A single-copy transgene insertion system to investigate KNL-1 microtubule-binding mutants

Prior work in the early C. elegans embryo established high-resolution imaging assays to monitor chromosome alignment and segregation, formation of load-bearing kinetochore–microtubule attachments, and spindle checkpoint signaling (Oegema et al., 2001; Desai et al., 2003; Cheeseman et al., 2004; Essex et al., 2009). Capitalizing on these assays, we analyzed KNL-1 mutants defective for microtubule binding. We used Mos single-copy insertion (Frøkjaer-Jensen et al., 2008) to integrate WT or mutant KNL-1^RR::mCherry transgenes (RR denoting RNAi resistant) at a defined genomic location (Fig. 1 F). Transgenes were engineered to be RNAi resistant by altering the nucleotide sequence of exon 4 without affecting coding information; injection of a double-stranded RNA (dsRNA) homologous to exon 4 enabled penetrant and selective depletion of endogenous KNL-1 (Fig. 1 G). KNL-1 localizes to kinetochores, and its depletion leads to 100% embryonic lethality and a first-division kinetochore-null phenotype, characterized by clustering of chromosomes from the two pronuclei, failure of chromosome segregation, and premature spindle pole separation (Desai et al., 2003). The transgene-encoded WT KNL-1^RR::mCherry localized similarly to endogenous KNL-1 and rescued the kinetochore-null phenotype and embryonic lethality caused by endogenous KNL-1 depletion (Fig. 1 [H–K] and Video 1). WT KNL-1^RR::mCherry also rescued lethality of the knl-1(ok3457) deletion mutant (Fig. S1, B and C). Thus, single-copy KNL-1^RR::mCherry transgene insertion provides a means to precisely compare engineered KNL-1 mutants after depletion of endogenous KNL-1.

KNL-1 microtubule-binding mutants support formation of normal load-bearing kinetochore–microtubule attachments

To determine the in vivo function of KNL-1 microtubule-binding activity, we generated strains with 4A or Δ9 mutant KNL-1^RR::mCherry transgenes integrated at the same genomic location as the WT transgene. Both mutant KNL-1 fusions localized to kinetochores at levels similar to the WT fusion (Fig. 2 A). To compare the ability of the WT and mutant transgenes to support chromosome alignment and segregation, we first crossed them into a strain expressing GFP fusions with histone H2b and γ-tubulin and then depleted endogenous KNL-1 and monitored the dynamics of chromosomes and spindle poles throughout the first division. Quantitative analysis of spindle pole separation serves as a readout for the formation of load-bearing kinetochore–microtubule attachments, as interactions between astral microtubules and the cortex generate pulling forces that are resisted by load-bearing kinetochore–microtubule attachments within the spindle (Fig. 2 B; Oegema et al., 2001; Desai et al., 2003).

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Figure 2.

Microtubule-binding mutants of KNL-1 do not affect formation of load-bearing attachments or chromosome segregation. (A) An image of a metaphase plate in living one-cell embryos from strains harboring the indicated KNL-1^RR::mCherry transgenes and depleted of endogenous KNL-1. Bar, 2 µm. (B) A schematic of analysis performed after crossing in GFP::histone H2b and GFP::γ-tubulin and depleting endogenous KNL-1. (C) Frames from time-lapse sequences of the first embryonic division for the indicated KNL-1 mutants. Bar, 3 µm. (D) Spindle pole separation kinetics for the indicated conditions. Error bars represent the SEM with a 95% confidence interval. The no transgene knl-1(RNAi) trace is reproduced from Fig. 1 K. (E) Timing of anaphase onset in the indicated conditions. The first visible sign of sister chromatid separation (based on the GFP::H2b signal) was scored as anaphase onset. The gray dashed line indicates the NEBD–anaphase interval for the WT transgene (reproduced from the inset in Fig. 1 K). Error bars represent the SEM with a 95% confidence interval. For comprehensive statistical analysis, see Table S1. (F) Embryonic viability analysis of KNL-1 microtubule-binding mutants. L4-stage worms were injected with dsRNA-targeting endogenous knl-1, and the embryos laid by the injected worms were collected 21–41 h after injection and scored for hatching to form larvae.

After endogenous KNL-1 depletion in the 4A and Δ9 microtubule-binding mutant strains, no significant defects in chromosome segregation were observed. In addition, the kinetics of spindle pole separation were normal until metaphase, but anaphase onset was delayed by 50 s (Fig. 2 [C–E] and Video 2). These observations suggested that the microtubule-binding activity of KNL-1 does not make a major contribution to the mechanics of chromosome segregation. Consistent with this, the 4A and Δ9 mutants rescued embryo viability after depletion of endogenous KNL-1 (Fig. 2 F), and both mutants rescued the lethality of the knl-1(ok3457) deletion allele (Fig. S1, B and C).

Mild defects in chromosome–microtubule attachment can be masked by the spindle checkpoint, which delays cell cycle progression until proper attachments are established. For both the 4A and Δ9 mutants, we observed a significant (P < 0.0001; Table S1) 50-s delay in anaphase onset (Fig. 2 E) that correlated with a plateau in the spindle pole separation profile at metaphase spindle length (Fig. 2 D, top graph). Therefore, we codepleted the checkpoint protein Mad2^MDF-2 together with endogenous KNL-1 to determine whether a mild attachment defect resulting from the 4A and Δ9 mutations was being masked by checkpoint activity. Although Mad2^MDF-2 depletion eliminated the 50-s delay, it did not lead to visible segregation defects or enhance embryonic lethality (note that in C. elegans, the one-cell embryo undergoes ~550 individual cell divisions before hatching, and chromosome segregation defects are associated with 100% penetrant embryonic lethality; Fig. 2, C–F). Cumulatively, the lack of a significant pole separation defect in the 4A and Δ9 mutants, the ability of these mutants to rescue a deletion allele of KNL-1, and their lack of phenotypic enhancement after Mad2^MDF-2 codepletion suggest that the microtubule-binding activity of KNL-1 does not play a significant role in the formation of load-bearing attachments during chromosome segregation.

KNL-1 microtubule-binding mutants extend the spindle checkpoint–dependent cell cycle delay induced by monopolar spindles

The spindle checkpoint does not act as a timer controlling mitotic duration in C. elegans embryos and is potentially only very mildly activated during the rapid embryonic divisions, as detectable kinetochore enrichment of checkpoint signaling proteins such as Mad1^MDF-1 and Mad2^MDF-2 is not observed during unperturbed mitosis (Essex et al., 2009). Therefore, we hypothesized that the anaphase onset delay observed with KNL-1 microtubule-binding mutants in the first cell division was a result of a compromised ability to silence a mild checkpoint signal. We previously developed controlled formation of monopolar spindles as a means to monitor checkpoint activation in the early embryo (Fig. 3 A; Essex et al., 2009); this assay circumvents the difficulty of treating normally impermeable embryos with microtubule-depolymerizing drugs. Second-division embryos with monopolar spindles exhibit a significant mitotic delay that requires the checkpoint pathway and the KMN network and correlates with visible transient enrichment of Mad2^MDF-2 on unattached kinetochores (Essex et al., 2009).

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Figure 3.

Microtubule-binding mutants of KNL-1 significantly extend the spindle checkpoint–mediated cell cycle delay induced by monopolar spindles. (A) A schematic of the monopolar spindle–based checkpoint signaling assay in C. elegans embryos. Depletion of the centrosome duplication kinase ZYG-1 generates monopolar spindles in the second division, which trigger a spindle checkpoint–dependent cell cycle delay. (B) Mean time from NEBD to chromosome decondensation in the AB cell for the indicated conditions. Error bars represent the SEM with a 95% confidence interval. The gray dashed line marks the duration of AB cell mitosis for the WT transgene, and the red dashed line marks the duration of AB cell mitosis induced by monopolar spindles in the same strain. For comprehensive statistical analysis, see Table S2. (C) Stills from time-lapse sequences of the AB cell monopolar division in worm strains coexpressing GFP::Mad2^MDF-2 and the indicated transgenes. Mad2^MDF-2 accumulation on unattached kinetochores is marked with open arrowheads. Bar, 3 µm.

To determine whether the 4A and Δ9 mutations affected checkpoint signaling, especially whether they impaired checkpoint silencing, we generated monopolar spindles and measured the interval from nuclear envelope breakdown (NEBD) to chromosome decondensation. A GFP fusion of cyclin B has been generated in C. elegans, but the fluorescent signal is lost at anaphase of oocyte meiosis I and is not subsequently regained (Liu et al., 2004), likely because of insufficient time to mature GFP fluorescence. It is for this reason that we use decondensation or onset of cortical contractility, coupled with Mad2^MDF-2 inhibition, to measure checkpoint-dependent cell cycle delays.

The transgene-encoded WT KNL-1::mCherry fusion supported normal checkpoint signaling (Fig. 3 B); the NEBD-to-decondensation interval was longer in the presence of monopolar spindles than in control cells with bipolar spindles, and this delay was abrogated by Mad2^MDF-2 depletion. In cells expressing the microtubule-binding 4A and Δ9 mutants, the delay induced by monopolar spindles was significantly longer compared with the delay observed with WT KNL-1 (P < 0.0001; Fig. 3 B). This extended delay is a result of persistent checkpoint activation, as codepletion of Mad2^MDF-2 caused the cell cycle timing to match that of control embryos with bipolar spindles (Fig. 3 B). Thus, perturbing KNL-1 microtubule binding extends the time that cells with monopolar spindles spend in a checkpoint-active state.

Confocal imaging of GFP::Mad2^MDF-2 on chromosomes on the monopolar spindles revealed that the extended delay induced by the 4A mutation correlated with a significantly extended residence time for GFP::Mad2^MDF-2 on kinetochores (WT residence time was 69 ± 13 s for 13 embryos, and 4A residence time was 118 ± 11 s for 11 embryos; P < 0.0001; Fig. 3 C). The extended persistence of Mad2^MDF-2 on the kinetochores of monopolar spindles suggests that the checkpoint silencing defect observed in KNL-1 microtubule-binding mutants is a result of a kinetochore-localized reaction rather than an effect on cytoplasmic regulation of the checkpoint signal. This analysis of cells with monopolar spindles collectively with the results in one-cell embryos with bipolar spindles—where KNL-1 microtubule-binding mutants do not affect load-bearing attachment formation but nonetheless exhibit checkpoint-dependent extended mitotic duration—suggests that KNL-1 microtubule-binding activity contributes to checkpoint silencing at kinetochores.

KNL-1 microtubule-binding mutants do not extend the cell cycle in the absence of microtubules

In cells with bipolar spindles, checkpoint silencing occurs after formation of kinetochore–microtubule attachments on both sister chromatids. In cells with monopolar spindles, the mechanism that controls the exit from the checkpoint-activated state is not clear; the involvement of KNL-1 microtubule binding in this mechanism led us to hypothesize that noncentrosomal microtubules that assemble in the vicinity of chromosomes may interact with kinetochores facing away from the single pole, and this in turn promotes exit from the checkpoint-active state. This hypothesis makes two predictions. First, the checkpoint delay induced by microtubule depolymerization should be longer than that induced by monopolar spindles, as there would be no microtubules present to promote mitotic exit. Second, the checkpoint delay induced by microtubule depolymerization should be the same for WT and microtubule-binding mutants of KNL-1, as there would be no microtubules present to allow WT KNL-1 microtubule-binding activity to accelerate checkpoint silencing relative to the microtubule-binding mutants.

To examine checkpoint signaling in the absence of microtubules, we used a recently developed method to add drugs in a timed manner to C. elegans embryos (Carvalho et al., 2011). In this approach, inhibition of the perm-1 gene, which is involved in generating the eggshell permeability barrier, is used to permeabilize the eggshell to small molecules (Fig. 4 A). The fragile permeabilized embryos are placed in wells of a microchamber, enabling nocodazole addition while filming. To enable comparison with the monopolar spindle assay, nocodazole addition was performed 1–2 min before NEBD in the AB cell of the two-cell–stage embryo. As nocodazole treatment causes chromosomes to collapse together, decondensation cannot be used to monitor mitotic duration. Therefore, we quantified the interval between NEBD and the onset of cortical contractility, an alternative hallmark of Cdk1 inactivation (Essex et al., 2009).

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Figure 4.

KNL-1 microtubule-binding mutants do not extend the cell cycle in the absence of microtubules. (A) A schematic of the experimental approach used to analyze mitotic duration in the AB cell after microtubule (MT) depolymerization. (B) Mean time from NEBD to the onset of cortical contractility for the indicated conditions. Error bars represent the SEM with a 95% confidence interval.

Treatment of embryos with nocodazole resulted in a longer cell cycle delay compared with that induced by the formation of monopolar spindles, suggesting that microtubules do promote exit from the checkpoint-active state induced by monopolar spindles (Fig. 4 B). Next, we compared mitotic duration in WT and 4A mutant KNL-1 in the absence of microtubules and found that there was no significant difference between them (Fig. 4 B). We performed this experiment in a strain expressing GFP:Mad2^MDF-2 to correlate the delay in mitotic exit with GFP:Mad2^MDF-2 localization; however, chromosome compaction after nocodazole addition and diffuse Mad2^MDF-2 signal in the nuclear environment prevented us from assessing Mad2^MDF-2 kinetochore localization.

Thus, inhibiting KNL-1 microtubule-binding activity extends the checkpoint-active state in cells with bipolar and monopolar spindles but not in cells lacking microtubules. This result suggests that the effect of mutating KNL-1–dependent microtubule-binding activity on checkpoint silencing is a microtubule-dependent reaction in vivo.

KNL-1 microtubule-binding mutants do not affect PP1 docking and vice versa

The basic patch implicated in microtubule binding is adjacent to the conserved KNL-1 PP1-docking motifs (Fig. 5 A). As PP1 and the KNL-1–PP1 interaction have been implicated in checkpoint silencing (Pinsky et al., 2009; Vanoosthuyse and Hardwick, 2009; Meadows et al., 2011; Rosenberg et al., 2011) and the microtubule-binding mutants of KNL-1 that we engineered affected checkpoint silencing, we tested whether the mutations that disrupted microtubule binding also perturbed PP1 docking. For this purpose, we established biochemical and yeast two-hybrid–based assays for the KNL-1–PP1 interaction. C. elegans has four PP1 catalytic subunits: GSP-1, -2, -3, and -4 (Fig. S3 A). GSP-1 and -2 are partially redundant and essential for viability (Hsu et al., 2000), whereas GSP-3 and -4 are expressed during spermatogenesis (Chu et al., 2006). KNL-1 interacts with GSP-1 and -2 but not GSP-3 or -4 (Fig. 5 B); biochemical analysis confirmed that the interactions are direct (Fig. 5 E). Consistent with prior work, mutation of RRVSF to RRASA abolished binding in both assays (Fig. 5, C–E; Liu et al., 2010). Mutation of SILK to SAAA had no detectable effect in the biochemical assay, but a defect was observed at high stringency in the two-hybrid assay (Fig. S3 C). Notably, the 4A and Δ9 mutants that perturb microtubule binding/bundling did not affect PP1 docking (Fig. 5, D and E). Conversely, the RRASA mutant that abolished PP1 docking did not affect microtubule binding or bundling (Fig. 5 F; Liu et al., 2010). These results demonstrate that, despite their close proximity in the KNL-1 primary sequence, microtubule binding and PP1 docking are separable activities.

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Figure 5.

Microtubule-binding mutants of KNL-1 do not affect the interaction with PP1 and vice versa. (A) The conserved PP1 (PP1c)-docking motif SILK-RRVSF is highlighted, and mutations generated in the docking motif are indicated. (B) KNL-1 interacts with the C. elegans PP1 homologs GSP-1 and -2 but not GSP-3 and -4 in yeast two-hybrid assays (see also Fig. S3, A and B). (C) KNL-1 interaction with GSP-1/2 requires the RRVSF motif (see also Figs. 6 D and S3 C). (D) The 4A and Δ9 mutants do not perturb interaction of KNL-1 with GSP-1 and -2 in yeast two-hybrid analysis. KBP-5, a KNL-1–binding protein that interacts with the C-terminal half of KNL-1, serves as a positive control. (E) Biochemical analysis of KNL-1 interaction with GSP-2. Indicated variants of KNL-1^1–505–6xHis immobilized on nickel agarose beads were incubated with GST–GSP-2, and bead-bound proteins were analyzed by SDS-PAGE. Molecular mass is indicated in kilodaltons. (F) PP1-docking mutant of KNL-1 interacts normally with microtubules. Assays using purified KNL-1^1–505 variants were performed as described in Fig. 1 (C and E). The WT data are reproduced from Fig. 1 E. Error bars represent the SEM with a 95% confidence interval. Bar, 10 µm.

PP1-docking mutants of KNL-1 delay the formation of load-bearing attachments and extend mitotic duration in cells with monopolar spindles

Next, we analyzed the in vivo phenotypes of a KNL-1 RRASA mutant, an SAAA mutant, and an SAAA;RRASA double mutant. All of these mutants localized normally to kinetochores in the absence of endogenous KNL-1 (Fig. 6 A). Three phenotypes were evident after endogenous KNL-1 depletion in the RRASA mutant: a kinetic delay in the formation of load-bearing kinetochore–microtubule attachments (Fig. 6, B and C), a delay in anaphase onset (Fig. 6 [B, C, and E] and Video 3), and partial embryonic lethality that decreased with time after dsRNA injection (Figs. 6 G and S2). The delay in load-bearing attachment formation is inferred from the pronounced bump in the pole-tracking analysis (Fig. 6 B); the front half of the bump represents the interval during which kinetochores provide no resistance to spindle pole separation induced by cortical pulling forces. Despite the delay in attachment formation, lagging anaphase chromosomes were not observed at appreciable frequency in the first division (unpublished data). The SAAA mutant, consistent with its mild effect on PP1 binding relative to the RRASA mutant, showed significantly milder phenotypes in vivo (Fig. 6, B, D, and E). Importantly, an SAAA;RRASA double mutant was quantitatively identical to the RRASA mutant in the pole-tracking assay (Fig. 6 B and Video 3) and in the extent to which it elongated the NEBD–anaphase onset interval in one-cell embryos (Fig. 6, B and E). Thus, the RRASA mutant alone behaves as a null for KNL-1–PP1 docking in vivo. In contrast to the RRASA and SAAA;RRASA mutants, depletion of the two catalytic PP1 subunits (GSP-1 and -2) resulted in highly pleiotropic defects in the early embryo (Fig. S4), likely reflecting multiple PP1 functions that are executed in association with different docking subunits.

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Figure 6.

PP1-docking mutants of KNL-1 exhibit kinetic defects in load-bearing attachment formation and are synthetically lethal with checkpoint inhibition. (A) An image of a metaphase plate in living one-cell embryos from strains harboring the indicated KNL-1^RR::mCherry transgenes and depleted of endogenous KNL-1; the control image is reproduced from Fig. 2 A. Bar, 2 µm. (B) Spindle pole separation kinetics for the indicated conditions. Error bars represent the SEM with a 95% confidence interval. The WT trace is reproduced from Fig. 2 D. (C) Frames from time-lapse sequences of the first embryonic division for the indicated KNL-1 mutants. Bar, 3 µm. (D) Biochemical analysis of KNL-1 interaction with GSP-2 performed as in Fig. 5 E. Molecular mass is indicated in kilodaltons. (E) NEBD–anaphase onset interval in one-cell stage embryos for the indicated conditions. See Table S1 for statistical analysis. (F) Spindle pole separation kinetics for the indicated conditions. Error bars represent the SEM with a 95% confidence interval. The WT trace is reproduced from Fig. 2 D. (G) Embryonic viability analysis for the indicated conditions. Lethality was measured during two intervals (21–41 and 42–64 h) after endogenous KNL-1 depletion. In the earlier time point, embryos are depleted of maternal load but potentially inherit some dsRNA that affects zygotic KNL-1 expression; at the later time point, the maternal load is depleted, but zygotic expression is likely unaffected (see the legend of Fig. S2). (H) NEBD–chromosome decondensation interval in AB cells with bipolar or monopolar spindles for the indicated conditions. The red dashed line marks the duration of AB cell mitosis induced by monopolar spindles in the same strain. See Table S2 for statistical analysis. (E and H) The gray dashed lines mark the duration of AB cell mitosis for the WT transgene. Error bars represent the SEM with a 95% confidence interval.

Codepletion of Mad2^MDF-2 reduced the significant anaphase onset delay in the RRASA mutant (as well as the SAAA;RRASA mutant) but did not restore anaphase timing to that in controls (Fig. 6 [E and F] and Table S1). The reason why perturbing PP1 docking on the KNL-1 N terminus extends the interval between NEBD and anaphase onset independently of checkpoint signaling is unclear.

Importantly, Mad2^MDF-2 codepletion severely enhanced the lethality of the RRASA mutant, indicating that checkpoint activation protects against the defect in load-bearing attachment formation created by perturbing PP1 docking on KNL-1 (Fig. 6 G). Thus, unlike the 4A and Δ9 microtubule-binding mutants, where attachment defects are not observed and checkpoint inhibition does not enhance lethality, the RRASA mutant exhibits enhanced lethality after checkpoint inactivation.

Next, we performed the monopolar spindle checkpoint assay for the RRASA mutant. The RRASA mutant exhibited a significantly extended cell cycle delay (Fig. 6 H) as well as increased residence of GFP::Mad2^MDF-2 at unattached kinetochores (Fig. S5). The extended delay was largely abrogated by Mad2^MDF-2 codepletion, indicating that it is a result of persistent checkpoint activation. As prior work in yeast has suggested a direct role for the KNL-1–PP1 interaction in checkpoint silencing (Meadows et al., 2011; Rosenberg et al., 2011), this observation is most consistent with the view that KNL-1–PP1 docking, in addition to affecting the kinetics of load-bearing attachment formation, participates in checkpoint silencing.

The second conserved microtubule-binding activity of the KMN network is dispensable for load-bearing microtubule attachment formation

KNL-1 depletion in C. elegans or mutation of the KNL-1 ortholog Spc105 in budding yeast severely inhibits the formation of load-bearing attachments (Desai et al., 2003; Pagliuca et al., 2009; Akiyoshi et al., 2010). In C. elegans, this is a result of an essential role for KNL-1 in outer kinetochore assembly (Desai et al., 2003). The initial hypothesis for the function of KNL-1 microtubule binding was that it synergized with the Ndc80 complex to generate a dynamic yet stable kinetochore–microtubule interface (Cheeseman et al., 2006). The data presented here indicate that this is not the case, as mutants that abrogate KNL-1 microtubule binding in vitro exhibit no significant defects in chromosome segregation or load-bearing attachment formation and are not enhanced by inhibiting checkpoint signaling. These results were in contrast to perturbing PP1 docking on KNL-1, which caused kinetic defects in load-bearing attachment formation and strong synergistic lethality with checkpoint inhibition. Thus, we conclude that microtubule binding by C. elegans KNL-1 does not make an essential contribution to the ability of kinetochores to form load-bearing attachments.

KNL-1 microtubule binding functions in checkpoint silencing at the kinetochore in the presence of microtubules

Although perturbing KNL-1 microtubule binding did not affect chromosome segregation, it led to a significantly increased monopolar spindle–induced cell cycle delay and an extended NEBD–anaphase duration in the presence of bipolar spindles. The full magnitude of both delays was a result of checkpoint signaling, and, on monopolar spindles, the delay was correlated with increased persistence of Mad2^MDF-2 at kinetochores. In light of the lack of detectable attachment or segregation defects, these observations suggested that KNL-1 microtubule binding participates in silencing the spindle checkpoint signal. The extended checkpoint signaling observed in KNL-1 microtubule-binding mutants is not a general feature of perturbing kinetochore–microtubule interactions, as mutations affecting Ndc80 complex microtubule binding do not extend the duration of the monopolar spindle–induced cell cycle delay (unpublished data). Notably, KNL-1 microtubule-binding mutants only exhibited an increased checkpoint-dependent cell cycle delay relative to WT KNL-1 in the presence of microtubules, suggesting that KNL-1 microtubule-binding activity participates in a microtubule-dependent manner in checkpoint silencing at the kinetochore. We propose that KNL-1 microtubule binding senses the presence of microtubules attached to the kinetochore, potentially via the closely associated Ndc80 complex, and relays their presence to shut off generation of the checkpoint signal (Fig. 7 F).

KNL-1 microtubule binding: A dynein-independent means for sensing presence of microtubules at kinetochores?

Work in Drosophila melanogaster and vertebrate cells has indicated that kinetochore-localized dynein motor contributes to checkpoint silencing by removing checkpoint proteins from kinetochores after microtubule attachment (Howell et al., 2001; Wojcik et al., 2001). However, many species, including fungi and higher plants, silence the checkpoint after microtubule attachment without the involvement of dynein. Recent work in human cells suggests that even in systems with dynein involvement, the checkpoint can be silenced via a dynein-independent pathway (Chan et al., 2009; Barisic et al., 2010; Gassmann et al., 2010). Microtubule-binding KNL-1 mutants do not affect dynein recruitment to unattached kinetochores, suggesting that the effect we observe is not a result of a loss of kinetochore dynein (unpublished data). As the microtubule-binding and PP1-docking mutants of KNL-1 have an additive effect on checkpoint silencing and prior work has implicated PP1 in dynein regulation at the kinetochore (Whyte et al., 2008), we speculate that KNL-1 microtubule binding may affect checkpoint silencing through a dynein-independent mechanism. As KNL-1 family proteins are present throughout eukaryotic evolution, including in species that do not have kinetochore dynein, KNL-1 microtubule-binding–based silencing may represent an evolutionarily ancient mechanism that has been enhanced/superseded by the dynein-based mechanism in animal cells. Testing this idea will require elucidating whether KNL-1 microtubule binding functions in parallel to or in the same pathway as dynein in checkpoint silencing, mapping and mutating KNL-1 microtubule-binding activity in yeast/higher plants that lack kinetochore dynein or dynein altogether, and analyzing mammalian Knl1 microtubule-binding mutants under conditions in which kinetochore dynein has been largely removed (Gassmann et al., 2010).

Protein purification and pull-down assays

KNL-1^1–505 and GSP-2 ORFs were amplified from N2 cDNA and cloned into pET21A and pGEX6P-1, respectively. Expression in Rosetta (DE3) pLysS Escherichia coli was induced with 0.3 mM IPTG for 4 h at 20°C, and 6xHis and GST purifications were performed using standard procedures. For KNL-1^1–505, imidazole-eluted protein was exchanged into SP buffer (30 mM MOPS, pH 7.0, 50 mM NaCl, 0.5 mM EDTA, and 2 mM β-mercaptoethanol), bound to HiTrap SP Sepharose (GE Healthcare), eluted with a gradient from 50 to 500 mM NaCl, and dialyzed to reduce the salt concentration to 150 mM NaCl or exchanged into BRB80 (80 mM Pipes, pH 6.8, 1 mM EGTA, and 1 mM MgCl₂) + 80 mM KCl. For GSP-2, glutathione-eluted protein was exchanged into 30 mM Hepes, pH 7.0, 50 mM NaCl, 0.1 mM EDTA, and 2 mM β-mercaptoethanol, bound to HiTrap SP Sepharose, and eluted with a gradient from 50 to 500 mM NaCl followed by dialysis to 100 mM NaCl.

For pull-down assays, 3 µg KNL-1^1–505 was immobilized on 20 µl Ni–nitrilotriacetic acid agarose resin (QIAGEN) for 1 h at 4°C. The resin was then washed three times (wash buffer: 50 mM Hepes, pH 7.5, 100 mM NaCl, 2 mM MnCl₂, 0.1 mM EDTA, 0.1% Tween 20, 30 mM imidazole, 10% glycerol, and 5 mM β-mercaptoethanol) and incubated for 1 h at room temperature with 12 µg of purified GST–GSP-2 protein in a 55-µl volume. The resin was then washed three times, eluted with sample buffer, and analyzed by SDS-PAGE.

Microtubule-bundling and bead assays

For microtubule-bundling assays, 1 µM taxol-stabilized rhodamine microtubules was mixed with 0.5 µM KNL-1^1–505 variants or control buffer. After 5 min, the mixture was imaged using a 60× 1.4 NA Plan Apochromat objective.

For the bead microtubule–binding assay, 3 µg of KNL-1^1–505 variants was incubated with 10 µl of 12-pM polystyrene beads (SVP-05-10; Spherotech, Inc.) coated with anti-6xHis antibody (no. 34440; QIAGEN) for 45 min at 4°C. After sonication in ice water for 2 min, the beads were flowed through a microscope flow chamber coated with taxol-stabilized rhodamine microtubules, 20 fields of view were imaged at 60× magnification, and images were analyzed to measure the mean number of beads bound per field.

We are indebted to Christian Frøkjær-Jensen and Erik Jorgensen (University of Utah, Salt Lake City, UT) for their development of Mos single-copy insertion and their generosity with reagents and expertise. We thank the C. elegans Gene Knockout Consortium for the knl-1(ok3457) mutant, Andy Powers and Chip Asbury for help with the bead assay, Sue Biggins and members of the Desai and Oegema laboratories for useful discussions, and Becky Green for comments on the manuscript.

This work was supported by a grant from the National Institutes of Health to A. Desai (GM074215). K. Oegema and A. Desai receive salary support from the Ludwig Institute for Cancer Research.