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Dynamics of chromatin, proteins, and bodies within thecell nucleusAndrew Belmont
A new and still evolving paradigm of a highly dynamic nucleus
has emerged in recent years. This paradigm includes an
inherently high turnover rate of histone and non-histone protein
modifications, targeted turnover and/or displacement of stable
core histone proteins, constant flux of macromolecules through
chromosomes and nuclear bodies, including transcription
factors and co-activators, and an energy-dependent facilitation
of nuclear-protein complex formation and disassembly. Also
included are fast, local movements of chromosomes, together
with slower but long-range movements of chromosomes and
nuclear bodies.
Addresses
Department of Cell and Structural Biology, University of Illinois,
Urbana-Champaign B107, Chemical and Life Science Laboratory,
601 South Goodwin Avenue, Urbana, IL 61801, USAe-mail: [email protected]
Current Opinion in Cell Biology 2003, 15:304310
This review comes from a themed issue on
Nucleus and gene expression
Edited by Jeanne Lawrence and Gordon Hager
0955-0674/03/$ see front matter
2003 Elsevier Science Ltd. All rights reserved.
DOI 10.1016/S0955-0674(03)00045-0
Abbreviations
ChIP chromatin immunoprecipitationER oestrogen receptorFRAP fluorescence recovery after photobleachingGFP green fluorescent protein
GR glucocorticoid receptor
HAT histone acetyltransferasePML promyelocytic leukaemiapol RNA polymerase
SMN survival motor neuron
Introduction
Over the past two decades, the dynamics of the cyto-skeleton and cytoplasmic organelles have emerged as
central themes in cell biology. By contrast, the interphase
nucleus appeared to be the last refuge for the concept of
stable cell structures and compartmentalisation. Trans-
mission light microscopy of living cells revealed nuclei
whose shape and internal substructures showed little
apparent movement. At the molecular level, epigenetic
programmes of gene activity, such as X chromosome
inactivation, were demonstrated as being stable, once
established, for many cell generations, even in the
absence of factors required for initiation of silencing,
with chromatin proteins such as histones showing little
turnover. Moreover, biochemical in vitro reconstitution
experiments led to sequential models of assembly for
transcription pre-initiation complexes, over time periods
of tens of minutes, with the idea that these complexes,
once assembled, would be stable for tens of minutes to
hours, as observed for transcription factorDNA interac-
tions in vitro. Here, I review recent experiments that are
beginning to reveal instead a highly dynamic cell nucleus
whose components are in constant flux and movement.
Dynamics of chromosomal proteinsDNA folding on the nucleosome surface inhibits binding
of most chromosomal proteins. Slow, intrinsic nucleo-
some dynamics occurs through a breathing mechanism,
most likely involving progressive uncoiling from the
DNA entry or exit points on the core particle, leading
to transient release of DNA from the histone surface [1].
Although this breathing allows DNA binding of a wide
range of proteins, the binding rate is low. Pioneer
transcription factors capable of binding to compacted
nucleosome arrays and directly remodelling these arrays
have been identified [2]. Other transcription factors that
bind early in the activation process lead to chromatinremodelling through recruitment of chromatin-remodel-
ling and histone-modification complexes. In both cases,
chromatin remodelling and histone modifications may,through unmasking binding sites, facilitate binding of
other regulatory proteins. Histone modifications can
recruit other specific proteins to the site, according, to
a combinatorial or histone code [3], while in some cases
changing structural properties of individual nucleosomes
or their folding into higher-order chromatin structures.
What is new over the past year is a growing appreciation for
the existence of global histone acetylation and deacetyla-
tion activity, acting over large regions of the genome in
addition to locally targeted histone-modification activities
[4]. This allows rapid reversal of targeted histone acet-
ylation or deacetylation at regulatory regions after removalof the targeting signal. In yeast, using a TetR-targeted
VP16 activation domain, recovery to baseline histone
acetylation levels occurred within 2 min after removal
of targeted histone acetyltransferases (HATs) and 6 min
for targeted Rpd3 histone deacetylase [5]. This implies
an inherently high turnover cycle of core-histone modifi-
cations, in turn implying a constitutive high capacity for
dynamic chromatin remodelling throughout the genome.
In higher eukaryotic cells, there are indications that
particular histone modifications might be targeted not
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only to local DNA regions surrounding regulatory regions
but to large genome regions as well, hundreds to thou-
sands of kilobases in size. This includes not only large
heterochromatin domains, enriched in repetitive seq-
uences, but also euchromatic regions; for example,
domains of increased histone H3 acetylation are observedwithin interphase nuclei [6]. Recently, functional geno-
mics analysis has revealed clustering of coordinately
expressed genes in organisms as diverse as Drosophila,
Caenorhabditis elegans and humans [79]. Therefore, it is
not clear to what degree these locally enriched chromo-
somal patches reflect targeting to large genome regions
per se, versus the integrated signal from several different
gene loci sharing similar activity states. However, one can
distinguish between these two possibilities in the case of
DNA repair. UV irradiation leads to nuclear foci enrichedin DNA-repair proteins; each individual focus represents
the site of a DNA break. One of the earliest markers for
these foci is Ser139 phosphorylation within 110 min in
the H2AX histone variant. Phosphorylation of H2AX
occurs in microscopically large foci corresponding to
megabasepair DNA domains [10]. Changes in chromatin
structure resulting from DNA breaks, rather than the
break itself, have been proposed as the initial signalling
event leading to this highly delocalised response [11].
Given the emergence of the central role of core-histone
modifications in regulating DNA function over the pastfew years, it is natural that the dynamics of these mod-
ifications have been the focus of much work. However,
the recent discovery of core-histone methylation as a key
biological regulatory mechanism for transcriptional acti-
vation and silencing [3], together with the failure toidentify any histone demethylases, has turned attention
back to what had been thought was minimal turnover of
the core histones themselves. In vitro experiments, how-
ever, have demonstrated displacement of H2AH2B
dimers during transcription [12], and in vivo fluores-
cence recovery after photobleaching (FRAP) experi-
ments with green fluorescent protein (GFP)-labelledhistones demonstrates increased turnover of H2A and
H2B relative to H3 and H4, which decreases with tran-
scriptional inhibition [13]. The discovery of a histone H3
variant, H3.3, which is assembled into chromatin in a
pathway independent of replication, has suggested a
possible mechanism for turnover of methylated histoneH3 [14]. Recent studies reveal a replication-independent
deposition of histone H3.3 over transcriptionally active
ribosomal genes [15].
Thus, there exists a fast turnover of histone modifications
such as acetylation and phosphorylation, on a timescale of
minutes, and a slow turnover of core histones and variant
core histones, on a timescale of hours, which may con-
tribute to a longer-term epigenetic memory. In some
species, DNA methylation might further extend this
epigenetic memory. In between these two timescales is
the flux of linker histones H1 and high-mobility group
protein (HMG) through both open and condensed chro-
matin on a timescale of tens of seconds [16], which, at
least in the case of histone H1, can be modified by
phosphorylation [17].
Interestingly, application of FRAP to topoisomerase II,
a component of the nuclear matrix and chromosome
scaffold, revealed rapid in vivo exchange rates on a time-
scale of just 1020 s within interphase nuclei and mitotic
chromosomes [18,19]. These results challenge the
proposed role of topoisomerase II as a structural compo-
nent of chromosomes.
Dynamics of the transcriptional machineryAn obvious researchdirection has been to temporally orderchromatin modification events during gene activation
at specific loci. Using chromatin immunoprecipitation
(ChIP), several groups have successfully demonstrated
a specific sequence of histone modifications, together
with recruitment of different chromatin regulatory com-
plexes and co-activators, over a timescale of minutes to
tens of minutes, as reviewed recently [20]. Although
several genes investigated show recruitment of the same
chromatin-modification complexes, their order of recruit-
ment may differ; for instance, SWISNF recruitment
follows GCN5 recruitment and histone H3 acetylation
for the interferon-b promoter, but precedes recruitmentof the GCN5-containing SAGA (Spt-Ada-Gcn5-acetyl-
transferase) complex for the yeast HO promoter. These
results might suggest that multiple paths can lead to the
same activated state. Alternatively, the temporal order
may matter very much, with different orders of recruit-ment resulting in physiologically relevant differences in
activation temporal profiles or susceptibility to epigenetic
influences. For example, many yeast genes expressed
near mitosis require SWISNF activity followed by
GCN5 action; in the absence of SWISNF remodelling,
mitotic chromosome condensation might block GCN5
activity [21].
Biochemical purification of chromatin regulatory com-
plexes as distinct structural entities in the megadalton
size range has raised the natural question of how such
large complexes access condensed chromatin. An immu-
nofluorescence approach was used to monitor co-activatorrecruitment and histone modifications following targeting
of the VP16 acidic activation domain to an engineered
heterochromatic chromosome region. Unexpectedly, dif-
ferent subcomponents of both SWISNF and HAT com-
plexes showed quite different temporal recruitment
patterns, with some subcomponents accumulating within
several minutes after VP16 targeting and others after tens
of minutes [22]. These results suggest that rather than
diffusing into condensed chromatin as intact entities,
large chromatin-modifying complexes might assemble
on the gene target, possibly with individual subunits or
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subcomplexes acting as pioneer factors, opening their
target for subsequent binding of the intact complex.
Similarly, despite purification of a large RNA polymerase
II (pol II) holoenzyme containing the mediator complex,
CBP, and other proteins, ChIP has shown separate timing
of pol II and mediator recruitment [20
] Sequentialrecruitment also has been suggested for components of
DNA-repair holocomplexes [23].
Bigger surprises have come from in vivo FRAP mea-
surements, which have demonstrated off rates for trans-
cription factors binding to their target sequences, and
co-activators binding to their target proteins, orders of
magnitude faster than in vitro measured rates [24]. Intra-
nuclear recovery half lives of seconds to tens of seconds
have been measured for glucocorticoid receptor (GR)[25] and the GR-interacting protein 1 (GRIP-1) co-acti-
vator [26] on a chromosome array containing a
GR-responsive MMTV promoter, leading to a hit-
and-run model of transcriptional activation. Similar
dynamics have been observed for intranuclear oestrogen
receptor (ER) and the steroid receptor co-activator 1
(SRC-1) binding to ER in the presence of ligand
[27,28], with ER becoming immobilised in the nucleo-
plasm after exposure to proteosome inhibitors or hor-
mone antagonist [28]. The general transcription factor
TFIIB shows rapid turnover in vivo over seconds, but
TATA-binding protein (TBP) turnover is relatively slowwith a FRAP half life of minutes [29]; however, ChIP
experiments reveal TBP dissociation coincident with
removal of targeted VP16, within the 12 min experi-
mental time resolution [5]. FRAP of pol II subunits
shows a predominant component which slowly ex-changes over 1020 min; this component corresponds
to elongating polymerases [26,30]. A faster component,
which recovers over seconds to tens of seconds, might
correspond to turnover at the promoter [26].
Recently, an elegant FRAP analysis of pol I subunits and
transcription factors revealed similarly rapid dynamicsover ribosomal genes [31]. On the basis of a combination
of experiments and data modelling, it was concluded that
the pol I holoenzyme did not diffuse to its target pro-
moters as a pre-assembled structural entity in vivo, but
rather assembled at the promoter. These data highlight
the current paradox implicit in trying to reconcile in vitroand in vivo results. Specifically, a GFP-tagged pol I
holoenzyme could be isolated as a distinct, stable struc-
tural entity from cells expressing an epitope-tagged sub-
unit, but in vivo subunits showed dynamics inconsistent
with the idea of a pre-assembled holoenzyme.
Oneexplanationfor this paradox would be if thedynamic
behaviour observed in vivo depended on specific, energy-
dependent mechanisms facilitating exchange and
turnover. Using an in vitro chromatin reconstitution
system, Hager and co-workers [32] demonstrated that
the rapid turnover of GR on the MMTV promoter is
ATP-dependent and can be reproduced at least partially
by purified SWISNF, suggesting a turnover mechanism
involving chromatin-remodelling complexes. An alter-
native mechanism has emerged from studies showing
repression of ER transcriptional activation by the mole-cular chaperone p23, suggesting a role for disassembly
complexes fostering active turnover of large transcrip-
tion complexes [33].
Dynamics of chromosomesOne might expect passive chromosome movements sim-
ply as the result of chromatin decondensation/condensa-
tion; however, recent results indicate rapid and active
chromosome movements. Using chromosomes tagged
with GFPlac-repressor bound to lac-operator repeats, arapid, but locally constrained, motion has been observed
in yeast, Drosophila and mammalian cells [34,35], This
motion is rapid so rapid that current sampling rates on
the order of tens to hundreds of milliseconds are still too
slow adequately to follow the motion but localised to
excursions of a few hundred nanometres. Sites associated
with the nuclear periphery or nucleolus show lower
amplitude movements [36]. The motion varies according
to the physiological state of the cell, and ATP-depletion
experiments suggest that the motion is energy-depen-
dent [37]. In yeast, motion is reduced in S-phase cells,
which might reflect associations of chromosomes withthe DNA replication machinery and/or elements of
nuclear structure [38].
In Drosophila and yeast, long-range motions over greater
distances are observed over a longer timescale. In Dro-sophila, this long-range motion is developmentally regu-
lated, disappearing during later stages of spermatocyte
differentiation [39]. Absence of long-range chromosome
movements appears to be the general rule in mammalian
tissue-culture cells. However, at specific times during S
phase, long-range motions of specific, engineered chro-
mosome regions have been observed [40,41].
In mammalian cells, targeting of the acidic activator VP1
changed the position of a peripheral chromosome site to
the interior in stable transformants [41]. More recent data
show this induced motion occurs at a defined time after
VP16 targeting independent of cell cycle position(C Chuang and A Belmont, unpublished data). In addi-
tion, uncoiling of large-scale chromatin structure is
observed with VP16 targeting [42], similar to that induced
by GR in a chromosome array containing a GR-responsive
promoter [43]. These changes in position and conforma-
tion may recapitulate physiological changes in chromo-
some position and conformation observed for wild-type
chromosomes. Peripheral versus interior localisation of
gene-poor versus gene-rich chromosomes have been
described, as have changes in chromosome positioning
as a function of cell cycle progression [44].
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Dynamics of nuclear bodiesLong-range chromosome movements naturally raise
questions concerning whether these movements are tar-
geted to specific nuclear bodies. The list of distinct
nuclear suborganelles or compartments is growing, with
the addition over the past year of clastosomes [45
] andparaspeckles [46] to the established list of interchroma-
tin granule clusters (speckles), PML (promyelocytic
leukaemia), SMN (survival motor neuron) and Cajal
bodies, gems and nucleoli, as well as centromeric hetero-
chromatin and the nuclear periphery. Apparent localisa-
tion of a new multiple bridging protein to connections
between gems and SMN bodies [47] could suggest
functional connections between several of these bodies.
There is a long-standing observation of the association of
many active genes with interchromatin granule clusters(splicing speckles, Sc-35 bodies); recent data indicate
association of mRNA from these genes with these bodies
[48]. A growing literature points to concentrations of
specific and general transcriptionfactors within PML and
Cajal bodies, as well as physiological regulation of this
targeting (reviewed by Isogai and Tijan, this issue), and
both the histone and U2 gene loci specifically associate
with Cajal bodies [49,50]. A clear demonstration of a
role of Cajal bodies in the maturation of certain proteins
and ribonucleoprotein complexes [51,52,53,54] has
recently emerged, as well as a protein maturation path-
way from speckles to nucleoli [55]. Rapid trafficking ofproteins between nuclear bodies and the nucleoplasm
has also been demonstrated recently [46,56].
In addition to protein trafficking into and out of nuclear
bodies, the bodies themselves move within the nucleus.PML bodies move discontinuously with stationary peri-
ods interspersed with long-range movements at maxi-
mum velocities of 18 mm/min [57]. Their movements
are decreased by ATP depletion and reduced by the
myosin inhibitor 2,3-butanedione monoxime. By contrast,
Cajal bodies show an increase in long-range diffusive
movements and a decrease in immobile periods corre-lated with chromatin associations within live cells after
ATP depletion or transcriptional inhibition, suggesting
that transient associations with chromatin might be active
processes dependent on transcription [58].
Conclusions and future directionsIn this review, I have discussed dynamics of nuclear
proteins, chromosome loci, and nuclear bodies as if they
were distinct physiological processes. There already
exist strong hints that these processes are connected
functionally. For instance, recent data point to long-
range associations of elements within the globin locus
control region with distant promoter regions [59].
However, these same regulatory elements also regulate
chromatin modifications, as well positioning of the globin
locus relative to centromeric heterochromatin [60].
A specific transcription factor essential for globin gene
activation initially localises to these same centromeric
regions, with delocalisation to the nucleoplasm coincid-
ing with movements of the locus to the nuclear interior
and gene activation [61]. In this single example of regu-
lated gene activation, we connect dynamics of chromatin
modification, both local and long-range chromatin move-ments, and transcription-factor trafficking. Similar con-
nections have emerged between compartmentalisation
of transcription repressors, gene silencing, and intranuc-
lear localisation of gene loci [62]. Our future challenge
will be to understand how these different dynamic pro-
cesses together accomplish regulation of nuclear pro-
cesses. This will require continued development of
new technological approaches suitable for in vivo obser-
vations of nuclear dynamics.
References and recommended readingPapers of particular interest, published within the annual period ofreview, have been highlighted as:
of special interestof outstanding interest
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This study demonstrates that spontaneous access of DNA bound tonucleosome core proteins is produced by DNA breathing, produced bychanges in nucleosome conformation, as opposed to nucleosome sliding.
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Cirillo LA, Lin FR, Cuesta I, Friedman D, Jarnik M, Zaret KS:Opening of compacted chromatin by early developmentaltranscription factors HNF3 (FoxA) and GATA-4. Mol Cell 2002,9:279-289.
Using in vitro reconstituted chromatin including linker histone H1, thispaper demonstrates the capability of certain transcription factors to bindand remodel nucleosome arrays without the help of ATP-dependentchromatin-modifying complexes.
3. Turner BM: Cellular memory and the histone code. Cell 2002,111:285-291.
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Robyr D, Suka Y, Xenarios I, Kurdistani SK, Wang A, Suka N,Grunstein M: Microarray deacetylation maps determinegenome-wide functions for yeast histone deacetylases.Cell 2002, 109:437-446.
Chromatin immunoprecipitation is used with DNA chip analysis to com-pare histone acetylation patterns between wild-type yeast cells andknockouts of specific histone deacetylases (HDACs). This approachidentifies large subsets of genes whose histone acetylation patterns,and expression, are regulated by specific HDACs. Interestingly, Hda1appears to act preferentially over large 1025 kb genomic domainsflanking telomeres.
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Katan-Khaykovich Y, Struhl K: Dynamics of global histoneacetylation and deacetylation in vivo: rapid restoration ofnormal histone acetylation status upon removal of activatorsand repressors. Genes Dev 2002, 16:743-752.
In this work in yeast, very rapid global acetylation or deacetylation back tosteady-state levels is observed within several minutes after removal oftargetedrepressor (Ume6) or activator (VP16 acidic activator), respectively.The authors also show that TATA-binding protein (TBP) binding correlatesclosely with activator binding, rather than with histone acetylation.
6. Hendzel MJ,Kruhlak MJ,Bazett-JonesDP: Organization of highlyacetylated chromatin around sites of heterogeneous nuclearRNA accumulation. Mol Biol Cell 1998, 9:2491-2507.
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9. Spellman PT, Rubin GM: Evidence for large domains of similarlyexpressed genes in the Drosophila genome. J Biol 2002, 1:5.
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In vitro transcription through nucleosome core particles by RNA poly-merase II leads to a loss of one H2AH2B dimer, providing a possiblemechanism for in vivo turnover of core histones H2A and H2B that isindependent of replication but linked to transcription.
13. Kimura H, Cook PR: Kinetics of core histones in living humancells: little exchange of H3 and H4 and some rapid exchange ofH2B. J Cell Biol 2001, 153:1341-1353.
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Ahmad K, Henikoff S: Histone H3 variants specify modesof chromatin assembly. Proc Natl Acad Sci USA 2002,99(Suppl 4):16477-16484.
The authors demonstrate replication-independent deposition for the two
Drosophila histone H3 variants H3.3 and Cid using a Drosophila cellline system. H3.3 marks active chromatin and Cid marks centromericregions. Both are incorporated into chromatin throughout the cell cycle,implying mechanisms for core-histone displacement and/or turnover.
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Ahmad K, Henikoff S: The histone variant H3.3 marks activechromatin by replication-independent nucleosome assembly.Mol Cell 2002, 9:1191-1200.
Replication-independent incorporation of H3.3 into the active ribosomalgene array was demonstrated. Under favourable growth conditions,additional ribosomal arrays became active and these were found toincorporate H3.3 and to show reductions of methylated histone H3.Replication-independent incorporation was shown to be conferred uponnormal histone H3 by amino acid changes towards H3.3.
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Catez F, Brown DT, Misteli T, Bustin M: Competition betweenhistone H1 and HMGN proteins for chromatin binding sites.EMBO Rep 2002, 3:760-766.
Fluorescence recovery after photobleaching analysis indicates competi-tion between HMGN (high-mobility group protein N) and histone H1 for
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Dou Y, Bowen J, Liu Y, Gorovsky MA: Phosphorylation and anATP-dependent process increase the dynamic exchange of H1in chromatin. J Cell Biol 2002, 158:1161-1170.
Linker-histone phosphorylation leads to a more rapid fluorescencerecovery after photobleaching, suggesting decreased binding of thephosphorylated protein. Interestingly, ATP depletion in vivo results indecreased linker-histone mobility.
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Christensen MO, Larsen MK, Barthelmes HU, Hock R, AndersenCL, Kjeldsen E, Knudsen BR, Westergaard O, Boege F, Mielke C:Dynamics of human DNA topoisomerases IIalpha and IIbeta inliving cells. J Cell Biol 2002, 157:31-44.
Topoisomerase II has been proposed to be a major component of thechromosome scaffold and also the nuclear matrix. However, in vivofluorescence recovery after photobleaching experiments indicate rapidturnover of topoisomerase II on a timescale of seconds in both nuclei andmitotic chromosomes. If a more stably bound topoisomerase II fraction ispresent as part of a stable structural scaffold, it must correspond to asmall fraction of the total enzyme.
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Tavormina PA, Come MG, Hudson JR, Mo YY, Beck WT,Gorbsky GJ: Rapid exchange of mammalian topoisomerase IIalpha at kinetochores and chromosome arms in mitosis.J Cell Biol 2002, 158:23-29.
This is an independent fluorescence recovery after photobleaching studyof topoisomerase IIin vivo. Theauthors come to a similar conclusion as inChristensen et al. [18], that there is rapid turnover in nuclei and mitoticchromosomes.
20.
Featherstone M: Coactivators in transcription initiation: here are your orders. Curr Opin Genet Dev 2002, 12:149-155.
This is a nice reviewof ordered recruitment of differentcomponents of thetranscriptional machinery after gene activation of specific loci. Data fromseveral sources argue against one-step recruitment of entire holo-enyzmes, as defined by biochemical isolation.
21. Krebs JE, Fry CJ, Samuels ML, Peterson CL: Global role forchromatin remodeling enzymes in mitotic gene expression.Cell 2000, 102:587-598.
22.
Memedula S, Belmont AS: Sequential recruitment of HAT andSWI/SNF components to condensed chromatin by VP16.Curr Biol 2003, 13:241-246.
Using a microscopy-based immunofluorescence approach, recruitment
of various SWISNF and histone acetyltransferase (HAT) components aremonitored, together with histone acetylation, after targeting of VP16acidic activator to a condensed chromosome region. Different subunitsof SWISNF and HAT complexes accumulate at the chromosome sitewith very different kinetics, arguing against a simple one-step recruitmentof large chromatin-modifying complexes to condensed chromatin.
23.
Essers J, Houtsmuller AB, van Veelen L, Paulusma C, Nigg AL,Pastink A, Vermeulen W, Hoeijmakers JH, Kanaar R: Nucleardynamics of RAD52 group homologous recombinationproteins in response to DNA damage. EMBO J 2002,21:2030-2037.
Using fluorescence recovery after photobleaching analysis, this studydemonstrates stable association of Rad51 with radiation-induced foci,but rapid turnover of Rad52 and Rad54. In cells not exposed to radiation,different turnover rates for these proteins argues against stable pre-existing holoenzymes.
24. Hager GL, Elbi C, Becker M: Protein dynamics in the nuclearcompartment. Curr Opin Genet Dev 2002, 12:137-141.
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Becker M, Baumann C, John S, Walker DA, Vigneron M, McNallyJG, Hager GL: Dynamic behavior of transcription factors on anatural promoter in living cells. EMBO Rep 2002, 3:1188-1194.
This is a continuation of groundbreaking work [25], providing additionalsupport for the hit-and-run mechanism underlying transcription factorbinding and action in vivo. Using the same chromosome array system asthat previously described by McNally et al. [25], the Hager group demon-strated rapid turnover of the co-activator GRIP-1 at similar rates toglucocorticoid receptor. By contrast, the RPB1 RNA polymerase subunitrecovers over a much longer timescale (13 min), consistent with theprocessive property of elongating polymerase. A faster recovering com-ponent suggests the possibility of more rapidturnover of promoter-boundRNA polymerase II. Accumulation of RNA polymerase II peaks at2030 min after ligand addition, dropping substantially afterwards.
27. Stenoien DL, Nye AC, Mancini MG, Patel K, Dutertre M, OMalleyBW, Smith CL, Belmont AS, Mancini MA: Ligand-mediatedassembly and real-time cellular dynamics of estrogen receptoralpha-coactivator complexes in living cells. Mol Cell Biol 2001,21:4404-4412.
28. Stenoien DL, Patel K, Mancini MG, Dutertre M, Smith CL, OMalleyBW, Mancini MA: FRAP reveals that mobility of oestrogenreceptor-alpha is ligand- and proteasome-dependent.Nat Cell Biol 2001, 3:15-23.
29.
Chen D, Hinkley CS, Henry RW, Huang S: TBP dynamics in livinghuman cells: constitutive association of TBP with mitoticchromosomes. Mol Biol Cell 2002, 13:276-284.
The authors describe fluorescence recovery after photobleaching (FRAP)analysis of TATA-binding protein (TBP) in mitotic and interphase cells. Norecovery of photobleaching is seen in mitotic chromosomes, demonstrat-ing stable association of TBP. Within interphase nuclei, TBP shows slowrecovery over a timescale of minutes (i.e. 120). This recovery wasindependent of transcriptional inhibition, suggesting that TBP is stablybound over multiple initiation rounds.
30.
Kimura H, Sugaya K, Cook PR: The transcription cycle of RNApolymerase II in living cells. J Cell Biol 2002, 159:777-782.
Fluorescence recovery after photobleaching analysis of the Large RNApolymeraseII (pol II) reveals a transientlyimmobile fraction correlated withtranscriptional elongation. Little pol II is immobile in the presence oftranscriptional inhibition, arguing against stable binding of a signi ficantfraction of enzyme in transcription factories.
31.
Dundr M, Hoffmann-Rohrer U, Hu Q, Grummt I, Rothblum LI,Phair RD, Misteli T: A kinetic framework for a mammalian RNApolymerase in vivo. Science 2002, 298:1623-1626.
Fluorescence recovery after photobleaching analysis of multiple compo-nents of the RNA polymerase I (pol I) transcriptional apparatus aredescribed. Recovery over tens of seconds was observed for severalpre-initiation factors. Four pol I subunits showed biphasic recovery
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curves, with a slow recovery phase associated with transcription elonga-tion. Differences in theearlyrapid recovery rates forthe four polI subunitswas used to argue against recruitment of a pre-assembled holoenzyme.
32.
Fletcher TM, Xiao N, Mautino G, Baumann CT, Wolford R,Warren BS, Hager GL: ATP-dependent mobilization of theglucocorticoid receptor during chromatin remodeling.Mol Cell Biol 2002, 22:3255-3263.
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