Cyclin B1–Cdk1 is active in both the nucleus and the cytoplasm in prophase

Because activating cyclin B1–Cdk1 immediately triggered its import into the nucleus, it was possible that this would deplete active cyclin B1–Cdk1 from the cytoplasm until NEBD. To test this, we compared the activation of cyclin B1–Cdk1 in the cytoplasm and the nucleus using modified versions of the cyclin B1–Cdk1 biosensor. We targeted the biosensor to the cytoplasm by tagging it with an NES and targeted it to chromosomes by fusing it with histone H2B (Fig. S1 A). Fluorescence loss in photobleaching and FRAP assays confirmed that the H2B fusion protein was stably attached to chromosomes and did not diffuse into the cytoplasm (Fig. S1 B and not depicted). Monitoring dividing cells by time-lapse microscopy showed that the change in FRET efficiency of the biosensor was very similar between the cytoplasm and the nucleus in all cells analyzed, indicating that cyclin B1–Cdk1 became progressively more active in both the cytoplasm and the nucleus as cells entered mitosis (n = 16 for three independent experiments; Fig. 3 and Video 3). These results agree with the conclusions of Lindqvist et al. (2007), who found by immunofluorescence analysis that some active cyclin B1–Cdk1 remains in the cytoplasm in prophase, whereas the rest moves into the nucleus. Thus, activation in the cytoplasm allied with the immediate and rapid nuclear import of a proportion of cyclin B1–Cdk1 inherently synchronizes mitotic events in the cytoplasm and the nucleus.

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Figure 3.

Active cyclin B1–Cdk1 kinase is located both in the cytoplasm and nucleus during prophase. HeLa cells coexpressing cytoplasmic and nuclear targeted cyclin B1–Cdk1 biosensors (fused to an NES and histone H2B, respectively) were recorded as cells entered mitosis (one image every 1 min 40 s). DIC (top), YFP emission (inverted grayscale; middle), and emission ratio (bottom) images are shown. Quantifications of the emission ratio in the whole cell, the nucleus, and the cytoplasm correspond to the cell displayed. Bar, 10 µm.

Cyclin B1–Cdk1 accumulation in the nucleus does not require Plk1 activity or phosphorylation on cyclin B1

The coupling between cyclin B1–Cdk1 activation and its rapid import into the nucleus appeared to be the key to synchronizing nuclear and cytoplasmic changes in cell architecture as cells entered mitosis, but the mechanism behind this was unclear. Our previous results indicated that it was caused by an increase in nuclear import generated by phosphorylation of the N terminus of cyclin B1 (Hagting et al., 1999), but other studies had concluded that phosphorylation, particularly phosphorylation by Plk1, inhibited the export of cyclin B1 (Yang et al., 1998; Toyoshima-Morimoto et al., 2001).

To establish the mechanism, we first addressed the importance of phosphorylation in the N terminus of cyclin B1. We compared the nuclear import in prophase of wild-type cyclin B1 with mutants carrying mutations to Ala in the autophosphorylation sites (Ser126 and 128), the consensus Plk1 site (Ser133), the NES (Ser147), or in all sites together (Ser116, 126, 128, 133, and 147). We first determined that the rate of cyclin B1–GFP import was not affected by the level of its expression (Fig. S2). When coexpressed in HeLa cells, the nuclear import in prophase of all the cyclin B1 mutants linked to EGFP was indistinguishable in timing or rate from wild-type cyclin B1–mCherry (Fig. 4 A, Fig. S3, and Video 4). Thus, neither autophosphorylation nor phosphorylation by Plk1 was required to trigger the nuclear accumulation of cyclin B1 in prophase. However, it was possible that Plk1 could trigger the nuclear accumulation of cyclin B1 by a mechanism separate from phosphorylating cyclin B1. Therefore, we tested the consequence of adding a potent Plk1 inhibitor (BI 2536) to cells in prophase. As shown in Fig. 4 B, inhibiting Plk1 had no effect on the nuclear accumulation of cyclin B1 (Fig. 4 B and Video 5), which is in agreement with a previous study (Lénárt et al., 2007). The potency of the inhibitor was confirmed because all cells were subsequently blocked in prometaphase after NEBD (Lénárt et al., 2007), and those cells in anaphase immediately failed cytokinesis when the inhibitor was added (Video 5; Burkard et al., 2007; Petronczki et al., 2007). Thus, we can conclude that Plk1 is not required to activate cyclin B1–Cdk1, for the nuclear import of cyclin B1, nor for the synchrony of nuclear and cytoplasmic changes in prophase.

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Figure 4.

Nuclear accumulation of cyclin B1 in prophase is independent of its phosphorylation and of Plk1 activity. (A) Cells coexpressing wild-type (WT) cyclin B1–mCherry and 5xA (Ser116, 126,128,133, and 147A)–cyclin B1–GFP were assayed, and the nuclear accumulation of the proteins was quantified. Mean curves of quantifications in different cells are displayed (n = 5). (B) Cells expressing wild-type cyclin B1–mCherry were assayed, and100 nM Plk1 inhibitor (BI 2536) was added during the nuclear import of cyclin B1 in prophase. Two examples are displayed (one image/minute). Error bars show SEM.

Previously, we and others have shown that substituting glutamic acids at the phosphorylation sites in the N terminus of cyclin B1 created a nuclear import signal and reduced the binding between cyclin B1 and the exportin Crm1 in vitro (Hagting et al., 1999; Yang et al., 2001). Therefore, we compared the behavior of these mutants with wild-type cyclin B1 through the cell cycle. First, we quantified optical sections obtained with a spinning-disk confocal microscope to measure their distribution between the cytoplasm and the nucleus in interphase. This showed that substitution with glutamic acids progressively displaced the equilibrium toward the nucleus, whereas replacing the Ser with Ala shifted the equilibrium toward the cytoplasm (Fig. 5 A).

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Figure 5.

The nuclear/cytoplasmic distribution of cyclin B1 is only affected by phosphorylation in its N terminus domain in interphase. (A) The nuclear/cytoplasmic ratio of wild-type (WT) cyclin B1–GFP, Ser126– and 128A–cyclin B1–GFP, Ser126–,128A–, and 133A–cyclin B1–GFP, 5xA (Ser116,126,128,133, and 147A)–cyclin B1–GFP, Ser126– and 128E–cyclin B1–GFP, Ser126–,128A–, and 133E–cyclin B1–GFP, and 5xE (Ser116,126,128,133, and 147E) cyclin B1–GFP was quantified on optical sections of asynchronous interphase HeLa cells. Mean values ± SEM and numbers of cells assayed are displayed. (B) Cells expressing either wild-type or 5xE cyclin B1–GFP were recorded. Real time changes in the nuclear/cytoplasmic ratio were quantified on optical sections as cells entered mitosis. Five examples are displayed for each experiment.

Next, we measured the change in distribution between the cytoplasm and nucleus of 5xE and wild-type cyclin B1 during prophase (Fig. 6 A). We observed that the timing and rate of nuclear accumulation were the same, although the degree of nuclear accumulation of the 5xE mutant was only ~60% of wild-type cyclin B1 (n = 11; Fig. 6 A). This might be a direct consequence of the difference in their equilibrium distribution between the cytoplasm and the nucleus in interphase (Fig. 5 A). Therefore, we measured the changes in the distribution between the cytoplasm and nucleus of wild-type and 5xE cyclin B1 on optical sections of cells progressing from G2 phase to mitosis (Fig. 5 B). Wild-type cyclin B1 changed from approximately five times more concentrated in the cytoplasm in late G2 phase to about two times more concentrated in the nucleus just before NEBD (cytoplasm/nucleus, ~0.4 ± 0.06; n = 6; Fig. 5 B, left). Thus, the cytoplasm still contained substantial levels of cyclin B1 at the end of prophase. Notably, the 5xE mutant cyclin B1 reached the same distribution just before NEBD (cytoplasm/nucleus, ~0.45 ± 0.07; n = 5; Fig. 5 B, right). Thus, we conclude that phosphorylation of the N-terminal region of cyclin B1 modulates its interaction with the import machinery, but this effect is only significant in interphase.

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Figure 6.

The nuclear accumulation of wild-type and 5xE mutant of cyclin B1 in prophase is concurrent. (A) The nuclear accumulation of cyclin B1 in prophase was assayed in cells coexpressing wild-type (WT) cyclin B1–mCherry and 5xE cyclin B1–GFP. DIC (top), mCherry (middle), and GFP (bottom) images are shown. The region of interest used for quantification of the nuclear signal is displayed. Mean curves of quantifications in different cells are displayed (n = 6). Bar, 10 µm. (B) Cells were assayed as in A, and 300 nM Cdk inhibitor (Cdk1/2 inhibitor III) was added during prophase. Two examples are displayed. Error bars show SEM.

Cyclin B1–Cdk1 activation stimulates its nuclear import rate

A previous study had reported that the nuclear accumulation of cyclin B1 was driven by phosphorylation in the NES at Ser147, inhibiting its nuclear export (Toyoshima-Morimoto et al., 2001). Therefore, we tested whether substituting Ser147 with glutamic acid inhibited cyclin B1 export at prophase. We added a Cdk inhibitor, as wild-type or the 5xE mutant cyclin B1 began to accumulate in the nucleus in prophase and observed that both forms of cyclin B1 were rapidly reexported and returned to the original equilibrium distribution with the same timing, provided that at the beginning of the experiment they had accumulated in the nucleus to a similar extent (Fig. 6 B, left). Wild-type cyclin B1 was exported more rapidly if its relative accumulation in the nucleus was higher than the 5xE mutant at the end of prophase (Fig. 6 B, right; and Video 6). Thus, the introduction of glutamic acids into the N terminus of cyclin B1 did not inhibit its nuclear export.

To determine whether cyclin B1–Cdk1 activation in prophase stimulated its nuclear import, inhibited its export, or both, we inhibited the Crm1 exportin with leptomycin B (LMB). In agreement with our previous results (Hagting et al., 1999), LMB induced a slow and progressive nuclear accumulation of wild-type cyclin B1 in G2 phase cells (Fig. 7 and Fig. S4), whereas under the same conditions, the 5xE mutant was imported about five times faster than wild type. This explained the difference observed in their relative nuclear/cytoplasmic ratios in interphase (Fig. 5 A). However, when cells treated with LMB reached prophase, the nuclear import rate of both wild-type and 5xE cyclin B1 suddenly increased by up to 40-fold (Fig. 7 and Fig. S4). The timing, rate, and extent of nuclear accumulation of cyclin B1 during prophase were very similar in the presence or absence of LMB (control cells, n = 26: increase in rate, 20–54-fold; mean, 38-fold; LMB-treated cells, n = 9: increase in rate, 24–55-fold; mean, 39-fold; Fig. S4). Thus, the nuclear accumulation of cyclin B1 in prophase is almost exclusively caused by a strong stimulation of its import rate.

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Figure 7.

Cyclin B1 nuclear import rate increases significantly at mitotic entry. Cells expressing either wild-type (WT) or 5xE cyclin B1–GFP were recorded, and the nuclear import of wild-type and 5xE cyclin B1 were quantified after treating G2 phase cells with 20 nM LMB to inhibit nuclear export. Note the sudden and strong increase of the wild-type cyclin B1 nuclear import rate when the cell enters mitosis.

Cyclin B1–Cdk1 activation regulates the nuclear transport machinery

Because the rapid rise in nuclear import of cyclin B1 in prophase was not caused by Plk1 or by phosphorylation of cyclin B1, we first investigated whether active cyclin B1–Cdk1 altered soluble import factors in prophase. Cyclin B1 binds directly to the N terminus of importin β1 (residues 1–462; Moore et al., 1999), and this domain contains a unique S/T-P sequence (Ser12) that is phosphorylated in mitotic cell extracts (Dephoure et al., 2008). Therefore, we asked whether this phosphorylation might induce the nuclear import of cyclin B1–Cdk1, but overexpressing wild-type importin β1 or mutants with this site changed to Ala or Glu did not alter the nuclear/cytoplasmic distribution of cyclin B1 in interphase (Fig. S5).

Several proteins of the nuclear pore complex are phosphorylated by cyclin B1–Cdk1 (Macaulay et al., 1995; Blethrow et al., 2008; for review see Kutay and Hetzer, 2008), making us consider the possibility that active cyclin B1–Cdk1 might modify the permeability (i.e., size exclusion limit) and/or functional properties of the nuclear pores. To assay this, we used two different markers: an importin β1–binding domain of importin α (IBB) and a Gly-rich sequence recognized by importin β2 (M9), both fused to GFP (Lénárt et al., 2003). We coexpressed IBB-GFP or GFP-M9 with wild-type cyclin B1–mCherry, assayed their localization by spinning-disk confocal microscopy, and quantified optical sections of cells as they entered mitosis. As shown in Fig. 8 A, IBB-GFP began to move into the cytoplasm in late prophase ~10 min before NEBD, most likely reflecting the increase in the size exclusion limit of nuclear pores (Lénárt et al., 2003), whereas cyclin B1 started to accumulate in the nucleus several minutes before this (cyclin B1, mean of 21 ± 5 min before NEBD; IBB, mean of 10 ± 1.5 min; n = 7; Video 7). Surprisingly, we observed a very similar timing between cyclin B1 accumulation in the nucleus and GFP-M9 export to the cytoplasm, although on average, cyclin B1 began to accumulate in the nucleus ~3 min before M9 began to move out (cyclin B1, mean of 26.5 ± 2 min before NEBD; M9, 24 ± 3 min before NEBD; n = 6; Fig. 8 B and Video 8).

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Figure 8.

The nuclear accumulation of cyclin B1–Cdk1 in prophase correlates with a change in nuclear transport machinery. (A and B) Cells coexpressing cyclin B1–mCherry and IBB-GFP (A) or GFP-M9 (B) were recorded as they entered mitosis (one image/30 s), and the import of cyclin B1 and export of IBB or M9 quantified on optical sections. (A) The mCherry (top) and GFP (bottom) images and mean curves of quantification of nuclear cyclin B1 and cytoplasmic IBB are shown. (B) mCherry (top), GFP (middle), and pseudo-colored GFP (rainbow look up table [LUT]; bottom) images and mean curves of quantification of the nuclear cyclin B1 and cytoplasmic M9 are displayed. Boxed images show the time point when cyclin B1 begins to enter the nucleus and GFP-M9 begins to exit. Error bars show SEM. Bars, 10 µm.

To confirm that the nuclear import of cyclin B1 was not caused by passive diffusion after partial disassembly of nuclear pores or rupture of the nuclear envelope, we coexpressed wild-type cyclin B1–GFP (cyclin B1–Cdk1–GFP complex, molecular weight of ~100) with mCherry–α-tubulin (mCherry-α–β-tubulin complex, molecular weight of ~120) and recorded their nuclear accumulation by spinning-disk confocal microscopy. We observed that in a similar manner to IBB-GFP, mCherry–α-tubulin began to move into the nucleus ~10 min before NEBD, whereas cyclin B1 started to be imported several minutes before this (cyclin B1, mean of 24 ± 4 min before NEBD; α-tubulin, average of 6 ± 2 min before NEBD; n = 10; Fig. 9 A and Video 9). Thus, our results strongly indicate that cyclin B1–Cdk1 activity alters the nuclear transport machinery early in prophase, possibly through the modification of functional properties of nuclear pores, thereby triggering the nuclear import of cyclin B1.

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Figure 9.

Cyclin B1 but not Cdc25C moves into the nucleus before the nuclear pores become permeable to tubulin. (A and B) Cells coexpressing cyclin B1–GFP and mCherry–α-tubulin (A) or cyclin B1–mCherry and GFP-Cdc25C (B) were recorded as they entered mitosis (one image/40 s and one image/30 s, respectively), and the import of cyclin B1, α-tubulin, and Cdc25C was quantified on optical sections. (A) The GFP (top) and mCherry (bottom) images and mean curves of quantification of nuclear cyclin B1 and α-tubulin are shown. (B) mCherry (top) and GFP (bottom) images and mean curves of quantification of the nuclear cyclin B1 and Cdc25C are displayed. Error bars show SEM. Bars, 10 µm.

Cyclin B1–Cdk1 is first activated in the cytoplasm

One strong implication of our work is that cyclin B1–Cdk1 must first be activated in the cytoplasm and not the nucleus in mammalian cells. Were it to be activated only in the nucleus, it is difficult to see how any cyclin B1–Cdk1 activity could reach the cytoplasm until NEBD because activation induces an ~40-fold increase in the rate of cyclin B1–Cdk1 nuclear import. In agreement with this, we observed that cytoplasmic events such as cell rounding and centrosome separation follow almost immediately after, and remain dependent on, cyclin B1–Cdk1 activation (Gavet and Pines, 2010). Moreover, the nuclear accumulation of cyclin B1 is not affected by inhibiting its export. In further support of this conclusion, antiphospho antibodies that only recognize active cyclin B1–Cdk1 first stain centrosomes in mammalian cells (Jackman et al., 2003), and cyclin B1–Cdk1 can be activated in the absence of nuclei in X. laevis oocytes (Hara et al., 1980; Pérez-Mongiovi et al., 2000). Thus, activation in the cytoplasm allied with the immediate and rapid nuclear import of a proportion but not all of cyclin B1–Cdk1 means that activating cyclin B1–Cdk1 inherently synchronizes mitotic events in the cytoplasm and nucleus.

Our finding that cyclin B1–Cdk1 nuclear import remains dependent on cyclin B1–Cdk1 activity until NEBD also means that any enzyme repressing its activity, such as Chk1, Wee1, or Myt1, would also prevent cyclin B1–Cdk1 import. It has been suggested that inhibiting cyclin B1–Cdk1 specifically in the nucleus or blocking its nuclear import could generate a cell in which cyclin B1–Cdk1 is only active in the cytoplasm. Three ways by which this has been reported to happen are (1) through activating the Chfr tumor suppressor, (2) overexpressing Wee1 in human cells, and (3) when the Grapes/Chk1 kinase is activated in Drosophila melanogaster syncitial embryos (Heald et al., 1993; Summers et al., 2005; Royou et al., 2008). Our results tend to preclude all three mechanisms.

(1) In the study on the effect of Chfr, cyclin B1–Cdk1 itself was not activated (Summers et al., 2005), and our results now show that there is, therefore, no need to postulate an extra effect on cyclin B1 import because activation and import are coupled. (2) We have measured the effect of overexpressing Wee1 using targeted FRET biosensors to monitor cyclin B1–Cdk1 activity in the nucleus and the cytoplasm but find that we cannot uncouple the activities in the two compartments, which is the result predicted by our finding that active cytoplasmic cyclin B1–Cdk1 enhances its nuclear import and that nuclear export is not inhibited during prophase. (3) A similar argument applies to the effect of Chk1, at least in human cells. Thus, at present, we have found no evidence that the nuclear import of cyclin B1 per se is subject to regulation aside from the controls on the activation of cyclin B1–Cdk1, nor is there any need to invoke such regulation.

Plk1 acts upstream of the nuclear import of cyclin B1–Cdk1

A prevailing idea concerning mitotic entry is that by phosphorylating cyclin B1 to block its nuclear export, the Plk1 kinase either triggers or is essential for the accumulation of cyclin B1 in the nucleus. But, in direct contradiction to this and in agreement with our previous conclusions (Jackman et al., 2003) and those of others (Lénárt et al., 2007), we have shown in this study that neither Plk1 activity nor the inhibition of nuclear export is required for the rapid accumulation of cyclin B1–Cdk1 in prophase. Indeed, we find no evidence that altering nuclear export rates contributes to the nuclear accumulation of cyclin B1–Cdk1. Instead, this can be entirely generated through an up to ~40-fold increase in its import rate. Because the rate of import is the same in the presence or absence of Plk1 activity but fewer cells enter mitosis in the absence of Plk1 (Gavet and Pines, 2010), our experiments indicate that Plk1 acts upstream of cyclin B1–Cdk1 nuclear import.

Phosphorylation of B1 alters its import rate but is only significant in interphase

Our earlier experiments indicated that substituting five conserved Ser in the N terminus of cyclin B1 with Ala appeared to prevent the protein from entering the nucleus until NEBD, whereas substituting them for glutamic acid created a nuclear import signal (Hagting et al., 1999). Since then, however, the development of highly sensitive charge-coupled device cameras and spinning-disk confocal microscopy have improved our ability to assay and quantify the beginning of cyclin B1 nuclear entry, and we now show that both the wild-type and the Ala mutants are imported much earlier (~20 min before NEBD) than we had previously thought. We have confirmed that glutamic acid substitutions do cause cyclin B1 to become more nuclear in interphase because they increase its rate of import by about fivefold. However, this does not significantly contribute to the ~40-fold increase in the import rate once cells activate cyclin B1–Cdk1. Thus, the phosphorylation of cyclin B1 might help its nuclear import but is not necessary at prophase.

Cell culture and synchronization

HeLa cells were cultured in advanced DME (Invitrogen) supplemented with 2% FBS (Invitrogen), 200 µM Glutamax-1, 100 U/ml penicillin, 100 µg/ml streptomycin, and 250 ng/ml fungizone at 37°C with 10% CO₂. For time-lapse imaging, cells were cultured on glass-bottom dishes (Willco Wells) precoated with fibronectin (Sigma-Aldrich) at 1 µg/cm² for 2 h before use. Cells were synchronized in S phase by addition of 2.5 mM thymidine for 24 h then released into fresh medium. Cells entered mitosis on average 12 h after release from the thymidine block.

Transfection

HeLa cells were electroporated (3 × 10⁶ cells; 10–20 µg DNA) at 250 V and 1,500 µF in 4-mm cuvettes using an electroporator (Easyject plus; Equibio). We estimated the transfection efficiency to be ~40–60% depending on the quality of DNA. Cells were immediately plated on coated glass-bottom dishes and synchronized 7 h after transfection by a thymidine block and release. Alternatively, asynchronous cells were recorded 24 h after transfection.

Time-lapse imaging

Time-lapse imaging was performed on cells in L15 medium with 10% serum at 37°C using either an epifluorescence microscope (Deltavision; Applied Precision) controlled by SoftWoRx software and equipped with an inverted microscope (IX70; Olympus), an electron multiplier charge-coupled device camera (Cascade II 512K; Photometrics), and a 40× UApo NA 1.35 objective (Olympus) or a spinning-disk system (Marianas; Intelligent Imaging Innovations) controlled by Slidebook software and comprising a microscope (Axiovert Observer Z1; Carl Zeiss, Inc.) equipped with a disk-scanning unit (CSU-X1 Nipkow; Yokogawa) and a camera (QuantEM 512SC; Yokogawa). FRET imaging was performed using a CFP/YFP filter set with CFPex 436/10, CFPem 470/30, YFPem 535/30, and a CFP/YFP/mCherry dichroic C85363 (Chroma Technology Corp.). The same exposure time was used for CFPex/CFPem and CFPex/YFPem (between 100 and 200 ms). All quantifications were performed using ImageJ software (National Institutes of Health). For emission ratio measurements, we used the following formulas: whole cell signal = sum of the intensity of the pixels for one cell; mean background signal = mean signal per pixel for a region selected just beside the cell; whole cell signal corrected = whole cell signal − (area for the selected cell × mean background); and emission ratio = whole cell YFP signal corrected/whole cell CFP signal corrected.

Intensity-modulated display representations were performed using MetaMorph software according to the manufacturer's recommendations (MDS Analytical Technologies). In brief, for each intensity-modulated display, we used eight color hues and 32 intensities, ranking from dark to bright per hue. The maximum and minimum values were fixed manually and are indicated on each figure. The color intensities displayed for each hue were determined automatically by the software using one of the CFPex/YFPem images as a reference (Tsien and Harootunian, 1990).

For quantifications of cytoplasmic/nuclear ratio, we designed a region of interest corresponding approximately to one third of the nucleus and excluding centrosomes (Fig. 6 A) that could be used to quantify the nuclear signal and the cytoplasmic signal close to the nucleus in the same cell. FRAP experiments were performed using a 488-nm laser at maximum power and an exposure time of 100 ms.

Constructs