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Centrosome age regulates kinetochore–microtubule stability and biases chromosome mis-segregation

  1. Ivana Gasic
  2. Purnima Nerurkar
  3. Patrick Meraldi  Is a corresponding author
  1. University of Geneva, Switzerland
  2. Eidgenössische Technische Hochschule Zürich, Switzerland
Research Article
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Cite this article as: eLife 2015;4:e07909 doi: 10.7554/eLife.07909

Abstract

The poles of the mitotic spindle contain one old and one young centrosome. In asymmetric stem cell divisions, the age of centrosomes affects their behaviour and their probability to remain in the stem cell. In contrast, in symmetric divisions, old and young centrosomes are thought to behave equally. This hypothesis is, however, untested. In this study, we show in symmetrically dividing human cells that kinetochore–microtubules associated to old centrosomes are more stable than those associated to young centrosomes, and that this difference favours the accumulation of premature end-on attachments that delay the alignment of polar chromosomes at old centrosomes. This differential microtubule stability depends on cenexin, a protein enriched on old centrosomes. It persists throughout mitosis, biasing chromosome segregation in anaphase by causing daughter cells with old centrosomes to retain non-disjoint chromosomes 85% of the time. We conclude that centrosome age imposes via cenexin a functional asymmetry on all mitotic spindles.

https://doi.org/10.7554/eLife.07909.001

eLife digest

Cells are able to copy their DNA and then divide to make two daughter cells that each have a complete set of genetic material. In animal cells, the DNA is arranged within structures called chromosomes and groups of proteins called centrosomes control the process that separates the chromosome copies as the cell divides.

Each cell starts off with one centrosome, but before it divides, this centrosome is copied so that the cell now has two centrosomes at opposite ends of the cell, one old and one new. Filaments called microtubules assemble from the centrosomes and attach to the chromosomes. The microtubules first align all the chromosomes in the middle of the cell before pulling them towards the centrosomes as the cell divides.

Some cells divide such that the two daughter cells are destined to take on different roles, for example, a stem cell may divide to produce one stem cell and one skin cell. The end of the dividing cell that will become the stem cell contains the older centrosome, while the half that forms the skin cell will receive the younger centrosome. Other cells in the body may divide to form daughter cells that have the same fate, known as symmetrical division. In these cases, it is thought that there is no difference between the behaviour of the old and young centrosomes, but this idea has never been tested.

Here, Gasic et al. studied symmetrical division of human cells using fluorescent tags that made it possible to tell the centrosomes apart. The experiments show that the microtubules that assemble from the older centrosome bind the chromosome more tightly than those that form from the younger centrosome. This delays the alignment of the chromosomes that are connected to the old centrosome, as this process requires a flexible attachment. Moreover, in case the two chromosome copies fail to separate properly as cells divide, the older centrosome is more likely to receive both chromosome copies at the expense of the other centrosome. A protein called cenexin is present at higher levels around older centrosomes than around younger ones and is responsible for this effect.

Gasic et al.'s findings show that the age of the centrosomes leads to asymmetry in all cell divisions, even those that produce cells that are destined to have the same role in an organism. The next challenge will be to understand whether this asymmetry has any consequences for cells, in particular cancer cells.

https://doi.org/10.7554/eLife.07909.002

Introduction

The bipolar spindle has a symmetric appearance; nevertheless it contains two centrosomes of different ages, as every centrosome is duplicated once during the cell cycle, resulting in the presence of an old and young centrosome at mitotic onset (Nigg and Stearns, 2011). In asymmetric stem cell divisions centrosome age differentially affects their capacity to nucleate microtubules and their positioning with respect to the polarity and cell division axis (Yamashita et al., 2007; Wang et al., 2009; Januschke et al., 2013). Stem cells stereotypically inherit the centrosome, which nucleates more microtubules, which in most cases is the old centrosome (except in fly neuroblast divisions, where stem cells retain the young active centrosome; Januschke et al., 2011, Conduit and Raff, 2010). Old centrosomes also co-segregate with the ciliary membrane in stem cell divisions, allowing daughter stem cells to form a primary cilium earlier than the differentiating daughter cells (Paridaen et al., 2013). In symmetric divisions, the old and young centrosomes can be differentiated at the ultra-structural level, and in terms of their microtubule-anchoring capacity during interphase (Rieder and Borisy, 1982; Piel et al., 2000). The oldest centriole within the old centrosome contains distal and subdistal appendages: the first are necessary for centrioles to become basal bodies that can contact the plasma membrane (Graser et al., 2007; Hoyer-Fender, 2010), while the latter are involved in the organization of the interphase microtubule network, due to the presence of ninein, a key microtubule-anchoring protein (Mogensen et al., 2000). Both structures require the presence of cenexin, the oldest known marker for old centrosomes and appendages (Lange and Gull, 1995; Ishikawa et al., 2005). Importantly, all these structural differences disappear progressively as cells enter mitosis; therefore, it is assumed that the old and the young centrosomes behave indistinguishably in symmetrically dividing cells, resulting in a symmetric bipolar spindle. This hypothesis has, however, never been directly tested at the functional level. Here, we tested whether centrosome age affects cell division in symmetrically dividing human cells, focusing on the ability of centrosomes to organize the alignment and segregation of sister-chromatids into two daughter cells.

Results

The first key task of the mitotic spindle is to bind to chromosomes via kinetochores and align them onto the metaphase plate (Kops et al., 2010). To distinguish between old and young centrosomes, we used untransformed hTert-RPE1 and transformed HeLa cell lines expressing eGFP-centrin1, a centriolar protein whose abundance correlates with centriole age, or an anti-cenexin antibody, a marker for old centrosomes (Figure 1A,B, Kuo et al., 2011; Lange and Gull, 1995). In the vast majority of the cases both markers were enriched at the same centriole pair, indicating a robust recognition of the old centrosomes (data not shown). To investigate whether half-spindles associated with old or new centrosomes align chromosomes with the same efficiency, we analyzed late prometaphase cells that contained few unaligned chromosomes. We found that 61.23% of the unaligned chromosomes were in the vicinity of old centrosomes in Hela-eGFP-centrin1 cells as opposed to 50% expected for an unbiased distribution, suggesting a difference in the efficiency of chromosome alignment (Figure 1C,D, statistical tests for the chromosome alignment assays throughout the study are shown in Table 1). As such unaligned chromosomes were rare, we also treated cells with 10 ng/ml nocodazole, a condition that moderately stabilizes microtubules (Vasquez et al., 1997), and that delays chromosome alignment, leading to 3–6 unaligned chromosomes per cell (Figure 1C). Unaligned chromosomes were again preferentially found in the vicinity of the old centrosomes in HeLa eGFP-centrin1 (63.9%) and hTert-RPE1-eGFP-centrin1 cells (71.8%), confirming the bias in chromosome alignment (Figure 1E). We found the same bias (67.8%) in wild-type nocodazole-treated hTert-RPE1 cells stained with cenexin, excluding any effect due to eGFP-centrin1 expression (Figure 1E). We conclude that the half-spindles associated to the old centrosomes accumulate more unaligned chromosomes or that unaligned chromosomes align less efficiently when bound to microtubules emanating from the old centrosomes.

Centrosome age affects chromosome alignment.

(A) HeLa-eGFP-centrin1 (green) cell stained for cenexin (red, old centrosome marker) and DAPI (blue, DNA). One spindle pole contains the old centriole (brightest centrin1 signal and cenexin positive) and an accompanying daughter centriole (dim signal), which together form the old centrosome. The other spindle pole contains the young centriole (intermediate centrin1 signal), which is also accompanied by a daughter centriole and which together form the young centrosome. Scale bar in all panels = 5 μm. (B) Amounts of eGFP-centrin1 on the old, young and daughter centrioles in HeLa-eGFP-centrin1 cells determined from 3 independent experiments in 140 cells. (C) Untreated HeLa-eGFP-centrin1 cell (upper panel) and hTert RPE-eGFP-centrin1 cell treated with 10 ng/ml nocodazole (lower panel) stained for CENP-A (kinetochore marker) and DAPI. Yellow arrowheads indicate unaligned chromosomes; white arrowheads old centrosomes. (D and E) Proportion of unaligned chromosomes at old centrosomes in HeLa-eGFP-centrin1 cells (D), and in RPE1 cells stained for cenexin, RPE1-eGFP-centrin1 cells and HeLa-eGFP-centrin1 cells treated with 10 ng/ml nocodazole (E). For experiment and cell numbers, and p-values see Table 1. For results of individual experiments see Figure 1—source data 1. Error bars indicate s.e.m. * indicates p ≤ 0.05 in Binomial test compared to random distribution, *** indicates p ≤ 0.01 in Binomial test compared to random distribution.

https://doi.org/10.7554/eLife.07909.003
Table 1

Percentage of unaligned chromosomes at old centrosomes

https://doi.org/10.7554/eLife.07909.005
ConditionN0 of repeatsN0 of cellsN0 of chromosomes% Chromosomes at old centrosomes2-tailed Binomial test p
HeLa-eGFP-centrin1 DMSO3339361.230.037
hTert-RPE1 10 ng/ml nocodazole316122767.808.08e-8
hTert-RPE1-eGFP-centrin1 10 ng/ml nocodazole712729571.811.0e-12
HeLa-eGFP-centrin1 10 ng/ml nocodazole35753263.911.42e-10
hTert-RPE1 Eg5 inhibition recovery35315674.368.9e-10
hTert-RPE1 nocodazole recovery35916461.580.00373
HeLa-eGFP-CENP-A/eGFP-centrin1 10 ng/ml nocodazole314294658.038.68e-07
HeLa-eGFP-CENP-A/eGFP-centrin1 Eg5 inhibition recovery56830661.110.000121
hTert-RPE1-eGFP-centrin1 siCtrl 10 ng/ml nocodazole39220668.451.26e-07
hTert-RPE1-eGFP-centrin1 siNinein 10 ng/ml nocodazole47716966.860.0000138
hTert-RPE1-eGFP-centrin1 CENP-E inhibitor36539349.87n.s*
hTert-RPE1-eGFP-centrin1 CENP-E inhibitor 10 ng/ml nocodazole35945952.29n.s*
hTert RPE-eGFP-centrin1 5 nM Taxol35010543.81n.s*
hTert RPE-eGFP-centrin1 siDsn1 10 ng/ml nocodazole34216243.82n.s*
hTert RPE-eGFP-centrin1 siNnf1 10 ng/ml nocodazole5214652.17n.s*
  1. *

    Non-significant.

The bias in unaligned chromosomes could reflect faster kinetics in the initial capture of sister-kinetochore pairs by old centrosomes, for example, because they are closer to kinetochores at nuclear envelope breakdown or because they mature—that is, acquire a high, mitotic microtubule-nucleating capacity—earlier. Alternatively, the bias could reflect a permanent difference between the two centrosomes to capture or to align chromosomes. To test whether at mitotic onset old centrosomes capture kinetochores faster because they are closer, we compared the distances between kinetochores and old and young centrosomes at mitotic onset, but found no difference (Figure 2A). We next forced hTert-RPE1 or HeLa cells to enter mitosis with monopolar spindles by treating them with monastrol, a reversible inhibitor of Eg5, the kinesin that separates centrosomes (Mayer et al., 1999). A monastrol washout led to bipolar spindles with few unaligned chromosomes, 74.4% of which were adjacent to old centrosomes, indicating that the bias is independent of the initial centrosome position (Figure 2B,C; and Figure 2—figure supplement 1). To test whether old centrosomes capture more kinetochores because they mature earlier, we treated cells with high doses of nocodazole (1 μg/ml), allowing them to enter mitosis without microtubules and to fully mature the two centrosomes (Khodjakov and Rieder, 1999), before washing out nocodazole for 1 hr: 62.6% of the unaligned chromosomes were adjacent to old centrosomes, indicating that the alignment bias reflects a permanent difference between the centrosomes that is independent of the initial conditions at mitotic onset (Figure 2B,C). The two washout experiments also confirmed that this bias does not require low nocodazole concentrations, since in both cases, cells were released in nocodazole-free medium.

Figure 2 with 2 supplements see all
The asymmetric distribution of unaligned chromosomes depends on CENP-E.

(A) An illustrative example of a HeLa-eGFP-centrin1/mCherry-CENP-A cell where the distances from centrosomes to kinetochores were measured 30 s before nuclear envelope breakdown (left top), assay to calculate the distances between kinetochores and centrosomes (left bottom), and distribution (right) of the measured distances. Values are determined from 24 cells and 1434 kinetochores in 6 independent experiments. White arrowheads indicate old centrosomes in all panels. Scale bars in all panels = 5 μm. (B) Proportion of unaligned chromosomes at the old centrosomes in hTert-RPE1-eGFP-centrin1 cells after indicated treatments. Error bars indicate s.e.m. *** indicates p ≤ 0.01 in Binomial test. (C) hTert-RPE1-eGFP-centrin1 cells stained for CENP-A and DAPI after indicated treatments. (D) Differences in the intensity of the microtubule asters in a re-nucleation assay at old and young centrosomes as shown in E, calculated in 32–49 cells in 3 independent experiments. Columns indicate the median, errors bars the 99% CI. Precise methodology is shown in Figure 2—figure supplement 2. (E) HeLa-eGFP-centrin1 or hTert-RPE1-eGFP-centrin1 cells stained for α-tubulin after a microtubule re-nucleation assay. (F) hTert-RPE1-eGFP-centrin1 cells treated with 10 ng/ml nocodazole and/or CENP-E inhibitor, and stained for CENP-A. Yellow arrowheads indicate unaligned kinetochores in the proximity of the young centrosome (G) Proportion of unaligned chromosomes at old centrosomes in hTert-RPE1-eGFP-centrin1 cells treated with 10 ng/ml nocodazole and/or CENP-E inhibitor. Error bars indicate s.e.m. *** indicates p ≤ 0.01 in Binomial test. For results of all individual experiments see Figure 2—source data 1.

https://doi.org/10.7554/eLife.07909.006

In asymmetric cell divisions the old and young centrosomes have different capacities to nucleate microtubules, providing a key clue for centrosome positioning and inheritance (Wang et al., 2009; Januschke et al., 2013; Lerit and Rusan, 2013). If microtubule nucleation from the two centrosomes also differed in symmetric cell division, this might allow one centrosome to capture more kinetochores. However, a microtubule re-nucleation assay revealed no difference in microtubule nucleation capacity between the two centrosomes in HeLa and RPE1 cells (Figure 2D,E), suggesting that the centrosomal microtubule nucleation capacity did not cause the biased distribution of unaligned chromosomes. To study if chromosome alignment process per se is asymmetric, we inhibited the Centromere-associated Protein E (CENP-E), the kinetochore-bound kinesin that aligns polar chromosomes by transporting them along existing spindle microtubules (Wood et al., 1997; Kapoor et al., 2006; Barisic et al., 2014). Partial CENP-E inhibition, yielding few polar chromosomes, abolished the bias in the distribution of unaligned chromosomes in the absence or presence of 10 ng/ml nocodazole, (Figure 2F,G, 49.87% and 52.29% respectively), indicating that the bias depends on CENP-E, and that chromosome alignment itself is biased by centrosome age.

The CENP-E-dependent alignment bias could be due to an asymmetric abundance of CENP-E; however, the levels of CENP-E on unaligned chromosomes associated to old or young centrosomes were equal (Figure 3A,B and Figure 3—figure supplement 1A). Alternatively, since CENP-E favours lateral kinetochore–microtubule attachments to transport unaligned chromosomes towards the metaphase plate (Kapoor et al., 2006), we reasoned that a difference in the types of kinetochore–microtubule attachments might bias the alignment of unaligned chromosomes: specifically end-on attachments might delay CENP-E driven chromosome alignment, by creating a poleward drag. Indeed, chromosomes that are not captured by microtubules emanating from both poles, bind laterally to microtubules from the closest pole, and are first driven to this pole in a dynein-dependent manner, before CENP-E aligns them on the metaphase plate (Barisic et al., 2014). During these movements, kinetochores can in some cases form end-on monotelic or syntelic attachments. These non-bipolar end-on attachments are normally destabilized in an Aurora-B-dependent manner (Hauf et al., 2003), favouring the formation of lateral kinetochore–microtubule attachments. However, if end-on attachments were to be more stable at one centrosome, they would delay this conversion and create a drag on the CENP-E driven alignment. To test this hypothesis, we visualized by 3D-high-resolution microscopy kinetochore–microtubule attachments of individual, single kinetochores on unaligned chromosomes in cells treated with 10 ng/ml nocodazole and fixed with glutaraldehyde. At old centrosomes nearly three times more individual kinetochores had end-on attachments (13.0% vs 4.7% at the young centrosome, p = 0.003 in paired t-test) and fewer lateral attachments (83.6% vs 89.1% at the young centrosomes; p = 0.0007 in paired t-test; overall p = 0.0024 in 2way-ANOVA-test; Figure 3C,D and Figure 3—figure supplement 2); the number of unattached kinetochores was higher at young centrosomes, even tough this difference was statistically not significant (p = 0.06 in paired t-test; Figure 3—figure supplement 2). This implied an overall higher stability of kinetochore–microtubules at old centrosomes. To confirm this result, we quantified the levels of tubulin acetylation on individual kinetochore–fibres of sister-kinetochore pairs aligned on the metaphase plate, as tubulin acetylation preferentially accumulates on stable microtubules (Webster and Borisy, 1989). This analysis revealed higher level of acetylation on kinetochore–microtubules associated with old centrosomes (median difference of 22% in tubulin acetylation at the plus ends of microtubules attached to sister-kinetochores vs 2% in CREST levels between sister-kinetochores, p < 0.0001 in Wilcoxon Signed Rank Test; Figure 3E,F and Figure 3—figure supplement 1C). In contrast, when we measured the levels of detyrosinated tubulin, a modification that has been linked to preferential CENP-E motor activity (Barisic et al., 2015), we found no difference (median difference of 0.3%; Figure 3F and Figure 3—figure supplement 1D). To also functionally confirm the difference in microtubule stability, metaphase cells were treated for 10 min with 0.5 mM Ca2+, a condition that gradually destabilizes microtubules, before fixing them with glutaraldehyde and staining for kinetochores and microtubules. While such a treatment did not reveal strong overall differences in the two half-spindles (Figure 3G), a detailed analysis of kinetochore–microtubule attachments of aligned sister-kinetochores revealed that kinetochores oriented towards young centrosomes were significantly more likely to have lost their attachment, than those oriented towards the old centrosome (56.8% vs 43.2%; p = 0.014 in paired t-test; Figure 3G,H). Together, these data indicated that the kinetochore–microtubules emanating from old centrosomes are more stable.

Figure 3 with 4 supplements see all
Kinetochore–microtubules bound to old centrosomes are more stable.

(A) hTert-RPE1-eGFP-centrin1 cells treated with 10 ng/ml nocodazole and stained for CENP-E and DAPI. White arrowheads indicate old centrosomes in all panels. Scale bars in all panels = 5 μm. (B) Differences in the abundance of CENP-E and CREST (kinetochore marker) at kinetochores bound to old and young centrosomes, calculated from 27 cells in 3 independent experiments. Columns indicate the median; error bars the 99% CI. (C) Immunofluorescence image of a HeLa-eGFP-centrin1/eGFP-CENP-A (green) cells treated with 10 ng/ml nocodazole, fixed with glutaraldehyde, and stained for α-tubulin (red) and DAPI (blue). Single kinetochores in every unaligned sister-kinetochore pair were classified as end-on attached, laterally attached or unattached. Inset 1 on the left shows an illustrative example of a kinetochore pair with one unattached and one end-on attached kinetochore; inset 2 on the right shows an illustrative example with 2 laterally attached kinetochores. (D) Quantification of individual end-on attached kinetochores at old and young centrosomes in HeLa-eGFP-centrin1/eGFP-CENP-A cells treated with 10 ng/ml nocodazole, 5 nM taxol and the indicated siRNAs and 10 ng/ml nocodazole. Percentages are based on 3 independent experiments with 29–50 cells. Error bars indicate s.e.m; *** indicates p ≤ 0.01 in paired t-test. (E) hTertRPE1-eGFP-centrin1 cells stained with anti-acetylated tubulin (red) and CREST (green) antibodies. Shown are total projections (upper panels) or maximum-intensity projections of 5–10 planes around the focal plane of interest (lower panels). White arrowheads indicate kinetochore–microtubules with stronger acetylation, yellow with weaker acetylation. Note that the white arrows are on the side of the old centrosome. (F) Differences in the abundance of acetylated tubulin on k-fibres of sister-kinetochores, and detyrosinated tubulin on the two spindle halves in hTertRPE1-eGFP-centrin1 cells, based on 3 independent experiments and 32–33 cells. Methodology is explained in Figure 3—figure supplement 2. Columns indicate the meadian, error bars the 99% CI. (G) HeLa-eGFP-centrin1/eGFP-CENP-A cells treated with 0.5 mM Ca2+ for 10 min stained for α-tubulin (red) and DAPI (blue). Shown are total projections (upper panels) or maximum-intensity projections of 5–10 planes around the focal plane of interest (lower panels). White arrowheads in zoom-ins indicate end-on attached kinetochores and yellow arrow the unattached kinetochore. (H) Percentage of unattached kinetochores oriented towards old or young poles based on 3 independent experiments and 33 cells. (I) hTert-RPE1-eGFP-centrin1 cells stained with CENP-A antibodies (red) and DAPI (blue) after treatment with 5 nM taxol or the indicated siRNAs and 10 ng/ml nocodazole. White arrowheads indicate old centrosome. (J) Proportion of unaligned chromosomes at old centrosome in hTert-RPE1-eGFP-centrin1 cells treated with 5 nM taxol or with 10 ng/ml nocodazole after the indicated siRNA treatment. Error bars indicate s.e.m; *** indicates p ≤ 0.01 in Binomial test. (K) Differences in the abundance of cenexin, phospho-Aurora-A, and Plk1 at old and young centrosomes in HeLa and hTert-RPE1-eGFP-centrin1 cells, based on 3 independent experiments and 41–113 cells. Methodology is explained in Figure 3—figure supplement 2. Columns indicate the median, error bars the 99% CI. (L) Proportion of unaligned chromosomes at old centrosome in HeLa-eGFP-centrin1 cells treated with Aurora-A or Plk1 inhibitors. Error bars indicate s.e.m; *** indicates p ≤ 0.01 in Binomial test. For results of all individual experiments see Figure 3—source data 1.

https://doi.org/10.7554/eLife.07909.010

To test if the difference in kinetochore–microtubule stability is at the origin of the alignment bias, we depleted the kinetochore proteins Dsn1 or Nnf1 (both Mis12 complex), or treated the cells with a low dose of taxol (5 nM). These conditions strongly destabilize kinetochore–microtubules (Dsn1 and Nnf1 depletion; (Kline et al., 2006), or hyperstabilize kinetochore–microtubules (taxol; Figure 3—figure supplement 3). Either treatment abolished the bias in alignment and equalized the number of end-on attached unaligned kinetochores (Figure 3D,I,J). In contrast, knock-down of the Mitotic Centromere-Associated Kinesin (MCAK), a microtubule depolymerase that is required for destabilization of erroneous kinetochore–microtubule attachments (Knowlton et al., 2006), whose depletion leads to mild microtubule stabilization in metaphase, but not in prometaphase ((Bakhoum et al., 2009) and Figure 3—figure supplement 3), did not change the bias in chromosome alignment (Figure 3I,J). This suggested that a massive stabilization or destabilization of all kinetochore–microtubules equilibrates the difference in kinetochore–microtubule stability and chromosome alignment, but that a mild stabilization does not change this bias. We conclude that the difference in kinetochore–microtubule stability biases chromosome alignment.

Which factors at centrosomes could generate an age-dependent difference in kinetochore–microtubule stability causing a bias in chromosome alignment? We first considered two centrosomal kinases, Aurora-A and Plk1, which can both affect kinetochore–microtubule stability (Liu et al., 2012; Bakhoum et al., 2014). We compared by quantitative immunofluorescence the levels of Plk1 or the activity of Aurora-A (with an antibody that is specific for active Aurora-A) at old and new centrosomes, to reveal a potential asymmetry in kinase levels/activity. While Plk1 was symmetrically distributed, we found a modest increase of active Aurora-A on old centrosomes in HeLa cells (Figure 3K and Figure 3—figure supplement 1B). This difference was, however, not present in RPE1 cells (Figure 3K); moreover inhibition of Aurora-A or Plk1 did not abolish the bias in chromosome alignment, indicating that it does not depend on these two kinases (Figure 3L). In a second step, we investigated the possible involvement of ninein, as it is essential for cell fate determination in asymmetric cell divisions of neuronal progenitors and preferentially localizes to old centrosomes in asymmetric cell division (Wang et al., 2009), and of cenexin itself, the classical marker for old centrosomes (Figure 3K—note that ninein levels could not be compared on old and young mitotic centrosomes, as it is only present at very low levels (Logarinho et al., 2012)). While ninein depletion had no effect on chromosome alignment, cenexin depletion randomized the distribution of unaligned chromosomes (Figure 3I,J). Furthermore, it also equalized the percentage of end-on attached kinetochores at unaligned chromosomes, indicating that cenexin affects kinetochore–microtubule stability (Figure 3D). We conclude that old centrosomes stabilize kinetochore–microtubules in a cenexin-dependent manner.

The ultimate function of the mitotic spindle is to accurately segregate sister chromatids. If kinetochore–microtubules bound to the old centrosomes were more stable, we predicted that this should affect the fate of chromosomes that fail to fully disjoin in anaphase; chromosome non-disjunction is a frequent cause of chromosome mis-segregation in cancer cells, that can be caused by various defects, such as stretches of unreplicated DNA, telomere fusions, or chromosome entanglements (Aguilera and García-Muse, 2013). To monitor the fate of such chromosome non-disjunction, we monitored by live-cell imaging HeLa-eGFP-centrin1/mCherry-CENP-A cells released for synchronization purpose from a monastrol arrest. As previously reported, this procedure produced a number of single lagging, most likely merotelic, kinetochores, whose exact fate could not be tracked. However, in addition in roughly 2–5% of anaphases, we observed the presence of two lagging kinetochores that moved in synchrony between the two daughter DNA masses, but were separated by several microns, suggesting a sister-kinetochore pair on a non-disjoint chromosome (Figure 4A and Video 1). This assumption was confirmed by high-resolution immunofluorescence imaging, as such kinetochore pairs were invariably connected by a DNA thread (see representative images in Figure 4B). In those instances where both sister-kinetochores segregated to the same daughter cell, we found a strong bias in chromosome mis-segregation as 18 out of the 21 analyzed kinetochore pairs co-segregated with the old centrosome (Figure 4C; Video 1; number of experiments, cells and statistical tests for all chromosome mis-segregation events are in Table 2). This suggested that non-disjoint chromosomes are preferentially pulled towards the old centrosomes, possibly due to a higher stability of the kinetochore–microtubules emanating from the old centrosomes, which in a tug-of-war would favour a destabilization and release of the kinetochore–microtubules bound to the young pole. To test this hypothesis, we treated cells with Nnf1 and Cenexin siRNAs, which had abolished the bias in chromosome alignment and the asymmetry in the percentage of end-on attached kinetochores. In both cases, the bias in chromosome mis-segregation was abolished (Figure 4C; Video 2 and 3); in contrast when we depleted MCAK, which did not abolish the bias in chromosome alignment, chromosome mis-segregation was still biased (Figure 4C). We conclude that the difference in the stability of kinetochore–microtubules bound to the old or the young centrosome persists in anaphase, and that this difference causes non-disjoint chromosomes to co-segregate with old centrosomes.

Non-disjoined chromosomes co-segregate with old centrosomes.

(A) Time lapse images of a HeLa-eGFP-centrin1/mCherry-CENP-A cell with a non-disjoined sister-kinetochore pair in anaphase. White arrowhead indicates the old centrosome, yellow arrowheads the non-disjoined sister-kinetochore pair. Scale bar = 10 μm. (B) Illustrative example of a HeLa-eGFP-centrin1/eGFP-CENP-A cell in anaphase stained for α-tubulin with a non-disjoined chromosome. Insets highlight the non-disjoined chromosomes. Scale bar = 5 μm. (C) Proportion of non-disjoined chromosomes that co-segregate with the old centrosomes in HeLa-eGFP-centrin1/mCherry-CENP-A cells treated with the indicated siRNAs. For statistics and number of experiments, see Table 2. (D) Proposed model of how old and new centrosomes differentially affect chromosome alignment and chromosome segregation via kinetochore–microtubule stability.

https://doi.org/10.7554/eLife.07909.016
Video 1
HeLa cell expressing mCherry-CENP-A (kinetochore marker in red) and eGFP-centrin1 (centrosome age marker in green) after a monastrol washout.

Note that the mis-segregating chromosome moves towards the brighter, old centrosome.

https://doi.org/10.7554/eLife.07909.017
Table 2

Percentage of mis-segregating chromosomes that co-segregate with the old centrosomes

https://doi.org/10.7554/eLife.07909.018
ConditionN0 of repeatsNo of cellsNo of chromosomesNo of chromosomes to the old centrosome2-tailed binomial test p
HeLa-eGFP-centrin1/ mCherry-CENP-A112121180.0015
HeLa-eGFP-centrin1/ mCherry-CENP-A siMCAK38880.0078
HeLa-eGFP-centrin1/ mCherry-CENP-A siNnf163030100.0990
HeLa-eGFP-centrin1/ mCherry-CENP-A siCenexin61616100.4545
Video 2
HeLa cell expressing mCherry-CENP-A (kinetochore marker in red) and eGFP-centrin1 (centrosome age marker in green) depleted of Nnf1, after a monastrol washout.

Note that the mis-segregating chromosome moves towards the dimmer, young centrosome.

https://doi.org/10.7554/eLife.07909.019
Video 3
HeLa cell expressing mCherry-CENP-A (kinetochore marker in red) and eGFP-centrin1 (centrosome age marker in green) depleted of Cenexin, after a monastrol washout.

Note that the mis-segregating chromosome moves towards the dimmer, young centrosome.

https://doi.org/10.7554/eLife.07909.020

Discussion

Our results demonstrate that even in symmetrically dividing cells the two half-spindles behave in an asymmetric manner, and that centrosome age imposes a functional asymmetry on all mitotic spindles (see model Figure 4D). We find that this asymmetry reflects a differential stability of kinetochore–microtubules that depends on the presence of cenexin at old centrosomes, indicating that its presence influences the relative stability of kinetochore–microtubules. Previous studies demonstrated that knock-out of cenexin does not impair mitotic progression; it contributes, however, to the stability of the centrosome-bound microtubules during interphase, consistent with our findings that cenexin affects mitotic microtubules (Ishikawa et al., 2005; Tateishi et al., 2013). Cenexin is known since a long time as a marker for old centrioles (Lange and Gull, 1995), yet the molecular mechanisms by which it affects microtubules are unclear. It is required for the formation of distal and sub-distal appendages on the oldest centriole, two structures that are essential for the formation of basal bodies during ciliogenesis (Ishikawa et al., 2005). These structures consist of more then ten different centrosomal proteins, including CEP164, CEP123, CEP83, SCLT1 and FBF1 at the distal appendages and ninein, centriolin, ε-tubulin, trichoplein and CEP170 at sub-distal appendages (Mogensen et al., 2000; Chang et al., 2003; Gromley et al., 2003; Guarguaglini et al., 2005; Graser et al., 2007; Ibi et al., 2011; Sillibourne et al., 2013; Tanos et al., 2013; Tateishi et al., 2013). Moreover Plk1 has been shown to bind Odf2 at centrosomes (Soung et al., 2009). Future work will thus have to evaluate whether the ability to stabilize microtubules is a direct function of cenexin, or a more general function of centriolar appendages. We speculate that cenexin or some of its associated centrosomal proteins might control microtubule stability via three potential mechanisms: first, they could directly interact with microtubule plus-ends, as has been seen for γ-tubulin (Bouissou et al., 2009); second, they could affect microtubule plus-end dynamics by controlling the dynamics of the minus-end, a type of regulation that has been seen in the context of poleward microtubule flux (Maddox et al., 2003; Ganem et al., 2005; Matos et al., 2009); third they could act via centrosomal protein kinases, as it has been recently shown as a proof-of-principle that centrosomal-bound Aurora-A has the ability to regulate kinetochore–microtubule attachments (Chmátal et al., 2015; Ye et al., 2015).

The asymmetry in kinetochore–microtubule stability persists throughout mitosis and directs the fate of non-disjoint chromosomes, which co-segregate mostly with the old centrosome. At present it is unclear whether this asymmetry serves a direct purpose during mitosis, or if it a consequence of a centrosome asymmetry that is required for a non-mitotic function, but which cells have to deal with during each cell division. Possible non-mitotic purposes of this asymmetry include the necessity to only have one centriole capable of generating the basal body of a cilium, or the requirement to only have one centriole capable of anchoring interphase microtubules (Piel et al., 2000; Nigg and Raff, 2009). A possible mitotic function for an asymmetric behaviour of centrosomes is that it might protect stem cells that inherit the old centrosome, from losing non-disjoint chromosomes. This would provide a selective advantage, as haplo-insufficiency is much more frequent than triplo-lethality at the level of single genes in animal cells (Lindsley et al., 1972; Torres et al., 2007). We speculate that the asymmetric distribution of non-disjoint chromosomes might, however, also favour the rapid acquisition of new traits by co-occurrence of chromosome gains in cancer-stem cells. Even though gain/loss of non-disjoint chromosomes is only one of the causes of chromosomal instability in cancer cells, chromosome gain of non-disjoint chromosomes would not be random, but a favoured behaviour in cancer stem cells. Such a potential bias should thus be considered when modelling chromosomal instability in aneuploid cancer cell populations. Consistent with this hypothesis we note that the analysis of large human cancer samples reveals that whole chromosome gains (or losses) co-occur at much higher frequencies than combined chromosome gains and losses (Ozery-Flato et al., 2011). This phenomenon was so far thought to be the result of an evolutionary pressure; we propose that an asymmetric chromosome mis-segregation in cancer stem cells might provide a direct mechanistic explanation for this behaviour. Moreover, it could suggest a more general asymmetry in chromosomal instability, pointing to the need to determine whether other forms of chromosome mis-segregation, such as gain/loss of merotelic chromosomes, depend on centrosome age or not.

Materials and methods

Cell culture, drug, and siRNA treatments

HeLa, hTert-RPE1, and hTert-RPE1-eGFP-centrin1 cells (kind gift of A. Khodjakov) were grown in Dulbecco's modified medium (DMEM) supplemented with 10% FCS, 100 U/ml penicillin, 100 mg/ml streptomycin, at 37°C with CO2 in a humidified incubator. HeLa-eGFP-centrin1 cells (kind gift of S. Doxsey) were further maintained in 500 μg/ml G418. HeLa-eGFP-centrin1/mCherry-CENP-A cells were generated by stably transfecting eGFP-centrin1 in HeLa-mCherry-CENP-A cells (kind gift of A. McAinsh, U. of Warwick); as HeLa-eGFP-centrin1/CENP-A-GFP, they were further supplemented with 500 μg/ml puromycin and 500 μg/ml G418. Live-cell imaging experiments were performed at 37°C in Lab-Tek II (Thermofisher, Switzerland) and Ibidi IV (Ibidi, Switzerland) chambers in L-15 medium supplemented with 10% FCS. To enrich for unaligned chromosomes, mitotic cells were removed by shake-off and the remaining cells treated with 10 ng/ml nocodazole for 2 hr. For nocodazole and monastrol washout experiments, cells were treated with either 1 μg/ml nocodazole or 100 nM monastrol for 4 hr (Sigma, Switzerland), washed twice with fresh medium and left to recover for 1 hr. Aurora-A was inhibited for 2 hr with 100 nM MLN8237 (Selleckchem.com, Switzerland), Plk1 for 2 hr with 100 nM BI2536 (Axon Lab AG, Switzerland), and CENP-E for 2 hr with 5 nM GSK-923295 (Chem Express, Switzerland). To stabilize microtubules, cells were treated with 5 nM Taxol (Sigma, Switzerland) for 2hr. To monitor anaphase cells, cells were released from a monastrol arrest and followed by live cell imaging. The following SiRNA oligonucleotides (Invitrogen and Thermofisher, Switzerland) were used: siControl (scrambled) (Mchedlishvili et al., 2012), siNinein (Logarinho et al., 2012), siDsn1 (Kline et al., 2006), siNnf1 (McAinsh et al., 2006), siMCAK (Ganem and Compton, 2004), siCenexin (OnTarget smart pool, L-017319-01-0005, Thermofisher); they were transfected using RNAi Max Lipofectamine (Invitrogen) and validated by immunofluorescence microscopy (Figure 3—figure supplement 4).

Quantitative immunofluorescence

Cells were fixed with methanol at −20°C for 6 min, or with 20 nM Pipes (pH 6.8), 10 mM EGTA, 1 mM MgCl2, 0.2% Triton X-100, 4% formaldehyde for 7 min at room temperature. For the microtubule nucleation assay, cells were incubated on ice for 1 hr before release in 37°C medium for 15 s for RPE-eGFP-centrin1 and 30 s for HeLa-eGFP-centrin1 cells. To image the attachment state of unaligned kinetochores, cells were rinsed with cytoskeleton buffer (10 mM MES, 150 mM NaCl, 5 mM MgCl2, 5 mM glucose) prior and after fixation with 3% formaldehyde, 0.1% Triton X-100, and 0.05% glutaraldehyde for 10 min at room temperature. To analyze tubulin acetylation and detyrosination, cells were fixed with 20 nM Pipes (pH 6.8), 10 mM EGTA, 1 mM MgCl2, 0.2% Triton X-100, 4% formaldehyde for 7 min at room temperature. To image the Calcium stability of kinetochore–microtubules, cells were treated with 0.5 nM CaCl2 dissolved in warm DMEM for 10 min at room temperature, rinsed with cytoskeleton buffer (10 mM MES, 150 mM NaCl, 5 mM MgCl2, 5 mM glucose) prior and after fixation with 3% formaldehyde, 0.1% Triton X-100 and 0.05% glutaraldehyde for 10 min at room temperature. Three-dimensional image stacks of mitotic cells were acquired in 0.1- or 0.2-μm steps using 100x and 60x NA 1.4 objectives on an Olympus DeltaVision microscope (GE Healthcare, Switzerland) equipped with DAPI/FITC/TRITC/CY5 filter set (Chroma, Bellow Falls, VT) and a CoolSNAP HQ camera (Roper Scientific, Tuscon USA). 3D image stacks were deconvolved with SoftWorx (GE Healthcare) and analyzed with SoftWorx, Imaris (Bitplane, Switzerland) or ImageJ. For the nucleation assay, deconvolved total projections were analyzed as shown in Figure 1B. For the attachment status of unaligned kinetochores, single kinetochores were analyzed in 3D reconstruction of several z-stacks, and classified as shown in Figure 3C. To analyze Calcium stability, individual kinetochores were displayed in single z-planes and classified as shown in Figure 3E. The difference in microtubule nucleation capacity at old and young centrosomes was calculated as shown in Figure 2—figure supplement 2. The differences in protein levels at centrosomes or unaligned kinetochores at old and young centrosomes were calculated as shown in Figure 3—figure supplement 1. Images were mounted as figures using Adobe Illustrator. Primary antibodies used were mouse anti-CENP-A (1:2000, Abcam, United Kingdom), mouse anti-α-tubulin (1:10′000, Sigma), mouse anti-acetylated tubulin (1:1000; Sigma), rabbit anti-detyrosinated tubulin (1:1000, Merck-Millipore, Switzerland), rabbit anti-α-tubulin (1:500, Abcam), human CREST (1:400, Antibodies Inc, Davis USA), rabbit anti-phosphoT288-Aurora-A (1:1000, Cell Signalling, Danvers USA), rabbit anti-Plk1 (1:1000, Abcam), rabbit anti-Cenexin (1:1000, Abcam), rabbit anti-Ninein (1:500, Abcam), rabbit anti-Nnf1 (1:1000, McAinsh et al., 2006), rabbit anti-Dsn1 (1:2000, kind gift of Iain Cheeseman Kline et al., 2006), rabbit anti-MCAK (1:1000, Amaro et al., 2010), and rabbit anti-CENP-E (1:1000, Meraldi et al., 2004). Cross-adsorbed secondary antibodies were used (Invitrogen).

Live imaging

To visually monitor the fate of non-disjoint chromosomes, Hela-eGFP-centrin1/mCherry-CENP-A cells were recorded every 2 or 4 min in 26 × 0.7-μm steps using a 60 × 1.4 NA objective on an Olympus DeltaVision microscope equipped with a GFP/mRFP filter set (Chroma) and a CoolSNAP HQ camera. To distinguish eGFP-centrin1 intensities in both experiments, reference images were taken at the end of the experiment, as three-dimensional stacks of 40 × 0.3-μm steps and a high-exposure times using the same objective and camera. Time-lapse movies were visualized in Imaris (Bitplane). To calculate distances, three-dimensional positions of the old and the young centriole and of all the kinetochores were detected using Imaris (Bitplane) and distances calculated with a custom MatLab function (see source code 1).

Statistical methods

To check for biased distribution of polar chromosomes, Binomial probability test with expected probability success on a single trial of 0.5 was used. To calculate 2-ANOVAs, medians and median confidence intervals, and run t-tests PRISM (GraphPad, La Jolla, CA) were used. Graphs were plotted in Excel (Microsoft, Redmond, WA) and PRISM and mounted in Adobe Illustrator (Adobe, Mountain View, CA).

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Decision letter

  1. Anna Akhmanova
    Reviewing Editor; Utrecht University, Netherlands

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for submitting your work entitled “Centrosome age regulates kinetochore microtubule stability and biases chromosome mis-segregation” for peer review at eLife. Your submission has been favorably evaluated by Tony Hunter (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors.

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

In this study, the authors test if centrosome age influences symmetric cell divisions using live and fixed cell analysis. The authors observed that unaligned chromosomes preferentially associate with old centrosomes with or without the nocodazole treatment to enhance the numbers of unaligned chromosomes. Using calcium treatment or RNAi depletion of proteins to perturb kinetochore microtubule stability, they go on to suggest that this bias occurs because kinetochore microtubule attachments are more stable at old centrosomes. Further, they demonstrate that the stability of these attachments also relies on the old centrosome protein cenexin. Finally, using live and fixed cell imaging, the authors show that this asymmetry contributes to a bias during chromosome non-disjunction with the daughter cell inheriting the old centrosome preferentially retaining the non-disjoined chromosomes. The observation that centrosome age influences chromosome alignment and imparts a functional asymmetry on mitotic spindles to influence the orientation of chromosome non-disjunction is quite novel and is a very interesting phenomenon.

Essential revisions:

1) The most important point: much stronger experimental support for the idea that centrosome age contributes to differences in kinetochore microtubule stability must be provided. One possibility would be to use microtubule photoactivation experiments to measure kinetochore microtubule stability in combination with a fluorescent marker of centrosme age, such as centrin 1. Perhaps detailed quantitative analysis of tubulin post-translational modifications could also be of some help here.

2) The underlying mechanism is mysterious - how do centrosomal proteins affect kinetochore fiber stability? Ideally, some additional experimental data on this point should be added. At the very least, a much better discussion of the possible mechanisms should be included.

3) In a broad sense, the data suggest that end-on kinetochore microtubule attachments precede sidewall microtubule attachments. But, that is counter to all other current models. How do the authors reconcile this discrepancy?

4) What is the difference between Figure 3C panels 1 and 2? Both are showing an end-on and unattached kinetochore. The cartoon diagram suggests that difference is the presence or absence of CENP-E but CENP-E localization is not determined in the experiment. The authors need to clarify their classification of attachment status. Also, the role of CENP-E is not clearly defined relative to microtubule dynamics, and this should be more clearly explained.

5) The authors present data suggesting that hyperstabilization by depletion of CENP-H abolishes the alignment bias but that “mild” stabilization by MCAK depletion does not. Can they provide a more quantitative assessment for the difference between CENP-H and MCAK depletion on microtubule stability (Figure 3)? It is unclear why MCAK depletion does not result in an unbiased distribution.

6) The authors state that in 2-5% of HeLa cells after monastrol wash-out they observe anaphases with non-disjoined chromosomes that preferentially segregate to the daughter cell with the old centrosome. However, the dominant defect reported by many labs for these types of washouts is lagging chromosomes (i.e. single chromatids) and not non-disjoined chromosomes. Do the authors observe lagging chromosomes and do these segregate preferentially to daughter cells with old centrosomes? Is the asymmetric segregation specific to non-disjoined chromosomes? How do the authors explain why the rate of non-disjoined chromosomes increases under these washout regimes?

7) It is not clear that the described phenomenon has a biological function. It might actually be just a consequence of the existing intrinsic centrosome asymmetry that cells have to deal with. This point deserves a better discussion.

8) The model figure needs revision. The authors claim that the two spindle halves have different kinetochore microtubule stability due to centrosome age but the diagram only shows the non-disjoined chromosome with different microtubule stability. Do they really mean that the difference in kinetochore microtubule stability is limited to just the lagging chromosomes? For this reason, it might be best to depict the half spindles differently.

9) The last paragraph of the Discussion randomly mentions cancer stem cells and chromosomal instability. This does not align with the rest of the text or data presented. There is nothing about cancer stem cells reported in the paper and chromosomal instability results from lagging chromosomes and not non-disjoined chromosomes. They authors need to revise the last paragraph.

https://doi.org/10.7554/eLife.07909.022

Author response

1) The most important point: much stronger experimental support for the idea that centrosome age contributes to differences in kinetochore microtubule stability must be provided. One possibility would be to use microtubule photoactivation experiments to measure kinetochore microtubule stability in combination with a fluorescent marker of centrosme age, such as centrin 1. Perhaps detailed quantitative analysis of tubulin post-translational modifications could also be of some help here.

We agree with the reviewers that this point is crucial to the paper and that any experimental support for our hypothesis strengthens this study. The reviewers suggested using microtubule photoactivation to directly compare the turnover of kinetochore-microtubules at the two half-spindles. For this present study the photoactivation assay is, however, unsuited for two reasons. First, photoactivation is effective at quantifying large differences in kinetochore-microtubule stability, but the method often shows a high variability and is therefore unlikely to reveal small differences. For example, after the depletion of the plus-end binding protein APC and after photoactivation Bakhoum et al found an increase of 30% in kinetochore-microtubule stability compared to control depleted cells (Bakhoum, Genovese, et al. 2009). This difference was, however, not significant, even though it is generally recognized that APC is an important microtubule plus-end regulator. This indicates that photoactivation cannot reproducibly reveal differences in k-fiber stability of 30% or less, which is more than the difference we would expect between the “old” and “young” half-spindle. The second caveat is specific to our particular experiment: the most sensitive tool to measure microtubule dynamics is a photoactivatable-GFP-α-tubulin, which would require a Cherry/RFP/Tomato-labeled version of centrin1. However, all our attempts to generate a red centrin1 that specifically labels centrioles in an age-dependent manner failed, as all constructs lead to protein aggregation of around the spindle poles (Alexey Khodjakov made very similar observations, and has never been able to produce a functional red Centrin1 construct). As an alternative, we have generated a cell line expressing the photo-convertible Eos-tubulin in conjunction with eGFP-centrin1; however the signal of Eos-tubulin masked the GFP-centrin1 signal at the poles, precluding any determination of centriole age.

To nevertheless test for a potential large difference in microtubule stability we used HeLa cells expressing PA-GFP-tubulin/H2B-RFP to “blindly” compare the dynamics at the two half-spindles, not knowing the age of the respective centrosome. We photoactivated GFP-tubulin along a one pixel wide line diagonally across the metaphase plate and monitored 3 photoactivated kinetochore-fibers in each spindle half. Our measurements revealed that the differences in dissipation of kinetochore-microtubules between the two half-spindles were as high as the differences amongst the k-fibers within a half-spindle, suggesting that any difference between the two half-spindles is likely to be small, and thus outside of the sensitivity range of the photoactivation assay.

As suggested by the reviewers we instead analyzed post-translational tubulin modifications as a read-out of kinetochore-microtubule stability. In particular, we focused on tubulin acetylation and detyrosination, two modifications that are associated with stable microtubules (Webster & Borisy 1989; Janke 2014). Whilst tubulin acetylation was easily detectable on k-fibers, tubulin, detyrosination was concentrated at spindle poles and diffusely present on the whole spindle (Figure 3-figure supplement 1). We therefore quantified tubulin acetylation at the plus-ends of k-fibers of individual sister-kinetochore pairs aligned on the metaphase plate, and the overall tubulin detyrosination in the two half-spindles (the heavily detyrosinated centrioles were excluded from our measurements). Our analysis revealed 20% more tubulin acetylation of kinetochore-fibers associated with the old centrosome. but no difference in tubulin detyrosination (Figure 3E and F and Figure 3-figure supplement 1). The strong difference in tubulin acetylation indicates that k-fibers ends associated to the old centrosome are more stable than the kinetochore-fibers associated with the young centrosome, fully supporting our hypothesis of a differential stability of k-fibers.

2) The underlying mechanism is mysterious – how do centrosomal proteins affect kinetochore fiber stability? Ideally, some additional experimental data on this point should be added. At the very least, a much better discussion of the possible mechanisms should be included.

We agree with the reviewers that the underlying mechanisms by which centrosomes affect kinetochore microtubule stability are mysterious and very interesting, but a better understanding of this phenomenon requires a thorough investigation that is beyond the scope of this revision. Nevertheless, as suggested by the reviewers we have added in the Discussion potential mechanisms by which centrosomal proteins could affect the dynamics of k-fiber plus-ends.

3) In a broad sense, the data suggest that end-on kinetochore microtubule attachments precede sidewall microtubule attachments. But, that is counter to all other current models. How do the authors reconcile this discrepancy?

We do not think that our data contradict the current models of how polar chromosomes are brought to the metaphase plate. There is a general consensus that kinetochores of chromosomes close to a spindle pole will first bind microtubules laterally, before dynein brings them closer to that same spindle pole. At this point the plus-end directed CENP-E will use lateral attachments to pre-existing k-fibers to move the unaligned chromosomes towards microtubule plus-ends to align them on the metaphase plate. Importantly, during the dynein-driven poleward motion, as they encounter a high density of microtubules, kinetochores may form a transient end-on attachment, resulting in monotelic or syntelic kinetochore-microtubule attachments. Monotelic and syntelic kinetochore-microtubule attachments are not frequent (less than 15% according to our data), as Aurora-B destabilizes these premature end-on attached kinetochores allowing them to revert to a lateral attachment, and thus favor CENP-E driven chromosome alignment onto the metaphase plate. Our data suggest that these transient end-on attachments close to spindle poles are more stable and therefore more persistent when kinetochores are bound to microtubules emanating from the old centrosomes; they remain however a transient state between lateral attachments, which is fully compatible with all existing models. To make these assumptions more explicit, we have now added 2 sentences paragraph 4 of the Results section.

4) What is the difference between Figure 3C panels 1 and 2? Both are showing an end-on and unattached kinetochore. The cartoon diagram suggests that difference is the presence or absence of CENP-E but CENP-E localization is not determined in the experiment. The authors need to clarify their classification of attachment status. Also, the role of CENP-E is not clearly defined relative to microtubule dynamics, and this should be more clearly explained.

We apologize for the confusion: there is no difference between panels 1 and 2 in Figure 3C, as in fact both contained one unattached and one end-on attached kinetochore. To improve the clarity of Figure 3C we have now removed panel 2, and point to the examples of end-on and unattached kinetochores in panel 1. To avoid further confusion, we have also removed CENP-E from the drawing, and rather illustrate how end-on attachments create a drag on CENP-E driven chromosome alignment in the final model in Figure 4. Indeed, our cartoon might have given the impression that the presence of CENP-E favors end-on attachment, which was not our message. Our model does not assume that CENP-E changes microtubule dynamics, but rather that it acts as a plus-end driven kinesin, as is widely recognized. We also have expanded the figure legend of Figure 3C to better explain our classification

5) The authors present data suggesting that hyperstabilization by depletion of CENP-H abolishes the alignment bias but that “mild” stabilization by MCAK depletion does not. Can they provide a more quantitative assessment for the difference between CENP-H and MCAK depletion on microtubule stability (Figure 3)? It is unclear why MCAK depletion does not result in an unbiased distribution.

With regard to the absence of an effect after MCAK depletion, we believe that it is consistent with the published data, as (Bakhoum, Thompson, et al. 2009) has shown that MCAK depletion has no effect on microtubule dynamics in prometaphase, which would be the time when we observe unaligned chromosomes. To strengthen this interpretation we also performed a cold stable assay (Figure 3–figure supplement 3), which confirmed that in contrast to Dsn1 depletion, MCAK depletion had no major effect on kinetochore microtubule stability.

With regard to CENP-H depletion, after further analysis we realize that the situation is more difficult to interpret than we originally assumed. Indeed, our previous publication (Amaro et al. 2010) showed that CENP-H depletion results in an ambivalent phenotype: a very dynamic plus-end that undergoes very rapid switches from microtubule polymerization to microtubule depolymerization, coupled to very stable k-fiber lattices. Even though CENP-H depletion reproducibly abolishes the bias in chromosome alignment, it is difficult to tell whether this is the result of the k-fiber stabilization or (more likely) the result of very dynamic plus-ends, which makes any interpretation hazardous. We therefore decided to remove the CENP-H treatment from this study and to replace it with low doses of taxol (5nM), as an unambiguous tool to stabilize k-fibers (Figure 3D, I and J and Figure 3–figure supplement 3). We confirm with a cold-stable assay that taxol stabilizes k-fibers, and we show that this stabilization abolishes the centrosome-age dependent bias in kinetochore-microtubule attachment and chromosome alignment. This perturbation confirms in a cleaner way that over- and de- stabilization of k-fibers is sufficient to erase the asymmetry induced by centrosome age.

6) The authors state that in 2-5% of HeLa cells after monastrol wash-out they observe anaphases with non-disjoined chromosomes that preferentially segregate to the daughter cell with the old centrosome. However, the dominant defect reported by many labs for these types of washouts is lagging chromosomes (i.e. single chromatids) and not non-disjoined chromosomes. Do the authors observe lagging chromosomes and do these segregate preferentially to daughter cells with old centrosomes? Is the asymmetric segregation specific to non-disjoined chromosomes? How do the authors explain why the rate of non-disjoined chromosomes increases under these washout regimes?

The main purpose of a monastrol-washout in the experiments shown in Figure 4 was to obtain a synchronous anaphase population, a fact that we now indicate more explicitly in the manuscript. As reported by many other laboratories after a monastrol washout in our eGFP-centrin1/mCherry-CENP-A cells we often saw single lagging kinetochores, which most likely reflect merotelic chromosomes. However, due to the fact that the mCherry-CENP-A signal is weak and that it tends to fade if one excites the fluorescence too often, we did not have sufficient temporal resolution to “track” the entire history of these single lagging kinetochores. Therefore we could not determine whether these single chromatids were segregated to the correct or incorrect daughter cell. This is now explained in the revised text. We agree that it would be interesting to know whether merotelically attached chromosomes also show a bias in chromosome mis-segregation, but since in most of the cases merotelics segregate normally (Cimini et al. 2004) and since such an investigation requires to know the precise history of those chromosomes, we believe that addressing this question will be challenging, as these are rare events, and that it would go beyond the time frame of this revision.

This is the reason why we focused only on those chromosomes where we were sure that both sister chromatids segregate to the same cell. As non-disjoined sister-chromatids remain attached to each other during anaphase, they move in a synchronous manner and their fate (correct segregation or mis-segregation) is easy to determine. We used monastrol washout as a way to synchronize cells, as such non-disjoined sister-chromatids are rare (2-5% of cells, with a chromosome mis-segregation event in 1-2% of the cells). Given the rarity of the event we have not been able to determine whether monastrol increases the frequency of such non-disjoined chromosomes, but so far we have not obtained any evidence for such an effect.

7) It is not clear that the described phenomenon has a biological function. It might actually be just a consequence of the existing intrinsic centrosome asymmetry that cells have to deal with. This point deserves a better discussion.

We agree, and have extended the Discussion to also mention the possibility that this asymmetry is related to a non-mitotic function.

8) The model figure needs revision. The authors claim that the two spindle halves have different kinetochore microtubule stability due to centrosome age but the diagram only shows the non-disjoined chromosome with different microtubule stability. Do they really mean that the difference in kinetochore microtubule stability is limited to just the lagging chromosomes? For this reason, it might be best to depict the half spindles differently.

We agree, and we have changed the model in Figure 4D to make all the microtubules emanating from the old centrosome more stable.

9) The last paragraph of the Discussion randomly mentions cancer stem cells and chromosomal instability. This does not align with the rest of the text or data presented. There is nothing about cancer stem cells reported in the paper and chromosomal instability results from lagging chromosomes and not non-disjoined chromosomes. They authors need to revise the last paragraph.

First, with regard to the origin of chromosomal instability (CIN), we agree with the reviewers that our last paragraph gave the unnecessary impression that CIN only arises due to non-disjoint chromosomes. However, at the same time we disagree with the reviewers that CIN exclusively results from lagging chromosomes, as non-disjoint chromosomes which can be frequent in cancer cells with replication defects, are clearly also able to lead to gain or loss of chromosomes (as visualized with our own experiments in Figure 4) or see (Aguilera & García-Muse 2013). We therefore have revised our last paragraph to reflect that CIN has multiple origins, including non-disjoint chromosomes and lagging chromosomes.

Second, we also agree that our data are not directly related to cancer stem cells, and that our causal chain of events linking asymmetric gain and loss of chromosomes in human cancer tissues, with inheritance of old centrosomes in (cancer) stem cells, and our observed biased gain of chromosomes in cells inheriting the old centrosomes, is speculative. But we also would argue that the discussion part of a publication should/can also include speculative parts, to stimulate future investigations. We have thus taken this point in consideration and have emphasized the speculative nature of our proposed hypothesis, while believing that such a speculative part is not out of place.

https://doi.org/10.7554/eLife.07909.023

Article and author information

Author details

  1. Ivana Gasic

    1. Department of Cellular Physiology and Metabolism, University of Geneva, Geneva, Switzerland
    2. Institute of Biochemistry, Eidgenössische Technische Hochschule Zürich, Zürich, Switzerland
    Contribution
    IG, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Purnima Nerurkar

    Institute of Biochemistry, Eidgenössische Technische Hochschule Zürich, Zürich, Switzerland
    Contribution
    PN, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  3. Patrick Meraldi

    Department of Cellular Physiology and Metabolism, University of Geneva, Geneva, Switzerland
    Contribution
    PM, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    Patrick.meraldi@unige.ch
    Competing interests
    The authors declare that no competing interests exist.

Funding

Schweizerische Nationalfonds zur Förderung der Wissenschaftlichen Forschung (31003A_141256/1)

  • Patrick Meraldi

Université de Genève

  • Patrick Meraldi

Louis-Jeantet Foundation

  • Patrick Meraldi

Eidgenössische Technische Hochschule Zürich

  • Patrick Meraldi

Boehringer Ingelheim Fonds

  • Ivana Gasic

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

Acknowledgements

We thank the LMC of the ETH Zurich and the Bioimaging Core Facility of the University of Geneva for microscopy support, A Khodjakov for hTertRPE1-EGFP-centrin1 cells (Wadsworth Centre, USA), I. Cheeseman for Dsn1 antibodies (MIT, USA), A McAinsh for HeLa mCherry-CENP-A cells (Univ. of Warwick, UK) and S Doxsey (UMass, USA) for HeLa-eGFP-centrin1 cells. We thank A McAinsh and J. Meadows (both Univ. of Warwick, UK), M Gotta (Univ. of Geneva, Switzerland), H Maiato (IBMC, Portugal), V Panse (ETHZ, Switzerland) and the Meraldi lab members for critical discussions, and CH Tan and F Heydenreich for their contribution to Figure 1B. PM was funded by an SNF-project grant, the ETH Zurich, the University of Geneva and the Louis-Jeantet Foundation, IG by a Böhringer Ingelheim fellowship. IG and PN are part of the MLS PhD School.

The project was initiated and conceived by IG and PM, and directed by PM All experiments and data analysis were carried out by IG PN. helped IG. in the initial analysis of chromosome alignment in human cells. IG and PM interpreted the data and wrote the manuscript.

Reviewing Editor

  1. Anna Akhmanova, Utrecht University, Netherlands

Publication history

  1. Received: April 2, 2015
  2. Accepted: August 18, 2015
  3. Accepted Manuscript published: August 19, 2015 (version 1)
  4. Version of Record published: September 23, 2015 (version 2)

Copyright

© 2015, Gasic et al.

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

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