Two Arabidopsis cyclin promoters mediate distinctive ... - Europe PMC

Proc. Natl. Acad. Sci. USA Vol. 93, pp. 4868-4872, May 1996 Cell Biology

Two Arabidopsis cyclin promoters mediate distinctive transcriptional oscillation in synchronized tobacco BY-2 cells (cell cycle/cyclin A/cyclin B/Agrobacterium/suspension culture)

ORIT SHAUL*, VLADIMIR MIRONOV*, SYLVIA BURSSENS*, MARC VAN MONTAGU*t, AND DIRK INZE*t *Laboratorium voor Genetica, Department of Genetics, Flanders Interuniversity Institute for Biotechnology, and Recherche Agronomique (France), Universiteit Gent, B-9000 Ghent, Belgium

*Laboratoire Associe de l'Institut National de la

Contributed by Marc Van Montagu, January 2, 1996

ABSTRACT Cyclins are cell cycle regulators whose proteins oscillate dramatically during the cell cycle. Cyclin steady-state mRNA levels also fluctuate, and there are indications that both their rate of transcription and mRNA stability are under cell cycle control. Here, we demonstrate the transcriptional regulation of higher eukaryote cyclins throughout the whole cell cycle with a high temporal resolution. The promoters of two Arabidopsis cyclins, cyc3aAt and cyclAt, mediated transcriptional oscillation of the 8-glucuronidase (gus) reporter gene in stably transformed tobacco BY-2 cell lines. The rate of transcription driven by the cyc3aAt promoter was very low during G1, slowly increased during the S phase, peaked at the G2 phase and G2-to-M transition, and was down-regulated before early metaphase. In contrast, the rate of the cyclAt-related transcription increased upon exit of the S phase, peaked at the G2-to-M transition and during mitosis, and decreased upon exit from the M phase. This study indicates that transcription mechanisms that seem to be conserved among species play a significant role in regulating the mRNA abundance of the plant cyclins. Furthermore, the transcription patterns of cyc3aAt and cyclAt were coherent with their slightly higher sequence similarity to the A and B groups of animal cyclins, respectively, suggesting that they may fulfill comparable roles during the cell cycle.

Cyclins are activators of specific serine/threonine protein kinases, termed CDKs, which drive progression of the eukaryotic cell cycle (reviewed in ref. 1). Based upon sequence analyses, most plant cyclins identified so far can be divided into those showing slightly higher sequence similarity to either the A or B groups of animal cyclins, but these similarities are not sufficient to exclusively assign them to either group (2, 3). Animal A- and B-type cyclins have distinct patterns of expression and fulfill different roles throughout the cell cycle (reviewed in ref. 4). As the roles of cyclins are far more understood in animals than in plants, further affiliation of a plant cyclin to a certain group of animal cyclins based on a similar expression pattern may give a clue to its function. The Arabidopsis cyclAt and cyc3aAt cyclin genes represent plant cyclins with slightly higher homology to the B and A groups of animal cyclins, respectively (2). Whole-mount in situ hybridization of Arabidopsis root tips treated with cell cycle blockers indicated that steady-state mRNA levels of cyclAt are high at early metaphase and low at early S phase, while the opposite was true for cyc3aAt (2). These data indicated that mRNA levels of cyc3aAt are increased in advance to that of cyclAt, but were not sufficient to give a complete picture of the expression pattern and transcriptional regulation of these cyclins throughout the whole cell cycle. Fluctuation in the steady-state mRNA level of a cell cycle gene may be regulated by a change in the rate of transcription,

or in mRNA stability, or both. Transcriptional regulation of cell cycle genes is best established in yeast (reviewed in ref. 5). Research in animals has indicated that the mRNA abundance of several mammalian S phase genes is only partly determined by transcriptional regulation (reviewed in ref. 6). For example, fluctuations of 10- to 20-fold in the mRNA levels of histone and thymidine kinase genes result from a 3- to 4-fold increase in the rate of their transcription, and up to a 10-fold variation in their mRNA stability (7, 8). Besides S phase genes, cell cycle-dependent transcription rates have also been demonstrated for the mammalian CDK1 protein kinase and the cdc25C regulatory phosphatase genes (9, 10). Among animal cyclins, transcriptional regulation throughout the cell cycle has only been elucidated for the human A, Bi, and D cyclin genes. Using cell cycle blockers, it was shown that the promoters of the human Bi and D cyclins mediated elevated transcription in cells arrested at the M and GI phases, respectively (11, 12). In synchronized human cells, the steady-state mRNA level of the A and Bi cyclins was 3-4 times higher in G2 compared with the GI phase, while at the same time their nuclear run-on transcription rates varied to a lesser extent (13, 14). It was recently shown that the half-life of the human cyclin Bl mRNA is 10 times longer at G2-to-M than at the GI phase (15). Reports on promoter-reporter gene analyses of animal cyclin genes in synchronously growing cells are scarce. Studies of synchronized, stably transformed human cell lines have indicated that transcription driven by the cyclin A and cyclin Bi promoters initiates at the onset and at the end of the S phase, respectively (16, 17). However, no data is provided on cyclin A promoter-related transcription following the S phase. Furthermore, the cyclin Bi promoter-related transcription failed to decline after mitosis, in contrast with the nuclear run-on transcription data (13). Although it has been demonstrated that several plant cell cycle genes exhibit variation in their mRNA abundance throughout the cell cycle (reviewed in ref. 18), the transcriptional regulation is not well studied. It was shown that the cyclAt promoter mediated an increased expression in oryzalintreated root tips (19). Promoter-reporter gene analyses in synchronized suspension cultures have only been performed for histone genes (20-22). In the present study, we aimed to analyze the transcriptional regulation of the promoters of the Arabidopsis cyclAt and cyc3aAt cyclin genes in synchronized suspension cultures, as the currently available cell cycle blockers can arrest cells only at early S phase or during mitosis, and therefore fail to detect events at other stages. Furthermore, accurate identification of the cell cycle phase at which these cyclins are expressed may elucidate their sequence-based affiliation to different groups of animal cyclins and possibly give a clue as to their function. The difficulty of synchronizing Arabidopsis suspension cultures represents a major limitation

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*To whom reprint requests should be addressed at: Laboratorium voor Genetica, Universiteit Gent, K. L. Ledeganckstraat 35, B-9000 Ghent, Belgium. 4868

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in the study of Arabidopsis cell-cycle genes. We therefore used a tobacco BY-2 cell line, in which the highest level of culture synchronization, compared with other plant or animal cell lines, has been achieved so far (23). We show here that the promoters of the two Arabidopsis cyclins confer differential, "A- and B-type" oscillation in the transcription of a reporter gene throughout the cell cycle of stably transformed, highly synchronized tobacco BY-2 cells.

MATERIALS AND METHODS Construction of Chimeric Genes. Construction of the cyclAt promoter-gus-3' octopine synthase gene in a pGUS1 cloning vector and a pGSV4 binary vector has been described (19). The cyc3aAt promoter was PCR amplified from a genomic clone (2) using two oligonucleotides, one spanning the methionine start codon and introducing a Nco I site (5'-GAAGCTCTAAC CATGGTTTCAAA-3'), and the other 1.5-kb upstream and introducing a Sal I site (5'-CTGAGAAGAGTCGACAAGATTCT-3'). The resulting fragment was digested with Sal I and Nco I and introduced into the same sites in the pGUS1 vector upstream from the gus gene. A fragment containing the cyc3aAt promoter-gus-3' octopine synthase gene was produced by digestion with Hind III and Xba I and ligated into the same sites of the pGSV4 binary vector. Binary vectors containing either the cyclAt or cyc3aAt promoter-gus fusion were immobilized into Agrobacterium tumefaciens C58C1RifR(pMP90). Maintenance and Stable Transformation of Tobacco BY-2 Cells. The tobacco BY-2 cell line (derived from Nicotiana tabacum cv. Bright Yellow 2) was maintained by weekly dilution essentially as described (23). For transformation, 1 ml of a 6-day-old BY-2 culture was cocultivated with 200 ,ul of an overnight-grown agrobacteria culture in a Falcon 10-cm Petri dish containing 12 ml fresh medium for 4 days at 28°C. Cells were washed using a 100 /LM mesh filter and grown in 50 ml fresh medium supplemented with 500 gg/ml carbenicillin and 10 ,ug/ml of the kanamycin analogue G-418 (Geneticin, Sigma). Only when the culture reached maximal density (similar to a stationary, 1-week-old culture; this usually took -3 weeks), 1 ml culture was transferred to a similar medium. After two additional 1 ml transfer cycles in medium containing 500 ,tg/ml carbenicillin, 10 ,tg/ml G-418, and 250 ,tg/ml vancomycin, cultures were propagated in an antibiotic-free medium, and elimination of agrobacteria was verified. Culture Synchronization and Arrest with Cell Cycle Blockers. Cells were synchronized as described by Nagata et al. (23). In brief, 10 ml of a stationary, 7-day-old culture was grown in 100 ml fresh medium containing 5 mg/l aphidicoline (Sigma) for 24 hr, and then the cells were washed extensively and grown in 100 ml fresh medium. For treatment with cell cycle blockers, stationary 7-day-old cells were diluted 10-fold and grown 24 hr in fresh medium containing 5 mg/l aphidicoline, 10 mM hydroxyurea (Sigma), or 15 ,uM oryzalin (Sigma). To determine the mitotic index, cells were stained with lactopropionic orcein (23). RNA Extraction and Northern Blot Analyses. Total RNA was prepared using the TRIZOL Reagent (GIBCO/BRL) according to the manufacturer's instructions, followed by precipitation in 2 M LiCl. RNA samples (20 gg) were treated for 20 min at 65°C with a 10 mM sodium-phosphate (NaPi) buffer (pH = 7.2) containing 50% (vol/vol) dimethylsulfoxide and 4% (vol/vol) glyoxal (Fluka). After electrophoresis in 1.5% agarose gels [in a 35 mM triethanolamine (Merck) buffer titrated to pH = 7.0 with H3PO4] and blotting on Hybond N+ membranes (Amersham), RNA was fixed by 10 min incubation with 50 mM NaOH. Membranes were stained in 0.02% methylene blue and 0.3 M sodium acetate (pH = 5.5), rinsed in H20, and destained in 0.1% SDS. Hybridization with [32P]-labeled probes, corresponding to the coding regions of the gus and Arabidopsis histone H4 genes, was performed at

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65°C in a 100 mM NaPi buffer (pH = 7.2) containing 8% SDS, 10% PEG 8000, 0.5 mM EDTA, 0.1 mg/ml salmon sperm DNA, and 0.25 mg/ml heparin.

RESULTS Establishment of Transformed BY-2 Suspension Cultures. To study the transcriptional regulation of the Arabidopsis cyclAt and cyc3aAt cyclin genes throughout the cell cycle, their promoters were fused to the reporter gene f3-glucuronidase (gus; uidA from Escherichia coli) (19, 24), and the chimeric genes were stably transformed into tobacco BY-2 suspension cultures (the transformed cultures are hereafter referred to as cyclAt and cyc3aAt cultures). Agrobacterium-mediated transformation of BY-2 cells and recovery of transformed calli has been described (25). We developed a protocol to directly

obtain transformed suspension cultures (see Materials and Methods), and Southern blot hybridization indicated that these cultures contain a heterogeneous mixture of transformed cells (data not shown). Histochemical analysis showed that all the cells in the transformed cultures (in contrast with control cells), exhibited ,B-glucuronidase activity, even after 1 year of growth in nonselective media. Synchronization of Transformed Cultures. Transformed cultures were synchronized by dilution of stationary cells in the presence of aphidicoline, which inhibits DNA polymerase and blocks cells at early S phase (see Materials and Methods). Cell cycle progression was followed by determining the mitotic index and by using the histone H4 gene as an S phase maker. It has been shown that in tobacco BY-2 cells, as with other plant cells, histone H4 genes are expressed throughout the S phase, and in cycling cells their transcript levels reflect the rate of DNA synthesis (26, 27). The generation times of transformed and nontransformed cultures were indistinguishable and reproducibly shorter than previously described for these cells (23). A typical experiment, in which cells synchronously released into the S phase were analyzed until the second M phase, is shown in Fig. 1. As the ,B-glucuronidase protein is very stable (21, 24), we used gus mRNA levels as transcription-rate marker. In both transformed cultures, stationary cells, which are arrested at the G, or Go phase (determined by flow cytometry and by absence of mitotic cells), exhibited very low levels of gus transcript, while nontransformed cultures did not exhibit any gus mRNA, even during rapid growth (data not shown). The rate of transcription governed by the cyclAt promoter was very low in cells arrested by aphidicoline at early S phase (0 hr), started to increase after exit from this phase, and peaked at the G2-to-M transition and during mitosis. Transcription rate decreased upon exit from the M phase, was low during the second S phase, and started to increase again upon exit from this phase. In contrast to cyclAt cultures, gus mRNA levels in aphidicoline-arrested cyc3aAt cultures were higher than the minimal level observed during subsequent synchronous growth. Transcription rate governed by the cyc3aAt promoter was reproducibly lower during the G, phase (8-10 hr) compared with cells arrested by aphidicoline at early S phase. It has been shown that aphidicoline, although reducing the rate of DNA synthesis, blocks BY-2 cells after the point where transcription of histone genes, and possibly other S phase genes, is induced (27). Thus, based solely on analyses before and next to aphidicoline release, it is hard to deduce the cell cycle stage at which transcription of the cyc3aAt-gus chimeric gene is induced. Analysis of the second, unperturbed S phase indicated that the transcription rate governed by the cyc3aAt promoter slowly increases upon entry into the S phase, and is significantly elevated towards the end of this phase. In contrast, cyclAt-related transcription was low throughout the whole S phase. Cyc3aAt-related transcription also peaked and decreased in advance to that of cyclAt. For cyc3aAt, transcrip-

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Proc. Natl. Acad. Sci. USA 93 (1996)

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FIG. 1. Expression of the cyclAt-gus (A and C) and cyc3aAt-gus (B and D) chimeric genes in stably transformed, synchronized BY-2 cells. Cells were synchronized by treatment with aphidicoline, and RNA was prepared from samples taken immediately before (O hr) and up to 18 hr after release. (A and B) RNA blots were stained with methylene blue (M.B.) to detect the total amount of RNA loaded, then washed and sequentially hybridized with thegus and histone H4 gene probes. (C and D) Mitotic index (A), and relative levels ofgus (m) and histone H4 (a) mRNA. Transcript levels were quantified from the blots shown in A and B using a PhosphorImager (Molecular Dynamics), and normalized to that of 0 hr (for histone H4, full-scale represents 2-fold difference). Note that the 11-hr sample in the cyclAt culture (A) was degraded, and therefore not included in the quantification in C.

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tion rate peaked during the G2 phase and G2-to-M transition, and decreased during mitosis (note the difference between the two cyclins at 6-7 hr after release, which correspond to the peak of mitotic index). The apparent increase in cyc3aAtrelated gus mRNA levels at the second G2 phase and G2-to-M transition compared with the first ones was not observed in all experiments, but the timing of the transcription peaks throughout the cell cycle was consistent. Arrest of Transformed Cultures by Cell Cycle Blockers. Transcription governed by the cyc3aAt and cyclAt promoters seems to shut off during and upon exit of mitosis, respectively. To characterize the exact stage in mitosis in which these events occur, and to compare the transcriptional regulation in the transformed cultures with steady-state mRNA levels of the same cyclins in Arabidopsis plants treated with cell cycle blockers (2), we applied various blockers to transformed cultures. Stationary-phase cells were diluted 10-fold and grown for 24 hr in the presence of aphidicoline, hydroxyurea (another early S phase blocker that inhibits ribonucleotide reductase), or oryzalin, which inhibits microtubule polymerization and arrests cells at early metaphase. The results of a typical experiment are shown in Fig. 2. Mitotic index and histone H4 mRNA levels were analyzed in order to assess the efficiency of culture arrest. As shown in Fig. 3, it was evident that upon oryzalin treatment, most mitotic cells were arrested at late prometaphase or early metaphase, because most of these cells exhibited scattered, condensed paired chromosomes. Cells at this stage were rarely observed in unarrested cultures, even at the peak of mitotic index, probably due to the short duration of this stage. Cells at anaphase, which were very abundant in

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A) cycling cells at the peak of mitotic index, could not be identified after oryzalin treatment. In cyclAt cultures, the level of gus mRNA increased upon oryzalin treatment (Fig. 2), further indicating that transcription governed by the cyclAt promoter is not yet switched off at early metaphase. In this culture, transcription rate was reduced by aphidicoline or hydroxyurea treatment (note that gus mRNA levels are normalized to that of a control nonarrested culture, in which the proportion of mitotic cells is low). In contrast, all the drugs used decreased the cyc3aAt promoterdriven transcription. This is in agreement with the finding in the synchronized cultures, indicating that the peak of cyc3aAt promoter-related transcription occurs at the G2 phase and G2-to-M transition. Cells at both stages are presumably more abundant in the control nonarrested culture than after treatment with any of the cell cycle blockers, demonstrating the necessity of using synchronized cultures to identify the peak in cyc3aAt transcription. In contrast to cyclAt, cyc3aAt promoterrelated gus mRNA levels were reproducibly lower upon oryzalin treatment than upon aphidicoline or hydroxyurea treatments, indicating that rate of transcription governed by the cyc3aAt promoter is lower at early metaphase than in early S phase.

DISCUSSION

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Con. Aphid. Hu Oryz. FIG. 2. Expression of the cyclAt-gus (A and C) and cyc3aAt-gus (B and D) chimeric genes in stably transformed BY-2 cells treated with cell cycle blockers. RNA was prepared from cells grown for 24 hr in fresh medium (Con.) or in medium containing aphidicoline (Aphid.), hydroxyurea (Hu), or oryzalin (Oryz.). (A and B) RNA blots were stained with methylene blue (M.B.) to detect the total amount of RNA loaded, then washed and sequentially hybridized with the gus and histone H4 gene probes. The mitotic index of the cultures is also indicated (%MI). (C and D) Levels of gus mRNA quantified from the blots shown inA and B using a PhosphorImager (Molecular Dynamics) and normalized to that of control cells.

Tobacco BY-2 suspension cultures can be highly synchronized, and regeneration of stably transformed cell lines is feasible. This makes BY-2 cells an ideal system for studying transcriptional regulation of plant cell cycle genes. The promoters of the Arabidopsis cyclAt 'and cyc3aAt genes mediated cell cycledependent transcriptional oscillation of the gus reporter gene in transformed tobacco BY-2 cell lines. The results presented here are in agreement with mRNA levels of the same cyclins in Arabidopsis plants treated with cell cycle blockers (2). This indicates that transcriptional regulation plays a significant role in determining the mRNA levels of these genes, and that regulatory elements seem to be conserved between species. The reported reduction in cyc3aAt mRNA levels in oryzalintreated root tips, analyzed by whole-mount in situ hybridization (2), was more pronounced than the reduction observed here in cyc3aAt-gus mRNA levels after oryzalin treatment. This difference is not due to the use of a heterologous system, since in root tips of Arabidopsis plants transformed with the same cyc3aAt-gus construct, histochemical f3-glucuronidase activity was not significantly reduced by oryzalin (S. Burssens, unpublished data). One explanation may be the involvement of regulation at the level of cyc3aAt mRNA stability. Alternatively, the promoter region used for studying transcriptional

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regulation may lack some regulatory regions, that are responsible for a stronger decrease in cyc3aAt mRNA levels in early metaphase. To obtain a full understanding of the mechanisms controlling plant cyclin expression it will be necessary to study other putative levels of regulation, such as control of mRNA stability, mRNA translation, and protein stability. The expression pattern of the Arabidopsis cyc3aAt and cyclAt genes seems to correspond to animal A- and B-type cyclins, respectively (see below). However, because of problems in obtaining highly synchronized animal cells, the temporal resolution of the analyses in the latter was lower. It was shown that the steady-state mRNA levels and nuclear run-on transcription rates of the human cyclin A and B1 genes peak at the G2-to-M transition and during the M phase, respectively (13, 14). Cyclin A mRNA was shown to decline more rapidly than that of cyclin Bi after release from a mitotic block by nocodazole (14). Studies in human cell lines, stably transformed with promoter-reporter gene constructs, showed that transcription driven by the cyclin A and cyclin B1 promoters initiates at the onset and at the end of the S phase, respectively, but failed to demonstrate subsequent events (16, 17). The data presented here indicate that the cyc3aAt and cyclAt genes are expressed at a cell cycle window that resemble that of the A and B classes of animal cyclins, respectively. Studies in synchronized alfalfa (Medicago sativa) cells, and double-target in situ hybridization of Antirrhinum and soybean (Glycine max) plants, demonstrated that mRNA levels of the alfalfa cycMs2 gene, the Antirrhinum cyclAm and cyc2Am genes, and the soybean cyc5Gm gene, all more homologous to B cyclins, are abundant in G2 and M phase cells (28-30). Using doubletarget in situ hybridization, it was recently concluded that mRNA of the soybean cyc3Gm gene, which is more related to animal A cyclins, is expressed in late S phase and G2 cells (30). Thus it seems that, although plant cyclins are much more homologous to each other than to animal or yeast cyclins, their sequence-based affiliation to the different groups of animal cyclins is coherent with their expression pattern, suggesting that they may carry out similar functions. Most plants analyzed so far possess cyclins from the two groups, further suggesting that this classification may not only reflect evolutionary divergence, but also conserved functions. Animal A- and B-type cyclins participate in distinct complexes that fulfill different roles during the cell cycle. Both A and B cyclins are required, although presumably carrying out different functions, at the G2-to-M transition, and their degradation is required for exit from mitosis (reviewed in ref. 4). A-type cyclins are also required for the onset of the S phase (31) and for the dependence of mitosis on completion of DNA replication (32). The question whether the roles of the plant "A- and B-type" cyclins is similar to those of the animal mitotic cyclins remains to be resolved. We thank the Tobacco Science Research Laboratory, Japan Tobacco Inc., for permitting us use of the tobacco BY-2 cell suspension and Claude Gigot for providing us this culture, Catherine Bergounioux for flow cytometry, Ben Scheres for the Arabidopsis histone H4 gene, Liz Corben for critical reading of the manuscript, Martine De Cock for help preparing it, and Christiane Germonprez, Rebecca Verbanck, and Karel Spruyt for help preparing the illustrations. This work was supported by grants from the Belgian Programme on Interuniversity Poles of Attraction (Prime Minister's Office, Science Policy Programming, No. 38), the Vlaams Actieprogramma Biotechnologie (ETC

Proc. Natl. Acad. Sci. USA 93 (1996) 002), the Korber Stiftung, and the Fonds voor Geneeskundig Wetenschappelijk Onderzoek (G.0121.96). O.S. is indebted to the Rothschild Foundation and the European Molecular Biology Organization for fellowships. D.I. is a Research Director of the Institut National de la Recherche Agronomique (France). 1. Pines, J. (1994) Semin. Cell Biol. 5, 399-408. 2. Ferreira, P., Hemerly, A., de Almeida Engler, J., Bergounioux, C., Burssens, S., Van Montagu, M., Engler, G. & Inze, D. (1994) Proc. Natl. Acad. Sci. USA 91, 11313-11317. 3. Renaudin, J.-P., Colasanti, J., Rime, H., Yuan, Z. & Sundaresan, V. (1994) Proc. Natl. Acad. Sci. USA 91, 7375-7379. 4. King, R. W., Jackson, P. K. & Kirschner, M. W. (1994) Cell 79, 563-571. 5. Koch, C. & Nasmyth, K. (1994) Curr. Opin. Cell Biol. 6, 451-459. 6. Naeve, G. S., Sharma, A. & Lee, A. S. (1991) Curr. Opin. Cell Biol. 3, 261-268. 7. Morris, T. D., Weber, L. A., Hickey, E., Stein, G. S. & Stein, J. L. (1991) Mol. Cell. Biol. 11, 544-553. 8. Coppock, D. L. & Pardee, A. B. (1987) Mol. Cell. Biol. 7, 2925-2932. 9. Welch, P. J. & Wang, J. Y. J. (1992) Proc. Natl. Acad. Sci. USA 89, 3093-3097. 10. Lucibello, F. C., Truss, M., Zwicker, J., Ehlert, F., Beato, M. & Muller, R. (1995) EMBO J. 14, 132-142. 11. Cogswell, J. P., Godlevski, M. M., Bonham, M., Bisi, J. & Babiss, L. (1995) Mol. Cell. Biol. 15, 2782-2790. 12. Lee, H.-H., Chiang, W.-H., Chiang, S.-H., Liu, Y.-C., Hwang, J. & Ng, S.-Y. (1995) Gene Expression 4, 95-109. 13. Pines, J. & Hunter, T. (1989) Cell 58, 833-846. 14. Pines, J. & Hunter, T. (1990) Nature (London) 46, 760-763. 15. Maity, A., McKenna, W. G. & Muschel, R. J. (1995) EMBO J. 14, 603-609. 16. Henglein, B., Chenivesse, X., Wang, J., Eick, D. & Brechot, C. (1994) Proc. Natl. Acad. Sci. USA 91, 5490-5494. 17. Piaggio, G., Farina, A., Perrotti, D., Manni, I., Fuschi, P., Sacchi, A. & Gaetano, C. (1995) Exp. Cell Res. 216, 396-402. 18. Jacobs, T. W. (1995)Annu. Rev. Plant Physiol. Plant Mol. Biol. 46, 317-339. 19. Ferreira, P. C. G., Hemerly, A. S., de Almeida Engler, J., Van Montagu, M., Engler, G. & Inze, D. (1994) Plant Cell 6, 17631774. 20. Lepetit, M., Ehling, M., Chaubet, N. & Gigot, C. (1992) Mol. Gen. Genet. 231, 276-285. 21. Ohtsubo, N., Nakayama, T., Terada, R., Shimamoto, K. & Iwabuchi, M. (1993) Plant Mol. Biol. 23, 553-565. 22. Kapros, T., Stefanov, I., Magyar, Z., Ocsovszky, I. & Dudits, D. (1993) In Vitro Cell. Dev. Biol. 29P, 27-32. 23. Nagata, T., Nemoto, Y. & Hasezawa, S. (1992) Int. Rev. Cytol. 132, 1-30. 24. Jefferson, R. A., Kavanagh, T. A. & Bevan, M. W. (1987) EMBO J. 6, 3901-3907. 25. An, G. (1985) Plant Physiol. 79, 568-570. 26. Mikami, K. & Iwabuchi, M. (1993) in Control of Plant Gene Expression, ed. Verma, D. P. S. (CRC, Boca Raton, FL), pp. 51-68. 27. Reichheld, J.-P., Sonobe, S., Clement, B., Chaubet, N. & Gigot, C. (1995) Plant J. 7, 245-252. 28. Hirt, H., Mink, M., Pfosser, M., Bogre, L., Gyorgyey, J., Jonak, C., Gartner, A., Dudits, D. & Heberle-Bors, E. (1992) Plant Cell 4, 1531-1538. 29. Fobert, P. R., Coen, E. S., Murphy, G. J. P. & Doonan, J. H. (1994) EMBO J. 13, 616-624. 30. Kouchi, H., Sekine, M. & Hata, S. (1995) Plant Cell 7, 1143-1155. 31. Girard, F., Strausfeld, U., Fernandez, A. & Lamb, N. J. C. (1991) Cell 67, 1169-1179. 32. Walker, D. H. & Maller, J. L. (1991) Nature (London) 354, 314-317.