Intersection of ChIP and FLIP, genomic methods to study the dynamics of the cohesin proteins
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- McNairn, A.J. & Gerton, J.L. Chromosome Res (2009) 17: 155. doi:10.1007/s10577-008-9007-9
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The evolutionarily conserved cohesin proteins Smc1, Smc3, Rad21 (Mcd1), and Scc3 function in the cohesin complex that provides the basis for chromosome cohesion and is involved in gene regulation. Understanding how these proteins link together the genome requires the use of whole-genome approaches to study the molecular mechanisms of these essential proteins. While chromatin immunoprecipitation followed by DNA microarray (ChIP-chip) studies have provided a snapshot in time of where these proteins associate with various genomes, the cohesin proteins are dynamic in their localization and interactions on chromatin. Study of the dynamic nature of these proteins requires approaches such as live cell imaging. We present evidence from fluorescence loss in photobleaching (FLIP) experiments in budding yeast that the decay constant of each cohesin subunit is ∼60–90 s in interphase. The decay constant on chromatin increases from G1 to S phase to metaphase, consistent with the interaction with chromatin becoming more stable once chromosomes are cohered. A small population of Smc3 at a position consistent with centromeric location has a longer decay constant than bulk Smc3. The characterization of the interaction of cohesin with chromatin, in terms of both its position and its dynamics, may be key to understanding how this protein complex contributes to chromosome segregation and gene regulation.
Keywordscohesin FLIP cell cycle SMC
enhanced green fluorescent protein
fluorescence loss in photobleaching
fluorescence recovery after photobleaching
fluorescence resonance energy transfer
mitotic chromosome determinant 1
precocious dissociation of sisters
sister chromatid cohesion
structural maintenance of chromosomes
From the time the chromosomes replicate until they separate at the metaphase-to-anaphase transition, sister chromatids are held together by a cohesive force. This cohesion is achieved via a protein complex known as the cohesin complex. The complex consists of four subunits, Smc1, Smc3, Scc3 (SA1, SA2), and Mcd1/Scc1 (Rad21), which are thought to form a ring that can hold sister chromatids together (Haering et al. 2008; Onn et al. 2008). However, many other proteins have been shown to be important for chromosome cohesion, including those involved in loading the cohesin complex onto DNA (Scc2, Scc4), those which couple cohesion to DNA replication (Eco1), those involved in maintaining cohesion (Pds5), and those involved in dissolving cohesion (separase, polo-like kinase). The collective activity of all of these factors can be thought of as the cohesin pathway. Most components of this pathway are evolutionarily conserved from budding yeast to human.
The question of where cohesin proteins localize within the genome is of foremost importance, as the binding of the cohesin complex to chromatin is critical for multiple cellular processes including chromosome cohesion, DNA damage repair, and proper gene regulation. The chromosomal sites at which cohesin associates with DNA have been mapped by ChIP-chip (chromatin immunoprecipitation followed by hybridization to DNA microarrays) in yeast, flies, and humans (Glynn et al. 2004; Lengronne et al. 2004; Misulovin et al. 2008; Parelho et al. 2008; Stedman 2008; Wendt et al. 2008) and by in-situ chromosome staining (McGuinness et al. 2005; Gomez et al. 2007). These studies revealed both similarities as well as disparities in the localization of cohesin proteins among different eukaryotes. One similarity is that centromeric regions show high levels of cohesin association, which presumably is important for chromosome segregation. Cohesin binding sites are also found throughout the chromosome arms spaced semi-periodically and can exhibit cell-specific chromosomal localizations that correlate with the epigenetic state of the chromatin. In budding yeast, cohesin is largely found in regions of convergent transcription and is absent from actively transcribed genes (Glynn et al. 2004; Lengronne et al. 2004). In contrast, fly cohesin binds preferentially to transcribed regions and overlaps with RNA polymerase II (Misulovin et al. 2008). In mammalian cells, most sites are in intergenic regions and introns (Wendt et al. 2008).
A number of observations suggest that although cohesin can reproducibly associate with the same regions of the genome each time it loads onto chromosomes, it does not recognize a specific sequence. In flies and yeast, there has been no motif significantly associated with the bound sequences (Glynn et al. 2004; Lengronne et al. 2004; Misulovin et al. 2008). In humans, cohesin binding along chromosome arms correlates with a consensus sequence that has been previously identified as the CTCF binding site (Parelho et al. 2008; Stedman et al. 2008; Wendt et al. 2008). CTCF is a conserved enhancer blocking protein found in vertebrates and is associated with a number of functions, including transcription, X-chromosome inactivation, and imprinting (Wallace and Felsenfeld 2007). The ability of CTCF to induce intra- and inter-chromosomal interactions may create regions of juxtaposed DNA to which cohesin may bind and stabilize. However, when CTCF is depleted, cohesin still appears to associate with DNA, but its binding is significantly delocalized, arguing that the site specificity is provided by the CTCF protein (Wendt et al. 2008). Thus, as is the case in lower eukaryotes, the cohesin complex itself is able to associate with DNA without sequence specificity in higher eukaryotes.
The position of the cohesin complex on DNA is altered under different gene expression programs. In yeasts, cohesin in chromosome arms is found predominantly in intergenic regions between convergent transcription units (Glynn et al. 2004; Lengronne et al. 2004). However, there are also some open reading frames where cohesin is found, of which some are inducible genes. When these genes are induced by heat shock, addition of galactose, or amino acid starvation, cohesin no longer localizes to these regions (Glynn et al. 2004; Lengronne et al. 2004; Bausch et al. 2007). Furthermore, when transcription is repressed, cohesin can return (Bausch et al. 2007). In flies, cohesin binds to the Abd-B homeobox gene in cells in which it is transcribed, but not in cells in which it is silenced (Misulovin et al. 2008). These results emphasize the lack of specificity for any particular DNA sequence, but also highlight that cohesin association with DNA is dynamic, depending on transcription.
The ChIP-chip experiments strongly suggest that the association of cohesin subunits with DNA is dynamic. The dynamics of the complex have been suggested to be controlled by a variety of proteins such as the maintenance subunit Pds5 (Losada et al. 2005) and the loading factor Scc2 (Rollins et al. 1999, 2004). For instance, in X. laevis egg extracts, mitotic chromosomes assembled in the absence of Pds5 have intact arm cohesion, but centromeric cohesion is loosened, although chromosomes retain an unusually high level of cohesin (Losada et al. 2005). Thus, Pds5 may play both positive and negative roles in sister chromatid cohesion, possibly by directly modulating the dynamic interaction of cohesin with chromatin. Alterations in the dynamics of the complex as caused by mutation of Scc2 have been suggested to affect the transcription of the cut locus in flies (Rollins et al. 2004). Understanding the dynamics of the complex may prove to be critical for understanding how the complex contributes to gene regulation.
Chromatin immunoprecipitation is a powerful method for gaining a high-resolution snap shot of where a protein of interest is located. The association of cohesin with a particular sequence can be followed at the resolution of a few hundred base pairs over a timescale of minutes to hours. However, this method is not optimal for capturing the behavior of a protein of interest on a very short timescale, such as seconds to minutes. Also, it measures where a protein of interest is located or enriched in a population of cells, but does not allow single cell measurements. A complementary approach to ChIP-chip experiments is live-imaging experiments. In these types of experiments, a protein of interest is fused to a fluorescent protein and the properties of the fusion protein can be assessed through bleaching the fluorescence with a laser and measuring how the protein behaves. These studies have the advantage of being able to monitor the behavior of a protein in a single cell over seconds to minutes (Rabut and Ellenberg 2005; Sprague and McNally 2005).
In order for sister chromatid cohesion to persist from the time sisters are replicated until mitosis, the linkage provided by cohesin would have to be stable over this period of time. However, many experiments have shown that cohesin can change locations, suggesting that its association with chromatin can be dynamic. In addition, a large fraction of cohesin on chromosomes arms dissociates during prophase in mammalian cells, while cohesin at the centromere remains until the metaphase-to-anaphase transition. In order to address how cohesin associates with chromatin during the cell cycle, Gerlich and colleagues performed a study in which a cohesin subunit, Scc3/SA1, was fused to EGFP and monitored by photobleaching and quantitative live-imaging by iFRAP (inverse fluorescence recovery after photobleaching) in the rat NRK cell line (Gerlich et al. 2006). They found three subpopulations of Scc3: (1) about 50% of Scc3 is soluble and highly dynamic, (2) about 30–50% is chromatin-bound with an exchange rate of 16 min, and (3) a fraction of 10–20%, present only in late S and G2 phase of the cell cycle, appears to be stably bound to chromatin, with an exchange rate of greater than 1 h (Gerlich et al. 2006). They speculate that the stably bound pool mediates sister chromatid cohesion. Consistent with this proposal, Bloom and colleagues have found evidence for a very stable cylindrical array of Smc3 in mitotic yeast nuclei that they propose organizes pericentric chromatin into an intramolecular loop (Yeh et al. 2008).
Since its discovery, cohesin has been studied intensively in budding yeast due to the ease of manipulation and the many tools available in this model organism. We have explored the dynamics of each cohesin subunit (Mcd1, Smc1, Smc3, and Scc3) and the accessory proteins Pds5 and Scc2 using fluorescence loss in photobleaching (FLIP) in budding yeast. Each protein was fused to GFP and its properties were monitored in interphase. We find that each subunit has a decay constant on the order of 1–1.5 min. The decay constant for Smc1 increases 4-fold from G1 phase to S phase to M-phase, consistent with the stability of the association with DNA increasing over the course of the cell cycle. Finally, a pool of Smc3 with slower kinetics may correspond to pericentric cohesin. These studies provide evidence that cohesin subunits are able to exchange on and off chromatin and potentially in and out of the complex in vivo on a timescale of much less than a cell cycle without the loss of chromosome cohesion.
Materials and methods
GFP-tagged yeast strains were obtained from the yeast GFP collection (Open Biosystems; Thermo Fisher Scientific, Huntsville, AL, USA). A dilution growth assay was used to determine whether the tagged proteins resulted in any defects in cell growth. For immunoprecipitation, whole-cell extracts were prepared as previously described (Camahort et al. 2007). Strains expressing Smc1-GFP, Smc1-GFP and Smc3-FLAG, Smc1-GFP and Mcd1-FLAG were utilized. The FLAG tags were introduced into the GFP strains using homologous recombination and standard transformation procedures. Immunoprecipitations were carried out overnight at 4°C using anti-FLAG M2 antibody (Sigma, St Louis, MO, USA). Immunocomplexes were precipitated using protein G agarose beads (GE Biosciences). Western blots were probed with an anti-GFP antibody (Clontech) to verify the expression of a full-length protein. The Smc3-GFP strain was transformed with a plasmid expressing Spc42-RFP (a gift from R. Li) in order to obtain the image in Fig. 3.
In fluorescence loss in photobleaching (FLIP) experiments, a single spot within the nucleus with a diameter of 0.2–0.5 μm was repeatedly bleached with one iteration of 100% power of a 488 nm laser line (50% output) using a Zeiss LSM510 Meta with 100X PlanFluor lens. The average diameter of a single z-axis plane of the yeast nucleus is ∼2–2.5 μm, resulting in approximately 10% of the nucleus being bleached with each pulse. Yeast strains were plated onto synthetic complete agar pads in hemi-spherical depression slides. Twenty pre-bleach images were collected and then images were captured before and after each bleach pulse, with 2.5 s between bleach pulses. After collecting 200 images, six separate measurements within the nucleus are taken to measure fluorescence loss (see Fig. 2B). Unbleached control cells were monitored for photobleaching due to image acquisition, and all measurements were normalized to the background fluorescence. Decay constants (the rate at which fluorescence is lost) for cohesin and control proteins were determined using OriginLab using nonlinear curve regression to a single exponential decay.
For cell cycle experiments, logarithmically growing yeast strains were synchronized using alpha-factor to achieve a G1-phase arrest and then released into either 0.2 μm hydroxyurea for S-phase arrest or 15 μg/ml nocodazole for mitotic arrest.
In vivo kinetics
In recent years, the development of optimized GFP variants and better microscope technology has led to the increased usage of photobleaching methods for study of the in vivo dynamics of chromatin-associated proteins. A large number of proteins have now been examined by photobleaching, demonstrating a wide variety of decay constants and differential kinetics (Phair et al. 2004). These methods have even been able to distinguish multiple populations of a tagged protein within the nucleus (Phair et al. 2004). An apparent trend seen in photobleaching experiments is that proteins involved in certain cellular processes, such as transcription or DNA replication, exhibit increased residence times when they are actively engaged (Dundr et al. 2002; Sporbert et al. 2002).
In addition to cohesin subunits, we used FLIP to examine the behavior of Dun1, a kinase involved in DNA damage response (Zhou and Elledge 1993). We examined the kinetics of the Dun1 protein to ensure that the power of the laser bleach pulse did not activate the DNA damage pathway. This is an important control, since the cohesin proteins have roles in DNA damage repair. The kinetics of Dun1 were unaffected by repeated bleach pulses, whereas the Dun1 kinetics change dramatically in response to hydroxyurea, which results in stalled forks and activation of the DNA damage response (decay constant increased from 113.54 ± 6.61 s to 246.16 ± 18.2 s). Thus, the bleach pulse alone is insufficient to activate the DNA damage response pathway.
Interphase kinetics of cohesin pathway proteinsa
Decay constant (s)
96 ± 3.5
55.4 ± 1.4
67.6 ± 1.8
91.6 ± 4.8
58.7 ± 3.3
70.9 ± 1.5
113.5 ± 6.6
Subnuclear populations of cohesin
Centromeric regions in yeast show high levels of cohesin association by ChIP-chip, whereas cohesin mapping at mammalian centromeres by ChIP has not yet been determined due to the highly repetitive alpha satellite DNA associated with mammalian centromeres. However, high levels of cohesin present at mammalian centromeres have been observed by microscopy (McGuinness et al. 2005; Gomez et al. 2007). Cohesin at the centromere may help resist the force of microtubules until all of the chromosomes are properly aligned, and this resistance may also assist in biorientation of sister chromatids.
The sites at which cohesin associates with the genome are of great interest, since they will very likely correlate with local sister cohesion and/or gene regulation. The high-resolution snapshot obtained from ChIP-chip studies enables us to appreciate that cohesin can be bound to a particular sequence in one transcriptional program and removed under a different transcriptional program. Thus, it is likely that cohesin can respond to changes in the chromatin environment. The dynamics of the association of cohesin with chromatin has been explored using live cell imaging. Photobleaching studies in mammalian cells have demonstrated that the cohesin subunits Scc3 and Mcd1 are capable of dynamically exchanging on and off the chromatin, with a decrease in rate as the cell cycle progresses (Gerlich et al. 2006). Here we confirm and extend these results for budding yeast, demonstrating a similar increase in the stability of the Smc1 association with chromatin over the cell cycle, which correlates well with the engagement of the cohesin complex in chromosome cohesion. We have examined the decay constant for all subunits of the complex and also for two accessory factors in the cohesin pathway. For each we measured a decay constant of ∼1–1.5 min, indicating the dynamic nature of these genome linker proteins. A small pool of pericentric Smc3 has slower kinetics. The picture of the cohesin pathway that emerges from the ChIP-chip and live cell imaging data suggests that neither alone will be sufficient to explain the molecular function of these proteins in gene regulation and cohesion. By combining the knowledge from these two types of experiments, a more holistic model of cohesin behavior may be elucidated.
Our results for the S. cerevisiae cohesin complex dynamics are mostly consistent with those previously reported for the human subunits (Gerlich et al. 2006). The immobile fraction, which they speculate corresponds to centromeric cohesin, had a residence time of several hours, or about 20% of the length of a cell cycle (24 h). Although we do not observe a pool of cohesin with this very long residence time, we do observe a pool of Smc3 that is less mobile and likely corresponds to pericentric cohesin. The freely diffusing population observed in the previous study, which we do not detect, may have been an artifact of the overexpression of the Scc3-GFP protein. Thus, the differential dynamics of pericentric cohesin as compared to arm cohesin, which appears to be evolutionarily conserved from yeast to human, might suggest that cohesin has different functions at these different locations. In addition, the large pool of cohesin subunits with short residence times suggests that cohesin subunits in a complex may be able to exchange, presumably without globally compromising sister cohesion.
Several factors have been identified that can alter the exchange rate of cohesin subunits. Depletion of Wapl, a protein that co-immunoprecipitates with the cohesin complex, by RNAi in HeLa cells results in an increase in the residence time of the ‘fast’ population of Scc1 (Kueng et al. 2006). In contrast, depletion of sororin reduces the amount of Smc1 that is stably bound to chromatin (Schmitz et al. 2007). Mutations in Scc2/Nipped-B result in the localized loss of cohesin and cohesin-mediated enhancer blocking at the cut gene locus in Drosophila (Rollins et al. 2004). Mutations in Scc2 are associated with the human disease Cornelia de Lange syndrome. It has been proposed that the mutations might alter the ability of the cohesin complex to interact with chromatin, which could affect local gene regulation (Dorsett 2004). It seems likely that elucidation of both the dynamics and the location of the cohesin subunits on DNA will provide the molecular clues needed to understand this human disease.
While it seems clear that cohesin can dynamically associate with chromatin and with specific regions of chromatin, the question remains as to what chromatin features cohesin recognizes and how its association influences gene regulation. Additional types of experiments will be necessary to address these questions. The use of genome conformation capture technology in specific cell types may prove useful for discerning which non-contiguous regions of the genome might be held together by cohesin to influence gene regulation. Genetic screens may also prove useful to identify pathways that act in parallel with cohesin for gene regulation.
We would like to thank Joel Schwartz, Scarlett Gard, and the Stowers Institute Imaging Center. We thank the Hudson Foundation for financial support.
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