Mitotic progression, arrest, exit or death is determined by centromere integrity and independent of de novo transcription

Recent studies have challenged the prevailing dogma that transcription is repressed during mitosis. Transcription was also proposed to sustain the spindle assembly checkpoint (SAC) for several hours in response to unattached kinetochores. Here we used live-cell imaging of human cells in culture, combined with RNA-seq and qPCR, to investigate the requirement for de novo transcription during mitosis. Under conditions of persistently unattached kinetochores, transcription inhibition with actinomycin D, or treatment with other DNA-intercalating drugs, delocalized the chromosomal passenger complex (CPC) protein Aurora B from centromeres, compromising SAC robustness and cell fate. However, we were unable to detect significant changes in transcript levels. Moreover, inhibition of transcription independently of DNA intercalation had no effect on SAC response, mitotic progression, exit or death. Mechanistically, we show that DNA intercalating agents reduce the interaction of the CPC with nucleosomes. Thus, the capacity of human cells to progress, sustain, exit or die in mitosis relies on centromere integrity, rather than de novo transcription.


Introduction
When cells commit to mitosis the nuclear envelope disassembles and chromatin organizes into highly condensed chromosomes. This causes the displacement of several transcription factors from DNA and the inactivation of the transcription machinery in a cell cycle-dependent manner (Gottesfeld and Forbes, 1997;Kim et al., 1997;Martínez-Balbás et al., 1995;Parsons and Spencer, 1997;Segil et al., 1996;Shermoen and O'Farrell, 1991;Spencer et al., 2000. Because the halflife of most transcripts exceeds the normal duration of mitosis (typically 30 min in human cells), the prevailing dogma was that transcription is largely repressed during mitosis in higher eukaryotes (Gottesfeld and Forbes, 1997;Konrad, 1963;Martínez-Balbás et al., 1995;Murphy and Attardi, 1973;Prescott and Bender, 1962;Rizkallah and Hurt, 2009;Spencer et al., 2000;Taylor, 1960). However, several recent studies have challenged this notion and provided evidence that the level of chromatin compaction on mitotic chromosomes is highly heterogeneous (Nishino et al., 2012), allowing accessibility of some transcription factors and chromatin-modifying enzymes (Burke et al., 2005;Chen et al., 2005;Dey et al., 2009;Egli et al., 2008;Gauthier-Rouviere et al., 1991;Michelotti et al., 1997;Segil et al., 1996;Yan et al., 2006). Noteworthy, a post-translationally modified form of RNA Polymerase II, which is normally associated with transcription elongation, was found to bind mitotic chromosomes at their centromeres (Chan et al., 2012;Dirks and Snaar, 1999;Liu et al., 2015;Molina et al., 2016), suggesting that some transcription is still taking place during mitosis.
Transcription might also play an instrumental role during a prolonged mitosis due to incapacity to satisfy the spindle assembly checkpoint (SAC), a signalling mechanism that can delay mitosis up to several hours in the presence of unattached kinetochores (Musacchio, 2015). In line with this hypothesis, the master mitotic regulator Cyclin B1 was proposed to be actively transcribed during mitosis and to be required to sustain a robust SAC response (Mena et al., 2010;Sciortino et al., 2001). Centromere assembly, which underlies the catalytic mechanism behind the SAC, has also been recently proposed to depend on transcription of centromeric α-satellite DNA and to be regulated by non-coding RNAs (Blower, 2016;Carone et al., 2009;Chan et al., 2012;Du et al., 2010;Grenfell et al., 2016;Li et al., 2008;Liu et al., 2015;Nakano et al., 2003;Pezer and Ugarković, 2008;Rošić et al., 2014;Topp et al., 2004;Wong et al., 2007;Zhang et al., 2005). In particular, the centromeric localization and activation of Aurora B, the catalytic subunit of the chromosomal passenger complex (CPC) required for SAC response and error correction during mitosis (Carmena et al., 2012), was shown to be dependent on centromeric transcription (Blower, 2016;Grenfell et al., 2016). However, whether centromeric transcription persists during mitosis remains controversial. Finally, a recent study using pulse-labelling of nascent transcripts and RNA-seq of mitotic-enriched cell populations reported that over 8000 transcripts are expressed de novo during mitosis and mitotic exit to prepare cells for the subsequent interphase (Palozola et al., 2017;Strzyz, 2017;Timmers and Verrijzer, 2017).
Although several pieces of evidence indicate that some level of transcription might still take place during mitosis, most of these conclusions were drawn based on the analysis of synchronized, yet impure, mitotic cell populations, fixed material and a diverse range of transcription inhibitors with imprecise temporal control of transcriptional repression. Moreover, it remains unclear whether transcription is required for normal mitotic progression and exit. For these reasons, we sought to investigate the requirement of de novo transcription during mitosis using direct live-cell imaging and a wide range of transcription inhibitors. We found that the capacity of human cells to progress, sustain, exit or die in mitosis is independent of de novo transcription. In agreement, quantitative RNA-seq and qPCR analysis failed to unveil potential transcripts produced de novo during mitosis. Finally, we uncovered that DNA-intercalating agents, which include well-established transcription inhibitors, such as actinomycin D, reduce the interaction of the CPC with nucleosomes and significantly compromise centromere integrity and function in a transcription-independent manner. Our findings raise awareness about the use of different transcription inhibitors in living cells and shed light on a long-lasting controversy.

Actinomycin D compromises spindle assembly checkpoint robustness in a transcription-independent manner
We reasoned that if de novo transcription takes place during a normal mitosis, its role would be exacerbated during a prolonged mitosis. To test this, we used livecell imaging to quantify the duration of mitosis in human HeLa cells treated with the microtubule depolymerizing drug nocodazole, which generates persistently unattached kinetochores that prevent SAC satisfaction. After 3 h in nocodazole, we directly monitored cells that became committed to mitosis, as determined by typical cell rounding, and inhibited transcription with the well-established DNAintercalating drug actinomycin D (Fig. 1A). Consistent with previous observations in fixed cells (Becker et al., 2010), we found that live cells treated with nocodazole remained in mitosis for 19.2±7.0 h (median ± SD, n=386 cells), whereas cells treated with nocodazole followed by actinomycin D persisted in mitosis for only 16.1±5.6 h (n=301 cells, p<0.0001, Mann-Whitney Rank Sum Test, Fig. 1B,C).
Thus, treatment with actinomycin D compromises the capacity of cells to sustain a prolonged mitotic delay and interferes with their fate.
The relationship between the capacity to sustain a robust SAC response and the subsequent cell fate after a mitotic delay has been shown to depend on the kinetics of anaphase-promoting complex (APC)-mediated Cyclin B1 degradation, with slower degradation promoting mitotic cell death, and faster degradation promoting mitotic slippage (Brito and Rieder, 2006;Gascoigne and Taylor, 2008). We therefore tested whether inhibition of Cyclin B1 proteolysis with the proteasome inhibitor MG132 could rescue the faster exit and different fate of cells treated with nocodazole and actinomycin D. As controls, we modulated mitotic duration either by inhibition of the SAC kinase Mps1 (to accelerate mitotic exit), or inhibition of the proteasome with MG132 (to delay mitotic exit) after nocodazole treatment. Accordingly, cells treated with nocodazole and MG132 increased the mitotic delay relative to cells treated only with nocodazole (from 19.2±7.0 h to 30.1±14.7 h, p<0.0001, Mann-Whitney Rank Sum Test, Fig. 1C), and caused cell death in mitosis in 100% of the cases (Fig. 1D). Conversely, acute SAC inactivation through Mps1 inhibition after nocodazole treatment significantly decreased the mitotic delay relative to nocodazole-treated cells (from 19.2±7.0 h to 5.4±2.6 h, p<0.0001, Mann-Whitney Rank Sum Test, Fig. 1C), and caused almost immediate mitotic slippage in 100% of the cells (Fig. 1D). Proteasome inhibition with MG132 reverted the Mps1 inhibition phenotype, increasing mitotic duration and switching cell fate from mitotic slippage to death (p<0.0001, Mann-Whitney Rank Sum Test, Fig. 1C,D). Most important, the ability of actinomycin D to reduce the mitotic delay and induce slippage in cells treated with nocodazole was rescued after inhibition of proteasome activity with MG132 (p 0.01, Mann-Whitney Rank Sum Test, Fig. 1C,D). Overall, these experiments suggest that actinomycin D compromises SAC response and the capacity to sustain a prolonged mitotic delay, in a proteasome-dependent manner.
The conclusions above are at odds with models in which SAC response depends on de novo transcription of Cyclin B1 during mitosis (Mena et al., 2010;Sciortino et al., 2001). To clarify this issue we used quantitative time-lapse fluorescence microscopy to monitor the levels and respective kinetics of degradation of Cyclin B1 tagged at its endogenous genomic locus with Venus (Collin et al., 2013) (Fig. 1E). In agreement with previous reports, Cyclin B1-Venus fluorescence decayed very slowly in the presence of nocodazole, likely due to residual APC activity (Fig. 1F) (Brito and Rieder, 2006). Treatment of nocodazole-arrested cells with actinomycin D caused a faster decline in Cyclin B1-Venus (p<0.0001, Analysis of Covariance (ANCOVA), Fig.1F). Interestingly, Cyclin B1-Venus levels in nocodazole-treated cells remained constant for several hours after inhibition of the proteasome with MG132, regardless of the presence of actinomycin D (p 0.05, ANCOVA, Fig.1F). We conclude that actinomycin D compromises SAC robustness and the capacity to prevent APC-mediated Cyclin B1 degradation, independently of de novo transcription.

Actinomycin D compromises the localization of Aurora B and Mad1 at centromeres/kinetochores
To determine how actinomycin D compromises SAC robustness during a prolonged mitosis, we investigated the localization of the SAC-related proteins Aurora B (at centromeres) and Mad1 (at unattached kinetochores). Accordingly, we added actinomycin D for 8 h in cells pre-treated with nocodazole, which was kept throughout the entire duration of the experiment. To assist in the identification of mitotic cells, while assessing Aurora B activity on chromosomes, we also investigated the phosphorylation of Histone H3 at Serine 10 [pH3S10; (Crosio et al., 2002)]. In contrast with nocodazole-treatment in which all cells showed high levels of pH3S10, addition of actinomycin D to nocodazole-treated cells reduced or abolished pH3S10 accumulation in approximately 20% of the cells ( Fig. 2A,B). Importantly, these cells still showed signs of chromosome condensation (as determined by DAPI staining of DNA) that were indistinguishable from neighbouring cells with normal levels of pH3S10 ( Fig.2A).
Moreover, the observed reduction/abolishment of pH3S10 was not associated with a pre-apoptotic state, as those cells were negative for a cleaved form of caspase 3 ( Fig. 2A). In line with the observed reduction/abolishment of pH3S10, these cells also showed abnormally low levels of total Aurora B and its active phosphorylated form on Threonine 232 (Yasui et al., 2004). Noteworthy, even in those nocodazole-and actinomycin D-treated cells that showed high levels of pH3S10, Aurora B, including its active form, was found displaced along the chromosome arms ( Fig.2A). Thus, actinomycin D compromises normal Aurora B localization and activity.
In order to get spatiotemporal insight into the effect of actinomycin D on Aurora B localization we monitored GFP-Aurora B by quantitative fluorescence live-cell microscopy in nocodazole-treated cells. Additionally, we further tested whether Aurora B localization was dependent on mitotic translation and on its own kinase activity by inhibiting protein synthesis with cycloheximide and treating cells with the Aurora B inhibitor ZM447439, respectively. We found that while Aurora B centromeric levels remained constant over a 5 h period in cells treated either with nocodazole alone or nocodazole and cycloheximide, actinomycin D caused the displacement of Aurora B from centromeres onto chromosome arms as soon as 2 h after addition of actinomycin D, leading to a 30% reduction of centromeric fluorescence after 4 h (p<0.0001, ANCOVA, Fig. S1A,B). Curiously, Aurora B inhibition led to an enrichment of its own levels at centromeres, without any detectable displacement (Fig. S1A,B). These observations were confirmed by quantitative fluorescence microscopy analysis in fixed cells (Fig. 2C,D and S2A-C). These data directly demonstrate that actinomycin D displaces Aurora B from centromeres.
Finally, we tested whether actinomycin D treatment compromised the localization of Mad1 at unattached kinetochores. We found that addition of actinomycin D for 4 h to nocodazole-treated cells led to a 45% reduction of Mad1 at the outer kinetochore (Fig. 2C,E). Similar findings have previously been reported for the SAC proteins Bub1 and BubR1 (Becker et al., 2010). Taken together, these experiments show that actinomycin D compromises the normal localization (and activity) of Aurora B at centromeres and SAC proteins to unattached kinetochores. They also offer a plausible explanation for how SAC robustness might be affected by actinomycin D treatment.

Actinomycin D does not affect total CPC protein levels nor the levels of mitotic transcripts
Western blot analysis revealed that Aurora B, as well as the total protein levels of the CPC regulatory subunits Survivin and Borealin remained unchanged relative to controls (including inhibition of translation during mitosis with cycloheximide or Aurora B inhibition with ZM447439) after 4 h treatment with actinomycin D (Fig.   3A,B). To test whether Aurora B was regulated by an effect of actinomycin D over mitotic transcription, we performed a comparative genome-wide RNA-seq analysis between cells treated either with nocodazole alone or nocodazole and actinomycin D. Importantly, only mitotic cells were analysed, as they were obtained by shake-off after nocodazole treatment and subsequently transferred to new culture flasks before addition of actinomycin D (Fig. 3A). Our quantitative transcriptome analysis did not reveal any significant changes either in annotated or novel assembled transcripts (including all biotypes) upon actinomycin D treatment, ruling out differential gene expression after transcription inhibition during mitosis. Importantly, we confirmed that Cyclin B1, as well as all key mitotic regulators, including Aurora B, were not de novo transcribed even during a prolonged mitosis ( Fig. 3C and Table 1).
Due to the low mappability of the centromere, RNA-seq reads could not be properly aligned to these regions. Consequently, the role of centromeric transcription, namely of regulatory non-coding RNAs, could not be properly assessed in the previous experiment. Nevertheless, we were able to directly measure by qPCR the expression of centromeric α-satellite RNA relative to a control house-keeping gene (GAPDH) in cells that had been arrested in mitosis with nocodazole, in the presence or absence of actinomycin D. We found that actinomycin D treatment did not reduce the level of centromeric α-satellite RNA Therefore, we reasoned that actinomycin D could displace Aurora B from centromeres, not by inhibiting mitotic transcription, but through its ability to intercalate with DNA. In line with this hypothesis, it was previously shown that actinomycin D, but not α-amanitin [a potent and selective inhibitor of RNA polymerase II that triggers the degradation of the largest RNA polymerase II subunit (Nguyen et al., 1996)], interferes with the normal centromeric localization of Aurora B (Becker et al., 2010). However, α-amanitin is a slow-uptake drug and inhibition of transcription was not confirmed in these studies. For this reason, we first compared the effect of α-amanitin with that of actinomycin D or triptolide, a fast and irreversible transcription inhibitor that prevents the formation of the "transcription bubble" without intercalating with DNA (Titov et al., 2011;Vispé et al., 2009). To determine the extent of transcription inhibition by these drugs, we used qPCR to amplify Mcl1, a short half-life transcript isolated from asynchronous cells after 4 h treatment either with α-amanitin, actinomycin D or triptolide. As suspected, we could not detect any significant effect of α-amanitin over Mcl1 transcript levels relative to controls (Fig. 4A). In sharp contrast, treatment with actinomycin D or triptolide resulted in a 5-fold decrease in Mcl1 transcript levels ( Fig. 4A). Finally, we confirmed that, within the time course of our experiment, triptolide, but not α-amanitin, caused the degradation of the large subunit of RNA Polymerase II (Fig. S3A). Thus, triptolide, but not α-amanitin, efficiently inhibits transcription in living cells and it does so independently of DNA intercalation.
Next, we investigated whether transcription inhibition with triptolide compromises SAC robustness in cells treated with nocodazole. We found no significant differences in mitotic duration between nocodazole-treated cells with or without triptolide (19.3±7.1 h vs 19.5±5.8 h, respectively, median ± SD, n=301 cells, p=0.65, Mann-Whitney Rank Sum Test, Fig.4B). Moreover, contrary to what was previously observed in the case of actinomycin D, we did not detect any effect of triptolide over cell fate after the mitotic delay imposed by nocodazole treatment and most cells died in mitosis (Fig. 4C). Triptolide also did not cause the displacement of Aurora B from centromeres in both fixed and live-cell assays ( Fig. 4D-G). These results, supported by our RNA-seq data, indicate that SAC robustness, Aurora B centromeric localization, as well as the capacity to die in mitosis after a prolonged mitotic delay is independent of de novo transcription.

DNA intercalation interferes with the interaction between the CPC and nucleosomes
Our previous results suggest that DNA intercalation, rather than transcriptional inhibition, causes the displacement of Aurora B from centromeres. To test this, we took advantage from another bona fide DNA intercalating compound, ethidium bromide, which has not been implicated in transcription inhibition. Indeed, we found that Mcl1 transcript levels were not significantly affected within 4 h of ethidium bromide treatment ( Fig 5A). Cells treated with nocodazole and ethidium bromide spent slightly less time in mitosis when compared to cells treated with nocodazole only, but the difference was not statistically relevant (19.2±7.0 h vs 18.1±5.9 h, median ± SD, n=386;151 cells/condition, p=0.13, Mann-Whitney Rank Sum Test, Fig.5B). However, similar to actinomycin D, ethidium bromide caused a significant effect over cell fate after the mitotic delay imposed by nocodazole, causing mitotic slippage in approximately 50% of the cells (Fig. 5C).
Ethidium bromide also displaced Aurora B from centromeres in fixed cells, although to a less extent when compared with actinomycin D (Fig. 5D,E). These data suggest that DNA intercalating drugs interfere with normal Aurora B localization at centromeres and compromise cell fate after a prolonged mitosis.
To directly test whether DNA intercalating drugs interfere with the capacity of Aurora B to associate with centromeric chromatin, we probed the ability of the The capacity to satisfy the spindle assembly checkpoint and resume mitosis and cytokinesis after a prolonged mitotic delay does not require de novo transcription Lastly, we tested whether the capacity to progress and exit mitosis requires de novo mitotic transcription. To assure an efficient transcriptional repression during mitosis, we performed a nocodazole washout experiment. Accordingly, HeLa cells were incubated for 5 h with nocodazole and the transcription inhibitors (actinomycin D or triptolide) were added in the last 2 h of the mitotic blockage.
Nocodazole was subsequently washed out and the cells were allowed to progress through mitosis only in the presence of the transcription inhibitors (Fig. 6A). Livecell imaging was used to directly determine the duration of mitosis, while assessing cell fate. We found that control and triptolide-treated cells took a similar time to exit mitosis after nocodazole washout (79.1 ± 28.7 min vs 86.2 ± 31.7 min, median ± SD, n = 116;113 cells/condition, p=0.11, Mann-Whitney Rank Sum Test,

Discussion
Whether transcription is permissive and required during mitosis is a recurrent debate in the field. The resolution of this long-lasting controversy has been hampered by the difficulty to efficiently and specifically repress transcription during mitosis, which often takes only 30 min in human cells. It has however been suggested that the master mitotic regulator Cyclin B1 is actively transcribed during mitosis (Sciortino et al., 2001) and de novo transcription is required to sustain a mitotic delay under conditions that prevent SAC satisfaction for several hours (Mena et al., 2010). Centromere/kinetochore assembly was also linked to active transcription (Carone et al., 2009;Chan et al., 2012;Du et al., 2010;Grenfell et al., 2016;Li et al., 2008;Liu et al., 2015;Nakano et al., 2003;Pezer and Ugarković, 2008;Rošić et al., 2014;Topp et al., 2004;Wong et al., 2007;Zhang et al., 2005), including the localization and activity of Aurora B, as shown in Xenopus egg extracts (Blower, 2016;Grenfell et al., 2016). More recently, an entire transcription program was proposed to remain constitutively active during mitosis and mitotic exit in human cells (Palozola et al., 2017;Strzyz, 2017; Timmers and Verrijzer, 2017), but whether this potential program is required for mitotic progression and exit was not elucidated. By combining direct live-cell imaging, while monitoring the efficiency and specificity of transcription inhibition at the whole genome level, we show that the capacity of human cells to sustain, progress, exit or die in mitosis does not require de novo transcription. Moreover, we demonstrate that commonly used transcription inhibitors, such as actinomycin D and α-amanitin show serious limitations for their use in live-cell experiments that aim to understand mitosis. Actinomycin D and other DNA intercalating agents caused partial dissociation of the CPC from nucleosomes, thereby compromising Aurora B centromeric localization and SAC response. On the other hand, αamanitin, as a slow uptake drug, failed to efficiently inhibit transcription even after several hours in mitosis. Most importantly, efficient inhibition of mitotic transcription independently of DNA intercalation using triptolide had no discernible effect over Aurora B centromeric localization or SAC response. We conclude that centromere integrity, not mitotic transcription, is required for proper localization and activity of Aurora B and to mount a robust SAC able to sustain mitosis in human cells for several hours in the event of unattached kinetochores.
In light of a recent study reporting the identification of over 900 nascent transcripts from allegedly metaphase cells (Palozola et al., 2017) [see also comments by (Strzyz, 2017;Timmers and Verrijzer, 2017)], it was surprising that our genome wide RNA-seq or qPCR analyses failed to reveal any significant alteration in gene expression, including Cyclin B1 and other mitotic genes, during a prolonged mitosis after transcription inhibition with actinomycin D. However, in the previous study, nocodazole-treated cells were only 95% pure and transcripts isolated 40 min after nocodazole washout might have derived from cells that had already exited mitosis and reached early G1 stage where transcription is expected to be permissive (Hsiung et al., 2016). Thus, the use of synchronized, yet impure, cell populations, as opposed to monitoring transcription inhibition only after cells commit to mitosis by direct live-cell imaging, might account for the differences observed between studies. Nevertheless, it remains possible that, as cells progress through mitosis and enter anaphase, de novo transcription starts to be permissive, despite its dispensability for the completion of and exit from mitosis.
Consistent with a global shut-down of transcription during prometaphase (the physiological equivalent of nocodazole-arrested cells), RNA Polymerase II is generally found displaced from chromatin (Hsiung et al., 2016), with the notorious exception of the centromeric region (Chan et al., 2012). Because recent studies have suggested that non-coding RNAs associate with centromere and kinetochore proteins, including CENP-A, CENP-C and Aurora B (Blower, 2016;Carone et al., 2009;Du et al., 2010;Ferri et al., 2009;Molina et al., 2017;Molina et al., 2016;Rošić et al., 2014;Wong et al., 2007), it remains possible that transcription of non-coding RNAs plays a role in centromere assembly and function. However, whether this occurs during mitosis or during G1, when centromere assembly takes place in mammalian cells (Jansen et al., 2007) is unclear. Centromeric transcription is involved in nucleosome disassembly during interphase (Boeger et al., 2003) to facilitate the replacement of histone H3 by CENP-A (McKittrick et al., 2004;Tagami et al., 2004). Intriguingly, the observation that the elongating form of RNA Polymerase II is present at centromeres during mitosis (Chan et al., 2012) suggest that some level of local transcription of noncoding RNAs might still take place. However, it should be noted that a recent study in S. pombe indicated that RNA Polymerase II stalls at centromeric DNA and the level of stalling is directly proportional to the level of cnp1p (centromerespecific histone H3) nucleosome assembly (Catania et al., 2015).
While we could not experimentally exclude that non-coding RNAs are being transcribed from the centromere during a prolonged mitosis, we failed to detect any significant change in the total levels of α-satellite RNA after transcription inhibition, in agreement with previous reports (Liu et al., 2015).
Indeed, experimental increase of α-satellite RNA levels reduced the binding of Sgo1 to nucleosomes (Liu et al., 2015) and led to problems in chromosome segregation (Chan et al., 2017;Zhu et al., 2011), suggesting that the levels of αsatellite RNA must be tightly regulated to ensure a faithful mitosis.
Overall, our results demonstrate that, regardless of our capacity to detect residual transcription that might still take place during mitosis, this is not required for human cells to sustain, progress, exit or die in mitosis.

Fluorescence quantification
For quantitative measurements, all images compared were acquired using identical acquisition settings. Image analysis was performed using ImageJ. Briefly, individual kinetochores were identified by ACA staining and marked by a region of interest (ROI). The average fluorescence intensity of signals (pixel gray levels) of pAurora B at the inner centromere and Mad1 at kinetochores were measured on the focused z plan, and the background fluorescence was measured outside the ROI and subtracted. Fluorescence were used as primary antibodies, and anti-rabbit and anti-mouse antibodies were used as secondary antibodies (1:5,000; Jackson ImmunoResearch Laboratories, Inc.).

RNA extraction and transcription analysis by RT-qPCR
Total RNA was isolated from interphase and mitotic HeLa cells (shake-off) using trizol (Life technologies). Reverse transcriptase reaction was performed with 500 ng of total RNA with iscript (Bio-rad) using the random primers and oligo(dTs) supplied, following the manufacturer's instructions. All primer sequences are in Supplemental Material. For each analysis, gapdh was used for normalization. RT-qPCRs were performed in the iCycler iQ5 Real-Time PCR Detection System (Bio-Rad Laboratories). The data obtained were analyzed using the Bio-Rad iQ5 Optical System Software v2.1 (BioRad Laboratories). The amplified products were additionally analyzed on agarose gel electrophoresis.

RNA library preparation for RNAome sequencing
Quantity and total RNA integrity was checked following isolation using an Agilent Technologies 2100 Bioanalyzer. Sample with RNA Integrity Number (RIN) value greater than 8 were selected. The RNA samples were prepared as described before in (Derks et al., 2015) with the following modifications. Five micrograms of total RNA were depleted according to the Illumina Ribo-zero magnetic protocol (www.illumina.com). The Ribo-zero treated RNA was purified using Agencourt RNAClean XP Beads. RNA was fragmented on a Covaris S220. One microliter of the sheared rRNA depleted RNA was loaded on an Agilent Technologies 21000 Bioanalyzer RNA Pico chip to determine successful ribosomal RNA depletion. The sheared RNA sample was concentrated in a Thermo scientific SPD100 speedvac to 5 µl and a sequencing library was prepared according to the Illumina TruSeq Small RNA protocol (www.illumina.com). In short, adapters are subsequently ligated to the 3'end and the 5'end of the RNA. The RNA is reverse transcribed, amplified by PCR and run on SDS-PAGE gel. RNA fragments between 140 -500 bp are cut out of the gel and purified. One microliter was loaded on an Agilent Technologies 2100 Bioanalyzer using a DNA 1000 assay to determine the library concentration and quality.

Bridge amplification and Sequencing by synthesis
Cluster generation was performed according to the Illumina TruSeq SR Cluster kit v2 (cBot) Reagents Preparation Guide (www.illumina.com). Briefly, six RNAome libraries were pooled together to get a stock of 2 nM. Five microliter of the 2 nM stock was denaturated with NaOH, diluted to 11 pM and hybridized onto the flowcell. The hybridized products were sequentially amplified, linearized and end-blocked according to the Illumina Single Read Multiplex Sequencing user guide. After hybridization of the sequencing primer, sequencing-by-synthesis was performed using the HiSeq 2000 with a 36-cycle protocol. The sequenced fragments were denaturated with NaOH using the HiSeq 2000 and the indexprimer was hybridized onto the fragments. The index was sequenced with a 6cycle protocol.

Bioinformatics Analysis
Raw FASTQ files were aligned to the human genome (NCBI/build37.1) and annotated using Gencode version 25. De novo transcriptome assembly was performed using Gencode annotation and the StringTie assembler (Pertea et al., 2015). Mapping of sequence reads was performed using STAR aligner and transcript detection and quantification was performed using Cufflinks.
Gene/transcript levels were compared between control and actinomycin Dtreated samples using the ballgown program (Frazee et al., 2014).

CPC-CTM expression and purification
Full length (f.l.) Survivin was cloned as a 3C-cleavable His-GFP tagged protein in a pRSET vector (Thermo Fisher Scientific), f.l. Borealin was cloned as a TEVcleavable His-tagged protein in a pETM vector (gift from C. Romier, IGBMC, Strasbourg), and INCENP1-58 was cloned as an untagged protein in a pMCNcs vector. The complex of f.l. Survivin, f.l. Borealin and INCENP1-58 was prepared by co-expressing the subunits in E. Coli strain BL21 pLysS with an overnight induction at 18 o C. Cells were lysed in lysis buffer containing 25 mM Hepes pH 7.5, 500 mM NaCl, 25 mM Imidazole, 2 mM -mercaptoethanol (Bme). The complex was purified by affinity chromatography using a HisTrap HP column (GE Healthcare). The protein-bound column was washed with lysis buffer, followed by 25 mM Hepes pH 7.5, 1 M NaCl, 50 mM KCl, 10 mM MgCl, 25 mM Imidazole, 2 mM ATP, 2 mM Bme and proteins were eluted using 25 mM Hepes pH 7.5, 500 mM NaCl, 500 mM Imidazole, 2 mM Bme. Tags were cleaved during an overnight incubation with 3C and TEV while dialysing against 25 mM Hepes pH 7.5, 150 mM NaCl, 4 mM dithiothreitol (DTT) at 4 o C. The complex was further purified by a cation exchange chromatography (HiTrap SP, GE Healthcare) followed by gel filtration using a Superdex 200 increase 10/300 column (GE Healthcare) equilibrated with 25 mM Hepes pH 8, 200 mM NaCl, 4 mM DTT.

Expression and purification of recombinant histones and refolding of histone octamers
Human H2A and H2B and Xenopus laevis H3 and H4 were purified as described before (Luger et al., 1999, Methods Mol Biol) with minor changes. LB media was used instead of 2X TY-AC media for expression of H2A, H2B and H3 in E. coli BL21 (DE3) pLysS cells and H4 in E. coli BL21 cells. The preparation of inclusion bodies was performed using a Dounce glass/glass homogenizer. After solubilisation of the inclusion bodies and the three-step dialysis against Urea dialysis buffer (7 M Urea, 100 mM NaCl, 10 mM Tris pH 8, 1 mM EDTA, 5 mM Bme), the sample was applied to a HiTrap Q anion exchange column and then to a HiTrap SP cation exchange column (GE Healthcare). The histones were eluted from the HiTrap SP column using a linear gradient from 100 mM to 1 M NaCl in 7 M Urea, 10 mM Tris pH 8, 1 mM EDTA and 1 mM DTT. After the ion exchange step, purified recombinant histones were dialysed against water with 5 mM Bme, lyophilized and stored at -80 o C. Refolding of histone octamers was performed as previously described (Luger et al., 1999, Methods Mol Biol). Briefly, lyophilized core histones were resuspended in unfolding buffer (7M Guanidine HCl, 20 mM Tris pH 7.5, 10 mM DTT) and mixed to equimolar ratios. The histone mix was then dialysed against three changes of 500ml refolding buffer (10 mM Tris pH 8, 2M NaCl, 1 mM EDTA, 5 mM Bme) and the octamers were purified by size exclusion chromatography using a Superdex 200 increase 10/300 column (GE Healthcare) equilibrated with refolding buffer. Pooled fractions with equimolar ratios of histones were stored at -80 o C.

Nucleosome core particle reconstitution and drug treatment
The 147bp 601 Widom positioning sequence was amplified from a pBS-601Widom vector with 5' IR700 labelled primers. Mononucleosome reconstitution was carried out using the salt gradient dialysis method (Luger et al., 1999, Methods Mol Biol). Histone octamers were added to DNA to a 0.9 molar ratio of octamer to DNA after adjusting the salt concentration to 2 M NaCl. The mixture was then dialysed against TE buffer (10 mM Tris pH 8, 1 mM EDTA, 50 mM NaCl) by gradually decreasing the ionic strength from 2 M to 50 mM NaCl over a period of 20 h using a peristaltic pump. Reconstituted NCPs were treated with DMSO, 80 µm Actinomycin D, 10 µm Triptolide or 25 µm Ethidium Bromide for 1 h at 4 o C before incubating them with the CPC-CTM recombinant protein.

Electrophoretic Mobility Shift Assay
Electrophoretic mobility assay (EMSA) was used to detect specific interaction between CPC and the nucleosome core particles (NCPs). 25 and 50 nM recombinant CPC-CTM was added to 5 nM IR700-labelled NCPs in reaction buffer (25 mM Hepes pH 7.5, 100 mM NaCl, 1 mM DTT, 10% Glycerol). Reactions were incubated 30 min at 4 o C and resolved in a 0.5 % agarose gel in 1X Tris-CAPS buffer (60 mM Tris, 40 mM CAPS pH 9.3). The fluorescent bound and unbound NCPs were detected with Odyssey CLx Infrared Imaging System (LI-COR Biosciences). The ratio of CPC-bound NCPs to free NCPs was quantified using the Image Studio software (LI-COR Biosciences). Signal at the 0 nM CPC lane was subtracted from all values to account for any effect on NCPs destabilization upon drug treatment (EtBr-treated nucleosomes showed a faint smear above the unbound-NCP band indicating a slight NCP destabilization upon EtBr treatment).

Statistical analysis.
To confirm that sampling is sufficient, we used two statistical tests: First, we performed a proportion test or unpaired Student's t-test to confirm using P-values that we have sampled sufficient number of cells or kinetochores for concluding on differences we report. Second, we measured standard error over mean (SEM) across experimental repeats or across cells and confirmed that the differences we report are not only based on differences in mean values, but also the spread