Augmin prevents merotelic attachments by promoting proper arrangement of bridging and kinetochore fibers

  1. Valentina Štimac
  2. Isabella Koprivec
  3. Martina Manenica
  4. Juraj Simunić
  5. Iva M Tolić  Is a corresponding author
  1. Division of Molecular Biology, Ruđer Bošković Institute, Croatia

Abstract

The human mitotic spindle is made of microtubules nucleated at centrosomes, at kinetochores, and from pre-existing microtubules by the augmin complex. However, it is unknown how the augmin-mediated nucleation affects distinct microtubule classes and thereby mitotic fidelity. Here, we use superresolution microscopy to analyze the previously indistinguishable microtubule arrangements within the crowded metaphase plate area and demonstrate that augmin is vital for the formation of uniformly arranged parallel units consisting of sister kinetochore fibers connected by a bridging fiber. This ordered geometry helps both prevent and resolve merotelic attachments. Whereas augmin-nucleated bridging fibers prevent merotelic attachments by creating a nearly parallel and highly bundled microtubule arrangement unfavorable for creating additional attachments, augmin-nucleated k-fibers produce robust force required to resolve errors during anaphase. STED microscopy revealed that bridging fibers were impaired twice as much as k-fibers following augmin depletion. The complete absence of bridging fibers from a significant portion of kinetochore pairs, especially in the inner part of the spindle, resulted in the specific reduction of the interkinetochore distance. Taken together, we propose a model where augmin promotes mitotic fidelity by generating assemblies consisting of bridging and kinetochore fibers that align sister kinetochores to face opposite poles, thereby preventing erroneous attachments.

Editor's evaluation

This manuscript investigates how the protein augmin contributes to correct spindle architecture and chromosome segregation during cell division in cultured human cells. High-quality super-resolution imaging and functional studies provide new insight into the mechanism of proper chromosome attachment to the spindle microtubules. This is an important paper that will be of interest to researchers interested in cell biology and cell biophysics.

https://doi.org/10.7554/eLife.83287.sa0

Introduction

The mitotic spindle has a sophisticated architecture that enables it to accurately segregate chromosomes during cell division. It consists of three major classes of microtubules: kinetochore microtubules that form kinetochore fibers (k-fibers) connecting chromosomes to the spindle pole through kinetochores, midplane-crossing microtubules that form antiparallel arrays in the central part of the spindle, and astral microtubules that extend from the spindle poles towards the cell cortex (McIntosh, 2016; O’Toole et al., 2020; Prosser and Pelletier, 2017). During metaphase and early anaphase, the majority of midplane-crossing microtubule bundles are laterally attached to a pair of sister k-fibers resembling a bridge between them, which is why they are called bridging fibers (Kajtez et al., 2016; Vukušić et al., 2017). These fibers balance the tension between sister kinetochores and maintain the curved shape of the metaphase spindle (Kajtez et al., 2016; Polak et al., 2017; Tolić and Pavin, 2016). In addition to linking sister k-fibers, some midplane-crossing microtubules can also form connections between neighboring k-fibers (O’Toole et al., 2020).

Spindle microtubules in human somatic cells are generated by several nucleation mechanisms, including centrosome-dependent and augmin-dependent nucleation (Kirschner and Mitchison, 1986; Pavin and Tolić, 2016; Petry, 2016; Prosser and Pelletier, 2017; Wu et al., 2008; Zhu et al., 2008), with an addition of chromatin- and kinetochore-dependent nucleation as a third mechanism that contributes to the directional formation of k-fibers (Maiato et al., 2004; Sikirzhytski et al., 2018; Tulu et al., 2006). Centrosome-dependent nucleation was long thought to be predominant in spindle assembly; however, numerous studies revealed that a significant number of microtubules also arise from pre-existing microtubules, through augmin, an eight-subunit protein complex that serves as a recruiter of the γ-tubulin ring complex (γ-TuRC) required for microtubule nucleation (David et al., 2019; Goshima et al., 2008; Kamasaki et al., 2013; Lawo et al., 2009; Song et al., 2018; Uehara et al., 2009). Augmin-nucleated microtubules grow at an angle of 0-30o relative to the pre-existing microtubule (Kamasaki et al., 2013; Petry et al., 2013; Verma and Maresca, 2019) and show a directional bias towards kinetochores, resulting in the preserved polarity of the spindle once the initial kinetochore-microtubule attachments form (David et al., 2019; Kamasaki et al., 2013). Depletion of augmin complex in different cell types results in impairment of microtubule bundles within the spindle accompanied by the formation of long, curved bundles on the spindle periphery, loss of spindle bipolarity, shorter interkinetochore distance, chromosome misalignment, mitotic delays, and a higher incidence of aneuploidy and cytokinesis failure (Almeida et al., 2022; Hayward et al., 2014; Uehara and Goshima, 2010; Uehara et al., 2009; Wu et al., 2008; Zhu et al., 2008). Of the eight subunits in the complex, the two directly interacting subunits HAUS6 (hDgt6/FAM29A) and HAUS8 (hDgt4/Hice1) have been extensively studied because of their ability to interact with a γ-TuRC adapter protein NEDD1 and pre-existing microtubules, respectively (Song et al., 2018; Uehara et al., 2009). While previous studies mainly focused on the effect of augmin on astral and kinetochore microtubules, the effect on midplane-crossing microtubules remains largely unexplored (Almeida et al., 2022; Hayward et al., 2014; Song et al., 2018; Uehara et al., 2009; Uehara et al., 2016; Uehara and Goshima, 2010; Wu et al., 2008; Zhu et al., 2008). Recent electron tomography work on spindles in human cells showed that ends of midplane-crossing microtubules interact with the wall of kinetochore microtubules (O’Toole et al., 2020), indicating that augmin-dependent nucleation might play an important role in their formation.

Augmin depletion has previously been linked to higher incidence of segregation errors (Wu et al., 2008) and the appearance of lagging chromosomes (Almeida et al., 2022; Viais et al., 2021), which were connected to impaired brain development in a recent study (Viais et al., 2021). Homozygous loss of HAUS6 subunit of the augmin complex was also seen in several cancer types, such as sarcomas, pancreatic adenocarcinomas, gliomas, and glioblastomas (ICGC/TCGA, 2020, retrieved by using cBioPortal Cerami et al., 2012; Gao et al., 2013). However, the origin of segregation errors in augmin depletion remains largely unexplored due to extensive mitotic delays often experienced by these cells (Wu et al., 2008).

To explore how the augmin-dependent microtubule nucleation affects functionally distinct microtubule bundles and thereby mitotic fidelity, we depleted augmin in hTERT-RPE1 and HeLa cells and imaged them using stimulated emission depletion (STED) (Hell and Wichmann, 1994; Klar and Hell, 1999) and confocal microscopy. We show that the augmin complex plays a vital role in the formation of highly organized microtubules required for mitotic fidelity. Augmin depletion leads to a three-fold increase in segregation errors when the checkpoint is weakened. A significant number of lagging chromosomes following augmin depletion have a reduced tension and a large tilt with respect to the spindle axis in metaphase, which may facilitate the formation of merotelic attachments within the disorganized spindle area of augmin-depleted cells. The appearance of severely disordered microtubules occurs along with the strong reduction in the number of proper bridging microtubules connecting sister k-fibers. Interestingly, the interkinetochore distance after augmin depletion was larger for kinetochore pairs with bridging fibers than for those without, indicating a specific effect of the augmin-generated bridging fibers on interkinetochore tension. Taken together, we propose that augmin affects mitotic fidelity by forming highly organized microtubule arrangements consisting of two sister k-fibers connected by a bridging fiber. In these arrangements oriented parallel to the spindle axis, augmin-nucleated bridging fibers prevent erroneous kinetochore-microtubule attachments during metaphase, while augmin-nucleated k-fibers resolve them in anaphase.

Results

Augmin is vital for the formation of uniformly arranged units consisting of two sister k-fibers connected by a bridging fiber

To overcome the limitations of confocal microscopy and explore the relationship between different classes of microtubules within the crowded metaphase plate area, we performed stimulated emission depletion (STED) superresolution imaging (Hell and Wichmann, 1994; Klar and Hell, 1999) of microtubules within the bipolar metaphase spindles of human cells (Figure 1A–B, Figure 1—figure supplement 1). This enabled us to easily distinguish between k-fibers that start at kinetochores and midplane-crossing microtubules that pass through the central part of the spindle. HAUS6 or HAUS8 components of the augmin complex were depleted by siRNA in hTERT-RPE1 (hereafter referred to as RPE1) cells stably expressing CENP-A-GFP and immunostained for tubulin (Figure 1A, Figure 1—figure supplement 1). Augmin depletion was confirmed using immunocytochemistry and western blot analysis (Figure 2—figure supplement 1A-CFigure 2—source data 1).

Figure 1 with 1 supplement see all
Augmin ensures the proper formation of entities consisting of bridging fibers that connect two sister k-fibers.

(A) STED superresolution images of microtubules immunostained for α-tubulin (gray) in control (left), HAUS6- (middle), and HAUS8-depleted (right) RPE1 cells stably expressing CENP-A-GFP (rainbow, confocal). Images show maximum intensity projections of 6 central z-planes of metaphase spindles. Kinetochores are color-coded for depth from blue to red with the Spectrum LUT in ImageJ. (B) Insets of STED superresolution images of microtubules (gray) in spindle midzones of control (left), HAUS6- (middle) and HAUS8-depleted (right) cells. Next to each image is the schematic representation of the microtubules in the midzone, with white lines representing k-fiber microtubules and yellow lines representing midplane-crossing microtubules. Images show a single z-plane and do not correspond to midzones of spindles in panel (A). All images are adjusted for clarity based on the intensity of astral microtubules in each image (see Materials and methods). Scale bars, 2 µm.

In control cells, the vast majority of midplane-crossing microtubule bundles laterally attached to a pair of sister k-fibers and formed a bridging fiber between them, consistent with previous findings (Kajtez et al., 2016; Vukušić et al., 2017). These bridging fibers were nearly-parallel with respect to one another and the spindle axis. Additionally, a small portion of midplane-crossing microtubules formed a secondary connection between a k-fiber on one side and a non-sister k-fiber on the other side (O’Toole et al., 2020). In comparison with untreated cells, midplane-crossing microtubules after HAUS6 or HAUS8 depletion extended at a variety of angles, and were wavy and disordered, particularly in the inner part of the spindle close to the pole-to-pole axis. Strikingly, midplane-crossing microtubules less often formed bridging fibers that connect to sister k-fibers in cells depleted of HAUS6 or HAUS8. Instead, they formed more complex arrangements, primarily consisting of one or more connections between various k-fibers within the metaphase spindle. This was contrary to k-fibers which, even though often missing a bridge between them, appeared relatively similar to those in control cells (Figure 1B, Figure 1—figure supplement 1). Taken together, augmin is vital for the proper organization of midplane-crossing microtubules into uniformly arranged bridging fibers that connect two sister k-fibers and extend nearly-parallel to the spindle axis.

Augmin helps both prevent and resolve segregation errors through joint action of bridging and k-fibers

The appearance of disordered midplane-crossing microtubules in the metaphase plate area of spindles without augmin prompted us to investigate whether these microtubules affect mitotic fidelity. Augmin depletion has previously been linked to higher incidence of segregation errors (Almeida et al., 2022; Viais et al., 2021; Wu et al., 2008), but their origin remained largely unexplored due to extensive mitotic delays in augmin-depleted cells (Wu et al., 2008). To avoid mitotic delays, we performed live-cell confocal imaging for which we codepleted the checkpoint protein Mad2 together with HAUS6 to induce anaphase onset, and used Mad2-depleted cells (Mayr et al., 2007), which have only a few segregation errors, as a control (Figure 2A, Videos 1 and 2). To explore the mechanistic origin of segregation errors, we divided them into three distinct groups: misaligned chromosomes in which both kinetochores were found outside the metaphase plate just before anaphase onset, lagging chromosomes in which the kinetochore is visibly stretched and positioned in the central part of the spindle while other kinetochores are already separating, and other less common and diverse errors (Figure 2A–B, See Materials and methods).

Figure 2 with 1 supplement see all
Augmin-nucleated midplane-crossing microtubules prevent kinetochore tilt and thus merotelic attachments.

(A) Time-lapse images of RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP (rainbow, confocal) in Mad2-depleted cells (left) and Mad2/HAUS6-codepleted cells (right). Yellow arrows represent lagging kinetochores and orange arrows misaligned kinetochores. Kinetochores are color-coded for depth from blue to red with the 16 Colors LUT in ImageJ. (B) The number of segregation errors per cell (top) and the percentage of errors causing aneuploidy (bottom) in Mad2-depleted cells (gray) and Mad2/HAUS6-codepleted cells (dark orange, light orange, dark yellow, light yellow). All segregation errors (dark orange) are divided into three groups: misaligned (light orange), lagging (dark yellow) and other (light yellow). Schematic representations next to the graph represent misaligned kinetochores (top) and lagging kinetochore (bottom). The number of errors in Mad2-depleted cells - in total 46 errors in 22 out of 31 cells; 10 misaligned kinetochore pairs in 9 out of 31 cells; 26 lagging kinetochores in 16 out of 31 cells; 10 other errors in 9 out of 31 cells. Number of errors in Mad2/HAUS6-codepleted cells - in total 172 errors in 25 out of 30 cells; 21 misaligned kinetochore pairs in 11 out of 30 cells; 78 lagging kinetochores in 23 out of 30 cells; 73 other errors in 20 out of 30 cells. Aneuploidy in Mad2-depleted cells - in total 11/46 errors in 22 out of 31 cells; 2/10 misaligned kinetochore pairs in 9 out of 31 cells; 6/26 lagging kinetochores in 16 out of 31 cells; 3/10 other errors in 9 out of 31 cells. Aneuploidy in Mad2/HAUS6-codepleted cells - in total 47/172 errors in 25 out of 30 cells; 17/21 misaligned kinetochore pairs in 11 out of 30 cells; 35/78 lagging kinetochores in 23 out of 30 cells; 35/73 other errors in 20 out of 30 cells. (C) The distance of closer kinetochore to the proximal pole and (D) further kinetochore to the distal pole of misaligned kinetochore pairs in time for Mad2-depleted (gray) and Mad2/HAUS6-codepleted cells (orange). Values are shown as mean (dark line) and SEM (shaded areas). The insets show the positions of kinetochores (yellow) with respect to spindle poles (gray). (E) Univariate scatter plot of the interkinetochore distance of error-free kinetochore pairs (gray) and misaligned kinetochore pairs (orange) in Mad2/HAUS6-codepleted cells. N=30 cells and 120 error-free kinetochore pairs in Mad2-depleted cells and N=30 cells and 21 misaligned kinetochore pairs in Mad2/HAUS6-codepleted cells. (F) The percentage of lagging kinetochores in Mad2-depleted (top) and Mad2/HAUS6-codepleted cells (bottom) divided by their location with respect to the pole-to-pole axis into inner and outer (schematic representation shown as inset, see Materials and methods). (G) Univariate scatter plot of the interkinetochore distance of error-free and lagging kinetochore pairs in Mad2-depleted (gray) and Mad2/HAUS6-codepleted cells (yellow). (H) Univariate scatter plot of the angle that the error-free and lagging kinetochore pairs form with the pole-to-pole axis (tilt) in Mad2-depleted (gray) and Mad2/HAUS6-codepleted cells (yellow). N=31 cells and 124 error-free and 26 lagging kinetochore pairs from Mad2-depleted cells. N=30 cells and 120 error-free and 78 lagging kinetochore pairs from Mad2/HAUS6-codepleted cells. (I) The correlation of the tilt and the interkinetochore distance for Mad2-depleted (gray) and Mad2/HAUS6-codepleted cells (yellow). Inset shows schematic representation of the tilt (kinetochores are shown in yellow and spindle poles in gray). (J) The insets of kinetochore pairs with merotelic attachments in RPE1 cells stably expressing CENP-A-GFP (yellow, confocal) and immunostained for α-tubulin (gray, STED). (K) The insets of kinetochore pairs from cells as in (J) but exposed to cold treatment. Images in (J) and (K) are smoothed with 0.5-mm-sigma Gaussian blur and adjusted for clarity (see Materials and methods). Schematic representations in (J) and (K) are shown above the insets for better visualization of merotelic microtubule attachments. (L) The schematic representations of a kinetochore pair with amphitelic attachment in metaphase that does not cause any segregation errors during anaphase when augmin is present (top) and a kinetochore pair with merotelic attachment in metaphase that ends up as the lagging kinetochore during anaphase when augmin is not present (bottom). (E, G and H) Boxes represent standard deviation (dark gray), 95% confidence interval of the mean (light gray) and mean value (black). All results were obtained from three independent experiments. Statistical analysis (B (top) and E) Mann–Whitney U test; (B (bottom) and F) Fisher’s exact test; (G and H) ANOVA with the post-hoc Tukey test; (I) linear regression; p-value legend: <0.0001 (****), 0.0001–0.001 (***), 0.001–0.01 (**), 0.01–0.05 (*), ≥0.05 (ns). Scale bars, 2 µm.

Figure 2—source data 1

Immunoblot analysis of HAUS6 siRNA treatment efficiency in RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP.

Full unedited blots of all three independent experiments are shown.

https://cdn.elifesciences.org/articles/83287/elife-83287-fig2-data1-v2.pdf
Video 1
RPE1 cell stably expressing CENP-A-GFP and Centrin1-GFP (16-colors) following Mad2 depletion.

Kinetochores are color-coded for depth from blue to red with the 16 Colors LUT and noise was processed with the Despeckle function in ImageJ. Scale bar, 2 µm.

Video 2
RPE1 cell stably expressing CENP-A-GFP and Centrin1-GFP (16-colors) following Mad2/HAUS6 codepletion.

Kinetochores are color-coded for depth from blue to red with the 16 Colors LUT and noise was processed with the Despeckle function in ImageJ. Scale bar, 2 µm.

The treatment with Mad2 siRNA resulted in a total of 1.5±0.2 segregation errors per cell (all data are given as mean ± SEM). However, the effect was significantly more severe when Mad2 was codepleted with HAUS6, resulting in a total of 5.7±0.8 segregation errors per cell (Figure 2B). Tracking of sister kinetochores in live-cell videos revealed that the number of misaligned kinetochore pairs per cell was similar in both Mad2 depletion and Mad2/HAUS6 codepletion (Figure 2B), which is why we presume they appeared due to Mad2 depletion independently of HAUS6 depletion. Nevertheless, they were much more likely to missegregate in cells without HAUS6, where 80 ± 9% of kinetochore pairs jointly segregated into the same cell, compared to only 20 ± 13% in control cells (Figure 2B). Interestingly, lagging kinetochores were both more frequent and more likely to missegregate in cells with Mad2/HAUS6 codepletion, in which there were 2.6±0.4 lagging kinetochores per cell, and 45 ± 6% of all lagging kinetochores ultimately missegregated. In contrast, there were only 0.8±0.2 lagging kinetochore pairs per cell in Mad2 depletion, and 77 ± 8% of them segregated correctly (Figure 2B). Finally, errors classified as others were more frequent in cells with Mad2/HAUS6 codepletion when compared to those with Mad2 depletion, but not more prone to missegregation (Figure 2B).

As misaligned chromosomes appeared equally frequently in Mad2 depletion and Mad2/HAUS6 codepletion, but differed in their ability to correctly segregate, we used them as a tool to isolate the role of augmin in resolving segregation errors during anaphase. Consistent with previous findings (Uehara et al., 2009), both poleward movement of kinetochores during anaphase A and spindle elongation during anaphase B were reduced following Mad2/HAUS6 codepletion (Figure 2—figure supplement 1D-F). To analyze in detail the movement of misaligned kinetochore pairs, we tracked both kinetochores with respect to the proximal and distal pole, that is the pole which is closer to the misaligned kinetochore pair or the pole which is further away from it, respectively. The kinetochore closer to the proximal pole approached the proximal pole during anaphase A both after Mad2 depletion and Mad2/HAUS6 codepletion, thereby moving towards the pole to which it should segregate in both cases (Figure 2C). However, the kinetochore further away from the proximal pole usually remained stagnant for a short period of time, and afterwards typically moved towards the distal pole and accurately segregated as a ‘lazy’ kinetochore (Sen et al., 2021) in Mad2-depleted cells. In contrast, the kinetochore further away from the proximal pole experienced a short stagnation period and then typically moved away from the pole to which it should segregate, thereby missegregating in Mad2/HAUS6-codepleted cells (Figure 2D, Figure 2—figure supplement 1G-H). As the interkinetochore distance of both correctly and incorrectly segregating misaligned kinetochore pairs in Mad2/HAUS6-codepleted cells was similar (Figure 2E), the absence of biorientation is unlikely to be the cause of missegregation for these kinetochores. Instead, missegregation likely occurs due to k-fibers with fewer microtubules (Almeida et al., 2022; Uehara et al., 2009; Zhu et al., 2008) creating insufficient force to move the kinetochore towards the distal pole (Dudka et al., 2018), against the movement of neighboring kinetochores and the corresponding chromosome mass which travel towards the proximal pole (Figure 2A, Video 2).

Whereas lagging chromosomes have been previously observed following augmin depletion (Almeida et al., 2022; Viais et al., 2021; Wu et al., 2008), their origin remains unknown. Because we observed that disorganized midplane-crossing microtubules were often concentrated in the inner part of the mitotic spindle near the main spindle axis upon augmin depletion (Figure 1B, Figure 1—figure supplement 1), we decided to investigate the spatial distribution (Figure 2F) of lagging kinetochores in Mad2-depleted and Mad2/HAUS6-codepleted cells to see if their increased number might be connected to this phenotype. In Mad2-depleted cells, all lagging kinetochore pairs were situated in the outer half of the spindle just before anaphase onset. Remarkably, in Mad2/HAUS6-codepleted cells, 19 ± 5% of all lagging kinetochore pairs were situated in the inner part of the spindle just before anaphase onset (Figure 2F), where disordered midplane-crossing microtubules most frequently appeared. Thus, organization of midplane-crossing microtubules into bridging fibers might play an important role in mitotic fidelity.

To further explore how this compromised spindle geometry affects lagging kinetochores, we measured their interkinetochore distance just before anaphase onset. The lagging kinetochore pairs in Mad2/HAUS6-codepleted cells had an interkinetochore distance of 0.89±0.03 µm, which was significantly smaller than the interkinetochore distance of 0.98±0.02 µm measured in error-free kinetochore pairs (Figure 2G). This was not the case for lagging kinetochore pairs in Mad2-depleted cells which had an interkinetochore distance of 1.10±0.05 µm, similar to the interkinetochore distance of 1.05±0.02 µm measured in error-free kinetochore pairs (Figure 2G). Distinctly reduced interkinetochore distance of lagging kinetochore pairs following Mad2/HAUS6 codepletion suggests that they appear due to compromised spindle architecture being unable to maintain adequate kinetochore tension.

As sister k-fibers in the disorganized spindle region were sometimes diagonally positioned with respect to the pole-to-pole axis, we decided to test if this tilt is also connected to the appearance of segregation errors by measuring the angle that either lagging or error-free kinetochores form with the spindle axis just before anaphase onset. As for the interkinetochore distance following Mad2/HAUS6 codepletion, lagging kinetochore pairs were different from error-free kinetochore pairs, the tilt of which was 19.7±1.6° and 10.9±0.8°, respectively (Figure 2H). In contrast, in Mad2-depleted cells with preserved spindle geometry there was no difference between lagging kinetochore pairs with the tilt of 8.2±1.5° and error-free kinetochore pairs with the tilt of 8.9±0.6°, which was also similar to the tilt of 10.9±0.8° measured for error-free kinetochore pairs following Mad2/HAUS6 codepletion (Figure 2H). The tilt of kinetochores inversely correlated with the interkinetochore distance in augmin depletion, but not in control cells (Figure 2I). These data indicate the importance of nearly parallel configuration of kinetochore pairs during metaphase for mitotic fidelity, and points to the augmin-specific cause of lagging kinetochores that likely arise due to compromised and tilted bundle architecture facilitating the formation of merotelic attachments. Indeed, we found merotelic attachments in HAUS6-depleted cells imaged using STED microscopy, with most kinetochores forming an attachment with the microtubule from the opposite side of the mitotic spindle, while missing a proper bridging fiber (Figure 2J). To further test the nature of the observed attachments, we combined STED microscopy with cold treatment to remove midplane-crossing microtubules and preserve only kinetochore microtubules (DeLuca et al., 2006; Sacristan et al., 2018; Silkworth et al., 2012), which allowed us to confirm their true merotelic nature (Figure 2K). Interestingly, our live-cell imaging experiments reveal that augmin is required not only to prevent the formation of merotelic attachments in metaphase, but also to resolve them in anaphase, as a larger percentage of lagging kinetochore pairs ends up missegregating in Mad2/HAUS6 codepletion than in Mad2 depletion (Figure 2B). This suggests that the insufficient force provided by k-fibers with fewer microtubules (Dudka et al., 2018), which is responsible for missegregation of misaligned kinetochore pairs following augmin depletion, also leads to inability to resolve merotelic attachments during anaphase.

Altogether, we propose that augmin ensures mitotic fidelity through the joint action of bridging and k-fibers. While augmin-nucleated bridging fibers prevent merotelic attachments by creating a nearly parallel and highly bundled spindle geometry unfavorable for creating additional attachments, augmin-nucleated k-fibers produce robust force required to resolve any potentially appearing errors during anaphase (Figure 2L).

Bridging fibers are predominantly generated through augmin-dependent nucleation

As our visual assessment revealed that spindles without augmin have disorganized arrangements of midplane-crossing microtubules and often lack proper bridging fibers (Figure 1B), which were also missing at kinetochore pairs that formed merotelic attachments (Figure 2J), we set out to analyze how augmin-dependent microtubule nucleation contributes to the formation of bridging fibers in immunostained RPE1 cells imaged using STED microscopy (Figure 3A–B). Bridging fibers were strictly defined as midplane-crossing microtubules that connect two sister k-fibers, whereas k-fibers were defined as microtubules that start at kinetochores (Figure 3C). We measured tubulin signal intensity of randomly selected bridging (Ib) and k-fibers (Ik) which had no other microtubules in their immediate neighborhood, in a small square region between two kinetochores or at the pole-side of kinetochore, respectively (Figure 3C, see Materials and methods). By using the resulting tubulin signal intensities, we first estimated the number of microtubules in the bridging fiber in untreated RPE1 cells. Electron tomography of spindles in RPE1 cells showed that k-fibers consist of nk = 12.6 ± 1.7 microtubules (O’Toole et al., 2020). Thus, the bridging fiber consists of nb = Ib × nkIk = 3.8 ± 0.4 microtubules (for the explanation of Ib and Ik calculations, see Materials and methods). The accuracy of this calculation was additionally tested by measuring the intensity of astral microtubules, which presumably consist of single microtubules (McDonald et al., 1992). Indeed, using the number of microtubules in the k-fiber, our measurement of astral microtubule intensities showed that the astral microtubules consist of na = 1.0 ± 0.1 microtubules (Figure 3D).

Figure 3 with 1 supplement see all
Augmin is crucial for the nucleation of bridging microtubules.

(A) The insets of kinetochore pairs in RPE1 cells stably expressing CENP-A-GFP (not shown) immunostained for α-tubulin (gray, STED) in control cells (left) and after HAUS6 (middle) or HAUS8 (right) depletion. The insets demonstrate kinetochore pairs with bridging fibers affected by HAUS6 or HAUS8 depletion compared to bridging fibers in control cells. The positions of kinetochores are marked with yellow circles. (B) The schematic representation of three possible pathways of microtubule nucleation: (1) centrosome-dependent (2) augmin-dependent and (3) chromatin- and kinetochore-dependent nucleation. The augmin complex is shown in yellow. (C) Top: the schematic representation of the mitotic spindle in metaphase and the method used to measure the tubulin intensity of the astral microtubules. Small square regions were measured on microtubules extending from the spindle pole, corresponding to astral microtubules. Their background was measured in the empty area between the two astral microtubules, and it was subtracted from astral microtubule intensity. Bottom: Schematic representation of the method used to measure the tubulin intensity of the bridging and k-fiber. Small square regions were measured between two kinetochores or right next to the kinetochore, corresponding to bridging and k-fibers, respectively. The intensity of k-fibers was measured as an average of two sister k-fibers, and the average value of the background within the spindle was subtracted from all measurements. Ia+bcg = intensity of astral microtubules with background, Ik+bcg = intensity of k-fibers with background, Ib+bcg = intensity of bridging microtubules with background, Ibcg = intensity of background. (D) Univariate scatter plot of tubulin signal intensities of astral microtubules in control cells (reference value, dark gray, Ia), and bridging fibers (Ib) and k-fibers (Ik) in control cells (gray), HAUS6- (yellow) and HAUS8-depleted cells (orange). (E) The reduction of tubulin signal in the bridging fiber (Ib) and the k-fiber (Ik) following HAUS6 (yellow) or HAUS8 (orange) depletion, values are shown as mean ± SEM. p-Values were calculated using the absolute values of tubulin signal intensity of bridging or k-fibers following HAUS6 or HAUS8 depletion, compared to the absolute values of tubulin signal intensity of corresponding fibers in control cells. (D) and (E) N=30 cells and 90 astral microtubules in control cells, 158 bridging and sister k-fibers in control and 180 bridging and sister k-fibers in HAUS6- and HAUS8 siRNA-treated cells. (F) STED superresolution images of microtubules stained for α-tubulin (gray) in RPE1 cells stably expressing CENP-A-GFP (rainbow, confocal) in control cells (left) and HAUS6 siRNA-treated cells (right) exposed to cold treatment. The images are maximum intensity projections and kinetochores are color-coded for depth from blue to red with the Spectrum LUT in ImageJ. (G) Univariate scatter plot of the tubulin signal intensities of k-fibers in control cells (gray) and upon HAUS6 depletion (yellow) in cells exposed to cold treatment. N=30 cells and 101 bundles in control cells and 102 bundles in HAUS6-depleted cells. (H) The fractions of kinetochore pairs with bridging fibers (dark gray) and with undetectable bridging fibers (light gray) in control, HAUS6- and HAUS8-depleted cells. Kinetochore pairs are divided based on their location in the spindle into outer and inner (See Materials and methods and Results). (D) and (G) Boxes represent standard deviation (dark gray), 95% confidence interval of the mean (light gray) and mean value (black). All results were obtained from three independent experiments. Statistical analysis (D and E) ANOVA with post-hoc Tukey test, (G) Mann–Whitney U test, (H) chi-square test; p-value legend:<0.0001 (****), 0.0001–0.001 (***), 0.001–0.01 (**), 0.01–0.05 (*), ≥0.05 (ns). All images are adjusted for clarity (see Materials and methods). Scale bars, 2 µm.

Quantification of STED images further revealed that HAUS6 depletion resulted in 68 ± 8% reduction of the bridging fiber signal intensity and 24 ± 6% reduction of the k-fiber signal intensity, with similar results obtained by HAUS8 depletion (Figure 3D–E). These data indicate that augmin depletion affects not only k-fibers, but even more so bridging fibers. The contribution of augmin to the nucleation of k-fibers was independently tested by measuring their intensity in spindles exposed to cold treatment in which bridging fibers are removed (Figure 3F). HAUS6 depletion resulted in a 37 ± 5% reduction of the k-fibers (Figure 3G), which is consistent with a previous study (Zhu et al., 2008) and comparable to values under non-cold conditions. Based on the measurements under non-cold conditions, we estimate that after HAUS6 depletion bridging fibers consist of 1.2±0.7 microtubules and k-fibers of 9.6±1.5 microtubules, which we interpret as microtubules nucleated in an augmin-independent manner. Thus, 2.6±0.7 microtubules in the bridging fiber and 3.0±0.9 microtubules in the k-fiber are nucleated in an augmin-dependent manner.

Remarkably, 41 ± 4% of all kinetochore pairs in HAUS6-depleted cells had no detectable bridging fibers, defined as those with the tubulin signal below the background signal (see Materials and methods), and consistent with results obtained using visual inspection (Figure 3—figure supplement 1A). The majority of kinetochore pairs without bridging fibers were located in the inner part of the mitotic spindle, where as much as 50 ± 5% of all kinetochore pairs had undetectable bridging fibers after augmin depletion, compared to only 27 ± 5% in the outer part (Figure 3H), thus pointing to an irregular and more complex spatial distribution of bridging fibers in the inner part of the spindles following augmin depletion. Similar results were obtained from superresolution imaging after HAUS8 depletion (Figure 3H). To further validate our results, we performed live-cell confocal imaging with SiR-tubulin (Lukinavičius et al., 2014) and analyzed the spindles by two independent methods (Figure 3—figure supplement 1A–H, See Materials and methods). Altogether, these results reveal that the augmin complex is a major nucleator of bridging fibers, whereas its contribution to the formation of k-fibers is significant but less prominent.

The compromised microtubule nucleation following augmin depletion led to the impairment of overall spindle geometry, creating a unique system where three main types of interactions between k-fibers and bridging fibers can be found within the same spindle: (1) sister k-fibers attached to bridging fibers, (2) sister k-fibers without a bridging fiber, and (3) solitary, long, interpolar bundles without associated kinetochores. This is in contrast with control cells, where the first group dominates and the other two groups are rarely found (Polak et al., 2017). To gain insight into the contribution of each of these functionally distinct microtubule bundles to the maintenance of spindle geometry, we traced the outermost bundles in HAUS6 siRNA-treated RPE1 cells imaged using STED microscopy and fitted a circle to the bundle outline (Figure 3—figure supplement 1, see Materials and methods). Whereas the bundles without kinetochores in HAUS6 siRNA-treated cells had a significantly longer contour when compared to all other bundle types (Figure 3—figure supplement 1J), k-fibers without bridging fibers in augmin-depleted cells had a significantly larger radius of curvature than any of the other bundle types in augmin-depleted or control cells (Figure 3—figure supplement 1K). Taken together, the outer interpolar bundles without associated kinetochores are excessively long and make the spindle wider, whereas k-fibers lacking a bridging fiber are overly straight, ultimately resulting in a diamond-like shape of the spindle. This change in spindle shape in the absence of proper bridging fibers is consistent with the prediction of our theoretical model (Kajtez et al., 2016) and previous experiments (Jagrić et al., 2021).

In addition to spindle architecture, compromised microtubule nucleation following augmin depletion also affected spindle dynamics, as poleward flux in U2OS cells stably expressing CENP-A-GFP, mCherry-tubulin and photoactivatable-GFP-α-tubulin was significantly reduced (Figure 3—figure supplement 1L-M), in agreement with findings in Indian Muntjac cells (Almeida et al., 2022). Recent speckle microscopy experiments in RPE1 cells, which were able to separate the effect of augmin on poleward flux of bridging and k-fibers, revealed that both k-fibers and the remaining bridging fibers were significantly slowed down (Risteski et al., 2022). Bridging fibers fluxed faster than k-fibers in control and augmin-depleted cells (Risteski et al., 2022), supporting the model in which poleward flux is largely driven by sliding apart of antiparallel microtubules (Brust-Mascher et al., 2009; Mitchison, 2005; Miyamoto et al., 2004). We propose that augmin depletion results in slower flux of bridging fibers because the remaining bridging microtubules are likely nucleated at the poles, where microtubule depolymerization mechanisms might curb poleward flux speed (Ganem et al., 2005). In contrast, PRC1 depletion does not affect the flux (Risteski et al., 2022; Steblyanko et al., 2020) even though it reduces bridging fibers (Kajtez et al., 2016; Polak et al., 2017), possibly because the remaining bridging microtubules are generated away from the poles via augmin and can thus flux freely. In sum, augmin ensures proper architecture and dynamics of the metaphase spindle largely through the nucleation of bridging fibers, which link sister k-fibers and ensure their proper shape and function.

Augmin-depleted spindles contain fewer overlap bundles, which have longer overlap regions and are located at the spindle periphery

Our finding that bridging fibers were more severely perturbed in the inner part of the spindle after augmin depletion prompted us to examine the spatial distribution of these midplane-crossing microtubules and their overlap regions throughout the spindle. We used protein regulator of cytokinesis 1 (PRC1) as a marker because it preferentially crosslinks overlap microtubules (Li et al., 2018; Mollinari et al., 2002), thus providing a specific label for bridging fibers (Polak et al., 2017). By taking a standard ‘side view’ of the spindle and rotating the 3D image stack of the spindle into an ‘end-on’ view, we were able to gain insight into the redistribution of bridging microtubules throughout the spindle cross-section in HeLa (Kajtez et al., 2016) and RPE1 (Asthana et al., 2021) cells stably expressing PRC1-GFP with and without MG-132 treatment (Figure 4A–B, Figure 4—figure supplement 1A). To compare their distribution to that of tubulin, we also rotated the 3D image stacks of the spindles in RPE1 cells stained with SiR-tubulin (Figure 4A–B, Figure 4—figure supplement 1B; Novak et al., 2018).

Figure 4 with 1 supplement see all
Augmin-depleted spindles have fewer bridging fibers, which have larger overlap length and are located at the spindle periphery.

(A) The four columns on the left represent live images of metaphase spindles in untreated HeLa or MG-132-treated RPE1 cells stably expressing PRC1-GFP (yellow) and stained with SiR-DNA (blue) in control cells (top rows) and after HAUS6 depletion (bottom rows). 1st and 2nd column: side view of the spindle; 3rd and 4th column: end-on view of the same spindle, showing a barrel-like arrangement of PRC1-labeled bundles after augmin depletion. Images on the right show the end-on view of RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP (both in blue) and stained with SiR-tubulin (gray) in control cells (top) and after HAUS6 depletion (bottom). Side views are sum intensity projections of 5 central z-slices (∆z=0.5 µm) in HeLa cells and 10 central z-slices for RPE1 cells. End-on views are sum projections of 10 central z-slices (∆z=0.083 µm) for HeLa and 20 central z-slices for RPE1 cells. (B) Left: schematic representations of different views of the spindle. Eye signs mark the angle for the side view (1) and the end-on view (2). Side view was used to measure the length of overlap regions (yellow) and end-on view to determine the number of bundles (yellow dots). Right: schematic representation of the end-on view of RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP (blue dots) and stained with SiR-tubulin (gray dots). (C) The reduction of the PRC1 signal in RPE1 cells treated with MG-132 measured in sum intensity projection of 10 central z-slices following HAUS6 depletion. Values are shown as mean ± SEM. P-values were calculated using the absolute values of PRC1 signal intensity following HAUS6 depletion (N=39 cells), compared to the absolute values of PRC1 signal intensity in control cells (N=32 cells). (D) Univariate scatter plot of the number of bundles in RPE1 cells treated with MG-132 counted in the end-on view of the spindle in control cells (gray) and HAUS6-depleted cells (yellow). N=32 control cells and N=39 HAUS6-depleted cells. (E) The PRC1-GFP intensity profiles in RPE1 cells treated with MG-132 measured in the end-on view of the spindle in control cells (gray) and after HAUS6 depletion (yellow). The blue line in the inset marks the measured region (width: 2.5 µm). Mean (thick lines) and SEM (shaded areas). (F) Univariate scatter plot of overlap length divided by spindle length in RPE1 cells treated with MG-132 measured in the side view of the spindle in control cells (gray) and HAUS6-depleted cells (yellow). N=75 bundles in 32 control cells and N=74 bundles from 39 HAUS6-depleted cells. (D and F) Boxes represent standard deviation (dark gray), 95% confidence interval of the mean (light gray) and mean value (black). All results were obtained from three independent experiments. Statistical analysis (C, D and F) Mann–Whitney U test; p-value legend:<0.0001 (****), 0.0001–0.001 (***), 0.001–0.01 (**), 0.01–0.05 (*), ≥0.05 (ns). All images are adjusted for clarity so that all PRC1 bundles are visible in each cell (see Materials and methods). Scale bars, 2 µm.

The signal intensity of PRC1-GFP bundles in RPE1 cells was reduced by 55 ± 4% following augmin depletion (Figure 4C). Consistently, the number of PRC1-labeled overlap bundles measured in an end-on view of spindles was almost halved; from 28±1–16±1 distinct bundles in control and HAUS6-depleted RPE1 cells, respectively (Figure 4D). Comparable trends were also observed in HeLa cells after depletion of HAUS6 or HAUS8 (Figure 4—figure supplement 1C-G).

The augmin-depleted cells showed a specific barrel-like distribution of the PRC1-GFP labeled bundles, with more overlap bundles being present around the perimeter of the spindle and fewer in the central part (Figure 4A ‘end-on view’ and 4D, Figure 4—figure supplement 1H-I). However, DNA was uniformly distributed throughout the spindle cross-section, both in augmin-depleted and control cells (Figure 4A ‘end-on view’). In agreement with this result, kinetochores and tubulin signal were also found uniformly distributed over the spindle cross-section (Figure 4A ‘end-on view’ of RPE1 cells). This observation indicates that k-fibers are present and roughly uniformly distributed throughout the spindle cross-section and is in agreement with our finding that augmin primarily affects bridging fibers, while k-fibers are less perturbed (Figure 3).

To explore the role of the observed overlap repositioning in defining the overall spindle geometry, we measured spindle width, the diameter of the metaphase plate, spindle length, and overlap length in RPE1 and HeLa cells (see Materials and methods). Despite the spindles being wider in both cell lines, the diameter of the metaphase plate was not larger, as the spindles widened due to the long, curved bundles without kinetochores (Figure 4—figure supplement 1J). While the spindles in RPE1 cells shortened following augmin depletion, those in HeLa cells were longer (Figure 4—figure supplement 1J), consistent with previous observations on Drosophila S2 cells and Xenopus egg extracts (Goshima et al., 2007; Petry et al., 2011). This difference in spindle length might be due to the overlaps remaining the same length after augmin depletion in RPE1 cells, while being longer and thereby able to push the spindle poles further apart in HeLa cells (Figure 4—figure supplement 1K). When both spindle length and overlap length were taken into account, the relative length of overlaps with respect to spindle length increased in RPE1 cells from 48±1%–57 ± 2% following augmin depletion (Figure 4F), comparable to the increase in HeLa cells (Figure 4—figure supplement 1L-M). Altogether, these results suggest that augmin regulates both the width and length of metaphase spindles, while also restricting the portion of spindle length occupied by overlap microtubules.

Interestingly, the long curved bundles characteristic for augmin depletion (Goshima et al., 2008; Uehara et al., 2016; Wu et al., 2008; Zhu et al., 2008) exhibited PRC1 signal along most of their length, suggesting that they consist of antiparallel microtubules, even though contrary to bridging fibers, they form away from the k-fibers and kinetochores (corresponding to long, curved bundles in Figure 1A and Figure 3—figure supplement 1). These bundles likely arose either due to PRC1 crosslinking excessively long astral microtubules that were now able to reach the spindle midzone or due to PRC1 activity combined with the excess of free tubulin present as a consequence of less tubulin being incorporated in bridging and k-fibers. Altogether, the data suggest that there was an overall redistribution of PRC1 within the spindle from a large number of relatively short overlaps to a small number of relatively long overlaps. These results were further corroborated by PRC1-antibody staining in unlabeled HeLa cells, which also showed a reduced number of elongated PRC1 signals along the curved outer bundles after augmin depletion (Figure 4—figure supplement 1F). Thus, without augmin, the spindles are wider and contain fewer overlaps, which occupy a larger portion of spindle length and tend to accumulate at the spindle periphery.

The interkinetochore distance decreases preferentially in the inner part of the spindle and at kinetochores with weaker bridging fibers after augmin depletion

The interkinetochore distance, which is a readout of interkinetochore tension (Waters et al., 1996), decreases after augmin depletion (Uehara et al., 2009; Zhu et al., 2008). Our measurements on RPE1 and HeLa cells also showed a reduced interkinetochore distance in augmin-depleted cells (Figure 5—figure supplement 1A). This reduction of interkinetochore tension may be due to weaker k-fibers (Uehara et al., 2009; Zhu et al., 2008). However, we noticed that the interkinetochore distance was smaller in the inner part of the spindle in augmin-depleted cells (Figure 5A–D, Figure 5—figure supplement 1B), where bridging fibers were most severely impaired (Figures 3H and 4A). This was not the case in control cells, which showed no difference in interkinetochore distance between the inner and the outer part of the spindle (Figure 5D, Figure 5—figure supplement 1B). These findings motivated us to investigate a potential link between the lack of proper bridging fibers and the interkinetochore tension. We thus divided kinetochore pairs in STED images into two groups: (1) those with a bridging fiber (i.e. signal intensity of the bridging fiber above the background signal), and (2) those with undetectable signal intensities at the expected locations of bridging fibers, which we for simplicity refer to as kinetochore pairs without bridging fibers (Figure 5B–C). Remarkably, kinetochore pairs without bridging fibers had a significantly smaller interkinetochore distance than kinetochore pairs with bridging fibers (Figure 5E).

Figure 5 with 1 supplement see all
The reduction of the interkinetochore distance after augmin depletion is related to the impairment of bridging fibers.

(A) STED superresolution images of microtubules immunostained for α-tubulin (gray) in control (left), HAUS6- (middle) and HAUS-8 depleted (right) RPE1 cells stably expressing CENP-A-GFP (rainbow, confocal). Images are maximum intensity projections and kinetochores are color-coded for depth from blue to red with the Spectrum LUT in ImageJ. (B) The schematic representation of a kinetochore pair (KCs) with (top) and without (bottom) bridging fiber (See Results). (C) Enlarged boxes show KCs with or without a bridging fiber in control (left), HAUS6- (middle), and HAUS8- (right) depleted RPE1 cells. Images represent single z-plane taken from spindles in (A) and smoothed with 0.75-mm-sigma Gaussian blur. Kinetochores are shown in yellow. (D) Univariate scatter plot of the interkinetochore distance in control (gray), HAUS6- (yellow), and HAUS8- (orange) depleted cells with kinetochore pairs divided based on their distance from the long (pole-to-pole) spindle axis (outer kinetochore pairs shown in darker colors and inner in lighter colors). N=30 cells in all three conditions; 78 and 80 outer and inner kinetochore pairs for control, respectively; 84 and 96 outer and inner kinetochore pairs for HAUS6 depletion, respectively; 88 and 92 outer and inner kinetochore pairs for HAUS8 depletion, respectively. (E) Univariate scatter plot of the interkinetochore distance in HAUS6- (yellow) and HAUS8- (orange) depleted cells. Kinetochore pairs are divided into two groups: with bridging fiber (darker colors) and without bridging fiber (lighter colors). N=30 cells in HAUS6/8-depleted cells; 106 pairs with and 74 kinetochore pairs without bridging fibers in cells following HAUS6 depletion, respectively; 110 and 70 kinetochore pairs with and without bridging fibers in cells following HAUS8 depletion, respectively. (F) The correlation of the interkinetochore distance and the k-fiber intensity for kinetochore pairs with (dark gray) and without (light gray) bridging fiber in HAUS6- (left) and HAUS8-depleted (right) cells. (G) Univariate scatter plot of the interkinetochore distance for HAUS6- (yellow) and HAUS8-depleted (orange) cells. Kinetochore pairs are divided into two groups: with bridging fiber (darker colors) and without bridging fiber (lighter colors), but in this case both groups have the same k-fiber intensity. N=27 kinetochore pairs with a bridging fiber and 18 kinetochore pairs without a bridging fiber in HAUS6-depleted cells, respectively; N=23 kinetochore pairs with and N=25 kinetochore pairs without a bridging fiber in HAUS8-depleted cells. (D, E and G) Boxes represent standard deviation (dark gray), 95% confidence interval of the mean (light gray) and mean value (black). All results were obtained from three independent experiments. Statistical analysis (D, E and G) t-test for samples that followed normal distribution or Mann–Whitney U test for samples that significantly departured from normality, determined using the Shapiro-Wilk test; (F) linear regression; p-value legend:<0.0001 (****), 0.0001–0.001 (***), 0.001–0.01 (**), 0.01–0.05 (*), ≥0.05 (ns). All images are adjusted for clarity based on the intensity of astral microtubules in each image (see Materials and methods). Scale bars, 2 µm.

Although this result suggests a role of bridging fibers in regulating interkinetochore distance, this effect may be indirect and arise due to k-fibers, askinetochore pairs that lacked a bridging fiber typically had thinner k-fibers than those with a bridging fiber in augmin-depleted cells (Figure 5—figure supplement 1C). Hence, we used several approaches to separate the contribution of bridging and k-fibers to the interkinetochore tension. First, we found that although the interkinetochore distance correlated both with bridging and k-fiber intensity after augmin depletion, the correlation with bridging fiber intensity was stronger (Figure 5—figure supplement 1D-E). Such correlations were absent in control cells (Figure 5—figure supplement 1D-E). To explore a specific contribution of k-fibers to the interkinetochore tension, we divided the kinetochore pairs in augmin-depleted cells into two subsets, those with and without bridging fibers, and found that the interkinetochore distance did not correlate with k-fiber intensity within each group (Figure 5F), which argues against the k-fiber intensity as a sole determinant of interkinetochore tension. In agreement with this, when we selected two subsets of kinetochore pairs with either very strong or very weak k-fiber intensity but with comparable bridging fiber intensities (Figure 5—figure supplement 1F-G), we found no difference in the interkinetochore distance between these subsets (Figure 5—figure supplement 1H). Finally, to examine a specific contribution of bridging fibers, we identified two subsets of kinetochore pairs with similar k-fiber intensity values, one of which had bridging fibers and the other which did not (Figure 5—figure supplement 1). We found that the interkinetochore distance was larger in the subset with bridging fibers than without (Figure 5G), indicating a specific effect of bridging fibers on interkinetochore tension. Analysis of live-cell confocal images of RPE1 cells yielded similar results (Figure 5—figure supplement 1J-M). Based on these data, we conclude that augmin has a significant role in regulating interkinetochore tension through the nucleation of bridging microtubules.

Discussion

In this paper, we propose a model of the metaphase spindle in which the bridging fiber, which laterally connects sister k-fibers, forms by augmin-based nucleation of microtubules along the existing microtubules. The newly nucleated microtubules in the bridging fiber create an antiparallel overlap in which the microtubules slide apart, generating a pushing force that the bridging fiber exerts on its k-fibers. In doing so, the bridging fiber works together with k-fibers to produce tension and maintain the appropriate orientation of sister kinetochores parallel to the spindle axis, thereby preventing merotelic attachments and ensuring mitotic fidelity (Figure 6).

A model of augmin-dependent nucleation of bridging microtubules with their contribution to mitotic fidelity.

(A) Bridging microtubules are to a large extent formed by augmin-dependent nucleation. They ensure the alignment of sister kinetochores parallel to the spindle axis and the interkinetochore tension. Augmin-nucleated microtubules (yellow) and microtubules nucleated through other pathways (gray) in bridging and k-fibers are shown together with the number of microtubules in each group, as estimated from HAUS6 depletion experiments (See Results). (B) Impaired structure of bridging fibers upon augmin depletion leads to weaker interkinetochore tension and increased tilt of the kinetochores, which puts kinetochores at risk of interacting with additional microtubules (red), resulting in merotelic attachments.

Our work shows that the depletion of the augmin complex by silencing the HAUS6 or HAUS8 subunits causes severe thinning of bridging fibers in metaphase spindles, combined with the appearance of wavy and disordered midplane-crossing microtubules. Thus, we conclude that the predominant nucleation of bridging microtubules by the augmin complex enables their directional bundling along the sister k-fibers, in agreement with previous observations on directionality of microtubule growth (David et al., 2019; Kamasaki et al., 2013). K-fibers were also thinner, though to a lesser extent, indicating that they are largely nucleated in an augmin-independent manner, at the centrosome or kinetochores and chromosomes. This is in agreement with previous electron microscopy studies of mammalian spindles, where k-fiber microtubules were observed to typically reach the centrosome, while sometimes also extending from the kinetochores with their minus ends within the spindle (Kiewisz et al., 2022; McDonald et al., 1992; Sikirzhytski et al., 2018), the latter likely representing a combination of microtubules nucleated either at the kinetochores or by the augmin complex. In contrast, most midplane-crossing microtubules, which likely correspond to bridging fibers, start at different points along the k-fiber (Mastronarde et al., 1993). Electron tomography of spindles in RPE1 cells confirmed this result by showing that microtubule minus ends are found along the k-fiber, less than 50 nm from the k-fiber wall and at a distance 2–4 µm from the pole (O’Toole et al., 2020). As we found that the same number of microtubules in bridging and k-fibers was nucleated by augmin, we propose that the existing microtubules orient the growth of augmin-nucleated microtubules (David et al., 2019; Kamasaki et al., 2013), which later become kinetochore microtubules if their plus end binds to the kinetochore or bridging microtubules if they grow past the kinetochore and interact with the bridging or kinetochore microtubules on the other side. However, as there are less microtubules in the bridging fiber to begin with, augmin-mediated nucleation becomes the predominant pathway of their nucleation.

Previous work showed that augmin depletion results in a decrease of interkinetochore distance (Uehara et al., 2009; Zhu et al., 2008), a readout of tension, but it remained unclear if this is due to impaired k-fibers or perturbation of other microtubules. Treatment of cells with a microtubule-destabilizing agent that results in thinner k-fibers causes a reduction of the interkinetochore tension (Dudka et al., 2018), supporting the former possibility. However, a similar effect on the interkinetochore tension was observed upon perturbation of the bridging fiber by removing the microtubule crosslinker PRC1 (Jagrić et al., 2021; Kajtez et al., 2016; Polak et al., 2017), in agreement with the latter possibility. When looking at a subset of kinetochore pairs that had a bridging fiber and those that did not, we found that the tension was more compromised in the latter group. Notably, tension was independent of the k-fiber thickness within each group and depended on the presence or the absence of bridging fibers even when the effect of k-fibers was excluded. Although our experiments cannot directly discern the exact contribution of the bridging and k-fiber impairment in the decrease of tension on kinetochores, they reveal that augmin-generated bridging microtubules have a significant and specific role in the maintenance of interkinetochore tension.

Considering the importance of the interkinetochore tension for the accuracy of cell division (Lampson and Grishchuk, 2017), the maintenance of tension by the bridging fiber might represent an important mechanism for silencing of the spindle assembly checkpoint (Musacchio and Salmon, 2007; Nicklas et al., 1995), supported by the fact augmin-depleted cells with characteristically compromised bridging fibers have extensive mitotic delays (Wu et al., 2008). However, recent work suggests that the reduction of interkinetochore tension caused by k-fiber thinning does not necessarily lead to checkpoint activation, but instead may sometimes result in reduced anaphase A speed caused by low microtubule occupancy, followed by a subsequent increase in lagging chromosomes (Dudka et al., 2018). As proteins involved in bridging fiber nucleation and crosslinking, including HAUS6, HAUS8, and PRC1 have all been linked to various types of cancer (ICGC/TCGA, 2020, retrieved by using cBioPortal Cerami et al., 2012; Gao et al., 2013), it is plausible that impairment of bridging fibers also leads to such increase in lagging chromosomes and contributes to aneuploidy in cancers. Indeed, once the checkpoint was weakened, lagging chromosomes that appeared following augmin depletion had a reduced interkinetochore distance, which was consistent with previous findings on lazy chromosomes (Sen et al., 2021). However, contrary to previous findings in cells with intact bridging fibers, in which there was no connection between lagging chromosomes and tilt of their kinetochores (Sen et al., 2021), we found that kinetochores of lagging chromosomes in augmin-depleted cells predominantly form a large tilt with respect to the spindle axis. This suggested that the lack of bridging fibers represents a specific mechanism of predisposing the kinetochore to merotelic attachments by tilting the kinetochores and exposing their surface to the microtubules from the opposite pole. The tilted kinetochore pair is more likely to encounter additional microtubules also because midplane-crossing microtubules that form following augmin depletion no longer exhibit directional growth, but are instead wavy and extend in various directions. Once a merotelic attachment forms, it may further amplify the existing tilt due to pulling by additional microtubules in a skewed direction. Moreover, erroneous attachments might be less likely to undergo error correction in augmin-depleted cells, as bridging fibers may serve as tracks that guide Aurora B within the Chromosome Passenger Complex towards the kinetochores to correct the errors (Matković et al., 2022 Preprint). In addition to predisposing the kinetochore to merotelic attachments through impaired bridging fibers, thinning of k-fibers upon augmin depletion might ultimately be responsible for the inability to resolve merotelic attachments, as strong k-fibers are necessary to ensure proper segregation of kinetochores during anaphase (Dudka et al., 2018). Altogether, our results reveal that augmin is the leading nucleator of bridging fibers and suggest a delicate interplay of bridging and k-fibers in the maintenance of spindle organization, kinetochore tension and mitotic fidelity.

Materials and methods

Cell lines

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Experiments were carried out using unlabeled human HeLa-TDS cells from the High-Throughput Technology Development Studio (MPI-CBG, Dresden); human HeLa-Kyoto BAC cells stably expressing PRC1-GFP (Poser et al., 2008), courtesy of Ina Poser and Tony Hyman (MPI-CBG, Dresden, Germany); human HeLa cells stably expressing CENP-A-GFP and Centrin1-GFP (Gasic et al., 2015), which were a gift from Emanuele Roscioli and Andrew McAinsh (University of Warwick, Coventry, UK); human HeLa-TDS cells stably expressing GFP-α-tubulin as described in our previous work Kajtez et al., 2016; human hTERT-RPE1 (hTERT immortalized retinal pigment epithelium) cells stably expressing CENP-A-GFP and human hTERT-RPE1 (hTERT immortalized retinal pigment epithelium) cells stably expressing both CENP-A-GFP and Centrin1-GFP (Magidson et al., 2011), a courtesy of Alexey Khodjakov (Wadsworth Center, New York State Department of Health, Albany, NY, USA); human RPE1 CRISPR-Cas9 cells stably expressing PRC1-GFP (Asthana et al., 2021), a gift from Thomas Surrey (Centre for Genomic Regulation, Barcelona, Spain); and human U2OS cells stably expressing CENP-A-GFP, mCherry-α-tubulin and PA-GFP-α-tubulin (Barisic et al., 2014), a gift from Marin Barišić (Danish Cancer Society Research Center, Copenhagen, Denmark).

Cell culture

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All cell lines were cultured in flasks in Dulbecco’s Modified Eagle’s Medium with 1 g/L D-glucose, pyruvate and L-glutamine (DMEM, Lonza, Basel, Switzerland), supplemented with 10% (vol/vol) heat-inactivated Fetal Bovine Serum (FBS, Sigma Aldrich, St. Louis, MO, USA) and penicillin (100 IU/mL)/streptomycin (100 mg/mL) solution (Lonza, Basel, Switzerland). For the selection of U2OS CENP-A-GFP mCherry-α-tubulin PA-GFP-α-tubulin, HeLa PRC1-GFP, HeLa CENP-A-GFP Centrin1-GFP and Hela GFP-α-tubulin cell lines, 50 μg/ml geneticin was added to the medium (Life Technologies, Waltham, MA, USA).

All cells were kept at 37 °C and 5% CO2 in a Galaxy 170 R humidified incubator (Eppendorf, Hamburg, Germany). They have also been regularly tested for mycoplasma contamination by examining the samples for extracellular DNA staining with SiR-DNA (100 nM, Spirochrome, Stein am Rhein, Switzerland) and Hoechst 33342 dye (1 drop/2 ml of NucBlue Live ReadyProbes Reagent, Thermo Fisher Scientific, Waltham, MA, USA) and have been confirmed to be mycoplasma free.

Sample preparation and RNAi transfection

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At 80% confluence, DMEM medium was removed from the flask and cells were washed with 5 ml of PBS. Then, 1 ml 1% trypsin/EDTA (Biochrom AG, Berlin, Germany) was added to the flask and cells were incubated at 37 °C and 5% CO2 in a humidified incubator for 5 min. After the incubation, trypsin was blocked by adding 2 ml of DMEM medium. For RNAi experiments, the cells were seeded to reach 60% confluence the next day and cultured on 35 mm uncoated dishes with 0.16–0.19 mm (1.5 coverglass) glass thickness (MatTek Corporation, Ashland, MA, USA) in 2 mL DMEM medium with previously described supplements. After one day of growth, cells were transfected with either targeting or non-targeting siRNA constructs which were diluted in OPTI-MEM medium (Life Technologies, Waltham, MA, US) to a final concentration of 20 nM for HAUS6 and HAUS8 in medium with cells. HAUS6 and HAUS8 siRNA transfections were performed 48 hr prior to imaging and cotransfection with 100 nM Mad2 siRNA was additionally performed 24 hr prior to imaging using Lipofectamine RNAiMAX Reagent (Life Technologies, Waltham, MA, US) according to the instructions provided by the manufacturer. After four hours of treatment, the medium was changed to the previously described DMEM medium. The constructs used were human HAUS6 siRNA (L-018372-01-0005, Dharmacon, Lafayette, CO, USA), human HAUS8 siRNA (L-031247-01-0005, Dharmacon, Lafayette, CO, USA), human Mad2 siRNA (L-003271-00-0010, Dharmacon, Lafayette, CO, USA) and control siRNA (D-001810-10-05, Dharmacon, Lafayette, CO, USA). Mad2 was chosen instead of MPS1 inhibitors as it induces less segregation errors per cell in control cells, as well as to avoid a biased approach in which the inhibitor would likely be added at a different time point during an hours-long error correction process that occurs in augmin-depleted cells.

When microtubules were visualized during live cell-imaging in cell lines without stable expression of tubulin, staining was performed to a final concentration of 100 nM with a far-red silicon rhodamine (SiR)-tubulin-670 dye (Spirochrome, Stein am Rhein, Switzerland), 45 min to 2 hr prior to imaging. As for DNA, either SiR-DNA or SPY-555-DNA (Spirochrome, Stein am Rhein, Switzerland) were used to a final concentration of 100 nM, 20 min to 2 hr prior to imaging. In order to avoid dye efflux, a broad-spectrum efflux pump inhibitor verapamil (Spirochrome, Stein am Rhein, Switzerland) was added at a final concentration of 0.5 μM to RPE1 cells along with tubulin and/or DNA dyes. As an additional control for metaphase cells, MG-132 (Sigma Aldrich, M7449-1ML, MO, USA) was added to the cells used for the measurement of spindle size for 30 min prior to imaging at a final concentration of 20 µM.

Immunofluorescence

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For confocal imaging, HeLa cells stably expressing PRC1-GFP were grown on glass-bottom dishes (14 mm, No. 1.5, MatTek Corporation) and fixed by 2 ml of ice-cold methanol for 1 min at –20 °C. Following fixation, cells were washed three times for 5 min with 1 ml of PBS and permeabilized with 0.5% Triton-X-100 in water for 15 min at a room temperature. This step was repeated twice when tubulin staining was performed. To block unspecific binding, cells were incubated in 1 ml of blocking buffer (1% normal goat serum (NGS)) for 1 hr at 4 °C. Cells were then washed three times for 5 min with 1 ml of PBS and incubated with 250 µl of primary antibody solution overnight at 4 °C. The primary antibodies used were as follows: rabbit polyclonal PRC1 (diluted 1:100, sc-8356, Santa Cruz Biotechnology, RRID:AB_2169665), rabbit polyclonal HAUS6 (diluted 1:250, ab-150806, Abcam), rabbit polyclonal HAUS8 (diluted 1:100, PA5-21331, Invitrogen, RRID:AB_11153508) and rat monoclonal tubulin (diluted 1:100, MA1-80017, Invitrogen, RRID:AB_2210201). After the incubation with a primary antibody, cells were washed 3 times for 5 min with 1 ml of PBS and then incubated with 250 µl of secondary antibody for 45 min at a room temperature. Alexa Fluor 488 and 594 (Abcam, ab150073, ab150076, ab150156) were used as secondary antibodies at a 1:1000 dilution for PRC1 staining, 1:500 dilution for HAUS6 and 1:250 for HAUS8 and tubulin staining. DAPI (1 µg/mL) was used for chromosome visualization.

For STED imaging, RPE1 cells stably expressing CENP-A-GFP were grown on glass-bottom dishes (14 mm, No. 1.5, MatTek Corporation), cell medium was removed, and cytoskeleton extraction buffer was added for 20 s to remove the components of the cytoplasm. Following extraction, cells were fixed in 3% paraformaldehyde and 0.1% glutaraldehyde solution for 10 min. To reduce the background fluorescence, quenching (100 mM glycine in PBS) and reduction (0.1% sodium borohydride in PBS) solution were added for 7 and 10 min, respectively. To prevent non-specific binding, cells were incubated in blocking/permeabilization buffer (2% normal goat serum and 0.5% Triton-X-100 in PBS) for 2 hr at 4 °C. Microtubules were then stained using a rat anti-tubulin primary antibody solution (diluted 1:500 in blocking/permeabilization buffer, MA1-80017, Invitrogen) with a 4 °C overnight incubation. The next day, cells were washed with PBS three times for 5 min. After washing, a secondary antibody Alexa Fluor 594 (dilution 1:1000, Abcam) was added and incubated for 1 hr at room temperature. Additionally, DAPI (1 µg/mL) was added and incubated for 15 min at room temperature to visualize chromosomes. Cells exposed to cold treatment were incubated with cold media on ice for 13 min prior to extraction of cytoplasmic components.

Immunoblotting

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RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP were grown on six-well plates (Greiner Bio-one) and subjected to HAUS6 or HAUS8 siRNA treatment as indicated before. Following transfection, cells were lysed in RIPA buffer (R0287, Sigma) containing 1 x protease inhibitor (5892970001, Roche), 1 x phosphatase inhibitor (4906837001, Roche) and 1 mM PMSF by two cycles of freezing and thawing in liquid nitrogen. Protein extracts were mixed with 2 x Laemlli sample buffer (S3401, Sigma) and heated at 95 °C for 10 min prior to SDS-PAGE. After protein transfer onto the nitrocellulose membrane (IB23002, Invitrogen) and blocking with blocking solution (5% bovine serum albumin and 0,1% Tween 20 in PBS) for 1 hr, membranes were incubated overnight at 4 °C with primary antibodies diluted in blocking solution. The primary antibodies used were as follows: rabbit polyclonal HAUS6 (diluted 1:1000, ab-150806, Abcam), rabbit polyclonal GAPDH (diluted 1:1000, ab9485, Abcam, RRID:AB_307275). Rabbit polyclonal HAUS8 antibody (diluted 1:1000, PA5-21331, Invitrogen, RRID:AB_11153508, and NBP2-42849, Novus Biologicals, RRID:AB_2665500) resulted in no detectable bands under these conditions. Membranes were washed with 0.1% Tween 20 in PBS, incubated for 1 hr with anti-rabbit HRP-conjugated secondary antibodies (dilution 1:10,000, ab6721, RRID:AB_955447) and visualized on the C-DiGit blot scanner (LI-COR, Bad Homburg, Germany) with WesternSure PREMIUM Chemiluminescent Substrate (926–95000, LI-COR).

Microscopy

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STED microscopy of fixed cells and live-cell imaging of anaphase were performed using an Expert Line easy3D STED microscope system (Abberior Instruments, Göttingen, Germany) with the 100 x/1.4NA UPLSAPO100x oil objective (Olympus, Tokio, Japan) and an avalanche photodiode (APD) detector. The 488 nm line was used for excitation in both cases, with the addition of the 561 nm line for excitation and the 775 nm laser line for depletion during STED superresolution imaging. Images were acquired using the Imspector software. The xy pixel size for fixed cells was 20 nm and 6 focal planes were acquired with 300 nm distance between planes were acquired. For confocal live-cell imaging of anaphase, the xy pixel size was 80 nm and 16 focal were acquired, with 1 µm distance between the planes and 30 s time intervals between different frames.

Other confocal images and videos were acquired using a previously described microscope setup (Buđa et al., 2017), consisting of a Bruker Opterra Multipoint Scanning Confocal Microscope (Bruker Nano Surfaces, Middleton, WI, USA), mounted on a Nikon Ti-E inverted microscope with a Nikon CFI Plan Apo VC 100 x/1.4 numerical aperture oil objective (Nikon, Tokyo, Japan). During live-cell imaging, cells were kept at 37 °C and 5% CO2 in Okolab Cage Incubator (Okolab, Pozzuoli, NA, Italy). To excite Hoechst, GFP, mCherry or SiR fluorescence, a 405 nm, 488 nm, 561 nm or 647 nm laser lines were used, respectively. Opterra Dichroic and Barrier Filter Set 405/488/561/640 enabled the separation of excitation light from the emitted fluorescence. Images were acquired using Evolve 512 Delta Electron Multiplying Charge Coupled Device (EMCCD) Camera (Photometrics, Tuscon, AZ, USA), with camera readout mode of 20 MHz. The xy pixel size was 83 nm. In all experiments where the whole spindle stack was imaged, z-stacks were acquired with unidirectional xyz scan mode at 37 focal planes and 0.5 μm distance between the planes. Photoactivation was performed perpendicular to pole-to-pole axis of metaphase spindles using a 405 nm laser diode (Coherent, Santa Clara, CA, USA) and a line pattern of 12 equally distributed points, with each point representing one laser hit. The interval between the points was 0.05ms and photoactivation area was set to 0.5 µm for each point. The interval between successive frames was set to 10 s and one central z-plane was imaged.

Image processing and data analysis

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In cells where HAUS6 or HAUS8 were silenced only bipolar metaphase spindles were imaged and analyzed, even though multipolar spindles were observed as reported previously (Lawo et al., 2009). All images were analyzed in Fiji/ImageJ (National Institutes of Health, Bethesda, MD, USA, RRID:SCR_003070). Raw images were used for quantification. In representative immunofluorescence images of augmin depletion (Figure 2—source data 1), all signals were adjusted equally in control and treated cells. However, due to severe reduction of bridging fibers in augmin-depleted spindles, the contrast in images for representation on figures was not always equally adjusted, as this led to important spindle structures being either highly oversaturated or barely visible. It was instead adjusted so that astral microtubules are similarly visible in control and augmin-depleted spindles in STED microscopy, or that all present bridging fibers are visible in confocal microscopy. These adjustments did not result in any structures being omitted or otherwise modified in a way that could lead to misrepresentation. MatLab (MathWorks, Natick, MA, USA, RRID:SCR_001622) was used for calculations and plotting and Rstudio (R Foundation for Statistical Computing, Vienna, Austria, RRID:SCR_000432) to transform the cells into an end-on view. Figures were assembled in Adobe Illustrator CS5 (Adobe Systems, Mountain View, CA, USA, RRID:SCR_010279).

Measuring the bridging fiber intensity

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For the analysis based on small square regions which was performed on both STED and confocal images, tubulin intensity in bridging and k-fibers was measured using either a 25x25 or 5x5 pixel Square tool (ImageJ) on STED and confocal images, respectively. To measure bridging fibers, the square was positioned on the fiber spanning the area between two kinetochores. K-fibers were measured right next to the kinetochores. The average intensity of the two sister k-fibers was used for further analyses. The background was measured using the same tool at several empty areas within the spindle and its average was subtracted from bridging (Ib = Ib+bcgIbcg) and k-fiber intensities (Ik = Ik+bcgIbcg). 10 randomly positioned empty areas were taken into account while calculating background for STED imaging, whereas 2 empty areas just above and below bridging fiber were taken into account while calculating background for confocal imaging due to different nature of the acquired images. All measurements were performed on randomly selected bundles in single z-planes, after determining that no other microtubules were crossing the area of measurement. In STED images, the intensity of astral microtubules was additionally measured using the 25x25 pixel Square tool (ImageJ). Their background was measured using the same tool in the empty area between the two astral microtubules and it was subtracted from astral microtubule intensity (Ia=Ia+bcg-Ibcg).

On top of that, tubulin intensities of the bridging fiber and k-fiber region in confocal images were measured in a single z plane using the Segmented Line tool by drawing a 5 pixel line (ImageJ) along the contour of k-fibers and the corresponding bridging fiber. Background was measured in the same z plane by drawing the 5 pixel thick line along the length of the metaphase midzone, as this represents the background signal caused by the presence of neighboring microtubules. The minimum value of the midzone intensity profile was subtracted from the intensity profiles of bundle contours. The minimum value of the mean tubulin intensity profile was set as a distance of zero μm and was selected as the center of the bridging fiber. The final intensity of a bridging fiber (Ib) was calculated as the mean value of intensities in the area 500 nm around the center of the bridging fiber. The final intensity of a k-fiber region (Ibk), which also includes the bridging fiber, was calculated as an average of two mean values of intensities in the area 500 nm around the distance of 1.5 μm away from the center of the bridging fiber. The intensity value of k-fibers alone (Ik) was then calculated as Ik = IbkIb.

Comparison of SiR- and GFP-tubulin signal

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For the comparison of SiR and GFP-tubulin signal in HeLa cells stably expressing GFP-α-tubulin, the previously described analysis was used. Tubulin intensity of the bridging fiber and k-fiber region was measured in a single z plane using the Segmented Line tool in ImageJ by drawing the 5 pixel line along the contour of k-fibers and the corresponding bridging fiber. Due to high noise, only outermost bundles were taken into analysis and the reduction was not calculated. The background was measured in the same z plane by drawing the 5x5 pixel square in the cytoplasm and was subtracted from the bundle intensity profiles. The minimum value of the intensity profile was set as a distance of zero μm and was selected as the center of the bridging fiber.

The interkinetochore distance

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To calculate the interkinetochore distance, two points were put on the centers of signal in each kinetochore pair using a Point tool in ImageJ. Additionally, two points were put on spindle poles and two points on the upper and lower edge of the metaphase plate. Interkinetochore distance and distance from spindle axes were calculated using a home-written MatLab script. Two-dimensional analysis was applied when all tracked kinetochore pairs resided within 2 μm around the central z-plane. Three-dimensional analysis was used when all kinetochore pairs in the spindle were taken into account, regardless of their position. In 3D analysis, the exact distance from the long spindle axis (c) was calculated using the Pythagorean theorem, where a=projected distance of a middle point between two sister kinetochores in a particular z-plane from the long spindle axis, b=distance between the central z-plane and the z-plane of the kinetochore pair x 0.81 (correction factor for the oil objective only, see Transformation of spindles into an end-on view). Kinetochore pairs were defined as those in the inner or the outer part of the spindle if their distance to the pole-to-pole axis was smaller or larger than the average distance of all tracked kinetochore pairs to the pole-to-pole axis, respectively. Additionally, kinetochores with visible bridging fibers and kinetochores with no visible bridging fibers were separately tracked and analyzed in confocal images of RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP, where the presence of bridging fibers was determined by measuring intensity profiles of the tubulin signal between two kinetochores.

Transformation of spindles into an end-on view

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Z-stacks of the mitotic spindles that were positioned horizontally were transformed into the end-on view using a home-written R script, based on the original version (Novak et al., 2018). Prior to transformation, a single-channel grayscale z-stack was rotated using ImageJ to make the long axis of the spindle parallel to the x-axis. Signal intensity at each pixel was used to obtain an end-on view of the spindles by applying the following transformation: I' (i · pixel size, j · pixel size, k · z-distance)=I (k · z-distance, I · pixel size, j · pixel size). A correction factor of 0.81 was used for the z-distance to correct for aberrations caused by the different refractive index mismatch of aqueous samples and immersion oil (Novak et al., 2018).

Number of bundles

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The number of bundles in HeLa and RPE1 PRC1-GFP cells was determined in an end-on view of the spindle by using sum intensity projections of 10 central z-planes covering 0.83 µm along the long spindle axis.

Spindle length, width, and metaphase plate diameter

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Spindle length, width and metaphase plate diameter were measured on maximum intensity projections of the side view z-stack of spindles. Spindle length was determined as a distance between the two poles. The position of the poles was determined as either the center of Centrin1 signal or the outermost points of the tubulin or PRC1 signal at the spindle pole. Spindle width was measured as the distance between two lines parallel to the long axis of the spindle and encompassing the outermost PRC1- or tubulin-labeled bundles. Additionally, in RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP the metaphase plate diameter was measured as the distance between the outermost kinetochore pairs, whereas in HeLa PRC1-GFP it was measured as the distance between the outermost chromosome ends.

Overlap length

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Overlap length was measured on sum intensity projections of 2–4 z-planes covering the entire bundle of interest, using ImageJ Segmented line tool by drawing a pole to pole line along the contour of PRC1-GFP and acquiring an intensity profile. The overlap length was defined as the length of the base of the PRC1-GFP intensity peak (Polak et al., 2017).

PRC1 intensity

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PRC1 intensity was measured in sum intensity projections of 10 central z-planes of the spindle. Total PRC1 signal in the cell was marked by using Polygon selection tool (ImageJ) and 5x5 Square tool was used to determine the background in the cytoplasm. The final intensity values were obtained using the following formula: PRC1 intensity = Integrated Density of the spindle – (Area of selected cell x mean fluorescence of background). Intensity profiles of PRC1-GFP were measured on the sum intensity projections of 10 central z-planes in an end-on view of the spindle by drawing a 50 pixel wide Straight line tool across the diameter of the spindle.

Spindle shape

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The shape of spindles was determined in ImageJ using a Point tool. Ten points were distributed throughout the bundle, with the first and last point positioned at the spindle poles. The images were rotated to make the long spindle axis parallel to the x axis. In control cells, only the outermost bundle was tracked. In HAUS6 siRNA-treated cells, three different groups of outermost bundles were tracked: bundles with visible bridging fibers, bundles with no visible bridging fibers and curved bundles extending far from the metaphase plate. Shape and curvature were calculated using a home-written MatLab script by fitting a circle to the tracked points along the bundle. Contour lengths of the bundles were measured by calculating the cumulative distance between the first and the last point of the tracked bundle.

Poleward flux rate

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For measuring the poleward flux rate, 10-pixel wide line was drawn from pole to pole along the bundle with photoactivation signal that lasted at least 5 time frames (40 s), using the Segmented Line tool in ImageJ. The position of photoactivated mark in each time frame was determined as the distance between the peaks of intensity profiles in photoactivation and SiR-tubulin channels for photoactivation mark and closer spindle pole, respectively. The analysis was performed on images processed with Gaussian Blur filter with Sigma set to 2 to improve the definition of the intensity profile peaks.

Tracking and classification of segregation errors

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Three types of segregation errors were analyzed in Mad2-depleted and Mad2/HAUS6-codepleted anaphase spindles: misaligned kinetochores, lagging kinetochores and other errors. Misaligned kinetochores were defined as those in which both kinetochores of the pair were situated outside the metaphase plate 30 s before anaphase onset. Lagging kinetochores were defined as those in which the CENP-A signal was visibly stretched and the kinetochore was situated in the central part of the spindle, outside the kinetochore mass that was moving towards the pole during anaphase. Finally, segregation errors classified as others included various kinetochores, precise classification of which was impossible using only CENP-A signal. These included kinetochores situated outside the moving kinetochore mass without stretched CENP-A signal, kinetochore pairs that remained completely unseparated for the whole duration of anaphase, as well as kinetochore pairs in which both kinetochores remained non-stretched and in the central part of the spindle despite the initial separation. Segregation errors were further divided based on their distance to the pole-to-pole axis into those in the inner and the outer part of the spindle. The distance of the kinetochore pair from the pole-to-pole axis was determined in 3D using a home written Matlab script, and the obtained value was then normalized to spindle half-width determined using maximum intensity projections in ImageJ. All kinetochores situated in the area less than 0.5 from the spindle axis 30 s before anaphase onset were defined as those in the inner part of the spindle and all kinetochore pairs in the area equal to or above the value of 0.5 were defined as kinetochores in the outer part of the spindle. All kinetochore pairs were manually tracked in time from just before anaphase onset until entering the daugther cell by using the Point tool in ImageJ. In images acquired using STED microscopy, merotelic attachments were defined as those in which one kinetochore forms attachments with microtubules from the opposite side of the spindle, with no visible microtubule signal just below or above the kinetochore.

Anaphase A and B speed

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To measure anaphase A speed, the coordinates of kinetochores and poles were tracked in time using the Point tool in ImageJ. The speed was calculated from the time point when the distance between the kinetochore and its closer pole started gradually decreasing. The slope of a line equation obtained from the linear fitting of distances over time was determined for every kinetochore as the anaphase A speed. Anaphase B speed was calculated in a manner similar to anaphase A, but instead of kinetochores, the positions of poles were tracked in time with the first time frame determined as the frame when the distance between two poles started gradually increasing.

Measuring the tilt of kinetochores at anaphase onset

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Anaphase onset was determined in a frame just before the interkinetochore distance of the tracked kinetochore pair started gradually increasing. The coordinates of kinetochore pairs that ended up as errors and of error-free kinetochore pairs were tracked along with the coordinates of spindle poles. The angle that the kinetochores form with the long spindle axis was calculated using a home written Matlab script.

Data availability

All source codes and source data have been deposited to the Dryad repository (https://doi.org/10.5061/dryad.fn2z34tz7).

The following data sets were generated
    1. Tolić IM
    (2022) Dryad Digital Repository
    Data from: Augmin prevents merotelic attachments by promoting proper arrangement of bridging and kinetochore fibers.
    https://doi.org/10.5061/dryad.fn2z34tz7

References

    1. McIntosh JR
    (2016) Mitosis
    Cold Spring Harb Perspect Biol 8:a023218.
    https://doi.org/10.1101/cshperspect.a023218
    1. Mitchison TJ
    (2005) Mechanism and function of poleward flux in Xenopus extract meiotic spindles
    Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 360:623–629.
    https://doi.org/10.1098/rstb.2004.1616

Decision letter

  1. Thomas Surrey
    Reviewing Editor; Centre for Genomic Regulation (CRG), Spain
  2. Anna Akhmanova
    Senior Editor; Utrecht University, Netherlands

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

[Editors' note: this paper was reviewed by Review Commons.]

https://doi.org/10.7554/eLife.83287.sa1

Author response

General Statements [optional]

We thank the reviewers for providing thoughtful and constructive feedback on our manuscript. Motivated by their comments, we decided to perform a full revision with extensive new experiments, which led to important novel findings and thus a significant revision of the manuscript. The main changes lie in that we performed STED super resolution imaging of augmin depleted spindles, which allowed us to analyze the previously indistinguishable microtubule arrangements and erroneous attachments, as well as to quantify k-fibers and bridging fibers with unprecedented accuracy. Moreover, a key part of the revision is a functional approach where we tested how augmin depletion affects chromosome segregation fidelity, as suggested by Reviewer 3. Surprisingly, we found that without augmin, the lack of bridging fibers results in kinetochore tilt and, consequently, merotelic attachments in metaphase. Our STED images provide the first clear demonstration of such attachments, and these new findings offer a mechanistic explanation of how augmin-dependent microtubule branching promotes mitotic fidelity.

The main conceptual advance of our work is that the augmin complex is crucial for the proper organization of midplane-crossing microtubules into uniformly arranged bridging fibers that connect two sister k-fibers and extend nearly-parallel to the spindle axis. We found that bridging fibers are often absent following augmin-depletion, and have a specific role in the maintenance of interkinetochore tension, as well as in promoting mitotic fidelity by preventing merotelic attachments. K-fibers were also impaired, but to a lesser extent, with their thinning amplifying the negative effect on mitotic fidelity by compromising resolution of merotelic attachments later in anaphase. Altogether, we propose a model where augmin promotes mitotic fidelity by generating assemblies consisting of bridging and kinetochore fibers that align sister kinetochores to face opposite poles, thereby preventing erroneous attachments.

We have now addressed the criticism of the reviewers by identifying the specific effects of bridging fibers upon augmin depletion and considerably enhanced the significance by exploring the effects on chromosome segregation fidelity, thereby transforming our paper from a descriptive to a mechanistic study. As such, the study is no longer of interest only to the mitotic field, but to a larger scientific community including cell biologists, molecular biologists and biophysicists interested in microtubules, mitotic and meiotic spindles, cell division, chromosome segregation, aneuploidy, cancer, and development, as well as scientists developing quantitative super resolution imaging protocols for imaging of cellular structures.

Reviewer #1 (Evidence, reproducibility and clarity (Required)):

In the manuscript "Augmin regulates kinetochore tension and spatial arrangement of spindle microtubules by nucleating bridging fibers", Manenica et al. explore the impact of augmin dependent microtubule nucleation on formation of a subset of spindle microtubules that bridge sister kinetochore fibers and investigate how this could affect the spindle forces and architecture. Using RNAi- and CRISPR-Cas9- based loss-of-function experimental approach, the authors propose that the bridging fibers are nucleated by augmin and that removal of augmin impairs proper spindle architecture, interkinetochore tension and microtubule poleward flux, specifically via its effect on the bridging fibers. Overall, the study is well designed and the manuscript well written. Expanding the knowledge on augmin contribution to the spindle functions and better understanding of the roles of bridging fibers would be important and of interest to cell biologists studying mitosis. Although this manuscript clearly shows that augmin depletion impairs the formation of bridging fibers (and other microtubules), the specific contribution of the bridging fibers to the augmin-dependent spindle functions is less clear.

We thank the reviewer for pointing out the quality of our experiment design and writing. We also appreciate the criticism regarding the exact contribution of bridging fibers to the spindle architecture, interkinetochore tension and microtubule poleward flux. We addressed this in our revised manuscript, where we studied the specific effect of bridging fibers on each of these phenotypes.

Using STED microscopy, we were now able to determine the specific effect of bridging fibers on spindle architecture by analyzing the previously indistinguishable microtubule arrangements within the crowded metaphase plate area and quantifying k-fibers and bridging fibers with unparalleled precision and accuracy. This type of analysis confirmed that augmin is vital for the organization of midplane-crossing microtubules into highly organized bridging fibers that connect sister kinetochore fibers and extend nearly-parallel to the spindle axis. Without augmin, proper bridging fibers were thin and often even absent, while the remaining midplane-crossing microtubules appear wavy and disorganized. K-fibers were also thinner, but to a lesser extent, as confirmed by two different types of analysis.

We clarified the contribution of bridging fibers to the interkinetochore tension using different types of analysis, which revealed that augmin-nucleated bridging microtubules have a specific role in the maintenance of interkinetochore tension. Indeed, the severe impairment of bridging fibers had a stronger effect on the interkinetochore distance than thinning of k-fibers following augmin depletion, and the specific effect of bridging fibers was demonstrated for kinetochores with the same k-fiber intensity.

As for poleward flux, a detailed analysis of poleward flux following augmin depletion, with a specific focus on the differences in bridging and k-fiber flux, is now published as a part of the recent Cell Reports paper from our lab (Risteski et al., 2022). Thus, this is no longer a main point of our manuscript, which now shifted its focus to more striking consequences of augmin depletion, particularly microtubule arrangements within the metaphase plate and mitotic fidelity.

The effect of augmin on mitotic fidelity, as suggested by Reviewer 3, was studied using sophisticated live-cell imaging and tracking protocols. We described the main results in the General Statement (see above), and dedicated an entire new section of the manuscript (pages 6-11 along with a new Figure 2) to these exciting new results that provide a mechanistic explanation of how augmin-dependent microtubule nucleation prevents merotelic attachments.

Major comments:

1) Using cold treatment-induced microtubule destabilization, Zhu et al. (JCB 2008) showed that augmin depletion affected exclusively kinetochore microtubules. Since the bridging microtubules are usually not visible in the cold-treated spindles (due to being less stable/cold resistant compared to the k-fibers), it is unlikely that the observed effects were mainly associated with the bridging fibers. Thus, it would be important to further clarify the respective contribution of augmin to the formation of k-fibers and the bridging fibers. The cold-treatment experiment performed by Zhu et al. could be used in RPE1 and HeLa-PRC1-GFP cells to address the contribution of augmin nucleation to kinetochore- vs. bridging microtubules from another angle.

Because of the above mentioned results by Zhu et al. it is difficult to grasp how augmin depletion could have a bigger effect on the bridging fibers than on the k-fibers, as concluded from the Figure 2C data. In fact, Figure 2A clearly shows a strong effect on k-fibers in spindles where the bridging fibers are reduced/missing.

Also, Figure 1 D and E suggest that HAUS8 siRNA exclusively affected the bridging fibers, leaving the k-fibers intact, which is again against the data reported in Zhu et al. 2008 and in contrast with the representing image shown in Figure 1B. Even if the RNAi was less efficient compared to HAUS6 RNAi, as the authors proposed, this could still not explain the observed discrepancy.

We thank the reviewer for this comment. To better address the contribution of augmin to either bridging or kinetochore microtubule nucleation, we repeated the experiments by using STED super-resolution microscopy on cells immunostained for tubulin, with or without HAUS6/8 depletion. STED imaging enabled a previously unattainable and exceptionally detailed visualization of k-fibers and bridging fibers (Figure 1 and Figure 3A), and subsequent analysis confirmed our initial result that augmin depletion has a larger effect on bridging fibers than on kfibers (Figure 3D,E). Importantly, due to the increased precision of our measurements when using STED images, the new results for HAUS6 siRNA and HAUS8 siRNA were now similar, and showed that the lack of augmin affects k-fibers to an extent comparable to Zhu et al., 2008 (Figure 3D,E). The data from the original manuscript, which were obtained by confocal microscopy and were therefore somewhat less precise due to the intrinsic limitations of the method, are now shown in Supplementary Figure S3.

To independently test the effect of augmin depletion on k-fibers, we also performed experiments with cold treatment as suggested, imaged the cells by STED microscopy, and analyzed the images as in Zhu et al., 2008 (Figure 3F,G). These experiments were done on RPE1 cells that stably express CENP-A-GFP. Cold treatment in HeLa-PRC1-GFP cells was no longer necessary, as STED imaging clearly showed the absence of bridging fibers in cold treated cells. The new results, that emphasise the effect on bridging fibers but no longer state that k-fibers are intact are described on page 12:

“Quantification of STED images further revealed that HAUS6 depletion resulted in 68 ± 8% reduction of the bridging fiber signal intensity and 24 ± 6% reduction of the k-fiber signal intensity, with similar results obtained by HAUS8 depletion (Figure 3D-E). These data indicate that augmin depletion affects not only k-fibers, but even more so bridging fibers. The contribution of augmin to the nucleation of k-fibers was independently tested by measuring their intensity in spindles exposed to cold treatment in which bridging fibers are removed (Figure 3F). HAUS6 depletion resulted in a 37 ± 5% reduction of the k-fibers (Figure 3G), which is consistent with a previous study (Zhu et al., 2008) and comparable to values under non-cold conditions.”

2) The authors showed that kinetochore pairs in the outer parts of Augmin-depleted spindles have larger inter-kinetochore distance compared to those in the inner parts of spindles. They indirectly related this to a predominant presence of the bridging fibers in the outer parts, concluding that augmin regulates inter-kinetochore tension via nucleation of the bridging fibers. A more direct way would be to show the eventual positive correlation between the inter-kinetochore distance and the bridging- and k- fibers intensity. Also, it would be nice to include the quantifications and correlation data for inter-kinetochore distance, distance from the spindle axis and the bridging- and k- fibers intensities for the control cells too.

We analyzed the new data and included the correlations in the altered manuscript. We explained the correlation data for the interkinetochore distance and the distance from the spindle axis as follows: “… we noticed that the interkinetochore distance was smaller in the inner part of the spindle in augmin-depleted cells (Figure 5A-D, Supplementary Figure S5B), where bridging fibers were most severely impaired (Figure 3H and 4A). This was not the case in control cells, which showed no difference in interkinetochore distance between the inner and the outer part of the spindle (Figure 5D, Supplementary Figure S5B).”

We also included the correlation data for the interkinetochore distance and the bridging and kfibers intensity: “… we found that although the interkinetochore distance correlated both with bridging and k-fiber intensity after augmin depletion, the correlation with bridging fiber intensity was stronger (Supplementary Figure S5D-E). Such correlations were absent in control cells (Supplementary Figure S5D-E).”

In addition, we performed yet another type of analysis where we divided kinetochore pairs into subsets with either comparable bridging fiber intensities and contrasting k-fiber intensities or comparable k-fiber intensities and contrasting bridging fiber intensities. We then compared the interkinetochore distance of kinetochore pairs within these subsets, as follows: "To explore a specific contribution of k-fibers to the interkinetochore tension, we divided the kinetochore pairs in augmin-depleted cells into two subsets, those with and without bridging fibers, and found that the interkinetochore distance did not correlate with k-fiber intensity within each group (Figure 5F), which argues against the k-fiber intensity as a sole determinant of interkinetochore tension. In agreement with this, when we selected two subsets of kinetochore pairs with either very strong or very weak k-fiber intensity but with comparable bridging fiber intensities (Supplementary Figure S5F-G), we found no difference in the interkinetochore distance between these subsets (Supplementary Figure S5H). Finally, to examine a specific contribution of bridging fibers, we identified two subsets of kinetochore pairs with similar k-fiber intensity values, one of which had bridging fibers and the other which did not (Supplementary Figure S5I). We found that the interkinetochore distance was larger in the subset with bridging fibers than without (Figure 5G), indicating a specific effect of bridging fibers on interkinetochore tension. Analysis of live-cell confocal images of RPE1 cells yielded similar results (Supplementary Figure S5J-M). Based on these data, we conclude that augmin has a significant role in regulating interkinetochore tension through the nucleation of bridging microtubules."

In sum, in the altered manuscript we conclude that both bridging and k-fibers regulate the kinetochore tension, and that in augmin-depleted cells there is indeed a specific effect of bridging fibers on interkinetochore tension, which is larger than the effect of k-fibers.

3) It is stated in the manuscript that the k-fibers without bridging fibers have shorter contour length compared to the k-fibers with bridging fibers, and that the curvature of k-fibers lacking the bridging fibers is drastically reduced. However, the data in Figure 5D and Table 1 show a slight effect on the contour length of the k-fibers lacking the bridging fibers compared to the ones containing the bridging fibers only in RPE1 siHAUS8 cells, while this effect seems to be missing in RPE1 HAUS8 KO cells, as well as in siHAUS6 in RPE1 and HeLa cells.

Figure 2 shows that the kinetochore pairs without the bridging fibers are located closer to the spindle axis. Thus, it is not clear whether the effect on curvature observed in the augmin depleted cells is independent of the position of kinetochore pairs within the spindle, as the spindle axis-proximal pairs would anyway have a bigger radius compared to the more distant ones.

As these analyses were previously performed on bundles stained with SiR-tubulin and using confocal microscopy, we have now determined their curvature on spindles immunostained for tubulin and imaged by STED microscopy, where the shape of these bundles can be determined more precisely both in control and HAUS6 depleted cells. In the revised manuscript, only those spindles were taken for further analysis (Supplementary Figure S3I-K), as this is no longer a major point in the revised manuscript which shifted its focus towards other more striking consequences of augmin depletion. The text was revised as follows: “Whereas the bundles without kinetochores in HAUS6 siRNA-treated cells had a significantly longer contour when compared to all other bundle types (Supplementary Figure S3J), k-fibers without bridging fibers in augmin-depleted cells had a significantly larger radius of curvature than any of the other bundle types in augmin-depleted or control cells (Supplementary Figure S3K). Taken together, the outer interpolar bundles without associated kinetochores are excessively long and make the spindle wider, whereas k-fibers lacking a bridging fiber are overly straight, ultimately resulting in a diamond-like shape of the spindle.”

As for previous Figure 2, in all experiments regarding shape, we only analyzed the outermost bundles, so the potential effect of the position of kinetochore pairs within the spindle can be excluded. We explained that in the Methods section and highlighted it in the caption of the Supplementary Figure 3I: “In control cells, only the outermost bundle was tracked. In HAUS6 siRNA treated cells, three different groups of outermost bundles were tracked: bundles with visible bridging fibers, bundles with no visible bridging fibers and curved bundles extending far from the metaphase plate” and “Examples of each bundle type are shown in insets. From left to right: the outermost bundle in control cells, the outermost bundle with a bridging fiber, the outermost bundle without a bridging fiber and the outermost bundle without kinetochores in HAUS6-depleted cells”, respectively.

4) The authors reported that augmin depletion impairs microtubule poleward flux and conclude that this happens exclusively due to the perturbation of bridging fibers. While the results from this and other studies clearly show that augmin depletion perturbs spindle microtubules in general, it is not clear whether this had a stronger effect on the bridging microtubules (see the comments in point 1). Thus, the impact of augmin depletion on kinetochore microtubules or other antiparallel microtubules within the spindle (e.g. the ones recently shown in O’Toole et al., MboC 2020) cannot be ruled out as a potential cause of the impaired microtubule flux. Also, Steblyanko et al. (EMBO J, 2020) showed that PRC1 depletion hadffectt on microtubule poleward flux in metaphase cells. Since it has been previously shown by the authors of this manuscript that PRC1 depletion disrupts the formation of bridging fibers, it is unlikely that the bridging fibers are the main cause of the augmin depletion-mediated effect on the microtubule flux.

We modified the text on poleward flux to include the contribution of both bridging and k-fibers. We also performed new experiments on U2OS cells and included references to the new work from our lab recently published in Cell Reports (Risteski et al., 2022), which was able to distinguish between the effect of augmin depletion on bridging and k-fiber flux. The effect of augmin on poleward flux is now only a minor point in our revised manuscript which mainly focuses on microtubule arrangements and mitotic fidelity.

However, as requested, we included a comment on PRC1 depletion: “Recent speckle microscopy experiments in RPE1 cells, which were able to separate the effect of augmin on poleward flux of bridging and k-fibers, revealed that both k-fibers and the remaining bridging fibers were significantly slowed down (Risteski et al., 2022). Bridging fibers fluxed faster than k-fibers in control and augmin-depleted cells (Risteski et al., 2022), supporting the model in which poleward flux is largely driven by sliding apart of antiparallel microtubules (Brust-Mascher et al., 2009; Mitchison, 2005; Miyamoto et al., 2004). We propose that augmin depletion results in slower flux of bridging fibers because the remaining bridging microtubules are likely nucleated at the poles, where microtubule depolymerization mechanisms might curb poleward flux speed (Ganem et al., 2005). In contrast, PRC1 depletion does not affect the flux (Risteski et al., 2021 Preprint; Steblyanko et al., 2020) even though it reduces bridging fibers (Kajtez et al., 2016; Polak et al., 2017), possibly because the remaining bridging microtubules are generated away from the poles via augmin and can thus flux freely.”

Minor comments:

1) Introduction: chromatin- and kinetochore- mediated generation of spindle microtubules are ignored when describing the origins of spindle microtubules in human somatic cells.

We included the chromatin- and kinetochore-mediated generation of spindle microtubules in the Introduction. We revised the text as follows: “Spindle microtubules in human somatic cells are generated by several nucleation mechanisms, including centrosome-dependent and augmin dependent nucleation (Kirschner and Mitchison, 1986; Pavin and Tolić, 2016; Petry, 2016; Prosser and Pelletier, 2017; Wu et al., 2008; Zhu et al., 2008), with an addition of chromatin- and kinetochore-dependent nucleation as a third mechanism that contributes to the directional formation of k-fibers (Maiato et al., 2004; Sikirzhytski et al., 2018; Tulu et al., 2006).”

2) The authors proposed less efficient HAUS8 depletion as a potential reason of discrepancy between the siHAUS6 and siHAUS8 results. This should be shown by Western blot, like it is presented for the RNAi efficiency of siHAUS6.

We agree with the reviewer that it would be best to include Western blot for the RNAi efficiency of siHAUS8. However, as we explained in the Methods section, commercially available HAUS8 antibodies resulted in no detectable bands in our hands, regardless of the modifications in the Western blot protocol. We explained this in the Methods section, as follows: “Rabbit polyclonal HAUS8 antibody (diluted 1:1000, PA5-21331, Invitrogen and NBP2-42849, Novus Biologicals) resulted in no detectable bands under these conditions”. For this reason, we performed immunocytochemistry to determine the efficiency of siHAUS8. The discrepancy was now also addressed as a part of our new super resolution STED analysis, which enabled increased precision and where depletion of HAUS6 and HAUS8 produced the same results.

3) The measurements of total PRC1 intensities are mentioned in the manuscript text, but not shown in the figures.

PRC1 measurements are now performed on both RPE1 and HeLa cells with corresponding graphs shown in Figure 4C and Supplementary Figure S4C.

4) Supplementary Videos 3 and 4 are wrongly annotated as Supplementary Videos 1 and 2 in the text.

As we have a new set of videos, this is no longer applicable.

5) Given the spindle length phenotypes are opposite in HeLa and RPE1 cells, in order to be consistent with the other experiments it would be better to perform the PRC1 measurements in RPE1 cells (e.g. using the anti-PRC1 antibody as shown in Supplementary Figure 3B).

We have now performed size measurements in all three cell lines: RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP, RPE1 cells stably expressing PRC1-GFP and HeLa cells stably expressing PRC1-GFP treated with MG-132. These results are now shown in Supplementary Figure S4J-K. The phenotypes remained the same as in the original experiments. We revised the text to better explain the observed differences as follows: “While the spindles in RPE1 cells shortened following augmin depletion, those in HeLa cells were longer (Supplementary Figure S4J), consistent with previous observations on Drosophila S2 cells and Xenopus egg extracts (Goshima et al., 2007; Petry et al., 2011). This difference in spindle length might be due to the overlaps remaining the same length after augmin depletion in RPE1 cells, while being longer and thereby able to push the spindle poles further apart in HeLa cells (Supplementary Figure S4K).”

Please note that due to the extensive amount of new data regarding microtubule arrangements and segregation errors obtained from STED microscopy and live-cell imaging of kinetochores over time, size measurements are now only a small part of our revised story and are, as such, no longer presented in the main figure.

6) Why are the microtubule flux rates for RPE1-PA-GFP-α-tubulin cells measured in this study largely different than the rates reported for the same cell line in Dudka et al., Nat Comms 2018 and Dudka et al., Curr Biol 2019? In order to better understand this difference and strengthen the microtubule flux data, it would be helpful to increase the experimental numbers to match the ones used in the mentioned studies.

We performed photoactivation experiments on a higher number of U2OS cells stably expressing CENP-A GFP, mCherry-tubulin and PA-tubulin (N = 30 measured photoactivation spots in 30 control and HAUS6-depleted cells, see Supplementary Figure S3L-M). U2OS cells with labelled kinetochores and tubulin were used to exclude the potential effects of SiR-tubulin on poleward flux, as well as to better determine the position of the metaphase plate. The results in control cells are comparable to the poleward flux measured in the same cell line (Steblyanko et al., 2020).

Similar to shape and size measurements, due to the extensive and exciting new data regarding microtubule arrangements and segregation errors and because poleward flux was studied into much more detail in a recent Cell Reports study from our lab, this is no longer a major point in our revised manuscript. Poleward flux, along with shape analysis, is now included in the Supplementary Figure 3, as a part of the section about spindle architecture and dynamics.

7) The number of cells used per each experiment should be clearly stated.

In all experiments included in the main figures, we have now performed 3 independent experiments with at least 10 cells each. The numbers are also clearly stated in the captions of figures for all experiments.

Reviewer #1 (Significance (Required)):

This study expands the analysis of augmin contribution to the spindle functions and focuses on its role in formation of the bridging fibers, which is of interest to cell biologists studying mitosis. It clearly shows that in addition to its effect on the k-fibers, augmin depletion also impairs the formation of bridging fibers. However, the exact contribution of the bridging fibers to the spindle functions affected by augmin depletion remains unclear.

We thank the reviewer for the thoughtful comments and are convinced that our new experiments which include the never-before-seen arrangements of microtubules, along with functional analysis of augmin-depleted cells, demonstrated that bridging fibers have a specific k-fiber independent effect on kinetochore tension and a specific role in preventing merotelic attachments.

Since our revised manuscript includes both the first quantitative analysis of the human spindle using STED microscopy, as well as new mechanisms by which augmin ensures mitotic fidelity, it is no longer of interest only to cell biologists studying mitosis but also to a wider scientific community, including cell biologists, molecular biologists and biophysicists interested in microtubules, mitotic and meiotic spindles, cell division, chromosome segregation, aneuploidy, cancer, and development, as well as scientists developing quantitative superresolution imaging protocols for imaging of cellular structures.

Reviewer #2 (Evidence, reproducibility and clarity (Required)):

Summary:

The authors found that the microtubules in the bridging fibres of the mitotic spindle in a human cell line are predominantly supplied via augmin-dependent nucleation. On the other hand, the contribution of augmin to kinetochore fibre formation is ~40%. Augmin-depleted cells showed reduced inter-kinetochore tension and slower poleward flux of spindle microtubules, suggesting that bridging fibres play a role in these events. This study expands our knowledge on the role of augmin and augmin-mediated microtubules in animal somatic cells.

We thank the reviewer for this accurate summary of our work. In addition to the findings nicely described here, we now have an entire new section on the role of augmin in mitotic fidelity, as suggested by Reviewer 3 and presented in the new Figure 2.

Major comments:

– Are the key conclusions convincing?

Yes.

– Should the authors qualify some of their claims as preliminary or speculative, or remove them altogether?

In the current manuscript, the slower flux is attributed solely to the lack of bridging fibres in the augmin-depleted cells. This is an overinterpretation, as the augmin's role in the spindle is not limited to generating bridging fibres.

We modified the text on poleward flux to include the contribution of both bridging and k-fibers. We also performed new experiments on U2OS cells and included references to the new work from our lab recently published in Cell Reports (Risteski et al., 2022), which was able to distinguish between the effect of augmin depletion on bridging and k-fiber flux. Because poleward flux was analyzed into much more detail in this study, and because we produced surprising new data regarding microtubule arrangements and segregation errors, poleward flux is no longer a major point in our revised manuscript. It is now included in the Supplementary Figure 3 along with shape analysis, as a part of the section about spindle architecture and dynamics.

The text about flux was modified as follows: “Recent speckle microscopy experiments in RPE1 cells, which were able to separate the effect of augmin on poleward flux of bridging and k-fibers, revealed that both k-fibers and the remaining bridging fibers were significantly slowed down (Risteski et al., 2022). Bridging fibers fluxed faster than k-fibers in control and augmin-depleted cells (Risteski et al., 2022), supporting the model in which poleward flux is largely driven by sliding apart of antiparallel microtubules (Brust-Mascher et al., 2009; Mitchison, 2005; Miyamoto et al., 2004). We propose that augmin depletion results in slower flux of bridging fibers because the remaining bridging microtubules are likely nucleated at the poles, where microtubule depolymerization mechanisms might curb poleward flux speed (Ganem et al., 2005). In contrast, PRC1 depletion does not affect the flux (Risteski et al., 2021 Preprint; Steblyanko et al., 2020) even though it reduces bridging fibers (Kajtez et al., 2016; Polak et al., 2017), possibly because the remaining bridging microtubules are generated away from the poles via augmin and can thus flux freely.”

– Would additional experiments be essential to support the claims of the paper? Request additional experiments only where necessary for the paper as it is, and do not ask authors to open new lines of experimentation.

No.

– Are the suggested experiments realistic in terms of time and resources? It would help if you could add an estimated cost and time investment for substantial experiments.

N/A

– Are the data and the methods presented in such a way that they can be reproduced

Yes.

– Are the experiments adequately replicated and statistical analysis adequate?

Yes.

Minor comments:

– Specific experimental issues that are easily addressable.

None.

– Are prior studies referenced appropriately?

Yes.

– Are the text and figures clear and accurate?

1) Page 15: "To determine the curvature of the bundles, we ….. with all other bundles types (Figure 5E)." – I could not understand this sentence well, and would like to ask for a revision.

The text has now been changed to: “To gain insight into the contribution of each of these functionally distinct microtubule bundles to the maintenance of spindle geometry, we traced the outermost bundles in HAUS6 siRNA treated RPE1 cells imaged using STED microscopy and fitted a circle to the bundle outline (Supplementary Figure S3I, see Methods)”.

2) The following words may be too strong:

Page 20: whereas k-fiber microtubules are "mainly" nucleated in an augmin-independent manner (could 61% contribution be called "mainly?").

We revised this sentence on page 24 as follows: “K-fibers were also thinner, though to a lesser extent, indicating that they are largely nucleated in an augmin-independent manner, at the centrosome or kinetochores and chromosomes.”

Page 21, bottom: "demonstrates".

As we changed this section of the manuscript, this is no longer applicable.

– Do you have suggestions that would help the authors improve the presentation of their data and conclusions?

No.

Reviewer #2 (Significance (Required)):

– Describe the nature and significance of the advance (e.g. conceptual, technical, clinical) for the field.

The presence of bridging fibres has been recognised for decades; however, until recently, little attention has been paid to this structure from a mechanistic and functional point of view. The Tolic lab has been shedding light on this structure for the past several years. The current study represents another step forward in the research of the origin and function of bridging fibres.

– Place the work in the context of the existing literature (provide references, where appropriate).

Augmin's critical contribution to microtubule nucleation in the human somatic spindle has been well documented, as cited by the authors. The current study is the first to show that augmin also contributes to bridging fibres. The >70% contribution may be more than expected, given that centrosomal microtubules frequently reach the spindle midzone.

Reduced inter-kinetochore tension has also been documented, but previous studies attributed this exclusively to reduced number of kinetochore microtubules. The current study has revised this view.

We thank the reviewer for correctly pointing out the main points of our original manuscript. These results are now strengthened by new experiments, and accompanied by STED microscopy imaging and a section about the role of augmin-nucleated microtubules in the maintenance of mitotic fidelity.

– State what audience might be interested in and influenced by the reported findings. Spindle researchers.

Since our revised manuscript includes both the first quantitative analysis of the human spindle using STED microscopy, as well as new mechanisms by which augmin ensures mitotic fidelity, it is no longer of interest only to spindle researchers but also to a wider scientific community, including cell biologists, molecular biologists and biophysicists interested in microtubules, mitotic and meiotic spindles, cell division, chromosome segregation, aneuploidy, cancer, and development, as well as scientists developing quantitative super resolution imaging protocols for imaging of cellular structures.

– Define your field of expertise with a few keywords to help the authors contextualize your point of view. Indicate if there are any parts of the paper that you do not have sufficient expertise to evaluate.

This review is written by a researcher who is familiar with the literature of the mitotic spindle.

We thank the reviewer for an accurate summary of our work and perceptive comments.

Reviewer #3 (Evidence, reproducibility and clarity (Required)):

In this study Manenica et study how the presence of the augmin complex affects the overall spindle architecture and the different types of spindle microtubules. The authors propose that depletion of augmin affects particularly bridging microtubules, leading to their disappearance on sister-kinetochores located in the central part of the metaphase plate.

Overall the manuscript is well written, clear and supported by excellent explanatory schemes. The main conclusion of the manuscript, i.e. that augmin plays an essential role in the formation of bridging microtubules is generally well supported by the data. A number of other conclusions, however, are less well supported by the data and would benefit from a number of additional experiments, repetitions or analysis. Specifically:

We thank the reviewer for complimenting the clarity of our manuscript, and the data supporting the main conclusion. Other conclusions, which the reviewer states were not equally well supported by the data, are now strengthened by a new set of experiments using STED microscopy, increased repetitions (at least 30 cells from 3 independent experiments) and multiple new analyses. In addition, our revised manuscript now also includes both the first quantitative analysis of the human spindle using STED microscopy, as well as an entire new section on the mechanisms by which augmin ensures mitotic fidelity, as thoughtfully suggested below.

1) Throughout all the figures the authors use a t-test, which is fine when comparing two conditions, but not for multiple experimental conditions. The authors should instead use an ANOVA test or apply a Bonferroni correction. This can strongly affect the significance of some of the reported results.

We agree and have performed either ANOVA with post-hoc Tukey test or Mann-Whitney U-test instead of t-test where appropriate throughout the whole manuscript. The statistical analyses that were used are clearly stated in the captions of the figures.

2) Another general concern is that the authors rely throughout the manuscript on live cell imaging data from few cells (5-10). Live cell imaging data has the advantage to avoid fixation artifacts, but the low sample size is a major concern, as for every experiment the authors rely on 10 cells (and for inter-kinetochore distances on 5) in three independent experiments overall. This means that two our of three of those independent experiments are based on 3 cells only, which is too low, given that siRNA depletions are known to be variable in their efficiencies. With such low number of cells, there is always the danger of an unconscious selection bias, which can skew a result. Just to take an example the spindle length, structure and density for HAUS-8 depleted RPE1 cells looks very different in the examples show in Figure 1B, 2A, or 5B.It is therefore essential to work with a higher sample size, at minimum 10 cells per independent experiment.

This is a valid critique, which we addressed by performing 3 independent experiments with at least 10 cells each for all of our new analyses included in the main figures.

3) Throughout all experiments the authors use 100nm Sir-Tubulin, which in our hands already leads to substantial changes in microtubule dynamics, as it stabilizes spindle microtubules. I understand why the authors did this, as they wanted to also stain for weak bridging fibers, but tt would be important to validate some of the obtained results with an independent approach, for example fixed-cell imaging and tubulin staining, to rule out artifacts introduced by SiR-tubulin.

We thank the reviewer for this suggestion. To validate our results, we performed tubulin immunostaining, as suggested. Moreover, we imaged the immunostained cells by using super resolution STED microscopy, which significantly increased the precision of our measurements, and included these results in the main figures (Figure 1, Figure 2J and K, Figure 3, Figure 5). The data obtained from cells stained with SiR-tubulin and imaged using live-cell confocal microscopy are now shown in Supplementary figures.

4) Figure 1: Given that the authors later report that augmin affects more strongly bridging fibers in the central part of the spindle, how were the values in terms of microtubule densities obtained in the experiments in Figure 1: only on the outer microtubules, or overall in the spindle?

Values in Figure 1 (now Figure 3) were obtained overall in the spindle. We described the selection in the text as follows: “We measured tubulin signal intensity of randomly selected bridging (Ib) and k-fibers (Ik) which had no other microtubules in their immediate neighborhood…”

5) Figure 2: the authors conclude that depletion of augmin has a much stronger effect on the bridging fibers located in the central part of the spindle. This is a very interesting result, but it begs the question as to the origin of this difference. If the authors analyze in control cells the interkinetochore distances and the density of the bridging fibers of the kinetochores located in the central part of the metaphase plate vs those located at the outer part of the plate, do they already see a difference? In other words, is the effect of augmin due to already weaker bridging fibers in the central part of the spindle, or is the depletion effect indeed specific for those bridging fibers located in the middle. This analysis should be possible with the existing data (+ a higher sample size)

This was now analyzed in the cells imaged using STED microscopy with increased resolution and a higher sample size. From our new data, it is clear that the depletion effect is indeed specific for bridging fibers located in the middle as there was no significant difference between the interkinetochore distance in the inner and the outer part of the spindles in control cells (Figure 5D and Supplementary Figure S5B). The same trend can also be seen for bridging fiber density (Figure 3H). We modified the text as follows: “However, we noticed that the interkinetochore distance was smaller in the inner part of the spindle in augmin-depleted cells (Figure 5A-D, Supplementary Figure S5B), where bridging fibers were most severely impaired (Figure 3H and 4A). This was not the case in control cells, which showed no difference in interkinetochore distance between the inner and the outer part of the spindle (Figure 5D, Supplementary Figure S5B).”

6) Figure 4: the authors study spindle width, length and diameter of the metaphase plate in a small number of cells (10). One concern is that these values might change as cells progress from late prometaphase to anaphase onset (metaphase plate width decreases for example). Given the low number of cells the authors do not know if they are comparing cells at similar mitotic times. To circumvent this issues, they could: either arrest the cells with MG132 for 1 hour, to obtain an endpoint, or record these different values as cells progress through mitosis and thus be able to compare similar conditions.

We agree with this suggestion, and we performed new experiments by arresting the cells with MG-132: “… in HeLa (Kajtez et al., 2016) and RPE1 (Asthana et al., 2021) cells stably expressing PRC1-GFP with and without MG-132 treatment (Figure 4A-B, Supplementary Figure S4A).” We measured spindle width, length and diameter of the metaphase plate in arrested RPE1 cells stably expressing CENP-A-GFP and Centrin1-GFP, RPE1 cells stably expressing PRC1-GFP, and HeLa cells stably expressing PRC1-GFP. We treated the cells with MG-132 for 30 minutes, as this was in our hands enough to arrest the cells, without causing other changes, e.g., problems with spindle orientation that occur after 1 hour of treatment. The results are now part of the Supplementary Figure S4 and are obtained from three independent experiments with at least 10 cells per experiment.

7) In the discussion the authors conclude that the longer bundles and the reduction in microtubule poleward flux is due to the absence of bridging microtubules. This is an over-interpretation as augmin could in theory affect these parameters independently of the bridging microtubules, longer bundles could be generally due to the reduced number of microtubules in the k-fibers and the bridging microtubules. A better control would be to affect bridging microtubules with an independent tool, such as PRC1 depletion, and to measure these paramenters in the same RPE1 cell line, since differences can arise from cell line to cell line as the authors also document in their study (for example spindle length in Figure 4).

We modified the Discussion based on new results, so these statements are now in the Results section. For the long, curved bundles, we modified the sentence as follows: “These bundles likely arose either due to PRC1 crosslinking excessively long astral microtubules that were now able to reach the spindle midzone or due to PRC1 activity combined with the excess of free tubulin present as a consequence of less tubulin being incorporated in bridging and k-fibers.”

Regarding the reduced poleward flux following augmin depletion, we revised the text as follows: “Recent speckle microscopy experiments in RPE1 cells, which were able to separate the effect of augmin on poleward flux of bridging and k-fibers, revealed that both k-fibers and the remaining bridging fibers were significantly slowed down (Risteski et al., 2021 Preprint). Bridging fibers fluxed faster than k-fibers in control and augmin-depleted cells (Risteski et al., 2021 Preprint), supporting the model in which poleward flux is largely driven by sliding apart of antiparallel microtubules (Brust-Mascher et al., 2009; Mitchison, 2005; Miyamoto et al., 2004). We propose that augmin depletion results in slower flux of bridging fibers because the remaining bridging microtubules are likely nucleated at the poles, where microtubule depolymerization mechanisms might curb poleward flux speed (Ganem et al., 2005). In contrast, PRC1 depletion does not affect the flux (Risteski et al., 2021 Preprint; Steblyanko et al., 2020) even though it reduces bridging fibers (Kajtez et al., 2016; Polak et al., 2017), possibly because the remaining bridging microtubules are generated away from the poles via augmin and can thus flux freely.”

However, due to the extensive and thrilling new data regarding microtubule arrangements and segregation errors and because poleward flux was studied into much more detail in a recent Cell Reports study from our lab (Risteski et al., 2022), this is no longer a major point in our revised manuscript. Poleward flux, along with shape analysis, is now included in the Supplementary Figure 3, as a part of the section about spindle architecture and dynamics.

Minor comment:

– The reported flux rate for control-depleted cells is substantially higher than the flux rates normally reported for human cells. This could be due to the experimental conditions (slight changes in temperature), but at minimum the authors should comment on this.

We performed photoactivation experiments on a higher number of U2OS cells stably expressing CENP-A GFP, mCherry-tubulin and PA-tubulin (N = 30 measured photoactivation spots in 30 control and HAUS6-depleted cells, see Supplementary Figure S3L-M). U2OS cells with labelled kinetochores and tubulin were used to exclude the potential effects of SiR-tubulin on poleward flux, as well as to better determine the position of the metaphase plate. The results in control cells are comparable to the poleward flux measured in the same cell line (Steblyanko et al., 2020).

Please note that, as previously stated, this is no longer a major point in our revised manuscript and can be found in Supplementary Figure 3.

Reviewer #3 (Significance (Required)):

The significance of the study is that the authors performed a detailed description of the effects of augmin depletion on the spindle architecture, in particular bridging fibers. Nevertheless, many of the reported results are already known (and as cited by the authors): the reduction in interkinetochore distances or the change in spindle architecture. The 3 main novel results, is the fact that augmin affects more bridging microtubules, particularly in the central part of the spindle, and that it also affect poleward microtubule flux, which limits the impact of this study to a specialized mitotic spindle audience. Nevertheless, if the authors address the reviewers concerns, this could be a nice, descriptive study for the mitotic field.

One way to expand the significance of this study would be to test how augmin depletion and the lack of bridging microtubules in the central part of the metaphase plate affects chromosome segregation. Does the specific absence of bridges in this part lead to more lagging chromosomes, chromosome segregation errors, or micronuclei amongst sister chromatids located in the central part of the spindle? Is there a differential anaphase A speed for those kinetochore vs those at the periphery that still are associated to bridging fibers? Such a functional approach could allow to highlight the most interesting aspect of this study, the spatial difference in the effects of augmin depletion. Such experiments would, however, not be part of a revision, but rather a substantial enhancement of the present study.

Patrick Meraldi

This is a great idea that we truly appreciate! We performed new experiments to study lagging chromosomes and indeed found that they were more often located in the inner part of the spindle in HAUS6-depleted than in control cells, which is likely due to the specific impairment of bridging fibers in that area. We also found that lagging chromosomes typically had a lower interkinetochore distance and a higher kinetochore tilt just before the onset of anaphase, which we interpret as a signature of perturbed bridging fibers. Using STED microscopy, we were able to clearly see merotelic attachments at kinetochores lacking a bridging fiber in metaphase following augmin depletion. Thus, we propose that augmin-nucleated bridging fibers prevent merotelic attachments by creating a nearly parallel and highly bundled microtubule arrangement unfavorable for creating additional attachments. On the other hand, augmin-nucleated k-fibers produce robust force required to resolve errors during anaphase. We dedicated an entire new section on pages 6-11 and a new Figure 2 to these exciting new results.

We agree with the Reviewer 3 that these results substantially increase the significance of this study, which is now no longer descriptive nor of interest only to the mitotic field, but offers a new mechanism that safeguards mitotic fidelity, which makes it of interest to a wider scientific community. This includes cell biologists, molecular biologists and biophysicists interested in microtubules, mitotic and meiotic spindles, cell division, chromosome segregation, aneuploidy, cancer, and development, as well as scientists developing quantitative superresolution imaging protocols for imaging of cellular structures.

We are thankful to the reviewer for raising such an interesting question, and motivating us to use a functional approach to study the mechanism by which the observed effects of augmin depletion compromise mitotic fidelity.

https://doi.org/10.7554/eLife.83287.sa2

Article and author information

Author details

  1. Valentina Štimac

    Division of Molecular Biology, Ruđer Bošković Institute, Zagreb, Croatia
    Contribution
    Conceptualization, Software, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft
    Contributed equally with
    Isabella Koprivec
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0398-5493
  2. Isabella Koprivec

    Division of Molecular Biology, Ruđer Bošković Institute, Zagreb, Croatia
    Contribution
    Conceptualization, Software, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft
    Contributed equally with
    Valentina Štimac
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6486-8261
  3. Martina Manenica

    Division of Molecular Biology, Ruđer Bošković Institute, Zagreb, Croatia
    Contribution
    Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
  4. Juraj Simunić

    Division of Molecular Biology, Ruđer Bošković Institute, Zagreb, Croatia
    Contribution
    Conceptualization, Supervision
    Competing interests
    No competing interests declared
  5. Iva M Tolić

    Division of Molecular Biology, Ruđer Bošković Institute, Zagreb, Croatia
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing – review and editing
    For correspondence
    tolic@irb.hr
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1305-7922

Funding

European Research Council (ERC Consolidator Grant 647077)

  • Iva M Tolić

Croatian Science Foundation Cooperation Programme (HRZZ project PZS-2019-02-7653)

  • Iva M Tolić

European Regional Development Fund (QuantiXLie Centre of Excellence (KK.01.1.1.01.0004))

  • Iva M Tolić

European Research Council (ERC Synergy Grant 855158)

  • Iva M Tolić

European Regional Development Fund (IPSted (KK.01.1.1.04.0057))

  • Iva M Tolić

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Alexey Khodjakov, Ina Poser and Tony Hyman, Thomas Surrey, Emanuele Roscioli, Andrew McAinsh, Mariola Chacon and Marin Barišić for the cell lines. We also thank Marko Šprem and Abberior team for help with developing microscopy protocols, Ivana Štimac and Marko Šprem for help with creating the MatLab and R scripts, Josip Čačković and Arian Ivec for technical assistance with the initial experiments and calculations, all members of Tolić and Pavin groups for discussions and advice, Ivana Šarić for the drawings. Funding This work was funded by the European Research Council (ERC Consolidator Grant, GA number 647077 and ERC Synergy Grant, GA number 855158), Croatian Science Foundation Cooperation Programme with Croatian Scientists in Diaspora “Research Cooperability” (HRZZ Project PZS-2019-02-7653), as well as QuantiXLie Centre of Excellence (Grant KK.01.1.1.01.0004) and IPSted (KK.01.1.1.04.0057) projects cofinanced by the Croatian Government and European Union through the European Regional Development Fund - the Competitiveness and Cohesion Operational Programme.

Senior Editor

  1. Anna Akhmanova, Utrecht University, Netherlands

Reviewing Editor

  1. Thomas Surrey, Centre for Genomic Regulation (CRG), Spain

Publication history

  1. Preprint posted: September 10, 2020 (view preprint)
  2. Received: September 7, 2022
  3. Accepted: September 26, 2022
  4. Accepted Manuscript published: October 21, 2022 (version 1)
  5. Version of Record published: November 7, 2022 (version 2)

Copyright

© 2022, Štimac, Koprivec et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Valentina Štimac
  2. Isabella Koprivec
  3. Martina Manenica
  4. Juraj Simunić
  5. Iva M Tolić
(2022)
Augmin prevents merotelic attachments by promoting proper arrangement of bridging and kinetochore fibers
eLife 11:e83287.
https://doi.org/10.7554/eLife.83287

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