As chromatin remodeling and transcription factor binding augment transcription and replication, and are themselves increased by these processes, a temporal.
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Oncogene (2001) 20, 3094 ± 3099 2001 Nature Publishing Group All rights reserved 0950 ± 9232/01 $15.00 www.nature.com/onc
Chromatin alteration, transcription and replication: What's the opening line to the story? Michelle Craig Barton*,1 and Alison J Crowe2 1
Department of Biochemistry and Molecular Biology, University of Texas, M.D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, Texas, TX 77030, USA; 2Department of Zoology, University of Washington, Box 351800, Seattle, Washington, WA 98195-1800, USA
Polymerase accessibility to chromatin is a limiting step in both RNA and DNA synthesis. Unwinding DNA and nucleosomes during polymerase complex binding and processing likely requires priming by chromatin restructuring. The initiating step in these processes remains an area of speculation. This review focuses on the physical handling of chromatin during transcription and replication, the fate of nucleosomes assembled on DNA during unwinding and processing the chromatin substrate, and how these alterations in chromatin structure may aect gene expression. Transcription or replication may alter chromatin structure during synthesis, enabling regulatory factor binding and, potentially, future rounds of transcription. As chromatin remodeling and transcription factor binding augment transcription and replication, and are themselves increased by these processes, a temporal model of structural alterations and gene activation is built that may be more circular than linear. Oncogene (2001) 20, 3094 ± 3099. Keywords: chromatin; replication; transcription; polymerase; remodeling Introduction A ¯urry of scienti®c attention has focused on chromatin remodeling as a requisite step in gene activation. Interactions between previously de®ned chromatin remodeling complexes and nucleosomes/ histones are linked to enhanced gene transcription (recently reviewed in Kingston and Narlikar, 1999; Peterson and Workman, 2000; Sudarsanam and Winston, 2000; Wole and Guschin, 2000), as well as increased DNA replication (Flanagan and Peterson, 1999; Hu et al., 1999; Li, 1999; Li et al., 1998). Both of these operations rely on large, megadalton-size complexes of proteins, physically processing a chromatin substrate by lumbering along a nucleosome-assembled DNA template or serving as nuclear matrix-associated factory sites of synthesis (reviewed in Cook, 1999). Could physical machination of chromatin in this way,
*Correspondence: MC Barton, Department of Biochemistry and Molecular Biology, Box 117, University of Texas, M.D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030, USA
alter its structure and aect replication and transcription eciency? We will consider the fate of nucleosomes and chromatin structure during transcription and replication, and review the consequences for gene expression. The substrate is chromatin Chromatin as a hindrance to transcription activation can inhibit binding of upstream (or downstream) regulatory proteins as well as assembly of a preinitiation complex. Once the polymerase complex is engaged, and an open complex formed, ecient elongation through the gene body requires unwrapping and decoding a nucleosome-assembled substrate. A polymerase complex, whether RNA or DNA synthesizing, must somehow gain access to the correct strand of DNA, reliably copy and transduce the information and continue along the template (reviewed in Kodadek, 1998). During elongation, the encroaching polymerase complex could either (a) force complete release of nucleosomes, with re-assembly occurring de novo in the wake of the processive complex, or (b) maintain interaction between at least one strand of DNA and nucleosomes or component histones in a type of ternary complex. This interaction could facilitate a continuum of regulation where a nucleosomal binding pattern is maintained or re-established quickly. However, the time and modi®cation processes required to reset chromatin to its ground state could provide a window of structural plasticity during DNA and/or RNA polymerase operations. The transcription elongation process is one of starts, stops and stutters; the consequences of which may be modulated by trans-acting elongation factors such as TFIIS, Elongin, ELL factors and others (reviewed in Conaway et al., 2000; Orphanides and Reinberg, 2000). Over a number of years, elegant studies by the Felsenfeld laboratory and their collaborators support a model where contact between histones and a DNA template is maintained throughout elongation (Bednar et al., 1999; Clark and Felsenfeld, 1992; Studitsky et al., 1994, 1995, 1997; Williamson and Felsenfeld, 1978). In vitro investigations with E. coli RNA polymerase, SP6 bacteriophage RNA polymerase, S. cerevisiae RNA polymerase III and histone/mononucleosome-associated
Gene activation by replication and transcription MC Barton and AJ Crowe
DNA templates of varying lengths and sequence composition, reveal that a nucleosome virtually steps around the processive polymerase. A nucleosome from the leading edge of the polymerase machinery is internally transferred via a DNA loop to promoterproximal DNA. Arrested, short-lived RNA polymerase/ DNA intermediates have been visualized by electron cryomicroscopy at the predicted positions along the DNA template. Traversing the intranucleosomal DNA loop substrate, RNA polymerases pause with a periodic 10-11 base pair frequency (Bednar et al., 1999). The elongating RNA polymerase II complex: Nimble dancer or chromatin snowplow? The collective eects of nucleosome-mediated pausing on a higher-order chromatin substrate are unknown. Multiple lines of evidence reveal that chromatin remodeling and/or histone modi®cation expedites the rather inecient transcription elongation on a multinucleosome DNA template. ATP-dependent chromatin remodeling complexes and histone modi®ers may render transcription more ecient by modifying or remodeling chromatin along the gene body or stabilizing nucleosome/loop formation (Brown and Kingston, 1997; Walia et al., 1998; reviewed in Workman and Kingston, 1998). Biochemical studies revealed the importance of the protein FACT (facilitates chromatin transcription), which was puri®ed by its ability to augment elongation processing of chromatin-assembled templates (Orphanides et al., 1998; reviewed in Orphanides and Reinberg, 2000). FACT works in a two-step, ordered combination with chromatin-remodeling factor RSF, which promotes transcription initiation. RSF-mediated nucleosome binding disruption, is localized to sites near the promoter while FACT acts more downstream, binding and modifying histones H2A and H2B interactions (LeRoy et al., 1998; Orphanides et al., 1999). Several proteins categorized originally by their roles in transcription elongation are also modi®ers of chromatin structure or associated with chromatin regulation. These include the elongation factor DSIF, a human homologue of yeast Spt4 and Spt5 proteins (Wada et al., 1998), and elongation factor TFIIS, which was associated with the SWI/SNF chromatin remodeling complex by a synthetic lethal screen in S. cerevisiae (Davie and Kane, 2000). In combination with disabled TFIIS activity (elongation promotion), chromatin remodeling and/or histone modi®cation is essential for transcription of a greater number of genes than seen with SWI/SNF disruption alone (Davie and Kane, 2000). Additionally, the SWI/SNF complex has a continued requirement in downstream events following transcription initiation, implying a potential role for chromatin remodeling during elongation (Biggar and Crabtree, 1999). Perhaps the most intriguing link between chromatin structure alteration and an elongating RNA polymerase complex is that remodeling and modi®er proteins have been identi®ed as members of the polymerase
holoenzyme complex. Various laboratories have puri®ed RNA polymerase II holoenzyme preparations with SWI/SNF complex members and histone acetyltransferases CBP and P/CAF among their components (Cho et al., 1998; Neish et al., 1998; Wilson et al., 1996). This puri®ed holoenzyme must necessarily be disengaged from its substrate, and not every biochemical preparation of the holoenzyme RNA polymerase II complex contains these remodeling factors (Cairns et al., 1996; Kershnar et al., 1998; Neish et al., 1998). However, additional studies have focused on isolation of an engaged, elongating form of an RNA polymerase II complex (Otero et al., 1999; Wittschieben et al., 1999). Associated with the hyperphosphorylated RNA polymerase II C-terminal domain (CTD) in a complex with yeast chromatin and RNA, is a transcription elongation factor called Elongator. One of the Elongator subunits is a novel and highly conserved histone acetyltransferase that is shared with the SAGA complex, and can acetylate all four histone proteins (Wittschieben et al., 1999, 2000). Chromatin remodeling proteins and histone modi®ers in alliance with RNA polymerase II evoke a previously used metaphor of an elongating RNA polymerase as a chromatin snowplow. Does chromatin structure remain altered in a measurable way in the wake of the snowplow? Are there opposing forces in equilibrium with remodeling factors and histone acetyltransferases that re-establish a resting state to transcribed chromatin after passage of a polymerase complex?
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The trail of transcription The relative depletion of histones H2A and H2B in transcribed chromatin has been invoked as intriguing evidence for nucleosome alteration post-transcription (Baer and Rhodes, 1983). Sogo and colleagues, using accessibility to psoralen crosslinking and 2D-gel eletrophoresis analysis, show that chromatin structure `opening' is initiated by RNA polymerase I transcribing ribosomal RNA gene repeats. The altered, active chromatin is disrupted locally, and occupies relatively short stretches of DNA along the path of the processing polymerase complex (Lucchini and Sogo, 1995). These results imply that active chromatin structure, which contains disrupted nucleosomes and modi®ed histones, is a short-lived intermediate that is not transmitted or copied during replication. Opposing forces stripping histones of their modi®cation and returning nucleosomes to full octamer composition must therefore be targeted to recently transcribed chromatin. However, timing may be very important in interactions between regulatory factors and modi®ed chromatin; and, dierent measurements of chromatin alteration induced by RNA polymerase II have revealed further information as well. More recent functional studies of RNA polymerase II action reveal that ongoing transcription may potentiate further chromatin activation (Gribnau et al., 2000). Oncogene
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Convincing evidence from Fraser and colleagues supports a model where transcription begat transcription, possibly by physically perturbing chromatin as a template (Ashe et al., 1997; Gribnau et al., 2000). Building on observations made in the 1970's that large, likely noncoding, transcripts exist across the globin genes of chick and murine loci (Bastos and Aviv, 1977; Imaizumi et al., 1973), these authors ®nd that intergenic transcription is unidirectional, regulated and initiates just prior to developmental activation of stage-speci®c globin genes. Mapping nascent RNA transcripts using in situ hybridization reveals three subdomains of intergenic transcription within the human b-globin locus (Gribnau et al., 2000). Termed domain transcripts, these RNAs initiate at speci®c, intergenic start sites, terminate within sharp boundaries of closed chromatin structure and are not dependent on concomitant globin gene-speci®c transcription. Further, deletion of a domain transcript initiation region results in considerable loss of downstream globin gene expression. Taken together, a picture is built of RNA polymerase II holoenzyme travel and transcription across a gene locus that opens chromatin structure for further alteration and speci®c, regulated gene expression (reviewed in Travers, 1999). Chromatin with potential How could `priming' transcription occur in a repressed chromatin domain? In a ®eld concerned with initiating, regulatory events in chromatin structural alteration and gene expression, transcription activating transcription poses a prototypical chicken-and-egg problem. The concept of `transcriptional competence' across an entire gene locus, distinct from transcriptional activity, is supported by structural studies of the chick b-globin locus. Locus-wide, tissue-speci®c sensitivity to DNaseI digestion, 5 ± 10-fold greater than ¯anking genomic DNA and globin loci in non-expressing tissues, comaps with broad, locus-wide histone hyperacetylation. These tissue-speci®c alterations in globin locus chromatin structure occur prior to more localized, developmentally regulated DNaseI hypersensitive sites where regulatory factors interact with sequence-speci®c DNA elements (Hebbes et al., 1988, 1994). Utilizing chromatin immunopreciptiation analysis of the S. cerevisiae genome, the Grunstein laboratory ®nds that widespread, global histone modi®cation exists across gene loci (Vogelauer et al., 2000). Unique to these analyses is the level of quantitation applied over a relatively broad region of the genome, and the manipulation of histone acetyltransferase and deacetylase expression levels by gene deletion. These authors ®nd that multiple chromosomal domains exhibit an equilibrium between acetylation and deacetylation, one that can be shifted rapidly but favors a general underacetylated condition. In a state dictated by a balance of enzymatic actions, chromatin structure may be more dynamic than previously considered. Targeting by a minimal number of DNA-bound regulatory
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factors could shift the equilibrium toward chromatin activation. Further analyses of yeast and mammalian genomes in this way should reveal whether all euchromatin can be characterized by histone modi®cation at a quanti®able level, in¯uenced by tissue-, developmental-, and ligand/signal-speci®c factors that tip the balance toward activation or repression of chromatin structure. The case for DNA polymerase: a matter of timing A recent review by Orphanides and Reinberg (Orphanides and Reinberg, 2000) discusses the concept of a `pioneer [RNA] polymerase'. This polymerase would be specially equipped to pry open unmodi®ed, fully repressed chromatin, between borders of transcription silencing, for future rounds of transcription. By modifying histones and remodeling chromatin structure with its appropriate protein subunits, the pioneer polymerase complex could initiate chromatin structure alteration to a level of readiness amenable to regulatory factor interactions with DNA. As noted by these same authors, there is evidence for primed chromatin structure (discussed above), but little experimental support for a special-forces RNA polymerase. Another candidate to consider for the title of pioneer polymerase, ful®lling the quali®cations of multi-subunit protein complex that traverses gene loci and manipulates chromatin, is DNA polymerase. The correlative links between DNA polymerase action and gene expression are numerous and attractive at several levels. Surveys of housekeeping genes, tissuespeci®c and developmentally expressed genes established the dogma that early replication (in S phase of the cell cycle) means active gene expression (Goldman et al., 1984). Comparison of a single gene in highly expressing cells versus non-expressing cells, such as bglobin in red blood cells versus HeLa cells, reveals that b-globin gene replication timing diers markedly between the two cell types. Further, during red cell development, the timing of b-globin replication switches concomitantly with developmental expression of the gene(s) (reviewed in Groudine et al., 1989; Groudine and Weintraub, 1981; Kitsberg et al., 1993). Replication late in the cell cycle correlates with silenced gene expression, such as the mammalian inactive X chromosome and yeast silenced mating type genes compared to their actively expressed counterparts, which are replicated early (reviewed in Fangman and Brewer, 1992; Raghuraman et al., 1997). Direct cause-and-eect between DNA replication and gene transcription activation is more dicult to establish. Recent analyses of inactive X chromosomes isolated from patients with ICF (Immunode®ciency, centromeric region instability, and facial anomalies) syndrome indicate that late replication is sucient to confer a silenced state even in the presence of transactivator binding (Hansen et al., 2000). ICF is caused by a point mutation in the methyl transferase gene DNMT3B. The resulting hypomethylation of
Gene activation by replication and transcription MC Barton and AJ Crowe
inactive X chromosomes and escape from silencing of X-inactivated genes occurs concomitantly with a shift in replication timing, linking the processes of replication and transcription. The switch from silenced to activated requires multiple steps, including advanced replication timing, hypomethylation, chromatin remodeling and transcription factor binding (Litt et al., 1997; Sasaki et al., 1992). Again, it is not clear in what order these steps must proceed but, with each progressive alteration, there is an increased probability that a gene will escape silencing. The consequences of replication DNA replication results in the transient disruption of 1 ± 2 nucleosomes at the replication fork; additionally, nucleosomes assembled de novo after replication fork passage may be transiently hyperacetylated and depleted in histones H2A and H2B (reviewed in Verreault, 2000; Wole, 1991). As a result, there may be a brief opportunity for activators and/or repressors to bind DNA regulatory sites normally occluded by nucleosomes, and establish chromatin structure that diers from the original replication substrate. Chromatin replication in synthetic nuclei directly links alterations in chromatin structure mediated by passage of a replication fork and activation of gene expression (Barton and Emerson, 1994; Crowe et al., 2000). Cytoplasmic Xenopus laevis egg extracts assemble chromatin and an encapsulating, nuclear membrane on DNA templates in vitro (Newport, 1987). Coupled transcription analyses of chick b-globin and mouse alpha-fetoprotein genes replicated in synthetic nuclei, demonstrates replication-dependent transcription activation in the presence of tissue- and developmentalspeci®c trans-activators. Chromatin assembled in the presence of trans-activators during primed, singlestranded DNA replication is activated for expression and repressed when uncoupled from replication (Almouzni et al., 1990; Kamakaka et al., 1993). Returning to the previous study of intergenic domain transcription across de®ned regions of the b-globin locus (Gribnau et al., 2000), a temporal order of transcription was established. In situ hybridization and quanti®ed measurements of nascent transcripts at speci®c times during the cell cycle reveals periodic domain transcription at the time of globin replication early in S phase, but also in late G1. The long-term eects on chromatin structure and globin gene-speci®c expression follow the early replication and transcription of locus subdomains, perhaps working in concert. Replication-independent gene activation Studies by Sogo and colleagues, using techniques of psoraren accessibility, crosslinking, 2-dimensional gel electrophoresis and electron microscopy, focus on the relationship between DNA replication and transcription of ribosomal RNA genes (Lucchini and Sogo,
1995; Muller et al., 2000). Several ®ndings are of interest: (1) there is no measurable dierence between the lengths of the short nucleosome-free regions at replication forks within transcriptionally active and transcriptionally inactive chromatin; (2) the amount of time it takes to re-assemble nucleosomes in the wake of DNA synthesis is also equal between the two types of chromatin; and (3) active origins of replication occur primarily downstream of an actively transcribing rRNA gene and not upstream. These authors further suggest that it is transcription factor binding at a rRNA enhancer, maintained in an open nucleosome-free structure, that activates a downstream replication origin and blocks replication fork movement upstream from the origin (Muller et al., 2000). The open chromatin structure of the rRNA enhancer does not appear to be dependent on ongoing transcription or replication, but may be highly sequence-directed. How these activities compare to the majority of RNA polymerase II-transcribed genes with ill-de®ned, broad regions of replication origin needs to be determined. A recent study of the yeast heat shock factor (HSF) transcriptional activator found that HSF binding to its nucleosomal site in vivo requires an S phase-speci®c event that is replication-independent (Venturi et al., 2000). These results indicate that, in addition to DNA replication, there are other cell cycle-linked cellular processes that facilitate factor binding in early S phase. There is precedent for cell cycle regulation of chromatin remodeling complexes: Human SWI/SNF complexes are inactivated during mitosis through phosphorylation, potentially to allow formation of a transcriptionally repressed chromatin structure (Sif et al., 1998). Inactivation of SWI/SNF during mitosis may be necessary to allow chromosome condensation to proceed in the absence of active remodeling. It is possible, therefore, that S phase-speci®c induction of SWI/SNF or related remodeling activities may enhance transcription factor binding during this critical stage of the cell cycle.
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The question remains Though DNA replication aords an opportunity for initiating changes in chromatin structure and gene expression as replication forks move through the genome, there is no unifying picture of DNA replication as the global, primary modi®er of chromatin structure. Like transcription activation, DNA replication can be ampli®ed through binding or tethering a diverse array of transcription factors at certain ARS (autonomously replicating sequence) elements in S. cerevisiae (Hu et al., 1999; Li, 1999; Li et al., 1998). Chromatin remodeling occurs at transcription factor-bound ARS elements, presumably through targeting of ATP-dependent chromatin remodeling complexes such as SWI/SNF (Flanagan and Peterson, 1999). We return again to the question of how these transcription factors gain accessibility to Oncogene
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repressed chromatin to begin the cycle of chromatin remodeling, replication and transcription. The ability of transcription factors to bind their sites when nucleosome-assembled is limited but highly variable (Adams and Workman, 1995). Proteins such as Swi5p may readily interact with chromatin and bind, targeting the SWI/SNF chromatin remodeling complex, which then interacts with the SAGA complex, permitting further interaction with transcription factors at the HO gene locus in yeast (Cosma et al., 1999). As other gene regulatory models are assessed for the temporal process of transcription activation (Dilworth
et al., 2000; Gribnau et al., 2000), there will likely emerge gene- or gene locus-speci®c initiators of a sequential order of stages in gene expression. It is likely that individual genes depend on highly regulated ways of accessing chromatin structure, the combination of which may be unique. An imbalance in any one of the means of altering chromatin structure, e.g. increased DNA replication, unregulated transcription or disrupted chromatin modi®er interactions, could feed into a cascade of gene regulatory dysfunction through escalating changes in chromatin structure.
References Adams CC and Workman JL. (1995). Mol. Cell. Biol., 15, 1405 ± 1421. Almouzni G, Mechali M and Wole AP. (1990). EMBO J., 9, 573 ± 582. Ashe JL, Monks J, Wijgerde M, Fraser P and Proudfoot NJ. (1997). Genes Dev., 11, 2494 ± 2509. Baer BW and Rhodes D. (1983). Nature, 301, 482 ± 488. Barton MC and Emerson BM. (1994). Genes Dev., 8, 2453 ± 2465. Bastos RM and Aviv H. (1977). Cell, 11, 641 ± 650. Bednar J, Studitsky VM, Grigoryev SA, Felsenfeld G and Woodcock CL. (1999). Mol. Cell, 4, 377 ± 386. Biggar SR and Crabtree GR. (1999). EMBO J., 18, 2254 ± 2264. Brown SA and Kingston RE. (1997). Genes Dev., 11, 3116 ± 3121. Cairns BR, Lorch Y, Li Y, Zhang M, Lacomis L, ErdjumentBromage H, Tempst P, Du J, Laurent B and Kornberg RD. (1996). Cell, 87, 1249 ± 1260. Cho H, Orphanides G, Sun X, Yang XJ, Ogryzko V, Lees E, Nakatani Y and Reinberg D. (1998). Mol. Cell. Biol., 18, 5355 ± 5363. Clark DJ and Felsenfeld G. (1992). Cell, 71, 11 ± 22. Conaway JW, Shilatifard A, Dvir A and Conaway RC. (2000). Trends Biochem. Sci., 25, 375 ± 380. Cook PR. (1999). Science, 284, 1790 ± 1795. Cosma MP, Tanaka T and Nasmyth K. (1999). Cell, 97, 299 ± 311. Crowe AJ, Piechan JL, Sang L and Barton MC. (2000). Mol. Cell. Biol., 20, 4169 ± 4180. Davie JK and Kane CM. (2000). Mol. Cell. Biol., 20, 5960 ± 5973. Dilworth FJ, Fromental-Ramain C, Yamamoto K and Chambon P. (2000). Mol. Cell, 6, 1049 ± 1058. Fangman WL and Brewer BJ. (1992). Cell, 71, 363 ± 366. Flanagan JF and Peterson CL. (1999). Nucleic Acids Res., 27, 2022 ± 2028. Goldman MA, Holmquist GP, Gray MC, Caston LA and Nag A. (1984). Science, 224, 686 ± 692. Gribnau J, Diderich K, Pruzina S, Calzolari R and Fraser P. (2000). Mol. Cell, 5, 377 ± 386. Groudine M, Forrester WC, Novak U and Epner E. (1989). Prog. Clin. Biol. Res., 316A, 15 ± 35. Groudine M and Weintraub H. (1981). Cell, 24, 393 ± 401. Hansen RS, Stoger R, Wijmenga C, Stanek AM, Can®eld TK, Luo P, Matarazzo MR, D'Espositio M, Feil R, Gimelli G, Weemaes CM, Laird CD and Gartler SM. (2000). Hum. Mol. Genet., 9, 2575 ± 2587. Hebbes TR, Clayton AL, Thorne AW and Crane-Robinson C. (1994). EMBO J., 13, 1823 ± 1830. Oncogene
Hebbes TR, Thome AW and Crane-Robinson C. (1988). EMBO J., 7, 1395 ± 1402. Hu YF, Hao ZL and Li R. (1999). Genes Dev., 13, 637 ± 642. Imaizumi T, Diggelmann H and Scherrer K. (1973). Proc. Natl. Acad. Sci. USA, 70, 1122 ± 1126. Kamakaka RT, Bulger M and Kadonaga JT. (1993). Genes Dev., 7, 1779 ± 1795. Kershnar E, Wu SY and Chiang CM. (1998). J. Biol. Chem., 273, 34444 ± 34453. Kingston RE and Narlikar GJ. (1999). Genes Dev., 13, 2339 ± 2352. Kitsberg D, Selig S, Keshet I and Cedar H. (1993). Nature, 366, 588 ± 590. Kodadek T. (1998). Trends Biochem. Sci., 23, 79 ± 83. LeRoy G, Orphanides G, Lane WS and Reinberg D. (1998). Science, 282, 1900 ± 1904. Li R. (1999). J. Biol. Chem., 274, 30310 ± 30314. Li R, Yu DS, Tanaka M, Zheng L, Berger SL and Stillman B. (1998). Mol. Cell. Biol., 18, 1296 ± 1302. Litt MD, Hansen RS, Hornstra IK, Gartler SM and Yang TP. (1997). J. Biol. Chem., 272, 14921 ± 14926. Lucchini R and Sogo JM. (1995). Nature, 374, 276 ± 280. Muller M, Lucchini R and Sogo JM. (2000). Mol. Cell., 5, 767 ± 777. Neish AS, Anderson SF, Schlegel BP, Wei W and Parvin JD. (1998). Nucleic Acids Res., 26, 847 ± 853. Newport J. (1987). Cell, 48, 205 ± 217. Orphanides G, LeRoy G, Chang CH, Luse DS and Reinberg D. (1998). Cell, 92, 105 ± 116. Orphanides G and Reinberg D. (2000). Nature, 407, 471 ± 475. Orphanides G, Wu WH, Lane WS, Hampsey M and Reinberg D. (1999). Nature, 400, 284 ± 288. Otero G, Fellows J, Li Y, de Bizemont T, Dirac AM, Gustafsson CM, Erdjument-Bromage H, Tempst P and Svejstrup JQ. (1999). Mol. Cell, 3, 109 ± 118. Peterson CL and Workman JL. (2000). Curr. Opin. Genet. Dev., 10, 187 ± 192. Raghuraman MK, Brewer BJ and Fangman WL. (1997). Science, 276, 806 ± 809. Sasaki T, Hansen RS and Gartler SM. (1992). Mol. Cell. Biol., 12, 3819 ± 3826. Sif S, Stukenberg PT, Kirschner MW and Kingston RE. (1998). Genes Dev., 12, 2842 ± 2851. Studitsky VM, Clark DJ and Felsenfeld G. (1994). Cell, 76, 371 ± 382. Studitsky VM, Clark DJ and Felsenfeld G. (1995). Cell, 83, 19 ± 27. Studitsky VM, Kassavetis GA, Geiduschek EP and Felsenfeld G. (1997). Science, 278, 1960 ± 1963.
Gene activation by replication and transcription MC Barton and AJ Crowe
Sudarsanam P and Winston F. (2000). Trends Genet., 16, 345 ± 351. Travers A. (1999). Proc. Natl. Acad. Sci. USA, 96, 13634 ± 13637. Venturi CB, Erkine AM and Gross DS. (2000). Mol. Cell. Biol., 20, 6435 ± 6448. Verreault A. (2000). Genes Dev., 14, 1430 ± 1438. Vogelauer M, Wu J, Suka N and Grunstein M. (2000). Nature, 408, 495 ± 498. Wada T, Takagi T, Yamaguchi Y, Ferdous A, Imai T, Hirose S, Sugimoto S, Yano K, Hartzog GA, Winston F, Buratowski S and Handa H. (1998). Genes Dev., 12, 343 ± 356. Walia H, Chen HY, Sun JM, Holth LT and Davie JR. (1998). J. Biol. Chem., 273, 14516 ± 14522.
Williamson P and Felsenfeld G. (1978). Biochemistry, 17, 5695 ± 5705. Wilson CJ, Chao DM, Imbalzano AN, Schnitzler GR, Kingston RE and Young RA. (1996). Cell, 84, 235 ± 244. Wittschieben BO, Fellows J, Du W, Stillman DJ and Svejstrup JQ. (2000). EMBO. J., 19, 3060 ± 3068. Wittschieben BO, Otero G, de Bizemont T, Fellows J, Erdjument-Bromage H, Ohba R, Li Y, Allis CD, Tempst P and Svejstrup JQ. (1999). Mol. Cell, 4, 123 ± 128. Wole AP. (1991). J. Cell. Sci., 99, 201 ± 206. Wole AP and Guschin D. (2000). J. Struct. Biol., 129, 102 ± 122. Workman JL and Kingston RE. (1998). Annu. Rev. Biochem., 67, 545 ± 579.
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