Tetraploidy can constitute a metastable intermediate between normal diploidy and oncogenic aneuploidy. Here, we show that the absence of p53 is not only permissive for the survival but also for multipolar asymmetric divisions of tetraploid cells, which lead to the generation of aneuploid cells with a near‐to‐diploid chromosome content. Multipolar mitoses (which reduce the tetraploid genome to a sub‐tetraploid state) are more frequent when p53 is downregulated and the product of the Mos oncogene is upregulated. Mos inhibits the coalescence of supernumerary centrosomes that allow for normal bipolar mitoses of tetraploid cells. In the absence of p53, Mos knockdown prevents multipolar mitoses and exerts genome‐stabilizing effects. These results elucidate the mechanisms through which asymmetric cell division drives chromosomal instability in tetraploid cells.
Tetraploid cells are detected in some precancerous lesions such as Barrett's oesophagus and cervical dysplasia, where their presence coexists with the loss of functional p53 (Heselmeyer et al, 1996; Maley et al, 2004). Owing to the increase in the number of chromosomes, perhaps coupled to changes in the geometry of the mitotic machinery (Storchova et al, 2006; Storchova and Kuffer, 2008), tetraploid cells frequently activate the DNA damage response and become genomically unstable. Thus, tetraploidy may be considered as a metastable state that links normal diploidy to cancer‐associated aneuploidy (Storchova and Pellman, 2004; Fujiwara et al, 2005; Margolis, 2005).
Numerous tumour suppressor genes including p53 (Margolis, 2005), BRCA1 (Schlegel et al, 2003), LATS2 (Aylon et al, 2006) and APC (Tighe et al, 2004) actively repress tetraploidy, meaning that their removal can either stimulate the spontaneous tetraploidization of cells or facilitate the survival of tetraploid cells generated upon cytokinesis or karyokinesis inhibition. The former has been shown for the knockdown of APC in cultured cells and for the conditional knockout of APC in small intestine epithelia in vivo (Caldwell et al, 2007). The latter has been demonstrated in cultured cancer cell lines that were depleted from p53 by gene knockout or RNA interference (Cross et al, 1995; Andreassen et al, 2001; Castedo et al, 2006a, 2006b), as well as in p53−/− primary mouse mammary epithelial cells (Fujiwara et al, 2005; Senovilla et al, 2009). Moreover, the expression of some oncogenes including Myc (Yin et al, 1999), Aurora‐A (Wang et al, 2006) and human papillomavirus (HPV)‐encoded E6 (Incassati et al, 2006) can stimulate tetraploidization.
The mechanisms through which tetraploidy favours oncogenesis are complex and have not yet been entirely elucidated. One single tetraploid cell can undergo multipolar mitosis, which often leads to the generation of three or more daughter cells (Storchova and Pellman, 2004). This process causes the near‐to‐stochastic distribution of chromosomes and hence is lethal for most daughter cells. Nullisomy (the total absence of one particular chromosome) and polysomy (the presence of extra copies of one chromosome), indeed, result in major genetic defects involving the incorrect assembly of multiprotein complexes and fatal linkage disequilibria, which are rarely compatible with cell survival (Zhivotovsky and Kroemer, 2004; Roumier et al, 2005; Ganem et al, 2007). Moreover, during mitosis, the presence of more than two centrosomes in tetraploid cells can also favour merotelic chromosome attachments and hence chromosomal lagging, which may favour chromosome loss or asymmetric distribution among daughter cells, even when the division is bipolar (Ganem et al, 2009).
Multipolar and asymmetric cell division, as they can result from tetraploidy (Levine et al, 1991; Ganem et al, 2007), are commonly observed in malignant lesions and have been suspected to contribute to oncogenesis for over a century (Boveri, 2008). Supernumerary centrosomes, as they are detected in malignant cells (Levine et al, 1991; D'Assoro et al, 2002), can be induced experimentally, and this reportedly suffices to trigger oncogenesis (Basto et al, 2008; Gergely and Basto, 2008). Moreover, ‘anisocytosis’ and ‘anisokaryosis’ (heterogeneity in cell size and nuclear size, respectively), which presumably result from asymmetric divisions, are well‐established histological hallmarks of malignancy (Boveri, 2008; Holland and Cleveland, 2009). It is interesting to note that in some cancer types (e.g., non‐small cell lung cancer), these morphological traits of malignancy correlate with the expression of one particular oncogene, Mos (Gorgoulis et al, 2001).
Mos (also called c‐Mos) is the first human oncogene cloned, and has been identified as the cellular homologue of the viral oncogene v‐Mos, which is encoded by the Moloney murine sarcoma virus (Oskarsson et al, 1980; Watson et al, 1982). The p39Mos protein (hereafter referred to as Mos) has been shown to stimulate the transformation of murine fibroblasts in vitro (Okazaki and Sagata, 1995; Fukasawa and Vande Woude, 1997). Moreover, transfection‐enforced overexpression of Mos reportedly inhibits mitotic progression (Wang et al, 1994) and causes the generation of binucleated cells due to the inhibition of cytokinesis (Okazaki et al, 1992; Fukasawa and Vande Woude, 1995). Apparently, Mos can stabilize another oncogene product, c‐Fos, (Okazaki and Sagata, 1995) and enhance the expression of cyclins (Rhodes et al, 1997), thereby stimulating cell proliferation. The genetic invalidation of Mos has no obvious phenotypic consequences in mice (Colledge et al, 1994; Hashimoto et al, 1994). However, although Mos−/− male mice exhibit normal reproduction rates, Mos−/− female are nearly infertile (Colledge et al, 1994; Hashimoto et al, 1994), in line with the fact that Mos is strictly necessary for the first meiotic division of oocytes (Sagata et al, 1989a), and then exerts a critical checkpoint function during metaphase II (Sagata et al, 1989b). Both the meiosis‐regulatory and the transforming effects of Mos require its serine–threonine kinase activity (Haccard et al, 1993; Okazaki and Sagata, 1995). Known Mos substrates include cyclin B2, tubulin and MEK1 (Roy et al, 1990; Zhou et al, 1991; Sagata, 1997), and the meiotic checkpoint function of Mos depends on its capacity to activate the mitogen‐activated protein kinase (MAPK) pathway (Haccard et al, 1993). Thus, very little is known about the role of Mos in somatic cells and on the mechanisms by which Mos can act as an oncogene.
Here, we developed a cellular model of aneuploidization in which p53−/− cells were driven into tetraploidy, which was followed by multipolar mitosis and re‐acquisition of a near‐to‐diploid chromosome content. We found that Mos was upregulated in p53‐deficient tetraploid cells and that it was strictly required for the occurrence of multipolar divisions, presumably because Mos acts as an inhibitor of centrosome coalescence.
Results and discussion
Generation of sub‐tetraploid derivatives from p53‐deficient tetraploid cells
Shortly (2 days) after a 48 h‐long treatment with the microtubule poison nocodazole or the cytokinesis inhibitor cytochalasin D, p53−/− human colon carcinoma HCT 116 cell cultures contained a higher fraction of polyploid cells (with a ⩾4n DNA content), yet a lower amount of dying and dead cells (exhibiting the dissipation of the mitochondrial transmembrane potential (ΔΨm) and the breakdown of plasma membranes, respectively) (Kroemer et al, 2007; Galluzzi et al, 2009) than their p53‐proficient counterparts (Figure 1A and Supplementary Figure S1). Moreover, fluorescence‐activated cell sorter (FACS)‐purified cells with an ∼8n DNA content formed colonies more efficiently when they did not express p53 than when they did so (Figure 1B), in line with the notion that p53 deficiency is permissive for the generation and survival of tetraploid cells (Castedo et al, 2006b). Cultures derived from FACS‐purified cells with an ∼8n DNA content were characterized by a 4n DNA content in the G1 phase of the cell cycle and by an 8n DNA content in the G2 and M phases, thereby exhibiting bona fide traits of tetraploidy. However, after several passages, p53−/− (but neither p21−/− nor Bax−/−) cultures progressively accumulated a population of cells with a ∼2n DNA content (Figure 1C and D). This was observed in six independent experiments in which the initial contamination with ∼2n cells (measured immediately after FACS purification) was undetectable. To understand the origin of such ∼2n population, we followed the fate of tetraploid cells 1 week after their generation by videomicroscopy, and found that p53−/− cells underwent multipolar (mostly tri‐ or tetrapolar) divisions—which are associated with Y‐ and X‐shaped metaphases (Figure 2A)—much more frequently than p53+/+ control cells (Figure 2B and Supplementary Videos 1 and 2). These results suggest that cells with a ∼2n DNA content (hereafter referred to as ‘sub‐tetraploid’) appearing at significant frequencies (⩾10%), as early as 15 days after tetraploidization, might result from a peculiar process of multipolar division. A significant percentage of daughter cells that originated from p53−/− tetraploid cells by multipolar mitosis could enter and terminate normal bipolar divisions (Figure 2C and D, and Supplementary Video 3), suggesting that such sub‐tetraploid cells can give rise to a new lineage. The reduction of the chromosomal content was significantly more frequent among p53−/− tetraploid cells than among control ones (Figures 1 and 2), and, in another cell line, was exacerbated by pharmacological inhibition of p53 by cyclic pifithrin‐α (Komarov et al, 1999) (Supplementary Figure S2). Thus, the tumour suppressor p53 reduces the probability of sub‐tetraploidy.
Sub‐tetraploidy is linked to centrosome defects and aneuploidy
Next, we generated multiple tetraploid clones from isogenic HCT 116 cells. As compared with wild type (WT), Bax−/−, p21−/− or apoptosis‐resistant (owing to the expression of the caspase inhibitor p35 from Baculovirus) tetraploid cells, p53−/− tetraploid clones displayed some difficulties in maintaining a stable tetraploid genome. Four weeks after cloning, indeed, only one‐third of p53−/− tetraploid clones still exhibited a clean tetraploid DNA content profile, whereas the other two‐thirds accumulated viable sub‐tetraploid populations (Figure 3A and B), first as rather broad shoulders (‘phase 1’) and later as sharper peaks (‘phase 2’) of sub‐tetraploid cells. Such sharper peaks presumably arise in cultures that are dominated by one or few viable sub‐clones with a comparable sub‐tetraploid DNA content. Phase 1 and phase 2 unstable sub‐tetraploid populations also arose after repeated transfection (approximately once every 5 days) with a small interfering RNA (siRNA) targeting p53 (Supplementary Figure S2). Chromosome counting confirmed that phase 2 unstable p53−/− tetraploid cells frequently contained a roughly diploid number of chromosomes (Figure 3C and D). Phase 1 unstable p53−/− tetraploid cells were characterized by a high frequency of aberrant mitoses that were either monopolar, bipolar characterized by lagging chromosomes or multipolar linked to supernumerary centrosomes. Multipolar mitoses were also more frequent among phase 2 unstable tetraploids as compared to WT or stable p53−/− tetraploids (Figure 3E). As compared to phase 1 cells, phase 2 cells proliferated more quickly, and among them the sub‐tetraploid population had shorter duplication times and shorter mitoses than the tetraploid one (Supplementary Figure S3). This explains the outgrowth of sub‐tetraploid cells over their tetraploid counterparts. When such sub‐tetraploid cells were exposed to nocodazole, they could again tetraploidize and then revert once more to sub‐tetraploidy, but this process of reversion was not accelerated (Supplementary Figure S4).
Fluorescence in situ hybridization (FISH) carried out during interphase proved that a large portion of sub‐tetraploid cells (which were FACS‐purified from unstable p53−/− tetraploid clones) was aneuploid (Figure 4A‐C). Thus, especially in phase 1 cultures, nullisomies (which are always lethal) were frequently detected (Figure 4C). Accordingly, most (>99%) of such phase 1 sub‐tetraploid cells failed to form stable offspring in clonogenic assays and died (Figure 4D). In phase 2 cultures, the frequency of aneuploid cells was lower, and nullisomies were infrequent (Figure 4C), presumably because viable cells (which efficiently form clones, Figure 4D) had been positively selected. To further explore the behaviour of sub‐tetraploid cells, we generated phase 1 and phase 2 clones from p53−/− tetraploid cells expressing histone H2B fused to the N‐terminus of green fluorescent protein (H2B–GFP, which labels chromatin in green and hence allows for monitoring chromosome movement), and followed the fate of their sub‐tetraploid derivatives after FACS purification (Figure 4E, F and Supplementary Figure S5, and Supplementary Videos 4, 5 and 6). Although phase 1 sub‐tetraploid cells rarely (∼3%) engaged in subsequent cycles of division, phase 2 sub‐tetraploid cells did so much more frequently (∼60%). Nonetheless, cell cycle blockade, mitosis without cytokinesis and cell death occurred more frequently among sub‐tetraploid cells than among their diploid progenitors (Figure 4E, F and Supplementary Figure S5, and Supplementary Videos 4, 5 and 6).
It is important to note that upon inoculation into immunodeficient (nu/nu) mice, tetraploid cells FACS‐purified from unstable tetraploid clones (and in particular established phase 2 cultures) formed tumours more rapidly than stable (p53−/− and p53+/+) tetraploid cells (Figure 5A), corroborating the idea that multipolar divisions might favour the selection of aggressive tumour variants. After in vivo selection, sub‐tetraploid tumour cells could be recovered at similar frequencies as after an equivalent period of in vitro culture (Figure 5B and C) and were particularly frequent among tumours that arose from unstable phase 2 clones. Thus, it appears that the tetraploidization of p53−/− cells can accelerate tumour progression in the context of the emergence of a sub‐tetraploid cancer cell population arising from multipolar mitoses.
Obligate contribution of Mos to the generation of sub‐tetraploid cells
Tetraploidization has been associated with the overexpression of meiosis‐specific proteins, in particular Mos (Kalejs et al, 2006). Accordingly, we observed an increase in Mos protein levels that was particularly pronounced in unstable clones (as compared with stable ones) (Figure 6A). We then assessed whether Mos overexpression (as induced by transfection with a polycystronic vector encoding Mos and GFP) might modify the behaviour of stable tetraploid cells. Although Mos‐transfected p53+/+ were unable to proliferate (Fukasawa and Vande Woude, 1997; Kalejs et al, 2006), p53−/− tetraploid cells survived Mos overexpression while exhibiting an unstable phenotype, associated with the gradual increase of sub‐tetraploid cells in the cultures (Figure 6B and Supplementary Figure S6). Accordingly, depletion of Mos with three different siRNAs had the opposite effect on unstable tetraploid cells and reduced their propensity to generate sub‐tetraploid offspring (Figures 6C and Supplementary Figure S6). Such Mos‐specific siRNAs had no anti‐proliferative effects, ruling out a trivial explanation for their tetraploidy‐stabilizing activity (Supplementary Figure S7). Moreover, retransfection of unstable tetraploid cells with a polycystronic construct coding for a non‐interferable Mos mutant and GFP annihilated the genome‐stabilizing effect of one of the Mos‐specific siRNAs, formally excluding off‐target effects of this siRNA (Figure 6D). These results could be recapitulated in a completely different experimental system, namely WT HCT 116 cells subjected to repetitive transient transfections with a p53‐specific siRNA alone or in combination with a Mos‐specific siRNA. In this setting, the emergence of a sub‐tetraploid population resulting from recurring depletion of p53 was prevented when Mos was concomitantly downregulated (Figure 6E).
In line with reports indicating that Mos can associate with centrosomes (Wang et al, 1994; Doxsey et al, 2005; Nogales‐Cadenas et al, 2009), we found that a Mos–GFP fusion protein localized to centrosomes in HCT 116 cells (Figure 7A and Supplementary Figure S8). To control specificity, we demonstrated that the expression of Mos–GFP (but not of GFP alone) was abolished by a Mos‐specific (but not by a control) siRNA (Supplementary Figure S9). Therefore, we investigated whether Mos might influence centrosome dynamics, and in particular whether Mos might affect centrosome clustering, one mechanism relying on the molecular motor HSET that allows for the maintenance of tetraploid genomes (Kwon et al, 2008). Upon siRNA‐mediated knockdown of HSET (Supplementary Figure S10), stable p53−/− tetraploid cell cultures progressively accumulated a sub‐tetraploid population and this effect was not counteracted by the simultaneous knockdown of Mos (Figure 7B). In phase 1 unstable p53−/− tetraploid cells, Mos depletion limited the generation of a sub‐tetraploid offspring (Figure 7B) while resulting in the conversion of multipolar to bipolar mitoses, in which supernumerary centrosomes either congressed to two centrosome pairs or lost their connection to the β‐tubulin network and were inactivated (Figure 7C, D and Supplementary Figure S8). In this setting, HSET depletion lead to an increase in multipolar divisions paralleled by a decrease in bipolar mitoses characterized by centrosome coalescence (Figure 7D), correlating with a further increase in the generation of sub‐tetraploid cells (Figure 7B). These effects were reduced by the simultaneous depletion of Mos (Figure 7B and D), in line with the hypothesis that Mos modulates centrosome dynamics, thereby affecting the frequency of multipolar mitoses in tetraploid cells. To directly evaluate the putative effects of Mos on centrosome clustering, we transiently transfected a (phase 2) unstable p53−/− tetraploid clone that had previously been engineered for the expression of GFP–H2B with a centrin–DsRed fusion‐encoding construct, allowing for the simultaneous monitoring of chromosomal and centrosomal dynamics by videomicroscopy (Figure 7E). Cells that were left untreated or were transfected with a control siRNA frequently underwent multipolar divisions, which were characterized by the presence of more than two centrin–DsRed‐positive dots. However, upon transfection with a Mos‐specific siRNA the frequency of multipolar (or failed) divisions dropped (Figure 7E and F). In line with immunofluorescence microscopy results (Figure 7C, D and Supplementary Figure S8), we observed an increase in the incidence of bipolar divisions that exhibited centrosome coalescence. Taken together, these data strongly suggest that the knockdown of Mos favours centrosome clustering, correlating with the suppression of multipolar mitosis.
On theoretical grounds, tetraploid cells may generate an aneuploid offspring by two non‐exclusive mechanisms. First, as tetraploid cells contain supernumerary centrosomes, they may enter multipolar mitosis, thereby dividing into more than two (usually three or four) daughter cells provided with randomly distributed chromosomes. Most of these cells would be non‐viable and would die during the subsequent interphase or after one additional round of (failing) mitosis. However, in exceptional circumstances, the overall genome composition of one daughter cell may be compatible with survival, and rare, particularly fit cells would then out compete their tetraploid parent, as it has been observed in cell cultures and in tumours. Alternatively, tetraploid cells might lose single (or few) chromosomes because of their merotelic attachment, which results from a ‘multipolar spindle intermediate’. In this case, the presence of supernumerary centrosomes enforces an initial multipolar metaphase followed by the clustering of all centrosomes in two poles, which results in a bipolar division, whereby the chromosome that is merotelically attached lags during anaphase and eventually is either lost or distributed into the wrong daughter cell. This mechanism might give rise to a gradual, progressive loss of chromosomes until the achievement of a sub‐tetraploid stage.
Irrespective of the molecular details underlying the aneuploidization of tetraploid cells, it appears that centrosome clustering constitutes a prime mechanism for the maintenance of tetraploid genomes. Supernumerary centrosomes found in tetraploid cells would indeed cause multipolar division or merotelic chromosome attachment (and hence chromosome loss), unless they aggregate in two discrete pools and form two microtubule‐organizing centres. This aggregation is under the control of molecular motors including HSET, and HSET depletion indeed compromises the stability of tetraploid cells (Kwon et al, 2008). As shown here, overexpression of Mos that occurs in p53−/− tetraploid cells also inhibits centrosome clustering, thereby favouring multipolar divisions and resulting in the reduction of the tetraploid genome to a sub‐tetraploid one.
Little information is available on the cross talk between p53 and Mos. In samples from non‐small cell lung carcinoma patients, p53 alterations were shown to correlate with high Mos expression, aneuploidy and tumour aggressiveness (Gorgoulis et al, 2001). As a possible scenario, the inactivation of p53 would favour the presence of supernumerary centrosomes (Fukasawa et al, 1996; Carroll et al, 1999) while augmenting the probability of cells to survive the tetraploidization process (Castedo et al, 2006a; Senovilla et al, 2009). Tetraploid cells that lack an intact p53 system would then upregulate Mos, in turn favouring aneuploidization. Beyond such an oncogenic cooperation at the functional level, additional interactions between p53 and Mos may be envisioned. Reportedly, p53‐deficient cells tolerate higher levels of Mos than their p53‐proficient counterparts (Fukasawa and Vande Woude, 1997; Kalejs et al, 2006), suggesting that the ‘oncogenic stress’ mediated by ectopic expression of this meiosis‐restricted protein may kill cells or arrest their cycle if the p53 system is intact. Thus, it is conceivable that the absence of p53 may be permissive for the upregulation of Mos (Fukasawa and Vande Woude, 1997; Gorgoulis et al, 2001; Kalejs et al, 2006), which must occur at the post‐transcriptional level, as Mos mRNA levels were not increased in tetraploid cells (data not shown). However, the exact molecular mechanisms explaining the unscheduled expression of Mos in tumour cells remain elusive.
The precise oncogenic mode of action of Mos is an ongoing conundrum. Enforced Mos expression in fibroblasts reportedly causes centrosome amplification (Saavedra et al, 1999). However, although siRNA‐mediated knockdown of Mos in tetraploid cells apparently did not decrease the number of centrosomes, Mos expression inversely correlated with the coalescence of supernumerary centrosomes, as if Mos acted as an inhibitor of centrosome clustering. Accordingly, the depletion of Mos favoured centrosome coalescence while switching multipolar to bipolar mitoses, and this effect was inhibited by the simultaneous knockdown of the kinesin HSET, which is indispensable for centrosome clustering. Based on these results, we conclude that Mos may exert its oncogenic activity, at least in part, through the induction of multipolar, asymmetric cell division. It remains to be determined whether the oncogenic activity of Mos is mediated by its direct interaction with centrosomes or by hitherto unknown effects on cellular morphology/dynamics that indirectly affect centrosome clustering.
Materials and methods
Unless otherwise indicated, media and supplements for cell culture were purchased from Gibco‐Invitrogen (Carlsbad, CA, USA), plasticware from Corning B.V. Life Sciences (Schiphol‐Rijk, The Netherlands), and chemicals from Sigma‐Aldrich (St Louis, MO, USA).
Cell lines, culture conditions and chemicals
Wild type, p53−/−, Bax−/− and p21−/− diploid human colon carcinoma HCT 116 cells (kindly provided by Bert Vogelstein), diploid HCT 116 cells stably transfected with a vector encoding the baculoviral caspase inhibitor p35 (Zhang et al, 2006), WT and p53−/− diploid HCT 116 cells transfected with a cDNA coding for the fusion between histone H2B and green fluorescent protein (H2B–GFP, PharMingen, San Diego, CA, USA), as well as WT diploid human colon carcinoma RKO cells were routinely maintained in McCoy's 5A medium (PAA Laboratories GmbH, Pasching, Austria) supplemented with 10% foetal calf serum (FCS) at 37 °C in a 5% CO2 atmosphere. H2B–GFP‐expressing cells were cultured in the presence of 20 μg/ml blasticidine. Cells were seeded onto the appropriate supports (6‐, 12‐, 24‐ or 96‐well plates, 35 or 100 mm Ø Petri dishes) 24 h before the beginning of the experiment. To generate tetraploid clones, parental diploid cells were treated for 48 h with 0.6 μg/ml cytochalasin D or 100 nM nocodazole and then cultured for 2 weeks in drug‐free culture medium, followed by staining with Hoechst 33342 (2 μM; Molecular Probes‐Invitrogen, Eugene, OR, USA) and cloning of cells characterized by an ∼8n DNA content on a FACSVantage cell sorter (BD Biosciences, San Jose, CA, USA), as previously described (Castedo et al, 2006b). Alternatively, tetraploid clones were isolated by the limiting dilution technique (Castedo et al, 2006b). To prevent drifts in cell composition, clones were split well before complete confluence. For each experiment, at least three different clones for each genotype and/or phenotype were used.
Plasmid transfection and RNA interference
A cDNA encoding for centrin derived from a human testis mRNA library (GenBank accession number BC029515.1) was amplified in a pCMV‐sport6 vector (Invitrogen) and then cloned into a pDsRed2‐C1 plasmid (Clontech Laboratories, Mountain View, CA, USA) as a BglII restriction fragment, generating a construct for the expression of a centrin‐DsRed chimera. The human Mos cDNA (Image Clone 40016104) was purchased from Geneservice (Nottingham, UK) within a pCR‐bluntII‐TOPO plasmid (Invitrogen). The Mos sequence was then transferred either to the polycystronic expression vector pIRES‐hrGFP2 (Agilent Technologies, Santa Clara, CA, USA) as a BamH1–Not1 restriction fragment, or to the pEGFP‐C3 plasmid (Clontech Laboratories) as a Xho1–BamH1 restriction fragment upon PCR amplification (primers 5′‐TATACTCGAGCCCTCGCCCCTGGCCC‐3′ and 5′‐TATAGGATCCTCAGCCGAGTTCAGCTTTCA‐3′, restriction sites are underlined). In order to generate a non‐interferable but functional Mos mutant, the Mos sequence was further modified from within the pIRES‐hrGFP2 vector to introduce the silent mutations ATCATA at position 619 (Ile207) and TTGCTA at position 622 (Leu208). Site‐directed mutagenesis was performed with the Quikchange Site‐Directed Mutagenesis Kit (Stratagene, La Jolla, CA, USA) and the primers 5′‐GGACCTGAAGCCCGCGAACATACTAATCAGTGAGCAGGATGTC‐3′ and 5′‐GACATCCTGCTCACTGATTAGTATGTTCGCGGGCTTCAGGTCC‐3′ (mutated nucleotides are underlined), according to the manufacturer's instructions. Briefly, PCR was employed to produce several copies of the entire plasmid including the desired mutations. The reaction mix was then incubated for 1 h at 37°C with the Dpn1 restriction enzyme, which cleaves only methylated DNA (in this case, the parental, non‐mutated molecules). Uncleaved (mutated) DNA was then used for transformation of Escherichia coli XL1‐blue competent cells (Stratagene).
Custom‐designed siRNAs duplexes targeting Mos (Mos_1 sense 5′‐GCCCGCGAACAUCUUGAUCdTdT‐3′; Mos_2 sense 5′‐GCCUAAAGCCGACAUUUAUdTdT‐3′) and p53 (p53_1 sense 5′‐GUGAGCGCUUCGAGAUGUUdTdT‐3′; p53_2 sense 5′‐GACUCCAGUGGUAAUCUACdTdT‐3′) (Gu et al, 2004) were purchased from Sigma‐Proligo (The Woodlands, TX, USA). Alternatively, siRNA duplexes for the downregulation of Mos (Mos_3) and HSET (HSET_1) (siGENOME Smart‐pool M‐003859 and L‐004958, respectively) were purchased from Dharmacon (Chicago, IL, USA). As a control, a non‐targeting siRNA with an sequence unrelated to both human and murine genomes was used (UNR, sense 5′‐GCCGGUAUGCCGGUUAAGUdTdT‐3′). HCT 116 cells in 12‐well plates were transfected with plasmids at 80% confluence by means of the Attractene® transfection reagent (Qiagen, Hilden, Germany) as recommended by the manufacturer, or with siRNAs at 30–40% confluence by means of the HiPerFect® transfection reagent (Qiagen), as previously described (Hoffmann et al, 2008). After 72 h, transfection efficiency was determined by immunoblotting (see below). When required, cells were transfected first with siRNAs and 24 h later with plasmids.
For the simultaneous quantification of DNA content (cell cycle profiling), plasma membrane integrity and mitochondrial transmembrane potential (ΔΨm), live cells were collected and stained with 2 μM Hoechst 33342 (Molecular Probes–Invitrogen), 1 μg/ml propidium iodide (PI, which only incorporates into dead cells) and 40 nM of the ΔΨm‐sensitive dye 3,3′‐dihexyloxacarbocyanine iodide (DiOC6(3), from Molecular Probes–Invitrogen) (Castedo et al, 2002; Galluzzi et al, 2009). To assess cell cycle distribution, cells were collected, fixed by gentle vortexing in ice‐cold 80% (v/v) ethanol (Carlo Erba Reagents, Milano, Italy) and stained with 50 μg/ml PI in 0.1% (w/v) d‐glucose in PBS supplemented with 1 μg/ml (w/v) RNAse A (Mouhamad et al, 2007; Mondragon et al, 2009). For the cell count assay, the same amount of FITC‐labeled beads (Becton Dickinson) was added to each sample and cells were counted until 5000 beads had passed through the FACS. Cell proliferation was determined by counting the cells every 24 h for 4 days upon the seeding. Cytofluorometric acquisition of blue (Hoechst 33342), red (PI) and green (DiOC6(3)) fluorescence were carried out by means of a FACSCalibur or a FACScan cytofluorometer (BD Biosciences) equipped with a 70 μm nozzle. Statistical analysis was carried out by using the CellQuest™ software (BD Biosciences), upon gating on the events characterized by normal forward scatter and side‐scatter parameters.
For videomicroscopy, HCT 116 cells stably transfected with the cDNA encoding for H2B–GFP were grown in 96‐well imaging plates (BD Biosciences) under standard culture conditions (37 °C, 5% CO2) and subjected to pulsed observations (every 10 min for up to 48 h) with a BD pathway 855 automated live cell microscope (BD Biosciences). Images were analyzed with the open source software Image J (freely available from the National Institute of Health, Bethesda, MD, USA at the address http://rsb.info.nih.gov/ij/).
Cell proliferation assays
Cell proliferation was measured by the fluorescein‐based dye 5‐(and‐6)‐carboxyfluorescein diacetate succinimidyl ester (CFSE) staining. Briefly, 1 × 106 cells were incubated with 2.5 μM CFSE in PBS for 10 min at 37°C. The labelling reaction was stopped by adding an equal volume of FCS and 5 × 104 cells were seeded in 12‐well plates. After 24 h, cells were transfected with siRNAs and grown in culture for additional 3–5 days. Cell proliferation was monitored daily on a FACSCalibur cytofluorometer (BD Biosciences) upon excitation at 488 nm and acquisition in the FL1 (green) channel, followed by data analysis by means of the CellQuest software (BD Biosciences).
Immunofluorescence microscopy determinations were performed as previously described (Vitale et al, 2007, 2008). Briefly, cells were fixed in 4% (w/v) paraformaldehyde in PBS, permeabilized with 0.1% SDS and immunostained with antibodies specific for γ‐ and β‐tubulin (both from Sigma‐Aldrich). Slides were then incubated with the appropriate Alexa Fluor® conjugates (Molecular Probes–Invitrogen) in the presence of 10 μM Hoechst 33342 (Molecular Probes–Invitrogen), which was used for nuclear counterstaining. Fluorescence and confocal fluorescence images were captured using an IRE2 microscope equipped with a DC300F camera (both from Leica Microsystems GmbH, Wetzlar, Germany) and a TSC‐SPE microscope (Leica Microsystems GmbH) equipped with a 63X/1.15 objective (Olympus America, Center Valley, PA, USA), respectively. Signals from different probes were acquired in sequential scan mode and image analysis was performed with the open source software Image J (National Institutes of Health).
HCT 116 cells were washed with cold PBS and lysed in a buffer containing 1% NP40, 20 mM HEPES (pH 7.9), 10 mM KCl, 1 mM EDTA, 10% glycerol, 1 mM orthovanadate, 1 mM PMSF, 1 mM dithiothreitol, 10 μg/ml aprotinin, 10 μg/ml leupeptin and 10 μg/ml pepstatin, as previously described (Zermati et al, 2007). Thereafter, protein extracts (50 μg per lane) were separated according to molecular weight on precast 4–12% SDS–PAGE gels (Invitrogen), followed by electrotransfer to Immobilon™ membranes (Sigma‐Aldrich) and immunoblotting with antibodies specific for HSET and Mos (both from Santa Cruz Biotechnology, Santa Cruz, CA, USA). Primary antibodies that specifically recognize glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) or β‐actin (both from Millipore‐Chemicon International, Temecula, CA, USA) were used as loading control. Finally membranes were incubated with appropriate goat IgG conjugated to horseradish peroxidase (Southern Biotech, Birmingham, AL, USA), followed by chemiluminescence detection with the SuperSignal West Pico® reagent and CL‐XPosure® X‐ray films (both from Thermo Scientific–Pierce, Rockford, IL, USA).
Cytogenetic analysis and FISH
Chromosome spreads were prepared by conventional procedures (Mouhamad et al, 2007). Briefly, HCT 116 clones were treated with 100 nM nocodazole for 8 h to enrich mitotic cells, then collected and subjected to hypotonic lysis by incubation in 75 μM KCl for 10 min at 37°C. After removal of hypotonic solution, cells were fixed in freshly prepared Carnoy solution (3:1 methanol:acetic acid) and stored at −20°C. Fixed cells were then dropped onto pre‐cooled Superfrost Plus glass microscope slides (Thermo Fisher Scientific, Waltham, MA, USA) and dried at room temperature. Chromosomes were stained with 100 ng/ml 4′,6 diamidino‐2‐phenylindole (DAPI, from Molecular Probes–Invitrogen) and mounted with glass coverslips in Vectashield H‐1000 mounting medium (Vector Laboratories, Burlingame, CA, USA). Finally, fluorescence images were visualized and captured using an IRE2 microscope equipped with a DC300F camera (both from Leica Microsystems GmbH). For each experimental condition, 100 cells from three independent experiments were analyzed. For FISH, freshly FACS‐purified diploid and sub‐tetraploid cells were dropped onto Superfrost Plus polylysine‐coated glass microscope slides (Thermo Fisher Scientific) and fixed in situ with 9:1 methanol:acetic acid for 5 min. Thereafter, cells were air‐dried overnight and hybridized with a commercial mixture of three probes (Abbott Laboratories, Abbott Park, IL, USA) that detect the centromeric region of chromosome 8 (labelled with FITC—green colour), chromosome 10 (labeled with rhodamine—red colour) and chromosome 18 (labeled with Aqua—blue colour). For each experimental conditions, 100–300 nuclei were surveyed.
Clonogenic survival assays
To evaluate clonogenic survival, freshly generated tetraploid and sub‐tetraploid cells were stained with 2 μM Hoechst 33342 (Molecular Probes–Invitrogen), FACS‐purified on a FACSVantage cell sorter (BD Biosciences), seeded at different concentrations (from 1 to 50 × 103 for well) in 6‐well plates, and cultured for up to 10 days under normal conditions. Colonies were then fixed/stained with an aqueous solution containing 0.25% (w/v) crystal violet, 70% (v/v) methanol and 3% (v/v) formaldehyde (Carlo Erba Reagents) and counted (Zhang et al, 2006). Only colonies made of ⩾30 cells were included in the quantification. For each treatment, the surviving fraction (SF) was estimated according to the formula: SF=(number of colonies formed/ number of cells seeded) × plating efficiency (defined as the ratio between the number of colonies formed and the number of cells seeded in control conditions).
In vivo xenograft model
Athymic nu/nu female mice (age=42 days, body weight=20 g, provided by the Institut Gustave Roussy (IGR) in‐house animal facility) were used throughout this study in strict compliance with widely accepted ethical guidelines for animal experimentation. Mice were kept in Makrolon® type III wire mesh laboratory cages (Charles River, Boston, MA, USA), under poor germ conditions at 24°C and 50–60% humidity, and were allowed for food and water ad libitum. Light cycle was artificially controlled to provide 14 h of light (from 0630 h to 2030 h). After 4 days of acclimation period, mice were subcutaneously xenografted with 2 × 106 WT or p53−/− tetraploid HCT 116 cells, as previously described (Vitale et al, 2007). Tumour growth was then assessed every 2–4 days with a standard caliper. For ethical reasons, mice were killed before tumours reached a volume of 3400 mm3 or a surface of 250 mm2. Statistical analysis was carried out by means of the SigmaStat package (Systat Software, San Jose, CA, USA). One‐way analysis of variance was carried out and statistical significance was determined by means of two‐tailed Student's t‐test (*P<0.05) in the context of a pairwise comparison procedure.
To recover tumour cells, mice were killed by cervical dislocation following the FELASA guidelines. Quickly after killing, tumour tissue was surgically removed and washed in RPMI 1640 culture medium supplemented with 100 U/ml penicillin G sodium, 100 μg/ml streptomycin sulphate and 10% FCS. Thereafter, a tissue fragment of ∼125 mm3 was cut into pieces of ∼1 mm3, which were dissociated by incubation with 0.05% trypsin for 45 min at 37°C. Every 15 min the disaggregation of cells was mechanically carried out. Finally, dissociated cells were washed once with RPMI 1640 supplemented with 100 U/ml penicillin G sodium, 100 μg/ml streptomycin sulphate and 10% FCS, and seeded into 25 cm2 flasks. Recovered tumour cells were routinely maintained under standard culture conditions (37°C, 5% CO2) in McCoy's 5A medium supplemented with 100 U/ml penicillin G sodium, 100 μg/ml streptomycin sulphate, 100 mM HEPES buffer, 1 mM sodium pyruvate and 10% FCS.
Unless otherwise specified, all experiments were carried out in triplicate parallel instances and independently repeated at least three times. Data were analyzed with Microsoft Excel (Microsoft, Redmond, WA, USA) and statistical significance was assessed by means of two‐tailed Student's t‐test (*P<0.05).
Supplementary data are available at The EMBO Journal Online (http://www.embojournal.org).
Conflict of Interest
The authors declare that they have no conflict of interest.
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Supplementary Figures and Video Legends
GK is supported by the Ligue Nationale contre le Cancer (Equipe labellisée), Agence Nationale pour la Recherche, European Commission (Apo‐Sys, ChemoRes, ApopTrain), Fondation pour la Recherche Médicale, Institut National du Cancer, Cancéropôle Ile‐de‐France. MC is supported by the Association pour la Recherche sur le Cancer (ARC). LS is supported by ApopTrain Marie Curie training network of the European Union and FRM, MM and SR‐V by FRM, LG by the Apo‐Sys consortium of the European Union, OK by EMBO. IV, LS, MJ, MM, OK, LN, AC, SRV, GM, DM, SV, NT, NJ, AV and MC carried out the experiments presented in this article. IV, LG, MC and GK prepared the figures and wrote/edited the paper.
↵† These authors share senior co‐authorship
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