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. Author manuscript; available in PMC: 2020 Dec 10.
Published in final edited form as: Methods Cell Biol. 2020 Feb 24;158:91–116. doi: 10.1016/bs.mcb.2020.01.004

A Comparative Analysis of Methods to Measure Kinetochore-Microtubule Attachment Stability

Jessica D Warren 1,2, Bernardo Orr 3,4, Duane A Compton 1,2
PMCID: PMC7727308  NIHMSID: NIHMS1649737  PMID: 32423652

Abstract

During mitosis, spindle microtubules dynamically attach to and detach from kinetochores in a precise and regulated fashion. To ensure mitotic fidelity, kinetochore-microtubule (k-MT) attachments must be stable enough to satisfy the spindle assembly checkpoint (SAC), but sufficiently unstable to facilitate the correction of mal-oriented attachments. Different methods are available to assess k-MT stability in both live and fixed cells, but a comparative survey of these methods has not yet been reported. Here, we evaluate several quantitative and semi-quantitative methods for determining k-MT stability and apply each technique to illustrate changes in spindle microtubule dynamics upon perturbation with physiologically relevant concentrations of microtubule stabilizing (Taxol) and destabilizing (UMK57 and nocodazole) compounds. We discuss the utility of each technique for defining specific features of spindle microtubule dynamics and k-MT attachment stability.

Keywords: Mitosis, Spindle, Chromosomes, Kinetochore, Microtubule, Tubulin

I. Introduction

The mitotic spindle apparatus is a microtubule-based structure responsible for chromosome segregation during cell division. Microtubules within the spindle are classified according to their plus-end interactions. Astral microtubules project toward the cell cortex, interpolar microtubules extend between the two spindle poles, and kinetochore microtubules (k-MTs) interact with kinetochore protein complexes via sidewall or end-on attachment. All spindle microtubules retain dynamic instability (T. Mitchison & Kirschner, 1984), albeit with differences. Non-k-MTs undergo rapid switches between growth and shrinkage, leading to high dynamicity, whereas k-MTs switch less frequently between these states, resulting in lower dynamicity (Cassimeris, Rieder, Rupp, & Salmon, 1990; T. J. Mitchison, 1988; Zhai, Kronebusch, & Borisy, 1995).

Two key features regulating the dynamicity of k-MTs are the rate of microtubule attachment and the rate of microtubule detachment from the kinetochore. The enhanced stability of k-MTs with respect to non-k-MTs is considered to be a consequence of their interaction with the kinetochore (Zhai et al., 1995) and many kinetochore proteins have been reported to modulate k-MT attachment/detachment rate (reviewed in Godek, Kabeche, & Compton, 2015). Attachments between kinetochores and microtubules must be dynamic in order to facilitate chromosome movement (Kops, Saurin, & Meraldi, 2010) and error correction (Bakhoum, Thompson, Manning, & Compton, 2009). Published work demonstrates that k-MT attachment stability is precisely regulated within a narrow range to ensure that chromosomes are segregated faithfully with each cell division (Bakhoum, Genovese, & Compton, 2009). Thus, methods to quantitatively analyze the stability of k-MT attachments are important to provide the research community with the capacity to evaluate the molecular mechanisms responsible for high-fidelity chromosome segregation.

The correction of erroneous k-MT attachments is essential for faithful chromosome segregation. A rate limiting step in the error correction process is the frequency with which microtubules detach from kinetochores (Nicklas & Ward, 1994). Because of this central importance of k-MT attachment stability in mitotic fidelity, various methods have been reported to assess the stability of this class of spindle microtubules in cultured cells using fluorescence microscopy. These methods differ in their ease of implementation and in the type of quantitative information they provide. Here, we have conducted a comparative analysis of each of the quantitative and semi-quantitative assays commonly used to measure k-MT attachment stability. We have tested each of these assays with established microtubule stabilizing (Taxol) and destabilizing (UMK57) compounds at physiologically relevant concentrations ((Orr, Talje, Liu, Kwok, & Compton, 2016; Yvon, Wadsworth, & Jordan, 1999; Zasadil et al., 2014). This comparative analysis identifies the merits and limitations of each method and should aid investigators in choosing the most appropriate assay for their research question. Furthermore, to provide an example of how these assays can be combined to better our understanding of how microtubule dynamics affect mitotic processes, we use these assays to document the effects of low dose nocodazole treatment on the mitotic spindle.

Although we restrict our comparisons to human cells, the methods described here should be suitable for evaluation of k-MT attachment stability in any cultured cells, under a variety of conditions of cellular perturbation. Unless otherwise stated, all cell lines were maintained at 37°C in a humidified atmosphere with 5% CO2 and were cultured in Dulbecco’s modified Eagle’s medium, (DMEM; Invitrogen) supplemented with 10% FBS (HyClone), 10 mM HEPES (GE Healthcare), 250 μg/L Amphotericin B (Sigma Aldrich), 50 U/mL penicillin (Mediatech) and 50 μg/mL streptomycin (Mediatech). All live cell imaging was performed in FluoroBrite DMEM (Gibco) supplemented with 10% FBS, 4mM L-glutamine (Thermo Fisher Scientific), 10mM HEPES, 250 μg/L Amphotericin B, 50 U/mL penicillin and 50 μg/mL streptomycin. UMK57 was used at 100 nM, as previously reported (Orr et al., 2016) and Taxol (Paclitaxel, Biotang) was used at 5 nM. All controls for drug treatment were performed using 0.1% DMSO. Importantly, Taxol and UMK57 treatment did not grossly alter spindle morphology (Figure 1).

Figure 1. Spindle Morphology.

Figure 1.

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Representative images (maximum intensity projections) of tubulin fluorescence intensity. U2OS cells were treated for 1 hour with 5 μM MG-132 before the addition of DMSO, 5nM Taxol, or 100nM UMK57 for 1 hour. Cells were fixed in 1% glutaraldehyde, as previously described (Thompson & Compton, 2011) and stained for α-tubulin (green) and DNA (blue). Scale bar, 5 μm.

II. Photoactivation of Spindle Microtubules

k-MTs continuously undergo cycles of detachment, depolymerization, polymerization, and re-attachment (Zhai et al., 1995). Detachment is assumed to be the rate limiting step in this cycle, since microtubule shrinkage and growth occur rapidly (T. Mitchison, Evans, Schulze, & Kirschner, 1986; T. Mitchison & Kirschner, 1984). In this way, k-MT stability is predominantly determined by the rate at which microtubules detach from kinetochores. Spindle microtubules may be selectively marked by photobleaching or photoactivation techniques; however, photoactivation of fluorescently-labeled tubulin (either chemically conjugated or expressed as GFP-tagged fusion protein) is an optimal method for the measurement of k-MT detachment rate (Kabeche & Compton, 2013; Zhai et al., 1995), since photobleached marks are quickly lost due to the rapid recovery of fluorescently labeled non-k-MTs (Cassimeris et al., 1990; Gorbsky, Simerly, Schatten, & Borisy, 1990; Saxton et al., 1984; Wadsworth & Salmon, 1986). By measuring fluorescence dissipation of a photoactivated spindle region, k-MT detachment rate can be quantified as a direct measure of k-MT stability. Since both k-MTs and non-k-MTs are fluorescently marked in this assay, the detachment rate for non-k-MTs can also be measured with appropriate reduction of image acquisition intervals. Additionally, the relative proportion of k-MT and non-k-MT populations within the spindle can be inferred from the slow and fast components of fluorescence decay curves, respectively (Zhai et al., 1995).

A. Image Acquisition and Data Analysis

The photoactivation assay was performed with U2OS (ATCC®, HTB-96) cells stably expressing photoactivatable GFP-α-tubulin (maintained in DMEM + 1 mg/mL G418 (InvivoGen)). Cells were treated with DMSO, Taxol, or UMK57 in FluoroBrite DMEM for 1 hour. 5 μM MG-132 (Sigma-Aldrich) was added prior to rose chamber assembly to prevent mitotic exit. Cells were maintained at 37°C during image acquisition. Images were acquired with a Plan Apo VC 100x, 1.4 NA, oil immersion objective (Nikon) using a QuorumWaveFX-X1 spinning disk confocal system on a Nikon Eclipse Ti microscope, equipped with an ILE laser source (Andor Technology), a Mosaic digital mirror (Andor Technology), and a Hamamatsu ImageEM camera. Mitotic cells were identified by differential interference contrast (DIC) microscopy and classified as prometaphase or metaphase on the basis of chromosome alignment. Photoactivation was performed with a 405 nm laser (35% power, 500ms pulse) on a rectangular region of interest over one half of the mitotic spindle. Fluorescence z-stacks with a 1 μm step size (7 slices) were captured every 15 seconds for 4 minutes. For increased precision in measuring the turnover of rapid MT populations, z-stacks can be acquired every 5 seconds.

Fluorescence dissipation after photoactivation (FDAPA) was quantified from maximum intensity projections using MetaMorph® software (Molecular Devices). Average pixel intensities were measured within an area surrounding the region of highest fluorescence intensity and background subtraction was performed using an equally sized area from the non-activated half-spindle at each time point. Fluorescence intensities were corrected for photobleaching using values of fluorescence loss obtained from cells photoactivated in the presence of 1 μM Taxol (where the photoactivated region did not dissipate). Fluorescence intensities were then normalized to the first time point after photoactivation for each cell. To measure microtubule stability, the average fluorescence intensity at each time point was fit to a two-phase exponential decay curve [F(t) = A1e−k1t + A2e−k2t] using GraphPad software, where F(t) is measured photoactivated fluorescence at time t, A1 represents the percentage of fluorescence from the fast decay process, and A2 represents the percentage of fluorescence from the slow decay process, with decay rates of k1 and k2, respectively (Zhai et al., 1995). For high stringency, we only consider curves with good fit (R2 ≥ 0.98). When the two-phase decay equation is solved, the rate constants and the percentage of fluorescence attributable to the fast (non-k-MT) and slow (k-MT) processes are obtained. The half-life (t1/2) for each process was calculated as ln2/k. The measured k-MT half-life represents the time it takes for half of the fluorescently labeled k-MTs to detach from kinetochores (Bakhoum, Genovese, et al., 2009).

B. Merits and Limitations

This method is suitable for measuring several parameters of spindle microtubule stability. k-MT half-life measurements are a direct readout of microtubule detachment rate and the photoactivation assay is sufficiently sensitive to discern significantly different increases or decreases in k-MT attachment stability. This method also provides the researcher with data for non-k-MT turnover, although the sensitivity for detecting changes in non-k-MT stability using this method remains to be determined.

Photoactivation detects the increase in k-MT stability between prometaphase and metaphase control cells (Figures 2E and 2G), as previously demonstrated (Kabeche & Compton, 2013). As shown in Figures 2A–2D, perturbation of k-MT dynamics through Taxol or UMK57 treatment alters the rate at which fluorescence is lost from the photoactivated region and redistributed to the rest of the spindle. Taxol treatment stabilizes k-MT attachments, as demonstrated by the 1.96 ± 0.28 minute and 1.60 ± 0.62 minute increase in k-MT half-life in prometaphase and metaphase, respectively (Figure 2E). The previously characterized MCAK agonist, UMK57, has been shown to decrease k-MT half-life by 1.60 ± 0.39 minutes in metaphase (Figure 2G; Orr et al., 2016). Although each compound alters metaphase k-MT dynamics by ~30% (either increased or decreased stability depending on which compound is applied), an important distinction between the treatments is their mode of action. Since Taxol binds directly to microtubules, both k-MTs and non-k-MTs are stabilized by Taxol treatment. However, UMK57 treatment is selective for k-MTs, since the compound targets MCAK, a kinesin which plays an important role at the centromere to depolymerize k-MTs (Manning et al., 2007; Wordeman & Mitchison, 1995). This distinction manifests as a shift in the percentage of photoactivated fluorescence intensity that is attributable to the slow decay process (Figures 3F and 3H). The percentage of fluorescence in the slow decaying population increased by ~30% in both prometaphase and metaphase after Taxol treatment (Figure 2F), while UMK57 treatment did not affect the percentage of fluorescence in the slow decaying population in metaphase (Figure 2H). However, UMK57 treatment did result in a small, but significant, increase in the percentage of fluorescence in the slow decaying population in prometaphase (Figure 2H).

Figure 2. Photoactivation of Spindle Microtubules.

Figure 2.

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A) Examples of normalized fluorescence dissipation after photoactivation in prometaphase for individual DMSO (black), Taxol (dark gray), or UMK57 (light gray)-treated cells that are representative of the average k-MT half-life in (E, G). B) Representative DIC and fluorescence images (maximum intensity projections) of photoactivated, prometaphase U2OS cells treated with DMSO, Taxol, or UMK57. Scale bar, 5 μm. pre-PA, pre-Photoactivation. C) Examples of normalized fluorescence dissipation after photoactivation in metaphase for individual DMSO (black), Taxol (dark gray), or UMK57 (light gray)-treated cells that are representative of the average k-MT half-life in (E, G). D) Representative DIC and fluorescence images of photoactivated, metaphase U2OS cells treated with DMSO, Taxol, or UMK57. Scale bar, 5 μm. pre-PA, pre-Photoactivation. E) Average k-MT half-life for DMSO (gray) or Taxol (black)-treated U2OS cells expressing photoactivatable GFP-α-tubulin. Error bars indicate SEM; n ≥ 20 cells per condition; *p ≤ 0.05; ***p ≤ 0.001 using two-tailed t test. F) Percentage of photoactivated fluorescence intensity attributable to the slow decay process. Data represent the mean percentage for DMSO (gray) or Taxol (black)-treated cells. Error bars indicate SEM; n ≥ 20 cells per condition; ***p ≤ 0.001 using two-tailed t test. G-H) Data reproduced from Cell Reports, 17(7), Orr, B., Talje, L., Liu, Z., Kwok, B. H., & Compton, D. A., Adaptive Resistance to an Inhibitor of Chromosomal Instability in Human Cancer Cells, 1755–1763, 2016, with permission from Elsevier. G) Average k-MT half-life for DMSO (gray) or UMK57 (black)-treated U2OS cells expressing photoactivatable GFP-α-tubulin. Error bars indicate SEM; n ≥ 12 cells per condition; *p ≤ 0.05; ***p ≤ 0.001 using two-tailed t test. H) Percentage of photoactivated fluorescence intensity attributable to the slow decay process. Data represent the mean percentage for DMSO (gray) or UMK57 (black)-treated cells. Error bars indicate SEM; n ≥ 12 cells per condition; *p ≤ 0.05 using two-tailed t test.

Figure 3. Nocodazole or Cold Shock.

Figure 3.

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A) Representative immunofluorescence images of prometaphase phenotypes observed in the nocodazole shock assay. Images represent maximum intensity projections of deconvolved z-stacks. U2OS cells fixed and stained for α-tubulin (green) and ACA (red). Scale bar, 10 μm. B) Stacked bar graphs display the percentage of prometaphase (left) and metaphase (right) phenotypes observed in the nocodazole shock assay. Microtubule levels were classified as either low polymer (white), high polymer (gray), or spindle-like (black). The high polymer and spindle-like categories were significantly different between the DMSO and Taxol treatments in both prometaphase and metaphase. The high polymer and spindle-like categories were significantly different between the DMSO and UMK57 treatments in prometaphase, but there was no statistically significant difference between the DMSO and UMK57 treatments in metaphase. Each category was analyzed with a two-tailed Fisher’s exact test (n = 109 cells per condition); **p ≤ 0.01; ***p ≤ 0.001; n.s., p ≥ 0.05. C) Representative immunofluorescence images of metaphase phenotypes observed in the cold shock assay. Images represent maximum intensity projections of deconvolved z-stacks. U2OS cells fixed and stained for α-tubulin (green) and ACA (red). Scale bar, 10 μm. D) Stacked bar graphs display the percentage of prometaphase (left) and metaphase (right) phenotypes observed in the cold shock assay. Microtubule levels were classified as either low polymer (white), high polymer (gray), or spindle-like (black). All three phenotypic categories were significantly different between the DMSO and Taxol treatments in both prometaphase and metaphase. There was no statistically significant difference between the DMSO and UMK57 treatments in either mitotic stage. Each category was analyzed with a two-tailed Fisher’s exact test (n ≥ 135 cells per condition); **p ≤ 0.01; ***p ≤ 0.001; n.s., p ≥ 0.05.

The limitation of this technique is that it can be technically challenging for certain cell types, such as those with small spindles. Furthermore, image acquisition and data analysis are both time- and labor-intensive. An attribute of this assay is that it can be used to measure rates of poleward microtubule flux within the spindle (Kabeche & Compton, 2013; T. J. Mitchison, 1989). Although it is not demonstrated here, the measurement of flux rate is straight-forward and can be calculated using the time-lapse images acquired after photoactivation.

III. Nocodazole or Cold Shock Assays

Unstable spindle microtubules disassemble when exposed to cold temperatures (Brinkley & Cartwright, 1975; Rieder, 1981) or a high dose of the tubulin-sequestering drug, nocodazole (Cassimeris et al., 1990; Gayek & Ohi, 2014). In these immunofluorescence based assays, the amount of tubulin polymer that remains after rapid cold or nocodazole treatment (shock) is an indirect readout for microtubule turnover, since short-lived microtubules are quickly depolymerized, while longer-lived microtubules (e.g. k-MTs) persist. The cold and nocodazole shock assays used here were adapted from the methods reported in Gayek & Ohi, 2014.

A. Methodology

For the nocodazole shock assay, U2OS cells were arrested in mitosis with 5 μM MG-132 for 2 hours before the addition of DMSO, Taxol, or UMK57 for 1 hour. Cells were then treated with media containing 5 μM nocodazole for 6 minutes and were permeabilized for 20 seconds at room temperature in PermFix (100 mM K 1,4-piperazinediethanesulfonic acid, pH 6.8, 0.2% Triton X-100, 10 mM K ethylene glycol tetraacetic acid, 1 mM MgCl2) before fixation. Cells were fixed in cold methanol for 10 minutes at −20°C. For the cold shock assay, U2OS cells were arrested in mitosis with 5 μM MG-132 for 2 hours before the addition of DMSO, Taxol, or UMK57 for 1 hour. The cell culture dish was then placed on ice for 29 minutes. Cells were permeabilized for 1 minute in PermFix before cold methanol fixation for 10 minutes at −20°C.

Fixed cells were washed 2 × 5 minutes with TBS-BSA (10 mM Tris, 150 mM NaCl, 10% BSA, and 0.1% NaN3) + 0.5% Triton X-100. Tubulin and ACA were visualized with α-tubulin DM1α (Sigma Aldrich; 1:4000) and ACA (anti-centromere antibody; Geisel School of Medicine; 1:2000) primary antibodies diluted in TBS-BSA + 0.1% Triton X-100. After 1 hour incubation, cells were washed 3 × 10 minutes with TBS-BSA + 0.1% Triton X-100. Cells were incubated for 1 hour with Alexa Fluor® 488 (Molecular Probes; 1:2000), Alexa Fluor® 594 (Molecular Probes; 1:2000), and DAPI (Molecular Probes, 400ng/mL) in TBS-BSA + 0.1% Triton X-100. Cells were washed 2 × 10 minutes in TBS-BSA + 0.1% Triton X-100 followed by a 10 minute wash with TBS-BSA. Coverslips were then mounted on slides with Pro-Long Gold Anti-Fade (Life Technologies).

B. Imaging Acquisition and Data Analysis

Images of prometaphase and metaphase spindles were acquired with a cooled charge-coupled device camera (Andor Technology) mounted on a Nikon Eclipse Ti microscope with a Plan Apo VC 60x, 1.4 NA, oil immersion objective (Nikon). Image series in the z-axis were obtained using 0.3 μm optical sections (23 slices). For both the nocodazole shock and cold stable assays, raw immunofluorescence images of spindles were classified as either “spindle-like”, “high polymer”, or “low polymer” (Gayek & Ohi, 2014). Image deconvolution and contrast enhancement for representative images was performed using AutoQuant X3 (Media Cybernetics) and Fiji (Schindelin et al., 2012).

C. Merits and Limitations

The amount of tubulin polymer was assessed in prometaphase and metaphase cells for both the nocodazole shock (Figures 3A and 3B) and cold shock (Figures 3C and 3D) assays. As observed in Figures 3A and 3C, the three-tier classification system for spindle structures is somewhat subjective. Therefore, it is critical for the researcher to adhere to strict classification criteria for each treatment. Additionally, it is important to keep incubation times constant across treatments, since the degree of polymer remaining is directly related to the amount of time cells are exposed to either nocodazole or cold temperature.

The nocodazole shock assay generated discernable differences between prometaphase and metaphase control cells, with more spindle-like structures in metaphase, consistent with increased microtubule stability (Figure 3B). Furthermore, Taxol treatment significantly increased the number of spindle-like cells observed in both prometaphase and metaphase, indicating a stabilization of k-MT attachments (Figure 3B). UMK57-treated cells also displayed increased microtubule polymer in prometaphase; however, the metaphase destabilization effect of UMK57 was not detected in this assay (Figure 3B).

The cold shock assay did not generate discernable differences between prometaphase and metaphase control cells (Figure 3D). Otherwise, the results of this assay were similar to those observed in the nocodazole shock assay (Figures 3C and 3D). The number of spindle-like cells was significantly increased in prometaphase and metaphase due to Taxol treatment, while cells treated with UMK57 were indistinguishable from DMSO-treated control cells.

The nocodazole shock and cold shock assays are easy to perform and the tubulin polymer classification system allows the researcher to assess large numbers of cells with ease. Both assays can be used to demonstrate significant differences in k-MT stability between cell lines (Gayek & Ohi, 2014). Care must be taken when using these semi-quantitative approaches because of the subjectivity in assigning cells to one of three categories. Also, the sensitivity of these assays is somewhat low. Cold shock was unable to detect the subtle increase in k-MT stability from prometaphase to metaphase and both the nocodazole and cold shock assays were unable to detect k-MT destabilization induced by UMK57 treatment.

IV. Calcium Treatment Assay

Calcium treatment can inhibit tubulin assembly and promote microtubule depolymerization (O’Brien, Salmon, & Erickson, 1997; Weisenberg & Deery, 1981). Similar to the cold and nocodazole shock assays, the calcium treatment assay takes advantage of the differential sensitivities of non-k-MTs and k-MTs to calcium-induced microtubule depolymerization. By quantifying whole spindle tubulin intensity after calcium treatment, the relative level of k-MT stability can be determined. The permeabilization and fixation protocols used here were performed as previously described (Thompson & Compton, 2011).

A. Methodology

Cells were treated for 2 hours with 5 μM MG-132 before the addition of DMSO, Taxol, or UMK57 for 1 hour. Cells were extracted in calcium buffer (100 mM pipes, 1 mM MgCl2, 1 mM CaCl2, and 0.5% Triton X-100 at pH 6.8) at room temperature for 5 minutes. Cells were then fixed in 1% glutaraldehyde in PBS for 10 minutes followed by 2 × 10 minute washes in 0.5 mg/mL NaBH4. Cells were stained for Tubulin (DM1α), and DNA, using the same immunofluorescence protocol described in section IIIA.

B. Image Acquisition and Data Analysis

Using a Nikon Eclipse Ti microscope with a Plan Apo VC 60x, 1.4 NA, oil immersion objective, images of bipolar spindles were acquired, where both spindle poles were located in the same focal plane. Image series in the z-axis (0.2 μm optical sections, 41 slices) were obtained using a cooled charge-coupled device camera (Andor Technology). Mean tubulin intensity of the whole spindle (minus background) was quantified from sum intensity projections of fluorescence z-stacks in Fiji. Mean background intensity was calculated for the region extending beyond the spindle, as previously described (Hoffman, Pearson, Yen, Howell, & Salmon, 2001).

C. Merits and Limitations

For the calcium treatment assay, tubulin intensity within the mitotic spindle was quantified after Taxol or UMK57 treatment (Figure 4). There was no discernable difference in mean tubulin intensity between prometaphase and metaphase cells. However, mean tubulin intensity was significantly increased in Taxol-treated prometaphase and metaphase cells (Figures 4A and 4C), consistent with increased microtubule stability. In contrast, there was no detectable difference in tubulin intensity after UMK57 treatment for either mitotic stage (Figures 4A and 4C).

Figure 4. Calcium Treatment.

Figure 4.

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A) Quantification of spindle intensity for prometaphase U2OS cells treated for 2 hours with 5 μM MG-132 before the addition of DMSO, Taxol, or UMK57 for 1 hour. Cells were permeabilized in calcium buffer for 5 minutes and fixed in glutaraldehyde, as described in section IVA. Whole spindle measurements were made on sum intensity projections of fluorescence z-stacks. Data represent the mean + SEM; n=10 cells per condition; ***p ≤ 0.001; n.s., p ≥ 0.05 using two-tailed t test. B) Representative images (maximum intensity projections) of U2OS cells quantified in (A) fixed and stained for α-tubulin (green) and DNA (blue). Scale bar, 10 μm. C) Quantification of spindle intensity for metaphase U2OS cells treated for 2 hours with 5 μM MG-132 before the addition of DMSO, Taxol, or UMK57 for 1 hour. Calcium treatment assay was performed as described in (A). Whole spindle measurements were made on sum intensity projections of fluorescence z-stacks. Data represent the mean + SEM; n=10 cells per condition; ***p ≤ 0.001; n.s., p ≥ 0.05 using two-tailed t test. D) Representative images (maximum intensity projections) of U2OS cells quantified in (C) fixed and stained for α-tubulin (green) and DNA (blue). Scale bar, 10 μm.

Similar to the nocodazole shock and cold shock assays, the calcium treatment assay was only sensitive enough to detect Taxol-induced changes in microtubule dynamics. This suggests that the nocodazole shock, cold shock, and calcium treatment assays are most effective when all microtubules are ubiquitously affected by a given treatment or when there is a significant shift in the stable/unstable microtubule populations, as observed in Figure 2F. When using the calcium treatment assay to compare k-MT stability across cell lines, it is important to control for variable spindle size and variable amounts of total tubulin between cell types.

Whole spindle, sum projection measurements were more accurate than bulk measurement of k-MT attachment sites or whole spindle, maximum intensity projection measurements (data not shown). Since U2OS cells are near triploid, the increased number of chromosomes in this cell line prevented us from measuring fluorescence intensity at individual k-fibers. Measuring the fluorescence intensity of individual k-fibers following calcium treatment has been used previously to assess microtubule occupancy at kinetochores and to infer k-MT stability from those measurements (Etemad et al., 2018; Zaytsev, Sundin, DeLuca, Grishchuk, & DeLuca, 2014). However, it is important to consider that microtubule occupancy and k-MT detachment rate are independent parameters (King & Nicklas, 2000). For example, it may be possible for kinetochores to have more (higher occupancy), but less stable microtubule attachments, or fewer (lower occupancy), but more stable microtubule attachments. The latter scenario is consistent with data reported from cells lacking the kinetochore protein CENP-E (Maffini et al., 2009; McEwen et al., 2001; Putkey et al., 2002).

V. Inter-kinetochore Distance and Spindle Length Measurements

Bioriented sister kinetochores are under tension when they are stably attached to microtubules emanating from opposite spindle poles. Microtubule pulling forces cause an increase in the distance between sister kinetochores (Waters, Skibbens, & Salmon, 1996). Inter-kinetochore distance (IKD) is frequently used as a readout for k-MT stability, since IKD increases as cells progress through mitosis (Waters, Skibbens, & Salmon, 1996), correlating with increases in k-MT half-life (Kabeche & Compton, 2013). It was also recently demonstrated that decreased IKD correlates with decreased k-fiber intensity (Zaytsev et al., 2014).

Similarly, changes in spindle length can also be indicative of changes in k-MT stability. The metaphase spindle is maintained at a steady-state length (Goshima, Wollman, Stuurman, Scholey, & Vale, 2005) by the concerted action of a number of different molecules (Goshima & Scholey, 2010). Microtubule dynamics play an important role in the establishment and maintenance of spindle length and perturbation of molecules which promote microtubule polymerization (Goshima et al., 2005) or depolymerization (Goshima et al., 2005; T. J. Mitchison et al., 2005) has been shown to cause spindle lengthening or shortening, respectively.

A. Methodology

U2OS cells were treated for 1 hour with DMSO, Taxol, or UMK57. Cells were then fixed with 3.5% paraformaldehyde (pH 6.8) for 10 minutes at room temperature and stained for Hec1 (Novus Biologicals; 1:1000), Tubulin (YL1/2) (Novus Biologicals; 1:2000), and DNA, using the same immunofluorescence protocol described in section IIIA.

B. Image Acquisition and Data Analysis

Images of prometaphase and metaphase spindles were acquired with a cooled charge-coupled device camera (Andor Technology) mounted on a Nikon Eclipse Ti microscope with a Plan Apo VC 60x, 1.4 NA, oil immersion objective (Nikon). Image series in the z-axis were obtained using 0.2 μm optical sections (41 slices) for IKD images or 0.3 μm optical sections (29 slices) for spindle length images. IKD and spindle length measurements were performed on raw immunofluorescence images using the measurement tool in Nikon Elements software. IKD was measured between the centroids of sister kinetochore (Hec1) spots located in the same focal plane. Spindle length was measured as the distance between focused microtubule minus ends in a metaphase, bipolar spindle, where both poles were located in the same focal plane. Image deconvolution and contrast enhancement for representative images was performed using AutoQuant X3 and Fiji.

C. Merits and Limitations

IKD was increased in metaphase cells relative to prometaphase cells. In response to microtubule stabilization through Taxol treatment, IKD was decreased by ~0.1 μm in prometaphase and metaphase cells (Figures 5A and 5B). Metaphase IKD also decreased in response to k-MT destabilization through treatment with UMK57, as previously reported (Figures 5A and 5C; Orr et al., 2016). These results reveal a paradox for using IKD to assess k-MT stability, since IKD decreased in response to treatments that stabilized or destabilized k-MT attachments. From IKD measurements alone, it is difficult to draw conclusions about how k-MT dynamics are affected by a given treatment. An additional limitation of this assay is that the upper and lower limits of inter-kinetochore stretch are determined by sister-chromatid cohesion and centromere geometry (Kitajima, Hauf, Ohsugi, Yamamoto, & Watanabe, 2005; Lončarek et al., 2007). Defects in centromere integrity can also result in altered inter-kinetochore distances which are not reflective of changes in k-MT stability (Merriman et al., 2013; Sapkota, Wasiak, Daum, & Gorbsky, 2018).

Figure 5. Inter-kinetochore Distance and Spindle Length.

Figure 5.

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A) Representative immunofluorescence images of prometaphase (left) and metaphase (right) U2OS cells treated for 1 hour with DMSO, Taxol, or UMK57. Images represent single slices of deconvolved fluorescence z-stacks. Cells were fixed and stained for DNA (blue) and Hec1 (red) to measure inter-kinetochore distance (IKD). Scale bar, 10 μm. B) Quantification of IKD as measured by distance between Hec1 staining on sister kinetochores in prometaphase (gray) and metaphase (black) cells treated with DMSO or Taxol for 1 hour. Black bars indicate the mean ± SEM; n=100 kinetochore pairs per condition; ***p ≤ 0.001 using two-tailed t test. C) Data reproduced from Cell Reports, 17(7), Orr, B., Talje, L., Liu, Z., Kwok, B. H., & Compton, D. A., Adaptive Resistance to an Inhibitor of Chromosomal Instability in Human Cancer Cells, 1755–1763, 2016, with permission from Elsevier. Quantification of IKD as measured by distance between Hec1 staining on unaligned (gray) or aligned (black) sister kinetochores in cells treated with DMSO or Taxol. Black bars indicate the mean ± SEM; n ≥ 150 kinetochore pairs per condition; *p ≤ 0.05; ***p ≤ 0.001 using two-tailed t test. D) Representative maximum intensity projections (deconvolved) of U2OS cells treated for 1 hour with DMSO, Taxol, or UMK57. Cells were fixed and stained for DNA (blue) and Tubulin (YL1/2) (green) to measure spindle length. Scale bar, 10 μm. E) Quantification of metaphase spindle length in DMSO, Taxol, or UMK57-treated cells. Data represent the mean + SEM; n ≥ 42 cells per condition; **p ≤ 0.01; n.s., p ≥ 0.05 using two-tailed t test.

It has been previously demonstrated that MCAK activity is responsible for limiting mitotic spindle length (T. J. Mitchison et al., 2005). In support of this model, UMK57 treatment caused a small, but significant, decrease in metaphase spindle length (Figure 5D). Taxol treatment had no effect on spindle length (Figure 5D). Under these conditions, microtubule stabilization does not lead to increased spindle length, since nanomolar concentrations of Taxol decrease microtubule turnover (Figure 2E), but do not enhance microtubule polymerization (Yvon et al., 1999). When using this assay as a readout for k-MT stability, it is important to note that, in addition to microtubule dynamics, a number of different mechanical forces are responsible for regulating metaphase spindle length (Dumont & Mitchison, 2009; Goshima & Scholey, 2010).

VI. Kinetochore Oscillatory Movement

Congressing chromosomes undergo oscillatory movements (Skibbens, Skeen, & Salmon, 1993), driven in part by dynamic switches between polymerizing and depolymerizing k-MTs (Inoué & Salmon, 1995). Microtubule polymerization is correlated with anti-poleward kinetochore movement, while microtubule depolymerization is correlated with poleward kinetochore movement. Fluorescently labeled kinetochore pairs can be tracked in live cells to measure velocity, switch rate, and oscillation amplitude (Bissonette & Stumpff, 2014). By examining these parameters, perturbations of k-MT stability can be detected.

A. Image Acquisition and Data Analysis

Kinetochore pair tracking was performed on U2OS cells stably expressing EGFP-CENP-A, generated by FuGENE 6 (Roche Diagnostics) plasmid transfection followed by 1 mg/mL G418 selection. Cells were treated with DMSO, Taxol, or UMK57 in FluoroBrite DMEM for 1 hour. Coverslips were then mounted in a rose chamber and maintained at 37°C during image acquisition. Live cell imaging was performed using a QuorumWaveFX-X1 spinning disk confocal system on a Nikon Eclipse Ti microscope, equipped with an ILE laser source (Andor Technology) and a Hamamatsu ImageEM camera. A Plan Apo VC 60x, 1.4 NA, oil immersion objective (Nikon) was used to image kinetochore oscillations. Mitotic cells were identified by DIC and fluorescence microscopy. Z-stacks with a 0.5 μm step size (13 slices) were acquired every 5 seconds.

Bi-oriented kinetochore pairs moving around the equator were selected for quantification. Kinetochore pairs were tracked manually in three dimensions using the Fiji plugin TrackMate (Tinevez et al., 2017). Poleward and anti-poleward velocity measurements were calculated for trajectories between directional switches [v = |(x2 − x1)/(t2 − t1)|], where x2 and x1 are the relative positions of a given kinetochore at time points t2 and t1. Deviation from average position (DAP) was calculated as the standard deviation of the distances between a kinetochore and its average position at every time point (Bissonette & Stumpff, 2014). Switch rate was calculated as the number of times a kinetochore changed direction during oscillatory movement divided by the amount of time the kinetochore was filmed. Kymographs were generated in Fiji using the KymographBuilder plugin.

B. Merits and Limitations

The oscillatory movements of kinetochore pairs were differentially altered by microtubule stabilization using Taxol and by k-MT destabilization using UMK57 (Figures 6A and 6B). The DAP increased in response to UMK57 treatment from 0.57 μm observed in control cells to 0.81 μm (Figure 6C). Conversely, the DAP decreased in response to Taxol treatment from 0.57 μm observed in control cells to 0.49 μm (Figure 6C). Consistent with fixed cells analyses (Figures 5B and 5C), IKD measured using live cell kinetochore tracking was decreased by treatment with either Taxol or UMK57 (Figure 6D). Poleward and anti-poleward velocities were not changed by Taxol treatment, but increased in cells treated with UMK57, indicating that elevated MCAK activity promotes k-MT depolymerization (Figure 6E). The switch rate of sister kinetochore pairs was unaffected by either treatment (Figure 6F).

Figure 6. Kinetochore Oscillatory Movement.

Figure 6.

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A) Distance versus time plots of representative kinetochore pairs from EGFP-CENP-A expressing U2OS cells treated with DMSO, Taxol, or UMK57. Relative position was measured as the distance from the metaphase plate (represented by the dotted line at y=0). B) Representative kymographs display CENP-A fluorescence over time. Vertical scale bar represents 1 minute; horizontal scale bar represents 1 μm. C) Deviation from average position (DAP) measurements of kinetochores from cells treated with DMSO, Taxol, or UMK57. Data represent mean + SEM; n=24 kinetochores per condition; *p ≤ 0.05, ***p ≤ 0.001 using two-tailed t test. D) IKD was measured as the distance between EGFP-CENP-A spots in U2OS cells treated with DMSO, Taxol, or UMK57. Average IKD was determined for individual kinetochore pairs oscillating for 500 seconds. Data represent the mean + SEM; n=12 kinetochore pairs per condition; *p ≤ 0.05; **p ≤ 0.01 using two-tailed t test. E) Poleward (left) and anti-poleward (right) velocity measurements of kinetochores from cells treated with DMSO, Taxol, or UMK57. Data represent mean + SEM; n ≥ 78 measurements, N=24 kinetochores per condition; **p ≤ 0.01 using two-tailed t test. F) Switch rate of kinetochores oscillating for 500 seconds. Data represent the mean + SEM; n=12 kinetochore pairs per condition analyzed by two-tailed t test.

Under certain conditions (UMK57 treatment), microtubule depolymerization rate (i.e. kinetochore velocity) and microtubule detachment rate (i.e. k-MT half-life) are directly correlated (Figures 6E and 2G). However, when microtubule detachment rate is significantly reduced by Taxol treatment (Figure 2E), microtubule depolymerization rate is unaffected (Figure 6E). These data demonstrate the need for additional studies to further our understanding of the relationship between k-MT stability and the k-MT depolymerization dynamics associated with chromosome motion.

VII. Application of Methods to Evaluate the Effects of Low Dose Nocodazole Treatment

Previous studies have demonstrated that nanomolar concentrations of nocodazole induce mitotic arrest by suppressing microtubule dynamics (Jordan, Thrower, & Wilson, 1992; Vasquez, Howell, Yvon, Wadsworth, & Cassimeris, 1997). In interphase cells, low dose nocodazole increases the amount of time microtubules spend in a ‘paused’ state. However, low dose nocodazole was also shown to increase microtubule catastrophe frequency (Vasquez et al., 1997). We sought to determine how k-MT stability and non-k-MT stability are affected by nanomolar concentrations of nocodazole during mitosis in human cells.

A. Non-k-MTs Display Increased Sensitivity to Nocodazole Treatment

In these experiments, non-transformed hTERT RPE1 (ATCC®, CRL-4000) cells were treated with nanomolar concentrations of nocodazole and k-MT stability was assessed by photoactivation. We did not observe any significant difference in prometaphase or metaphase k-MT half-life after low dose nocodazole treatment (Figure 7A). However, non-k-MTs appeared to be more sensitive to this treatment, as non-k-MT half-life was significantly decreased in cells treated with 33 nM, 44 nM, or 66 nM nocodazole (Figure 7B). The percentage of fluorescence in the fast decaying population significantly decreased with increasing nocodazole concentrations (Figure 7C), suggesting that non-k-MTs are preferentially depolymerized.

Figure 7. Differential Sensitivities of k-MTs and non-k-MTs to Low Dose Nocodazole Treatment.

Figure 7.

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A) Average k-MT half-life for DMSO or nocodazole-treated RPE1 cells expressing photoactivatable GFP-α-tubulin. Error bars indicate SEM; n ≥ 10 cells per condition; n.s., p ≥ 0.05 using two-tailed t test. B) Average non-k-MT half-life for DMSO or nocodazole-treated RPE1 cells expressing photoactivatable GFP-α-tubulin. Error bars indicate SEM; n ≥ 10 cells per condition; ***p ≤ 0.001 using two-tailed t test. C) Percentage of photoactivated fluorescence intensity attributable to the fast decay process. Data represent the mean percentage for DMSO or nocodazole-treated cells. Error bars indicate SEM; n ≥ 10 cells per condition; ***p ≤ 0.001 using two-tailed t test. D) Quantification of spindle intensity for metaphase RPE1 cells treated with DMSO or low doses of nocodazole for 1 hour. Cells were fixed in 1% glutaraldehyde, as previously described (Thompson & Compton, 2011). Whole spindle measurements were made on sum intensity projections of fluorescence z-stacks. Data represent the mean ± SEM; n≥18 cells per condition; ***p ≤ 0.001 using two-tailed t test. E) Quantification of spindle intensity for metaphase RPE1 cells treated with DMSO or low doses of nocodazole for 1 hour. Cells were permeabilized in calcium buffer for 5 minutes and fixed in glutaraldehyde, as described in section IVA. Whole spindle measurements were made on sum intensity projections of fluorescence z-stacks. Data represent the mean ± SEM; n≥16 cells per condition; n.s., p ≥ 0.05 using two-tailed t test. F) Quantification of spindle length in RPE1 cells treated for 1 hour with DMSO or low doses of nocodazole. Data represent mean + SEM; n≥15 cells per condition; **p ≤ 0.01; ***p ≤ 0.001 using two-tailed t test. G) Quantification of IKD as measured by distance between Hec1 staining on sister kinetochores in metaphase RPE1 cells treated with DMSO or low doses of nocodazole for 1 hour. Black bars indicate the mean ± SEM; n≥130 kinetochore pairs and N≥16 cells per condition; *p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001 using two-tailed t test. H) Percentage of metaphase cells with at least one Mad2-positive kinetochore. RPE1 Venus-Mad2 cells treated for 2 hours with 5 μM MG-132 before the addition of DMSO or low doses of nocodazole for 1 hour. Analysis performed on MTSB-extracted cells fixed in 3.5% paraformaldehyde and stained with α-GFP (Abcam 1:1000) to detect Venus-Mad2 (n>130 cells per condition; ***p<0.001 using two-tailed Fisher’s Exact Test). I) Quantification of Mad2 fluorescence intensity relative to Hec1 intensity in metaphase RPE1 Venus-Mad2 cells. Quantification performed on MTSB-extracted cells fixed in 3.5% paraformaldehyde and stained with α-Hec1 and α-GFP to detect Venus-Mad2 (n≥43 kinetochores and N≥10 cells per condition; **p<0.05 using two-tailed t test). J) Live cell imaging of RPE1 Venus-Mad2 cells treated with either DMSO or 33 nM nocodazole for 1 hour prior to image acquisition. Nocodazole treatment leads to increased mitotic duration (left) and Mad2 persistence (right). Mitotic duration measured as the time from nuclear envelope breakdown to anaphase onset. Duration of Mad2 signal was measured as the time from NEB to Mad2 disappearance. Each spot represents a single cell; black bars indicate the mean ± SEM; n ≥ 7 cells per condition; ***p ≤ 0.001 using two-tailed t test.

To test this possibility, we measured tubulin intensity within the mitotic spindle after glutaraldehyde fixation (Figure 7D). Mean tubulin intensity was significantly decreased by nanomolar concentrations of nocodazole. Interestingly, the amount of calcium-stable tubulin polymer was unchanged (Figure 7E), consistent with the differential sensitivity of non-k-MTs and k-MTs to low dose nocodazole treatment. Metaphase spindle length was also decreased by low dose nocodazole treatment (Figure 7F). As previously discussed in section VC, multiple factors are known to influence spindle length; however, spindle length may be decreased in nocodazole-treated cells due to depolymerization and loss of interpolar non-k-MTs.

We also observed that nanomolar concentrations of nocodazole caused a decrease in IKD (Figure 7G). The loss of tension between sister kinetochores may be due to a loss of bridging fibers (Milas & Tolić, 2016; Vukušić et al., 2017) or decreased microtubule occupancy at the kinetochore, similar to what was reported for Hec1 mutants (Etemad et al., 2018; Zaytsev et al., 2014). Measurement of fluorescence intensity at individual k-fibers or electron microscopy would be required to test this hypothesis. Reduced microtubule occupancy at specific kinetochores may also explain the observed increase in the number of Mad2-positive kinetochores (Figure 7H) and the increase in the amount of kinetochore-localized Mad2 after nocodazole treatment (Figure 7I). It has been previously demonstrated that attachment defects activate the spindle assembly checkpoint (Etemad, Kuijt, & Kops, 2015; Etemad et al., 2018; Magidson et al., 2016; Tauchman, Boehm, & DeLuca, 2015) and the mitotic delay we observe suggests that the spindle assembly checkpoint is activated in these nocodazole treated cells (Figure 7J).

VIII. Concluding Remarks

In summary, the protocols outlined in this chapter are several of the most commonly used quantitative and semi-quantitative methods for determining k-MT stability. Our quantitative evaluation of the effects of low dose nocodazole treatment provides a proof of principle for how these assays can be combined to determine how the alteration of microtubule dynamics influences mitotic spindle function.

Although we provide a comprehensive analysis of most methods, our study was not exhaustive. Quantification of k-fiber fluorescence intensity (Etemad et al., 2018; Zaytsev et al., 2014) and NDC80-microtubule binding (Yoo et al., 2018) have been successfully used to assess microtubule occupancy at the kinetochore as a readout for k-MT attachment stability. Advanced imaging techniques such as electron microscopy (Cimini et al., 2001; King & Nicklas, 2000; Nixon et al., 2017) have also been used to determine microtubule occupancy. However, as previously discussed in section IVC, although microtubule occupancy and k-MT stability are related, they are separable parameters of k-MT attachment (King & Nicklas, 2000; Maffini et al., 2009).

The methods reviewed here can be applied to many different cell types and can be used in a multitude of research applications. These methods vary in their ease of implementation, their ability to detect changes in prometaphase and metaphase k-MT stability, and their sensitivity to detect subtle changes in k-MT attachment stability induced by treatment with Taxol and UMK57. The photoactivation technique is most challenging to implement, but is recommended whenever possible, since this assay is highly sensitive and appears to provide the richest data (i.e. quantitative detachment rates and stable vs. unstable population percentages). If this assay is not feasible due to technical limitations, the best alternative would be to use a combination of specific assays to reveal how a given treatment may be affecting microtubule dynamics.

Acknowledgements

We wish to thank current and past members of the Compton lab as well as other members of the mitosis field, whose extensive efforts contributed to the development and refinement of these methods. RPE1 Venus-Mad2 cells were a gift from Johnathan Pines and the photoactivatable GFP-α-tubulin plasmid was a gift from Alexey Khodjakov. This work has been supported by National Institutes of Health grant R37GM051542 to D.A. Compton.

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