NuMA recruits dynein activity to microtubule minus-ends at mitosis.

To build the spindle at mitosis, motors exert spatially regulated forces on microtubules. We know that dynein pulls on mammalian spindle microtubule minus-ends, and this localized activity at ends is predicted to allow dynein to cluster microtubules into poles. How dynein becomes enriched at minus-ends is not known. Here, we use quantitative imaging and laser ablation to show that NuMA targets dynactin to minus-ends, localizing dynein activity there. NuMA is recruited to new minus-ends independently of dynein and more quickly than dynactin; both NuMA and dynactin display specific, steady-state binding at minus-ends. NuMA localization to minus-ends involves a C-terminal region outside NuMA's canonical microtubule-binding domain and is independent of minus-end binders γ-TuRC, CAMSAP1, and KANSL1/3. Both NuMA's minus-end-binding and dynein-dynactin-binding modules are required to rescue focused, bipolar spindle organization. Thus, NuMA may serve as a mitosis-specific minus-end cargo adaptor, targeting dynein activity to minus-ends to cluster spindle microtubules into poles.

Introduction

Each time a cell divides, molecular motors help remodel the microtubule cytoskeleton into a bipolar assembly of microtubules called the spindle. The spindle’s bipolar architecture is essential to its function – accurate chromosome segregation. In mammalian spindles, microtubule plus-ends mechanically couple to chromosomes, while microtubule minus-ends are focused into two poles that dictate where chromosomes are transported at anaphase. To build the spindle’s architecture, motors must exert spatially regulated forces on microtubules. Microtubule ends offer platforms for such localized regulation, since they are structurally distinct. Indeed, motor recruitment and activity at plus-ends is well-documented (Wu et al., 2006), but motor regulation at minus-ends is less well understood.

Dynein is a minus-end-directed motor which slides parallel spindle microtubules to focus their minus-ends into spindle poles (Heald et al., 1996; Verde et al., 1991), working in complex with its adaptor dynactin and the microtubule-binding protein NuMA (Gaglio et al., 1996; Merdes et al., 1996). The clustering of parallel microtubules into poles presents a geometric problem when forces are indiscriminately applied all along microtubules: inversely oriented motors between parallel microtubules will oppose each other, resulting in gridlock, unless symmetry is broken by dynein enrichment at microtubule minus-ends (Hyman and Karsenti, 1996; McIntosh et al., 1969; Surrey et al., 2001). In computational models, localizing a minus-end-directed motor at microtubule ends permits microtubule clustering into asters or poles (Foster et al., 2015; Goshima et al., 2005; Nedelec and Surrey, 2001; Surrey et al., 2001) and the emergence of a robust steady-state spindle length (Burbank et al., 2007). More recently, experimental work has shown that dynein-dynactin and NuMA do indeed selectively localize to spindle minus-ends, with dynein pulling on them after kinetochore-fiber (k-fiber) ablation in mammalian spindles (Elting et al., 2014; Sikirzhytski et al., 2014). This is consistent with suggestions that dynein and NuMA capture and pull on distal k-fiber minus-ends in monopolar spindles (Khodjakov et al., 2003). Altogether, these findings demonstrate the importance (in silico) and existence (in vivo) of localized dynein activity at spindle microtubule minus-ends.

How dynein becomes localized at minus-ends remains an open question. Dynein may be enriched near minus-ends because it walks toward them on microtubules and piles up when it runs out of track; indeed, pile-up of dynein has been observed at minus-ends in vitro (McKenney et al., 2014; Soundararajan and Bullock, 2014) and can drive minus-end clustering (Tan et al., 2017). Alternatively, the exposed α-tubulin interface at minus-ends is structurally distinct and could bind an adaptor protein that specifically recruits dynein, analogous to recruitment at canonical dynein cargoes like organelles (Kardon and Vale, 2009). NuMA can target dynein-dynactin to the cell cortex (Lechler and Fuchs, 2005; Nguyen-Ngoc et al., 2007) and thus could be one such adaptor. However, in vitro NuMA has shown no direct affinity for minus-ends specifically, binding all along the microtubule lattice (Du et al., 2002; Forth et al., 2014; Haren and Merdes, 2002) or at both ends (Seldin et al., 2016), unlike three proteins known to interact directly with minus-ends at mitosis: γ-TuRC (Zheng et al., 1995), CAMSAP1 (Akhmanova and Hoogenraad, 2015; Hendershott and Vale, 2014; Jiang et al., 2014) and KANSL1/3 (Meunier et al., 2015). In cells, NuMA is thought to require dynein activity to carry it to minus-ends and spindle poles (Merdes et al., 2000), where it anchors spindle microtubules (Dionne et al., 1999; Gaglio et al., 1995; Heald et al., 1997; Silk et al., 2009). Thus, it remains unclear whether dynein-dynactin and NuMA have specific binding sites at minus-ends, and if so, whether they are recruited by known minus-end binders. Finally, knowing how dynein is targeted to minus-ends would make it possible to test the in vivo role of minus-end-localized – compared to indiscriminately-localized – forces in spindle organization.

Here, we use laser ablation in cells to create new, isolated minus-ends in the mammalian spindle, and quantitative imaging to map protein recruitment to these ends and its mechanistic basis. We demonstrate that NuMA binds at minus-ends independently of dynein, and that NuMA targets dynactin – and thereby dynein activity – to spindle minus-ends. This challenges the prevailing model that dynein delivers NuMA to spindle minus-ends and poles (Merdes et al., 2000; Radulescu and Cleveland, 2010). NuMA localization to minus-ends is independent of known direct minus-end binders γ-TuRC, CAMSAP1, and KANSL1/3, and it involves both NuMA’s canonical microtubule-binding domain and an additional region of its C-terminus. Thus, NuMA – which is sequestered in the nucleus at interphase (Lydersen and Pettijohn, 1980) – may serve as a mitosis-specific minus-end cargo adaptor, recruiting dynein activity to spindle minus-ends. Both NuMA’s minus-end-binding domain and dynactin-binding domain are required for correct spindle architecture, supporting long-standing in silico predictions that localizing dynein to minus-ends enables effective clustering of parallel microtubules into poles. These findings identify a mechanism for mitosis-specific recruitment of dynein to microtubule minus-ends and, more broadly, illustrate how spatial regulation of local forces may give rise to larger scale cytoskeletal architectures.

Results

Dynactin and NuMA display mitosis-specific, steady-state binding at microtubule minus-ends

To visualize the spatial targeting of the dynein-dynactin complex to microtubule minus-ends – which are normally buried in dense mammalian spindles – we used nocodazole washout and laser ablation to create resolvable minus-ends in mitotic cells. First, to determine whether dynein-dynactin and NuMA localize to individual microtubule minus-ends, we treated mammalian PtK2 and RPE1 cells with the microtubule-depolymerizing drug nocodazole and fixed cells 6–8 min after drug washout to capture acentrosomal microtubules with clearly visible plus- and minus-ends (Figure 1A). p150 (p150, a dynactin subunit) and NuMA strongly co-localized at one end of these individual microtubules (Figure 1A), with a clear binding preference for minus-ends over the microtubule lattice or the plus-end when polarity was marked by EB1 (Figure 1B). Interestingly, in prophase cells before nuclear envelope breakdown, p150 localized predominantly to plus-ends rather than minus-ends (Figure 1B; Figure 1—figure supplement 1), consistent with dynactin’s interphase localization (Vaughan et al., 1999). Thus, nuclear envelope breakdown (NEB) confers dynactin’s preference for minus-ends, suggesting regulated, mitosis-specific spatial targeting.

Figure 1.

Dynactin and NuMA display specific, steady-state binding at mitotic minus-ends.

See also Figure 1—figure supplement 1 and Videos 1–3. (A) Representative immunofluorescence image showing co-localization of NuMA (green) and p150 (dynactin subunit; cyan) at microtubule minus-ends in mitotic PtK2 cells (post-NEB) fixed after washout of 5 μM nocodazole. Scale bar, 10 μm. Inset: zoom of white box, with 1 μm scale bar. (B) Representative immunofluorescence images of mitotic RPE1 cells, processed as in (A). After nuclear envelope breakdown (post-NEB), EB1 (green) and p150 (cyan) localize to opposite microtubule ends. In prophase cells (pre-NEB), p150 instead co-localizes with EB1 at plus-ends. Dashed white circles highlight ends. Scale bar, 1 μm. Graph displays mean percentage ±SEM of p150 at each location within one cell for n = 72 microtubules, 8 cells (pre-NEB); n = 57 microtubules, 16 cells (post-NEB). (C–E) Representative time-lapse live images and kymographs of PtK2 cells expressing (C) GFP-Arp1A, (D) GFP-NuMA, or (E) GFP-CAMSAP1. Arp1A, NuMA, and CAMSAP1 are all recruited to k-fiber minus-ends (arrowheads) created by ablation (‘X’s) and move with them as dynein pulls them poleward (Elting et al., 2014). Time is in min:s, with the frame captured immediately following ablation set to 00:00. ‘*’ marks plus-end of ablated k-fiber, and ‘o’ marks its sister. Scale bar, 5 μm. Kymographs (right) are along poleward paths (dashed lines). (F) Plot of mean normalized GFP intensity and SEM (shading) over time at ablation-created minus-ends for GFP-Arp1A, GFP-NuMA, and GFP-CAMSAP1. Time = 0 s at the first frame following ablation. n = 10 ablations, 7 cells (Arp1A); n = 18 ablations, 7 cells (NuMA); n = 13 ablations, 7 cells (CAMSAP1). (G) Time from ablation to half maximum GFP intensity, calculated for each individual ablation (see Materials and methods) and then averaged for data in (F). Error bars show SEM. p-Values calculated by Tukey post hoc test after one-way ANOVA (F(2,38) = 9.26, p=0.0005). (H) Time to half-maximum GFP-Arp1A or GFP-NuMA intensity at ablation-created minus-ends as a function of length of ablation-created k-fiber stubs. Correlation coefficient = −0.62, p=0.06, n = 10 (Arp1A); correlation coefficient = 0.44, p=0.07, n = 18 (NuMA).

Immunofluorescence showing localization of p150 (dynactin subunit; magenta) and NuMA (red) within asters formed after washout of 5 μM nocodazole in RPE1 cells. Before nuclear envelope breakdown (pre-NEB), NuMA is sequestered in the nucleus and p150 is visible at aster plus-ends (arrowheads), facing outward. After nuclear envelope breakdown (post-NEB), both NuMA and p150 concentrate at aster centers (arrowhead) where minus-ends are. (Similar to Figure 1B, where individual microtubules in the cell periphery with resolvable plus- and minus-ends were analyzed instead of the asters we show here.) Scale bar, 5 μm.

Figure 1—figure supplement 1.

Dynactin localization shifts from plus-ends to minus-ends upon nuclear envelope breakdown.

Immunofluorescence showing localization of p150 (dynactin subunit; magenta) and NuMA (red) within asters formed after washout of 5 μM nocodazole in RPE1 cells. Before nuclear envelope breakdown (pre-NEB), NuMA is sequestered in the nucleus and p150 is visible at aster plus-ends (arrowheads), facing outward. After nuclear envelope breakdown (post-NEB), both NuMA and p150 concentrate at aster centers (arrowhead) where minus-ends are. (Similar to Figure 1B, where individual microtubules in the cell periphery with resolvable plus- and minus-ends were analyzed instead of the asters we show here.) Scale bar, 5 μm.

Video 1.

Dynactin is robustly recruited to new spindle microtubule minus-ends.

See also Figure 1C. Live confocal imaging of a PtK2 cell expressing GFP-Arp1A, with spindle microtubules labeled by siR-tubulin. GFP-Arp1A is recruited to k-fiber minus-ends (arrowhead) created by ablation (‘X’) and moves with them as dynein pulls them poleward (Elting et al., 2014). Time is in min:s, with the frame captured immediately following ablation set to 00:00. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 μm.

Video 3.

CAMSAP1 is robustly recruited to new spindle microtubule minus-ends.

See also Figure 1E. Live confocal imaging of a PtK2 cell expressing GFP-CAMSAP1, with spindle microtubules labeled by siR-tubulin. GFP-CAMSAP1 is immediately recruited to k-fiber minus-ends (arrowhead) created by ablation (‘X’) and moves with them as dynein pulls them poleward. Time is in min:s, with the frame captured immediately following ablation set to 00:00. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 μm.

Second, we sought to test whether dynactin and NuMA have finite binding sites at minus-ends by measuring the kinetics of their recruitment. To do so, we used laser ablation of k-fibers in PtK2 cells to create new minus-ends within the spindle body (Figure 1C–E). By spatially and temporally synchronizing the creation of a bundle of minus-ends, laser ablation allowed for dynamic measurements of the recruitment to minus-ends of GFP-tagged dynactin (Arp1A) and NuMA and comparison to a direct minus-end binding protein, CAMSAP1. High dynein background in the cytoplasm prevented accurate recruitment kinetics measurements of the motor itself. Dynactin, NuMA, and CAMSAP1 robustly recognized new microtubule minus-ends within the spindle (Figure 1C–E; Videos 1–3). The kinetics of dynactin and NuMA recruitment to minus-ends were distinct from CAMSAP1, which reached a max intensity after approximately 5–10 s, but then decreased in intensity as if its binding sites at the minus-end were gradually obstructed (Figure 1E–G). Unlike CAMSAP1, dynactin and NuMA intensities reached a stable plateau, suggesting that binding saturates, reaching a steady-state. This finite binding is consistent with a truly end-specific localization (rather than localization along the unbounded lattice near the minus-end). In addition, the rate of NuMA or dynactin accumulation at new minus-ends did not correlate with the length of k-fiber stubs created by ablation (Figure 1H), which could indicate that recruitment rate is set by the number of individual microtubule minus-ends (which is similar across k-fibers [McEwen et al., 1997]) rather than k-fiber length.

NuMA is robustly recruited to new spindle microtubule minus-ends.

See also Figure 1D. Live confocal imaging of a PtK2 cell expressing GFP-NuMA, with spindle microtubules labeled by siR-tubulin. GFP-NuMA is quickly recruited to k-fiber minus-ends (arrowhead) created by ablation (‘X’) and moves with them as dynein pulls them poleward (Elting et al., 2014). Time is in min:s, with the frame captured immediately following ablation set to 00:00. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 μm.

Interestingly, NuMA intensity at new minus-ends increased at a faster rate and saturated sooner than dynactin (Figure 1F,G). This observation hints at a model in which NuMA targets dynein-dynactin to minus-ends, and it is less easily consistent with the idea that dynein delivers NuMA to minus-ends. If true, this hypothesis would explain why dynactin’s minus-end preference arises after NEB, when NuMA is released from the nucleus into the mitotic cytoplasm.

NuMA localizes to minus-ends independently of dynein

The finding that NuMA localizes to minus-ends more quickly than dynactin could suggest that NuMA directly or indirectly recognizes the exposed α-tubulin structure of minus-ends and subsequently recruits dynein-dynactin (‘Structural Recognition,’ Figure 2A). Alternatively, dynein-dynactin could walk toward minus-ends, carrying NuMA, and pile up or dwell there (‘Walk and Pile-up,’ Figure 2B). To test the ‘Walk and Pile-up’ model, we inhibited dynein-dynactin in PtK2 spindles by over-expressing p50 (dynamitin), resulting in fully unfocused k-fiber arrays (Figure 2C). The ‘Walk and Pile-up’ model predicts that in the absence of dynein-dynactin transport, NuMA – which requires dynactin for its interaction with dynein (Merdes et al., 2000) – should not reach minus-ends. Instead, GFP-NuMA was robustly recruited to minus-ends created by k-fiber ablation (Figure 2C; Video 4), indicating that NuMA can localize to minus-ends independently of dynein activity. Similarly, NuMA was recruited to ablation-created minus-ends when dynein was inhibited by p150-CC1 over-expression (Figure 2—figure supplement 1A,B). Thus, NuMA localizes to minus-ends without dynein carrying it there – consistent with early observations in extract asters and spindles (Gaglio et al., 1996; Heald et al., 1997) but in contrast to the prevailing view that dynein delivers NuMA to minus-ends (Merdes et al., 2000; Radulescu and Cleveland, 2010). Slower NuMA accumulation kinetics after p50 overexpression suggest that dynein-dynactin-NuMA complex formation may aid rapid NuMA recruitment, but dynein ‘Walk and Pile-up’ alone cannot explain NuMA’s minus-end affinity (Figure 2D,E).

Figure 2.

NuMA can localize to minus-ends independently of dynein.

See also Figure 2—figure supplement 1 and Video 4. (A–B) Models for targeting dynein to minus-ends. Model A, Structural Recognition: Cytoplasmic molecules may recognize the exposed α-tubulin interface at minus-ends and recruit dynein. Model B, Walk and Pile-up: Minus-end-directed dynein may walk to minus-ends and pile up. (C) Representative time-lapse live images and kymograph of PtK2 cell expressing GFP-NuMA, in which dynein-dynactin is inhibited by overexpression of p50 (dynamitin). K-fibers are unfocused and splayed, but NuMA is still robustly recruited to k-fiber minus-ends (arrowheads) created by ablation (‘X’). Time is in min:s, with frame captured immediately following ablation set to 00:00. ‘*’ marks plus-end of ablated k-fiber, and ‘o’ marks its sister. Scale bars, 5 µm. Kymograph (right) taken along dashed line path. (D) Plot of mean normalized GFP-NuMA intensity and SEM (shading) over time at ablation-created minus-ends. Time = 0 s at the first frame following ablation. n = 18 ablations, 7 cells for control; n = 14 ablations, 8 cells for dynein-inhibited (p50 overexpression). (E) Time from ablation to half maximum GFP-NuMA intensity, calculated for each individual ablation (see Methods) and then averaged for data in (D). Error bars show SEM. (F) Representative immunofluorescence images of inducible-Cas9 dynein heavy chain (DHC)-knockout HeLa cells (McKinley and Cheeseman, 2017) showing robust localization of NuMA and p150 (dynactin) at minus-ends when DHC is deleted. Scale bar, 5 μm.

(A) Live images of GFP-α-tubulin immediately before and after k-fiber ablation at targeted sites (red ‘X’s) in a PtK2 cell in which dynein cargo-binding is inhibited by transfection of the dominant-negative p150-CC1 fragment (Quintyne and Schroer, 2002). Scale bar, 5 μm. (B) Immunofluorescence image of NuMA (magenta) and α-tubulin (yellow) in cell from (A), fixed after ablation. NuMA (arrowheads) localizes to new minus ends. Scale bar, 5 μm. (C) Western blot showing >65% depletion (normalized to α-tubulin) of dynein intermediate chain (DIC) protein at the cell population level after dynein heavy chain (DHC) depletion using inducible-Cas9 DHC-knockout HeLa cells (McKinley and Cheeseman, 2017). DIC depletion has been shown to correlate with DHC depletion (Levy and Holzbaur, 2008). Because knockout was heterogeneous across the population, dynein loss within individual cells chosen for analysis was always verified by immunofluorescence, as in (D), or by spindle phenotype. (D) Immunofluorescence images of inducible DHC-knockout HeLa cells show robust localization of NuMA at minus-ends after DHC knockout, in the absence of dynein (dynein intermediate chain; DIC). Scale bar, 5 μm. (E) Plot of mean normalized GFP-NuMA intensity and SEM (shading) over time at ablation-created minus-ends in HeLa cells with and without dynein deletion (DHC KO). Time = 0 s at the first frame following ablation (‘X’). n = 12 ablations, 7 cells for control; n = 10 ablations, 8 cells for DHC KO. (F) Time from ablation to half maximum GFP-NuMA intensity, calculated for each individual ablation (see Methods) and then averaged for data in (E). Error bars show SEM.

Video 4.

NuMA quickly localizes to k-fiber minus-ends despite dynein-dynactin inhibition.

See also Figure 2C. Live confocal imaging of a PtK2 cell expressing GFP-NuMA, in which dynein-dynactin is inhibited by overexpression of p50 (dynamitin). K-fibers are unfocused and splayed due to dynein-dynactin inhibition, but NuMA is still robustly recruited to k-fiber minus-ends (arrowheads) created by ablation (‘X’). Time is in min:s, with frame captured immediately following ablation set to 00:00. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 µm.

Figure 2—figure supplement 1.

NuMA localizes to minus-ends despite dynein inhibition or knockout.

(A) Live images of GFP-α-tubulin immediately before and after k-fiber ablation at targeted sites (red ‘X’s) in a PtK2 cell in which dynein cargo-binding is inhibited by transfection of the dominant-negative p150-CC1 fragment (Quintyne and Schroer, 2002). Scale bar, 5 μm. (B) Immunofluorescence image of NuMA (magenta) and α-tubulin (yellow) in cell from (A), fixed after ablation. NuMA (arrowheads) localizes to new minus ends. Scale bar, 5 μm. (C) Western blot showing >65% depletion (normalized to α-tubulin) of dynein intermediate chain (DIC) protein at the cell population level after dynein heavy chain (DHC) depletion using inducible-Cas9 DHC-knockout HeLa cells (McKinley and Cheeseman, 2017). DIC depletion has been shown to correlate with DHC depletion (Levy and Holzbaur, 2008). Because knockout was heterogeneous across the population, dynein loss within individual cells chosen for analysis was always verified by immunofluorescence, as in (D), or by spindle phenotype. (D) Immunofluorescence images of inducible DHC-knockout HeLa cells show robust localization of NuMA at minus-ends after DHC knockout, in the absence of dynein (dynein intermediate chain; DIC). Scale bar, 5 μm. (E) Plot of mean normalized GFP-NuMA intensity and SEM (shading) over time at ablation-created minus-ends in HeLa cells with and without dynein deletion (DHC KO). Time = 0 s at the first frame following ablation (‘X’). n = 12 ablations, 7 cells for control; n = 10 ablations, 8 cells for DHC KO. (F) Time from ablation to half maximum GFP-NuMA intensity, calculated for each individual ablation (see Methods) and then averaged for data in (E). Error bars show SEM.

To confirm that NuMA localizes to minus-ends even after genetic dynein deletion, we used an inducible CRISPR/Cas9 HeLa cell line to knock out dynein’s heavy chain (DHC) (McKinley and Cheeseman, 2017). After dynein knockout (Figure 2—figure supplement 1C,D), both NuMA and dynactin localized robustly to k-fiber minus-ends (Figure 2F; Figure 2—figure supplement 1E,F). Together, these data indicate that NuMA can bind to minus-ends, either directly or indirectly, in the absence of dynein and are consistent with NuMA-mediated recognition of minus-ends (‘Structural Recognition,’ Figure 2A). In addition, they suggest that NuMA may interact with and target dynactin to minus-ends even in the absence of dynein, when dynein-dynactin complexes cannot form and dynein motility cannot deliver dynactin close to minus-ends.

NuMA is required for dynein activity and dynactin localization at minus-ends

If NuMA does target dynein-dynactin to minus-ends, localizing force there, we would expect NuMA to be required for dynein to transport minus-ends created by k-fiber ablation (Elting et al., 2014; Sikirzhytski et al., 2014). To test this hypothesis, we made inducible CRISPR/Cas9 RPE1 cell lines to knock out NuMA (Figure 3—figure supplement 1) (McKinley et al., 2015; Wang et al., 2015). Indeed, when NuMA was knocked out – causing elongated, heterogeneously disorganized spindles with detached centrosomes – ablation-created minus-ends were no longer transported toward poles by dynein (Figure 3A–C; Video 5). These data are consistent with a previous finding that NuMA antibody injection prevents distal k-fiber looping events in monopolar spindles, in which dynein likely pulls on free k-fiber minus-ends (Khodjakov et al., 2003). Thus, NuMA is required for dynein activity at spindle microtubule minus-ends.

Figure 3—figure supplement 1.

Inducible CRISPR/Cas9 NuMA knockout.

(A) Schematic of inducible CRISPR/Cas9 knockout system used (McKinley et al., 2015; Wang et al., 2015). Cell lines created have CRISPR/Cas9 and guide RNA stably integrated, but Cas9 expression is only induced upon doxycycline addition 4 days before imaging or analysis, allowing for genetic manipulation of essential mitotic genes. (B) Western blot showing >90% depletion (normalized to actin) of NuMA protein at the cell population level after knockout. Whenever possible, complete NuMA loss within individual cells analyzed was verified by immunofluorescence.

Figure 3.

NuMA is required for force generation at minus-ends.

See also Figure 3—figure supplement 1 and Video 5. (A) Representative time-lapse live images of RPE1 cell in which NuMA has been knocked out using an inducible Cas9 system. GFP-CAMSAP1 is expressed to label minus-ends. After ablation (‘X’), the k-fiber minus-end (arrowhead) is not transported poleward by dynein and remains detached from the spindle. Time is in min:s, with the frame captured immediately following ablation set to 00:00 s. Plus-end of ablated k-fiber is marked by ‘*’. Scale bars, 5 μm. (B) Cartoon: In the absence of NuMA, force generation at ablation-created minus-ends is not observed. (C) Movement of ablation-created minus-ends (marked by GFP-CAMSAP1) relative to the pole. In control cells (green traces), minus-ends are transported toward the pole by dynein at consistent speeds, but this transport is lost when NuMA is knocked out (purple traces). n = 19 ablations, 8 cells (control); n = 20 ablations, 6 cells (NuMA knockout).

(A) Schematic of inducible CRISPR/Cas9 knockout system used (McKinley et al., 2015; Wang et al., 2015). Cell lines created have CRISPR/Cas9 and guide RNA stably integrated, but Cas9 expression is only induced upon doxycycline addition 4 days before imaging or analysis, allowing for genetic manipulation of essential mitotic genes. (B) Western blot showing >90% depletion (normalized to actin) of NuMA protein at the cell population level after knockout. Whenever possible, complete NuMA loss within individual cells analyzed was verified by immunofluorescence.

Video 5.

NuMA is required for transport of minus-ends by dynein.

See also Figure 3A. Live confocal imaging of a RPE1 cell in which NuMA has been knocked out using an inducible Cas9 system, and GFP-CAMSAP1 is expressed to label minus-ends. After ablation (‘X’), the k-fiber minus-end (arrowhead) is not transported poleward by dynein and remains detached from the disorganized spindle. Time is in min:s, with the frame captured immediately following ablation set to 00:00 s. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 μm.

Dynein activity at minus-ends could require NuMA because NuMA modulates dynein-dynactin’s ability to pull on microtubules, or because NuMA localizes dynein to minus-ends. Given the findings in Figures 1 and 2, we hypothesized that NuMA localizes dynein activity by recruiting dynactin to minus-ends. Indeed, after NuMA knockout, dynactin (GFP-Arp1A) was no longer detectable at k-fiber minus-ends created by laser ablation (Figure 4A; Video 6). Immunofluorescence experiments confirmed that in the absence of NuMA, dynactin remained on the spindle but no longer localized to minus-ends (Figure 4B). Interestingly, dynein’s localization within the spindle (labeled using an antibody against dynein intermediate chain) was less minus-end-specific than dynactin’s, both before and after NuMA deletion (Figure 4C).

Figure 4.

NuMA recruits dynactin to mitotic minus-ends.

See also Video 6. (A) Representative time-lapse live images and kymograph of RPE1 cell in which NuMA has been knocked out using an inducible Cas9 system. 2xGFP-Arp1A recruitment was never observable at minus-ends (arrowheads) created by ablation (‘X’) (n = 16 ablations, 6 cells). Time is in min:s, with frame captured immediately following ablation set to 00:00. Plus-end of ablated k-fiber is marked by ‘*’. Scale bars, 2 µm. Kymographs (right) taken along dashed line path. (B) Representative immunofluorescence images of control and NuMA-knockout RPE1 spindles, showing a loss of p150 (dynactin) at minus-ends in the absence of NuMA. Scale bars, 5 μm. (C) Representative immunofluorescence images of control and NuMA-knockout RPE1 spindles. Dynein localizes along spindle microtubules in both conditions; its distribution is less noticeably altered by NuMA loss. Scale bars, 5 μm. (D) Representative immunofluorescence images of mitotic RPE1 cells (post-NEB) fixed after washout of 5 μM nocodazole, as in Figure 1A,B. In control cells, p150 (dynactin; purple) localizes to microtubule minus-ends, opposite EB1 (red). When NuMA is knocked out, p150 co-localizes with EB1 at plus-ends. Dashed white circles highlight ends. Scale bars, 2 μm. Graph displays mean percentage ±SEM of p150 at each location within each cell, for n = 72 microtubules, 8 cells (control); n = 72 microtubules, 11 cells (NuMA knockout).

Video 6.

Dynactin is not recruited to minus-ends in the absence of NuMA.

See also Figure 4A. Live confocal imaging of a RPE1 cell in which NuMA has been knocked out using an inducible Cas9 system. In the absence of NuMA, 2xGFP-Arp1A does not localize to k-fiber minus-ends, and its recruitment is not detectable at new minus-ends (arrowheads) created by ablation (‘X’). Note that mislocalized 2xGFP-Arp1A puncta diffuse randomly within the spindle, including within microns of the new minus-end, but do not remain there. Time is in min:s, with frame captured immediately following ablation set to 00:00. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 µm.

In addition, nocodazole washout experiments revealed that dynactin (p150) localization to individual microtubule minus-ends was lost upon NuMA deletion (Figure 4D). Instead, dynactin frequently co-localized with EB1 at plus-ends, similar to its interphase and pre-NEB localization pattern (Figure 4D; Figure 1B). Thus, the data indicate that NuMA is required for the transport of minus-ends by dynein because NuMA localizes to minus-ends and recruits dynactin there.

NuMA localizes to minus-ends independently of known minus-end binders

How is NuMA targeted to minus-ends, independently of dynein? In vitro, canonical microtubule-binding regions of NuMA have shown no preference for minus-ends relative to the lattice or plus-end of purified microtubules (Du et al., 2002; Forth et al., 2014; Haren and Merdes, 2002; Seldin et al., 2016). Given this lack of minus-end-specific binding in vitro, we hypothesized that NuMA is indirectly recruited to spindle minus-ends by one of three known direct minus-end binders active at mitosis: γ-TuRC (Zheng et al., 1995), CAMSAP1 (Hendershott and Vale, 2014; Jiang et al., 2014) (Figure 1; CAMSAP2 and 3 are phosphorylated at mitosis and no longer interact with microtubules [Jiang et al., 2014]), and KANSL1/3 (Meunier et al., 2015). To test this hypothesis, we treated RPE1 cells with 30 μM gatastatin to block γ-TuRC binding (Chinen et al., 2015) (Figure 5A; Figure 5—figure supplement 1A,B). We also made inducible CRISPR/Cas9 RPE1 cell lines to knock out CAMSAP1 or KANSL1 (Figure 5—figure supplement 1C,D), as KANSL1 depletion has been shown to delocalize the entire KANSL complex (Meunier et al., 2015). CAMSAP1 knockout caused a small reduction in spindle length (Figure 5—figure supplement 1E), consistent with the spindle minus-end protecting function ascribed to its Drosophila homolog, Patronin (Goodwin and Vale, 2010). To our surprise, however, none of these perturbations qualitatively altered NuMA localization at spindle poles (Figure 5A). To check for a more subtle contribution of γ-TuRC, CAMSAP1, or KANSL1 to NuMA localization at minus-ends, we performed k-fiber ablations after 30 μM gatastatin treatment, CAMSAP1 knockout, or KANSL1 knockout and quantified GFP-NuMA recruitment kinetics at new minus-ends. NuMA recruitment to new minus-ends remained robust, and recruitment timescales were statistically indistinguishable from control (Figure 5B–C). Thus, the data indicate that the direct mitotic minus-end binders γ-TuRC, CAMSAP1, and KANSL1/3 are not responsible for NuMA’s localization to spindle microtubule minus-ends.

Figure 5.

NuMA localizes to minus-ends independently of known minus-end binding proteins.

See also Figure 5—figure supplement 1. (A) Schematic of hypothesis that a minus-end binding protein recruits NuMA. Instead, representative immunofluorescence images show unchanged NuMA localization in control RPE1 cells and RPE1 cells in which direct mitotic minus-end binders are inhibited (30 μM gatastatin to inhibit γ-tubulin) or knocked out (CAMSAP1, KANSL1). Scale bar, 5 μm. (B) Plot of mean normalized GFP-NuMA intensity and SEM (shading) over time at ablation-created minus-ends. Time = 0 s at the first frame following ablation. n = 18 ablations, 7 cells (control); n = 9 ablations, 8 cells (+gatastatin); n = 10 ablations, 5 cells (CAMSAP1 knockout); n = 8 ablations, 4 cells (KANSL1 knockout). (C) Time from ablation to half maximum GFP-NuMA intensity was calculated for each individual ablation (see Materials and methods) and then averaged for data in (B). Error bars show SEM. ‘n.s.’ indicates no statistically significant difference. One-way ANOVA: F(3,42) = 0.45, p=0.72. (D–G) 3D STORM of NuMA at PtK2 k-fiber minus-ends created by ablation. (D) K-fibers were cut using ablation (red ‘X’), and cells were immediately fixed for immunofluorescence. Individual ablated spindles were imaged by spinning-disk confocal (E) and then by 3D STORM (F,G). In two right-most panels (G), structures are colored according to position in the Z-axis (red = +300 nm, blue = −300 nm). Scale bars: 5 μm in (D,E); 1 μm in (F); 100 nm in (G). (E) Distances between neighboring ‘nodes’ of NuMA (e.g., ‘*’ here and in (G)) are ~50–150 nm, consistent with measured spacing between individual microtubules within PtK2 k-fibers (McDonald et al., 1992). n = 32 nodes, 4 ablations.

(A) Mean spindle length ± SEM in control RPE1 cells compared to cells treated with 30 μM gatastatin (γ-tubulin inhibitor) for 25 min. The mean length of gatastatin-treated spindles was reduced by 23%, very similar to previously reported measurements after γ-tubulin inhibition (Chinen et al., 2015). n = 24 control cells; n = 25 gatastatin-treated cells from three separate experiments. (B) Microtubule re-growth assay in STLC-arrested monopolar spindles after cold treatment (bottom panels) shows that 30 μM gatastatin blocks γ-TuRC’s ability to bind α-tubulin to nucleate microtubules. Because gatastatin specifically blocks γ-tubulin binding, not α- or β-tubulin, it does not depolymerize existing microtubule networks (top panels). Scale bar, 5 μm. (C–D) Western blot showing > 85% depletion (normalized to actin) of (B) CAMSAP1 and (C) KANSL1 protein at the cell population level after knockout (KO). Whenever possible, complete protein loss within individual cells analyzed was verified by immunofluorescence. (E) Mean spindle length ± SEM in control RPE1 cells (no doxycycline added) and CAMSAP1 knockout cells (after Cas9 induction by doxycycline). Complete CAMSAP1 loss was verified by immunofluorescence for all cells analyzed. n = 11 control cells; n = 12 CAMSAP1 knockout cells from one experiment.

Figure 5—figure supplement 1.

Characterization of gatastatin treatment, CAMSAP1 knockout, and KANSL1 knockout.

(A) Mean spindle length ± SEM in control RPE1 cells compared to cells treated with 30 μM gatastatin (γ-tubulin inhibitor) for 25 min. The mean length of gatastatin-treated spindles was reduced by 23%, very similar to previously reported measurements after γ-tubulin inhibition (Chinen et al., 2015). n = 24 control cells; n = 25 gatastatin-treated cells from three separate experiments. (B) Microtubule re-growth assay in STLC-arrested monopolar spindles after cold treatment (bottom panels) shows that 30 μM gatastatin blocks γ-TuRC’s ability to bind α-tubulin to nucleate microtubules. Because gatastatin specifically blocks γ-tubulin binding, not α- or β-tubulin, it does not depolymerize existing microtubule networks (top panels). Scale bar, 5 μm. (C–D) Western blot showing > 85% depletion (normalized to actin) of (B) CAMSAP1 and (C) KANSL1 protein at the cell population level after knockout (KO). Whenever possible, complete protein loss within individual cells analyzed was verified by immunofluorescence. (E) Mean spindle length ± SEM in control RPE1 cells (no doxycycline added) and CAMSAP1 knockout cells (after Cas9 induction by doxycycline). Complete CAMSAP1 loss was verified by immunofluorescence for all cells analyzed. n = 11 control cells; n = 12 CAMSAP1 knockout cells from one experiment.

Given this lack of involvement of direct minus-end binding proteins, we sought to confirm using super-resolution microscopy that NuMA specifically localizes at individual spindle microtubule minus-ends. Three-dimensional stochastic optical reconstruction microscopy (3D STORM [Huang et al., 2008; Rust et al., 2006]) of NuMA at k-fiber minus-ends created by ablation revealed organized clusters of NuMA puncta, rather than a lawn of molecules along the lattice near the minus-end (Figure 5D–G). The spacing between puncta was consistent with the distance between individual microtubules within k-fibers (Figure 5H) as measured by electron microscopy in the same cell type (PtK2) (McDonald et al., 1992), supporting the idea that these NuMA puncta are built on individual microtubule minus-ends. A yet undiscovered minus-end binding protein may recruit NuMA; alternatively, NuMA may have direct minus-end-specific binding ability that has not been recapitulated in vitro.

NuMA function requires both minus-end-recognition and dynactin-recruitment modules

To define the NuMA domain required for spindle minus-end localization, we performed rescue experiments with different NuMA truncations in the NuMA-knockout background (Figure 6A). This avoids the C-terminus-mediated oligomerization (Harborth et al., 1999) with endogenous protein that can complicate interpretations of localization. In this NuMA-knockout background, full-length NuMA (‘FL’) localized to spindle minus-ends, as did a bonsai construct (‘N-C’) with most of the central coiled-coil removed (Figure 6B). To our surprise, ‘N-C’ rescued spindle architecture as effectively as full-length NuMA, indicating that the extraordinary length of NuMA protein is not essential to its function in spindle structure (Figure 6C). Importantly, NuMA’s C-terminus (‘C’) alone localized to minus-ends even in the absence of endogenous full-length protein (Figure 6B). Its intensity at minus-ends was less striking than that of full-length or N-C protein at poles, likely because minus-ends are not as densely concentrated in disorganized spindles, and perhaps because NuMA’s N-terminus facilitates formation of higher order NuMA assemblies. However, the preference of NuMA ‘C’ for spindle ends was clear. Because dynein-dynactin interacts with NuMA’s N-terminus (Kotak et al., 2012), the minus-end localization of NuMA’s C-terminus provides further support for dynein-independent minus-end recognition.

Figure 6.

NuMA function requires both minus-end-recognition and dynactin-recruitment modules.

See also Table 1, Figure 6—figure supplement 1, and Video 7. (A) Schematic maps of NuMA truncations and chimeras used. ‘FL’ indicates full-length NuMA. ‘N-C,’ ‘C,’ ‘C-tail1,’ and ‘C-tail2’ are NuMA truncations as indicated. ‘N-Tau’ includes the microtubule binding domain of Tau (orange). Dynein-dynactin bind in the first 705 amino acids of NuMA (Kotak et al., 2012). ‘C-tail2’ was previously implicated in microtubule (MT) binding (Chang et al., 2017; Du et al., 2002; Gallini et al., 2016; Haren and Merdes, 2002). (B) Representative immunofluorescence images showing localization of GFP-tagged NuMA truncations and chimeras expressed in RPE1 cells in which endogenous NuMA has been knocked out. Canonical microtubule binding domains in tail2 localize all along spindle microtubules, while the addition of tail1 to tail2A confers minus-end localization. Note that NuMA antibody does not recognize NuMA’s C-terminus and only weakly recognizes its N-terminus. Scale bar, 5 μm. (C) Graph of the mean percentage of cells ± SEM for each condition from data in (B) that display bipolar spindles with focused poles (light gray) compared to disorganized spindle architecture characteristic of NuMA knockout (detached centrosomes, loss of two focused poles; dark gray). n = 19 (‘FL’); n = 32 (‘N-C’); n = 20 (‘C’); n = 16 (‘N-Tau’) from three to five independent experiments. p-Values calculated by Tukey post-hoc test after one-way ANOVA (F(3,10) = 267, p<0.00001). (D) Representative time-lapse live images and kymograph of RPE1 cell expressing GFP-C-tail1+2A in a NuMA-knockout background. After ablation (‘X’), C-tail1+2A is recruited to k-fiber minus-ends (arrowhead). The k-fiber stub slowly polymerizes, but its minus-end is not transported by dynein and remains detached from the spindle. Time is in min:s, with the frame captured immediately following ablation set to 00:00 s. Scale bars, 2 μm. Kymograph (right) taken along dashed line path. (E) Representative immunofluorescence images showing localization of dynactin (p150) after NuMA knockout and rescue with GFP-tagged NuMA truncations and chimeras in RPE1 cells. Constructs containing NuMA’s N-terminus recruit p150. Scale bar, 5 μm.

(A) Schematic maps of additional NuMA C-terminal tail truncations. (B) Representative immunofluorescence images showing localization of GFP-tagged NuMA truncations expressed in RPE1 cells in which endogenous NuMA has been knocked out. C-tail1 does not localize to microtubules and C-tail2A localizes all along spindle microtubules, while in combination (‘C-tail1+2A’, Figure 6) they localized at minus-ends. C-tail2B localizes to the cortex but not to microtubules. Scale bar, 5 μm.

Table 1.

NuMA’s pole-focusing ability correlates with dynactin- and minus-end-binding.

See also Figure 6 and Figure 6—figure supplement 1.

Knockout and rescue with NuMA version:	Recruits dynactin	Localizes to lattice (no end preference)	Localizes to preferentially to minus-ends	Rescues pole focusing
FL	+	-	+	+
N-C	+	-	+	+
N-Tau	+	+	-	-
C	-	-	+	-
C-tail1+2A	n.d.	-	+	-
C-tail2	n.d.	+	-	-

n.d.=not done, but unlikely to associate with dynactin given that the full C-terminus does not.

Figure 6—figure supplement 1.

NuMA ‘C-tail1’ and ‘C-tail2A’ alone do not localize to minus-ends.

(A) Schematic maps of additional NuMA C-terminal tail truncations. (B) Representative immunofluorescence images showing localization of GFP-tagged NuMA truncations expressed in RPE1 cells in which endogenous NuMA has been knocked out. C-tail1 does not localize to microtubules and C-tail2A localizes all along spindle microtubules, while in combination (‘C-tail1+2A’, Figure 6) they localized at minus-ends. C-tail2B localizes to the cortex but not to microtubules. Scale bar, 5 μm.

Video 7.

The ‘tail1+2A’ region of NuMA’s C-terminus is sufficient for minus-end localization.

See also Figure 6D. Live confocal imaging of a RPE1 cell expressing GFP-C-tail1+2A in a NuMA knockout background. After ablation (‘X’), GFP-C-tail1+2A is recruited to k-fiber minus-ends (arrowhead). The k-fiber stub slowly polymerizes, but its minus-end is not transported by dynein and remains detached from the spindle. Note that C-tail1+2A does not rescue spindle architecture; video shows one confocal slice of a three-dimensionally disorganized spindle. Also note that bright GFP signal from a neighboring interphase cell is noticeable on the left side of the video. Time is in min:s, with the frame captured immediately following ablation set to 00:00 s. Plus-end of ablated k-fiber is marked by ‘*’. Scale bar, 5 μm.

We sought to more closely define which sections within NuMA’s C-terminus (a.a. 1701–2115) are involved in microtubule minus-end localization. The second half of the C-terminus (‘C-tail2’, a.a. 1882–2115), which contains residues previously implicated in NuMA-microtubule interactions (Chang et al., 2017; Du et al., 2002; Gallini et al., 2016; Haren and Merdes, 2002), localized all along spindle microtubules with no minus-end preference (Figure 6B). Similarly, C-tail2A (a.a. 1882–1981) bound along the lattice, while C-tail1 (a.a. 1701–1881) did not bind microtubules (Figure 6—figure supplement 1). However, in combination (‘C-tail1+2A’, a.a. 1701–1981) they localized at minus-ends (Figure 6B). Indeed, the C-tail1+2A region was sufficient for recruitment to new minus-ends created by ablation, even in the absence of endogenous NuMA (Figure 6D; Video 7). Which pieces of the tail1+2A region are specifically required for minus-end localization, and what they each do, will require further dissection. Interestingly, related NuMA residues were recently shown to bind at both plus- and minus-ends and play a role in spindle orientation (Seldin et al., 2016). In sum, our data indicate that NuMA’s localization to spindle microtubule minus-ends is independent of dynein, independent of known direct minus-end binding proteins, and mediated by its C-terminal residues 1701–1981 (‘C-tail1+2A’).

Importantly, NuMA’s C-terminus (‘C’) localized to minus-ends but could not rescue proper pole focusing or spindle architecture, unlike full-length protein or N-C (Figure 6C). This suggests that NuMA’s function in spindle organization requires both minus-end binding (via its C-terminus) and the ability to recruit dynactin to minus-ends (via its N-terminus) (Kotak et al., 2012). Consistent with this hypothesis, NuMA’s C-terminus (‘C’) was unable to recruit dynactin to minus-ends, while its N-terminus and C-terminus fused (‘N-C’) did (Figure 6E; Table 1).

To test whether recruiting dynein-dynactin to the sides of microtubules was sufficient for proper microtubule clustering into poles, we fused NuMA’s N-terminus to the Tau microtubule-binding domain (‘N-Tau’). N-Tau localized along the length of spindle microtubules and recruited dynactin there (Figure 6E). Despite combining dynein-dynactin binding and microtubule lattice binding, N-Tau was not enriched at minus-ends and was unable to rescue spindle architecture (Figure 6C; Table 1).

Like a traditional cargo adaptor, NuMA may target force to spindle minus-ends using a cargo (minus-end) binding module (‘C’) and a dynactin-recruitment module (‘N’). Furthermore, the inability of N-Tau to rescue spindle architecture in the absence of endogenous NuMA suggests that specifically targeting dynein-dynactin to minus-ends, not just all along spindle microtubules as N-Tau does, is critical for organizing a focused, bipolar spindle.

Discussion

NuMA targets dynactin to minus-ends, spatially regulating dynein activity at mitosis

Our data indicate that NuMA localizes to spindle microtubule minus-ends independently of dynein and minus-end binding proteins γ-TuRC, CAMSAP1, and KANSL1/3. NuMA then targets dynactin to minus-ends, localizing dynein motor activity there. In addition, targeting dynein to the end of its track could permit amplification by motor pile-up, as NuMA at minus-ends captures processive dynein complexes. Altogether, our findings are consistent with a model in which NuMA confers minus-end targeting of the dynein-dynactin complex upon nuclear envelope breakdown (NEB), when NuMA is released from the nucleus. Indeed, active microtubule clustering by dynein is first observed coincident with NEB (Rusan et al., 2002). Thus, NuMA may provide both spatial (minus-end-specific) and temporal (mitosis-specific) regulation of dynein-powered force (Figure 7A).

Figure 7.

Model: NuMA spatially targets dynein activity to minus-ends at mitosis to ensure minus-end clustering into poles.

(A) Our findings suggest that NuMA confers minus-end targeting to the dynein-dynactin complex upon NEB, when NuMA is released from the nucleus. NuMA localization at minus-ends is mediated by a region of its C-terminus (amino acids 1701–1981, in red) that includes tail1 in addition to lattice-binding domain tail2A. NuMA targets dynactin to minus-ends, localizing dynein motor activity. (B) Model: Without minus-end targeting, dynein molecules acting on the red microtubule oppose each other, resulting in gridlock (left). NuMA-mediated targeting of force to minus-ends allows for productive clustering of the red microtubule into the pole (right). Altogether, the data support a model for mammalian spindle organization in which targeting poleward force to microtubule minus-ends specifically – by NuMA-mediated dynactin recruitment – provides robust clustering of microtubules into a focused, bipolar spindle.

The data also indicate that NuMA’s minus-end localization at mitosis is mediated by the tail1+2A region of its C-terminus, while previously identified microtubule-binding domains tail2A or tail2B (Du et al., 2002; Gallini et al., 2016; Haren and Merdes, 2002) are not sufficient. We propose three hypotheses for tail1+2A-mediated minus-end recognition. First, a yet-unidentified minus-end binding protein or the minus-end-directed kinesin HSET (Gaglio et al., 1996; Mountain et al., 1999) could localize NuMA via an interaction that requires this longer segment of the NuMA tail. During the preparation of this manuscript, ASPM was identified as a novel mammalian spindle minus-end binder, but ASPM deletion does not affect NuMA localization (Jiang et al., 2017). Second, NuMA may recognize both plus- and minus-ends via microtubule curvature sensing, as previously proposed (Seldin et al., 2016), and yet be actively excluded from plus-ends within the spindle by other proteins. Third, NuMA tail1+2A may bind minus-ends directly in cells, perhaps using post-translational modifications (Compton and Luo, 1995; Gallini et al., 2016; Magescas et al., 2017; Yan et al., 2015) not previously recapitulated in vitro.

Function of force at spindle microtubule minus-ends

Models of spindle assembly in silico predict that enriching a minus-end-directed motor at microtubule ends can break gridlock between parallel microtubules and allow robust minus-end clustering into poles (Burbank et al., 2007; Goshima et al., 2005; Hyman and Karsenti, 1996; Surrey et al., 2001). However, a lack of mechanistic information and tools made it difficult to test such hypotheses in vivo. The present study reveals a mechanism for direct recruitment of the motor to minus-ends, via NuMA, and is consistent with the prediction that targeting dynein-dynactin to the sides of microtubules is not sufficient for robust spindle organization. Both the minus-end-binding module (‘C’) and dynein-dynactin-binding module (‘N’) of NuMA are required for bipolar focusing, while the distance between them is not critical for function. Fusing the dynein-dynactin-binding module to the Tau microtubule binding domain is not sufficient, suggesting a requirement for minus-end-specific forces (Figure 6; Table 1). However, we cannot formally exclude that features of the NuMA C-terminus (missing from ‘N-Tau’) other than minus-end binding enable rescue of pole focusing, or that while N-Tau recruits dynactin (Figure 6D), it cannot effectively couple to dynein. Altogether, the work is consistent with a model for mammalian spindle organization in which targeting poleward force to microtubule minus-ends (by NuMA-mediated dynactin recruitment) is critical for organizing microtubules into a focused, bipolar array (Figure 7B). More broadly, the emergence of pole architecture at mitosis illustrates how spatial regulation of molecular-scale activities (like NuMA localizing dynein activity at individual minus-ends) can give rise to complex and diverse cellular-scale structures.

NuMA as mitotic dynein adaptor or activator

The data suggest that NuMA in the mammalian spindle body may function as a traditional cargo adaptor, with a cargo (minus-end)-binding module and a motor-binding module. In the spindle body context, microtubule minus-ends are the cargo, just as the cortex can be framed as cargo for cortical dynein – analogous to more canonical interphase cargo like membranous vesicles or organelles. This framework raises the question of whether NuMA additionally serves as a dynein activator, inducing highly processive motility like interphase dynein adaptors can (McKenney et al., 2014; Schlager et al., 2014).

Dynein’s localization in mammalian mitosis is more ubiquitous than that of dynactin (Figure 4C; Figure 2—figure supplement 1C); high cytoplasmic levels of dynein prevented us from detecting clear dynein enrichment at ablation-created minus-ends, for example (data not shown), and dynein localization is not as altered by NuMA knockout as dynactin localization (Figure 4C). These observations could be explained if NuMA and dynactin at minus-ends selectively activate dynein there – through a conformational shift that renders the motor more processive, for example – rather than simply selectively localizing it there. In other words, dynein’s localization within the spindle body may be less tightly regulated than its activity. The loss of dynein activity at minus-ends observed after NuMA knockout (Figure 3C) may stem from a lack of dynein activation (without NuMA and dynactin present at minus-ends) rather than a lack of dynein enrichment.

Unlike known dynein activators, NuMA is thought to not only homodimerize but also oligomerize into higher order assemblies (Harborth et al., 1999; Saredi et al., 1997). NuMA’s oligomerization ability suggests that it could assemble teams of motors and invites a comparison to motor-clustering in microdomains on large cellular cargoes, like phagosomes (Rai et al., 2016). The increased force and processivity provided by teams of NuMA-dynactin-dynein complexes on mitotic minus-ends could enable transport and clustering of minus-ends despite high loads and friction created by dense microtubule crosslinking and – in the case of k-fiber minus-ends – coupling to chromosomes.

Materials and methods

Cell culture and transfection

PtK2 cells were cultured in MEM (11095; Thermo Fisher, Waltham, MA) supplemented with sodium pyruvate (11360; Thermo Fisher), nonessential amino acids (11140; Thermo Fisher), penicillin/streptomycin, and 10% heat-inactivated fetal bovine serum (FBS) (10438; Thermo Fisher). RPE1 and HeLa cells were cultured in DMEM/F12 with GlutaMAX (10565018; Thermo Fisher) supplemented with penicillin/streptomycin and 10% FBS. For Tet-on inducible CRISPR-Cas9 cell lines, tetracycline-screened FBS (SH30070.03T; Hyclone Labs, Logan, UT) was used. Cell lines were not STR-profiled for authentication. All cell lines tested negative for mycoplasma. Cells were maintained at 37°C and 5% CO2 and were transfected with DNA using ViaFect (E4981; Promega, Madison, WI) 48 hr (RPE1/HeLa) or 72 hr (PtK2) before imaging.

Inducible CRISPR-Cas9 knockout cells

sgRNAs were designed against 5’ exons of NuMA, CAMSAP1, and KANSL1 using http://crispr.mit.edu. sgRNAs are listed in Key Resource Table. The plasmid used to express sgRNAs under control of the hU6 promoter (pLenti-sgRNA) was a gift from T. Wang, D. Sabatini, and E. Lander (Whitehead/Broad/MIT). An RPE1 cell line containing doxycycline-inducible human codon-optimized spCas9 was a gift from I. Cheeseman (Whitehead/MIT) and was generated as described in (McKinley et al., 2015) using a derivative of the transposon described in (Wang et al., 2014). We infected this inducible-spCas9 RPE1 cell line with each pLenti-sgRNA as described in (Wang et al., 2015) using virus expressed in HEK293T cells and 10 μg/mL polybrene and selected with 6 μg/mL puromycin. For each targeted gene, we tested 3 independent sgRNA sequences, each of which generated indistinguishable spindle phenotypes (data not shown), and picked one line for subsequent studies. Four days before each experiment, spCas9 expression was induced with 1 μM doxycycline hyclate.

Live imaging and laser ablation

For live imaging, cells were plated on glass-bottom 35 mm dishes coated with poly-D-lysine (MatTek Corporation, Ashland, MA) and imaged in a stage-top humidified incubation chamber (Tokai Hit, Fujinomiya-shi, Japan) maintained at 30°C and 5% CO2. To visualize tubulin, 100 nM siR-Tubulin dye (Cytoskeleton, Inc., Denver, CO) was added 2 hr prior to imaging, along with 10 µM verapamil (Cytoskeleton, Inc.). Under these conditions, there was no detected defect in spindle appearance or microtubule dynamics. As described elsewhere (Elting et al., 2014), cells were imaged using a spinning disk confocal inverted microscope (Eclipse Ti-E; Nikon Instruments, Melville, NY) with a 100 × 1.45 Ph3 oil objective through a 1.5X lens, operated by MetaMorph (7.7.8.0; Molecular Devices, Sunnyvale, CA). Laser ablation (30 3-ns pulses at 20 Hz) with 551 nm light was performed using the galvo-controlled MicroPoint Laser System (Andor, Belfast, UK). For laser ablation experiments, images were acquired more slowly prior to ablation and more rapidly after ablation (typically 7 s prior and 3.5 s after ablation).

Immunofluorescence and antibodies

For immunofluorescence, cells were plated on #1.5 25 mm coverslips coated with 1 mg/mL poly-L-lysine. Cells were fixed with 95% methanol + 5 mM EGTA at −20°C for 3 min, washed with TBS-T (0.1% Triton-X-100 in TBS), and blocked with 2% BSA in TBS-T for 1 hr. Primary and secondary antibodies were diluted in TBS-T+2% BSA and incubated with cells overnight at 4°C (primary) or for 20 min at room temperature (secondary). DNA was labeled with Hoescht 33342 (Sigma, St. Louis, MO) before cells were mounted in ProLongGold Antifade (P36934; Thermo Fisher). Cells were imaged using the spinning disk confocal microscope described above. Antibodies: mouse anti-α-tubulin DM1α (T6199; Sigma), rabbit anti-α-tubulin (ab18251; Abcam, Cambridge, UK), rabbit anti-NuMA (NB500-174; Novus Biologicals, Littleton, CO), mouse anti-p150-Glued (610473; BD Biosciences, San Jose, CA), mouse anti-α-tubulin DM1α conjugated to Alexa488 (8058S; Cell Signaling, Danvers, MA), mouse anti-dynein intermediate chain (MAB1618MI; Millipore, Billerica, MA), rabbit anti-EB1 (sc-15347; Santa Cruz Biotechnology, Dallas, TX), rabbit anti-KANSL1 (PAB20355; Abnova, Taipei City, Taiwan), rabbit anti-CAMSAP1 (NBP1-26645; Novus Biologicals), mouse anti-actin (MAB1501; Millipore), rabbit anti-γ-tubulin (T3559; Sigma), and camel nanobody against GFP coupled to Atto488 (gba-488; ChromoTek, Hauppauge, NY).

STORM

PtK2 cells expressing GFP-α-tubulin (gift of A. Khodjakov, Wadsworth Center) were plated on photo-etched, gridded coverslips (G490; ProSciTech, Kirwan, Australia) coated with 1 mg/mL poly-L-lysine (P-1524; Sigma) and imaged at 29–30°C in a homemade heated aluminum coverslip holder using the confocal microscope and ablation system described above. 20–30 s after k-fiber ablation, imaging media was replaced with fixative (as above) chilled to −80°C, and the coverslip holder was placed on ice for 1 min. Cells were incubated with 3% BSA in PBS for 1 hr at room temperature, and then with primary antibodies overnight at 4°C. Secondary antibodies ((anti-mouse Cy3B; Jackson Immunoresearch, West Grove, PA); anti-rabbit AF647 (Life Tech, Carlsbad,CA)) were incubated for 30 min at room temperature. Antibody incubations were followed by four washes with 0.2% BSA in PBS. Samples were stored in PBS during confocal imaging, and coverslip grid was used to re-find the individual ablated cell. For 3D STORM imaging, samples were mounted on glass slides with a standard STORM imaging buffer consisting of 5% (w/v) glucose, 100 mM cysteamine, 0.8 mg/mL glucose oxidase, and 40 µg/mL catalase in 1M Tris-HCI (pH 7.5) (Huang et al., 2008; Rust et al., 2006). Coverslips were sealed using Cytoseal 60. STORM imaging was performed on a homebuilt setup based on a modified Nikon Eclipse Ti-E inverted fluorescence microscope using a Nikon CFI Plan Apo λ 100x oil immersion objective (NA 1.45). Dye molecules were photoswitched to the dark state and imaged using either 647- or 560 nm lasers (MPB Communications, Montreal, CAN); these lasers were passed through an acousto-optic tunable filter and introduced through an optical fiber into the back focal plane of the microscope and onto the sample at intensities of ~2 kW cm−2. A translation stage was used to shift the laser beams towards the edge of the objective so that light reached the sample at incident angles slightly smaller than the critical angle of the glass-water interface. A 405 nm laser was used concurrently with either the 647- or 560 nm lasers to reactivate fluorophores into the emitting state. The power of the 405 nm laser (typical range 0–1 W cm−2) was adjusted during image acquisition so that at any given instant, only a small, optically resolvable fraction of the fluorophores in the sample were in the emitting state. Emission was recorded with an Andor iXon Ultra 897 EM-CCD camera at a framerate of 220 Hz, for a total of ~120,000 frames per image. For 3D STORM imaging, a cylindrical lens of focal length 1 m was inserted into the imaging path so that images of single molecules were elongated in opposite directions for molecules on the proximal and distal sides of the focal plane (Huang et al., 2008). Two-color imaging was performed via sequential imaging of targets labeled by AF647 and Cy3B. The raw STORM data was analyzed according to previously described methods (Huang et al., 2008; Rust et al., 2006).

Drug treatment and microtubule re-growth

To inhibit γ-tubulin, 30 μM gatastatin (gift of Takeo Usui and Ichiro Hayakawa, University of Tsukuba and Okayama University, respectively) (Chinen et al., 2015) was added 25–60 min before imaging (Figure 5A–C, Figure 5—figure supplement 1A). To test microtubule re-growth in the presence of gatastatin (Figure 5—figure supplement 1B), all cells were treated with 10 μM STLC for 6 hr to create monopolar spindles. For cold treatment, cells were then placed on ice; after 1 hr, 30 μM gatastatin or equivalent (0.1%) DMSO was added. After 10 more minutes on ice, cells were moved to room temperature for 1 min before fixation. Control cells (no ice) were incubated in media containing 30 μM gatastatin or DMSO for 11 min at room temperature before fixation. For all microtubule re-growth experiments after nocodazole washout, cells were treated with 5 μM nocodazole (M1404; Sigma) for 15 min at 37°C. After three washes, cells were incubated at room temperature for 6–8 min before fixation and immunofluorescence.

Plasmids

2xGFP-Arp1A was made by inserting EGFP from pEGFP-N1 (Clontech, Takara Bio USA, Mountain View, CA) by Gibson assembly between GFP and Arp1A of GFP-Arp1A (human Arp1A in a pBABE variant, Addgene 4432; gift from I. Cheeseman, Whitehead Institute) (Kiyomitsu and Cheeseman, 2012). 2x-GFP-Arp1A localized correctly to kinetochores and poles, and spindle organization was unperturbed. To make Cas9-resistant GFP-NuMA (‘GFP-NuMA_resistant’), full-length human NuMA (NM_006185.3) with silent mutations (5’-GTGTCAGAGAGACTGGACTTT-3’ mutated to 5’-GTTAGTGAACGCTTGGATTTT-3’, preserving amino acids 57–62 of NP_006176.2 (‘VSERLD’)) was synthesized and cloned (Epoch Life Science, Missouri City, TX) into pEGFP-N1 at BglII and EcoRI sites. Subsequent truncations of NuMA (‘N-C’, ‘C’, ‘C-tail1’, ‘C-tail2’, ‘C-tail2A’, ‘C-tail2B’) were synthesized and cloned (Epoch Life Science) into ‘GFP-NuMA_resistant’ at BglII and HindIII sites. To make GFP-N-Tau, NuMA amino acids 1–1410 from ‘GFP-NuMA_resistant’ followed by a flexible linker and MAPTau (NM_01684.1) from pmEmerald-MAPTau-C-10 (gift from M. Davidson, Florida State University) were synthesized and cloned (Epoch Life Science) into ‘GFP-NuMA_resistant’ at HindIII and SalI sites. Other plasmids used: DsRed-p150217-548 (CC1; amino acids 217–548 of chicken p150 in pDsRed-N1, Clontech, gift from T. Schroer, Johns Hopkins University) (Quintyne and Schroer, 2002); mCherry-p50 (chicken p50 in mCherry-C1, Clontech, gift from M. Moffert and T. Schroer, Johns Hopkins University) (Shrum et al., 2009); GFP-NuMA (human NuMA in pEGFP-N1, Clontech, gift from D. Compton, Dartmouth Medical School) (Kisurina-Evgenieva et al., 2004); GFP-CAMSAP1 (human CAMSAP1 in pEGFP-C1, Clontech, gift from A. Akhmanova, Utrecht University) (Jiang et al., 2014).

Data analysis

To determine the percentage of p150 at plus-ends vs. minus-ends (Figure 1B, Figure 4D), we used single microtubules where both ends were clearly visible. We found that EB1 consistently labeled just one end, the plus-end. We determined p150 localization relative to the EB1-labeled plus-end and calculated the percentage of p150 at each location within each cell. Percentages for multiple cells were averaged for Figures 1B and 4D. Pre-NEB cells were distinguished from post-NEB cells by the exclusion of microtubules from the nucleus, circle-shaped chromosome packing in the nucleus, and, when possible, NuMA localization within the nucleus.

Kymographs of GFP-Arp1A, GFP-NuMA, and GFP-CAMSAP1 puncta and pole position over time (Figure 1C–E, Figure 2C, Figure 6D) were generated in ImageJ (Version 2.0.0/1.51 hr). To measure GFP intensity at ablation-created minus-ends over time (Figure 1F, Figure 2D, Figure 2—figure supplement 1E, Figure 5B), we used a home-written Matlab (R2012a Version 7.4) program to integrate GFP intensity within a 1.4 μm-diameter circle centered on the manually-tracked k-fiber minus-end, and to measure local background intensity within a surrounding 2.7 μm-diameter ‘donut’. Code is available at https://github.com/chueschen/IntensityAtMinusEnd (Hueschen, 2017; copy archived at https://github.com/elifesciences-publications/IntensityAtMinusEnd). After background subtraction, the intensity measured at the cut site during the three frames before ablation (k-fiber intensity) was averaged and set to zero. For NuMA and Arp1A, we then fit a sigmoid function ( where y = intensity and x = time) to each trace, normalized to plateau height (a = 1), and solved for the time b at which y = 0.5*a to determine time to half-maximum intensity (Figure 1G, Figure 2E, Figure 2—figure supplement 1F, Figure 5C). For CAMSAP1, we normalized each trace to peak height (mean intensity from t = 5 s to t = 20 s) and found the first point at which intensity passed 0.5 to determine time to half-maximum intensity. Finally, to generate mean intensity traces, data from all traces were collected into 5 s wide bins in time and their intensities were averaged. Stub length (distance between k-fiber plus- and minus-ends, Figure 1H) was measured in ImageJ at the first frame following ablation.

Minus-end position data (Figure 3C) were generated by manual tracking of ablation-created k-fiber minus-ends (marked by GFP-CAMSAP1) and spindle poles in time-lapse videos, using a second home-written Matlab program (Elting et al., 2014). Nearest neighbor distances between NuMA puncta in STORM imaging (Figure 5H) were measured as the center-to-center distance from each NuMA puncta to its nearest neighboring puncta. NuMA truncation rescue capability (Figure 6C) reports the percentage of bipolar spindles with two focused poles compared to disorganized spindle architecture characteristic of NuMA knockout (detached centrosomes, loss of k-fiber focusing into two poles). Percentage was calculated for each experiment (n = 3–5 experiments) and then averaged.

Statistics

All data are expressed as average ±standard error of the mean (SEM). Calculations of correlation coefficients (Pearson’s r) and p values were performed in Matlab. One-way ANOVA and Tukey post-hoc tests (Figures 1G, 5C and 6C) were performed in Microsoft Excel and Matlab. All other p values were calculated using two-tailed unpaired t-tests with GraphPad Software. Quoted n’s are described in more detail in Figure Legends, but in general refer to individual biological structures analyzed (biological replicates, for example, individual spindle lengths, individual k-fiber ablations, etc.).

Image presentation

Time-lapse images (Figures 1C–E, 2C, 3A, 4A, 5D and 6D) show a single spinning disk confocal slice, as do immunofluorescence images of microtubule re-growth after nocodazole or cold treatment (Figures 1A–B and 4D, Figure 5—figure supplement 1B) and post-ablation confocal immunofluorescence images (Figure 5E, Figure 2—figure supplement 1A–B). 3D STORM images (Figure 5F–G) show a single 600 nm slice in Z. Immunofluorescence images of spindles (Figures 2F, 4B, C, 5A, 6B and E and Figure 1—figure supplement 1, Figure 2—figure supplement 1D, Figure 6—figure supplement 1B) show max intensity projections (1–2 μm in Z) of spinning disk confocal Z-stacks.

Video preparation

Videos show a single spinning disk confocal Z-slice imaged over time and were formatted for publication using ImageJ and set to play at 35x relative to real time. Videos were corrected to play at a constant frame rate, even when the acquisition rate was not constant.

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "NuMA Targets Dynein to Microtubule Minus-Ends at Mitosis" for consideration by eLife. Your article has been favorably evaluated by Anna Akhmanova (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors. The following individual involved in review of your submission has agreed to reveal their identity: Reto Gassmann (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

In this manuscript, Hueschen and co-workers use microscopy, laser ablation and molecular genetic techniques to investigate NuMA's role in regulating dynein and dynactin within the mitotic spindle. Their findings include the observation that NuMA localizes rapidly to newly created microtubule minus ends, and that this behaviour does not appear to depend on dynein, dynactin, or known minus-end binders γ-TuRC, CAMSAP1, and KANSL1/3. Conversely, the localization of dynactin (and, by inference, dynein) does depend on NuMA. The experiments suggest a specific region of NuMA autonomously recognizes minus ends, thereby recruiting dynein activity there. This work provides interesting evidence regarding the hierarchy of minus end association between NuMA, dynein, and dynactin, while raising a number of mechanistic puzzles and experimental queries.

This manuscript exhibits rigor in quantifying the localization kinetics of microtubule minus-end-binding proteins and in using knockout cell lines to perform clean molecular replacement experiments. The results are well presented with sound statistical analysis and fully support the authors' conclusions. The central idea of the study – NuMA targets dynein motor activity specifically to microtubule minus ends, and not vice versa as previously thought – is an important novel insight that will be of great interest to motor biochemists, developmental biologists and cell biologists.

Essential revisions:

1) Minus-end recognition

In several places in the manuscript, it is stated that NuMA "recognizes" minus ends. To many, this will imply a direct interaction, which is not explicitly tested in this work. The authors state that previous in vitro experiments with related NuMA fragments have not shown specific minus-end localization. An in vitro microtubule-binding experiment with the exact 'C-tail1+2A' construct used in this work would greatly strengthen the authors conclusions. Alternatively, instances of "recognizes" should be softened to "localizes to" or another expression that does not imply direct binding.

Further, discussion of the Sedlin et al. (2016) study could be expanded on. For example, if the 1811-1985 region of NuMA localizes to plus and minus ends as that paper suggests, is it conceivable that cellular factors are required to prevent plus-end binding? The notion of microtubule curvature recognition by NuMA is also raised in that paper. The authors should comment on these points.

2) NuMA localization under conditions of dynein inhibition or knockout

To investigate whether NuMA is localized to minus ends by direct recruitment or dynein transport, dynein activity is perturbed by p50 overexpression, p150 overexpression, and knockout of the dynein heavy chain. However, the kinetics of NuMA recruitment are only probed in detail in the p50 overexpression case. This provides the interesting observation that NuMA can still localize to minus ends when dynein activity is perturbed, but apparently with slower kinetics, suggesting that both direct recruitment and transport may contribute to NuMA localization. However, as the kinetic traces are noisy, it would be important to show the same trend in at least one of the other dynein inhibition conditions (i.e. perform laser ablation experiments, ideally in the dynein heavy chain knockout cell line).

The key point here is to strengthen the conclusion that dynein transport alone does not target NuMA to minus ends.

3) Figure 1: NuMA is shown to reach the spindle poles before dynactin, suggesting it recruits dynein/dynactin, not the other way round. There is no measurement of dynein. The authors should explain why dynein was not looked at.

4) Figure 2C reports that a DHC KO still leads to Numa and dynactin localization. This suggests that dynactin can be localized by NuMA independently of dynein. An important control is to show that the dynein level is completely knocked down by Western blot.

5) Figure 4 shows how NuMA targets dynactin. It would be helpful to the reader to include the dynein localization (currently in Figure 4—figure supplement 1) in this main figure. It explains why the authors focus on dynactin localization and do not directly address dynein localization.

6) Gatasin treatment

Do the authors have data to establish how effectively γ-TuRC was removed from minus ends by this treatment? Spindle length measurements do not probe this directly.

7) Super-resolution microscopy

A control with another minus-end binder (e.g. CAMSAP) needs to be included to provide convincing evidence for "organized clusters of NuMA puncta", rather than a lawn of molecules along the lattice near the minus-end.

8) Title: NuMA Targets Dynein to Microtubule Minus-Ends at Mitosis

There is a disconnect between title and the experiments in the study, as the localization data pertain mainly to dynactin rather than dynein. "Dynein" should be replaced with "Dynein and Dynactin" or "dynein activity".

There may also be some confusion with the use of "Targets" as some will feel dynein naturally targets minus end via its motor activity. Replacing with "Recruits" or "Localizes" may avoid this confusion.

9) The Materials and methods suggest that T-tests were used for statistical comparisons. However, because multiple comparisons are made, ANOVA and a suitable post-hoc test seems more appropriate and avoids the risk of Type I errors with multiple T-tests.

10) A 2xGFP-Arp1A construct is generated. Please state how the functionality of this construct in the dynactin complex was verified.

11) Figure 1A. As no EB1 signal is shown, the microtubule polarity is not clear in the example shown.

Also, please comment on the fidelity of EB1 labelling plus ends of acentrosomal microtubules i.e. do these microtubules ever have EB1 signal at both ends?

12) N-Tau experiments

The conclusion that dynein needs to be localized at the minus end, rather along the length of the microtubule, is based on use of an N-Tau construct. Please comment on caveats associated with this construct. For example, is the affinity between Tau and the microtubule load bearing? What is the expression level of the N-Tau construct compared to endogenous NuMA and other constructs?"

Essential revisions:

1) Minus-end recognition

In several places in the manuscript, it is stated that NuMA "recognizes" minus ends. To many, this will imply a direct interaction, which is not explicitly tested in this work. The authors state that previous in vitro experiments with related NuMA fragments have not shown specific minus-end localization. An in vitro microtubule-binding experiment with the exact 'C-tail1+2A' construct used in this work would greatly strengthen the authors conclusions. Alternatively, instances of "recognizes" should be softened to "localizes to" or another expression that does not imply direct binding.

Further, discussion of the Sedlin et al. (2016) study could be expanded on. For example, if the 1811-1985 region of NuMA localizes to plus and minus ends as that paper suggests, is it conceivable that cellular factors are required to prevent plus-end binding? The notion of microtubule curvature recognition by NuMA is also raised in that paper. The authors should comment on these points.

We appreciate the feedback that our language in certain places implied knowledge of a direct interaction between NuMA and minus-ends. We have carefully reviewed the text with this concern in mind and have softened our language to better convey our intended message: that our data do not distinguish between direct or indirect recruitment of NuMA to minus-ends (subsection “NuMA localizes to minus-ends independently of dynein”, last paragraph; subsection “NuMA is required for dynein activity and dynactin localization at minus-ends”, last paragraph; subsection “NuMA recognizes localizes to minus-ends independently of known minus-end binders”; subsection “NuMA function requires both minus-end-recognition and dynactin-recruitment modules”, second paragraph; subsection “NuMA targets dynactin to minus-ends, spatially regulating dynein activity at mitosis”, last paragraph; Figure 6B legend). We also thank the reviewers for the suggestion to expand our discussion of the Seldin et al. (2016) study. We have done so in the last paragraph of the subsection “NuMA targets dynactin to minus-ends, spatially regulating dynein activity at mitosis”.

2) NuMA localization under conditions of dynein inhibition or knockout

To investigate whether NuMA is localized to minus ends by direct recruitment or dynein transport, dynein activity is perturbed by p50 overexpression, p150 overexpression, and knockout of the dynein heavy chain. However, the kinetics of NuMA recruitment are only probed in detail in the p50 overexpression case. This provides the interesting observation that NuMA can still localize to minus ends when dynein activity is perturbed, but apparently with slower kinetics, suggesting that both direct recruitment and transport may contribute to NuMA localization. However, as the kinetic traces are noisy, it would be important to show the same trend in at least one of the other dynein inhibition conditions (i.e. perform laser ablation experiments, ideally in the dynein heavy chain knockout cell line).

The key point here is to strengthen the conclusion that dynein transport alone does not target NuMA to minus ends.

As suggested, we have performed laser ablation experiments in the inducible dynein heavy chain (DHC) knockout HeLa cell line. We measured GFP-NuMA recruitment to new minus-ends with and without deletion of DHC, and the data show the same trends as with and without p50 overexpression in PtK2 cells (Figure 2C-E). NuMA is robustly recruited to HeLa k-fiber minus-ends even after DHC deletion, but recruitment is slightly delayed. We have added these data to Figure 2—figure supplement 1. We thank the reviewers for strengthening the manuscript with this suggestion.

3) Figure 1: NuMA is shown to reach the spindle poles before dynactin, suggesting it recruits dynein/dynactin, not the other way round. There is no measurement of dynein. The authors should explain why dynein was not looked at.

We have added an explanation as suggested (subsection “Dynactin and NuMA display mitosis-specific, steady-state binding at microtubule minus-ends”, second paragraph). Thank you for this feedback. In brief, high cytoplasmic levels of dynein result in variable and low signal-to-background imaging of the motor on microtubules. This precluded accurate measurements of dynein’s dynamics.

4) Figure 2C reports that a DHC KO still leads to Numa and dynactin localization. This suggests that dynactin can be localized by NuMA independently of dynein. An important control is to show that the dynein level is completely knocked down by Western blot.

We agree with the reviewers and have added a western blot demonstrating dynein depletion after inducible DHC KO to Figure 2—figure supplement 1C. Furthermore, dynein depletion was always verified in individual cells analyzed – by spindle phenotype or, whenever possible, by immunofluorescence labeling of dynein.

5) Figure 4 shows how NuMA targets dynactin. It would be helpful to the reader to include the dynein localization (currently in Figure 4—figure supplement 1) in this main figure. It explains why the authors focus on dynactin localization and do not directly address dynein localization.

We thank the reviewers for this feedback and have moved the dynein localization into the main Figure 4, as suggested.

6) Gatasin treatment

Do the authors have data to establish how effectively γ-TuRC was removed from minus ends by this treatment? Spindle length measurements do not probe this directly.

To more directly test how effectively gatastatin blocks γ-TuRC’s interaction with minus-ends, we performed a microtubule regrowth assay in the presence of 30 μM gatastatin or equivalent DMSO (as in Chinen et al., Nat Comm 2016). After cold treatment, 30 μM gatastatin dramatically blocked γ-TuRC’s ability to bind α-tubulin to nucleate new microtubules (bottom panels, below). Gatastatin did not depolymerize existing microtubule networks (top panels), demonstrating its specificity in binding γ-tubulin, not α- or β-tubulin. We thank the reviewers for suggesting this important control and have included these data in Figure 5—figure supplement 1B. Details of the re-growth assay have been added to the manuscript’s Materials and methods.

7) Super-resolution microscopy

A control with another minus-end binder (e.g. CAMSAP) needs to be included to provide convincing evidence for "organized clusters of NuMA puncta", rather than a lawn of molecules along the lattice near the minus-end.

We thank the reviewers for this suggestion. We have performed 3D STORM of CAMSAP at PtK2 k-fiber minus-ends created by ablation, as done for NuMA in Figure 5D-H. In Author response image 1, we show examples of k-fiber minus-ends (white boxes) created by ablation and imaged by 3D STORM after immunofluorescence labeling of NuMA (top) or CAMSAP (bottom). In middle panels, structures are colored according to position in the Z-axis. Cartoons depict orientation of k-fiber and minus-end. Like NuMA, CAMSAP forms organized clusters of puncta, and distances between neighboring ‘nodes’ of both CAMSAP and NuMA (e.g., ‘*’) are ~50-150 nm (n = 32 nodes, 4 ablations for NuMA; n = 28 nodes, 3 ablations for CAMSAP). This spacing is consistent with measured spacing between individual microtubules within PtK2 k-fibers (McDonald et al., J. Cell Biol. 1992). These data support the idea of “organized clusters of NuMA puncta, rather than a lawn of molecules along the lattice near the minus-end”.

Author response image 1.

Unfortunately, limited signal from these minus-ends structures meant that we could only detect NuMA or CAMSAP clearly when labeled with the high-photon-yield and low-duty-cycle dye Alexa 647; this precluded two-color imaging of NuMA and CAMSAP co-localization in the same cell. In addition, while the large size and flat shape of mitotic PtK2 (rat kangaroo) cells made STORM experiments possible, commercially available CAMSAP1 antibodies raised against human antigen did not label any PtK2 proteins. The images in Author response image 1 show immunofluorescence staining with a CAMSAP2 antibody (17880-1-AP; Proteintech). In human cells (e.g., RPE1), this CAMSAP2 antibody does not label spindles, as human CAMSAP2 is phosphorylated at mitosis and does not interact with microtubules (Jiang et al., Dev Cell2014). Although these data support our claims, we do not include them in the manuscript due to the above limitations – and to avoid potential confusion looking forward.

8) Title: NuMA Targets Dynein to Microtubule Minus-Ends at Mitosis

There is a disconnect between title and the experiments in the study, as the localization data pertain mainly to dynactin rather than dynein. "Dynein" should be replaced with "Dynein and Dynactin" or "dynein activity".

There may also be some confusion with the use of "Targets" as some will feel dynein naturally targets minus end via its motor activity. Replacing with "Recruits" or "Localizes" may avoid this confusion.

We appreciate this feedback and have changed the title accordingly: NuMA Recruits Dynein Activity to Minus-Ends at Mitosis.

9) The Materials and methods suggest that T-tests were used for statistical comparisons. However, because multiple comparisons are made, ANOVA and a suitable post-hoc test seems more appropriate and avoids the risk of Type I errors with multiple T-tests.

We thank the reviewer for this suggestion. We conducted ANOVA analyses on all data in which multiple comparisons are made: Figure 1G (Arp1A, NuMA, and CAMSAP1 recruitment timescales to minus-ends after ablation); Figure 5C (NuMA recruitment timescales after inhibition or deletion of minus-end binding proteins); and, Figure 6C (spindle organization after rescue with NuMA truncations). Results are reported in Figure panels 1G, 5C, 6C and their corresponding figure legends, as well as below.

Figure 1G: To compare the timescales of Arp1A, NuMA, and CAMSAP1 recruitment to new minus-ends, we used a one-way ANOVA, which revealed a statistically significant difference between groups (F(2,38) = 9.26, p = 0.0005). A Tukey post hoc test showed a significant difference between recruitment timescales for Arp1A vs. NuMA (p = 0.03) and Arp1A vs. CAMSAP1 (p = 0.001). NuMA vs. CAMSAP1 differed at p = 0.10.

Figure 5C: A one-way ANOVA showed no statistically significant difference between NuMA recruitment timescales after gatastatin treatment, CAMSAP1 knockout, or KANSL1 knockout: F(3,42) = 0.45, p = 0.72.

Figure 6C: A one-way ANOVA revealed statistically significant differences in the percentage of cells showing focused, bipolar spindle architecture after rescue experiments with different NuMA truncations (F(3,10) = 267, p <0.00001). A Tukey post hoc test showed a significant difference between recruitment timescales for FL or N-C vs. C (p = 0.001) and FL or N-C vs. N-Tau (p = 0.001).

10) A 2xGFP-Arp1A construct is generated. Please state how the functionality of this construct in the dynactin complex was verified.

The 2xGFP-Arp1A construct localized correctly within the spindle (e.g., at poles and brightly at kinetochores without complete end-on attachments), just like 1xGFP-Arp1A, and it did not perturb spindle organization. We have added a sentence stating this in the Materials and methods (subsection “Plasmids”).

11) Figure 1A. As no EB1 signal is shown, the microtubule polarity is not clear in the example shown.

Also, please comment on the fidelity of EB1 labelling plus ends of acentrosomal microtubules i.e. do these microtubules ever have EB1 signal at both ends?

We thank the reviewers for this feedback. We have changed the corresponding text to make it clear that Figure 1B, not 1A, provides the information about microtubule polarity (subsection “Dynactin and NuMA display mitosis-specific, steady-state binding at microtubule minus-ends”, first paragraph). Interestingly, we see EB1 label plus-ends, not minus-ends, of single acentrosomal microtubules with high fidelity at mitosis. (Small bundles of microtubules, on the other hand, sometimes show EB1 labeling at both ends, presumably because they contain microtubules of mixed polarity.) In agreement with this specificity, we have never been able to detect evidence of microtubule polymerization at minus-ends after ablation in mitosis, at least in PtK2 cells. We have added a clarifying note on this topic to the Materials and methods (subsection “Data Analysis”, first paragraph).

12) N-Tau experiments

The conclusion that dynein needs to be localized at the minus end, rather along the length of the microtubule, is based on use of an N-Tau construct. Please comment on caveats associated with this construct. For example, is the affinity between Tau and the microtubule load bearing? What is the expression level of the N-Tau construct compared to endogenous NuMA and other constructs?"

We have added a discussion of this caveat: “we cannot formally exclude […] that while N-Tau recruits dynactin (Figure 6D), it cannot effectively couple to dynein.” In addition, other experimental and computational studies (e.g., Elie et al., Sci. Rep. 2015; Ahmadzadeh et al., Biophys. J.2015) suggest that Tau’s interaction with the microtubule can bear load.

To quantify the expression level of the GFP-N-Tau construct compared to full-length (FL) GFP-NuMA, we measured the average GFP intensity within the spindle region (normalized to cytoplasmic background intensity) for a confocal slice through the spindle center, for all cells used in our analysis of spindle organization (Figure 6C). Unfortunately, more ideal comparisons of integrated intensity within the whole spindle volume were not possible, given how these data were acquired. However, these average intensity measurements (Author response image 2) suggest that N-Tau expression was comparable or higher than full-length NuMA. Each data point represents an individual cell; n = 19 for FL NuMA and n = 16 for N-Tau. Thus, N-Tau’s inability to rescue spindle organization is not due to low protein expression. Comparison to endogenous levels was not possible, as the polyclonal NuMA antibody binds with unknown stoichiometry to N-Tau vs. NuMA. Due to these limitations, we do not report this data in the manuscript.

Author response image 2.

Key resources table

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Cell line (Potorous tridactylus) (male)	PtK2	T. Mitchison	ATCC Cat#CCL-56; RRID: CVCL_0514	kidney epithelial
Cell line (P. tridactylus) (male)	PtK2 GFP-tubulin	PMID: 12604591		kidney epithelial, stably expressing GFP-α-tubulin
Cell line (Homo sapiens) (female)	RPE1	ATCC	ATCC Cat#CRL-4000; RRID: CVCL_4388	retina, epithelial
Cell line (H. sapiens) (female)	RPE1 NuMA knockout	this paper		RPE1 with stably integrated spCas9 (Tet-On promoter) and NuMA sgRNA #2
Cell line (H. sapiens) (female)	RPE1 CAMSAP1 knockout	this paper		RPE1 with stably integrated spCas9 (Tet-On promoter) and CAMSAP1 sgRNA #1
Cell line (H. sapiens) (female)	RPE1 KANSL1 knockout	this paper		RPE1 with stably integrated spCas9 (Tet-On promoter) and KANSL1 sgRNA #3
Cell line (H. sapiens) (female)	HeLa dynein heavy chain (DHC) knockout	PMID: 28216383	cmk1a DYNC1H1 sgD1.1	RPE1 with stably integrated spCas9 (Tet-On promoter) and DHC sgRNA
Recombinant DNA reagent (plasmid)	GFP-Arp1A	I. Cheeseman	Addgene 4432	Progenitor: pBABE variant
Recombinant DNA reagent (plasmid)	2xGFP-Arp1A	this paper		Progenitor: GFP-Arp1A
Recombinant DNA reagent (plasmid)	DsRed-p150217-548 (CC1)	PMID: 12391026		Progenitor: pDsRed-N1 (Clontech)
Recombinant DNA reagent (plasmid)	mCherry-p50	PMID: 19196984		Progenitor: mCherry-C1 (Clontech)
Recombinant DNA reagent (plasmid)	GFP-CAMSAP1	PMID: 24486153		Progenitor: pEGFP-C1 (Clontech)
Recombinant DNA reagent (plasmid)	GFP-NuMA	PMID: 15561764		Progenitor: pEGFP-N1 (Clontech)
Recombinant DNA reagent (plasmid)	GFP-NuMA_resistant	this paper		Progenitor: GFP-NuMA. Invisible to NuMA sgRNA #2. NM_006185.3
Recombinant DNA reagent (plasmid)	GFP-NuMA 'N-C'	this paper		Progenitor: GFP-NuMA_resistant
Recombinant DNA reagent (plasmid)	GFP-NuMA 'C'	this paper		Progenitor: GFP-NuMA_resistant
Recombinant DNA reagent (plasmid)	GFP-NuMA 'C-tail1'	this paper		Progenitor: GFP-NuMA_resistant
Recombinant DNA reagent (plasmid)	GFP-NuMA 'C-tail2'	this paper		Progenitor: GFP-NuMA_resistant
Recombinant DNA reagent (plasmid)	GFP-NuMA 'C-tail2A'	this paper		Progenitor: GFP-NuMA_resistant
Recombinant DNA reagent (plasmid)	GFP-NuMA 'C-tail2B'	this paper		Progenitor: GFP-NuMA_resistant
Recombinant DNA reagent (plasmid)	GFP-N-Tau	this paper		Progenitors: GFP-NuMA_resistant; pmEmerald-MAPTau-C10 (M.Davidson)
Antibody	anti-α-tubulin (DM1α; mouse)	Sigma	Sigma T6199	IF (1:1000); WB (1:5000). RRID: AB_477583
Antibody	anti-α-tubulin (DM1α; mouse) conjugated to AF488	Cell Signaling	Cell Signaling 8058S	IF (1:200). RRID: AB_10860077
Antibody	anti-α-tubulin (rabbit)	Abcam	Abcam ab18251	IF (1:500). RRID: AB_2210057
Antibody	anti-NuMA (rabbit)	Novus Biologicals	Novus Biologicals NB500-174	IF (1:400); WB (1:1000). RRID: AB_10002562
Antibody	anti-p150-Glued (mouse)	BD Biosciences	BD Biosciences 610473	IF (1:400). RRID: AB_397845
Antibody	anti-dynein intermediate chain (mouse)	Millipore	Millipore MAB1618MI	IF (1:250); WB (1:250). RRID: AB_2246059
Antibody	anti-EB1 (rabbit)	Santa Cruz Biotech	Santa Cruz sc-15347	IF (1:100). RRID: AB_2141629
Antibody	anti-γ-tubulin (rabbit)	Sigma	Sigma T3559	IF (1:500). RRID: AB_477575
Antibody	anti-KANSL1 (rabbit)	Abnova	Abnova PAB20355	WB (1:500). RRID: AB_10984400
Antibody	anti-CAMSAP1 (rabbit)	Novus Biologicals	Novus Biologicals NBP1-26645	WB (1:500). RRID: AB_1852845
Antibody	anti-GFP (camel) conjugated to Atto488	ChromoTek	ChromoTek gba-488	IF (1:500). RRID: AB_2631434
Dye	siR-tubulin	Cytoskeleton, Inc.	Cytoskeleton Inc. CYSC-002	100 nM
Drug	verapamil	Cytoskeleton, Inc.	Cytoskeleton Inc. CYSC-002	10 μM
Drug	gatastatin	PMID: 26503935		30 μM
Drug	nocodazole	Sigma	Sigma M1404	5 μM
Drug	STLC	Sigma	Sigma 164739	10 μM
Sequence-based reagent	NuMA sgRNA #1	this paper		5'-ATGACACTCCACGCCACCCG-3'
Sequence-based reagent	NuMA sgRNA #2*	this paper		5'-AAGTCCAGTCTCTCTGACAC-3'
Sequence-based reagent	NuMA sgRNA #3	this paper		5'-ACAGCAAATCTTGAAGCAGC-3'
Sequence-based reagent	CAMSAP1 sgRNA #1*	this paper		5'-GCCGCGTCGTAGCGGTCCAG-3'
Sequence-based reagent	CAMSAP1 sgRNA #2	this paper		5'-CCGACAGTCTGTATAATATT-3'
Sequence-based reagent	CAMSAP1 sgRNA #3	this paper		5'CCGAATATTATACAGACTGT-3'
Sequence-based reagent	KANSL1 sgRNA #1	this paper		5'-GAGCCAGTTTGAACCGGATA-3'
Sequence-based reagent	KANSL1 sgRNA #2	this paper		5'-ACACCATATCCGGTTCAAAC-3'
Sequence-based reagent	KANSL1 sgRNA #3*	this paper		5'-GAGCCCGTTTTCCCCCATTG-3'

*Guide RNA expressed by cell line used for subsequent experiments, following initial verification of consistent spindle phenotypes.