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. Author manuscript; available in PMC: 2009 Sep 28.
Published in final edited form as: Cell. 1990 May 4;61(3):535–547. doi: 10.1016/0092-8674(90)90535-m

The Roles of Drosophila Cyclins A and B in Mitotic Control

Christian F Lehner 1,*, Patrick H O’Farrell 1
PMCID: PMC2753431  NIHMSID: NIHMS142631  PMID: 2139805

Summary

We have cloned, sequenced, and characterized the expression of a Drosophila cyclin B gene. The independent evolutionary conservation of A- and B-type cyclins implies that they have distinct roles. Indeed, in mutant embryos deficient in cyclin A, cells that accumulate only cyclin B do not enter mitosis. Thus, in vivo, cyclin B is not sufficient for mitosis. Furthermore, we find that the two cyclins are coexpressed in all proliferating cells throughout development. Though lacking a formal demonstration that cyclin B is essential as it is in other organisms, we propose that each of these proteins fulfills a distinct and essential role in the cell cycle.

Introduction

In all eukaryotes, mitotic processes are controlled by a 34 kd kinase that was initially identified in Schizosaccharomyces pombe as the product of the cdc2+ gene (Simanis and Nurse, 1986). This p34cdc2 kinase is activated just before cells enter mitosis (Draetta and Beach, 1988; Moreno et al., 1989). Histone H1 appears to be one of its substrates (Arion et al., 1988; Labbé et al., 1988; Brizuela et al., 1989; Langan et al., 1989). In addition, the activated kinase is thought to start a complex kinase activation cascade that directs the cell into mitosis. In support of its proposed master role in initiating mitosis, the p34cdc2 kinase has been found in all preparations of MPF (maturation- or mitosis-promoting factor) that have been purified using biological assays for mitosis-promoting activity (Dunphy et al., 1988; Gautier et al., 1988; Labbé et al., 1989a, 1989b). While entry into mitosis requires activation of the p34cdc2 kinase, exit depends on its inactivation (Murray et al., 1989).

The accurate regulation of the p34cdc2 kinase involves a number of regulators. Among them, cyclin proteins have properties particularly pertinent to both activation and inactivation of the p34cdc2 kinase. Cyclin proteins are known to be physically associated with the p34cdc2 kinase (Booher et al., 1989; Draetta et al., 1989; Labbé et al., 1989b; Meijer et al., 1989; Pines and Hunter, 1989). In the absence of cyclin protein expression, p34cdc2 kinase activity (H1 kinase activity and MPF activity) is undetectable and cells fail to enter mitosis (Booher and Beach, 1988; Minshull et al., 1989; Murray and Kirschner, 1989; Murray et al., 1989). These observations are consistent with the idea that cyclin proteins are subunits that bind and activate the p34cdc2 kinase. Importantly, cyclin proteins are not continuously present during the cell cycle but are rapidly and completely degraded during each metaphase (Evans et al., 1983; Standart et al., 1987). This degradation has been shown to be required for the completion of mitosis and might be responsible for the observed inactivation of the p34cdc2 kinase during mitosis (Murray et al., 1989). After mitosis, cyclin proteins reaccumulate to attain maximal concentrations just before mitosis (Evans et al., 1983), when the p34cdc2 kinase activity is also maximal. Although cyclin proteins are clearly important regulators of the p34cdc2 kinase, the sudden premitotic increase in p34cdc2 kinase activity is not simply a reflection of the gradual increase in the abundance of cyclin proteins (Meijer et al., 1989; Pines and Hunter, 1989). Other regulators are known to participate in the regulation of the p34cdc2 kinase, but their physiological roles have yet to be established (Russell and Nurse, 1986; Hayles et al., 1986; Russell and Nurse, 1987a, 1987b; Moreno et al., 1989; Booher et al., 1989).

Cyclins as well as other regulators controlling entry into mitosis have been highly conserved in evolution. Homologs of cyclin proteins originally identified in clams, sea urchins, and starfish have been found in yeast, Xenopus, humans, and Drosophila (Evans et al., 1983; Swenson et al., 1986; Standart et al., 1987; Pines and Hunt, 1987; Booher and Beach, 1988; Hagan et al., 1988; Lehner and O’Farrell, 1989; Pines and Hunter, 1989; Westendorf et al., 1989; Whitfield et al., 1989). In clams, two different but highly homologous cyclins, cyclin A and cyclin B, have been identified (Evans et al., 1983; Swenson et al., 1986; Westendorf et al., 1989). Interestingly, homologs of both cyclin A and cyclin B have been found in Xenopus (T. Hunt, personal communication) and Drosophila (Whitfield et al., 1989; this report).

The fact that these two cyclin types have been conserved in evolution suggests that they have different roles. Available evidence, however, fails to identify distinct functions for the two cyclin types. After injection of synthetic mRNAs into Xenopus oocytes–the only functional assay that has been performed with both cyclin types–either clam cyclin A or cyclin B mRNAs induce Oocyte maturation (Swenson et al., 1986; Westendorf et al., 1989). Moreover, in cyclin-depleted Xenopus egg extracts, addition of a B-type sea urchin cyclin is sufficient to restore cell cycle progression in vitro (Murray and Kirschner, 1989). In Drosophila, in situ hybridization experiments indicated stage- and tissue-specific differences in the expression of the two cyclin types (Whitfield et al., 1989), This observation suggested a specialization of cyclin types for particular cell types and thus offered a possible explanation for why two functionally redundant cyclin types might have been conserved in evolution.

The results presented here, however, lead to contrary conclusions. The levels of cyclin B mRNA do not directly reflect the levels of cyclin B accumulation. Cyclin A and cyclin B proteins are coexpressed in all tissues and stages analyzed. Moreover, analysis of cyclin B expression in mutant, cyclin A–deficient embryos clearly indicates that cyclins A and B are not functionally redundant. We suggest that cyclins A and B are specialized to fulfill distinct roles, both fundamental for mitotic control in all cells.

Results

Cyclin Genes in Drosophila

We previously isolated a Drosophila gene encoding a cyclin homologous to clam cyclin A and, to a lesser extent, to cyclin B (Lehner and O’Farrell, 1989). To identify additional cyclin genes, we designed two primers recognizing conserved sequences found in all cyclin cDNAs isolated so far (see Experimental Procedures). These primers were used for polymerase chain reaction (PCR) experiments (data not shown). When total genomic DNA from Drosophila served as a template, PCR products were resolved on a gel as two bands, 260 bp and 210 bp in size. The larger PCR product appears to be derived from the cyclin A gene, since a 260 bp product was also observed when cloned genomic cyclin A sequences were used as a template.

The following evidence demonstrates that the smaller PCR product (210 bp) amplified from total genomic DNA was derived from a cyclin B gene. The DNA in this smaller band was gel purified and used as a probe to isolate cDNAs. The sequence of the longest, apparently full-length cDNA (2691 bp) and the deduced amino acid sequence of the encoded long open reading frame are shown in Figure 1. In addition to the open reading frame that starts at an ATG in a context matching the consensus for translational start sites (Cavener, 1987), the cDNA has a polyadenylation site followed by a poly(A) tail. As revealed by homology searches, the encoded polypeptide is most similar to B-type cyclins from clam, Xenopus, sea urchin, and humans (Pines and Hunt, 1987; Minshull et al., 1989; Pines and Hunter, 1989; Westendorf et al., 1989).

Figure 1. Nucleotide Sequence and Predicted Amino Acid Sequence of a Drosophila Cyclin B cDNA.

Figure 1

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The nucleotide sequence of a Drosophila cyclin B cDNA (2691 bp) has a long open reading frame encoding a protein of 530 amino acids. The predicted amino acid sequence is shown above the nucleotide sequence. The putative initiating ATG is underlined along with flanking bases matching the consensus sequence for translational start sites in Drosophila (Cavener, 1987). The putative polyadenylation signal, followed by a poly(A) tail, is also underlined.

A comparison of the conserved C-terminal domains of the cyclins from clam and Drosophila, the only two species for which both A- and B-type sequences have been published (Swenson et al., 1986; Westendorf et al., 1989; Lehner and O’Farrell, 1989; this report), demonstrates that cyclin A and cyclin B have been independently conserved in evolution (Figure 2) and confirms previous findings based on the analysis of frog cyclins (T. Hunt et al., personal communication). In the conserved region shown in Figure 2, the A-type cyclins of the two evolutionary distant species clam and Drosophila are 45% identical; the two B-type cyclins are 47% identical. The extent of identity is significantly lower when A-type cyclins are compared with B-type cyclins (clam A and clam B, 36%; Drosophila A and Drosophila B, 31%; clam A and Drosophila B, 29%; Drosophila A and clam B, 33%).

Figure 2. Amino Acid Sequence Comparison of Drosophila and Clam Cyclins.

Figure 2

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The amino acid sequences of the conserved cyclin domains of Drosophila cyclin B (D.B), clam cyclin B (C.B), Drosophila cyclin A (D.A), and clam cyclin A (C.A) are aligned. Gaps introduced for optimal alignment are indicated by dashes. Regions are boxed if the two A-type cyclins or the two B-type cyclins have similar amino acids (identical or conserved replacements using the following grouping: A,L,V,I,M; K,R; D,E; S,T; N,Q; Y,F). Positions similar in at least three sequences are marked with asterisks. Boldface indicates sequence identities in at least three of the four sequences. The clam sequences are according to Swenson et al. (1986) and Westendorf et al. (1989).

The size of the PCR products generated from the cyclin A gene (260 bp) and the cyclin B gene (210 bp) indicated that the former but not the latter contained a small intron in the amplified region: 210 bp fragments were obtained irrespective of whether cloned genomic cyclin B sequences, cyclin B cDNA, or cyclin A cDNA was used as the template. In the PCR products obtained with total genomic DNA, we detected no sequences other than those derived from the identified cyclin A and cyclin B genes (data not shown). On Southern blots, cross-hybridization between the two cyclin genes was observed only at very low stringency, even when probes from the most conserved regions where used. Under these conditions additional faint signals were also detected on genomic blots. Therefore we cannot exclude the possibility that additional cyclin genes are present, although we did not detect any as PCR products.

Expression and Distribution of Cyclin B mRNA

The expression of cyclin B was analyzed by Northern blot and in situ hybridization experiments. On Northern blots probed with a cyclin B cDNA probe, an abundant maternal mRNA (2.7 kb) was detected in total RNA from early embryos that had not yet started zygotic transcription (Figure 3, lane 1). A much lower signal was observed in RNA isolated from embryos that had completed the early, syncytial divisions (Figure 3, lane 2). Subsequently, zygotic transcription led to reaccumulation of relatively high levels of mRNA in older embryos (Figure 3, lane 3). The developmental profile of cyclin B mRNA abundance was very similar to the one observed with cyclin A probes (Lehner and O’Farrell, 1989). The abundance of cyclin transcripts mirrored the rate of proliferation. High levels were present during the rapid syncytial cell cycles (Figure 3, lane 1) and during the later embryonic cell divisions (Figure 3, lane 3). Lower levels were found before gastrulation when the nuclei became cellularized and no cell divisions occurred (Figure 3, lane 2), and in older embryos where only a few cells continued to divide (Figure 3, lane 4).

Figure 3. Cyclin B mRNA Levels during Embryonic Cell Cycle Progression.

Figure 3

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Total RNA from an equal number of early embryos during the syncytial cleavage divisions (lane 1), during interphase of Cycle 14 (lane 2), during cycle 15 and 16 (lane 3), and at a stage after cell divisions had become restricted mainly to the nervous system (lane 4) were probed on Northern blots with the cyclin B cDNA. The estimated size (kb) of the cyclin mRNA is indicated at left. Embryos were collected and aged at 25°C for the times indicated and staged under the microscope before isolation of RNA. Reprobing the filter with a probe specific for ribosomal protein rp49 (O’Connell and Rosbash, 1984) revealed roughly equal signal intensities in all four lanes (not shown).

Using a recently developed in situ hybridization method that relies on cytochemical detection of hybrids (Tautz and Pfeiffle, 1989), we analyzed the localization of cyclin B mRNA (Figure 4). In preblastoderm embryos, cyclin B mRNA was present throughout the cytoplasm and at a much higher level at the posterior end (Figure 4A). Because of the abundance of cyclin B mRNA, this posterior concentration was most evident if the color reaction used for detection of the hybridized probe was terminated early (Figures 4A–4D). During nuclear migration, the mRNA cap at the posterior end was incorporated into the forming pole cells and the dispersed fraction of the cyclin B mRNA appeared to migrate in association with the nuclei to the periphery (Figure 4B). In the pole cells (the prospective germ cells), cyclin B mRNA appeared to be sequestered into cytoplasmic granules (Figure 4C). In somatic cells, signal intensities became progressively weaker during the blastoderm stages. If the color reaction was terminated early, no signal was apparent in somatic cells (Figure 4D). However, if the color reaction was allowed to proceed longer (Figures 4E–4I), specific labeling was clearly detectable in the cytoplasm of somatic cells (Figure 4E). Therefore, cyclin B mRNA is not exclusively localized to the posterior pole. In contrast to cyclin B mRNA, cyclin A mRNA was found to be uniformly distributed (not shown). During gastrulation, signal intensities in somatic cells increased again and cyclin B mRNA was found throughout the embryo (Figure 4F). In germband-retracting embryos, label was observed in the developing nervous system, but no longer in epidermal cells that had completed their developmental program of cell divisions (Figure 4G).

Figure 4. Localization of Cyclin B mRNA during Drosophila Embryogenesis.

Figure 4

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Cyclin B mRNA was localized by a color reaction after in situ hybridization with a digoxigeninylated cyclin B cDNA probe and incubation with alkaline phosphatase–conjugated anti-digoxigenin antibodies. Early embryos were staged after double labeling with the DNA stain Hoechst 33528. The DNA labeling is shown only in (C), which shows a combination of fluorescence and bright-field images.

(A)Embryo before nuclear migration (cycle 3). Maximal signal intensities were found in a cap at the posterior end. In addition, signal intensities increased gradually toward the anterior end.

(B)Embryo during nuclear migration (cycle 8). Cyclin B mRNA migrated in close association with the nuclei toward the periphery, and the signal at the polar cap was included in the polar buds.

(C)Pole cells (cycle 14). After formation of the pole cells, cyclin B mRNA appeared to be sequestered into cytoplasmic granules clustered around the nuclei.

(D)Somatic cells (cycle 14). No signal was detected in somatic cells if the color reaction was terminated early, as in (A)–(D).

(E) Somatic cells (cycle 14). A specific signal in the cytoplasm of somatic cells was detected after prolonged color reaction as shown in (E)–(I).

(F) Embryo during cycle 15. Signal was observed throughout the embryo except in the amnioserosa (labeled A).

(G) Region from an embryo after cycle 16. No signal was detected in epidermal cells (between arrows) that had completed their developmental division program. Signal was present, however, in the nervous system (labeled N) where cell divisions continue.

(H) Same region as in (G) but from an embryo homozygous for a null mutation in the cyclin A gene (neo114). As in wild-type embryos (G), cyclin B transcripts were no longer detected in epidermal cells during germband retraction. Mutant embryos were identified on the basis of their reduced nuclear densities as revealed by double labeling with the DNA stain Hoechst 33528 (not shown; see Lehner and O’Farrell, 1989).

(I) Same as in (H) but hybridized with a cyclin A cDNA probe demonstrating the lack of zygotic cyclin A expression in homozygous neo114 embryos. In all panels except (C), anterior is left and ventral down. (C) shows the pole cells at the posterior end.

Characterization of Anti-Cyclin Antibodies

To analyze the expression of cyclin B protein, we raised antibodies against a bacterially produced cyclin B fusion protein. Because we intended to use these antibodies for double-labeling experiments in combination with the previously obtained rabbit antibodies against cyclin A (Lehner and O’Farrell, 1989), antibodies were produced in mice. In addition, it was important to rule out cross-reactivity. To this end, the specificity of the anti-cyclin antibodies was analyzed by immunoprecipitation and immunoblotting experiments. For immunoprecipitation experiments, radiolabeled full-length cyclin A (Figure 5A, lane 1) and cyclin B (Figure 5A, lane 2) were synthesized in transcription–translation reactions. The two cyclins were mixed (Figure 5A, lane 3) and the affinity-purified antibodies against cyclin A or the antiserum against cyclin B was used for immunoprecipitation from this mixture. The anti-cyclin A antibodies reacted only with cyclin A (Figure 5A, lane 4), and the anti-cyclin B antibodies reacted only with cyclin B (Figure 5A, lane 5). No immunoprecipitation was observed with the mouse preimmune serum (Figure 5A, lane 6).

Figure 5. Characterization of Anti-Cyclin Antibodies.

Figure 5

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Immunoprecipitation experiments. Radiolabeled cyclin A (lane 1) and cyclin B (lane 2) were obtained after in vitro transcription–translation reactions. The two proteins were mixed (lane 3), and immunoprecipitations were done using affinity–purified rabbit antibodies against cyclin A (lane 4), a mouse antiserum against cyclin B (lane 5), or the corresponding mouse preimmune serum (lane 6). Proteins were resolved on a 7.5% SDS–polyacrylamide gel and visualized by autoradiography. Only the region containing cyclin A and cyclin B is shown. The estimated sizes (kd) of cyclin A and cyclin B are indicated at left. Immunoblotting experiments. Total protein extracts from 0–1 hr Drosophila embryos were resolved on a 10% SDS–polyacrylamide gel and stained with Coomassie blue (lane 2) or transferred to nitrocellulose (lanes 3–5) and probed with anti-cyclin A antibodies (lane 3), the mouse antiserum against cyclin B (lane 4), or the corresponding mouse preimmune serum (lane 5). Sizes (kd) of the marker proteins in lane 1 are indicated at left.

The specificity of the antibodies was further analyzed by immunoblotting experiments. As shown previously (Lehner and O’Farrell, 1989), the anti-cyclin A antibodies detected a closely spaced doublet (59 and 61 kd) in total protein from early Drosophila embryos (Figure 5B, lane 3). The 59 kd form has the same gel electrophoretic mobility as cyclin A obtained in transcription–translation reactions and most likely represents unmodified cyclin A, while the 61 kd form appears to be modified cyclin A. The anti-cyclin B antiserum reacted exclusively with a protein of the same size (64 kd) as the one obtained in transcription–translation reactions (Figure 5B, lane 4). The specificity of the antibodies was further demonstrated by the results of immunofluorescent stainings of mutant embryos unable to express cyclin A zygotically (Lehner and O’Farrell, 1989; and see below). All these observations demonstrate that the anti-cyclin antibodies are specific for either cyclin A or cyclin B; no cross-reaction is evident.

Expression of Cyclin B Protein

As expected from the high abundance of maternal cyclin B mRNA, an intense immunofluorescent signal was detected in early embryos (not shown). However, the enrichment of cyclin B mRNA at the posterior pole and in the pole cells was not paralleled by a corresponding distribution of cyclin B protein. Anti-cyclin B labeling appeared uniform in early embryos. Moreover, in double-labeling experiments the same ratio of immunofluorescent signal was observed in somatic cells and pole cells with both anti-cyclin A and anti-cyclin B antibodies (see Figure 7A), indicating that cyclin B has the same distribution as cyclin A, which is translated from a uniformly distributed mRNA (not shown). These observations led us to suspect that the localized fraction of cyclin B mRNA might not be readily available for translation.

Figure 7. Cyclin B Distribution during Embryogenesis.

Figure 7

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Embryos were stained with the antiserum against cyclin B followed by rhodamine-conjugated secondary antibodies (A, B, D). Double labeling with anti-cyclin A antibodies and fluorescein-conjugated secondary antibodies is shown in (C) and in the inset in (A). Embryos at the following stages are shown:

(A) Onset of mitosis 14. Uniform cyclin B labeling is present in the cytoplasm. The same ratio between labeling intensity in somatic cells and pole cells (indicated by arrows) is observed with both anti-cyclin B antibodies and anti-cyclin A antibodies (inset).

(B) Late in mitosis 14. Most cells have completed mitosis 14 and are not labeled with anti-cyclin B antibodies. Cells in the neurogenic region (N), in the amnioserosa (A), and in the posterior midgut (P) have not yet entered mitosis 14 and are intensely labeled.

(C, D) After mitosis 16. Most cells have completed their developmental program of cell divisions and are no longer labeled. Cells in the central and peripheral nervous system that undergo additional mitotic divisions are labeled. Double labeling with anti-cyclin A(C) and anti-cyclin B antibodies (D) indicates that the two cyclins are coexpressed. The insets show a region of the embryo at a higher magnification. In all panels, anterior is left, ventral down.

Interestingly, as seen for cyclin A (Lehner and O’Farrell, 1989), there was no obvious cell cycle–dependent degradation of cyclin B during the early syncytial cell cycles (mitoses 1–9) and only partial degradation in the cortical layer after nuclear migration (mitoses 10–13). Degradation of cyclins during the early syncytial stages and the role of degradation for this syncytial cell cycle progression will be considered elsewhere. Here we focus on the accumulation and degradation of cyclin B in the somatic cells during postcellularization embryonic divisions.

Before the first cell division (mitosis 14), cyclin B accumulates in the cytoplasm and immunofluorescent labeling is found to be excluded from the nucleus (Figure 7A and interphase cells in Figures 6A and 6B [see i]). In addition to the cytoplasmic labeling, however, a single dot in the nucleus was also labeled (Figure 6A, arrowhead). Two nuclear dots were observed immediately before entry into mitosis. In control experiments with the preimmune serum, no labeling was observed (not shown). The intensity of this intranuclear labeling varied in different experiments, indicating that fixation conditions might not be optimal. No intranuclear labeling was observed with anti-cyclin A antibodies (Lehner and O’Farrell, 1989), even in double-labeling experiments in which strong labeling of intranuclear dots was observed with anti-cyclin B antibodies (not shown). In prophase, labeling of the nuclear dots was no longer detectable and diffuse labeling was found predominantly in the region of the condensing chromatin (Figures 6A and 6B, cell marked p). By the end of metaphase, labeling was almost undetectable (Figures 6A and 6B, cell marked m). In anaphase and in subsequent stages of mitosis, cyclin B was no longer detectable (Figures 6A and 6B, cell marked a). In the subsequent interphase, labeling in the cytoplasm and in the intranuclear dots reaccumulated gradually.

Figure 6. Intracellular Localization of Cyclin B during Interphase and Mitosis.

Figure 6

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Embryos were labeled with the antiserum against cyclin B(A) and with the DNA stain Hoechst 33258 (B). During interphase (i), the cytoplasm and intranuclear dots (arrowhead) were stained. In prophase (p), staining concentrated in the region of the condensing chromatin. Labeling became weak late in metaphase (m) and was undetectable in anaphase (a). In double-labeling experiments using rabbit anti-cyclin A(C), mouse anti-cyclin B antibodies (D), and Hoechst 33258 (E), cyclin A was undetectable in late metaphase cells (arrowheads) in which cyclin B was still detected. Primary antibodies were detected with rhodamine-conjugated (A, D) or fluorescein-conjugated (C) secondary antibodies.

Cyclin A is cytoplasmic during interphase, relocates to the nucleus during prophase, and is degraded during metaphase (Lehner and O’Farrell, 1989). Based on double-labeling experiments using anti-cyclin A and anti-cyclin B antibodies, the time course of degradation appears to be different for cyclin A and cyclin B. Cyclin A was no longer detectable at the end of metaphase (Figure 6C, arrowheads). In contrast, cyclin B was still detectable at this stage (Figure 6D, arrowheads). These observations suggest that cyclin A might be degraded before cyclin B, and it appears that this difference has been conserved in evolution since the same difference was observed in clam (Evans et al., 1983; Luca and Ruderman, 1989).

In an in vitro cell cycle system derived from Xenopus eggs, the rate of cyclin accumulation was found to determine the time of mitotic events. Whereas high rates of accumulation induced mitotic events early, low rates resulted in a delayed onset of mitotic events (Murray and Kirschner, 1989). However, based on immunofluorescence experiments, differential rates of cyclin B accumulation are not involved in controlling the timing of the first asynchronous mitosis in Drosophila embryos (mitosis 14). Before mitosis 14, cyclin B accumulated at the same rate in all cells throughout the embryo (Figure 7A) despite the fact that cells enter mitosis 14 asynchronously in a pattern highly reproducible from embryo to embryo (Foe, 1989). Cells that had passed metaphase were easily recognized, however, because they no longer stained with anti-cyclin antibodies. Thus as illustrated in Figure 7B, the patterned cell divisions were readily visualized. Accumulation and degradation of cyclin B were also found to accompany subsequent cell cycles (see below).

After mitosis 16, most cells terminate mitotic proliferation. In the peripheral and central nervous system, however, cell divisions continue and an intricate pattern of cyclin B expression is observed (Figure 7D). In contrast, no cyclin B signal was detected in cells that had completed their developmental program of cell division. These observations indicate that cyclin B expression is correlated with proliferation.

Double-labeling experiments using anti-cyclin A and anti-cyclin B antibodies indicated that the two cyclins are coexpressed even in late embryos where cyclins are expressed in intricate patterns (Figures 7C and 7D). The same pattern revealed by anti-cyclin B antibodies (Figure 7D) was also revealed by the anti-cyclin A antibodies (Figure 7C). At the higher magnification shown in the insets, co-expression can be seen at the single-cell level, though relative levels of cyclin A and cyclin B expression differ from cell to cell. Coexpression of cyclins A and B was also observed in imaginal discs and brains of third-instar larvae (not shown).

Cyclin B Expression and Mitosis in Cyclin A–Deficient Mutants

The identification of null mutations in the cyclin A gene (Lehner and O’Farrell, 1989) allowed us to ask whether cyclin B expression is sufficient to trigger entry into mitosis. We showed previously that cells in the dorsolateral epidermis of mutant embryos that are unable to express cyclin A zygotically progress through the first 15 rounds of mitosis, presumably owing to maternally derived cyclin A. In these mutant embryos, however, epidermal cells never entered into the 16th mitosis (Lehner and O’Farrell, 1989). The anti-cyclin B antibodies provide a probe to examine divisions in such mutant embryos. During interphase preceding mitosis 15, when cyclin A accumulation was nearly undetectable in mutant embryos (compare Figures 8A and 8C), cyclin B accumulation was normal (compare Figures 8B and 8D). Surprisingly, based on the observed patterns of cyclin B staining (Figures 8B and 8D) and the distribution of mitotic figures (not shown), mitosis 15 appeared to proceed normally despite the very low levels of cyclin A.

Figure 8. Cyclin Expression during Mitosis 15 and 16 in Wild-Type and Mutant Embryos.

Figure 8

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Wild-type embryos (A, B, E, F) and embryos homozygous for a null mutation in the cyclin A gene (C, D, G, H) during mitosis 15 (A–D) and mitosis 16 (E–H) are shown after double labeling with anti-cyclin A (A, C, E, G) and anti-cyclin B antibodies (B, D, F, H). For further explanations see text. In all panels anterior is left, ventral down.

After the degradation of the cyclins during mitosis 15, cyclin A accumulation was no longer detectable in mutant embryos (Figure 8G), yet cyclin B accumulated normally (Figure 8H). In contrast to mitosis 15, the cells of the mutant dorsolateral epidermis never entered mitosis 16 and mitotic figures could never be detected (not shown). In wild-type embryos of the same age, mitotic figures were readily observed. In addition, mitosis 16 was readily visualized in wild-type embryos by both anti-cyclin A and anti-cyclin B stainings (Figures 8E and 8F): sharp boundaries separated labeled cells that had not yet divided from unlabeled cells that had already passed metaphase 16. In contrast to wild-type embryos, mutant embryos showed more uniform staining with anti-cyclin B antibodies; abrupt transitions from labeled to unlabeled cells were not evident (Figure 8H). These observations indicate that in mutant embryos cells are blocked prior to mitosis despite the accumulation of cyclin B. Thus cyclin B accumulation without cyclin A is not sufficient for entry into mitosis.

The abrupt disappearance of cyclin B labeling that normally accompanies mitosis did not occur in the cyclin A mutants arrested in interphase 16. Similarly, string (stg) mutant embryos that are arrested in interphase of cycle 14 sustain high levels of both cyclin A and cyclin B (C. F. L. and P. H. O., unpublished data). All of these observations are consistent with the idea that abrupt mitotic degradation of cyclins is dependent on mitotic processes.

Interestingly, although abrupt disappearance of cyclin B labeling did not occur in cyclin A mutant cells arrested in interphase 16, cyclin B labeling nonetheless disappeared slowly from the mutant epidermis during subsequent development and became confined to the developing central and peripheral nervous system (not shown). In situ hybridization experiments suggested that these changes in the pattern of cyclin B expression result from controls at the transcriptional level. In both wild-type and mutant embryos, cyclin B transcripts faded from epidermal cells at an age where epidermal cells of wild-type embryos had completed their developmental program of divisions (Figures 4G and 4H). In contrast, high levels of cyclin B transcripts persisted in cells of the developing nervous system in both mutant and wild-type embryos (Figures 4G and 4H). These observations suggested that the developmental regulation of cyclin B gene transcription is not affected by the fact that cells in the mutant embryos stop division prematurely.

Discussion

The identification of both an A-type and a B-type cyclin in Drosophila demonstrates that the two cyclin types originally identified in clam have been independently conserved in evolution. To study the roles of these highly related mitotic regulators, we have analyzed their subcellular location and expression in wild-type and cyclin A mutant embryos.

Cyclin A and Cyclin B Are Coexpressed

Work from D. Glover’s laboratory has suggested that the two Drosophila cyclin mRNAs are differentially expressed during development. In their in situ hybridization experiments, no cyclin B mRNA was detected in somatic cells either during the cellular blastoderm stage or in imaginal discs of third-instar larvae. Reciprocally, abundant cyclin B mRNA but no cyclin A mRNA was found in male larval gonads (Whitfield et al., 1989). These observations suggested that A- and B-type cyclins have tissue-specific roles. Our results do not agree with this conclusion. While we see a higher concentration of cyclin B mRNA in germ cell precursors, low levels of cyclin B mRNA were readily detected in somatic cells at the cellular blastoderm stage using a sensitive whole-mount in situ hybridization protocol (Tautz and Pfeiffle, 1989). Furthermore, our antibodies have allowed a careful comparison of the distribution of cyclin A and cyclin B protein throughout Drosophila development. Our results indicate that the two proteins are coexpressed in proliferating cells during all stages. Cyclin B was clearly present in somatic cells of embryos at the blastoderm and postblastoderm stages (Figure 7A) and was found in discs. In addition, cyclin A was readily detected in gonads (S. DiNardo, personal communication). It appears that the discrepancies have two origins: first, the previous study failed to detect the lower levels of cyclin B mRNA present in somatic cells, and second, the differences in cyclin B mRNA levels are not associated with a comparable difference in levels of cyclin B protein.

Our in situ hybridization experiments, however, fully confirmed the finding that cyclin B mRNA but not cyclin A mRNA is considerably concentrated in the pole cells (Whitfield et al., 1989). In addition, our results demonstrate that a higher cyclin B mRNA concentration is found in a posterior cap long before the onset of nuclear migration and before substantial zygotic transcription. The maternal cyclin B mRNA in this posterior cap is segregated into the pole cells and subsequently localized into compact cytoplasmic granules. While we do not at present know the nature of these granules, to explain our failure to detect increased levels of cyclin B protein in pole cells, we suggest that the cyclin B mRNA sequestered in these granules might not be available for translation.

Cyclin A and Cyclin B Are Functionally Distinct

The independent evolutionary conservation of cyclin A and cyclin B strongly suggests that the two cyclin types have at least somewhat different functional roles. But do the cyclins play independent roles in the control of mitosis in each cell, or do they have special and unique roles at different times of development or in different tissues? The observed coexpression of the different cyclins suggest that they both have roles in every cell cycle. Indeed, analysis of the phenotype associated with null mutations in the cyclin A gene has demonstrated that cyclin A accumulation is required in epidermal cells to allow mitosis 16 (Lehner and O’Farrell, 1989). Moreover, the demonstration here that cyclin B accumulates in these cyclin A–deficient cells shows that cyclin B is not sufficient to allow entry into mitosis.

In contrast to our results, experiments with Xenopus egg extracts have indicated that the accumulation of a B-type cyclin is sufficient to trigger mitotic events in the absence of cyclin A (Murray and Kirschner, 1989). This apparent discrepancy might be explained in one of the following ways. Cell cycle regulation in the first embryonic cell cycles in frog embryos and consequently in these extracts might be special in that the requirement for cyclin A accumulation is bypassed at this stage. Indeed, at least two regulatory mechanisms (checkpoints; Hartwell and Weinert, 1989) are known to be bypassed during these specialized, early cell cycles: Entry into mitosis is not dependent on complete replication of chromosomal DNA, and these cycles continue even in the presence of microtubule poisons that block mitotic spindle assembly (Kimelman et al., 1987). Alternatively, cyclin B might have been overexpressed in the in vitro experiments, and over-expression of cyclin B might render entry into mitosis independent of cyclin A accumulation. Finally, the preparation of the extract might artificially produce the observed independence of cell cycle progression from cyclin A accumulation.

Several observations favor an important quantitative consideration. mRNA injection experiments with Xenopus oocytes have clearly shown that cyclin A and cyclin B act synergistically. A combination of clam cyclin A and cyclin B mRNA is over 20-fold more effective in inducing maturation than either mRNA alone (Westendorf et al., 1989). Moreover, in mutant Drosophila embryos deficient for cyclin A production, epidermal cells can progress through mitosis 15 normally despite a large quantitative reduction in cyclin A levels. Both these observations indicate that the dose requirement for cyclin A is extremely low in the presence of cyclin B. Therefore, residual, low levels of cyclin A accumulating in the in vitro extracts might in fact allow entry into mitosis in combination with cyclin B.

The Rate of Cyclin Accumulation Does Not Time Entry into Mitosis

In vitro experiments with Xenopus egg extracts indicated that the rate of cyclin accumulation could determine the time of entry into mitosis (Murray and Kirschner, 1989). However, the rate-limiting component of the mitotic trigger might be different in vivo or at different stages of development. Indeed, our observations demonstrate that neither cyclin A nor cyclin B levels govern the timing of entry into mitosis 14, the first asynchronous mitosis in Drosophila embryos (Lehner and O’Farrell, 1989; this report). Furthermore, analysis of the stg gene in Drosophila has strongly suggested that the stg gene product is rate limiting for entry into mitosis 14 (Edgar and O’Farrell, 1989). The stg gene product is required for entry into mitosis and encodes a protein homologous to the mitotic inducer encoded by the cdc25 gene in S. pombe (Russell and Nurse, 1986). Moreover, before mitosis 14, stg transcripts were found to accumulate in a pattern that anticipates the pattern of the subsequent mitosis, and recent experiments involving premature, ectopic expression of stg before mitosis 14 have demonstrated that the stg gene product is rate limiting for entry into mitosis (B. Edgar and P. H. O., unpublished data).

Roles of the Cyclins in Control of Mitosis

Our results allow us to conclude that cyclin A is essential for entry into mitosis 16 and that cyclin B expression at normal levels cannot substitute for cyclin A. Unfortunately, because we do not at present have a cyclin B mutant, we cannot demonstrate that cyclin B is essential. In Xenopus egg extracts, however, antisense oligonucleotide–mediated ablation of cyclin B mRNAs blocks mitotic events (Minshull et al., 1989). Similarly, disruption of the S. pombe cdc13 gene, which encodes a cyclin protein that is clearly more homologous to cyclin B than to cyclin A, leads to G2 arrest (Booher and Beach, 1988). These results demonstrate that cyclin B is required for mitosis in other species. We consequently suggest that both cyclin A and cyclin B play distinct and essential roles in the control of mitosis.

Subcellular Localization of Cyclin A and Cyclin B

Consistent with the proposed functional nonidentity of cyclin A and B, our immunofluorescent localization has revealed subtle differences in the subcellular distribution of cyclin A and cyclin B during both interphase and mitosis. During interphase, a fraction of cyclin B but not cyclin A appears to be associated with intranuclear dots. Further experiments will be required to rigorously demonstrate that the intranuclear labeling represents cyclin B. Interestingly, however, anti-cyclin B labeling in clam has been found not only in the cytoplasm but also in the nucleolus (see Westendorf et al., 1989), and in S. pombe the cdc13+ gene product has been reported to accumulate exclusively within the nucleus and restricted to a region that stains only very weakly with DNA stains (Booher et al., 1989). In S. pombe, cytoplasmic cdc13+ gene product might not have been detected for the same technical reasons that prevented immunofluorescent detection of the p34cdc2 kinase during the G1 phase (Booher et al., 1989). Therefore, cyclin B localization might in fact be more conserved in evolution than is currently apparent.

The majority of the p34cdc2 kinase appears to be located in the nucleus in both S. pombe and HeLa cells (Booher et al,, 1989; Riabowol et al., 1989). The differences in the intracellular distribution of the cyclins and the p34cdc2 kinase in higher eukaryotes raise the question of when and where these proteins associate. In this regard, the observed redistribution of cyclin proteins to the region of the condensing chromatin early in prophase is interesting. Since the p34cdc2 kinase is not activated until late in G2, this relocalization might reflect formation of a p34cdc2 kinase–cyclin complex.

During metaphase both cyclin A and cyclin B appear to be completely degraded, and as has been found in clam (Evans et al., 1983; Luca and Ruderman, 1989), cyclin A appears to be degraded more rapidly than cyclin B. As shown in our immunofluorescence experiments, cyclin A degradation is completed within metaphase and cyclin B disappears around the metaphase–anaphase transition. Clearly, however, our findings are based solely on comparisons of immunofluorescent signal intensities, and these may not reflect protein levels linearily. Nonetheless, our observations suggest that the temporal difference between cyclin A and cyclin B degradation has been conserved in evolution and is likely to be functionally relevant.

In conclusion, our results indicate that cyclin A and cyclin B have distinct and essential roles in the control of mitosis.

Experimental Procedures

Fly Strains

Wild-type flies were from the Sevelen strain. The characterization of the fly lines carrying mutations in the cyclin A gene has been described previously (Lehner and O’Farrell, 1989). The line used in this study (neo114) was generated by P element mutagenesis by Cooley et al. (1988).

PCR Experiments

The primers used in PCR experiments (Saiki et al., 1985) were designed to anneal to the cDNA sequences encoding the amino acids MRSILIDW and KYEEMPPE, respectively. The exact sequences were as follows: primer 1, GCAGGATCCATGCG(C/G/T)GCCAT(T/C)(T/C)(T/C)-T(C/G/T)AT(T/C)TGG; primer 2, TC(G/T)GG(G/T)GG(A/G)TA(G/T/C)-ATCTCCTC(A/G)TACTT. Ten micrograms of total genomic DNA, 1 ng of λ DNA from a genomic clone containing cyclin A gene sequences (kindly provided by H. Vässin and Y.-N. Jan), and 0.1 ng of a Bluescript plasmid with a full-length cyclin B cDNA insert or of a plasmid containing genomic cyclin B sequences (kindly provided by T. Jongens, B. Hay and Y.-N. Jan) were used as templates in 100 μl reactions containing 10 mM Tris–HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.01% (w/v) gelatin, 0.2 mM dATP dGTP, dCTP, and dTTP, 5 μM primer 1, 5 μM primer 2, and 2.5 U of Taq polymerase. After an initial 2 min at 94°C, amplification was allowed for 25 cycles (1 min at 94°C, 1 min at 55°C, and 3 min at 72°C).

cDNA Isolation and Sequencing

DNA amplified in PCR experiments was gel isolated, labeled by random priming (Hodgson and Fisk, 1987), and used to probe a λgt10 cDNA library made with poly(A)+ RNA from 3–12 hr embryos (Poole et al., 1985). Sixteen positive plaques were purified, and insert sizes were determined in PCR experiments using primers flanking the EcoRI cloning site in λgt10. The largest insert was cut out from the recombinant phage and cloned into M13 vectors (Norrander et al., 1983). The DNA sequence was determined using a Sequenase kit (United States Biochemical Corp.).

Northern Blots

Northern blots were probed with RNA probes synthesized with T7 polymerase from a linearized Bluescript plasmid containing the cyclin B cDNA as described by Stroeher et al. (1988). Nylon filters were hybridized for 14 hr at 58°C in 50% formamide, 5× SSC, 50 mM NaPO4 (pH 7.0), 5× Denhardt’s solution, 0.1% SDS, 200 (μg/ml salmon sperm DNA. The filters were washed twice for 5 min at room temperature in 2× SSC, 0.1% SDS, and four times at 65°C in 0.2× SSC, 0.1% SDS.

Immunological Methods

To obtain an immunogen for the production of anti-cyclin B antibodies, two primers were synthesized and used in PCR experiments for amplification of a part of the cyclin B cDNA (base pairs 872–2539) and for introduction of a BamHI site at the 5′ end and a HindIII site at the 3′ end. The amplified DNA was digested with BamHI and HindIII and cloned into the corresponding sites of the pAR3040 vector (Studier and Moffat, 1986). The fusion protein encoded by this construct was produced in E. coli and isolated by preparative gel electrophoresis. Mice were immunized intraperitoneally with 150 μg of fusion protein emulsified in Freund’s complete adjuvant and boosted twice with 100 μg of fusion protein in Freund’s incomplete adjuvant.

The antisera were tested by immunoprecipitation assays: mRNA was synthesized in vitro with T3 polymerase from a linearized Blue-script plasmid with a cyclin B or a cyclin A cDNA insert. The synthetic mRNA was subsequently translated in a reticulocyte lysate (Promega) in the presence of [36S]methionine to yield full-length cyclin A or cyclin B protein. Aliquots of the in vitro translation reactions were mixed and diluted 100-fold in RIPA buffer (Lehner et al., 1986), and 2 μl of anti-cyclin B antiserum or affinity-purified rabbit antibodies against Drosophila cyclin A (Lehner and O’Farrell, 1989) was added. Immune complexes were isolated as described previously (Lehner et al., 1986).

Immunoblotting and immunofluorescence experiments were performed as described previously (Lehner et al., 1986; Lehner and O’Farrell, 1989).

In Situ Hybridization Experiments

In situ hybridization to polytene chromosomes of third-instar larvae was done as described previously (Lehner and O’Farrell, 1989). The cyclin B gene was mapped to the chromosomal region 59A. The same chromosomal localization has been reported for the cyclin B gene identified by Whitfield et al. (1989).

In situ hybridization experiments to detect mRNA were done according to Tautz and Pfeiffle (1989) with minor modifications. Embryos were dechorionated, fixed, and devitellinized as described by Edgar and O’Farrell (1989). Subsequently, embryos were rinsed in methanol followed by ethanol. After a rinse in ethanol-xylene (1:1), embryos were soaked in xylene for several hours. Xylene was replaced by ethanol-xylene (1:1) followed by 100% ethanol. After several rinses in methanol, embryos were rinsed once in 50% methanol, 50% PBT (PBT: PBS containing 0.1% [w/v] Tween 20) and postfixed for 20 min in 5% formaldehyde in PBT. Fixative was removed by three rinses in PBT, and embryos were incubated for 4 min in PBT containing 50 μg/ml proteinase K. Protease digestion was stopped by washing twice for 2 min in PBT containing 2 mg/ml glycine. After two additional rinses in PBT, embryos were again postfixed for 20 min in 5% formaldehyde in PBT. Fixative was removed by five washes (2 min each) in PBT. Embryos were then incubated for 2 min in PBT-hybridization solution (1:1) followed by an additional 2 min in hybridization solution (50% formamide, 5× SSC, 100 μg/ml sonicated and boiled salmon sperm DNA, 100 μg/ml tRNA, 50 μg/ml heparin, 0.1% Tween 20). After prehybridization for 1 hr at 45°C, embryos were hybridized overnight at 45°C in hybridization solution containing 0.1 μg/ml heat-denatured probe.

Probes were prepared from cDNA inserts that had been digested with Alul. One hundred nanograms of DNA was combined with 0.1 mg of random primers (6-mers, Pharmacia), and H2O was added to 14 μl. The DNA was denatured by a 5 min incubation in boiling water followed by chilling in a mixture of salt water and ice. Two microliters of 10x Vogel buffer (1 M PIPES [pH 6.6], 50 mM MgCl2, 100 mM 2-mercaptoethanol), 2.0 μl of a deoxynucleotide mix containing digoxigenin-dUTP (Genius kit, Boehringer), and 5–10 U of Klenow fragment were added. After an overnight incubation at 14°C, an additional 5–10 U of Klenow fragment was added and the incubation was continued at room temperature for 4 hr. The reaction was stopped by adding EDTA to 50 mM and incubating at 65°C for 10 min. Eighty microliters of H2O, 50 μg of tRNA, and LiCl (300 mM final concentration) were added and the probe was precipitated with ethanol.

After hybridization, excess probe was removed by 20 min washes at 45°C in hybridization solution, in PBT-hybridization solution (1:1), and in PBT (five times). For probe detection, embryos were incubated for 1 hr at room temperature in PBT containing alkaline phosphatase-conjugated anti-digoxigenin antibodies (Genius kit, Boehringer) that had been preadsorbed previously with fixed embryos for several hours. Excess antibodies were removed by four washes in PBT (20 min, room temperature). Before the final color reaction, embryos were rinsed twice in 100 mM Tris (pH 9.5), 100 mM NaCl, 50 mM MgCI2, 1 mM levamisole (Sigma), 0.1% Tween 20. Color development was done in 1 ml of the same buffer containing 4.5 μl of NBT and 3.5 μl of X-phosphate as provided in the Genius kit (Boehringer). The color reaction was stopped by rinsing several times in PBT; after staining in Hoechst 33258 (1 μg/ml), embryos were mounted in 80% glycerol.

Acknowledgments

We are indebted to H. Vässin, T. Jongens, B. Hay, and Y.-N. Jan for providing genomic clones of the cyclin genes, communicating results prior to publication, and helpful discussions. We also would like to thank D. Lakich, B. Edgar, H. Richardson, and J.-P. Vincent for critical reading of the manuscript. This work was supported by a fellowship from the Swiss National Science Foundation (to C. F. L.) and by a National Science Foundation grant (to P. H. O.).

Footnotes

GenBank Accession Number

The accession number for the Drosophila cyclin B sequence is M33192.

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