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Mol Biol Cell. 2012 June 15; 23(12): 2319–2326.
PMCID: PMC3374750

Constitutive dynein activity in she1 mutants reveals differences in microtubule attachment at the yeast spindle pole body

Kerry S. Bloom, Monitoring Editor
University of North Carolina

Abstract

The organization of microtubules is determined in most cells by a microtubule-organizing center, which nucleates microtubule assembly and anchors their minus ends. In Saccharomyces cerevisiae cells lacking She1, cytoplasmic microtubules detach from the spindle pole body at high rates. Increased rates of detachment depend on dynein activity, supporting previous evidence that She1 inhibits dynein. Detachment rates are higher in G1 than in metaphase cells, and we show that this is primarily due to differences in the strengths of microtubule attachment to the spindle pole body during these stages of the cell cycle. The minus ends of detached microtubules are stabilized by the presence of γ-tubulin and Spc72, a protein that tethers the γ-tubulin complex to the spindle pole body. A Spc72–Kar1 fusion protein suppresses detachment in G1 cells, indicating that the interaction between these two proteins is critical to microtubule anchoring. Overexpression of She1 inhibits the loading of dynactin components, but not dynein, onto microtubule plus ends. In addition, She1 binds directly to microtubules in vitro, so it may compete with dynactin for access to microtubules. Overall, these results indicate that inhibition of dynein activity by She1 is important to prevent excessive detachment of cytoplasmic microtubules, particularly in G1 cells.

INTRODUCTION

Proper function of microtubules depends on their correct organization within cells. In most cells, microtubules are organized by the microtubule-organizing center (MTOC), which nucleates microtubule assembly. Microtubule plus ends extend outward from the MTOC, creating a polarized array of microtubules that the cell uses for the directional transport of vesicles, organelles, and chromosomes (reviewed in Desai and Mitchison, 1997 blue right-pointing triangle). Because many of these transport events involve the movement of large cargoes, they must generate considerable force. For example, in yeast, single microtubules are used to pull the nucleus toward the bud neck and chromosomes toward the spindle poles (O'Toole et al., 1999 blue right-pointing triangle). Thus the anchoring of microtubules to the MTOC must be strong enough to withstand these forces.

The MTOC in the budding yeast, Saccharomyces cerevisiae, is the spindle pole body (SPB), an ~0.5-GDa structure that is embedded in the nuclear membrane (reviewed in Jaspersen and Winey, 2004 blue right-pointing triangle). The SPB is a trilaminar, disk-shaped structure: a central plaque spans the nuclear envelope, an outer plaque faces the cytoplasm, and an inner plaque faces the nucleoplasm. Early in the cell cycle, a structure termed the half-bridge is adjacent to one side of the central plaque. Subsequent SPB duplication produces side-by-side SPBs that are separated by a bridge that is twice the size of the half-bridge. SPB separation severs the bridge, and SPBs with their associated half-bridges move to opposite sides of the nuclear envelope to form the spindle poles.

The γ-tubulin complex nucleates microtubules at the SPB (Marschall et al., 1996 blue right-pointing triangle; Spang et al., 1996 blue right-pointing triangle). In yeast, the γ-tubulin complex contains the γ-tubulin protein Tub4, Spc97, and Spc98 (Knop and Schiebel, 1997 blue right-pointing triangle). In the cytoplasm, this complex is linked to the SPB through Spc72 (Knop and Schiebel, 1998 blue right-pointing triangle). In G1 cells, Spc72 is primarily unphosphorylated and binds to Kar1, which is located in the half-bridge. As cells enter S phase and proceed through mitosis, Spc72 becomes phosphorylated and binds preferentially to Nud1, a component of the outer plaque (Pereira et al., 1999 blue right-pointing triangle). Therefore, cytoplasmic microtubules primarily arise from the half-bridge during G1 and from the outer plaque during mitosis.

The primary role of cytoplasmic microtubules in yeast is to orient the spindle. Two pathways contribute to spindle orientation. Early in the cell cycle a complex of Bim1, Kar9, and Myo2 associates with the plus end of the cytoplasmic microtubule (Korinek et al., 2000 blue right-pointing triangle; Lee et al., 2000 blue right-pointing triangle; Yin et al., 2000 blue right-pointing triangle). Myo2 is a myosin V protein that translocates along polarized actin cables into the bud, thereby pulling the microtubule and its attached SPB toward the bud neck (Yin et al., 2000 blue right-pointing triangle; Hwang et al., 2003 blue right-pointing triangle). This event places the metaphase spindle adjacent to the bud neck. Later in the cell cycle, a second pathway involving dynein ensures that the elongating spindle passes through the bud neck and into the bud (Moore et al., 2009 blue right-pointing triangle). During anaphase, dynactin is recruited to the cytoplasmic microtubule plus end, where it activates dynein. Dynein then interacts with Num1, a protein bound to the bud cortex. Here the minus end–directed motor activity of dynein reels in the cytoplasmic microtubule, thereby pulling the attached SPB into the bud (Farkasovsky and Küntzel, 1995 blue right-pointing triangle; Heil-Chapdelaine et al., 2000 blue right-pointing triangle; Lee et al., 2005 blue right-pointing triangle; Markus and Lee, 2011 blue right-pointing triangle).

To ensure the proper timing of dynein function, its activity is restricted to a small portion of the cell cycle around anaphase (reviewed in Moore et al., 2009 blue right-pointing triangle). Dynein is found on cytoplasmic microtubules during the majority of the cell cycle, so the timing of dynein activity is believed to depend on the loading of the dynactin complex, which becomes enriched at cytoplasmic microtubule plus ends during anaphase (Woodruff et al., 2009 blue right-pointing triangle). She1 inhibits the loading of the dynactin complex onto microtubule ends at other stages of the cell cycle (Woodruff et al., 2009 blue right-pointing triangle). In this study, we show that She1 activity is important to prevent high rates of cytoplasmic microtubule detachment from the SPB. We characterize the molecular nature of these detachment events, show that their frequency depends on the way in which cytoplasmic microtubules are anchored to the SPB, and propose a model for She1 action.

RESULTS

Detachment of cytoplasmic microtubules from the SBP in she1[increment] mutants depends on the cell cycle and dynein activity

We noticed that cytoplasmic microtubules in she1[increment] cells frequently detached from their anchor point at the SPB and moved freely around the cell periphery before depolymerizing (Figure 1A and Supplemental Video S1). Similar cytoplasmic microtubule detachment from the SPB was previously observed in cells containing cnm67Δ or SPC72stu2Δ mutations, which affect the integrity of the SPB outer plaque (Hoepfner et al., 2000 blue right-pointing triangle; Usui et al., 2003 blue right-pointing triangle). We quantified this effect by determining the fraction of cytoplasmic microtubules that detach from the SPB per minute. In asynchronously growing wild-type cells only 0.02% of microtubules detach (Figure 1B). In contrast, in asynchronously growing she1[increment] cells 0.7% of microtubules detach.

FIGURE 1:
she1[increment] increases the rate of cytoplasmic microtubule detachment from the SPB. (A) Time-lapse images of a G1-arrested she1Δ cell expressing GFP-Tub1. The yellow arrowheads point to the plus end and the green arrowheads point to the minus ...

Further observation of microtubule detachment in asynchronous cultures revealed that the majority of these events occurred in cells that were growing early in the cell cycle, before the formation of a bipolar spindle. To measure this difference, we created uniform populations of cells by arresting them either in G1, by exposure to α-factor, or in metaphase, by depletion of Cdc20. During G1 arrest, 0.1% of microtubules detach in wild-type cells and 1.5% of microtubules detach in she1[increment] cells (Figure 1B). During metaphase arrest, 0.02% of microtubules detach in wild-type cells and 0.2% of microtubules detach in she1[increment] cells. Thus, in wild-type and she1[increment] cells microtubule detachment is five- and eightfold more frequent, respectively, in G1 than in metaphase. In G1 and metaphase cells, microtubule detachment is 15- and 10-fold more frequent, respectively, in she1[increment] cells than in wild-type cells.

Woodruff et al. (2009) blue right-pointing triangle reported that She1 limits dynein activity to anaphase by inhibiting the recruitment of dynactin to cytoplasmic microtubule ends at other points in the cell cycle. This result suggests that the increased microtubule detachment observed in she1[increment] cells is likely due to untimely dynein activity. To test this possibility, we measured microtubule detachment in cells lacking the dynactin complex protein Nip100, which is essential for dynein activity. Microtubule detachment rates in she1[increment] nip100[increment] cells were even less than those in wild-type cells for asynchronous, G1, and metaphase populations (Figure 1B). Thus the increased frequency of microtubule detachment in she1[increment] cells depends on dynein activity.

Detachment rate depends on the site of cytoplasmic microtubule anchorage

We were curious as to why the microtubule detachment rate differed between G1 and metaphase. In cycling cells, cytoplasmic microtubules originate from both the outer plaque and half-bridge during the early portion of the cell cycle but extend exclusively from the outer plaque once the spindle has formed (Byers and Goetsch, 1975 blue right-pointing triangle; O'Toole et al., 1999 blue right-pointing triangle; Pereira et al., 1999 blue right-pointing triangle). In cells arrested in G1 by treatment with α-factor, cytoplasmic microtubules originate only from the half-bridge (Figure 2A). Hence, cells arrested in G1 by exposure to α-factor and in metaphase by depletion of Cdc20 contain cytoplasmic microtubules that originate only from the half-bridge and outer plaque, respectively. Because cytoplasmic microtubules originate from two alternate locations in G1 and metaphase cells, detachment rates could reflect differences in the strengths of attachment of microtubules to the SPB. An alternative explanation is that the amount of force pulling on cytoplasmic microtubules could differ during G1 and metaphase. To distinguish between these two possibilities, we wanted to compare detachment rates from these two locations in the same cell cycle state. To this end, we created a strain in which cytoplasmic microtubules are nucleated from the outer plaque even when arrested in G1. The kar1-Δ15 mutation deletes the portion of Kar1 that binds Spc72 and thus eliminates cytoplasmic microtubule nucleation from the half-bridge (Vallen et al., 1992 blue right-pointing triangle; Pereira et al., 1999 blue right-pointing triangle). In metaphase cells, when cytoplasmic microtubules normally nucleate from the outer plaque, we predicted that the kar1-Δ15 mutation should have little effect on cytoplasmic microtubule detachment, and this is what we observed for SHE1 and she1[increment] cells (Figure 2, C and D).

FIGURE 2:
(A) In wild-type (KAR1) G1 cells, Spc72 binds to Kar1 and microtubules are nucleated from the half-bridge. Graph shows cytoplasmic microtubule detachment rate in KAR1 (CUY2015) and KAR1 she1[increment] (CUY2016) G1 cells. (B) The kar1-[increment]15 mutation ...

In G1 cells, we observed about half the normal number of cytoplasmic microtubules in kar1-Δ15 and kar1-Δ15 she1[increment] cells (2.1 microtubules per wild-type cell and 1.0 microtubule per kar1-[increment]15 cell). These cytoplasmic microtubules presumably arise from the outer plaque where Spc72 binds to Nud1. At this stage of the cell cycle, when Spc72 binding to Kar1 is normally favored, binding to Nud1 is not optimal (Pereira et al., 1999 blue right-pointing triangle). In addition to reduced cytoplasmic microtubule numbers, we observed a significant number of cytoplasmic microtubules in kar1-Δ15 and kar1-Δ15 she1Δ cells that are not attached to the SPB (0.5 ± 0.1 and 0.6 ± 0.1 per cell, respectively). Although these might have detached from the SPB, it seems more likely they are nucleated by cytoplasmic aggregates of Spc72 that fail to bind to the SPB (Vallen et al., 1992 blue right-pointing triangle; Pereira et al., 1999 blue right-pointing triangle). The kar1-Δ15 mutation also increased the rate of cytoplasmic microtubule detachment threefold above that observed in wild-type cells. Although we do not know the mechanism of this increased rate of release, we refer to it as spontaneous release because it does not depend on dynein activity; introducing the nip100[increment] mutation does not lower cytoplasmic microtubule detachment in kar1-Δ15 cells (Supplemental Table S1).

Using the kar1-[increment]15 mutation, we could measure the effect of she1[increment] on microtubule detachment from both the half-bridge (she1[increment] cells) and the outer plaque (kar1-[increment]15 she1[increment] cells) at the same point in the cell cycle (G1). In G1 kar1-Δ15 she1Δ cells, the cytoplasmic microtubule detachment rate is nearly fourfold lower than in she1[increment] cells (Figure 2, A and B). This rate is similar to the spontaneous release rate observed in kar1-[increment]15 cells, indicating that she1[increment] has relatively little effect in kar1-[increment]15 cells. The nip100[increment] mutation does not lower microtubule detachment in kar-15[increment] she1[increment] cells, indicating that this residual release is not due to dynein activity (Supplemental Table S1). Thus microtubule attachment to the outer plaque, even under suboptimal conditions, is stronger than the normal microtubule attachment to the half-bridge.

Detached microtubules possess γ-tubulin and Spc72

We next set out to establish the place at which cytoplasmic microtubule detachment occurs. The first aim was to find whether cytoplasmic microtubules broke somewhere along the length of the polymer or whether they were pulled intact from the SPB. If the latter were true, it might be possible to observe anchoring proteins from the γ-tubulin complex or even the SPB on the minus ends of detached cytoplasmic microtubules. We initially imaged she1[increment] cells in G1 expressing mCherry-Tub1 and green fluorescent protein (GFP)–Tub4 (Figure 3A). Of the 72 cytoplasmic microtubules observed detaching from the SPB, 62 (86%) had visible GFP-Tub4 at the end detaching from the SPB. We also imaged cytoplasmic microtubule detachment in she1[increment] cells in G1 expressing mCherry-Tub1 and Spc72-GFP (Figure 3A). Spc72-GFP was observed on the ends of 21 of 26 (81%) detached cytoplasmic microtubules. On the other hand, we never observed Spc42 on the ends of detached cytoplasmic microtubules in cells expressing GFP-Tub1 and Spc42-mRFP. This result was expected, since Spc42 is in the central core of the SPB. In summary, these results indicate that detached cytoplasmic microtubules contain γ-tubulin and Spc72 at their minus ends, indicating that the break must occur somewhere on the SPB-proximal side of Spc72.

FIGURE 3:
Detached cytoplasmic microtubules contain γ-tubulin and Spc72. (A) Detached cytoplasmic microtubules in G1-arrested she1Δ cells expressing mCherry-Tub1 and GFP-Tub4 (left, CUY2028) or mCherry-Tub1 and Spc72-GFP (right, CUY2037). Arrowheads ...

We also measured the lifetime of 10 detached cytoplasmic microtubules in she1[increment] cells expressing mCherry-Tub1 and GFP-Tub4; 4 of these are plotted in Figure 3B. The GFP-Tub4 decoration remained on the minus ends of detached microtubules for 69 ± 38 s. During this time some microtubules elongated, some shortened, and some remained fairly constant in length. After loss of the GFP-Tub4 decoration, microtubules inevitably shortened and disappeared in 17 ± 7 s. Thus the minus ends of detached cytoplasmic microtubules appear to be stabilized initially against depolymerization by the presence of a cap of γ-tubulin and associated proteins. Loss of this cap results in rapid depolymerization of the microtubule.

We next examined whether the break point in G1 cells was between Spc72 and Kar1. Spc721-276–Kar1192-433 is a fusion protein that combines the γ-tubulin complex–binding portion of Spc72 and the half-bridge-binding region of Kar1 (Pereira et al., 1999 blue right-pointing triangle). This construct was expressed in a strain lacking the native Spc72 and Kar1 proteins, so that the only source of these proteins is the fusion protein (Figure 3C). Thus all cytoplasmic microtubules in these cells are anchored at the half-bridge through the Spc72–Kar1 fusion protein. Addition of the Spc72–Kar1 fusion protein lowers the rates of cytoplasmic microtubule detachment twofold in wild-type cells and 11-fold in she1[increment] cells (Figure 3D). Because the fusion protein substantially reduces cytoplasmic microtubule detachment, we conclude that the interaction between Spc72 and Kar1 is the linkage that is normally broken during this process in G1 cells.

We used a second fusion protein to examine whether the break point in metaphase cells was between Spc72 and Cnm67. Spc721-276–Cnm671-581 is a fusion protein that combines the Tub4-binding region of Spc72 and the outer plaque–binding portion of Cnm67 (Gruneberg et al., 2000 blue right-pointing triangle). This fusion protein bypasses the need for Nud1 that normally bridges these two proteins. Cells expressing the Spc72–Cnm67 fusion lack the native Spc72, so the only source of cytoplasmic microtubule anchoring is through the fusion protein located in the outer plaque (Figure 3C). The presence of the Spc72–Cnm67 fusion protein did not reduce the rate of cytoplasmic microtubule detachment; in fact detachment rates rose fivefold in wild-type cells and 1.3-fold in she1[increment] cells (Figure 3D). Thus we cannot conclude that it is the linkage between Spc72 and Cnm67 that is normally broken during cytoplasmic microtubule detachment in metaphase cells.

Of interest, the Spc72–Kar1 fusion did not reduce cytoplasmic microtubule detachment rates in metaphase-arrested cells; in fact these rates rose 17-fold in wild-type and twofold in she1Δ cells (Figure 3D). Thus, even the enhanced stability provided by this fusion protein at the half-bridge is still less than that provided by the normal outer plaque connection in metaphase cells. In addition, the Spc72–Cnm67 fusion reduced the rate of cytoplasmic microtubule detachment in G1 cells: twofold for wild-type cells and ninefold for she1[increment] cells (Figure 3D). This indicates that even the weakened outer plaque connection provided by this fusion protein is still stronger than the normal half-bridge connection. Both of these results support the conclusion of the preceding section that the connection of cytoplasmic microtubules to the outer plaque is stronger than that at the half-bridge.

Overexpression of She1 inhibits dynactin complex localization to microtubule ends

Our observation that excessive cytoplasmic microtubule detachment in she1[increment] cells depends on dynein activity is consistent with previous work suggesting that She1 negatively regulates dynein activity (Woodruff et al., 2009 blue right-pointing triangle). Thus we hypothesized that over­expression of She1 would produce a phenotype similar to that caused by inhibiting dynein activity—specifically, the elongation of anaphase spindles within the mother cell. To test this, we imaged cells expressing She1 from the highly efficient GAL1/10 promoter. We quantified the fraction of midanaphase spindles (between 3 and 6 μm) that were incorrectly oriented with both spindle poles in the mother cell. Cells overexpressing She1 had a level of spindle misorientation similar to that in dyn1[increment] and nip100[increment] cells (Figure 4A). Overexpressing She1 in dyn1[increment] or nip100[increment] cells did not increase their frequency of spindle misorientation, indicating that the She1 acts through the dynein pathway.

FIGURE 4:
Overexpression of She1 inhibits localization but not assembly of the dynactin complex. Cells contained a SHE1 construct expressed from the GAL1/10 promoter on a plasmid. SHE1 was overexpressed by shifting cells to galactose-containing medium for 4 h. ...

To investigate the mechanism of She1 inhibition of dynein activity, we overexpressed She1 in cells expressing GFP-tagged dynactin complex proteins and mCherry-Tub1 and scored cells for GFP signal at cytoplasmic microtubule plus ends and the SPB. For cells expressing Nip100-GFP, Arp1-GFP, and Jnm1-GFP, She1 overexpression decreased the number of cells with plus-end labeling (nine-, seven-, and eightfold, respectively) and increased the number of cells with labeling only at the SPB (seven-, 17-, and fourfold, respectively; Figure 4, B and C). The overall number of cells that displayed a GFP signal was relatively unchanged. Overexpression of She1 did not substantially affect the localization of Dyn1-GFP (Figure 4, B and C).

One possible mechanism by which She1 could inhibit dynactin loading onto microtubule plus ends is by interfering with dynactin assembly. To test this possibility, we overexpressed She1-GFP and assessed the interaction between dynactin complex proteins by coimmunoprecipitation. Binding between Arp10-3HA and Nip100-13Myc, Nip100-3HA and Ldb18-13Myc, Jnm1-3HA and Arp1-13Myc, and Jnm1-3HA and Nip100-13Myc was not disrupted in cells overexpressing She1-GFP (Figure 4D). Thus, She1 does not appear to interfere with dynactin assembly. In addition, She1-GFP was not found in any of the immunoprecipitates, indicating that overexpression of She1-GFP does not disturb dynactin localization through direct interaction with its subunits (Supplemental Figure S1).

She1 associates directly with microtubules

As previously reported, She1-GFP localizes to mitotic spindles, cytoplasmic microtubules, and a ring structure at the bud neck (Wong et al., 2007 blue right-pointing triangle; Woodruff et al., 2009 blue right-pointing triangle). We assessed She1's ability to associate with microtubules by incubating whole-cell extracts of a strain expressing She1-13Myc with preassembled, Taxol-stabilized bovine microtubules. The microtubules were then spun down and the amount of She1-13Myc in the pellet and supernatant compared. She1-13Myc bound to microtubules in a microtubule concentration–dependent manner (Figure 5A). The apparent dissociation constant, equal to the concentration of polymerized tubulin required to cosediment half of the She1-13Myc, is ~2 μM. This association between She1 and microtubules could be direct or could be mediated by another protein in the cell extract. To determine whether She1 can bind directly to microtubules, we purified GST-She1 from bacterial cells and incubated it with Taxol-stabilized microtubules. GST-She1 bound to microtubules with an apparent dissociation constant of ~1 μM (Figure 5B). Thus She1 has the ability to bind to microtubules directly and with an affinity similar to that of other microtubule-associated proteins (Blake-Hodek et al., 2010 blue right-pointing triangle).

FIGURE 5:
She1 associates with microtubules. (A) Lysate from cells expressing She1-13Myc (CUY1865) was incubated with various amounts of preassembled microtubules, the mixture centrifuged, and the supernatant (S) and pellet (P) fractions analyzed by Western blotting ...

DISCUSSION

We found that in cells lacking She1, cytoplasmic microtubules detach from the SPB at greatly elevated rates. This phenotype has also been noted by Markus et al. (2011) blue right-pointing triangle. Microtubule detachment requires dynein activity; simultaneous loss of the dynactin protein Nip100 completely eliminates the she1[increment] effect. This result agrees with previous reports indicating that She1 is a negative regulator of dynein function. This conclusion is also supported by our data showing that overexpression of She1 produces a defect in spindle orientation identical to that caused by loss of dynein or Nip100.

To determine how She1 inhibits dynein activity, we examined the localization of dynein and dynactin complex proteins in cells overexpressing She1. Whereas She1 overexpression has little effect on dynein localization, it greatly reduces the localization of dynactin proteins to cytoplasmic microtubule plus ends. This agrees with previous reports that loss of She1 allows dynactin components to localize to cytoplasmic microtubule plus ends at earlier points in the cell cycle (Woodruff et al., 2009 blue right-pointing triangle) and in greater number (Markus et al., 2011 blue right-pointing triangle). She1 overexpression does not affect the integrity of the dynactin complex, supporting the previous suggestion that She1 interferes with the interaction between dynein and dynactin (Woodruff et al., 2009 blue right-pointing triangle; Markus et al., 2011 blue right-pointing triangle). However, She1 did not coprecipitate with the dynactin complex, indicating that any interaction is transient. She1 was shown previously to associate with cytoplasmic microtubules during G1 and preanaphase, when dynein is inactive, but not during anaphase, when dynein is active (Woodruff et al., 2009 blue right-pointing triangle). Our results show that She1 can bind directly to microtubules, leading to the possibility that She1 inhibits dynein activity by competing with dynactin for access to microtubules.

Although dynein activity is down-regulated during G1, it is still the major cause of microtubule detachment during this portion of the cell cycle; loss of Nip100 causes a fourfold decrease in microtubule detachment in G1 cells. On the other hand, Myo2, the type V myosin protein that is responsible for directing cytoplasmic microtubules to the bud in the early stages of the cell cycle and might therefore be expected to play a role in microtubule detachment in G1 cells, has relatively little effect on microtubule detachment. Even if we attribute all of the microtubule detachment in nip100[increment] cells to Myo2 activity, it still amounts to only 25% of the rate in wild-type cells. In G1 cells lacking She1, which may contain maximal dynein activity, 95% of microtubule detachment can be attributed to dynein activity (as determined by comparing detachment in she1[increment] vs. she1[increment] nip100[increment] cells). These results likely reflect the fact that dynein pulls harder on cytoplasmic microtubules than Myo2; in vitro studies show that cytoplasmic dynein generates more than twice the force of myosin V (Gennerich et al., 2007 blue right-pointing triangle; Mehta et al., 1999 blue right-pointing triangle).

Although dynein pulls on the SPB-distal plus ends of cytoplasmic microtubules, the breakage event occurs at the SPB. Detached microtubules always appear to be full length and most contain γ-tubulin and Spc72 at their minus ends. Because Spc72 binds Spc97 and Spc98 but not Tub4 (Knop and Schiebel, 1998 blue right-pointing triangle), we assume that the entire γ-tubulin complex is present. Of interest, the γ-tubulin complex appears sufficient to stabilize the minus ends of detached microtubules, because these microtubules disappear rapidly after loss of this structure. In G1-arrested cells, Spc72 is primarily unphosphorylated and binds to Kar1 in the half-bridge. A Spc72–Kar1 protein fusion greatly reduces microtubule detachment in G1 cells, indicating that the Spc72–Kar1 interaction is the weak link at the half-bridge. In metaphase cells, Spc72 is phosphorylated and binds to Nud1, which in turn binds Cnm67, in the outer plaque. However, fusing Spc72 to Cnm67 did not reduce microtubule detachment from the outer plaque. This could be taken as evidence that the Spc72 attachment is not the weak link in the outer plaque. On the other hand, one could simply argue that the Spc72–Cnm67 fusion damages the outer plaque sufficiently to allow microtubule detachment by an aberrant mechanism.

Microtubule detachment rates are higher in G1 cells than in M-phase cells. Three lines of evidence indicate that this difference is due to a weaker microtubule attachment to the SPB half-bridge, rather than a stronger pulling force, in G1 cells. 1) We used the kar1-[increment]15 mutation to restrict microtubule nucleation to the outer plaque. This allowed us to measure the effect of she1[increment] on microtubule detachment from both the half-bridge (she1[increment] cells) and the outer plaque (kar1-[increment]15 she1[increment] cells) at the same point in the cell cycle (G1). The microtubule detachment rate from the half-bridge was fourfold higher than from the outer plaque. 2) We used the Spc72–Kar1 protein fusion to direct microtubule attachment to the half-bridge. This strengthens microtubule attachment to the half-bridge, but even this strengthened attachment is weaker than normal outer plaque attachment. Similarly, we used the Spc72-Cnm67 fusion protein to direct microtubule attachment to the outer plaque. This weakens microtubule attachment at the outer plaque, but even this weakened attachment is stronger than the normal half-bridge attachment. 3) If we assume that dynein is maximally active in she1[increment] cells, then the pulling force should be equivalent in G1 and M phase she1[increment] cells; however, microtubule detachment is eightfold more frequent in G1 cells. Thus we conclude that microtubule attachments to the outer plaque are stronger than those at the half-bridge.

It is not entirely clear why yeast nucleate cytoplasmic microtubules from two distinct locations on the SPB. Nucleation from both sites is not essential for vegetative growth; both the Spc72–Kar1 fusion, which confines nucleation to the half-bridge, and the Spc72–Cnm67 fusion, which confines nucleation to the outer plaque, allow cells to grow (Pereira et al., 1999 blue right-pointing triangle; Gruneberg et al., 2000 blue right-pointing triangle). However, microtubules coming from the half-bridge are essential for karyogamy (Conde and Fink, 1976 blue right-pointing triangle; Pereira et al., 1999 blue right-pointing triangle) and, because mating occurs only between cells arrested in G1 by mating factors, it follows that the half-bridge would be the predominant site of microtubule nucleation in cycling G1 cells. The switch to the outer plaque as cells enter mitosis may be made in part to withstand the pulling forces exerted by dynein, which is most active in the later portions of the cell cycle.

MATERIALS AND METHODS

Yeast strains and plasmids

Yeast strains used in this study are S288C or derivatives and are listed in Supplemental Table S2. Plasmids used are listed in Supplemental Table S3.

MATa cells were arrested in G1 by adding 3 μg/ml α-factor to the medium for 3 h. Cells containing PMET3-CDC20-3HA were arrested in metaphase by adding 20 μg/ml methionine to the medium for 3 h. She1 was overexpressed in cells containing PGAL1/10-SHE1, PGAL1/10-SHE1-GFP, or PGAL1/10-SHE1-13MYC by growing cultures to log phase in 2% raffinose medium for several generations and then shifting them to 2% galactose medium for 4–6 h.

Microscopy and image analysis

Images were obtained using a spinning disk confocal system as previously described (Huang and Huffaker, 2006 blue right-pointing triangle). All images are maximum-intensity projections of z-series stacks ([increment]z = 0.7 μm). Analysis was performed using ImageJ (National Institutes of Health, Bethesda, MD). The mean values and standard deviations of microtubule detachment were determined using the Poisson approximation to the binomial distribution.

Microtubule cosedimentation assays

For assays using whole-cell extracts, log-phase yeast cells expressing She1-13Myc (CUY1865) were centrifuged for 5 min at 2000 rpm, washed with lysis buffer (80 mM 1,4-piperazinediethanesulfonic acid [PIPES], 1 mM ethylene glycol tetraacetic acid [EGTA], 1 mM MgSO4, 5% glycerol, 100 mM KCl, 0.25% Brij-35, 2 mM dithiothreitol [DTT], pH 6.8), and resuspended in lysis buffer containing 20 μM Taxol, 1 mM phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, and 10 μg/ml pepstatin. Cells were lysed in a bead beater, and lysates were precleared by centrifugation at 350,000 × g for 5 min at 4°C. Tubulin (Cytoskeleton, Denver, CO) was diluted to 1 mg/ml in PEM-DGT (100 mM PIPES, 2 mM EGTA, 1 mM MgCl2, 20 μM Taxol, 1 mM GTP, and 4 mM DTT) and precleared by centrifugation at 150,000 × g for 5 min at 4°C. The supernatant was incubated at 37°C for 20 min to allow microtubule polymerization, followed by centrifugation at 35,000 × g for 20 min at room temperature. The microtubule pellet was resuspended in PEM-DGT to a final concentration of 5 mg/ml. Various amounts of microtubules were added to 20 μl of precleared yeast cell lysate and incubated for 20 min on ice. Samples were centrifuged at 175,000 × g for 10 min at 4°C. Supernatant and pellet fractions were analyzed by SDS–PAGE and Western blotting using 9E10 anti-Myc (Covance, Princeton, NJ) and DM1α anti-tubulin (Sigma-Aldrich, St. Louis, MO) antibodies.

For assays using purified protein, She1–glutathione S-transferase (GST) was expressed in BL21 Escherichia coli cells containing plasmid pXX6. Cells were grown at 26°C in 1 l of Luria–Bertani broth containing 50 μg/ml kanamycin and then induced by adding 50 μM isopropyl-β-d-thiogalactoside for 4 h. Cells were then spun down, washed with cold phosphate-buffered saline (PBS), and resuspended in 8.5 ml of cold PBS containing EDTA-free protease inhibitors (Roche, Basel, Switzerland). Cells were lysed by the addition of 10 μl of 100 mg/ml lysozyme for 30 min, followed by sonication. Triton X-100 was added to 1% and the mixture rocked at 4°C for 20 min. Cell debris was pelleted at 12,000 × g for 25 min at 4°C. The supernatant was incubated with glutathione–Sepharose beads for 2.5 h at 4°C. Beads were then pelleted, washed four times with cold PBS, resuspended in 350 μl of 80 mM reduced glutathione in PBS (pH 7), and rocked for 16 h at 4°C. Purified GST-She1 was then obtained by adding the mixture to a minichromatography column (Bio-Rad, Hercules, CA) and spinning at 5000 × g for 2 min. GST-She1 was incubated with assembled microtubules at room temperature for 5 min and then spun at 175,000 × g for 40 min. Supernatant and pellet fractions were analyzed by SDS–PAGE and Western blotting using anti-GST antibodies.

Coimmunoprecipitations

Coimmunoprecipitations were performed as described previously (Wolyniak et al., 2006 blue right-pointing triangle), except that anti-hemagglutinin (HA) affinity gel or anti-Myc affinity gel (Sigma-Aldrich) was used to precipitate epitope-tagged proteins from cell lysates. Western blots were probed with either 9E10 anti-Myc or HA.11 anti-HA antibodies (Covance).

Supplementary Material

Supplemental Materials:

Acknowledgments

We thank Mark Rose (Princeton University, Princeton, NJ), Elmar Schiebel (Zentrum für Molekulare Biologie der Universität Heidelberg, Heidelberg, Germany), and Dean Dawson (Oklahoma Medical Research Foundation, Oklahoma City, OK) for yeast strains and plasmids. This work was supported by National Institutes of Health Grant GM40479 (to T.C.H.). I.A.A. was funded by National Institutes of Health Predoctoral Grant GM073576.

Abbreviations used:

GFP
green fluorescent protein
MTOC
microtubule-organizing center
SPB
spindle pole body

Footnotes

This article was published online ahead of print in MBoC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E12-03-0223) on April 25, 2012.

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