microscope, the cells were fixed and embedded as described and sectioned at a nominal thickness of 1 pm. About 10 sections were picked up on a carbon and Formvafl-coated slot grid, stained with uranyl acetate and lead citrate for about 30 min each at room
McIntosh et al.: Mechanism of Mitosis
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temperature, and observed in a J E M 1000 (operating at an accelerating voltage of 1,000 kV) . Polarization optical measurements were made with a Brace-Kohler compensator and Zeiss polarization optics. Photographs were taken o n Kodak tri-X film developed in Diafinem. Anaphase motions were recorded o n Plus-X reversal film with a Bolex movie camera and Zeiss differential interference contrast optics. W e prepared repolymerization-competent microtubule protein, using a modification of the method of Olmsted and Borisy." The brains of I-3-day-old chicks were homogenized at 0" C in 0.1 M PIPES buffer at p H 6.9 that contained 1 m M E G T A and 2.5 m M G T P . T h e homogenate was spun at 35,000 X g for 30 min at 0" C; the supernatant was collected and made 30% in glycerol, then warmed to 37" C for 25 min, and spun at 35,000 X g for 1 hour at 35" C. T h e pellet was resuspended in 0.1 M PIPES at p H 6.9, 1 m M EDTA, 1 m M G T P at 37"C, and immediately pelleted again at the same temperature. This material was frozen in liquid nitrogen and stored at -70" C for u p to 1 week. A frozen pellet was homogenized in glycerol-free buffer with 1 m M G T P at 0" C, extracted for 1 hour and spun at 35,000 X g f o r 30 min. This supernatant was our standard tubulin preparation. Its purity was assessed by electrophoresis in 0.1% sodium dodecyl sulfate, and its capacity to polymerize was measured viscometrically and by direct view of negatively stained specimens in the electron microscope. To study the motion of chromosomes in lysed cells, we made coverslip preparations as described above and selected a cell with a suitable spindle. T h e wax seal was then broken, and a preparation of tubulin a n d / o r C x with 0.02-0.2% Triton@ X-100 ( T x ) added, was drawn under the coverslip with filter paper. RESULTS Stability
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When mammalian cells at metaphase are lysed with 0.01 %-0.1% T x in a buffer that supports brain tubulin polymerization in vitro, the spindle as seen in polarization optics disappears.Z5 As the tubulin concentration is raised, the rate of disappearance decreases (FIGURE 1 ). At high concentration, the spindle birefringence ( B R ) will increase. If the preparation of tubulin is warmed t o 37" C for 25 min prior to its use for lysis, then the spindle BR stays approximately constant for all concentrations of tubulin that we have tried above 1 mg/ml. Changes in spindle BR at lysis are apparently dependent upon the concentration of microtubule subunits, not upon the total concentration of tubulin present in the lysing solution. W e have determined that the increase in spindle BR is accompanied by a significant increase in the number of tubules per spindle (FIGURE 2). While trying to correlate BR with microtubule number, we observed, as others have before,l32 O' that spindle BR can drop significantly during fixation f o r electron microscopy. Fixation in the glutaraldehyde-Carbowax mixture described above will generally preserve spindle BR in mammalian cells for the first 20 min after the addition of fixative. Higher concentrations of Cx will sometimes promote an increase in spindle BR during fixation. Multiple experiments show,
Annals New York Academy of Sciences
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FIGURE1. (a) A graph of BR versus time after lysis for PtK, cells treated with 0.1 M PIPES buffer (pH 6.9), 1 mM EGTA, 2 mM GTP, and 0.1% Tx,c onta hhg difl'erent concentrations of tubulin: (m)~ 0 . 4 4mg/ml; ( 0 )=0.88 mg/ml; ( A ) = 1.75 mg/ml; (0) ~ 3 . mg/ml; 5 and (0) =7.0 mg/ml. All preparations were placed at 37" C for 5 min prior t o their use as a lysing solution. FIGUREl b shows BR versus time for cells lysed in different concentrations of tubulin in the above buffer and warmed to 37" C for 25 min prior to use in lysis: ( 0 ) = 0 . 7 5 mg/ml; ( O ) = 1.5 mg/ml; and ( A ) = 3 . 0 mg/ml. The error margin is the same for both graphs, and is a statement of the range of the reproducibility of multiple measurement of the same point.
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FIGURE 2. Qualitative correlations between spindle BR and spindle tubule number. FIGURES 2a, b, and c are polarization optical and electron-microscopic pictures of a cell lysed in an equilibrium mixture of tubules and tubulin. Quantitative data are 3 and 4 (cell 71). FIGURES 2d, e, and f are similar pictures of presented in FIGURES a cell lysed in recently warmed tubulin. (This is cell 76 in FIGURES 3 and 4.) Light micrographs, x 800; electron micrographs, x 48,000.
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FIGURE 3. A graph of BR versus time, which shows the variation of BR with fixation. Three lysed cells are monitored with different concentrations of tubulin in the lysing solution, to show BR changes with time both before and during fixation. The =Cell 76 lysed in about 10 mg/ following cells are fixed in glutaraldehyde-Cx. (0) ml recently warmed tubulin; ( 0 )=cell 77 lysed in about 8 mg/ml recently warmed tubulin; ( A ) =cell 71 lysed in about 8 mg/ml tubulin, prewarmed for 15 min; 70, lysed in about 8 mg/ml tubulin warmed for 15 min and fixed in phos(.)=cell phate-buffered glutaraldehyde; ( 0 )=cell 87, an unlysed cell fixed in glutaraldehyde-
cx.
however, that occasional cells will retain their BR in phosphate-buffered glutaraldehyde, and occasional cells will lose BR during fixation in the glutaraldehyde-Cx mixture. These results are independent of whether or not the cell has been lysed prior to fixation (FIGURE 3 ) . FIGURE 4 shows the microtubule distribution profiles for some of the cells whose BR records are presented in FIGURE 3. Our results may be summarized as follows. When BR increases, the number of tubules in the spindle increases. The relation between the two is not linear, even when BR is kept constant during glutaraldehyde fixation. If BR is allowed to vary during fixation, the relation between BR before fixation and the number of tubules as seen in the electron microscope is not significantly altered, which indicates that loss of BR during glutaraldehyde fixation is not accompanied by significant loss in the number of microtubules. These statements apply both to lysed and unlysed cells (FIGURES 3 and 4). When cells are lysed in high concentrations of microtubule subunits (5-15 mg/ml, warmed from 0" C to 37" C immediately before use in lysis), the average tubule length as well as the average tubule number increases. FIGURE 5a is a metaphase cell lysed in a mixture of tubules and tubule subunits that is approximately at equilibrium (4 mg/ml warmed for 25 min, viscosity5b is a cell lysed in recently warmed tubulin. In FIGURE plateaued). FIGURE 5b there are tubules that appear to skewer the chromosomes and continue in
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straight lines for distances that exceed the spindle length, while in FIGURE Sa, the spindle shape is well preserved. This hypertrophy of number and length is not found in all spindle tubules. We have counted the number of tubules attached to individual kinetochores in normal, fixed cells and in augmented spindles. At metaphase, the chromosomes of one PtK, cell showed a n average of 31 10 tubules per kinetochore ( N = 8 ) ; that cell had a maximum of 1,575 tubules. A spindle from the same experiment as seen in FIGURE 5b showed a maximum of 6,970 tubules, but only 30 2 8 tubules per kinetochore ( N = 4). Thus the increase in tubule number is exclusively of nonchromosomal tubules in our cells. We have previously observed that the chromosome tubules are the most stable in the spindle with respect to extraction at low tubulin concentrations,2s which indicates that both their polymerization and depolymerization rates in v i m are substantially lower than those of the nonchromosome tubules.
*
Functional Stability of the Spindle
The motile capacity of the spindle is considerably more labile than the structure of the spindle as seen in the light and electron microscopes. The BR of the spindle will vary u p or down with tubulin concentration after lysis in
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Section Number FIGURE 4. Microtubule number versus position along the spindle axis measured in numbers of sections. Each section is about 900 A thick. These data correspond to 3. (.)=Cell 76; ( O ) = some of the cells whose BR records are shown in FIGURE cell 77; (A)=cell 71; (.)=cell 70. The BR increase observed with lysis in high concentrations of tubule subunits is reflected in the increase in microtubule number.
FIGURE 5. (a) A cell lysed in an equilibrium mixture of tubules and tubulin. The 5b is a cell from the sections for both 5a and 5b are about 1 gm thick. FIGURE experiment that produced cell 76. The quadrupling of the normal tubule number is accompanied by an increase in average tubule length and an extension of tubules out beyond the normal domain of the spindle. Presumably the length increase accounts for the image in FIGURE, 2 in which the chromosomes are riddled with tubules, for the peak rather than a trough in the middle of the top microtubule distribution curve of FIGURE 4, and for the tubules that radiate from the poles out of the spindle 5b. x 18,700. in FIGURE
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0.1% Tx, but chromosome motion will generally cease in detergent solutions stronger than 0.03%. In 0.03% Tx, 1 mM MgCI,, 1 mM each ATP and GTP, 1 mM dithiothreitol, and about 4 mglml tubulin brought to approximate equilibrium, the chromosomes will continue to move for some time after addition of the detergent (FIGURE 6 ) . Their rate of motion is less than the physiological rate by a factor of 3-5, but movement occurs, and the cell appears to be well lysed. Addition of 2-5% Cx to the lysing solution makes the chromosome motion considerably more rapid, but it also interferes with lysis and introduces the possibility that the cell membrane is not truly disrupted. Since Cx in PIPES buffer will stabilize a ~pindle,’~ we have tried to develop a Tx-Cx lysing solution without tubulin which would give reproducible lysis and nucleotide-dependent motion of the chromosomes, in order to facilitate experiments on the nature of the enzyme system of anaphase. These effort have been only partially successful. Treatment of the cells with 0.125% Tx and 5% Cx in 0.1 M PIPES buffer will induce extensive surface blebbing of the cell. The cell processes withdraw, the mitochondria spherulate, the spindle poles become distinct, and the cytoplasm pales in about 45 sec. If the solution is left on the cell for a few min, the spindle fibers become visible in the light microscope, and the chromosomes will no longer move. If, on the other hand, the lysing solution is replaced by about 2% Cx and 0.1-2.5 mM ATP and MgCI,, the the chromosomes will continue to move (FIGURE 7 ) . Cells that are fixed while the chromosomes are moving in this solution and viewed in the electron microscope do not look extracted. The chromosomes have an unusual texture, the cell surface is pitted, and the mitochondria are peculiar, but clearly the cell is not extensively lysed (FIGURE 7b). Nonetheless, if NaF is added with the lysing solution, the chromosomes stop rapidly, while N a F added to the medium without detergent has no effect upon chromosome motion (FIGURE 7a). These results indicate that chromosome motion can occur in cells lysed in different ways, but the data are too sparse to allow us to conclude anything about the nature of the mitotic motors. Structural Studies on Fixed Cells
Several recent studies of spindle structure have employed the method of counting the numbers of microtubules per spindle cross-section as a function of position on the spindle axis at different times during anaphase, to see how the spindle tubules are redistributed.Sn-?3 The available studies on mammalian cells are in disagreement on one important point: Brinkley and Cartwright 3n found that the distribution of tubules in the anaphase interzone did not change with time, while in our earlier work, we found that it did.3l We have reinvestigated this point using CHO and PtK, cells, and have confirmed our findings with HeLa and WI-38 (FIGURES 8 and 9 ) . In early anaphase, the number of tubules in the interzone does not vary with position along the spindle axis, while in late anaphase and telophase, we find, with Brinkley et C Z ~ .that , ~ ~there is a larger number of tubules at the middle of the interzone than on either side.31 As before, however, the number of tubules in the middle of the interzone is not twice the number on either side, as might be expected from the interdigitation of two families of tubules, nor is it uniform; the ratios are about 1:1.5: 1. We are attempting to clarify the structure of the middle of the interzone by reconstructing it in 3 dimensions, using serial cross-sections to track the
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FIGURE 6. Chromosome separation versus time during anaphase in PtKl cells. In 6a ( A ) = a r e a normal, unlysed cell; (A)=& cell lysed as described in the text; A = a n observation made after lysis had begun. The bar on the ordinate shows 6b is the cell recorded in FIGURE 6a t e metaphase length of both spindles. FIGURE while at metaphase; FIGURE 6c is after the onset of anaphase, but before lysis; FIGURE 6d is just after lysis, and corresponds to the boxed triangle in FIGURE 6a. FIGURE 6c is at 2224 sec. x 1,360.
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FIGURE 7. (a) A graph of chromosome separation versus time. ( A ) = A PtKl cell in tissue culture medium (5 m M NaF was added prior to the boxed triangle); (.)=a cell treated with lysing solution prior to the first boxed square, and treated with 2.5% Cx, 2.5 mM ATP and MgCL in PIPES prior to the second boxed square; ( A ) = a cell treated with lysing solution prior to the first boxed triangle and with 2.5% Cx, 2.5 mM ATP and MgCL, and 5 mM NaF prior to the second boxed triangle. FIGURE 7b is a cell treated exactly as the cell represented by (B) in FIGURE 7a. but fixed during chromosome motion. x 13,500.
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FIGURE 8. ( a ) The microtubule distribution profile of a CHO cell fixed during early anaphase. The diagram shows the approximate distribution of the chromosomes. The C’s show where the centrioles lay. (The dotted C indicates that our sectioning started too late; this position is an estimate, based upon the convergence of the spindle tubules.) FIGURE 8b is a CHO cell later in anaphase. FIGURE 8c shows data from the midregion of the interzone of a CHO telophase.
41 8
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McIntosh et al.: Mechanism of Mitosis
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Section Number FIGURE 9. ( a ) The microtubule distribution profile from an early anaphase PtKl 9b shows similar data from a telophase PtK, cell. ( 0 )=data; (- - - -) = cell. FIGURE a rectangle whose width equals the length of any single dark-staining portion of the midbody microtubules. This is thought to correspond to the length of the zone of overlap. The height of the rectangle was calculated in such a way that the area under the rectangle is equal to the area under the curve defined by the data. This graphic ruse pulls the zones of persumptive overlap into register and shows that the data are consistent with a sloppy region of interdigitation.
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tubules. This method will be presented in detail elsewhere, but our preliminary results resolve the question of the structure of the late anaphase interzone. FIGURE 10 shows the output of our computer-facilitated tubule-tracking study. These data demonstrate that the bulk of the tubules in the middle of the interzone are interdigitating, and that the reason why no 1:2:1 ratios have been observed by direct counts prior to late telophase31 is that the zone of overlap is sloppy. Because tubule tracking is tedious, and because it is inherently suspect due to the large number of correspondences that must be assigned to link up successive images of the same tubule, we have sought an alternate approach to the problem of anaphase spindle structure. Longitudinal thick sections of fixed cells are not useful, because there is so much matrix material in the mammalian spindle that the microtubules are essentially invisible in the high-voltage electron microscope. We have studied the structure of isolated mammalian spindles in collaboration with Sisken,3* but while these images are graphic, they are difficult to interpret in detail, both because the structure contains so many tubules (FIGURE 11) , and because anaphase spindles generally fall apart during preparation. If, however, cells are grown on coverslips, lysed with Tx into an equilibrium mixture of tubules and tubule subunits, then fixed, embedded, and thick-sectioned parallel to their spindle axes, one reaches a satisfactory combination of extracting nontubule material from the spindle and viewing only enough tubules to be comprehensible. The resulting micrographs are most impressive by far when presented in stereo (which will not be attempted here). Even without that perspective, however, we can detect several features of the spindle tubule redistribution during anaphase. Metaphase
The metaphase plate is usually crowded with chromosomes that obscure the spindle tubules. Occasional cells will by chance have few chromosomes showing in the section that contains the spindle poles (FIGURE 12). These images show many tubules near the poles, which end as they approach the metaphase plate. Stereo viewing shows that the majority of these ends are at the surface of the section, but some tubules end within the section before reaching the metaphase plate, some end at the metaphase plate, and some extend a considerable distance into the opposite half-spindle. The images show that the lengths of the tubules that do not end on a chromosome are heterodisperse. The lengths of chromosome-connected tubules are also varied even within a single bundle. Many do not reach from the kinetochore to the pole. These observations are consistent with the distribution profiles at metaphase, which show as few as 150 tubules immediately inside the spindle pole, but the number rises to about 1500 near the metaphase plate. The ending of nonchromosomal tubules short of the metaphase plate is also corroborated by the tubule counts. PtK, cells have about 11 chromosomes, with about 31 tubules per kinetochore. One of our normal metaphase distributions shows 1540 tubules maximum and about 900 minimum at the metaphase plate. If this drop were due entirely to chromosome tubules, there would have to be more than 50 tubules per kinetochore, rather than 31. Therefore some nonchromosomal tubules must end before they reach the spindle equator. FIGURE 13 is a diagram of our current understanding of the arrangement of microtubules in the metaphase spindle.
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FIGURE 10. ( a ) A computer-constructed drawing of the digitized positions of a bundle of microtubules from the interzone of a PtK, telophase. The two polygons linked by a line served as references for the preliminary alignment of this section with the next. ( b ) A set of correspondences between digitized tubule positions on adjacent sections. (c) A projection of tubule positions. The width of the bundle has been increased by a factor of 4, to improve the visibility of the tubules. (d) A drawing of tubule positions; the abcissa is distance along the spindle axis, the ordinate has no significance. These are data from 40 sections. There are many tubules that begin and end within this length along the spindle; these fragments have been set below as 10e and f are histograms of microtubule number versus posiseparate data. FIGURES tion, as labeled beside the figures.
FIGURE 11. A HeLa metaphase spindle isolated in hexylene glycol by J. Siskin. After isolation, it was fixed in glutaraldehyde and OsO,, dehydrated in acetone, and critical-point-dried in CO,, then micrographed in the IEM 1000. x 13,600. FIGURE 12. A high-voltage electron micrograph of a metaphase PtK, cell lysed in an equilibrium mixture of tubules and tubulin and fixed in glutaraldehyde Cx, then thicksectioned. The section is about 1 p m thick. The lack of chromosomes at the metaphase plate is fortuitous; the chromosomes lay in the adjacent sections on either side. x 10,400.
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Early anaphase in a fixed P t K , cell is shown in FIGURE 14. The chromosomes have moved nearer to the poles; a larger fraction of the chromosomeconnected tubules now appear to run all the way to the poles, as if the chromosome fiber had been pushed poleward. The interzone tubules can clearly be seen. Midanaphase is shown in FIGURE 15. T h e chromosomes have moved a little closer to the poles, and the spindle has elongated as the sister chromatids have moved apart. The interzone is now well defined, and substantial interzone tubule bundles are evident. These appear to be of uniform diameter and uniform tubule number, consistent with the tubule distribution profiles at midanaphase. By late anaphase, the chromosome tubules have shortened, and the chromosomes have drawn up around the poles (FIGURE 16). Tubule bundles still run the extent of the now much elongated interzone. A special region can be detected at the midregion of the interzone: the dark-staining tubule clusters are the stem 3fi The stem bodies form when the spindle is just about twice its metaphase length. FIGURES 17, 18, and 19 are higher-magnification pictures of interzone fibers from mid- and late anaphase and from telophase, showing 20 shows our understanding the changes in their appearance with time. FIGURE of the microtubule arrangement in a mammalian cell at late anaphase.
FIGURE 13. A diagram that shows our understanding of the arrangement of spindle tubules at metaphase in PtKI. Only half the spindle is shown, for the sake of simplicity.
DISCUSSION At the onset of anaphase, sister chromatids part, and sister chromosome fibers are transported in opposite directions. Subsequently, the chromosome fibers shorten. This decrease in length could be due to a loss of subunits at either the polar or the chromosomal end of the tubules that make up the bundle. Since anaphase chromosomes have been seen to stay at rest with respect to a variety of markers thought to be identified with the chromosome fibers,". :ji it is probable that the chromosome fiber is losing subunits from its polar end.2o The motions of the nonchromosomal tubules are not so easy to determine (for want of useful markers such as the chromosome, which display the position of tubule ends). Our observations with electron microscopy are completely consistent with the idea that microtubules of essentially fixed lengths slide over one another during the initial chromosome separation. They d o not, however, prove that the nonchromosomal tubules slide. Since we have used the ends of tubules as markers for tubule positions, one can imagine that a tubule might appear to slide because it adds subunits at one end and loses them at the other (FIGURE 21). O u r data d o not exclude this possibility, but the hypothesis does
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Annals New York Academy of Sciences
FIGURE14. Early anaphase in PtK,. PtKi. X 10,400.
x
10,400. FIGURE15. Mid anaphase in
FIGURE 16. Late anaphase in PtK,. x 5,600. FIGURE 17. An interzone fiber from the cell shown in FIGURE 15. x 32,000. FIGURE 18. An interzone fiber from the cell shown in FIGURE 16. x 32,000. FIGURE 19. A thin section of a midbody isolated from HeLa cells. Thick sections of the midbody are hard to visualize because of the tight packing of the tubules, but this thin section indicates overlapping tubules. x 32,000.
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Annals New York Academy of Sciences FIGURE 20. A diagram that shows our understanding of the arrangement of spindle tubules :it late anaphase in mammalian cells.
introduce several complications into mitotic mechanisms: (1 ) A given nonchromosomal tubule must gain subunits at one end and lose them at the other, while the chromosome tubules are only losing subunits. ( 2 ) The loss of subunits must stop just as that particular tubule end has come into the right register with others, so that an approximately constant segment of tubule overlap is achieved at telophase. ( 3 ) Since spindle elongation will continue in some cells after the zone of overlap is established,3O the net addition of subunits at the growing end of a given tubule must continue after the net loss of subunits has stopped. The observations of the motions of nonchromosomal tubules during anaphase are more easily explained by sliding: a tubule that is in equilibrium with its environment is pushed along its own axis by hypothetical sliding forces. The cessation of sliding with a limited remaining overlap is easy to understand if the sliding forces are due to tubule-tubule interactions. Subsequent growth of the spindle is presumably due to the addition of tubule subunits onto these interzone fibers from the material released by disassembly of chromosome fibers. We therefore take these data as evidence that tubules slide. Due to the lack of an apparent polarity in spindle tubules as seen in the electron microscope, it is not possible to say in which direction the spindle tubules slide, if indeed they do. Our structural investigations can contribute nothing to an understanding of what might make the tubules move. The studies by Salmon and InouC support the idea that tubule disassembly is rate-limiting during anaphase in marine e g g ~ , ~398 ,but there are still many candidates for the role of mitotic motor. In our opinion, the two enzyme systems most likely to drive the chromosomes are the dynein of cilia and flagella or the actomyosin of muscle. There is at present only limited evidence that dynein occurs in spindles, and other spindle ATPases have been found.8 There is some evidence that actinlike fibers occur in spindles,40 but that point is still in question. Moreover, one needs to have more evidence than simply the presence or absence of a given component in the
FIGURE21. Two ways to move a microtuble, both of which will appear as sliding.
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spindle in order t o assign it a functional role; one would like to know the enzymology of the machinery that actually does the work of moving the chromosomes. We are hopeful that our in vitro system for chromosome movement will provide the experimental handle to answer this question in the not too f a r distant future.
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