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Vol. 18, Issue 12, 4847-4858, December 2007
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Department of Cell Biology, Harvard Medical School, Boston, MA 02115
Submitted June 20, 2006;
Revised August 9, 2007;
Accepted September 6, 2007
Monitoring Editor: Ted Salmon
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). In human cells, at least four different CDKs along with four cyclin classes regulate cell division (for review, see Sanchez and Dynlacht, 2005). In late G1, D-type cyclins activate CDK4/6, promoting phosphorylation of Rb, activation of E2F-dependent transcription, and DNA synthesis. In S phase, A- and E-type cyclins bind and activate CDK2, which phosphorylates substrates that induce centrosome duplication and fire origins of replication (Sherr, 2000). In mitosis, A- and B-type cyclins bind and activate CDK1 to phosphorylate proteins that mediate the dramatic changes in nuclear and cytoskeletal architecture required for chromosome segregation (Nigg et al., 1996). Even though all CDKs phosphorylate a consensus motif consisting of S/TPxK/R (Nigg, 1991), different cyclin–CDK complexes phosphorylate distinct substrates, raising the question of how substrate selection is achieved by different cyclin–CDK complexes.
Cyclins play an important role in modulating the substrate selectivity of a particular cyclin–CDK pair. B-type cyclins bestow a high intrinsic activity to CDKs resulting in the phosphorylation of any substrate containing the canonical S/TPxK/R motif (Loog and Morgan, 2005). Alternatively, A- and E-type cyclins rely on a hydrophobic patch (HP) to recruit S/TPxK/R-containing substrates to its CDK partner for phosphorylation (Schulman et al., 1998; Brown et al., 1999). This HP interacts with substrates containing an RxL/Cy motif, which is often found in substrates whose phosphorylation occurs during S phase (Schulman et al., 1998; Cross and Jacobson, 2000). HP mutations in the budding yeast S phase cyclin Clb5, but not the M phase cyclin Clb2, abolish substrate binding and phosphorylation (Cross and Jacobson, 2000; Loog and Morgan, 2005). Recent studies of human cyclin B1 suggest that residues neighboring the HP are structurally distinct from those in A- and E-type cyclins and that they may actually inhibit interactions with RxL/Cy-containing substrates (Brown et al., 2007; Petri et al., 2007). Nevertheless, the HP is highly conserved among B-type cyclins, suggesting it plays some functional role. Consistent with this notion, HP mutations in the budding yeast mitotic cyclin Clb2 disrupt its localization to the bud neck (Bailly et al., 2003), suggesting that the HP may be conserved because it plays a critical role in mediating localization of B-type cyclins.
Additional evidence suggests that the capacity of cyclin B1 to promote CDK1-dependent events at mitotic entry may rely on its subcellular localization. At mitotic entry, cyclin B1-CDK1 promotes chromosome condensation, nuclear lamina solubilization, mitotic aster assembly, and Golgi breakdown, whereas cyclin B2-CDK1 can only induce Golgi disassembly (Draviam et al., 2001). These distinct functions of cyclin B1 and cyclin B2 to induce different events of mitosis correspond well with localization of the proteins. During interphase, cyclins B1 and B2 are both concentrated in the cytoplasm; cyclin B1 shuttles in between the nucleus and the cytoplasm, whereas cyclin B2 is mainly associated with the Golgi (Jackman et al., 1995). At prophase, cyclin B1 accumulates in the nucleus (Pines and Hunter, 1994; Hagting et al., 1998) and then to condensed chromatin, spindle microtubules, centrosomes, and chromatin during prometaphase (Pines and Hunter, 1991), corresponding to localization of known CDK1 substrates, including nuclear lamins (Enoch et al., 1991; Luscher et al., 1991), microtubule-associated proteins (Ookata et al., 1995; Charrasse et al., 2000) and the chromatin-associated protein RCC1 (Arnaoutov and Dasso, 2003). In contrast, cyclin B2 remains associated with the disassembled Golgi during mitosis. Domain mapping studies have suggested that the N-terminal domain is important in dictating the distinct functions of B-type cyclins. Replacement of the cyclin B1 N-terminal domain with that of cyclin B2 is sufficient to mislocalize the chimeric protein to the Golgi (Draviam et al., 2001). This chimeric protein retains the ability to induce Golgi disassembly, but it loses the ability to induce chromosome condensation, nuclear lamina solubilization, and mitotic aster assembly (Draviam et al., 2001), demonstrating that subcellular localization of different B-type cyclins is a critical determinant of their ability to properly promote the events of mitosis. Given the potential importance of localization in modulating the biological consequences of cyclin B1-CDK1 activity, we set out to identify the sequences in cyclin B1 that mediate its localization to specific structures during mitosis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Adenovirus Construction
All adenoviruses were constructed using the AdEasy system (He et al., 1998). Enhanced green fluorescent protein (EGFP) from pEGFP-N1 (Clontech, Palo Alto, CA) was subcloned into pShuttle-CMV to create pShuttle-CMV-EGFP. Full-length human cyclin B1 was amplified by polymerase chin reaction (PCR) with pfu polymerase (Stratagene, La Jolla, CA) and subcloned into pShuttle-CMV-EGFP to make the cyclin B1-EGFP fusion construct (FL1-433). To generate the FL
DB adenovirus, DNA encoding residues 1-41 and 51-433 of cyclin B1 was PCR amplified separately and cloned sequentially, yielding a construct that lacked the nine-amino acid Destruction box (D-box). To construct NT1-107, NT1-166, NT1-210, NT1-311, CT108-433, CT166-433, and CT211-433, DNA encoding the appropriate residues was PCR amplified, subcloned into pEGFP-N1 in frame, and then subcloned into pShuttle-CMV. The FLY170A and CT108-433, Y170A viruses were generated using the GeneTailor mutagenesis kit and primers designed to mutate the tyrosine at position 170 to an alanine (Invitrogen). DNA sequencing of both strands confirmed constructs. Adenoviral constructs were generated by recombination in BJ5183-AD1 cells (Stratagene), and infective adenoviruses were produced in 293 cells as described previously (He et al., 1998).
Time-Lapse Imaging and Microscopy
BS-C-1 cells were seeded onto no. 1.5 25-mm poly-L-lysine–coated glass coverslips or into glass-bottomed microwell dishes (MatTek, Ashland, MA). Adenovirus was added to BS-C-1 cells at the time of plating. Cells were imaged 16-18 h after infection. Mounted coverslips were imaged in a heated stage chamber at 37°C (20/20 Technology, Wilmington, NC), and microwell dishes were imaged in an enclosed incubation chamber at 37°C (Solent Scientific, Portsmouth, United Kingdom). Cells were cultured in CO2-independent L15 medium for the 20/20 system or in CO2-dependent DMEM for the Solent system. Similar results were obtained on both systems. During the course of the experiments presented in this manuscript, at least 50 FL1-433–expressing cells were imaged. At least 30 cells were imaged expressing NT1-107, NT1-166, CT108-433, and FLY170A. Images were acquired using MetaMorph imaging software (Molecular Devices, Sunnyvale, CA) on an inverted TE2000 microscope (Nikon, Melville, NY) by using a 100x 1.4 numerical aperture objective equipped with Yokogawa spinning disk confocal (PerkinElmer Life and Analytical Sciences, Boston, MA) and Orca-ER digital camera (Hamamatsu, Bridgewater, NJ). For time-lapse experiments, images were collected every 15–45 s, by using exposure times between 800 and 2000 ms, with illumination light shuttered between acquisitions. A 2 x 2 binning was used for all live-cell imaging. Figures were produced using Adobe PhotoShop 7.0 (Adobe Microsystems, Mountain View, CA).
Antibodies and Chemicals
Two of five antibodies tested reacted with endogenous cyclin B1 at the kinetochore: GNS1 and H-433 from Santa Cruz Biotechnology (Santa Cruz, CA). Antibodies that did not show kinetochore localization were H-20 and D-11 from Santa Cruz Biotechnology and Ab-3 from Calbiochem (San Diego, CA). All cyclin B1 antibodies were used a 1:1000 dilution, except the H-433 directly conjugated to fluorescein isothiocyanate (FITC) (sc-752 FITC), which was used at 1:100. Human anti-centromeric antibodies were used at a 1:250 dilution (Antibodies, Davis, CA). Hec1 antibody was used at 1:1000 (BD Biosciences Transduction Laboratories, Lexington, KY), Mad2 antibody was used at 1:100 (Abcam, Cambridge, MA), T9026 tubulin antibody was used at 1:500 (Sigma-Aldrich, St. Louis, MO), and Aurora B antibody was used at 1:500 (BD Biosciences, Franklin Lakes, NJ). Goat anti-mouse, goat anti-human, and donkey anti-rabbit secondary antibodies conjugated to Alexa-488, Alexa-568, or Alexa-647 were used at 1:1000 (Invitrogen). Monastrol was a gift from Tim Mitchison (Harvard Medical School, Boston), and it was used at a final concentration of 100 µg/ml. Nocodazole, taxol and MG132 were obtained from Sigma-Aldrich.
Immunofluorescence
Before immunostaining, 37°C media was exchanged for ice-cold media, and samples were incubated at 4°C for 10 min. After cold treatment, staining was performed as described previously (Waters et al., 1998). Cells were prepermeabilized in PHEM [60 mM piperazine-N,N'-bis(2-ethanesulfonic acid), 25 mM HEPES, 10 mM EGTA, and 4 mM MgSO4] with 0.5% Triton X-100 for 5 min, fixed in 1% formaldehyde in phosphate-buffered saline (PBS) for 15 min, and thoroughly rinsed in PBS with 0.05% Tween 20 (PBST). Cells were blocked in PBS plus 5% newborn calf serum for 1 h. All antibodies were diluted in blocking solution and staining was carried out at 37°C. Samples were washed after each step with PBST. DNA was stained using 2 µg/ml Hoechst 33342 (Invitrogen) in PBS for 2 min. Coverslips were mounted in 0.1 M n-propyl gallate (Sigma-Aldrich) in 90% glycerol in PBS. Z-series were acquired at 0.3-µm steps by using the same microscope as described for time-lapse imaging. Deconvolutions of Z-series were performed using AutoDeblur, version 8.0 (AutoQuant, Troy, NY).
Small Interfering RNA (siRNA) Transfection
HeLa cells were plated in Opti-MEM with 1% fetal bovine serum (Invitrogen) onto sterile glass coverslips at a density of 1 x 105 cells per well. Twenty-four hours after plating, cells were washed three times with Opti-MEM without FBS. GeneSilencer was used to transfect HeLa cells with modifications from the manufacturer's protocol (Genlantis, San Diego, CA). Opti-MEM, GTS diluent (GTS Gene Therapy Systems, San Diego, CA), Gene Silencer, and siRNA were mixed directly and incubated at room temperature for 15 min and then added to the cells. After 5 h of incubation, the media were replaced with DMEM plus 10% FBS. Control siRNA (Dharmacon RNA Technologies, Lafayette, CO, catalog. No. D-001210-01-20), Hec1 siRNA (Invitrogen, catalog no. 1299003; oligo HSS115931) or Mad2 siRNA (Dharmacon RNA Technologies, catalog no. KINRA-000007) were added to the cells at a final concentration of 25 nM. A mock transfection control was included in all experiments.
Immunoprecipitation
BS-C-1 cells were infected and plated at a density of 6 x 106 cells per 10-cm dish. Eighteen hours after infection, cells were washed with PBS and incubated with 1 ml of 20 mM Tris, 150 nM NaCl, 1.5 mM EDTA, 5 mM EGTA, and 0.5% NP-40 (TBST) for 30 min at 4°C. Cells were collected and centrifuged at 15,700 x g for 30 min at 4°C. Supernatant was transferred to a fresh tube, combined with 2 µl of anti-green fluorescent protein (GFP) (Ab290) (Abcam), and rotated at 4°C for 90 min. Washed and equilibrated protein A-Sepharose was added to each reaction and incubated overnight at 4°C. Reactions were washed three times with TBST and resolved by SDS-polyacrylamide gel electrophoresis. Immunoprecipitated proteins were detected by immunoblot by using cdc2 (Santa Cruz Biotechnology; sc-8395) or cyclin B1 (H-433) at 1:500 dilution or GFP antibody at 1:2000 dilution. Horseradish peroxidase-conjugated secondary antibodies were used at 1:5000 dilutions (Jackson ImmunoResearch Laboratories, West Grove, PA).
Quantitation of Kinetochore Signal
Z-series with a 0.3-µm step size were acquired of live, mitotic cells. Using MetaMorph imaging software, integrated intensity of individual kinetochores was collected from the median plane of the stack and from the three planes above and below the median plane. Fixed regions were placed over each kinetochore in the plane, in addition to a region outside the cell, the cytoplasm, and on chromatin. Chromatin associated signal was subtracted from kinetochore signal to generate normalized intensity values. Integrated intensity from 89 to 277 pairs was collected from a minimum of five different cells to calculate the normalized integrated intensity for the kinetochore signal. Confidence levels were calculated using Excel (Microsoft, Redmond, WA). Line scans measuring centrosome-associated integrated intensity were generated using MetaMorph imaging software.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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) (Figure 1). Spinning disk confocal microscopy also revealed the presence of cytoplasmic, punctate foci of FL1-433 in interphase (Figure 1A, asterisks). The size and number of these foci did not vary with viral titer, and they did not colocalize with lysosomes (data not shown). It has been reported that inactive cyclin B1–Cdc2 complexes form aggregates in starfish oocytes (Terasaki et al., 2003), and these foci may represent a similar form of sequestered cyclin B1. When using spinning disk confocal microscopy, we also observed a novel localization of FL1-433 to kinetochores during prometaphase (Figure 1, A–D, and Supplemental Video 1). This kinetochore population could first be observed at nuclear envelope breakdown (NEB) (Figure 1B). As cells progressed to metaphase, FL1-433 signal remained at kinetochores of chromosomes positioned near centrosomes, or on those clearly unattached to spindle microtubules (Figure 1, C and D, arrows). FL1-433 signal at kinetochores of chromosomes aligned at the metaphase plate was not detectable above the background of the chromatin-associated signal (Figure 1E). Local background subtraction methods (Hoffman et al., 2001) indicated that the overall decrease in FL1-433 signal intensity at kinetochores between the first indication of spindle microtubule attachment and the completion of congression was approximately fourfold (Supplemental Video 1; data not shown). After metaphase completion, we observed degradation of cyclin B1 as revealed by the global reduction of EGFP signal (Figure 1F).
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To confirm the live-cell imaging data, we determined the localization of endogenous cyclin B1 in BS-C-1 cells by indirect immunofluorescence. In these experiments, endogenous chromatin- and spindle microtubule-associated cyclin B1 was not preserved, but centrosome- and kinetochore-associated cyclin B1 was present (Figure 2). Punctate signal, which colocalized with anticentromeric antigen (ACA) signal, was observed in prometaphase cells labeled with cyclin B1 antibody confirming the localization of endogenous cyclin B1 to kinetochores (Figure 2, B and C, arrows). Preabsorption of primary antibodies with recombinant cyclin B1-CDK1 eliminated kinetochore-associated signal (Supplemental Figure S2). Consistent with live-cell imaging results, prometaphase kinetochores demonstrated robust cyclin B1 signal (Figure 2C, arrows), whereas kinetochores of fully congressed chromosomes exhibited almost no detectable signal (Figure 2, E and F). In anaphase cells, ACA signal persisted (Figure 2G), whereas cyclin B1 was undetectable (Figure 2H). Localization of endogenous cyclin B1 to kinetochores was also observed in HCT 116, MCF7, and MCF10A cell lines (data not shown), demonstrating that this novel localization is not unique to BS-C-1 cells. It is important to note that only two of five antibodies tested recognized the endogenous kinetochore population, which may explain the failure to observe cyclin B1 at kinetochores in previous fixed cell studies (Pines and Hunter, 1991; Jackman et al., 1995, 2003; Draviam et al., 2001). Both antibodies recognized epitopes in the N-terminal domain of cyclin B1, suggesting that C-terminal epitopes may become masked upon kinetochore association.
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). To determine whether FL1-433 kinetochore localization was sensitive to tension generation, we treated cells with taxol. Taxol stabilizes microtubules, thereby alleviating tension at kinetochores, but it does not ablate kinetochore–microtubule attachments (Waters et al., 1998). Metaphase-stage cells treated with taxol displayed shrinkage of the mitotic spindle but no FL1-433 relocalization to kinetochores (Figure 3C). This was also confirmed in fixed cells (Figure 3D). Together, these results show that the absence of tension is insufficient to promote cyclin B1 kinetochore relocalization, whereas lack of microtubule attachment to kinetochores causes robust cyclin B1 relocalization.
We also treated cells with monastrol, an inhibitor of the motor protein Eg5 (Mayer et al., 1999). We found that monastrol did not disrupt FL1-433 localization to kinetochores, chromatin, centrosomes, or spindle microtubules (Figure 3E), demonstrating that mitotic localization of cyclin B1 is not dependent on Eg5. Over time, kinetochore association of FL1-433 signal coalesced from two points into a single point, a pattern that remained for the duration of the experiment (Figure 3, E and F, arrows; and Supplemental Video 2). Signal coalescence is consistent with the stable attachment of microtubules to one of the kinetochores in a chromatid pair, as expected in prolonged monastrol exposure (Kapoor et al., 2000).
Cyclin B1 Localization Dynamics Are Independent of the Destruction Box
The destruction box (D-box) is a short-conserved sequence essential for recognition of cyclin B1 by the anaphase-promoting complex/cyclosome (APC/C) (Pfleger et al., 2001; Yamano et al., 2004). This D-box is required for degradation of cyclin B1 at the metaphase-to-anaphase transition. To assess the role of D-box–dependent degradation in cyclin B1 localization, we imaged a mutant in which the nine-amino acid D-box was deleted (FLDB). We observed that the dynamics of FLDB localization were identical to FL1-433: the protein was cytoplasmic during interphase and localized to chromatin, centrosomes, spindle microtubules, and kinetochores during mitosis (Figure 3, G and H). Furthermore, FLDB was delocalized from kinetochores at metaphase (Figure 3I), indicating that cyclin B1 dissociation does not depend on the APC/C. This protein failed to be degraded at the metaphase–anaphase transition, confirming that it is resistant to APC/C-mediated proteolysis (data not shown). Treatment with a proteasome inhibitor (MG132) did not perturb kinetochore association or dissociation of wild-type cyclin B1 (data not shown), demonstrating that the dissociation mechanism is also proteasome independent.
Cyclin B1 Localizes to the Outer Kinetochore
To better characterize the localization of cyclin B1-CDK1 at mitotic kinetochores, we compared its localization to that of ACA, Aurora B, Hec1, and Mad2 by immunofluorescence in BS-C-1 and HeLa cells. Relative localization of cyclin B1 was determined by line scan analysis through single kinetochore pairs in prometaphase cells. We found that localization of cyclin B1 in HeLa cells is similar to that observed in BS-C-1 cells (Figure 4). This analysis showed that cyclin B1 localized to the outer kinetochore, colocalizing with the outer kinetochore proteins Hec1 and Mad2 (Figure 4, A–D). Cyclin B1 also localized just outside of ACA signal in both cell types (Figure 4, E and F). In both cell types, cyclin B1 did not localize with Aurora B, a chromosomal passenger protein that resides in the inner centromere (Figure 4, G and H).
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). All cells that displayed this disorganized morphology were depleted of Hec1 as measured by immunofluorescence (Figure 5H), and they also showed depletion of cyclin B1 at the kinetochore (Figure 5I), although ACA staining was unperturbed (Figure 5G). Nocodazole treatment was insufficient to relocalize cyclin B1 to kinetochores (data not shown). Specificity of the antibody staining was confirmed by examining anaphase cells in control knockdown experiments, where we observed persistence of Hec1 kinetochore staining, but no cyclin B1 staining (Supplemental Figure 3). Furthermore, the level of Hec1 depletion correlated with cyclin B1 depletion on kinetochores in populations of cells treated with Hec1 siRNA (Supplemental Figure 4). Because Hec1 knockdown is known to displace multiple outer kinetochore proteins, including Mad2 (Martin-Lluesma et al., 2002; DeLuca et al., 2003, Lin et al., 2006), we next tested whether depletion of Mad2 was sufficient to inhibit cyclin B localization (Figure 5, K–T). Cells transfected with Mad2 siRNA were arrested in mitosis with 100 nM nocodazole and 2 µM MG132 for 1 h. Immunofluorescence of endogenous protein demonstrated that in the absence of Mad2 (Figure 5R), cyclin B1 localization at kinetochores was reduced (Figure 5S). Control experiments using primary and secondary antibodies separately demonstrated the specificity of the signal and the absence of bleed through (data not shown). These data show that cyclin B1 localization depends on Mad2 and other proteins removed upon Hec1 depletion.
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and Figure 7A), localization of NT1-107 must depend solely on cyclin B1 itself rather than indirect association mediated by CDK1. During interphase, NT1-107 localization was predominantly nuclear, presumably due to its lack of a cytoplasmic retention signal (CRS) (Figure 6B). Extension of NT1-107 to include the CRS region (NT1-166) restored its cytoplasmic localization during interphase (Figure 6D) but not its localization to centrosomes or kinetochores in mitosis (Figure 6C), demonstrating that the CRS is not a critical determinant of cyclin B1 localization during mitosis.
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CDK1 Association Is Not Necessary for Cyclin B1 Localization
Because the C-terminal domain of cyclin B1 is sufficient to bind and activate CDK1 (Murray et al., 1989; Nugent et al., 1991), we asked whether CDK1 association was the principal mechanism of FL1-433 and CT108-433 localization to centrosomes, spindle microtubules, and kinetochores. We generated a single amino-acid point mutation at residue 170 (FLY170A) that was predicted to abolish CDK1 binding. Point mutations at this conserved tyrosine residue ablate the ability of cyclin B2 to bind CDK1 (Goda et al., 2001). To determine whether the mutant proteins bound CDK1, we infected BS-C-1 cells with adenovirus expressing FL1-433 or mutant versions of cyclin B1 and we used a GFP antibody to immunoprecipitate EGFP-fusion proteins. By this analysis, FL1-433 and CT108-433 bound CDK1, whereas NT1-107 and FLY170A did not (Figure 6A). Therefore, FLY170A was used to determine the requirement for CDK1 binding in localization of cyclin B1 to mitotic structures.
Time-lapse, spinning disk confocal microscopy showed that FLY170A was cytoplasmic during interphase and translocated to the nucleus before NEB (data not shown). On entry into mitosis, FLY170A localized to kinetochores, chromatin, spindle microtubules, and centrosomes (Figure 7C) in a manner that was indistinguishable from FL1-433 (Figure 7B). Loss of kinetochore signal during metaphase congression was also observed (Supplemental Video 4), indicating that CDK1 association is not required for cyclin B1 dissociation from kinetochores after microtubule attachment, or degradation of cyclin B1 at the metaphase-to-anaphase transition. Nocodazole treatment stimulated localization of FLY170A to kinetochores (data not shown). These results demonstrate that cyclin B1 can mediate its own dynamic localization during mitosis, independent of CDK1 interaction.
Although qualitative inspection showed no difference between FL1-433 and FLY170A localization, we quantitatively measured kinetochore-associated signal to validate this conclusion. Quantitation confirmed that in unperturbed cells, there was no significant difference between FL1-433, FLY170A, or CT108-433 signal at kinetochores in prometaphase (Figure 7D). When cells were arrested for at least 30 min with 100 nM nocodazole, a statistically significant increase in the kinetochore-associated signal of FL1-433 and FLY170A was observed (Figure 7D). This is consistent with literature showing that some kinetochore-associated proteins accumulate during a nocodazole arrest (Hoffman et al., 2001). However, CT108-433 kinetochore-associated signal was not increased after nocodazole-induced arrest (Figure 7D). These data suggest that although the N-terminal domain of cyclin B1 is not essential for kinetochore localization, it may play a role in stimulating association at the kinetochore under conditions of nocodazole arrest.
The normal localization of FLY170A demonstrates that cyclin B1 does not require CDK1 binding for localization to any mitotic structures. Therefore, we predicted that association of CT108-433 to kinetochores, spindle microtubules and centrosomes would also be independent of CDK1 binding. To test this idea, we generated a version of CT108-433 containing the Y170A mutation (CT108-433, Y170A). As expected, we found that this mutation abolished the interaction with CDK1 (Figure 7A). Surprisingly, CT108-433, Y170A failed to localize to kinetochores, although it remained at centrosomes and spindle microtubules (Figure 7F). Nocodazole was insufficient to localize CT108-433, Y170A to kinetochores (data not shown). These data show that CDK1 association is required for the kinetochore localization of cyclin B1 when its N-terminal domain is truncated, indicating that cyclin B1 may also be recruited to the kinetochore by a CDK1-dependent mechanism.
The Cyclin Box, but Not CDK1 Binding, Is Required for Centrosome Localization
Cyclin B1 localization to centrosomes has been proposed to be a site of cyclin B1–CDK1 activation that may regulate mitotic entry (Jackman et al., 2003). To identify sequence elements required for cyclin B1 localization to the centrosome, we measured the integrated intensity of EGFP signal across the length of the spindle and through both centrosomes by line scan in mitotic cells expressing various cyclin B1–EGFP constructs. By this method, FL1-433 and CT108-433 signal showed two peaks of centrosome-associated signal; for FL1-433, this centrosome-associated signal was roughly twice as bright as that associated with chromatin, spindle microtubules, or the cytoplasm (Figure 8A, arrows). Confirming the specificity of this analysis, NT1-107 showed no peaks of centrosome signal (Figure 8B), consistent with our qualitative analysis. FLY170A and CT108-433, Y170A localized efficiently (Figure 8, C and D, arrows), demonstrating that CDK1 interaction is not required for localization to centrosomes, even when the N-terminal domain is removed. This pattern of localization demonstrates that the mechanism of cyclin B1 localization to centrosomes is distinct from that of localization to kinetochores, where CDK1 binding is essential upon removal of the N-terminal domain (Figure 7F).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Hagting et al., 1998; Clute and Pines, 1999). Spinning disk confocal microscopy also revealed novel localization of FL1-433 at kinetochores (Figure 1), which was confirmed by immunostaining of endogenous cyclin B1 protein (Figures 2 and 4). Kinetochore localization is sensitive to the state of microtubule attachment. Cyclin B1 is found at unattached kinetochores during prometaphase and after microtubule depolymerization with nocodazole (Figure 3). Consistent with our findings, the binding partner of cyclin B1, CDK1, has been shown to localize to kinetochores in mitosis (Rattner et al., 1990). Our data provide evidence that the CDK1 population at kinetochores is in fact active and that the abundance of the complex varies during mitotic progression.
Our mutagenesis data suggest that either cyclin B1 or CDK1 can independently recruit the cyclin B1–CDK1 complex to unattached kinetochores. Cyclin B1 can localize to kinetochores independently of CDK1 association, because a point mutation that abolishes CDK1 association (Y170A) does not perturb the normal dynamics of kinetochore association and dissociation (Figure 7). This is consistent with previous studies showing that mutations in Clb2 that ablate Cdc28 binding do not alter its normal localization to the bud neck, spindle pole body, or mitotic spindle (Bailly et al., 2003). Our data also suggest that sequences in both the N- and C-terminal domains of cyclin B1 are required for kinetochore localization in the absence of CDK1 binding, because the N-terminal domain is insufficient to localize to kinetochores (NT1-311), and the C-terminal domain fails to associate with kinetochores if it cannot bind CDK1 (CT108-433, Y170A). Together, these findings demonstrate that cyclin B1 can interact directly with components of the kinetochore.
Our data also reveal that CDK1 can target the cyclin B1–CDK1 complex to kinetochores. This mechanism is demonstrated by the localization of cyclin B1 mutants lacking the N-terminal domain. Although the CDK1-binding CT108-433 construct shows normal localization and delocalization from kinetochores (Figure 6 and Supplemental Video 3), a construct that does not bind CDK1 (CT108-433, Y170A) fails to localize to kinetochores (Figure 7). Furthermore, any construct able to bind CDK1 was also found to localize to kinetochores (Figure 9), suggesting that CDK1 binding may be sufficient for kinetochore localization. We propose that cyclin B1 association can render the entire cyclin B1–CDK1 complex sensitive to the state of microtubule attachment, perhaps through dynein-dependent transport of the complex used by spindle checkpoint proteins to leave the kinetochore (Hoffman et al., 2001; Howell et al., 2001). In this proposed model, active removal of cyclin B1 upon microtubule attachment would inactivate the kinetochore population of CDK1.
We found that cyclin B1 colocalizes with the Hec1 and Mad2 at the outer kinetochore during prometaphase (Figure 4). In addition, siRNA experiments show that either Hec1 or Mad2 depletion alters the normal kinetochore localization of cyclin B1 (Figure 5), demonstrating that both of these proteins must play a role in targeting cyclin B1 to the kinetochore. Although Hec1 depletion alters Mad1, Mad2, Zwint, Zw10, and Mps1 localization (Martin-Lluesma et al., 2002; DeLuca et al., 2003, Lin et al., 2006), Mad2 depletion has never been shown to alter the localization of other checkpoint proteins (Chung and Chen, 2002; Orr et al., 2007), and the relative dependencies of other checkpoint proteins on Mad2 kinetochore association are not sufficiently understood. Therefore, our data showing that cyclin B1 levels are reduced after Mad2 depletion is mechanistically intriguing. However, because Mad2 depletion results in a more modest effect on cyclin B1 than Hec1 depletion, we propose that additional proteins other than Mad2 and also affected by Hec1 depletion, play a role in targeting cyclin B1 to the kinetochore. Importantly, our data rule out a mechanism in which Cdc20 or APC/C-association alone recruit cyclin B1 to kinetochores (Acquaviva et al., 2004), because nondegradable versions of cyclin B1 showed normal dynamics of kinetochore localization and delocalization (Figure 3).
Our identification of cyclin B1 at unattached kinetochores corroborates previous studies proposing a positive role of CDK1 in spindle checkpoint activity in budding yeast (Kitazono and Kron, 2002; Kitazono et al., 2003) and in vertebrates (Chung and Chen, 2003; D'Angiolella et al., 2003). One potential function of kinetochore-associated cyclin B1-CDK1 could be to promote inhibition of Cdc20 by Mad2. Cdc20 contains two putative Cdk1 phosphorylation sites (Yudkovsky et al., 2000), and Cdc20 phosphorylation is required for Mad2 binding, as nonphosphorylatable versions of Cdc20 are resistant to Mad2 binding and inhibition (Chung and Chen, 2003; D'Angiolella et al., 2003). Cdc20 localizes to kinetochores from prophase through anaphase, but its association is highly dynamic, with binding times ranging from 1 to 5 s to 21 to 23 s (Kallio et al., 2002; Howell et al., 2004). We propose that localization of cyclin B1-CDK1 to unattached kinetochores could ensure the local phosphorylation of Cdc20, facilitating its binding to the active conformation of Mad2 generated at the kinetochore. In addition, other kinetochore proteins are known substrates of the cyclin B1–CDK1 complex, including INCENP (Goto et al., 2006), Bub1 (Qi et al., 2006), and APC/C subunits (Golan et al., 2002; Kraft et al., 2003). Therefore, recruitment of cyclin B1-CDK1 to unattached kinetochores could affect the activity of other components of the spindle checkpoint or regulate other processes that are essential for chromosome congression and anaphase onset.
The N-Terminal Domain Targets Cyclin B1 to Chromatin
The first 107 residues of cyclin B1 are necessary and sufficient for chromatin localization, providing a mechanism to concentrate CDK1 near chromatin-associated substrates. Because the N-terminal domain of cyclin B1 cannot bind CDK1, and the C-terminal CDK1-binding domain cannot localize to chromatin, we conclude that chromatin localization occurs independently of CDK1 binding. Possible CDK1 substrates on chromatin include condensins (Kimura et al., 1998), RCC1 and histone H1 (Arnaoutov and Dasso, 2003; Hutchins et al., 2004).
Besides targeting cyclins for APC/C-dependent degradation, other functions for the N-terminal domain of cyclin B1 have not been described. We found that removal of the conserved D-box, a motif required for interactions with Cdc20 and the APC/C (Yamano et al., 2004), did not alter chromatin localization, demonstrating that undefined sequences within the N-terminal domain are necessary for chromatin localization. The N-terminal domain is highly basic and could nonspecifically promote association with negatively charged DNA. However, cyclin A, which has access to condensed chromatin during early stages of mitosis and a similar isoelectric point, does not localize to chromatin (den Elzen and Pines, 2001). Therefore, it is likely that specific sequences within the N-terminal domain of cyclin B1, rather than charge-based mechanisms, are critical to chromatin localization.
The C-Terminal Domain Localizes Cyclin B1 to Centrosomes
Our data indicate that localization of cyclin B1 to centrosomes, like the chromatin population, is mediated directly by cyclin B1, rather than through a CDK1-dependent mechanism. Mutants of cyclin B1 that do not bind to CDK1 showed normal localization to centrosomes (Figures 8 and 9). Unexpectedly, the determinants of localization to chromatin and centrosomes are independent and separable. Whereas the N-terminal domain is necessary and sufficient for chromatin localization, the C-terminal domain of cyclin B1 is the critical mediator of centrosome association. Residues critical for centrosome localization lie within the cyclin box, because extension of the N-terminal domain to include the cyclin box (NT1-311) was sufficient to restore localization to centrosomes (Figure 8). Although the cyclin box plays an essential role in binding to CDK1, our findings indicate that its role in mediating centrosome association is independent of CDK1 binding. We propose that sequences specifically in the hydrophobic patch, which includes the MRAIL motif (positions 201–205) and a series of other conserved residues (W208, Q211, V212, L244, and V272) may be the critical determinants of cyclin B1 localization to centrosomes. The hydrophobic patch lies on a face of cyclin B1 opposite the face required for CDK1 association, explaining how it might mediate centrosome association independent of CDK1 binding. A fusion construct containing only residues 211-311 did not localize to any structure or compartment properly (data not shown), consistent with a requirement for the entire MRAIL motif and all HP residues. Consistent with our proposed HP-dependent mechanism of localization, HP mutations in Clb2 of budding yeast perturb localization to the bud neck (Bailly et al., 2003). However, because these same mutations in Clb2 do not disrupt localization to the spindle pole body, HP residues may have different roles in determining cyclin B localization in different organisms.
The Role of Cyclin B1 Localization in CDK1 Action
Our analysis shows that cyclin B1 is localized to different mitotic structures through distinct and separable mechanisms. The N-terminal domain of cyclin B1 is necessary and sufficient for chromatin localization, whereas the C-terminal domain including the HP residues mediates centrosome localization. In both cases, localization can occur independently of CDK1 binding. The requirements for kinetochore localization are more complex, because cyclin B1 localizes to kinetochores through both direct and indirect (CDK1-dependent) mechanisms.
The existence of distinct cyclin B1–CDK1 recruitment mechanisms has important implications for understanding the specificity of substrate phosphorylation during mitosis and also for regulation of cyclin B1-CDK1. For example, the affinity of cyclin B1 binding sites on chromatin, kinetochores, and centrosomes may be distinct. Therefore, cyclin B1 may be recruited to high-affinity sites during early stages of mitosis and be recruited to lower affinity sites only when cyclin B1 levels increase later in mitosis. The existence of distinct recruitment mechanisms also permits differential regulation of recruitment of cyclin B1 to each structure during mitosis. This principle is illustrated by our observation that kinetochore-associated cyclin B1 is regulated by the state of microtubule attachment, whereas levels of chromatin and centrosome-associated cyclin B1 remain steady until degradation at anaphase onset. Distinct recruitment pathways may also enable differential regulation of cyclin B1–CDK1 activity at different mitotic structures. It has been shown that the initial site of cyclin B1–CDK1 activation is at the centrosome (Jackman et al., 2003). This centrosome-localized activation relies on Plk1-dependent phosphorylation and Cdc25B-dependent dephosphorylation of cyclin B1-CDK1 (Jackman et al., 2003; Lindqvist et al., 2005). The other isoforms of Cdc25 have different patterns of localization, and they do not promote cyclin B1-CDK1 activation as efficiently in vivo (Dalal et al., 1999; Lindqvist et al., 2004, 2005; Kallstrom et al., 2005). Therefore, the existence of a specific centrosome-recruitment sequence on cyclin B1 may help concentrate the players necessary for efficient cyclin B1–CDK1 activation during mitotic entry.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
* Present address: Genome and Proteome Sciences, Novartis Institutes for Biomedical Research, 250 Massachusetts Ave., Cambridge, MA 02139.
Address correspondence to: Randall W. King (randy_king{at}hms.harvard.edu).
Abbreviations used: APC/C, anaphase-promoting complex/cyclosome; ACA, anti-centromeric antigen; CRS, cytoplasmic retention signal; D-box, Destruction box; HP, hydrophobic patch; NEB, nuclear envelope breakdown.
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-GFP). CDK1 associates with FL1-433 and CT108-433 but not with NT1-107, FLY170A or CT108-433, Y170A (
