Download Distinct Classes of Mitotic Cyclins Are Differentially

The Plant Cell, Vol. 7, 1143-1155, August 1995 O 1995 American Society of Plant Physiologists
Distinct Classes of Mitotic Cyclins Are Differentially
Expressed in the Soybean Shoot Apex during the Cell Cycle
Hiroshi Kouchi,ail Masami Sekine,b and Shingo Hataa92
a
Department of Applied Physiology, National lnstitute of Agrobiological Resources, Tsukuba, lbaraki, 305 Japan
Department of Biotechnology, Osaka University, Suita, Osaka, 565 Japan
Protein phosphorylation by the complexes of cyclin and cyclin-dependent kinase plays a key role in cell cycle progression
in all eukaryotes. The amplification by polymerase chain reaction of a cyclin box from developing root nodules and root
apices of soybean showed the expression of a number of different molecular species of mitotic cyclins in plant meristems,
and they were classified into five distinct groups based on their sequence similarities. The complete soybean cyclin cDNAs,
cyclGm to cycSGm, corresponding to each group were isolated, and their predicted amino acid sequences showed clear
similarities to mitotic cyclins identified from various organisms. These genes are expressed predominantly in such
meristematic tissues as root and shoot apices and young developing nodules. Double-target in situ hybridization involving histone H4 as an S-phase marker allowed us to estimate the phases during which these cyclin genes are abundantly
expressed. The results indicated that cyc5Gm is expressed in GP-to-Mphases and cyc3Gm is expressed from late S-toG2 phases. These expression patterns, together with the sequence criteria, strongly suggest that cyc3Gm and cyc5Gm
encode the plant cognates for A- and B-type cyclins, respectively. In addition, the expression of cyclGm was restricted
during a short period in S phase, suggesting that it belongs to a nove1 class of plant cyclins. Sequence comparison of
18 plant mitotic cyclins cloned thus far showed that they can be divided into four distinct structural groups with different
functions in cell cycle progression.
INTRODUCTION
The cell division cycle in eukaryotes is controlled by common
mechanisms in which heterodimeric protein kinases play the
central role. These protein kinases consist of the catalytic
subunit, cyclin-dependent kinases (cdks), and their regulatory
subunit, cyclins. The association of the cdks with specific cyclins, together with phosphorylation and dephosphorylation
of cdks, is required for most eukaryotic cell cycle transitions
(for reviews, see Norbury and Nurse, 1992; King et al., 1994;
Nurse, 1994).
Recent advances in the study of the eukaryotic cell cycle
have shown the presence of a number of structurally homologous cdks and cyclins in higher eukaryotes. Among them, the
role of the association between p34CdC2,
the 34-kD protein kinase encoded by the fission yeast cdc2 homologs, and mitotic
cyclins is the best understood. Cyclins were first identified in
marine invertebrates (Evans et al., 1983; Swenson et al., 1986);
the p34cdc2-cyclincomplexes must function for cells to undergo mitosis, that is, to enter into M phase. The M-to-G,phase transition is accompanied by the dissociation of these
complexes and the breakdown of cyclins (for review, see Hunt,
1991; Surana et al., 1993; King et al., 1994).
To whom correspondence should be addressed.,
* Current address: Faculty of Agriculture, Kyoto University, Kitashirakawa-Oiwakecho, Sakyo-ku, Kyoto, 606 Japan.
The animal mitotic cyclins have been subdivided into A- and
B-type cyclins, according to differences in their sequences and
expression patterns during the cell cycle. Although A- and
B-type cyclins share many structural and functional similarities, they do not appear to be redundant in cell cycle regulation
in many vertebrates. 60th have been implicated in the G2-toM-phase transition, activation of the p34cdc2protein kinase,
and induction of the entry into M phase (Minshull et al., 1989).
However, A-type cyclins appear to piay a role in DNA replication during S phase, in addition to their role in progression
through GP phase (Girard et al., 1991; Pagano et al., 1992;
Zindy et al., 1992). It has also been suggested that the substrate specificity of cyclin-cdk complexes is differentially
modulated by A- and B-type cyclins (Peeper et al., 1993). In
addition, it has been suggested that even at the Gz-toM-phase transition, A- and B-type cyclins carry out different
functions, because in Drosophila, the B-type cyclin is unable
to compensate functionally for the loss of the A-type cyclin
(Lehner and OFarrell, 1990). The S-phasefunction of the A-type
cyclin probably involves the formation of a complex with
p33cdk2,which is structurally related to but functionally distinct
from p34cdc2,and has been implicated in G, and S phases
(Pines and Hunter, 1990; Pagano et al., 1992). In Saccharomyces cerevisiae, no cyclin A structural counterpart has been
identified. However, CLB5 and CLB6 have been suggested
1144
The Plant Cell
to be functionally equivalent with A-type cyclins, because they
operate in the timing and execution of S phase (Kühne and
Linder, 1993; Schwob and Nasmyth, 1993).
The other subfamilies of cyclins have been identified as G1
cyclins. They are required for completion of G1 phase and
commitment to the entry into S phase. CLN1, CLN2, and CLN3
from S. cerevisiae were the first G1 cyclins to be identified
(Nash et al., 1988; for review, see Reed, 1991). Thus far, mammalian C-, D-, and E-type cyclins have been identified as G1
cyclins (Lew et al., 1991; for review, see Sherr, 1993, 1994).
In S. cerevisiae, CLNs and mitotic cyclins share p34CDC28 as
the only catalytic subunit, whereas mammalian D- and E-type
cyclins seem to form complexes with cdks other than p34cdc2
(Pines, 1993; Sherr, 1993,1994). With the exception of the human E-type cyclin (Lew et al., 1991), these G1 cyclins share
no or very limited structural similarity with mitotic cyclins.
Higher plants grow by continuously differentiating new organs postembryonically from meristem cells, according to
interna1developmental programs or as a result of environmental
stimuli. New organs are normally initiated by the division of
specific groups of cells even in mature plants, and only a few
phytohormones are involved in these processes. Therefore,
higher plants and animals must differ significantly in the control of the cell division cycle (Jacobs, 1992). We first identified
plant cyclins from soybean and carrot (Hata et al., 1991). Subsequently, a number of cyclins have been described from a
wide variety of plants, that is, Arabidopsis (Hemerly et al., 1992;
Ferreira et al., 1994), alfalfa (Hirt et al., 1992), Antirrhinum
(Fobert et al., 1994), maize (Renaudin et al., 1994), and tobacco
(WSetiady, M. Sekine, N. Hariguchi, T. Yamamoto, K. Yoshida,
H. Kouchi, and A. Shinmyo, unpublished data). These cyclins
all share conserved “cyclin box” structures, and some of them
are able to induce maturation of Xenopus oocytes (Hata et al.,
1991; Renaudin et al., 1994) or complement a CLN mutant of
S. cerevisiae (Y.Y. Setiady, M. Sekine, N. Hariguchi, T.
Yamamoto, K. Yoshida, H. Kouchi, and A. Shinmyo, unpublished data). In addition, G1 cyclin candidates related to cyclin
D have recently been isolated from Arabidopsis, and they complement S.cerevisiae CLN mutants (Soni et al., 1995). A number
of functional and/or structural homologs of cdks have also been
isolated from higher plants (Colasanti et al., 1991; Feiler and
Jacobs, 1991; Ferreiraet al., 1991; Hashimotoet al., 1991; Hata,
1991; Hirayama et al., 1991; Hirt et al., 1991, 1993; Mia0 et al.,
1993).
These results show that a common mechanism of eukaryotic cell cycle regulation is probably conserved in higher plant
cells. However, the available data on regulatory mechanisms
of the cell division cycle in higher plants are still very limited.
In plants, the cell division cycle can be arrested in either the
G1 or the G2 phase; cells arrested at G1or G2 have the capacity to reenter the cell division cycle (van’t Hof, 1985; Bergounioux
et al., 1992). Thus, the availability of mitotic cyclins as well as
of G1 cyclins may be critical for regulating differentiation
and/or dedifferentiation processes in plants. Further characterization of plant cyclins will provide more clues to the
molecular mechanisms that control developmental processes
in higher plants. In this study, we show that soybean plants
express a number of different cyclins in the meristematic tissues. Among the five complete cDNAs isolated, three have
A-like structures and the other two have B-like structures.
Analyses of their expression in shoot meristems by in situ hybridization demonstrated that the transcription of these distinct
classes of cyclin genes is regulated differentially during the
cell cycle in plant meristems.
RESULTS
Cloning of Cyclin cDNAs
Degenerated oligonucleotide primers were designed for a
highly conserved region of published amino acid sequences
of mitotic cyclins (Hata et al., 1991). Polymerase chain reaction (PCR) using these primers with the cDNAs from young
developing nodules or root tips resulted in amplification of
cDNA fragments with the expected length (185 bp). These fragments were subcloned, and 30 clones were sequenced. As
a consequence, 29 clones showed the conserved amino acid
sequences typical of mitotic cyclins (data not shown). These
clones were tentatively divided into five groups (designated
groups I toV) according to structural similarities. Groups I, II,
and III contain four, three, and two different molecular species, respectively, whereas groups IV and V each consist of
only one molecular species. The group V species has a nucleotide sequence identicalto s73-6, which previously was identified
from soybean nodules (Hata et al., 1991). Therefore, we attempted to isolate full-length clones for groups I to IV by
screening the nodule cDNA library using PCR products of each
group as probes. Cloning of cDNAs of groups I and II from
the nodule cDNA library was not successful, probably due to
the Iow leve1 of expression of these cyclins in intact tissues.
However, they could be isolated from a cDNA library of
suspension-cultured cells. Finally, we obtained four soybean
cyclin cDNA clones (cyclGm, cycPGM, cycSGM, and cyc4Gm)
corresponding to groups I, II, III, and IV, respectively. Their
deduced amino acid sequences are shown in Figure 1 with
Cyc5Gm, which formerly was named S13-6 (Hata et al., 1991).
Highly conserved amino acids in the cyclin box are well conserved in all sequences; moreover, all sequences contain a
conserved destruction box in the N-terminal region that is
responsible for ubiquitin-mediated proteolysis.
conserved amino acids for A- and 6-type cyclins are shown
above the soybean cyclin sequences in Figure 1 (Hata et al.,
1991). These conserved amino acids appear to be distributed
almost randomly among the five soybean cyclins, and it was
not easy to assign them to either of the phylogenetic types.
However, it was possible to classify them into two groups based
on the homology over the entire amino acid sequences; one
includes CyclGm, Cyc2Gm, and CycBGm, and the other includes Cyc4Gm and Cyc5Gm. The amino acid identities are
45 to 50% between CyclGm, Cyc2Gm, and Cyc3Gm, and 55%
1145
Differential Expression of Plant Cyclins
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Figure 1. Deduced Amino Acid Sequences for Soybean Cyclins.
Conserved amino acids in A- and 8-type cyclins (A-con. and B-con., respectively) are shown above the soybean cyclins. The destruction box
is underlined with a thick bar, and the cyclin box is bordered by the symbols >> and <<. Residues identical in all soybean cyclin sequences
are indicated by asterisks. The region corresponding to the PCR products is bordered by the symbols > and <. The sequence motifs referred
to in the text are underlined. Gaps (indicated by dashes) were introduced to optimize the alignment. The nucleotide sequences of qclGm, W C ~ G ~ ,
cyc3Gm, and cyc4Gm have been submitted to the GenBank, DDBJ, and EMBL nucleotide sequence data bases with the accession numbers
D50868, D50869, D50870, and 050871, respectively. cyc5Gp (X62820) was formerly named s73-6 (Hata et al., 1991).
1146
The Plant Cell
between Cyc4Gm and CycSGm. However, the identities between these two groups are only 23 to 29%. The amino acids
conserved commonly in A- and B-type cyclins are highly conserved in all soybean cyclin sequences (81 to 94%). The amino
acids conserved distinctively in A-type cyclins are conserved
63 to 73% in CydGm, Cyc2Gm, and CycSGm, whereas they
are conserved 51 and 37% for Cyc4Gm and CycSGm, respectively. On the other hand, the amino acid consensus only for
B-type cyclins is less well conserved in all five sequences, but
they accounted for 23,41, and 23% for CydGm, Cyc2Gm, and
CycSGm, respectively, and 43 and 45% for Cyc4Gm and
CycSGm. In addition, the motif EVXEEY(K/R)L in the cyclin
box typical of A-type cyclins is perfectly conserved in CydGm
to CycSGm. Although the motif typical of B-type cyclins,
FLRRXSK, is conserved more loosely in all five sequences,
the motif HX(K/R)F, which is highly conserved in yeast B-type
cyclins (Kiihne and Linder, 1993), is present in both Cyc4Gm
and CycSGm. Thus, CydGm, Cyc2Gm, and CycSGm resemble A- rather than B-type cyclins, whereas Cyc4Gm and
CycSGm more closely resemble B-type cyclins. However, these
sequence criteria were not enough to assign them definitely
to either of the phylogenetic types (Hata et at., 1991; Ferreira
et al., 1994).
The nucleotide sequence homologies among these five cyclins are 40 to 60%, and they do not cross-hybridize each other
under the standard high-stringency conditions. Therefore, we
used the entire cDNA inserts for DNA and RNA gel blot analyses. Figure 2 shows gel blot hybridization of soybean genomic
DNA with each cyclin cDNA insert. The results indicated totally discrete patterns of hybridization, and, therefore, these
five cyclin genes are not tandemly arranged.
cydGm cyc2Gm cycSGm cyc4Gm cycSGm
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Figure 3. RNA Gel Blot Analysis of Total RNA Isolated from Various
Tissues of Soybean Seedlings with Cyclin cDNAs.
Total RNAs (10 ng) were electrophoresed on a 1% agarose gel, transferred to nitrocellulose membranes, and probed with the cDNA inserts
indicated. The numerals within parentheses indicate the length of the
transcripts in kilobases. Lane 1, shoot apex; lane 2, root tips; lane 3,
nodulated root segments at 4 days after B. japonicum inoculation; lane
4, stems; and lane 5, nodules at 9 days after inoculation.
2.3-
Expression of Cyclin Genes in Various Tissues
1.050.770.56-
Figure 2. DNA Gel Blot Analysis of Genomic DNA Isolated from Soybean Hypocotyls.
Ten micrograms of DNA was digested to completion with EcoRI (E)
or Hindlll (H), electrophoresed on a 0.8% agarose gel, transferred to
nylon membranes, and probed with the cDNA inserts indicated. Numbers at left indicate the lengths in kilobases.
Figure 3 shows the results of gel blot hybridization of total
RNAs from various organs of soybean seedlings. All five cyclin
transcripts were predominantly expressed in the meristematic
organs, such as shoot apices, root tips, and young developing
nodules, but at very low levels in stems (control nonmeristematic organs), indicating that the expression of these genes
is closely related to cell division. These patterns of cyclin
expression contrasted with that of cdc2, which was highly expressed in stems as well as in meristematic organs. The cdc2
probe that we used has a sequence 99% identical with cdc2-S5
(Miao et al., 1993) containing a PSTAIRE domain. Recently,
Differential Expression of Plant Cyclins
a nove1 member of the cdc2 gene family has been described
that does not contain a perfectly conserved PSTAIRE domain
and shows the cell cycle-dependent expression pattern (Forbert
et al., 1994). The unusually large sizeof cyc2Gm mRNA is due
to the presence of a M O 0 nucleotide noncoding sequence at
the 5' end. The transcript levels of cyc5Gm were highest in
all organs, whereas cyc7Gm and cyc2Gm transcripts were at
very low levels. The transcripts of cyclGm and cyc5Gm appeared to be most abundant in shoot apices, whereas those
of cyc3Gm were more abundant in root tips and nodules. However, these differences are not enough to demonstrate the
differentialexpressionof cyclins betweendifferent meristematic
tissues, and we conclude that these five cyclin genes are all
expressed in the meristematic organs of soybean seedlings.
Cell Cycle-Specific Expression of Cyclin Genes
In situ hybridization of soybean shoot apices with histone H4,
cyc7Gn-1, cyc3Gm, and cyc5Gm is shown in Figures 4A to 4D,
respectively. Experiments with cyc2Gm and cyc4Gm were unsuccessful because of the extremely low hybridizationsignals.
The scattered signal appearance in isolated cells in the
meristematic region is a characteristic feature of cell cycle-specific genes (Fobert et al., 1994). The signals appeared randomly
in isolated cells in the apical meristems, vascular cambium,
and developing leaves. The frequency of labeled cells and the
signal intensity were highest for histone H4, intermediate for
cyc5Gm and cyc3Gm, and very low for cyc7Gm. In particular,
the hybridization signals for cyc7Gm appeared in only a small
fraction of the cells in the shoot apex, indicating expression
during a short period of the cell cycle. The differences in the
frequencies and relativesignal intensities betweenthese genes
were almost consistent with the results of the RNA gel blot
hybridization analysis (Figure 3).
Differential Expression of Cyclin Genes during
the Cell Cycle
To examine the interrelationships among expression of these
genes, we performed double-target in situ hybridization using
a combination of any two probes involving histone H4, according to the method developed by Fobert et al. (1994). Because
transcripts of histone genes accumulate preferentially and
abundantly in S phase (Nakayama and Iwabuchi, 1993),they
provide a good control to mark S-phase cells. The microtome
sections of shoot apices were hybridized with a mixture of
digoxigenin- and fluorescein-labeled probes, and the signals
were detected sequentially using anti-digoxigenin and antifluorescein antibodies conjugated with alkaline phosphatase
in combination with different color substrates (see Methods).
A typical result for a combination of cyc5Gm and histone H4
is shown in Figures 5A and 58. The sections were hybridized
with a mixture of digoxigenin-labeled cyc5Gm probe and
1147
fluorescein-labeled histone H4 probe. Hybridization signals for
cyc5Gm were first detected using anti-digoxigenin-alkaline
phosphatase conjugate with the Fast Red substrate (Figure
5A). After heat inactivationof alkaline phosphatase conjugated
to anti-digoxigenin,signals for histone H4 were visualized using
anti-fluorescein-alkaline phosphatase conjugate with nitro blue
tetrazolium salt/5-bromo-4chloro-3-indolyl phosphate toluidinium salt (Figure 5B). Signals for the cyc5Gm probe appeared
in cells completely different from those containing signals for
the histone H4 probe. We examined more than 250 cells from
a number of independent sections showing the cyc5Gm signals, and we found that only 2% of them expressed histone H4.
Similar experiments were performed for combinations of
cyc3Gm and histone H4 (Figures 6A and 6B) and combinations of cyc3Gm and cyc5Gm (Figures 6C and 6D). The
expression of cyc3Gm overlapped that of histone H4 in part
(9% of 250 cells with cyc3Gm signals showed histone H4 expression), whereas it overlapped with that of cyc5Gm in a much
higher proportion(41% of 255 cells). The results of experiments
that used cyc7Gm were contrasted with results of those that
used cyc3Gm. The cells with cyc7Gm signals were completely
included in those with histone H4 signals (Figures 7A and 76),
whereas essentially no cells expressed both cyc7Gm and
cyc5Gm (Figures 7C and 7D).
Figures 8A to 8F show the results of in situ hybridization
with %-labeled probes for histone H4, cyc5Gm, and cyc3Gm.
The sections were counterstained with toluidine blue, and
dividing cells at metaphase and early telophase could be distinguished. However, it was difficult to distinguish the cells at
prophase from those at interphase due to the limitation of
toluidine blue staining. Signals for histone H4 were very strong,
but dividing cells at metaphase and early telophase contained
no hybridization signal (Figure 8A). In contrast, the signals for
cyc5Gm were clearly detectable in cells at metaphaseand interphase (and prophase). During early telophase, the cells showed
weak signals that were significantlyabove the background leve1
(Figures 8B and 8C). For cyc3Gm, the hybridization signals
were barely detectable in dividing cells at metaphase and early
telophase and appeared only at interphase (Figures 8D to 8F).
DlSCUSSlON
In this study, we describe the characterization of five distinct
mitotic cyclins from soybean plants, cyc7Gm to cyc5Gm. The
expression patterns of three of these, cyclGm, cyc3Gm, and
cyc5Gm, were analyzed in further detail by double-target in
situ hybridization involving histone H4 as an S-phase marker.
As described first by Fobert et al. (1994), the transcripts of these
cell cycle-related genes appeared in isolated cells dispersed
in the meristematic tissues (Figures 4A to 4D), indicating that
they are expressed during limited periods in the cell cycle. Similar dispersed patterns of expression of cyc2Ms and histone
H4 have been described in nodule meristems of alfalfa (Yang
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Figure 4. Localization of the Transcripts of Cell Cycle-Related Genes in Soybean Shoot Apices.
Hybridization signals are visible as purple or dark blue color development. The sense strand control hybridization gave no signal (data not shown).
(A) Histone H4.
(B) cycSGm.
(C) cyc3Gm.
(D) cydGm. Bar = 200 urn.
Differential Expression of Plant Cyclins
1149
Figure 5. Double-Labeling of a Soybean Axillary Bud for cycSGm and Histone H4.
The paraffin-embedded sections were hybridized with the mixtures of digoxigenin- and fluorescein-labeled antisense RNAs.
(A) Detection of hybridization signals with the digoxigenin-labeled cycSGm probe (red).
(B) The same section after detection of the fluorescein-labeled histone H4 probe (black). Bar = 25 urn.
et al., 1994). The frequencies of cells labeled by these probes
should reflect the relative length of the phases in the cell cycle
during which they are abundantly expressed. The frequencies
of cells expressing cycSGm and cycSGm were considerably
less than the frequencies of those expressing histone H4, and
the expression of cydGm was found in an extremely small number of cells in the shoot apex. Although the signal intensities
of in situ hybridization are not always parallel with the abundance of the mRNAs in vivo, the results show that the
expression of cydGm is restricted to a very short period in
the cell cycle.
Double-target in situ hybridization showed more precisely
the relative timing of the expression of these genes. cycSGm
was expressed in cells different from those accumulating histone H4 mRNAs, indicating that the expression of cycSGm is
temporally separated from S phase (Figures 5A and 5B). Moreover, transcripts of cycSGm were detected in dividing cells at
metaphase and early telophase as well as in cells at interphase
(Figures 8B and 8C). Therefore, assuming that these cell cycle
genes are expressed once per cell cycle, it is most likely that
cycSGm is expressed during G2 and early M phases. Similar
results were obtained for two Antirrhinum cyclins, cydAm and
cyc2Am, which have high homology with cycSGm (Fobert et
al., 1994).
Based on the G2- and M-phase-specific expression of
cycSGm, we can deduce the phases during which cydGm and
cycSGm are expressed. Transcripts of cyc3Gm were detected
in a small fraction of the cells that expressed histone H4 and
in a much greater proportion of the cells that expressed cycSGm
(Figures 6A to 6D). In addition, they were barely detected at
metaphase and telophase (Figures 8D to 8F). A simple explanation for these results is that cycSGm is expressed predominantly
during late S and G2 phases. In contrast with the expression
pattern of cycSGm, cydGm expression appeared to be restricted to S phase and did not overlap that of cycSGm (Figures
7A to 7D). We conclude that, taken together with the large difference in the frequencies of cells labeled with cydGm and
histone H4, the transcripts of cydGm become abundant during a very short period in S phase.
Several authors have described the sequences of A-like and
B-like cyclins from higher plants (Hataetal., 1991; Ferreiraet
al., 1994; Renaudin et al., 1994). In this study, we describe
three A-like and two B-like cyclin sequences from soybean
plants. Although these sequences show significant similarities with both A- and B-type cyclins, their similarities with amino
acids conserved distinctly in A- and B-type cyclins are not as
high, making it difficult to assign them to either the A- or B-type
cyclin group based on sequence motifs alone. It has been
demonstrated that A-type cyclins are synthesized and destroyed
earlier in the cell cycle than are B-type cyclins (Minshull et al.,
1990; Pines and Hunter, 1990). Double-target in situ hybridization studies with cycSGm, cycSGm, and histone H4 clearly
indicate that cycSGm is expressed at a much earlier phase
of the cell cycle than is cycSGm. Therefore, the results
presented in this study strongly suggest that cycSGm and
cyc5Gm are A- and B-type cyclins, respectively, based on their
differential expression patterns during the cell cycle, in addition to the sequence criteria.
Furthermore, we have recently isolated three mitotic cyclin
cDNAs from tobacco, Ntcyc25, Ntcyc27, and Ntcyc29, that are
homologous with cycSGm, cyc2Gm, and cycSGm, respectively
(YY. Setiady, M. Sekine, N. Hariguchi, T. Yamamoto, K. Yoshida,
H. Kouchi, and A. Shinmyo, unpublished data). Their sequence
identities across ~170 amino acids of the cyclin box are 88%
1150
The Plant Cell
Figure 6. Double-Labeling of Shoot Apices with cyc3Gm and Histone H4, and with cyc3Gm and cycSGm.
The arrows indicate the positions of cells with hybridization signals from both probes.
(A) and (B) Detection of hybridization signals with the digoxigenin-labeled cycSGm probe (red) followed by that of the fluorescein-labeled histone
H4 probe (black in [B]).
(C) and (D) Detection of hybridization signals with the digoxigenin-labeled cyc3Gm probe (red) followed by that of the fluorescein-labeled cycSGm
probe (black in [D]). Bar = 25 urn.
for Ntcyc25 and Cyc3Gm, 86% for Ntcyc27 and Cyc2Gm, and
77% for Ntcyc29 and CycSGm. Their expression patterns in
the highly synchronized culture of tobacco suspension cells
clearly indicate that A/fcyc25 and A/fcyc27 are expressed predominantly during S-to-G2 phases, whereas Ntcyc29 is
expressed during G2-to-M phases. These findings again suggest that the classes of cyclins represented by cycSGm and
cycSGm are, respectively, the A- and B-type cyclin cognates
from higher plants.
The expression pattern of cydGm is quite unlike that of
cycSGm and cycSGm. The sequence of cyclGm is more closely
related to A- than to B-type cyclins, but expression is restricted
to a short period of S phase and does not appear to be associated with G2 phase. Therefore, cydGm might represent
a novel class of cyclins with a function in DNA replication during S phase.
Since the first evidence of plant cyclins (Hata et al., 1991),
18 plant cyclins have been described, including the four soybean cyclins newly identified in this study. Renaudin et al. (1994)
and Ferreira et al. (1994) have shown that plant mitotic cyclins
can be divided into at least three groups, one consisting of
A-like cyclins and the other two of B-like cyclins. More sequence
data are now available, particularly for A-like cyclins, and we
constructed a phylogenetic tree of these cyclins from a multiple sequence alignment of the cyclin box (Figure 9). This
revealed that the plant cyclins identified so far can be divided
into two distinct groups. One presumably includes A-type and
the other B-type cyclins. Each group appears to be subdivided
into two classes, which we tentatively designated a1, a2, (31,
and (52, according to the terminology proposed by Soni et al.
(1995). (31 and P2 cyclins both have a well-conserved HX(K/R)FL
motif within the cyclin box, but a more upstream conserved
Differential Expression of Plant Cyclins
(S/T)(S/T)VL(S/T)ARSKAACG motif (Renaudin et al., 1994) was
found only in P1 cyclins. a1 and a2 cyclins have a conserved
EVXEEY(K/R)L motif or closely related sequences within the
cyclin box, which is typical for A-type cyclins. They also contain a few short conserved sequences upstream of the cyclin
box. There are only two a1 cyclins, and one of them (C13-1)
is the product of a cDNA truncated in the N terminus (Hata
et al., 1991), so it is not possible to characterize sequence
criteria to distinguish between a1 and a2. However, cydGm
shows a quite different expression pattern from that of cyc3Gm
and should therefore be classified as a novel class distinct from
other A-type cyclins.
Among these tentative classes, the soybean cyclins we have
identified fall into three distinct classes, a1, a2, and [31, all of
which are expressed in different meristematic organs of soybean plants. Several distinct cyclins identified from single plant
species, such as maize (CydaZm, CydbZm, Cyc2Zm, and
1151
Cyc3Zm) and Arabidopsis (CydAt, Cyc2aAt, Cyc2bAt,
CycSaAt, and Cyc3bAt), also fall into different classes. Therefore, these classes are suggested not only to reflect the genetic
divergence between plant species but also to represent classes
of cyclins that have different functions in the plant cell cycle.
The previous observations for cell cycle-specific expression
of a few of these plant cyclins also support this explanation.
Two Antirrhinum cyclins, cyclAm and cyc2Am, which would
belong to class (31, showed an expression pattern essentially
the same as that of cycSGm (Fobert et al., 1994). Furthermore,
cycSaAt and cycSbAt of Arabidopsis, which would belong to
class 0.2, have been suggested to be expressed earlier in the
cell cycle than cyc2aAt and cyc2bAt, which would belong to
class (32 (Ferreira et al., 1994).
The results presented in this study clearly demonstrate that
the transcription of distinct classes of mitotic cyclin genes is
regulated differentially in the plant cell cycle. These genes are
Figure 7. Double-Labeling of Shoot Apices with cydGm and Histone H4, and with cydGm and cycSGm.
The arrows indicate the positions of cells with hybridization signals from both probes.
(A) and (B) Detection of hybridization signals with the digoxigenin-labeled cydGm probe (red) followed by that of the fluorescein-labeled histone
H4 probe (black in [B]).
(C) and (D) Detection of hybridization signals with the digoxigenin-labeled cydGm probe (red) followed by that of the fluorescein-labeled cycSGm
probe (black in [D]). Bar = 25 urn.
The Plant Cell
1152
expressed in all meristematic tissues at different times during
the cell cycle, suggesting that they each have essential and
distinct functions in cell cycle progression in plant meristems.
Recently, Ferreira et al. (1994) have reported the presence of
an organ-specific cyclin in Arabidopsis. They have shown that
cyc2bAt mRNAs are present only in roots, whereas another
closely related gene, cyc2aAt, is expressed in all plant organs;
both of them would be classified as B-type (p2) cyclins by our
criteria. Furthermore, two closely related cdc2 genes, cdc2S5 and cdc2-S6, have been suggested to be expressed
differentially in various organs of soybean (Miao et al., 1993).
S
f
4k
Our study showed no significant specificity of expression of
the five distinct soybean cyclins in different meristematic organs and tissues. However, sequencing of the PCR products
revealed the presence of a number of different molecular species of cyclin mRNAs, particularly A-type cyclins (data not
shown). Because our hybridization analysis might not be appropriate to distinguish such individual mRNA species belonging
to the same group, use of 3' noncoding regions as probes
and/or of reverse transcription-PCR with primers specific to
individual sequences may be necessary. It would be intriguing to analyze, in greater detail, the differential expression of
*< '*.
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Figure 8. In Situ Hybridization of Shoot Apical Regions with 35S-Labeled Probes of Histone H4, cycSGm, and cyc3Gm.
The hybridization signals are visible as black dots. Metaphase (m) and early telophase (t) cells can be seen. Sections were poststained with
toluidme blue.
(A) Hybridization with histone H4. Bar = 20 urn.
(B) and (C) Hybridization with cyc5Gm. Metaphase and early telophase cells exhibit positive signals. Apparently nondividing cells also exhibit
strong signals (indicated by arrowheads). Bars = 20 urn.
(D) to (F) Hybridization with cyc3Gm. Arrowheads indicate cells with positive signals. Bars = 10 urn.
Differential Expression of Plant Cyclins
a1
Cyc2Gm
cyc3aAt
Cyc3bAt
Cyc3Gm
Cyc2Zm
a2
=
Cyc2aAt
Cyc2bAt
fl
al., 1989). Oligo(dT)-primed double-strandedcDNAs were synthesized
from 2 to 5 pg of poly(A)+RNA using a cDNA synthesis kit (Pharmacia Biotechnology), and cDNA libraries were constructed in hgtlO
phages, as described by Kouchi and Hata (1993). Genomic DNA was
obtained from etiolated hypocotyls of 7-day-oldsoybean seedlings, according to the method of Rogers and Bendich (1988).
Polymerase Chain Reaction and Cloning of Cyclin cDNAs
Cyc4Gm
CycSGm
CyclAt
Cycl aZm
Cycl bZm
1153
Pl
P2
Cyc2Ms
Cyc3Zm
Figure 9. Phylogenetic Tree of Mitotic Cyclins from Higher Plants.
The tree was depicted from a sequence alignment of the cyclin box
indicated in Figure 1, according to the method of Higgins and Sharp
(1988). Length of the horizontal lines reflects divergence. The sources
of published sequences of plant cyclins are as follows: C131 from carrot
(Hata et al., 1991); CycSaAt, Cyc3bAt, CycSaAt, Cyc2bAt, and CyclAt
from Arabidopsis (Hemerly et al., 1992; Ferreira et al., 1994); CyclAm
and Cyc2Am from Antirrhinum(Fobert et al., 1994); CyclaZm, CyclbZm,
CycPZm, and Cyc3Zm from maize (Renaudin et al., 1994); Cyc2Ms
from alfalfa (Hirt et al., 1992). The five soybean cyclins described in
this paper are indicated in boldface.
these cyclin genes in different organs or tissues andlor in response to environmental stimuli, such as infection with
Rhizobium. Such analysis may provide more insights into how
these cell cycle-related genes are involved in plant developmental programs.
Degenerated oligonucleotide primers were designed for highly conserved regions of published amino acid sequences of mitotic cyclins.
Their sequences are 5’ATI(C/T)Tl~lGATTGG(C/T)TIGTI(G/C)A(A/G)GT-3‘ (sense) and 5’-TCTGG(T/G)GG(A/G)TA(G/C)AT(C/T)TC(C/T)TCATATTT-3‘ (antisense), representing amino acid sequences of ILVDWLV(E/Q)V and KYEE(I/M)YPP(D/E), respectively (Hata et al.,
1991). Deoxyinosinetriphosphate(designatedI in the primer sequence)
was used to decrease the degree of degeneration. The polymerase
chain reaction (PCR) amplification was performedwith oligo(dT)-primed
cDNA prepared from poly(A)+ RNAs of young, developing soybean
root nodules and root tips using Tth DNA polymerase (Toyobo, Tokyo).
The primer concentrationswere 2 pM, and the cycling conditions were
30 cycles at 94°C (1 min), 45% (1.5 min), and 72% (2 min). The amplified fragments (185 bp) were isolated from the agarose gel, cloned
into a pBluescript II SK+ plasmid vector (Stratagene), and sequenced.
The selected PCR products were labeled by random priming with
32P-dCTPand used to screen the cDNA libraries of 9-day-old nodules
and of suspension-cultured cells by standard procedures (Sambrook
et al., 1989). Hybridization was carried out in 50% formamide, 4.8 x
SSC (1 x SSC is 0.15 M NaCI, 0.015 M trisodium citrate, pH 7.0), 5 x
Denhardt’s solution (1 x Denhardt’s solution is 0.020/0 Ficoll, 0.02%
polyvinylpyrrolidone, 0.02% BSA), 0.5% SDS, 50 mM Hepes, pH 7.3,
and 100 pglmL denatured salmon sperm DNA at 42% for >16 hr. The
filter was washed in 2 x SSC, 0.5% SDS for 30 min at room temperature, and then in 0.5 x SSC and 0.1% SDS at 55OC four times for 10
min each. lnserts of the purified phages were subcloned into pBluescript
II SK+ plasmids for further characterization.
RNA and DNA Transfer Blot Analysis
Agarose gel electrophoresis of total RNA and digested genomic DNA
was performed using standard procedures (Sambrook et al., 1989).
The RNA and DNA in the gel were blotted onto nitrocellulose membranes and hybridized with 32P-labeledcDNA inserts. The conditions
of hybridization and subsequent washings were the same as those
given for the screening of cDNA libraries.
METHODS
DNA Sequencing
Plant Growth and Library Construction
Soybean (Glycine max var Akisengoku) seedlings were grown with or
without inoculation with Bfadyrhizobium japonicum, as described by
Kouchi and Hata (1993). Suspension-culturedcells of soybean, initiated
from hypocotyls of seedlings in the presence of 2,4-dichlorophenoxyacetic acid, were cultured in 85 medium (Gamborg, 1975) for 2 years.
Total RNA was isolated from the tissues or cells by guanidium thiocyanate extraction, followed by centrifugation on cesium chloride, and
the poly(A)+RNA-enriched fraction was selected by oligo(dT)-cellulose column chromatography by standard procedures (Sambrook et
The cDNA inserts subcloned into pBluescript plasmids were sequenced
by the dideoxy chain termination method (Sanger et al., 1977) using
an automatic sequencer (Model37OA; Applied Biosystems,Foster City,
CA). 60th strands were entirely sequenced.
In Situ Hybridization
Preparation of digoxigenin-labeled antisense RNA probes was as described by Kouchi and Hata (1993). The RNA probes corresponding
to entire cDNA sequences (poly[A] tails were removed if present) were
1154
The Plant Cell
synthesized using T3 or T7 RNA polymerase (Stratagene) and employed after partia1hydrolysis with sodium carbonate. The sense probes
were also prepared and used as negative controls. Shoot apices of
7-day-old soybean seedlingswere fixed in 4% (w/v) paraformaldehyde
and 0.25% glutaraldehyde in 10 mM sodium phosphate buffer, pH 7.2,
containing 100 mM NaCI, dehydrated through a graded ethanol series, and embedded in paraffin wax as described by Kouchi and Hata
(1993). The sections (&pm thick) were attached to poly-L-lysine-coated
slides. In situ hybridization procedures were the same as those described by Kouchi and Hata (1993), except for a minor modification
in which washes with 2 x SSC and 1 x SSC containing 50% formamide were simply replaced by those with 2 x SSC and 0.5 x SSC
without formamide, respectively.
The method of double-target in situ hybridization was essentially
the same as the method developed by Fobert et al. (1994). Fluoresceinlabeled RNA probes were prepared using the same protocol as that
given for the preparation of digoxigenin-labeledprobes using fluorescein-11-rUTP (Boehringer Mannheim) instead of digoxigenin-11-rUTP.
The sections were hybridized with a mixture of two probes; one was
labeledwith digoxigenin and the other with fluorescein. After hybridization, the digoxigenin-labeledprobe was detectedwith anti-digoxigeninalkaline phosphatase in combination with Fast Red TRlnaphtol AS-MX
(Sigma). The slides were mountedwith glycerin, and microphotcgraphs
were taken. The coverslips were then removed, and the slides were
incubated in 2 x SSC at 68OC for 3 hr to inactivate the alkaline phosphatase. The second, fluorescein-labeled probe was detected with
the anti-fluorescein-alkaline phosphatase conjugate (Boehringer
Mannheim; 1:lOOO diluted) in combinationwith nitro blue tetrazolium
salt/5-brom~-chloro-3-indolyl
phosphate toluidinium salt (Kouchi and
Hata, 1993). Then the slides were mounted with glycerin, and photographs were taken. In some cases, the slides were further dehydrated
through a graded ethanol series and mounted with Eukitt (O. Kindler,
Germany). This step enabled us to confirm the overlappingof signals
from digoxigenin- and fluorescein-labeled probes in the same cell, because the precipitation from Fast Red in the first round of detection
diminished completely in ethanol.
The in situ hybridization procedure with 35S-labeledRNA probes
was as described by van de Wiel et al. (1990). The sections were counterstained with toluidine blue to visualize dividing cells.
ACKNOWLEDGMENTS
We thank Miki Maeda, Norimitsu Hariguchi, and TakuoYamamotofor
their technical assistance. We also thank Drs. Wei-Cai Yang and Ton
Bisseling for their suggestions and help with the in situ hybridization
experiments with 3%-labeled probes and for providing the soybean
histone H4 cDNA. This research was supported by a grant-in-aid(No.
BMP-IV-95-1-1)to H.K.from the Ministry of Agriculture, Forestry, and
Fisheries of Japan.
Received March 21, 1995; accepted June 21, 1995.
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DOI 10.1105/tpc.7.8.1143
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