![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Molecular and Cellular Biology, March 2006, p. 2456-2466, Vol. 26, No. 6
0270-7306/06/$08.00+0 doi:10.1128/MCB.26.6.2456-2466.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Department of Biology, Duke University, Durham, North Carolina,1 Department of Molecular Biology, The Scripps Research Institute, La Jolla, California2
Received 18 November 2004/ Returned for modification 13 December 2004/ Accepted 23 December 2005
 |
ABSTRACT |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
mutants but not in clb6 mutants. We have examined the basis for the differential functions of Clb5 and Clb6 and determined that unlike Clb5, which is stable until mitosis, Clb6 is degraded rapidly at the G1/S border. N-terminal deletions of CLB6 were hyperstabilized, suggesting that the sequences responsible for directing the destruction of Clb6 reside in the N terminus. Clb6 lacks the destruction box motif responsible for the anaphase promoting complex-mediated destruction of Clb5 but contains putative Cdc4 degron motifs in the N terminus. Clb6 was hyperstabilized in cdc34-3 and cdc4-3 mutants at restrictive temperatures and when S/T-P phosphorylation sites in the N terminus were mutated to nonphosphorylatable residues. Efficient degradation of Clb6 requires the activities of both Cdc28 and Pho85. Finally, hyperstabilized Clb6 expressed from the CLB6 promoter rescued the slow S-phase defect exhibited by clb5 cells. Taken together, these findings suggest that the SCFCdc4 ubiquitin ligase complex regulates Clb6 turnover and that the functional differences exhibited by Clb5 and Clb6 arise from the distinct mechanisms controlling their stability. |
INTRODUCTION |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Transition from G1 into S phase is triggered by the S-phase cyclins Clb5 and Clb6 (1, 8, 23, 35). Clb5 and Clb6 are approximately 40% identical at the amino acid sequence level and are transcribed in a pulse in G1 (8, 23, 35). Transcription is controlled by similar regulatory elements in the promoters of both CLB5 and CLB6 (28). Although CLB5 and CLB6 are transcribed in G1, Clb5/Cdc28 (and presumably Clb6/Cdc28) complexes are not activated until the G1/S border when the Clb-specific Cdk inhibitor, Sic1, is degraded by the action of the ubiquitin ligase complex, SCFCdc4 (Skp1/Cullin/F-box) (33, 34, 41, 42).
Clb5 and Clb6 appear to play redundant roles in the initiation of S phase. Either Clb5 or Clb6 can promote entry into S phase on schedule (1, 8, 35). However, Clb5 but not Clb6 is required for timely progression through S phase (8, 35). Further studies showed that clb5 cells could not efficiently trigger DNA replication from replication origins that fire late in S phase but that replication initiation from late-firing origins in clb6 cells was unperturbed (5).
Phenotypic differences between clb5 and clb6 mutants were also observed in the regulation of spindle pole body duplication (17) and DNA replication (S. B. Haase and S. I. Reed, unpublished data). Moreover, Clb6 but not Clb5 is required to regulate the nuclear localization of the transcription factor Swi6 (12).
The differential phenotypes observed for clb5 and clb6 mutants could be explained by differences in posttranscriptional regulation, protein function, or both regulation and function. Here, we report that the stability of Clb5 and Clb6 is differentially regulated. Clb5 has been shown to persist throughout S phase and is marked for destruction in mitosis by the action of the ubiquitin ligase complex APCCdc20(anaphase- promoting complex) (20, 21, 36). We present evidence that Clb6 is targeted for destruction at the G1/S border by the activity of a functionally distinct ubiquitin ligase complex, SCFCdc4. We also show that the differential regulation of Clb5 and Clb6 protein stability can account for the differences observed in the ability of these S-phase cyclins to promote timely progression through S phase.
|
MATERIALS AND METHODS |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
ns) (31) except where noted. Relevant genotypes of strains are described in Table 1. CLB6 and CLB5 were tagged at their respective genomic loci by PCR-mediated homologous recombination (oligonucleotides are listed in Table 2) using pFA6KANMX2 (43) as a template. Haploid CLB6-9MYC-KAN cells (SBY464) were mated to haploid CLB5-9MYC-KAN cells (SBY466). Diploids were then sporulated to obtain haploid cells bearing both the CLB5-9MYC and the CLB6-9MYC alleles (SBY468).
|
|
pCRII-CLB6-HA3-URA3 was constructed by cloning the URA3 open reading frame from pUCHURA3 into pCRII-CLB6-HA3 on HindIII ends. pUCHURA3 was constructed by PCR amplification of URA3 from yeast genomic DNA and ligation into the HindIII site of pUCH (18). PCR-mediated gene replacement was used to replace the CLB6 open reading frame with mutated CLB6 alleles. Oligonucleotides described in Table 2 were used to amplify CLB6-HA3-URA3 and its derivatives from pCRII-URA3 vectors. The resultant PCR products were integrated into yeast cells by homologous recombination. Mutations were verified by sequencing PCR fragments generated from genomic DNA. All gene deletions were made by PCR-mediated gene replacement using oligonucleotides listed in Table 2. The kanamycin resistance gene was amplified from pFA6KANMX2 with oligonucleotides complementary to sequence flanking the gene being replaced on their 3' ends. PCR products were then integrated into yeast cells by homologous recombination, and transformants were selected by screening for kanamycin resistance.
Cell growth and arrest conditions.
Yeast cultures were grown in YEP medium (1% yeast extract, 2% Bacto peptone, 0.001% adenine, 0.001% uracil) supplemented with 2% sugar (dextrose, sucrose, galactose, or raffinose). Cells were grown at 30°C except as noted. Temperature-sensitive mutants were grown at 25°C and shifted to 34°C for arrest, except where otherwise noted. Mating pheromone synchrony experiments were performed by adding alpha factor (
-factor) (2 µg/ml -factor for BAR1 strains and 40 to 60 ng/ml for bar1 strains) to the growth medium for 2 h. Synchronous populations were released from the arrest into medium without -factor.
Protein stability assays. GAL1-driven expression of all CLB6 alleles was induced by adding 10% galactose solution to mid-log-phase cultures in YEP-sucrose or YEP-raffinose to a final concentration of 2% galactose or by harvesting cells by centrifugation and resuspending them in YEP-galactose (except as noted). Galactose concentrations and pulse durations were adjusted as necessary to achieve protein expression levels similar to that of endogenous Clb6 at the G1/S border. In order to terminate transcription and translation, cells were harvested and resuspended in equal volumes of YEP-dextrose medium containing 1 mg/ml cycloheximide at t = 0 min. Proteins were extracted from whole-cell lysates, and results were analyzed by Western blotting as described previously (17).
Phosphatase assay. Clb6-HA expression was induced in YEP-galactose medium for 1.5 h. Whole-cell lysates were prepared as described above and then treated with phosphatase. Phosphatase reaction mixtures were assembled as recommended by the vendor and either left on ice for 45 min or incubated at 30°C for 45 min. Where indicated, 400 U of lambda phosphatase was used per reaction, and 10 mM sodium orthovanadate and 50 mM sodium fluoride were used to inhibit phosphatase activity.
Flow cytometry. Flow cytometric analysis of DNA content was performed as described previously (16). The percentage of cells in S phase was determined by first fitting normal curves to the 1C and 2C peaks and integrating under these curves to quantitate the percentage of cells in G1 and G2/M phases. The remaining cells were judged to be in S phase.
|
RESULTS |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and clb6 mutants exhibit distinct phenotypes, we began by examining the expression of the two proteins in a synchronous population of cells as they traversed the cell cycle.
Both Clb5 and Clb6 were fused at the C terminus with a nine-myc tag at their respective genomic loci. CLB5-9MYC CLB6-9MYC cells were synchronized in G1 by treatment with the mating pheromone -factor and then released into the cell cycle. Clb5 and Clb6 protein levels were then monitored at several time points by Western blotting (Fig. 1A). Cell cycle progression was assessed by monitoring budding index and DNA content (Fig. 1B).
|
-factor arrest and persisted until 60 min after release, when cells entered mitosis (Fig. 1A and B). Clb6 also appeared in G1 but disappeared rapidly between 20 min and 30 min after release (Fig. 1A). Analysis of budding index and DNA content (Fig. 1B) indicates that Clb6 disappears near the G1/S transition. Because CLB5 and CLB6 are transcribed simultaneously in G1, these observations suggest that the turnover of Clb5 and Clb6 is regulated differently. The N terminus of Clb6 confers instability on the protein. Clb5 and Clb6 are closely related at the protein sequence level. The proteins are approximately 40% identical throughout their lengths but show the highest degree of homology and identity in their C-terminal regions (Fig. 2A). The degron sequence motif recognized by the APCCdc20 (RRALTDVPVN) has been mapped to the N-terminal region of Clb5 (20, 21, 36). Although Clb6 and Clb5 share some homology in this region, a critical amino acid change in the degron sequence (RXXL to KXXL) (Fig. 2A) is likely to prevent recognition of Clb6 by the APC. Since degradation signals for Clb5 are located in the N terminus and Clb5 and Clb6 protein sequences are the most divergent in this region, we reasoned that degradation signals specific for Clb6 might also reside in its N-terminal region.
|
Wild-type Clb6 appears to be very unstable in asynchronous cells, exhibiting a half-life of less than 5 min. The Clb641 deletion is significantly stabilized compared to wild-type Clb6, indicating that sequences important for degradation are contained in the first 41 amino acids. However, Clb6100 and Clb6135 appear to be more stable than Clb641, suggesting that additional degron sequences exist downstream. We have shown that expression of Clb6100 is capable of promoting S phase in clb1,2,3,4,5,6 cells, indicating that Clb6100 can still bind Cdc28 and activate DNA replication (data not shown) and suggesting that hyperstabilization is not the indirect result of a failure to form normal cyclin/Cdk complexes.
Clb6 is hyperstabilized in SCF mutants and is ubiquitinated. Examination of the N terminus of Clb6 revealed sequences similar to SCFCdc4 phospho-degron motifs found in the Cdk inhibitor Sic1 (29), including three potential Cdk phosphorylation sites (no SP or TP sequences are found in Clb5). Moreover, like Sic1 (34), Clb6 is destroyed at the G1/S transition. Taken together, these observations suggest that SCF complexes may regulate Clb6 stability. Therefore, we tested the stability of Clb6 in cells bearing temperature-sensitive mutations in SCF components.
Using the GAL1 promoter shutoff protocol described in the legend to Fig. 2, we found that Clb6 is significantly stabilized in cdc34-3 sic1 and cdc4-3 sic1 cells at restrictive temperatures, compared to that in wild-type cells at the same temperature (Fig. 3A and B). These findings suggest that SCFCdc4 complexes regulate the turnover of Clb6.
|
) in otherwise wild-type cells does not affect the stability of Clb6 (Fig. 3C), indicating that the hyperstabilization of Clb6 in cdc4-3 sic1 and cdc34-3 sic1 mutants is not an indirect effect of the loss of Sic1.
Clb6 does not appear to be fully stabilized in cdc34-3 sic1 mutants (compare Fig. 3B to Fig. 2B), which might suggest that other mechanisms are required for efficient Clb6 degradation. However, we have observed that cdc34-3 SIC1 cells can enter S phase (data not shown), suggesting that the cdc34-3 allele is partially functional even at 35°C. Furthermore, we have determined that Clb6 is significantly less stable at higher temperatures, even in wild-type cells (see Fig. S1 in the supplemental material). Thus, the general instability of Clb6 at high temperatures is likely to offset the stabilizing effects of the SCFts mutants. Consistent with these observations, the stability of Clb6 in cdc4-3 sic1 cells at 33°C (a partially restrictive temperature for the cdc4-3 allele) is comparable to that in the N-terminal deletions (compare Fig. 3A to Fig. 2B).
If Clb6 is a direct target of SCFcdc4, then it should be ubiquitinated and destroyed by the proteasome. Thus, inhibition of the proteasome should stabilize Clb6. Using the GAL1 promoter shutoff protocol described above, we found that HA-tagged Clb6 is stabilized in cells treated with the proteasome inhibitor MG132 (Fig. 3D). Ubiquitinated proteins are highly unstable and normally appear as a "ladder" of slow-mobility species on sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels. A prominent ladder of slower-mobility Clb6 species, indicative of ubiquitination, was observed on long exposures of Western blots in extracts from cells treated with MG132 (Fig. 3E). Moreover, the intensity of slower-mobility bands was also increased in extracts that were treated with the deubiquitinase inhibitor N-ethylmaleimide (NEM). Slower-mobility forms were not observed in untreated cells (Fig. 3E).
Clb6 is a phospho-protein, and mutation of putative phosphorylation sites leads to hyperstabilization. The SCF complex normally recognizes phosphorylated substrates (37, 44). Thus, we first determined whether Clb6 is a phospho-protein. Slower-mobility forms of Clb6 were observed under a number of conditions (see Fig. 2 and 3). In order to determine whether the slower-mobility forms of Clb6 represented phosphorylated species, we treated cell lysates with lambda phosphatase. Slow-mobility forms of Clb6 disappeared in lysates treated with phosphatase but remained in untreated lysates and in lysates containing phosphatase and phosphatase inhibitors (Fig. 4A).
|
3P) was the most stable, exhibiting a half-life comparable to that of the N-terminal deletions (Fig. 4B and C). These results suggest that all three putative phosphorylation sites contribute to the instability of Clb6. SCF-regulated proteins, such as Sic1 in budding yeast (29) and cyclin E in human cells (39, 46), exhibit similar behaviors when putative phosphorylation sites are mutated. In both cases, mutation of a single phosphorylation site has some marginal stabilizing effect, but mutation of multiple sites is required for maximum stabilization. Both Cdc28 and Pho85 contribute to the destruction of Clb6. Many SCF targets in both yeast and mammalian cells are phosphorylated by cyclin/Cdk complexes (19, 24, 37, 44, 46) and are hyperstabilized when cyclin/Cdk activities are inhibited (46). Therefore, we asked whether inhibition of cyclin/Cdk activity would stabilize Clb6. Using the GAL1 promoter shutoff protocol described above, we examined Clb6 stability in cells bearing a cdc28as-1 allele that can be inhibited by the ATP analogue 1-NM-PP1 (Fig. 5A) (2).
|
3P was significantly more stable than wild-type Clb6 in cdc28as-1 cells treated with 1-NM-PP1 (data not shown), confirming that these putative phosphorylation sites are important for the degradation of Clb6. Taken together, these results suggest that an additional kinase is required for efficient degradation of Clb6.
Efficient degradation of the SCFcdc4 target, Sic1, requires the combined action of Cdc28 and Pho85 kinases (30). We therefore examined the stability of Clb6 in cells with the PHO85 gene deleted, both alone (Fig. 5B) and in combination with the cdc28as-1 allele. Again, Clb6 was only partially stabilized in pho85 cells (Fig. 5B) and in pho85 cdc28as-1 double mutants in the absence of 1-NM-PP1 (Fig. 5C). However, Clb6 was significantly hyperstabilized in pho85 cdc28as-1 double mutants in the presence of 1-NM-PP1 (Fig. 5C). These results suggest that Pho85 and Cdc28 have redundant functions in the regulation of Clb6 stability.
Clb63P expressed from the endogenous CLB6 promoter is not destroyed at the G1/S border and suppresses the S-phase defect of clb5 cells.
Having established that mutation of putative phosphorylation sites stabilizes Clb6, we examined whether Clb63P was destroyed at the G1/S border. We replaced the endogenous wild-type copy of CLB6 with an HA-tagged CLB63P allele in a strain with the CLB5 gene deleted (clb5). clb5 CLB63P cells were arrested in G1 with -factor. The synchronized cell population was then released into the cell cycle and sampled at various time points after release for the analysis of both Clb6 protein levels and DNA content (Fig. 6B and C). Results were compared to those of clb5 cells bearing a wild-type allele of CLB6 tagged with HA (Fig. 6A and C). In the clb5 CLB6-HA control strain, Clb6 protein appeared in late G1, disappeared at the G1/S border, and reappeared in G1 of the next cell cycle (Fig. 6A). Clb63P also appeared in late G1, but Clb63P levels persisted beyond the G1/S border, remaining high throughout S phase (Fig. 6B). Thus, mutation of the phosphorylation sites bypassed the controls that normally trigger the destruction of Clb6 at the G1/S border.
|
3P persisted throughout the time course, protein levels dropped somewhat around the time of mitosis (Fig. 6B). From this experiment, we could not determine whether the decrease in Clb63P levels was cell cycle regulated (e.g., an APC-mediated event). Therefore, we performed a similar G1 arrest/release experiment but released clb5 CLB63P cells into medium containing nocodazole in order to arrest cells before anaphase and prevent any APC-mediated destruction of Clb63P. We found that Clb63P levels declined in a continuous manner in nocodazole-arrested cells (see Fig. S2 in the supplemental material), suggesting that the decrease in Clb63P levels observed in cycling cells (Fig. 6B) is not due to a regulated mitotic destruction mechanism.
As mentioned above, cells with the CLB5 gene deleted normally progress slowly through S phase compared to wild-type or clb6 cells. Because Clb63P persisted throughout S phase, we were able to determine whether the inability of Clb6 to promote normal S-phase kinetics on its own was related to its destruction at the G1/S transition. S-phase kinetics of clb5 cells bearing wild-type CLB6 or the CLB63P allele in the synchrony/release experiments shown in Fig. 6A and B were analyzed by flow cytometry (Fig. 6C).
A synchronous population of clb5 cells bearing wild-type CLB6 moved slowly through S phase (Fig. 6C). Cells entered S phase on schedule (coincident with bud emergence), approximately 20 min after release from the -factor arrest. Cells then progressed through the middle of S phase slowly, with DNA contents intermediate between 1C and 2C at the 25-min and 30-min time points. The majority of the population reached 2C content of DNA between 35 min and 40 min.
clb5 CLB63P cells entered S phase with similar kinetics, indicating that the phosphorylation site mutation is competent for initiating S phase on schedule. However, the clb5 CLB63P mutants completed S phase significantly faster than clb5 CLB6 cells. Though clb5 CLB63P cells appeared to release from the G1 arrest somewhat less synchronously than the clb5 CLB6 cells, the bulk of the population had nearly completed S phase by 30 min. Furthermore, a significant fraction of clb5 CLB63P cells completed mitosis and reached G1 (1C DNA content and unbudded) again at 60 min, while very few clb5 CLB6 cells reappeared in G1 until 75 min. These results suggest that the inability of Clb6 to promote normal S-phase kinetics in the absence of Clb5 is due to its destruction at the G1/S border.
When Clb63P was expressed from the CLB6 promoter in wild-type cells, we observed no increase in the speed of S phase (data not shown). Thus, the rescue of the slow S-phase defect observed in clb5 mutants is likely to be a suppression of the defect in the clb5 mutant rather than a general increase in S-phase velocity.
We also examined S-phase kinetics in asynchronous cell populations by flow cytometric analysis of DNA content (Fig. 6D). In all strains shown, Clb6 and Clb6 mutants are epitope tagged with HA. As shown previously (8, 23, 35), a significant increase in the population of S-phase cells is observed in clb5 cells compared to that in wild-type cells, indicative of a slow S phase. The cell cycle profile of an asynchronous population of clb5 CLB63P cells (13.6% S-phase cells) looks similar to that of the wild-type cells (12.6% S-phase cells) and has noticeably fewer cells in S phase than clb5 CLB6 cells (22.9% S-phase cells). These results are consistent with the finding in synchronous populations (Fig. 6C) that Clb63P can suppress the slow S-phase defect of clb5 cells. Furthermore, asynchronous wild-type cells expressing Clb63P exhibit a cell cycle distribution (12.4% S-phase) similar to that of cells expressing wild-type CLB6 (Fig. 6D), indicating that Clb63P does not accelerate cells through S phase in wild-type cells.
The decrease in the S-phase fraction observed for asynchronous clb5 CLB63P cells compared to that for clb5 CLB6 cells (Fig. 6D) does not necessarily reflect a more rapid progression through S phase. A similar cell cycle profile could result from extending the time required to traverse both G1 and G2/M phases. However, no evidence for slow progression through G1 or S phase was observed in synchronized clb5 CLB63P cell populations (Fig. 6C). In fact, clb5 CLB63P cells entered S phase at the same time and returned to G1 and S phase on the second cycle more rapidly than did clb5 CLB6 cells (Fig. 6C), indicating no delay in progression through G1 or G2/M.
|
DISCUSSION |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and clb6 mutants are related to differential posttranscriptional regulation of the two proteins. Our findings indicate that unlike Clb5, which persists until anaphase, Clb6 is destroyed at the G1/S border via N-terminal motifs by the SCFCdc4 complex. Three putative Cdk phosphorylation sites in Clb6 contribute to its instability, and both Cdc28 and the related kinase Pho85 promote the degradation of Clb6. The observation that Clb6 is not quite as stable in pho85 cdc28as-1 double mutants (see Fig. 5C) as in the triple phosphorylation site mutant (see Fig. 5C and Fig. 4B and C) indicates that an additional kinase(s) may play a minor redundant role in the phosphorylation of Clb6. The differential regulation of Clb5 and Clb6 stability appears to have arisen from the acquisition of SCF degron sequences in Clb6, as well as a mutation at a critical residue in the destruction box (D-box) (RXXL to KXXL). However, in Drosophila melanogaster, the Pim protein apparently has a functional D-box with a KXXL sequence (25), suggesting that the D-box in Clb6 could still be functional. Furthermore, the stability of the S-phase cyclin in Schizosaccharomyces pombe, Cig2, appears to be controlled collaboratively by the SCF and APC ligase complexes (47). We found that Clb6 was not measurably stabilized in cells treated with nocodazole or in cells bearing a temperature-sensitive allele of APC2 (see Fig. S2 in the supplemental material). Taken together, these findings suggest that the APC plays a very minor role, if any, in regulating the stability of Clb6.
Differential regulation of S-phase cyclins and the progression of S phase. Entry into S phase is triggered by the destruction of Clb-specific Cdk inhibitor Sic1 at the G1/S border (29, 33, 41, 42). Our findings indicate that Clb6 is also destroyed very close to the G1/S border. Because Clb6 can promote timely S-phase entry in the absence of Clb5, active Clb6/Cdc28 complexes must be present at G1/S, but such complexes are likely to exist for only a brief period.
Although Clb6-associated kinase is present for only a very short period at the G1/S border, the conservation of Clb6 among the Saccharomyces species (see below) suggests an important function. Clb6 has been shown to be involved in the regulation of Swi6 localization (12), although this regulation does not appear to be critical for cell cycle progression. It has also been reported that clb6 mutants actually grow and divide faster than wild-type cells (1), which would suggest that cells bearing a wild-type allele of CLB6 should be at a selective disadvantage. For now, the conserved function of Clb6 remains a mystery.
Although either Clb5 or Clb6 can trigger the timely entry into S phase, Clb6 cannot promote normal progression through S phase without Clb5. The basis for the S-phase defect in clb5 cells is not yet fully understood. Nonetheless, a defect in late origin firing in clb5 cells has been reported (5) and could account for the significant slowing of DNA replication in clb5 mutants. The fact that Clb6 is normally destroyed at the G1/S border suggests that this defect is simply due to the lack of Clb6 availability late in S phase. Consistent with this hypothesis, Clb/Cdc28-dependent Cdc45 loading at late-firing origins occurs late in S phase (48, 49). Moreover, overexpression of Clb6 both promotes normal S-phase kinetics and restores the firing of late origins in clb5 mutants (13), suggesting that slow progression of cells through S phase in these cells is in fact linked to the defect in late origin firing.
Like Clb5 and Clb6, the related mitotic cyclins Clb1 and Clb2 also appear to have evolved divergent functions. Clb2 but not Clb1 is required for efficient mitotic progression in vegetative cells (10, 31), and Clb1 is essential for meiotic progression while Clb2 is dispensable (14). The basis for the functional diversity in this case appears to be differential transcription during mitosis and meiosis (14). It is interesting that both Clb1/Clb2 and Clb5/Clb6 diverged with respect to regulation of protein expression.
Gene deletion studies indicate a significant functional redundancy between the members of the B-cyclin family. In an extreme example, the essential functions of all six B-type cyclins in a sextuple deletion strain can be rescued by expressing Clb1 constitutively (15), arguing that differential expression patterns are the major distinguishing feature of B-cyclin functions. However, biochemical studies indicate that cyclin/Cdk complexes can phosphorylate a broad spectrum of Cdk targets but that distinct complexes vary in their intrinsic activities and substrate affinities (26). Thus, the specialization of cyclins for diverse cell cycle functions may reflect a suite of evolutionary changes that affect cyclin expression as well as localization, activity, and substrate affinity.
Why are Clb5 and Clb6 differentially regulated? Our observations suggest that Clb5 and Clb6 are functionally similar. Why then is it important for Clb6 to be destroyed at the G1/S border while Clb5 persists until mitosis?
The destruction of Clb6 at the G1/S transition may be conserved through the evolution of Saccharomyces species. Protein sequence alignments with Clb6 orthologs from other Saccharomyces species (S. bayanus, S. mikatae, and S. paradoxus) indicate that all three putative Cdk phosphorylation sites and the sequences surrounding them are highly conserved (Fig. 7A) (3, 22). Clb5 orthologs are highly conserved in the region of the D-box degron sequences that mediates destruction by the APC. These observations suggest that the distinct regulatory processes for Clb5 and Clb6 are evolutionarily conserved.
|
It has been reported that Clb6 inhibits progression through G2 (1). Thus, destruction of Clb6 at the G1/S border may be required for timely progression through G2. However, we have shown that expression of the CLB63P mutant from the endogenous CLB6 promoter in clb5 cells (Fig. 6C and D) or wild-type cells (data not shown) causes no measurable change in cell division time or defects in cell cycle distribution (Fig. 6D) as measured by flow cytometry. It is likely that Clb6 levels in the constitutive overexpression experiment reported by Basco et al. are significantly higher than the levels of Clb63P expressed from the CLB6 promoter (which are similar to wild-type levels) and that the significant G2 delay they observed is a result of elevated Clb6 levels rather than the persistence of Clb6 late in the cell cycle. Furthermore, Basco et al. favored a model in which Clb6 exerts its inhibitory effect on G2, at least in part, via regulation of CDC6 transcription (1). Because CDC6 is transcribed in G1, our finding that Clb6 is destroyed at the G1/S border is not necessarily inconsistent with the premise that Clb6 influences progression through G2 by affecting CDC6 transcription. However, our findings do indicate that destruction of Clb6 at the G1/S border is not likely to be necessary for maintaining normal cell cycle kinetics through G2/M.
Clues to the importance of destroying Clb6 at the G1/S border may be found in studies of the mammalian S-phase cyclin cyclin E. In many respects, budding yeast Clb6 and mammalian cyclin E are strikingly similar. Like Clb6, cyclin E is first expressed in G1 and then destroyed soon after cells traverse the G1/S transition (7). Cyclin E contains two consensus Cdk sites, and its stability is also regulated by the SCFhCdc4 complex (39).
Improper regulation of cyclin E expression and turnover appears to be associated with tumorigenesis and genome instability. Mutations in hCdc4 in a breast cancer cell line have been correlated with high expression levels of cyclin E (39). In fact, cyclin E levels are elevated in many tumor types (6, 32), and deregulated cyclin E expression has been shown to promote chromosome instability (38). However, the mechanism for the induction of chromosome instability by deregulated expression of cyclin E has not yet been fully elucidated.
Reminiscent of Clb5, mammalian cyclin A is expressed at the G1/S border (9) and persists until mitosis when it is degraded by the APC (11). Thus, there appear to be parallel regulatory mechanisms controlling the persistence of S-phase cyclins (Clb5 
cyclin A and Clb6 cyclin E) in both yeast and mammalian cells. The biological significance of the differential regulatory mechanisms for cyclin E and cyclin A has yet to be determined.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
Supplemental material for this article may be found at http://mcb.asm.org/.
|
REFERENCES |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
2. Bishop, A. C., J. A. Ubersax, D. T. Petsch, D. P. Matheos, N. S. Gray, J. Blethrow, E. Shimizu, J. Z. Tsien, P. G. Schultz, M. D. Rose, J. L. Wood, D. O. Morgan, and K. M. Shokat. 2000. A chemical switch for inhibitor-sensitive alleles of any protein kinase. Nature 407:395-401.[CrossRef][Medline]
3. Cliften, P., P. Sudarsanam, A. Desikan, L. Fulton, B. Fulton, J. Majors, R. Waterston, B. A. Cohen, and M. Johnston. 2003. Finding functional features in Saccharomyces genomes by phylogenetic footprinting. Science 301:71-76.
4. Dietrich, F. S., S. Voegeli, S. Brachat, A. Lerch, K. Gates, S. Steiner, C. Mohr, R. Pohlmann, P. Luedi, S. Choi, R. A. Wing, A. Flavier, T. D. Gaffney, and P. Philippsen. 2004. The Ashbya gossypii genome as a tool for mapping the ancient Saccharomyces cerevisiae genome. Science 304:304-307.
5. Donaldson, A. D., M. K. Raghuraman, K. L. Friedman, F. R. Cross, B. J. Brewer, and W. L. Fangman. 1998. CLB5-dependent activation of late replication origins in S. cerevisiae. Mol. Cell 2:173-182.[CrossRef][Medline]
6. Donnellan, R., and R. Chetty. 1999. Cyclin E in human cancer. FASEB J. 13:773-780.
7. Ekholm, S. V., and S. I. Reed. 2000. Regulation of G(1) cyclin-dependent kinases in the mammalian cell cycle. Curr. Opin. Cell Biol. 12:676-684.[CrossRef][Medline]
8. Epstein, C. B., and F. R. Cross. 1992. CLB5: a novel B cyclin from budding yeast with a role in S phase. Genes Dev. 6:1695-1706.
9. Erlandsson, F., C. Linnman, S. Ekholm, E. Bengtsson, and A. Zetterberg. 2000. A detailed analysis of cyclin A accumulation at the G(1)/S border in normal and transformed cells. Exp. Cell Res. 259:86-95.[CrossRef][Medline]
10. Fitch, I., C. Dahmann, U. Surana, A. Amon, K. Nasmyth, L. Goetsch, B. Byers, and B. Futcher. 1992. Characterization of four B-type cyclin genes of the budding yeast Saccharomyces cerevisiae. Mol. Biol. Cell 3:805-818.[Abstract]
11. Geley, S., E. Kramer, C. Gieffers, J. Gannon, J. M. Peters, and T. Hunt. 2001. Anaphase-promoting complex/cyclosome-dependent proteolysis of human cyclin A starts at the beginning of mitosis and is not subject to the spindle assembly checkpoint. J. Cell Biol. 153:137-148.
12. Geymonat, M., A. Spanos, G. P. Wells, S. J. Smerdon, and S. G. Sedgwick. 2004. Clb6/Cdc28 and Cdc14 regulate phosphorylation status and cellular localization of Swi6. Mol. Cell. Biol. 24:2277-2285.
13. Gibson, D. G., J. G. Aparicio, F. Hu, and O. M. Aparicio. 2004. Diminished S-phase cyclin-dependent kinase function elicits vital Rad53-dependent checkpoint responses in Saccharomyces cerevisiae. Mol. Cell. Biol. 24:10208-10222.
14. Grandin, N., and S. I. Reed. 1993. Differential function and expression of Saccharomyces cerevisiae B-type cyclins in mitosis and meiosis. Mol. Cell. Biol. 13:2113-2125.
15. Haase, S. B., and S. I. Reed. 1999. Evidence that a free-running oscillator drives G1 events in the budding yeast cell cycle. Nature 401:394-397.[CrossRef][Medline]
16. Haase, S. B., and S. I. Reed. 2002. Improved flow cytometric analysis of the budding yeast cell cycle. Cell Cycle 1:132-136.[Medline]
17. Haase, S. B., M. Winey, and S. I. Reed. 2001. Multi-step control of spindle pole body duplication by cyclin-dependent kinase. Nat. Cell Biol. 3:38-42.[CrossRef][Medline]
18. Heinzel, S. S., P. J. Krysan, M. P. Calos, and R. B. DuBridge. 1988. Use of simian virus 40 replication to amplify Epstein-Barr virus shuttle vectors in human cells. J. Virol. 62:3738-3746.
19. Henchoz, S., Y. Chi, B. Catarin, I. Herskowitz, R. J. Deshaies, and M. Peter. 1997. Phosphorylation- and ubiquitin-dependent degradation of the cyclin-dependent kinase inhibitor Far1p in budding yeast. Genes Dev. 11:3046-3060.
20. Irniger, S., and K. Nasmyth. 1997. The anaphase-promoting complex is required in G1 arrested yeast cells to inhibit B-type cyclin accumulation and to prevent uncontrolled entry into S-phase. J. Cell Sci. 110:1523-1531.[Abstract]
21. Jacobson, M. D., S. Gray, M. Yuste-Rojas, and F. R. Cross. 2000. Testing cyclin specificity in the exit from mitosis. Mol. Cell. Biol. 20:4483-4493.
22. Kellis, M., N. Patterson, M. Endrizzi, B. Birren, and E. S. Lander. 2003. Sequencing and comparison of yeast species to identify genes and regulatory elements. Nature 423:241-254.[CrossRef][Medline]
23. Kuhne, C., and P. Linder. 1993. A new pair of B-type cyclins from Saccharomyces cerevisiae that function early in the cell cycle. EMBO J. 12:3437-3447.[Medline]
24. Lanker, S., M. H. Valdivieso, and C. Wittenberg. 1996. Rapid degradation of the G1 cyclin Cln2 induced by CDK-dependent phosphorylation. Science 271:1597-1601.[Abstract]
25. Leismann, O., A. Herzig, S. Heidmann, and C. F. Lehner. 2000. Degradation of Drosophila PIM regulates sister chromatid separation during mitosis. Genes Dev. 14:2192-2205.
26. Loog, M., and D. O. Morgan. 2005. Cyclin specificity in the phosphorylation of cyclin-dependent kinase substrates. Nature 434:104-108.[CrossRef][Medline]
27. Mathias, N., S. L. Johnson, M. Winey, A. E. Adams, L. Goetsch, J. R. Pringle, B. Byers, and M. G. Goebl. 1996. Cdc53p acts in concert with Cdc4p and Cdc34p to control the G1-to-S-phase transition and identifies a conserved family of proteins. Mol. Cell. Biol. 16:6634-6643.[Abstract]
28. Moll, T., E. Schwob, C. Koch, A. Moore, H. Auer, and K. Nasmyth. 1993. Transcription factors important for starting the cell cycle in yeast. Philos. Trans. R. Soc. Lond. B Biol. Sci. 340:351-360.[Medline]
29. Nash, P., X. Tang, S. Orlicky, Q. Chen, F. B. Gertler, M. D. Mendenhall, F. Sicheri, T. Pawson, and M. Tyers. 2001. Multisite phosphorylation of a CDK inhibitor sets a threshold for the onset of DNA replication. Nature 414:514-521.[CrossRef][Medline]
30. Nishizawa, M., M. Kawasumi, M. Fujino, and A. Toh-e. 1998. Phosphorylation of Sic1, a cyclin-dependent kinase (Cdk) inhibitor, by Cdk including Pho85 kinase is required for its prompt degradation. Mol. Biol. Cell 9:2393-2405.
31. Richardson, H., D. J. Lew, M. Henze, K. Sugimoto, and S. I. Reed. 1992. Cyclin-B homologs in Saccharomyces cerevisiae function in S phase and in G2. Genes Dev. 6:2021-2034.
32. Sandhu, C., and J. Slingerland. 2000. Deregulation of the cell cycle in cancer. Cancer Detect. Prev. 24:107-118.[Medline]
33. Schneider, B. L., Q. H. Yang, and A. B. Futcher. 1996. Linkage of replication to start by the Cdk inhibitor Sic1. Science 272:560-562.[Abstract]
34. Schwob, E., T. Bohm, M. D. Mendenhall, and K. Nasmyth. 1994. The B-type cyclin kinase inhibitor p40SIC1 controls the G1 to S transition in S. cerevisiae. Cell 79:233-244.[CrossRef][Medline]
35. Schwob, E., and K. Nasmyth. 1993. CLB5 and CLB6, a new pair of B cyclins involved in DNA replication in Saccharomyces cerevisiae. Genes Dev. 7:1160-1175.
36. Shirayama, M., A. Toth, M. Galova, and K. Nasmyth. 1999. APC(Cdc20) promotes exit from mitosis by destroying the anaphase inhibitor Pds1 and cyclin Clb5. Nature 402:203-207.[CrossRef][Medline]
37. Skowyra, D., K. L. Craig, M. Tyers, S. J. Elledge, and J. W. Harper. 1997. F-box proteins are receptors that recruit phosphorylated substrates to the SCF ubiquitin-ligase complex. Cell 91:209-219.[CrossRef][Medline]
38. Spruck, C. H., K. A. Won, and S. I. Reed. 1999. Deregulated cyclin E induces chromosome instability. Nature 401:297-300.[CrossRef][Medline]
39. Strohmaier, H., C. H. Spruck, P. Kaiser, K. A. Won, O. Sangfelt, and S. I. Reed. 2001. Human F-box protein hCdc4 targets cyclin E for proteolysis and is mutated in a breast cancer cell line. Nature 413:316-322.[CrossRef][Medline]
40. Stueland, C. S., D. J. Lew, and S. I. Reed. 1993. Full activation of p34CDC28 histone H1 kinase activity is unable to promote entry into mitosis in checkpoint-arrested cells of the yeast Saccharomyces cerevisiae. Mol. Cell. Biol. 13:3744-3755.
41. Verma, R., R. S. Annan, M. J. Huddleston, S. A. Carr, G. Reynard, and R. J. Deshaies. 1997. Phosphorylation of Sic1p by G1 Cdk required for its degradation and entry into S phase. Science 278:455-460.
42. Verma, R., R. M. Feldman, and R. J. Deshaies. 1997. SIC1 is ubiquitinated in vitro by a pathway that requires CDC4, CDC34, and cyclin/CDK activities. Mol. Biol. Cell 8:1427-1437.[Abstract]
43. Wach, A., A. Brachat, R. Pohlmann, and P. Philippsen. 1994. New heterologous modules for classical or PCR-based gene disruptions in Saccharomyces cerevisiae. Yeast 10:1793-1808.[CrossRef][Medline]
44. Willems, A. R., T. Goh, L. Taylor, I. Chernushevich, A. Shevchenko, and M. Tyers. 1999. SCF ubiquitin protein ligases and phosphorylation-dependent proteolysis. Philos. Trans. R. Soc. Lond. B Biol. Sci. 354:1533-1550.[CrossRef][Medline]
45. Wolfe, K. H., and D. C. Shields. 1997. Molecular evidence for an ancient duplication of the entire yeast genome. Nature 387:708-713.[CrossRef][Medline]
46. Won, K. A., and S. I. Reed. 1996. Activation of cyclin E/CDK2 is coupled to site-specific autophosphorylation and ubiquitin-dependent degradation of cyclin E. EMBO J. 15:4182-4193.[Medline]
47. Yamano, H., K. Kominami, C. Harrison, K. Kitamura, S. Katayama, S. Dhut, T. Hunt, and T. Toda. 2004. Requirement of the SCFPop1/Pop2 ubiquitin ligase for degradation of the fission yeast S phase cyclin Cig2. J. Biol. Chem. 279:18974-18980.
48. Zou, L., and B. Stillman. 2000. Assembly of a complex containing Cdc45p, replication protein A, and Mcm2p at replication origins controlled by S-phase cyclin-dependent kinases and Cdc7p-Dbf4p kinase. Mol. Cell. Biol. 20:3086-3096.
49. Zou, L., and B. Stillman. 1998. Formation of a preinitiation complex by S-phase cyclin CDK-dependent loading of Cdc45p onto chromatin. Science 280:593-596.
This article has been cited by other articles:
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
