MOLECULAR AND CELLULAR BIOLOGY, Nov. 2006, p. 7913–7928
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Vol. 26, No. 21
Negative and Positive Regulation of Gene Expression by Mouse
Histone Deacetylase 1?
Gordin Zupkovitz,1Julia Tischler,1† Markus Posch,2‡ Iwona Sadzak,1§ Katrin Ramsauer,3
Gerda Egger,1¶ Reinhard Grausenburger,1Norbert Schweifer,2Susanna Chiocca,4
Thomas Decker,3and Christian Seiser1*
Max F. Perutz Laboratories, Department of Medical Biochemistry, Medical University of Vienna, Vienna Biocenter, A-1030 Vienna,
Austria1; Boehringer Ingelheim Austria, A-1121 Vienna, Austria2; Max F. Perutz Laboratories, Institute of Microbiology and
Genetics, University of Vienna, Vienna Biocenter, A-1030 Vienna, Austria3; and European Institute of Oncology,
Department of Experimental Oncology, 20141 Milan, Italy4
Received 6 July 2006/Returned for modification 17 July 2006/Accepted 21 August 2006
Histone deacetylases (HDACs) catalyze the removal of acetyl groups from core histones. Because of their
capacity to induce local condensation of chromatin, HDACs are generally considered repressors of transcrip-
tion. In this report, we analyzed the role of the class I histone deacetylase HDAC1 as a transcriptional regulator
by comparing the expression profiles of wild-type and HDAC1-deficient embryonic stem cells. A specific subset
of mouse genes (7%) was deregulated in the absence of HDAC1. We identified several putative tumor sup-
pressors (JunB, Prss11, and Plagl1) and imprinted genes (Igf2, H19, and p57) as novel HDAC1 targets. The
majority of HDAC1 target genes showed reduced expression accompanied by recruitment of HDAC1 and local
reduction in histone acetylation at regulatory regions. At some target genes, the related deacetylase HDAC2
partially masks the loss of HDAC1. A second group of genes was found to be downregulated in HDAC1-
deficient cells, predominantly by additional recruitment of HDAC2 in the absence of HDAC1. Finally, a small
set of genes (Gja1, Irf1, and Gbp2) was found to require HDAC activity and recruitment of HDAC1 for their
transcriptional activation. Our study reveals a regulatory cross talk between HDAC1 and HDAC2 and a novel
function for HDAC1 as a transcriptional coactivator.
The DNA of eukaryotic cells is compacted by basic histone
proteins in a highly organized structure called chromatin. The
nucleosome, the basic unit of chromatin, consists of 147 base
pairs of DNA wrapped around the histone octamer, composed
of two copies of each of the four core histones, H2A, H2B, H3,
and H4 (78). Although the structure of the core nucleosome is
well defined, the basic N-terminal histone tails protrude from
the core nucleosome and show no defined structure (38, 39).
These histone tail domains are subject to posttranslational
modifications, such as acetylation, methylation, phosphoryla-
tion, and ubiquitination (recently reviewed in reference 41).
These modifications affect various biological processes, includ-
ing the transcription of chromatin-embedded genes. Recent
observations indicate that histone modifications occur interde-
pendently and create a pattern that might modulate the affinity
of histone-binding proteins. These findings are the basis of the
histone code hypothesis (21, 35, 67, 75). An alternative way of
explaining the cooperation of multiple histone modifications is
the recently proposed chromatin signaling network model (64).
A correlation between histone acetylation and increased
gene expression was discovered earlier on (3). According to the
current model, the acetylation of lysine residues within the
histone tails neutralizes the positive charge of ε-amino groups
and thereby reduces the interaction between the N-terminal
tails of histones and the negatively charged DNA. Acetylation
at the N termini of core histones is therefore believed to induce
the local opening of chromatin structures. In addition, acety-
lated histone tails are specifically recognized and bound by
bromodomain-containing proteins, such as components of the
basal transcription machinery or histone acetyltransferases
(HATs) (85). Reversible histone acetylation is controlled by
histone acetyltransferases, which usually act as transcriptional
coactivators, and histone deacetylases (HDACs), which re-
press transcription. Activator complexes containing HAT ac-
tivity have been shown to contribute to transcriptional activa-
tion by recruitment of general transcription factors and RNA
polymerase II (7, 74). In contrast, recruitment of repressor
complexes with HDAC activity is considered to lead to
deacetylation of histones, stabilization of nucleosome struc-
ture, and formation of a repressive chromatin state.
During the last decade, more than a dozen histone deacety-
lases have been identified in mammalian cells. Based on se-
quence similarities, HDACs are divided into four functional
classes: class I (HDAC1, HDAC2, HDAC3, and HDAC8),
class II (HDAC4, HDAC5, HDAC6, HDAC7, HDAC9, and
* Corresponding author. Mailing address: Max F. Perutz Laborato-
ries, Department of Medical Biochemistry, Medical University of
Vienna, Vienna Biocenter, Dr. Bohr-Gasse 9/2, A-1030 Vienna, Austria.
Phone: 431 4277 61770. Fax: 431 4277 9617. E-mail: christian.seiser
† Present address: The Wellcome Trust Sanger Institute, Hinxton,
Cambridge CB10 1SA, United Kingdom.
‡ Present address: Wellcome Trust Biocentre, University of Dundee,
Dundee DD1 5EH, United Kingdom.
§ Present address: Max F. Perutz Laboratories, Institute of Micro-
biology and Genetics, University of Vienna, Vienna Biocenter, A-1030
¶ Present address: Department of Biochemistry and Molecular Bi-
ology, USC/Norris Comprehensive Cancer Center, Keck School of
Medicine of the University of Southern California, Los Angeles, CA
?Published ahead of print on 28 August 2006.
HDAC10), class III (SIRT1 to SIRT7), and the recently de-
scribed class IV of HDACs, which consists of HDAC11-related
enzymes (28, 29). The class I enzyme HDAC1 belongs to an
ancient family of highly conserved enzymes and was the first
protein shown to have histone deacetylating activity in mam-
mals (reviewed in reference 46). Human HDAC1 was purified
and cloned by an affinity purification approach (73) and was
shown to share significant homology with the previously iden-
tified Saccharomyces cerevisiae transcriptional regulator Rpd3/
Sdi2/Sds6 (51, 80–82). In mouse cells, expression of the
HDAC1 gene is stimulated by growth factors (5) and con-
trolled by its own product in a negative feedback loop (32, 65).
The enzyme plays an important role in various biological pro-
cesses, such as cell cycle progression, cell proliferation, and
differentiation (46). The HDAC1 mouse knockout (KO) has
also revealed the essential function of this deacetylase in em-
bryonic development (36). HDAC1 is a nuclear protein and
can heterodimerize with the closely related deacetylase
HDAC2 (31, 72). Both enzymes are found in three major
multiprotein complexes, named Sin3, NuRD, and CoREST (2,
29). HDAC1 can repress gene transcription either directly or
as part of these multiprotein complexes when recruited by a
variety of transcriptional regulators, including SP1/SP3, YY1,
unliganded nuclear receptors, the pocket proteins pRB, p107,
and p130, and the tumor suppressor p53 (13, 52).
HDAC inhibitors have been shown to induce cell cycle ar-
rest, differentiation, or apoptosis in tumor cells, and some of
these compounds are currently tested as antitumor drugs in
clinical trials (19, 20, 44). These inhibitors affect the catalytic
activity of most class I and class II deacetylases. However, little
is known about the individual roles of mammalian deacetylases
in transcriptional control and the relevant target enzymes for
HDAC inhibitors as antitumor drugs. To unravel the role
of HDAC1 as a transcriptional regulator, we identified puta-
tive HDAC1 target genes by comparing the gene expression
profiles of wild-type and HDAC1-deficient embryonic stem
(ES) cells. A restricted subset of mouse genes involved in
biological processes, such as growth control, cell communica-
tion, and transcriptional regulation, was found to be reversibly
deregulated upon the loss of HDAC1. By chromatin immuno-
precipitation (ChIP) assays, we observed the presence of the
HDAC1 enzyme associated with a reduction in histone acety-
lation levels at the regulatory regions, thereby providing com-
pelling evidence for a direct regulation of these genes by
HDAC1. In addition, our data revealed a delicate balance
between the class I enzymes HDAC1 and HDAC2 in the
regulation of common target genes. Finally, we also identified
a novel function for HDAC1 as a positive regulator of gene
MATERIALS AND METHODS
Cell culture. HDAC1 wild-type and homozygous mutant ES cells (36) were
cultivated in M15 medium supplemented with antibiotics and 15% (vol/vol) fetal
calf serum and either supplemented with 103U/ml of leukemia inhibitory factor
on gelatinized culture dishes or without leukemia inhibitory factor on feeder cell
layers. All ES cell experiments were performed with cell lines obtained from
littermates (36). To stably express wild-type HDAC1 and the myc-tagged, enzy-
matically inactive HDAC1-H140/141A mutant in ES cell lines, the ES cell-
specific expression vectors pMSCVpuro-HDAC1 and pMSCVpuro-mut were
linearized and electroporated into ES cells by using a Bio-Rad Gene Pulser II
with 0.4-cm-wide sterile cuvettes (165-2088; Bio-Rad), with a single pulse at 230
V and 500 ?F. Electroporated ES cells were incubated for 5 min at room
temperature and then plated onto puromycin-resistant feeder cells. After 24 h,
puromycin (5 ?g/ml) was applied to select for targeted clones. The selection
procedure was continued for 14 days, and single clones were picked, expanded,
and analyzed for expression of HDAC1. The human osteosarcoma cell line
U2OS was kept in Dulbecco’s modified Eagle’s medium supplemented with 10%
(vol/vol) fetal calf serum and antibiotics. U2OS cells were stably transfected with
pcDNA3-Prss11 (56) (kindly provided by M. Kawaichi, Takayama) or pcDNA3
empty vector by using Lipofectamine 2000 (Invitrogen) according to the manu-
facturer’s instructions. Transfected cells were split after 24 h and grown in the
presence of 360 ?g/ml of G418 for selection. After 10 days, single clones were
picked and analyzed for Prss11 expression by real-time reverse transcription
(RT)-PCR. Trichostatin A (TSA) was obtained from Wako Pure Chemical
Affymetrix analysis. Total RNA (approximately 5 ?g) was used to synthesize
double-stranded cDNA by using a custom Superscript double-stranded cDNA
synthesis kit (Invitrogen, Karlsruhe, Germany). Biotin-labeled cRNA was then
prepared from this template by using an Enzo BioArray high-yield RNA tran-
script labeling kit (Affymetrix, High Wycombe, United Kingdom), and unincor-
porated nucleotides were removed using RNeasy columns (QIAGEN, Hilden,
Germany). Hybridization, washing, and fluorescence staining of the Affymetrix
GeneChip murine genome MG-U74Av2 array (Affymetrix Inc., Santa Clara,
CA) were carried out according to the manufacturer’s instructions (GeneChip
Expression Analysis Technical Manual; Affymetrix). All experiments were per-
formed in triplicate with independently extracted RNAs. Data analysis was
performed by means of a comparison matrix, with control (wild-type) experi-
ments as a background, for generation of expression signal log ratios with basis
2 and subsequently changes (n-fold) between wild-type and knockout samples,
using Microarray Suite software version 5.0 (Affymetrix). Next, mean changes
(n-fold) and relative standard deviations between experiments were calculated
and genes with changes (n-fold) of ?2 or ?0.5 and relative standard deviations
of ?0.6 were selected for further analysis. The association of genes with partic-
ular functions, pathways, and diseases was analyzed through the use of Ingenuity
Pathways Analysis (Ingenuity Systems). Gene networks are ranked according to
their scores. Calculations for network scores are based on the hypergeometric
distribution calculated via the computationally efficient Fisher exact test for
two-by-two contingency tables. The significance value associated with functions
and pathways is a measure of how likely it is that genes from the data set file
participate in that function. The significance is expressed as a P value, which is
calculated using the right-tailed Fisher exact test.
Protein analysis and antibodies. Whole-cell protein extraction, histone isola-
tion, and Western blot analysis were performed as previously described (5, 36,
59). The following antibodies were used for protein detection on immunoblots
and for chromatin immunoprecipitation assays: HDAC1 (polyclonal rabbit an-
tibody and monoclonal mouse antibody), HDAC2 (polyclonal rabbit antibody
and monoclonal mouse antibody), acetyl histone-H3, acetyl histone-H4, acetyl
K9-H3 from Upstate, and the C terminus of histone H3 (Abcam). Polyclonal
trimethyl K9-H3 and trimethyl K27-H3 antibodies were kind gifts from T. Jenu-
wein (59). The ?-actin protein was visualized with a monoclonal antibody (AC-
74; Sigma). The proliferation marker Ki67 was detected with the monoclonal
Ki67 antigen antibody (Novo Castra) by indirect immunofluorescence micros-
copy (Zeiss Axiovert 135TV microscope) as previously described (72). Nuclear
DNA was visualized with 4?,6-diamidino-2-phenylindole (DAPI).
Immunoprecipitation assays. Immunoprecipitation assays were performed as
previously described (18). For combined analysis of proteins and associated
deacetylase activity, the precipitated material was resuspended in 50 ?l of ex-
traction buffer (20 mM Tris-HCl [pH 8.0], 100 mM NaCl, 1 mM EDTA, 0.5%
Nonidet P-40, 1 mM phenylmethylsulfonyl fluoride, 2 mM dithiothreitol, Roche
Diagnostics complete protease inhibitor cocktail), and 30-?l aliquots were ex-
amined for protein expression on Western blots. The remaining 20-?l aliquot
was assayed for HDAC activity. Histone deacetylase activity assays were done as
described previously (5, 37).
Chromatin immunoprecipitation assays. Chromatin immunoprecipitation as-
says were carried out as described previously (8, 11, 65) with some modifications.
Chromatin was cross-linked for 10 min by using formaldehyde and then soni-
cated. Equal amounts of sonicated chromatin were diluted 10-fold and precipi-
tated overnight with the following antibodies: HDAC1, HDAC2, acetyl histone-
H3, acetyl histone-H4, acetyl K9-H3, trimethyl K9-H3, trimethyl K27-H3, and
preimmune serum as a control. The chromatin-antibody complexes were isolated
by incubation with 30-?l protein A-Sepharose beads (50% slurry, 100 ?g/ml
salmon sperm DNA, 500 ?g/ml bovine serum albumin) while rocking at 4°C for
2 hours. The beads were harvested and washed as described previously (65).
Chromatin-antibody complexes were eluted from the protein A-Sepharose beads
7914 ZUPKOVITZ ET AL.MOL. CELL. BIOL.
by addition of 2% sodium dodecyl sulfate, 0.1 M NaHCO3, and 10 mM dithio-
threitol. Cross-linking was reversed by addition of a 0.05 volume of 4 M NaCl and
incubation of the eluted samples for 6 h at 65°C. The DNA was extracted with
phenol-chloroform, precipitated with ethanol, and dissolved in water.
PCR analysis of immunoprecipitated DNA. All PCRs were performed on an
iCycler (Bio-Rad) by using Promega PCR Master Mix. The linear range for each
primer pair was determined empirically using different amounts of genomic
DNA. PCRs with 1:40 dilutions of genomic DNA (input) were carried out along
with the immunoprecipitated DNA. PCR products were resolved on 2% aga-
rose–Tris-acetate-EDTA gels. Primer sequences are available upon request.
RNA isolation, Northern blotting, and real-time RT-PCR analysis. Total cel-
lular RNA was isolated with TRIzol reagent (GibcoBRL) as recommended by
the manufacturer. Northern blot hybridization was performed by the sandwich
method as previously described (68). For cDNA, 1 ?g of total RNA was reverse
transcribed with an iScript cDNA synthesis kit (Bio-Rad). Real-time RT-PCRs
were performed with 0.5 ?l of the RT reaction mixture by the iCycler iQ system
(Bio-Rad), using SYBR green (Molecular probes) for labeling. Primer sequences
are available upon request.
Microarray data accession number. The microarray data have been deposited
in the Gene Expression Omnibus (GEO) public database (http://www.ncbi.nlm
.nih.gov/geo/) under the accession number GSE 5583.
HDAC1 regulates a specific subset of genes in mouse ES
cells. To understand the role of HDAC1 at the cellular level,
we previously generated wild-type and HDAC1-deficient
mouse ES cells by blastocyst outgrowth experiments with lit-
termates (36). The total cellular HDAC activity in ES cells is
significantly reduced upon the loss of HDAC1, indicating that
this enzyme is one of the major deacetylases in these cells.
Given the important role of HDAC1 as a chromatin-modifying
enzyme, we wanted to determine on a genome-wide basis
whether the loss of HDAC1 could affect gene expression in
mouse cells. We analyzed total RNA from wild-type and
HDAC1 KO ES cells on MGU74Av2 Affymetrix GeneChip
microarrays, representing about 10,000 genes with known func-
tions and expressed sequencing tags. Experiments were per-
formed in triplicate with three independently isolated sets of
RNA. Significantly differentially regulated genes were identi-
fied by comparative analysis using Microarray Suite software
version 5.0 (Affymetrix) as described in Materials and Meth-
ods. The criterion for the selection of potential target genes
was a more-than-twofold difference in expression, based on the
mean value for all three independent experiments. Interest-
ingly, only approximately 7% of all genes present on the array
were deregulated in the absence of HDAC1, suggesting that
HDAC1 controls the transcription of a specific subset of mam-
malian genes (Fig. 1A). Moreover, of the more than 600 de-
regulated genes, nearly two-thirds showed increased expres-
sion levels in HDAC1 KO cells. This finding supports the
generally accepted idea that HDAC1 acts mainly as a tran-
scriptional repressor. However, a significant proportion of
genes showed a more-than-twofold decrease in expression in
HDAC1-deficient cells, suggesting a potential role for HDAC1
in the activation of these genes.
Potential HDAC1 target genes were analyzed for networks
and molecular functions through the use of Ingenuity Pathways
Analysis (Ingenuity Systems). In agreement with a proposed
role for HDAC1 as a regulator of many biological processes,
HDAC1 targets were identified as components of molecular
networks for cell-to-cell signaling, cellular movement, cell
death, gene expression, cell morphology, and cancer (Fig. 1B
and C). Classification according to physiological system devel-
opment and function revealed a potential role for HDAC1
target genes in the development of the hematological system,
skeletal and muscle systems, and immune and nervous systems
and in corresponding diseases (data not shown).
Given the growth-inhibitory effects of HDAC inhibitors, the
impaired proliferation of HDAC1 KO cells points towards
HDAC1 as one of the possible targets for these tumor drugs
(36). In this context, it is remarkable that several tumor-related
genes, such as those encoding JunB, Prss11, Plagl1, Apc2,
metallothionein 1, and metallothionein 2, were found in our
screen (see also Discussion). In addition, a relatively high num-
ber of imprinted genes were found to be either upregulated
(the Peg3, H19, and Plagl1 genes) or downregulated (the Igf2
and p57 genes) in this screen. Of the 35 imprinted genes
present on the array, which represent about 55% of all known
imprinted genes in the mouse (16, 49), 10 were deregulated in
the absence of HDAC1 (data not shown). Most of these genes
have been implicated in proliferation and growth control. In-
triguingly, chromosome 7, which contains several clusters of
imprinted genes, showed a significant deviation from the ex-
pected chromosomal distribution of deregulated genes, with
25% more deregulated genes than expected (Fig. 1D). In gen-
eral, the chromosomal distribution of deregulated genes (i.e.,
the number of genes found to be deregulated on a specific
chromosome relative to the total number of deregulated
genes) was similar to the chromosomal distribution of genes
present on the array and the chromosomal distribution of
genes found in the whole mouse genome (Fig. 1D).
The deregulation of HDAC1 target genes is reversible. As a
first step in the analysis of putative HDAC1 target genes, we
wanted to validate the results from the DNA array screen. To
this end, we isolated total RNA from wild-type and HDAC1
KO ES cells and analyzed a subset of randomly picked poten-
tial target genes by real-time RT-PCR and Northern blot anal-
yses. Of the 33 deregulated genes examined, 16 upregulated and
12 downregulated genes were successfully confirmed (Fig. 2).
Three genes (the Apobec2, Gata4, and Gjb3 genes) did not
show the expected result, while two genes (the Rb1 and Rfx2
genes) were upregulated below the defined threshold level.
Next, we stably transfected HDAC1 KO cells with a retro-
virus-derived ES cell expression vector encoding HDAC1, or
with the empty expression vector as a control, to test whether
the deregulation of putative target genes could be reverted.
Among several clones originating from HDAC1-deficient cells
transfected with the HDAC1 encoding vector, two cell lines
(KO-reA and KO-reB), which expressed HDAC1 protein al-
most at wild-type levels, were chosen for further analyses.
HDAC1-null cells transfected with the empty vector showed
no changes in HDAC1 and HDAC2 expression or in total
HDAC activity compared to untransfected cells (Fig. 3A and
data not shown). We have previously shown that the expression
of the closely related deacetylase HDAC2 is increased upon
deletion of HDAC1 (36) (Fig. 3A). Interestingly, the reintro-
duction of HDAC1 into KO cells led to a significant reduction
in HDAC2 expression and completely restored total histone
deacetylase activity (Fig. 3A). To analyze HDAC2-associated
HDAC1 protein levels, we immunoprecipitated HDAC2 from
extracts of wild-type, HDAC1 KO, and reintroduced cell lines
(KO-reA and KO-reB) and visualized precipitated HDAC1
and HDAC2 on immunoblots. As shown in Fig. 3B, the
VOL. 26, 2006 TRANSCRIPTIONAL REGULATION BY HDAC17915
FIG. 1. DNA array analysis of wild-type versus HDAC1-deficient ES cell lines. (A) Summary of DNA array results. Total numbers of genes
found on Affymetrix GeneChip Mu74 microarray analysis are presented as nonregulated (black) and deregulated (light gray) genes. The dark gray
7916 ZUPKOVITZ ET AL.MOL. CELL. BIOL.
amounts of HDAC2-associated HDAC1 protein in wild-type,
KO-reA, and KO-reB cells were comparable. Similarly, the
increased HDAC2-associated deacetylase activities in KO cells
were reduced to wild-type-cell levels upon reintroduction of
HDAC1. Taken together, these data show that the increase in
HDAC2 expression levels and associated activity and the re-
duction in total deacetylase activity observed in HDAC1 KO
cells are reversible.
We compared the mRNA expression levels of potential tar-
get genes in reintroduced KO ES cell lines with the respective
wild-type and KO controls by real-time RT-PCR and Northern
blot analyses (Fig. 3C and D). The expression levels of Prss11,
p21, Mt1, Mt2, Ass1, and JunB, which was found to be up-
regulated in KO ES cells, showed significant decreases upon
HDAC1 reintroduction in two independent cell lines (Fig. 3C).
The only exception was the Apc2 gene, whose expression was
threefold induced in KO cells (Fig. 2) but did not significantly
respond to reexpression of HDAC1 (data not shown). At the
moment, the reason for this observation is unclear. However,
the Apc2 gene was the only gene in this set of HDAC1 targets
that misses a CpG island within its promoter region.
Furthermore, mRNA levels of Igf2 and Dnmt3a, which were
downregulated in KO cells, were restored almost to wild-type-
cell levels after reintroduction of HDAC1 in both cell lines
(Fig. 3D). The fact that p57 expression was reestablished in
KO-reB cells but not in KO-reA cells might be due to the
complex interplay of allele-specific repressive chromatin mod-
ifications that is linked to the regulation of imprinted genes
(see Discussion). Taken together, these data suggest that
HDAC1 can reversibly regulate the expression of a specific
subset of mouse genes.
Cross talk between the class I deacetylases HDAC1 and
HDAC2. Next, we wanted to know whether the enzymatic
activity of HDAC1 is required for the regulation of putative
target genes. For that purpose, we expressed an inactive
HDAC1 protein in HDAC1-null cells. The mutation of histi-
dines 140/141 into alanines in the catalytic domain leads to a
complete loss of HDAC1 enzymatic activity without provoking
conformational changes in the protein (31, 72). Wild-type and
HDAC1 KO cells were stably transfected with a retroviral
expression vector encoding the myc-tagged HDAC1-H140/
141A mutant or the empty vector as a control. Interestingly, of
several dozens of clones analyzed by indirect immunofluores-
cence analysis and immunoblotting (data not shown), only one
showed a clearly detectable signal for the mutant HDAC1
protein. This clone, referred to as KO-mut, expressed
bar represents the number of positively regulated genes, and the white bar represents the number of negatively regulated genes in HDAC1-
deficient ES cells. (B) Molecular networks of putative HDAC1 target genes. Putative HDAC1 target genes were overlaid onto a global molecular
network developed from information contained in the Ingenuity Pathways Knowledge Base. Networks of putative HDAC1 target genes were then
algorithmically generated based on their connectivity. The networks are ranked according to their scores, and the five highest-ranking networks
of upregulated and downregulated genes are displayed. The numbers of genes in each network are shown in brackets. (C) Functional classification
of HDAC1 target genes. The probe sets corresponding to all deregulated transcripts were analyzed using Ingenuity Pathways Analysis (Ingenuity
Systems) for molecular and cellular functions. Different categories are ranked according to the numbers of associated genes. Gray bars represent
genes that are upregulated in HDAC1 KO cells, and white bars represent genes that are downregulated in HDAC1 KO cells. The probability that
each biological function and/or disease assigned to the data set is due to chance alone is indicated as a P value for each category (shown on the
right-hand side of each bar). (D) Chromosomal distribution of deregulated genes. Black bars, total numbers of genes on the indicated chromo-
somes relative to the total number of genes in the mouse genome; white bars, numbers of genes on the indicated chromosomes present on the array
relative to the total number of genes present on the array; gray bars, numbers of deregulated genes on the indicated chromosomes relative to the
total number of deregulated genes. Chromosomes X and Y are excluded, since the cell lines were of opposite sexes.
FIG. 2. Validation of the DNA array screen. (A) SYBR green real-time RT-PCR analysis of upregulated genes. Total RNA isolated from
wild-type (WT) and HDAC1-deficient ES cells was reverse transcribed, and PCR was performed with primers specific for the cDNA of indicated
genes. Gene expression levels were normalized to tubulin ?1 levels and shown as expression levels in HDAC1-null cells relative to expression levels
in wild-type cells (arbitrarily set to 1). (B) SYBR green real-time RT-PCR analysis of downregulated genes. The analysis was performed as
described for panel A.
VOL. 26, 2006 TRANSCRIPTIONAL REGULATION BY HDAC17917
HDAC1-H140/141A at a relatively low level compared to the
levels of HDAC1 in the wild-type control (Fig. 4A, left panel),
suggesting that constant high expression of the inactive
HDAC1 mutant might interfere with proliferation of ES cells.
Hence, this clone was taken for further analyses. Remarkably,
the total HDAC activity in the HDAC1 mutant-expressing
cells was about 10% lower than the activity in KO control cells
(Fig. 4A, right panel). This might be due to the slightly reduced
HDAC2 expression in HDAC1-H140/141A-expressing cells
and/or to a negative effect of the mutant HDAC1 protein on
HDAC2 activity. Alternatively, HDAC2 might act as an im-
postor in the absence of HDAC1, a mechanism that has pre-
viously been described for certain mitogen-activated protein
Since HDAC1 and HDAC2 proteins are known to copre-
cipitate and are usually found in the same complexes (2, 29),
we wanted to test whether the HDAC1-H140/141A mutant
also interacted with HDAC2. Thus, HDAC2 was precipitated
from extracts prepared from vector-transfected wild-type cells,
vector-transfected KO cells, and KO cells expressing HDAC1
mutant protein. Subsequent immunoblot analysis with the
HDAC2 antibody showed an expression pattern similar to that
observed for whole-cell lysates (Fig. 4A), with a weak HDAC2
signal in wild-type cells, a strong HDAC2 signal in KO cells,
and slightly reduced amounts of precipitated HDAC2 upon
expression of the H140/141A mutant in KO cells (Fig. 4B).
Both wild-type and mutant HDAC1 proteins efficiently asso-
ciated with HDAC2, as shown by immunoblot analysis of
HDAC2 immunoprecipitates with the HDAC1 antibody (Fig.
4B). Consistent with previous findings (36), HDAC2-associ-
ated activity was significantly increased in HDAC1 KO cells,
indicating that despite its habitual heterodimerization with
HDAC1, HDAC2 can also act as an active deacetylase in the
absence of HDAC1 (Fig. 4B, right panel). Upon the expression
of the HDAC1-H140/141A mutant, HDAC2 activity levels in
HDAC1-null ES cells dropped by 20% (Fig. 4B, right panel).
All together, these data imply that HDAC1-H140/141A ex-
pression in HDAC1 KO cells leads to reduced HDAC2 ex-
pression and HDAC2-associated activity and to a decrease in
total HDAC activity.
FIG. 3. The deregulation of HDAC1 target genes in HDAC1-null cells is reversible. (A) Whole-cell extracts were prepared from two different
clones of HDAC1-transfected KO ES cells (KO-reA and KO-reB) and respective wild-type (WT-vec) and KO (KO-vec) controls. Extracts were
analyzed for expression of HDAC1, HDAC2, and ?-actin on a Western blot (left panel) and for deacetylase activity as described in Materials and
Methods (right panel). Data presented for relative HDAC activities per hour per 10 ?g protein are mean values for three independent experiments.
(B) Equal amounts of the extracts described in panel A were precipitated with HDAC2-specific antibodies, and the immunoprecipitates were
examined for the presence of HDAC1 and HDAC2 (left panel) and associated deacetylase activity (right panel). Data presented are representative
of three independent experiments. (C) SYBR green real-time RT-PCR analysis of mRNA expression in the cell lines described for panel A. Gene
expression levels were normalized to tubulin ?1 levels and are shown relative to expression levels in the wild-type control. Data presented are mean
values for three independent experiments. (D) Northern blot analysis of total RNA isolated from the same cells as used for panel A probed with
cDNA fragments of the indicated genes. Methylene blue staining of 18S RNA illustrates the equal loading of the samples.
7918 ZUPKOVITZ ET AL.MOL. CELL. BIOL.
To analyze how the HDAC1-H140/141A mutant affects the
transcription of putative HDAC1 target genes, we examined
the expression of selected target genes by quantitative real-
time RT-PCR (Fig. 4C). In contrast to that of the HDAC1
wild-type protein, expression of the inactive mutant did not
restore normal expression of putative HDAC1 target genes.
Most of the genes analyzed, e.g., those encoding Mt2, Prss11,
p21, Mt1, Plagl1, and Ass1, showed no change or a moderate
increase in mRNA levels in HDAC1-H140/141A-expressing
cells compared to those in the KO control. Strikingly, two of
the genes analyzed, the Apc2 and JunB genes, had three- to
four-times-higher expression levels in KO-mut cells. Given the
fact that the inactive HDAC1 mutant reduces the enzymatic
activity of HDAC2, the strong increases in the expression lev-
els of these two genes suggest that, in the absence of HDAC1,
HDAC2 might be involved in their regulation.
HDAC inhibitor treatment reveals two distinct groups of
HDAC1 target genes. In order to determine whether other
histone deacetylases are involved in the regulation of putative
HDAC1 target genes, we next analyzed the responsiveness of
putative HDAC1 target genes to the deacetylase inhibitor
TSA, a general inhibitor of most class I and class II HDACs
(43, 86). To this end, wild-type and HDAC1-deficient ES cells
were treated for 12 h with the solvent dimethyl sulfoxide
(DMSO) or increasing concentrations of TSA. The expression
of a panel of putative HDAC1 target genes was then analyzed
by real-time RT-PCR. The known TSA responsive gene
HDAC1 (26, 32, 65) showed a dose-dependent increase in
FIG. 4. Expression of an enzymatically inactive HDAC1 mutant induces a subset of HDAC1 target genes. (A) Left panel, Western blot analysis
of whole-cell extracts prepared from HDAC1-H140/141A mutant-expressing HDAC1 KO cells (KO-mut) and respective wild-type (WT-vec) and
KO (KO-vec) controls. The blot was incubated sequentially with antibodies specific for HDAC1, HDAC2, and ?-actin, respectively. Right panel,
equal amounts of the extracts described for the left panel were analyzed for total deacetylase activity. Data presented for relative HDAC activities
are mean values for three independent experiments. (B) Left panel, Western blot analysis of HDAC2 immunoprecipitates prepared from the
protein extracts described for panel A with antibodies specific for HDAC1 and HDAC2. Right panel, comparison of HDAC2-associated
deacetylase activities after immunoprecipitation with an HDAC2-specific antibody from the whole-cell extracts described for panel A. Data
presented are representative of three independent experiments. (C) SYBR green real-time RT-PCR analysis of target genes. Total RNA isolated
from KO-mut cells and vector-transfected KO cells was analyzed by quantitative RT-PCR for the expression of the indicated genes. Expression
levels (n-fold) were normalized to expression levels in KO-vec cells and presented as mean values for three experiments.
VOL. 26, 2006 TRANSCRIPTIONAL REGULATION BY HDAC17919
expression in wild-type ES cells (Fig. 5A). As expected for
HDAC1 target genes, all tested genes displayed increased ex-
pression in response to TSA treatment in wild-type cells (Fig.
5B-I). Furthermore, in the absence of HDAC1, all tested tar-
get genes were less sensitive to the deacetylase inhibitor. These
results confirm a crucial role for HDAC1 and its enzymatic
activity in the regulation of these genes. Interestingly, in
HDAC1 KO cells, genes such as those encoding Prss11, p21,
Ass1, and Mt2 (Fig. 5B to E) showed only moderate response
(two- to threefold) to TSA, suggesting that the major histone
deacetylase involved in their regulation is HDAC1. In contrast,
four other genes, the Mt1, JunB, Apc2, and Plagl1 genes,
displayed a strong response to TSA (8- to 22-fold) in HDAC1
KO cells (Fig. 5F to I), indicating that these genes are regu-
lated by other HDACs as well. As shown above, the expression
of Apc2 and JunB was significantly enhanced in HDAC1 KO
cells upon the expression of the enzymatically inactive mutant,
i.e., under conditions where HDAC2-associated deacetylase
activity was reduced (Fig. 4C). These findings strongly suggest
that HDAC2 is involved in the regulation of Apc2 and JunB.
Another putative HDAC1 target gene, the Plagl1 gene,
showed a different response; it was not induced by the inactive
HDAC1 mutant but showed a dramatic increase in expression
upon TSA treatment. A possible explanation is that Plagl1
might be regulated by other histone deacetylases, which are
sensitive to TSA but unaffected by the expression of the inac-
tive HDAC1 mutant. Moreover, since the Plagl1 gene is an
imprinted gene, the regulation of its expression might be more
complex. Taken together, these data imply that HDAC1 puta-
tive target genes can be divided into two groups: genes that are
regulated mainly by HDAC1 (e.g., the Prss11 and Ass1 genes)
and genes, such as the JunB and Apc2 genes, that are regulated
by HDAC1 and other histone deacetylases.
Repression of target genes is associated with the presence of
HDAC1 and histone deacetylation. To test whether the regu-
lation of the identified target genes involves direct recruitment
of HDAC1, we analyzed two genes of each group in ChIP
assays. Chromatin isolated from wild-type and HDAC1-defi-
cient ES cells was immunoprecipitated with antibodies specific
for HDAC1 and HDAC2 or with an unspecific antibody as a
control. The amount of immunoprecipitated DNA was ana-
lyzed by semiquantitative PCR using primers specific for the
proximal promoters (within the first 500 bp upstream of the
first exon) and intragenic regions (within 1,000 to 1,500 bp
downstream of the first exon) of individual genes. In wild-type
ES cells, HDAC1 was present on the analyzed regulatory re-
gions of all tested genes, with the exception of the proximal
promoter region of Apc2 (Fig. 6). However, it is likely that
other regions within the Apc2 gene are important for the
HDAC1-dependent regulation of this gene, since HDAC1 was
detected at the Apc2 intragenic region within 1,000 bp of exon
1. In contrast, HDAC1 was absent from control genes, such as
those encoding ?-actin and GAPDH (data not shown). All
together, these data provide strong evidence for a direct in-
volvement of HDAC1 in the regulation of Prss11, Ass1, JunB,
In addition, we analyzed the presence of the HDAC1 ho-
mologue HDAC2 at the promoter of the putative target genes.
Here, it is important to note that the HDAC2 antibody is more
efficient than the HDAC1 antibody in ChIP assays and gives
stronger signals. Therefore, it is not possible to directly com-
pare the amounts of recruited HDAC1 and HDAC2. As shown
in Fig. 6, HDAC2 was present on all genes tested in this
experiment. The amounts of recruited HDAC2 stayed un-
changed on the Prss11 and Ass1 promoters in the absence of
HDAC1. These data suggest that HDAC1 is the predominant
deacetylase regulating Prss11 and Ass1 and that HDAC2 can-
not replace HDAC1 as a repressor of these genes. HDAC1 and
HDAC2 are probably recruited by different transcription fac-
tors and/or by complexes in which HDAC2 does not compen-
sate for the loss of HDAC1. This idea is also supported by the
weak responses of Prss11 and Ass1 to TSA in HDAC1-null
cells (Fig. 5B and C).
In contrast, HDAC2 recruitment was highly increased at the
JunB and Apc2 promoters in HDAC1 knockout cells. This is
consistent with the finding that these genes respond robustly to
FIG. 5. TSA positively regulates the expression of potential HDAC1 target genes. SYBR green real-time RT-PCR analysis of wild-type (WT)
and KO ES cells treated for 12 h with DMSO or 5, 10, 20, and 40 ng/ml (16.5, 33.1, 66.1, and 132.3 nM, respectively) of TSA. Target gene expression
levels, normalized to tubulin ?1 expression levels, are shown relative to expression levels in DMSO-treated wild-type cells (arbitrarily set to 1). The
data presented are mean values for three independent experiments.
7920ZUPKOVITZ ET AL.MOL. CELL. BIOL.
TSA even in the absence of HDAC1 (Fig. 5G and H). We
therefore conclude that both HDAC1 and HDAC2 are in-
volved in the repression of these genes, probably within the
same complexes, where the increased presence of HDAC2 can
partially compensate for the loss of HDAC1 or act as an
impostor. This idea is also supported by the fact that the
expression of the inactive HDAC1 mutant activates JunB and
Apc2 in KO cells, either by negatively affecting HDAC2 activ-
ity or by replacing HDAC2 in putative regulatory complexes.
To examine the effect of the loss of HDAC1 activity on local
histone modifications, we next examined the acetylation levels
of histones H3 and H4 at the chosen HDAC1 target genes in
wild-type and HDAC1-deficient cells. In addition, we analyzed
these genes for the presence of the repressive marks trimethyl
K9 and trimethyl K27 at histone H3. To test whether the
absence of HDAC1 and increased transcription affect the nu-
cleosome density, we also performed ChIP analyses with an
antibody that recognizes the C terminus of histone H3. As
shown in Fig. 6, most of the target genes (with the exception of
the Apc2 promoter) showed slightly reduced nucleosome den-
sities at both the promoters and the intragenic regions in
HDAC1 KO cells. Despite the small reduction in associated
histone proteins upon the loss of HDAC1, we consistently
observed increased acetylation levels of histones H3 and H4 on
nucleosomes associated with the promoters and intragenic re-
gions of all tested genes (Fig. 6). Furthermore, the analysis of
modifications of specific lysine residues on the HDAC1 target
gene promoters revealed a strong hyperacetylation of lysine 9
on histone H3 within all tested target gene regions. Taking into
account the slight reduction in nucleosome density, we ob-
served at the same time the loss of the trimethylation marks on
both lysine 9 and lysine 27 on histone H3 in KO cells.
Taken together, these data show a strong correlation be-
tween the presence of HDAC1 and decreased histone acety-
lation at specific genes, indicating that these are indeed direct
HDAC1 target genes. Our results also emphasize the role of
HDAC1 as an epigenetic regulator, by showing that the loss of
this enzyme can provoke changes not only on its direct sub-
strates, acetylated histones, but also on substrates of other
An HDAC1 target gene, the Prss11 gene, encodes a regulator
of cell proliferation. An HDAC1 target gene, the Prss11 gene,
is a putative tumor suppressor gene in ovarian cancer (12) and
was shown to be consistently upregulated upon the loss of
HDAC1 expression in a variety of cell types, including mouse
fibroblasts and human tumor cells (G. Zupkovitz, S. Chiocca,
and C. Seiser, unpublished observations). Remarkably, small
interfering RNA-mediated ablation of HDAC1 impaired the
proliferation of the human osteosarcoma cell line U2OS, sug-
gesting that HDAC1 might be a relevant target for HDAC
inhibitors as tumor drugs (S. Senese, K. Zaragoza-Dorr, S.
Minardi, L. Bernard, G. F. Draetta, M. Alcalay, C. Seiser, and
S. Chiocca, submitted for publication). To test whether en-
hanced expression of Prss11 would affect the proliferation
rates of human tumor cells, U2OS cells were stably transfected
with a Prss11 expression vector. Vector-transfected control
cells and Prss11-overexpressing single clones were analyzed in
proliferation assays and for expression of the proliferation
marker Ki67. As shown in Fig. 7, both cell numbers and per-
centages of Ki67 positive cells were significantly reduced in
Prss11-overexpressing tumor cells. Thus, the HDAC1 target
gene Prss11 gene encodes a negative regulator of cell prolif-
eration and its overexpression in the absence of HDAC1 might
contribute to the proliferation phenotype observed in HDAC1
KO ES cells and HDAC1-deficient tumor cells.
HDAC1 as a positive regulator of gene expression. Our
analysis of gene expression in wild-type and HDAC1 KO ES
cells revealed that in addition to the 4% of genes upregulated,
3% of genes were downregulated in the absence of HDAC1.
These data suggest that HDAC1 can positively regulate a sig-
nificant portion of murine genes (Fig. 1A). The activating
effect of HDAC1 on gene expression can be a direct conse-
quence of HDAC1 recruitment to a target gene or an indirect
effect due to the increased expression and/or activity of tran-
scriptional repressors upon the loss of HDAC1. For instance,
overexpression of HDAC2 in HDAC1 KO ES cells might
FIG. 6. Absence of HDAC1 on the regulatory regions of target genes is associated with hyperacetylation of histones. Formaldehyde-cross-
linked chromatin from wild-type (WT) and HDAC1 KO cells was immunoprecipitated with control (con), HDAC1, HDAC2, acetylated H3
(AcH3), acetylated H4 (AcH4), acetylated lysine 9 H3 (AcK9H3), trimethylated lysine 9 H3 (3MeK9H3), trimethylated lysine 27 H3 (3MeK27H3),
and C-terminal H3 antibodies (cH3). DNA isolated from immunoprecipitated fractions and total input chromatin was analyzed by semiquantitative
PCR specific for the indicated regions. These results are representative of three independent experiments.
VOL. 26, 2006 TRANSCRIPTIONAL REGULATION BY HDAC17921
cause the repression of specific genes in the absence of
HDAC1 by a compensatory mechanism. To distinguish be-
tween these possibilities, we analyzed several putative target
genes that were downregulated in HDAC1 KO cells for their
responsiveness to TSA. Genes that require HDAC1 for their
activation should be negatively regulated by the deacetylase
inhibitor in wild-type cells. In contrast, genes that are re-
pressed by increased levels of HDAC2 in HDAC1 KO cells
should be reactivated by TSA.
Four putative target genes, the Edg1, Efnb2, Ehd1, and Gja1
genes, showed significant downregulation in HDAC1 KO cells
(2.5- to 5-fold) when tested by real-time RT-PCR analysis (Fig.
8A). The expression of three of these genes (the Efnb2, Ehd1,
and Edg1 genes) was induced by TSA in HDAC1-null cells,
indicating that these genes are negatively regulated by HDACs
in the absence of HDAC1. Efnb2 and Ehd1 showed no signif-
icant response to the deacetylase inhibitor in wild-type cells,
suggesting that their expression is normally not controlled by
TSA-sensitive deacetylases. In contrast, higher concentrations
of TSA stimulated Edg1 expression in wild-type cells and
HDAC1 KO ES cells, indicating that this gene is negatively
regulated by HDACs in both cell types.
Therefore, we examined the Edg1 gene for the presence of
HDAC1 and HDAC2 in ChIP assays. As shown in Fig. 8B,
HDAC1 and small amounts of HDAC2 were detectable at the
Edg1 promoter in wild-type ES cells. The recruitment of
HDAC2 was significantly increased upon the loss of HDAC1,
which correlated with the almost complete loss of Edg1 ex-
pression. Surprisingly, histone acetylation at the Edg1 pro-
moter was not affected by the absence of HDAC1, as signals
for both acetylated histone H3 and acetylated histone H4 and
the C terminus of histone H3 (nucleosome density) were all
reduced to similar extents (Fig. 8B). These results indicate that
additional deacetylase substrates might participate in the reg-
ulation of Edg1. Interestingly, K9 trimethylation on histone H3
seemed to be linked to the presence of HDAC1, whereas K27
trimethylation on histone H3 was not significantly changed in
HDAC1 KO cells. Taken together, these data suggest that the
expression levels of Edg1, Efnb2, and Ehd1 are reduced in
HDAC1-null cells by a compensatory mechanism that involves
the activity of other histone deacetylases, such as HDAC2.
In contrast to those of the target genes described above,
expression levels of Gja1 were reduced not only upon the loss
of HDAC1 but also in a dose-dependent manner by TSA (Fig.
8A). As demonstrated by ChIP analyses, HDAC1 was present
at the Gja1 promoter in wild-type cells, while HDAC2 was
recruited mainly in HDAC1-null cells (Fig. 8B). Given that
Gja1 expression is further reduced by TSA treatment of
HDAC1 KO cells, HDAC2 might to some extent compensate
for the loss of HDAC1. Analysis of chromatin modifications at
the Gja1 promoter revealed that the acetylations of histone H4
and K9 at histone H3 were slightly reduced in the absence of
HDAC1 (Fig. 8B), while the trimethylation of H3-K9 was
increased in HDAC1 KO cells (Fig. 8B). Thus, the reduced
expression of Gja1 correlated with the increased presence of
the repressive K9 trimethylation mark on histone H3. The
fact that histone acetylation on the Gja1 promoter is only
slightly affected suggests that for the regulation of this gene,
other proteins are the relevant targets for HDAC1. In sum-
mary, the Gja1 gene represents an unusual HDAC1 target
gene that requires the presence of HDAC1 and its enzy-
matic activity for activation.
HDAC1 and its activity are required for the activation of
IFN target genes. Next, we asked whether the positive role of
HDAC1 in the regulation of gene expression is a more general
FIG. 7. The HDAC1 target gene Prss1 gene negatively affects pro-
liferation in human tumor cells. (A) Equal numbers (120,000) of un-
transfected, empty-vector-transfected (VecA and VecB), and Prss11-
overexpressing (Prss11A, Prss11B, and Prss11C) U2OS cells were
plated in triplicate on 10-cm dishes. Cell numbers were determined
after 3 days by using a Casy cell counter. The insert shows mean values
for three vector-transfected cell lines (U2OS?vec) and three Prss11-
overexpressing cell lines (U2OS?Prss11). (B) Ki67 staining of the
same cell lines as used for panel A. Ki67 was visualized by indirect
fluorescence microscopy, and nuclear DNA was stained with DAPI.
Data are presented as percentages of Ki67-positive cells per 100
counted cells. The insert shows mean values for Ki67-positive cells for
cell lines transfected with empty-vector (U2OS?vec) and Prss11-over-
expressing (U2OS?Prss11) cell lines.
7922 ZUPKOVITZ ET AL.MOL. CELL. BIOL.
phenomenon. Several reports suggest that histone deacetylase
activity is necessary for activation of interferon (IFN) target
genes, probably through involvement of HDAC1 (9, 54, 63).
Interferon stimulation leads to activation of the JAK/STAT
pathway, resulting in tyrosine phosphorylation of STAT1 and
STAT2. Homodimerization of phosphorylated STAT1 or het-
erodimerization of phosphorylated STAT1 and STAT2 in-
duces their translocation into the nucleus and the activation of
their target genes (for a review, see reference 15). To test a
direct involvement for HDAC1 in the interferon response, we
analyzed the expression of IFN target genes in wild-type and
HDAC1 KO ES cells. Pilot experiments showed that IFN
treatment induced STAT1 phosphorylation in both wild-type
and HDAC1-null ES cells (data not shown). We next stimu-
lated wild-type and HDAC1 KO ES cells with IFN-? in the
presence or absence of TSA and analyzed the expression of
two known IFN-? target genes, the Irf1 and Gbp2 genes. Real-
time RT-PCR analysis demonstrated the transcriptional acti-
vation of both genes in response to IFN-? in wild-type cells
(Fig. 9A). In contrast, the presence of the deacetylase inhibitor
TSA during IFN-? stimulation inhibited the induction of both
genes. Interestingly, the interferon-dependent activation levels
of both Irf1 and Gbp2 were significantly reduced in HDAC1
KO ES cells. These data strongly support the idea that histone
deacetylase activity, and in particular HDAC1 activity, is im-
portant for the induction of certain interferon target genes.
To test whether the activation of specific IFN target genes
requires direct recruitment of HDAC1 to their promoters, we
performed ChIP assays with wild-type and HDAC1 KO ES
cells before and after stimulation with IFN-? and in the pres-
ence or absence of TSA, using specific antibodies for HDAC1,
the C terminus of histone H3, and acetylated histones H3 and
H4. In addition, we monitored the presence of trimethylated
K9 and K27 at histone H3 at both target promoters. The
immunoprecipitated DNA was further analyzed by semiquan-
titative PCR or real-time PCR with primers specific for the
IFN-?-activated site of the Irf1 promoter and the proximal
promoter region of Gbp2. As shown in Fig. 9B, HDAC1 was
absent from both promoters in unstimulated wild-type cells but
was recruited in response to IFN-? stimulation. While TSA
treatment abolished the IFN-?-dependent induction of both
genes, it did not affect the presence of HDAC1 on the Irf1
promoter in IFN-?-treated wild-type cells, suggesting that the
activity of recruited HDAC1 is essential for gene activation.
Small amounts of HDAC1 were also found on both promoters
upon treatment with TSA alone. This might be due to the
previously demonstrated induction of HDAC1 expression by
TSA (32, 65) or to increased recruitment caused by TSA-
mediated acetylation of HDAC1 (60).
As shown in ChIP assays with the C-terminal H3 antibody,
the nucleosome density at the Irf1 promoter was slightly re-
duced in the presence of TSA and was unchanged at the Gbp2
promoter in both wild-type and HDAC1 KO cells under all
conditions tested. Histone acetylation was mostly unaffected
upon IFN treatment, with the exception of H4 acetylation at
the Irf1 promoter in wild-type cells, which was decreased upon
recruitment of HDAC1. Treatment with TSA led uniformly to
hyperacetylation of histone H4 on both promoters. Since there
was no consistent reduction of histone acetylation during the
activation of these target genes (Fig. 9B), we conclude that the
crucial substrates for HDAC1 seem to be nonhistone proteins.
Remarkably, IFN-dependent induction of both genes in
wild-type cells correlated with reduced K9 trimethylation of
histone H3 (Fig. 9B). In HDAC1 KO cells, levels of H3-K9
FIG. 8. Positive impact of HDAC1 on the expression of a subset of target genes. (A) Real-time RT-PCR analysis of the expression of Edg1,
Efnb2, Ehd1, and Gja1 in wild-type (WT) and HDAC1 KO ES cells upon treatment for 12 h with solvent (DMSO) or different concentrations of
TSA. Normalized gene expression levels are shown relative to expression levels in DMSO-treated cells. (B) ChIP analysis of the Edg1 and Gja1
promoters in wild-type and HDAC1 KO cells. Formaldehyde-cross-linked chromatin was immunoprecipitated with control antibodies (con) or
antibodies specific for HDAC1, HDAC2, acetylated H3 (AcH3), acetylated H4 (AcH4), or the C terminus of histone H3 (cH3). DNA isolated from
immunoprecipitated material was analyzed by semiquantitative PCR with primers specific for the respective promoter regions. These results are
representative of three independent experiments.
VOL. 26, 2006 TRANSCRIPTIONAL REGULATION BY HDAC17923
trimethylation at both promoters were relatively low and not
affected by IFN or TSA treatments, with the exception of the
Gbp2 promoter under TSA treatment. Methylation of H3-K27
was reduced on both promoters upon IFN or TSA treatment of
wild-type cells and was generally lower in HDAC1 KO cells.
Thus, activation of both target genes was consistently accom-
panied by the gradual loss of repressive chromatin marks. All
together, these data strongly suggest that HDAC1 and histone
deacetylase activity are necessary for the induction of certain
IFN target genes.
HDAC1 as transcriptional regulator of a specific subset of
mouse genes. In this study, we have analyzed the role of
HDAC1 as transcriptional regulator in mouse ES cells. Several
studies have investigated the effect of histone deacetylase in-
hibitors on gene expression (19, 20), but this is to our knowl-
edge the first analysis of the regulatory function of a single
mammalian deacetylase using a genome-wide approach. We
show here that a specific subset (7%) of mouse genes is de-
regulated in the absence of HDAC1. The number of putative
HDAC1 targets might be in fact underestimated, since
HDAC2 can partially compensate for the loss of HDAC1 as a
transcriptional regulator (see below). Compared to the num-
bers of genes found to be deregulated by deacetylase inhibitors
(2 to 8%), the number of HDAC1 target genes is relatively
high (24, 42, 48, 79). This fact indicates that HDAC1 is a key
player in the regulation of gene expression in ES cells and is in
accordance with the changes observed in HDAC1-deficient ES
cells, such as significantly decreased cellular HDAC activity
and hyperacetylation of a subset of core histones (36). Fur-
thermore, all analyzed HDAC1 target genes were sensitive to
the deacetylase inhibitor TSA, thus supporting the idea that
the enzymatic activity of HDAC1 is important for the regula-
tion of target genes.
Biological function of HDAC1 target genes. Database anal-
ysis of HDAC1 target genes showed that the deacetylase con-
trols the expression of genes involved in a variety of biological
processes. Based on the phenotype of HDAC1-deficient em-
bryos and ES cells, HDAC1 has been implicated in prolifera-
tion control. Along this line, a significant fraction of HDAC1
target genes is involved in growth control and cell communi-
cation. In particular, several genes with proposed tumor sup-
pressor activity (the JunB, Plagl1, Apc2, metallothionein 1,
metallothionein 2, and Prss11 genes) are regulated by HDAC1.
For instance, JunB is downregulated in several human tumors
(10, 57) and was shown to suppress cell proliferation by tran-
scriptional activation of p16 (58). The Plagl1 gene, which en-
codes a growth suppressor, is frequently silenced in ovarian
FIG. 9. HDAC1 activity is required for the induction of specific IFN target genes in ES cells. (A) Real-time RT-PCR analysis of the expression
of Irf1 and Gbp2 in wild-type (WT) and HDAC1 KO ES cells. Cells were treated for 1 h with solvent (mock) or with 10 ?g/ml IFN-? (IFN?), 20
ng/ml (33.1 nM) TSA alone (TSA), or IFN-? and TSA together (IFN?TSA). Expression levels of Irf1 and Gbp2 were normalized to tubulin ?1
levels and are shown relative to the expression levels in wild-type ES cells. Data presented are mean values for three independent experiments.
(B) Presence of HDAC1 and hypoacetylation of histone H4 on Irf1 promoter are associated with its IFN-? induction. Formaldehyde-cross-linked
chromatin from wild-type and HDAC1 KO ES cells treated as described for panel A was immunoprecipitated with control antibodies (con) or
antibodies specific for HDAC1, acetyl-H3 (AcH3), acetyl-H4 (AcH4), or the C terminus of histone H3 (cH3). DNA isolated from immunopre-
cipitated fractions was analyzed by semiquantitative PCR specific for proximal promoter regions of the Irf1 (left) and Gbp2 (right) genes.
7924 ZUPKOVITZ ET AL.MOL. CELL. BIOL.
and breast cancer cells (1, 6), the adenomatous polyposis coli
(APC)-like APC2 gene encodes a putative tumor suppressor
(34), and the genes for Mt1 and Mt2 are repressed in some
metastatic tumors (87).
The Prss11 gene was originally isolated as a gene whose
expression was downregulated in a human fibroblast cell line
after transformation with simian virus 40 (89). Repression of
human Prss11 has been repeatedly observed in ovarian cancers
(66) and melanomas (4), in close correlation with the malig-
nant progression and metastasis of these tumors. We show
here that Prss11 is negatively regulated by HDAC1 and that its
overexpression significantly impairs proliferation in human tu-
mor cells. These findings suggest that HDAC1 is one of the
relevant target enzymes for HDAC inhibitors as tumor drugs.
This idea is also supported by the finding that HDAC1 inacti-
vation induced apoptosis in human tumor cells (Senese et al.,
submitted). We have recently established an HDAC1 mouse
tumor model and will test the function of some of the above-
mentioned genes for their relevance as HDAC1-regulated tu-
Another interesting finding is the identification of several
imprinted genes as HDAC1 targets. It is important to note that
the allele-specific silencing of some of these imprinted genes
occurs only upon differentiation. However, the observed de-
regulation of specific imprinted genes is not due to unsched-
uled ES cell differentiation in the absence of HDAC1, since
virtually all wild-type and HDAC1 KO cells are positive for
Oct4, a marker for undifferentiated ES cells (R. Brunmeir and
C. Seiser, unpublished results). A hallmark of imprinted genes
is the presence of differentially methylated CpG islands known
as differentially methylated regions. Preliminary data suggest
that HDAC1 might be required for the methylation of specific
differentially methylated regions (G. Egger, unpublished ob-
servations). DNA methylation is closely linked to histone
deacetylation, since class I deacetylases, including HDAC1,
were found to associate with both methyl-binding proteins and
methyltransferases (22, 23, 50, 88). In accordance with these
findings, deacetylase inhibitors have been previously shown to
affect the expression of several imprinted HDAC1 target
genes, including the H19, Igf2, and p57 genes (25, 26). To
better understand the role of HDAC1 in the regulation of
imprinted genes, it will be necessary to establish a system
suitable for the study of allele-specific gene expression in the
presence and absence of HDAC1.
Consequences of HDAC1 recruitment. ChIP analysis re-
vealed that HDAC1 is recruited to a specific set of target
promoters. Similarly, the homologous yeast deacetylase Rpd3p
was found to be associated with a specific class of target genes,
while the HATs Gcn5 and Esa1 are generally recruited to the
promoters of active protein-coding genes (61). In addition,
HDAC1 was recruited to the 5? intragenic regions of all inves-
tigated target genes. A detailed ChIP analysis throughout the
murine HDAC1 gene, whose expression is under the control of
HDAC1, showed a predominant localization of HDAC1 at the
5? region of this gene (65; G. Zupkovitz, unpublished results).
These findings corroborate the results from many laborato-
ries showing the recruitment of HDAC1 by specific tran-
scription factors to promoters and enhancers of target genes
(46). Noticeably, the loss of HDAC1 led to reduced nucleo-
some density on most of the HDAC1 target promoters,
suggesting a crucial role for the deacetylase for chromatin
condensation at these genes. Accordingly, the occupancy of
target genes by HDAC1, with the exception of positively
regulated target genes (see below), resulted in the reduced
acetylation of histones H3 and H4.
Interestingly, the presence of other epigenetic marks, namely,
trimethylation at K9 and K27 of histone H3, was reduced at
HDAC1 target genes in the absence of HDAC1. Concerning
histone modifications on target promoters, these results are in
agreement with several reports describing the association of
HDAC1 with histone H3 methyltransferases, including the K9
methyltransferase Suv39h1 and the histone H3-modifying poly-
comb complex PRC2 (14, 77). However, in contrast to a pre-
vious report that showed a link between K9 trimethylation and
transcriptional elongation (76), we observed reduced K9 tri-
methylation at intragenic regions at genes that were induced in
the absence of HDAC1. This might be explained by the com-
plex cross talk between enzymes that control histone acetyla-
tion and methylation. For instance, HDAC1 was shown to
interact not only with a K9 methyltransferase but also with the
recently identified K9-demethylating enzyme JMJD2A (83). In
addition, the recruitment and expression of JMJD2A might be
regulated in an HDAC-dependent manner (27). We are cur-
rently performing a chromosome-wide ChIP-on-chip analysis
for mouse HDAC1 to understand in more detail the conse-
quences of HDAC1 recruitment on histone acetylation and
other chromatin modifications.
Functional links between HDAC1 and HDAC2. Expression
of the related class I enzyme HDAC2 was reliably upregulated
upon inactivation of HDAC1 in all cell systems that we tested,
including mouse ES cells, fibroblasts, T cells, and human tumor
cells (36; unpublished data; Senese et al., submitted). Interest-
ingly, only 7% of the mouse HDAC1 target genes are found to
be deregulated in human tumor cells missing HDAC1 (Senese
et al., submitted). However, when HDAC1 and HDAC2 were
simultaneously inactivated, more than 20% of the murine
HDAC1 targets were differentially expressed in U2OS cells.
Increased amounts of HDAC2 might partially compensate for
the loss of HDAC1 as a transcriptional repressor. This idea is
supported by several findings in the present study. For in-
stance, a group of genes, including the JunB and Apc2 genes,
showed a significant sensitivity towards the HDAC inhibitor
TSA even in the absence of HDAC1 (Fig. 5), suggesting that
they are regulated also by other HDACs. Accordingly, ChIP
assays demonstrated increased recruitment of HDAC2 as a
consequence of the loss of HDAC1 (Fig. 6). Such genes are
most probably regulated by the recruitment of repressor com-
plexes that contain increased amounts of HDAC2 in the ab-
sence of HDAC1, which mask the actual repressive capacity of
HDAC1 for these target genes. In fact, HDAC2 might act as
an impostor (40) by replacing HDAC1 as a component of
certain repressor complexes in HDAC1 KO cells. In agree-
ment with this idea, the expression of an enzymatically inactive
HDAC1 mutant led to increased expression of JunB and Apc2
in HDAC1-null cells (Fig. 4C). It is likely that for some other
genes, HDAC2 can fully compensate for the lost repressor
function of HDAC1 in KO cells. These genes were not de-
tected in our screen but might be identified in an HDAC1
VOL. 26, 2006 TRANSCRIPTIONAL REGULATION BY HDAC17925
HDAC1 as a positive regulator of transcription. Finally, we
have also characterized genes that display reduced expression
levels in HDAC1-deficient cells. For one group of genes, the
decrease in expression in the absence of HDAC1 was rescued
by TSA treatment (Fig. 8A, Efnb2, Ehd1, and Edg1), indicat-
ing that these genes are repressed by other deacetylases in
HDAC1 KO cells. In accordance with this idea, large amounts
of HDAC2 were associated with these genes in the absence of
HDAC1 but not in wild-type cells. In parallel with reduced
expression levels, the acetylation levels of associated histones
were also decreased. Thus, the loss of HDAC1 indirectly leads
to the repression of this group of genes due to the enhanced
presence of HDAC2. Efnb2, a ligand of the ephrin receptor
tyrosine kinase that is required for hippocampal plasticity (30),
was found to be a prognostic marker for neuroblastomas (70,
71). Ehd1, a member of the eps15 homology domain-contain-
ing family (47), was shown to participate in the endocytosis of
the insulin-like growth factor 1 receptor (62), and Edg1 is a
G-protein-coupled receptor for sphingosine-1-phosphate (33)
that seems to stimulate cell migration and metastasis (84).
The most surprising finding of this study was the identifica-
tion of genes that require HDAC1 directly for their activation.
In contrast to all the other HDAC1 target genes, these genes
showed a negative response to TSA (Fig. 8A, Gja1, and Fig.
9A). HDAC1 was recruited either constitutively (Gja1) or, in
the case of the Irf1 and Gbp2 IFN-responsive genes, in re-
sponse to the IFN signal, strongly suggesting that these genes
are direct HDAC1 targets. Gja1 is the major protein of gap
junctions in the heart and seems to display differential re-
sponses to deacetylase inhibitors in normal versus transformed
cells (55). The Irf1 gene was shown to be a tumor susceptibility
gene which encodes a protein with tumor suppressor-like func-
tion (reviewed in reference 69). In contrast to those of the
tumor suppressors p21 and Prss11, which are negatively regu-
lated by HDAC1, Irf1 expression levels seem to be dependent
on HDAC1. The HDAC1 homologue Rpd3p was originally
identified as a factor required for both activation and repres-
sion of yeast genes (82) and plays a positive role in the activa-
tion of osmoresponsive promoters (17). In mammalian cells,
HDAC1 was recently identified as a coactivator of the glu-
cocorticoid receptor (60). Furthermore, histone deacetylase
activity was shown to be required for the proper activation of
IFN-responsive genes and HDAC1 was implicated in this pro-
cess (reviewed in reference 53). We provide the first evidence
for the regulated HDAC1 recruitment to the promoters of Irf1
and Gbp2 during transcriptional induction by IFN-?. The
mechanism of HDAC1-dependent activation of target genes is
not entirely clarified. On the one hand, HDAC1 might be
required predominantly for the deacetylation of specific tran-
scription regulators that trigger their activation. This idea is
supported by the fact that histone acetylation at the positively
regulated Gja1, Irf1, and Gbp2 target genes is mostly unaf-
fected by the presence of HDAC1 (Fig. 8 and 9). On the other
hand, the stimulation of HDAC1 target genes could be accom-
panied by waves of histone acetylation and deacetylation that
are controlled by both HATs and HDACs. Similar cyclic
changes in the acetylation of core histones have been previ-
ously observed during the hormone-dependent activation of
genes (45, 60). Taken together, the results presented in our
study demonstrate a major function for HDAC1 as a transcrip-
tional regulator in mouse ES cells.
We thank G. Weitzer for his continuous help with the ES cell work
and D. Meunier, R. Brunmeir, and E. Simboeck for helpful discus-
sions. We are grateful to G. Brosch, Medical University of Innsbruck,
for providing histones for HDAC assays, to T. Jenuwein for histone
antibodies, and to M. Kawaichi for plasmids.
This work was supported by the Austrian Science Fund (FWF
P16443-B04) and the GEN-AU project Epigenetic Plasticity of the
Mammalian Genome (Federal Ministry for Education, Science and
Culture). G.Z. was a fellow of the Vienna Biocenter Ph.D. program
(Austrian Science Fund).
1. Abdollahi, A., D. Pisarcik, D. Roberts, J. Weinstein, P. Cairns, and T. C.
Hamilton. 2003. LOT1 (PLAGL1/ZAC1), the candidate tumor suppressor
gene at chromosome 6q24-25, is epigenetically regulated in cancer. J. Biol.
2. Ahringer, J. 2000. NuRD and SIN3 histone deacetylase complexes in devel-
opment. Trends Genet. 16:351–356.
3. Allfrey, V. G., R. Faulkner, and A. E. Mirsky. 1964. Acetylation and meth-
ylation of histones and their possible role in the regulation of RNA synthesis.
Proc. Natl. Acad. Sci. USA 51:786–794.
4. Baldi, A., A. De Luca, M. Morini, T. Battista, A. Felsani, F. Baldi, C.
Catricala, A. Amantea, D. M. Noonan, A. Albini, P. G. Natali, D. Lombardi,
and M. G. Paggi. 2002. The HtrA1 serine protease is down-regulated during
human melanoma progression and represses growth of metastatic melanoma
cells. Oncogene 21:6684–6688.
5. Bartl, S., J. Taplick, G. Lagger, H. Khier, K. Kuchler, and C. Seiser. 1997.
Identification of mouse histone deacetylase 1 as a growth factor-inducible
gene. Mol. Cell. Biol. 17:5033–5043.
6. Bilanges, B., A. Varrault, E. Basyuk, C. Rodriguez, A. Mazumdar, C. Pan-
taloni, J. Bockaert, C. Theillet, D. Spengler, and L. Journot. 1999. Loss of
expression of the candidate tumor suppressor gene ZAC in breast cancer cell
lines and primary tumors. Oncogene 18:3979–3988.
7. 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.
8. Chadee, D. N., M. J. Hendzel, C. P. Tylipski, C. D. Allis, D. P. Bazett Jones,
J. A. Wright, and J. R. Davie. 1999. Increased Ser-10 phosphorylation of
histone H3 in mitogen-stimulated and oncogene-transformed mouse fibro-
blasts. J. Biol. Chem. 274:24914–24920.
9. Chang, H. M., M. Paulson, M. Holko, C. M. Rice, B. R. Williams, I. Marie,
and D. E. Levy. 2004. Induction of interferon-stimulated gene expression and
antiviral responses require protein deacetylase activity. Proc. Natl. Acad. Sci.
10. Chang, Y. S., K. T. Yeh, M. Y. Yang, T. C. Liu, S. F. Lin, W. L. Chan, and
J. G. Chang. 2005. Abnormal expression of JUNB gene in hepatocellular
carcinoma. Oncol. Rep. 13:433–438.
11. Cheung, P., K. G. Tanner, W. L. Cheung, P. Sassone Corsi, J. M. Denu, and
C. D. Allis. 2000. Synergistic coupling of histone H3 phosphorylation and
acetylation in response to epidermal growth factor stimulation. Mol. Cell
12. Chien, J., J. Staub, S. I. Hu, M. R. Erickson-Johnson, F. J. Couch, D. I.
Smith, R. M. Crowl, S. H. Kaufmann, and V. Shridhar. 2004. A candidate
tumor suppressor HtrA1 is downregulated in ovarian cancer. Oncogene
13. Cress, W. D., and E. Seto. 2000. Histone deacetylases, transcriptional con-
trol, and cancer. J. Cell. Physiol. 184:1–16.
14. Czermin, B., G. Schotta, B. B. Hulsmann, A. Brehm, P. B. Becker, G. Reuter,
and A. Imhof. 2001. Physical and functional association of SU(VAR)3-9 and
HDAC1 in Drosophila. EMBO Rep. 2:915–919.
15. Decker, T., S. Stockinger, M. Karaghiosoff, M. Muller, and P. Kovarik. 2002.
IFNs and STATs in innate immunity to microorganisms. J. Clin. Investig.
16. Delaval, K., and R. Feil. 2004. Epigenetic regulation of mammalian genomic
imprinting. Curr. Opin. Genet. Dev. 14:188–195.
17. De Nadal, E., M. Zapater, P. M. Alepuz, L. Sumoy, G. Mas, and F. Posas.
2004. The MAPK Hog1 recruits Rpd3 histone deacetylase to activate osmo-
responsive genes. Nature 427:370–374.
18. Doetzlhofer, A., H. Rotheneder, G. Lagger, M. Koranda, V. Kurtev, G.
Brosch, E. Wintersberger, and C. Seiser. 1999. Histone deacetylase 1 can
repress transcription by binding to Sp1. Mol. Cell. Biol. 19:5504–5511.
19. Dokmanovic, M., and P. A. Marks. 2005. Prospects: histone deacetylase
inhibitors. J. Cell. Biochem. 96:293–304.
20. Drummond, D. C., C. O. Noble, D. B. Kirpotin, Z. Guo, G. K. Scott, and C. C.
Benz. 2005. Clinical development of histone deacetylase inhibitors as anti-
cancer agents. Annu. Rev. Pharmacol. Toxicol. 45:495–528.
7926 ZUPKOVITZ ET AL.MOL. CELL. BIOL.
21. Fischle, W., Y. Wang, and C. D. Allis. 2003. Histone and chromatin cross-
talk. Curr. Opin. Cell Biol. 15:172–183.
22. Fuks, F., W. A. Burgers, A. Brehm, L. Hughes Davies, and T. Kouzarides.
2000. DNA methyltransferase Dnmt1 associates with histone deacetylase
activity. Nat. Genet. 24:88–91.
23. Fuks, F., W. A. Burgers, N. Godin, M. Kasai, and T. Kouzarides. 2001.
Dnmt3a binds deacetylases and is recruited by a sequence-specific repressor
to silence transcription. EMBO J. 20:2536–2544.
24. Gius, D., H. Cui, C. M. Bradbury, J. Cook, D. K. Smart, S. Zhao, L. Young,
S. A. Brandenburg, Y. Hu, K. S. Bisht, A. S. Ho, D. Mattson, L. Sun, P. J.
Munson, E. Y. Chuang, J. B. Mitchell, and A. P. Feinberg. 2004. Distinct
effects on gene expression of chemical and genetic manipulation of the
cancer epigenome revealed by a multimodality approach. Cancer Cell 6:361–
25. Grandjean, V., L. O’Neill, T. Sado, B. Turner, and A. Ferguson-Smith. 2001.
Relationship between DNA methylation, histone H4 acetylation and gene
expression in the mouse imprinted Igf2-H19 domain. FEBS Lett. 488:165–
26. Gray, S. G., and T. J. Ekstrom. 1998. Effects of cell density and trichostatin
A on the expression of HDAC1 and p57Kip2 in Hep 3B cells. Biochem.
Biophys. Res. Commun. 245:423–427.
27. Gray, S. G., A. H. Iglesias, F. Lizcano, R. Villanueva, S. Camelo, H. Jingu,
B. T. Teh, N. Koibuchi, W. W. Chin, E. Kokkotou, and F. Dangond. 2005.
Functional characterization of JMJD2A, a histone deacetylase- and retino-
blastoma-binding protein. J. Biol. Chem. 280:28507–28518.
28. Gregoretti, I. V., Y. M. Lee, and H. V. Goodson. 2004. Molecular evolution
of the histone deacetylase family: functional implications of phylogenetic
analysis. J. Mol. Biol. 338:17–31.
29. Grozinger, C. M., and S. L. Schreiber. 2002. Deacetylase enzymes: biological
functions and the use of small-molecule inhibitors. Chem. Biol. 9:3–16.
30. Grunwald, I. C., M. Korte, G. Adelmann, A. Plueck, K. Kullander, R. H.
Adams, M. Frotscher, T. Bonhoeffer, and R. Klein. 2004. Hippocampal
plasticity requires postsynaptic ephrinBs. Nat. Neurosci. 7:33–40.
31. Hassig, C. A., J. K. Tong, T. C. Fleischer, T. Owa, P. G. Grable, D. E. Ayer,
and S. L. Schreiber. 1998. A role for histone deacetylase activity in HDAC1-
mediated transcriptional repression. Proc. Natl. Acad. Sci. USA 95:3519–
32. Hauser, C., B. Schuettengruber, S. Bartl, G. Lagger, and C. Seiser. 2002.
Activation of the mouse histone deacetylase 1 gene by cooperative histone
phosphorylation and acetylation. Mol. Cell. Biol. 22:7820–7830.
33. Hobson, J. P., H. M. Rosenfeldt, L. S. Barak, A. Olivera, S. Poulton, M. G.
Caron, S. Milstien, and S. Spiegel. 2001. Role of the sphingosine-1-phos-
phate receptor EDG-1 in PDGF-induced cell motility. Science 291:1800–
34. Jarrett, C. R., J. Blancato, T. Cao, D. S. Bressette, M. Cepeda, P. E. Young,
C. R. King, and S. W. Byers. 2001. Human APC2 localization and allelic
imbalance. Cancer Res. 61:7978–7984.
35. Jenuwein, T., and C. D. Allis. 2001. Translating the histone code. Science
36. Lagger, G., D. O’Carroll, M. Rembold, H. Khier, J. Tischler, G. Weitzer, B.
Schuettengruber, C. Hauser, R. Brunmeir, T. Jenuwein, and C. Seiser. 2002.
Essential function of histone deacetylase 1 in proliferation control and CDK
inhibitor repression. EMBO J. 21:2672–2681.
37. Lechner, T., A. Lusser, G. Brosch, A. Eberharter, M. Goralik Schramel, and
P. Loidl. 1996. A comparative study of histone deacetylases of plant, fungal
and vertebrate cells. Biochim. Biophys. Acta 1296:181–188.
38. Luger, K., A. W. Mader, R. K. Richmond, D. F. Sargent, and T. J. Richmond.
1997. Crystal structure of the nucleosome core particle at 2.8 Å resolution.
39. Luger, K., and T. J. Richmond. 1998. The histone tails of the nucleosome.
Curr. Opin. Genet. Dev. 8:140–146.
40. Madhani, H. D., and G. R. Fink. 1998. The riddle of MAP kinase signaling
specificity. Trends Genet. 14:151–155.
41. Margueron, R., P. Trojer, and D. Reinberg. 2005. The key to development:
interpreting the histone code? Curr. Opin. Genet. Dev. 15:163–176.
42. Mariadason, J. M., C. Nicholas, K. E. L’Italien, M. Zhuang, H. J. Smartt,
B. G. Heerdt, W. Yang, G. A. Corner, A. J. Wilson, L. Klampfer, D. Arango,
and L. H. Augenlicht. 2005. Gene expression profiling of intestinal epithelial
cell maturation along the crypt-villus axis. Gastroenterology 128:1081–1088.
43. Marks, P., R. A. Rifkind, V. M. Richon, R. Breslow, T. Miller, and W. K.
Kelly. 2001. Histone deacetylases and cancer: causes and therapies. Nat.
Rev. Cancer 1:194–202.
44. Mei, S., A. D. Ho, and U. Mahlknecht. 2004. Role of histone deacetylase
inhibitors in the treatment of cancer (review). Int. J. Oncol. 25:1509–1519.
45. Metivier, R., G. Penot, M. R. Hubner, G. Reid, H. Brand, M. Kos, and F.
Gannon. 2003. Estrogen receptor-alpha directs ordered, cyclical, and com-
binatorial recruitment of cofactors on a natural target promoter. Cell 115:
46. Meunier, D., and C. Seiser. 2006. Histone deacetylase 1. In E. Verdin (ed.),
Histone deacetylases: transcriptional regulation and other cellular functions.
Humana Press Inc., Totowa, N.J.
47. Mintz, L., E. Galperin, M. Pasmanik-Chor, S. Tulzinsky, Y. Bromberg, C. A.
Kozak, A. Joyner, A. Fein, and M. Horowitz. 1999. EHD1—an EH-domain-
containing protein with a specific expression pattern. Genomics 59:66–76.
48. Mitsiades, C. S., N. S. Mitsiades, C. J. McMullan, V. Poulaki, R. Shringar-
pure, T. Hideshima, M. Akiyama, D. Chauhan, N. Munshi, X. Gu, C. Bailey,
M. Joseph, T. A. Libermann, V. M. Richon, P. A. Marks, and K. C. Ander-
son. 2003. Transcriptional signature of histone deacetylase inhibition in
multiple myeloma: biological and clinical implications. Proc. Natl. Acad. Sci.
49. Morison, I. M., J. P. Ramsay, and H. G. Spencer. 2005. A census of mam-
malian imprinting. Trends Genet. 21:457–465.
50. Nan, X., H. H. Ng, C. A. Johnson, C. D. Laherty, B. M. Turner, R. N.
Eisenman, and A. Bird. 1998. Transcriptional repression by the methyl-CpG-
binding protein MeCP2 involves a histone deacetylase complex. Nature
51. Nasmyth, K., D. Stillman, and D. Kipling. 1987. Both positive and negative
regulators of HO transcription are required for mother-cell-specific mating-
type switching in yeast. Cell 48:579–587.
52. Ng, H. H., and A. Bird. 2000. Histone deacetylases: silencers for hire. Trends
Biochem. Sci. 25:121–126.
53. Nusinzon, I., and C. M. Horvath. 2005. Histone deacetylases as transcrip-
tional activators? Role reversal in inducible gene regulation. Sci. STKE
54. Nusinzon, I., and C. M. Horvath. 2003. Interferon-stimulated transcription
and innate antiviral immunity require deacetylase activity and histone
deacetylase 1. Proc. Natl. Acad. Sci. USA 100:14742–14747.
55. Ogawa, T., T. Hayashi, M. Tokunou, K. Nakachi, J. E. Trosko, C. C. Chang,
and N. Yorioka. 2005. Suberoylanilide hydroxamic acid enhances gap junc-
tional intercellular communication via acetylation of histone containing con-
nexin 43 gene locus. Cancer Res. 65:9771–9778.
56. Oka, C., R. Tsujimoto, M. Kajikawa, K. Koshiba-Takeuchi, J. Ina, M. Yano,
A. Tsuchiya, Y. Ueta, A. Soma, H. Kanda, M. Matsumoto, and M. Kawaichi.
2004. HtrA1 serine protease inhibits signaling mediated by Tgfbeta family
proteins. Development 131:1041–1053.
57. Passegue, E., W. Jochum, M. Schorpp-Kistner, U. Mohle-Steinlein, and E. F.
Wagner. 2001. Chronic myeloid leukemia with increased granulocyte pro-
genitors in mice lacking junB expression in the myeloid lineage. Cell 104:
58. Passegue, E., and E. F. Wagner. 2000. JunB suppresses cell proliferation
by transcriptional activation of p16(INK4a) expression. EMBO J. 19:2969–
59. Peters, A. H., S. Kubicek, K. Mechtler, R. J. O’Sullivan, A. A. Derijck, L.
Perez-Burgos, A. Kohlmaier, S. Opravil, M. Tachibana, Y. Shinkai, J. H.
Martens, and T. Jenuwein. 2003. Partitioning and plasticity of repressive
histone methylation states in mammalian chromatin. Mol. Cell 12:1577–
60. Qiu, Y., Y. Zhao, M. Becker, S. John, B. S. Parekh, S. Huang, A. Hendar-
wanto, E. D. Martinez, Y. Chen, H. Lu, N. L. Adkins, D. A. Stavreva, M.
Wiench, P. T. Georgel, R. L. Schiltz, and G. L. Hager. 2006. HDAC1
acetylation is linked to progressive modulation of steroid receptor-induced
gene transcription. Mol. Cell 22:669–679.
61. Robert, F., D. K. Pokholok, N. M. Hannett, N. J. Rinaldi, M. Chandy, A.
Rolfe, J. L. Workman, D. K. Gifford, and R. A. Young. 2004. Global position
and recruitment of HATs and HDACs in the yeast genome. Mol. Cell
62. Rotem-Yehudar, R., E. Galperin, and M. Horowitz. 2001. Association of
insulin-like growth factor 1 receptor with EHD1 and SNAP29. J. Biol. Chem.
63. Sakamoto, S., R. Potla, and A. C. Larner. 2004. Histone deacetylase activity
is required to recruit RNA polymerase II to the promoters of selected
interferon-stimulated early response genes. J. Biol. Chem. 279:40362–40367.
64. Schreiber, S. L., and B. E. Bernstein. 2002. Signaling network model of
chromatin. Cell 111:771–778.
65. Schuettengruber, B., E. Simboeck, H. Khier, and C. Seiser. 2003. Autoreg-
ulation of mouse histone deacetylase 1 expression. Mol. Cell. Biol. 23:6993–
66. Shridhar, V., A. Sen, J. Chien, J. Staub, R. Avula, S. Kovats, J. Lee, J. Lillie,
and D. I. Smith. 2002. Identification of underexpressed genes in early- and
late-stage primary ovarian tumors by suppression subtraction hybridization.
Cancer Res. 62:262–270.
67. Strahl, B. D., and C. D. Allis. 2000. The language of covalent histone
modifications. Nature 403(6765):41–45.
68. Sutterluety, H., S. Bartl, J. Karlseder, E. Wintersberger, and C. Seiser. 1996.
Carboxy-terminal residues of mouse thymidine kinase are essential for the
rapid degradation in quiescent cells. J. Mol. Biol. 259:383–392.
69. Tanaka, N., and T. Taniguchi. 2000. The interferon regulatory factors and
oncogenesis. Semin. Cancer Biol. 10:73–81.
70. Tang, X. X., M. E. Robinson, J. S. Riceberg, D. Y. Kim, B. Kung, T. B. Titus,
S. Hayashi, A. W. Flake, D. Carpentieri, and N. Ikegaki. 2004. Favorable
neuroblastoma genes and molecular therapeutics of neuroblastoma. Clin.
Cancer Res. 10:5837–5844.
71. Tang, X. X., H. Zhao, M. E. Robinson, B. Cohen, A. Cnaan, W. London, S. L.
Cohn, N. K. Cheung, G. M. Brodeur, A. E. Evans, and N. Ikegaki. 2000.
VOL. 26, 2006TRANSCRIPTIONAL REGULATION BY HDAC17927