3′ non-translated sequences in Drosophila cyclin B transcripts direct posterior pole accumulation late in oogenesis and peri-nuclear association in syncytial embryos

0 Cancer Research Campaign Laboratories, Cell Cycle Genetics Group, Department of Biochemistry, The University , Dundee, DD1 4HN, Scotland - We have characterised forms of the Drosophila cyclin B transcript that differ as a result of a splicing event which removes a nucleotide segment from the 3 untranslated region. In oogenesis, both cyclin A RNA and a shorter form of the cyclin B transcript are seen in the cells of the germarium that are undergoing mitosis. The shorter cyclin B transcript alone is then detectable in the presumptive oocyte until stages 7-8 of oogenesis. Both cyclin A RNA and a longer form of the cyclin B RNA are then synthesised in the nurse cells during stages 9-11, to be deposited in the oocyte during stages 11-12. These transcripts become evenly distributed throughout the oocyte cytoplasm but, in addition, those of cyclin B become conThe mitotic cyclins, proteins first identified in the eggs of marine invertebrates, are characterised by changes in their abundance during the cell cycle; they accumulate in interphase and are abruptly degraded in mitosis (Evans et al., 1983; Standart et al., 1987; Swenson et al., 1986). Cyclins are required to complex with and thus activate the major cell cycle regulatory protein kinase p34 cdc2 (Swenson et al., 1987; Draetta et al., 1989; Meijer et al., 1989; Solomon et al., 1990; Parker et al., 1991; Gautier and Maller, 1991). The active kinase complex brings about the phosphorylation of a variety of substrates including histone H1, microtubules, centrosomes and nuclear lamins. These phosphorylation events are among those required for chromosome condensation, organisation of the mitotic spindle and breakdown of the nuclear envelope prior to entry into mitosis (reviewed by Moreno and Nurse, 1990). Inactivation of the p34cdc2 kinase is mediated by abrupt degradation of the cyclins. Addition of a cyclin B mRNA encoding a truncated form of the protein, resistant to proteolytic cleavage, prevents exit from mitosis in Xenopus oocyte extract showing that cyclin degradation is necessary for progress through the cell cycle (Murray and Kirschner, 1989; Murray et al., 1989). p34cdc2 is part of a conserved mechanism that regulates centrated at the posterior pole. Examination of the distributions of RNAs transcribed from chimeric cyclin genes indicates that sequences in the 3 untranslated region of the larger cyclin B RNA are required both for it to become concentrated at the posterior pole and to direct those transcripts in the body of the syncytial embryo to their peri-nuclear localisation. These sequences are disrupted by the splicing event which generates smaller cyclin B transcripts. the G2-M transition. Thus, the cdc2 gene product of S.pombe may be functionally replaced by the Saccha romyces cerevisiae, Drosophila or human homologues (Beach et al., 1982; Jimenez et al., 1990; Lehner and OFarrell, 1990b; Lee and Nurse, 1987). The A and B type cyclin genes are similarly conserved and have been cloned and sequenced from a number of higher eukaryotes including Drosophila, sea urchins, clams, Xenopus and humans (Lehner and OFarrell, 1989, 1990a; Whitfield et al., 1989; Pines and Hunt, 1987; Swenson et al., 1986; Westendorf et al., 1989; Minshull et al., 1989, 1990; Pines and Hunter, 1989, 1990). A comparison of the cyclin sequences in a wide range of organisms points towards the conservation of the distinct A and B types of cyclin suggesting these molecules have differing roles in the cell cycle. In Drosophila this is supported by the finding that embryos having mutations in the cyclin A gene arrest development in the cell cycles that follow cellularisation, once the maternal contribution to the embryo has been exhausted. Hence cyclin B cannot substitute for cyclin A function (Lehner and OFarrell, 1989). Examination of the behaviour of cyclin A and B proteins in cellularised Drosophila embryos and larval brains reveals differences in the timing of their accumulation and breakdown (Whitfield et al., 1990). This may reflect the differential timing of the activation of the p34cdc2 kinase associated with either cyclin A or cyclin B as observed in Xenopus cell-free systems (Minshull et al., 1990). Pines and Hunter (1991) have also found that the two cyclins accumulate in different sub-cellular compartments in mammalian cells. In the syncytial Drosophila embryo, cyclin A appears to shuttle between an association with chromatin and the cytoplasm, whereas cyclin B is localised to the region at which nuclear envelope breakdown begins and is subsequently associated with polar microtubules of the mitotic spindle (Maldonado-Codina and Glover, 1992). An abundant maternal supply of both cyclin A and B transcripts is present in the unfertilized Drosophila egg. Unlike the uniformly distributed cyclin A transcripts, those of cyclin B are concentrated at the posterior pole of egg. During embryogenesis cyclin B mRNA becomes incorporated into the progenitors of the germ-line, the pole-cells (Whitfield et al., 1989; Lehner and OFarrell, 1990a; Raff et al., 1990). Cyclin B transcripts also become more closely concentrated around the somatic nuclei than those of cyclin A in a manner that requires the integrity of microtubules. In order to establish and maintain the posterior pole localisation, a component of the posterior cytoplasm is required. The distribution of cyclin B transcripts in a variety of mutant embryos that fail to form pole cells suggests that this localisation requires a polar granule associated component (Raff et al., 1990). In this paper we examine the distribution patterns of cyclin A and B transcripts during oogenesis and identify a signal in the cyclin B transcript required for both the posterior pole and nuclear localisation. Materials and methods Hybridisation was to poly (A)+ RNA immobilised on nitrocellulose filters (gift of Dr. K. OHare) with probes made by random oligo-labelling (Feinburg and Vogelstein, 1983) of cDNA fragments shown in Fig 2. Hybridisation was carried out at 42C for 24 hours in 50% formamide, 0.75 M NaCl, 0.15 M Tris-HCl pH 8, 10 mM EDTA, 200 mg/ml denatured salmon sperm DNA, 0.5% SDS, 36 mM Na 2HPO4, 4 mM NaH 2PO4, 5 Denhardts solution. Filters were washed in 5 Denhardts, 0.3 M NaCl, 60 mM TrisHCl pH 8, 4 mM EDTA, 2 30 minutes followed by 75 mM NaCl, 15 mM Tris-HCl, 1 mM EDTA, 3 10 minutes. Filters were exposed for 7 days at - 70C with intensifying screens. In situ hybridisation to whole mount embryos and ovaries In situ hybridisation was carried out according to the method of Tautz and Pfeifle (1989). Ovaries were dissected in 0.7% NaCl and placed directly into fixative (4% paraformaldehyde in PBS; phosphate buffered saline, 120 mM NaCl, 10 mM sodium phosphate, with an equal volume of heptane). Embryos were also treated using this fixative after dechorionation in 50% hypochlorite. Following fixation, embryos were devitellinised by vigorous shaking in 1:1 heptane:methanol (Mitchison and Se (...truncated)