MOLECULAR AND CELLULAR BIOLOGY, Jan. 2008, p. 435–447
Copyright © 2008, American Society for Microbiology. All Rights Reserved.
Vol. 28, No. 1
Transcriptional Activation of Histone Genes Requires NPAT-Dependent
Recruitment of TRRAP-Tip60 Complex to Histone Promoters
during the G1/S Phase Transition?‡
Michael DeRan,1,2† Mary Pulvino,1† Eriko Greene,1§ Chuan Su,1¶ and Jiyong Zhao1*
Department of Biomedical Genetics1and Department of Biochemistry and Biophysics,2University of
Rochester Medical Center, Rochester New York 14642
Received 6 April 2007/Returned for modification 15 June 2007/Accepted 11 October 2007
Transcriptional activation of histone subtypes is coordinately regulated and tightly coupled with the onset
of DNA replication during S-phase entry. The underlying molecular mechanisms for such coordination and
coupling are not well understood. The cyclin E-Cdk2 substrate NPAT has been shown to play an essential role
in the transcriptional activation of histone genes at the G1/S-phase transition. Here, we show that NPAT
interacts with components of the Tip60 histone acetyltransferase complex through a novel amino acid motif,
which is functionally conserved in E2F and adenovirus E1A proteins. In addition, we demonstrate that
transformation/transactivation domain-associated protein (TRRAP) and Tip60, two components of the Tip60
complex, associate with histone gene promoters at the G1/S-phase boundary in an NPAT-dependent manner.
In correlation with the association of the TRRAP-Tip60 complex, histone H4 acetylation at histone gene
promoters increases at the G1/S-phase transition, and this increase involves NPAT function. Suppression of
TRRAP or Tip60 expression by RNA interference inhibits histone gene activation. Thus, our data support a
model in which NPAT recruits the TRRAP-Tip60 complex to histone gene promoters to coordinate the
transcriptional activation of multiple histone genes during the G1/S-phase transition.
Histone proteins are integral components of eukaryotic
chromatin and play crucial roles in virtually all cellular pro-
cesses that involve chromosomal DNA, such as DNA replica-
tion, transcription, DNA repair, recombination, and chromo-
some segregation (27, 28). The bulk of the histones are
assembled with genomic DNA into chromosomes, and the
biosynthesis of multiple histone subtypes (replication-depen-
dent histones) is tightly coordinated and coupled with DNA
synthesis during the S phase of the cell cycle (20, 36, 45, 51).
Perturbation of such coordination and coupling can lead to the
loss of chromosomes, DNA damage, and developmental arrest
(39, 55, 62, 64), underscoring the importance of the proper
regulation of these events.
The rate of histone synthesis in S phase is regulated at both
the transcriptional and the posttranscriptional levels (20, 36,
45, 51). Histone gene transcription increases from 3- to 10-fold
as cells enter S phase (20). The transcription of each histone
subtype (H1, H2A, H2B, H3, and H4) in S phase is likely
regulated by proteins or protein complexes that interact di-
rectly with the subtype-specific regulatory elements (SSREs) in
the promoters of replication-dependent histone genes (20, 45).
Indeed, it has been shown that Oct1 and its coactivator com-
plex OCA-S interact with the H2B SSRE to activate H2B
transcription, while HiNF-P interacts with the H4 SSRE to
stimulate H4 expression (14, 40, 67). Components of OCA-S
include nuclear p38/glyceraldehyde-3-phosphate dehydroge-
nase (GAPDH) and lactate dehydrogenase, and the activity of
OCA-S is regulated by NAD?and NADH, suggesting a link
between the histone gene transcription and the cellular meta-
bolic state/redox status (67). The molecular mechanism under-
lying the regulation of H2B transcription by Oct1/OCA-S and
cellular metabolic/redox states, however, is not yet clear. The
mechanism of HiNF-P action is also not fully understood.
Additional protein factors, such as YY1, HIRA, FLASH, and
BZAP45, have also been implicated in the regulation of his-
tone gene transcription with unknown molecular mechanisms
(3, 12, 19, 31, 41, 61).
We and others have demonstrated previously that the cyclin
E-Cdk2 substrate NPAT associates with histone gene promot-
ers in S phase (66, 67). The overexpression of NPAT activates
promoters of multiple histone genes through the SSREs within
the promoters (66). The suppression of NPAT expression
through RNA interference or conditional knockout impedes
expression of all histone subtypes (16, 63). The promoter DNA
sequences of different histone subtypes are quite divergent,
and direct DNA binding by NPAT has not been detected.
Therefore, it was proposed that coordination of the transcrip-
tion of multiple histone subtypes by NPAT probably occurs
through the interaction of NPAT with factors that regulate
transcription of the individual subtypes (66). Indeed, both the
physical and the functional interactions of NPAT with Oct-1/
OCA-S and HiNF-P have been demonstrated (40, 67). Addi-
tionally, it has been shown that both the association of NPAT
with histone promoters and the NPAT-mediated histone tran-
* Corresponding author. Mailing address: Department of Biomedi-
cal Genetics, University of Rochester Medical Center, 601 Elmwood
Avenue, Rochester, NY 14642. Phone: (585) 273-1453. Fax: (585)
273-1450. E-mail: Jiyong_zhao@urmc.rochester.edu.
† These authors contributed equally to this work.
§ Present address: Sidney Kimmel Comprehensive Cancer Center,
Johns Hopkins University, Baltimore, MD 21231.
¶ Present address: Department of Parasitology, Nanjing Medical
University, Nanjing, Jiangsu 210029, People’s Republic of China.
‡ Supplemental material for this article may be found at http://mcb
?Published ahead of print on 29 October 2007.
scriptional activation are regulated by cyclin E-Cdk2 phosphor-
ylation (34, 53, 66). Hence, NPAT functions as a key global
regulator of coordinated transcriptional activation of multiple
histone subtypes during the G1/S-phase transition and links the
cell cycle machinery to the regulation of histone gene expres-
sion. In addition to histone gene expression, NPAT has been
shown to play a critical role in S-phase entry (16, 63, 65). Thus,
NPAT may also play a role in coupling histone expression with
the onset of DNA synthesis.
Despite recent advances in our understanding of the regu-
lation of histone gene transcription, the molecular mechanisms
underlying NPAT function, as well as the coordinated tran-
scriptional activation of histone subtypes and the coupling of
histone gene expression with DNA replication, have remained
largely unknown. Given the importance of NPAT in histone
gene transcription, an understanding of NPAT function would
likely advance the elucidation of these mechanisms.
The transformation/transactivation domain-associated pro-
tein (TRRAP) was initially identified as a factor that interacts
with the N terminus of c-Myc, as well as with the transactiva-
tion domain of E2F (37). TRRAP is an essential cofactor for
oncogenic transformation by c-Myc and E1A through its direct
interaction with these proteins (7, 37, 46). TRRAP also func-
tions as an essential cofactor for E2F-mediated transcriptional
activation (30). TRRAP, as well as its Saccharomyces cerevisiae
homolog Tra1, has been shown to be a key component of
several multiprotein histone acetyltransferase (HAT) com-
plexes (4, 6, 8, 24, 35, 52, 58) and is likely involved in the
recruitment of one or more of these HAT complexes to target
gene promoters through its interaction with transcriptional
activators, such as c-Myc, E2Fs, and p53 (2, 5, 15, 38, 56). In
addition to transcription-related functions, the TRRAP-con-
taining HAT complexes are also directly involved in double-
stranded DNA break repair (6, 24, 42, 47, 50). Thus, TRRAP
plays crucial roles in both the proliferation control and the
maintenance of genomic integrity.
To elucidate the molecular mechanism by which NPAT reg-
ulates transcriptional activation of histone genes, we carried
out a systematic deletion analysis to identify the domain(s)
critical for NPAT function. Here, we describe the identifica-
tion of a novel amino acid sequence motif termed the DLFD
motif, which is conserved in the NPAT, E2F, and E1A proteins
and is involved in the functions of these proteins. Our data,
together with previous observations, indicate that the DLFD
motif is required for the interaction of NPAT, and likely that
of E2F and E1A, with the cofactor TRRAP. Moreover, our
results show that TRRAP and Tip60 are recruited to histone
gene promoters at the G1/S-phase boundary by NPAT and are
required for the transcriptional activation of histone genes.
Consistent with the NPAT-dependent recruitment of TRRAP-
Tip60, histone H4 acetylation at histone promoters also in-
creases during the G1/S-phase transition in an apparently
NPAT-dependent manner. These results suggest a mechanism
by which the transcriptional activation of histone genes is co-
ordinately regulated during S-phase entry.
MATERIALS AND METHODS
Cell culture, transfection, immunoprecipitation, luciferase reporter assays,
Northern blotting, and antibodies. Human osteosarcoma U2OS cells and human
embryonic kidney 293T (HEK293T) cells were purchased from ATCC and cul-
tured in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal
bovine serum (FBS). Human colon carcinoma HCT116 cells carrying one con-
ditional allele of NPAT and one wild-type allele (referred to as NPATflox/?)
were generously provided by Wade Harper. Transient transfections were carried
out either by calcium precipitation or by using FuGENE6 (Roche) as described
previously (65, 66). Luciferase reporter assays, chromatin immunoprecipitation
(ChIP) assays, and Northern blotting were performed essentially as described
previously (16, 53, 65, 66).
Antibodies specific for the NPAT protein were described previously (65, 66).
Antibodies against TRRAP (catalog no. T-17), Tip60 (catalog no. N-17 and
K-17), and the GAL4 DNA binding domain (catalog no. RK5C1) were from
Santa Cruz Biotechnology. Antibodies for ?-tubulin (catalog no. GTU88) and
Flag tag (catalog no. M2) were from Sigma. Antibody specific for histone H4
acetylated at lysines 5, 8, 12, and 16 (catalog no. 06-866) was from Upstate
Expression plasmids and construction of fusion proteins. Plasmids carrying
the wild-type NPAT cDNA sequence were described previously (65, 66). For the
construction of mammalian expression plasmids encoding the yeast GAL4 DNA
binding domain fused with various NPAT sequences, DNA fragments encoding
the indicated NPAT sequences were generated either by restriction enzyme
digestion or by PCR and cloned in frame into the vector pFA-CMV (Stratagene).
Bacterial glutathione S-transferase (GST) fusion constructs were generated by
cloning the indicated NPAT fragments, prepared by PCR, into the vector pGEX
(Amersham). The point mutation constructs were generated by PCR and cloned
into appropriate expression plasmids. The PCR products were sequenced to
confirm that the correct DNA sequences were obtained.
Preparation of GST-NPAT fusion proteins. GST fusion proteins containing
either the wild-type or the mutant transactivation domain of NPAT were ex-
pressed in Escherichia coli BL21 cells. The bacterial cells were lysed by sonica-
tion. The lysates were incubated with glutathione-agarose beads (Sigma). After
the GST fusion proteins were washed with a buffer of 50 mM Tris-HCl (pH 8.0),
1 M NaCl, 0.5 mM phenylmethylsulfonyl fluoride (PMSF), they were eluted from
the beads with reduced glutathione (Sigma) in phosphate-buffered saline. The
eluted proteins were then loaded onto to an SP-Sepharose column (Amersham)
equilibrated with SP buffer (5 mM phosphate [pH 7.0], 1 mM EDTA, 5%
glycerol, 0.5 mM PMSF). The flowthrough was loaded onto a DEAE-Sepharose
column (Amersham) equilibrated with buffer D (20 mM Tris-HCl [pH 8.0 ], 100
mM NaCl, 1 mM EDTA, 5% glycerol, 0.5 mM PMSF). The proteins were eluted
from the column by a 100 mM to 1 M NaCl gradient in D buffer. Fractions
containing the fusion proteins were combined and dialyzed against phosphate-
Purification of nuclear proteins that interact with the transactivation domain
of NPAT. HeLa S3 cells (from the National Cell Culture Center) were lysed in
buffer A (10 mM HEPES [pH 7.9 ], 10 mM KCl, 10 mM EDTA, 0.4% NP-40;
Panomics) and centrifuged to collect the nuclei. The nuclei were treated with
buffer B (20 mM HEPES [pH 7.9 ], 400 mM NaCl, 1 mM EDTA, 10% glycerol;
Panomics) and centrifuged to collect nuclear proteins in the supernatant. To
remove proteins that interact with GST or with the NPAT transactivation do-
main independently of the DLFD motif, the nuclear extract was precleared by
incubation with a mutant NPAT transactivation domain fusion protein (amino
acids [aa] 262 to 329 or aa 262 to 350 [AAA]) and glutathione beads. The
precleared nuclear extract was then incubated with GST-NPAT aa 262 to 338 or
GST-NPAT aa 262 to 350 and glutathione beads. The beads were washed with
buffer B with 0.1% NP-40. The proteins associated with the GST-NPAT fusion
were eluted from the beads with 0.2% Sarkosyl in buffer B (43). The eluted
proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electro-
phoresis (SDS-PAGE) and stained with colloidal Coomassie blue (Invitrogen).
Protein bands specific for samples purified from the GST-NPAT aa 262 to 338 or
from GST-NPAT aa 262 to 350 beads were excised for liquid chromatography
tandem mass spectrometry (LC-MS/MS) analysis at Taplin Biological Mass
Spectrometry Facility of Harvard Medical School.
RNA interference by shRNA and quantitative real-time PCR analyses. For
generating small hairpin RNAs (shRNAs) targeting human TRRAP mRNA, two
sequences (shTRRAP-1, GGCTCGAAAGGAATGGATCC; and shTRRAP-2,
GGCGACTACCTGGAGAACAT) were cloned into the pBS/U6 vector (54). The
pBS/U6 plasmid expressing an shRNA specific for the firefly luciferase mRNA
sequence (ACAAGACAATTGCACTGA) (57) was used as a control with North-
ern blotting experiments. The pBS/U6/Ski plasmid expressing an shRNA specific for
the mouse Ski mRNA sequence (GCTGGAGGCAGAGTTGGAG) was also used
as a control. For the Tip60 knockdown experiment, we employed two pLKO.1
constructs expressing Tip60-specific shRNAs, the RNA interference Consortium
clones TRCN0000020314 (shTip60-1) and TRCN0000020315 (shTip60-2), targeting
the sequences CGTCCATTACATTGACTTCAA and CCTCAATCTCATCAAC
436DERAN ET AL.MOL. CELL. BIOL.
TACTA, respectively. The pLKO.1 vector expressing a scrambled sequence (GTT
CTCCGAACGTGTCACG) was used as a control.
After the knockdown, Tip60 mRNA levels were measured by using real-time
quantitative PCR (qPCR). RNA was isolated using TRIzol (Invitrogen) reagent and
then used as a template for cDNA synthesis by Moloney murine leukemia virus
reverse transcriptase (Invitrogen) as described by the manufacturer. qPCRs using iQ
SYBR green Supermix (Bio-Rad) were performed in triplicate with either the
Tip60-specific primers TCCAGGCAATGAGATTTACCG and TCTTATGGTCA
AGGAAACACTTGG or the GAPDH-specific primers CATGGGTGTGAACCA
TGAGA and CAGTGATGGCATGGACTGTG. Gene expression was normalized
ChIP assays. The ChIP assays were carried out essentially as described previously
(66). Briefly, HCT116 NPATflox/?cells were cross-linked with 1% formaldehyde for
10 min at room temperature. After undergoing cross-linking, the cells were lysed,
and the chromatin was sonicated into 400- to 1,000-bp fragments, using a Branson
450 sonicator. The lysates were cleared by centrifugation at 11,000 rpm for 5 min at
room temperature. A fraction of the lysates was saved as an input control, and the
remaining lysates were diluted 10-fold with the dilution buffer. The diluted lysates
were precleared with protein G beads (protein G-Sepharose; Amersham) and then
incubated with antibodies specific for either NPAT, TRRAP, TIP60, acetylated H4,
or the Flag tag and 15 ?l of protein G-Sepharose at 4°C. Immunoprecipitated
samples were washed three times with the dilution buffer and twice with a wash
solution (50 mM Tris-HCl [pH 8.0], 500 mM NaCl, 1 mM EDTA, 0.5% Triton
X-100) and then eluted with 1% SDS and 10 mM Tris-EDTA buffer at 65°C. The
samples were treated with proteinase K at 50°C for 1 h. The input control samples
were treated with 10 ?g RNase at 37°C for 3 h and then 100 ?g proteinase K at 50°C
for 1 h. Reversion of cross-linking was carried out at 65°C overnight. DNA was
phenol-chloroform extracted and ethanol precipitated. The DNA samples were
subsequently quantitated by real-time PCR with SYBR green. Data analysis was
performed using the method described previously (33). For the amplification of H4
sequences, the primers (5?-CTATTTCGGTTTGGCCCTTT-3? and 5?-CTGAGGC
AGCGCCTTTATAC-3?) were used to cover about a 120-bp promoter region of the
H4/e gene. For the amplification of the H2B sequence, the previously described
primer set (66), which covers a 180-bp promoter sequence upstream of the initiation
codon ATG, was used.
NPAT contains a transactivation domain with a motif con-
served in the E2F and E1A proteins. In order to elucidate the
mechanism by which NPAT coordinates histone gene tran-
scription during the G1/S-phase transition, we set out to iden-
tify the domain(s) in NPAT that is required for transcriptional
activation. Short deletions from either the N terminus or the C
terminus of NPAT abolished its ability to activate histone
promoters (data not shown) (60), hampering the analysis of the
internal domain(s) required for histone gene expression by the
use of histone promoter-reporter assays. To circumvent this
difficulty and to facilitate the identification of the domain in-
volved in the transactivation, we fused full-length NPAT or
various NPAT terminal deletions with the DNA binding do-
main (DBD) of the yeast transcriptional activator GAL4 pro-
tein and tested the fusions’ abilities to activate a GAL4-respon-
sive reporter. Fusion proteins containing either full-length
NPAT or regions encompassing aa 262 to 350 of NPAT exhib-
ited significant transcriptional activity (Fig. 1A and B), sug-
gesting that a transcriptional activation domain resides in this
region. The fusion of aa 262 to 350 of NPAT with the DBD
activated the GAL4 reporter dramatically (Fig. 1C), confirm-
ing that this region carries a transcriptional activation function.
To further define the transcriptional activation domain, we
generated additional deletion constructs and tested their abil-
ities to activate transcription. The domain that consists of aa
262 to 338 functions as a potent transcriptional activator (for
convenience, it was referred to as the transactivation domain of
NPAT). In contrast, the fusion containing aa 262 to 329,
though expressed at a level similar to that of the fusion con-
taining aa 262 to 338, has a greatly decreased ability to activate
transcription (Fig. 1C). These data indicate that aa 329 to 338
are crucial for transcriptional activation.
The sequence aa 329 to 338 of NPAT contains a sequence
motif that is also present in the transcriptional activation domain
of E2Fs (9, 26, 49), as well as in the conserved region 1 (CR1) of
the adenovirus E1A protein that was shown to be essential for
E1A-mediated transformation (10, 11) (Fig. 1D). The most con-
served residues in this motif are the amino acids DLFD, and thus
we refer to this element as the DLFD motif. Previous studies
suggest that the region containing the DLFD motif is involved in
E2F and E1A function (7, 13). To assess the significance of the
DLFD motif in NPAT-mediated transcriptional activation, we
replaced the amino acids LFD with alanine residues in the NPAT
(262-350)-GAL4-DBD fusion protein. This replacement had no
effect on the expression of the fusion protein but severely com-
promised its transcriptional activation ability (Fig. 1C), suggesting
that the DLFD motif is critical for the function of the NPAT
transactivation domain. To determine whether the region consist-
ing of aa 262 to 350 is also required for NPAT to activate histone
350 deleted. This mutant NPAT was unable to activate a histone
H4 promoter reporter, indicating that these residues are essential
for histone gene activation by NPAT (Fig. 1E). To further dem-
onstrate the role of the DLFD motif in NPAT-mediated histone
gene activation, we changed the amino acids LFD to AAA in the
full-length NPAT and tested histone promoter activation by the
mutant NPAT. Similar to the GAL4 DBD fusion proteins, mu-
tation of the LFD sequences in NPAT, which had no effect on
NPAT expression, resulted in a marked decrease in histone pro-
moter activation (Fig. 1E). These results indicate that the DLFD
motif is critical for NPAT to regulate histone gene activation.
Thus, the DLFD motif may represent an important functional
domain conserved in NPAT, E2F, and E1A.
TRRAP interacts with the transactivation domain of NPAT
via the DLFD motif. Having identified a transactivation domain
in NPAT that contains a conserved functional DLFD motif, we
sought next to identify proteins that interact with this transacti-
vation domain through the DLFD motif. In order to facilitate this
investigation, two wild-type and two mutant forms of the trans-
activation domain were constructed and fused to GST (Fig. 2A).
These GST fusion proteins were purified from bacteria and sub-
sequently incubated with HeLa nuclear extract. Proteins that in-
teracted with these NPAT transactivation domain fusion proteins
were affinity purified, separated by SDS-PAGE, and visualized by
Coomassie staining. The staining revealed that the wild-type
weak or no interaction with the mutant transactivation domain
(Fig. 2B, compare lanes 3 and 4 with lanes 1 and 2). The identities
of the purified interacting proteins were determined by mass
This analysis identified a number of proteins that appear to
interact with the NPAT transactivation domain through the
DLFD motif. Among the proteins identified were YY1 and
BZAP45 (Fig. 2B), which are proteins that had previously been
implicated in the regulation of histone gene transcription (12, 31,
41, 61). Mass spectrometric analysis also revealed the presence of
heat shock protein 70 (Hsp70), which was previously shown to be
a component of the OCA-S coactivator complex (67). These find-
VOL. 28, 2008NPAT RECRUITS TRRAP-Tip60 COMPLEX437
ings suggest that this approach identifies authentic proteins in-
volved in the transcriptional regulation of histone genes. In addi-
tion to these previously described regulators, we identified several
components of the Tip60/hNuA4 histone acetyltransferase com-
plex, TRRAP, Tip48, and Tip49 (6, 8) (Fig. 2B). Interestingly,
TRRAP has previously been characterized as a cofactor that is
Moreover, the DLFD motifs in E2F and E1A are apparently
involved in their interactions with TRRAP (7, 30). Hence,
TRRAP may also be an important cofactor for NPAT-mediated
transactivation, and its interaction with NPAT is likely mediated
through the DLFD motif.
To confirm the interaction between NPAT and TRRAP, ob-
served with the initial mass spectrometry analysis, the GST fusion
proteins were once again incubated with HeLa nuclear extract,
and the interaction of TRRAP with the GST-fusion proteins was
determined by Western blotting analysis with a TRRAP-specific
antibody. As shown in Fig. 2C, the wild-type NPAT transactiva-
tion domain, but not the mutant domain, interacts with TRRAP,
validating the observation that TRRAP interacts with the trans-
FIG. 1. Identification of a functional DLFD motif. (A) Deletion analysis of the GAL4-NPAT fusion proteins. pFA-CMV vector, which expresses the
GAL4 DBD, or a pFA-CMV plasmid that expresses the indicated DBD-NPAT fusion protein was transfected together with the GAL4-responsive
was also cotransfected. Twenty-four hours posttransfection, the cells were lysed and the activities of luciferase and ?-galactosidase were assayed. The
relative activity was calculated as previously described (66). The mean results and standard deviations from three independent experiments are depicted.
(B) Schematic representation and summary of results shown in panel A. The positions of the first and last amino acids of the fused NPAT sequences are
indicated on the left. Full-length NPAT contains 1,427 amino acids. The dotted lines indicate the region of NPAT required for transactivation. (C) The
DLFD sequence is required for transactivation. The indicated GAL4 DBD-NPAT fusion proteins were tested for their ability to activate the pFR-Luc
reporter. The expression levels of the fusion proteins were analyzed by Western blotting using an antibody specific for the GAL4-DBD. The Western blot
of ?-tubulin was used as an equal loading control. (D) Conservation of the DLFD motif in NPAT, E2F and E1A proteins. The sequences surrounding
the amino acids DLFD from human NPAT, human and Drosophila E2F and several human adenovirus E1A proteins are aligned. Identical amino acid
residues are shaded black, and similar residues are shaded gray. The numbers indicate the locations of the amino acids in NPAT. (E) Essential role for
the DLFD motif in NPAT-mediated histone promoter activation. Full-length wild-type NPAT, a mutant NPAT with the deletion of amino acid residues
262 to 350, and a mutant NPAT with the LFD sequence in the transactivation domain replaced by AAA sequence were tested for their ability to activate
a histone H4 promoter (pGLH4(65)) as previously described (66). The expression of the wild-type and mutant NPAT was analyzed by Western blotting
with an NPAT-specific antibody. The Western blot of ?-tubulin was used as a loading control.
438DERAN ET AL. MOL. CELL. BIOL.
FIG. 2. Interaction of TRRAP with NPAT. (A) Schematic representation of GST-NPAT fusion proteins used for affinity purification. (B). Identi-
fication of nuclear proteins interacting with functional transactivation domain of NPAT. HeLa nuclear extract was incubated with the indicated GST
fusion proteins bound to the glutathione-agarose beads. Proteins associated with the fusion proteins were eluted and separated on an 8% SDS-PAGE
gel. The proteins were visualized by colloidal Coomassie blue staining. The proteins were then identified by LC-MS/MS. (C) Interaction of TRRAP with
NPAT transactivation domain requires the DLFD motif. The indicated GST fusion proteins were incubated with HeLa nuclear extract. Interaction of
TRRAP with the fusion proteins was analyzed by Western blotting using a TRRAP-specific antibody (top panel). Ten percent of the input was loaded
for analysis of the TRRAP level in HeLa nuclear extract. (Lower panel) a Coomassie blue-stained SDS-PAGE gel indicating equal amounts of GST
with a monoclonal antibody specific for NPAT (DH3) or HA tag (12CA5). The immunoprecipitates were analyzed by Western blotting with a
TRRAP-specific antibody. Five percent of the input was loaded for analysis of TRRAP in HeLa nuclear extract. One hundred percent of the
immunoprecipitates was loaded for the analysis. (E) Confirmation of in vivo interaction of TRRAP with NPAT. HCT116 NPATflox/?cells were infected
with either Ad-LacZ or Ad-Cre (Baylor Vector Development Laboratory) as indicated. Three days after infection, the whole cell lysates were
immunoprecipitated with an NPAT-specific antibody. The presence of TRRAP in the immunoprecipitates was analyzed by Western blotting with a
TRRAP-specific antibody. The input lanes were loaded with 2% (for TRRAP) and 5% (for NPAT) of the starting material used for the immunopre-
cipitation. One hundred percent of the immunoprecipitated material was analyzed for IP samples. Under the experimental conditions, the NPAT
knockout efficiency is about 80% as previously reported (63).
activation domain of NPAT via the DLFD motif. To determine
whether endogenous NPAT and TRRAP interact, nuclear ex-
tracts prepared from HeLa cells were subjected to immunopre-
cipitation with an NPAT-specific antibody. TRRAP protein can
be readily detected in the immunoprecipitates of the NPAT an-
tibody but not in the immunoprecipitates of a control antibody
(Fig. 2D), indicating that NPAT associates with TRRAP in vivo.
Endogenous NPAT in anti-TRRAP immunoprecipitates was not
detected, likely due to the low affinity of the available TRRAP
antibody (data not shown). To verify that the coimmunoprecipi-
tation of TRRAP results from the in vivo interaction between
TRRAP and NPAT, rather than examine nonspecific immuno-
precipitation by the anti-NPAT antibody, we compared coimmu-
noprecipitations of TRRAP by the NPAT-specific antibodies in
both the wild-type NPAT and the HCT116 conditionally null
NPAT cells. These latter cells (referred to as NPATflox/?cells)
carry one mutant NPAT allele and one wild-type allele with exon
2 flanked by loxP sites. Infection of the NPATflox/?cells with
adenovirus that expresses Cre recombinase (Ad-Cre) induces the
excision of exon 2 of the wild-type allele, generating NPAT-
deficient cells (63). As shown in Fig. 2E, the amount of TRRAP
that coimmunoprecipitated is reduced in the Ad-Cre-infected
NPATflox/?cells, where NPAT levels are decreased but not com-
pletely depleted under the experimental conditions. Coimmuno-
precipitation of TRRAP with NPAT is due to the specific inter-
action of these two proteins, rather than a nonspecific association
of these proteins with an antibody, as the antibody specific for the
hemagglutinin (HA) tag immunoprecipitates neither NPAT nor
TRRAP (data not shown). Together, these data show that NPAT
associates with TRRAP in vivo, likely through the DLFD motif.
NPAT recruits TRRAP to histone promoters during the
G1/S-phase transition. We and others have previously shown
that NPAT physically associates with histone promoters in vivo
and that this association increases upon S-phase entry (66, 67). As
NPAT interacts with TRRAP, which has been shown to be re-
cruited by Myc and E2F to their respective target promoters (15,
38, 46, 56), it is possible that NPAT may recruit TRRAP to
histone promoters to activate histone gene transcription during
the G1/S-phase transition. To test this possibility, we investigated
NPATflox/?cells under different conditions by ChIP assays. Sim-
ilar to the cell cycle-dependent association of NPAT with the
histone promoters (Fig. 3A) (67), the association of TRRAP with
a histone H4 promoter increases about 10-fold in early S-phase
cells compared to that with quiescent cells (Fig. 3A). In contrast,
the association of TRRAP with the H4 promoter in vivo is com-
promised in the NPAT-deficient (Ad-Cre-infected) cells. Thus,
TRRAP associates with histone promoters in S phase, and this
association depends on NPAT. Under the experimental condi-
tions used, the NPAT knockout efficiency is approximately 80 to
85% (63) Fig. 2 (and data not shown). The small increase in the
association of TRRAP with the H4 promoter in the Ad-Cre-
infected cells upon S-phase entry is likely due to the effect of the
remaining NPAT protein. To determine more precisely the cell
cycle stage at which TRRAP becomes associated with histone
promoters, we analyzed the association of TRRAP with histone
promoters at additional time points during the G1-to-S-phase
transition. As shown in Fig. 3B and Table 1, the association of
TRRAP with the H4 promoter in vivo occurs at the G1/S-phase
boundary, as does the association of NPAT with the H4 pro-
moter. The association of TRRAP and that of NPAT with a
histone H2B promoter follow virtually identical kinetics (Fig. 3C).
Thus, the associations of both NPAT and TRRAP with histone
promoters occur during the G1/S-phase transition, a point when
S-phase-specific histone gene transcription is initiated. Again, the
increased association of TRRAP with the histone promoters re-
lies on NPAT (Fig. 3B and C). Thus, TRRAP associates with
histone promoters in vivo at the G1/S-phase boundary and in an
Since NPAT is required for S-phase entry, the decreased asso-
ciation of TRRAP with the histone promoters in NPAT-deficient
cells could result from a cell cycle block at the G1/S-phase tran-
sition, rather than from the direct requirement for NPAT to
recruit TRRAP to these promoters. To distinguish these two
possibilities, the cells were treated with aphidicolin, a DNA poly-
merase-specific inhibitor, to block S-phase entry, and the associ-
ation of TRRAP with the H2B and H4 promoters was then
analyzed. In the presence of aphidicolin, both the control (Ad-
LacZ-infected) and the NPAT-deficient (Ad-Cre-infected) cells
were blocked at the G1/S-phase boundary (Table 2) (53). Similar
to the results with untreated cells (Fig. 2A and B), the association
of TRRAP with the H2B and H4 promoters was markedly de-
creased in Ad-Cre-infected (NPAT-deficient) cells compared
with that in the Ad-LacZ-infected cells (Fig. 3D), indicating that
the association of TRRAP with histone promoters requires
NPAT directly. These results, together with the above observa-
tion that NPAT-dependent association of TRRAP with histone
promoters is greatly increased in late G1phase (Fig. 3A and B),
demonstrate that NPAT recruits TRRAP to histone gene pro-
moters during the G1-to-S-phase transition.
NPAT-dependent association of Tip60 with histone promot-
ers. TRRAP is a component of several multiprotein histone
acetyltransferase complexes and is involved in the recruitment of
these HAT complexes to chromatin (5, 6, 8, 38, 46). Thus, the
association of TRRAP with histone promoters suggests that one
or more HAT complexes also associate with the histone promot-
ers. Since the transactivation domain of NPAT interacts with
other components of the Tip60 complex, Tip48 and Tip49 (Fig.
2B), we hypothesized that NPAT may interact with the Tip60
complex and recruit Tip60 to histone promoters during the G1/
S-phase transition. To test this hypothesis, we investigated the
association of Tip60 with histone promoters by using ChIP assays.
The results shown in Fig. 4 indicate that, similar to TRRAP,
Tip60 associates with histone promoters in an NPAT-dependent
fashion. Furthermore, the NPAT-dependent association of Tip60
with histone promoters likely results from a direct requirement
that NPAT recruit Tip60 to the histone promoters, rather than
from an indirect cell cycle effect, as the experiments were carried
out in the presence of aphidicolin as described above. Thus, our
data suggest that NPAT recruits Tip60, and likely the Tip60 HAT
complex as well, to the histone gene promoters at the G1/S-phase
Effect of NPAT on histone acetylation at the histone gene
promoters. Since it appears that NPAT recruits Tip60 to his-
tone promoters, NPAT may regulate histone acetylation at
histone promoters. To test this possibility, we investigated his-
tone acetylation of chromatin surrounding the sites where
NPAT associates with an H4 promoter. It had been reported
previously that Tip60, like its yeast homolog Esa1, preferen-
tially acetylates histone H4 (1, 24), and thus, we focused our
440DERAN ET AL.MOL. CELL. BIOL.
study on H4 acetylation. Similar to the associations of NPAT,
TRRAP, and Tip60 to the H4 promoter, the acetylation of his-
tone H4 is greatly increased at the G1/S-phase boundary (Fig. 5A
and Table 1). This increase in H4 acetylation is partially inhibited
in the NPAT-deficient cells (Fig. 5A and see Discussion), sug-
gesting that NPAT plays a role in the H4 acetylation at the
histone promoters. Again, the NPAT-dependent H4 acetylation
is independent of S-phase entry (Fig. 5B), reminiscent of the
NPAT-dependent association of TRRAP and Tip60 to histone
promoters. Thus, histone H4 acetylation relies on NPAT and
correlates with the association of TRRAP and Tip60 at histone
promoters during the G1-to-S-phase transition.
FIG. 3. NPAT recruits TRRAP to histone promoters. (A) NPAT-dependent association of TRRAP with a histone H4 promoter at early S phase.
(Left panel) HCT116 NPATflox/?cells were infected with either Ad-LacZ or Ad-Cre as indicated. Sixty hours after infection, the cells were starved for
36 h in medium containing 0.1% FBS. The cells were then stimulated to enter the cell cycle with 10% FBS. The cell cycle distribution of the cells at the
indicated times was analyzed as previously described (53, 66). (Right panel) ChIP assays on serum-starved (0 h) or early S-phase cells (12 h). The
cross-linked chromatin was immunoprecipitated with the indicated antibodies, and the presence of the H4 promoter DNA sequence in the immuno-
precipitates was analyzed as described in Materials and Methods. The antibody specific for the Flag tag was used as a negative control. (B) NPAT-
dependent association of TRRAP to an H4 promoter at the G1/S-phase boundary. The ChIP assays were carried out as described for panel A, except
that the cells were harvested at 0, 8 and 10 h after serum stimulation. The amounts of DNA immunoprecipitated at different time points were compared,
relative to the amounts at 0 h set as 1. (C) NPAT-dependent association of TRRAP to an H2B promoter at the G1/S-phase boundary. The ChIP assays
were performed as described for panel B, except that primers specific for an H2B promoter were used. (D) NPAT-dependent, but S-phase-independent
association of TRRAP with the H4 and H2B promoters. The NPATflox/?cells were treated as described for panel A, except that 5 ?g/ml aphidicolin was
added into the medium at the same time when the cells were stimulated with 10% FBS to block S-phase entry (53). The cells were harvested 18 h after
serum and aphidicolin addition, and the ChIP assays were carried out as described for panels B and C. Under these experimental conditions, the levels
of association of NPAT and TRRAP with histone promoters are 50 to 55% of those observed at 12 h after serum starvation.
TABLE 1. Cell cycle distribution before and after serum stimulationa
Cell distribution (%) at the indicated time (h) of stimulation
LacZ CreLacZ CreLacZCre LacZ Cre
aHCT 116 NPATflox/?cells were treated as described in the legend to Fig. 3B. At the indicated times, the distribution of cells in the cell cycle was analyzed as
described in the legend to Fig. 3A.
VOL. 28, 2008NPAT RECRUITS TRRAP-Tip60 COMPLEX441
TRRAP and Tip60 are required for histone promoter acti-
vation. Our observations that TRRAP interacts with NPAT
and associates with histone promoters in vivo in an NPAT-
dependent manner suggest that TRRAP also functions as a
cofactor for histone gene transcription. To test this idea di-
rectly, we examined the requirement for TRRAP in histone
gene promoter activation. For this purpose, we designed two
mRNA. Using these constructs, we were able to achieve a mod-
erate knockdown of TRRAP expression (50 to 60%) (Fig. 6A).
Interestingly, TRRAP knockdown resulted in a significant reduc-
tion of promoter activity from both the H4 and the H2B promot-
ers (Fig. 6B and C), while it had no effect on either the CMV
promoter or mutant histone promoters (see Fig. S1 in the sup-
plemental material). These data suggest that TRRAP is required
for the activation of multiple histone gene promoters. The obser-
vation that a moderate depletion of TRRAP protein by TRRAP-
specific shRNA results in a significant inhibition of histone pro-
moter activation raises the possibility that TRRAP is a limiting
factor for histone gene activation.
As it was previously shown that the knockout of TRRAP
leads to proliferation arrest (23), one possible scenario would
be that the decrease observed for the histone promoter activity
when TRRAP expression is perturbed might result from a cell
cycle effect rather than from a direct requirement for TRRAP
in the activation of histone gene transcription. To assess this
possibility, we examined the cell cycle distribution in cells that
express the TRRAP-specific shRNAs in parallel with the anal-
ysis of the histone promoter activation. Flow cytometry anal-
ysis revealed that the S-phase populations of cells transfected
with the TRRAP shRNA constructs were similar to that of the
vector-transfected cells (Table 3). This observation indicates
that the decreased levels of histone promoter activity in cells
with perturbed TRRAP expression are not a result of cell cycle
arrest but rather are due to the requirement for TRRAP in
histone gene promoter activation.
To determine whether TRRAP is required for endogenous
histone gene expression, we examined the levels of histone
mRNAs in TRRAP-knocked-down cells by Northern blotting
35% reduction in the levels of endogenous histone mRNAs (Fig.
6D). This effect is less than that seen with H4 and H2B promoter-
reporter assays (Fig. 6B and C). However, this is not unexpected,
as it is known that posttranscriptional mechanisms also play a
major role in the regulation of histone mRNA levels in mamma-
lian cells (20, 36, 45). Together, these data indicate that TRRAP
is a critical cofactor for histone gene transcription.
Similar to TRRAP, Tip60 also was observed to associate with
histone promoters. Therefore, we investigated whether Tip60 is
involved in histone promoter activation. We employed two
shRNAs targeting two distinct sequences in Tip60 mRNA to
knock down the expression of Tip60 (Fig. 7A). As shown in Fig.
7B and C, the suppression of Tip60 expression inhibits both the
H2B and the H4 promoter activities, suggesting that Tip60 is also
required for histone gene transcriptional activation.
The work presented here provides mechanistic insights into
the coordinated transcriptional activation of multiple histone
FIG. 4. NPAT-dependent association of Tip60 with histone pro-
moters. The treatment of NPATflox/?cells and ChIP assays with a
Tip60-specific antibody were carried out as described for Fig. 3D.
FIG. 5. NPAT-dependent increase in histone H4 acetylation at
histone promoters at the G1/S-phase boundary. (A) Increase in histone
H4 acetylation at the G1/S-phase boundary. The treatment of the cells
and ChIP analysis using an antibody specific for acetylated histone H4
were performed as described for Fig. 3B. (B) S-phase-independent
increase in histone H4 acetylation. The treatment of cells and the ChIP
assays with an antibody specific for acetylated histone H4 were carried
out as described for Fig. 3D.
TABLE 2. Cell cycle distribution in the presence of aphidicolina
Cell distribution (%) at the indicated cycle
aHCT 116 NPATflox/?cells were treated as described in the legend to Fig. 3D.
The cell cycle distribution in the presence of aphidicolin was analyzed as de-
scribed in the legend to Fig. 3A.
442DERAN ET AL.MOL. CELL. BIOL.
subtypes by NPAT at the G1/S-phase transition. Our data in-
dicate that the TRRAP-Tip60 complex interacts with NPAT
and becomes associated with histone promoters at the G1/S-
phase boundary in an NPAT-dependent manner. Moreover,
consistent with the presence of the TRRAP-Tip60 HAT com-
plex at histone promoters, histone H4 acetylation, which is
associated with transcriptional activation (25, 59), increases at
histone promoters during the G1/S-phase transition. This in-
FIG. 6. TRRAP is required for transcriptional activation of histone genes. (A) Knockdown of TRRAP expression by TRRAP-specific shRNAs.
HEK293T cells were transfected with pBS/U6 vector (control) or pBS/U6 plasmids expressing shRNAs specific for human TRRAP mRNA
(shTRRRAP1 and shTRRAP2, respectively). pBabe-puro that carries a puromycin resistance gene was also cotransfected for selection with puromycin.
harvested and expression of TRRAP was analyzed by Western blotting using a TRRAP-specific antibody. (B) Effect of TRRAP knockdown on H4
promoter activation. 293T cells were transfected with an H4 promoter-luciferase reporter (66), together with the pBS/U6 vector, pBS/U6 expressing a
mouse Ski-specific shRNA or pBS/U6 expressing a TRRAP-specific shRNA. To normalize for transfection efficiency, the cells were also cotransfected
with pCMV-LacZ. At the indicated time after transfection, the cells were harvested and the activities of luciferase and ?-galactosidase were assayed as
previously described (66). (C) Effect of TRRAP knockdown on H2B promoter activation. Activation of an H2B promoter (pGLH2B) (66) in cells
transfected with the vector or plasmids expressing TRRAP-specific shRNAs or mouse Ski-specific shRNA was assayed as described for panel B.
(D) Effect of TRRAP knockdown on the expression of endogenous histones. HEK293T cells were transfected with pBS/U6 expressing shRNAs specific
for the luciferase gene (control) or TRRAP (shTRRAP-1 and shTRRAP-2). pBabe-puro was also cotransfected for selection. Twenty-four hours after
transfection, puromycin was added into the medium to select for transfected cells. Two days after puromycin selection, the cells were harvested and
mRNA levels of multiple histone genes were analyzed by Northern blotting, and the hybridization signals were quantitated using a phosphorimager as
previously described (16). The levels of GAPDH were used as loading controls. The relative levels of the indicated histone mRNA in shTRRAP-
expressing cells compared with those in control cells, after normalization with GAPDH signal, are shown below each Northern blot.
TABLE 3. Cell cycle distribution of cells transfected with indicated shRNA constructsa
Cell distribution (%) ? SD at the indicated time
Day 2Day 3
Control shTRRAP-1shTRRAP-2Control shTRRAP-1shTRRAP-2
37.1 ? 3.8
40.5 ? 2.1
22.4 ? 3.0
35.4 ? 6.4
39.5 ? 2.2
25.0 ? 4.6
27.4 ? 3.9
45.7 ? 4.8
26.9 ? 4.2
34.2 ? 3.0
44.7 ? 1.2
21.2 ? 2.1
37.4 ? 2.1
37.4 ? 5.1
25.3 ? 3.7
30.7 ? 7.1
37.5 ? 6.1
31.8 ? 11.0
a293T cells were transfected as described in the legend to Fig. 6B. The distribution of cells in the cell cycle was analyzed as described in the legend to Fig. 3A. The
mean results and standard deviations from three independent experiments are shown.
VOL. 28, 2008NPAT RECRUITS TRRAP-Tip60 COMPLEX443
FIG. 7. Tip60 is required for histone promoter activation. (A) Tip60 expression is reduced by two Tip60-specific shRNA constructs. U2OS cells
were transfected with pLKO.1 constructs carrying Tip60-specific shRNA sequences or a control nonspecific scrambled sequence. pLKO.1 carries
a puromycin resistance gene allowing for selection of transfected cells. Twenty-four hours after transfection, puromycin was added to culture media
and transfected cells were selected. The cells were harvested three days after selection. Relative Tip60 mRNA levels were determined by real-time
qPCR. The qPCR data were normalized to expression of GAPDH. (B) Effect of Tip60 knockdown on H2B promoter activation. U2OS cells were
transfected with an H2B promoter luciferase reporter (66), together with pLKO.1/Scramble (shScr) or pLKO.1 expressing shRNAs specific for
human Tip60 mRNA (shTip60-1 and shTip60-2, respectively). To normalize for transfection efficiency, the cells were cotransfected with pCMV-
LacZ. At the indicated time after transfection, the cells were harvested and the activities of luciferase and ?-galactosidase were assayed as
described previously (66). (C) Effect of Tip60 knockdown on H4 promoter activation. Activation of an H4 promoter luciferase reporter (66) in cells
transfected with pLKO.1/Scramble (shScr), pLKO.1/shTip60-1 or pLKO.1/shTip60-2 was assayed as described for panel B.
444DERAN ET AL.MOL. CELL. BIOL.
crease in histone H4 acetylation also depends on NPAT. Sup-
pression of TRRAP or Tip60 expression through RNA inter-
ference leads to the inhibition of histone gene transcriptional
activation. These results, together with the previous observa-
tions that the association of NPAT with histone promoters as
well as NPAT-mediated histone promoter activation is regu-
lated by cyclin E-Cdk2 phosphorylation (34, 53, 66), suggest
there is a mechanism underlying the coordinated transcription
of histone subtypes during the G1/S-phase transition (Fig. 8):
NPAT becomes phosphorylated by cyclin E-Cdk2 at the G1/S-
phase boundary, and this phosphorylation promotes the asso-
ciation of NPAT with histone promoters, likely through its
interactions with the subtype-specific factors, such as Oct1/
OCA-S and HiNF-P, which bind to specific DNA sequences
(e.g., SSREs) in the promoters of the different histone gene
subtypes. As a result, a TRRAP-containing HAT complex(s),
for example, the Tip60 HAT complex, is recruited to the pro-
moters of multiple histone genes. This HAT complex recruit-
ment in turn leads to histone acetylation and, subsequently,
transcriptional activation of multiple histone promoters.
It was previously shown that the phosphorylation of NPAT
by cyclin E-Cdk2 regulates its activity in histone gene activa-
tion (34, 66). It appears that cyclin E-Cdk2 modulates NPAT
function by regulating the localization of NPAT at histone
gene clusters (53). It is not clear whether cyclin E-Cdk2 regu-
lates NPAT through additional mechanisms. Our preliminary
results indicate that NPAT interacts with TRRAP throughout
the cell cycle, suggesting that the interaction of NPAT with
TRRAP may be independent of cyclin E-Cdk2 activity.
The results shown in Fig. 3 and 4 suggest that the majority of
TRRAP/Tip60 recruitment to histone gene promoters occurs
immediately prior to S-phase entry. It is possible that TRRAP/
Tip60 recruitment may involve a two-step mechanism in which
additional recruitment of these factors takes place in S phase,
as intermediate levels of association of NPAT and TRRAP
with histone gene promoters were observed in the presence of
aphidicolin. In addition to NPAT, the recruitment of the
TRRAP/Tip60 complex to histone gene promoters may also
involve other histone gene transcription factors.
According to the model proposed in Fig. 8, one might expect
the NPAT (the LFD-to-AAA) mutant to be dominant nega-
tive. Our results, however, indicate that this mutant apparently
has no inhibitory activity on histone promoter activation (Fig.
1E). The exact reason why the NPAT (AAA) mutant protein
fails to function as a dominant-negative mutant is not clear.
One possible explanation is that one or more of the proteins
that interact with the NPAT transactivation domain may be
involved in stabilizing the interaction of NPAT with histone
promoters. Without this stabilizing interaction, the presence of
the mutant at the promoter may be merely transient, thus
resulting in a failure of the NPAT (AAA) mutant to be dom-
In this study, we have identified a domain in NPAT that
possesses intrinsic transactivation potential. Interestingly, this
domain, referred to as the transactivation domain of NPAT,
contains a DLFD motif that is required for NPAT-mediated
transcriptional activation and is functionally conserved in E2F
and adenovirus E1A proteins. Our results clearly demonstrate
that the DLFD motif is crucial for the interaction of NPAT
with TRRAP, as well as for NPAT-mediated transcriptional
activation (Fig. 1 and 2). The DLFD motif also appears to be
crucial for the interaction of several E2F proteins with TRRAP
and for their transcriptional activation function. It was previ-
ously observed that deletion of the DLFD sequence in a trans-
activation domain of E2F1 (residues 389 to 422) fused to the
GAL4 DNA-binding domain resulted in an almost complete
loss of its transcriptional activation capability (13). Consistent
with this observation, the LFD-to-AAA mutation in the tran-
scriptional activation domain of E2F3 (residues 391 to 465)
results in the loss of transactivation when the mutant domain is
fused to the GAL4 DBD (our unpublished observation).
Moreover, the replacement of the LFD sequence with AAA in
the transactivation domain abolishes the interaction of E2F3
with TRRAP (our unpublished observation). It was reported
that the last seven amino acids of E2F4, which include the
second aspartic acid residue in the DLFD motif, are critical for
its interaction with TRRAP and E2F4-mediated reporter ac-
tivation (30). E1A may also utilize the DLFD motif to interact
with TRRAP. It was shown that the deletion of E1A from the
CR1 region, which includes the DLFD motif, abolishes both
TRRAP binding and transformation (7). Thus, the DLFD mo-
tif functions as a TRRAP-interacting module that is conserved
in NPAT, E2F, and E1A proteins. Several other TRRAP-
interacting proteins, such as c-Myc, p53, and BRCA1 (2, 37,
44), apparently lack the DLFD motif and therefore likely in-
teract with TRRAP through a different sequence motif(s). The
existence of multiple TRRAP-interacting motifs may allow the
recruitment of TRRAP-containing complexes by distinct fac-
tors to be differentially regulated. It is interesting to note that
the DLFD motif is also part of the sequences in E2F proteins
shown to interact with the retinoblastoma protein pRB (13, 21,
22, 32, 48). Hence, pRB may inhibit E2F function by prevent-
FIG. 8. A model for coordinated transcriptional activation of histone subtypes by cyclin E-Cdk2 substrate NPAT. SSRE, subtype-specific
regulator element; SSBP, proteins, such as Oct-1 and HiNF-P that directly bind SSRE elements within the promoters of a histone subtype; TFs,
transcription factors, P, phosphorylation. See text for details.
VOL. 28, 2008 NPAT RECRUITS TRRAP-Tip60 COMPLEX445
ing the association of E2F with TRRAP-containing HAT com-
TRRAP has been shown to be a component of a number of
HAT complexes, including the GCN5/PCAF and Tip60 com-
plexes (6, 8). We focused on the TRRAP-Tip60 HAT complex
in this study because we observed an interaction between the
NPAT transactivation domain and two other components of
the Tip60 HAT complex, Tip48 and Tip49, in our initial mass
spectrometric analysis (Fig. 2B). It is possible that, similar to
E2F and c-Myc, which recruit Tip60 as well as GCN5 com-
plexes to their target promoters, NPAT may interact with and
recruit additional TRRAP-containing HAT complexes to his-
tone promoters in vivo. Compared with the NPATflox/?cells
infected with Ad-LacZ, the Ad-Cre-infected NPATflox/?cells
showed only a moderate (30 to 55%) reduction in histone H4
acetylation at histone promoters at the G1/S-phase boundary.
This might be due to the fact that some residual NPAT protein
remains in these cells and can still recruit the TRRAP-Tip60
complex to the histone promoters (Fig. 2E, 3, and 4). Alter-
natively, other protein factors might also recruit a HAT com-
plex (or complexes) to the histone gene promoters to induce
histone acetylation in concert with, but independent of, NPAT.
Since proteins other than histones can also be the substrates of
HATs (17, 18), the NPAT-recruited HAT(s) may also play a
role in histone gene transcription by acetylating nonhistone
proteins at histone promoters.
In addition to components of the Tip60 complex (Fig. 2B),
NPAT appears to interact with YY1, BZAP45, and Hsp70,
which have been implicated in histone gene transcription (12,
31, 41, 61, 67). Although an in vivo interaction of NPAT with
these proteins remains to be determined, the observation
raises the possibility that these proteins may participate in
regulation of histone gene transcription through their cooper-
ation with NPAT. The transactivation domain of NPAT ap-
parently interacts with a number of additional proteins (Fig.
2B), which have not been shown to be involved in histone gene
transcription. Further studies are needed to determine their
interactions with NPAT in vivo, as well as their roles in tran-
scriptional activation of histone genes. Such studies may shed
new light on the coordinated regulation of histone gene tran-
We thank Hartmut Land and Dirk Bohmann for helpful discussions
and critical reading of the manuscript. We also thank Wade Harper for
providing HCT116 NPATflox/?cells, Yang Shi for providing the
pBS/U6 plasmid, Dennis McCance for providing the pBS/U6/Luc plas-
mid, Yin Sun for providing the pBS/U6/Ski plasmid, and Luojing Chen
for providing the pLKO.1 nonspecific control. We are grateful to Peter
Keng for help with FACS analysis, Anders Naar for suggestions on
protein purification, and Taplin Biological Spectrometry Facility for
mass spectrometric analysis.
This work was supported by NIH grant R01 GM65814 to J.Z.
1. Allard, S., R. T. Utley, J. Savard, A. Clarke, P. Grant, C. J. Brandl, L. Pillus,
J. L. Workman, and J. Cote. 1999. NuA4, an essential transcription adaptor/
histone H4 acetyltransferase complex containing Esa1p and the ATM-re-
lated cofactor Tra1p. EMBO J. 18:5108–5119.
2. Ard, P. G., C. Chatterjee, S. Kunjibettu, L. R. Adside, L. E. Gralinski, and
S. B. McMahon. 2002. Transcriptional regulation of the mdm2 oncogene by
p53 requires TRRAP acetyltransferase complexes. Mol. Cell. Biol. 22:5650–
3. Barcaroli, D., L. Bongiorno-Borbone, A. Terrinoni, T. G. Hofmann, M.
Rossi, R. A. Knight, A. G. Matera, G. Melino, and V. De Laurenzi. 2006.
FLASH is required for histone transcription and S-phase progression. Proc.
Natl. Acad. Sci. USA 103:14808–14812.
4. Brand, M., K. Yamamoto, A. Staub, and L. Tora. 1999. Identification of
TATA-binding protein-free TAFII-containing complex subunits suggests a
role in nucleosome acetylation and signal transduction. J. Biol. Chem. 274:
5. Brown, C. E., L. Howe, K. Sousa, S. C. Alley, M. J. Carrozza, S. Tan, and
J. L. Workman. 2001. Recruitment of HAT complexes by direct activator
interactions with the ATM-related Tra1 subunit. Science 292:2333–2337.
6. Carrozza, M. J., R. T. Utley, J. L. Workman, and J. Cote. 2003. The diverse
functions of histone acetyltransferase complexes. Trends Genet. 19:321–329.
7. Deleu, L., S. Shellard, K. Alevizopoulos, B. Amati, and H. Land. 2001.
Recruitment of TRRAP required for oncogenic transformation by E1A.
8. Doyon, Y., and J. Cote. 2004. The highly conserved and multifunctional
NuA4 HAT complex. Curr. Opin. Genet. Dev. 14:147–154.
9. Dyson, N. 1998. The regulation of E2F by pRB-family proteins. Genes Dev.
10. Dyson, N., P. Guida, C. McCall, and E. Harlow. 1992. Adenovirus E1A
makes two distinct contacts with the retinoblastoma protein. J. Virol. 66:
11. Dyson, N., and E. Harlow. 1992. Adenovirus E1A targets key regulators of
cell proliferation. Cancer Surv. 12:161–195.
12. Eliassen, K. A., A. Baldwin, E. M. Sikorski, and M. M. Hurt. 1998. Role for
a YY1-binding element in replication-dependent mouse histone gene ex-
pression. Mol. Cell. Biol. 18:7106–7118.
13. Flemington, E. K., S. H. Speck, and W. G. Kaelin, Jr. 1993. E2F-1-mediated
transactivation is inhibited by complex formation with the retinoblastoma
susceptibility gene product. Proc. Natl. Acad. Sci. USA 90:6914–6918.
14. Fletcher, C., N. Heintz, and R. G. Roeder. 1987. Purification and character-
ization of OTF-1, a transcription factor regulating cell cycle expression of a
human histone H2b gene. Cell 51:773–781.
15. Frank, S. R., T. Parisi, S. Taubert, P. Fernandez, M. Fuchs, H. M. Chan,
D. M. Livingston, and B. Amati. 2003. MYC recruits the TIP60 histone
acetyltransferase complex to chromatin. EMBO Rep. 4:575–580.
16. Gao, G., A. P. Bracken, K. Burkard, D. Pasini, M. Classon, C. Attwooll, M.
Sagara, T. Imai, K. Helin, and J. Zhao. 2003. NPAT expression is regulated
by E2F and is essential for cell cycle progression. Mol. Cell. Biol. 23:2821–
17. Glozak, M. A., N. Sengupta, X. Zhang, and E. Seto. 2005. Acetylation and
deacetylation of non-histone proteins. Gene 363:15–23.
18. Gu, W., and R. G. Roeder. 1997. Activation of p53 sequence-specific DNA
binding by acetylation of the p53 C-terminal domain. Cell 90:595–606.
19. Hall, C., D. M. Nelson, X. Ye, K. Baker, J. A. DeCaprio, S. Seeholzer, M.
Lipinski, and P. D. Adams. 2001. HIRA, the human homologue of yeast
Hir1p and Hir2p, is a novel cyclin-cdk2 substrate whose expression blocks
S-phase progression. Mol. Cell. Biol. 21:1854–1865.
20. Heintz, N. 1991. The regulation of histone gene expression during the cell
cycle. Biochim. Biophys. Acta 1088:327–339.
21. Helin, K., E. Harlow, and A. Fattaey. 1993. Inhibition of E2F-1 transactiva-
tion by direct binding of the retinoblastoma protein. Mol. Cell. Biol. 13:
22. Helin, K., J. A. Lees, M. Vidal, N. Dyson, E. Harlow, and A. Fattaey. 1992.
A cDNA encoding a pRB-binding protein with properties of the transcrip-
tion factor E2F. Cell 70:337–350.
23. Herceg, Z., W. Hulla, D. Gell, C. Cuenin, M. Lleonart, S. Jackson, and Z. Q.
Wang. 2001. Disruption of Trrap causes early embryonic lethality and defects
in cell cycle progression. Nat. Genet. 29:206–211.
24. Ikura, T., V. V. Ogryzko, M. Grigoriev, R. Groisman, J. Wang, M. Horikoshi,
R. Scully, J. Qin, and Y. Nakatani. 2000. Involvement of the TIP60 histone
acetylase complex in DNA repair and apoptosis. Cell 102:463–473.
25. Jenuwein, T., and C. D. Allis. 2001. Translating the histone code. Science
26. Kaelin, W. G., Jr., W. Krek, W. R. Sellers, J. A. DeCaprio, F. Ajchenbaum,
C. S. Fuchs, T. Chittenden, Y. Li, P. J. Farnham, M. A. Blanar, et al. 1992.
Expression cloning of a cDNA encoding a retinoblastoma-binding protein
with E2F-like properties. Cell 70:351–364.
27. Khorasanizadeh, S. 2004. The nucleosome: from genomic organization to
genomic regulation. Cell 116:259–272.
28. Kornberg, R. D., and Y. Lorch. 1999. Twenty-five years of the nucleosome,
fundamental particle of the eukaryote chromosome. Cell 98:285–294.
29. Lang, S. E., and P. Hearing. 2003. The adenovirus E1A oncoprotein recruits
the cellular TRRAP/GCN5 histone acetyltransferase complex. Oncogene
30. Lang, S. E., S. B. McMahon, M. D. Cole, and P. Hearing. 2001. E2F tran-
scriptional activation requires TRRAP and GCN5 cofactors. J. Biol. Chem.
31. Last, T. J., A. J. van Wijnen, M. J. Birnbaum, G. S. Stein, and J. L. Stein.
1999. Multiple interactions of the transcription factor YY1 with human
histone H4 gene regulatory elements. J. Cell. Biochem. 72:507–516.
32. Lee, C., J. H. Chang, H. S. Lee, and Y. Cho. 2002. Structural basis for the
446DERAN ET AL.MOL. CELL. BIOL.
recognition of the E2F transactivation domain by the retinoblastoma tumor Download full-text
suppressor. Genes Dev. 16:3199–3212.
33. Livak, K. J., and T. D. Schmittgen. 2001. Analysis of relative gene expression
data using real-time quantitative PCR and the 2(-delta delta C(T)) method.
34. Ma, T., B. A. Van Tine, Y. Wei, M. D. Garrett, D. Nelson, P. D. Adams, J.
Wang, J. Qin, L. T. Chow, and J. W. Harper. 2000. Cell cycle-regulated
phosphorylation of p220(NPAT) by cyclin E/Cdk2 in Cajal bodies promotes
histone gene transcription. Genes Dev. 14:2298–2313.
35. Martinez, E., V. B. Palhan, A. Tjernberg, E. S. Lymar, A. M. Gamper, T. K.
Kundu, B. T. Chait, and R. G. Roeder. 2001. Human STAGA complex is a
chromatin-acetylating transcription coactivator that interacts with pre-
mRNA splicing and DNA damage-binding factors in vivo. Mol. Cell. Biol.
36. Marzluff, W. F., and R. J. Duronio. 2002. Histone mRNA expression: mul-
tiple levels of cell cycle regulation and important developmental conse-
quences. Curr. Opin. Cell Biol. 14:692–699.
37. McMahon, S. B., H. A. Van Buskirk, K. A. Dugan, T. D. Copeland, and M. D.
Cole. 1998. The novel ATM-related protein TRRAP is an essential cofactor
for the c-Myc and E2F oncoproteins. Cell 94:363–374.
38. McMahon, S. B., M. A. Wood, and M. D. Cole. 2000. The essential cofactor
TRRAP recruits the histone acetyltransferase hGCN5 to c-Myc. Mol. Cell.
39. Meeks-Wagner, D., and L. H. Hartwell. 1986. Normal stoichiometry of his-
tone dimer sets is necessary for high fidelity of mitotic chromosome trans-
mission. Cell 44:43–52.
40. Miele, A., C. D. Braastad, W. F. Holmes, P. Mitra, R. Medina, R. Xie, S. K.
Zaidi, X. Ye, Y. Wei, J. W. Harper, A. J. van Wijnen, J. L. Stein, and G. S.
Stein. 2005. HiNF-P directly links the cyclin E/CDK2/p220NPATpathway to
histone H4 gene regulation at the G1/S phase cell cycle transition. Mol. Cell.
41. Mitra, P., P. S. Vaughan, J. L. Stein, G. S. Stein, and A. J. van Wijnen. 2001.
Purification and functional analysis of a novel leucine-zipper/nucleotide-fold
protein, BZAP45, stimulating cell cycle regulated histone H4 gene transcrip-
tion. Biochemistry 40:10693–10699.
42. Murr, R., J. I. Loizou, Y. G. Yang, C. Cuenin, H. Li, Z. Q. Wang, and Z.
Herceg. 2006. Histone acetylation by Trrap-Tip60 modulates loading of
repair proteins and repair of DNA double-strand breaks. Nat. Cell Biol.
43. Naar, A. M., P. A. Beaurang, S. Zhou, S. Abraham, W. Solomon, and R.
Tjian. 1999. Composite co-activator ARC mediates chromatin-directed tran-
scriptional activation. Nature 398:828–832.
44. Oishi, H., H. Kitagawa, O. Wada, S. Takezawa, L. Tora, M. Kouzu-Fujita, I.
Takada, T. Yano, J. Yanagisawa, and S. Kato. 2006. An hGCN5/TRRAP
histone acetyltransferase complex co-activates BRCA1 transactivation func-
tion through histone modification. J. Biol. Chem. 281:20–26.
45. Osley, M. A. 1991. The regulation of histone synthesis in the cell cycle. Annu.
Rev. Biochem. 60:827–861.
46. Park, J., S. Kunjibettu, S. B. McMahon, and M. D. Cole. 2001. The ATM-
related domain of TRRAP is required for histone acetyltransferase recruit-
ment and Myc-dependent oncogenesis. Genes Dev. 15:1619–1624.
47. Robert, F., S. Hardy, Z. Nagy, C. Baldeyron, R. Murr, U. Dery, J. Y. Masson,
D. Papadopoulo, Z. Herceg, and L. Tora. 2006. The transcriptional histone
acetyltransferase cofactor TRRAP associates with the MRN repair complex
and plays a role in DNA double-strand break repair. Mol. Cell. Biol. 26:
48. Shan, B., T. Durfee, and W. H. Lee. 1996. Disruption of RB/E2F-1 interac-
tion by single point mutations in E2F-1 enhances S-phase entry and apop-
tosis. Proc. Natl. Acad. Sci. USA 93:679–684.
49. Shan, B., X. Zhu, P. L. Chen, T. Durfee, Y. Yang, D. Sharp, and W. H. Lee.
1992. Molecular cloning of cellular genes encoding retinoblastoma-associ-
ated proteins: identification of a gene with properties of the transcription
factor E2F. Mol. Cell. Biol. 12:5620–5631.
50. Squatrito, M., C. Gorrini, and B. Amati. 2006. Tip60 in DNA damage
response and growth control: many tricks in one HAT. Trends Cell Biol.
51. Stein, G. S., J. L. Stein, A. J. Van Wijnen, and J. B. Lian. 1996. Transcrip-
tional control of cell cycle progression: the histone gene is a paradigm for the
G1/S phase and proliferation/differentiation transitions. Cell Biol. Int. 20:
52. Sterner, D. E., and S. L. Berger. 2000. Acetylation of histones and transcrip-
tion-related factors. Microbiol. Mol. Biol. Rev. 64:435–459.
53. Su, C., G. Gao, S. Schneider, C. Helt, C. Weiss, M. A. O’Reilly, D. Bohmann,
and J. Zhao. 2004. DNA damage induces downregulation of histone gene
expression through the G1 checkpoint pathway. EMBO J. 23:1133–1143.
54. Sui, G., C. Soohoo, B. el Affar, F. Gay, Y. Shi, W. C. Forrester, and Y. Shi.
2002. A DNA vector-based RNAi technology to suppress gene expression in
mammalian cells. Proc. Natl. Acad. Sci. USA 99:5515–5520.
55. Sullivan, E., C. Santiago, E. D. Parker, Z. Dominski, X. Yang, D. J. Lanzotti,
T. C. Ingledue, W. F. Marzluff, and R. J. Duronio. 2001. Drosophila stem
loop binding protein coordinates accumulation of mature histone mRNA
with cell cycle progression. Genes Dev. 15:173–187.
56. Taubert, S., C. Gorrini, S. R. Frank, T. Parisi, M. Fuchs, H. M. Chan, D. M.
Livingston, and B. Amati. 2004. E2F-dependent histone acetylation and
recruitment of the Tip60 acetyltransferase complex to chromatin in late G1.
Mol. Cell. Biol. 24:4546–4556.
57. Thrash, B. R., C. W. Menges, R. H. Pierce, and D. J. McCance. 2006. AKT1
provides an essential survival signal required for differentiation and stratifi-
cation of primary human keratinocytes. J. Biol. Chem. 281:12155–12162.
58. Vassilev, A., J. Yamauchi, T. Kotani, C. Prives, M. L. Avantaggiati, J. Qin,
and Y. Nakatani. 1998. The 400 kDa subunit of the PCAF histone acetylase
complex belongs to the ATM superfamily. Mol. Cell. 2:869–875.
59. Wang, Y., W. Fischle, W. Cheung, S. Jacobs, S. Khorasanizadeh, and C. D.
Allis. 2004. Beyond the double helix: writing and reading the histone code.
Novartis Found. Symp. 259:3–17. (Discussion, 17–21 and 163–169.)
60. Wei, Y., J. Jin, and J. W. Harper. 2003. The cyclin E/Cdk2 substrate and
Cajal body component p220NPATactivates histone transcription through a
novel LisH-like domain. Mol. Cell. Biol. 23:3669–3680.
61. Wu, F., and A. S. Lee. 2001. YY1 as a regulator of replication-dependent
hamster histone H3.2 promoter and an interactive partner of AP-2. J. Biol.
62. Ye, X., A. A. Franco, H. Santos, D. M. Nelson, P. D. Kaufman, and P. D.
Adams. 2003. Defective S phase chromatin assembly causes DNA damage,
activation of the S phase checkpoint, and S phase arrest. Mol. Cell. 11:341–
63. Ye, X., Y. Wei, G. Nalepa, and J. W. Harper. 2003. The cyclin E/Cdk2
substrate p220NPATis required for S-phase entry, histone gene expression,
and Cajal body maintenance in human somatic cells. Mol. Cell. Biol. 23:
64. Zhao, J. 2004. Coordination of DNA synthesis and histone gene expression
during normal cell cycle progression and after DNA damage. Cell Cycle
65. Zhao, J., B. Dynlacht, T. Imai, T. Hori, and E. Harlow. 1998. Expression of
NPAT, a novel substrate of cyclin E-CDK2, promotes S-phase entry. Genes
66. Zhao, J., B. K. Kennedy, B. D. Lawrence, D. A. Barbie, A. G. Matera, J. A.
Fletcher, and E. Harlow. 2000. NPAT links cyclin E-Cdk2 to the regulation
of replication-dependent histone gene transcription. Genes Dev. 14:2283–
67. Zheng, L., R. G. Roeder, and Y. Luo. 2003. S phase activation of the histone
H2B promoter by OCA-S, a coactivator complex that contains GAPDH as a
key component. Cell 114:255–266.
VOL. 28, 2008 NPAT RECRUITS TRRAP-Tip60 COMPLEX447