Jcb In India Case Study Ppt

Long-range organization of chromosome XII between anchoring elements (TEL, CEN, and NUC)

Global architecture of chromosome XII was first investigated in living yeast cells by monitoring the position of 15 loci distributed along the chromosome. Using targeted homologous recombination of fluorescent operator-repressor system (FROS), 12 independent insertions distributed every ∼100 kb were generated along the non-rDNA region of chromosome XII (Fig. 1 A). We also labeled three individual loci within the rDNA region at known distances from the CEN (Fig. 1; see Materials and methods).

Figure 1.

Color-coded statistical mapping of positions of 15 loci along chromosome XII. (A) Schematic representation of chromosome XII with the 15 FROS-labeled loci. Note that the rDNA of 1.8 Mb is depicted as a 1-Mb segment. Loci studied by particle tracking are marked with asterisks. (B) Spatial distributions of each locus are represented using gene maps, color-coded heat maps determined by the percentile of the distribution (Berger et al., 2008; Therizols et al., 2010). (top) The dashed yellow circle, the red circle, and the small red dot depict the median NE, the median NUC, and the median location of the nucleolar center, respectively. (bottom) The gene maps are represented with the genomic position indicated on top (N represents the number of nuclei analyzed). Note that the position along the genome of the three rDNA loci has been determined by I-SceI cleavage. The data shown are from a single representative experiment out of three repeats.

We chose to represent the position of a locus as a probability distribution relative to nuclear and nucleolar centers in living cells (Berger et al., 2008; Therizols et al., 2010). The spatial repartition of loci positions was assayed in a large number of cells in interphase (>1,500), and the data were represented in a color-coded statistical map of loci positions in which the percentage in an enclosing contour represents the probability of finding a locus inside (Fig. 1 B). The radius of the nuclei and their morphology, characterized by the distance between the nuclear and the nucleolar centers, were similar in all strains (see Fig. S1). We also selected FROS strains with comparable nuclear (3.4 ± 0.09 µm3) and nucleolar volumes (1.37 ± 0.08 µm3) so as to overlay the positions of loci in the same map (see Materials and methods and Video 1).

The visual inspection of statistical maps shows that the position of loci along chromosome XII is consistent with a path dictated by the genomic coordinates of each locus and the segmentation of the chromosomes between three structuring elements CEN, TEL, and NUC (Fig. 1 B and Video 1). We then wished to assess whether chromosome XII folding could be predicted by nuclear models based on polymer physics (Wong et al., 2012). The median distance to the nuclear and nucleolar centers was plotted as a function of the genomic coordinates of each locus (box plots in Fig. 2, A and B; see Materials and methods), and these data compared well with the model predictions (Wong et al., 2012; Fig. 2, A and B, black line). We noted that the fit to the model prediction was poorer for genomic positions 450–1,050 kb, i.e., from NUC to the right TEL (see, e.g., the anomalous dynamics of the rDNA in Fig. 4).

Figure 2.

The position of loci along chromosome XII relative to the NUC and the NE are predicted by computational modeling. (A and B) The distance of the locus to the nuclear center (A) and to the nucleolar center (B) is plotted versus its genomic position. Yellow and red ellipsoids depict the NE and NUC, respectively, and the black line represents the distance (left). The median distance is shown with box plots for the 15 loci described in Fig. 1 and for four RNA Pol III genes (white and green datasets). The median distance of chromosome XII loci to the nuclear center and to the centroid of the rDNA segment from a computational model of chromosome XII (Wong et al., 2012) is shown with solid black lines. (C) The gene map of loci t(P(UGG)L, tA(UGC)L, SNR6, and t(L(UAA)L relative to interpolated positions −58, 65, 216, and 647 kb (which correspond to their genomic coordinates) reveals global agreement between the model and measurements (see Video 2). The experimentally determined gene map (top) is compared with the interpolated position (bottom). The dashed yellow circle, the curved broken red line, and the small red dot depict the median NE, the median NUC, and the median location of the nucleolar center, respectively. (D) The yeast U6 small nuclear RNA (snRNA) gene SNR6 is unaffected by FROS insertion. Locations of important elements are indicated relative to the transcriptional start site (bent arrow) as +1 (top). Indirect end-labeling analysis of the chromatin structure on SNR6 is shown on the bottom-right panel. Ellipses on the right mark the positioned nucleosomes in the corresponding lanes with wild-type unmodified (U) or FROS tagged (T) strains. Note that the apparent hyper-sensitive site (asterisk) detected upon FROS insertion corresponds to a HindIII site introduced along with tetO repeat. SNR6 expression is not affected by FROS insertion. Transcript levels in untagged and FROS-tagged strains were normalized against the U4 transcript used as an internal control. The mean and scatter of RNA levels estimated from three independent experiments are plotted (error bars; bottom left).

The smooth variations of the median positions between consecutive loci suggested that the statistical maps could be interpolated with a genomic resolution much finer than 100 kb. We thus computed statistical maps every kilobase along chromosome XII (Video 2; see Materials and methods), and we challenged their relevance by comparing them to the positions of four RNA Pol III–transcribed genes along chromosome XII (tP(UGG), tA(UGC), tL(UAA), and SNR6); their genomic positions correspond to the green box plots of Fig. 2 (A and B). Experimental and interpolated statistical maps are represented in the top and bottom half of Fig. 2 C, respectively. If SNR6 shows a good agreement with interpolation, mild or strong discrepancies are observed, respectively, for tA(UGC), tL(UAA), or tP(UGG). Such data argue for a local effect of Pol III genes on chromosome structure, and differ from colocalization assays, in which Pol III–transcribed genes were organized in clusters close to CEN or NUC (Thompson et al., 2003; Duan et al., 2010). We suggest that the distribution of genes is primarily influenced by their genomic position, and only locally by sequence-specific physical interactions (Fig. 2, A and B). Note that we checked that the FROS insertion did not affect chromatin structure by analyzing the inner structure of SNR6 using in situ indirect-end labeling (IEL; Fig. 2 D). Our results show that nucleosome positioning within the upstream and transcribed regions is similar in the control and the labeled strain (Fig. 2 D). Moreover, the transcription level of SNR6 was not significantly modified by FROS labeling (Fig. 2 D). In conclusion, experimental data and predictions from the computational model are largely compatible, which suggests that the folding of chromosome XII is mainly dictated by local attachments to the three main nuclear structures NUC, TEL, and CEN.

Major transcriptional reprogramming has little effect on chromosome XII structure

To investigate further whether molecular interactions driven by transcription played a role in global chromosome architecture, we probed the structure of chromosome XII by statistical mapping after a major transcriptional repression (Fig. 3). Cells were treated with rapamycin, an inhibitor of the conserved protein kinase complex TORC1 (target of rapamycin; Loewith et al., 2002), which induces a global modification of transcription, mimicking nutrient deprivation characterized by transcriptional repression of Pol II ribosomal protein genes, rRNA 5S genes transcribed by Pol III, and a repression of Pol I activity (Hardwick et al., 1999; Wullschleger et al., 2006). After 20 min of rapamycin treatment, the mean nucleolar volume decreased by 37%, but the nucleus remained nearly constant in size (4% increase in volume), in agreement with previous studies (Therizols et al., 2010). The statistical maps of 12 loci positions were determined in nontreated versus rapamycin-treated cells, and are represented, respectively, in the top half and bottom half of each gene map (Fig. 3). Nucleolar size reduction generated a global shift of all loci toward the NUC. However, we did not detect major modifications of internal chromosome folding at the population level, thus supporting our model in which chromosome architecture is mainly determined by the biophysical properties of chromosomes and volume exclusion rather than by molecular interactions mediated by transcription.

Figure 3.

Color-coded statistical mapping of positions of 12 loci along chromosome XII after rapamycin treatment. (A) Schematic representation of chromosome XII with the 12 FROS-labeled loci using the centromere as an origin. (B) The spatial distributions of each locus (bottom of the gene map representations) are compared with loci position in rapamycin-treated cells (top of the gene map representations). The dashed yellow circle, the curved broken red line, and the small red dot depict the median NE, the median NUC, and the median location of the nucleolar center, respectively.

Chromatin motion of chromosome XII

We investigated the dynamics of chromosome XII and compared these data to predictions of polymer models. The motion of 10 loci (Fig. 1 A, asterisks) was studied by recording their trajectories in 2D rather than in 3D to increase the acquisition speed and reduce phototoxicity. Normal cell growth was observed during up to 2 h of acquisition (Video 3). A broad temporal range (from 0.2 to 400 s) was probed using five distinct inter-frame intervals of 190 ms, 360 ms for all loci, and 1, 1.5, and 10 s for three loci. Dedicated software was developed to perform systematic trajectory analysis (Fig. S2, A–C; See Materials and methods). This software allowed us to extract the mean square displacement (MSD), which is the mean of the squared travel distances after a given time lag, for every locus, and to compare the dynamics in the case of central, peripheral, or nucleolar-localized loci (Fig. S2). Trajectories during which deformations of the nucleus or detection of drifts of the nuclear center occurred (e.g. during mitosis, see Video 3) were disregarded.

Several studies suggested that chromatin motion was determined by a normal diffusive behavior in a restrained volume (MSD∼t in the small time limit, and MSD∼a in the long time limit, with a being the radius of constraint; Marshall et al., 1997; Heun et al., 2001; Meister et al., 2010). To display MSD in the short and long time regimes, we used linear and logarithmic representations (Fig. 4 and Fig. S3). Loci −30, 680, and rDNA were chosen as representative examples (Fig. 4). We observed that MSD curves exhibited a power-law scaling response expressed as MSD∼Γ × tα, with an anomalous parameter α of ∼0.5 ± 0.07 (Fig. 4 B) and an amplitude of the motion Γ of ∼0.01 µm2 × s−0.5. Interestingly, the anomalous diffusive response is consistent with the Rouse polymer model that describes the movements of polymer segments based on their elastic interactions and on viscous frictions (De Gennes, 1979). This model is expected to apply to dense polymer solutions (De Gennes, 1979), and it was recently observed that the bacterial genome behaves as predicted from this model (Weber et al., 2010a,b). It also applies to the molecular dynamics simulations of the yeast genome (Rosa and Everaers, 2008), thus strengthening our view that chromosome properties in yeast fit with polymer models.

Figure 4.

Loci motion is slowed down at NUC, and homogeneous elsewhere. (A) The temporal evolution of the MSD is plotted at position 680 kb, −30 kb (for central localization), and at rDNA (nucleolar localization) in log-log (top) or linear (bottom) scale. MSD extracted from time lapse of 0.19-, 0.36-, 1-, 1.5-, and 10-s inter-frames are depicted in different colors. The curves are fitted with an anomalous diffusion model (solid lines), showing that the anomaly parameter is 0.5 ± 0.07. Note that the movements of rDNA are slow in comparison to those of the locus at 680 or −30 kb. The black line represents the fit to the dataset with a power-law scaling of 0.55. (B) The plot represents the anomaly parameter versus the genomic position, and shows the different dynamics in NUC for the 10 analyzed loci. (C) Spatial fluctuations of the 10 analyzed loci are compared by measuring the amplitude of the power-law scaling response using a model with Γt0.5. (D) The gene territory, defined by the volume occupied by 50% of the gene population (the green isocontour in gene maps) expressed in cubic micrometers is measured as a function of the genomic position. Gene territory experimental errors were determined by measuring gene territories from three samplings of the full dataset.

We further developed the description of chromatin dynamics at a short time interval along chromosome XII. We compared the amplitude of the MSD traces by plotting Γ for every locus (Fig. 4 C and Fig. S3), showing that the chromatin movements are homogeneous for most of the loci with a value in the range ∼0.010–0.015 µm2 × s−0.5. These results suggest a homogeneous behavior of chromatin throughout chromosome XII, except for rDNA. In addition, chromatin dynamics were not different for a locus with a central or peripheral localization (Fig. S3), and the amplitude of fluctuations Γ was moderately reduced by ∼20% for loci with nucleolar versus central localization (Fig. S3), which suggests that nucleolar proximity tends to restrain chromatin motion of non-rDNA loci. The investigation of MSD in the long time limit, best viewed in linear representation, revealed a deviation from the Rouse regimen MSDΓ × t0.5. As previously suggested (Meister et al., 2010), the apparent confinement in the MSD is related to loci diffusing in a restrained volume. Space visited after 200 s (∼0.15 µm2, ∼0.25 µm2, and ∼0.3 µm2 for the rDNA, locus −30 kb, and locus 680 kb, respectively), can be assigned to an explored volume of 0.57, 1.02, and 1.34 µm3 for the three loci (Meister et al., 2010). These values are in excellent agreement with the dimensions of gene territories, as defined by the volume in which 50% of the gene positions are detected in statistical maps (Fig. 4 D; rDNA, locus −30 kb, and locus 680 kb, respectively, at 0.8, 1.1, and 1.2 µm3). This analysis also shows that one locus can explore the entire statistical map within a few minutes.

Finally, the motility of three chromosomal loci in the rDNA was investigated (Fig. 4), showing an MSD with two distinct slopes: increasingly slowly for t < 5 s (α = ∼0.25) and more abruptly after 5 s (α = ∼0.7). These dynamics are not consistent with the Rouse model (Fig. 4 A, black lines). In these polymer models, several specific properties of the rDNA were disregarded, including among others the dynamics associated with transcriptional activity, the depletion of nucleosomes from actively transcribed rDNA (Conconi et al., 1989), or the possible local tethering of rDNA via CLIP proteins (Mekhail et al., 2008; Mekhail and Moazed, 2010). This problem should be considered in future molecular dynamics simulations.

To conclude, our systematic analysis of position and motion of loci along chromosome XII showed that its inner structure mainly consists of constrained chromosome arms anchored to nuclear elements (SPB, NUC, and NE), as expected from recent computational models (Tjong et al., 2012; Wong et al., 2012). We also demonstrated that the Rouse polymer model accurately describes the motion of chromatin loci except for the rDNA (Rosa and Everaers, 2008), the biophysical properties of which remain to be investigated.

We may finally speculate on whether the properties of chromosomes observed in yeast are relevant to metazoans. Our results suggest that chromosomes should be viewed as anchored polymers in a dense environment. Lamin associated domains (LADs) and nucleolar tethering domains (NADs), which appear to be distributed in large regions of the metazoan genome (Guelen et al., 2008; Németh et al., 2010; van Koningsbruggen et al., 2010), may provide anchoring regions analogous to NUC, CEN, and TEL. We believe that the relevance of the Rouse model between anchoring regions should be studied, and spatial references should be proposed to construct statistical maps to investigate chromosome folding properties.

Abstract

The kinetochore forms a dynamic interface with microtubules from the mitotic spindle. Live-cell light microscopy–based observations on the dynamic structural changes within the kinetochore suggest that molecular rearrangements within the kinetochore occur upon microtubule interaction. However, the source of these rearrangements is still unclear. In this paper, we analyze vertebrate kinetochore ultrastructure by immunoelectron microscopy (EM) in the presence or absence of tension from spindle microtubules. We found that the inner kinetochore region defined by CENP-A, CENP-C, CENP-R, and the C-terminal domain of CENP-T is deformed in the presence of tension, whereas the outer kinetochore region defined by Ndc80, Mis12, and CENP-E is not stretched even under tension. Importantly, based on EM, fluorescence microscopy, and in vitro analyses, we demonstrated that the N and C termini of CENP-T undergo a tension-dependent separation, suggesting that CENP-T elongation is at least partly responsible for changes in the shape of the inner kinetochore.

Introduction

Faithful chromosome segregation during mitosis is essential for the accurate transmission of genetic material. The kinetochore forms a dynamic interface with microtubules from the mitotic spindle to facilitate chromosome segregation (Cheeseman and Desai, 2008). Importantly, kinetochore–microtubule attachments are capable of generating as much as 700 pN of force (Nicklas, 1988), which functions to both move chromosomes and to generate signals that report on proper attachment status. As might be predicted based on this large amount of force generated at kinetochores, live-cell light microscopy–based observations have suggested that structural rearrangements within the kinetochore occur in the presence of tension (Maresca and Salmon, 2009; Uchida et al., 2009). These works also suggested that these rearrangements are essential for release from a spindle assembly checkpoint–dependent cell cycle arrest. However, the molecular source of these rearrangements is still unclear, as these changes were only evaluated as an increase in intrakinetochore distance (the distance between the inner and outer kinetochore) by light microscopy.

To examine the molecular rearrangements within the kinetochore in detail, it is essential to observe changes in kinetochore ultrastructure using EM. Early EM studies of vertebrate cells revealed that the kinetochore has a trilaminar morphology, including an inner and outer plate (Brinkley and Stubblefield, 1966; Jokelainen, 1967; Comings and Okada, 1971; Rieder, 1982; McEwen et al., 2007). Microtubules attach directly to the outer plate (Brinkley and Stubblefield, 1966; Jokelainen, 1967; Comings and Okada, 1971; McEwen et al., 1998), whereas chromatin-bound proteins, such as CENP-C, localize to the inner plate (Saitoh et al., 1992).

To analyze the structural dynamics of the kinetochore that occur in response to force-dependent changes during mitosis, we combined EM analysis with specific observations of representative kinetochore markers using immuno-EM. We have previously identified multiple kinetochore components and generated a toolkit of antibodies suitable for analyzing the kinetochore ultrastructure (Okada et al., 2006; Hori et al., 2008a; Amano et al., 2009; Ohta et al., 2010). Based on our immuno-EM analyses, we found that the inner kinetochore region is dramatically deformed in the presence of tension from spindle microtubules. In addition, we found that the N and C termini of CENP-T undergo a tension-dependent separation based on EM, fluorescence microscopy, and in vitro analyses, suggesting that CENP-T elongation is responsible for changes in the shape of the inner kinetochore. Thus, this work defines a key force-dependent molecular change within the kinetochore.

Results

High resolution mapping of proteins within the kinetochore by immuno-EM

To conduct a comprehensive analysis of the tension-dependent structural changes that occur within the kinetochore, we began by analyzing the kinetochore structure of mitotic chromosomes in chicken DT40 cells in the absence of microtubules (using treatment with the microtubule-depolymerizing drug nocodazole at 500 ng/ml for 3 h). After a conventional fixation with glutaraldehyde and staining with uranyl acetate and lead citrate, we imaged 170-nm-thick serial sections for individual mitotic cells. We observed clear electron-dense kinetochore outer plates in the region of the primary constriction of each chromosome (Fig. 1 A). To define the structure of each kinetochore, we measured the distance between the paired outer kinetochore plates visualized in Fig. 1 A. We also measured the length of the chromatin region at the primary constriction. The mean distance between the paired outer kinetochore plates was 754 nm, the mean length of the chromatin region was 637 nm, and the mean width of the outer plate was 41 nm (Fig. 1, B and C). To conduct an unbiased analysis of this EM data, we used digital images and obtained distribution curves of signal intensity. Using the signal intensity values, we measured the size of the outer plates (Figs. 1, B and C; and S1).

Figure 1.

Ultrastructure of the kinetochore in DT40 cells. (A) An image of a chromosome with paired outer plates from DT40 cells observed by EM. The arrowhead shows the outer plate. (B) Distribution of data for the width of the outer plate of 131 kinetochores. The method describing how the size of the outer plate was measured is described in Fig. S1. (C) Summary for the size of the kinetochore region in DT40 cells treated with nocodazole and fixed by glutaraldehyde. We chose chromosomes with the sister kinetochore plates and measured the length and width of the plates by the method described in Fig. S1. The mean of the length and width of the outer region is ∼227 nm (227 ± 70.5 nm) and ∼41 nm (41 ± 4.6 nm), respectively. (D) Immuno-EM images of nocodazole-treated DT40 cells using anti–CENP-E, anti-Mad2, anti-Ndc80, anti-Mis12, anti–CENP-C, anti–CENP-R, anti–CENP-T, and anti-HA (for detection of CENP-A–HA) antibodies. Signals are shown as gold labeling. Orange lines show the distribution of background signals. (E) Measurement of the sister kinetochore distance. Chromosomes with the sister kinetochore plates that were parallel to the plane of one section were chosen. Signal intensities of the red line were measured. The regions between two pink dashed lines show the regions with positive signals (higher than background signals). The yellow line shows the maximum of background signals. Signal intensities of the orange line on the chromosome arm were measured as background signals. The sister kinetochore region is defined by two peaks in the graph for the red line. The distance between two peaks is defined as the distance between the sister kinetochores. (F) Position of each protein in kinetochores. The half-distance between the sister kinetochore signals for each protein was plotted in the graph. CENP-A and CENP-T are located in the most internal region. Mis12, Ndc80, Mad2, and CENP-E are located near the outer plate. Yellow boxes indicate the regions of higher magnification on the right. Bars, 250 nm.

Using these conditions in DT40 cells, it was not possible to visualize clear trilaminar plates, necessitating the use of molecular markers to analyze kinetochore structure. Therefore, we next probed nocodazole-arrested cells with primary antibodies against CENP-C, CENP-E, CENP-R, CENP-T, Mad2, Mis12, or Ndc80 (Fukagawa et al., 1999; Hori et al., 2003, 2008a,b; Kline et al., 2006) and gold-labeled secondary antibodies. Specific information regarding these antibodies, including the antigens used to generate them, is summarized in Materials and methods. In each case, we succeeded in visualizing the labeled proteins with gold particles at DT40 kinetochores (Fig. 1 D). We also stained DT40 cells expressing CENP-A–HA with anti-HA antibodies to visualize CENP-A at kinetochores, as our existing CENP-A antibody was not functional for immuno-EM (Fig. 1 D). We next measured the distance between the two paired foci on sister kinetochores for the eight kinetochore proteins shown in Fig. 1 D. For these measurements, we used a line scan for digital EM images and obtained a signal distribution for the gold particles with two peaks (Fig. 1 E). Then, we measured the distance between the two peaks. The position of each kinetochore protein relative to the center of the chromosome is defined as half of the sister kinetochore distance. These measurements are summarized in Figs. 1 F and S1 E. The mean position of the outer plate was ∼377 ± 50 nm from the center of chromosome, which corresponds to the half-distance of the outer surface of the outer plate. CENP-E, Mad2, Mis12, and Ndc80 were located outside or near the outer plate (Fig. 1 F), which is consistent with previous studies in human cells (Cooke et al., 1997; DeLuca et al., 2005). In contrast, CENP-A, CENP-C, CENP-R, and CENP-T were located internally to the outer plate (Fig. 1 F). CENP-C was previously mapped to the inner kinetochore region using immuno-EM analysis in human cells (Saitoh et al., 1992). We recently demonstrated that CENP-T binds centromere DNA directly (Hori et al., 2008a), and kinetochore single-molecule high resolution colocalization analysis in human cells also suggests that CENP-A, CENP-C, and CENP-T are close to centromeric chromatin (Wan et al., 2009). Thus, our EM analysis, combined with previous studies, demonstrates that CENP-A, CENP-C, CENP-R, and CENP-T are located at the inner kinetochore and that CENP-E, Mad2, Mis12, and Ndc80 are located at the outer kinetochore (Fig. 1 F).

The width of the inner kinetochore region, but not the outer kinetochore, is stretched in the presence of spindle microtubules

The aforementioned studies define a baseline state of kinetochore structure in DT40 cells in the absence of microtubules. To analyze the structural changes that occur in response to tension from microtubule attachments, we next performed EM analysis on cells treated with the proteasome inhibitor MG132, which inhibits the degradation of anaphase-promoting complex/cyclosome substrates, preventing anaphase onset. We confirmed that chromosomes aligned at the metaphase plate with kinetochore-bound microtubules in cells treated with MG132 (unpublished data). We then performed immuno-EM to visualize the localization of diverse inner and outer kinetochore proteins (Figs. 2 and 3). The distance between sister kinetochores (both inner and outer kinetochores) was increased in cells treated with MG132 relative to cells treated with nocodazole (a 180-nm increase for the CENP-T–CENP-T distance; a 160-nm increase for the Ndc80–Ndc80 distance), suggesting that MG132-treated cells generate tension from spindle microtubules. Strikingly, the rectangular-like shape of the inner kinetochore structure occupied by CENP-A, CENP-C, CENP-R, and CENP-T in cells treated with nocodazole was deformed into an ovallike shape in cells treated with MG132 (Fig. 2). Although our antibodies are polyclonal antibodies and it is hard to determine whether there are specific epitopes within each protein, we found that our existing CENP-T antibodies recognized the C-terminal region but not the N terminus (unpublished data). We measured the length and width of the kinetochore region occupied by gold particles. A detailed method for these measurements is described in Fig. S2. The distribution of the inner kinetochore region occupied by the C terminus of CENP-T was significantly altered in the presence of microtubules with a length of 164 ± 40 nm (n = 100) and a width of 86 ± 26 nm (n = 100) in cells treated with MG132 compared with a length of 228 ± 35 nm (n = 100) and a width of 61 ± 18 nm (n = 100) in cells treated with nocodazole (Fig. 2, B and C). This change is statistically significant based on a Student’s t test (Fig. 2). Immuno-EM against CENP-C, CENP-R, and CENP-A–HA revealed similar structural changes in the inner kinetochore in cells treated with MG132 (Fig. 2, D–L). In contrast, the width of the rectangular outer kinetochore region visualized with antibodies against Ndc80, Mis12, and CENP-E is not stretched in cells treated with MG132 or nocodazole (Fig. 3, A–I), whereas the length of the outer kinetochore structure was decreased in cells treated with MG132 (Fig. 2, B, E, and H). The measurements for the length and width of the inner and outer kinetochore in nocodazole- or MG132-treated cells are summarized in Fig. 3 J. In total, these data indicate that the width of the inner kinetochore region, but not the outer kinetochore, is stretched in the presence of spindle microtubules.

Figure 2.

The structural deformation of the inner kinetochore when cells are treated with MG132. (A) Immuno-EM images with anti–CENP-T antibodies in DT40 cells treated with nocodazole (Noc) or MG132. (B) Measurement of the length of the inner kinetochore containing CENP-T in cells treated with nocodazole (n = 100) or MG132 (n = 100). (C) Measurement of the width of the inner kinetochore containing CENP-T in cells treated with nocodazole (n = 100) or MG132 (n = 100). (D) Immuno-EM images with anti–CENP-C antibodies in DT40 cells treated with nocodazole or MG132. (E) Measurement of the length of the inner kinetochore containing CENP-C in cells treated with nocodazole (n = 100) or MG132 (n = 100). (F) Measurement of the width of the inner kinetochore containing CENP-C in cells treated with nocodazole (n = 100) or MG132 (n = 100). (G) Immuno-EM images with anti-HA antibodies in CENP-A–HA-expressing DT40 cells treated with nocodazole or MG132. (H) Measurement of the length of the inner kinetochore containing CENP-A–HA in cells treated with nocodazole (n = 70) or MG132 (n = 70). (I) Measurement of the width of the inner kinetochore containing CENP-A–HA in cells treated with nocodazole (n = 70) or MG132 (n = 70). (J) Immuno-EM images with anti–CENP-R antibodies in DT40 cells treated with nocodazole or MG132. (K) Measurement of the length of the inner kinetochore containing CENP-R in cells treated with nocodazole (n = 100) or MG132 (n = 100). (L) Measurement of the width of the inner kinetochore containing CENP-R in cells treated with nocodazole (n = 100) or MG132 (n = 100). Error bars show SDs, and the significance of these measurements was estimated by the Student’s t test (indicated by asterisks). Yellow boxes indicate the regions of higher magnification on the right. Bars, 250 nm.

Figure 3.

The distribution of outer kinetochore proteins in the absence or presence of spindle microtubules. (A) Immuno-EM images with anti-Ndc80 antibodies in DT40 cells treated with nocodazole (Noc) or MG132. (B) Measurement of the length of the outer kinetochore containing Ndc80 in DT40 cells treated with nocodazole (n = 104) or MG132 (n = 118). (C) Measurement of the width of the outer kinetochore containing Ndc80 in DT40 cells treated with nocodazole (n = 104) or MG132 (n = 118). (D) Immuno-EM images with anti-Mis12 antibodies in DT40 cells treated with nocodazole or MG132. (E) Measurement of the length of the outer kinetochore containing Mis12 in DT40 cells treated with nocodazole (n = 100) or MG132 (n = 100). (F) Measurement of the width of the outer kinetochore containing Mis12 in DT40 cells treated with nocodazole (n = 100) or MG132 (n = 100). (G) Immuno-EM images with anti–CENP-E antibodies in DT40 cells treated with nocodazole or MG132. (H) Measurement of the length of the outer kinetochore containing CENP-E in DT40 cells treated with nocodazole (n = 100) or MG132 (n = 100). (I) Measurement of the width of the outer kinetochore containing CENP-E in DT40 cells treated with nocodazole (n = 100) or MG132 (n = 100). The width is not increased and significantly decreased. (J) The structural change of the inner kinetochore in the presence of microtubules. The inner kinetochore was changed to the ovallike structure from the rectangular structure, and the width of the inner kinetochore (magenta) was significantly stretched. In contrast, the width of the outer kinetochore (blue) was not stretched even in presence of microtubules. Error bars show SDs, and the significance of the length measurements was estimated by the Student’s t test (indicated by asterisks), whereas it was not estimated by the Student’s t test for the width. Yellow boxes indicate the regions of higher magnification on the right. Bars, 250 nm.

The stretched distribution of the inner kinetochore depends on tension from spindle microtubules

Our aforementioned analysis demonstrates that inner kinetochore distribution is altered by the presence of spindle microtubules. Although this is likely caused by the force exerted by these microtubules on bioriented kinetochores, we next sought to define the activities responsible for this deformation using specific perturbations that affect kinetochore function. We first tested DT40 cells depleted for the outer kinetochore protein Ndc80. We confirmed that Ndc80 is undetectable by Western blot analysis after the addition of tetracycline to Ndc80 conditional knockout cells (Hori et al., 2003). In addition, Nuf2, another subunit of Ndc80, is degraded in Ndc80-deficient cells, suggesting that complex formation of the Ndc80 complex is compromised (Hori et al., 2003). We previously observed that Ndc80 depletion compromises proper attachment of microtubules to kinetochores and arrests cells at mitosis (Hori et al., 2003). Importantly, depletion of Ndc80 prevented the stretched distribution of the inner kinetochore even in cells blocked in mitosis with MG132 (Fig. 4, A–C). Thus, the lack of proper attachments prevents the deformation of the inner kinetochore.

Figure 4.

The proper binding of Ndc80 with the microtubule is essential for the structural deformation within the inner kinetochore. (A) Immuno-EM images with anti–CENP-T antibodies in Ndc80-deficient cells treated with nocodazole (Noc) or MG132. The structural deformation of the inner kinetochore was not observed in Ndc80-deficient cells treated with MG132. (B) Measurement of the length of the inner kinetochore containing CENP-T in Ndc80-deficient cells treated with nocodazole (n = 70) or MG132 (n = 70). (C) Measurement of the width of the inner kinetochore containing CENP-T in Ndc80-deficient cells treated with nocodazole (n = 70) or MG132 (n = 70). (D) Images of the outer plates in wild-type, Ndc80-deficient, and Ndc80 D4 mutant cells. The arrows show the outer plates. (E) The size of the outer plate (means ± SD) in wild-type and Ndc80 D4 mutant cells. (F) Immuno-EM images with anti–CENP-T antibodies in Ndc80 D4 mutant cells treated with nocodazole or MG132. (G) Measurement of the length of the inner kinetochore containing with CENP-T in Ndc80 D4 mutant cells treated with nocodazole (n = 70) or MG132 (n = 70) and untreated cells (n = 70). (H) Measurement of the width of the inner kinetochore containing CENP-T in Ndc80 D4 mutant cells treated with nocodazole (n = 70) or MG132 (n = 70) and untreated cells (n = 70). (I) Immunofluorescence observation using antibodies against CENP-T and α-tubulin in Ndc80 D4 mutant cells treated with MG132. Error bars show SDs, and the significant difference was not detected. Green and magenta rectangles indicate distribution in cells. Yellow boxes indicate the regions of higher magnification on the right. Bars: (A, D, and F) 250 nm; (I) 10 µm.

Although Ndc80 depletion prevents the formation of kinetochore–microtubule attachments, completely eliminating Ndc80 also alters the structure of the kinetochore outer plate (Fig. 4 D; DeLuca et al., 2005). To distinguish between these two functions, we used a conditional replacement mutant of Ndc80 in which four Aurora B phosphorylation sites were replaced with aspartic acid to mimic constitutive phosphorylation (D4 mutant; Welburn et al., 2010). The Ndc80 D4 mutant retains the structural contribution of the Ndc80 complex to the outer kinetochore but reduces interactions with microtubules (Cheeseman et al., 2006; DeLuca et al., 2006; Guimaraes et al., 2008; Miller et al., 2008; Welburn et al., 2010). We confirmed that the outer plate is formed properly in the Ndc80 D4 mutant cells by EM based on its morphology (Fig. 4, D and E). When Ndc80 D4 mutant cells were treated with MG132, we did not detect a deformation of the inner kinetochore compared with cells treated with nocodazole (Fig. 4, F–H). Our results, combined with the previous biochemical data, indicate that proper binding of Ndc80 to microtubules is essential for the structural change of the inner kinetochore. Although the outer plate is formed and microtubules appear to attach to the outer plate in the Ndc80 D4 mutant cells (Fig. 4 I), we found that the distance between sister kinetochores was only slightly increased in the Ndc80 D4 mutant cells treated with MG132 relative to cells treated with nocodazole (a 25-nm increase compared with a 180-nm increase in wild-type cells), suggesting that there is no tension or weak tension in Ndc80 D4 mutant cells.

We next observed the inner kinetochore distribution in conditional mutant cells for the inner kinetochore protein CENP-H. Although the length of the inner kinetochore region was reduced in CENP-H–deficient cells, the width of the rectangular inner kinetochore structure was not stretched in CENP-H–deficient cells treated with MG132 (Fig. 5, A–C), indicating that the structural deformation of the inner kinetochore from the rectangular to the ovallike shape cannot occur in CENP-H–deficient cells even in the presence of microtubules. In CENP-H–deficient cells, the localization of a subset of outer kinetochore proteins is altered (Hori et al., 2003), and the plate structure of outer kinetochores is disorganized (Liu et al., 2006; Hori et al., 2008a), suggesting that proper attachment of microtubules to kinetochores is perturbed similar to Ndc80-deficient cells.

Figure 5.

The distribution of inner kinetochore proteins in various knockout cell lines. (A) Immuno-EM images with anti–CENP-T antibodies in CENP-H–deficient cells treated with nocodazole (Noc) or MG132. Although the size of the inner kinetochore in the CENP-H–deficient cells was reduced (length of 120 nm) compared with that of wild-type cells (length of 220 nm; see panel B), the width of the rectangular inner kinetochore structure was not changed in cells treated with MG132, indicating that the structural deformation of the inner kinetochore was not observed in CENP-H–deficient cells treated with MG132. (B) Measurement of the length of the inner kinetochore containing CENP-T in CENP-H–deficient cells treated with nocodazole (n = 70) or MG132 (n = 70). (C) Measurement of the width of the inner kinetochore containing CENP-T in CENP-H–deficient cells treated with nocodazole (n = 70) or MG132 (n = 70). (B and C) The significant difference was not detected. (D) Immuno-EM images with anti–CENP-T antibodies in CENP-C–deficient cells treated with nocodazole or MG132. The size of inner kinetochore of CENP-C–deficient cells was smaller (length of 160 nm). However, the structural deformation of the inner kinetochore was observed in CENP-C–deficient cells treated with MG132. (E) Measurement of the length of the inner kinetochore containing CENP-T in CENP-C–deficient cells treated with nocodazole (n = 70) or MG132 (n = 70). (F) Measurement of the width of the inner kinetochore containing CENP-T in CENP-C–deficient cells treated with nocodazole (n = 70) or MG132 (n = 70). (G) Immuno-EM images with anti–CENP-T antibodies in CENP-S–deficient cells treated with nocodazole or MG132. The structural deformation of the inner kinetochore was observed in CENP-S–deficient cells treated with MG132. (H) Measurement of the length of the inner kinetochore containing CENP-T in CENP-S–deficient cells treated with nocodazole (n = 113) or MG132 (n = 113). (I) Measurement of the width of the inner kinetochore containing CENP-T in CENP-S–deficient cells treated with nocodazole (n = 113) or MG132 (n = 113). (E, F, H, and I) The significance of these measurements was estimated by the Student’s t test (indicated by asterisks). (J) Summary of results for the structural deformation in the inner kinetochore in various cell lines. When tension from spindle microtubules was produced, the structural deformation in the inner kinetochore was observed. When we observed that the distance between the paired sister kinetochores increased >100 nm in cells treated with MG132 relative to cells treated with nocodazole, we judged that tension occurs (+). Therefore, although we observed 4- and 25-nm increases in Ndc80-deficient and Ndc80 D4 cells, respectively, we judged that these values meant that tension does not occur (−). Yellow boxes indicate the regions of higher magnification on the right. Error bars show SDs. Bars, 250 nm.

Although our analysis suggests that both proper kinetochore structure and the presence of microtubules are necessary for the structural deformation of the inner kinetochore, it is possible that the stretched distribution of the inner kinetochore reflects the physical movement of chromosomes but not the tension-derived force from spindle microtubules. To test this possibility, we used Rad21/Scc1-deficient DT40 cells in which cohesion is disrupted and tension, but not kinetochore–microtubule attachments, is eliminated (Sonoda et al., 2001). Even when Rad21-deficient cells are treated with nocodazole, sister chromatids separate, and the distance between sister kinetochores is increased relative to wild-type cells (Fig. 6, A and B). However, the width of the inner kinetochore is not stretched in Rad21-deficient cells (Fig. 6), suggesting that the physical attachment of kinetochores to microtubules and the movement of the chromosomes are not sufficient for the structural change of the inner kinetochore.

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