The Plant Cell, Vol. 10, 2063–2075, December 1998, www.plantcell.org © 1998 American Society of Plant Physiologists
Cell Cycle–Dependent Proteolysis in Plants: Identification of
the Destruction Box Pathway and Metaphase Arrest
Produced by the Proteasome Inhibitor MG132
Pascal Genschik,1 Marie Claire Criqui, Yves Parmentier, Aude Derevier, and Jacqueline Fleck
Institut de Biologie Moléculaire des Plantes du CNRS, 12 rue du Général Zimmer, 67084 Strasbourg Cédex, France
It is widely assumed that mitotic cyclins are rapidly degraded during anaphase, leading to the inactivation of the cell cycle–dependent protein kinase Cdc2 and allowing exit from mitosis. The proteolysis of mitotic cyclins is ubiquitin/26S
proteasome mediated and requires the presence of the destruction box motif at the N terminus of the proteins. As a
first attempt to study cyclin proteolysis during the plant cell cycle, we investigated the stability of fusion proteins in
which the N-terminal domains of an A-type and a B-type tobacco mitotic cyclin were fused in frame with the chloramphenicol acetyltransferase (CAT ) reporter gene and constitutively expressed in transformed tobacco BY2 cells. For
both cyclin types, the N-terminal domains led the chimeric cyclin–CAT fusion proteins to oscillate in a cell cycle–specific manner. Mutations within the destruction box abolished cell cycle–specific proteolysis. Although both fusion proteins were degraded after metaphase, cyclin A–CAT proteolysis was turned off during S phase, whereas that of cyclin
B–CAT was turned off only during the late G2 phase. Thus, we demonstrated that mitotic cyclins in plants are subjected
to post-translational control (e.g., proteolysis). Moreover, we showed that the proteasome inhibitor MG132 blocks BY2
cells during metaphase in a reversible way. During this mitotic arrest, both cyclin–CAT fusion proteins remained stable.
INTRODUCTION
The cell division cycle is a sequential process that permits
the replication of the genome, the segregation of chromosomes to two daughter nuclei, and finally cytokinesis. Progression through the cell cycle involves many proteins,
which are regulated both at the transcriptional and posttranslational levels. Today, it has become clear that the
timed destruction of key proteins plays an essential role in
cell cycle progression. The first protein shown to be subjected to cell cycle–specific proteolysis was cyclin B (Evans
et al., 1983). For many other proteins, cell cycle–specific
proteolysis has been reported: among them are the G 1 cyclins and the mitotic cyclins, the cyclin-dependent kinase
inhibitors (CKIs), proteins involved in sister chromatid separation, and spindle components (reviewed in Peter and
Herskowitz, 1994; Murray, 1995; King et al., 1996; Hoyt,
1997). The degradation of cell cycle regulator proteins is not
just a way to break down a protein whose function is
achieved at a specific step of the cell cycle, but it is also a
way to directly control the cell cycle. Thus, the highly regulated proteolysis of CKIs allows DNA replication to begin,
and degradation of proteins involved in sister chromatid
separation is required at the onset of anaphase.
Proteins subjected to degradation are marked with ubiquitin tags and subsequently are targeted to the degradative
1 To whom correspondence should be addressed. E-mail Pascal.
[email protected]; fax 33-3-88-61-44-42.
action of the 26S proteasome. The ubiquitin/26S proteasome proteolytic pathway is highly conserved in eukaryotes
and is involved in many other important cellular functions
aside from cell cycle progression (reviewed in Hochstrasser,
1995). Degradation via this pathway is a two-step process:
the protein is first tagged by the covalent attachment of
ubiquitin; subsequently, it is degraded by a multicatalytic
protease complex called the 26S proteasome. Conjugation
of ubiquitin to the protein involves a cascade of three enzymes: E1, E2, and E3. The E1 (ubiquitin-activating) enzyme
forms a high-energy bond with ubiquitin, which is then
transesterified to a ubiquitin-conjugating enzyme (E2). The
transfer of ubiquitin to the target protein substrate usually
requires ubiquitin ligase activity (E3).
Proteolysis during mitosis requires specific E3 activity located on a large complex called the APC or cyclosome (reviewed in King et al., 1996; Townsley and Ruderman, 1998).
At the onset of and during anaphase, several key proteins are
degraded, for example, mitotic cyclins (reviewed in Murray,
1995), the Saccharomyces cerevisiae anaphase inhibitor
Pds1p (Cohen-Fix et al., 1996), the Schizosaccharomyces
pombe anaphase inhibitor Cut2p (Funabiki et al., 1996), the
cohesion protein Scc1p (Michaelis et al., 1997), and a protein associated with the mitotic spindle Ase1p (Juang et al.,
1997). Although the proteolysis of all these proteins is mediated by the APC complex, the timing of their respective
degradation differs, indicating a further element in the
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complexity of the degradation machinery. In vertebrates, cyclin A is degraded during metaphase, whereas cyclin B is
degraded later during mitosis (Minshull et al., 1990; Whitfield
et al., 1990; Hunt et al., 1992); in budding yeast, Pds1p proteolysis precedes that of cyclin Clb2p (Cohen-Fix et al.,
1996). Thus, to complete the degradation of an APC substrate at the appropriate time, other proteins are involved in
the recognition of the substrates and the regulation of the
proteolytic machinery.
The only structural motif identified in APC substrates is a
nine–amino acid sequence called the destruction box (D
box) motif in the N-terminal domain of the targeted proteins.
This motif seems to be critical for the stability of the proteins
because mutations in or removal of the box stabilizes cyclins A and B in cell-free systems (Glotzer et al., 1991; Luca
et al., 1991; Lorca et al., 1992a; Holloway et al., 1993; van
der Velden and Lohka, 1993; Brandeis and Hunt, 1996), in
transfected and stably transformed vertebrate cells (Gallant
and Nigg, 1992; Brandeis and Hunt, 1996), in frog oocytes
(Murray et al., 1989; Luca et al., 1991), in Drosophila embryos (Rimmington et al., 1994; Sigrist et al., 1995), in budding yeast (Surana et al., 1993; Amon et al., 1994), and in
fission yeast (Yamano et al., 1996). Furthermore, replacement of the D box sequences of Xenopus B1 and B2 cyclins
by the cyclin A1 D box sequence stabilizes the proteins in
the frog egg extracts, indicating that D box signals may not
be interchangeable (Klotzbücher et al., 1996). Deletion of the
N-terminal domain of Cut2p, which contains two sequences
similar to the cyclin D box, stabilizes the protein, and it was
shown that both sequences are required for proteolysis during anaphase (Funabiki et al., 1996). Degradation of Pds1p,
the S. cerevisiae ortholog of Cut2p, also requires an intact D
box (Cohen-Fix et al., 1996). Finally, the Ase1 protein contains five sequences similar to the D box motif, and mutation
of one of them stabilizes the protein (Juang et al., 1997).
No data on proteolysis during the cell cycle have been reported thus far for plants. Significantly, however, a putative
D box is conserved in all mitotic plant cyclins (reviewed in
Plesse et al., 1998; Renaudin et al., 1998). Moreover, by using indirect immunofluorescence analysis, it was shown recently that the level of maize cyclin 1b declines dramatically
during anaphase in root tip cells (Mews et al., 1997), supporting the notion of B-type cyclin destruction during mitosis.
As a first step to study cyclin proteolysis in plants, we investigated the stability of fusion proteins in which the N-terminal domains of tobacco mitotic cyclins were fused in
frame with the chloramphenicol acetyltransferase ( CAT ) reporter gene. We decided to use two mitotic cyclins: the
A-type cyclin (CycA105; Reichheld et al., 1996) and the B-type
cyclin (NTCYC1; Qin et al., 1995). Cyclin NTCYC1 belongs
to the CycB1 group and was renamed Nicta;CycB1;1, and
cyclin CycA105 belongs to the CycA3 group and was renamed Nicta;CycA3;1 according to the classification of
Renaudin et al. (1998). The two cyclins are expressed at different times during the cell cycle. A-type cyclin mRNAs accumulate from S phase until the end of the G 2 phase,
whereas B-type cyclin mRNAs were found to accumulate
exclusively from late G2 to early mitosis (Setiady et al., 1995;
Qin et al., 1996; Reichheld et al., 1996). The N-terminal domains of both cyclins were fused to a reporter gene under
the control of the cauliflower mosaic virus (CaMV) 35S constitutive promoter, and stably transformed tobacco BY2 cell
lines were established. We demonstrate that the levels of
the chimeric proteins oscillate during the cell cycle and that
this oscillation is dependent on an intact D box. Furthermore, we used the proteasome inhibitor carbobenzoxylleucinyl-leucinyl-leucinal (MG132) and show that it induces
cell cycle arrest in tobacco BY2 cells during metaphase and
inhibition of the proteolysis of chimeric cyclin–CAT fusion
proteins.
RESULTS
Establishing Stably Transformed Tobacco BY2 Cell
Lines Constitutively Expressing the Chimeric
Cyclin–CAT Fusion Proteins
To study mitotic cyclin protein oscillations during the cell cycle, we fused the N terminus of an A-type cyclin (Nicta;
CycA3;1) and a B-type cyclin (Nicta;CycB1;1) to the bacterial CAT reporter enzyme (constructs CycA105–CAT and CycB–
CAT; Figure 1A). Both cyclins carry the D box motif (RxxLxx[L/I]xN) in their N-terminal domain (where R and L residues are highly conserved and x stands for any amino acid).
The dependence of reporter protein turnover on the intact D
box was also investigated. In constructs CycA105mutDbox–
CAT and CycBmutDbox–CAT (Figure 1A), we mutated the
two highly conserved amino acids of the D box motifs from
RxxLxx(L/I)xN to GxxVxx(L/I)xN. In vertebrates, it has been
shown that these two mutations abolish the mitotic instability of cyclin B1 (Brandeis and Hunt, 1996). The chimeric
genes were cloned into a binary vector under the control of
the constitutive CaMV 35S promoter rather than the endogenous cyclin promoters to avoid transcriptional regulation of
the constructs during the cell cycle. The constructs were introduced into tobacco BY2 cells by Agrobacterium-mediated transformation.
For all constructs, two to three independently transformed
cell lines were established and propagated in liquid medium.
After several subcultures, the transformant growth curves
were indistinguishable from that of the untransformed BY2
cell culture (shown in Figure 1B with one transgenic cell line
per construct), and CAT mRNAs accumulated to the same
level in asynchronously growing cell cultures (shown in Figure 1C with one of the CAT control transgenic cell lines). The
accumulation level of CAT fusion proteins in the different
transgenic cell lines showed a broad range of variation (5 to
2000 pg per 50 mg of total protein; data not shown). For example, the CAT fusion protein levels determined at mid-log
growth phase from three different CycB–CAT transgenic
Cell Cycle–Dependent Proteolysis in Plants
2065
cell lines (CycB–CAT;10oct, CycB–CAT;22sept, and CycB–
CAT;24oct) were 4, 30, and 400 pg per 50 mg of total protein, respectively. Nevertheless, despite this variation, the
growth curves of all transgenic lines were comparable with
those of the wild-type BY2 cell culture, indicating that the
expression of the chimeric protein is not toxic and does not
disturb the cell cycle.
After synchronization of the CAT control transgenic cell
culture by a 24-hr treatment with aphidicolin (an inhibitor of
DNA polymerase), mRNA accumulation under the control of
the CaMV 35S promoter was constant throughout the cell
cycle, as shown by RNA gel blot analysis (Figure 2B). Rehybridization of the same blot with the histone H4 probe and
the mitotic cyclin B probe, respectively, showed the characteristic S and M phase mRNA accumulation patterns. Recently, Ito et al. (1998) also reported the constitutive
expression of the firefly luciferase gene under the control of
the CaMV 35S promoter throughout the BY2 cell cycle. All
synchronized cell cultures were analyzed by RNA gel blotting and always showed the same constitutive expression of
the chimeric constructs throughout the cell cycle (data not
shown). Total protein was extracted from synchronized CAT
control cultures, and the CAT protein level was determined.
As expected, no significant changes of the CAT protein level
were observed during the cell cycle (Figure 2A).
Proteolysis of the Chimeric Cyclin–CAT Proteins
throughout the Cell Cycle
In a synchronized CycB–CAT cell culture, the chimeric protein started to accumulate during the G 2 phase of the cell
cycle and reached its maximum level 1 hr earlier than the
peak in the mitotic index (Figure 3). At this time, many of the
cells passed metaphase. The decrease in protein level paralleled the decline in the mitotic index and both stayed very
low at the G1-to-S9 transition (S9 indicates the second DNA
replication phase). The difference in chimeric protein levels between mitosis and interphase was z10-fold. For CycA105–
CAT constructs, the chimeric protein levels decreased similarly at the end of mitosis. However, the level of CycA105–
CAT protein was always higher during S phase (directly after
aphidicolin release) when compared with the level during G1
phase (13 to 16 hr) and was high during G 2. In addition,
Figure 1. Growth Curves and Gene Expression Analysis of Stably
Transformed Tobacco BY2 Cell Cultures.
(A) Schematic structure of the constructs inserted between the
BamHI and SacI restriction sites of the binary vector pBI121.1. The
constructs were put under the control of the CaMV 35S promoter.
Open boxes represent the N-terminal domain of the cyclins, and
gray boxes indicate the CAT sequence. Vertical black rectangles indicate the mutated D box.
(B) Growth curves of the cell suspension cultures determined as the
log of fresh weight obtained from 3 mL of culture.
(C) Gel blot analysis of RNAs extracted at different times during the
culture of a CAT contol line. Twenty micrograms of total RNA was
separated by electrophoresis on an agarose–formaldehyde gel,
transferred to a nylon membrane, and hybridized successively with
the indicated probes.
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Figure 2. Accumulation Patterns of the CAT Protein and mRNAs during the Cell Cycle for the CAT Control Transgenic BY2 Cell Line.
(A) After aphidicolin removal, the progression throughout the cell cycle was monitored by 3H-thymidine incorporation (solid circles) and
mitotic index determination (open diamonds). The CAT protein level
was determined at 1-hr intervals (solid squares). This experiment
was performed twice with the cell line CAT control;24oct, and both
experiments gave the same result.
(B) Gel blot analysis of RNAs extracted during the synchronization
experiment at S, G2, M, and G1 phases of the cell cycle. Twenty micrograms of total RNA was separated by electrophoresis on an agarose–formaldehyde gel, transferred to a nylon membrane, and
hybridized successively with different probes as indicated.
during S9 phase, the CycA105–CAT chimeric protein level
started to increase again (earlier than CycB–CAT protein).
Thus, although both fusion proteins are very unstable at the
end of mitosis, their accumulation pattern during the cell cycle is different. For constructs CycBmutDbox–CAT and
CycA105mutDbox–CAT, mutations inside the D box motif
clearly abolished the cell cycle–specific oscillations of the
fusion proteins (Figure 3). To address the problem of reproducibility, we performed the synchronization experiments
with at least two independently transformed cell lines per
construct. For constructs CycB–CAT and CycA105–CAT, we
used three independently transformed cell lines. For a given
construct, we observed that the oscillation patterns during
the cell cycle of the ectopically expressed fusion protein
were identical in high- or low-expressing transgenic cell cultures (data not shown).
To confirm that the proteolysis of CycA105–CAT fusion
protein is already turned off during S phase, we followed the
accumulation of the CycA105–CAT protein from 14 hr after
aphidicolin release until 25 hr (roughly from G1 to the second
mitosis; Figure 4A). In this synchronization experiment, the
mitotic indexes for the first and second mitotic peak were 40%
(data not shown) and 20% at 9 and 25 hr, respectively, after
aphidicolin release. The level of CycA105–CAT protein during the first mitosis reached a value of 22 pg per 50 mg of total protein (data not shown). Under these conditions, the level
of CycA105–CAT protein stayed low during G 1 phase and
started to accumulate during mid-S9 phase. In contrast to the
cyclin A construct, the CycB–CAT fusion protein was present at low levels in a similar experiment, indicating that proteolysis of the B-type cyclin is maintained during DNA
replication (data not shown). These results indicate that proteolysis of the A-type cyclin–CAT chimeric protein is turned
off during S phase, which is earlier than for the cyclin B construct.
Unfortunately, we could not determine exactly when during mitosis the cyclin fusion protein proteolysis is turned on.
From different synchronization experiments, we found that
degradation of both CycA105–CAT and CycB–CAT fusion
proteins only started when anaphase figures appeared (data
not shown). Furthermore, we checked the stability of the
chimeric proteins in metaphase-arrested cells. The two
CycA105–CAT and CycB–CAT cultures were first synchronized with aphidicolin and then treated with the antitubulin
drug oryzalin during early mitosis (when z5 to 10% of
prophases could be counted) and reached a metaphase arrest of z40% (Figure 4B). Under these conditions, after 4 hr
of culture in the presence of oryzalin, .95% of the mitotic
cells were in metaphase (the remainder being in prophase).
Thus, we could show that in both cultures, the fusion proteins accumulated and were stable in the metaphaseblocked cells. Similar results were obtained when we used
the antitubulin drug propyzamide (data not shown). From
these observations, we can only conclude that cyclin fusion
protein degradation is a postmetaphase event occurring
during anaphase.
Cell Cycle–Dependent Proteolysis in Plants
The Proteasome Inhibitor MG132 Induces Metaphase
Arrest in Tobacco BY2 Cell Cultures
Taken together, the results presented above indicate that
plant mitotic cyclins are degraded by the D box pathway.
Because the final stage in degradation of D box–containing
proteins involves the 26S proteasome, we decided to test
the effect of MG132, which is a very efficient proteasome inhibitor in mammalian cell cultures (Rock et al., 1994).
MG132 has no effect on wild-type yeast cells because they
are impermeant to this agent; experimentation requires the
use of mutant strains with enhanced permeability to the
drug (Lee and Goldberg, 1996). Here, we demonstrate that
MG132 can be used in plant cell suspension cultures at a fi-
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nal concentration of 100 mM. The concentration effect and
cytotoxicity of the drug for plant cells will be described elsewhere (M.C. Criqui, Y. Parmentier, A.C. Schmit, and P.
Genschik, manuscript in preparation).
To examine the mitotic effect of MG132 on plant cells,
BY2 cells were first synchronized with aphidicolin, and the
drug was added during G2 phase (at 4.5 hr after aphidicolin
release, as soon as the first prophases were observed) or
early mitosis (at 6.5 hr after aphidicolin release, when z10%
of mitotic cells, mostly in prophase, were observed). As a
control, cells were subcultured in the presence of DMSO
alone; no effect on mitotic progression could be detected
(Figure 5A). In the presence of MG132, the BY2 cells could
clearly enter mitosis and progress through prophase, but
Figure 3. Accumulation Patterns of the Cyclin–CAT Fusion Proteins during the Cell Cycle.
The progression through the cell cycle was monitored by 3H-thymidine incorporation and mitotic index determination. The curves presented in
this figure were obtained with cell lines CycB–CAT;24oct (CycB–CAT), CycBmutDbox–CAT;10oct (CycBmutDbox–CAT), CycA105–CAT;22sept
(CycA105–CAT), and CycA105mutDbox–CAT;24oct (CycA105mutDbox–CAT). Symbols are as given for Figure 2.
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Figure 4. Off and On of the Proteolytic Machinery during the Cell
Cycle.
(A) Stability of the CycA105–CAT fusion protein from G1 to the second mitosis. The progression through the cell cycle was monitored
by 3H-thymidine incorporation and mitotic index determination. This
synchronization experiment was conducted with the cell line
CycA105–CAT;10oct (CycA105–CAT). Symbols are as given for Figure 2.
(B) Stability of the cyclin–CAT fusion proteins in metaphase-arrested
cells. The level of the fusion proteins was determined in S phase (1
hr after aphidicolin removal), in early mitosis (7 hr after aphidicolin
removal; the time when oryzalin was added), and at different times
during the culture in the presence of the antitubulin drug. In these
experiments, we used cell lines CycB–CAT;10oct (CycB–CAT) and
CycA105–CAT;10oct (CycA105–CAT).
they were arrested at metaphase with fully condensed chromosomes congregating at the metaphase plate (Figures 5B
and 6A). If MG132 was added at 6.5 hr of culture, almost
50% of the cells could be blocked in metaphase. Immunofluorescent staining of the microtubules in drug-treated cells
showed that metaphase spindles and the interphase microtubules appeared to be normal (Figures 6E and 6F)
compared with the untreated control BY2 cells (Figures 6C
and 6D).
We also assayed histone H1 kinase activity in protein extracts from control and MG132-treated cells. Protein samples from untreated and drug-treated cells (at 8.5 and 13.5 hr
after aphidicolin release; indicated by an asterisk in Figure
5B) were extracted by using a p13 suc1–Sepharose matrix,
which is commonly used to isolate cyclin-dependent kinase
(CDK)–cyclin complexes (Labbé et al., 1991). The activity of
the bound kinases was monitored by the in vitro phosphorylation of the histone H1 protein, which was used as a substrate (Figure 5C). In contrast to untreated cells, in which H1
kinase activity decreased at 13.5 hr of culture, activity remained elevated in the blocked cells. H1 kinase activity was
already higher at 8.5 hr in the MG132-treated cells, in which
the percentage of cells in prophase and metaphase was
higher than in the control.
The addition of MG132 later during mitosis, at a time
when most of the cells had passed metaphase, showed aberrant anaphase activity and also cells with very enlarged
nuclei, which often underwent amitotic divisions (Figure 6B),
indicating that 26S-dependent proteolysis is also involved in
other steps during mitosis and not just the metaphase-toanaphase transition. The effect of the drug on those cells is
currently under investigation. We also used another peptide
aldehyde, carbobenzoxyl-leucinyl-leucinyl-norvalinal (MG115),
and found that in BY2 cell cultures, the drug had the same
effect, although MG132 was slightly more effective (data not
shown).
MG132 is a peptide aldehyde that functions as a substrate analog. To determine whether metaphase arrest induced by the drug could be reversed, we added 100 mM
MG132 to aphidicolin-synchronized cells during early mitosis. After 4 hr of subculturing in the presence of the drug,
50% of the cells were arrested during metaphase. When
subjected to extensive washes, the cells were able to exit
mitosis after 4 hr (Figure 5D). For comparison, after the
metaphase arrest produced by the antitubulin drug propyzamide, the cells exited mitosis in ,2 hr. However, even
though metaphase arrest induced by the drug is reversible,
many aberrant anaphase figures were observed (data not
shown).
MG132 Stabilizes the Chimeric Cyclin–CAT
Fusion Proteins
We examined the effect of the proteasome inhibitor on
chimeric cyclin–CAT protein stability. CycB–CAT and CycA105–
Cell Cycle–Dependent Proteolysis in Plants
CAT transgenic cell cultures were first blocked with aphidicolin. In early mitosis after aphidicolin release (when z5 to
10% of prophases were observed), half of the cultures were
treated with MG132. Mitotic indexes and CAT protein levels
were determined at 1-hr intervals in cell cultures either treated
or not with MG132 (Figure 7A). As reported above, the transition from prophase to metaphase was not affected by the
2069
drug, and the treated cells accumulated during metaphase.
For both cultures, the cyclin–CAT proteins accumulated and
remained stable during drug treatment. However, the level
of the cyclin–CAT fusion protein was always lower in the
MG132-treated cells than in the controls. RNA gel blot analysis indicated that the drug has an effect on transcription or
RNA stability because the cyclin–CAT mRNA level was
Figure 5. Metaphase Arrest in MG132-Treated Synchronized BY2 Cells.
(A) The mitotic index throughout the cell cycle in aphidicolin-synchronized BY2 cells in the presence (1) and absence (2) of 0.2% DMSO. Arrows indicate the times after aphidicolin release at which MG132 was added (4.5 hr [1] or 6.5 hr [2]) .
(B) Percentage of mitotic cells (in prophase, metaphase, anaphase, and telophase) in untreated and MG132-treated BY2 cell cultures. The asterisks indicate the times chosen to assay histone H1 kinase activity. The 1 and 2 indicate the times at which MG132 was added (see [A]).
(C) Histone H1 kinase assays were carried out with p13suc1–Sepharose—bound protein fractions from untreated (2) and drug-treated (1) cell
cultures. Histone H1 loading was controlled by Coomassie Brilliant Blue R 250 staining (data not shown). Phosphorylated histone H1 was visualized by autoradiography and estimated by direct Cerenkov counting of the labeled bands.
(D) Percentage of mitotic cells after MG132 release.
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already lower 3 hr after the drug treatment (Figure 7B). Furthermore, we obtained a similar mRNA accumulation pattern
when we used the constitutively expressed Arabidopsis
EF-Tu elongation factor as a probe (data not shown).
Nevertheless, MG132 does not affect the level of all mRNAs
equally, because inhibition of the 26S proteasome by this drug
induces accumulation of the polyubiquitin Ubi.U4 mRNA. Recently, an increase of heat shock hsp70 mRNAs was also
found in HepG2 cells treated with different proteasome inhibitors (Zhou et al., 1996), and induction of heat shock proteins after drug treatment was reported both in mammalian
and yeast cells (Bush et al., 1997; Lee and Goldberg, 1998).
However, the polyubiquitin Ubi.U4 gene is not readily heat
shock inducible (Genschik et al., 1994a), and it is not clear
whether the signaling that activates this gene uses the same
pathway as that which activates the hsp genes. Nonetheless, our results indicate that a defect in proteolysis by the
26S proteasome can lead to the activation of at least one
component of this proteolytic pathway (e.g., a polyubiquitin
gene).
Figure 6. Microscopic Observations of the Effects of MG132 on Mitosis in BY2 Cells.
(A) Metaphase-arrested cells after MG132 treatment stained with
49,6-diamidino-2-phenylindole (DAPI).
(B) Aberrant mitotic figures visualized with DAPI when MG132 was
added late in mitosis.
(C) and (D) DAPI and immunofluorescence staining of microtubules,
respectively, of a control BY2 cell in metaphase.
(E) and (F) DAPI and immunofluorescence staining of microtubules,
respectively, of two metaphase-arrested cells after MG132 treatment.
DISCUSSION
In recent years, the molecular characterization of plant
genes involved in cell cycle control (e.g., CDKs, cyclins,
CKIs, and retinoblastoma protein homologs) suggests that
the basic mechanisms controlling the cell cycle are also
highly conserved in plants. However, until now, the function
of proteolysis during plant cell cycle progression has not
been addressed. In this study, we used two approaches: (1)
we analyzed the stability of mitotic cyclin–CAT fusion proteins in synchronized BY2 cell cultures; and (2) we studied
the effect of the proteasome inhibitor MG132 on mitosis.
Previously, it was shown that the highly conserved D box
motif RxxLxxIxN in the N terminus of mitotic cyclins and other
APC substrates is important for their degradation (see the
Introduction). In this study, we showed the function of two different tobacco mitotic cyclin N-terminal domains in cell cycle–dependent proteolysis. In the constructs tested, the cyclin
core regions were replaced by the CAT reporter protein,
thereby preventing the chimeric protein from interacting with
CDK proteins. For the Xenopus cyclins A and B2, it was previously shown that mutations affecting the binding of CDK
stabilize these cyclins, even if the D box is intact (Stewart et
al., 1994; van der Velden and Lohka, 1994), indicating that proteolysis of those cyclins requires their binding to CDK1. Nevertheless, this interaction is not required for Xenopus cyclin B1
and sea urchin cyclin B (Glotzer et al., 1991; Holloway et al.,
1993; Stewart et al., 1994; van der Velden and Lohka, 1994).
Moreover, fusion of the 105 N-terminal residues of mouse
cyclin B1 to the CAT reporter enzyme led to the degradation of
chimeric proteins at the end of mitosis in stably transformed
NIH3T3 cells, as occurred with the endogenous B-type cyclin
(Brandeis and Hunt, 1996). Curiously, a similar experiment in
which the 86 N-terminal residues of mouse cyclin B2 were
fused to the CAT reporter enzyme gave the same results
(Brandeis and Hunt, 1996). Our data clearly indicate that all
of the information necessary for cell cycle–specific breakdown for both tobacco A-type and B-type cyclins is located
in the N-terminal domains of the proteins and that interaction with a CDK partner is not required for degradation of the
chimeric proteins. In addition, the degradation of the chimeric
proteins is dependent on an intact D box. Thus, we demonstrate that in plants as well as in animals, a proteolytic pathway is activated at the end of metaphase and that this
pathway recognizes the highly conserved D box motif.
The accumulation pattern of the CycB–CAT fusion protein
during the BY2 cell cycle is similar to the expression pattern
of the tobacco B-type cyclin genes (Setiady et al., 1995; Qin
et al., 1996; Reichheld et al., 1996; results not shown), despite the constitutive expression of the chimeric construct.
Thus, if the chimeric fusion protein reflects the endogenous
tobacco B-type cyclin level, as is the case for the mouse B1
cyclin (see above), then the protein accumulates during late
G2 phase and is degraded after metaphase. The window of
stability of the CycA105–CAT fusion protein is wider than
Cell Cycle–Dependent Proteolysis in Plants
2071
that for the B-type cyclin. During S9 phase, the chimeric protein level started to increase again and was high throughout
the G2 phase. The endogenous mRNAs of this cyclin also accumulate during the S and G2 phases (Reichheld et al., 1996).
Thus, the patterns of protein stability for both cyclins during
the cell cycle are consistent with the expression patterns of
their respective genes.
In Xenopus extracts, it has been shown that human cyclin
B—but not cyclin A—is able to turn on the proteolytic machinery (Lorca et al., 1992b). This activation is most probably controlled by the phosphorylation of APC and is not
directly induced by cyclin B–Cdc2; however, at least one
downstream kinase is required for phosphorylation (reviewed
in Townsley and Ruderman, 1998). Recently, evidence was
provided that the mouse Polo-like kinase activates APC by
phosphorylation after the kinase itself is activated during
early mitosis by cyclin B–Cdc2 (Kotani et al., 1998). The
rapid increase in accumulation of the CycB–CAT fusion protein just before the D box pathway is activated suggests that
this may also be the case for plants.
Even if the stability of the plant mitotic cyclins during the cell
cycle is similar to that in the animal model, there are at least
two important differences. (1) The tobacco cyclin B fu sion
protein accumulates only late during G2 phase, and the window of stability is very sharp, whereas animal mitotic B-type
cyclins are already starting to accumulate during S phase
(Pines and Hunter, 1989; Jackman et al., 1995; Brandeis and
Hunt, 1996). Meanwhile, the CycA105–CAT protein has already accumulated during S phase, at a time when CycB–
CAT degradation occurs. The differences in stability between both fusion proteins during S phase and early G 2
phase could be explained by the fact that the CycA105–CAT
construct may carry information at its N terminus that protects the chimeric protein from degradation. Alternately, the
CycB–CAT fusion protein may be a much better substrate
for proteolysis than is the CycA105–CAT protein at a time
when the proteolytic machinery is not fully active. (2) In animal cells, degradation of cyclin A occurs during metaphase,
which is earlier than for cyclin B. Cyclin B is degraded during
anaphase, and the spindle assembly checkpoint cannot prevent its destruction (Minshull et al., 1990; Whitfield et al.,
1990; Hunt et al., 1992). In BY2 cells, however, the degradation of both cyclin A and B fusion proteins starts only after
metaphase, and their proteolysis was not detected in
Figure 7. Accumulation Patterns of the Cyclin–CAT Protein and
mRNAs during the Cell Cycle in Untreated and MG132-Treated
CycB–CAT and CycA105–CAT Cell Cultures.
(A) Progression throughout the cell cycle was monitored by mitotic
index determination (open diamonds) in untreated ( 2MG132) and
drug-treated (1MG132) cell cultures. The fusion protein level is indicated by solid squares. The arrows indicate the times at which 100 mM
of MG132 was added. In the treated cultures, cells arrested exclusively in metaphase. In these experiments, we used cell lines CycB–
CAT;10oct (CycB–CAT) and CycA105–CAT;10oct (CycA105–CAT).
(B) Gel blot analysis of RNAs extracted at different times from untreated (2) and MG132-treated (1) CycB–CAT cell cultures. The 0,
1, 3, 5, and 8 hr correspond, respectively, to 6.5, 7.5, 9.5, 11.5, and
14.5 hr after aphidicolin release. Twenty micrograms of total RNA
was separated by electrophoresis on an agarose–formaldehyde gel,
transferred to a nylon membrane, and hybridized successively with
the indicated probes. The same results were obtained with untreated and drug-treated CycA105–CAT cell cultures (data not
shown).
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propyzamide- and oryzalin-treated cells arrested in metaphase. Unfortunately, we could not determine exactly the
point during mitosis at which cyclin proteolysis is turned on.
Curiously, with plants, protein localization experiments
have indicated that the maize cyclin Zeama;CycAI;1, which
is an A-type cyclin belonging to the cycA1 group, is not degraded during anaphase but instead becomes strongly associated with microtubules in the phragmoplast (Mews et
al., 1997). Thus, degradation of certain cyclins during the
plant cell cycle may not be required, which suggests that
these proteins might be involved in late mitotic functions.
Furthermore, specific antibodies raised against three different tobacco A-type cyclins from group CycA3 (including
Nicta;CycA3;1 presented in this work) were used to quantify
the relative abundance of the cyclins on immunoblots and also
indicated a constitutive protein level during the BY2 cell cycle
(B. Combettes and N. Chaubet, personal communication).
These observations raise at least two questions. Why do
A-type cyclins carry the D box motif if they are not subjected
to cell cycle–dependent proteolysis? Why is the N terminus
of Nicta;CycA3;1 cyclin able to target a reporter protein to
postmetaphase-specific degradation? If A-type cyclins in plants
are stable during the cell cycle, then the discrepancy between our results and the localization experiments could be
explained by the lack of interaction between the CycA105–
CAT fusion protein and its kinase partner. Without this
interaction, the CycA105–CAT fusion protein would not be
protected from degradation and would be recognized like
other D box–containing proteins (e.g., cyclin B), which would
be degraded at the metaphase-to-anaphase transition. However, protection solely mediated by association with the kinase is unlikely because for Xenopus cyclins A and B2, the
interaction of these cyclins with the CDK1 kinase does not
protect the cyclins from proteolysis but rather is absolutely
required for their degradation (Stewart et al., 1994; van der
Velden and Lohka, 1994). On the other hand, A-type cyclins
may be protected from degradation by their cellular relocalization after metaphase, and this would necessarily involve
the interaction of the cyclin with its kinase partner. In this
case, our chimeric protein would be incorrectly located and
exposed to the D box proteolytic pathway. However, the cellular localization during the cell cycle of the D box pathway
proteolytic machinery (e.g., APC or the 26S proteasome
complexes) is poorly understood, and it will be of interest to
perform this kind of study in the future.
It is well established that the degradation of the cell cycle
regulatory proteins carrying the D box is carried out by the
26S proteasome in both animal and yeast cells. Thus, we
tested the effects of the tripeptide proteasome inhibitor
MG132 on mitosis. When the drug was added late in the G 2
phase, the BY2 cells were still able to break down the nuclear envelope, set up the mitotic spindles, and congregate
the highly condensed chromosomes to the metaphase
plates. For both cyclin A– and cyclin B–CAT chimeric proteins, the drug led to their stabilization. In mammalian cells,
the same drug led also to mitotic arrest at metaphase with
high H1 kinase activity and stabilization of cyclin B1
(Sherwood et al., 1993). However, the metaphase arrest of
the drug in both mammalian and plant cells cannot be ascribed to cyclin B stabilization because sister chromatid separation can proceed while mitotic CDK is active (Holloway et
al., 1993; Surana et al., 1993). Thus, the metaphase arrest
provoked by MG132 in BY2 cells suggests strongly the existence of a plant protein whose degradation by the proteasome at the metaphase-to-anaphase transition is absolutely
required for sister chromatid separation.
METHODS
Procedures
Unless stated otherwise, all procedures for manipulating DNA and
RNA were conducted according to Sambrook et al. (1989) and
Ausubel et al. (1994).
Proteasome Inhibitors
Carbobenzoxyl-leucinyl-leucinyl-norvalinal (MG115) and carbobenzoxyl-leucinyl-leucinyl-leucinal (MG132) were provided by PEPTIDES
International, Inc. (Louisville, KY). The drugs were dissolved in DMSO
and were never kept for .1 month at 2208C.
Cyclin–Chloramphenicol Acetyltransferase Fusion Constructs
XbaI and SacI sites were introduced 59 and 39, respectively, of the
chloramphenicol acetyltransferase (CAT ) coding sequence from the
Escherichia coli enzyme by polymerase chain reaction (PCR)–based
site-directed mutagenesis. The PCR reaction was performed using
oligonucleotide 1 (59-CTAAGGAATCTAGAATGGAGAAAAAAATC-39)
and oligonucleotide 2 (59-GCACCAAGAGCTCCCTTAAAAAAATTACG-39) as the upstream and downstream primers, respectively, and
plasmid pRT99-CAT as the template (Töpfer et al., 1988). The XbaISacI fragment was cloned into the pBluescript II SK1 (Stratagene, La
Jolla, CA) vector. The same strategy was used to introduce BamHI
and XbaI sites upstream and inside of, respectively, the coding
regions of tobacco cyclin A (CycA105, renamed Nicta;CycA3;1;
Reichheld et al., 1996) and tobacco cyclin B (NTCYC1, renamed
Nicta;CycB1;1; Qin et al., 1995). For cyclin A, we used oligonucleotide 3 (59-TACCCCGGATCCAGTTGTTTGAATG-39) and oligonucleotide 4 (59-TCATTCGTCTAGAAGGAGTCACATC-39); for cyclin B, we
used oligonucleotide 5 (59-CAAGAAGGATCCCTTCAAATGGATAAC-39)
and oligonucleotide 6 (59-ACTTTGGTCTAGAGGAAAGTCCAC-39) as
upstream and downstream primers, respectively.
The PCR reaction led to the amplification of the coding region covering the first 137 amino acids of cyclin A and the first 134 amino acids of cyclin B, roughly from Met (at position 11) to the cyclin box.
Both PCR fragments were cloned into the pBluescript II SK1 vector
and designated pSKCycA105 and pSKCycB, respectively. The cyclin
BamHI-XbaI fragments were subcloned into pSKCAT, leading to
constructs pSKCycA105-CAT and pSKCycB-CAT, in which the cyclin coding regions were in frame with the CAT coding region. In constructs pSKCycA105mutDbox and pSKCycBmutDbox, we mutated
Cell Cycle–Dependent Proteolysis in Plants
two highly conserved amino acids in the destruction box (D box)
motif from RxxLxx(L/I)xN to GxxVxx(L/I)xN by PCR using oligonucleotide 7 (59-ACTGAAAGGAGTCGTGGTTGGCGAG-39) and oligonucleotide 8 (59-AGTGGATCATTTCCATTGGAGG-39) for cyclin A and
oligonucleotide 9 (59-AAGAAATGGACGTGCTGTTGGAGAC-39) and
oligonucleotide 10 (59-CCATCGGCTTGTGCATTTTTCTG-39), respectively, for cyclin B. All PCR constructs were sequenced on both
strands. The BamHI-SacI DNA fragments from each of the five constructs (pSKCAT, pSKCycA105-CAT, pSKCycA105mutDbox-CAT,
pSKCycB-CAT, and pSKCycBmutDbox-CAT) were subcloned into
the binary pBI121.1 vector (Clontech, Palo Alto, CA), replacing the
b-glucuronidase reporter gene.
BY2 Cell Culture, Transformation, and Synchronization
A rapidly growing suspension culture of tobacco BY2 cells (Nicotiana
tabacum cv Bright Yellow 2) was maintained by weekly dilution
(1.5:100) of cells into fresh medium modified according to Nagata et
al. (1992) and cultured at 278C and 130 rpm in the dark. The plasmids
described above were introduced by electroporation into the disarmed Agrobacterium tumefaciens strain LBA4404. BY2 cells were
stably transformed as follows. Four milliliters of a 3-day-old BY2 culture was cocultivated with 100 mL of an Agrobacterium culture in
Petri dishes in the dark for 2 days at 278C. Cells were washed three
times by centrifugation and were grown either in 40 mL of fresh medium or plated on solid medium, both supplemented with carbenicillin (500 mg/mL) and kanamycin (100 mg/mL). When the cultures
reached maximal density (after 1 week), 2 mL of the cultures was
transferred to 80 mL of fresh medium containing carbenicillin and
kanamycin. The cultures were then subjected to four or six rounds of
subculturing until they reached the growth rate of the untransformed
BY2 cell culture. When the transformants were plated on solid medium, 3 weeks of growth were required before calli could be recovered. Between 50 and 300 calli were pooled in liquid medium and
processed as described above.
Tobacco BY2 cells were synchronized according to Nagata et al.
(1992). Briefly, transgenic and control cell suspensions at stationary
phase (7 days old) were subcultured for 24 hr in a medium containing
aphidicolin (3 mg/mL; Sigma), washed with 1 L of sucrose solution at
40 g/L, and then suspended in 100 mL of medium modified according to Nagata et al. (1992). DNA synthesis was determined by subculturing 1 mL from the cell suspensions in the presence of 1.5 mCi 3H-TTP
(94 Ci/mmol; Amersham Corp.) for 30 min at 278C on a rotary shaker.
After extensive washing of the cells, DNA was extracted, and estimation of 3H-thymidine incorporation was carried out as described previously (Genschik et al., 1994b). For mitotic index determination,
cells were stained with 1 mg/mL 49,6-diamidino-2-phenylindole (DAPI;
Sigma) in the presence of 0.2% Triton X-100; interphase, prophase,
metaphase, anaphase, and telophase figures were determined for at
least 600 cells by using UV light microscopy. When analysis of late
(i.e., postmitotic) stages was needed, 1.54 mg/mL of propyzamide
(Sumitomo Chemical Co., Osaka, Japan) or 3.46 mg/mL of oryzalin
was added before the preprophase stage, z5 to 6 hr after aphidicolin
release, maintained for 4 hr, and removed by extensive washes.
RNA Extraction and Gel Blotting
Total RNA from the cell cultures was isolated as described by
Verwoerd et al. (1989). RNA (20 mg per lane) was separated on a
2073
formaldehyde–agarose gel, blotted onto Hybond N (Amersham
Corp.) nylon membrane by capillary transfer using 20 3 SSPE (1 3
SSPE is 0.18 M NaCl, 0.01 M NaH2PO4, and 0.001 M Na2-EDTA, pH
7.7), and UV cross-linked to the membrane. The integrity and the
amount of RNA applied to each lane were verified by control hybridizations using a tomato 25S rRNA probe (Kiss et al., 1989). The histone
H4 probe corresponds to the 196-bp restriction fragment AccI-DdeI
of the coding region of the gene H4A748 (Chaboute et al., 1987). The
probe specific for the polyubiquitin gene Ubi.U4 corresponds to the
626-bp restriction fragment SalI-HindIII of gene Ubi.U4 (Genschik et
al., 1994a). The cyclin B probe corresponds to a Nicta;CycB1;1
cDNA (Qin et al., 1995). The CAT probe corresponds to the complete
CAT coding sequence (Töpfer et al., 1988). The probes were labeled
with a–32P-dCTP (3000 Ci/mmol; Amersham Corp.) by the random
priming method (Feinberg and Vogelstein, 1983). RNA gel blots were
hybridized overnight at 428C in 5 3 SSPE, 50% formamide, 10%
dextran sulfate, 1% SDS, and 50 mg of denatured salmon sperm
DNA. The blots were subsequently washed in 2 3 SSC (1 3 SSC is
0.15 M NaCl and 0.015 M sodium citrate) and 0.1% SDS for 30 min
at 428C and in 0.2 3 SSC and 0.1% SDS for 30 min at 428C.
Protein Extraction, CAT Immunoassay, and Histone H1
Kinase Activity
BY2 cells were homogenized with a Dounce tissue grinder (Wheaton,
Millville, NJ) in extraction buffer (25 mM Tris-HCl, pH 7.5, 15 mM
MgCl2, 15 mM EGTA, 150 mM NaCl, 0.1% Tween-20, 1 mM DT T,
and complete protease inhibitor cocktail mix [Boehringer Mannheim])
and centrifuged at 20,000g. The protein content was determined by
using the Bio-Rad protein assay kit. CAT protein content was determined by using the CAT ELISA kit (Boehringer Mannheim). No crossreacting protein was found in the extracts from nontransformed BY2
cell cultures. For histone H1 kinase activity, all extracts were prepared as described above in the presence of 5 mM NaF. One hundred micrograms of total protein was incubated with 50 mL of 25%
(v/v) p13suc1–Sepharose beads (kindly provided by Lásló Bögre, Institut für Mikrobiologie und Genetik, Vienna, Austria) for 2 hr on a rotary
shaker at 48C. After incubation, the washing conditions of the beads
and the kinase reaction were as published by Magyar et al. (1993).
Immunofluorescence Analysis
Immunofluorescence analysis was done according to Chang-Jie and
Sonobe (1993). Antibodies used were a mouse antibody raised
against tubulin (Amersham Corp.) diluted 1:3000 and fluorescein
isothiocyanate–conjugated goat antibody (Jackson Immunoresearch
Laboratories, West Grove, PA) raised against mouse IgG diluted
1:250.
ACKNOWLEDGMENTS
We thank Catherine Bergounioux for the cyclin Nicta;CycB1;1 cDNA
and Nicole Chaubet for the cyclin Nicta;CycA3;1 cDNA; Bruno
Combettes and Nicole Chaubet for sharing unpublished results;
Tobacco Science Research Laboratory, Japan Tobacco, Inc., for allowing us to use the TBY2 cell suspension; Anne C. Schmit for help
2074
The Plant Cell
with immunofluorescent microtubule staining; Philippe Hammann for
DNA sequencing; and Leon Otten and Dennis Francis for critical
reading of the manuscript.
Received July 6, 1998; accepted September 22, 1998.
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