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Department of Biochemistry, Institute of Developmental Biology, Russian Academy of Sciences, Moscow, Russia,1 Kimmel Cancer Center and Department of Microbiology and Immunology, Thomas Jefferson University, Philadelphia, Pennsylvania,2 Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot, Israel3
Received 26 October 2004/ Accepted 29 November 2004
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ABSTRACT |
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INTRODUCTION |
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130-amino-acid (aa) sequence which was initially identified in the protein products of three regulatory genes in Drosophila, whose names account for the name SET, i.e, SU(VAR)3-9, enhancer of Zeste [E(Z)], and trithorax (Trx). SET domains are usually located at the carboxyl termini of proteins, although in some proteins, such as ASH1 (another regulator belonging to the TrxG family), the motif is located in the middle of the protein (Fig. 1A), and in the proteins RIZ and BLIMP this motif maps to the N termini. In addition to the highly conserved SET motif, less conserved 50- to 80-aa pre-SET and post-SET regions may be present at the amino and carboxyl boundaries, respectively, of the SET domain (Fig. 1A) (reviewed in reference 12). Genes encoding SET domains are widely represented in eukaryotic genomes (>300 database entries). Based on the homology of their SET motifs, the most characterized human proteins were classified into four major families, SET1, SET2, SUV39, and RIZ (15). A recently discovered function of the SET domain is the methylation of lysine residues in nucleosomal histones. SET domains of different proteins target specific lysines, and these modifications have different consequences on gene expression. For example, the methylation of Lys 4 of histone H3 is usually associated with activation, while the methylation of H3 Lys 9 or Lys 27 is associated with silencing (reviewed in reference 28).
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Although the methylation of histone residues is widely believed to be the central function of the SET domains, important aspects of this process, such as how SET proteins are recruited in the first place and how the histone modifications survive replication, are not understood. Here we describe another biochemical feature associated with SET domain proteins which might play a role in the aforementioned processes and in other processes. A motif present within the general area of the SET domains of several proteins binds single-stranded and supercoiled DNAs as well as RNA. This binding is very tight so that it will last and interfere with nucleosome assembly.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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CAT (22) were used as single- and double-stranded DNA probes, respectively, for gel retardation assays. Protein procedures. Recombinant glutathione S-transferase (GST) fusion proteins were expressed in Escherichia coli strain Origami B (Novagen), purified by standard procedures (including immobilization on glutathione-Sepharose), and analyzed in sodium dodecyl sulfate (SDS)-13% polyacrylamide gels. In some cases, to remove bound endogenous nucleic acids, we additionally treated the immobilized GST-SET polypeptides with DNase I and RNase A and washed them with HEMG buffer (50 mM HEPES-KOH [pH 7.6], 0.2 mM EDTA, 25 mM MgCl2, 20% glycerol [vol/vol], 1 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride) containing 500 mM NaCl, 0.1% NP-40, and 2 to 3 M urea. GST pull-down experiments were performed in EX protein buffer (17) (10 mM HEPES-KOH [pH 7.6], 10 mM KCl, 1.5 mM MgCl2, 0.5 mM EGTA, 10% glycerol [vol/vol], 1 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride) containing the desired amounts of NaCl (indicated in the figure legends) and 0.05% NP-40. Loading was followed by a series of washes with HEMG buffer containing 500 mM NaCl and 0.1% NP-40. Bound DNAs were isolated by phenol extraction of the glutathione-Sepharose beads and were analyzed in 1% agarose gels.
Coupled in vitro transcription and GST pull-down assay. The transcription of pGEM Express positive control template DNA (Promega) was performed directly in the buffer for the GST pull-down assay, supplemented with a complete or partial set of transcription components (a 0.1 mM concentration of each ribonucleoside triphosphate, 1 U each of SP6, T3, and T7 RNA polymerases [Promega], 1 U of RNasin [Boehringer], 3 U each of RNases A and H [Boehringer], and 3 U of DNase I [Boehringer]) as indicated in Fig. 3.
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Modeling of secondary structures. The utilization of Expasy modeling software allows analyses of two proteins side-by-side or of several proteins simultaneously. The modeling templates, which were all SET-domain proteins with known three-dimensional (3D) structures, included CLR4 (NCBI no. mvxA and lmvhA), DIM5 (NCBI no. 1pegA, 1pegB, and 1m19A) and SET7 (1MUF, 1NGA, and 1NGC). For simultaneous threading of an ssDNA binding sequence into the three templates, we used SWISS-PdbViewer or the automated homology modeling servers SWISS-Model and Geno 3D (5). Protein structures were visualized with the RasMol viewer in the color mode "chain" or "structure," which showed chains of proteins in different colors or structural elements in different colors, respectively.
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RESULTS |
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We commonly used a commercial 100-bp DNA ladder (Fisher), either intact or denatured by heating at 98°C in Tris-EDTA buffer, as a dsDNA and ssDNA probe. The choice of DNA source was not critical, since the testing of several other ss- or dsDNA substrates yielded similar results (not shown). We also compared SET binding to the negatively supercoiled versus linear forms of DNA. DNA helixes of circular closed DNA molecules can have a deficiency or excess of coils compared to a linear DNA of the same size. These under- or overwound DNA states are referred to as negatively or positively supercoiled DNA, respectively. The typical plasmid DNA isolated from a bacterial source is negatively supercoiled at a rate of one supercoil per 200 bp of DNA. Negatively or positively supercoiled DNA molecules tend to relieve arising torsional tensions either by changing their helical twist (decreasing or increasing it, respectively) and/or by writhing of the axis. Thus, negative, in contrast to positive, superhelical stress is known to destabilize the DNA duplex, resulting in locally unwounded regions of DNA (reviewed in reference 6). As a control, we used relaxed forms of plasmids, in which superhelical stress was removed by a treatment with topoisomerase I (Topo I).
ALL-1, also termed MLL, HRX, or HTRX, is the human homologue of Drosophila TRX and acts as a positive regulator of HOX genes (26, 40). SET fragments of the ALL-1 and E. coli SSB proteins were expressed as GST fusions in bacteria (Fig. 1B), and their binding with various DNA substrates was assessed in several assays under stringent washing conditions for DNA-protein complexes by use of a buffer containing 500 mM NaCl and 0.1% NP-40.
Like the case for the E. coli SSB protein, the 151- and 400-aa C-terminal fragments of ALL-1 associated tightly with immobilized single-stranded, but not double-stranded, DNA (Fig. 1B). When immobilized on a glutathione-Sepharose matrix via a GST moiety (Fig. 1C), the SET region of ALL-1 efficiently retained ssDNA fragments, whereas its binding to dsDNA was barely detectable (the input amounts of the ds- and ssDNA were equalized for DNA content, but since ethidium bromide binds ssDNA less efficiently than dsDNA, lanes with ssDNA appear to be underloaded). In contrast to the case for the GST-SSB protein of E. coli, ALL-1 C-terminal tails bound the negatively supercoiled form of the plasmid (Fig. 1C, bottom panel, lanes 5 to 8 versus lanes 3 and 4). Complexes of GST-SET with ssDNA could be further eluted with reduced glutathione (Fig. 1D), indicating that the physical association was not due to a nonspecific aggregation on the surface of the Sepharose matrix. Gel retardation assays performed with physiological ionic strengths (140 to 150 mM NaCl) also revealed a higher affinity of the ALL-1 SET region for ss- versus dsDNA (Fig. 1E).
We noted that at low ionic strengths (below 100 mM NaCl), ALL-1 SET fragments and some preparations of EcSSB bound small amounts of dsDNA (not shown), indicating that the difference in the affinities of SET fragments and EcSSB for dsDNA was quantitative rather than qualitative. The glutathione-Sepharose matrix or immobilized GST retained neither ss- nor dsDNA (not shown).
The SET regions of several other SET-containing proteins, including Drosophila E(Z), TRX, and ASH1, human ALR, the E(Z)-related Arabidopsis CLF, and yeast SET1 and SET2, also bound single-stranded and supercoiled DNAs (not shown; see Fig. 2 for detailed analyses of some of these), suggesting that such DNA binding may be a general feature of SET regions. Altogether, the tested proteins encompassed members of the SET domain families SET1 and SET2 (15). Interestingly, the corresponding polypeptides of two representatives of the SUV39 family, Drosophila SU(VAR)3-9 and human G9a, did not bind to supercoiled (not shown) and ssDNA (Fig. 2).
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We wondered how strong the binding was between the SET-containing polypeptides and supercoiled or ssDNAs. The ALL-1 SET region was bound to both templates and subjected to washings at increasing stringencies (Fig. 1G, middle and bottom panels). We found that the polypeptide was still bound in the presence of 4.5 M urea and 3.5 M NaCl (or even 5 M NaCl [not shown]), suggesting a strong association.
We next demonstrated that even though SET binding to supercoiled DNA was very stable, it was reversible: DNA supercoiling was essential not only for establishing but also for maintaining binding to the ALL-1 SET domain (Fig. 1H). To this end, the GST-tagged 220-aa ALL-1 C-terminal SET domain was immobilized on glutathione-Sepharose and assayed for binding to the negatively supercoiled, linear, and relaxed forms of three plasmids (3, 4.5, and 6 kb). Only the torsionally stressed plasmids remained bound to ALL-1 SET after washing with 0.5 M NaCl-0.1% NP-40 buffer (Fig. 1H, lanes 10 to 12 versus lanes 13 to 18). When the bound supercoiled plasmids were subsequently linearized or relaxed by treatment with a restriction enzyme or with Topo I, most of the plasmid DNA was eluted in the wash buffer, whereas unmodified supercoiled plasmids remained bound to immobilized ALL-1 SET (Fig. 1H, compare lanes 19, 21, and 23 to lanes 25, 27, 29, 31, 33, and 35). Thus, the DNA-protein complexes dissociate upon relief from torsional tension.
Mapping the ssDNA-binding motifs within SET regions. We applied deletion analysis to map the ssDNA recognition motifs within several representative SET-domain proteins. These included ALL-1 and its Drosophila homologue TRX (Fig. 2A and B), Drosophila E(Z) and the related plant protein CLF (Fig. 2D and E), and Drosophila ASH1 (Fig. 2C). The black bars at the tops of the schemes indicate the smallest polypeptides capable of binding to ssDNA with an affinity comparable to that of the full-length SET regions. The data in Fig. 2 show that the ALL-1 and TRX ssDNA-binding domains are located at the pre-SET-SET boundaries. The corresponding domains of E(Z) and CLF were mapped to the pre-SET region, and E(Z) had a second binding domain within pre-SET-SET. The ASH1 protein, in which the SET domain is in the middle of the protein, contains two ssDNA-binding domains, at the pre-SET-SET and SET-post-SET boundaries (Fig. 2C). In some instances, polypeptides spanning the minimal binding sequences together with some additional sequences did not bind DNA [T11 of TRX, E7 of E(Z), and A2,3 of ASH1]. This was likely due to the polypeptides assuming steric conformations which prevented DNA binding. No binding of ssDNA was observed for the SET regions of SU(VAR) 3-9 and G9a (Fig. 2F) with polypeptides spanning SET and extending upstream.
The ssDNA-binding motif associates with in vitro-transcribed DNA and with nascent RNA. A major physiological process that causes an opening of the DNA duplex is the traversing of DNA by RNA polymerases. To assess if transcription of a DNA template is sufficient for making it available for an interaction with the SET region, we performed a coupled in vitro transcription-GST pull-down assay by using phage RNA polymerases and an appropriate DNA template. The latter, the pGEM Express positive control template (Promega), contains two promoters for each of the T3, T7, and SP6 polymerases which generate RNAs of 250 and 1,525, 1,065 and 2,346, and 1,787 and 2,566 bases, respectively. In the absence of ongoing transcription, the interaction of the DNA template and the 220-aa ALL-1 C-terminal SET fragment occurred only at background levels (Fig. 3A, lanes 2 to 7). However, if a full set of reaction components was supplied to allow transcription, the DNA template was efficiently associated with the ALL-1 SET region, as was the nascent RNA species (lanes 8 to 11). Transcribed DNA and the RNA product interacted with the polypeptide independently of each other, as indicated by the selective removal of DNA (lanes 16 to 19) or RNA (lanes 12 to 15) after a treatment with DNase I or a mixture of RNases A and H, respectively (we took advantage of RNase H's ability to destroy RNA within RNA-DNA hybrids). We did not test whether processing by eukaryotic RNA polymerases can also induce an association of DNA with the polypeptide. However, the basic principles of eukaryotic and prokaryotic transcription are well conserved, and it can be presumed that the SET region is associated with transcribing eukaryotic genes as well. We tested whether SET regions of some other proteins could also bind transcribed DNA structures. Indeed, SET-containing fragments of TRX, ALR, and ASH1, but not SU(VAR)3-9, also exhibited binding to in vitro-transcribed DNA (Fig. 3B). Note that SET polypeptides isolated from bacteria are found associated with various amounts of nucleic acids which can be removed by incubation with DNase and RNase (not shown). The associated nucleic acids do not appear to impair histone lysine methyltransferase activity (26).
The ssDNA-binding motif is not displaced during in vitro assembly of nucleosomes and prevents the formation of regular nucleosome arrays. Within the cell nucleus, DNA is exposed to interactions with numerous factors, some of which result in very stable associations. For example, most eukaryotic DNA is found in complex with histones, forming chromatin structures (10) that can be destroyed only by treatments with high concentrations of salt or ionic detergents. We tested whether nucleosomes would exclude or remove bound ALL-1 SET regions from the template DNA during assembly (Fig. 3C). The template DNA used for this experiment contained single-stranded regions generated by heating of the DNA to temperatures sufficient to open the easily melting sequences but not the entire DNA duplex (21). Such transiently melted DNA regions usually renature quickly and do not significantly affect nucleosome assembly, but they appear to be sufficient to provide strong binding of the ALL-1 SET region (Fig. 3C, left panel). The GST-tagged 220- and 350-aa ALL-1 SET fragments were incubated with such partially melted DNA and, as a control, intact double-stranded DNAs, and subsequent nucleosome assembly was performed as previously described (18) by use of a Drosophila embryo extract and an ATP regeneration system (Fig. 3C, right panel). Under these conditions, the ATP-dependent spacing activities of the extract generate continuous arrays of dynamic nucleosomes with regular spacing (19). Upon the completion of assembly, the integrity of the nucleosomal arrays was assayed by the use of staphylococcal nuclease, which cleaves regularly spaced nucleosome arrays to produce a typical ladder of the oligonucleosomal DNA fragments. The digestion of chromatin assembled on the intact dsDNA resulted in a set of well-resolved DNA fragments (Fig. 3C, right panel, lanes 2 to 9), implying that preincubation with GST-SET or GST polypeptides did not interfere with nucleosome assembly. However, when the DNA template was partially melted and preincubated with GST-ALL-1 SET fragments (but not with GST alone) prior to the assembly reaction, the nucleosome ladder was significantly less pronounced (compare lanes 11 to 13 with lanes 14 to 17). This result suggests that the association between the SET fragments and ssDNA survives assembly, preventing the formation of regular nucleosome arrays. Thus, the binding of SET-domain proteins to ssDNA efficiently competes with the extremely strong affinity between nucleosomal proteins and DNA.
The homeotic mutation trxZ11 impairs binding of the TRX SET region to ssDNA. The TRX SET domain spans the well-studied developmental mutation trxZ11, in which a highly conserved glycine (G3601) is replaced with serine (34). This mutation results in homeotic transformations in flies that are heterozygous for trxZ11 and causes lethality at the pupal stages in hemizygous animals (4). To assess whether this mutation affects the binding of the TRX SET region to ssDNA, we applied a pull-down assay to compare the activities of the GST-tagged TRX SET fragments derived from the trxZ11 and wild-type proteins (Fig. 4). trxZ11 polypeptides, but not their wild-type counterparts, showed progressively reduced abilities to bind ssDNA when the ionic strength of the binding buffer was increased from 125 to 175 mM. This effect was apparent with the 385- and 151-aa TRX SET regions and became striking (Fig. 4, bottom panel) when an 81-residue TRX polypeptide which was the smallest to bind ssDNA was used (composed of pre-SET and a small region of SET).
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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When possible functions of SSBLS are considered, one potential role will involve the recruitment of SET proteins to their targets. The mechanism(s) associated with this process is largely unknown. Recent results with fission yeast, plants, flies, and vertebrates (8, 29, 31, 38, 39) suggest that short interfering RNAs (siRNAs) function to recruit H3 Lys9 methyltransferases to specific loci to direct heterochromatin assembly. It has been speculated that the protein complex RISC/RITS, which contains siRNA, associates with the H3 Lys9 methyltransferase and is targeted to specific chromosome regions through base pairing of the former to single-stranded DNA bubbles or nascent transcripts. In principle, this model can be extended to other histone methyltransferases that act in the local decondensation or condensation of euchromatin [e.g., ALL-1, ASH1, and E(Z)]. A motif within SET proteins which binds single-stranded nucleic acids may facilitate the targeting and anchoring of the RISC/RITS complex to specific chromosome regions. Moreover, since each RISC contains only one of the two strands of the siRNA duplex (25), the SET-domain proteins might physically link to RISC through the binding of SSBLS to complex-associated single-strand RNA.
A role for transcription of a target gene in the recruitment of a SET polypeptide was also deduced from our recent work, in which we found that the TRX SET region encoded by a transgene was recruited in vivo to the heat shock genes only during active transcription of the latter (33). Moreover, the TRX SET domain is indispensable for recruiting the TRX protein and the components of the TAC1 multiprotein complex to heat shock genes (33). Furthermore, several reports have indicated that initial transcription of the regulatory region of the Drosophila homeotic gene ultrabithorax is required for establishing the maintenance state (and presumably the corresponding covalent histone modifications) of this gene's transcriptional activity (3, 11, 30). These recent findings are consistent with the ability of SSBLS to bind transcribed DNA (and RNA) as well as with its ability to interact in a reversible way with torsionally stressed DNA, which is considered a hallmark of functionally active DNA structures (6, 16, 21). Another possible role for SSBLS is to prevent the formation of regular nucleosome arrays in transcribed regions so as to maintain the required level of transcription. Finally, the capacity of SET-domain proteins to bind ssDNA may play a mechanistic role in the inheritance of histone covalent modifications. One can imagine a scenario in which, during progression of the replication fork, SET-domain proteins dissociate from the parental DNA strands to be quickly recruited to the newly synthesized DNA due to the strong interaction between the SET regions and ssDNA. In this scenario, SET-domain proteins play an active role not only in marking histones for transcription and silencing states, but also in propagating these states in the progeny cells.
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ACKNOWLEDGMENTS |
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This study was supported by NCI grant CA50507, grants from the Israeli Science Foundation, the Israel Cancer Research fund, and U.S.-Israel BSF, and grants to W.K. from the Institute of Developmental Biology (Russian Academy of Science) and the Leukemia Research Foundation (Evanston, Ill.).
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FOOTNOTES |
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REFERENCES |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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2. Alvares-Venegas, R., and Z. Avramova. 2002. SET-domain proteins of the Su(var)3-9, E(z) and trithorax families. Gene 285:25-37.[CrossRef][Medline]
3. Bender, W., and D. P. Fitzgerald. 2002. Transcription activates repressed domains in the Drosophila birthorax complex. Development 129:4923-4930.
4. Breen, T. R. 1999. Mutant alleles of the Drosophila trithorax gene produce common and unusual homeotic and other developmental phenotypes. Genetics 152:319-344.
5. Combet, C., M. Jambon, G. Deleage, and G. Geourjon. 2002. Geno3D: automatic comparative molecular modelling of protein. Bioinformatics 18:213-214.
6. Esposito, F., and R. R. Sinden. 1988. DNA supercoiling and eukaryotic gene expression. Oxf. Surv. Eukaryot. Genes 5:1-50.[Medline]
7. Francis, N. J., and R. E. Kingston. 2001. Mechanisms of transcriptional memory. Nat. Rev. Mol. Cell Biol. 2:409-421.[CrossRef][Medline]
8. Fugakawa, T., M. Nogami, M. Yoshikawa, M. Ikeno, T. Okazaki, T. Nakayama, and M. Oshimura. 2004. Dicer is essential for formation of the heterochromatin structure in vertebrate cells. Nat. Cell Biol. 6:784-791.[CrossRef][Medline]
9. Hamiche, A., V. Carot, M. Alilat, F. De Lucia, M. F. O'Donohue, B. Revet, and A. Prunell. 1996. Interaction of the histone (H3-H4)2 tetramer of the nucleosome with positively supercoiled DNA minicircles: potential flipping of the protein from a left- to a right-handed superhelical form. Proc. Natl. Acad. Sci. USA 93:7588-7593.
10. Hayes, J. J., and J. C. Hansen. 2001. Nucleosomes and the chromatin fiber. Curr. Opin. Genet. Dev. 11:124-129.[CrossRef][Medline]
11. Hogga, I., and F. Karch. 2002. Transcription through the Iab-7 cis-regulatory domain of the bithorax complex interferes with polycomb mediated silencing. Development 129:4915-4955.
12. Jenuwein, T., G. Laible, R. Dorn, and G. Reuter. 1998. SET domain proteins modulate chromatin domains in eu- and heterochromatin. Cell. Mol. Life Sci. 54:80-93.[CrossRef][Medline]
13. Kastani, K. R., J. J. Arredondo, A. J. Kal, and P. C. Verrijzer. 2001. A homeotic mutation in the trithorax SET domain implies histone binding. Genes Dev. 15:2197-2202.
14. Kerr, I. D., R. I. Wadsworth, L. Cubeddu, W. Blankenfeldt, J. H. Naismith, and M. F. White. 2003. Insights into ssDNA recognition by the OB fold from a structural and thermodynamic study of SSB protein. EMBO J. 22:2561-2570.[CrossRef][Medline]
15. Kouzarides, T. 2002. Histone methylation in transcriptional control. Curr. Opin. Genet. Dev. 12:198-209.[CrossRef][Medline]
16. Krajewski, W. A. 1996. Enhancement of transcription by short alternating C.G tracts incorporated within a Rous sarcoma virus-based chimeric promoter: in vivo studies. Mol. Gen. Genet. 252:249-254.[Medline]
17. Krajewski, W. A. 2000. Histone hyperacetylation facilitates chromatin remodeling in a Drosophila embryo cell-free system. Mol. Gen. Genet. 263:38-47.[CrossRef][Medline]
18. Krajewski, W. A. 2002. Histone acetylation status and DNA sequence modulate ATP-dependent nucleosome repositioning. J. Biol. Chem. 277:14509-14513.
19. Krajewski, W. A., and P. B. Becker. 1999. Reconstitution and analysis of hyperacetylated chromatin. Methods Mol. Biol. 119:207-217.[Medline]
20. Krajewski, W. A., and S. V. Razin. 1993. DNA-protein interactions and spatial organization of DNA. Mol. Biol. Rep. 18:167-175.[CrossRef][Medline]
21. Krajewski, W. A., and S. V. Razin. 1992. Organization of specific DNA sequence elements in the region of the replication origin and matrix attachment site in the chicken alpha-globin gene domain. Mol. Gen. Genet. 235:381-388.[CrossRef][Medline]
22. Krajewski, W. A., and K. A. W. Lee. 1994. A monomeric derivative of the cellular transcription factor CREB functions as a constitutive activator. Mol. Cell. Biol. 14:7204-7210.
23. Lohman, T. M., and M. E. Ferrari. 1994. Escherichia coli single-stranded DNA-binding protein: multiple DNA-binding modes and cooperativities. Annu. Rev. Biochem. 63:527-570.[Medline]
24. Mahmoudi, T., and C. P. Verrijzer. 2001. Chromatin silencing and activation by polycomb and trithorax group proteins. Oncogene 20:3055-3066.[CrossRef][Medline]
25. Martinez, J., A. Patkaniowska, H. Urlaub, R. Lührmann, and T. Tuschl. 2002. Single-stranded antisense siRNAs guide target RNA cleavage in RNAi. Cell 110:563-574.[CrossRef][Medline]
26. Nakamura, T., T. Mori, S. Tada, W. Krajewski, T. Rozovskaia, R. Wassel, G. Dubois, A. Mazo, C. M. Croce, and E. Canaani. 2002. ALL-1 is histone methyltransferase that assembles a supercomplex of proteins involved in transcriptional regulation. Mol. Cell 10:1119-1128.[CrossRef][Medline]
27. Nislow, C., E. Ray, and L. Pillus. 1997. SET1, a yeast member of the trithorax family, functions in transcriptional silencing and diverse cellular processes. Mol. Biol. Cell 8:2421-2436.
28. Orlando, V. 2003. Polycomb, epigenomes and control of cell identity. Cell 112:599-606.[CrossRef][Medline]
29. Pal-Bhadra, M., B. A. Leibovitch, S. G. Gandhi, M. Rao, U. Bhadra, J. A. Birchler, and S. C. Elgin. 2004. Heterochromatic silencing and HP1 localization in Drosophila are dependent on the RNAi machinery. Science 303:669-672.
30. Rank, G., M. Prestel, and R. Paro. 2002. Transcription through intergenic chromosomal memory elements of the Drosophila bithorax complex correlates with an epigenetic switch. Mol. Cell. Biol. 22:8026-8034.
31. Schramke, V., and R. Allshire. 2003. Hairpin RNAs and retrotransposon LTRs effect RNAi and chromatin-based gene silencing. Science 301:1069-1074.
32. Simon, J. A., and J. W. Tamkun. 2002. Programming off and on states in chromatin: mechanisms of polycomb and trithorax group complexes. Curr. Opin. Genet. Dev. 12:210-218.[CrossRef][Medline]
33. Smith, S. T., S. Petruck, Y. Sedkov, E. Cho, S. Tillib, E. Canaani, and A. Mazo. 2004. Modulation of heat shock gene expression by the TAC1 chromatin modifying complex. Nat. Cell Biol. 6:162-167.[CrossRef][Medline]
34. Stassen, M. J., D. Bailey, S. Nelson, V. Chinwalla, and P. J. Harte. 1995. The Drosophila trithorax proteins contain a novel variant of the nuclear receptor type DNA binding domain and an ancient conserved motif found in other chromosomal proteins. Mech. Dev. 52:209-223.[CrossRef][Medline]
35. Strahl, B. D., and C. D. Allis. 2002. The language of covalent histone modifications. Nature 403:41-45.
36. Theobald, D. L., R. M. Mitton-Fry, and D. S. Wuttke. 2003. Nucleic acid recognition by OB-fold proteins. Annu. Rev. Biophys. Biomol. Struct. 32:115-133.[CrossRef][Medline]
37. Turner, B. M. 2000. Histone acetylation and an epigenetic code. Bioessays 22:836-845.[CrossRef][Medline]
38. Verdel, A., S. Jia, S. Gerber, T. Sugiyama, S. Gygi, S. I. Grewal, and D. Moazed. 2004. RNAi-mediated targeting of heterochromatin by the RITS complex. Science 303:672-676.
39. Volpe, T. A., C. Kidner, I. M. Hall, G. Teng, S. I. Grewal, and R. A. Martienssen. 2002. Regulation of heterochromatic silencing and histone H3 lysine 9 methylation by RNAi. Science 297:1833-1837.
40. Yu, B. D., R. D. Hanson, J. L. Hess, S. E. Horning, and S. J. Korsmeyer. 1998. MLL, a mammalian trithorax-group gene, functions as a transcriptional maintenance factor in morphogenesis. Proc. Natl. Acad. Sci. USA 95:10632-10636.
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