Gene transcription is a dynamic process involving a continuous
assembly and disassembly of the transcription complexes on gene
promoters (Hager et al., 2009; Sikorski and Buratowski, 2009).
Live-cell imaging combined with fluorescence recovery after
photobleaching (FRAP) has revealed a high mobility and transient
chromatin interactions for a variety of transcription factors involved
in RNA polymerase II (pol II)-mediated transcription (Farla et al.,
2004; Farla et al., 2005; Hager et al., 2009; Hoogstraten et al., 2002;
Karpova et al., 2008; McNally et al., 2000; Mueller et al., 2008;
Muller et al., 2001; Sprouse et al., 2008; Stenoien et al., 2001; van
Royen et al., 2009). The Drosophilaheat shock factor is an exception
to this and shows a slow recovery under heat shock conditions (Yao
et al., 2006).
Tagging the largest catalytic subunit of pol II with green
fluorescent protein (GFP) revealed that ~75% of this enzyme
moved rapidly in living cells, whereas 25% was transiently
immobile and active in transcription, as this fraction disappeared
after transcription inhibition (Kimura et al., 2002). Kinetic
measurements of the different steps in the pol II transcription cycle
indicate that formation of the pre-initiation complex (PIC) is very
dynamic and rather inefficient, with only a small fraction of the
initiated pol II proceeding to elongation (Darzacq et al., 2007).
This is in agreement with recent findings of promoter-proximal
pausing of pol II as a rather general step in the transition to
productive elongation (Core and Lis, 2008). Visualization of native
mRNA transcripts indicates that elongating pol II proceeds with a
rate ranging from 1.5-2.0 kb/minute (Boireau et al., 2007; Yao et
al., 2007) to 4.3 kb/minute (Darzacq et al., 2007).
Detailed analysis of the pol I transcription complex shows that
pol I is dynamically imported into the nucleolus (Dundr et al.,
2002) and when higher levels of transcription are required, assembly
of pol I complexes is more efficient and the rate of entry into
elongation is elevated (Gorski et al., 2008).
The TATA-binding protein (TBP) plays a central role in
eukaryotic transcription by all three RNA polymerases and resides
in at least four transcription complexes: TFIID, B-TFIID, SL1 and
TFIIIB (for reviews, see Burley and Roeder, 1996; Muller et al.,
2007; Pereira et al., 2003; Thomas and Chiang, 2006). In TFIID,
TBP is associated with 13-14 evolutionary conserved TBP-
associated factors (TAFs) (Tora, 1992). B-TFIID represents the
most abundant TBP-containing complex and consists of TBP and
a single TAF, BTAF1, which is a conserved member of the SNF2
ATPases family (Pereira et al., 2003). In cellular lysates TBP is
stably associated in the TFIID, TFIIIB and SL1 complexes, but
TBP and BTAF1 are exchanging rapidly (Mousson et al., 2008).
DNA-bound TBP provides the platform for association of basal
transcription factors and pol II to direct formation of a functional
PIC. TBP-DNA complexes can also be recognized efficiently by
the negative co-factor NC2, which can inhibit in vitro transcription
by blocking association of the basal factors TFIIA and TFIIB
(Goppelt et al., 1996; Inostroza et al., 1992; Meisterernst et al.,
1991). Interestingly, both biochemical and genomic experiments
point to a functional interplay between BTAF1 (or its yeast
orthologue Mot1p) and NC2 (Geisberg et al., 2002; Hsu et al.,
2008; Klejman et al., 2004; van Werven et al., 2008).
TBP dynamics have been studied by FRAP both in living human
cells (HeLa) (Chen et al., 2002), and in yeast (Sprouse et al.,
2008), with strikingly different results. In the HeLa cell system, a
slow recovery (>20 minutes) was measured, indicating a large
immobile fraction of TBP, which most probably involves chromatin
binding (Chen et al., 2002). This is in contrast to yeast TBP, which
Chromatin interaction of TATA-binding protein is
dynamically regulated in human cells
Petra de Graaf1, Florence Mousson1, Bart Geverts2, Elisabeth Scheer3, Laszlo Tora3, Adriaan B. Houtsmuller2
and H. Th. Marc Timmers1,*
1Department of Physiological Chemistry and Netherlands Proteomic Center, University Medical Centre Utrecht, Universiteitsweg 100,
3584 CG Utrecht, Netherlands
2Department of Pathology, Josephine Nefkens Institute, Erasmus MC, 3000 CA Rotterdam, Netherlands
3Department of Functional Genomics, Institut de Génétique et de Biologie Moléculaire et Cellulaire (IGBMC), UMR 7104 CNRS, UdS,
INSERM U964, BP 10142, F-67404 ILLKIRCH Cedex, CU de Strasbourg, France
*Author for correspondence (firstname.lastname@example.org)
Accepted 27 April 2010
Journal of Cell Science 123, 2663-2671
© 2010. Published by The Company of Biologists Ltd
Gene transcription in mammalian cells is a dynamic process involving regulated assembly of transcription complexes on chromatin in
which the TATA-binding protein (TBP) plays a central role. Here, we investigate the dynamic behaviour of TBP by a combination of
fluorescence recovery after photobleaching (FRAP) and biochemical assays using human cell lines of different origin. The majority
of nucleoplasmic TBP and other TFIID subunits associate with chromatin in a highly dynamic manner. TBP dynamics are regulated
by the joint action of the SNF2-related BTAF1 protein and the NC2 complex. Strikingly, both BTAF1 and NC2 predominantly affect
TBP dissociation rates, leaving the association rate unchanged. Chromatin immunoprecipitation shows that BTAF1 negatively regulates
TBP and NC2 binding to active promoters. Our results support a model for a BTAF1-mediated release of TBP-NC2 complexes from
Key words: Basal transcription factors, Gene expression, FRAP, TFIID, TBP
Journal of Cell Science
shows a remarkable fast FRAP (less than 15 seconds), but not as
rapid as TFIIB or TAF1 (Sprouse et al., 2008). Besides different
model systems, another difference between the two studies is the
method of expressing GFP-tagged TBP. In the human system, a
transient overexpression system was used, whereas in yeast the
genes for TBP, TFIIB and TAF1 were replaced by fluorescently
Here, we investigate the dynamics of TBP in living human cells
by using a retroviral transduction to express stably GFP-TBP to
near-endogenous levels. Analysis of these cell lines by fluorescence
recovery after photobleaching reveals that the major fraction of
TBP fluorescence recovers rapidly, which is indicative of a dynamic
interaction with chromatin. In a combination of siRNA-mediated
knockdown and transient overexpression experiments, we find that
the NC2 complex stabilizes chromatin association of TBP, whereas
BTAF1 promotes its release. Chromatin immunoprecipitation
experiments confirm that BTAF1 controls the promoter-bound
fraction of TBP and NC2. Our findings support a model for a
dynamic control of basal transcription factor complexes to allow
rapid transcriptional responses of cells to internal and external cues.
GFP-TBP is partly bound to chromatin in living cells
To study TBP dynamics in vivo, we generated cell lines expressing
GFP-tagged human TBP by retroviral transduction. Confocal
microscope images of monoclonal cell lines derived from human
U2OS osteosarcoma or HeLa cervix carcinoma cells showed that,
as expected, GFP-TBP is present in the nucleus and its pattern
overlaps with DNA staining (Fig. 1A). To ensure that the GFP
moiety does not affect TBP function, extracts of the generated cell
lines were analyzed for expression and incorporation of GFP-TBP
into TBP-containing complexes. Gel filtration and
2664 Journal of Cell Science 123 (15)
immunoprecipitation analyses indicated that GFP-TBP is efficiently
incorporated in both B-TFIID and TFIID complexes
(supplementary material Fig. S1A-C). Furthermore, the analyses
show that GFP-TBP is expressed at near endogenous levels in
HeLa- and U2OS-derived cell lines. We used the GFP-TBP cell
lines to analyze the behaviour of TBP by fluorescent recovery after
photobleaching (FRAP). As controls, we compared FRAP curves
of GFP-TBP with those of GFP fused to histone H2B (H2B-GFP)
or to the SV40 nuclear localization signal (NLS-GFP). FRAP
experiments were performed by bleaching a narrow strip (~0.75
m) covering the nucleoplasm across the width of the nucleus and
recording the recovery of fluorescence in this strip every 21
mseconds (van Royen et al., 2009; Xouri et al., 2007). As expected,
the recovery of NLS-GFP fluorescence was very rapid and no
immobile fraction could be detected, whereas H2B-GFP was largely
immobile (80-90%) in the strip-FRAP experiments. GFP-TBP
fluorescence in both HeLa- and U2OS-derived cells displayed a
rapid initial recovery followed by a slower subsequent phase (Fig.
1B). A sequence of frames depicting the whole nucleus after
bleaching shows similar results: no recovery for H2B-GFP, partial
recovery for GFP-TBP and full recovery for NLS-GFP
(supplementary material Fig. S2). To analyze experimental FRAP
curves, a large database was constructed, which contained FRAP
curves generated by Monte Carlo computer simulations (see
Materials and Methods for a detailed description). Each curve in
this database represents a scenario with a specific diffusion
coefficient and a specific konand koffrepresenting the rate at which
proteins bind to and release from immobile DNA (e.g. Xouri et al.,
2007). Each experimental FRAP curve was compared with the
computer-generated curves of the database using standard ordinary
least square fitting. From this, the ten best-fitting curves were
taken to calculate the average diffusion coefficients and rate
Fig. 1. Dynamics of TBP and TFIID-TAFs in living cells.
(A)Confocal microscopic images of GFP-TBP-expressing
HeLa (left) or U2OS (right) cell lines indicate nuclear
localization of GFP-TBP. GFP signals are in green (upper
panels) and DAPI signals for DNA-staining in blue (lower
panels). Scale bars, 5m. (B)HeLa tk–and U2OS cells
expressing GFP-TBP (respectively light and dark blue), NLS-
GFP (respectively pink and purple) or H2B-GFP (respectively
light and dark green) were analyzed for FRAP by CLSM. Prior
to photo-bleaching, a strip spanning the nucleus was scanned
at low laser power. After this, the ROI was scanned every 21
mseconds until 38 seconds after bleaching. Raw data were
normalized for fluorescence intensity before bleaching and an
average of 10 nuclei was plotted. (C)ts013 cells expressing
GFP-TAF1 and U2OS cells expressing GFP-TAF5 were
analyzed for FRAP as described in B. U2OS cells expressing
NLS-GFP and GFP-TBP were included as a reference.
Incorporation of the tagged TAFs into TFIID was shown by
immunoprecipitation experiments (supplementary material Fig.
S1D,E). (D)U2OS GFP-TBP cells were transfected with RNAi
oligos as indicated and analyzed for FRAP as described in B.
Transfection efficiency of the RNAi oligos was analysed by
co-transfecting siGLO red (supplementary material Fig. S3).
(E)Knockdown efficiency was analyzed in total lysates of the
transfected cells as described in D by immunoblotting with
TAF1, BTAF1, TAF5, GAPDH and GFP antibodies.
Knockdown of TAF3 was efficient as measured by RT-PCR
(data not shown).
Journal of Cell Science
constants. The values for diffusion coefficient and rate constants
determined in this way are summarized for all experiments in
supplementary material Table S1. Note that on- and off-rate
constants correspond directly to the (transiently-) immobile fraction
(see Materials and Methods). The HeLa- and U2OS-derived GFP-
TBP cell lines yielded similar results. In both cell lines, ~70% of
the GFP-TBP recovers rapidly and the characteristic time spent in
the subsequent transiently immobile state was 2-3 minutes
(supplementary material Table S1).
TFIID components interact very transiently with chromatin
These observations raise the issue of the dynamics of the TFIID
complex, consisting of TBP and 13-14 TAFs (Burley and Roeder,
1996; Muller et al., 2007; Thomas and Chiang, 2006). To investigate
this, we tagged different TFIID-TAFs with GFP. First, GFP-tagged
TAF1 was stably expressed in the hamster ts013 cell line, which
expresses a temperature-sensitive mutant of TAF1 (Sekiguchi et
al., 1991). The majority of GFP-TAF1 recovered rapidly after
bleaching with a small transiently immobile fraction (~18%) (Fig.
1C; supplementary material Table S1). It is important to note that
FRAP of GFP-TAF1 is slower than the NLS-GFP, indicating a
short residence time to chromatin. Second, a GFP-tagged version
of TAF5, one of the structural TAFs (Cler et al., 2009; Leurent et
al., 2004), was stably expressed in U2OS cells. TAF5 exhibited a
similar recovery to TAF1 in the FRAP analysis (Fig. 1C;
supplementary material Table S1). Thus, the TAFs of TFIID are
highly mobile in vivo, with a small proportion displaying short
interaction times with chromatin (1-2 sec, supplementary material
Effect of TAFs and BTAF1 expression on TBP dynamics
Given the association of TBP with the TFIID complex, we
investigated whether TAFs determine the dynamic interaction of
TBP with chromatin. We selected three different TFIID TAFs for
siRNA-mediated knockdown: TAF1, which interacts with TBP
(Takada et al., 1992); TAF3, which anchors TFIID to methylated
histones (Vermeulen et al., 2007); and the structural TAF5 (Leurent
et al., 2004). To ensure that the single cells analyzed by FRAP
were transfected by siRNAs, a transfection indicator was included
as described in the Materials and Methods (supplementary material
Fig. S3). The majority of cells were transfected, as also indicated
by a reduction in protein levels (Fig. 1E). Knockdown of TAF1,
TAF3 or TAF5 expression did not significantly change the dynamics
of TBP (Fig. 1D). Similarly, a reduced expression of the TBP-
associating factor BRF1 from the TFIIIB complex involved in pol
III transcription did not influence TBP dynamics (supplementary
material Fig. S4). By contrast, siRNA knockdown of BTAF1, the
single TAF in the B-TFIID complex (Timmers et al., 1992), reduced
the mobile fraction of TBP from 70% to less than 50% (Fig. 1D;
supplementary material Table S1). Similar results were obtained
upon inducible shRNA knockdown of BTAF1 (supplementary
material Fig. S5). From these experimental FRAP curves, both the
dissociation rate constant koffand association rate constant konwere
determined (see Materials and Methods for a detailed description).
Interestingly, koff decreased when BTAF1 was downregulated,
whereas kondid not change significantly (supplementary material
Table S1), suggesting that BTAF1 regulates the release of TBP
from chromatin and not its association. To further investigate the
role of BTAF1 in TBP-chromatin interaction, we determined the
mobility of TBP upon BTAF1 overexpression. To this end, GFP-
TBP was transfected with or without HA-tagged BTAF1 in 293T
Regulation of TBP dynamics by BTAF1/NC2
cells and the resulting cells were subjected to FRAP analysis.
Overexpression of BTAF1 increased the mobile fraction of GFP-
TBP (Fig. 2A,B).
Having established that BTAF1 function is important for TBP
dynamics, we investigated involvement of BTAF1 in global
chromatin association of TBP by biochemical fractionation of cell
extracts. Therefore, nuclear proteins were selectively extracted
from inducible BTAF1 knockdown cells and analyzed by
immunoblotting. The chromatin-bound fraction of TBP was
specifically increased upon reduction of BTAF1 expression (Fig.
3). As expected, histone H4 could be detected only in the chromatin-
enriched fractions. Little effect was observed on the distribution of
the TAF1 and TAF5 subunits of TFIID, indicating that BTAF1
regulates chromatin binding of TBP without affecting the TAF
subunits of TFIID. This hypothesis is strengthened by the
observation that the dynamic behaviour of TAF5 is not affected by
BTAF1 knockdown (data not shown).
Fig. 2. Chromatin association of GFP-TBP is differentially regulated by
NC2 complex and BTAF1. (A)HEK293T cells were transfected with
pEGFP-TBP (25 ng) and subsequently pMT2-SM HA-BTAF1 wt (2g),
pCMV HA-NC2 (250 ng), pCMV FLAG-NC2 (250 ng) as indicated. Mock
DNA (pMT2-SM) was added to have equal amounts of plasmid per
transfection. Cells were analyzed for FRAP 20 hours after transfection.
Quantification of FRAP signals was performed as described in Fig. 1B.
(B)Expression levels of the exogenous protein as described in A was analyzed
in total lysates by immunoblotting with HA, FLAG and GFP antibodies.
(C)U2OS GFP-TBP-expressing cells were transfected with RNAi oligos as
indicated and analyzed for FRAP as described in Fig. 1B. (D)Knockdown
efficiency was analyzed in total lysates of the transfected cells as described in
C by immunoblotting with BTAF1, NC2, GAPDH, GFP and TAF5
antibodies. (E)U2OS GFP-TBP expressing cells were transfected with RNAi
oligos as indicated and analyzed for FRAP as described in Fig. 1B.
(F)Knockdown efficiency was analyzed in total lysates of the transfected cells
as described in E by immunoblotting with BTAF1, NC2, GAPDH and GFP
Journal of Cell Science
NC2 regulates TBP-chromatin interaction
The negative co-factor NC2 interacts with DNA-bound TBP with
a high affinity (Gilfillan et al., 2005). Several observations indicate
that BTAF1 and its yeast ortholog Mot1p overlap in function with
NC2 (Geisberg et al., 2002; Klejman et al., 2004; van Werven et
al., 2008). Thus, NC2 may also play a role in regulation of TBP
interaction to chromatin in human cells. Interestingly, knockdown
of NC2 expression increases the fraction of mobile TBP molecules
in the GFP-TBP U2OS cells (Fig. 2C). Remarkably, knockdown of
the NC2 subunit also reduced the expression of the NC2 subunit
in total lysates (Fig. 2D). This strengthens the notion that NC2
and NC2 act as a complex to regulate TBP. As shown before,
BTAF1 knockdown reduced the mobile fraction (supplementary
material Table S1). The opposing effects of NC2 and BTAF1 on
TBP binding to chromatin are corroborated by the combined
knockdown of NC2 and BTAF1, which reverted the dynamics of
TBP to that of untreated cells (Fig. 2E,F). These results predict that
overexpression of the NC2 complex may increase chromatin
association of TBP in cells. Indeed, the fraction of mobile TBP is
reduced only when both subunits of NC2 are overexpressed (Fig.
2666 Journal of Cell Science 123 (15)
2A), which again indicates that they act as a complex. Furthermore,
we could show that overexpression of TBP increased the chromatin
binding of both GFP-NC2 and GFP-NC2 (Fig. 5), which
strengthens our hypothesis that NC2 controls the binding of TBP
This set of experiments allowed us to also determine the effect
of NC2 on the kon and koff of the TBP-chromatin interaction.
Strikingly, only the koffchanges after reduction or overexpression
of NC2 complex (Fig. 4A; supplementary material Table S1),
whereas the kondoes not change significantly (Fig. 4B). Comparison
with the effects of BTAF1 (Fig. 4A) reveals that BTAF1 and NC2
have opposite effects on the dissociation rate of TBP from
chromatin. We used our computer modelling to illustrate the effect
of altering konor koffon FRAP curves. The FRAP curves depicted
in Figs 1 and 2 clearly fit better to curves representing scenarios
where only koffis varied (Fig. 4C) compared with scenarios where
only konchanges (Fig. 4D).
BTAF1 regulates TBP and NC2 levels at promoters
FRAP analysis indicated that BTAF1 and the NC2 complex act in
concert to regulate the dynamic behaviour of TBP in cells. To
explore the influence of BTAF1 on promoter binding of TBP and
NC2, we performed chromatin immunoprecipitation (ChIP)
experiments in cells with a reduced BTAF1 expression. We selected
a set of highly expressed genes for this analysis. As expected, TBP
binding peaks at the transcription start sites (TSS) of the genes
encoding ribosomal proteins RPL31, RPL34 and RPS10, and the
shared promoter of the divergent histone H2BJ and H2AG genes
(Fig. 6B; supplementary material Fig. S6). BTAF1 knockdown
results in increased TBP binding at the promoter of all three loci.
This is further supported by ChIP sequencing data for TBP,
comparing GAPDH with BTAF1 knockdown using siRNA
(Johannes et al., 2010). Of the genomic regions increased in TBP
binding by BTAF1 knockdown 70% correspond to promoter
regions. This indicates that the effect of BTAF1 on TBP binding is
very general to pol II promoters (Johannes et al., 2010).
Interestingly, the ChIP experiments in Fig. 6C show a strong
Fig. 3. BTAF1 knockdown increases chromatin association of TBP.
Nuclear fractions were prepared from HeLa TRsshBTAF1 cells treated with or
without doxycycline for 8 days as described in the Materials and Methods.
Soluble fraction, washes and soluble chromatin fraction were analyzed
immunoblotting with TBP, BTAF1, TAF1, TAF5 and histone H4 antibodies.
Fig. 4. BTAF1 and NC2 affect dissociation rates of TBP
from chromatin. (A,B)Graphs showing the dissociation and
association rates of TBP (konand koff), as determined by
computer modelling-based analysis, after either
downregulation or overexpression of BTAF1 (green squares),
NC2 or NC2 (yellow squares), or NC2 and NC (red
squares) compared with untreated cells (blue squares). In
addition, the variation of individual control experiments
performed in parallel with downregulation and overexpression
experiments is shown (blue circles). The experiments in which
either NC2 or NC2 expression level was manipulated were
averaged for clarity as they showed similar reproducible
results. Mean and standard deviations (as shown with the error
bars) were calculated from 2-10 biological independent
experiments. (C)FRAP curves representing a scenario where
konwas constant (0.004 s–1) and koffwas varied (0.016, 0.008,
0.004 and 0.002 s–1), resulting in both increase of immobile
fraction and residence time. (D)FRAP curves representing a
scenario where koffwas constant (0.008 s–1) and konwas varied
(0.002, 0.004 and 0.016 s–1). As residence time depends on koff
only, in this scenario only the immobile fraction varies with
Journal of Cell Science
increase of promoter binding by NC2 after reduction of BTAF1
expression (see also supplementary material Fig. S6C). These data
support a model in which BTAF1 counteracts the effect of NC2,
as observed in the FRAP experiments. This is agreement with
biochemical experiments in yeast, which suggested that Mot1p-
NC2-TBP-DNA represents a transcriptionally silent complex (van
Werven et al., 2008). To analyze whether the increased association
of TBP and NC2 has transcriptional consequences pol II binding
was measured by ChIP. Binding of pol II is not altered after BTAF1
knockdown (Fig. 6D). To confirm this, mRNA expression of these
genes was analyzed by reverse transcription qPCR. The increased
binding of TBP did not enhance mRNA levels; RPL31, RPS10,
H2A and H2B mRNAs are not influenced by BTAF1 knockdown,
whereas RPL34 expression is reduced to ~60% (Fig. 6E;
supplementary material Fig. S6D). The limited effect of BTAF1 on
transcription is consistent with mRNA profiling of BTAF1
knockdown cells (data not shown). Taken together, these analyses
indicate that an increased association of TBP and NC2 to promoters
does not lead to increased mRNA transcript levels.
In this study, we combined live-cell imaging and biochemical
approaches to investigate the molecular mechanisms involved in
the dynamic behaviour of human basal transcription factor TBP.
FRAP experiments show that a significant fraction of TBP is
transiently immobilized on the chromatin, but the majority of TBP
is mobile. Part of this mobile pool may represent the TFIID
complex. The TAF1 and TAF5 subunits of TFIID interact only
shortly with chromatin (1-2 seconds) and most of these proteins
are represented in the mobile fraction. The mobile fraction of TBP
is regulated by BTAF1 and by NC2 expression. Either
downregulation of BTAF1 or overexpression of NC2 decreases the
mobile fraction and increases the part of TBP, which is transiently
immobile on chromatin. Conversely, knockdown of NC2 or
overexpression of BTAF1 increases the mobile fraction.
Interestingly, computer-based analysis of the FRAP data suggests
that the observed changes in TBP dynamics by BTAF1 and NC2
depend only on changes in dissociation of TBP from the chromatin,
leaving association rates unchanged.
The results of our study are largely similar to recent findings in
yeast, which indicated that TBP is a rather mobile protein (Sprouse
et al., 2008). By contrast, in a previous study of human cells, Chen
et al. reported a large immobile fraction of GFP-TBP upon transient
overexpression, which displayed residence times of ~20 minutes
(Chen et al., 2002). ATP depletion increased the chromatin binding
of transfected GFP-TBP even further, which would be consistent
with involvement of the ATPase function of BTAF1. Besides
technical differences in the FRAP setup, we propose that the
differences between the studies relate to the overexpression versus
stable expression of GFP-TBP as overexpression of TBP could
titrate out BTAF1 and/or NC2. Indeed, when we transiently express
GFP-TBP in U2OS cells, we observe an increase in chromatin
binding when transfecting higher amounts of GFP-TBP
(supplementary material Fig. S7). Additional support for this is
that, in contrast to Chen et al. (Chen et al., 2002), we do not
observe GFP-TBP localization to the mitotic chromosomes, which
is consistent with findings with endogenous proteins [Kieffer-
Kwon et al. (Kieffer-Kwon et al., 2004) and data not shown].
Although the TBP, BTAF1/Mot1p and NC2 proteins are highly
conserved throughout evolution, the molecular mechanisms by
which they perform their function might be slightly different
Regulation of TBP dynamics by BTAF1/NC2
between organisms. In yeast cells, TBP-YFP fluorescence fully
recovered within 15 seconds after bleaching with no detectable
immobile fraction (Sprouse et al., 2008). Importantly, TBP-YFP
was expressed from the endogenous locus. Interestingly, the
dynamics of yeast TBP are reduced in a mot1 mutant (Sprouse et
al., 2008). No effect on TBP mobility was observed in a yeast
NC2 (bur6) mutant strain, whereas reduction of TAF1 (taf1)
slightly increased TBP mobility (Sprouse et al., 2008). By contrast,
we observed in human cells that TBP dynamics are differentially
regulated by BTAF1 and NC2, whereas reduced expression of
TAF1 or other TFIID-TAFs has no effect on the dynamic association
of TBP with the chromatin.
The heterodimeric NC2 repressor complex has been shown in
vitro to increase binding of TBP to promoter and non-promoter
DNA, and to stabilize TBP-DNA complexes in general (Gilfillan
et al., 2005; Goppelt et al., 1996). Experiments in vitro indicated
that human BTAF1 and NC2 can compete for binding to DNA-
bound TBP (Klejman et al., 2004). In addition, genome-wide
location analyses in yeast indicate that Mot1p and NC2 bind to an
overlapping set of promoters (van Werven et al., 2008). Biochemical
analysis of chromatin complexes resulted in isolation of a stable
Mot1p-NC2-TBP-TATA complex (van Werven et al., 2008), which
could be disrupted by the addition of ATP. By contrast, analyses of
human chromatin failed to identify a stable BTAF1-NC2-TBP-
DNA complex [Gilfillan et al. (Gilfillan et al., 2005) and data not
Our observations that modulating NC2 expression selectively
affects the dissociation constant of TBP (Fig. 4) fits with the
Fig. 5. Chromatin association of GFP-NC2 is regulated by TBP.
(A)HEK293T cells were transfected with pEGFP-NC2 (100 ng) and pCMV-
flag-NC2 (200 ng) with or without pMT2-SM HA-TBP (1.7g). Mock DNA
(pMT2-SM) was added to ensure equal amounts of plasmid were present per
transfection. Cells were analyzed for FRAP 20 hours after transfection.
(B)HEK293T cells were transfected with pEGFP-NC2 (100 ng) and pCMV-
HA-NC2 (200 ng) with or without pMT2-SM HA-TBP (1.7g). Mock DNA
(pMT2-SM) was added to have equal amounts of plasmid per transfection.
Cells were analyzed for FRAP 20 hours after transfection. (C)Protein
expression was analyzed in total lysates of the transfected cells as described in
A,B by immunoblotting with TBP, NC2, FLAG and GFP antibodies.
Journal of Cell Science
observation that only DNA-bound TBP is a substrate for NC2. Our
experiments in yeast (van Werven et al., 2008) combined with
other studies (Meisterernst et al., 1991; Inostroza et al., 1992)
indicate that this complex is transcriptionally inactive. Our data
suggest that the immobilized TBP is locked in an NC2 complex. It
has been proposed that TBP-NC2 slides along the DNA fibre
(Schluesche et al., 2007). This sliding of the transcriptionally
inactive NC2-TBP complex (Schluesche et al., 2007) could clear
the core promoter for formation of a functional preinitiation
complex. This is supported by the ChIP analysis (Fig. 6), which
showed an increased NC2 occupancy on promoters upon BTAF1
knockdown. Although NC2 binding is increased around the
transcription start site, this does not affect transcription as shown
by the pol II ChIP and RT-PCR analysis. Consistent with this is the
observation that ChIP experiments with mot1 mutant cells show an
increased TBP and NC2 association at a specific set of promoters
(Dasgupta et al., 2005; Geisberg et al., 2002), as observed in the
ChIP experiments under BTAF1 knockdown conditions (Fig. 6).
These observations suggest co-occurrence of NC2-TBP repressive
complexes and TBP in functional transcription complex on the
same chromatin fragment. We propose that NC2-TBP slides away
from the core promoter and does not obstruct association of active
In general, several studies now indicate that basal transcription
factor complexes are highly dynamic in vivo (Darzacq et al., 2007;
Hager et al., 2009; Hoogstraten et al., 2002; Sprouse et al., 2008;
van Werven et al., 2009). In addition, it has been shown that many
2668Journal of Cell Science 123 (15)
gene-specific transcription factors have relative short DNA
residence times (Bosisio et al., 2006; Farla et al., 2004; McNally
et al., 2000; Mueller et al., 2008; Muller et al., 2001). We now
report that TFIID-TAFs are also mobile and display short residence
times on chromosomal DNA of 1-2 seconds (Fig. 1C). Although a
fraction of TBP is bound to chromatin with a relative long residence
time, most of TBP (~70%) is freely mobile. Together, our results
add to previously published observations (Darzacq et al., 2007)
and further suggest that the pol II PICs are very dynamic structures
and that TBP bound to pol II promoters is rapidly exchanging (van
Werven et al., 2009). This challenges models of pol II mRNA
factories consisting of tightly bound and rather immobile protein
assemblies (Sutherland and Bickmore, 2009), but rather support a
model where transcription sites are rapidly assembling and
disassembling entities (Lebedeva et al., 2005). Importantly, this
kinetic model allows for a rapid response to changes in mRNA
requirement of cells. Furthermore, the antagonistic regulation of
TBP-chromatin interactions by BTAF1 and NC2 fits to models of
transcriptional bursts (Chubb et al., 2006) in which promoters can
switch rapidly between transcriptionally active and inactive states.
We propose that this switching is intimately linked to NC2 and
BTAF1 regulation of TBP dynamics.
Materials and Methods
The monoclonal TBP antibody 1F8, the rabbit polyclonal antibody PF299 and the
mouse polyclonal antibodies C-BTAF1 (blanc-o and L.O.R.O.) against BTAF1 have
been described previously (Pereira et al., 2004). The monoclonal TBP antibodies
Fig. 6. BTAF1 regulates TBP and NC2 levels at
promoters. (A)Schematic representation of the genes rpl31,
rpl34 and the hist1h2bjh2ag locus. The numbers correspond to
the graphs represented in B-D. (B)Chromatin
immunoprecipitation (ChIP) from HeLa tk–cells transfected
with siRNA targeting BTAF1 or control siRNA followed by
quantitative PCR. Signals are plotted as percentage of input
for precipitation with TBP (SL30) antibody. The data shown
are representative of a triplicate. (C)ChIP followed from
HeLa tk–cells transfected with siRNA targeting BTAF1 or
control siRNA by quantitative PCR. Signals are plotted as
percentage of input for precipitation with NC2 antibody. The
data shown are representative of a triplicate. (D)ChIP
followed from HeLa tk–cells transfected with siRNA targeting
BTAF1 or control siRNA by quantitative PCR. Signals are
plotted as percentage of input for precipitation with pol II
(8WG16) antibody. The data shown are representative of a
duplicate. (E)cDNA was prepared from HeLa tk–cells
transfected with siRNA targeting BTAF1 or control siRNA.
mRNA levels of RPL31, RPL34, H2A, H2B, GAPDH and
BTAF1 were analyzed by quantitative PCR. Depicted is an
average of three biological independent experiments.
(F)Protein levels were analyzed in total lysates of the
transfected cells as described in B by immunoblotting with
BTAF1, TBP, NC2 and tubulin antibodies.
Journal of Cell Science
Regulation of TBP dynamics by BTAF1/NC2
SL30 (Ruppert et al., 1996), NC2 antibodies 4G7 (Gilfillan et al., 2005) and
polyclonal TAF5 antibody (Christova and Oelgeschlager, 2002) have been described.
The monoclonal GFP antibody (mix of clones 7.1 and 13.1) was from Roche Applied
Science and the monoclonal tubulin antibody (DM1A) was obtained from
Calbiochem. Histone H4 (ab31827) and TAF1 (6B3) antibodies were obtained from
Abcam and Upstate, respectively. GAPDH antibodies (6C5) were obtained from
Millipore. The monoclonal HA antibody was purified from the hybridoma cell line
The constructs pEGFP-TBP, pTER, pCAG-TRsand pcDNA4-TO luciferase (Chen
et al., 2002; van de Wetering et al., 2003) have been described previously. To
generate a retroviral expression vector for N-terminal or C-terminal fusion proteins,
the GFP ORF was amplified by PCR and inserted into the pBabe puro plasmid,
resulting in pBabe-puro GFP-N-term and pBabe-puro GFP C-term. The plasmids
were modified to act as Gateway destination plasmids by introducing a blunt-ended
cassette containing attR sites flanking the ccdB gene and the chloramphenicol
resistance gene from RfC (for the N-terminal fusion construct) or RfA (for the C-
terminal fusion construct) resulting in pBabe-puro GFP-Nterm Dest GW and pBabe-
puro GFP-Cterm Dest GW, respectively. The cloning junctions were verified by
Human TBP, TAF1, TAF5, NC2, NC2, BTAF1 and Histone H2B were amplified
using appropriate oligonuclotides flanked with attB1 or attB2 sequences. The
amplified sequences was cloned into pDONR201 (Invitrogen) using a standard BP
reaction to generate pENTR-N-TBP, pENTR-N-TAF1, pENTR-N-TAF5, pENTR-
N-NC2, pENTR-N-NC2, pENTR-N-BTAF1 and pENTR-C-H2B, respectively.
pBabe GFP-TBP, pBabe GFP-TAF1, pBabe GFP-TAF5, pBabe GFP-NC2 and
pBabe GFP-NC2 were generated in a LR reaction with pBabe puro GFP-Nterm
Dest GW and, respectively, pENTR-N-TBP, pENTR-N-TAF1, pENTR-N-TAF5,
pENTR-N-NC2 and pENTR-N-NC2. pBabe H2B-GFP was generated in a LR
reaction with pBabe puro GFP-Cterm Dest GW and pENTR-C-H2B. pBabe NLS-
GFP was generated by conventional cloning of an oligonucleotide encoding an
optimal translational initiation site followed by the SV40 NLS (PKKKRKV) in
pBabe-puro GFP-C-term. NC2 and NC2 were amplified using appropriate
oligonucleotides flanked with restriction sites and inserted into, respectively,
pcDNA3.1 HA and pcDNA3.1 Flag by conventional cloning to obtain pCMV HA
NC2 and pCMV Flag NC2. pMT2-SM HA was modified to act as a destination
plasmid by introducing the RfC cassette to yield pMT2-SM HA Dest GW. To yield
the mammalian expression constructs, pMT2-SM HA-BTAF1 and pMT2-SM HA
TBP pMT2-SM HA Dest GW was recombined in a standard LR reaction with either
pENTR-N-BTAF1 or pENTR-N-TBPwt.
pTER shBTAF1 constructs were generated by inserting an dsDNA-oligo encoding
a RNA short hairpin targeting knock-down of BTAF1 into the BglII/HindIII sites of
the pTER vector (van de Wetering et al., 2003). Sequences of used oligonucleotides
are available upon request.
Cell lines and transient transfection
HeLa tk–, U2OS and HEK293T cells were grown in Dulbecco’s modified Eagle’s
medium (DMEM) supplemented with 10% foetal calf serum, L-glutamine and
antibiotics at 37°C. Mutant hamster ts13 cells (Sekiguchi et al., 1991) were grown
in DMEM with high glucose (4.5 mg/ml) supplemented with 10% foetal calf serum,
L-glutamine and antibiotics at 34°C or 39.5°C. Transient transfection of plasmid
DNA was performed 6 hours after seeding using FuGene6 transfection reagent
according to the manufacturer’s instructions (Roche Applied Science). Transient
siRNA transfection was performed 18-24 hours after seeding using Dharmafect 1
reagent (Dharmacon). siGLO Red Transfection Indicator (D-001630-02, Dharmacon)
was included in the siRNA transfection and revealed a transfection efficiency of at
least 95% (supplementary material Fig. S3).
Generation of stable cell lines
pBabe constructs were used to generate retrovirus particles using the Phoenix
ecotropic packaging cells. HeLa or U2OS target cells were electroporated with an
expression plasmid for the ecotropic receptor 24 hours before infection. For infection,
cells were exposed to a 1:1 dilution of virus supernatant and fresh medium in
presence of polybrene (5 g/ml). Infected cells were split 24 hours after infection
and subjected to puromycin selection (0.5 mg/l for HeLa, 1.0 mg/l for U2OS).
Monoclonal colonies were isolated, propagated and tested for expression both by
fluorescent microscopy and immunoblot analysis.
The Tet repressor expressing construct pCAG-TRswas transfected in U2OS and
HeLa cells using the calcium phosphate method. Hygromycin-resistant clones were
tested by transient transfection of the luciferase construct TO Luc as described (van
de Wetering et al., 2003). HeLa TRsA7 and U2OS TRsC2.1 were used to generate
BTAF1 inducible knock-down cell lines by transfecting pTER shBTAF1 using
Dharmafect reagent 1 according to the manufacturer (Dharmacon). Cells were
selected using zeocin and hygromycin. Individual clones were isolated and tested for
efficient knockdown of BTAF1. After transient transfection of the ecotropic receptor
BTAF1, knock-down cell lines were infected with retroviruses expressing GFP-TBP,
NLS-GFP or H2B-GFP as described above.
Immunoprecipitation and immunoblotting
Cells were lysed in IP lysis buffer [50 mM Tris-HCl (pH 8.0), 150 mM KCl, 5 mM
MgCl2, 0.5 mM EDTA, 0.1% NP40, 10% glycerol and protease inhibitors]. Cleared
lysate was added to antibody-coupled protein A beads (anti-BTAF1 Pf299) or
Dynabeads Sheep-anti-mouse (GFP antibody or anti-TBP SL30). Precipitated proteins
were separated on 10% SDS-PAGE and blotted onto PVDF membranes. The
membrane was developed with the appropriate antibodies and ECL (Pierce).
Nuclear extract was prepared from HeLa GFP-TBP cell lines as described (Dignam
et al., 1983) and dialysed against buffer A [20 mM HEPES-KOH (pH 7.9), 20%
glycerol, 1 mM EDTA, 1 mM DTT, 1 mM PMSF, 400 mM KCl, 0.1% Triton X-
100]. Nuclear extract was applied on a Superose 12 column and eluted with buffer
A. Aliquots of the 0.4 ml fractions were separated on a 10% SDS-polyacrylamide
gel and analyzed by immunoblotting.
Cell imaging and FRAP studies were performed using a Zeiss 510 META confocal
LSM using the 488 nm laser line of a 200 mW Argon laser with tube current set at
6.1-6.3 A. All images and FRAP results were obtained using a 40? oil immersion
lens (NA1.3) using filters that pass emission light between 505 and 530 nm. Strip-
FRAP was performed as described (Farla et al., 2004; van Royen et al., 2009).
Briefly, a narrow strip (~0.75 m) spanning the width of the nucleus was monitored
every 21 ms using 0.2-0.5% laser power of the 488 nm laser line, an intensity at
which no significant monitor bleaching was observed. After 4 seconds the strip was
quickly bleached for 42 mseconds at maximum laser power. Subsequently, recovery
of fluorescence was monitored at 21 ms intervals for 40 seconds. For quantitative
analyses, fluorescence intensity in the strip was expressed relatively to the
fluorescence before bleaching. For each protein the average of 10 nuclei were taken.
Quantitative FRAP analysis
Computer modelling used to generate FRAP curves for fitting was based on Monte
Carlo simulation of diffusion and binding to immobile elements (representing
chromatin binding) in an ellipsoidal volume (representing the nucleus) (Farla et al.,
2004; Farla et al., 2005; Xouri et al., 2007). Bleaching simulation was based on
experimentally derived three-dimensional laser intensity profiles, which determined
the probability for each molecule to become bleached considering their 3D position
relative to the laser beam. Diffusion was simulated at each new time step t + ?t by
deriving a new position (xt+?t, yt+?t, zt+?t) for all mobile molecules from their current
position (xt, yt, zt) by xt+?txt+ G(r1), yt+?tyt+ G(r2), and zt+?tzt+ G(r3), where ri
is a random number (0 ≤ ri≤ 1) chosen from a uniform distribution, and G(ri) is an
inversed cumulative Gaussian distribution with 0 and s22D?t, where D is the
diffusion coefficient. Immobilization was derived from simple binding kinetics:
kon/koffFimm / (1–Fimm), where Fimm is the fraction of immobile molecules. The
probability per unit time to be released from the immobile state was given by
Pmobilisekoff1 / Timm, where Timm is the characteristic time spent in immobile
complexes expressed in unit time steps. The probability per unit time for each
mobile particle to become immobilized (representing chromatin-binding) was defined
as Pimmobilisekonkoff. Fimm / (1–Fimm), where koff1 / Timm. Note that kon and koff in this
model are effective rate constants with dimension s–1.
In all simulations, the size of the ellipsoid was based on the size of the measured
nuclei, and the region used in the measurements determined the size of the simulated
bleach region. The laser intensity profile using the simulation of the bleaching step
was derived from confocal images stacks of chemically fixed nuclei containing GFP
that were exposed to a stationary laser beam at various intensities and varying
exposure times. The unit time step ?t corresponded to the experimental sample rate
of 21 mseconds.
For quantitative analysis of the FRAP data, raw FRAP curves were normalized to
pre-bleach values and the best fitting curves (by ordinary least squares) were selected
from a large set of computer simulated FRAP curves in which three parameters
representing mobility properties were varied: diffusion rate (ranging from 0.04 to 25
m2/s), immobile fraction (0, 10, 20, …, 90%) and time spent in immobile state (2,
4, 8, 16, 32, 64, 128, 256, 512, 1024, ? s). Because individual curves generated by
Monte Carlo modelling, in contrast to analytically derived curves, show the slight
variation typical for diffusion of a limited amount of molecules in a small volume
(i.e. the exact, relatively small number of molecules in a small volume varies, a
phenomenon on which, for example, fluorescence correlation spectroscopy is based),
we did not use the best-fitting curve only, but took the ten best-fitting curves and
calculated the average diffusion coefficients and rate constants corresponding to
HeLa TRsshBTAF1 cells were grown for 8 days with or without doxycycline (1
mg/l). Cells were trypsinised and resuspended in hypotonic lysis buffer [10 mM
HEPES-KOH (pH 8.0), 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT and 0.5 mM
PMSF]. Cells were lysed using a 27G syringe. One volume of 2? whole cell
extraction buffer was added [40 mM HEPES-KOH (pH 8.0), 1.5 mM MgCl2, 4 mM
EDTA, 50% glycerol, 1.2 M NaCl, 0.5 mM DTT and protease inhibitors] and lysates
were tumbled for 30 minutes at 4°C followed by centrifugation for 45 minutes at
Journal of Cell Science
2670 Journal of Cell Science 123 (15)
90,000 g (TLA100.2) to yield the soluble fraction. High salt buffer [20 mM HEPES-
KOH (pH 7.9), 600 mM NaCl, 340 mM sucrose, 0.1% Triton X-100, 1 mM EDTA
and 1 mM -mercaptoethanol] was added to the pellet. Samples were homogenized
and tumbled for 30 minutes at 4°C followed by centrifugation for 30 minutes at
90,000 g to yield wash 1. This wash was repeated a second time (wash 2). One pellet
volume of nucleosome isolation buffer [20 mM HEPES-KOH (pH 7.9), 250 mM
sucrose, 10 mM MgCl2, 3 mM CaCl2, 0.1% Triton X-100 and 5 mM -
mercaptoethanol] was added to the pellet. Samples were prewarmed for 5 minutes
at 37°C. Micrococcal nuclease (10 U/ml) was added and the samples were incubated
for an additional 30 minutes at 37°C. Reaction was stopped with 10 mM EGTA and
cooled on ice. Soluble chromatin fraction was isolated by centrifugation for 5
minutes at 10,000 g in a table top centrifuge.
Subconfluent cultures of HeLa tk–cells were crosslinked by addition of 1%
formaldehyde in PBS for 10 minutes at 37°C. Cells were lysed in buffer [50 mM
Tris-HCl (pH 7.9), 1% SDS, 10 mM EDTA, 1 mM DTT, and protease inhibitors].
The lysate was sonicated six times for 30 seconds in a Bioruptor (Diagenode,
Belgium) resulting in DNA fragments of 200 to 600 bp. Soluble material was
supplemented with 0.1% Triton X-100 and 0.1% Na-DOC and incubated for 6 hours
with Dynabeads coupled to antibody of interest. Samples were processed as previously
described (Vermeulen et al., 2007). Binding to promoter and ORF regions of RPL31,
RPL34, RPS10 and H2BJ-H2AG locus DNA was measured by quantitative PCR
(Chromo4-equipped PCR cycler 5MJ Research, Bio-Rad) and normalized against
input samples from the same experiment. Primer sequences are available upon
Gene expression analysis
Total RNA was extracted using the RNeasy kit (Qiagen) including a DNase treatment
step. Total RNA (250 ng) was used for cDNA synthesis (Superscript II, Invitrogen).
Expression of RPL31, RPL34, RPS10, H2B, H2A, BTAF1 and GAPDH were analyzed
by quantitative PCR and normalized against a standard reference cDNA from
untreated HeLa tk–cells.
We thank M. Meisterernst, I. Davidson, N. Hernandez, H.
Stunnenberg and T. Oelgeschlager for antibodies; M. van de Wetering
and D. Chen for plasmids; L. Kleij and H.A.A.M. van Teeffelen for
technical assistance; Folkert van Werven, Pim Pijnappel and Markus
Kleinschmidt for critically reading of the manuscript; and the Timmers
group for valuable discussions. This work was supported by grants
from the European community (STREP LSHG-CT-2004-502950;
HPRN-CT 00504228 and EUTRACC LSHG-CT-2007-037445), the
Netherlands Organization for Scientific research (ALW 855.01.077/03-
DYNA-F-03 and CW 700.57.302), the Netherlands Proteomics Center,
ANR (05-BLAN-0396-01; Regulome), INCA (2008-UBICAN), AICR
(09-0258) and by the Human Frontier Science Program (LT-
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