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Correspondence to Don W. Cleveland: dcleveland{at}ucsd.edu
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satellite DNA (Cleveland et al., 2003; Amor et al., 2004b; Carroll and Straight, 2006).
Despite the prevalence of centromeres at adenine-thymine–rich repetitive satellite DNA, the DNA sequences themselves appear to play a nonessential role in centromere specification. This is most clearly exemplified by the characterization of human neocentromeres. In these rare but naturally occurring patient cases, a specific centromere has relocated to another site on the chromosome without any apparent DNA rearrangements, concomitant with vacating the original satellite–containing locus (Amor and Choo, 2002; Amor et al., 2004a; Ventura et al., 2004). This shows that DNA sequences normally associated with centromeres are neither necessary nor sufficient to promote centromere propagation and that maintenance of centromeres is determined predominantly in an epigenetic manner.
Centromere protein A (CENP-A) is a conserved histone H3 variant that replaces canonical H3 specifically at centromeres in all known eukaryotes (Palmer et al., 1987; Meluh et al., 1998; Henikoff et al., 2000; Oegema et al., 2001) and has been shown to be required for the localization of nearly all other centromere and kinetochore components (Howman et al., 2000; Oegema et al., 2001; Goshima et al., 2003; Amor et al., 2004a; Regnier et al., 2005; Foltz et al., 2006; Liu et al., 2006). We have recently shown that the loop1 and 2 helix of the CENP-A histone fold domain is responsible for forming a rigid/inaccessible interface with histone H4 and that this region, when transplanted into canonical histone H3, confers centromere targeting (Black et al., 2004, 2007a) and provides an essential function of CENP-A (Black et al., 2007b). CENP-A chromatin directly recruits a six-component CENP-A nucleosome-associated complex (CENP-ANAC) that forms the foundation for the assembly of other centromere components and the kinetochore during mitosis (Foltz et al., 2006). The existence of a CENP-A–directed centromere-specific chromatin structure makes CENP-A a prime candidate for the epigenetic propagation of centromere identity. This directly implies that CENP-A propagation at the centromere is a partially or completely self-directed process. It is, however, unknown how CENP-A is discriminated from canonical histone H3 and how its specific incorporation at centromeric nucleosomes is achieved.
Earlier models have suggested that differences in timing of replication of centromeric DNA versus the genome overall may provide a temporal window permissive for CENP-A loading (O'Keefe et al., 1992; Csink and Henikoff, 1998). However, this appears not to be the case, as replication of centromeric DNA is not restricted to a specific time during S phase (Shelby et al., 2000; Sullivan and Karpen, 2001). Alternatively, CENP-A loading could be separate from assembly of canonical histones altogether by allowing CENP-A loading outside S phase. Indeed, DNA replication is not required for CENP-A assembly and CENP-A mRNA, and protein levels peak only after S phase during late G2 phase, consistent with a disconnect between the timing of CENP-A and H3 assembly (Shelby et al., 1997, 2000). Whether propagation of centromeric chromatin and general chromatin is indeed temporally distinct and how and when CENP-A nucleosomes turn overis not known. This we now test by developing and exploiting a novel, covalent fluorescent pulse-labeling strategy with SNAP tagging.
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CENP-A is stably associated with centromeres across the cell cycle
To determine turnover of CENP-A at centromeres, cells were synchronized at the G1–S boundary by tandem treatments with thymidine. CENP-A bound to unreplicated centromeres was pulse labeled with TMR-Star and chased for up to two cell cycles (Fig. 1 A and Fig. 2 A).
Consistent with earlier reports on total CENP-A levels that had indicated slow protein turnover (Shelby et al., 2000; Regnier et al., 2005), centromere duplication in the initial round of DNA synthesis produced a 60 ± 14% reduction in intensity of TMR-Star–labeled CENP-A–SNAP at individual centromeres by the first mitosis and through the subsequent G1 (Fig. 2). After a second cycle of DNA replication, the previously labeled, centromere-bound CENP-A–SNAP was diminished to 25 ± 5% of its initial level, whereas the total number of fluorescent centromeres positive per cell remained unchanged throughout the experiment (Fig. 2). Thus, despite continued synthesis of both SNAP-tagged and endogenous CENP-A, CENP-A already loaded into centromeric chromatin by late G1 is redistributed to, and retained by, daughter centromeres.
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11 h after release from thymidine and concomitant with entry into G1 (Fig. 3 C).
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To time the arrival of CENP-A–SNAP at the centromere more accurately, we followed live cells containing a pool of TMR-Star–labeled but nonassembled CENP-A–SNAP from metaphase through early G1 (Fig. 3 D). TMR-Star labeling of live cells resulted in the nonspecific retention near the cell periphery (presumably in internal membranes) of a proportion of the dye, a proportion that is removed during normal fixation and washing conditions. Nevertheless, no TMR-Star signal could be detected specifically at centromeres in metaphase (Fig. 3 E). However, assembly of nascent CENP-A–SNAP could be detected as early as 50 min after anaphase onset and continued for several hours in early G1 (Fig. 3 E and Video 1, available at http://www.jcb.org/cgi/content/full/jcb.200701066/DC1).
CENP-A assembly occurs exclusively during G1
Next, we determined whether loading of CENP-A is unique to the early hours of G1 or whether loading is also permissive at any other point in the cell cycle, including the possibility of a secondary CENP-A loading stage, as has been suggested in fission yeast (Takahashi et al., 2005). CENP-A–SNAP in mitotic cells arrested with nocodazole treatment was initially quenched with nonfluorescent BG, and a G1 phase cell population was generated by release from nocodazole arrest. The CENP-A–SNAP pool produced during mid-to-late G1 was labeled and monitored for timing of centromeric deposition (Fig. 4).
DNA content was assayed by FACS to verify cell cycle position (unpublished data). No assembly of CENP-A–SNAP–TMR-Star was detectable at centromeres during the subsequent S, G2, or M phases (Fig. 4 B). However, fluorescent CENP-A from the prior G1 assembled into the new daughter centromeres after exit from this subsequent mitosis (Fig. 4 B). Thus, despite the presence of a stable noncentromeric CENP-A pool, no loading occurred at any stage of the cell cycle before the following G1 phase (10–18 h after CENP-A synthesis and labeling).
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3.4- and 2.5-fold, respectively; Fig. 5), findings only consistent with CENP-A loading in G1 rather than before mitosis.
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It should be noted that because of the short time cells spend in mitosis (1 h) and the inherent spread in synchrony as cells transverse across the cell cycle, an early G1 phase cell population will invariably contain a fraction of cells that are in G2. (In this case, 33% of the H3-CFP G1 population had in fact not yet reached G1 by the time cells were fused.) Therefore, in all fusions, a spread of heterokaryons loading CENP-A–SNAP at centromeres at one, both, or neither of the nuclei is expected. Nevertheless, despite this inherently imperfect synchrony, a striking finding was that in binucleate heterokaryons derived from fusion between cell populations enriched in G1 and G2, most (66%) G1 cell–derived nuclei (H3-CFP marked) recruited CENP-A–SNAP to centromeres to levels indistinguishable from surrounding nonfused G1 cells. In contrast, no heterokaryons were found that had assembled CENP-A in both nuclei, indicating that G2-derived nuclei, although sharing the same cytoplasm with a CENP-A–assembling G1-derived nucleus, did not assemble CENP-A (Fig. 6, B and C), despite the presence of fluorescently labeled CENP-A–SNAP. The frequency of heterokaryons loading CENP-A–SNAP in one or neither nucleus corresponded to the frequency of H3-CFP G1 and G2 cells at the time of fusion (Fig. 6 C, arrows), indicating that in heterokaryons the G1- and G2-derived nuclei are neither inducing nor inhibiting CENP-A assembly in the other nucleus. Therefore, the early G1 cell cycle state that is permissive for CENP-A assembly does not directly dictate the ability to load CENP-A. Rather, passage through mitosis is crucial to allow CENP-A assembly as cells enter G1.
Microtubule attachment is not required for CENP-A assembly in G1
Our experiments suggest that mitosis is a key cell cycle determinant in initiating CENP-A loading. To exclude the possibility that proficiency for CENP-A loading is determined by a "timing" mechanism rather than actual mitotic passage and G1 entry, cells were arrested using nocodazole to produce a nascent unloaded pool of CENP-A–SNAP in mitosis. Nocodazole-treated cells never assembled CENP-A–SNAP, even by the time 94% of control cells had reentered G1 and loaded CENP-A–SNAP (Fig. 4 B), reaffirming the notion that exit from mitosis is required for CENP-A loading.
Multiple processes occur during mitosis that might act to trigger new CENP-A nucleosome recruitment. These include chromatin stretching, which occurs during metaphase and has been proposed as a mechanism for the exchange of histone H3 for CENP-A–containing nucleosomes (Ahmad and Henikoff, 2002; Mellone and Allshire, 2003; Carroll and Straight, 2006). Although the concept of functional reinforcement of centromere location that is part of this model is appealing, no experimental evidence has been generated in support for such a mechanism. Alternatively, DNA decondensation or the presence of other mitotic kinetochore components may be integral to triggering the process of centromere assembly.
To test the tension-dependent CENP-A loading model, cells were produced that completed mitosis in the absence of microtubule attachment (and therefore microtubule-mediated chromatin stretching). To do this, cells were depleted of BubR1 with transcription-mediated short hairpin RNA and treated with nocodazole to block microtubule assembly, and CENP-A loading was assessed (Fig. 7 A). Under these conditions, cells enter mitosis without spindle assembly or kinetochore attachment, but quickly exit without the BubR1-dependent mitotic checkpoint (Kops et al., 2004).
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50-h time frame of our experiments. (An added benefit of this outcome is demonstration that the covalent SNAP-BG binding is indeed irreversible.) Moreover, the ability to differentially label SNAP protein pools synthesized at different times allows direct assessment of the fate of nascent proteins, including the turnover rates of proteins at the same cellular location, but assembled at different times.
Little CENP-A turnover across the cell cycle, but assembly in G1 phase
Using the SNAP tag approach, nearly all centromeric CENP-A is shown to remain centromere associated even during centromeric DNA replication, consistent with a role for CENP-A as an epigenetic marker maintaining centromere identity though cell division (Vafa and Sullivan, 1997; Warburton et al., 1997; Black et al., 2004, 2007a). More surprisingly, loading of newly synthesized CENP-A occurs in a discrete cell cycle window in early G1. A mitosis intervening between centromere DNA replication and new CENP-A loading is a prerequisite for CENP-A assembly. Although earlier work suggested that CENP-A may load in G2 phase based on an increase in overall CENP-A protein levels at this time (Shelby et al., 2000), our direct visualization with the SNAP tag has demonstrated that, despite its continued expression throughout the cell cycle, newly made CENP-A is accumulated in a nuclear, but noncentromeric, form before mitosis. The abrupt onset of CENP-A assembly at centromeres initiating at the end of mitosis firmly supports a model in which loading of CENP-A requires one or more key events during mitosis that may include nuclear envelope breakdown or chromatin decondensation, thereby allowing potential CENP-A assembly factors access to centromeric chromatin. Alternatively, assembly may be dependent on mitotic modification of CENP-A itself, which creates an environment that is permissive for subsequent CENP-A loading.
Although passage through mitosis itself is a strict requirement for CENP-A loading, microtubule attachment at kinetochores has no apparent role in CENP-A assembly, in contrast to previous proposals (Ahmad and Henikoff, 2002; Mellone and Allshire, 2003; Carroll and Straight, 2006). It remains possible, however, that components of the greater centromere/kinetochore affect CENP-A loading or stabilization after loading. Defects in structural centromere proteins have been shown to affect CENP-A levels (Kline et al., 2006; Okada et al., 2006). It is therefore possible that components of the centromere (which themselves are dependent on CENP-A for their localization) recruit new CENP-A or parts of the loading machinery. This would serve a direct epigenetic feedback between active centromeres and the propagation of new centromeric chromatin.
Finally, propagation of CENP-A chromatin may await the availability of an active loading factor or an adaptor molecule at centromeres. The recently identified hMis18, hMis18ß, and M18BP1/hsKNL2 proteins, of which the M18BP1/hsKNL2 Myb domain–containing protein is an evolutionary conserved component, have been proposed to be required for CENP-A loading (Fujita et al., 2007; see Maddox et al. on p. 757 of this issue). Strikingly, all these proteins display a pattern of centromere localization coincident with CENP-A assembly (from anaphase through early G1). Thus, recruitment of hMis18, hMis18ß, and M18BP1/hsKNL2 at the centromere during late anaphase could be dictated by a modification of centromeric chromatin coincident with mitotic exit or may represent a component of the CENP-A loading machinery that is itself activated during mitotic exit.
Implications for CENP-A assembly and centromeric chromatin structure
The sudden onset of CENP-A assembly exclusively after reentry into G1, but not in mitosis, carries with it two important implications for epigenetic centromere inheritance. First, a requirement for a subsequent mitosis as a prerequisite for loading of CENP-A onto previously replicated centromeric DNA intrinsically couples centromere replication and maturation to cell cycle progression. Second, loading of new CENP-A after mitosis dictates that centromeres and the kinetochores assembled on them proceed through mitosis with only half the complement of CENP-A. During S phase, CENP-A protein is redistributed among sister centromeres, leaving vacant DNA sequences that are not replenished by CENP-A but are most likely occupied by typical histone H3.1–containing nucleosomes, which are available in excess during DNA replication. Indeed, histone H3–containing nucleosomes have been detected on mitotic centromeres interspersed with CENP-A–containing nucleosomes and have been shown to occupy centromeric chromatin when CENP-A levels are depleted (Blower et al., 2002; Sullivan and Karpen, 2004). Our work now indicates that the mixed chromatin state generated in S phase does not represent an intermediate state of centromeric chromatin where canonical nucleosomes serve as transient placeholders but that in fact this centromeric chromatin composition is what promotes kinetochore formation during mitosis (Fig. 8).
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-tubulin were generated by stable integration via Moloney murine leukemia retroviral delivery essentially as described previously (Shah et al., 2004; CFP–-tubulin retroviral construct was provided by J. Shah, Harvard Medical School, Boston, MA). Cells stably expressing the CENP-A–SNAP fusion protein were selected by blasticidin S (5 µg/ml; Calbiochem) and were isolated and individually sorted by flow cytometry. The resulting monoclonal lines were expanded and examined by fluorescence microscopy after TMR-Star labeling and by Western blot to identify lines expressing proper levels of the CENP-A–SNAP fusion protein. Clone 23 (Fig. S1) was used for all experiments in this paper unless stated otherwise. Ratio of CENP-A–SNAP levels to endogenous CENP-A in parental HeLa cells is 0.7:1. H3-CFP or CFP–-tubulin cell lines were isolated by puromycin selection (1 µg/ml; Calbiochem). BubRI or control short hairpin RNA producing pSUPER plasmids and transfection procedures were as described previously (Kops et al., 2004).
SNAP quench and pulse labeling
SNAP tag activity in cells was quenched by addition of 20 µM O6-BG (BG-block; Covalys) in complete growth medium for 30 min at 37°C or pulse labeled with 2 µM TMR-Star (Covalys) in complete growth medium for 15 min at 37°C. After quenching or pulse labeling, cells were washed twice with prewarmed PBS, after which cells were reincubated in complete medium to allow excess compound to diffuse from cells. After 30 min, cells were washed again twice in PBS followed by reincubation in complete medium.
Cell synchronization
Unless stated otherwise in figures or legends, HeLa cells were treated with 2 mM thymidine in complete medium for 17 h, washed twice in PBS, and released in complete medium containing 24 µM deoxycytidine for 9 h followed by addition of thymidine to a final concentration of 2 mM for 17 h, after which cells were released again into complete medium containing 24 µM deoxycytidine and assayed. Nocodazole was used at 100 ng/ml.
Immunofluorescence
Cells were grown and SNAP assayed on glass coverslips followed by fixation and processed for immunofluorescence using standard procedures. Cells were not preextracted before fixation. Anti–CENP-A (a gift from K. Yoda, Nagoya University, Nagoya, Japan) was used at a dilution of 1:100, anti–CENP-C (a gift from W. Earnshaw, University of Edinburgh, Edinburgh, UK) sera was used at a dilution of 1:1,000, and anti-Mad1 (a gift from A. Musacchio, European Institute of Oncology, Milan, Italy) tissue culture supernatant was used at a dilution of 1:20. YL1/2 -tubulin (Serotec) was used at a dilution of 1:2,500. Anti-HA11 (Covance Research Products, Inc.) was used at a dilution of 1:1,000. Donkey secondary antibodies (anti-mouse Cy5- or FITC-conjugated and anti-rabbit FITC-conjugated) were obtained from Jackson Immunoresearch Laboratories. Samples were stained with DAPI before mounting in ProLong (Invitrogen).
Cell–cell fusions
Double thymidine–arrested H3-CFP and CFP-tubulin–expressing CENP-A–SNAP cells were SNAP-quenched with BG-block followed by release either directly or after 4 h to generate out-of-phase populations. After release of the trailing population, cells were coseeded on 18 x 18 mm uncoated coverslips. 5 h after seeding, coverslips were washed once in prewarmed PBS and incubated cell-side down in a 100 µl PEG-1500 (Roche) for 30 s followed by addition of 500 µl PBS and three subsequent washes in PBS. Coverslips were returned to complete medium containing 100 ng/ml nocodazole to prevent G2 cells from entering G1 and were TMR-Star labeled and fixed 4 h after PEG fusion.
Microscopy
Digital images were captured using a DeltaVision RT system (Applied Precision) controlling an interline charge-coupled device camera (Coolsnap; Roper) mounted on an inverted microscope (IX-70; Olympus). For each sample, images were collected at 1x binning using a 100x oil objective at 0.2 µm z sections spanning the entire nucleus and were subsequently deconvolved, and maximum signals were projected as 2D images using softWoRx (Applied Precision; all images are deconvolved except those shown in Fig. S2). For quantification, images were converted to unscaled TIFF images. Centromere signal intensity was determined using MetaMorph (Molecular Devices) by measuring integrated fluorescence intensity within an 8 x 8 pixel square. Background signal was subtracted from an area within the nucleus not containing centromeres. For live-cell imaging, cells were grown on 22 x 22 mm glass coverslips transfected with YFP–CENP-C (Shah et al., 2004) using Effectene (QIAGEN) 48 h before SNAP labeling, after which coverslips were mounted on a slide separated in a double-stick tape chamber in phenol red–free CO2 independent DME (Invitrogen) containing 0.5 U/ml of the oxygen-scavenging enzyme, Oxyrase (Oxyrase, Inc.), and sealed with a 1:1:1 mixture of vasalin, lanolin, and paraffin.
Images were acquired at 2x binning using a 60x oil objective for TMR-Star and YFP, as well as differential interference contrast at 10-min intervals. For each time point, 5 x 1 µm z sections were acquired for fluorescence images, and a single differential interference contrast image was acquired at the middle z position. Stacks were deconvolved, and maximum intensity was projected using softWoRx and assembled into a paneled video using MetaMorph.
Immunoblots
Whole cell extracts equivalent to 50,000 cells were separated by SDS-PAGE and transferred to nitrocellulose. Blots were probed by human anti-centromere serum (Antibodies, Inc.) at a dilution of 1:300.
Online supplemental material
Fig. S1 shows the cell cycle–dependent CENP-A assembly of independently established cell lines expressing different levels of CENP-A–SNAP. Fig. S2 shows the diffuse nuclear localization of noncentromere-loaded CENP-A–SNAP in G2 cells. Fig. S3 shows evidence for CENP-A–SNAP loading coincident with cytokinesis and nuclear envelope reformation. Video 1 shows a time lapse of early G1 CENP-A–SNAP assembly at centromeres corresponding to stills shown in Fig. 3 E. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200701066/DC1.
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Acknowledgments |
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This work has been supported by grants GM29513 and GM74150 from the National Institutes of Health (NIH) to D.W. Cleveland. The Neuroscience Microscopy Shared Facility is supported by grant NS047101 from the National Institute of Neurological Disorders and Stroke. L.E.T. Jansen has been supported by a postdoctoral fellowship from Philip Morris USA, Inc. B.E. Black has been supported by a postdoctoral fellowship from the American Cancer Society and by a Career Award in the Biomedical Sciences from the Burroughs Welcome Fund. D.R. Foltz has been supported by a postdoctoral fellowship from NIH and The Leukemia & Lymphoma Society. D.W. Cleveland receives salary support from the Ludwig Institute for Cancer Research.
Submitted: 11 January 2007
Accepted: 7 February 2007
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