Correspondence to Roeland W. Dirks: r.w.dirks{at}lumc.nl
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Abbreviations used in this paper: 5-aza-C, 5-aza-2'deoxycytidine; BP, bandpass; dn, double null; FLIM, fluorescence lifetime imaging microscopy; FRET, fluorescence resonance energy transfer; HP1, heterochromatin protein 1; PMEF, primary mouse embryonic fibroblast; TSA, trichostatin A.
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Introduction |
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The histone methyltransferase SUV39H1 has been implicated to play an essential role in the initial steps of heterochromatin formation in mammals by selective methylation of H3K9. In particular, trimethylation of H3K9 is mediated by SUV39H1 and proved to be essential in the establishment of constitutive heterochromatin at pericentromeric and telomeric regions in the genome, both of which consist of tandem repeated DNA sequences (Peters et al., 2003; Garcia-Cao et al., 2004). Mice that were deficient for SUV39H1 were shown to display impaired pericentric H3K9 methylation and chromosomal instability (Peters et al., 2001). Furthermore, loss of methylated H3 tails was shown to be accompanied by delocalization of HP1 from pericentric heterochromatin (Maison et al., 2002). In Drosophila melanogaster, SU(VAR)3-9 is the homologue of human SUV39H1 and of mouse Suv39h1 and Suv39h2. Similar to mouse mutants, D. melanogaster SU(VAR)3-9null mutants revealed a strong reduction in H3K9 methylation and an almost complete loss of HP1 from chromocenter heterochromatin (Schotta et al., 2002). Together, these findings underscore the essential role of SUV39H1 methyltransferase activity in establishing heterochromatin.
The catalytic methyltransferase activity of SUV39H1 has been mapped to the conserved COOH-terminal SET domain and uses monomethylated H3K9 as a substrate (Peters et al., 2003). The first 44 amino acids at the NH2 terminus of SUV39H1 form an interaction domain for the chromoshadow domain of HP1 and are, together with the adjacent chromodomain, required for binding to heterochromatin (Melcher et al., 2000). Interestingly, in HP1-deficient D. melanogaster, salivary gland nuclei SU(VAR)3-9 was shown to be lost from chromocenter heterochromatin and to spread along euchromatic regions, suggesting that an interaction between SU(VAR)3-9 and HP1 is essential for the association of SU(VAR)3-9 with centromeric heterochromatin (Schotta et al., 2002). However, the exact mechanism by which SUV39H1 interacts with heterochromatin is poorly understood.
Recently, the initial step in centromeric heterochromatin formation was suggested to be mediated by direct or indirect binding of SUV39H1 to components of an RNA interference pathway (Maison et al., 2002), but the mechanism by which SUV39H1 is initially recruited to centromeric repeat units is still enigmatic. Also, it is unclear whether SUV39H1 interacts only temporally with chromatin to methylate histone H3K9 or participates in a more stable multiprotein complex together with HP1 or other chromatin proteins to support a stable heterochromatic, pericentromeric chromatin structure. To gain further insight into the roles that SUV39H1 plays in heterochromatin, we investigated the in vivo kinetics of SUV39H1 in human osteosarcoma U2OS and in mouse NIH3T3 cells. Using FRAP analysis, we showed that SUV39H1 has a significantly slower exchange rate and a larger immobile fraction in heterochromatic regions compared with HP1ß. Our results suggest that a substantial fraction of SUV39H1 is immobile at pericentromeric heterochromatin, at least on the time scale of our experiments, and may, therefore, play a structural role. Furthermore, our data indicate that at least a part of the dynamic fraction of SUV39H1 is recruited to heterochromatin independently from HP1 binding. In addition, interactions between various SUV39H1 deletion mutants and HP1ß were analyzed in vivo by photobleaching and fluorescence resonance energy transfer (FRET) techniques to characterize the binding of HP1ß to SUV39H1 at heterochromatin.
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Results |
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The presence of an immobile or considerably less mobile fraction of SUV39H1 in heterochromatin was further investigated by performing two successive FRAP measurements on the same heterochromatic domain in NIH3T3 cells. EYFP-SUV39H1 was photobleached in a heterochromatic domain, and, after a recovery period of 450 s, the same area was photobleached for the second time, after which the fluorescence recovery was measured again. During the second FRAP, an immobile fraction would be invisible because it was bleached during the first FRAP. As shown in Fig. 3 E, after the second bleach, a full recovery of fluorescence was measured, which was 70% of the initial amount of fluorescence that was measured before the first photobleaching. Thus, this experiment confirmed the existence of an immobile fraction, at least on the time scale of the experiment.
Because various lines of evidence suggest that SUV39H1 interacts with HP1 proteins (Aagaard et al., 1999; Melcher et al., 2000) and that HP1 proteins are very mobile in the nucleus of living cells (Cheutin et al., 2003; Festenstein et al., 2003), we sought to compare the dynamic properties of SUV39H1 with those of HP1 proteins in NIH3T3 cells. Fusion constructs of HP1 and HP1ß with EYFP were made and transiently transfected into cells. Fusion proteins were shown to localize to the same nuclear sites as their endogenous counterparts, as revealed by immunocytochemistry with specific anti-HP1 antibodies. Cells showing moderate expression levels were selected, and, upon bleaching of small areas inside the nucleus that corresponded to heterochromatin and euchromatin, recovery of fluorescence was recorded. Consistent with previous data, HP1 proteins revealed a very dynamic behavior in NIH3T3 cells (Fig. 3 B). For EYFP-HP1ß, we measured a t1/2 of
4.2 s and a maximum fluorescence recovery of
95%, which was reached after 60 s from bleaching (Fig. 3 C and Table I). Similar kinetics of EYFP-HP1ß was measured when CFP-SUV39H1 and EYFP-HP1ß were coexpressed in the same cells (unpublished data). Together, these results suggest that SUV39H1 is more stably bound to chromatin than HP1 proteins and that a significant fraction of SUV39H1 is recruited to chromatin in an independent fashion or at least is not in a stable complex together with HP1 proteins.
The SET domain with adjacent regions mediates stable binding of SUV39H1 to heterochromatin
Three distinct protein domains have been identified in SUV39H1, of which aa 344 at the NH2 terminus form the HP1ß interaction surface that, together with the adjacent chromodomain (aa 4488), direct accumulation at heterochromatic regions (Melcher et al., 2000). The COOH-terminal SET domain (aa 249412) has been shown to be responsible for H3K9 methylation but has also been suggested to modulate heterochromatin association of SUV39H1 (Melcher et al., 2000; Lachner et al., 2001). To investigate how the different domains contribute to the kinetic behavior of SUV39H1 at chromatin in vivo, we generated various deletion mutants of SUV39H1 according to Melcher et al. (2000), fused them to EYFP, and transiently expressed them in NIH3T3 cells. A cartoon depicting the deletion mutants is shown in Fig. 4. First, we analyzed the spatial distribution of fusion proteins in the cell nucleus. Cells moderately expressing EYFPSUV39H1-SET (aa 3249 of Suv39H1 fused to EYFP) or EYFPSUV39H1-Nchromo (aa 3118 of Suv39H1 fused to EYFP) revealed the characteristic heterochromatic localization pattern that has also been observed for the full-length protein and after staining with DAPI or HP1 antibodies (Fig. 5 A, top). However, when expressed in U2OS cells, slightly different localization patterns were observed. The patterns seemed more dotlike when compared with localization of the full-length fusion protein (Fig. 5, C and D, top). Nevertheless, the expressed EYFPSUV39H1-
SET and EYFPSUV39H1-Nchromo fusion proteins were of the correct size, as determined by Western blotting (Fig. 5 B). Confocal analysis of U2OS cells expressing the fusion proteins that were stained with an anticentromere antibody showed EYFPSUV39H1-
SET (Fig. 5 C, bottom) or EYFPSUV39H1-Nchromo (Fig. 5 D, bottom) localization only at the centromeres. This localization in interphase nuclei is consistent with the finding that SUV39H1-
SET and SUV39H1-Nchromo localize more exclusively to the centromeres of metaphase chromosomes (Melcher et al., 2000).
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SUV39H1 dynamics at centromeric heterochromatin is independent of its methyltransferase activity
Our data and that of others suggest that targeting of SUV39H1 to centromeric heterochromatin is mediated by the HP1-binding domain together with the chromodomain and that this targeting is even more profound in absence of the SET domain (Melcher et al., 2000). Meanwhile, a SUV39H1 mutant lacking the SET domain becomes more mobile at centromeric heterochromatin. This suggests that the SET domain is not essential for recruitment, but it still may play a role in stabilizing the interaction of full-length SUV39H1 with centromeric heterochromatin. To investigate whether it is the methyltransferase enzymatic activity of the SET domain that mediates SUV39H1 binding to heterochromatin, we analyzed the mobility of two point mutants; one with impaired methyltransferase activity (SUV39H1-H324L) and one with enhanced methyltransferase activity (SUV39H1-H320R). Both mutants were fused to EYFP and revealed the same localization as EYFP-SUV39H1 when transiently transfected into NIH3T3 cells. FRAP analysis of both point mutants revealed that the fluorescence recovery times and the mobile fractions at heterochromatin did not significantly deviate from that of wild-type SUV39H1 (Fig. 7), indicating that the binding of SUV39H1 is not mediated by its methyltransferase enzymatic activity.
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U2OS cells were transiently cotransfected with EYFP-HP1ß and ECFP-SUV39H1, EYFP-HP1ß and ECFPSUV39H1-SET, or with EYFP-HP1ß and ECFPSUV39H1-
N89. As a negative control, cells were transfected with ECFP-SUV39H1 only. For each combination, 1630 cells were analyzed using the three-filter method as well as the FLIM method. Cells expressing EYFP-HP1ß together with ECFP-SUV39H1 or ECFPSUV39H1-
SET revealed sensitized emission (Fig. 8, A and B), a significant decrease in mean donor lifetime, respectively, and 2.2 and 2.1 against 2.5 ns measured in control cells (Table II). Spatial analysis revealed most prominent FRET at heterochromatic areas (Fig. 8, A and B). Cells that were cotransfected with EYFP-HP1ß and ECFPSUV39H1-
N89 did not show sensitized emission (Fig. 8 C) or a decrease in donor lifetime (Table II). These results indicate that SUV39H1 and SUV39H1-
SET, but not SUV39H1-
N89, interact with HP1ß in living cells. Moreover, the data suggest that the potential role of the SET domain in binding of SUV39H1 to heterochromatin is not mediated by an interaction with HP1.
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DNA demethylation increases SUV39H1 mobility
Evidence is accumulating that a tight interplay exists between DNA methylation and histone modification. Interactions of SUV39H1 with different DNA methyltransferases (Fuks et al., 2003) as well as with methylcytosine guanine dinucleotidebinding domain proteins (Fujita et al., 2003) have been described previously. To explore the role of DNA methylation in strengthening the interaction between SUV39H1 and heterochromatin, we treated cells with the DNA demethylating agent 5-aza-2'deoxycytidine (5-aza-C). Cell sorting analysis revealed that this treatment resulted in some decrease of cells that are in S or G2 phase of the cell cycle (9 vs. 20% in untreated cells), with a vast majority of cells in G1 (90 vs. 78%). Furthermore, we showed by Western blotting that this treatment had no significant influence on the trimethylation of H3K9 or acyetylation of histones (see Fig. 10 D). Next, we analyzed the mobility of EYFP-SUV39H1 by FRAP, and the resulting curves show that DNA demethylation increases the mobility of SUV39H1 (t1/2 of 14.6 s compared with
19.0 s in untreated cells; Fig. 9). This suggests that the interaction of SUV39H1 with components of the DNA methylation machinery or with the unmethylated DNA itself has changed. However, the mobility of SUV39H1 in 5-aza-Ctreated cells was still less than determined for SUV39H1-
SET in untreated cells, indicating that these interactions are not the sole mechanisms by which SUV39H1 binds to pericentromeric heterochromatin. Furthermore, the observation that EYFPSUV39H1-
SET becomes more mobile in 5-aza-Ctreated cells compared with nontreated cells (Fig. 9) suggests that it is not the SET domain of SUV39H1 that mediates the interaction with components of the DNA methylation machinery or with DNA itself.
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Trichostatin A (TSA) treatment affects the binding of SUV39H1 but not SUV39H1-SET to chromatin
Heterochromatin is not only characterized by hypermethylation but also by hypoacetylation of histones. The hyperacetylation of histones is a hallmark of transcriptionally active genes and has been suggested to prevent methylation of histone H3K9 and, thus, heterochromatin formation (Jenuwein and Allis, 2001). Therefore, histone deacetylases might be active at heterochromatin to maintain a hypoacetylated state. Treatment of cells with the histone deacetylase inhibitor TSA has been shown to result in the dissociation of HP1 from pericentromeric heterochromatin domains, probably by preventing methylation of H3 by SUV39H1 (Taddei et al., 2001). To test whether the in vivo kinetics of SUV39H1 binding to chromatin alters as a consequence of hyperacetylation, NIH3T3 cells expressing EYFP-SUV39H1 were treated with TSA. As shown by Western blotting, this treatment indeed resulted in an increased amount of acetylated histones and in a decrease in trimethylated H3K9 (Fig. 10 D). Cell sorting analysis revealed that this treatment resulted in some decrease of cells that were in S or G2 phase of the cell cycle (12 vs. 20% in untreated cells), with a vast majority of cells in G1 (87 vs. 78%). TSA-treated cells revealed a less pronounced localization of EYFP-SUV39H1 at heterochromatic regions, which is consistent with previous data (Maison et al., 2002; Cheutin et al., 2003). Furthermore, FRAP analysis revealed a faster fluorescence recovery of EYFP-SUV39H1 in TSA-treated cells (t1/2 of 7.7 s) compared with untreated cells (t1/2 of
19.0 s). Also, the total mobile fraction of EYFP-SUV39H1 at chromatin increased from
70 to
90% (Fig. 10, A and C, and Table I). TSA treatment had no significant effect on the kinetics of SUV39H1-
SET (t1/2 of
8.6 s). However, consistent with previous data (Cheutin et al., 2003), EYFP-HP1ß mobility increased after TSA treatment (Fig. 10 B) to a t1/2 of
1.8 s compared with 4.2 s in untreated cells (Fig. 10 C). These data show that hyperacetylation of histones reduces the binding of EYFP-SUV39H1 and EYFP-HP1ß to heterochromatin but does not prevent a transient SET-independent interaction of EYFP-SUV39H1 with chromatin.
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Discussion |
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Initially, HP1 had been considered a stable component of heterochromatin in order to maintain a compact, inaccessible chromatin conformation that would exclude transcriptional activity. This concept has changed because HP1 proteins were shown by FRAP analysis to be highly mobile in living cells (Cheutin et al., 2003; Festenstein et al., 2003). Recently, various other structural chromatin-binding proteins were shown to interact transiently, indicating that transient interactions are a common feature of chromatin proteins (Christensen et al., 2002; Harrer et al., 2004; Phair et al., 2004). These findings led to the suggestion that the dynamic nature of architectural chromatin-associated proteins would be essential to regulate gene-related processes (Harrer et al., 2004; Phair et al., 2004). This idea is consistent with recent findings showing that condensed chromatin domains are accessible for high molecular weight dextrans (Verschure et al., 2003) as well as for gene regulatory and chromatin proteins (Chen et al., 2005). Our observation that a substantial fraction of SUV39H1 is stably associated with chromatin during an extended period of time suggests, however, that not all chromatin-associated proteins are necessarily in continuous flux with the nucleoplasm. The significant immobile fraction of SUV39H1 might be indicative of chromatin conformations that are not prone to processes regulating transcriptional activity but that play a role in the structural organization of the cell nucleus. This might be particularly true for centromeric and telomeric regions, which have been implicated to play roles in spatial nuclear organization and generally show constrained movements in live cells (Molenaar et al., 2003). Consistent with this idea is the observation that changes in DNA methylation and histone acetylation have significant effects on the architecture of interphase chromosome arms but not on the positioning of centromeres and telomeres in wheat cell nuclei (Santos et al., 2002).
SUV39H1, however, is not only found at constitutive silent regions but also at genomic regions displaying transcriptional activity (Greil et al., 2003). Furthermore, transcriptionally inactive genes are found in open chromatin structures, whereas transcriptionally active genes are found in domains of compact chromatin (Gilbert et al., 2004). Hence, the classical strict distinction between heterochromatin (that it is supposed to be transcriptionally inactive) and euchromatin (that it is supposed to be transcriptionally active) does not exist. Because it is not possible to spatially define and distinguish small transcriptionally active and inactive regions in living cells expressing EYFP-SUV39H1, it was not possible to reproducibly analyze and compare the dynamic properties of SUV39H1 association at sites that were distinct from the relatively large heterochromatin domains. Therefore, the FRAP data obtained from nuclear regions that were adjacent to the pericentromeric heterochromatin in NIH3T3 cells is likely a reflection of SUV39H1 mobility rates at various levels of chromatin compaction. Nevertheless, the fact that we, on average, measured higher mobility rates in regions that we defined as euchromatic suggests that SUV39H1 is more dynamic at less condensed chromatin than at pericentromeric heterochromatin.
Recently, a slow moving fraction of HP1 has been identified at pericentromeric heterochromatin by using fluorescence correlation microscopy and FRAP (Schmiedeberg et al., 2004). However, this fraction appears to be relatively small (7% in NIH3T3 cells) when compared with the relatively large immobile fraction of SUV39H1 (30% in NIH3T3 cells) that we measured. Furthermore, the fast fluorescence recovery rate of HP1 (t1/2 of
4 s) compared with the slow recovery rate of SUV39H1 (t1/2 of
19 s) support the idea that both proteins are not necessarily recruited as a complex to chromatin, which is in contradiction with current models (Lachner et al., 2001; Maison and Almouzni, 2004). Our findings and the data of others are consistent with a model that SUV39H1 is a structural component of heterochromatin and is able to recruit HP1 by direct proteinprotein interaction (Stewart et al., 2005). Interestingly, another SET domaincontaining protein, multiple myeloma SET II, has recently been shown to bind to a nucleoplasmic structure (possibly chromatin) with even higher affinity than SUV39H1 (Keats et al., 2005). This finding suggests that the tight association of SUV30H1 with chromatin is a more common feature that is shared by other SET proteins. However, multiple myeloma SET II belongs to another subfamily than does SUV39, whose function is unknown and may not even possess histone methyltransferase activity.
To identify the protein domain that is responsible for stable binding of the immobile population of SUV39H1 to pericentromeric heterochromatin, we characterized the exchange rate of various deletion mutants of SUV39H1 at pericentromeric heterochromatin in living cells by FRAP analysis. This analysis revealed that in addition to the NH2 terminus and the adjacent chromodomain of SUV39H1, the SET domain also plays a role in chromatin binding. Consistent with previous data (Melcher et al., 2000), we observed that SUV39H1-SET localization was mainly restricted to pericentromeric heterochromatin regions compared with wide-spread chromatin associations of full-length SUV39H1. This suggests that the SET domain plays a role in chromatin association. Our finding that SUV39H1-
SET is highly mobile at the pericentromeric heterochromatin regions in living cells reinforces this suggestion and indicates that the SET domain mediates the stable interaction of SUV39H1 with pericentromeric chromatin, whereas the other domains are responsible for the recruitment of SUV39H1 to chromatin. Furthermore, FRAP analysis of the point mutants SUV39H1-H324L and SUV39H1-H320R revealed that it is not the histone methyltransferase activity of the SET domain that stabilizes binding of SUV39H1. This is consistent with previous localization data showing that heterochromatin association of SUV39H1 can be uncoupled from its intrinsic histone methyltransferase activity (Lachner et al., 2001).
The mechanism that mediates the stable interaction of SUV39H1 via the SET domain with pericentromeric chromatin remains elusive. It has been described previously that SUV39H1 not only interacts with HP1 (Aagaard et al., 1999) but also interacts with components of the DNA methylation pathway (Fujita et al., 2003; Fuks et al., 2003) and with histone deacetylases (Vaute et al., 2002). Our FRET analysis reveals that SUV39H1 and HP1 indeed interact at pericentromeric chromatin in live cells but that this interaction does not mediate the stable binding of SUV39H1 with heterochromatin. Furthermore, we could find no indication that binding of the immobile SUV39H1 population to pericentromeric chromatin is mediated by DNA methylation. Interestingly, treatment of cells with the deacetylation inhibitor TSA resulted in a higher mobility of SUV39H1 and in a significant reduction of the immobile SUV39H1 fraction at pericentromeric heterochromatin, whereas the kinetic behavior of SUV39H1-SET did not change. This suggests that histone deacetylase activity is required for the stable interaction of SUV39H1 to pericentromeric chromatin via the SET domain. Remarkably, exposure of HeLa cells to TSA was previously reported to relocate centromeres from a more internal position to the nuclear periphery (Taddei et al., 2001; Maison et al., 2002), supporting the idea that the immobile population of SUV39H1 may contribute to maintaining nuclear architecture. By immunoprecipitation and activity assays, it has been demonstrated that the NH2-terminal part of SUV39H1 can interact with histone deacetylase 1 and 2 (Vaute et al., 2002). This interaction, which might be direct, may help to recruit SUV39H1 to heterochromatin but does not, however, explain the role of the SET domain in maintaining an immobile population of SUV39H1. Recent studies indicate that a yet unidentified RNA may contribute to HP1 and may also contribute to SUV39H1 binding to pericentromeric heterochromatin (Maison et al., 2002; Muchardt et al., 2002). We are currently investigating the interesting possibility that an RNA component contributes to the establishment of an immobile population of SUV39H1 at pericentromeric heterochromatin.
In summary, our data have important implications for understanding the mechanisms through which SUV39H1 exerts its biological functions. It is intriguing that the SET domain not only displays histone methyltransferase activity but also plays a role in the binding of SUV39H1 to heterochromatin. In particular, the SET domain is likely to be involved in stabilizing the association of a population of SUV39H1 proteins to heterochromatin. We speculate that this stable, immobile population of SUV39H1 plays an essential role in the maintenance of a functional nuclear architecture and may also act as an interaction platform for other heterochromatin proteins. Future studies will address how the stable association of SUV39H1 at heterochromatin is maintained. This is particularly relevant because the SET domain of SUV39H1 has also been identified as being a proteinprotein interaction domain with possible implications in cancer development (Schneider et al., 2002). For example, the SET domain was shown to interact with Sbf-1, a protein with the ability to transform fibroblasts (Cui et al., 1998). It can be speculated that this interaction may interfere with the functioning of SUV39H1 either by deregulating methyltransferase activity or possibly by direct destabilization of the chromatin conformation by weakening the binding of SUV39H1 to chromatin.
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Materials and methods |
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HP1ß was amplified with the forward primer 5'-GCGCGGTACCCATGGGGAAAAAACAAAACAAG-3' containing the KpnI site and the reverse primer 5'-GCGCCCCGGGCGTTCTTGTCATCTTTTT-3' containing the XmaI site and was cloned in the KpnI-XmaI fragment of pECFP-N1 and pEYFP-N1 (CLONTECH Laboratories, Inc.). Two SUV39H1 point mutants were made using the QuikChange II Site-Directed Mutagenesis Kit (Stratagene). For generating point mutant SUV39H11-H324L, we used the sense primer 5'-CTATGGCAACATCTCCCGCTTTGTCAACCACAGTTG-3' and the antisense primer 5'-CAACTGTGGTTGACAAAGCGGGAGATGTTGCCATAG-3'. For generating SUV39H1-H320R, the sense primer was 5'-CATCTCCCACTTTGTCAACCTCAGTTGTGACCCCAAC-3' and the antisense primer was 5'-GTTGGGGTCACAACTGAGGTTGACAAAGTGGGAGATG-3'. All constructs were verified by sequencing.
Cell culture and transfection
U2OS cells and NIH3T3 mouse fibroblast cells were cultured on 3.5-cm glass bottom petri dishes (MatTek) in DME without phenol red and containing 1.0 mg/ml glucose, 4% FBS, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin buffered with 25 mM Hepes, pH 7.2 (all from Invitrogen). Wild-type and Suv39h dn PMEFs (provided by T. Jenuwein, Research Institute of Molecular Pathology, Vienna, Austria) were cultured as described previously (Lehnertz et al., 2003). Transient transfections were performed at 7080% confluence using 1 µl LipofectAMINE 2000 (Invitrogen) and 0.751.5 µg DNA. For TSA treatment, cells were incubated with 50 ng/ml TSA (Sigma-Aldrich) for 1822 h. For 5-aza-C treatment, cells were incubated with 5 µM 5-aza-C (Sigma-Aldrich) for 4854 h.
Immunocytochemistry
Antibodies that were used in this study are listed as follows: human anticentromere (Antibodies, Inc.), mouse antiheterochromatin protein 1 and ß (both from Euromedex), rabbit anti2x-trimethylated H3K9 (provided by T. Jenuwein), and the appropriate secondary antibodies. These antibodies were diluted 1:100, 1:500, 1:500, and 1:250, respectively, in TBS containing 0.5% (wt/vol) blocking reagent (Roche) and 0.1% Tween 20.
Cells that were grown on microscopic glass slides were washed in PBS and fixed in 2% formaldehyde in PBS for 5 min for incubation with the anticentromere antibody or with 4% formaldehyde and 0.5% Triton X-100 (Sigma-Aldrich) in 0.1x PBS for 5 min. Subsequently, cells were permeabilized in PBS containing 1% Triton X-100 for 15 min, washed three times in PBS, and washed once in TBS containing 0.1% Tween 20. Then, cells were incubated with the first antibody for 45 min for the anticentromere or for 3 h at 37°C followed by three washes in TBS/0.1% Tween 20. Finally, cells were incubated with the secondary antibody for 45 min at 37°C, washed in TBS/0.1% Tween 20, and mounted in Citifluor (Agar Scientific, Ltd.) containing 400 µg/ml DAPI (Sigma-Aldrich).
Cell sorting
Cells were stained with propidium iodide according to the DNAcon3 kit (DakoCytomation). Cells were sorted in FACS (Becton Dickinson) and analyzed with Cell Quest software (BD Biosciences).
Protein blot analysis
Cells were lysed in NuPAGE LDS Sample Preparation Buffer (Invitrogen). Protein samples were then size fractionated on Novex 412% BisTris gradient gels using MOPS buffer (Invitrogen) and were subsequently transferred onto Hybond-C extra membranes (GE Healthcare) using a submarine system (Invitrogen). Blots were stained for total protein using Ponceau S (Sigma-Aldrich). After blocking with PBS containing 0.1% Tween 20 and 5% milk powder, the membranes were incubated with anti-GFP antibody (1:500; Roche), anti-H3Ac antibody (1:1,000; Upstate Biotechnology), and anti2x-trimethylated H3K9 (1:500). The secondary antibodies that were used were antirabbit (1:2,000) and antimouse (1:5,000) HRP-conjugated antibodies (Pierce Chemical Co.). Bound antibodies were detected by chemiluminescence using ECL Plus (GE Healthcare).
Microscopy
Wide-field microscopy was performed on a microscope (model DMRXA; Leica) equipped with a 100-W mercury arc lamp and a 100x NA 1.3 plan Apo objective. Colocalization studies were performed on a confocal microscope system (model TCS/SP2; Leica). Image stacks were acquired with a 100x NA 1.4 plan Apo objective and were analyzed with Leica confocal software.
FRAP
Photobleaching experiments were performed on a confocal microscope. The temperature of the cells was maintained at 37°C by a heating ring surrounding the culture chamber (Harvard App., Inc.) and a microscope objective heater (Bioptechs). A 100x NA 1.4 plan Apo lens and a 514-nm laser line was used for photobleaching. Selected areas were subjected to five excitation pulses of 500 ms each at high laser power. Images were collected at time intervals of 450 ms from 5 s for 1.510 min after bleaching. The first postbleach image was acquired 2 s after photobleaching. To better quantify the recovery of fast moving SUV39H1 deletion mutants, selected areas were bleached once for 500 ms, and the first postbleach image was acquired after 500 ms. Quantitation of the fluorescent recovery was performed using Leica confocal software and Microsoft Excel. The recovery curves were corrected for background, fluorescence fading, and decrease in fluorescence during photobleaching. The t1/2 value was defined as the time required for reaching half-maximum recovery and was calculated from the corrected recovery curves.
FRET imaging
FRET was measured using a microscope (Axiovert 135 TV; Carl Zeiss MicroImaging, Inc.) equipped with a 100-W mercury arc lamp, a 100x plan Neofluar NA 1.3 phase objective, and a CCD camera (Micromax; Princeton Instruments). The spectral FRET measurements using three-filter sets were performed essentially as described by Xia and Liu (2001). CFP images were obtained with a filter set containing a 436/20-nm bandpass (BP) excitation filter, a 510-nm dichroic mirror, and a 480/200-nm BP emission filter. YFP images were obtained with a filter set containing a 500/20-nm BP filter, a 520-nm dichroic mirror, and a 535/30-nm BP emission filter (all from Chroma Technology Corp.). FRET images were acquired with a filter set containing a 436/20-nm BP excitation filter, a 510-nm dichroic mirror, and a 535/30-nm BP emission filter. The set of three images was corrected for background and pixel shift. Images were further processed, and the relative FRET efficiencies were calculated on a pixel-by-pixel basis. For the calculation of relative FRET values, the formula as described previously by Xia and Liu (2001) was used:
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FLIM was performed essentially as described previously (Gadella, 1999) using a microscope (Axiovert 135 TV; Carl Zeiss MicroImaging, Inc.) equipped with a 100-W mercury arc lamp, a 100x plan Neofluar NA 1.3 phase objective, and an image intensifier (model C5825; Hamamatsu) modulated at 40,000 MHz. ECFP was excited with a 458-nm argon/krypton laser line (Innova 70C; Coherent) modulated at 80,000 MHz using an acousto-optic modulator and a filter set containing a 458/10-mn BP excitation filter, a 470-nm dichroic mirror, and a 480/20-nm BP emission filter. 20 phase images (one every 36°C; 10 forward and 10 back to correct for bleaching) were acquired using a CCD camera (CoolSNAP fx; Photometrics). Reference phase settings and modulation were set every 15 min using a freshly prepared solution of 100 nM rhodamine 6G in 100% ethanol with a single component lifetime of 4.0 ns. Fluorescence lifetime values and
mod were calculated with the equations
= (1/
) tan
and
M = (1/
)
(1/M2 1). Lifetime values were calculated for each pixel and were displayed in pseudocolors.
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Acknowledgments |
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This work was partially supported by Cyttron (grant BSIK03036).
Submitted: 25 February 2005
Accepted: 1 July 2005
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References |
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