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Chromatin movement visualized with photoactivable GFP-labeled histone H4
Differentiation (2008) 76:83–90 DOI: 10.1111/j.1432-0436.2007.00234.x
r 2007, Copyright the Authors Journal compilation r 2007, International Society of Differentiation
O RI G INA L AR T I C L E
Karien Wiesmeijer . Ilke M. Krouwels . Hans J. Tanke . Roeland W. Dirks
Chromatin movement visualized with photoactivable GFP-labeled histone H4
Received May 9, 2007; accepted in revised form August 22, 2007
Abstract The cell nucleus is highly organized with chromosomes occupying discrete, partially overlapping territories, and proteins that localize to specific nuclear compartments. This spatial organization of the nucleus is considered to be dynamic in response to environmental and cellular conditions to support changes in transcriptional programs. Chromatin, however, is relatively immobile when analyzed in living cells and shows a constrained Brownian type of movement. A possible explanation for this relative immobility is that chromatin interacts with a nuclear matrix structure and/or with nuclear compartments. Here, we explore the use of photoactivatable GFP fused to histone H4 as a potential tool to analyze the mobility of chromatin at various nuclear compartments. Selective photoactivation of photoactivatable-GFP at defined nuclear regions was achieved by two-photon excitation with 820 nm light. Nuclear speckles, which are considered storage sites of splicing factors, were visualized by coexpression of a fluorescent protein fused to splicing factor SF2/ASF. The results reveal a constrained chromatin motion, which is not affected by transcriptional inhibition, and suggests an intimate interaction of chromatin with speckles. Key words chromatin dynamics fluorescent protein two-photon microscopy photoactivation speckles live cell imaging
Karien Wiesmeijer Ilke M. Krouwels Hans J. Tanke . ) Roeland W. Dirks (* Department of Molecular Cell Biology Leiden University Medical Center Postal zone S1-P P.O. Box 9600, 2300 RC Leiden, The Netherlands Tel: 131 715269222 E-mail: [email protected]
Introduction The mammalian interphase nucleus is highly organized, which is reflected by the organization of chromosomes in discrete chromosome territories during interphase and the localization of many nuclear proteins to discrete nuclear compartments. Chromosome territories vary in size and shape, intermingle to some extent, and their spatial positioning in the nucleus appears to be nonrandom as gene-poor chromosomes locate toward the nuclear periphery and gene-rich chromosomes locate toward the nuclear interior (Croft et al., 1999; Boyle et al., 2001; Cremer et al., 2001, 2006; Branco and Pombo, 2006). This spatial three-dimensional positioning correlates well with the preferred peripheral localization of transcriptional silent chromatin in the cell nucleus (Andrulis et al., 1998) but also with local gene density (Goetze et al., 2007; Ku¨pper et al., 2007). In addition, specific spatial relationships among individual chromosome territories have been shown. As a consequence, translocations may occur more frequently among certain chromosome pairs than others (Roix et al., 2003). This spatial three-dimensional organization of chromosome territories is not static but dynamic and changes due to dramatic alterations in gene activity as a result of cell proliferation or differentiation (Bridger et al., 2000; Kim et al., 2004). However, other data indicate that chromosomes are predominantly randomly organized (Cornforth et al., 2002), suggesting that a strict order of chromosome neighbors is not required for proper cell functioning. Within individual chromosome territories, a variety of levels of chromatin organization can be discriminated, ranging from more condensed compact regions to open unfolded structures (van Driel et al., 2003). Some of the unfolded chromatin creates loops that emanate from the chromosome territory in a transcription-dependent way. The functional significance of such loops
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is not yet clear but they may provide genes that are positioned at these loops a better access to components of the transcription machinery (Volpi et al., 2000; Mahy et al., 2002). Transcriptional activity, however, is not limited to genes that have a position at the border or even outside the chromosome territory or subchromosomal domain. Active genes are also found distributed throughout chromosome territories. Nevertheless, it is assumed that some transcriptionally active genes are collectively arranged in functional transcription units or factories where their activities are controlled in a coordinated manner (Chakalova et al., 2005). Hence, more important than being at the edge or interior of a chromosome territory is the spatial positioning of genes in transcription-competent environments. For a number of genes, it was shown that their spatial localization in the nucleus changed after altering their transcriptional activity (Zink et al., 2004). Also, the positioning of transcriptionally active genes close to centromeric or peripheral heterochromatin leads to gene silencing (Brown et al., 1999; Francastel et al., 1999; Li et al., 2001). Thus, the spatial positioning of at least a subset genes is most likely dynamic in nature and is possibly directed by cellular activities, including transcription, RNA processing, and DNA repair. Consequently, nuclear positioning is a critical determinant of gene regulation (Kosak and Groudine, 2002; Cremer et al., 2004; Lanctot et al., 2007). An interesting question, yet to be answered, is whether associations of chromosome loci with nuclear compartments, like Cajal bodies, PML bodies, and speckles, could possibly contribute to the establishment of a nuclear organization and possibly to some nuclear functions as well. Histone genes as well as snRNA and snoRNA loci were shown to associate with Cajal bodies (Smith et al., 1995; Smith and Lawrence, 2000; Frey and Matera, 2001; Shopland et al., 2001) and consistent with these findings, also many active, non-characterized, transcription sites were shown to associate with Cajal bodies (Kielich et al., 2002), suggesting that these bodies play a more general role in facilitating transcription or RNA processing. Other chromatin regions were shown to associate preferentially with PML nuclear bodies rather than with Cajal bodies. Like Cajal bodies, however, there seems to be a preference for transcriptionally active chromatin regions to associate with PML bodies (LaMorte et al., 1998; Boisvert et al., 2000; Kielich et al., 2002; Wang et al., 2004). Interestingly, chromatin seems to establish true physical connections with PML bodies as shown for 10-nm chromatin fibers that make physical connections to the cores of PML bodies (Eskiw et al., 2004). Therefore, it has been proposed that PML bodies may serve as anchoring sites for euchromatin (Eskiw et al., 2004), consistent with the observation that PML bodies are nearly immobile in the nucleus (Wiesmeijer et al., 2002).
Most prevalent associations of gene loci with a nuclear compartment have, however, been found for nuclear speckles. Speckles, or SC-35 domains, are distinct irregular-shaped compartments containing poly(A) RNA, various RNA processing factors as well as many other factors as revealed by proteomic analysis (Saitoh et al., 2004). Many different genes and gene-rich R-bands on chromosomes have been found in association with speckles, which themselves are immobile in the nucleus (Shopland et al., 2003). Interestingly, also gene transcripts were shown to access these speckle domains, suggesting that these speckles play a role in RNA processing (Hattinger et al., 2002; Shopland et al., 2002). Together, these studies indicate that nuclear bodies function in close association with chromatin by which it may regulate nuclear functions. However, the nature and dynamics of associations between nuclear compartments and chromatin in live cells have thus far been poorly studied, which is also because of technical limitations. In this study, we show the applicability of photoactivatable GFP (PA-GFP) to study the movement of chromatin in the cell nucleus, also in relation to different nuclear compartments. For this purpose, we expressed histone H4 fused to PA-GFP in combination with a fusion protein that localizes specifically to a certain nuclear compartment in U2OS cells. Activation of PA-GFP is achieved by irradiation with 408 nm light or by two-photon excitation at 820 nm, leading to a threefold increase in fluorescence upon excitation with 488 nm light (Patterson and Lippincott-Schwartz, 2002). Chromatin movement is registered by taking time-lapse images at regular time intervals with 488 nm light. The advantage of the two-photon approach is that the PAGFP fusion protein can be irradiated selectively within a defined region or spot that is positioned anywhere in the three-dimensional space of a cell and that any phototoxicity in cells associated with single photon activation can be ruled out (Post et al., 2005).
Methods Generation of constructs The histone H4 (Z46261) sequence was amplified from a cDNA that was generated from human osteosarcoma cells (U2OS), using the forward primer 5 0 -GCGCGCGGTACCATGTCTGGTAGAGGCAAAGG-3 0 containing the KpnI site and the reverse primer 5 0 -GCGCGCCCCGGGTCAGCCACCAAAGCCGTACA-30 containing the XmaI site. Purified PCR fragments were inserted in-frame into the KpnI-XmaI fragment of pPA-GFP-C1 (Patterson and Lippincott-Schwartz, 2002). The coding sequence of SF2/ASF was originally cloned into the pEGFP-C1 vector (Molenaar et al., 2004) and subcloned into DsRed-Express (Clontech, Mountain View, CA). The plasmid EYFP-SUV39H1 has been described before (Krouwels et al., 2005).
85 Cell culture and transfection U2OS (human osteosarcoma) cells were cultured in 3.5 cm glass bottom Petri dishes (MatTek Corporation, Ashland, MA) in Dulbecco’s modified Eagle’s medium medium without phenol red, containing 1.0 mg/ml glucose, 4% fetal bovine serum, 2 mM glutamine, 100 U/ml penicillin, and 100 mg/ml streptomycin buffered with 25 mM Hepes to pH 7.2 (all from Invitrogen, Carlsbad, CA). Transient transfections were performed when cells were approximately at 70%–80% confluency using 1.5 ml Lipofectamine 2000 (Invitrogen) and 0.75–1.5 mg of DNA. To inhibit histone deacetylation, cells were incubated with 50 ng/ ml trichostatin A (TSA) (Sigma-Aldrich, St. Louis, MO) for 18– 22 hr. Transcription inhibition was accomplished by treating the cells with 30 mg/ml 5,6-dichloro-1--D-ribofuranosylbenzimidazole (DRB, Sigma-Aldrich) for 2 hr. DNA damage was introduced by incubating cells with 0.02% methyl methane-sulfonate (MMS) for time periods between 45 min and 2 hr (MMS, Sigma-Aldrich). A potential polymerization of nuclear actin was prevented by incubating cells with 4 mM latrunculin (Calbiochem, Merck Chemicals Ltd., Beeston, Nottingham, UK) for 2 hr at 371C.
Photoactivation and imaging of PA-GFP Photoactivation and subsequent time-lapse imaging were performed with a laser scanning confocal microscope (Leica TCS SP2, Leica Microsystems, Mannheim, Germany). The TCS SP2 is equipped with a two-photon Millenia Vs and a Tsunami laser (both Spectra-Physics, Mountain View, CA). During the course of the experiments, the temperature of the cells was maintained at 371C by a heating ring surrounding the culture chamber (Harvard App. Inc., Holliston, MA) and by an objective heater (Pecon GmbH, Erbach, Germany). The PA-GFP was activated by two-photon excitation using 820 nm light. The Millenia laser was set at 5 W. Following photoactivation, subsequent time-lapse images were taken with a 100 NA 1.4PL APO objective lens and 488 nm laser excitation. Fluorescence intensities and fluorescence distribution characteristics were measured using the Leica SP2 software.
Results Photoactivation of PA-GFP-histone H4 in U2OS cells To explore the potential of PA-GFP-histone H4 as a marker to study chromatin dynamics in the cell nucleus of living human cells, we transiently expressed this protein in U2OS cells. Following selective photoactivation of PA-GFP-histone H4 in the nucleus within a defined line with a mean width of 1 mm and in a selected image plane with 820 nm two-photon light, image stacks were taken at 1–10 min time intervals with 488 nm excitation light. Just before photoactivation, only a very low level of green fluorescence emission light was observed in PA-GFP-histone H4-expressing cells under 488 nm excitation (Fig. 1A, pre-activation), which is consistent with previous data (Patterson and Lippincott-Schwartz, 2002). Shortly after photoactivation, the selected line in the nucleus revealed a high level of fluorescence at 488 nm excitation (Fig. 1A, post-activation). The subsequent time-lapse images, which were taken each 10 min over a period of 1 hr, revealed that the line changed its shape, implying that chromatin gradually
dispersed away from the selected photoactivated line. Rather than a diffuse staining, defined structural chromatin conformations were observed as shown in Figure 1A. The maximum increase in width of the activated area that we measured after analyzing 17 cells at 2 hr after photoactivation was 2 mm. These observations confirm the incorporation of the transiently expressed PA-GFP-histone H4 into the chromatin structure because unincorporated fluorescent molecules would diffuse rapidly throughout the nucleoplasm. Both thread-like structures as well as more condensed focilike chromatin conformations were shown to move away from the photoactivated lines, suggesting that chromatin with different degrees of folding is equally mobile. The width of the stripe, however, did not significantly increase between 30 min and 6 hr after photoactivation, indicating that the movement of chromatin is constrained, which is consistent with previous observations (Gasser, 2002). However, we cannot exclude the possibility that some large-scale movements are taking place as well and that these are not detected due to resolution and sensitivity limits, which are inherent to this form of light microscopy. Some longrange directional movements of chromatin have been observed recently by tracking an integrated 256 copy lac operator repeat sequence (Chuang et al., 2006). Next, we wished to investigate whether the movement of chromatin is dependent on the transcriptional activity of a cell. Cells expressing PA-GFP-histone H4 were treated with the RNA polymerase II transcription inhibitor DRB and a defined line was photoactivated in the nucleus, resulting in bright fluorescent chromatin in this region at 488 nm excitation. Cells were monitored for 2 hr, taking images every 10 min. After 25 min, we could measure a small but insignificant difference in the width of the photoactivated line when comparing untreated with DRB-treated cells, indicating that small-scale chromatin movements are not dependent on ongoing transcription. Because chromatin mobility could possibly be dependent on cell cycle stage, we analyzed chromatin mobility in cells expressing both PA-GFP-histone H4 and proliferating cell nuclear antigen (PCNA)-CFP. PCNA is a component of the replication machinery and can be used as a cell cycle marker in living cells when fused to a fluorescent protein (Leonhardt et al., 2000). Following photoactivation of PA-GFP-histone H4, we observed no significant differences in chromatin movement between cells in G1, S, and G2 (result not shown).
Chromatin movement at nuclear compartments To investigate whether PA-GFP-histone H4 can be used as a tool to analyze chromatin mobility next to nuclear bodies, we selectively photoactivated chromatin that is immediately surrounding nucleoli or nuclear speckles.
86 Fig. 1. Chromatin movement in a U2OS cell expressing PA-GFP-histone H4. (A) After photoactivation of a line-shaped region at 820 nm in the nucleus of two cells using two-photon excitation, a fluorescent line is visible when the cells are imaged using 488 nm laser light. A subsequent time-lapse image that has been taken at 1 hr after photoactivation shows a more dispersed distribution of chromatin near the photoactivated region. A magnification of the region indicated by the box shows various levels of chromatin compaction. (B) A similar movement of chromatin is observed in PA-GFP-histone H4-expressing cells that have been treated with the transcription inhibitor DRB. Scale bar 5 10 mm.
Nucleoli could be clearly distinguished from the nucleoplasm by Nomarski imaging and by the absence of any fluorescent signal in PA-GFP-histone H4-transfected cells. As illustrated in Figure 2, chromatin that surrounded nucleoli could be selectively photoactivated using two-photon excitation and monitored over time by taking time-lapse images every 30 min up to 6 hr after photoactivation. The images show that although chromatin becomes more dispersed, it remains positioned near the photoactivated region. Some chromatin, however, remained in close association with the nucleolus, even at 6 hr after photoactivation. Similar observations were made for chromatin that surround nuclear speckles in cells expressing both PA-GFP-histone H4 and SF2/ASF-DsRED-Express. Chromatin was shown to move only within a confined region while a fraction of the photoactivated chromatin remained in close contact with speckle domains during the time course of the time-lapse experiment (Fig. 3A). Consistent with the idea that speckles do not contain DNA, we did not observe chromatin moving into speckles. Next, we wished to investigate whether this association of chromatin with speckle domains requires transcriptional activity. Cells expressing both PA-GFP histone H4 and SF2/ASF-DsRed-Express were incubated with DRB and a region surrounding a speckle was photoactivated. Subsequent time-lapse images re-
vealed that chromatin still became dispersed and that some chromatin remained in association with the speckle domains (Fig. 3B). Also, when chromatin within a line-shaped region just next to speckles was photoactivated, we observed that some chromatin remained in close association with speckles during the course of the experiment (Fig. 3C). These observations suggest that chromatin is not retracted from these domains when genes are transcriptionally silenced.
Fig. 2. Photoactivated PA-GFP-histone H4 reveals chromatin mobility next to nucleoli. After photoactivation of chromatin at regions that surround nucleoli, time-lapse images were taken every 30 min. The image taken at 6 hr after photoactivation shows a more
dispersed distribution of chromatin compared with the image taken just after photoactivation. A magnification of the indicated box shows the presence of various levels of chromatin compaction. The nucleoli are visible as black holes and are indicated by arrows.
Impact of different chromatin features on chromatin dynamics Thus far, our results suggest that neither the movement nor the association of chromatin with nuclear speckles is significantly influenced by changes in transcriptional activity in the chosen model system (U2OS cells). To investigate whether other factors may have an impact on chromatin movement, we imaged the movement of photoactivated chromatin in cells that were treated with one of the following agents: TSA, a histone deacetylase inhibitor; latrunculin, an inhibitor of nuclear actin polymerization; and MMS, a DNA-alkylating agent. Remarkably, none of these treatments revealed a significant change in chromatin mobility compared with control cells (Figs. 4A–4C). However, cells treated
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Fig. 3. Time-lapse images show chromatin movement at nuclear speckles in U2OS cells expressing PA-GFP-histone H4, SF2/ASFDsRED-Express, and proliferating cell nuclear antigen (PCNA)-CFP. (A) Time-lapse images show the dispersal of chromatin next to a speckle. The single confocal sections in the upper row show the homogeneous distribution of PCNA (cyan), indicating that this cell is in G1 or a G2, and the distribution of PA-GFP-histone H4 at 488 nm light (green). The images in the lower row show the distribution of photoactivated PA-GFPhistone H4 (green) relative to speckles (red). (B) Single confocal sections showing the expression of CFP-PCNA and the distribution of photoactivated PA-GFP-histone H4 relative to speckles (red) in a G1 or G2 cell that has been treated with the transcription inhibitor 5,6dichloro-1--D-ribofuranosylbenzimidazole. (C) A line, crossing the cell nucleus and hitting nuclear speckles (shown in red) was photoactivated at 820 nm using a two-photon laser and time-lapse images were recorded at 488 nm light. The single confocal sections suggest a tight association of chromatin (green) with nuclear speckles (red). The localization pattern of PCNA shows that the cell that has been photoactivated is either in G1 or in G2.
with latrunculin revealed a change in cellular and nuclear morphology, which made it more difficult to interpret the movement of photoactivated chromatin regions in time-lapse experiments. Because our previous data suggested that binding of the histone methyltransferase SUV39H to chromatin could possibly lead to a stabilization of heterochromatin (Krouwels et al., 2005), we analyzed the movement of photoactivated chromatin in cells expressing both PA-GFP histone H4 and CFP-SUV39H. The resulting time-lapse images revealed no significant change in chromatin movement compared with control cells expressing PA-GFP histone H4 only (Figs. 4D,4E). Hence, the binding of SUV39H to chromatin apparently does not lead to a more constrained chromatin movement. Expression of CFP-SUV39H is shown in Figure 4F. Together, these data suggest that changes in chromatin modifications have no significant impact on general chromatin mobility in U2OS cells.
Discussion In the interphase nucleus, chromatin is organized in higher order structures that are thought to be important to control gene activity. Heterochromatic structures, which are heavily methylated, are associated with gene transcriptional inactivity, and euchromatic structures, characterized by histone acetylation, are associated with gene expression. In addition, the spatial positioning of genomic regions in the nucleus has been linked to gene activity, suggesting the existence of micro-domains favoring transcription and possibly RNA processing. In order to adapt to changing conditions, one would expect that the spatial organization of the nucleus is dynamic including that of chromatin. Although various approaches have been used to study the dynamic nature of chromatin in the nucleus of living cells (Dirks and Tanke, 2006), most of them have the disadvantage that either only a small defined chromatin region can be analyzed or that only total chromatin can be analyzed.
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Fig. 4. Artificially induced changes in some chromatin properties have no significant impact on general chromatin mobility in U2OS cells. (A–C) Single confocal sections of cells that were imaged in the presence of the drugs trichostatin A, latrunculin, or methyl methanesulfonate and do not reveal a significant change in chromatin mobility following photoactivation compared with untreated cells (see Fig. 1). (D) Chromatin movement is recorded in a cell expressing both CFP-SUV39H and PA-GFP-histone H4. After photoactivation, chromatin is shown to disperse from the fluorescent line that has been recorded immediately after photoactivation by irradiation with 488 nm light. (E) A magnification of the indicated box at 120 min reveals some higher order chromatin structures. (F) This confocal section of the same cell as in D shows the expression of CFP-SUV39H.
These limitations can be overcome by the expression of a histone-PA GFP fusion protein and its subsequent photoactivation using one- or two-photon laser excitation (Post et al., 2005; Kruhlak et al., 2006). Because this fusion protein will in principle become incorporated into all nucleosomes and thereby into the full chromatin complement, any region of interest in a cell nucleus can be selectively photoactivated and monitored over a given time period. The advantage of two-photon activation rather than one-photon activation is that a defined volume within a three-dimensional space can be photoactivated and subsequently analyzed (Schneider et al., 2005). In this study, we show that this approach is of great potential interest for the analysis of chromatin move-
ment in the cell nucleus, in particular, where it concerns the potential association of chromatin with other nuclear compartments. Essential in this respect is that histone H4 becomes stably incorporated into nucleosomes and shows little exchange with the unbound pool of histone H4 proteins present in the surrounding nucleoplasm (Kimura and Cook, 2001). In this study, we monitored cells generally for up to 6 hr after photoactivation, but we also monitored cells for more extended time-periods and even overnight. Depending on the frequency by which the images were acquired, we observed some decrease in the fluorescence intensity of the photoactivated regions when cells were sampled for several hours. Nevertheless, the photoactivated regions could still be identified even at 16 hr after photoactiva-
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tion. When bleaching is limited, the degree of phototoxicity is expectedly low, an essential condition to ensure that what is being observed closely resembles physiological chromatin behavior. Indeed, we observed that cell division is not affected by photoactivation and subsequent live-cell imaging indicating limited phototoxicity. Hence, photoactivated PA-GFP-histone H4 provides a long-lasting mark for chromatin regions that have been selected and photoactivated. Our data show that chromatin movement is constrained, which is in agreement with numerous other studies (for a review, see Gasser, 2002). Interestingly, following photoactivation, some small chromatin foci (generally between 0.55 and 0.65 mm in size) could be discriminated, suggesting that chromatin is not completely unfolded when moving. These chromatin foci, which probably represent chromatin at different levels of compaction, are mobile but do not move over large distances. Chromatin motion is probably restricted by the physical structure of the chromatin molecule itself as well as by the presence of neighboring structures, which may either be chromatin or another nuclear compartment. When we analyzed the movement of chromatin in the direct vicinity of nucleoli or speckles, we observed no significant differences with chromatin movement at other nuclear regions. Some chromatin, however, maintained a close association with the nucleolus and revealed little mobility for at least several hours. This is in agreement with the observation that DNA loci containing a lac operator repeat sequences are less mobile at nucleoli than at other nucleoplasmic loci (Chubb et al., 2002). Similarly, some chromatin remained firmly associated with speckles. This firm association suggests that some chromatin might be physically attached to speckles. Consistent with this idea is the observation that some genes are preferentially associated with speckles and that transcripts of some of these genes are funneled in these domains (Smith et al., 1999; Johnson et al., 2000). Transcription inhibition, however, did not result in a detachment of chromatin from speckles, indicating that the association of chromatin with nuclear speckles is not dependent on ongoing transcription. Also, when we analyzed the mobility of chromatin in DRB-treated cells at different nuclear regions, we observed no significant difference compared with untreated cells, indicating that chromatin movements over short distances are not influenced by changes in transcriptional activity. Consistent with our data, it has recently been shown that the mobility of loci-containing lac operator repeats is independent of transcriptional activity (Mearini and Fackelmayer, 2006). Interestingly, the mobility of chromatin seems to be insensitive to changes in some properties of chromatin in general. Changes in the histone code did not reveal any influence on chromatin mobility. Also, a treatment of cells with the nuclear actin-depolymerizing drug latrunculin did not reveal significant changes in chromatin movement.
Together, these data suggest that chromatin mobility is largely restricted by the core structure of the chromatin fiber that may form, together with neighboring chromatin fibers, a nuclear scaffold. The association of chromatin with other nuclear compartments like nucleoli, speckles, and lamina may further impose restrictions on chromatin movement and may support the organization of chromatin in the cell nucleus as originally proposed by Abney et al., 1997). Acknowledgments We thank Dr. Jennifer Lippincott-Schwartz for kindly providing the PA-GFP-C1 plasmid and Dr. Cristina Cardoso for providing the GFP-PCNA plasmid. This work was supported by Cyttron grant no. BSIK03036.
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Mahy, N.L., Perry, P.E. and Bickmore, W.A. (2002) Gene density and transcription influence the localization of chromatin outside of chromosome territories detectable by FISH. J Cell Biol 159:753–763. Mearini, G. and Fackelmayer F, .O. (2006) Local chromatin mobility is independent of transcriptional activity. Cell Cycle 5:1989–1995. Molenaar, C., Abdulle, A., Gena, A., Tanke, H.J. and Dirks, R.W. (2004) Poly(A) rnas roam the cell nucleus and pass through speckle domains in transcriptionally active and inactive cells. J Cell Biol 165:191–202. Patterson, G.H. and Lippincott-Schwartz, J. (2002) A photoactivatable GFP for selective photolabeling of proteins and cells. Science 297:1873–1877. Post, J.N., Lidke, K.A., Rieger, B. and Arndt-Jovin, D.J. (2005) One—and two-photon photoactivation of a paGFP-fusion protein in live Drosophila embryos. FEBS Lett 579:325–330. Roix, J.J., McQueen, P.G., Munson, P.J., Parada, L.A. and Misteli, T. (2003) Spatial proximity of translocation-prone gene loci in human lymphomas. Nat Genet 34:287–291. Saitoh, N., Spahr, C.S., Patterson, S.D., Bubulya, P., Neuwald, A.F. and Spector, D.L. (2004) Proteomic analysis of interchromatin granule clusters. Mol Biol Cell 15:3876–3890. Schneider, M., Barozzi, S., Testa, I., Faretta, M. and Diaspro, A. (2005) Two-photon activation and excitation properties of PAGFP in the 720–920 nm range. Biophys J 89:1346–1352. Shopland, L.S., Byron, M., Stein, J.L., Lian, J.B., Stein, G.S. and Lawrence, J.B. (2001) Replication-dependent histone gene expresson is related to Cajal body (CB) association but does not require sustained CB contact. Mol Biol Cell 12:565–576. Shopland, L.S., Johnson, C.V., Byron, M., McNeil, J. and Lawrence, J.B. (2003) Clustering of multiple specific genes and generich R-bands around SC-35 domains: evidence for local euchromatic neighborhoods. J Cell Biol 162:981–990. Shopland, L.S., Johnson, C.V. and Lawrence, J.B. (2002) Evidence that all SC-35 domains contain mRNAs and that transcripts can be structurally constrained within these domains. J Struct Biol 140:131–139. Smith, K.P., Carter, K.C., Johnson, C.V. and Lawrence, J.B. (1995) U2 and U1 snRNA gene loci associate with coiled bodies. J Cell Biochem 59:473–485. Smith, K.P. and Lawrence, J.B. (2000) Interactions of U2 gene loci and their nuclear transcripts with Cajal (coiled) bodies: evidence for preU2 within Cajal bodies. Mol Biol Cell 11:2987–2998. Smith, K.P., Moen, P.T., Wydner, K.L., Coleman, J.R. and Lawrence, J.B. (1999) Processing of endogenous pre-mrnas in association with SC-35 domains is gene specific. J Cell Biol 144:617– 629. van Driel, R., Fransz, P.F. and Verschure, P.J. (2003) The eukaryotic genome: a system regulated at different hierarchical levels. J Cell Sci 116:4067–4075. Volpi, E.V., Chevret, E., Jones, T., Vatcheva, R., Williamson, J., Beck, S., Campbell, R.D., Goldsworthy, M., Powis, S.H., Ragoussis, J., Trowsdale, J. and Sheer, D. (2000) Large-scale chromatin organization of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei. J Cell Sci 113:1565–1576. Wang, J., Shiels, C., Sasieni, P., Wu, P.J., Islam, S.A., Freemont, P.S. and Sheer, D. (2004) Promyelocytic leukemia nuclear bodies associate with transcriptionally active genomic regions. J Cell Biol 164:515–526. Wiesmeijer, K., Molenaar, C., Bekeer, I.M.L.A., Tanke, H.J. and Dirks, R.W. (2002) Mobile foci of Sp100 do not contain PML: PML bodies are immobile but PML and Sp100 proteins are not. J Struct Biol 140:180–188. Zink, D., Amaral, M.D., Englmann, A., Lang, S., Clarke, L.A., Rudolph, C., Alt, F., Luther, K., Braz, C., Sadoni, N., Rosenecker, J. and Schindelhauer, D. (2004) Transcription-dependent spatial arrangements of CFTR and adjacent genes in human cell nuclei. J Cell Biol 166:815–825.
r 2007, Copyright the Authors Journal compilation r 2007, International Society of Differentiation
O RI G INA L AR T I C L E
Karien Wiesmeijer . Ilke M. Krouwels . Hans J. Tanke . Roeland W. Dirks
Chromatin movement visualized with photoactivable GFP-labeled histone H4
Received May 9, 2007; accepted in revised form August 22, 2007
Abstract The cell nucleus is highly organized with chromosomes occupying discrete, partially overlapping territories, and proteins that localize to specific nuclear compartments. This spatial organization of the nucleus is considered to be dynamic in response to environmental and cellular conditions to support changes in transcriptional programs. Chromatin, however, is relatively immobile when analyzed in living cells and shows a constrained Brownian type of movement. A possible explanation for this relative immobility is that chromatin interacts with a nuclear matrix structure and/or with nuclear compartments. Here, we explore the use of photoactivatable GFP fused to histone H4 as a potential tool to analyze the mobility of chromatin at various nuclear compartments. Selective photoactivation of photoactivatable-GFP at defined nuclear regions was achieved by two-photon excitation with 820 nm light. Nuclear speckles, which are considered storage sites of splicing factors, were visualized by coexpression of a fluorescent protein fused to splicing factor SF2/ASF. The results reveal a constrained chromatin motion, which is not affected by transcriptional inhibition, and suggests an intimate interaction of chromatin with speckles. Key words chromatin dynamics fluorescent protein two-photon microscopy photoactivation speckles live cell imaging
Karien Wiesmeijer Ilke M. Krouwels Hans J. Tanke . ) Roeland W. Dirks (* Department of Molecular Cell Biology Leiden University Medical Center Postal zone S1-P P.O. Box 9600, 2300 RC Leiden, The Netherlands Tel: 131 715269222 E-mail: [email protected]
Introduction The mammalian interphase nucleus is highly organized, which is reflected by the organization of chromosomes in discrete chromosome territories during interphase and the localization of many nuclear proteins to discrete nuclear compartments. Chromosome territories vary in size and shape, intermingle to some extent, and their spatial positioning in the nucleus appears to be nonrandom as gene-poor chromosomes locate toward the nuclear periphery and gene-rich chromosomes locate toward the nuclear interior (Croft et al., 1999; Boyle et al., 2001; Cremer et al., 2001, 2006; Branco and Pombo, 2006). This spatial three-dimensional positioning correlates well with the preferred peripheral localization of transcriptional silent chromatin in the cell nucleus (Andrulis et al., 1998) but also with local gene density (Goetze et al., 2007; Ku¨pper et al., 2007). In addition, specific spatial relationships among individual chromosome territories have been shown. As a consequence, translocations may occur more frequently among certain chromosome pairs than others (Roix et al., 2003). This spatial three-dimensional organization of chromosome territories is not static but dynamic and changes due to dramatic alterations in gene activity as a result of cell proliferation or differentiation (Bridger et al., 2000; Kim et al., 2004). However, other data indicate that chromosomes are predominantly randomly organized (Cornforth et al., 2002), suggesting that a strict order of chromosome neighbors is not required for proper cell functioning. Within individual chromosome territories, a variety of levels of chromatin organization can be discriminated, ranging from more condensed compact regions to open unfolded structures (van Driel et al., 2003). Some of the unfolded chromatin creates loops that emanate from the chromosome territory in a transcription-dependent way. The functional significance of such loops
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is not yet clear but they may provide genes that are positioned at these loops a better access to components of the transcription machinery (Volpi et al., 2000; Mahy et al., 2002). Transcriptional activity, however, is not limited to genes that have a position at the border or even outside the chromosome territory or subchromosomal domain. Active genes are also found distributed throughout chromosome territories. Nevertheless, it is assumed that some transcriptionally active genes are collectively arranged in functional transcription units or factories where their activities are controlled in a coordinated manner (Chakalova et al., 2005). Hence, more important than being at the edge or interior of a chromosome territory is the spatial positioning of genes in transcription-competent environments. For a number of genes, it was shown that their spatial localization in the nucleus changed after altering their transcriptional activity (Zink et al., 2004). Also, the positioning of transcriptionally active genes close to centromeric or peripheral heterochromatin leads to gene silencing (Brown et al., 1999; Francastel et al., 1999; Li et al., 2001). Thus, the spatial positioning of at least a subset genes is most likely dynamic in nature and is possibly directed by cellular activities, including transcription, RNA processing, and DNA repair. Consequently, nuclear positioning is a critical determinant of gene regulation (Kosak and Groudine, 2002; Cremer et al., 2004; Lanctot et al., 2007). An interesting question, yet to be answered, is whether associations of chromosome loci with nuclear compartments, like Cajal bodies, PML bodies, and speckles, could possibly contribute to the establishment of a nuclear organization and possibly to some nuclear functions as well. Histone genes as well as snRNA and snoRNA loci were shown to associate with Cajal bodies (Smith et al., 1995; Smith and Lawrence, 2000; Frey and Matera, 2001; Shopland et al., 2001) and consistent with these findings, also many active, non-characterized, transcription sites were shown to associate with Cajal bodies (Kielich et al., 2002), suggesting that these bodies play a more general role in facilitating transcription or RNA processing. Other chromatin regions were shown to associate preferentially with PML nuclear bodies rather than with Cajal bodies. Like Cajal bodies, however, there seems to be a preference for transcriptionally active chromatin regions to associate with PML bodies (LaMorte et al., 1998; Boisvert et al., 2000; Kielich et al., 2002; Wang et al., 2004). Interestingly, chromatin seems to establish true physical connections with PML bodies as shown for 10-nm chromatin fibers that make physical connections to the cores of PML bodies (Eskiw et al., 2004). Therefore, it has been proposed that PML bodies may serve as anchoring sites for euchromatin (Eskiw et al., 2004), consistent with the observation that PML bodies are nearly immobile in the nucleus (Wiesmeijer et al., 2002).
Most prevalent associations of gene loci with a nuclear compartment have, however, been found for nuclear speckles. Speckles, or SC-35 domains, are distinct irregular-shaped compartments containing poly(A) RNA, various RNA processing factors as well as many other factors as revealed by proteomic analysis (Saitoh et al., 2004). Many different genes and gene-rich R-bands on chromosomes have been found in association with speckles, which themselves are immobile in the nucleus (Shopland et al., 2003). Interestingly, also gene transcripts were shown to access these speckle domains, suggesting that these speckles play a role in RNA processing (Hattinger et al., 2002; Shopland et al., 2002). Together, these studies indicate that nuclear bodies function in close association with chromatin by which it may regulate nuclear functions. However, the nature and dynamics of associations between nuclear compartments and chromatin in live cells have thus far been poorly studied, which is also because of technical limitations. In this study, we show the applicability of photoactivatable GFP (PA-GFP) to study the movement of chromatin in the cell nucleus, also in relation to different nuclear compartments. For this purpose, we expressed histone H4 fused to PA-GFP in combination with a fusion protein that localizes specifically to a certain nuclear compartment in U2OS cells. Activation of PA-GFP is achieved by irradiation with 408 nm light or by two-photon excitation at 820 nm, leading to a threefold increase in fluorescence upon excitation with 488 nm light (Patterson and Lippincott-Schwartz, 2002). Chromatin movement is registered by taking time-lapse images at regular time intervals with 488 nm light. The advantage of the two-photon approach is that the PAGFP fusion protein can be irradiated selectively within a defined region or spot that is positioned anywhere in the three-dimensional space of a cell and that any phototoxicity in cells associated with single photon activation can be ruled out (Post et al., 2005).
Methods Generation of constructs The histone H4 (Z46261) sequence was amplified from a cDNA that was generated from human osteosarcoma cells (U2OS), using the forward primer 5 0 -GCGCGCGGTACCATGTCTGGTAGAGGCAAAGG-3 0 containing the KpnI site and the reverse primer 5 0 -GCGCGCCCCGGGTCAGCCACCAAAGCCGTACA-30 containing the XmaI site. Purified PCR fragments were inserted in-frame into the KpnI-XmaI fragment of pPA-GFP-C1 (Patterson and Lippincott-Schwartz, 2002). The coding sequence of SF2/ASF was originally cloned into the pEGFP-C1 vector (Molenaar et al., 2004) and subcloned into DsRed-Express (Clontech, Mountain View, CA). The plasmid EYFP-SUV39H1 has been described before (Krouwels et al., 2005).
85 Cell culture and transfection U2OS (human osteosarcoma) cells were cultured in 3.5 cm glass bottom Petri dishes (MatTek Corporation, Ashland, MA) in Dulbecco’s modified Eagle’s medium medium without phenol red, containing 1.0 mg/ml glucose, 4% fetal bovine serum, 2 mM glutamine, 100 U/ml penicillin, and 100 mg/ml streptomycin buffered with 25 mM Hepes to pH 7.2 (all from Invitrogen, Carlsbad, CA). Transient transfections were performed when cells were approximately at 70%–80% confluency using 1.5 ml Lipofectamine 2000 (Invitrogen) and 0.75–1.5 mg of DNA. To inhibit histone deacetylation, cells were incubated with 50 ng/ ml trichostatin A (TSA) (Sigma-Aldrich, St. Louis, MO) for 18– 22 hr. Transcription inhibition was accomplished by treating the cells with 30 mg/ml 5,6-dichloro-1--D-ribofuranosylbenzimidazole (DRB, Sigma-Aldrich) for 2 hr. DNA damage was introduced by incubating cells with 0.02% methyl methane-sulfonate (MMS) for time periods between 45 min and 2 hr (MMS, Sigma-Aldrich). A potential polymerization of nuclear actin was prevented by incubating cells with 4 mM latrunculin (Calbiochem, Merck Chemicals Ltd., Beeston, Nottingham, UK) for 2 hr at 371C.
Photoactivation and imaging of PA-GFP Photoactivation and subsequent time-lapse imaging were performed with a laser scanning confocal microscope (Leica TCS SP2, Leica Microsystems, Mannheim, Germany). The TCS SP2 is equipped with a two-photon Millenia Vs and a Tsunami laser (both Spectra-Physics, Mountain View, CA). During the course of the experiments, the temperature of the cells was maintained at 371C by a heating ring surrounding the culture chamber (Harvard App. Inc., Holliston, MA) and by an objective heater (Pecon GmbH, Erbach, Germany). The PA-GFP was activated by two-photon excitation using 820 nm light. The Millenia laser was set at 5 W. Following photoactivation, subsequent time-lapse images were taken with a 100 NA 1.4PL APO objective lens and 488 nm laser excitation. Fluorescence intensities and fluorescence distribution characteristics were measured using the Leica SP2 software.
Results Photoactivation of PA-GFP-histone H4 in U2OS cells To explore the potential of PA-GFP-histone H4 as a marker to study chromatin dynamics in the cell nucleus of living human cells, we transiently expressed this protein in U2OS cells. Following selective photoactivation of PA-GFP-histone H4 in the nucleus within a defined line with a mean width of 1 mm and in a selected image plane with 820 nm two-photon light, image stacks were taken at 1–10 min time intervals with 488 nm excitation light. Just before photoactivation, only a very low level of green fluorescence emission light was observed in PA-GFP-histone H4-expressing cells under 488 nm excitation (Fig. 1A, pre-activation), which is consistent with previous data (Patterson and Lippincott-Schwartz, 2002). Shortly after photoactivation, the selected line in the nucleus revealed a high level of fluorescence at 488 nm excitation (Fig. 1A, post-activation). The subsequent time-lapse images, which were taken each 10 min over a period of 1 hr, revealed that the line changed its shape, implying that chromatin gradually
dispersed away from the selected photoactivated line. Rather than a diffuse staining, defined structural chromatin conformations were observed as shown in Figure 1A. The maximum increase in width of the activated area that we measured after analyzing 17 cells at 2 hr after photoactivation was 2 mm. These observations confirm the incorporation of the transiently expressed PA-GFP-histone H4 into the chromatin structure because unincorporated fluorescent molecules would diffuse rapidly throughout the nucleoplasm. Both thread-like structures as well as more condensed focilike chromatin conformations were shown to move away from the photoactivated lines, suggesting that chromatin with different degrees of folding is equally mobile. The width of the stripe, however, did not significantly increase between 30 min and 6 hr after photoactivation, indicating that the movement of chromatin is constrained, which is consistent with previous observations (Gasser, 2002). However, we cannot exclude the possibility that some large-scale movements are taking place as well and that these are not detected due to resolution and sensitivity limits, which are inherent to this form of light microscopy. Some longrange directional movements of chromatin have been observed recently by tracking an integrated 256 copy lac operator repeat sequence (Chuang et al., 2006). Next, we wished to investigate whether the movement of chromatin is dependent on the transcriptional activity of a cell. Cells expressing PA-GFP-histone H4 were treated with the RNA polymerase II transcription inhibitor DRB and a defined line was photoactivated in the nucleus, resulting in bright fluorescent chromatin in this region at 488 nm excitation. Cells were monitored for 2 hr, taking images every 10 min. After 25 min, we could measure a small but insignificant difference in the width of the photoactivated line when comparing untreated with DRB-treated cells, indicating that small-scale chromatin movements are not dependent on ongoing transcription. Because chromatin mobility could possibly be dependent on cell cycle stage, we analyzed chromatin mobility in cells expressing both PA-GFP-histone H4 and proliferating cell nuclear antigen (PCNA)-CFP. PCNA is a component of the replication machinery and can be used as a cell cycle marker in living cells when fused to a fluorescent protein (Leonhardt et al., 2000). Following photoactivation of PA-GFP-histone H4, we observed no significant differences in chromatin movement between cells in G1, S, and G2 (result not shown).
Chromatin movement at nuclear compartments To investigate whether PA-GFP-histone H4 can be used as a tool to analyze chromatin mobility next to nuclear bodies, we selectively photoactivated chromatin that is immediately surrounding nucleoli or nuclear speckles.
86 Fig. 1. Chromatin movement in a U2OS cell expressing PA-GFP-histone H4. (A) After photoactivation of a line-shaped region at 820 nm in the nucleus of two cells using two-photon excitation, a fluorescent line is visible when the cells are imaged using 488 nm laser light. A subsequent time-lapse image that has been taken at 1 hr after photoactivation shows a more dispersed distribution of chromatin near the photoactivated region. A magnification of the region indicated by the box shows various levels of chromatin compaction. (B) A similar movement of chromatin is observed in PA-GFP-histone H4-expressing cells that have been treated with the transcription inhibitor DRB. Scale bar 5 10 mm.
Nucleoli could be clearly distinguished from the nucleoplasm by Nomarski imaging and by the absence of any fluorescent signal in PA-GFP-histone H4-transfected cells. As illustrated in Figure 2, chromatin that surrounded nucleoli could be selectively photoactivated using two-photon excitation and monitored over time by taking time-lapse images every 30 min up to 6 hr after photoactivation. The images show that although chromatin becomes more dispersed, it remains positioned near the photoactivated region. Some chromatin, however, remained in close association with the nucleolus, even at 6 hr after photoactivation. Similar observations were made for chromatin that surround nuclear speckles in cells expressing both PA-GFP-histone H4 and SF2/ASF-DsRED-Express. Chromatin was shown to move only within a confined region while a fraction of the photoactivated chromatin remained in close contact with speckle domains during the time course of the time-lapse experiment (Fig. 3A). Consistent with the idea that speckles do not contain DNA, we did not observe chromatin moving into speckles. Next, we wished to investigate whether this association of chromatin with speckle domains requires transcriptional activity. Cells expressing both PA-GFP histone H4 and SF2/ASF-DsRed-Express were incubated with DRB and a region surrounding a speckle was photoactivated. Subsequent time-lapse images re-
vealed that chromatin still became dispersed and that some chromatin remained in association with the speckle domains (Fig. 3B). Also, when chromatin within a line-shaped region just next to speckles was photoactivated, we observed that some chromatin remained in close association with speckles during the course of the experiment (Fig. 3C). These observations suggest that chromatin is not retracted from these domains when genes are transcriptionally silenced.
Fig. 2. Photoactivated PA-GFP-histone H4 reveals chromatin mobility next to nucleoli. After photoactivation of chromatin at regions that surround nucleoli, time-lapse images were taken every 30 min. The image taken at 6 hr after photoactivation shows a more
dispersed distribution of chromatin compared with the image taken just after photoactivation. A magnification of the indicated box shows the presence of various levels of chromatin compaction. The nucleoli are visible as black holes and are indicated by arrows.
Impact of different chromatin features on chromatin dynamics Thus far, our results suggest that neither the movement nor the association of chromatin with nuclear speckles is significantly influenced by changes in transcriptional activity in the chosen model system (U2OS cells). To investigate whether other factors may have an impact on chromatin movement, we imaged the movement of photoactivated chromatin in cells that were treated with one of the following agents: TSA, a histone deacetylase inhibitor; latrunculin, an inhibitor of nuclear actin polymerization; and MMS, a DNA-alkylating agent. Remarkably, none of these treatments revealed a significant change in chromatin mobility compared with control cells (Figs. 4A–4C). However, cells treated
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Fig. 3. Time-lapse images show chromatin movement at nuclear speckles in U2OS cells expressing PA-GFP-histone H4, SF2/ASFDsRED-Express, and proliferating cell nuclear antigen (PCNA)-CFP. (A) Time-lapse images show the dispersal of chromatin next to a speckle. The single confocal sections in the upper row show the homogeneous distribution of PCNA (cyan), indicating that this cell is in G1 or a G2, and the distribution of PA-GFP-histone H4 at 488 nm light (green). The images in the lower row show the distribution of photoactivated PA-GFPhistone H4 (green) relative to speckles (red). (B) Single confocal sections showing the expression of CFP-PCNA and the distribution of photoactivated PA-GFP-histone H4 relative to speckles (red) in a G1 or G2 cell that has been treated with the transcription inhibitor 5,6dichloro-1--D-ribofuranosylbenzimidazole. (C) A line, crossing the cell nucleus and hitting nuclear speckles (shown in red) was photoactivated at 820 nm using a two-photon laser and time-lapse images were recorded at 488 nm light. The single confocal sections suggest a tight association of chromatin (green) with nuclear speckles (red). The localization pattern of PCNA shows that the cell that has been photoactivated is either in G1 or in G2.
with latrunculin revealed a change in cellular and nuclear morphology, which made it more difficult to interpret the movement of photoactivated chromatin regions in time-lapse experiments. Because our previous data suggested that binding of the histone methyltransferase SUV39H to chromatin could possibly lead to a stabilization of heterochromatin (Krouwels et al., 2005), we analyzed the movement of photoactivated chromatin in cells expressing both PA-GFP histone H4 and CFP-SUV39H. The resulting time-lapse images revealed no significant change in chromatin movement compared with control cells expressing PA-GFP histone H4 only (Figs. 4D,4E). Hence, the binding of SUV39H to chromatin apparently does not lead to a more constrained chromatin movement. Expression of CFP-SUV39H is shown in Figure 4F. Together, these data suggest that changes in chromatin modifications have no significant impact on general chromatin mobility in U2OS cells.
Discussion In the interphase nucleus, chromatin is organized in higher order structures that are thought to be important to control gene activity. Heterochromatic structures, which are heavily methylated, are associated with gene transcriptional inactivity, and euchromatic structures, characterized by histone acetylation, are associated with gene expression. In addition, the spatial positioning of genomic regions in the nucleus has been linked to gene activity, suggesting the existence of micro-domains favoring transcription and possibly RNA processing. In order to adapt to changing conditions, one would expect that the spatial organization of the nucleus is dynamic including that of chromatin. Although various approaches have been used to study the dynamic nature of chromatin in the nucleus of living cells (Dirks and Tanke, 2006), most of them have the disadvantage that either only a small defined chromatin region can be analyzed or that only total chromatin can be analyzed.
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Fig. 4. Artificially induced changes in some chromatin properties have no significant impact on general chromatin mobility in U2OS cells. (A–C) Single confocal sections of cells that were imaged in the presence of the drugs trichostatin A, latrunculin, or methyl methanesulfonate and do not reveal a significant change in chromatin mobility following photoactivation compared with untreated cells (see Fig. 1). (D) Chromatin movement is recorded in a cell expressing both CFP-SUV39H and PA-GFP-histone H4. After photoactivation, chromatin is shown to disperse from the fluorescent line that has been recorded immediately after photoactivation by irradiation with 488 nm light. (E) A magnification of the indicated box at 120 min reveals some higher order chromatin structures. (F) This confocal section of the same cell as in D shows the expression of CFP-SUV39H.
These limitations can be overcome by the expression of a histone-PA GFP fusion protein and its subsequent photoactivation using one- or two-photon laser excitation (Post et al., 2005; Kruhlak et al., 2006). Because this fusion protein will in principle become incorporated into all nucleosomes and thereby into the full chromatin complement, any region of interest in a cell nucleus can be selectively photoactivated and monitored over a given time period. The advantage of two-photon activation rather than one-photon activation is that a defined volume within a three-dimensional space can be photoactivated and subsequently analyzed (Schneider et al., 2005). In this study, we show that this approach is of great potential interest for the analysis of chromatin move-
ment in the cell nucleus, in particular, where it concerns the potential association of chromatin with other nuclear compartments. Essential in this respect is that histone H4 becomes stably incorporated into nucleosomes and shows little exchange with the unbound pool of histone H4 proteins present in the surrounding nucleoplasm (Kimura and Cook, 2001). In this study, we monitored cells generally for up to 6 hr after photoactivation, but we also monitored cells for more extended time-periods and even overnight. Depending on the frequency by which the images were acquired, we observed some decrease in the fluorescence intensity of the photoactivated regions when cells were sampled for several hours. Nevertheless, the photoactivated regions could still be identified even at 16 hr after photoactiva-
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tion. When bleaching is limited, the degree of phototoxicity is expectedly low, an essential condition to ensure that what is being observed closely resembles physiological chromatin behavior. Indeed, we observed that cell division is not affected by photoactivation and subsequent live-cell imaging indicating limited phototoxicity. Hence, photoactivated PA-GFP-histone H4 provides a long-lasting mark for chromatin regions that have been selected and photoactivated. Our data show that chromatin movement is constrained, which is in agreement with numerous other studies (for a review, see Gasser, 2002). Interestingly, following photoactivation, some small chromatin foci (generally between 0.55 and 0.65 mm in size) could be discriminated, suggesting that chromatin is not completely unfolded when moving. These chromatin foci, which probably represent chromatin at different levels of compaction, are mobile but do not move over large distances. Chromatin motion is probably restricted by the physical structure of the chromatin molecule itself as well as by the presence of neighboring structures, which may either be chromatin or another nuclear compartment. When we analyzed the movement of chromatin in the direct vicinity of nucleoli or speckles, we observed no significant differences with chromatin movement at other nuclear regions. Some chromatin, however, maintained a close association with the nucleolus and revealed little mobility for at least several hours. This is in agreement with the observation that DNA loci containing a lac operator repeat sequences are less mobile at nucleoli than at other nucleoplasmic loci (Chubb et al., 2002). Similarly, some chromatin remained firmly associated with speckles. This firm association suggests that some chromatin might be physically attached to speckles. Consistent with this idea is the observation that some genes are preferentially associated with speckles and that transcripts of some of these genes are funneled in these domains (Smith et al., 1999; Johnson et al., 2000). Transcription inhibition, however, did not result in a detachment of chromatin from speckles, indicating that the association of chromatin with nuclear speckles is not dependent on ongoing transcription. Also, when we analyzed the mobility of chromatin in DRB-treated cells at different nuclear regions, we observed no significant difference compared with untreated cells, indicating that chromatin movements over short distances are not influenced by changes in transcriptional activity. Consistent with our data, it has recently been shown that the mobility of loci-containing lac operator repeats is independent of transcriptional activity (Mearini and Fackelmayer, 2006). Interestingly, the mobility of chromatin seems to be insensitive to changes in some properties of chromatin in general. Changes in the histone code did not reveal any influence on chromatin mobility. Also, a treatment of cells with the nuclear actin-depolymerizing drug latrunculin did not reveal significant changes in chromatin movement.
Together, these data suggest that chromatin mobility is largely restricted by the core structure of the chromatin fiber that may form, together with neighboring chromatin fibers, a nuclear scaffold. The association of chromatin with other nuclear compartments like nucleoli, speckles, and lamina may further impose restrictions on chromatin movement and may support the organization of chromatin in the cell nucleus as originally proposed by Abney et al., 1997). Acknowledgments We thank Dr. Jennifer Lippincott-Schwartz for kindly providing the PA-GFP-C1 plasmid and Dr. Cristina Cardoso for providing the GFP-PCNA plasmid. This work was supported by Cyttron grant no. BSIK03036.
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