The Spindle Apparatus and Cytokinesis in Spermatocytes of the Silkworm B. mori

Spermatocytes cultured underneath a layer of halocarbon oil were relatively large (∼33 μm in diameter), optically clear, and amenable to micromanipulation. Digital-enhanced polarization microscopy on cells from late metaphase to early anaphase revealed a short but robust spindle (Figure 1A and Video S1; n = 9), whose relatively small size provided ample space for manipulation of spindle microtubules. To visualize the dynamics of microtubules and actin filaments during cytokinesis, we microinjected metaphase-stage spermatocytes with rhodamine tubulin and low-level Alexa Fluor 488 phalloidin (Figure 1B and Video S2; n = 4). The fluorescently labeled cytoskeleton was imaged using spinning disc confocal microscopy. Furrow initiation occurred during early anaphase after actin aggregates emerged at the interdigitating microtubule plus ends at the equator (Figure 1B, between 0 and 4). The aggregates gradually enlarged before coalescing with cortical actin filaments to assemble the contractile ring that constricts the cell (Figure 1B, 4–45). The time from anaphase onset to cleavage was not significantly different for the uninjected control cells (48.4 ± 2.5 min) and the injected, double-labeled cells (49.5 ± 5.0 min), suggesting that the low-level phallodin was not perturbing actin function.

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Figure 1. Cytokinesis of Silkworm Spermatocytes

(A) Polarization micrograph of a dividing spermatocyte.

(B) Confocal micrographs of a spermatocyte microinjected with rhodamine tubulin (microtubules false colored green), and low-level Alexa Fluor 488 phalloidin (actin false colored red). Actin aggregates appeared at the equatorial microtubule plus ends during early anaphase (0–20), then fused into a contractile ring that bisected the cell (4–45).

(C) Astral flagella, as well as naturally, asymmetrically positioned asters, were visible in dividing spermatocytes. Astral flagella were visible only in fixed, immunostained cells (a–c). Astral microtubules were prominent in fixed anaphase spermatocytes (e.g., b). One aster was loosely attached to the spindle (arrow). Occasionally, the asters (arrows) in fixed (a and c) and live (d and e) spermatocytes (arrows) were naturally positioned on the same side of the spindle. a and d, metaphase; b, c and e. anaphase. Fixed cells were double-immunolabeled for actin and tubulin; live cells were labeled as in (B).

Time in min. Time 0 in (A and B), anaphase onset. Bars, 10 μm.

https://doi.org/10.1371/journal.pbio.0060209.g001

Image stacks of fixed, immunostained cells revealed dense arrays of astral microtubules radiating from the centrosomes to the polar cortex (Figure 1C, a–c; n = 15). Notably, asters in both fixed (Figure 1C, a–c) and live (Figure 1C, d and e; n = 15; also in Figure 1B, 0) cells appeared to be attached only loosely to the spindle, which is a natural phenomenon in silkworm spermatocytes [31]. Presumably due to the centrioles' motile flagellar axonemes (Figure 1C, a–c; as seen in immunostained cells), both asters were mobile within the cell. In approximately 15% of the cells, the asters even moved to the same side or the same pole of the spindle, as documented in metaphase (Figure 1C, a and d) and anaphase (Figure 1C, c and e) stage cells. This natural phenomenon makes silkworm spermatocytes an ideal system for separating the role of asters from that of the central spindle in cytokinesis.

Evidence for Polar Relaxation as Gauged by Redistribution of Cortical Actin

Cortical flow of actin filaments driven by spindle microtubules.

In cells in which both asters (Figure 2A, arrows) happened to be located at the same pole of the spindle, cortical actin filaments appeared to flow to the opposite side of the cell during anaphase (Figure 2A, 0–6.3, and Video S3; n = 4. Note: panels were chosen to highlight cortical flow; these focal planes are not optimal for viewing actin aggregates). In such cells, the midzone of the central spindle was never aligned with the cell's equator, and these cells divided asymmetrically around the equator of the shifted spindle (Figure 2B and Video S4; n = 6). Our observation suggested that astral microtubules could relax the polar cortex by excluding cortical actin filaments from the pole. However, since the asymmetric furrow was aligned with the spindle equator, it was also possible that the spindle midzone signaled the band of cortex that encircled the spindle equator, stimulating recruitment of actin to the furrow. To discern between these alternatives, we needed to assess whether actin could be redistributed within the cortex in the absence of an intact central spindle. Thus, we used a microneedle to push the spindle poles together in anaphase, mechanically remodeling the spindle apparatus. We refer to this operation as “collapsing” the spindle. During the remodeling process, the spindle was simultaneously relocated to a region of the cell previously unexposed to any furrow cues (e.g., nascent actin aggregates). By collapsing the spindle, we accomplished three objectives. First, the act of collapsing served to reorganize the central spindle, destroying its equatorial configuration. This likely impeded its ability to stimulate the cortex in its new location, preventing the formation of a complete, circular furrow. Second, relocation of the spindle presumably prevented deposition of additional furrow cues at the original cortical site, and depending on the timing of the move, may well have prevented deposition of any cues at the original location. Third, relocation and collapsing of the spindle allowed us to induce controlled exclusion of actin filaments from a region of our choice; relocation brought the spindle close enough to the cortex to initiate cortical flow, whereas collapsing exposed or freed some microtubule plus ends that originally interdigitated with the opposite half of the spindle, potentially altering their dynamic properties (see below).

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Figure 2. Cortical Actin Filaments Were Excluded from the Polar Cortex Bordering the Asymmetrically Distributed Asters

(A) Both asters (arrows) were naturally positioned at the upper pole of the spindle. Cortical actin filaments flowed towards the lower polar cortex during early anaphase. Cells in Figure 2 were labeled as in Figure 1B. Time 0 depicts anaphase onset.

(B) Cortical actin filaments, excluded by asymmetrically distributed asters (arrows), assembled a contractile ring around the equator of the naturally shifted spindle. Actin aggregates were also present by later anaphase.

Time in min. Time 0, anaphase onset. Bars, 10 μm.

https://doi.org/10.1371/journal.pbio.0060209.g002

Shortly after anaphase onset (Figure 3A, 0, and Video S5; n = 12), we pushed the spindle apparatus with a microneedle to an arbitrary region of the cell cortex while collapsing it as described (Figure 3A, 3). Shortly thereafter, cortical actin filaments began to flow away from the spindle microtubules to the opposite side of the cell (Figure 3A, 10). Because actin filaments are a major component of actomyosin contractile elements, their exclusion from the cortex near the spindle would presumably result in cortical relaxation. This microtubule-driven actin flow resulted in asymmetric distribution of cortical actin filaments (Figure 3A, 10) and initiation of contractile ring assembly (Figure 3A, 16, and Video S5) around the presumptive, newly defined zone of microtubule overlap [32]. The microtubule-driven actin flow persisted into telophase, as shown in similar experiments in which cells were manipulated while undergoing cytokinesis (Figure 3B and Video S6; n = 26). This resulted in asymmetric placement of the contractile ring (Figure 3B, 18–37). Because randomly dislocated asters (Figure 2) and arbitrarily repositioned, collapsed spindles (Figure 3A and 3B) caused cortical actin flow, we hypothesized that cortical relaxation could be induced at any region of the cell cortex by microtubules from any source. To test this idea, we mechanically remodeled cells by placing both asters in one region of the cell, separated from the asterless, collapsed spindle, as depicted in the schematic in Figure 3C (Figure 3C, 5–12, and Video S7; n = 8). As expected, the contractile ring assembled between the two noninterdigitating structures due to actin exclusion from both “poles” (Figure 3C, 5–66, and Video S7).

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Figure 3. Cortical Flow of Actin Filaments Was Driven by Spindle Microtubules

(A) During anaphase (0), when the spindle apparatus was collapsed, and pushed with a microneedle to an arbitrary region of the cell cortex (3), actin filaments flowed to the opposite side of the cell (3–25). This redistribution of actin filaments by spindle microtubules resulted in asymmetric cell division (71).

(B) Repositioning and reorganization of the spindle (2) during telophase resulted in a similar scenario, preceded by regression of the original furrow.

(C) An anaphase cell (with only one aster visible in this focal plane; 0) was mechanically remodeled by moving spindle microtubules to the upper portion of the cell and collapsing the spindle (shown at top of cell, 5–21). The collapsed spindle was manipulated to detach and relocate first one of its asters (dot in center of cell, 5) and then its second aster (to left of first aster; both asters marked by arrows, 12–21). Possibly due to actin exclusion (5–21) by microtubules from both structures, the contractile ring formed between the collapsed spindle and the pair of detached asters (21–66). This is shown in the schematic diagram (representing the cell in 21).

(D) A schematic of the set-up, with fluorescence images of microtubule-driven actin flow blocked by a microneedle. The schematic shows the cell's outline, the location of the needle (gray), and the direction of flow of cortical actin (red), as induced by the collapsed, repositioned spindle (green, at top). The manipulation needle (arrows), which indented but did not pierce the plasma membrane, locally intercepted the actin flow; brighter fluorescence accumulated on the side of the needle that faced the repositioned spindle (1.7–4.5). The three large red objects (3.4–4.5, upper right) are cell division scars, i.e., remnants from former cytokinetic rings. These scar structures can also move around within the cortex during cortical flow. Note: despite removal of the holding needle, the spindle is not depicted as a giant aster, since the reorganization from monopolar spindle to aster takes at least several minutes. Cells in (A–D) were labeled as in Figure 1B. Time in min. Time 0 in (A), (C), and (D), mid to late anaphase; in (B), telophase. Bars, 10 μm.

https://doi.org/10.1371/journal.pbio.0060209.g003

Cortical stiffness correlated with local density of cortical actin.

We had been using local density of cortical actin to infer cortical stiffness, and we needed to test the validity of this assumption. To assay the degree of cortical stiffness in a region of high actin density relative to one of low actin density, cortical flow of actin filaments was induced using a repositioned, collapsed spindle to generate a density differential. After cortical flow was induced, the cortex farthest from the spindle (high actin density) was probed by depression with the side of a flexible microneedle. As predicted, the needle tip was bent by the cortex (Figure 4; n = 7). When the cortex on the opposite side of the cell (low actin density) was probed using the same needle, the cortex was deformed, while the needle remained unbent (Figure 4; same seven cells). (Similar results were obtained using grasshopper spermatocytes; unpublished data (n = 6).) Thus, we concluded that regions with a high density of cortical actin are indeed stiffer (i.e., less relaxed) than regions of low density, justifying our use of the term “polar relaxation” to describe redistribution of actin via cortical flow.

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Figure 4. Cortical Stiffness Correlates with Local Density of Cortical Actin

(A) After the spindle was repositioned and collapsed to induce cortical flow, we tested the stiffness of the actin-dense region of the cortex. (a) The needle (marked with asterisks) in its initial position, tangential to the actin-rich cortex. (b) After the needle was pushed against the cortex, the stiff cortex deflected the needle.

(B) After rotating the cell in (A) by 180°, the cortical stiffness of the actin-depleted region of the cortex was tested. (a) The needle (marked with asterisks) in its initial position, tangential to actin-depleted cortex. (b) After the needle was pushed against the cortex, the relaxed cortex was deformed by the needle without deflecting it. F-actin was labeled using low-level Alexa Fluor 488 phalloidin to monitor flow. Bars, 10 μm.

https://doi.org/10.1371/journal.pbio.0060209.g004

Evidence for Equatorial Stimulation as Gauged by De Novo Assembly of Actin

De novo assembly and delivery of actin aggregates mediated by microtubule plus ends.

Two sources of actin may contribute to contractile ring assembly: pre-existing actin filaments from the cell cortex and the cytoplasm, and actin aggregates that are assembled de novo at the equator. We have demonstrated that microtubule-driven cortical flow provides a mechanism by which cortical actin is excluded from the polar cortex. (Polar microtubules may also play a role in clearing cytoplasmic actin filaments from the polar region. See legend to Video S4.) We hypothesized that microtubules might stimulate de novo actin assembly at the equator, since the speckles of actin fluorescence that were detected at equatorial microtubule plus ends increased in number and size as anaphase progressed (Figure 1B). To detect de novo actin assembly, we monitored its dynamics starting at the metaphase-anaphase transition in cells with labeled microtubules and actin filaments (Figure 5A and Video S9; n = 15). Before onset of anaphase, actin fluorescence was absent from the spindle equator where the aligned metaphase chromosomes were located (Figure 5A, 0). As the cell entered anaphase (Figure 5A, 2), speckles of actin fluorescence soon emerged at the spindle midzone where microtubule plus ends overlapped (Figure 5A, 2–4). The speckles gradually grew into bigger aggregates as anaphase proceeded (Figure 5A, 4–10, and Video S9). The de novo emergence and accumulation of actin fluorescence was readily apparent when actin alone was visualized, using the rhodamine channel (Figure 5A, insets). Notably, nascent actin aggregates were assembled across the entire midzone of the central spindle, and delivered laterally to the incipient contractile ring by microtubules (Figure 5B and Video S10; n = 13). This novel means of delivery was driven by a structural reorganization of each spindle half: microtubules became more focused at their minus ends (at the spindle poles), while splaying out toward the equatorial cortex at their plus ends, thus enlarging the spindle midzone region. These splaying microtubules were organized into bundles, some of which were coated with an actin aggregate at the plus ends of the bundle, enabling its delivery to the cortex. Lateral transport appeared to begin around the time of furrow initiation, while cortical flow was already in progress. However, we could not definitively rule out an earlier starting point, since small numbers of peripheral microtubules could have escaped detection. Moreover, it was problematic to define a precise endpoint, as the trajectory of the outwardly splaying spindle was confounded by the ingressing furrow.

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Figure 5. Interdigitating Microtubule Plus Ends Hosted De Novo Actin Assembly and Delivered Actin Aggregates to Equator

(A) Emergence (0–2) and growth (2–10) of nascent actin fluorescence at the equatorial microtubule plus-end overlap region reflect de novo assembly of actin aggregates.

(B) A splaying microtubule bundle (arrowheads) delivered the actin aggregate into the ingressing furrow. Cells in (A) and (B) were labeled as in Figure 1B.

(C) De novo assembly of actin aggregates was not inhibited by microtubule stabilization (compare to (A) and Figure 1B). Microtubules were labeled and stabilized by Oregon green paclitaxel, and actin filaments were labeled by rhodamine phalloidin. Insets: actin channel. Time in min. Time 0 in (A and C), immediately before anaphase onset; in (B), late anaphase. Bars, 10 μm.

https://doi.org/10.1371/journal.pbio.0060209.g005

Polar Relaxation Was Dependent on Microtubule Dynamics, whereas Equatorial Stimulation Was Largely Independent

Lateral transport of de novo–assembled actin aggregates by paclitaxel-stabilized microtubules.

To further test how central spindle microtubules deliver de novo–assembled actin aggregates to the cell cortex, we remodeled these actin-tipped microtubules with a microneedle to expose their plus ends to the cortex (Figure 6). After anaphase onset, a central spindle (Figure 6A, a) was collapsed by pushing the spindle poles together (Figure 6A, b). Microtubules from the collapsed spindle reorganized into two lateral bundles that extended from the centrosomes outward, toward opposite sides of the cell cortex, with each bundle containing microtubules from both original half spindles (Figure 6A, c). Over time, the lateral spindle could reorganize into a monopolar spindle (Figure 6A, d) if the chromosomes and centrosomes were held together with a microneedle, or into a giant aster, if the needle was removed (Figure 6A, e). To prevent the formation of new regions of microtubule overlap [32], and to investigate dependence on dynamics, we stabilized microtubules with Oregon green paclitaxel (Figure 6B, n = 4 slowly evolving spindles; 6C, n = 12 lateral spindles; 6D, n = 6 monopolar spindles; and 6E and 6F, n = 7 giant asters).

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Figure 6. Redistribution of Cortical Actin, but Not Equatorial Stimulation, Was Dependent on Microtubule Dynamics

(A) A schematic showing remodeling of the central spindle by micromanipulation. A post-anaphase spindle (a, green) was collapsed using a micromanipulation needle (b, gray). The manipulation created two lateral microtubule bundles, with the exposed microtubule plus ends of the bundles pointing toward opposite sides of the cortex. The interior ends of the bundles flanked chromosomes (c, blue), kinetochores (c, purple) and centrosomes (c, orange). If the collapsed spindle was brought close to the cortex and held in place by a microneedle, it reorganized into a monopolar spindle (d). If the holding needle was removed, the spindle usually reorganized into a giant aster (e).

(B) Actin exclusion was inhibited when the remodeled spindle was stabilized by paclitaxel. Neither the stabilized lateral spindle (8) nor the evolving monopolar spindle (12–22) induced unidirectional flow of actin filaments (compare to Figure 3). Cells in (B–F) were labeled as in Figure 5C.

(C) Actin aggregates were assembled at the plus ends of stabilized, laterally oriented microtubule bundles (0–2, arrowheads), and were delivered to the non-equatorial cortex where splaying microtubule bundles were in contact with the cortex (2–12, arrows).

(D) A monopolar spindle (2) gradually splayed its microtubule bundles toward the cell cortex (2–21), which delivered actin aggregates (2, arrowheads) and induced a furrow (11–21, arrows). The furrow eventually regressed (61).

(E) Tracking of an actin aggregate delivered from the plus ends of one microtubule bundle to the cell cortex. The giant aster had chromosomes in its center and microtubule plus ends pointing outward, as in (A, panel e). When the aster was placed near the cell cortex using a microneedle, an actin aggregate moved away from the microtubule bundle plus ends and merged into the cortex (box, 0–8.8).

(F) The region of interest in (E) is shown with additional intervening time points to highlight the increasing cortical fluorescence caused by merging of actin aggregates. Time in min. Time 0 in (B), (D), and (E), mid to late anaphase; in (C), telophase; in (F), identical to 0 in (E). Bars, 10 μm.

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We mechanically positioned the remodeled and stabilized spindle so that its exposed microtubule plus ends, which were coated with the de novo–assembled actin aggregates, were removed from the original equatorial cortex and thus from any cues already present there. In all spindle configurations tested (Figure 6A, b–e), actin aggregates were delivered by the bundled microtubules to the nearby cortex. Notably, microtubule splay in a lateral spindle gradually brought microtubule plus ends close to the cell cortex, and hence allowed the delivery of actin aggregates (Figure 6C and Video S13). Furrow initiation could occur in such manipulated cells, provided the actin aggregates were delivered to a cortical zone that encircled the cell (Figure 6D, arrows). Using time-lapse imaging, we followed a giant aster positioned near the cell periphery (viewed from above, as in Figure 6A, e) and documented its delivery of an actin aggregate from the plus end of a microtubule bundle to the cell cortex (Figure 6E and 6F and Videos S14 and S15). The accumulation of actin aggregates made the cell cortex significantly brighter over time (Figure 6F, 0 onward), indicating that delivery of actin aggregates can proceed even in the absence of dynamic microtubules, so long as splay of central spindle microtubules is not blocked.

In the final stage of lateral transport, actin aggregates had to be released from the tips of the microtubule bundles to allow their delivery to the cortex. To examine this process in greater detail, the red and green channels were analyzed individually for signs of costaining at the tips of microtubule bundles (in transit from the central spindle to the cortex). The signal from the microtubules at the peripheral ends of the bundles was much weaker than the signal from the actin; thus, the costained regions appeared red rather than yellow in the merged images shown in Figure 6. However, close analysis revealed a correlation between the length of the costained region of the bundle and the distance from bundle tip to cortex (Table 1). This distance was actually measured from the distal end of the actin aggregate rather than from the distal end of the bundle itself. The length of the actin aggregate remained fairly constant—approximately 0.8 to 0.9 μm (Table 1). When a bundle was more than about 1.2 μm from the cortex, at least a portion of the aggregate (perhaps close to the entire aggregate) overlapped with the tip of the bundle. As the bundle, with its associated cargo, came within about 0.41–1.2 μm of the cortex, it was no longer possible for the entire length of the aggregate to fully overlap with the bundle, even taking into account the uncertainty of the measurements. In other words, the aggregate appeared to be “sliding” off the tip of the bundle, toward the cortex. Finally, as the actin-tipped bundle approached within 0.4 μm of the cortex, i.e., closer than the length of the aggregate itself, the aggregate was released completely from the bundle, and the overlap dropped to zero.

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Table 1.

Measurements of Actin Aggregate-Tipped Single Bundles

https://doi.org/10.1371/journal.pbio.0060209.t001

Equatorial Stimulation, but Not Polar Relaxation, Was Dependent on RhoA Activity

We wanted to determine whether de novo assembly of actin aggregates at the plus ends of central spindle microtubules requires RhoA as a regulator. To do so, we microinjected late metaphase cells with C3 ribosyltransferase, a botulinum toxin that specifically inhibits Rho by ADP ribosylation [33]. We hypothesized that this inhibition would disrupt de novo actin assembly at the spindle midzone, without affecting cortical flow of actin. As expected, the C3 transferase-treated cell continued to divide but failed to accumulate actin aggregates in the central spindle (Figure 7). As seen in optical sections of control cells (n = 15), actin accumulated both at the equatorial cortex (Figure 7A, top and bottom) and at microtubule plus ends within the central spindle (Figure 7A, middle). In contrast, C3 transferase-treated cells (n = 9) accumulated actin at the equatorial cortex (Figure 7B, top and bottom), but not within the central spindle (Figure 7B, middle). This result implies that RhoA inactivation inhibits actin assembly at the plus ends of central spindle microtubules (i.e., the initial step of equatorial stimulation), but does not inhibit relocation of cortical actin from the poles to the equatorial cortex (i.e., polar relaxation).

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Figure 7. Equatorial Stimulation, but Not Redistribution of Cortical Actin, Was Dependent on RhoA Activity

(A) A cytokinetic control cell, optically sectioned lengthwise, displayed actin aggregates at the microtubule plus ends of its spindle, both at the equatorial cortex (top and bottom) and in the central spindle (middle). Cells in (A and B) were labeled as in Figure 1B.

(B) In a cytokinetic cell microinjected with C3 transferase to inhibit RhoA, actin aggregates at the microtubule plus ends of the central spindle were absent.

(C) Despite inhibition of RhoA, actin flow was induced (2.5–11), following manipulation of the collapsed spindle to one side of the cell (2.5). Excluded filaments assembled into a contractile ring (35). Cell was labeled as in Figure 1B, and microinjected with C3 transferase.

(D) In a cell treated with paclitaxel and C3 transferase, the furrow failed to initiate due to inhibition of both pathways for redistribution of actin. Actin filaments were scattered in the cytoplasm long after the cell entered anaphase, as determined by the presence of splaying microtubules (17–73). (D) was labeled as in Figure 5C. Time in min. Time 0 in (C), late anaphase; in (D), metaphase-anaphase transition. Bars, 10 μm.

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Additional tests confirmed that in C3 transferase-treated cells, actin filaments were still excluded from the cortex adjacent to the dislocated spindle and incorporated into the contractile ring (Figure 7C, n = 8). However, when late metaphase cells were microinjected with C3 transferase, and subsequently treated with paclitaxel at anaphase onset, both pathways for actin redistribution were affected (Figure 7D, n = 5); specifically, delivery to the equatorial cortex from both the polar cortex and the midzone was impeded. Cells failed to divide long after entering anaphase (Figure 7D, 0–73 min), due to inhibition of both de novo assembly of nascent actin aggregates, and relocation of cortical actin filaments to the contractile ring.

Polar Relaxation and Equatorial Stimulation Both Induce Contractile Ring Assembly and Cell Cleavage

By mechanically manipulating the spindles of double-labeled silkworm spermatocytes, we show how polar relaxation (as gauged by redistribution of cortical actin) and equatorial stimulation of the cortex (as gauged by de novo actin assembly and transport) both contribute to contractile ring assembly and furrow induction. We demonstrate that spindle microtubules deliver both inhibitory and stimulatory signals to the cell cortex during furrow formation, depending on their configuration and location within the cell. The dynamic astral microtubules drive actin filaments from the polar region to the equatorial cortex, which relaxes the polar cortex, while increasing cortical tension at the actin-dense equator. Meanwhile, bundles of interdigitating microtubules across the entire central spindle stimulate de novo assembly of actin aggregates at their overlapping plus ends, and deliver the aggregates to the equatorial cortex. These dual signaling mechanisms ensure that both cortical actin and central spindle-associated actin are delivered to the equatorial cortex, resulting in a robust contractile ring, and providing redundancy to safeguard an essential cellular process. Videos S4 and S9, as well as previous studies [34,35], provide hints that cytoplasmic actin filaments (perhaps vesicle bound; [29]) may constitute a third source of actin that could ultimately be delivered to the equatorial cortex by microtubules.

We show that in the absence of equatorial stimulation, contractile ring assembly and furrowing can still be induced by means of actin redistribution driven by cortical flow (Figure 3A and 3B and Figure 7C). We accomplished this by displacing the entire spindle apparatus, bearing “stimulatory” furrow cues that were reorganized (via spindle collapse), or C3 transferase-suppressed, to an arbitrary region of the cortex. The existence of a cortical flow of actin filaments away from the displaced spindle microtubules was established by our ability to intercept the flow locally, using a microneedle (Figure 3D). We found that the flow of cortical actin during cytokinesis can be induced by any microtubules positioned near the cortex, provided their plus ends are dynamic and accessible (i.e., neither interdigitating nor otherwise bundled or stabilized). We hypothesize that collapsing the spindle can mechanically free a subset of plus ends; this could allow those formerly interdigitating central spindle microtubules to become more dynamic and to take on properties held by astral microtubules, such as the ability to contact the cell cortex and induce cortical flow of actin filaments. Interdigitating microtubules in an intact central spindle would be able to contact the cortex only laterally, and as incorrectly oriented bundles, which should not be conducive for promoting cortical flow. As predicted by this finding, furrow formation can occur between a collapsed asterless spindle and its two detached asters in a remodeled cell (Figure 3C). We have also shown that this microtubule-driven actin flow begins shortly after anaphase onset and persists through telophase (Figure 3A and 3B), contributing to both contractile ring assembly at the equator and furrow inhibition at the poles throughout the furrow induction and ingression stages. Cortical flow during cytokinesis has previously been inferred or observed for contractile elements including myosin II [28,36,37] and pre-existing actin filaments [17,35,38], as well as for a membrane-bound receptor-ligand complex, membrane domains, and cell surface proteins [39–41]. As the movement of certain surface proteins mirrors that of cortical actin, it has been suggested that the actin network could serve as a scaffold for these proteins, thus engineering their co-transport to the furrow region [41].

How might microtubules interact with actin filaments to effect their exclusion from the poles? Since paclitaxel inhibits cortical flow (Figure 6B–6D), actin filaments are likely to be driven from the polar region towards the equatorial cortex by elongating microtubules. Following anaphase onset, the plus ends of astral microtubules can elongate toward and contact the cortex [20,42]. Such microtubules would be anchored to the spindle pole at their minus ends, and conceivably constrained at their plus ends by indirect attachment to a mobile cortical component, namely filamentous actin (analogous to cortical capture via Bim1/Kar9/Myo2 [43,44]). Given the microtubule's length, flexibility, geometrical orientation within the cell, and constraints on its direction of movement, its elongation may be sufficient to drive cortical actin toward the equator. Elongation of tethered microtubules against a fixed barrier can produce force of up to 10 pN, as gauged by microtubule buckling [45]. Presumably, this force could be harnessed to allow tip-attached cortical actin to “hitchhike” to the equatorial cortex, where it would be released, possibly by binding to myosin II [38] or anillin [46]. Alternatively, or in addition, actin filaments could be transported along the subset of microtubules that are contiguous to the cortex, and longitudinally aligned, with appropriate polarity. Motor proteins would provide the driving force for transport, with microtubule elongation serving only to extend the track toward the equator. Actin filaments could either be cargo for a plus-end–directed kinesin (attached via an adaptor protein), or else the actin could be coupled to the microtubule track via kinesin/myosinV-type hetero-motors (as modeled in [44,47]), and slide along the stationary microtubule toward the equatorial cortex, with kinesin providing directionality for the flow. The tracks could consist of the cortically apposed section of centrosome-attached astral microtubules and/or those released from the centrosomes [42,48–50] that become cortically apposed [34,47,51,52].

We also show that actin aggregates can be assembled de novo within the central spindle in the absence of cortical-flow based redistribution of actin, by using paclitaxel to impede the flow. Similarly, paclitaxel-stabilized microtubules can induce a cleavage furrow in mammalian cells [53], perhaps implying mechanistic conservation. We demonstrate that cortical stimulation can be induced at any region in the cortex by microtubule plus ends of the central spindle (Figure 6). Our data reveal that de novo–assembled cytoplasmic actin aggregates can comprise a supply of actin sufficient to form the contractile ring and induce cell cleavage (Figure 6D). Newly assembled actin aggregates are precursors of the contractile ring, since they ultimately merge with the ring (Figure 5). The existence of de novo assembly of actin during contractile ring formation is consistent with evidence from diverse cell types, including grasshopper spermatocytes (unpublished data), yeast [54,55], Xenopus eggs [56], and mammalian cells [17,38]; in addition, there is precedent for the presence of actin nucleating protein formin at microtubule plus ends [57]. Assembly of actin aggregates is sensitive to the Rho GTPase inhibitor, C3 transferase (Figure 6B, middle), supporting the existence of a microtubule-dependent zone of active RhoA during cleavage plane specification [5,23,58–62]. Our findings also raise the possibility that equatorial stimulation signals could be transported along stationary central spindle microtubule tracks from the spindle pole region to the midzone (Figure 5B), before ultimately reaching the equatorial cortex, perhaps via lateral transport.

Actin aggregates are assembled across the entire spindle midzone (Figure 5A and 5C); hence they must be transported laterally from the plus ends of the bundled microtubules to the equatorial cortex. We have captured this novel means of delivery using time-lapse sequences (Figures 5B, 6E, and 6F). Bundles of central spindle microtubules, some tipped with actin aggregates, splay laterally outward as the spindle changes shape, thereby conveying their cargo to the incipient furrow (Figure 5B). The actin cargo remains associated with the tips of the microtubule bundles until the bundles are within about 0.4 μm of the cortex, less than the length of the aggregate itself (Table 1). At this point, the microtubule-associated aggregates appear to be released from the microtubule bundles, in that the costaining at the microtubule tips disappears. The dissociation of the actin from the bundles may be mediated by kinesins bound to the bundle, as the aggregate appears to slide off the end of the bundle as it approaches the cortex (Table 1). This transport of actin aggregates may be largely independent of microtubule dynamics; when a post-anaphase, paclitaxel-stabilized spindle with mechanically exposed, bundled plus ends is brought close to the cortex by micromanipulation, it remains functional for delivery of actin aggregates to the cortex (Figure 6E and 6F). The induction of an ectopic furrow following cortical deposition of actin aggregates from microtubule plus ends (Figure 6D) indicates that actin-tipped microtubules may contain sufficient furrow constituents (i.e., structural constituents along with any co-transported stimulatory cues) to permit contractile ring assembly. This idea is consistent with work by Hu et al., showing that the tips of monopolar spindles contain critical stimulatory components (e.g., the Rho GEF Ect2) [21].

We propose that in the bipolar spindle, other spatially dynamic midzone components may also utilize this microtubule-based lateral transport mechanism—either co-transported with actin aggregates, or else delivered via a distinct set of midzone microtubules. By ensuring timely relocation of cytokinetic components, lateral transport could play a role in processes such as midzone signaling of the equatorial cortex, assembly and maintenance of the contractile ring and abscission. For example, the midzone component centralspindlin becomes localized to the equatorial cortex during anaphase. Bundled interdigitating microtubules, tipped with centralspindlin, may become docked at the contractile ring via Anillin [46,63]. We suggest that these tipped microtubules may originate from the central spindle and deliver their cargo to the cortex via lateral transport. Similarly, lateral transport could provide a means of delivery to the cortex for a recently characterized subset of vesicles. These vesicles are associated both with actin and with the central spindle, and are thought to contribute to furrow ingression [29]. In fixed cells, the authors saw thin microtubule bundles potentially connecting the central spindle with the periphery [29], consistent with a lateral transport mechanism. As another example, by early anaphase, the chromosomal passenger complex (CPC) is at the mitotic spindle midzone, whereas later in anaphase it is also present at the equatorial cortex (reviewed in [64]), perhaps arriving there by lateral transport. Interestingly, in Dictyostelium, the CPC component INCENP has been shown to bind actin [65]. It is also worth determining whether a subset of RhoA may reach the equatorial cortex via lateral transport, given that RhoA localization may depend on microtubule organization [58].

Model for Microtubule Induction of Contractile Ring Assembly

Our results explain how a single spindle component, the microtubule, can play opposing roles in two complementary pathways of contractile ring assembly. The seemingly incongruous stimulatory and inhibitory effects on the cortex are produced by different configurations of microtubules that populate different regions of the cytokinetic cell. We propose a microtubule induction model to describe how microtubules perform their dual signaling at the poles and the equator to induce contractile ring formation and cleavage furrow initiation. During early anaphase, dynamic astral microtubules at the poles inhibit ectopic furrow formation by excluding pre-existing actin filaments from the poles (Figure 8A, red dotted arrows and red lines at cortex), ultimately resulting in actin accumulation at the incipient equatorial furrow. Meanwhile, at the plus ends of the relatively stable central spindle microtubules, actin aggregates are assembled de novo, and enlarge over time (Figure 8A, red lines in midzone). As the cell progresses toward telophase, preexisting cortical actin filaments continue to flow from the polar cortex to the equator (Figure 8B, red dotted arrows). At the same time, the plus ends of bundled central spindle microtubules splay laterally to deliver the actin aggregates to the equatorial cortex (Figure 8B, red solid arrows). The newly delivered aggregates coalesce with the actin excluded from the polar cortex to assemble the contractile ring (Figure 8C).

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Figure 8. Model for Spindle Microtubule Induction of Contractile Ring Assembly

(A) During furrow induction and ingression, dynamic astral microtubules exclude pre-existing actin filaments from the spindle pole (red dotted arrows).

(B) This exclusion results in mass migration of filaments to the equator. Meanwhile, overlapping spindle microtubule plus ends at the equator promote de novo assembly of actin aggregates, which are delivered (small red arrows) to the equatorial cortex by splaying bundles of central spindle microtubules.

(C) Actin filaments excluded from the polar region coalesce with actin aggregates transported from the central spindle to the equatorial cortex, for assembly of the contractile ring.

https://doi.org/10.1371/journal.pbio.0060209.g008

In summary, we show that actin is redistributed into the contractile ring via two microtubule-dependent pathways that coexist during cytokinesis of the silkworm spermatocyte: equatorial stimulation and polar relaxation. These dual signaling pathways ensure the fidelity of cytokinesis, which fails only if both mechanisms are inhibited (Figure 7D). With some modification, our model could be generalized to multiple cell types. For example, cytokinetic cells of varying sizes and spindle geometries might rely more heavily on one mechanism than the other to achieve a global balance between tension and relaxation. We have demonstrated that the entire cell cortex is potentially responsive to both stimulatory and inhibitory cues from microtubules during cytokinesis in silkworm spermatocytes. How might the cell restrict the contractile ring to a localized band of equatorial cortex? By stimulating only the equatorial cortex while concomitantly relaxing other regions of the cortex, a cell could ensure cytokinetic fidelity by building in redundancy. Polar relaxation could be thought to include a component of equatorial stimulation in that pre-existing actin filaments are transported to the equatorial cortex. Equatorial stimulation, with its de novo actin assembly, recruits cytoplasmic G-actin to the equator. Hence, the seemingly disparate models ultimately result in redistribution of all available actin resources to the equator for ring assembly.

Silkworm as a Model Organism

Our work with spermatocytes of the silkworm B. mori has enabled us to document a number of novel events essential for understanding contractile ring assembly. These spermatocytes are highly amenable to mechanical manipulations and imaging techniques, making them ideal for cytokinetic studies. Equally important, the silkworm genome has been sequenced [66], paving the way for molecular and genetic studies, such as RNAi inhibition of particular genes. Thus, we believe the silkworm spermatocyte has great potential as a model system for research in cell biology, cell physiology, and developmental biology—given its innate attributes coupled with the power of molecular genetics.

Primary cell culture.

The spermatocytes of silkworm Bombyx mori were isolated from the testes of a 5th instar larva and spread under inert halocarbon oil (400 oil, Halocarbon Products) to form a monolayer of cells in a glass chamber slide. Only cells in the first meiotic division were used.

Microscopy.

Cells were observed with an inverted Zeiss Axiovert 135 microscope modified for both digital-enhanced polarization [6] and spinning disc confocal microscopy (CARV, Carl Zeiss). The microscope is equipped with a 1.4 NA achromatic-aplanatic condenser and a 1.45 NA/100X α-Plan objective (Carl Zeiss). An EM-CCD digital camera (Hamamatsu C9100–12), Simple PCI software (C-image), and Photoshop software (Adobe Systems) were used to record and process images. All images in a given category “n” are fully representative of all unshown data in that category.

Micromanipulation.

Micromanipulation needles were pulled from glass tubing (outer diameter: 1.0 mm; inner diameter: 0.58 mm, World Precision Instruments) as described [67] using a microforge (Narishige, Model MF-830) and maneuvered with a Burleigh MIS-5000 series piezoelectric micromanipulator [6].

Microinjection and preparation of injectants.

Glass tubing (outer diameter: 1.0 mm; inner diameter: 0.75 mm) with an internal capillary (World Precision Instruments) was pulled on a Flaming/Brown P-87 micropipette puller (Sutter Instrument Company) to produce micropipettes with a tip diameter ∼ 0.1 μm. The injectant was back loaded into the micropipette using a Hamilton syringe. Microinjection was conducted at 60 psi using a custom-built pneumatic injector maneuvered with a Burleigh MIS-5000 series piezoelectric micromanipulator.

Alexa Fluor 568 tubulin. Alexa Fluor 568 (Invitrogen) was conjugated to porcine tubulin (Cytoskeleton) following the protocol adapted from Peloquin et al. [68]. The labeled tubulin was resuspended in an injection buffer (20 mM PIPES, 0.25 mM MgSO₄, 1 mM EGTA, 1 mM GTP, pH 6.8) to a final concentration of 3 mg/ml for microinjection.

Phalloidin. Alexa Fluor 488 phalloidin or rhodamine phalloidin (Invitrogen) in methanol was concentrated using a SpeedVac concentrator (Savant) and resuspended in the injection buffer to a final concentration of ∼6.6 μM for microinjection. Although the intracellular concentration of phalloidin was low, controls were performed to ensure that the phalloidin did not adversely affect the function of the actin filaments. The timing of cell division was not slowed down in a phalloidin-treated cell relative to uninjected controls (as in Table S1).

C3 transferase. C3 transferase (Cytoskeleton) was stored as 1 mg/ml aliquots at −70 °C in a buffer containing 500 mM imidazole (pH 7.5), 50 mM Tris HCl (pH 7.5), 1.0 mM MgCl₂, 200 mM NaCl, 5% sucrose, and 1% dextran. Immediately before microinjection, the C3 transferase was mixed with Alexa Fluor 568 tubulin and Alexa Fluor 488 phalloidin to a final concentration of 0.5 mg/ml in the micropipette.

Live cell labeling with the tubulin tracker and Hoechst stain.

Tubulin Tracker (Invitrogen) was used as directed, with the Oregon Green 488 Taxol (paclitaxel) diluted to 50 μM in insect Ringer's solution. The diluted Tubulin Tracker was micropipetted around the target cells, resulting in a further dilution of at least 1000 fold. Hoechst 33342 (Invitrogen) was stored as 10 mg/ml stock aliquots at −20 °C and diluted in Tubulin Tracker buffer to 0.5 mg/ml before micropipetting.

Immunofluorescence microscopy.

Silkworm spermatocytes were fixed and stained as previously described [6]. Microtubules were stained with tubulin primary antibody (Chemicon) and Alexa Fluor 488 conjugated secondary antibody (Invitrogen); actin filaments, with rhodamine phalloidin (Invitrogen); and chromosomes, with DAPI.

Labeling color scheme.

In all figures, microtubules are shown in green, and actin in red (false colored, if necessary). Chromosomes, when labeled, are blue (Hoechst for live cells, DAPI for fixed).

Measurement of cortical stiffness.

The degree of stiffness at one region of the spermatocyte cortex relative to another region was assayed as follows. The spindle of a cell in anaphase was collapsed (by pushing the spindle poles together) and mechanically repositioned near the cortex to induce cortical flow of actin filaments. After waiting 5–10 min to allow for redistribution of actin, the first region of interest was probed with the side of a flexible microneedle, tangential to the cortex. Depending on the flexural rigidity of the needle and the local cortical stiffness, a given needle would either deform the cortex by flattening it, without deflecting the needle, or the cortex would bend the needle. The cell was repositioned (by rotating the microscope slide by 180°) to bring the second region of interest in proximity with the needle, and then reprobed.