Nuclear tension controls mitotic entry by regulating cyclin B1 nuclear translocation

Cell-cycle progression is regulated by cyclins and their associated kinases. One such complex, composed of cyclin B1-CDK1, is responsible for regulating entry into mitosis. The biochemical mechanisms that regulate mitotic entry have been extensively studied in the past (for review, see Lindqvist et al. [2009]). For most of the cell cycle, the cyclin B1-CDK1 complex is inactive, due to low cyclin B1 expression levels and its mainly cytoplasmic localization (Hagting et al., 1998; Toyoshima et al., 1998). As cells transit from S to G2, cyclin B1 expression gradually increases (Akopyan et al., 2014; Feringa et al., 2016; Pines and Hunter, 1989). This results in cyclin B1 binding to CDK1. This cyclin B1-CDK1 complex is kept in an inactive state due to Myt1- and Wee1-mediated phosphorylations of CDK1 on residues T14 and Y15 (Lindqvist et al., 2009). Through the action of Cdc25 phosphatases, these inhibitory phosphorylations are removed, leading to complex activation. The active cyclin B1-CDK1 complex then stimulates its own nuclear import (Gavet and Pines, 2010) through the nuclear pore complexes (NPCs), resulting in chromosome condensation (Abe et al., 2011) and nuclear envelope permeabilization (NEP; Dantas et al., 2021). Consequently, the cyclin B1-CDK1 complex has been proposed to effectively synchronize cytoplasmic and nuclear events (Gavet and Pines, 2010), crucial for mitotic entry and efficient spindle assembly.

The influence of mechanical forces on the cell cycle and some of its key regulators has received considerable attention over the last few years (Huang et al., 1998; Klein et al., 2009; Lancaster et al., 2013; Benham-Pyle et al., 2015; Gudipaty et al., 2017; Uroz et al., 2018; Aureille et al., 2019). Experiments performed in isolated cells and epithelial layers have demonstrated that mechanical forces can stimulate the G1-S transition (Huang et al., 1998; Benham-Pyle et al., 2015; Uroz et al., 2018; Vianay et al., 2018; Aureille et al., 2019) by controlling specific transcriptional programs (Huang et al., 1998; Benham-Pyle et al., 2015; Aureille et al., 2019). This is likely due to forces imposed on the nucleus (Lombardi and Lammerding, 2011; Arsenovic et al., 2016) that induce its flattening (Elosegui-Artola et al., 2017; Aureille et al., 2019; Lomakin et al., 2020), facilitating the nuclear accumulation of transcription factors (TFs; Elosegui-Artola et al., 2017; Jacchetti et al., 2021), changing the organization of both chromatin (Nava et al., 2020) and the nuclear envelope (NE; Swift et al., 2013) or altering cell contractility (Lomakin et al., 2020; Venturini et al., 2020).

Evidence for mechanical regulation during other cell-cycle phases is more limited. Recently, mechanical stretching was proposed to trigger the G2-M transition by activating Piezo 1 (Gudipaty et al., 2017). Moreover, we and others have shown that cell traction forces decrease during the G2-M transition (Uroz et al., 2018; Vianay et al., 2018; Nunes et al., 2020), to allow mitotic cell rounding and efficient cell division (Lancaster et al., 2013; Nunes et al., 2020). Although these findings highlight the interplay between physical forces and cell proliferation, it remains unknown whether the main biochemical events that occur during the G2-M transition are sensitive to mechanical cues. Importantly, whether the spatial and temporal behavior of the cyclin B1-CDK1 complex responds to mechanical forces and contributes to ensure timely and efficient cell division remains unknown. Here, we investigate whether and how cyclin B1 responds to physical forces during the G2-M transition. We show that nuclear deformation triggers a contractility-mediated mechanism that facilitates the translocation of cyclin B1 to the nucleus, setting the timing of mitotic entry.

We started by investigating whether the G2-M transition is sensitive to the degree of cell adhesion. We seeded RPE-1 cells expressing H2B-GFP/tubulin-RFP in non-adherent, hydrophobic conditions (PLL-g-PEG). These cells were trypsinized, placed on a pLL-g-PEG-coated 35 mm² dish and immediately placed on the microscope for high spatiotemporal resolution imaging. We selected individual cells that were in late G2/early prophase, based on the presence of separated centrosomes and condensed chromosomes, and determined their capacity to enter mitosis. During a period of at least 3 h of imaging, these cells failed to enter mitosis (Fig. 1, A and C; n = 11), confirming that mitotic entry requires cell adhesion. Inversely, adherent cells seeded on fibronectin (FBN), readily entered mitosis (Fig. 1, B and C; n = 23), within the same time frame. We then tested whether mechanically stimulating cells in non-adherent conditions was sufficient to induce mitotic entry. For this purpose, we used a dynamic cell confinement device (Le Berre et al., 2014), which gives us temporal control over the process. Confinement was achieved by imposing on cells a fixed height of 8 μm with a rigid polydimethylsiloxane (PDMS)-coated glass slide (Fig. S1 A). Upon acute confinement, the cells that were seeded on PLL-g-PEG regained the ability to enter mitosis (Fig. 1, A–D; n = 12; ***, P < 0.001), indicating that mechanical confinement is sufficient to overcome the lack of cell adhesion. Under confinement conditions, these cells entered mitosis within ∼260 ± 129 s (mean ± SD) after stimulation, ruling out that this event was due to increased cyclin B1 transcription, as previously described (Gudipaty et al., 2017). One alternative hypothesis is that physical confinement accelerates mitotic entry by inducing a premature transport of cyclin B1 to the nucleus, as previously proposed for YAP or MyoD (Elosegui-Artola et al., 2017; Jacchetti et al., 2021). To confirm this is the case, we monitored the dynamics of nuclear accumulation of endogenous cyclin B1 tagged with Venus in RPE-1 cells (Collin et al., 2013). We specifically selected cells in late G2/early prophase, based on the presence of high cytoplasmic levels of cyclin B1 (Fig. 1 E; ***, P < 0.001), the absence of nuclear cyclin B1, and the separation of centrosomes (Fig. 1 F), all of which were previously determined as reliable markers for this cell-cycle stage (Akopyan et al., 2014; Feringa et al., 2016). We then measured nuclear cyclin B1 translocation in normal and confined conditions (Fig. 1, G–I). Next, we normalized our measurements to the lowest fluorescence intensity levels of nuclear cyclin B1 (corresponding to a value of 1) and quantified its increase as cyclin B1 translocated into the nucleus, up until the moment of tubulin nuclear entry, which is defined as time 0. This last event marks the loss of the nuclear barrier function, which is defined as NEP. By analyzing the patterns of nuclear cyclin B1 translocation, we determined that, in comparison with unmanipulated cells, mechanical stimulation triggered a fast nuclear accumulation of cyclin B1 (Fig. 1, J–M; ***, P < 0.001), as well as an increased nuclear-to-cytoplasmic (N/C) cyclin B1 ratio just prior to NEP (Fig. 1 N; ***, P < 0.001). This change in the N/C ratio strengthens the hypothesis that cyclin B1 translocation is a mechanosensitive event. Indeed, recent evidence has shown that the transport of high-molecular weight proteins (>40 kD) across the NPCs is sensitive to mechanical stimulation, with a preference for increased nuclear import, which is reflected in elevated N/C ratios (Andreu et al., 2022). However, we cannot rule out that confinement could act on other aspects of the nucleocytoplasmic transport, such as the rate of export. Indeed, we observed that following treatment with leptomycin B (LMB), which prevents the nuclear export of cyclin B1 (Yang et al., 1998; Yang et al., 2001), the difference in the N/C ratio is lost, when comparing confined and non-confined cells (Fig. 1 N; n.s., not significant).

Next, to assess the kinetics of translocation, we aligned cyclin B1 fluorescence levels to the lowest value for both experimental groups (Fig. 1 M) and performed an exponential fitting. We determined the translocation to occur within 478 ± 102 s, with a half-time of 331 s for unmanipulated cells. Strikingly, in confined cells, translocation occurred within 101 ± 12 s, with a half-time of 70 s, which resulted in a faster mitotic entry (Fig. 1, J–L). This was not due to a rupture of the nucleus, as we could not detect any obvious association of GFP-cGAS with DNA (Fig. S1, B and D) or leakage of NLS-GFP into the cytoplasm (Fig. S1, C, E, and F), when cells were confined to a height of 8 μm, which we routinely used for our experimental setup. However, confining cells to a height of 3 μm did increase association of cGAS with DNA (Fig. S1, B and D) and resulted in NLS-GFP leakage into the cytoplasm (Fig. S1, C and G). These results, together with the observed delay between cyclin B1 accumulation and tubulin translocation to the nucleus (Fig. 1, K and L), strongly suggest that the nuclear barrier function remains intact after an 8 μm confinement. Instead, as previously reported (Lomakin et al., 2020), confinement promoted an unfolding of the NE which could be readily observed (Fig. S1, H and I; ***, P < 0.001) and resulted in an increase in the distance between neighboring NPCs (Fig. S1, J and K; ***, P < 0.001), when compared to unconfined cells. Importantly, this unfolding was also observed in unmanipulated cells as they progress from interphase to prophase (Fig. S1 K; ***, P < 0.001). Because DNA damage can affect nuclear cyclin B1 accumulation (Toyoshima et al., 1998), we measured the levels of histone γ-H2AX, to determine if our experimental setup could be inducing DNA damage, thus affecting cyclin B1 nuclear accumulation. We did not observe significant changes in the levels of γ-H2AX foci (Fig. S1, L and M; ***, P < 0.001), suggesting that this process is independent of DNA damage.

Our data suggest that the mechanical environment might affect cyclin B1 nucleoplasmic translocation. To test this, we seeded cells on a soft hydrogel (5 kPa) or on a rigid glass, inducing low or high cellular tension, respectively. As predicted, cells on glass were more efficient in cyclin B1 nuclear shuttling, than cells on a soft gel (Fig. S2, A and B; ***, P < 0.001). This mechanical stimulation of cyclin B1 translocation was further confirmed by treating RPE-1 cells with a hypotonic shock (Fig. S2, C and D), known to induce cell and nuclear membrane tension (Bakkenist and Kastan, 2003; Kumar et al., 2014), without generating apparent DNA damage (Bakkenist and Kastan, 2003). As expected, the hypotonic shock induced a faster translocation of cyclin B1 into the nucleus, when compared to controls (Fig. S2, C–F; ***, P < 0.001). Confinement of HeLa cells yielded similar faster translocation of cyclin B1 into the nucleus (Fig. S2, G–I; **, P < 0.01), suggesting a conservation of this mechanism. Nevertheless, it should be noted that our confinement setup does not allow distinction between cytoplasmic and nuclear compression and, therefore, we cannot rule out that confinement may trigger additional cytoplasmic mechanosensitive pathways that could contribute to the G2/M transition.

During mitotic entry, cells reorganize their cytoskeleton and round up (Maddox and Burridge, 2003), decreasing the traction forces exerted by cells on the extracellular environment (Nunes et al., 2020). To test whether the process of cell rounding interferes with cyclin B1 translocation, we expressed a mutant form of the GTPase Rap1 (Rap1Q63E, hereafter named Rap1*), which effectively blocks focal adhesion disassembly and prevents cell rounding. Expression of Rap1* did not alter cyclin B1 translocation, even though cell rounding was efficiently blocked, as measured by cell membrane eccentricity (Nunes et al., 2020; Fig. S3, A and B; ***, P < 0.001). Although we cannot completely rule out that cells with different rounding properties could exhibit changes in cyclin B1 translocation kinetics, our data are indicative that this process seems to be independent of cell rounding.

Next, we set out to determine how this confinement-induced cyclin B1 translocation depended on the classical cyclin B1 import pathway. Because translocation in unconfined situations relies on cyclin B1-CDK1 activation (Gavet and Pines, 2010), we imaged cells treated with the CDK1 inhibitor RO-3306, with or without confinement (CDK1i; Fig. 2, A–C). As expected, CDK1 inhibition effectively blocked cyclin B1 translocation to the nucleus (Fig. 2, B and F). Interestingly, confinement was sufficient to overcome the inhibition of CDK1 and force translocation of cyclin B1 to the nucleus (Fig. 2, C and F; ***, P < 0.001). However, these cells failed to enter mitosis, as NEP was blocked by CDK1 inhibition (Heald and McKeon, 1990; Peter et al., 1990). This observation further strengthens the idea that confinement per se does not affect the nucleus barrier function. Accumulation of cyclin B1 relies on a balance between its nuclear import and export. Export is regulated by cyclin B1 binding to the exportin Crm1 (Yang et al., 2001), whereas import is dependent on binding to importin β (Moore et al., 1999) and is greatly enhanced by phosphorylation of cyclin B1 on its CRS sequence (Li et al., 1997; Hagting et al., 1999). We started by treating cells with importazole, to inhibit importin β function. This treatment efficiently blocked cyclin B1 nuclear translocation, even in confinement conditions (Fig. 2, D and G), indicating that the accelerated translocation of cyclin B1 to the nucleus cannot be explained by an increased diffusive shuttling alone, but by an active process, dependent on importin β. A similar block in cyclin B1 translocation was observed when cells expressing a mutant form of cyclin B1 with its five CRS phosphorylation sites mutated to alanines (cyclin B1-5A-GFP; Hagting et al., 1999), were subjected to confinement (Fig. S3, C and D). On the other hand, when nuclear export was inhibited by treatment with LMB, we observed significant decrease in the nuclear accumulation of cyclin B1 when compared to controls (Fig. 2, E and H; and Fig. S3 E), in accordance with previous reports (Santos et al., 2012). This effect was only partially rescued by confinement (Fig. 2 H; ***, P < 0.001). In addition to the mechanisms described above, Plk1 activity has also been recently proposed to regulate mitotic entry and cyclin B1-CDK1 activity (Gheghiani et al., 2017). To determine if confinement-induced translocation was dependent on Plk1 activity, we treated RPE-1 cells with 200 nM of the Plk1 inhibitor BI2536 for 2 h (Plk1i), a dose previously reported to block mitotic entry (Gheghiani et al., 2017). Accordingly, this treatment prevented cells from accumulating cyclin B1 in the nucleus and entering mitosis (Fig. S3, F–I). Strikingly, confining Plk1i cells was sufficient to rescue the accumulation of cyclin B1 and allow mitotic entry (Fig. S3, F–I; ***, P < 0.001). It should be noted that confined, Plk1i-treated cells that failed to enter mitosis also did not accumulate cyclin B1 in the nucleus (Fig. S3, H and I). Overall, our data strongly suggest that a mechanical signal acts in parallel with the biochemical pathways to help regulate the timing of cyclin B1 nuclear accumulation and control mitotic entry, in a process that requires phosphorylation of cyclin B1 in the CRS and binding to importin β.

We then sought to identify potential mechanosensing mechanisms involved in the transport of cyclin B1 to the nucleus. Mechanical forces generated by the cytoskeleton are transmitted to the nucleus through the linker of cytoskeleton and nucleoskeleton (LINC) complex (Lombardi and Lammerding, 2011). To address how force transmission might affect cyclin B1 nuclear translocation, we exogenously expressed a dominant-negative form of nesprin tagged with RFP (DN-KASH-RFP) that prevents its binding to SUN proteins and blocks force propagation (Lombardi and Lammerding, 2011). This was performed in two parallel experiments: firstly, we expressed DN-KASH-RFP in the RPE-1 cyclin B1–Venus cell line. This allowed us to specifically select cells that were expressing the KASH construct (Fig. S4 A) and measure cyclin B1 translocation (Fig. S4 B). Secondly, we expressed the same construct in the RPE-1 cyclin B1–Venus/tubulin-RFP that was used in our previous experiments, so that we could directly compare the effects on cyclin B1 translocation. Importantly, expressing DN-KASH-RFP in either cell line yielded similar results regarding cyclin B1 translocation (Fig. S4 B; ***, P < 0.001; n.s., not significant), with a significant delay in cyclin B1 nuclear accumulation (Fig. 3, A and I; ***, P < 0.001). These results demonstrate that the effect of confinement was mediated by force transmission to the nucleus. Similarly, we also observed delays in cyclin B1 translocation following inhibition of ROCK with Y-27632 (Fig. 3, C and J; **, P < 0.01), depletion of ROCK1 with shRNA (Fig. 3, F and K; ***, P < 0.001), myosin II inhibition with para-nitro-blebbistatin (p-N-blebb; Fig. 3, E and L; **, P < 0.01), inhibition of myosin light chain kinase (MLCK) with ML-7 (Fig. 3 K), and actin depolymerization with cytochalasin D (CytoD; Fig. 3, G and M; ***, P < 0.001). Conversely, microtubule depolymerization with nocodazole did not affect cyclin B1 translocation (Noc; Fig. 3 H; n.s., not significant). Importantly, confinement was able to fully rescue cyclin B1 accumulation that was lost upon expression of DN-KASH (Fig. 3, B and I; ***, P < 0.001) or treatment with Y27632 (Fig. 3, D and J), and partially rescued accumulation following actin (Fig. 3, G and M; ***, P < 0.001) or myosin (Fig. 3, E and L; **, P < 0.01) inhibitions. Overall, these experiments identify actomyosin-dependent force transmission to the nucleus as important players in facilitating cyclin B1 nuclear translocation.

Our results indicate that during the G2-M transition, the NE unfolds an event that can be exacerbated by mechanical confinement (Fig. S1). Moreover, we showed that actomyosin contractility facilitates the translocation of cyclin B1 into the nucleus (Fig. 3). Since increased actomyosin contractility was recently shown to induce NE unfolding (Lomakin et al., 2020; Venturini et al., 2020), we tested whether such a mechanism also acted during the G2-M transition. To do so, we evaluated the nuclear irregularity index (NII) of interphase and prophase nuclei using fixed-cell analysis. This parameter was calculated as 1-solidity (solidity is defined as nucleus area/nucleus convex area). Our results confirm a decrease in NII in prophase cells, when compared to interphase, indicating an unfolding of the NE (Fig. 4, A–C; ***, P < 0.001). Nuclear unfolding was previously associated with increased nuclear tension (Enyedi et al., 2016), which triggers the recruitment and activation of the calcium-dependent, nucleoplasmic phospholipase cPLA2 to the NE. Active cPLA2 was proposed to stimulate actomyosin contractility, creating a positive feedback loop (Lomakin et al., 2020; Venturini et al., 2020). If indeed prophase nuclei are under increased tension, it is possible that cPLA2 is recruited to the NE at this stage. Accordingly, we found that cPLA2 is recruited to the NE during prophase, similarly to what happens in confined interphase cells (Lomakin et al., 2020; Venturini et al., 2020; white arrows, Fig. 4, D and E; ***, P < 0.001), suggestive of increased NE tension and cPLA2 activation at this stage. While our data show that NE unfolding during prophase can recruit cPLA2, it does not explain how the NE unfolds in the first place. To determine this, we analyzed the NII of cells expressing DN-KASH or treated with actomyosin inhibitors. All treatments led to an increase in NII, which was reverted upon confinement (Fig. 4, F and G; ***, P < 0.001). This confinement-generated decrease in NII likely reflects an unfolding of the NE, which is evident from the images of confined nuclei (Fig. 4, F and L) as well as the increased NPC-NPC distance (Fig. S1; ***, P < 0.001). Overall, this suggests that actomyosin activity transmits forces to the NE through the LINC complex, leading to nuclear unfolding. To further characterize the mechanism behind nuclear unfolding and cPLA2 recruitment, we then inhibited cPLA2 activity with AAOCF3 and analyzed the changes in NII. Indeed, interfering with cPLA2 activity significantly increased NII (Fig. 4 H; ***, P < 0.001), although to a lesser extent than actomyosin inhibition (0.024 ± 0.019 for AAOCF3, 0.035 ± 0.015 for Y27632; P = 0.007 and 0.033 ± 0.016 for p-N-blebb; P = 0.019). As expected, inhibition of cPLA2 also led to a decrease of its association with the NE, even after confinement (Fig. 4, I and J; ***, P < 0.001). Importantly, confinement was able to induce nuclear unfolding and increase NPC distance, even after cPLA2 inhibition (Fig. 4, I–K; ***, P < 0.001). It should be noted that AAOCF3 treatment alone does not decrease NPC distance below those observed for control cells (Fig. S1 K and Fig. 4 K; 0.350 ± 0.05 μm for controls vs. 0.364 ± 0.08 μm for AAOCF3-treated cells).

So far, our results indicate that actomyosin activity leads to increased nuclear unfolding and cPLA2 NE recruitment. Whether this is reflective of increased nuclear tension, it remained unknown. For that purpose, we sought to interfere with nuclear tension, independently of actomyosin activity, by overexpressing lamin B receptor (LBR). Overexpression of LBR is known to cause NE folding (Gravemann et al., 2010), decrease nuclear tension, and significantly affect cytoplasmic calcium levels and production of arachidonic acid, key components of the cPLA2 signaling pathway (Lomakin et al., 2020). As expected, overexpression of LBR resulted in a significant increase in NII (Fig. 4, H and I; ***, P < 0.001) and a consequent decrease in the levels of cPLA2 on the NE (Fig. 4, I and J; ***, P < 0.001). Collectively, these data indicate that actomyosin contractility triggers NE unfolding and increased tension during prophase. This was further confirmed by blocking actomyosin activity or expressing DN-KASH, which significantly decreased cPLA2 accumulation on the NE (Fig. 4, F, L, and M; ***, P < 0.001), and could be rescued by confinement (Fig. 4, L and M; ***, P < 0.001). Taken together, this indicates that an intact connection between the cytoskeleton and nucleus is required to set off a tension-dependent signal that results in cPLA2 recruitment to the NE.

Overall, we concluded that an increase in actomyosin contractility during prophase is required to unfold the NE, increasing nuclear tension, and leading to cPLA2 recruitment. The question remains of the functional relevance of cPLA2 NE recruitment. If cPLA2 is functionally important to facilitate cyclin B1 translocation, inhibiting its activity should result in a delay in cyclin B1 nuclear accumulation. Indeed, inhibition of cPLA2 activity with AAOCF3 led to a significant decrease in cyclin B1 nucleoplasmic shuttling (Fig. 5, A and B; **, P < 0.01; ***, P < 0.001). To further understand how cPLA2 interfered with cyclin B1 translocation, we analyzed the levels of CDK1 Y15 phosphorylation and cyclin B1, following cPLA2 inhibition. Phosphorylation of CDK1 on T14/Y15 is well known to prevent mitotic entry (Parker et al., 1992; Kornbluth et al., 1994; Mueller et al., 1995). We reasoned that if cPLA2 affected the pathway controlling CDK1 activation, we should see an increase in this inhibitory phosphorylation, following treatment with AAOCF3. Accordingly, treatment with AAOCF3 increased the levels of CDK1 Y15, when compared to controls for both synchronized (1 for controls vs. 1.7 for AAOCF3-treated cells) and asynchronous cells (1 for controls vs. 1.7 for AAOCF3-treated cells), but not for CDK1i cells (1 for CDK1i vs. 0.9 for CDK1i + AAOCF3), without affecting total CDK1 levels (Fig. S4 D). These results suggest that inhibition of cPLA2, through a yet unknown mechanism, delays activation of the cyclin B1-CDK1 complex, which could partly explain the delay in nuclear translocation of cyclin B1 observed upon treatment with AAOCF3 (Fig. 5, A and B).

Since the release of internal Ca²⁺ stores triggers cPLA2 NE recruitment and activation (Enyedi et al., 2016), we next decided to acutely interfere with the release of Ca²⁺ using BAPTA-AM + 2APB. Strikingly, this treatment also decreased cyclin B1 nuclear translocation (Fig. 5, C and D; ***, P < 0.001), as anticipated. While we cannot rule out that interfering with calcium release might affect other cellular processes, the use of BAPTA to disrupt internal calcium release during prophase has been previously described (Kao et al., 1990) and should not affect NEP at the concentrations used in our study. Moreover, we sought to minimize possible side effects by adding the drugs acutely in late G2. Similarly to the BAPTA treatment, decreasing nuclear stiffness by overexpression of LBR also significantly delayed cyclin B1 nuclear translocation (Fig. 5, E and F; ***, P < 0.001) and decreased cPLA2 loading on the NE (Fig. 4, I and J). Remarkably, confinement was able to stimulate cyclin B1 translocation, when cPLA2 activity or calcium release was inhibited (Fig. 5, A–D; ***, P < 0.001). This likely occurs due to confinement-induced unfolding of the NE (Fig. S1), that is sufficient to bypass the pharmacological inhibition of contractility and still induce an increase in NPC distance (Fig. 5, G and H; ***, P < 0.001). Together, these observations support a working model for the mechanical regulation of mitotic entry based on actomyosin activity, that triggers nuclear unfolding and increases tension on the nucleus, leading to cPLA2 activation. This facilitates cyclin B1 transport across the NPCs, increasing its nuclear accumulation.

Nuclear translocation of cyclin B1 sets the time for the G2-M transition (Strauss et al., 2018) and is essential for preventing untimely mitotic entry, which results in chromosome segregation errors (Furuno et al., 1999). Similarly, confining cells throughout mitosis also contribute to the occurrence of segregation errors (Tse et al., 2012; Lancaster et al., 2013). Whether a short confinement during prophase only, which is sufficient to induce premature cyclin B1 translocation and NEP, results in chromosome segregation errors remains unknown. To test this, we subjected cells in prophase to a short confinement, which was released shortly after NEP (Fig. 6 A). This approach should induce mitotic entry, while still providing enough volume for the spindle to assemble unconstrained (Lancaster et al., 2013). Cells were then allowed to progress through mitosis unperturbed, so that we could determine mitotic timings, as well as the rate of chromosome missegregation. Notably, a significant proportion of cells that were subjected to short confinement entered mitosis with incomplete centrosome separation (Fig. 6, A–D). This condition has been shown, by us and others, to increase the frequency of mitotic errors, in particular the occurrence of lagging chromosomes (Silkworth et al., 2012; Nunes et al., 2020), by favoring the establishment of erroneous kinetochore-microtubule attachments. Importantly, our acute confinement resulted in increased chromosome segregation errors (Fig. 6 E; *, P < 0.05) and a slight mitotic delay (Fig. 6 F; 24 ± 7 min for controls vs. 36 ± 20 min for confined cells; *, P < 0.05), when compared to unconfined cells. We propose these mitotic errors are triggered by the acute confinement that accelerates NEP, before cells had time to organize a mitotic spindle. To test this, we decided to generate an artificial rupture of the NE with laser microsurgery (Schweizer et al., 2015), allowing cyclin B1 and tubulin to enter the nuclear space and anticipating mitotic entry (Fig. 6 G). Using this approach, we triggered immediate mitotic entry, which was sufficient to increase chromosome missegregation events (white arrowhead, Fig. 6, G and H; **, P = 0.02) and induce a slight mitotic delay (Fig. 6 I). Together, these experiments demonstrate that untimely mitotic entry through acute mechanical confinement during the G2-M transition can have deleterious downstream consequences for chromosome segregation.

The biochemical regulation of the G2-M transition has been extensively studied (Pines and Hunter, 1989; Li et al., 1997; Hagting et al., 1998; Hagting et al., 1999; Toyoshima et al., 1998; Gavet and Pines, 2010). A master regulator of this transition is the complex composed of cyclin B1-CDK1, whose activity must be tightly regulated. During interphase, cyclin B1 is expressed at low levels and localizes mainly to the cytoplasm (Hagting et al., 1998; Toyoshima et al., 1998). As cells transit from S to G2, cyclin B1 levels progressively increase (Akopyan et al., 2014; Feringa et al., 2016; Pines and Hunter, 1989), leading to cyclin binding to CDK1 in late G2. Once this occurs, the complex is inactivated by Myt1- and Wee1-mediated phosphorylation of CDK1 on residues T14 and Y15 (Lindqvist, 2010). When these inhibitory phosphorylations are removed by Cdc25 phosphatases, the complex becomes activated, rapidly stimulating its own nuclear import (Gavet and Pines, 2010). In addition to the removal of inhibitory phosphorylations on CDK1, additional mechanisms ensure the nuclear translocation of cyclin B1. These involve phosphorylation of the CRS sequence in cyclin B1 (Li et al., 1997; Hagting et al., 1999) and binding to importin β in a Ran-independent manner (Moore et al., 1999; Takizawa et al., 1999). Once inside the nucleus, the cyclin B1-CDK1 complex then triggers NPC disassembly by phosphorylating nucleoporin Nup53 (Linder et al., 2017) and is involved in the phosphorylation and subsequent disassembly of the nuclear lamina (Heald and McKeon, 1990). This mechanism allows a fast redistribution of the cyclin B1–CDK1 complexes between the cytoplasm and the nucleus, ensuring the spatiotemporal coordination of all necessary steps leading up to mitotic entry.

So far, the contribution of mechanical forces for this essential step of the cell cycle was little explored. Recent evidence proposed that during the G2-M transition, mechanical stretch activated Piezo 1, ultimately leading to cyclin B1 transcription and consequent mitotic entry (Gudipaty et al., 2017). Here, we propose a nongenetic, mechanical pathway based on nuclear tension that acts during the G2-M transition, impacting cyclin B1 translocation and NEP. In agreement with this model, we showed that this process requires force transmission to the nucleus through the LINC complex (Fig. 3, A, B, and I). Interestingly, the LINC complex was previously proposed to play a role in early spindle assembly, by facilitating chromosome alignment in a myosin II-dependent manner (Booth et al., 2019) and assisting in centrosome positioning (Stiff et al., 2020). Our work now proposes an additional role for the LINC complex- and actomyosin activity in facilitating cyclin B1 translocation (Fig. 3). In addition, we also demonstrated that decreasing nuclear tension by overexpression of LBR significantly impaired nuclear cyclin B1 uptake. Taken together, these observations suggest that nuclear force transmission and sensing could be an important player for early mitotic events. One key aspect that remains to be determined is how these forces crosstalk with the biochemical pathways that control cyclin B1 translocation. Our data clearly show that both importin β (Fig. 2 G) and CRS phosphorylation (Fig. S3, C and D) are essential for force-mediated cyclin B1 translocation. Moreover, inhibition of cPLA2 increases the levels of inactive CDK1 (Fig. S3 L). Therefore, while forces seem to accelerate nuclear entry of cyclin B1, they cannot bypass the biochemical requirements for cyclin B1–CDK1 translocation. How and when physical forces might affect the biochemical pathways regulating the G2-M transition is an interesting question for future studies.

Physical forces acting on the nucleus can trigger NE unfolding and increase nuclear tension (Lomakin et al., 2020; Venturini et al., 2020). Here, we demonstrate that nuclear unfolding is a process that normally occurs in prophase cells (Figs. 4 and S1), similarly to previous observations in G2 cells (Lomakin et al., 2020). This unfolding increases cPLA2 recruitment to the NE (Fig. 4, D and E), indicative of cPLA2 activation (Enyedi et al., 2016; Lomakin et al., 2020; Venturini et al., 2020) and likely reflects higher nuclear tension during the G2-M transition. So how does this increased tension affect cyclin B1 transport? Recent work showed that imposing forces on the nucleus is sufficient to drive nuclear import (Andreu et al., 2022) and decrease the restriction to nuclear transport, due to NPC deformation (Elosegui-Artola et al., 2017). It is tempting to speculate that during the G2-M transition, this increased nuclear tension could be sufficient to deform the nucleus and NPCs, leading to faster cyclin B1 transport across the NE, as was proposed for YAP or MyoD (Elosegui-Artola et al., 2017; Jacchetti et al., 2021). In fact, a recent report showed that NPCs can deform in vivo, and facilitate transport across the NE (Zimmerli et al., 2021). Such a model, based on force-induced modifications in the stringency of NPCs could also explain how cyclin B1 enters the nucleus even when CDK1 is inhibited (Gavet and Pines, 2010) and why CDK1-inhibited cells show only partial rescue of nuclear cyclin B1 under confinement (Fig. 2 F). Overall, we propose this mechanical pathway cooperates with the classical cyclin B1 transport machinery to fine-tune NEP according to the cellular tension state, thus ensuring timely and accurate cell division.

Establishing mechanical forces as an important player in cyclin B1 nuclear translocation and mitotic entry raises the interesting possibility that the nucleus might act as a sensor (Lomakin et al., 2020; Venturini et al., 2020) for external forces, regulating cell-cycle progression and cell division to control tissue growth and avoid over-proliferation.

Cell lines used were cultured in DMEM (Gibco, Life Technologies) supplemented with 10% FBS (Gibco, Life Technologies) and kept in culture in a 37°C humidified incubator with 5% CO₂. RPE-1 parental, and RPE-1 cell lines expressing exogenous H2B-GFP and tubulin-mRFP were already available in our lab. The RPE-1 cell line expressing endogenously tagged cyclin B1–Venus was a gift from Jonathon Pines (Collin et al., 2013). The RPE-1 cell line expressing endogenous cyclin B1–Venus/tubulin-mRFP was generated in our lab by transduction with lentiviral vectors containing pRRL-mRFP-α-tubulin, as previously described (Nunes et al., 2020). In brief, HEK293T cells at a 50–70% confluence were co-transfected with lentiviral packaging vectors (16.6 μg of Pax2, 5.6 μg of pMD2, and 22.3 μg of LV-tubulin-mRFP), using 5 μl of Lipofectamin 2000 (Life Technologies). Approximately 4–5 d after the transduction, the virus-containing supernatant was collected, filtered, and stored at −80°C. The RPE-1 cyclin B1–Venus cells were infected with virus particles together with polybrene (1:1,000) in standard culture media for 24 h. Approximately 2–3 d after the infection, the cells expressing tubulin-mRFP were isolated by fluorescence-activated cell sorting (FACS; FACS Aria II). To deplete ROCK1, a shRNAi-LV vector (kind gift from João Relvas, i3S, Porto, Portugal) was co-transfected using a lentiviral packaging vector, as explained above. The viral particles were then transduced in the RPE-1 Tub-mRFP cell line to generate stable cells depleted of ROCK1. The RPE-1 cell line expressing exougenous cGAS-GFP was a kind gift from Matthieu Piel (Institut Curie, Paris, France). The RPE-1 cell line expressing GFP-NLS/tubulin-mRFP was created by lentiviral transduction using a pCDH-NLS-copGFP-EF1-BlastiS plasmid (#132772; Addgene), as described above. The RPE-1 cell lines expressing exogenous LBR-mCherry and endogenous cyclin B1–Venus/LBR-mCherry were generated by lentiviral transfection as described above, with the plasmid pWPT LBR-mCherry (kind gift from Stephen Royle, Warwick Medical School, Warwick, UK).

CDK1 inhibitor (RO-3306; sc-358700A; Santa Cruz Biotechnology) was used at a concentration of 9 μM for 16 h. Importazole (kind gift from Helder Maiato) was added to the cells at a final concentration of 40 μM 2 h before the experiment, ROCK inhibitor (Y-27632; cat. no. Y0503; Sigma-Aldrich) was used at 5 μM for 30 min. To interfere with cPLA2 activity, AACOCF3 was used at 20 μM (cat. no. 1462; TOCRIS) for 30 min. To block the release of calcium ions from internal cellular stores, BAPTA-AM and 2APB (ab120503 and ab120124, respectively; Abcam) were used at 10 μM for 15–30 min. Myosin activity was perturbed using p-nitro-blebbistatin at 50 μM for 30 min (cat. no. 1621326-32-6; MotorPharma). MLCK activity was blocked using ML-7 (I2764; Merck) at 50 μM for 30 min. To interfere with the actin cytoskeleton, we used cytochalasin D at 0.5 μM (TO-1233; Biogen Científica) for 30 min. To perturb microtubules, nocodazole (M1404; Merck) was used at 3.3 μM for 30 min. Plk1 inhibitor (BI2536; AXON 1129; Axon Med Chem) was used at 200 mM for 2 h. To induce DNA damage, cells were treated with 1 μM of Etoposide (S1225; Selleck Chemicals CO.) for 2 h before fixation. Cell synchronization was performed by incubating with 2 mM thymidine (T1895; Sigma-Aldrich) for 16 h, followed by a 10 h release. A second block was performed for another 16 h, followed by a 6–8 h release. Control cells were treated with either DMSO (D4540; Sigma-Aldrich) or mock transfected with Lipofectamin 2000 (cat. no. 11668019; Invitrogen), as explained in the text.

To perform the hypotonic shock, RPE-1 cells expressing cyclin B1–Venus and tubulin-mRFP were seeded as described above. When cells were at the microscope, MiliQ water was added to the imaging medium (1:5 dilution).

Cells were transfected with the plasmids encoding the DN-KASH or the Rap1Q63E (Rap1*) mutant using Lipofectamin 2000. Specifically, 5 μl of Lipofectamin 2000 and 0.5 μg of plasmid DNA were diluted separated and incubated in OPTIMEM (Alfagene) for 30 min. The mixture was then added to confluent cells cultured and incubated for 6 h in reduced serum medium (DMEM with 5% FBS). Cells were analyzed 48 h after transfection.

Between 12 and 24 h before the experiments, 1.5 × 10⁵ cells were seeded on fluorodishes (WPI) coated with FBN (25 μg/ml; F1141; Sigma-Aldrich). Shortly before each experiment, DMEM 10% FBS medium was changed to Leibovitz’s-L15 medium (Life Technologies), supplemented with 10% FBS and Antibiotic-Antimycotic 100X (AAS; Life Technologies). Live-cell imaging experiments were performed at 37°C, using temperature-controlled Nikon TE2000 microscopes equipped with a modified Yokogawa CSU-X1 spinning-disk head (Yokogawa Electric), an electron multiplying iXon + DU-897 EM-CCD camera (Andor) and a filter wheel. Three laser lines were used to excite 488, 561, and 647 nm, and all the experiments were done with an immersion oil, 60× 1.4NA Plan-Apo DIC objective (Nikon). Image acquisition was controlled by NIS Elements AR software. Images with 17–21 z-stacks (0.5 μm step) were collected with a 20 s interval.

Cell extracts were collected after trypsinization and centrifuged at 1,200 rpm for 5 min, washed and resuspended in 30–50 μl of lysis buffer (20 nM HEPES/KOH, pH 7.9, 1 mM EDTA, pH 8, 150 mM NaCl, 0.5% NP40, 10% glycerol, and 1:50 protease inhibitor; 1:100 Phenylmethylsulfonul fluoride). The cells were then flash-frozen in liquid nitrogen and kept on ice for 30 min. After centrifugation at 14,000 rpm for 8 min at 4°C, the supernatant was collected, and protein concentration determined using the Bradford protein assay (Bio-Rad). The proteins were run on a 10% SDS-PAGE gel (50 μg/lane for all blots) and transferred using a wet blot apparatus for 1.5 h at 70 V, with constant amperage. For detection of total CDK1, the proteins were run on a 10% SDS-PAGE gel (5 μg/lane) and transferred using the Trans-Blot Turbo Transfer System blot apparatus for 10 min. Later, the membranes were blocked with 5% milk in Tris-buffered saline (TBS) with 0.1% Tween-20 (TBS-T) for 1 h at room temperature (RT). The primary antibodies used were rabbit anti-vinculin (1:1,000, cat. no. 700062; Thermo Fisher Scientific), rat anti-tyrosinated α-tubulin (1:500, MCA77G; Bio-Rad), mouse anti-cyclin B1 (1: 1,000; cat. no. 4135; Cell Signaling Technology), mouse anti-p34/Cdc2 (B-6; 1:500; cat. no. sc-8395; Santa Cruz Biotechnology), and rabbit anti-phospho Cdc2 Tyr15 (1:1,000; cat. no. 4539T; Cell Signaling Technology). All primary antibodies were incubated overnight at 4°C with shaking. After three washes in TBS-T, the membranes were incubated with the secondary antibody for 1 h at RT. The secondary antibodies used was anti-rabbit-HRP at 1:5,000 (cat. no. 111-005-003; Jackson Immuno Research) or 1:2,500 when detecting total CDK1. After several washes with TBS-T, the detection was performed with Clarity Western ECL Substrate (cat. no. 1705060; Bio-Rad), using a Bio-Rad ChemiDoc XRS + imaging system.

For dynamic confinement experiments, we adapted a cell confiner as previously described (Le Berre et al., 2014), using a custom-designed polydimethylsiloxane (PDMS, RTV615, GE) layout to fit a 35-mm fluorodish. A suction cup was custom-made with a 10/1 mixture (w/w PDMS A/crosslinker B), baked on an 80°C hot plate for 1 h and left to dry overnight before unmoulding. The confinement slide was polymerized on 10 mm round coverslips. These round coverslips were first treated with air plasma for 2 min (Zepto system, Diener Electronics) and incubated with a 0.3% Bind-Silane (M6514; Sigma-Aldrich)/5% acetic acid solution in ethanol. Then, the coverslips were rinsed with ethanol and left to dry. A gel with ∼15 kPa stiffness was prepared using an acrylamide (Bio-Rad)/bisacrylamide (Bio-Rad) mix. The mixture was added to the coverslips and allowed to polymerize. After polymerization, gels were hydrated with PBS and incubated with cell culture medium for at least 30 min. The confinement slide was then attached to the PDMS suction cup described above and connected to a vacuum generator apparatus (Elveflow).

For static confinement experiments, we used a commercially available 6-well confinement device (4DCell) with custom-designed confinement slides. The confinement slide was polymerized in PDMS on a round 10-mm standard microscope coverslip and designed with a regular holes array (diameter 449 μm, 1 mm spacing). Briefly, after activating the coverslip in a plasma chamber (Diener Electronics) for 2 min, this coverslip then was used to press a PDMS drop on top of the wafer, to obtain a thin layer. After baking at 95°C on a hot plate for 15 min, excess PDMS was removed. Isopropanol was used to peel off the glass slide with the PDMS pillars from the wafer. Microfabricated coverslips with confining pillars (8 μm height) were then attached to PDMS spacers that were stuck on a 6-well plate lid (4DCell).

For CH-STED microscopy, cells were grown as described above. Parental RPE-1 cells were seeded in the day before the experiment in coverslips coated with FBN. After fixation with 4% paraformaldehyde in cytoskeleton buffer, the cells were extracted in PBS with 0.5% Triton-X100 (Sigma-Aldrich). The coverslips were incubated with the primary antibodies (rabbit anti-TPR, 1:100, NB100-2867; and mouse anti-NUPs, 1:100, 24609; Abcam) in blocking solution overnight at 4°C. After washing with PBS-0.1% Triton-X, the coverslips were incubated with a 1:100 dilution of secondary antibodies (Abberior anti-rabbit STAR 580, cat. no. 2-0012-005-8, and Abberior anti-mouse STAR 635P, cat. no. 2-0002-007-5) at RT for 1 h. Later, coverslips were washed in PBS with 0.1% Triton-X100 and sealed on a glass slide using mounting medium (20 nM Tris pH 8, 0.5 N-propyl gallate, and 90% glycerol).

The images were acquired with an Abberior Instruments “Expert Line” gated-STED coupled to a Nikon Ti microscope. For all the acquisitions, we used an oil-immersion, 60× 1.4NA Plan-Apo objective (Nikon, Lambda Series) and pinhole size of 0.8 Airy units. The CH-STED technique creates an orthogonal direction on the STED parametric space that enables the independent tuning of both resolution and contrast using only one depletion beam in a standard STED setup (circular polarization based).

For the immunofluorescence experiments, cells were grown as previously described. RPE-1 parental cells were seeded in the day before the experiment in coverslips coated with FBN. After fixation with 4% paraformaldehyde in cytoskeleton buffer (274 mM NaCl, 2.2 mM Na₂HPO₄, 10 mM KCL, 0.8 mM KH₂PO₄, 4 mM EDTA, 4 mM MgCl₂, and 10 mM glucose, pH 6.1), cells were extracted with PBS-0.5% Triton-X100 (Sigma-Aldrich) following three washes (5 min each) with PBS-0.1% Triton-X100 and a 30 min incubation in blocking solution (10% FBS in 10% Triton-X100 in PBS). The coverslips were incubated with the primary antibodies (rabbit anti-cPLA2, 1:100, #2832; Cell Signaling; mouse anti-LaminA/C, 1:500, ABCAM ab8994; rat anti-tyrosinated α-tubulin 1:500, MCA77G; Bio-Rad; mouse γ-H2AX, 1:2,000, 05-636; Milipore) in blocking solution for 1 h at RT. After washing with PBS-0.1%Tríton-X for 5 min, the coverslips were incubated with the secondary antibodies (AlexaFluor 488, 568 and 647, 1:2,000; Invitrogen) at RT for 1 h. Later, coverslips were washed, three times, in PBS with 0.1% Triton-X100 and once with PBS. Images were acquired using an AxioImager z1 with a 63×, Plan oil DIC objective lens, 1.4NA (from Carl Zeiss), coupled with a CCD camera (ORCA-R2; Hamamatsu Photonics) and the Zen software (Carl Zeiss).

For the quantifications of cyclin B1 levels, images were analyzed using ImageJ. A small square region of interest (ROI) was defined, and cyclin B1 fluorescence intensity measured, throughout time in the cell nucleus. The same ROI was used to measure the background outside the cell area. All fluorescence intensity values were then background-corrected and the values were normalized to the lowest nuclear cyclin B1 level. To obtain a measure of cyclin B1 translocation rate, cyclin B1 fluorescence values were aligned relative to the lowest value, which was defined as time zero. In alternative, normalized fluorescence intensity values were normalized to the time of NEP.

For quantification of cPLA2 fluorescence intensity on the NE, images were analyzed using ImageJ. A defined ROI was used to measure the fluorescence intensity values in five different regions outside the cells, which was then used to calculate the average background levels. Afterwards, a different ROI was used to measure fluorescence intensity in the nucleoplasm and at the NE. Images were background-subtracted and integrated fluorescence density was measured using the ImageJ Measure tool. cPLA2 enrichment at the NE was calculated by obtaining ratio between fluorescence in the NE and in nucleoplasm.

NII was used to estimate the overall folding of the nucleus. To do so, we first obtained 2D images of the medial section of the nucleus using an anti-Lamin A/C antibody. These images were processed to obtain the nuclear area and convex area using ImageJ. Nuclear solidity was then calculated as area/convex area. Nuclear Irregularity Index was defined as 1-nuclear solidity.

A computational algorithm was developed in MATLAB (v2018b; The MathWorks, Inc.) to quantify compression-induced topological changes in the nuclear pores, within the nuclear membrane. For the analyses performed in this study, we used a method focused on estimating changes on the average inter-distance between TPR and a mixture of proteins that compose the NPC, respectively, tagged with Abberior anti-rabbit STAR 580 and anti-mouse STAR 635P. Spatial periodicity on staining (either TPR or NPC) was estimated through a spatial Fast-Fourier Transform (FFT) operation on membrane cross-section images. As an alternative, and as validation, intensity profile autocorrelation was also used to assess the spatial periodicity. The entire length of the nuclear membrane was previously scanned to select only segments with low curvatures and the “Straighten” tool (Schindelin et al., 2012) was used prior to the FFT/autocorrelation operations.

To perform a detailed quantitative analysis of centrosome positioning and movement and cell rounding, we used a previously described, custom-designed MATLAB (v2018b; The MathWorks Inc) script (Nunes et al., 2020). The algorithm used as a specific workflow designed for centrosome tracking in a 3D space having in consideration a pixel size of 0.176 μm and a z-step of 0.5 μm. The nucleus shape was reconstructed using H2B-GFP as marker, and cell shape was reconstructed using tubulin-RFP as a marker. Using this tool, we were able to correlate the angle between the centrosomes and the nucleus, as well as cell rounding during mitotic entry.

Laser microsurgery was performed with a doubled-frequency laser (FQ-500-532; Elforlight) coupled with an inverted microscope (TE2000U; Nikon), using a 100× 1.4NA, plan-apochromatic DIC objective lens and equipped with an iXonEM + EM-CD camera (Andor Technology). To induce a break on the NE, we used 8 consecutive pulses, with a pulse energy of 3–5 μJ and an interval of around 10 ns.

Three to six independent experiments were used for statistical analysis. When data are represented as box-whisker plots, the box size represents 75% of the population and the line inside the box represents the median of the sample. The size of the bars (whiskers) represents the maximum (in the upper quartile) and the minimum (in the lower quartile) values. Normality of the samples was assessed using the Kolmogorov–Smirnov test. Statistical analysis for multiple group comparison was performed using a parametric one-way ANOVA when the samples had a normal distribution. Otherwise, multiple group comparison was performed using a nonparametric ANOVA (Kruskal–Wallis). Multiple comparisons were analyzed using either post-hoc Student-Newman-Keuls (parametric) or Dunn’s (nonparametric) tests. When only two experimental groups were compared, we used either a parametric two-sided t test or a nonparametric Mann–Whitney test. Comparison for multiple time-course datasets was carried out using an ANOVA Repeated Measures, when the samples had a normal distribution. Otherwise, group comparison was carried out using Repeated Measures ANOVA on Ranks. No power calculations were used. All statistical analyses were performed using SigmaStat 3.5 (Systat Software, Inc.).

Fig. S1 contains the schematics of the dynamic cell confiner device and shows data regarding the integrity of the NE following confinement. Fig. S1 also shows evidence of nuclear unfolding upon confinement using live-cell imaging and CH-STED super resolution. Fig. S2 shows that cyclin B1 translocation is sensitive to the stiffness of the external environment. Fig. S2 also shows that cyclin B1 translocation is sensitive to alternative mechanical stimulation such as a hypotonic shock and demonstrates that mechanical stimulation of cyclin B1 translocation is maintained in HeLa cells. Fig. S3 shows that cyclin B1 translocation is insensitive to mitotic cell rounding and that expression of cyclin B1-5A-GFP blocks cyclin B1 translocation even under confinement. Fig. S3 also shows the effects of Plk1 inhibition on cyclin B1 translocation. Fig. S4 shows expression of DN-KASH-RFP and compares cyclin B1 translocation in two different cell lines expressing DN-KASH-RFP. It also shows the increase in phospho-CDK1 (Y15) levels after AAOCF3 treatment.

All custom-designed computational tools used in this manuscript are available from the corresponding author upon reasonable request.

The data that support the findings included in this manuscript are available from the corresponding author upon reasonable request. All cell lines and reagents generated for this study are available from the corresponding author upon request.

The authors would like to thank Jonathon Pines (The Institute of Cancer Research, London, UK) for the gift of the RPE-1 cyclin B1–Venus and HeLa cyclin B1–Venus cell lines and the plasmid for expression of cyclin B1-5A-GFP. The authors thank Matthieu Piel for the RPE-1 cGAS-GFP cell line. The authors thank Stephen Royle for the plasmid pWPT LBR-mCherry. The authors thank Joana Lima for performing the western blot to detect CDK1 levels in Fig. S4. The authors thank Dr. Buzz Baum, Dr. Alexis Lomakin, and members of the Ferreira and Maiato labs for critical reading of the manuscript.

This work was funded by Portuguese funds through FCT—Fundação para a Ciência e a Tecnologia/Ministério da Ciência, Tecnologia e Ensino Superior in the framework of the project PTDC/BIA-CEL/6740/2020. M. Dantas is supported by grant PD/BD/135548/2018 from the BiotechHealth FCT-funded PhD program. Work in the Maiato lab is funded by the European Research Council (ERC) consolidator grant CODECHECK, under the European Union’s Horizon 2020 research and innovation program (grant agreement 681443), Fundação para a Ciência e a Tecnologia of Portugal (PTDC/MED-ONC/3479/2020), and the NORTE-01-0145-FEDER-000051 project supported by NORTE 2020 under the PORTUGAL 2020 Partnership Agreement through the European Regional Development Fund.

The authors declare no competing financial interests.

Author contributions: M. Dantas performed experimental work, analyzed data, prepared figures, and jointly wrote the manuscript. J.G. Ferreira provided the conceptual framework, analyzed data, prepared figures, secured funding, and jointly wrote the manuscript. H. Maiato provided access to essential equipment, reviewed, and edited the manuscript. A. Oliveira and P. Aguiar developed MATLAB computational tools, reviewed and edited the manuscript.