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Micromanipulation of Chromosomes in Mitotic Vertebrate Tissue Cells: Tension Controls the State of Kinetochore Movement
EXPERIMENTAL CELL RESEARCH ARTICLE NO.
235, 314–324 (1997)
EX973691
Micromanipulation of Chromosomes in Mitotic Vertebrate Tissue Cells: Tension Controls the State of Kinetochore Movement Robert V. Skibbens1 and E. D. Salmon2 Department of Biology, University of North Carolina, Chapel Hill, North Carolina 27599-3280
INTRODUCTION In mitotic vertebrate tissue cells, chromosome congression to the spindle equator in prometaphase and segregation to the poles in anaphase depend on the movements of kinetochores at their kinetochore microtubule attachment sites. To test if kinetochores sense tension to control their states of movement poleward (P) and away from the pole (AP), we applied an external force to the spindle in preanaphase newt epithelial cells by stretching chromosome arms with microneedles. For monooriented chromosomes (only one kinetochore fiber), an abrupt stretch of an arm away from the attached pole induced the single attached kinetochore to persist in AP movement at about 2 mm/ min velocity, resulting in chromosome movement away from the pole. When the stretch was reduced or the needle removed, the kinetochore switched to P movement at about 2 mm/min and pulled the chromosome back to near the premanipulation position within the spindle. For bioriented chromosomes (sister kinetochores attached to opposite poles) near the spindle equator, stretching one arm toward a pole placed the kinetochore facing away from the direction of stretch under tension and the sister facing toward the stretch under reduced tension or compression. Kinetochores under increased tension exhibited prolonged AP movement while kinetochores under reduced tension or compression exhibited prolonged P movement, moving the centromeres at about 2 mm/min velocities off the metaphase plate in the direction of stretch. Removing the needle resulted in centromere movement back to near the spindle equator at similar velocities. These results show that tension controls the direction of kinetochore movement and associated kinetochore microtubule assembly/disassembly to position centromeres within the spindle of vertebrate tissue cells. High tension induces persistent AP movement while low tension induces persistent P movement. The velocity of P and AP movement appears to be load independent and governed by the molecular mechanisms which attach kinetochores to the dynamic ends of kinetochore microtubules. q 1997 Academic Press
1 Current address: Johns Hopkins School of Medicine, Department of Molecular Biology and Genetics, Baltimore, MD 21205. 2 To whom correspondence and reprint requests should be addressed. Fax: (919) 962-1625. E-mail: [email protected].
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0014-4827/97 $25.00 Copyright q 1997 by Academic Press All rights of reproduction in any form reserved.
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During mitosis, kinetochores attach chromosomes to spindle poles by capturing and stabilizing microtubule plus ends. In vertebrate tissue cells studied to date [1– 13 and references therein], kinetochores exhibit ‘‘directional instability,’’ abruptly switching between constant velocity (about 2 mm/min) states of movement poleward (P) or away from the pole (AP). In newt and PtK1 cells, fluorescent marks on the lattice of kinetochore microtubules move slowly poleward at about 0.5 mm/min or less [14, 15]. Thus, during P movement, kinetochore microtubule shortening occurs predominantly at the kinetochore and, during AP movement, kinetochore microtubule growth occurs exclusively at the kinetochore. Kinetochore P movement stretches its centromere chromatin poleward until the kinetochore abruptly switches to AP movement. AP movement reduces centromere stretch until the kinetochore switches back to P movement [4, 5]. The molecular mechanisms that attach kinetochores to the dynamic plus ends of kinetochore microtubules and produce the constant velocity states of kinetochore P and AP movement are not understood. P movement produces pulling forces [reviewed in 8, 9], but AP movement occurs mainly under tension and rarely pushes [4, 7, 12]. There is evidence that both plus- and minus-end-directed microtubule motor proteins bound to kinetochores as well as the shortening and growth phases of plus end microtubule dynamic instability are involved [9, 16, 17]. A key issue in mitosis is what attached kinetochores sense to control the durations of their P and AP states of movement. This question is important because the congression of sister chromatid pairs to the spindle equator in prometaphase and their segregation to opposite poles in anaphase depends on the regulation of the relative durations of P and AP kinetochore movement [4]. Chromosome congression to the equator in prometaphase occurs only when the sister kinetochore facing the equator persists in P movement toward the equator and the sister facing away from the equator persists in AP movement toward the equator; net movement of the centromere does not occur when both attached sisters
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move P or AP at the same time [4]. In anaphase, the segregation of separated sisters to the poles occurs only when the kinetochores persist in P movement, pulling their chromosome arms poleward [4, 6]. There is evidence that kinetochore switching between P and AP states depends on kinetochore phosphorylation [17], the number of kinetochore microtubules [18], or factors which affect microtubule assembly [9]. In addition, several reports indicate that kinetochores primarly sense tension to control the durations of their P and AP states [4–9, 19–22]. For example, on average, kinetochores on bioriented chromosomes at the metaphase plate are under tension because their centromere chromatin is stretched beyond their unattached rest length [7, 13, 19]. Reducing kinetochore tension on a bioriented chromosome by either ablating the sister [2, 19, 20] or ablating the centromereic chromatin between sister kinetochores results in prolonged P movement of the undamaged kinetochore at velocities similar to kinetochore poleward motion in anaphase [6, 12]. A kinetochore tensiometer model has been proposed for vertebrate tissue cells where the direction, but not the velocity, of kinetochore movement is sensitive to tension: higher tension increases the probability of the kinetochore switching from P and persisting in AP movement while low tension or compression increases the probability of switching from AP and persisting in P movement [4–8]. During mitosis, changes in kinetochore tension are produced by the relative motions of sister kinetochores and their chromosome arms. Poleward movement of chromosome arms is resisted by spindle polar microtubule arrays which push the arms away from the pole [1–8, 12]. These ‘‘polar ejection forces’’ are thought to increase in strength as microtubule density increases closer to the poles. Ejection forces may be produced by chromosome-associated microtubule motor proteins (chromokinesins) or the growth dynamics of the polar microtubule arrays and contribute significantly to the tension which biases a kinetochore into AP movement near the pole [2, 3, 12, 23, 24]. In this paper, we tested the kinetochore tensiometer model in mitotic newt lung cells by examining how kinetochores respond—in terms of direction, duration, and velocity of movement—to the application of a force external from the spindle produced by stretching a chromosome arm with a microneedle. Previously, such studies were performed by Nicklas [21, 22], but they were performed only in grasshopper spermatocytes and have not been tested in vertebrate mitotic cell systems. Several lines of evidence indicate that kinetochores in invertebrate meiotic cells behave differently than kinetochores in vertebrate mitotic cells [25] (i.e., the kinetochores in grasshopper spermatocytes move more slowly (0.5 mm/min) and without the amplitude of oscillation found in vertebrate tissue cells [1–12, 19, 21]). Furthermore, sister kinetochores are paired and act in
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concert in meiosis I cells while sister kinetochores are separate and act independently in mitotic cells [4, 6]. Therefore, it was important to test the kinetochore tensiometer model in a vertebrate cell system. Our micromanipulation studies on newt cells show that the direction of kinetochore movement is sensitive to tension as predicted by the kinetochore tensiometer model [6] and Nicklas’ [21, 22] micromanipulation in meiotic grasshopper spermatocytes. The constant velocities of kinetochore AP movement during mitosis indicate that kinetochores normally govern their AP velocity [4, 5, 12]. Our micromanipulation results show that the kinetochore is able to govern the rate at which it moves AP in response to a substantial pulling force. MATERIAL AND METHODS Tissue culture. Newt lung cultures were prepared as described [2, 26]. Oregon newts (Taricha granulosa) were obtained from Charles Sultan (Nashville, TN), the lungs dissected from the animal, minced, washed three times in calcium-free/magnesium-free Hanks Ringer solution for amphibians, trypsinized for 5 minutes, and placed in L15 media supplemented with fetal bovine serum and antibiotics. Lung tissue explants were cultured overnight, placed in Rose chambers, and incubated 10 to 15 days at room temperature (22–267C). For experimentation, Rose chambers were disassembled and the coverslips containing monolayers were treated as described below. Micromanipulation chambers. Prior to observation, a Rose chamber was disassembled and the coverslip inverted onto two metal plates (1.5 1 3 1 0.0615 in with a 20-mm-diameter central hole) previously sealed together with vacuum grease. The coverslip was affixed to one face of the metal plate with a thin layer of grease, sealed with VALAP (1:1:1 of Vaseline, lanolin, and paraffin), and the whole structure inverted so that the coverslip formed the floor of the micromanipulation chamber. The double thickness of metal allowed for a maximum volume of fresh medium (about 600 ml) to be placed over the explant and still allowed a thin layer of mineral oil to be layered over the medium. Mineral oil proved nontoxic to cells and prolonged cell viability by delaying changes in osmolarity caused by media evaporation. The mineral oil overlay also provided a flat surface so image quality was maintained. Micromanipulation needles. Needles appropriate for micromanipulation were generated in a three-step process using custom glass tubing (Pyrex: o.d. 0.8 mm, i.d. 0.6 mm, length 100 mm—no internal filament) obtained from Drummund Scientific Co. (Broomall, PA). First, a 457 angle in the filament was placed near one end of the pipette using a fine butane flame and forceps. The region of the pipette just beyond the bend was pulled over the flame with forceps to form a fine tip. This tip was clipped to an appropriate length (tip distance over 4–5 mm normal from the cylinder axis interfered with proper positioning of the microscope condenser). In the second step, the tip of the needle was sealed and finely tapered using a Microforge De Fonbrune, Series A, No. 101 (equipped with 151 oculars) at a heat setting of about 6–8. This filament was too long and flexible to encourage chromosome movement through the stiff cell cortex or intermediate filament cage (see Results). The flexible portion of the needle tip was broken off using the cold Microforge heating element. In the third and final step, the needle was pulled out to a very fine but inflexible short tip using the Microforge at a low heat setting (0.0–0.5). Final needle selection was performed using a Zeiss Neofluar 25X 0.6NA Phase 2 objective on the IM stand. Micromanipulation. Needle control was achieved using a Narishige pneumatic micromanipulation device assembled onto a crossbar affixed to the condenser/illumination tower of the Zeiss IM microscope stand. Direct coupling of the micromanipulation device to the
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illumination tower was critical for dampening needle vibrations. Fine control was achieved with the Narishige hanging micromanipulation joystick placed off the vibration isolation platform. Phase microscopy and video. Images of newt lung cells were obtained using a Zeiss IM stand assembled on top of a pneumatic vibration isolation platform and equipped with a 60W Tungsten bulb, high NA Phase condenser complete with a Phase 3 annulus, Nikon 100X (NA 1.4) Phase 4 objective, and a 41 projection lens. Images were collected using a DAGE-MTI NC-67MDX Newvicon camera and recorded in real time onto 3/4 inch format Umatic video tape using a Sony VO-5800H video cassette recorder. Selected sequences were time lapsed (1 frame every 2 s) into a Panasonic TQ2028F Optical Memory Disc Recorder (OMDR) for analysis. Tracking and analyses. Tracking and analyses were performed as previously described [4, 6]. Computer-generated cursors were superimposed on images containing the objects to be tracked. Gray values for each pixel in an 8X8 array for each cursor were used as a template to search consecutive images on the OMDR. The computer recorded frame number, X and Y coordinates for each object, and time, which were later converted to distance-versus-time plots and analyzed using the in-house program, Single Frame Movement (SFM). For calculations of chromatin deformation, distances between the kinetochore region and microneedle insertion site were directly measured to accurately depict chromatin stretch regardless of orientation to the spindle axis. Images of newt lung cells were transferred from the OMDR as TIFF files and image contrast and brightness corrected using Adobe Photoshop. Those images were transferred to Deneba Canvas as PICT files for labeling and reconstruction of small regions lost from the field of view. Final figures were printed using a Tectronix’ Phaser IISDX printer. Cell viability. Ensuring that cells remained viable throughout the micromanipulation procedure was of utmost concern. Ordinarily, cell death was easily detected during manipulation because the spindle and chromosomes were rapidly extruded from the ruptured cell. In subtler cases, cell death was apparent by an increase in the phase density of the chromosomes. Ultimately, we determined whether a cell survived manipulation by time-lapsing the video record of chromosome movement onto the OMDR. Only cells in which neighboring kinetochores actively oscillated during manipulation were included in this study.
RESULTS
Micromanipulation inside Vertebrate Mitotic Cells Manipulating mitotic newt lung chromosomes with microneedles without lysing the cell was much more difficult than expected based on Nicklas’ elegant studies using grasshopper spermatocytes [21, 22]. Microneedle tip movement was greatly restricted, presumably by the intermediate filament meshwork that forms a ‘‘cage’’ around the spindle [27], the rigidity of the cell cortex associated with the plasma membrane, or both. As a result, microneedles had to be extremely stiff to move through the cytoplasm and stretch the chromosome arms, yet be very sharp so that a single chromosome arm could be snagged. Typically, movement of the microneedle in the spindle region resulted in cell lysis. Consequently, manipulations were limited to chromosomes nearest the upper surface of the spindle so that the danger of puncturing the cell membrane with the microneedle was minimized. Of the 60/ mitotic newt lung cells in which chromosome manipulations were attempted, only 9 cells sur-
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vived the procedure of snagging a chromosome arm with the microneedle and, once snagged, stretching the arm. Cell death was usually observed by extrusion of the spindle from the lysed cell (see Material and Methods). Of these 9 surviving cells, 2 had manipulation experiments performed such that the chromosomes were pulled normal to the spindle axis. Although these centromeres responded to the added tension by moving in the direction of the stretched chromosome arm, the geometry of the pull relative to the spindle and resulting centromere movement made quantitative analysis difficult. Far cells were amenable to analysis such that both the centromere and pole remained visible by phase-contrast microscopy when a chromosome arm was stretched in a direction roughly parallel to the spindle axis. Experimentally Induced Tension Results in Kinetochore AP Movement for both Monoand Bioriented Chromosomes In the first example, one arm of a monooriented chromosome in a cell containing a monopolar spindle was stretched away from the pole (Fig. 1). Prior to manipulation (Fig. 1, 0 s), the target monooriented chromosome oscillated toward and away from the spindle pole, indicating that one kinetochore was tethered to the pole by kinetochore microtubules [2, 5]. A microneedle was lowered into the cell, deforming but not penetrating the cell membrane, and used to snag the chromosome between the sister chromatid arms (Fig. 1, 8 s). The microneedle was moved from a position 15.5 mm away from the spindle pole to 23 mm away from the spindle pole (Fig. 2 6–180 s), stretching the chromosome arm and exerting a firm and persistent tension at the kinetochore attachment site (Fig. 1, 94 s). The stretch of the arm (a 9.4-mm region of the arm between the kinetochore and microneedle was stretched 7.6 mm to approximately 17 mm) is clearly visible as a narrowing in the upper arm of the chromosome (Fig. 1, 94 s). While this tension was maintained, the centromere moved at 2.0 mm/min about 4 mm further away from the pole (Fig. 2, 6–140 s). Neighboring chromosomes continued to actively oscillate toward and away from the spindle pole and appeared completely unaffected by this manipulation procedure. Just prior to removal of the microneedle (180 s), the chromosome switched to P motility. After microneedle removal, the centromere continued to move at 1.9 mm/min (Fig. 2, 142–288 s) back toward the spindle pole to within 1 mm of its premanipulation position (Fig. 1, 288 s). In this experiment, the chromosome arm did not retract back to its premanipulation length after removal of the microneedle. In the second example, one arm of a bioriented chromosome near the metaphase plate of a bipolar spindle was stretched toward the spindle pole on the right side
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of the cell (Fig. 3). Prior to manipulation (Figs. 3 and 4), the centromere of the target bioriented chromosome actively oscillated several micrometers back and forth near the metaphase plate. A microneedle was lowered
FIG. 2. Distance-versus-time plot of centromere and microneedle movements shown in Fig. 1. The positions of the kinetochore (edge of the centromere that appeared to resist movement away from the pole (diamonds)), microneedle (asterisks), and hole in the chromosome arm (circles) produced by the microneedle were plotted relative to the spindle pole of the cell and superimposed onto one time axis. Vertical dashed lines indicate duration when tension (T) was exerted along the length of the chromosome arm.
FIG. 1. Phase-contrast micrograph sequence shows that a stretch of chromosome arm away from the spindle pole induces kinetochore AP motion in a monopolar spindle. See experimental details in text. Arrowhead, edge of the centromere (kinetochore) oriented toward the spindle pole; open arrow, monopolar spindle pole; curved arrow, microneedle. The hole between the sister chromatid arms left by the microneedle (removed at 180 s) is visible near the end of the upper arm. Bar, 10 mm.
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into the cell and used to snag one of the chromosome arms as their centromere was moving toward the left pole. The microneedle was briskly moved in an opposite direction roughly parallel to but slightly below the spindle axis (Fig. 4A), from a position 22.6 mm away from the left spindle pole to a position 30.6 mm away from the left spindle pole (Fig. 4B, 290–405 s) and away from the equator (Fig. 3, 290 s). The stretch of the arm (a 4.1-mm region of the arm between the kinetochore and the microneedle was stretched 8 mm to 12.4 mm) is clearly visible as a narrowing between the needle insertion point and the centromere (Fig. 3, 290 s) and placed the kinetochore and associated kinetochore fiber facing away from the microneedle under tension. The stretch of the chromosome arm also pulled the kinetochore nearest the microneedle toward the right pole and into its kinetochore fiber. After the chromosome arm was stretched, the centromere persisted in moving at nearly the same velocity as before the stretch toward the left pole for about 20 s before abruptly switching to AP movement in the direction of stretch. While the stretch of the chromosome arm was maintained, the centromere moved (one kinetochore moving AP, the sister moving P) at 1.95 mm/min about 4 mm (Fig. 4B, 290–405 s) further away from the left spindle pole and equator and in the direction of the stretched chromo-
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some arm, decreasing the stretch of the arm (Fig. 3, 450 s). Neighboring kinetochores actively oscillated and were unaffected by the manipulation procedure. Upon removal of the microneedle (405 s), the chromosome arm recoiled/refolded (124–205 s) and the centromere moved at approximately 1.97 mm/min (Fig. 4,
FIG. 4. Distance-versus-time plot of centromere and microneedle movements shown in Fig. 3. (A) Diagram shows the geometry of arm stretch, relative to the spindle axis, for the bipolar spindle shown in Fig. 3. Sister kinetochores (boxes labeled ‘‘1’’ and ‘‘2’’ denote leading and lagging edge of the centromere) are tethered to their respective poles (P1 and P2) by kinetochore microtubules (lines). The microneedle snagged a portion of the chromosome arm (circle) that was stretched by microneedle movement (arrow) in a direction away from the left spindle pole and equator, slightly below the spindle axis, placing the kinetochore facing away from the stretched arm under tension and the kinetochore facing toward the stretched arm under tension. (B) Distance-versus-time plot of the movements of kinetochore ‘‘1’’ (diamonds), the microneedle (asterisks), and the hole in the chromosome arm (circles) produced by the microneedle, relative to the left spindle pole of the cell. See text for details. Vertical dashed lines indicate the period when tension (T) was exerted along the length of the chromosome arm.
FIG. 3. Phase-contrast micrographs show centromere movement induced by stretching one arm of a bioriented chromosome toward one pole in a bipolar spindle. For clarity, only the left kinetochore or edge of centromere (arrowhead) that led or resisted centromere movement poleward or resisted chromosome stretch, the left spindle pole (open arrow), and the microneedle (curved arrow) are labeled. Time is given in s from the graph in Fig. 4B. Bar, 5 mm.
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450–600 s) back to the metaphase plate to within 0.5 mm of its premanipulation position (Fig. 3, 336 s). The third example also involves stretching one arm of a bioriented chromosome near the metaphase plate of a bipolar spindle (Fig. 5). Prior to manipulation (Fig. 5, 0 s), the target bioriented chromosome actively oscillated several micrometers back and forth at the metaphase plate at a position about 21.5 mm from the left spindle pole. As before, a microneedle was lowered into
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the cell and used to snag the chromosome between the sister chromatid arms. The microneedle was moved from a position 24.5 mm away from the left spindle pole to about 31.2 mm in a direction toward and below the spindle pole on the right side of the cell (Fig. 6A) and away from the left spindle pole and equator. The stretch of the chromosome arm (about 3.8 mm of chromatin between the kinetochore and needle site was stretched 6.2 mm to 10 mm) is evident (Fig. 5, 36 s) and placed the kinetochore facing away from the needle under tension. While stretch of the arm was maintained, the centromere moved (one kinetochore moving
FIG. 6. Distance-versus-time plot of centromere and microneedle movements for the cell shown in Fig. 5. (A) Diagram shows the geometry of microneedle stretching of the chromosome arm relative to the spindle axis. Symbols in schematic same are as in Fig. 4a. Although the kinetochore facing away from the stretched arm is under tension, it is unclear from the geometry of the arm stretch, relative to the spindle axis, to what extent the kinetochore nearest the microneedle was compressed. (B) Distance-versus-time plot of kinetochore (diamonds), microneedle (asterisks), and hole in the chromosome arm (circles) produced by microneedle movement, relative to the left spindle pole of cell shown in Fig. 5. Vertical dashed lines indicate the period when tension (T) was exerted along the length of the chromosome arm.
FIG. 5. Phase-contrast micrographs show that stretching the chromosome arm of a bioriented chromosome induced centromere movement. For clarity, only the left kinetochore (edge of the centromere that appeared to resist the stretch of the chromosome arm (arrowhead)), microneedle (curved arrow), and left spindle pole (open arrow) are labeled. Bar, 10 mm.
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AP, the sister moving P) at 0.9 mm/min (Fig. 6, 30–250 s) 3 mm further away from the equator and left spindle pole and moved toward the stretched chromosome arm (Fig. 5, 250 s). The chromosome arm recoiled after removal of the microneedle to approximately its original length (250 s) and the centromere moved at 1.0 mm/ min (Fig. 6, 252–416 s) back to the metaphase plate to within 0.5 mm of its premanipulation position (Fig. 5, 416 s). In a final example, a monooriented chromosome was detached from a monopolar spindle and the movement of the previously tethered kinetochore followed (Figs. 7 and 8) to contrast the movements of kinetochores still associated with microtubule plus ends (Figs. 1 –
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FIG. 8. Distance-versus-time plot shows centromere motility for the cell shown in Fig. 7. See text for details. Vertical dashed lines indicate periods when either detachment (D) procedures were performed or when tension (T) was exerted along the length of the chromosome arm.
6). A chromosome suitable for manipulation in a monopolar spindle was selected (Fig. 7). Prior to manipulation (Fig. 7, 0 s), this chromosome, like its neighboring chromosomes, actively oscillated near a position 19 mm from the spindle pole. A microneedle was lowered into the cell and used to snag the chromosome arm within 2 mm from the kinetochore. The microneedle was briskly moved 1– 3 mm toward the pole and then firmly moved about 8 mm further away from the pole (Fig. 7, 18 s). This procedure completely untethered the kinetochore from the spindle pole: the chromatin region between the kinetochore and microneedle insertion site was not stretched during chromosome movement away from the pole. In addition, the centromere followed the microneedle tip AP at 20 mm/min (Fig. 8, 2– 16 s) —a velocity about an order of magnitude faster than the normal rate of AP movement for kinetochores associated with microtubule plus ends [4 –7]. The microneedle was maintained in the cell. After a short lag period, the chromosome reattached to the spindle. Kinetochore P motion disentangled the chromosome from the microneedle and moved the chromosome at 3.5 mm/ min (Fig. 8, 28– 140 s) about 6 mm back toward the pole and cluster of monooriented chromosomes (Fig. 7, 122 s). The microneedle was removed at 66 s (first arrow in Fig. 8). This detachment procedure was repeated on the same chromosome. After a short poleward shove of the microneedle, the centromere was again rapidly (20 mm/ min) moved about 4 mm further away from the spindle pole (Figs. 7, 202 s and 8, 176–196 s). As in the first
FIG. 7. Phase-contrast micrographs show the manipulation procedure used to detach a monooriented chromosome from the spindle
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apparatus. See text for details. Note that the chromosome rotated after each detachment such that the centromere no longer appeared oriented toward the spindle pole. Bar, 5 mm.
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manipulation, the length of chromatin between the kinetochore and the microneedle insertion point did not change between the initial detachment and movement away from the spindle pole (i.e., the arm was not stretched). The microneedle was maintained in the cell for an extended period of time (Fig. 8, 144–414 s). After a brief lag period, the kinetochore reattached to the spindle and moved the chromosome at 3.3 mm/min (Fig. 8, 244–328 s) about 6.5 mm back toward the pole. In this experiment, however, the chromosome arm was anchored by the microneedle and became stretched by kinetochore P movement. While the chromosome arm was maximally stretched, the kinetochore stopped its net poleward migration and oscillated at a new position about 18–19 mm from the pole (330–412 s) until removal of the microneedle. These oscillations were difficult to track in our phase-contrast images and do not appear in Fig. 8. After removal of the microneedle, (414 s, second arrow in Fig. 8), the chromosome moved, on average, at 1 mm/min (Fig. 8, 414–560 s) about 3 mm back toward the pole (Fig. 7, 494 s). This chromosome arm was snagged a third time. During this experiment, the needle was not pushed toward the pole prior to stretching the chromosome arm but directly moved 5.4 mm further away from the pole. The kinetochore-to-pole linkage was not disrupted: movement of the microneedle away from the pole stretched the chromosome arm (3.6 mm of chromatin between the kinetochore and microneedle insertion point was stretched 5 mm to a length of 8.6 mm; Fig. 7, 676 s). While this stretch was maintained, the kinetochore moved at approximately 4.8 mm/min (Fig. 8, 598– 630 s) to about 5 mm further from the pole, at which point in time the cell membrane ruptured (not shown). DISCUSSION
Assessing our Manipulation Results How do we know that attached kinetochores of target chromosomes remained tethered during the manipulation-induced movements? First, the chromosome arms remained stretched during kinetochore AP movement. Second, the attached kinetochores moved AP at about 2 mm/min (Table 1), rates typical of kinetochores attached to microtubule plus ends in newt lung cells [4–8] and easily distinguishable from the rapid (20 mm/ min) manipulation-induced motion of kinetochores detached from the spindle (Table 1, Fig. 8, 2–16 and 176–196 s). In experiment 4, AP velocities of tethered kinetochores that were detached first and then allowed to reassociate with the spindle apparatus were slightly higher than typical kinetochore velocities. A similar result was previously reported by Nicklas [22] for chromosomes in grasshopper spermatocytes, and the reasons for this slightly higher velocity are unknown. In experiments 2 and 3, we conclude that both sister ki-
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TABLE 1 Velocities for Manipulated Chromosomes Velocity AP
Cell Cell Cell Cell
1 2 3 4I II
Cell 4 I II
P
Kinetochores attached to microtubules 2.0 1.9 1.95 2.0 0.9 1.0 4.8 3.5 N/A 3.3 Kinetochores detached from microtubules 20a N/A 20a N/A
Note. Summary of kinetochore velocities during manipulations in cells 1–4. Note that in cell 4, the kinetochore was first detached (two detachment events, I and II) from the pole and allowed to reattach and move to obtain AP and P velocities (see text for details). Velocity: average velocity for net kinetochore AP and P displacements, relative to the pole, during the course of the manipulation experiment. a Velocity of centromere AP movement by microneedle after detachment procedure (see Discussion).
netochores of the target bioriented chromosomes remained tethered during manipulations for two additional reasons. First, stretch-induced centromere movements off the metaphase plate were accompanied by oscillations in centromere stretch. These oscillations are clear when viewing the video records and are normally produced by the relative motion of attached sister kinetochores [4]. Second, upon release of the chromosome arm, the bioriented centromeres returned to their premanipulation position near the metaphase plate at rates typical of the congression of bioriented chromosomes. How do we know the kinetochore ‘‘response’’ to manipulation was real and not simply the coincidence of a spontaneous oscillation in the direction of stretch? For each of our experiments, increased tension (Table 1) resulted in centromere movement in the direction of the stretched chromosome arm (Figs. 1–8). When the microneedle was removed or the chromosome stretch decreased by AP movement, the centromere moved back to near its original average position near the metaphase plate. One way to unambiguously demonstrate that micromanipulation affected chromosome movement would be to induce extremely long movements to outside the spindle volume by repeatedly moving the microneedle away from the pole to maintain tension along the stretched arm as the kinetochore moved AP. Although prolonged chromosome movement has been successfully induced in meiotic invertebrate cells [21, 22], extensive microneedle tip displacements without cell lysis proved impossible for mitotic vertebrate cells. Nonetheless, our results show that the av-
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erage manipulation-induced net kinetochore displacement obtained in this study was 4 mm, a distance that is almost twice the average amplitude of normal kinetochore oscillations [4, 6, 7]. In Figs. 3 and 4, the kinetochore facing away from the direction of stretch did not immediately switch to AP movement but persisted for about 20 s at the same P velocity in spite of the higher tension produced by the stretch of the chromosome arm. This ‘‘lag’’ may be indicative of a ‘‘hysteresis’’ in the tension sensing mechanisms at the kinetochore which allows kinetochores to oscillate between P and AP movement over a similar tension range, switching from P to AP at higher tensions and from AP to P at lower tension [8]. Thus, our data show that elevated tension prolongs AP movement, but there is insufficient data to adequately test the role of tension in inducing kinetochore switching from P to AP movement. Finally, how do the normal mechanisms that govern kinetochore position from the pole relate to the tension generated by micromanipulation of the chromosome arms? The results of experiment 4 are particularly relevant to this question. The position where a kinetochore oscillated after it reconnected with polar spindle microtubules was changed simply by anchoring the chromosome arm with a microneedle before reattachment. After attachment, the kinetochore initially exhibited P movement, but began oscillating between phases of AP and P movement at a distance further from the pole than occurred without anchoring the arm. At this position, the tension generated by chromosome stretch was primarily supplied by the motion of the kinetochore and resisted mainly by the microneedle and not by the polar ejection forces on the arms. Removal of the microneedle eliminated this source of tension and allowed the kinetochore to pull the chromosome arms further into the spindle. We conclude from these results that vertebrate kinetochores oscillate at positions that can be defined by a force balance mechanism. Estimation of Forces Exerted during Manipulation We were unable to directly measure the force exerted by micromanipulation via microneedle tip deflection as was done by Nicklas [22, 28]. In mitotic newt lung cells, force was required to both stretch the chromosome arm and overcome a strong resistance presumably produced by the intermediate filament cage surrounding the spindle volume [27] and the stiff cell cortex associated with vertebrate cell plasma membranes. Furthermore, the chromosome region between the kinetochore and the microneedle insertion point (the stretched region) did not always rapidly recoil when the microneedle was removed, as reported by Nicklas for grasshopper spermatocytes. Often, the stretched chromatin region either appeared to slowly ‘‘refold’’ over 80/ s (Figs. 4B and 6B) or did not resume its premanipulation length
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during the course of the experiment (Fig. 2). This slow refolding may be similar to the reduction in chromosome length that occurs after the premetaphase stretch reported by Hughes-Schrader for mantids [29] and plasmids [30]. On the other hand, the stretched arm may have become entangled in the intermediate filament cage or retraction of the stretched arm may have been retarded by polar ejection forces. Optical traps with a strength of 3 mdyn (30 pN) have been used to measure the force produced by individual myosin [36, 37] and kinesin motors [42, 43]. Optical traps will move severed acentric chromosome fragments at velocities of 180 mm/min through the peripheral spindle region in newt lung cells [33]. However, an optical trap of similar force was insufficient to significantly stretch chromosome arms or affect centromere position within the metaphase spindle in newt lung cells (Skibbens, Salmon, and Sheetz, unpublished observations). The inability of 30 pN tension to induce prolonged kinetochore AP movement indicates that much higher tensions are required (see 31 for estimates between 90 and 1950 pN based on chromosome stretch). The Kinetochore-Tensiometer Model In this study, we provide direct evidence for vertebrate tissue cells that elevated tension (produced by stretched arms in our experiments) induces prolonged kinetochore AP movement while low tension (or compression) induces prolonged P movement at the constant velocities typical of kinetochore P and AP movement during mitosis. These findings are predicted by the kinetochore tensiometer model. They also provide direct evidence for another prediction of the model: that forces external from kinetochores capable of altering centromere stretch (as proposed for polar ejection forces that act on chromosome arms) [1–3, 5, 8, 23, 24] can bias chromosome movement toward and away from the spindle pole and determine chromosome position within the spindle. Our results in newt cells are consistent with those described by Nicklas [21, 22] for micromanipulation experiments in meiotic grasshopper cells: tension induced chromosomes to move at velocities typical of prometaphase chromosome movement toward the direction of stretch and off the spindle equator, independent of ‘‘load.’’ When the stretch was released by removing the microneedle, chromosomes recongressed at normal velocities toward the spindle equator. Since grasshopper meiotic kinetochores and newt mitotic kinetochores change their direction of movement in response to tension in a similar way, the kinetochore tensiometer model may be characteristic of eukaryotic kinetochores in general. The molecular mechanisms that kinetochores use to govern velocity, switch direction of movement, and sense tension remain important issues to be resolved.
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The P state of the kinetochore is a force-producing state which stretches the centromere and produces tension. The constant velocity of kinetochore P motility has been proposed to be limited, independent of changes in ‘‘load’’ that increase or decrease centromere stretch, by the slow speed of minus-directed microtubule motor proteins which couple the kinetochore to shortening kinetochore microtubule plus ends or by factors that limit the rate of kinetochore microtubule disassembly [8, 9, 17, 38]. The AP state of kinetochore movement rarely if ever generates pushing forces [4, 7, 12, 13]; once kinetochores attach to microtubule plus ends, centromeres usually remain stretched beyond their rest lengths as they oscillate between P and AP movements during mitosis [7]. Khodjakov and Rieder [12] concluded from studies of chromosome congression in Ptk1 cells that the AP state is not a force-generating state. Instead, they termed it an ‘‘N’’ state, for neutral or non-force-generating. However, our results reveal that kinetochore AP movement is tightly governed and load or tension independent. Previous studies have shown that the velocity of kinetochore movement is often constant as it reduces centromere stretch generated by the preceding kinetochore P movement [4, 5, 6]. Similarly, the rate of AP movement appears unaffected when kinetochore load is reduced by removing the bulk of the chromosome arms [6]. Our micromanipulation results show that AP velocity does not increase when a region of the chromosome arm is stretched to twice its rest length. We conclude that the kinetochore AP motility state governs the rate of movement and resists both polar ejection forces near the spindle pole and centromere tension. AP velocity may be rate limited by the proteins (perhaps microtubule motor proteins) that couple kinetochores to growing microtubule plus ends or other factors that limit the rate of kinetochore microtubule plus end assembly [6, 9, 17, 38]. The molecular mechanisms at the kinetochore that sense tension and control switching between P and AP states are poorly understood. In vitro studies on isolated CHO cell chromosomes show that increased phosphorylation results in plus-end-directed microtubule gliding across kinetochores while reduced phosphorylation results in gliding in the opposite direction [17]. However, these biochemical changes have not been successfully linked to changes in kinetochore motility in vivo although recent evidence indicates that kinetochore components are differentially phosphorylated in mitosis [38, 42–45]. So far, changes in the phosphorylation state of the kinetochore have been strongly correlated with the role of kinetochores in the spindle assembly cell cycle checkpoint for anaphase onset and not with the direction of kinetochore movement [17, 39–41 and reviewed by 46]. Nevertheless, our manipulation results show that tension regulates kinetochore directional instability and predict that tension alters the
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biochemical state of the kinetochore to control the direction of kinetochore motion. We thank Dr. Bruce Nicklas and Susan Ward, whose patient nurturing allowed for easy access into the world of micromanipulation and needle pulling. We also thank Steve Parsons, Phong Tran, Jen Waters, and Vicki Skeen for their support, enthusiasm, and editing during the course of this project. Special thanks go to Liz Blanton for helpful discussions of force analysis and figure preparation, to V.S. for filling in, and to Dr. Meg Kenna for editing early versions of the manuscript. This work was supported by a grant from the National Institutes of Health (GM24364).
REFERENCES 1. Bajer, A. S. (1982) J. Cell Biol. 93, 33–48. 2. Rieder, C. L., Davison, E. A., Jensen, L. C. W., Cassimeris, L., and Salmon, E. D. (1986) J. Cell Biol. 103, 581–591. 3. Ault, J. G., Demarco, A. J., Salmon, E. D., and Rieder, C. L. (1991) J. Cell Sci. 99, 701–710. 4. Skibbens, R. V., Skeen, V. P., and Salmon, E. D. (1993) J. Cell Biol. 122, 859–875. 5. Cassimeris, L., Rieder, C. L., and Salmon, E. D. (1994) J. Cell Sci. 107, 285–297. 6. Skibbens, R. V., Rieder, C. L., and Salmon, E. D. (1995) J. Cell Sci. 108, 2537–2548. 7. Waters, J. C., Skibbens, R. V., and Salmon, E. D. (1996) J. Cell Sci. 109, 2823–2831. 8. Rieder, C. L., and Salmon, E. D. (1994) J. Cell Biol. 124, 223– 233. 9. Inoue´, S., and Salmon, E. D. (1995) Mol. Biol. Cell 6, 1619– 1640. 10. Roos, U.-P. (1976) Chromosoma 54, 363–385. 11. Wise, D., Cassimeris, L., Rieder, C. L., Wadsworth, P., and Salmon, E. D. (1991) Cell Motil. Cytoskel. 18, 1–12. 12. Khodjakov, A., and Rieder, C. L. (1996) J. Cell Biol. 135, 1–13. 13. Shelby, R. D., Hahn, K. M., and Sullivan, K. F. (1996) J. Cell Biol. 135, 545–557. 14. Mitchison, T. J., and Salmon, E. D. (1992) J. Cell Biol. 119, 569–582. 15. Zhai, Y., and Borisy, G. G. (1995) J. Cell Biol. 131, 721–734. 16. Lombrillo, V. A., Nislow, C, Yen, T. J., Gelgand, V. I., and McIntosh, J. R. (1995) J. Cell Biol. 128, 107–115. 17. Hyman, A. A., and Mitchison, T. J. (1991) Nature 351, 206– 211. 18. Hyman, A. A., and Mitchison, T. J. (1991) Cold Spring Harbor Symp. Quant. Biol. 56, 745–750. 19. Hays, T., and Salmon, E. D. (1990) J. Cell Biol. 110, 391–404. 20. McNeill, P. A., and Berns, M. W. (1981) J. Cell Biol. 88, 543– 553. 21. Nicklas, R. B. (1977) in Mitosis Facts and Questions (Little, Paweletz, Petzelt, Ponstingl, Schroeter, and Zimmermann, Eds.), pp. 150–155, Springer–Verlag, New York. 22. Nicklas, R. B. (1988) Annu. Rev. Biophys. Chem. 17, 431–449. 23. Fuller, M. T. (1995) Cell 81, 5–8. 24. Vernos, I., and Karsenti, E. (1996) Curr. Opin. Cell Biol. 8, 4– 9. 25. Rieder, C. L., Ault, J. G., Eichenlaub-Ritter, U., and Sluder, G. (1993) in Chromosome Segregation and Aneuploidy (Vig, B. K., and Kappas, A., Eds.), Springer–Verlag, New York. 26. Rieder, C. L., and Hard, R. (1990) Int. Rev. Cytol. 122, 153– 220.
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27. Mandeville, E. C., and Rieder, C. L. (1990) Cell Motil. Cytoskel. 15, 111–120. 28. Nicklas, R. B. (1983) J. Cell Biol. 97, 542–548. 29. Hughes-Schrader, S. (1943) Biol. Bull. 85, 265–300. 30. Hughes-Schrader, S. (1946) Chromosoma 3, 1–21. 31. Skibbens, R. V. (1994) Ph.D. Dissertation at University of North Carolina, Chapel Hill. 32. Alexander, S. P., and Rieder, C. L. (1991) J. Cell Biol. 113, 805– 815. 33. Liang, H., Wright, W. H., Rieder, C. L., Salmon, E. D., Profeta, G., Andrews, J., Liu, Y., Sonek, G. J., and Berns, M. W. (1994) Exp. Cell Res. 213, 308–312. 34. Finer, J. T., Simmons, R. M., and Spudich, J. A. (1994) Nature 368, 113–119. 35. Vale, R. D. (1994) Cell 78, 733–737. 36. Svoboda, K., Schmidt, C. F., Schnapp, B. J., and Block, S. M. (1993) Nature 365, 721–727.
37. Svoboda, K., and Block, S. M. (1994) Cell 77, 773–784. 38. Desai, A., and Mitchison, T. J. (1995) J. Cell Biol. 128, 1–4. 39. Gorbsky, G. H., and Ricketts, W. A. (1993) J. Cell Biol. 122, 1311–1321. 40. Davis, A., Sage, C. R., Dougherty, C., and Farrell, K. W. (1994) Science 264, 839–842. 41. McIntosh, J. R. (1994) in Microtubules (Hyams, J., and Lloyd, C., Eds.), Wiley-Liss, New York. 42. Rattner, J. B., Lew, J., and Wang, J-L. (1990) Cell Motil. Cytoskel. 17, 277. 43. Shero, J. H., and Hieter, P. (1991) Genes Dev. 5, 549–560. 44. Nicklas, R. B., Krawitz, L. E., and Ward, S. C. (1993) J. Cell Sci. 104, 961–973. 45. Nicklas, R. B., Ward, S. C., and Gorbsky, G. J. (1995) J. Cell Biol. 130, 929–939. 46. Rutner, A. D., and Murray, A. W. (1996) Curr. Opin. Cell Biol. 8, 773–780.
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Micromanipulation of Chromosomes in Mitotic Vertebrate Tissue Cells: Tension Controls the State of Kinetochore Movement Robert V. Skibbens1 and E. D. Salmon2 Department of Biology, University of North Carolina, Chapel Hill, North Carolina 27599-3280
INTRODUCTION In mitotic vertebrate tissue cells, chromosome congression to the spindle equator in prometaphase and segregation to the poles in anaphase depend on the movements of kinetochores at their kinetochore microtubule attachment sites. To test if kinetochores sense tension to control their states of movement poleward (P) and away from the pole (AP), we applied an external force to the spindle in preanaphase newt epithelial cells by stretching chromosome arms with microneedles. For monooriented chromosomes (only one kinetochore fiber), an abrupt stretch of an arm away from the attached pole induced the single attached kinetochore to persist in AP movement at about 2 mm/ min velocity, resulting in chromosome movement away from the pole. When the stretch was reduced or the needle removed, the kinetochore switched to P movement at about 2 mm/min and pulled the chromosome back to near the premanipulation position within the spindle. For bioriented chromosomes (sister kinetochores attached to opposite poles) near the spindle equator, stretching one arm toward a pole placed the kinetochore facing away from the direction of stretch under tension and the sister facing toward the stretch under reduced tension or compression. Kinetochores under increased tension exhibited prolonged AP movement while kinetochores under reduced tension or compression exhibited prolonged P movement, moving the centromeres at about 2 mm/min velocities off the metaphase plate in the direction of stretch. Removing the needle resulted in centromere movement back to near the spindle equator at similar velocities. These results show that tension controls the direction of kinetochore movement and associated kinetochore microtubule assembly/disassembly to position centromeres within the spindle of vertebrate tissue cells. High tension induces persistent AP movement while low tension induces persistent P movement. The velocity of P and AP movement appears to be load independent and governed by the molecular mechanisms which attach kinetochores to the dynamic ends of kinetochore microtubules. q 1997 Academic Press
1 Current address: Johns Hopkins School of Medicine, Department of Molecular Biology and Genetics, Baltimore, MD 21205. 2 To whom correspondence and reprint requests should be addressed. Fax: (919) 962-1625. E-mail: [email protected].
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0014-4827/97 $25.00 Copyright q 1997 by Academic Press All rights of reproduction in any form reserved.
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During mitosis, kinetochores attach chromosomes to spindle poles by capturing and stabilizing microtubule plus ends. In vertebrate tissue cells studied to date [1– 13 and references therein], kinetochores exhibit ‘‘directional instability,’’ abruptly switching between constant velocity (about 2 mm/min) states of movement poleward (P) or away from the pole (AP). In newt and PtK1 cells, fluorescent marks on the lattice of kinetochore microtubules move slowly poleward at about 0.5 mm/min or less [14, 15]. Thus, during P movement, kinetochore microtubule shortening occurs predominantly at the kinetochore and, during AP movement, kinetochore microtubule growth occurs exclusively at the kinetochore. Kinetochore P movement stretches its centromere chromatin poleward until the kinetochore abruptly switches to AP movement. AP movement reduces centromere stretch until the kinetochore switches back to P movement [4, 5]. The molecular mechanisms that attach kinetochores to the dynamic plus ends of kinetochore microtubules and produce the constant velocity states of kinetochore P and AP movement are not understood. P movement produces pulling forces [reviewed in 8, 9], but AP movement occurs mainly under tension and rarely pushes [4, 7, 12]. There is evidence that both plus- and minus-end-directed microtubule motor proteins bound to kinetochores as well as the shortening and growth phases of plus end microtubule dynamic instability are involved [9, 16, 17]. A key issue in mitosis is what attached kinetochores sense to control the durations of their P and AP states of movement. This question is important because the congression of sister chromatid pairs to the spindle equator in prometaphase and their segregation to opposite poles in anaphase depends on the regulation of the relative durations of P and AP kinetochore movement [4]. Chromosome congression to the equator in prometaphase occurs only when the sister kinetochore facing the equator persists in P movement toward the equator and the sister facing away from the equator persists in AP movement toward the equator; net movement of the centromere does not occur when both attached sisters
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move P or AP at the same time [4]. In anaphase, the segregation of separated sisters to the poles occurs only when the kinetochores persist in P movement, pulling their chromosome arms poleward [4, 6]. There is evidence that kinetochore switching between P and AP states depends on kinetochore phosphorylation [17], the number of kinetochore microtubules [18], or factors which affect microtubule assembly [9]. In addition, several reports indicate that kinetochores primarly sense tension to control the durations of their P and AP states [4–9, 19–22]. For example, on average, kinetochores on bioriented chromosomes at the metaphase plate are under tension because their centromere chromatin is stretched beyond their unattached rest length [7, 13, 19]. Reducing kinetochore tension on a bioriented chromosome by either ablating the sister [2, 19, 20] or ablating the centromereic chromatin between sister kinetochores results in prolonged P movement of the undamaged kinetochore at velocities similar to kinetochore poleward motion in anaphase [6, 12]. A kinetochore tensiometer model has been proposed for vertebrate tissue cells where the direction, but not the velocity, of kinetochore movement is sensitive to tension: higher tension increases the probability of the kinetochore switching from P and persisting in AP movement while low tension or compression increases the probability of switching from AP and persisting in P movement [4–8]. During mitosis, changes in kinetochore tension are produced by the relative motions of sister kinetochores and their chromosome arms. Poleward movement of chromosome arms is resisted by spindle polar microtubule arrays which push the arms away from the pole [1–8, 12]. These ‘‘polar ejection forces’’ are thought to increase in strength as microtubule density increases closer to the poles. Ejection forces may be produced by chromosome-associated microtubule motor proteins (chromokinesins) or the growth dynamics of the polar microtubule arrays and contribute significantly to the tension which biases a kinetochore into AP movement near the pole [2, 3, 12, 23, 24]. In this paper, we tested the kinetochore tensiometer model in mitotic newt lung cells by examining how kinetochores respond—in terms of direction, duration, and velocity of movement—to the application of a force external from the spindle produced by stretching a chromosome arm with a microneedle. Previously, such studies were performed by Nicklas [21, 22], but they were performed only in grasshopper spermatocytes and have not been tested in vertebrate mitotic cell systems. Several lines of evidence indicate that kinetochores in invertebrate meiotic cells behave differently than kinetochores in vertebrate mitotic cells [25] (i.e., the kinetochores in grasshopper spermatocytes move more slowly (0.5 mm/min) and without the amplitude of oscillation found in vertebrate tissue cells [1–12, 19, 21]). Furthermore, sister kinetochores are paired and act in
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concert in meiosis I cells while sister kinetochores are separate and act independently in mitotic cells [4, 6]. Therefore, it was important to test the kinetochore tensiometer model in a vertebrate cell system. Our micromanipulation studies on newt cells show that the direction of kinetochore movement is sensitive to tension as predicted by the kinetochore tensiometer model [6] and Nicklas’ [21, 22] micromanipulation in meiotic grasshopper spermatocytes. The constant velocities of kinetochore AP movement during mitosis indicate that kinetochores normally govern their AP velocity [4, 5, 12]. Our micromanipulation results show that the kinetochore is able to govern the rate at which it moves AP in response to a substantial pulling force. MATERIAL AND METHODS Tissue culture. Newt lung cultures were prepared as described [2, 26]. Oregon newts (Taricha granulosa) were obtained from Charles Sultan (Nashville, TN), the lungs dissected from the animal, minced, washed three times in calcium-free/magnesium-free Hanks Ringer solution for amphibians, trypsinized for 5 minutes, and placed in L15 media supplemented with fetal bovine serum and antibiotics. Lung tissue explants were cultured overnight, placed in Rose chambers, and incubated 10 to 15 days at room temperature (22–267C). For experimentation, Rose chambers were disassembled and the coverslips containing monolayers were treated as described below. Micromanipulation chambers. Prior to observation, a Rose chamber was disassembled and the coverslip inverted onto two metal plates (1.5 1 3 1 0.0615 in with a 20-mm-diameter central hole) previously sealed together with vacuum grease. The coverslip was affixed to one face of the metal plate with a thin layer of grease, sealed with VALAP (1:1:1 of Vaseline, lanolin, and paraffin), and the whole structure inverted so that the coverslip formed the floor of the micromanipulation chamber. The double thickness of metal allowed for a maximum volume of fresh medium (about 600 ml) to be placed over the explant and still allowed a thin layer of mineral oil to be layered over the medium. Mineral oil proved nontoxic to cells and prolonged cell viability by delaying changes in osmolarity caused by media evaporation. The mineral oil overlay also provided a flat surface so image quality was maintained. Micromanipulation needles. Needles appropriate for micromanipulation were generated in a three-step process using custom glass tubing (Pyrex: o.d. 0.8 mm, i.d. 0.6 mm, length 100 mm—no internal filament) obtained from Drummund Scientific Co. (Broomall, PA). First, a 457 angle in the filament was placed near one end of the pipette using a fine butane flame and forceps. The region of the pipette just beyond the bend was pulled over the flame with forceps to form a fine tip. This tip was clipped to an appropriate length (tip distance over 4–5 mm normal from the cylinder axis interfered with proper positioning of the microscope condenser). In the second step, the tip of the needle was sealed and finely tapered using a Microforge De Fonbrune, Series A, No. 101 (equipped with 151 oculars) at a heat setting of about 6–8. This filament was too long and flexible to encourage chromosome movement through the stiff cell cortex or intermediate filament cage (see Results). The flexible portion of the needle tip was broken off using the cold Microforge heating element. In the third and final step, the needle was pulled out to a very fine but inflexible short tip using the Microforge at a low heat setting (0.0–0.5). Final needle selection was performed using a Zeiss Neofluar 25X 0.6NA Phase 2 objective on the IM stand. Micromanipulation. Needle control was achieved using a Narishige pneumatic micromanipulation device assembled onto a crossbar affixed to the condenser/illumination tower of the Zeiss IM microscope stand. Direct coupling of the micromanipulation device to the
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illumination tower was critical for dampening needle vibrations. Fine control was achieved with the Narishige hanging micromanipulation joystick placed off the vibration isolation platform. Phase microscopy and video. Images of newt lung cells were obtained using a Zeiss IM stand assembled on top of a pneumatic vibration isolation platform and equipped with a 60W Tungsten bulb, high NA Phase condenser complete with a Phase 3 annulus, Nikon 100X (NA 1.4) Phase 4 objective, and a 41 projection lens. Images were collected using a DAGE-MTI NC-67MDX Newvicon camera and recorded in real time onto 3/4 inch format Umatic video tape using a Sony VO-5800H video cassette recorder. Selected sequences were time lapsed (1 frame every 2 s) into a Panasonic TQ2028F Optical Memory Disc Recorder (OMDR) for analysis. Tracking and analyses. Tracking and analyses were performed as previously described [4, 6]. Computer-generated cursors were superimposed on images containing the objects to be tracked. Gray values for each pixel in an 8X8 array for each cursor were used as a template to search consecutive images on the OMDR. The computer recorded frame number, X and Y coordinates for each object, and time, which were later converted to distance-versus-time plots and analyzed using the in-house program, Single Frame Movement (SFM). For calculations of chromatin deformation, distances between the kinetochore region and microneedle insertion site were directly measured to accurately depict chromatin stretch regardless of orientation to the spindle axis. Images of newt lung cells were transferred from the OMDR as TIFF files and image contrast and brightness corrected using Adobe Photoshop. Those images were transferred to Deneba Canvas as PICT files for labeling and reconstruction of small regions lost from the field of view. Final figures were printed using a Tectronix’ Phaser IISDX printer. Cell viability. Ensuring that cells remained viable throughout the micromanipulation procedure was of utmost concern. Ordinarily, cell death was easily detected during manipulation because the spindle and chromosomes were rapidly extruded from the ruptured cell. In subtler cases, cell death was apparent by an increase in the phase density of the chromosomes. Ultimately, we determined whether a cell survived manipulation by time-lapsing the video record of chromosome movement onto the OMDR. Only cells in which neighboring kinetochores actively oscillated during manipulation were included in this study.
RESULTS
Micromanipulation inside Vertebrate Mitotic Cells Manipulating mitotic newt lung chromosomes with microneedles without lysing the cell was much more difficult than expected based on Nicklas’ elegant studies using grasshopper spermatocytes [21, 22]. Microneedle tip movement was greatly restricted, presumably by the intermediate filament meshwork that forms a ‘‘cage’’ around the spindle [27], the rigidity of the cell cortex associated with the plasma membrane, or both. As a result, microneedles had to be extremely stiff to move through the cytoplasm and stretch the chromosome arms, yet be very sharp so that a single chromosome arm could be snagged. Typically, movement of the microneedle in the spindle region resulted in cell lysis. Consequently, manipulations were limited to chromosomes nearest the upper surface of the spindle so that the danger of puncturing the cell membrane with the microneedle was minimized. Of the 60/ mitotic newt lung cells in which chromosome manipulations were attempted, only 9 cells sur-
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vived the procedure of snagging a chromosome arm with the microneedle and, once snagged, stretching the arm. Cell death was usually observed by extrusion of the spindle from the lysed cell (see Material and Methods). Of these 9 surviving cells, 2 had manipulation experiments performed such that the chromosomes were pulled normal to the spindle axis. Although these centromeres responded to the added tension by moving in the direction of the stretched chromosome arm, the geometry of the pull relative to the spindle and resulting centromere movement made quantitative analysis difficult. Far cells were amenable to analysis such that both the centromere and pole remained visible by phase-contrast microscopy when a chromosome arm was stretched in a direction roughly parallel to the spindle axis. Experimentally Induced Tension Results in Kinetochore AP Movement for both Monoand Bioriented Chromosomes In the first example, one arm of a monooriented chromosome in a cell containing a monopolar spindle was stretched away from the pole (Fig. 1). Prior to manipulation (Fig. 1, 0 s), the target monooriented chromosome oscillated toward and away from the spindle pole, indicating that one kinetochore was tethered to the pole by kinetochore microtubules [2, 5]. A microneedle was lowered into the cell, deforming but not penetrating the cell membrane, and used to snag the chromosome between the sister chromatid arms (Fig. 1, 8 s). The microneedle was moved from a position 15.5 mm away from the spindle pole to 23 mm away from the spindle pole (Fig. 2 6–180 s), stretching the chromosome arm and exerting a firm and persistent tension at the kinetochore attachment site (Fig. 1, 94 s). The stretch of the arm (a 9.4-mm region of the arm between the kinetochore and microneedle was stretched 7.6 mm to approximately 17 mm) is clearly visible as a narrowing in the upper arm of the chromosome (Fig. 1, 94 s). While this tension was maintained, the centromere moved at 2.0 mm/min about 4 mm further away from the pole (Fig. 2, 6–140 s). Neighboring chromosomes continued to actively oscillate toward and away from the spindle pole and appeared completely unaffected by this manipulation procedure. Just prior to removal of the microneedle (180 s), the chromosome switched to P motility. After microneedle removal, the centromere continued to move at 1.9 mm/min (Fig. 2, 142–288 s) back toward the spindle pole to within 1 mm of its premanipulation position (Fig. 1, 288 s). In this experiment, the chromosome arm did not retract back to its premanipulation length after removal of the microneedle. In the second example, one arm of a bioriented chromosome near the metaphase plate of a bipolar spindle was stretched toward the spindle pole on the right side
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of the cell (Fig. 3). Prior to manipulation (Figs. 3 and 4), the centromere of the target bioriented chromosome actively oscillated several micrometers back and forth near the metaphase plate. A microneedle was lowered
FIG. 2. Distance-versus-time plot of centromere and microneedle movements shown in Fig. 1. The positions of the kinetochore (edge of the centromere that appeared to resist movement away from the pole (diamonds)), microneedle (asterisks), and hole in the chromosome arm (circles) produced by the microneedle were plotted relative to the spindle pole of the cell and superimposed onto one time axis. Vertical dashed lines indicate duration when tension (T) was exerted along the length of the chromosome arm.
FIG. 1. Phase-contrast micrograph sequence shows that a stretch of chromosome arm away from the spindle pole induces kinetochore AP motion in a monopolar spindle. See experimental details in text. Arrowhead, edge of the centromere (kinetochore) oriented toward the spindle pole; open arrow, monopolar spindle pole; curved arrow, microneedle. The hole between the sister chromatid arms left by the microneedle (removed at 180 s) is visible near the end of the upper arm. Bar, 10 mm.
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into the cell and used to snag one of the chromosome arms as their centromere was moving toward the left pole. The microneedle was briskly moved in an opposite direction roughly parallel to but slightly below the spindle axis (Fig. 4A), from a position 22.6 mm away from the left spindle pole to a position 30.6 mm away from the left spindle pole (Fig. 4B, 290–405 s) and away from the equator (Fig. 3, 290 s). The stretch of the arm (a 4.1-mm region of the arm between the kinetochore and the microneedle was stretched 8 mm to 12.4 mm) is clearly visible as a narrowing between the needle insertion point and the centromere (Fig. 3, 290 s) and placed the kinetochore and associated kinetochore fiber facing away from the microneedle under tension. The stretch of the chromosome arm also pulled the kinetochore nearest the microneedle toward the right pole and into its kinetochore fiber. After the chromosome arm was stretched, the centromere persisted in moving at nearly the same velocity as before the stretch toward the left pole for about 20 s before abruptly switching to AP movement in the direction of stretch. While the stretch of the chromosome arm was maintained, the centromere moved (one kinetochore moving AP, the sister moving P) at 1.95 mm/min about 4 mm (Fig. 4B, 290–405 s) further away from the left spindle pole and equator and in the direction of the stretched chromo-
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some arm, decreasing the stretch of the arm (Fig. 3, 450 s). Neighboring kinetochores actively oscillated and were unaffected by the manipulation procedure. Upon removal of the microneedle (405 s), the chromosome arm recoiled/refolded (124–205 s) and the centromere moved at approximately 1.97 mm/min (Fig. 4,
FIG. 4. Distance-versus-time plot of centromere and microneedle movements shown in Fig. 3. (A) Diagram shows the geometry of arm stretch, relative to the spindle axis, for the bipolar spindle shown in Fig. 3. Sister kinetochores (boxes labeled ‘‘1’’ and ‘‘2’’ denote leading and lagging edge of the centromere) are tethered to their respective poles (P1 and P2) by kinetochore microtubules (lines). The microneedle snagged a portion of the chromosome arm (circle) that was stretched by microneedle movement (arrow) in a direction away from the left spindle pole and equator, slightly below the spindle axis, placing the kinetochore facing away from the stretched arm under tension and the kinetochore facing toward the stretched arm under tension. (B) Distance-versus-time plot of the movements of kinetochore ‘‘1’’ (diamonds), the microneedle (asterisks), and the hole in the chromosome arm (circles) produced by the microneedle, relative to the left spindle pole of the cell. See text for details. Vertical dashed lines indicate the period when tension (T) was exerted along the length of the chromosome arm.
FIG. 3. Phase-contrast micrographs show centromere movement induced by stretching one arm of a bioriented chromosome toward one pole in a bipolar spindle. For clarity, only the left kinetochore or edge of centromere (arrowhead) that led or resisted centromere movement poleward or resisted chromosome stretch, the left spindle pole (open arrow), and the microneedle (curved arrow) are labeled. Time is given in s from the graph in Fig. 4B. Bar, 5 mm.
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450–600 s) back to the metaphase plate to within 0.5 mm of its premanipulation position (Fig. 3, 336 s). The third example also involves stretching one arm of a bioriented chromosome near the metaphase plate of a bipolar spindle (Fig. 5). Prior to manipulation (Fig. 5, 0 s), the target bioriented chromosome actively oscillated several micrometers back and forth at the metaphase plate at a position about 21.5 mm from the left spindle pole. As before, a microneedle was lowered into
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the cell and used to snag the chromosome between the sister chromatid arms. The microneedle was moved from a position 24.5 mm away from the left spindle pole to about 31.2 mm in a direction toward and below the spindle pole on the right side of the cell (Fig. 6A) and away from the left spindle pole and equator. The stretch of the chromosome arm (about 3.8 mm of chromatin between the kinetochore and needle site was stretched 6.2 mm to 10 mm) is evident (Fig. 5, 36 s) and placed the kinetochore facing away from the needle under tension. While stretch of the arm was maintained, the centromere moved (one kinetochore moving
FIG. 6. Distance-versus-time plot of centromere and microneedle movements for the cell shown in Fig. 5. (A) Diagram shows the geometry of microneedle stretching of the chromosome arm relative to the spindle axis. Symbols in schematic same are as in Fig. 4a. Although the kinetochore facing away from the stretched arm is under tension, it is unclear from the geometry of the arm stretch, relative to the spindle axis, to what extent the kinetochore nearest the microneedle was compressed. (B) Distance-versus-time plot of kinetochore (diamonds), microneedle (asterisks), and hole in the chromosome arm (circles) produced by microneedle movement, relative to the left spindle pole of cell shown in Fig. 5. Vertical dashed lines indicate the period when tension (T) was exerted along the length of the chromosome arm.
FIG. 5. Phase-contrast micrographs show that stretching the chromosome arm of a bioriented chromosome induced centromere movement. For clarity, only the left kinetochore (edge of the centromere that appeared to resist the stretch of the chromosome arm (arrowhead)), microneedle (curved arrow), and left spindle pole (open arrow) are labeled. Bar, 10 mm.
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AP, the sister moving P) at 0.9 mm/min (Fig. 6, 30–250 s) 3 mm further away from the equator and left spindle pole and moved toward the stretched chromosome arm (Fig. 5, 250 s). The chromosome arm recoiled after removal of the microneedle to approximately its original length (250 s) and the centromere moved at 1.0 mm/ min (Fig. 6, 252–416 s) back to the metaphase plate to within 0.5 mm of its premanipulation position (Fig. 5, 416 s). In a final example, a monooriented chromosome was detached from a monopolar spindle and the movement of the previously tethered kinetochore followed (Figs. 7 and 8) to contrast the movements of kinetochores still associated with microtubule plus ends (Figs. 1 –
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FIG. 8. Distance-versus-time plot shows centromere motility for the cell shown in Fig. 7. See text for details. Vertical dashed lines indicate periods when either detachment (D) procedures were performed or when tension (T) was exerted along the length of the chromosome arm.
6). A chromosome suitable for manipulation in a monopolar spindle was selected (Fig. 7). Prior to manipulation (Fig. 7, 0 s), this chromosome, like its neighboring chromosomes, actively oscillated near a position 19 mm from the spindle pole. A microneedle was lowered into the cell and used to snag the chromosome arm within 2 mm from the kinetochore. The microneedle was briskly moved 1– 3 mm toward the pole and then firmly moved about 8 mm further away from the pole (Fig. 7, 18 s). This procedure completely untethered the kinetochore from the spindle pole: the chromatin region between the kinetochore and microneedle insertion site was not stretched during chromosome movement away from the pole. In addition, the centromere followed the microneedle tip AP at 20 mm/min (Fig. 8, 2– 16 s) —a velocity about an order of magnitude faster than the normal rate of AP movement for kinetochores associated with microtubule plus ends [4 –7]. The microneedle was maintained in the cell. After a short lag period, the chromosome reattached to the spindle. Kinetochore P motion disentangled the chromosome from the microneedle and moved the chromosome at 3.5 mm/ min (Fig. 8, 28– 140 s) about 6 mm back toward the pole and cluster of monooriented chromosomes (Fig. 7, 122 s). The microneedle was removed at 66 s (first arrow in Fig. 8). This detachment procedure was repeated on the same chromosome. After a short poleward shove of the microneedle, the centromere was again rapidly (20 mm/ min) moved about 4 mm further away from the spindle pole (Figs. 7, 202 s and 8, 176–196 s). As in the first
FIG. 7. Phase-contrast micrographs show the manipulation procedure used to detach a monooriented chromosome from the spindle
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apparatus. See text for details. Note that the chromosome rotated after each detachment such that the centromere no longer appeared oriented toward the spindle pole. Bar, 5 mm.
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manipulation, the length of chromatin between the kinetochore and the microneedle insertion point did not change between the initial detachment and movement away from the spindle pole (i.e., the arm was not stretched). The microneedle was maintained in the cell for an extended period of time (Fig. 8, 144–414 s). After a brief lag period, the kinetochore reattached to the spindle and moved the chromosome at 3.3 mm/min (Fig. 8, 244–328 s) about 6.5 mm back toward the pole. In this experiment, however, the chromosome arm was anchored by the microneedle and became stretched by kinetochore P movement. While the chromosome arm was maximally stretched, the kinetochore stopped its net poleward migration and oscillated at a new position about 18–19 mm from the pole (330–412 s) until removal of the microneedle. These oscillations were difficult to track in our phase-contrast images and do not appear in Fig. 8. After removal of the microneedle, (414 s, second arrow in Fig. 8), the chromosome moved, on average, at 1 mm/min (Fig. 8, 414–560 s) about 3 mm back toward the pole (Fig. 7, 494 s). This chromosome arm was snagged a third time. During this experiment, the needle was not pushed toward the pole prior to stretching the chromosome arm but directly moved 5.4 mm further away from the pole. The kinetochore-to-pole linkage was not disrupted: movement of the microneedle away from the pole stretched the chromosome arm (3.6 mm of chromatin between the kinetochore and microneedle insertion point was stretched 5 mm to a length of 8.6 mm; Fig. 7, 676 s). While this stretch was maintained, the kinetochore moved at approximately 4.8 mm/min (Fig. 8, 598– 630 s) to about 5 mm further from the pole, at which point in time the cell membrane ruptured (not shown). DISCUSSION
Assessing our Manipulation Results How do we know that attached kinetochores of target chromosomes remained tethered during the manipulation-induced movements? First, the chromosome arms remained stretched during kinetochore AP movement. Second, the attached kinetochores moved AP at about 2 mm/min (Table 1), rates typical of kinetochores attached to microtubule plus ends in newt lung cells [4–8] and easily distinguishable from the rapid (20 mm/ min) manipulation-induced motion of kinetochores detached from the spindle (Table 1, Fig. 8, 2–16 and 176–196 s). In experiment 4, AP velocities of tethered kinetochores that were detached first and then allowed to reassociate with the spindle apparatus were slightly higher than typical kinetochore velocities. A similar result was previously reported by Nicklas [22] for chromosomes in grasshopper spermatocytes, and the reasons for this slightly higher velocity are unknown. In experiments 2 and 3, we conclude that both sister ki-
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TABLE 1 Velocities for Manipulated Chromosomes Velocity AP
Cell Cell Cell Cell
1 2 3 4I II
Cell 4 I II
P
Kinetochores attached to microtubules 2.0 1.9 1.95 2.0 0.9 1.0 4.8 3.5 N/A 3.3 Kinetochores detached from microtubules 20a N/A 20a N/A
Note. Summary of kinetochore velocities during manipulations in cells 1–4. Note that in cell 4, the kinetochore was first detached (two detachment events, I and II) from the pole and allowed to reattach and move to obtain AP and P velocities (see text for details). Velocity: average velocity for net kinetochore AP and P displacements, relative to the pole, during the course of the manipulation experiment. a Velocity of centromere AP movement by microneedle after detachment procedure (see Discussion).
netochores of the target bioriented chromosomes remained tethered during manipulations for two additional reasons. First, stretch-induced centromere movements off the metaphase plate were accompanied by oscillations in centromere stretch. These oscillations are clear when viewing the video records and are normally produced by the relative motion of attached sister kinetochores [4]. Second, upon release of the chromosome arm, the bioriented centromeres returned to their premanipulation position near the metaphase plate at rates typical of the congression of bioriented chromosomes. How do we know the kinetochore ‘‘response’’ to manipulation was real and not simply the coincidence of a spontaneous oscillation in the direction of stretch? For each of our experiments, increased tension (Table 1) resulted in centromere movement in the direction of the stretched chromosome arm (Figs. 1–8). When the microneedle was removed or the chromosome stretch decreased by AP movement, the centromere moved back to near its original average position near the metaphase plate. One way to unambiguously demonstrate that micromanipulation affected chromosome movement would be to induce extremely long movements to outside the spindle volume by repeatedly moving the microneedle away from the pole to maintain tension along the stretched arm as the kinetochore moved AP. Although prolonged chromosome movement has been successfully induced in meiotic invertebrate cells [21, 22], extensive microneedle tip displacements without cell lysis proved impossible for mitotic vertebrate cells. Nonetheless, our results show that the av-
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erage manipulation-induced net kinetochore displacement obtained in this study was 4 mm, a distance that is almost twice the average amplitude of normal kinetochore oscillations [4, 6, 7]. In Figs. 3 and 4, the kinetochore facing away from the direction of stretch did not immediately switch to AP movement but persisted for about 20 s at the same P velocity in spite of the higher tension produced by the stretch of the chromosome arm. This ‘‘lag’’ may be indicative of a ‘‘hysteresis’’ in the tension sensing mechanisms at the kinetochore which allows kinetochores to oscillate between P and AP movement over a similar tension range, switching from P to AP at higher tensions and from AP to P at lower tension [8]. Thus, our data show that elevated tension prolongs AP movement, but there is insufficient data to adequately test the role of tension in inducing kinetochore switching from P to AP movement. Finally, how do the normal mechanisms that govern kinetochore position from the pole relate to the tension generated by micromanipulation of the chromosome arms? The results of experiment 4 are particularly relevant to this question. The position where a kinetochore oscillated after it reconnected with polar spindle microtubules was changed simply by anchoring the chromosome arm with a microneedle before reattachment. After attachment, the kinetochore initially exhibited P movement, but began oscillating between phases of AP and P movement at a distance further from the pole than occurred without anchoring the arm. At this position, the tension generated by chromosome stretch was primarily supplied by the motion of the kinetochore and resisted mainly by the microneedle and not by the polar ejection forces on the arms. Removal of the microneedle eliminated this source of tension and allowed the kinetochore to pull the chromosome arms further into the spindle. We conclude from these results that vertebrate kinetochores oscillate at positions that can be defined by a force balance mechanism. Estimation of Forces Exerted during Manipulation We were unable to directly measure the force exerted by micromanipulation via microneedle tip deflection as was done by Nicklas [22, 28]. In mitotic newt lung cells, force was required to both stretch the chromosome arm and overcome a strong resistance presumably produced by the intermediate filament cage surrounding the spindle volume [27] and the stiff cell cortex associated with vertebrate cell plasma membranes. Furthermore, the chromosome region between the kinetochore and the microneedle insertion point (the stretched region) did not always rapidly recoil when the microneedle was removed, as reported by Nicklas for grasshopper spermatocytes. Often, the stretched chromatin region either appeared to slowly ‘‘refold’’ over 80/ s (Figs. 4B and 6B) or did not resume its premanipulation length
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during the course of the experiment (Fig. 2). This slow refolding may be similar to the reduction in chromosome length that occurs after the premetaphase stretch reported by Hughes-Schrader for mantids [29] and plasmids [30]. On the other hand, the stretched arm may have become entangled in the intermediate filament cage or retraction of the stretched arm may have been retarded by polar ejection forces. Optical traps with a strength of 3 mdyn (30 pN) have been used to measure the force produced by individual myosin [36, 37] and kinesin motors [42, 43]. Optical traps will move severed acentric chromosome fragments at velocities of 180 mm/min through the peripheral spindle region in newt lung cells [33]. However, an optical trap of similar force was insufficient to significantly stretch chromosome arms or affect centromere position within the metaphase spindle in newt lung cells (Skibbens, Salmon, and Sheetz, unpublished observations). The inability of 30 pN tension to induce prolonged kinetochore AP movement indicates that much higher tensions are required (see 31 for estimates between 90 and 1950 pN based on chromosome stretch). The Kinetochore-Tensiometer Model In this study, we provide direct evidence for vertebrate tissue cells that elevated tension (produced by stretched arms in our experiments) induces prolonged kinetochore AP movement while low tension (or compression) induces prolonged P movement at the constant velocities typical of kinetochore P and AP movement during mitosis. These findings are predicted by the kinetochore tensiometer model. They also provide direct evidence for another prediction of the model: that forces external from kinetochores capable of altering centromere stretch (as proposed for polar ejection forces that act on chromosome arms) [1–3, 5, 8, 23, 24] can bias chromosome movement toward and away from the spindle pole and determine chromosome position within the spindle. Our results in newt cells are consistent with those described by Nicklas [21, 22] for micromanipulation experiments in meiotic grasshopper cells: tension induced chromosomes to move at velocities typical of prometaphase chromosome movement toward the direction of stretch and off the spindle equator, independent of ‘‘load.’’ When the stretch was released by removing the microneedle, chromosomes recongressed at normal velocities toward the spindle equator. Since grasshopper meiotic kinetochores and newt mitotic kinetochores change their direction of movement in response to tension in a similar way, the kinetochore tensiometer model may be characteristic of eukaryotic kinetochores in general. The molecular mechanisms that kinetochores use to govern velocity, switch direction of movement, and sense tension remain important issues to be resolved.
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The P state of the kinetochore is a force-producing state which stretches the centromere and produces tension. The constant velocity of kinetochore P motility has been proposed to be limited, independent of changes in ‘‘load’’ that increase or decrease centromere stretch, by the slow speed of minus-directed microtubule motor proteins which couple the kinetochore to shortening kinetochore microtubule plus ends or by factors that limit the rate of kinetochore microtubule disassembly [8, 9, 17, 38]. The AP state of kinetochore movement rarely if ever generates pushing forces [4, 7, 12, 13]; once kinetochores attach to microtubule plus ends, centromeres usually remain stretched beyond their rest lengths as they oscillate between P and AP movements during mitosis [7]. Khodjakov and Rieder [12] concluded from studies of chromosome congression in Ptk1 cells that the AP state is not a force-generating state. Instead, they termed it an ‘‘N’’ state, for neutral or non-force-generating. However, our results reveal that kinetochore AP movement is tightly governed and load or tension independent. Previous studies have shown that the velocity of kinetochore movement is often constant as it reduces centromere stretch generated by the preceding kinetochore P movement [4, 5, 6]. Similarly, the rate of AP movement appears unaffected when kinetochore load is reduced by removing the bulk of the chromosome arms [6]. Our micromanipulation results show that AP velocity does not increase when a region of the chromosome arm is stretched to twice its rest length. We conclude that the kinetochore AP motility state governs the rate of movement and resists both polar ejection forces near the spindle pole and centromere tension. AP velocity may be rate limited by the proteins (perhaps microtubule motor proteins) that couple kinetochores to growing microtubule plus ends or other factors that limit the rate of kinetochore microtubule plus end assembly [6, 9, 17, 38]. The molecular mechanisms at the kinetochore that sense tension and control switching between P and AP states are poorly understood. In vitro studies on isolated CHO cell chromosomes show that increased phosphorylation results in plus-end-directed microtubule gliding across kinetochores while reduced phosphorylation results in gliding in the opposite direction [17]. However, these biochemical changes have not been successfully linked to changes in kinetochore motility in vivo although recent evidence indicates that kinetochore components are differentially phosphorylated in mitosis [38, 42–45]. So far, changes in the phosphorylation state of the kinetochore have been strongly correlated with the role of kinetochores in the spindle assembly cell cycle checkpoint for anaphase onset and not with the direction of kinetochore movement [17, 39–41 and reviewed by 46]. Nevertheless, our manipulation results show that tension regulates kinetochore directional instability and predict that tension alters the
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biochemical state of the kinetochore to control the direction of kinetochore motion. We thank Dr. Bruce Nicklas and Susan Ward, whose patient nurturing allowed for easy access into the world of micromanipulation and needle pulling. We also thank Steve Parsons, Phong Tran, Jen Waters, and Vicki Skeen for their support, enthusiasm, and editing during the course of this project. Special thanks go to Liz Blanton for helpful discussions of force analysis and figure preparation, to V.S. for filling in, and to Dr. Meg Kenna for editing early versions of the manuscript. This work was supported by a grant from the National Institutes of Health (GM24364).
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37. Svoboda, K., and Block, S. M. (1994) Cell 77, 773–784. 38. Desai, A., and Mitchison, T. J. (1995) J. Cell Biol. 128, 1–4. 39. Gorbsky, G. H., and Ricketts, W. A. (1993) J. Cell Biol. 122, 1311–1321. 40. Davis, A., Sage, C. R., Dougherty, C., and Farrell, K. W. (1994) Science 264, 839–842. 41. McIntosh, J. R. (1994) in Microtubules (Hyams, J., and Lloyd, C., Eds.), Wiley-Liss, New York. 42. Rattner, J. B., Lew, J., and Wang, J-L. (1990) Cell Motil. Cytoskel. 17, 277. 43. Shero, J. H., and Hieter, P. (1991) Genes Dev. 5, 549–560. 44. Nicklas, R. B., Krawitz, L. E., and Ward, S. C. (1993) J. Cell Sci. 104, 961–973. 45. Nicklas, R. B., Ward, S. C., and Gorbsky, G. J. (1995) J. Cell Biol. 130, 929–939. 46. Rutner, A. D., and Murray, A. W. (1996) Curr. Opin. Cell Biol. 8, 773–780.
Received January 24, 1997 Revised version received June 24, 1997
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