A plant cyclin B2 is degraded early in mitosis and its ectopic expression shortens G2-phase and alleviates the DNA-damage checkpoint

Progress through the eukaryotic cell cycle is regulated by sequential waves of different cyclin and cyclin-dependent kinase (Cdk) activities(Pines and Rieder, 2001). Both in plants and in animals, several classes of Cdks exist, which act in combination with different cyclins at the major control steps of the cell cycle (Joubes et al., 2000). In animals, S phase is induced by Cdk2 bound to the S-phase-specific E- and A-type cyclins, and M phase is triggered by Cdk1 associated with the mitotic A- and B-type cyclins (Pines,1999). While entry into mitosis is mainly controlled by cyclin A-associated Cdk1 kinase, final commitment to mitosis is mediated by the rapid activation of cyclin B-Cdk1 (Pines and Rieder, 2001). This process is highly regulated and involves several levels of control, such as phosphorylation and dephosphorylation events on conserved sites of the Cdk kinase, binding of inhibitors, and the subcellular localization of the Cdk-cyclin complex(Ohi and Gould, 1999).

The first functional evidence that Cdks are pivotal for mitotic control in plants came from experiments in which microinjection of an active Cdk complex induced entry into mitosis (Hush et al.,1996). The plant functional homologue of the animal Cdk1 is named CdkA and has a protein kinase activity that peaks at the G1-S and the G2-M transitions (Bögre et al.,1997). Treatment of cells with roscovitine, a drug that most specifically inhibits CdkA, blocks cell cycle progression both in G1 and G2 phases, further indicating multiple roles for plant CdkA during G1 and G2 phases (Binarova et al., 1998). Another group of plant Cdks is divided into two subgroups, CdkB1 and CdkB2,and exists only in plants (Joubes et al.,2000). The tightly regulated expression and activity of B-type Cdks, as well as the block in G2 phase by the expression of a dominant-negative mutant form of CdkB1, suggest a specific function during G2-phase and mitosis (Magyar et al.,1997; Porceddu et al.,1999; Porceddu et al.,2001).

There are three A-type and two B-type cyclin groups in plants(Renaudin et al., 1998). In synchronized BY2 cells cyclin A3 is expressed during an earlier time window,from the G1-S transition until onset of mitosis compared with cyclin A1 and A2, which are expressed during mid S phase until midmitosis(Reichheld et al., 1996). Cyclin A2-associated kinase activity peaks both in S phase and mitosis in synchronized alfalfa cells (Roudier et al., 2000). In animals a single A-type cyclin associates with different Cdks during S phase and mitosis(Resnitzky et al., 1995; den Elzen and Pines, 2001),while in plants two A-type cyclins, the tobacco cyclin A1 and the alfalfa cyclin A2, were found only associated with CdkA both in vitro(Nakagami et al., 1999), and in the yeast two-bybrid system (Roudier et al., 2000).

The expression of the B1- and B2-type plant cyclins is tightly controlled and restricted to late G2 and M phases(Hirt et al., 1992; Fobert et al., 1994). A tobacco Cyclin B1-GFP fusion was found to be distributed between the cytoplasm and nucleus in interphase and to associate with the chromatin during prophase and metaphase until it is degraded at the onset of anaphase(Criqui et al., 2000; Criqui et al., 2001). These authors also found that ectopically expressed cyclin B1 did not alter mitotic progression in cultured cells, while expression of cyclin B1 in Arabidopsis was shown to stimulate root formation by producing more cells as building blocks for tissue enlargement(Doerner et al., 1996). Evidence for specialized functions of closely related plant B-type cyclins came from the finding that ectopic expression of the Arabidopsis cyclin B1;2, but not of cyclin B1;1 inhibited endoreduplication and induced cell divisions, resulting in multicellular trichomes(Schnittger et al., 2002). Recently, we presented evidence for the existence of a topoisomerase II-dependent checkpoint in plant cells by showing that the ectopic expression of Cyclin B2 alone is able to override the checkpoint and induce cells to enter into mitosis without delay(Gimenez-Abian et al.,2002).

Plant B2-type cyclins are known to be expressed from late G2 phase to mitosis (Hirt et al., 1992; Ferreira et al., 1994) but the role and localization of their protein has not been studied. We show that the cyclin B2 protein is localized to the nucleus and its expression is restricted to a short time window during early prophase by its proteolysis. We found that the induction of ectopic cyclin B2 expression during G2-phase drives cells to enter mitosis earlier and allows caffeine and okadaic acid to overcome a replication checkpoint block induced by hydroxyurea. Thus, ectopic cyclin B2 expression interferes with the control of progression from G2 to M phase,which probably causes the developmental abnormalities observed in transgenic plants.

The alfalfa Medsa;CycB2;2 gene was amplified by PCR, creating a KpnI site at the N terminus and NotI and XbaI restriction sites at the C terminus and cloned into the BinHygTX binary vector(Weinmann et al., 1994) as a KpnI-XbaI fragment. The triple HA epitope(Tyers et al., 1993) and GFP sequences (Sheen et al., 1995)were inserted as NotI-NotI fragments. Transgenic plants were obtained by Agrobacterium-mediated leaf disk transformation with the tobacco line Samsung Tb-Tet(Weinmann et al., 1994). Suspension cultures were initiated by placing stem segments of transgenic CycB2-HA and CycB2-GFP plants on 1 mg/l 2,4-D containing MS solid medium and the resulting calli were transferred after 1 month into liquid LS-modified medium (Sano et al., 1999)supplemented with 40 mg/l hygromycin B.

Synchronization was carried out by diluting a 7-day-old culture 1:5 and adding 10 mg/l aphidicolin (Sigma) 8 hours later for 16 hours. After removal of the aphidicolin by washing cells five times with medium, chlortetracycline(Sigma) was added at a concentration of 0.1 mg/l to induce cycB2-expression. Drugs were added 8 hours after aphidicolin removal in G2 phase. The following drug concentrations were used in these experiments: 10 μM amiprophos methyl, 50 μM taxol, 10 μM propyzamide, 100 μM roscovitine (a gift from M. Strnad, Olomouc, Czech Republic), 100 μM MG 132 (Calbiochem), 5μM lactacystin (Affinity, UK) and 1 μM epoxomicin (Affiniti, UK). To follow cell cycle progression, flow cytometric analysis was performed as described (Bögre et al.,1997) using a PAS2 flow cytometer (Partec, Münster, Germany). For mitotic index, cells were fixed in a 3:1 (v/v) ethanol/acetic acid mixture, then washed with 70% (v/v) ethanol. The DNA was stained with 1μg/ml DAPI and observed by epifluorescence microscopy.

A stationary phase cell culture (7-day-old culture) was diluted 1:10 with fresh medium, and bromo-deoxyuridine labeling reagent was added immediately at the dilution as recommended in the cell proliferation kit (RPN20) (Amersham Pharmacia Biotech). Cells were grown for 15 hours and samples were taken every 3 hours, fixed and processed for immunostaining for incorporated bromo-deoxyuridine (BrdU) using monoclonal antibody to BrdU (Amersham Pharmacia Biotech) or Cycb2-HA using HA.11 monoclonal antibody (BabCO,Richmond, California). For each timepoint, 1000 cells were analyzed for BrdU or HA-staining.

For indirect immunofluorescence, cells were fixed and stained as previously described (Bögre et al.,1997). The HA.11 monoclonal antibody (BabCO, Richmond, California)was used in a 1:1000 dilution and with the secondary anti-mouse Cy3-conjugated antibody (Sigma) at a dilution of 1:200. DNA was stained with 1 μg/ml DAPI in PBS. Microtubule staining was performed using an anti α-tubulin mouse monoclonal antibody DM1A (Sigma) at a dilution of 1:200, and anti mouse FITC-conjugated secondary antibody (Sigma). For GFP observation, a drop of cell suspension was transferred on a slide, carefully covered with a coverslip, and observed with an upright fluorescence microscope (Axioplan 2;Zeiss, Jena, Germany) equipped with a GFP filter (HQ480/20X; HQ510/20M; AF Analysentechnik, Jena, Germany). Typical exposure times were in a range of few seconds. Images were taken using a cooled charge-coupled device black-and-white digital camera (SPOT-2; Diagnostic Instruments, Burroughs,Michigan) and Metaview imaging software (Diagnostic Instruments, Burroughs,Michigan).

Northern blots were performed as described(Hirt et al., 1992). Proteins on western blots were detected by using anti-rabbit PSTAIRE antibody(Bögre et al., 1997), the HA.11 monoclonal antibody (BabCO, Richmond, California) or the polyclonal rabbit anti N-terminal cyclin B1;1 antibody (kindly provided by Pascal Genschik, Strasbourg, France). Cdk activities were measured in the samples after immunoprecipitation with the HA.11 monoclonal antibody or after binding to p13^suc1, as described previously(Bögre et al., 1997).

Leaf disks with 1 cm diameter were excised from sterile plants and placed on MS medium (Sigma) with 0.5/0.1 (rooting), 0.1/0.5 (shooting) and 0.5/0.5(callusing) naphtalene acetic acid (NAA)/6-benzylaminopurine (BAP)-containing media with and without 0.1 mg/l Cl-tetracycline for 3 weeks, when roots on 10 inocula for each treatments were counted and plates were photographed.

We studied the role of cyclin B2 during plant cell cycle progression by ectopically expressing an alfalfa cyclin B2 gene (Medsa;cycB2;2) in transgenic tobacco plants. As the expression of cycB2-HA from a constitutive promoter severely inhibited shoot and root development, and did not allow the regeneration of plants from transformed leaf disks, we generated plants in which the expression of cycB2 can be induced by tetracycline analogues (Weinmann et al.,1994). To follow the expression of the introduced gene, it was tagged with either the epitope haemagglutinin antigen (cycB2-HA) or with the green fluorescent protein (cycB2-GFP). Five independent lines were obtained with the cycB2-HA construct and named CycB2-HA-1 to 5, and five independent lines harboring the cycB2-GFP construct were obtained and named CycB2-GFP-1 to 5. Northern blot analysis revealed that in four of these lines the cycB2-HA expression was increased 50- to 1000-fold in detached leaves, which were incubated in 0.1 mg/l Cl-tetracycline for 24 hours (Fig. 1A). One of the lines showed a low level of constitutive expression (see lane 1 in Fig. 1A) and this line was retarded in both shoot and especially root growth.

To facilitate the studies of cell cycle progression, we generated cell cultures from the CycB2-HA and CycB2-GFP transgenic plants, which displayed tight transcriptional regulation by Cl-tetracycline. In cultured cells derived from line CycB2-HA-2 the cycB2-HA transcript was clearly detectable 10 minutes after adding Cl-tetracycline, and reached its maximal level of expression within 1 hour (Fig. 1B). We established the maximal induction of CycB2-HA protein at 0.01 mg/ml Cl-tetracycline (Fig. 1C). Correspondingly, in vitro kinase assays after immunoprecipitation using an anti HA antibody revealed that the CycB2-HA protein associated with and activated a histone H1 kinase (Fig. 1D), indicating that the cyclin moiety in these fusion proteins retained its normal function to bind and activate Cdk. The level of cycB2-HA mRNA expression was slightly lower than that of the endogenous mitotic cyclin Nt;tCycB1;1 (Fig. 2B). In view of the lack of a cyclin B2-specific antibody, we could not determine the endogenous cyclin B2 levels, but we found that the CycB2-HA-associated histone H1 kinase activity reached only around 20% of the total cellular Cdk activity purified from these cells by binding to p13^suc1(Fig. 1D). Thus, it is likely that the cyclin B2 protein was not overproduced, but its expression was temporally unscheduled.

To assess the consequences of altered cyclin B2 expression on cell cycle regulation, we synchronized the CycB2-HA-2 cell culture by aphidicolin treatment, which blocks cell cycle progression specifically during S phase. Following removal of the inhibitor, we induced the expression of CycB2-HA at various time points during the cell cycle. We found that ectopic expression of CycB2-HA during either S or early G2 phase did not initiate mitosis. However,cells entered mitosis 2 hours earlier compared with control cells when expression of CycB2-HA was ectopically induced at mid G2 phase(Fig. 2). Several lines of evidence supported the finding of this advanced entry into mitosis by 2 hours. First, the mitotic index peaked at 10 hours in cells ectopically expressing CycB2-HA, instead of 12 hours in control cells(Fig. 2A). Second, monitoring cell cycle progression by flow cytometry revealed that after cells had passed through mitosis, the percentage of cells with a G1 DNA increased 2 hours earlier in CycB2-HA expressing cells (data not shown). Third, we followed the mRNA expression of the tobacco cyclin B1 and of histone H4,as stage-specific markers for mitosis and S phase, respectively. Mirroring the results of mitotic index above, the peak of the endogenous cyclin B1 expression also appeared 2 hours earlier in tetracycline-treated cells(Fig. 2B).

Because microtubule structures are highly specific landmarks for the different stages of mitosis, we used these features to further refine and confirm the above interpretation. The pre-prophase band (PPB) appears between late G2 phase and late prophase, the spindle structures during mitosis, and the phragmoplast marks the cytokinetic plate during late anaphase and telophase (Staiger and Lloyd,1991). We quantified the relative proportion of PPB, spindle and phragmoplast visualized by tubulin immunostaining in synchronized cells expressing cycB2-HA. Disassembly of the PPB, which occurred 2 hours earlier than in control cells, was the most pronounced effect of ectopic CycB2-HA expression (Fig. 2C).

Blocking DNA synthesis arrests cells before mitosis, and blocks the expression of mitotic A- and B-type cyclins in plants(Renaudin et al., 1998). First, we tested whether ectopic cyclin B2 expression could induce S-phase-arrested cells to enter into mitosis. The induction of CycB2-HA expression in asynchronously dividing cells by treatment with tetracycline for 16 hours slightly increased the mitotic index(Fig. 3A, lanes 1 and 7) and this effect was abolished when cells were treated at the same time with HU(Fig. 3A, lanes 4 and 10). Thus, cyclin B2 alone is unable to override the HU-activated checkpoint block and we found a similar result in aphidicolin treated cells.

Drugs such as caffeine or okadaic acid are known to cancel the DNA-replication checkpoint in yeast and animal cells, but were found to be ineffective in plant cells (Amino and Nagata, 1996; Pelayo et al.,2001). A reason for this could be that in plants mitotic cyclins are not expressed in S-phase-arrested cells. Thus, to gain further evidence that CycB2 is a potent inducer for mitosis, we tested whether its ectopic expression could help caffeine or okadaic acid to override the S-phase block. In an asynchronously growing control culture the mitotic index was determined to be 7% and the addition of caffeine or okadaic acid for 1 hour did not modify the percentage of mitotic cells. The addition of hydroxyurea (HU) for 16 hours, which blocks cells in S-phase, decreased the mitotic index (MI) and this low MI was not increased by a subsequent 1 hour treatment with caffeine or okadaic acid (Fig. 3A, left panel). Thus, caffeine or okadaic acid were unable to override the HU-induced S-phase block, which is in agreement with the results found before in onion root cells (Amino and Nagata,1996; Pelayo et al.,2001). However, when caffeine or okadaic acid were added to the HU-blocked cells expressing CycB2-HA, the mitotic indices were raised to a level of around 50 and 25%, respectively, when compared with control cells without HU. Caffeine was slightly more effective than okadaic acid in stimulating mitosis in HU blocked cells, together with ectopic CycB2-HA expression (Fig. 3A, right panel). Thus, CycB2-HA expression allowed caffeine or okadaic acid to override replication checkpoints in S phase.

To address whether caffeine and okadaic acid stimulated cells to enter into mitosis by elevating the CycB2-HA-associated Cdk activity in HU-blocked cells,we immunopurified the CycB2-HA-associated kinase complex with an antibody directed against the HA-tag and measured its histone H1-kinase activity. In the absence of tetracycline, no CycB2-HA protein or CycB2-HA-associated Cdk activity were found, while the Cdk amounts, as detected by the PSTAIRE antibody, were constant in these extracts(Fig. 3B, left panel). In the presence of tetracycline, CycB2-HA protein expression was increased by equal amounts after all treatments, showing that the tetracycline-induction of CycB2-HA expression was similar in all the treatments. We found that the CycB2-HA-associated Cdk activity in cells treated with tetracycline was increased after treatment with caffeine and okadaic acid. The addition of HU abolished the CycB2-HA-associated Cdk activity. In agreement with its ability to induce mitosis in HU-blocked cells, caffeine and okadaic acid elevated the activity of CycB2-associated Cdk in HU-treated cells(Fig. 3B, right panel). Okadaic acid was again less effective in raising the Cdk activity than was caffeine. Thus, we conclude that the activity of CycB2-associated Cdk is inhibited in the presence of HU, and this inhibition is cancelled by treatment with caffeine and okadaic acid, which supports our findings that caffeine or okadaic acid together with CycB2 expression override the replication-checkpoint in plant cells.

To learn more about how cyclin B2 promotes the G2-M transition, we determined its localization in cultured cells by indirect immunofluorescence using an anti HA-antibody (Fig. 4). In interphase cells CycB2-HA was localized to the nucleus. In late G2-phase cells, which display a pre-prophase band, CycB2-HA was still present within the nucleus, but it was absent in pro-metaphase cells with condensed chromatin and visible spindle microtubules(Fig. 4A, PM). This suggests that CycB2-HA is degraded as cells progress from pre-prophase to pro-metaphase. No CycB2-HA staining was observed in cells in meta-, ana- or in telophase, and it was missing in around 40% of interphase cells(Fig. 4A).

To confirm the localization of the cyclin B2 protein in live cells, we used a cell line expressing a cyclin B2-GFP fusion. Similar to the CycB2-HA protein visualized by immunostaining with the HA antibody, CycB2-GFP was also found within the nucleus in interphase cells and it was never found in mitotic cells(Fig. 4B, left panel). We were unable to follow the degradation of CycB2-GFP by time-lapse microscopy,because UV irradiation of cells during late G2-phase blocked cell cycle progression, presumably by activating a prophase checkpoint, similar to that described in animal cells (Rieder and Cole, 1998).

In an asynchronously growing culture around 60% of interphase cells displayed CycB2-HA or CycB2-GFP staining within the nucleus after treatment with tetracycline. To define more precisely at what stage of interphase cyclin B2 was stable, we blocked cell cycle progression at the G1-S transition by treatment with aphidicolin. Both CycB2-HA and CycB2-GFP signals were detectable in around 90% of aphidicolin-blocked cells. After release from the block, a high proportion of cells (around 90%) remained positive for nuclear CycB2-GFP or CycB2-HA signals, which decreased when cells progressed through mitosis (data not shown). Thus, CycB2-HA and CycB2-GFP were stable during S and G2 phases, and became unstable as cells entered mitosis and G1 phase. To map the time window within G1 during which CycB2-HA was stable, we observed stationary phase cells arrested with a G1 DNA content. In these cells we never found CycB2 protein expression, indicating that it is unstable or not translated (Fig. 4C). Adding back fresh medium reinitiated the cell cycle and cells reached S phase within 0 to 15 hours, which was shown by monitoring incorporation of BrdU. CycB2-HA protein reappeared at around 6-9 hours in a large proportion of cells(Fig. 4C). Thus, there is a transition point within G1 when cyclin B2 becomes stabilized, similar to B-type cyclins of yeast and animal cells(Amon et al., 1993; Brandeis and Hunt, 1996).

We analyzed the timing of the initiation of CycB2-HA degradation by using drugs that inhibited cell cycle progression at specific time points. Inhibition of Cdk activity by adding roscovitine to synchronized cells during mid G2 phase (6 hours after the release from aphidicolin treatment) blocks cells at late G2 phase and G2-M interface, characterized by a fully developed prophase spindle and condensed chromatin surrounded by a persistent nuclear envelope (Binarova et al.,1998). Inducing such a G2-arrest kept CycB2-HA intact within the nucleus, while in control cells CycB2-HA became degraded after prophase(Fig. 5A). Thus, the degradation of cyclin B2 requires either an active Cdk or nuclear envelope breakdown.

Activation of the spindle checkpoint by treatment of cells with microtubule disrupting drugs leads to a metaphase arrest and stabilization of cyclin B1(Criqui et al., 2000). To study whether the spindle checkpoint might influence cyclin B2 degradation in plants, we treated the synchronized CycB2-HA-2 cells in late G2-phase with the microtubule stabilizing drug taxol, as well as with the microtubule disrupting drug amiprophos methyl (APM), and followed CycB2-HA localization with immunostaining. Both APM and taxol arrested a large proportion of cells in metaphase, without any detectable CycB2-HA-derived signal(Fig. 5B,C).

To confirm that CycB2 is unstable in metaphase-arrested cells, and that the signal is not masked and therefore undetectable by immunofluorescence, we used the CycB2-GFP cell line and followed the CycB2-GFP fluorescence in living cells during activation of the spindle checkpoint by depolymerizing microtubules with propyzamide (Fig. 6A). Again, we found that the CycB2-GFP-derived fluorescence was clearly present in propyzamide-treated interphase cells, but it was missing in metaphase-arrested cells (Fig. 6A, right panel, arrow). When the same treatment was applied to a cell line expressing a cyclin B1-GFP fusion, we found that the GFP signal was strongly present in association with chromosomes in metaphase-arrested cells(Fig. 6A, left panel) as it was reported previously (Criqui et al.,2000; Criqui et al.,2001).

To further ascertain that the CycB2-HA protein is indeed decreased when the spindle checkpoint has been activated and that cyclin B1 is stabilized in these cells, we analyzed protein extracts of synchronized cells by immunoblotting using the anti HA-antibody for detection of CycB2-HA protein and the anti cyclin B1;1 antibody for detection of endogenous cyclin B1(Criqui et al., 2000). In control cells, both the expression of CycB2-HA and cyclin B1 increased during early mitosis, reaching the maximum at 9 hours after aphidicolin removal, from which point both cyclin levels declined rapidly(Fig. 7). Treatment with propyzamide at 7 hours during G2 phase did not affect the levels of CycB2-HA protein, and it decreased at 15 hours to a similar level as in the control cells (Fig. 7A). Contrary to this, the endogenous cyclin B1 protein level remained high in propyzamide-treated cells (Fig. 7B). This strongly suggests, that cyclin B1, but not cyclin B2 is stabilized by the spindle-checkpoint in plants.

Degradation of A- and B-type cyclins is blocked by proteosome inhibitors such as MG 132, epoxomicin or lactacystin, both in plant and in animal cells and they arrest cells with over-condensed metaphase chromosomes(Genschik et al., 1998). Surprisingly, we found that the CycB2-HA derived signal was barely detectable in MG 132-treated cells (Fig. 5C).

To confirm that proteosome inhibitors differently affect B-type cyclin levels in vivo, we treated CycB1-GFP- and CycB2-GFP-expressing cells with MG 132, as well as with two other inhibitors: epoxomicin and lactacystin. Similar to our findings by immunolocalization in MG 132-treated cells, CycB2-GFP was barely detectable, while CycB1-GFP was strongly present and localized to the condensed chromosomes in cells which were arrested in metaphase after treatment with epoxomicin (Fig. 6B), MG 132 or lactacystin (data not shown).

Immunodetection of the CycB2-HA and the endogenous cyclin B1 protein levels further substantiated the above results. We treated cells before mitosis, at 7 hours after aphidicoline release with epoxomicin, lactacystin or MG 132, and prepared extracts from cells at 15 hours for immunoblotting. We found that CycB2-HA protein levels were unaffected by the proteosome inhibitors, and were as low in the treated cells as in the control cells(Fig. 7A), while the cyclin B1 protein levels increased (Fig. 7B).

In contrast to wild-type cells, we found that CycB2-HA-expressing cells arrested in the presence of MG 132 at a stage when chromosomes were not aligned along the metaphase plate (Fig. 5C). In untreated cells, which ectopically expressed CycB2-HA, we observed a slight increase in the percentage of metaphase cells compared with control cells, but we never found abnormalities in chromosome alignment. This indicates that increased levels of cyclin B2 during prophase might interfere with chromosome alignment at the metaphase plate. No such abnormalities were observed in control cells or CycB1-overexpressing cells treated with the proteosome inhibitors.

We initially observed, that constitutive expression of CycB2 severely retards plant growth with most pronounced effects on root development. To investigate how ectopic expression of CycB2 could interfere with shoot and root regeneration, we used a transgenic line with Cl-tetracycline inducible cycB2-HA expression. Detached leaves from transgenic plants were placed on Cl-tetracycline-containing medium, in combination with different ratios of cytokinin and auxin. It is well established that high ratios of cytokinin to auxin favor shoot regeneration, while the reverse would privilege root growth(Sugiyama, 1999). Our results show that ectopic expression of CycB2 specifically blocked root development in a medium containing high auxin to cytokinin ratios, conditions in which root development is normally encouraged (Fig. 8A). However, the initiation of shoot development in these transgenic plants was not detectably modified on high ratios of cytokinin to auxin containing medium (data not shown).

To determine whether the inhibition of root development can be traced to an altered cell cycle progression, we measured the distribution of cells with a G1 and G2 DNA content. Most cells in leaves without hormone treatment have G1 DNA content, irrespective of tetracycline-induced cycB2-HA expression (data not shown). Control experiments, in which leaves were incubated on auxin-containing medium but without induction of cycB2-HA expression, showed that after 3 days the cell cycle was initiated in a proportion of cells (as judged from BrdU incorporation for S-phase and observing mitotic figures in leaf pieces) and 30% of the cells displayed a G2 DNA content. By contrast, in leaves that were incubated with auxin while the expression of cycB2-HA was induced, only about 15% of the cells displayed G2 DNA content after 3 days(Fig. 8B), while similar percentages of S-phase and mitosis, as compared with control samples, were observed. The reduced proportion of cells with G2 DNA content indicates that,as in the previous experiments with cultured cells, the G2-phase is accelerated upon ectopic expression of cycB2 in the presence of auxin.

In plants, cell division can be influenced by growth or developmental signals not only in G1 phase, but also during G2 phase, but the molecular mechanisms for the control of G2 progression is poorly understood. We show that ectopic expression of a B2-type cyclin interferes with the timing of entry into mitosis and affects plant development.

The plant cyclin B2 appears to play a similar role to cyclin A in animal cells during entry into mitosis. In animals, microinjection of exogenous cyclin A into G2 cells rapidly induces chromosome condensation, while inhibition of cyclin A-Cdk-complex by p21^cip1/waf1 prevents G2 cells from entering prophase and induces early prophase cells to return back to interphase (Furuno et al.,1999). Irradiation-induced DNA damage identifies a late G2 control point in animal cells, from where the chromosomes return to an uncondensed interphase stage (Rieder and Cole,1998). An analogous G2-checkpoint was observed in onion cells,where chromosome condensation is reversed by inhibiting protein synthesis before mid-prophase (Gracia-Herdugo et al., 1974). This reversible condensation phase coincides with the timing of cyclin A-associated Cdk1 activation in animal cells (Clute and Pines, 1999), and our findings suggest that it might be a cyclin B2-associated Cdk kinase in plant cells. Although animal cyclin A is expressed during a broad time window and has a role both in S-phase and during entry into mitosis (Rosenblatt et al.,1992), plant cyclin B2 expression is confined to late G2-phase and early mitosis. The existence of several subgroups of plant A- and B-type cyclins, which are specifically expressed during S, G2 or mitotic phases,suggests a high degree of specialization in cyclin function in plants(Renaudin et al., 1998).

The PPB is a microtubule structure formed in late G2-phase, and it disassembles as cells enter mitosis in late prophase(Utrilla et al., 1993). Further evidence that ectopic cyclin B2 expression affects the G2 to M phase transition is given by the lower percentage of PPBs in these cells. This indicates that either the time window during which the PPB is present becomes shorter or a cyclin B2-associated Cdk activity directly induces the disassembly of the PPB. It has been shown that microinjections of an active Cdk complex into plant cells can cause the disassembly of PPB and nuclear envelope breakdown (Hush et al.,1996).

Ectopic Cyclin B2 expression could not initiate mitosis in S or early G2 phase. In yeast and animals, entry into mitosis is regulated by the phosphorylation of the Thr₁₄ and Tyr₁₅ inhibitory sites of CDK (Walworth, 2001). The existence of phosphorylation-mediated Cdk-regulatory pathways during plant mitosis is indicated by the presence of a wee1-kinase homolog in maize and Arabidopsis, which inactivates Cdks through its phosphorylation at the conserved Tyr₁₅ residue(Sun et al., 1999). Surprisingly, no apparent homologs of the cdc25 phosphatase have been found in the fully sequenced genome of Arabidopsis, but dephosphorylation of Tyr₁₅ is tightly regulated in response to cytokinin or during stress treatment(Reichheld et al., 1999). Expression of S. pombe cdc25 in tobacco can induce cells to resume cell divisions that have been arrested in G2 phase by cytokinin starvation(John, 1996; Zhang et al., 1996). Constitutive expression of cdc25 in tobacco plants leads to a number of developmental abnormalities, such as precocious flowering, aberrant leaves and flowers, and a smaller size of dividing cells in lateral roots compared with control plants (Bell et al.,1993).

There is a conserved DNA-synthesis checkpoint-signaling pathway in yeast and animal cells that regulates CDK activity through the phosphorylation of the Thr₁₄ and Tyr₁₅ inhibitory sites. Caffeine is a drug that inhibits checkpoint function in animal cells(Schlegel and Pardee, 1986; Andreassen and Margolis, 1992; Moser et al., 2000), such as the replication checkpoint during S phase(Schlegel and Pardee, 1986)and the DNA damage checkpoint in G2 phase(Blasina et al., 1999). In plants, caffeine is able to override the G2 DNA damage checkpoint(Hartley-Asp et al., 1980; González-Fernández et al.,1985) but it is unable to cancel replication checkpoints in S phase (Amino and Nagata, 1996; Pelayo et al., 2001). Okadaic acid, a protein phosphatase 2A inhibitor, induces premature mitosis in animal cells in a similar way to caffeine (Ghosh et al., 1996). Correspondingly, endothal, another PP2A-specific phosphatase inhibitor, was found to prematurely activate a plant mitotic CDK resulting in microtubule and chromosome condensation abnormalities(Ayaydin et al., 2000). Caffeine was found to releases the checkpoint-induced block in G2 phase by inhibiting the ATM kinase, which works upstream of the two checkpoint kinases:Chk1 and Chk2 (Blasina et al.,1999; Brondello et al.,1999). In a current model the Chk1 kinase directly inhibits the Cdc25 phosphatase and thus stops activation of CDK by keeping it in a phosphorylated state at the Thr₁₄ and Tyr₁₅ inhibitory sites. The animal target of caffeine, the ATM kinase, has been conserved in plants (Garcia et al., 2000),but little is understood about its function during checkpoint pathways in plants.

We show that, in hydroxyurea-treated cells, cyclin B2 expression stimulated the entry into mitosis only when the replication checkpoint pathway was compromised by caffeine or okadaic acid. Moreover, under these conditions, the CycB2-associated Cdk activity was also induced. This suggests that at least one of the targets of the replication checkpoint in plants is the CycB2-associated Cdk. In animal cells, the nuclear-localized cyclin A stimulates entry into mitosis (Furuno et al., 1999), and cyclin B1 is able to abolish the replication checkpoint in response to caffeine when experimentally altered to localize to the nucleus (Tam et al., 1995; Toyoshima et al., 1998). The plant cyclin B2 is nuclear, and during a period of S phase, we found the localization of CycB2 to be speckled within the nucleus (M. W.,unpublished).

In animal cells, A-type cyclins are degraded in pro-metaphase, while B-type cyclins undergo proteolysis in metaphase. Degradation of both cyclins requires the anaphase-promoting complex (APC). Although the APC is activated already during prometaphase, destruction of cyclin B in animal cells appears to be selectively inhibited by the spindle assembly checkpoint, resulting in stabilization of cyclin B until all chromosomes are aligned along the metaphase plate (Geley et al.,2001). Both cyclin types contain the conserved destruction box motif recognized by the APC, but in A-type cyclins, this motif is not sufficient to trigger proteolysis during prophase. Rather, prophase destruction of cyclin A is mediated by an extended destruction box including multiple overlapping elements (Kaspar et al., 2001). Plant cyclin A and B degradation during mitosis is also dependent on their conserved destruction box motif and likely to be mediated by the APC (Genschik et al.,1998). Similar to animal cyclins, plant cyclin A is undetectable in metaphase (Roudier et al.,2000), while plant cyclin B1 is present in metaphase and its degradation is inhibited upon activation of the spindle assembly checkpoint(Criqui et al., 2000). The plant cyc B2;2 genes are grouped together with B-type cyclins, based on similarities within the cyclin box, but the arrangement of the N-terminal destruction box motifs of the plant cyclin B2 sequence better resembles that of animal A-type cyclins which have multiple destruction box elements. Our studies indicate that the plant cyclin B2 protein uses a similar degradation mechanism as animal A-type cyclins and that it is degraded during late prophase, again similar to animal cyclin A. Treatment of cells with the proteosome inhibitors MG 132, epoxomicin or lactacystin arrested cells in metaphase, as expected, but surprisingly, cyclin B2 protein was undetectable by indirect immunofluorescence, suggesting that it is degraded in spite of the presence of the inhibitors in these cells. To exclude the possibility that the cyclin B2 was undetectable due to technical difficulties, such as masking the epitope, we repeated these experiments by treating cells expressing CycB1-GFP or CycB2-GFP with the same proteosome inhibitors and followed GFP fluorescence in living cells. Furthermore, we directly compared the protein level of CycB2-HA with that of endogenous cyclin B1 in proteosome-inhibitor-treated cells by immunoblotting. Both experiments led to the same conclusion: the degradation of cyclin B2 is hardly affected by treatments with proteosome inhibitors, while cyclin B1 destruction is readily blocked and cells are arrested in metaphase. These results indicate that cyclin B2 is either not targeted by the proteosome at pro-metaphase and proteolysed by some other mechanism or its destruction is more effective than that of other substrates,including cyclin B1. Recently, it was shown that cyclin A and cyclin F are degraded by proteolysis independent of proteosome in animal cells(Welm et al., 2002; Fung et al., 2002). Collectively, the data showing that cyclin B2 is only present until pro-metaphase promote our conclusion that plant cyclin B2 functions during early mitosis.

The expression of cyclin B2 to a level not significantly higher than that of endogenous cyclin somewhat delayed metaphase, without affecting subsequent chromosome alignment along the metaphase plate. Compromising proteolysis in these cells by treatment with proteosome inhibitors resulted in an accumulation of cells with chromosomes that were not correctly aligned at the metaphase plate. By contrast, MG 132-treatment of wild-type cells arrests cells in metaphase with normal spindle and chromosome alignment(Genschik et al., 1998). This shows that Cyclin B2 ectopic expression, together with impaired proteolysis synergistically affects chromosome alignment during metaphase. A similar effect has been reported in animal cells upon overexpression of cyclin A during late G2-phase and mitosis (den Elzen and Pines, 2001). This again suggests that the plant cyclin B2 has analogous functions to animal cyclin A in regulating mitosis.

Proper chromosome alignment at the metaphase plate is controlled by a complex anchor of many proteins associated with chromosomes mainly at their kinetochore/centomeric region. Kinetochore localization of phosphoepitopes recognized by the MPM2 antibody suggests that the function of these proteins is regulated by mitosis-specific phosphorylation in plants(Binarova et al., 1993). In Drosophila, Cyclin A inhibits chromosome disjunction by acting synergistically with other proteins involved in this process(Sigrist et al., 1995; Parry and O'Farrell, 2001). Among these are the Drosophila securin Pim, which stabilizes sister chromosome cohesion until it is degraded at the metaphase to anaphase transition (Leismann et al.,2000), and the Cdk inhibitor Rux, which interacts genetically and physically with Cyclin A and inhibits its in vitro kinase activity(Foley et al., 1999).

The increased number of metaphase cells observed upon ectopic cyclin B2 expression suggests that in plants B2-type cyclins might have a similar role to cyclin A in animal cells, delaying the progression through metaphase.

We found that cyclin B2 was non-detectable in stationary phase cells, which are arrested in G1, and begins to accumulate before cells enter S phase. This confirms the results obtained in yeast and animal cells(Amon et al., 1994; Brandeis and Hunt, 1996; Huang et al., 2001) and shows that APC activity is switched off during late G1 to S phase in plants. In yeast and animal cells, APC activity is required during G1 phase to prevent expression of proteins that may interfere with cellular polarization events or lead to premature DNA replication or disturb spindle assembly. Before mitosis,the APC must be turned off to allow mitotic substrates to accumulate. Inactivation of the APC has been linked to mid-G1, which is corresponding to the start in yeast, and is suggested to depend on raising G1 cyclin activity(Amon et al., 1994). Recently,it was shown that during a normal cell cycle the APC is inactivated in a graded manner, and that its complete inactivation occurs in S phase and requires S phase cyclins (Huang et al.,2001).

We found that ectopic expression of cyclin B2 in leaf segments interferes with root regeneration induced by auxin. This re-differentiation process is initiated in a single differentiated cell within the vascular bundle and was mapped by hormone shift-experiments to a relatively short time window, about 24 hours, during which auxin induces the cell to re-enter cell division and re-polarize (Attfield and Evans,1991). The majority of leaf cells enter the cell cycle from the G1 phase, and a 24 hour period might coincide with the time, when a proportion of cells are in G2-phase. Shortening of G2-phase may disrupt some events, such as cell polarization, which is required for the determination of the root fate. Timing of cell cycle phases was shown to be an important factor in determining polarized growth in yeast (Lew and Reed,1993). In plants, the timing of the presence of the PPB, which is particularly altered upon ectopic CycB2-HA expression, could be an important factor for the orientation of cell division. In line with this is a recent report which implicates Cdk1 as an important factor in regulating the orientation of asymmetric cell divisions in Drosophila(Tio et al., 2001). Our results suggest that enhancing the expression of one of the key cell cycle regulators, a cyclin B2 gene, is sufficient to prevent formation of a root meristem under conditions when it is normally formed.

M.W. was supported by a PhD fellowship from the Austrian Academy of Sciences. This work was supported by an Austrian Förderung der Wissenschaftlichen Forschung grant to E.H.-B.; by European Union Framework V project ECCO Grant QLRT-1999-00454 to E.H.-B., L.B. and P.B.; by a Biotechnology and Biological Science Research Council grant (111/P133340) to L.B.; by grant LN00A081 from the Czech Ministry of Education; by a collaborative Wellcome Trust grant to P.B. and L.B.; and by a Spanish Ministerio de Ciencia y Tecnologia grant (BMC2001-2195) to C.T. Moreover, we thank Gireg Weingartner for his patience and Giovanna Vinti, among others, for taking care of him.