Download Chromatin Regulators and Transcriptional Control of Drosophila

Chromatin Regulators and
Transcriptional Control of
Drosophila Development
Qi Dai
Stockholm University
The cover shows transgenic Drosophila embryos hybridized with a lacZ RNA
probe. The lacZ reporter gene is driven by a modified rho neuroectoderm enhancer (NEE) with synthetic Snail binding sites. The endogenous Snail transcription factor represses reporter gene expression in the wild-type (wt) embryo,
but fails to repress in an embryo devoid of the co-repressor Ebi (ebi).
Qi Dai, Stockholm 2007
ISBN (978-91-7155-547-2)
Printed in Sweden by US-AB, Stockholm 2007
Distributor: Wenner-Gren Institute, Division of Developmental Biology
To my beloved family
Abstract
The development of a multicellular organism is programmed by complex
temporal and spatial patterns of gene expression. In eukaryotic cells, DNA is
packaged by histone proteins into chromatin. Chromatin regulators affect
gene expression by influencing access of proteins to DNA and often function
as transcription co-factors. The interplay between transcriptional activators
and repressors is modulated by co-factors at cis-regulatory DNA modules
(CRMs) to generate precise gene expression patterns.
In this study, we have investigated the in vivo function of four co-factors,
dAda2b, Reptin, Ebi and Brakeless during development of the fruit fly Drosophila melanogaster. dAda2b and Reptin belong to histone acetyl transferase (HAT) complexes, a SAGA-like complex and the Tip60 complex,
respectively. We generated dAda2b mutants and found that lack of dAda2b
strongly affects global histone acetylation and viability. dAda2b mutants are
hyper-sensitive to irradiation-induced DNA damage. Similarly, yeast SAGA
mutants are sensitive to the genetoxic agent MMS. We propose that Ada2
and its associated complexes may be involved in DNA repair.
Reptin belongs to several chromatin regulatory complexes including the
Tip60 HAT complex which is actively involved in transcription, DNA repair
and other cellular processes. Our studies revealed new roles of Reptin and
other Tip60 complex components in Polycomb Group mediated repression
and heterochromatin formation, thereby promoting generation of silent
chromatin.
During embryogenesis, transcriptional repressors establish localized and
tissue-specific patterns of gene expression. Their activities often utilize
ubiquitously distributed co-repressors. In this thesis, we identified two novel
co-repressors in the early embryo, Ebi and Brakeless. Ebi genetically and
physically interacts with the Snail repressor. The Ebi-interaction motif in the
Snail protein is essential for Snail function in vivo and is evolutionarily conserved in insects. We further demonstrated that Ebi associates with histone
deacetylase 3 (HDAC3) and that histone deacetylation is part of the mechanism by which Snail mediates transcriptional repression.
We isolated Brakeless in a genetic screen for novel regulators of gene expression during embryogenesis. We found that mutation of brakeless impairs
function of the Tailless repressor. Brakeless and Tailless bind to each other
and interact genetically. Brakeless also associates with Atrophin, another
Tailless corepressor, and both are recruited to a Tailless-regulated gene in
embryos where they function together in Tailless-mediated repression.
In summary, transcription co-factors, including chromatin regulators, are
selectively required in distinct processes during development.
List of papers
I
Dai Qi, Jan Larsson, and Mattias Mannervik (2004) Drosophila
Ada2b is required for viability and normal histone H3 acetylation.
Mol Cell Biol. 24(18):8080-9.
II
Dai Qi, Haining Jin, Tobias Lilja, and Mattias Mannervik (2006)
Drosophila Reptin and other TIP60 complex components promote generation of silent chromatin. Genetics. 174(1):241-51.
III
Dai Qi, Mattias Bergman, Hitoshi Aihara, Yutaka Nibu, and
Mattias Mannervik (2007) Drosophila Ebi mediates Snaildependent transcriptional repression through HDAC3-induced
histone deacetylation. (Submitted).
IV
Achim Haecker, Dai Qi, Tobias Lilja, Bernard Moussian, Luiz
Paulo Andrioli, Stefan Luschnig, and Mattias Mannervik (2007)
Drosophila Brakeless interacts with Atrophin and is required for
Tailless-mediated transcriptional repression in early embryos.
PLoS Biol. 5(6):1298-1308.
Articles are reprinted with permission from publishers; American Society for
Microbiology, Genetics Society of America and Public Library of Science
Abbreviations
Ada2
AP axis
ATM
ATR
Atro
Bks
CRM
CtBP
DV axis
Gro
GTF
H3K9
HAT
Hb
HDAC
HMT
HP1
Knirps
Kruppel
NCoR
PcG
PEV
PRC1
Rho
Sim
SMRT
Sog
TF
Tip60
Tll
TrxG
Adaptor protein 2
Anterior-posterior axis
Ataxia-telangiectasia mutated
ATM and Rad-3 related
Atrophin
Brakeless
Cis regulatory module
C-terminal binding protein
Dorsal-ventral axis
Groucho
General transcription factor
Histone H3 lysine 9
Histone acetyl transferase
Hunchback
Histone deacetylase
Histone methyl transferase
Heterochromatin protein 1
Kni
Kr
Nuclear receptor corepressor
Polycomb group
Position effect variegation
Polycomb repressive complex 1
Rhomboid
Single-minded
Silencing mediator for retinoic acid and thyroid
hormone receptors
Short-gastrulation
Transcription factor
Tat interaction protein 60kD
Tailless
Trithorax group
Contents
Introduction................................................................................................... 11
Transcription machinery in eukaryotes......................................................................... 11
Chromatin and its function ............................................................................................ 12
Drosophila as a model system ..................................................................... 13
The life cycle of Drosophila........................................................................................... 13
Early embryonic development of Drosophila ................................................................ 14
Dorsal-ventral patterning ......................................................................................... 15
Snail in gastrulation and mesoderm formation .................................................. 16
Anterior-posterior patterning.................................................................................... 17
Histone modifications ................................................................................... 20
Functions of histone modifications ............................................................................... 21
Histone acetylation and its function .............................................................................. 22
Histone acetylation in gene regulation.......................................................................... 22
Histone acetylation in DNA repair................................................................................. 23
Histone acetyl-transferase complexes.......................................................................... 25
GNAT complexes .................................................................................................... 26
Ada2 proteins..................................................................................................... 27
Tip60 HAT complexes ............................................................................................. 28
Reptin................................................................................................................. 29
Histone deacetylases and their associated complexes................................................ 31
NCoR/SMRT/HDAC3 complex................................................................................ 32
Epigenetic control ......................................................................................................... 33
Heterochromatin and position effect variegation..................................................... 33
Polycomb group and Trithorax group...................................................................... 34
Co-regulators................................................................................................ 36
CtBP.............................................................................................................................. 38
Groucho ........................................................................................................................ 38
Atrophin and Brakeless................................................................................................. 39
Aim of this thesis .......................................................................................... 41
Results and discussion of project................................................................. 42
Paper I .......................................................................................................................... 42
Paper II ......................................................................................................................... 43
Paper III ........................................................................................................................ 44
Paper IV ........................................................................................................................ 45
Conclusion and Perspectives ....................................................................... 47
Acknowledgements ...................................................................................... 48
References ................................................................................................... 50
Introduction
A human being contains around 200 different cell types, all of which arise
from a single cell, the fertilized egg. This is probably the major issue that
fascinates developmental biologists, how a single-cell egg can develop into a
mature adult organism. The developmental process is directed by genetic
information that is stored in the fertilized egg. In the cell, DNA molecules
carry the genetic information units, called genes. Genes are first transcribed
into RNAs which subsequently deliver (translate) the genetic code to proteins, the fundamental performers of most functions in the cell. This is called
the central dogma of molecular biology and is valid in most organisms
except RNA viruses. By controlling where and when proteins are synthesized, genes control development. In other words, the genetically identical
cells come to differ from one another by expressing distinct sets of genes
during development.
Transcription machinery in eukaryotes
TF
TF
Chromatin
regulators
Mediator
B
A
D
TATA
F
RNA pol II
EH
Figure 1. Typical transcription machinery of eukaryotic genes. RNA Pol II binds
to the promoter together with GTFs (TFIIA, B, D, E, F and H). TATA box is the
recognition site by a subunit of TFIID, TATA box binding protein (TBP). Transcription factors (TFs) bind CRMs in DNA. The mediator complexes bridge GTFs
and TFs. Chromatin regulators change the status of chromatin.
Gene expression can be regulated at several steps, such as transcriptional
initiation, transcriptional elongation, RNA splicing and translation. Most of
eukaryotic genes are transcribed by RNA polymerase II (Pol II). RNA poly11
merase II recognizes transcriptional start sites with help from general transcription factors (GTFs) consisting of TFIID, TFIIA, TFIIB, TFIIE, TFIIF
and TFIIH. Additionally, the typical transcription machinery includes mediator complexes, accessory complexes including chromatin regulators and
transcription factors (TFs) (Figure 1) (reviewed in Hahn, 2004). TFs bind to
DNA and function as either activators that help transcription initiation or
repressors that hinder transcription initiation.
Chromatin and its function
The genome of a eukaryote is wrapped in proteins called histones to form
nucleosomes. One nucleosome contains a histone octamer (an H3/H4
tetramer and two H2A/H2B dimers) wrapped around 147 base pairs of DNA
(Luger et al., 1997). Numerous nucleosomes build up chromatin. Chromatin
is packaged into chromosomes. So the transcription machinery is present
with a partially concealed substrate. Changing the accessibility of DNA for
the transcription machinery is an important mean for regulating gene expression. Histones contribute to the function of chromatin through at least three
principles: First, chromatin-remodeling factors, such as the SWI/SNF complexes alter the position or properties of the nucleosomes by affecting the
stability of histone-DNA interactions (Peterson, 2002). Second, the modifications of histones regulate accessibility of chromatin for the cellular machinery (Strahl and Allis, 2000). Finally, the exchange of core histones with
specialized histone variants, such as H2AX and H3.3, affect the composition
and function of nucleosomes (Smith, 2002).
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Drosophila as a model system
The basic principle of development and many developmental genes are conserved in different organisms. We used the fruitfly Drosophila melanogaster
as a model system to investigate gene function in development. The fruitfly
has lots of basic advantages, such as inexpensive culturing condition, rapid
life cycle, large amount of progenies, and ease of manipulation. It also offers
many sophisticated tools of classical and molecular genetics. In Drosophila,
it is possible to genetically investigate the function of endogenously and
exogenously introduced genes in a way that is impossible in humans and
impractical in mice. For example, large scale screens using chemicalinduced mutagenesis can be used to analyze gene functions involved in specific developmental and cellular processes. Transposon-based technologies
are used to introduce transgenes, cause lethal mutations and screen for gene
expression patterns. It is also very efficient to create mosaic clones in somatic tissue as well as in the germ line in Drosophila. Furthermore, the
genes of interest can be inactivated and misexpressed in a tissue of choice.
Taken together, Drosophila provides us a very useful system to study the in
vivo function of regulators in gene expression.
The life cycle of Drosophila
The Drosophila life cycle starts from a very rapid embryonic stage (0-24
hours after fertilization (AF)), followed by three larval stages: the first instar
L1 (24-48 hours AF), the second instar L2 (48-72 hours) and the third instar
L3 (72-120 hours AF), and then the pupa stage (metamorphosis) (day 6-9
AF). After pupariation, the animal develops into an adult fly. The early Drosophila embryo is one of the best-characterized developmental systems and
provides a particularly useful framework for the analysis of gene function
(see below). During larval development, one of the notable events is the
growth of imaginal discs. They are initially small and composed of fewer
than 100 cells in the mid-stage embryo but contain tens of thousands of cells
in the mature larva. There is a pair of discs for every set of appendages (for
example, labia, antennae, eyes, wings, three pairs of legs and genitalia).
Imaginal discs differentiate into their appropriate adult structures during
metamorphosis. Another useful system are the polytene chromosomes in the
13
salivary glands in the third instar larva. The polytene chromosomes result
from successive rounds of replication without nuclear division or cytokinesis.
They are particularly useful in studying chromosome structure and localization of chromatin binding proteins.
Early embryonic development of Drosophila
B
amnioserosa
A
Dorsal
zen
Dorsal ectoderm
Acron
Telson
rho
sog
Dorsal gradient
Ventral ectoderm
Posterior
Anterior
Sna
Ventral
Head
Mesoderm
Thorax
Tail
Abdomen
Figure 2. The dorsal-ventral axis and anterior-posterior axis of Drosophila. A.The
schematic picture shows a section view of a pre-cellularization stage embryo. The
nuclear gradient of the Dorsal protein specifies four regions along DV axis from ventral to dorsal side: the mesoderm, ventral ectoderm, dorsal ectoderm and amnioserosa.
sna, rho, sog and zen are representative genes regulated by Dorsal in these regions. B.
Picture of cuticle preparation of a first instar larva. From anterior to posterior, the
main body of the animal is divided into the head, thorax and abdomen. The terminal
structures include acron and telson.
Before fertilization, many important genes are transcribed to mRNAs and/or
translated to proteins by the mother and then these gene products are deposited into the egg. In this way, the mother fly has stored enough material in
the egg to initiate the early embryonic development (reviewed in St Johnston
and Nusslein-Volhard, 1992). After fertilization, the diploid, zygotic nucleus
undergoes a series of nearly synchronous divisions which results in a
syncytium, meaning a single cell with multiple nuclei. Then most of the
resulting nuclei begin to migrate to the periphery. The end result of the migration and more divisions is the formation of a monolayer of approximately
6,000 nuclei surrounding the central yolk. During the following one hour,
from 2 to 3 hours after fertilization, cell membranes form between adjacent
nuclei. This process is called cellularization. During this short period, each
nucleus is rapidly specified to follow a particular pathway of differentiation.
The dorsal-ventral (DV) axis and anterior-posterior (AP) axis are both determined by this time. Along the DV axis, the embryo becomes divided up
into four regions: from ventral to dorsal these are the mesoderm, ventral
ectoderm (neurogenic ectoderm), dorsal ectoderm and the amnioserosa (Fig-
14
ure 2A). Along the AP axis, the embryo is divided into three broad regions
which will become the head, thorax and abdomen (Figure 2B).
Dorsal-ventral patterning
Dorsal-ventral patterning is implemented by the graded distribution of a
maternal transcription factor, Dorsal, and culminates in the differentiation of
specialized tissues (reviewed in St Johnston and Nusslein-Volhard, 1992).
Approximately 3-5 hours after fertilization, each of these tissues generates
multiple cell types.
The key player in this process is the Dorsal protein. It is deposited by the
mother and initially distributed throughout the cytoplasm of the unfertilized
egg. After fertilization, the Dorsal protein forms a nuclear gradient along
dorsal-ventral axis with the highest concentration in the ventral most region
and provides positional information for the determination of different cell
fates. The generation of the nuclear gradient of Dorsal originates from
oogenesis and involves at least 17 maternal factors (reviewed in Moussian
and Roth, 2005). The procedure can be generally divided into two steps.
First, a gradient of the activated Spaetzle ligand is established from ventral
to dorsal in the extracellular matrix (Morisato, 2001; Roth, 2003). Second,
Spaetzle binds to the cell surface receptor Toll and thus triggers another signaling cascade inside the cell, which finally induces the translocation of
Dorsal from the cytoplasm into nuclei (Bergmann et al., 1996; Towb et al.,
1998; Yang and Steward, 1997).
Once this nuclear gradient is achieved, Dorsal functions as a transcriptional regulator, leading to the differential expression of nearly 50 genes
across the dorsal-ventral axis (Stathopoulos et al., 2002). Roughly half the
genes encode transcriptional factors, whereas the other half encodes signaling components. They play fundamental roles in embryogenesis. The Dorsal
gradient specifies multiple thresholds of gene activation. High levels of Dorsal activate the transcription of twist and snail in the ventral-most cells,
which invaginate into the embryo to form mesoderm (Rusch and Levine,
1996). In the lateral region, intermediate levels of Dorsal activate genes (e.g.
rhomboid (rho)) that are required for the formation of ventral neuroectoderm.
In the more lateral region, low levels of Dorsal are sufficient to activate the
short-gastrulation (sog) gene in both the ventral and dorsal neurogenic ectoderm. The neuroectoderm gives rise to the ventral nerve cord and the ventral
epidermis. In principle, rho and sog can also be activated by high levels of
Dorsal. However, their expression is precluded from the ventral most cells
due to repression by Snail. How Dorsal produces so many different transcription outputs is extensively studied (Stathopoulos and Levine, 2002). It
was suggested that the nature of the Dorsal binding sites (affinity, number)
15
determines the binding efficiency and thus determines how much nuclear
Dorsal is sufficient. For example, the twist and snail cis regulatory modules
(CRM) only contain low-affinity binding sites, so they need highest level of
Dorsal. However, the rho CRM contains one high-affinity site and a few
low-affinity sites, which allow both high and intermediate levels of Dorsal to
activate. Finally, the sog CRM contains four evenly-spaced optimal sites,
which allows lowest levels of Dorsal to activate transcription. The combinatory role with other factors is also involved in regulating Dorsal target genes.
Twist protein regulates nearly half of the known Dorsal target genes synergistically once it is synthesized in the presumptive mesoderm. Snail also
regulates a large number of Dorsal target genes, but functions as a repressor
(Leptin, 1991). In this way, the broadly distributed activator and the localized repressor set up restricted expression of their target genes.
Dorsal protein can repress transcription as well (Jiang et al., 1993; Pan
and Courey, 1992). Some genes, like zerknullt (zen) and decapentaplegic
(dpp), are expressed in the dorsal portion of the embryo and are required for
the establishment of the amnioserosa and the dorsal epidermis. They are
repressed by Dorsal in the ventrolateral and ventral cells. The CRMs of these
genes contain not only high affinity Dorsal binding sites but also AT-rich
sites which are bound by the Cut and Dead-ringer proteins (Chen et al.,
1998). These proteins cooperatively recruit the Groucho co-repressor, mediating transcriptional repression (Valentine et al., 1998). Another factor,
Capicua, was also shown to be involved in the repression of zen (Jimenez et
al., 2000).
Snail in gastrulation and mesoderm formation
Gastrulation is the first morphogenetic process in embryonic development,
during which cells of the blastula undergo rearrangement and move to the
correct positions to set up the three germ layers (ectoderm, mesoderm and
endoderm). This requires the coordination of cell-fate determination, cellcycle control, individual cell-shape changes, and movement of groups of
cells (Ip and Gridley, 2002; Leptin, 1999; Leptin et al., 1992). During cellularization, the expression of Snail is sharply restricted to the 18 cell-width
ventral domain mainly due to the cooperative function between Dorsal and
Twist. These Snail-expressing cells correspond precisely to the presumptive
mesoderm. Either expanding or reducing the expression of Snail correlates
with changes in ventral cell invagination and the formation of future mesoderm (Leptin et al., 1992). Genetic rescue by Snail in a twist mutant can
promote ventral-cell invagination but not vice versa, indicating Snail has a
more direct role in regulating downstream events leading to gastrulation (Ip
et al., 1994). It was proposed that Snail can regulate two separate sets of
target genes, based on the fact that some snail hypomorphic mutants show
16
phenotypes on derepression of neuroectodermal genes and mesoderm differentiation, but not on ventral cell invagination. Snail represses neuroectodermal genes such as single-minded (sim), rho, brinker (brk), lethal of scute and
so on by direct binding to their enhancers. And it also promotes the expression of mesodermal genes such as dGATAb and zfh-1 to help the establishment of the mesodermal cell fate. The second set of genes (e.g. folded gastrulation) control the cell shape changes and cell movements during gastrulation (reviewed in Ip and Gridley, 2002). However, deregulation of individual Snail target genes does not interfere with mesoderm cell fate or
invagination, suggesting that the cumulative effect of regulating all these
genes is essential for gastrulation (Hemavathy et al., 1997).
The molecular function of the Snail repressor depends on two functional
domains, a zinc-finger DNA-binding domain in the C-terminus and a repression domain in the N-terminus (Boulay et al., 1987; Hemavathy et al., 1997).
The repression domain contains two conserved motifs that allow interaction
with the co-repressor dCtBP (Drosophila homolog of C-terminal binding
protein) (Nibu et al., 1998a). Our studies demonstrated that the full repression activity of Snail requires another co-repressor, Ebi, which interacts with
Snail through a conserved motif in the N-terminus of Snail protein (Paper
III).
Anterior-posterior patterning
The Drosophila larva has a very obvious feature, the regular segmentaion of
the larval cuticle along the AP axis, each segment carrying cuticular structures that define it as, for example, thorax or abdomen (Figure 2B). Similar
to DV patterning, the determination of AP axis is initiated by maternal factors and then carried out by a segmentation hierarchy of zygotic transcription
factors (reviewed in Niessing et al., 1997). The whole procedure requires
three classes of maternal determinants in the egg that control anterior patterning, posterior patterning and terminal patterning of the embryo respectively (reviewed in St Johnston and Nusslein-Volhard, 1992). The hierarchy
produces broad bands of gap gene expression, and the striped patterns of
pair-rule genes that subsequently control the segmental organization determined by segment polarity genes (Figure 3). By this time, the cell fates can
be determined at single-cell resolution along the AP axis.
Bicoid is the maternal determinant that patterns the head and the thorax
of the embryo. Bicoid protein forms gradient from anterior to posterior. Another maternal factor, Caudal, patterns the posterior part of the embryo. A
Caudal protein gradient is established from posterior to anterior. Bicoid and
Caudal initiate the activation of zygotic gap genes including orthodenticle,
hunchback (hb), Kruppel (Kr), knirps (kni) and giant, all of which code for
transcription factors (Ip et al., 1992; Lebrecht et al., 2005; Ochoa-Espinosa
et al., 2005; Rivera-Pomar et al., 1995). A steep gradient of Hb protein is
17
established from anterior to posterior. The Hb gradient, together with Bicoid,
activates or represses other gap genes. Interactions between gap genes also
help to sharpen and establish their border of expression. For example, the
anterior border of kni is set by Hb, and its posterior border is limited by Giant and Tailless (Tll) (Rivera-Pomar 1995). The anterior boundary of Kr
expression is repressed by Hb, but its posterior boundary repressed by Kni,
Giant and Tll (Hoch et al., 1992). The expression of tll is not repressed by
other gap genes. Instead, it is a downstream gene in response to Torso
signaling.
Bcd
Cad
Maternal effects
Gap genes
gt, Kr, kni, tll etc
Pair-rule genes
eve, ftz, odd, hairy, etc
Segment polarity genes
en, wg, hh, etc
Figure 3. Segmentation is achieved by a hierarchy of maternal and zygotic transcription factors. Bicoid (Bcd) and Caudal (Cad) form morphogen gradients
with highest level at anterior and posterior respectively. They activate the first
set of zygotic genes, the gap genes, which are expressed in rather broad regions.
Gap gene products define the periodical expression patterns of pair-rule genes,
whose products subsequently determine the expression of segment polarity
genes. Pair-rule genes are expressed in seven stripes, wheras segment polarity
genes are expressed in fourteen stripes.
The terminal regions are specified by a third group of maternal genes in
which a key factor is called Torso. Torso is a receptor tyrosine kinase uniformly distributed throughout the fertilized egg plasma membrane. But it is
only activated at the two ends of the embryo through binding to its ligand.
This signal induces the activation of zygotic genes, such as tll and huckebein,
at both poles, thus defining the two extremities of the embryo (reviewed in
Niessing et al., 1997).
Different concentrations and combinations of the gap genes set up the localized striped pattern of pair-rule genes (Jackle et al., 1992; Niessing et al.,
18
1997), including even-skipped (eve) and hairy. Pair-rule genes are expressed
in seven stripes, with a periodicity corresponding to alternate parasegments.
For example, eve defines odd-numbered parasegments, whereas ftz defines
even-numbered parasgments. The striped expression of primary pair-rule
genes requires the regulation by separate CRMs, working independently in
the regulatory region (Small et al., 1992). Each CRM contains multiple binding sites for both activators and repressors. A famous example is the regulation of eve whose regulatory region contains 5 separate CRMs, together producing the seven stripes. This enhancer autonomy is partially due to shortrange repression, meaning that a repressor bound to one enhancer does not
interfere with another (more discussion later). Pair-rule genes also regulate
the expression of each other, thereby establishing a refinement of the stripe
pattern (reviewed in Niessing et al., 1997).
The segment-polarity genes are activated by products of pair-rule genes
and are involved in stablizing the parasegment boundary and setting up a
signaling center at the boundary that patterns the segments (Nasiadka et al.,
2000; Nasiadka and Krause, 1999). Well-studied segment polarity genes
include engrailed (en), wingless and hedgehog. En is a transcription factor,
whereas Wingless and Hedgehog are signaling molecules.
The segment identity is specified by homeotic selector genes (Homeotic
genes). The two Homeotic gene clusters in Drosophila are called the Bithorax and Antennapedia gene complexes (Regulski et al., 1985; Wedeen et al.,
1986). The Bithorax complex is responsible for diversification of the posterior segments 5-12 and the Antennapedia complex for that of the anterior
segments 1-5. These genes are activated by gap genes and pair-rule genes at
an early embryonic stage but their expression pattern are maintained by
Polycomb Group genes (PcG) and Trithorax Group (trxG) genes later in
development (Breen and Harte, 1991; Harding and Levine, 1988; LaJeunesse
and Shearn, 1995; Simon et al., 1992).
19
Histone modifications
Histone modifications play important roles in different biological processes
including transcription regulation, DNA repair, replication and heterochromatin formation (reviewed in Kouzarides, 2007). Their functions depend on
the type of chemical modifications, the location in the histone octamer and
even the combinatorial effects of multiple modifications. The classic modifications include lysine acetylation, lysine and arginine methylation, serine
phosphorylation and lysine ubiquitylation. Recent studies have discovered
more modifications that include lysine sumoylation, ADP ribosylation, arginine deimination and proline isomeration (reviewed in Kouzarides, 2007).
The possible sites for modification in histones are numerous (Figure 4), and
the forms of modification are also various such as mono-, di-, or trimethyl
for lysines and mono- or di- (asymmetric or symmetric) for arginines. Most
modification sites were identified in the flexible N-terminal tails of histones,
such as histone H3 Lysine 4 (H3K4), H3K9, H3K14, H4K5, H4K8, and so
on (Figure 4). Interestingly, recent studies suggested that residues within
core histones can also be modified (e.g., H3K56 and H3K79) (Xu 2005).
Furthermore, most of these modifications, such as acetylation, phosphorylation and methylation, are all reversible. This fact of complexity gives histone
modifications enormous potential for functional responses.
P
Ac
U
K
36
K
119
Ac Ac Ac Ac
H2A
S
1
K
5
M
K K K
9 13 15
Ac Ac P Ac
Ac
M Ac
U
K
5
K
20
K K
23 24
K
120
M P M Ac Ac P Ac
Ac M Ac
Ac M M P
Is
M M Is Ac
R T K
2 3 4
K R K
K R K S
14 17 18 23 26 27 28
M
Ac
Ac
P
30
K K P K
36 37 38 56
H2B
K S K
12 14 15
M
M
H3
RK S T
8 9 10 11
M
P M Ac
Ac
Ac
S
1
K
8
K
12
H4
R
3
K
5
K
16
K
20
Figure 4. Most of histone modifications, including lysine methylation, arginine methylation, lysine acetylation, serine phosphorylation, lysine ubiquitylation and
proline isomeration on histone H2A, H2B, H3 and H4, are depicted.
20
Functions of histone modifications
There are two mechanisms by which histone modifications contribute to
chromatin function. First, some modifications (e.g. acetylation) reduce the
positive charge of histones, in principle resulting in the alteration of the
chromatin structure directly. Second, the modifications can generate binding
surfaces for non-histone proteins. Specifically, methylated lysines can be
recognized and bound by chromo-like domains (chromo, tudor, MBT) and
PHD domains, acetylated lysines can be recognized by bromodomains, and
phosphorylation is recognized by a domain within 14-3-3 proteins (Figure 5).
These nonhistone proteins often possess enzymatic activities which can further modify chromatin. For example, many histone acetyltransferases (HATs)
contain a bromodomain, facilitating the maintenance of the acetylated chromatin by further modifying regions that are already acetylated. Similarly,
histone methylases (HMTs) often contain a chromodomain, which is important for the maintenance of methylated nucleosomes. Other proteins containing these domains may not be histone modifying enzymes but instead deliver
enzymes to histones. For example, the Polycomb protein binds to H3K27me
through its chromodomain and is associated with ubiquitin ligase (Ring1A)
activity specific for H2A (de Napoles et al., 2004; Fang et al., 2004). Heterochromatin protein1 (HP1) binds H3K9me and is associated with
Su(var)3-9, a HMT for H3K9. It was also suggested that histone modifications can prevent the targeting of nonhistone protein onto chromatin
(Margueron et al., 2005). Moreover, one modification, can affect another
modification, positively or negatively. The phosphorylation of H3S10 disrupts the binding of HP1 to methylated H3K9 (Fischle et al., 2005) but facilitates the binding of GCN5 acetyltransferase to acetylated H3K9
(Clements et al., 2003). The effect of histone modifications seems to be context dependent. Methylation of H3K36 has a positive effect when it is present in the coding region and a negative effect when in the promoter (Landry
et al., 2003; Li et al., 2007a; Strahl et al., 2002).
Bdf1
H4
Bromo
Ac
K
8
Brd2
Taf1
Bromo
Bromo
Ac
K
12
Ac
K
16
JMJD2A CRB2
Chromo
Me
Tudor
Me
K
36
Pc
Rsc4
14-3-3
Chromo Bromo
Me
K
27
HP1
Chromo
Ac
P
K
14
S
10
Me
K
9
Chromo
Me
PH
D
BPTF
TAF3
K
20
CHD1
K
4
M
BT
Eaf3
H3
L(3)MBTL
ING2
Figure 5. The known modification sites for protein domains in non-histone proteins.
Proteins containing the bromodomain bind to acetylated lysine. Proteins containing
a chromodomain, Tudor domain, PHD domain or MBT domain bind to lysine methylation. 14-3-3 protein binds to phosphorylated serine.
21
The number, variety and interdependence of histone modifications led to the
histone code hypothesis, multiple histone modifications, acting in a combinatorial or sequential fashion on one or multiple histone tails, specify unique
downstream functions (Strahl and Allis, 2000). It has provided a framework
for the interpretation of many discoveries regarding the mechanisms by
which chromatin functions. However, the fact that the function of histone
modifications is highly context dependent suggests the code cannot be universal.
Histone acetylation and its function
Histone acetylation is one of the best-characterized histone modifications, an
event with a big impact on chromatin structure and function. It is possibly
involved in all chromatin-based processes, such as gene transcription, gene
silencing, DNA repair, DNA replication and chromosome condensation.
Acetylation can occur on all core histones (Figure 4) and is carried out by
HATs, which catalyze the transfer of an acetyl group from acetyl-CoA to the
lysine -amino groups on the N-terminal tails of histones. This process is
reversible. Histone deacetylases (HDACs) remove the acetyl group from
acetylated lysines.
Histone acetylation in gene regulation
The correlation between histone acetylation and transcription activation has
been established for a long time based on several facts. First, acetylation has
the most potential to unfold chromatin by neutralizing the basic charge of the
lysine, which makes DNA more accessible for targeting. Second, many
HATs were identified as co-activators, such as GCN5 and CBP/P300
(Bantignies et al., 2000; Brownell et al., 1996). Third, the actively transcribed regions tend to be hyperacetylated, whereas transcriptionally silent
regions tend to be hypoacetylated. The most-characterized sites are within
the N-terminal tail of the histones, such as H3K9, H3K14, H4K5, H4K8,
H4K12, and H4K16 (Figure 4). Some other sites were recently identified,
such as H3K18, whose pattern tightly correlates with gene activation in prostate cancer tissue (Seligson et al., 2005).
Histone deacetylation makes chromatin more compacted and generally
correlates with transcription repression. Consistently, many HDACs are part
of corepressor complexes such as Rpd3 (reduced potassium dependency 3)
in the Sin3-Rpd3 complex and HDAC3 in the NCoR/SMRT complex. However, recent evidence, that hypoacetylation of H4K16 and H2BK11/16 is
linked to transcription activation, challenges this over-simplified view
(Kurdistani et al., 2004; Wiren et al., 2005).
22
Histone acetylation/deacetylation also regulates gene expression through
gene silencing which involves heterochromatin establishment and DNA methylation. Histone deacetylation could be involved in both cases. In budding
yeast, HDACs are recruited to unacetylated histones in silenced regions, thus
reinforcing and maintaining the deacetylated state in heterochromatin
(Braunstein et al., 1996). How heterochromatin is established is not fully
understood. However, it has been shown that the involvement of H3K9 methylation and heterochromatin protein 1 (HP1) is important. Additionally, it
may involve the changes of several other histone marks (reviewed in Ebert et
al., 2006), such as H3K4 demethylation (Rudolph et al., 2007), H3S10
dephosphorylation (Zhang et al., 2006c), H3K9 deacetylation, H4K12 acetylation (Swaminathan et al., 2005). In mammalian cells, HDACs can be recruited to methyl groups attached to the cytosine residues within CpG islands via proteins such as methyl-CpG binding proteins and methyl-CpGbinding-domain-containing proteins, or via the DNA methylases, resulting in
tightly-packed chromatin and gene silencing.
Histone acetylation in DNA repair
DNA double strand breaks (DSBs) cause genetic instabilities and gene mutations if they are not accurately repaired. The response of eukaryotic cells to
DSBs is highly conserved and involves both checkpoint functions that arrest
the cell cycle, allowing time for repair to occur, and repair functions that
directly fix the break (reviewed in Khanna and Jackson, 2001). Defects in
DNA repair result in the accumulation of DNA damage and consequently
lead to apoptosis or enhanced carcinogenesis. DSBs are repaired through two
major pathways, homologous recombination (HR) and non-homologous end
joining (NHEJ) (reviewed in Bernstein et al., 2002; Khanna and Jackson,
2001). The DNA damage response pathway includes a series of events: detection by sensor proteins, delivery by transducer proteins and downstream
actions by effecter proteins (Figure 6A). Ataxia-telangiectasia mutated
(ATM) and ATM and Rad-3 related (ATR) kinases are transducer proteins
and belong to the phosphoinositide 3-kinase related kinase family (PIKK).
Upon DNA damage, ATM and ATR get auto-phosphorylated and activate
the second group of kinases, Chk2 and Chk1, which subsequently phosphorylate effecter proteins. ATM is actively involved in DNA repair, checkpoint response and apoptosis through modifying some key factors in these
processes (e.g. BRCA1, Chk2, p53) (Banin et al., 1998; Cortez et al., 1999;
Matsuoka et al., 1998). Chk2 can also phosphorylate the signal effector, p53,
to induce apoptosis and G1/S checkpoint (Hirao et al., 2000). p53 is a tumor
suppressor gene which is mutated in more than half of all human cancers
(Greenblatt et al., 1994). The phosphorylation stabilizes and stimulates p53
activity. Once activated, it functions as a transcription factor and regulates
23
expression of genes which are involved in apoptosis and in DNA repair
(reviewed in Oren, 2003). A p53 homolog is absent in yeast. The Drosophila
homolog of p53 is only involved in apoptosis but not in cycle arrest upon
DNA damage (Figure 6B) (Brodsky et al., 2004; Lee et al., 2003). Another
early response to DSBs is the accumulation and the spreading of phosphorylated histone variant H2AX (fly H2Av) and yeast H2A around the break site.
The phosphorylation is possibly mediated by ATM as well (Rogakou et al.,
1998; van Attikum and Gasser, 2005).
IR
B
A
DSB
DSB
Sensors
P
Rad17, Rad9, Rad1, Hus1,
MRN complex
P
Chk2
P
Effectors
p53
P
Mei-41(ATR)
dATM
Transducers ATM, ATR, DNA-PK
CDC25
H2Av
P
Mus304
P
Grp(Chk1)
P
P53
reaper, grim, hid, skl
Cell cycle arrest
DNA repair
Apoptosis
Apoptosis
G2 arrest
Figure 6. A. The general organization of the DNA-damage response pathway. It is
composed of sensor proteins, transducer proteins and effector proteins. The effectors subsequently activate signaling pathways for DNA repair, cell cycle arrest and
apoptosis. B. IR induced G2 arrest and apoptosis in Drosophila. Cell cycle arrest
and apoptosis pathways are rather independent. IR induced apoptosis is initiated by
dATM. dATM gets auto-phosphorylated, which subsequently activates Chk2.
dATM and Chk2 activate p53 by phosphorylation. p53 activates its target genes,
such as reaper, grim, hid (head involution defect) and skl (sickle), whose activities
are responsible for apoptosis. In parallel, G2 arrest is prominently implemented by
Mei-41 (dATR), Mus 304 (dATRIP) and Grp (Chk1). dATM and Chk2 may also
play roles in the initiation of G2 arrest.
Chromatin dynamics need to be adjusted to facilitate the recruitment of the
repair machinery to the break sites. Chromatin remodeling and histone modifications can make the lesion more accessible for damage response machinery (reviewed in Peterson and Cote, 2004; van Attikum and Gasser, 2005;
Wurtele and Verreault, 2006). It was suggested that chromatin remodeling
complexes including the RSC (remodels the structure of chromatin) complex,
the SWI/SNF complex and the INO80 complex play roles at multiple steps
during the detection and repair of DSBs (reviewed in Downs et al., 2007). In
addition to the phosphorylation of H2AX (H2Av in fly, H2A in yeast), several other histone modifications are involved in DNA repair (Peterson and
24
Cote, 2004; van Attikum and Gasser, 2005; Wurtele and Verreault, 2006).
The acetylation of conserved lysine residues in the N-terminal tails of H3
and H4 are important for DNA-damage responses (Bird et al., 2002; Downs
et al., 2007; Tamburini and Tyler, 2005). In yeast, histone H3 acetylation by
the Hat1(Qin and Parthun, 2006) complex and Gcn5 (Teng et al., 2002) and
H4 by the NuA4 (nucleosome acetyltransferase of H4) complex is impotant
for DNA repair (Bird et al., 2002). In mammalian cells, the Tip60 HAT
complex is recruited to sites of DSBs, where they induce H4 acetylation and
HR (Murr et al., 2006). The Drosophila Tip60 complex specifically acetylates phosphorylated H2Av, which is a prerequisite for the exchange with
unmodified H2A during DNA repair (Kusch et al., 2004). Not only was the
initial recruitment of HATs observed on DSBs, but also the subsequent association of several HDACs with chromatin near DSBs (Tamburini and Tyler,
2005). Possibly HDACs are required for the restoration of chromatin higherorder structure following DSB repair.
Histone acetyl-transferase complexes
The histone acetyltransferases are divided into five families (Figure 7), including the Gcn5-related N-acetyltransferases (GNATs); the MYST (for
MOZ, Ybf2/Sas3, Sas2 and Tip60)-related HATs; p300/CBP HATs; the
general transcription factor HATs, and the nuclear hormone-related HATs
SRC1 and ACTR (SRC3) (reviewed in Roth et al., 2001).
HAT
HDAC
Five families: GCN5
Three types:
MYST
Type I:
HDAC1, 2, 3, 8
Type II:
HDAC4, 5, 7, 9
HDAC6, 10
CBP
SRC-1
TAF1
Type III: Sir2 family
Figure 7. Histone acetyl transferases and histone deacetylases. Histone acetyl transferases are classified into five families. Histone deacetylases are classified into three
families. Histone acetylation generally correlates with more opened chromatin which
facilitates transcription. By contrast, deacetylation state of histone results in more
closed chromatin which inhibits transcription.
25
GNAT complexes
All the GNAT complexes contain HATs that show sequence and structural
similarity to Gcn5 (reviewed in Roth et al., 2001). They commonly share
two functional conserved domains: a HAT domain that encompasses three
regions of conserved amino acid sequence spanning approximately eighty
residues, and a C-terminal bromodomain that is believed to interact with
acetylated lysines. The metazoan members of the family share an additional
domain which is not found in yeast, the N-terminal PCAF region (reviewed
in Carrozza et al., 2003). Gcn5 relatives have been identified in virtually all
eukaryotes. Besides Gcn5 homologs, this family also includes the chromatinassembly-related Hat1, the elongator complex subunit Elp3, the mediator
complex subunits Nut1 and Hpa2.
Recombinant Gcn5 mainly acetylates free histone H3, but exhibits little
HAT activity when using nucleosomal histones as substrates, suggesting that
incorporation into complexes is required for Gcn5 to obtain the capacity to
acetylate nucleosomes. The most characterized GNAT complexes are yeast
SAGA (for Spt-Ada-Gcn5-acetyltransferase) and the ADA complex. Gcn5
and the accessory proteins Ada2 and Ada3 form the catalytic core of these
complexes and enable them to acetylate nucleosomes. Previous data also
indicated that incorporation of Gcn5 into complexes alters lysine specificity.
Gcn5 alone acetylates mainly H3K14 on free histones, but SAGA acetylates
lysines 9, 14, 18 and 23, and ADA acetylates 9, 14 and 18 (Grant et al.,
1999).
GNAT complexes were identified from different organisms and they have
similar composition and functions, but the complexes from higher metazoans
are more diverse than those in yeast. One Drosophila SAGA-like complex
has been identified. Three known mammalian SAGA-like complexes are
called PCAF, STAGA and TFTC (Ogryzko, 2000). The SAGA-like complexes basically contain: (i) Proteins for HAT activity (Ada2, Ada3, and
Ada4/Gcn5) and complex assembly (Ada1 and Ada5/Spt20), which were
isolated in a genetic screen as proteins interacting with the activators yeast
Gcn4 and the herpes simplex virus activation domain VP16; (ii) Proteins for
TBP binding, the Spt proteins, (Spt3, Spt7, and Spt8), which were initially
identified as suppressors of transcription initiation defects caused by promoter insertions of the Ty transposable element. (iii) A subset of TBPassociated factors (TAFs) (TAF5, TAF6, TAF9, TAF10 and TAF12). (iv)
Activator interaction protein Tra1, an ATM/PI-3 kinase-related homologue
of the human TRRAP protein and (v) proteins for deubiquitylation (Ubp8
and Sgf11), mediating the deubiquitylation of H2B (Shukla et al., 2006).
Several other SAGA-associated proteins were identified and characterized
recently. Sgf73p facilitates formation of the preinitiation complex assembly
at promoters (Shukla et al., 2006). The mRNA export factor Sus1 is involved
in SAGA-mediated H2B deubiquitinylation through its interaction with
26
Ubp8 and Sgf11 (Kohler et al., 2006). Both normal and polyglutamine- expanded ataxin-7 are components of TFTC-type GCN5 histone acetyltransferase- containing complexes (Helmlinger et al., 2006). SAGA-related complexes can function as co-activators through the interactions with TBP and
acidic activators, such as GCN4, Pho4 and Gal4 (Barbaric et al., 2003; Kuo
et al., 2000; Larschan and Winston, 2001). Recent studies by Govind et al.
suggested that SAGA also occupies coding sequences and contributes to H3
acetylation, H3K4 methylation and even nucleosome eviction (Govind et al.,
2007). Intriguingly, the SAGA complex has been implicated in transcriptional repression by the Arg/Mcm1 repressor complex (Ricci et al., 2002).
Mammalian TFTC complex exhibits increased binding to UV-damaged
DNA in vitro, implicating the function of GNAT complexes in DNA repair
(Brand et al., 2001).
GCN5 related proteins play critical roles during development. GCN5-null
mice are embryonic lethal with the lack of somite formation. The deletion of
PCAF, however, does not cause lethality, presumably due to compensation
by GCN5. Animals carrying deletions in both genes die earlier than GCN5null animals, indicating that PCAF does have distinct functions from GCN5
in early development (Xu et al., 2000; Yamauchi et al., 2000). dGCN5 null
mutants in Drosophila have defects in both oogenesis and metamorphosis,
while hypomorphic alleles of dGCN5 affect the formation of adult appendages and cuticle (Carre et al., 2005). Mutations in Arabidopsis GCN5 also
result in developmental defects (Vlachonasios et al., 2003). GCN5 possesses
both HAT-dependent and independent functions, as indicated by the fact that
GCN5 null mice die much earlier with increased apoptosis than GCN5 HAT
mutant mice with defects in neural tube closure but with no abnormal apoptosis (Bu et al., 2007). GCN5 also targets non-histone proteins as substrates
which are involved in diverse cellular processes. For example, PCAF acetylates high mobility group protein (HMG)-A1 that positively regulates transcription of the interferon gene (Munshi et al., 2001), the oncoprotein cMyc, and Ku70 that is involved in DNA repair through the non-homologous
end-joining (NHEJ) pathway (reviewed in Glozak et al., 2005).
Ada2 proteins
The Ada2 protein was first identified from yeast, in which its inactivation
could relieve the toxicity caused by overexpression of the strong transcriptional activation domain in the viral VP16 protein (Berger et al., 1992). Subsequently, Ada2 proteins were shown to exist in all known Gcn5 containing
complexes and to be required for the assembly of these complexes. In contrast with the single Ada2 gene present in Saccharomyces cerevisiae, the
Arabidopsis thaliana, fly and human genome all contain two genes, encoding two Ada2 proteins, Ada2a and Ada2b (Barlev et al., 2003; Kusch et al.,
2003; Muratoglu et al., 2003; Stockinger et al., 2001). However, Ada2a and
Ada2b are present in different complexes which possess different functional
27
activities. For example, in Drosophila, the SAGA-like complex contains
dAda2b and acetylates histone H3K9 and H3K14 (Pankotai et al., 2005; Qi
et al., 2004). However, the ATAC (Ada Two A Containing) complex can
acetylate histone H4K5 and H4K12 (Guelman et al., 2006; Pankotai et al.,
2005). Ada2 homologues share several conserved regions including a ZZ
domain, a SANT domain and three ADA boxes (Kusch et al., 2003). The ZZ
domain is a potential zinc-binding motif and is required for the interaction
with Gcn5 (Candau and Berger, 1996). The SANT (SWI3, Ada2, N-CoR and
TFIIIB) domain of Ada2 is able to associate with chromatin and is essential
for the catalytic activity of Gcn5 (Boyer et al., 2002).
Tip60 HAT complexes
Tip60 (Tat-interactive protein 60kDa) is one of the best characterised MYST
proteins whose family members function in a broad range of biological processes, such as gene regulation, dosage compensation, DNA damage repair
and tumourigenesis (reviewed in Utley and Cote, 2003). Tip60 homologues
have been identified in various organisms from yeast to human. They share
at least two functional domains, an N-terminal chromodomain and a Cterminal MYST catalytic domain. Recombinant Tip60 acetylates core histones H2A, H3 and H4 in vitro (Kimura and Horikoshi, 1998; Yamamoto
and Horikoshi, 1997). When assembled in multiprotein complex, Tip60 can
modify nucleosomal H2A and H4 (Ikura et al., 2000). Tip60, like GCN5 and
CBP, can also acetylate non-histone proteins, including androgen receptor,
upstream binding transcription factor, c-Myc, forkhead family protein
FOXP3 and so on (Li et al., 2007a; Sapountzi et al., 2006).
Tip60
Tip60
E(Pc)
E(Pc) ING3
ING3
TRRAP
TRRAP
Domino
Domino
DMAP1 Reptin Pontin
BAF53
BAF53 DMAP1 Reptin Pontin
Eaf6
Actin
Brd8
Actin Eaf6
Brd8
GAS41
GAS41 MRG15
MRG15
Figure 8. The composition of Tip60 complex. Tip60, E(Pc) and ING3 constitutes a
core enzymatic sub-complex.
The composition of the Tip60 complex is conserved between Drosophila
and human. It was suggested that the metazoan Tip60 complex is a hybrid of
the yeast NuA4 and SWR1 complexes (Doyon et al., 2004). Three subunits,
Tip60 (Esa1), EPC1 (yeast homolog EPL1, Drosophila homolog E(Pc) (Enhancer of Polycomb)) and ING3 (yeast Yng2, Drosophila dIng3) form a core
28
enzymatic subcomplex and exist as a distinct entity responsible for the
global non-targeted acetylation of chromatin in vivo (Boudreault et al., 2003).
Drosophila E(Pc) is a suppressor of position-effect variegation. Domino/p400 (Swr1), an ATPase possessing chromatin remodeling activity
(Mizuguchi et al., 2004), is able to exchange phosphorylated H2Av for unmodified H2Av after DNA damage (Kusch et al., 2004). Mortality factor 4
related gene 15 (MRG15) (Eaf3) contains a chromodomain and can recruit
HDAC1 to chromatin in the Rpd3S complex (Carrozza et al., 2005). The
TRRAP (transformation/transcription domain-associated protein, yeast Tra1)
protein is a common subunit in SAGA-related complexes and interacts with
transcription factors. Pontin (Tip49a, RuvBL1) and Reptin (Tip49b, RuvBL2)
are putative helicases, related to the bacterial repair protein RuvB (reviewed
in Gallant, 2007).
Previous studies have indicated two major aspects of roles of the Tip60
complexes, in transcription and in DNA DSBs response. In a lot of cases,
Tip60 was associated with transcription activation. Examples of Tip60coactivated transcription factors are androgen receptor (AR) (Gaughan et al.,
2002), NF- B (Baek et al., 2002; Kim et al., 2005), c-myc (Patel et al., 2004),
E2F1 (Taubert et al., 2004), p53 (Doyon et al., 2004) and so on. However,
Tip60 was also shown to have transcriptional repression activity. For example, Tip60 corepresses CREB (cAMP response element binding protein) by
direct binding to CREB and repressing CREB phosphorylation by protein
kinase A (Gavaravarapu and Kamine, 2000). Tip60 is believed to recruit
HDAC7 to STAT3 (signal transducer and activator of transcription 3) regulated promoters and thus represses transcription (Xiao et al., 2003). Tip60
promotes the acetylation of FOXP3 protein and interacts with HDAC7,
which are both required for the repressor function (Li et al., 2007b). The role
of Tip60 complexes in DNA repair has also been well studied. In yeast cells,
after DNA damage NuA4 complex is recruited to the vicinity of DNA lesions, which promotes the recruitment of the chromatin remodeling complexes INO80 and SWR1 (Downs et al., 2004). In Drosophila, Tip60 acetylates phosphorylated H2Av and Domino exchanges the acetylated and phosphorylated H2Av with unmodified H2Av, which is required for DSB repair
(Kusch et al., 2004). In mammalian cells, the Tip60 complex is involved in
the DNA damage-induced H4 hyperacetylation and homologous recombination (HR) (Ikura et al., 2000; Murr et al., 2006). In a word, this subfamily of
MYST complexes play multiple roles in a diverse variety of cellular pathways, involving not only normal cellular conditions but also abnormal processes including viral infection, neurodegernerative disease and cancer
(reviewed in Sapountzi et al., 2006).
Reptin
Reptin and the related Pontin protein are members of the AAA+ family of
helicases (ATPases associated with diverse cellular activities) (reviewed in
29
Ogura and Wilkinson, 2001). Reptin and Pontin can form a dodecamer consisting of two differently shaped, juxtaposed hexamers (Puri et al., 2007).
Their ATPase activities are interdependent as either protein alone is almost
inactive (Makino et al., 1999). Reptin and Pontin were found to be integral
subunits of different chromatin-modifying complexes, including the Tip60
complex, the INO80 complex and the Swr1 (SRCAP in metazoan) complex
(Kobor et al., 2004; Krogan et al., 2003; Mizuguchi et al., 2004; Shen et al.,
2000). Drosophila Reptin was co-purified with Polycomb repressive complex 1 (PRC1) which maintains the repressive state of HOX genes during
development (Saurin et al., 2001). Integration in these complexes indicates
potential roles of Reptin and/or Pontin in diverse cellular processes, including the response to DNA double-strand breaks and transcription regulation
(reviewed in Gallant, 2007). Reptin and Pontin were also identified by their
physical interaction with the transcription-associated protein -catenin
(Bauer et al., 2000), with the transcription factors TBP (Bauer et al., 1998;
Ohdate et al., 2003), Myc (Wood et al., 2000), E2F1 (Dugan et al., 2002)
and ATF2 (Activating transcription factor 2) (only Reptin) (Cho et al., 2001).
Along with components of the Tip60 complex, Reptin and/or Pontin were
shown to bind to the promoters of transcriptional targets of c-myc, E2F1 and
NF- B (Frank et al., 2003; Kim et al., 2005; Taubert et al., 2004). Interestingly, Reptin and Pontin may play similar or antagonistic roles in a contextdependent manner. For example, both Reptin and Pontin enhance the repression function of c-myc in the control of cell proliferation in Drosophila and
Xenopus (Bellosta et al., 2005; Etard et al., 2005). However, in the cases of
-catenin and NF- B, they have opposite activities. Drosophila and zebrafish Reptin act as transcriptional repressors , whereas Pontin are activators in
-Catenin/Wnt signaling (Bauer et al., 2000; Rottbauer et al., 2002). Similar
activities were observed in a human cell line in the regulation of a NF- B
p50 target, KAI1(Kim et al., 2005). KAI1 is a tetraspanin protein inhibiting
tumor metastasis. At the regulatory region of KAI1, p50 recruits the
Tip60/Pontin coactivator for transcriptional activation, whereas it recruits the
-catenin/Reptin corepressor for repression. In metastatic cells, KAI1 is
downregulated due to increased -catenin and decreased Tip60. Reptin/ catenin mediated repression partially depends on HDAC1. Reptin is sumoylated on lysine 456. It was suggested that sumoylation of Reptin stimulates
the repressive potential of Reptin by promoting the nuclear localization of
Reptin and by increasing its interaction with HDAC1 (Kim et al., 2006).
These studies indicate an additional function of Reptin independent of the
Tip60 complex.
30
Histone deacetylases and their associated complexes
The histone deacetylases (HDACs) are conserved proteins in different organisms. They are classified into three subfamilies (Figure 7): class I, class II
and Sir2 related HDACs, based on their protein sequence similarity to yeast
Rpd3, HDA1 and Sir2 respectively. Class I and class II proteins are evolutionarily related and share a common enzymatic mechanism, the zinccatalyzed hydrolysis of the acetyl-lysine amide bond. Class III proteins are
functionally different and catalyze the transfer of the acetyl group onto the
sugar moiety of NAD (reviewed in Blander and Guarente, 2004). HDACs
have diverse functions through multiple mechanisms: the incorporation into
different complexes, the modulation of deacetylase activity by proteinprotein interactions or by post-translational modifications, tissue specific
pattern, variable subcellular localizations and splice variants (reviewed in de
Ruijter et al., 2003). The substrates of HDACs are not only histones, but also
non-histone proteins, including p53, E2F, -tubulin, MyoD and MEF2
(Gregoire et al., 2007; Hubbert et al., 2002; Juan et al., 2000).
Class I HDACs include HDAC1 (Rpd3), 2, 3 and 8. HDAC1 and HDAC2
are closely related and present in common co-repressor complexes, such as
Sin3, NuRD (nucleosome remodeling and deacetylating) and Co-REST
complexes (Kasten et al., 1997; You et al., 2001; Zhang et al., 1999). These
complexes are widely involved in transcription regulation and developmental control. HDAC1 and HDAC2 are also regulated by phosphorylation,
which promotes both deacetylase activity and complex formation
(Galasinski et al., 2002). HDAC3, however, is stably associated with
NCoR/SMRT (nuclear receptor corepressor/ silencing mediator for retinoic
acid and thyroid hormone receptors) corepressor complexes. HDAC8 was
recently identified and its function remains to be discovered (Buggy et al.,
2000).
Class II HDACs include HDAC4, 5, 6, 7, 9 and 10. The subgroup containing HDAC4, 5, 7 and 9 is able to shuttle in and out of the nucleus in response to certain cellular signals (reviewed in Verdin et al., 2003). Upon
phosphorylation by several kinases, they are confined to the cytosol by interaction with 14-3-3 proteins. In the nucleus they associate with transcription
factors, for example the MEF and Runx families (Vega et al., 2004; Zhang et
al., 2002a). HDAC6 is able to deacetylate Tubulin and Hsp90, suggesting its
special role in cell motility and protein folding (Hubbert et al., 2002; Kawaguchi et al., 2003; Matsuyama et al., 2002). HDAC10 is the most recently
discovered member of the class II HDACs. It was suggested that it associates
with many other HDACs so it might function as a recruiter rather than as a
deacetylase although it does show deacetylase activity in vitro (de Ruijter et
al., 2003). HDAC11 is more closely related to the class I HDACs, but no
clear classification has been made due to the low similarity of the overall
sequence (Gao et al., 2002).
31
Class III HDACs are homologous to yeast Sir2 (Silent information regulator) protein that is known to be required for silencing of the mating type
loci and of telomeres. Recent evidence suggests that Sir2 and its related proteins are involved in various biological pathways including longevity, apoptosis, and skeletal myogenesis (Fulco et al., 2003; Guarente, 2001; Luo et al.,
2001).
NCoR/SMRT/HDAC3 complex
NCoR and the related protein SMRT (SMRTER in Drosophila) are involved
in repression by unliganded thyroid hormone and retinoic acid receptors (TR
and RAR) (Chen and Evans, 1995; Horlein et al., 1995). In addition, they are
recruited by several unrelated transcription repressors such as Notch binding
protein CBF-1 (Kao et al., 1998) and homeodomain proteins including Rpx2,
Pit, and Pbx (Asahara et al., 1999; Xu et al., 1998) in mammals.
NCoR/SMRT complexes additionally include GPS2 (G protein pathway
suppressor 2), HDAC3, TBL1 (Transducin like-1) and TBLR1 (TBL related-1) proteins (Guenther et al., 2000; Li et al., 2000; Zhang et al., 2002a).
NCoR and SMRT are highly related but distinct proteins, sharing similar
domain structure and function. They both contain a conserved deacetylaseactivating domain (DAD) that is essential for HDAC3 activation. Additionally, NCoR/SMRT can transiently recruit other HDACs (e.g. HDAC1-2) for
full repression activity (Guenther et al., 2000). SMRTER, the Drosophila
ortholog of mammalian SMRT, was identified through its interaction with
ecdyson receptor (Tsai et al., 1999). TBL1 is an F box/WD-40-containing
protein, originally cloned in relationship to an X-linked human disorder
called Ocular Albinism with late-onset Sensorineural Deafness (OASD)
(Bassi et al., 1999). WD-repeat domains are also found in two other wellstudied corepressors, Drosophila Groucho (Fisher and Caudy, 1998) and
yeast Tup1 (Wahi et al., 1998). In mammalian cells, TBL1 and its related
TBLR1 protein share very high homology, and they have overlapping functions but distinct specificity (Perissi et al., 2004; Yoon et al., 2005). TBL1
and TBLR1 can directly interact with hypoacetylated histones H2B and H4
(Yoon et al., 2005), which is important for the stable association of
NCoR/SMRT complexes with chromatin (Yoon et al., 2003; Yoon et al.,
2005). Interestingly, the presence of the F-box domains in TBL1, TBLR1
and their homologs implies a second function of this protein family, the ability to recruit the ubiquitin machinery. TBL1 associates with Siah-1 E3 ligase
and SIP (Siah-interacting protein), promoting -catenin degradation linked to
p53 responses (Matsuzawa and Reed, 2001). It was suggested that TBL1 and
TBLR1 recruiting ubiquitin/19S proteaseome complexes to nuclear receptorregulated targets is required for a corepressor/coactivator exchange upon
ligand binding (Perissi et al., 2004). Ebi, the fly homolog of TBL1, was first
identified as a downstream regulator of epidermal growth factor receptor
32
(EGFR) signaling during eye development (Dong et al., 1999). The
Ebi/SMRTER complex mediates cross-talk between EGFR and Notch signaling by regulating Delta expression in the eye (Tsuda et al., 2002)., This is
achieved by Ebi/SMRTER complex regulating expression of the charlatan
gene together with the transcription factor Suppressor of Hairless (Su(H))
(Tsuda et al., 2006). Additionally, Ebi collaborates with Sina (Seven in absentia) E3 ligase and the adaptor Phyllopod, inducing poly-ubiquitylation
and degradation of Tramtrack during eye development (Boulton et al., 2000;
Dong et al., 1999; Li et al., 2002).
Epigenetic control
In contrast to the term genetic , meaning information coded by DNA sequence, a definition of epigenetic is nuclear inheritance which is not based
on differences in DNA sequence (Holliday, 1994). When gene expression is
under epigenetic control, the expression pattern can be kept in a stable state
during cell division, meaning heritable . There are a number of phenomena
that involve epigenetic control, including position effect variegation (PEV)
in Drosophila, imprinting, X chromosome inactivation in female mammals,
heterochromatin formation, nuclear reprogramming, and gene silencing.
Some histone modifications and DNA methylation are related to epigenetic
phenomena, so they are collectively termed as epigenetic marks (reviewed in
Berger, 2007). DNA methylation is found in several organisms and it occurs
frequently in CpG sites in the DNA sequence. Methylated DNA can interfere
with transcription factor binding and can recruit repressor proteins which
specifically bind to methylated CpG. DNA methylation plays a critical role
in imprinting and X chromosome inactivation in mammals. Several histone
methylation sites are linked to heterochromatin formation, such as methylation on H3K9, H3K27 and H4K20. Methylation of H3K27 is involved in
gene silencing controlled by the Polycomb groups of proteins in Drosophila.
However, it is still unclear how epigenetic persistence of chromatin states is
achieved and which modifications are truly heritable.
Heterochromatin and position effect variegation
Heterochromatin was named for its appearance under the light microscope: it
appears dense compared to other chromatin, the euchromatin. Heterochromatin is usually located in the centromeres and the telomeres, and is required
for chromosome integrity. These regions contain many repetitious sequences
but few genes. The histones in heterochromatin are often hypoacetylated and
the residue H3K9 is methylated (Stewart et al., 2005). When boundary elements between heterochromatin and euchromatin are removed, heterochromatin packaging is able to spread into euchromatin (reviewed in Grewal and
33
Jia, 2007). This was firstly noticed in a phenomenon, position effect variegation (PEV), in Drosophila. There when a reporter gene (e.g. the white gene)
is juxtaposed with heterochromatin through chromosome rearrangement or
transposition, the expression of the reporter gene will show a variegated
pattern. PEV has been a critical tool for identifying factors involved in heterochromatin formation, particularly the investigation of the suppressors of
PEV (Su(var) genes). The identified genes typically encode either heterochromatin components or enzymes that control changes in the histone modifications. For example, Su(var)3-9, a H3K9 methyl transferase, work together with HP1, a methyl binder, and Su(var)3-7 (Jaquet et al., 2002) in a
complex to maintain stable gene silencing in heterochromatin (reviewed in
Ebert et al., 2006). It was suggested that the establishment of heterochromatin in gene silencing requires a sequential set of reactions: histone demethylase (LSD1) removes the mark of active genes, methylated H3K4,
HDACs (HDAC1) deacetylates H3K9, Su(var) 3-9 methylates H3K9, HP1
binds to methylated H3K9 and finally recruit more histone methyl transferase to make more heterochromatin marks (Grewal and Jia, 2007). Studies
from fission yeast discovered the involvement of the RNAi machinery in the
nucleation of heterochromatin (Hall et al., 2002). siRNA generated by the
RNAi machinery is able to target histone-modifying proteins (e.g. clr 4, homolog of Su(var)3-9) to chromatin which will subsequently methylate H3K9
(Verdel and Moazed, 2005). The methylation of H3K9 not only recruits
Swi6 (homolog of HP1) and other factors but also stabilize the association of
the RNAi complexes with chromatin, facilitating the spreading of heterochromatin. Similarly, recent investigations also implicate the RNAi system
in heterochromatic silencing in Drosophila (Pal-Bhadra et al., 2004).
Polycomb group and Trithorax group
Polycomb Group (PcG) proteins and Trithorax Group (TrxG) proteins are
conserved transcriptional regulators essential for maintaining repressed and
active expression patterns of critical genes respectively during development.
For example, PcG genes are necessary for silencing the Ubx gene in wing
discs and the Scr gene in second and third thoracic discs in Drosophila
(Beuchle et al., 2001; Ringrose and Paro, 2004). In addition, they are involved in cell proliferation, stem cell identity, cancer, genomic imprinting in
plants and mammals, and X inactivation (reviewed in Schuettengruber et al.,
2007). The DNA elements that recruit PcG and TrxG factors to chromatin
are called PcG and trxG response elements (PREs and TREs) respectively.
Three PcG complexes have been identified, PRC1, PRC2 and PhoRC. In
Drosophila, the core components of PRC2 include E(z) (Enhancer of zeste),
Esc (Extra sex comb), Su(z)12 (Suppressor of zeste 12) and Nurf55. E(z) is a
HMT and able to tri-methylate H3K27 (Czermin et al., 2002), a mark recognized and bound by Pc (Polycomb) that is a core subunit in the PRC1 com34
plex. PRC1 additionally contains Ph (Polyhomeotic), Psc (Posterior sex
combs) and dRing. dRing was shown to be a E3 ubiquitin ligase that
monoubiquitylates histone H2A (Wang et al., 2004). The PhoRC complex
was newly identified and contains the DNA-binding protein Pho (Pleiohomeotic) and dSfmbt (Scm-related gene containing four malignant brain
tumor domains) protein which binds to mono-and dimethylated H3K9 and
H4K20 (Klymenko et al., 2006).
TrxG genes are not always acting in complexes, but there are a few trxG
complexes classified. The Drosophila SWI/SNF chromatin remodeling
complex, also called the BRM complex, contains the proteins BRM, MOIRA,
OSA and SNR1. Trx (Trithorax), a H3K4 HMT, constitutes a TAC1 complex together with CBP (Petruk et al., 2001). Ash1, another H3K4 HMT,
genetically and physically interacts with CBP as well (Bantignies et al., 2000;
Byrd and Shearn, 2003; Petruk et al., 2001). The trxG protein Ash2 is also
present in a complex but its partners haven ¯t been i dentifi ed(Papoulas et al.,
1998). It is notable that the functions of PcG and trxG proteins involve several histone modifications, such as H3K27me3, H3K4me3, H2BK123ub1
and H2A119ub1 (Schuettengruber et al., 2007). Recently, it was shown that
noncoding RNAs also contribute to PcG protein mediated gene silencing
(reviewed in Costa, 2005).
35
Co-regulators
Transcription co-regulators are recruited to DNA by transcription factors and
thus they cooperate to integrate complex regulatory information. In general,
there are three means by which coregulators modulate transcription. First,
they change the accessibility of chromatin for RNA polymerase II and other
general transcription factors (GTFs). This class of proteins includes histone
modification enzymes and chromatin remodeling factors. Second, they interact directly with GTFs, such as the TBP associated factors (TAFs) and the
mediator complexes. Finally, they post-translationally modify transcription
factors or GTFs. Examples are HATs, HDACs, and protein kinases. These
modifications can either promote or disrupt the function of transcription
factors. Some coregulators are integrated into complexes which constitute
multiple functional units. This fulfills the requirements for complex developmental programs. Notably, one coregulator complex can be used by different transcription factors and one transcription factor can recruit distinct
classes of coregulator complexes in a context dependent manner (Rosenfeld
et al., 2006). Co-activators are recruited by activators, aiding transcription. A
number of co-activators have been extensively studied (e.g. CBP and SAGA,
reviewed in Carrozza et al., 2003; Daniel and Grant, 2007; Mannervik et al.,
1999). By contrast, co-repressors interact with repressors, inhibiting transcription. Besides NCoR/SMRT corepressor complexes, well-known corepressors include yeast Tup1 protein (reviewed in Malave and Dent, 2006),
Groucho (reviewed in Buscarlet and Stifani, 2007; Chen and Courey, 2000),
CtBP (reviewed in Chinnadurai, 2002; Mannervik et al., 1999) and
Sin3/Rpd3 complexes (reviewed in Ayer, 1999; Schreiber-Agus and DePinho, 1998; Silverstein and Ekwall, 2005).
Based on transcriptional studies in transgenic embryos, the Drosophila
repressors have been grouped into two different types: the short-range repressors work over short distances (less than 100bp) from the activator sites
or the core promoter and the long range repressors function over long distances (more than 1000 bp) (reviewed in Courey and Jia, 2001; Mannervik et
al., 1999). For example, there are three binding clusters for Kr in the eve
stripe 2 CRM. Two of them directly overlap the sites of Bicoid activator, and
may function through competition. On the third site, Kr can recruit the
dCtBP co-repressor that helps to inhibit the action of Bicoid bound nearby.
On the other hand, the long-range repressors, such as Hairy and Dorsal, are
able to suppress the activity of multiple enhancers. The mechanism of long36
range and short-range repression has not been fully understood. Three
mechanisms were proposed to interpret how repressors work: competing for
binding sites with activatiors; antagonizing the activity of the activatorcoactivator complex (quenching); inhibiting the activity of a transcription
complex at the core promoter (direct repression). Quenching and direct repression mechanisms involve the recruitment of co-factors. All of the identified short-range repressors appear to mediate their repression dependent on
dCtBP (Nibu and Levine, 2001; Nibu et al., 1998a; Zhang and Levine, 1999).
In contrast, two long-range repressors require Groucho (Barolo and Levine,
1997). Table 1 summarizes repressors and co-repressors in pre-cellular embryos.
Table 1. The known co-repressors of repressors in pre-cellular Drosophila embryos
Corepressor
Repressor
Family
Range
Reference
dCtBP
Kr
Zinc finger
short
(Nibu et al., 1998a)
Kni
Nuclear receptor
short
(Nibu et al., 1998a)
Giant
bZIP
short
(Nibu and Levine, 2001;
Strunk et al., 2001)
Snail
Zinc finger
short
(Nibu et al., 1998a)
Ebi
Snail
Zinc finger
short
(paper III)
Groucho
Hairy
bHLH
Long
(Paroush et al., 1994)
Dorsal
Rel
Long
(Dubnicoff et al., 1997)
Huckebein
Zinc finger
?
(Goldstein et al., 1999)
Capicua
HMG-box
?
(Jimenez et al., 2000)
Odd-skipped
Zinc finger
?
(Goldstein et al., 2005)
Eve
Homeodomain
?
(Kobayashi et al., 2001)
Runt
Runt
?
(Aronson et al., 1997)
Atrophin
Tailless
Nuclear receptor
?
(Wang et al., 2006)
Brakeless
Tailless
Nuclear receptor
?
(Paper IV)
37
CtBP
The C-terminal-binding protein (CtBP) was originally identified as a cellular
protein that interacts with a C-terminal motif (PXDLSXK) of adenovirus
E1A oncoproteins (Boyd et al., 1993). CtBP family proteins are highly conserved among invertebrates and vertebrates. They share a high degree of
amino acid homology with NAD-dependent 2-hydroxy acid dehydrogenases,
so it is conceivable that they may mediate repression through their enzymatic
activity (Kumar et al., 2002). The biochemical and genetic studies of Drosophila CtBP (dCtBP) have revealed that dCtBP functions as a transcriptional
corepressor (Nibu et al., 1998b; Poortinga et al., 1998). Kr, Knirps and Snail
interact with dCtBP through conserved CtBP-interaction motifs (Nibu et al.,
1998b). However, a similar interaction motif has not been found in the Giant
protein sequence (Nibu and Levine, 2001). Interestingly, these repressors
harbor both dCtBP-dependent and dCtBP-independent activities (Keller et
al., 2000; La Rosee-Borggreve et al., 1999; Strunk et al., 2001). Hairy uses
Groucho for its repression acitivity but it contains a divergent CtBPinteracting motif (PLSLV) which may antagonize the activity of Groucho
(Zhang and Levine, 1999). In addition, dCtBP is involved in signaling pathways, including the Notch and Wnt pathways. Specifically, dCtBP can be
recruited by Hairless to Supressor of Hairless (Su(H) targeted promoters,
repressing Notch target genes (Barolo et al., 2002; Morel et al., 2001).
dCtBP was shown to associate with Wnt response elements and was able to
repress Wnt target genes (Fang et al., 2006). The precise mechanism by
which CtBP proteins mediate transcriptional repression remains to be elucidated. Human CtBP has been reported to associate with HDACs (Sundqvist
et al., 1998). However, CtBP-mediated repression of certain promoters
seems to be sensitive to HDAC inhibitor trichostatin A (TSA), while others
are not (Chinnadurai, 2002). In Drosophila, the function of dCtBP is not
impaired in Rpd3 mutant embryos (Mannervik and Levine, 1999). Furthermore, dCtBP-mediated repression in cell culture is insensitive to TSA (Ryu
and Arnosti, 2003). So the involvement of HDACs in dCtBP function is not
clear.
Groucho
The Drosophila Groucho (Gro) protein was identified as a corepressor of
Hairy, a bHLH (basic helix-loop-helix) repressor that is essential for segmentation and neurogenesis (Paroush et al., 1994). Since then, more repressors have been identified to interact with Gro. These factors include Dorsal
(Dubnicoff et al., 1997; Flores-Saaib et al., 2001), Runt (Aronson et al.,
1997), Eve (Kobayashi et al., 2001), Odd-skipped (Goldstein et al., 2005),
En (Jimenez et al., 1997), Huckebein (Goldstein et al., 1999), Brinker
38
(Hasson et al., 2001) and Capicua (Jimenez et al., 2000). It was shown that
the recruitment of Gro by certain transcription factors is through the WRPWlike motifs (e.g. WRPW in Hairy, WRPY in Runt) or the engrailed homology domain 1 (EH-1) motifs (e.g. FSISNIL in Engrailed) (reviewed in Buscarlet and Stifani, 2007; Chen and Courey, 2000). The human homologs of
Gro have been named Transducin-like enhancer of split (TLE) proteins
(Stifani et al., 1992). The Gro/TLE proteins contain two conserved domains,
a WD-repeat domain and a glutamine-rich (Q) domain, both of which are
required for the repression function. Specifically, the WD-repeat domain can
mediate protein-protein interactions (Chen and Courey, 2000; Jimenez et al.,
1997) and the Q domain mediates homotetramerization and higher-order
oligomerization (Chen et al., 1998; Song et al., 2004). It was suggested that
the oligomerization of Gro and the ability to interact with histones and Rpd3
may contribute to the long-range repression activity of Gro (Chen et al.,
1999; Courey and Jia, 2001; Song et al., 2004). Studies in Drosophila indicated that Gro may function to convert an activator into a repressor. For example, Dorsal activates ventral-and-lateral- expressed genes (e.g. twist, rho,
and sog) but it represses dorsal-expressing genes (e.g. dpp, zen and tld). Its
repression activity requires the association with Cut and Dead Ringer (Dri),
which will together facilitate the recruitment of Gro onto the target promoter
(Valentine et al., 1998). Another example is dTcf (Drosophila homolog of T
cell factor) in Wingless signaling. In the absence of Wingless, dTcf is converted to a repressor, which involves the recruitment of Gro (Cavallo et al.,
1998; Levanon et al., 1998).
Atrophin and Brakeless
Atrophin proteins represent a new class of nuclear receptor corepressors
(Wang et al., 2006; Zhang et al., 2006a). Human Atrophin1 (ATN1) and
Atrophin2 (ATN2, RERE) are polyglutamine (poly-Q) proteins and expansion of the poly-Q domain of ATN1 causes the neurodegenerative disease
Dentatorubral-pallidoluysian atrophy (DRPLA) (Kanazawa, 1999). Mouse
and zebrafish Atrophin2 recruit HDAC1/2 and are involved in Fgf8 signaling during embryogenesis (Asai et al., 2006; Plaster et al., 2007; Zoltewicz
et al., 2004). Mouse Atrophin1 is not required for viability (Shen et al., 2007)
but interacts with TLX, which may contribute to prevent retinal malformation and degeneration during development (Zhang et al., 2006a). The TLX
repressor is the homolog of Drosophila Tll repressor and both belong to the
NR2 subfamily of orphan nuclear receptors (reviewed in Moran and Jimenez,
2006). Mutations in Drosophila atrophin cause a variety of patterning defects (Erkner et al., 2002; Fanto et al., 2003; Kankel et al., 2004; Zhang et al.,
2002b). It has been suggested that Drosophila Atrophin is recruited to DNA
by Tll and represses transcription by interacting with HDAC1 and HDAC2
39
(Wang et al., 2006). Tll specifies the terminal structures of embryo by repressing transcription of genes such as Kr and kni. Atrophin was also shown
to interact with Eve and Hkb (Zhang et al., 2002b). Additionally, Atrophin
has been classed as a member of the Trithorax Group of genes (Kankel et al.,
2004). Atrophin has been shown to associate with the atypical cadherin Fat
and is required for establishment of planar cell polarity in the eye and wing
(Fanto et al., 2003). Recently, Atrophin was shown to genetically interact
with Groucho and Brakeless (Wehn and Campbell, 2006).
Brakeless (Bks) is also known as Scribbler, and as Master of thickveins in
Drosophila. Previous studies have discovered three kinds of mutant phenotypes: 1) a behavioral locomotor phenotype in larvae (Yang et al., 2000); 2)
deregulation of expression of the thickveins (tkv) gene in the wing imaginal
disc (Funakoshi et al., 2001); 3) defects in axonal guidance in the eye due to
the derepression of the runt gene in R2 and R5 photo receptors (Kaminker et
al., 2002; Rao et al., 2000; Senti et al., 2000). Bks encodes a nuclear protein.
The facts that Bks genetically interacts with Atrophin and Groucho and that
bks mutants have similar phenotypes to atrophin mutants, imply its function
in transcriptional repression (Wehn and Campbell, 2006). The bks gene locus encodes two isoforms of proteins, BksA and BksB. They share the Nterminal 929 amino acids, but BksB is longer and has a 1373-residue Cterminal extension (Rao et al., 2000; Senti et al., 2000). There is a zinc finger
domain as well as a glutamine-rich domain in the unique part of BksB. BksA
can rescue the behavioral phenotype but only partially rescue the axon guidance phenotype. However, BksB fully rescued the defect in axon guidance
(Senti et al., 2000; Yang et al., 2000) This result indicates that these two
isoforms have non-overlapping function. The human homologs of Bks are
two putative zinc finger proteins, ZFP608 and ZFP609. Recently, it was
suggested that ZFP608 may play an inhibitory role in rag1 (recombinationactivating gene1) and rag2 expression (Zhang et al., 2006b).
40
Aim of this thesis
The purpose of this thesis has been to understand the in vivo function of
transcriptional co-regulators, using Drosophila melanogaster as a model.
These co-regulators, associated with DNA binding transcription factors,
regulate gene expression and thus affect the developmental process.
dAda2b and Reptin belong to HAT complexes, whereas Ebi associates
with HDAC3. This class of proteins may affect transcription by changing the
accessibility of chromatin. Brakeless is a nuclear protein with previously
unknown function that we isolated from a genetic screen. Our aim was to
investigate the function of these factors and their partners in Drosophila
development.
41
Results and discussion of project
Paper I
The SAGA complex is evolutionarily conserved and contains a trimeric subcomplex which consists of Gcn5, Ada2 and Ada3. There are two gene loci
encoding two Ada2 proteins in the Drosophila genome. Although dAda2a
and dAda2b are both associated with Gcn5, they exist in different complexes.
dAda2b is present in a SAGA-like complex (Kusch et al., 2003; Muratoglu
et al., 2003). We generated dAda2b mutant flies from a P-element transposon line by using imprecise excision. The homozygous mutant animals die
during the early pupa stage without any gross morphological abnormalities.
We showed that dAda2b is able to interact with Gcn5 in vitro and that
dAda2b and Gcn5 are expressed in many of the same tissues. Since yeast
Ada2 is required for maintaining Gcn5 in the SAGA complex and stimulates
the catalytic activity of Gcn5, we performed immunostainings of Drosophila
embryos and polytene chromosomes in salivary glands to investigate histone
acetylation level in dAda2b mutants by using antibodies against different
acetylated forms of histone H3 and histone H4. Consistent with previous
data from yeast which showed that Gcn5 preferentially acetylates histone H3,
we found that the acetylation of histone H3 on lysine 9 and lysine 14 is dramatically reduced in homozygous dAda2b mutants but that of histone H4 on
tested lysines is normal. In addition, chromatin acetylation is globally affected by the dAda2b mutation, rather than affecting a subset of gene loci.
We conclude that dAda2b is required for viability and normal H3 acetylation
levels either through affecting the catalytic activity of Gcn5 or proper targeting of Gcn5 to chromatin.
It has been shown that the activation domain in p53 can interact with both
mammalian and Drosophila SAGA complexes (Candau et al., 1997; Kusch
et al., 2003; Utley et al., 1998). In order to investigate whether dAda2b contributes to p53 function in vivo, we analyzed the p53-dependent induction of
apoptosis and reaper (rpr) gene expression in response to X-rays. Contradictory to the in vitro results, p53-induced apoptosis and reaper gene expression in response to DNA damage are not impaired in dAda2b mutants, indicating that the dAda2b/Gcn5 subcomplex of SAGA is not necessary for p53
function in vivo. Unexpectedly, increased apoptosis was observed after
42
treatment with gamma radiation in dAda2 mutants. However, when we induced reaper expression by using a reaper transgene instead of by DNA
damage, we did not detect any excessive apoptosis in dAda2b mutants. Further, we generated a dAda2b and p53 double mutant to examine whether the
increased apoptosis is dependent on p53. The double mutants did not show
an increase in apoptosis following irradiation, suggesting that the excessive
apoptosis in dAda2b single mutants is p53 dependent. We propose that increased apoptosis in dAda2b mutants could result from inefficient DNA repair, forcing more cells into apoptosis. If this is true, the function may be
evolutionarily conserved. When we tested yeast strains lacking Ada2p,
Ada3p or Gcn5p proteins on plates containing the DNA damaging agent
MMS, they all showed sensitivity to MMS. Taken together, our results show
that Ada2 and its associated proteins are sensitive to genotoxic agents in
both yeast and flies, indicating that an Ada2-Gcn5 complex may facilitate
efficient DNA repair or the generation of a DNA damage signal.
Paper II
Reptin was shown to co-purify with the Polycomb complex PRC1 in Drosophila (Saurin et al., 2001), prompting us to test if the biochemical interaction
with PRC1 was accompanied by a genetic interaction. We examined three
components of the PRC1 complex, Pc, Psc and Ph, and two components of
the E(z)-Esc complex (also called PRC2 complex), Esc and Pcl. As shown in
paper II, reptin genetically interacts with members from the PRC1 complex
to regulate expression of the Hox gene Scr (Sex comb reduced). Additionally,
reptin interacts with Pcl from the PRC2 complex. A lacZ transgene driven
by the IDE enhancer and a PRE from another Hox gene, Ubx, is derepressed
in reptin mutant clones in the wing imaginal disc. However, the endogenous
Ubx gene is still silent in the reptin mutant clones in the wing discs, suggesting that Reptin is not essential for the expression of Ubx and that repression
of Ubx gene is not as sensitive to the loss of Reptin as the Ubx PRE. reptin
mutants suppress PEV, which is another feature different from most PcG
genes. So we do not consider reptin a bona fide PcG gene. Since Reptin is
present in a Tip60 complex in mammals and recently was shown to be a
component of a Drosophila Tip60 complex (Doyon et al., 2004; Kusch et al.,
2004), it is possible that the genetic interactions with PcG genes are through
the presence of Reptin in the Tip60 complex. In fact, genes encoding three
other components in the Tip60 complex, E(Pc), domino and dMRG15, share
with reptin the ability to genetically interact with PcG genes and suppress
PEV. The Tip60 complex possesses at least two enzymatic activities, a HAT
activity associated with Tip60 and an ATPase activity associated with Domino. Domino protein is similar to P400 and to SRCAP in mammals and to
Swr1 in yeast (Eissenberg et al., 2005). This class of proteins has histone
43
exchange activities (Kobor et al., 2004; Krogan et al., 2003; Mizuguchi et al.,
2004). For example, Swr1 exchange H2A for the variant histone H2A.Z in
nucleosomes (Mizuguchi et al., 2004). The fly Domino protein exchanges
phosphorylated and acetylated H2Av for unmodified H2Av after DNA damage (Kusch et al., 2004).
What can be the basis for the genetic interaction between Tip60 components and PcG genes and that for the function in silent chromatin formation?
One possibility is that the Tip60 complex regulates PcG expression. However, we did not observe reduced Pc expression in reptin mutant embryos.
Another possibility is that the enzymatic activities of the Tip60 complex
cooperate with PcG genes to mediate transcriptional silencing. Since binding
of Pc to polytene chromosomes is abolished in H2Av mutant animals
(Swaminathan et al., 2005), histone variant exchange might cause the genetic
interaction with PRC1. However, we found that binding of PcG proteins to
polytene chromosomes is unaffected in domino mutant larvae. Htz1 is involved in the spreading of silenced chromatin (Babiarz et al., 2006; Meneghini et al., 2003). However, no change was found in binding of H2Av to
polytene chromosomes from domino mutant larvae. It is possible that PRC1mediated H2A ubiquitylation helps to recruit the Tip60 complex, whose
histone acetylation or histone exchange activity assists in transcriptional
repression. Alternatively, histone acetylation or exchange facilitates binding
of the PRC1 complex to the PREs. Crosstalk among different histone modifications may also contribute to the function of the Tip60 complex in heterochromatin.
Paper III
The Snail-family of repressors play central roles in morphogenesis by inducing mesoderm formation in several organisms, including flies and mammals
(reviewed in Nieto, 2002).We identified Ebi as an essential co-repressor for
the Drosophila Snail protein. In ebi mutant embryos, Snail-target genes are
de-repressed in the presumptive mesoderm. Ebi and Snail interact both genetically and physically. We further identified an evolutionarily conserved
interaction motif in Snail that is necessary for Ebi binding and for the repression activity of Snail. Ebi functions independently of another Snail corepressor, dCtBP, based on several lines of evidence. First, dCtBP protein
level is not affected in ebi mutant embryos. Second, the ebi mutation did not
affect the function of the Kr and Kni repressors, which also requires dCtBP.
Third, the Ebi-interaction domain (Snail 1-40) does not bind to dCtBP but
still has repression activity in S2 cells and in transgenic embryos. Fourth,
removing the Ebi-interacting motif from the Snail protein abolishes its repression activity. Our experiments show that repression of several Snailtarget genes requires both dCtBP and Ebi, and that Snail recruits both CtBP
44
and Ebi to the same rho enhancer in the presumptive mesoderm. Since the
ebi mutation prevents repression of rho, sog and sim but not brk, the ebi
mutant behaves as a snail hypomorph. Moreover, the Ebi-dependent repression domain, Snail 1-40, has weaker activity than the full-length repression
domain (Snail 1-245) in transgenic embryos. Taken together, these results
suggest that Snail requires both Ebi and dCtBP for full repressor activity.
HDAC3
Ebi
Y--CPLKKRP
Sna
a.a. 245
P-DLS-K
P-DLS-R
CtBP
CtBP
Zinc fingers
a.a. 390
DNA-binding
Figure 9. The interactions of Snail with Ebi and CtBP co-repressors. Schematic
drawing of Snail protein shows the C-terminal zinc-finger DNA binding domain
and N-terminal repression domain. Ebi interacts with Snail through the motif Y-CPLKKRP. CtBP interacts with Snail through two motifs, P-DLS-K and P-DLSR. Ebi and HDAC3 are present in the same complex and HDAC activity is part of
mechanism by which Snail mediates repression.
In mammals, the Ebi homolog TBL1 is part of the N-CoR/SMRT/HDAC3
co-repressor complex. Histone deacetylation is functionally linked to the
activity of this complex. Our studies found that Drosophila HDAC3 and Ebi
associate and that both are required for Snail repressor function in S2 cells,
as determined by RNAi and inhibition of HDAC activity. In the early embryo, Ebi is recruited to a Snail target gene in a Snail-dependent manner,
which coincides with histone hypoacetylation. These results indicate that
histone deacetylation is part of the mechanism by which Snail mediates repression and is a repression mechanism in early Drosophila development
(Figure 9).
Paper IV
From a screen for maternal factors that control patterning of Drosophila
embryos, we selected mutants which specifically affect gene expression in
the early embryo. We found that embryo segmentation is severely affected in
bks germline clone mutants, likely resulting from the derepression of two
gap genes, kni and Kr. However, mutations in bks do not affect the expression of other gap genes that are known to control the transcription of kni and
Kr, indicating that Bks may be required for function of one or more gap pro45
teins. We examined the function of Kni, Kr, Gt, Hb and Tll in bks mutants.
All of them except Tll function normally, suggesting that Bks is specifically
required for Tll-mediated repression. Additionally, we found that Bks genetically and physically interacts with Tll. Bks is recruited to the CRMs of
Tll target genes, and represses transcription when tethered to DNA. Tll was
recently shown to utilize Atrophin as a co-repressor (Wang et al., 2006). We
show that both Bks and Atrophin are recruited to the kni CRM. Additionally,
we demonstrate that Atrophin and Bks interact in vitro and that they can be
co-immunoprecipitated from S2 cells. The interaction is conserved between
fly and human as the human ZFP608 (homolog of Bks) protein interacts with
human Atrophin1. We propose that Bks and Atrophin function together as a
co-repressor complex and the complex may function throughout development as bks mutants share similar phenotypes with atrophin mutants in several developmental processes (Wehn and Campbell, 2006). Tll may simultaneously or separately interact with Bks and Atrophin. In either case, both
Bks and Atrophin are required for full Tll activity. However, overexpressing
Tll from a heat-shock promoter can repress the posterior kni stripe in both
wild-type and bks mutant embryos. This result indicates that Bks activity is
dispensable at high enough level of Tll. We believe that Bks and Atrophin
are cooperating as Tll co-repressors, so that Tll function is only partially
impaired by the absence of either one. Interaction with HDACs may be part
of the mechanism by which Atrophin mediates repression. We found that
Bks-mediated repression in S2 cells is insensitive to the HDAC inhibitor
TSA (Trichostatin A). So it is possible that Bks could repress transcription
through a separate mechanism.
46
Conclusion and Perspectives
Gene regulation in multi-cellular organisms is a key process to implement
the genetic program that guides development. In the Drosophila embryo,
zygotically expressed transcription factors require maternal co-regulators for
their activity. In this thesis, we have studied the in vivo function of four cofactors, dAda2b, Reptin, Bks and Ebi in Drosophila development. Although
these proteins are widely expressed, it appears that they take part in specific
cellular and developmental processes. dAda2b mutation strongly affects
global histone acetylation in the central nervous system (CNS) during the
embryonic stage and that on polytene chromosomes at the third instar larva
stage. dAda2b mutants are hyper-sensitive to irradiation-induced DNA damage and yeast Ada2 and other SAGA complex components are sensitive to
the DNA-damage agent MMS. This conserved feature indicates a role of
Ada2 and its associated SAGA complex in DNA repair. It would be interesting to find out the precise mechanism. Our studies revealed a new role of
Reptin and other TIP60 HAT complex components in PcG-mediated repression and in formation of silent chromatin. How does a HAT complex influence gene silencing? Again the molecular basis needs to be explored. To
establish localized and tissue-specific patterns of gene expression, transcriptional repressors require co-regulators for their full activities. Here we identified Ebi and Bks as novel co-repressors of Snail and Tailless respectively.
We further found that histone deacetylation is necessary for Snail-mediated
repression. This is the first evidence that histone deacetylation may be involved in cell fate specification during Drosophila embryonic development.
Chromatin regulators are actively involved in transcription. However, investigation of their roles in development is far from complete. Our studies
give insight into certain aspects of the function of these proteins in Drosophila development.
47
Acknowledgements
During these years in Sweden, I have received tremendous help and support
from my family, friends and colleagues. I would like to acknowledge
My supervisor Mattias Mannervik, for giving me your invaluable scientific
guidance throughout my studies, your continuous support and encouragement as well as your great patience and kindness.
Professor Christos Samakovlis, for giving me good advice on my work and
my career and always being there when we need help.
My former supervisor, Xiushan Wu, for bringing me to Sweden, being supportive to my decisions and giving me many good suggestions.
My former colleagues, Tobias, for taking a lot of general responsibilities in
the lab and translating Swedish documents for me; Mattias Bergman, for
helping with the Ebi project and many funny talks; Achim, for the pleasant
discussions about projects and movies; Haojiang and Haining, for all useful
information and help; my present colleagues, Filip and Fang, for being in
the lab with me during my last stay.
Monika, for taking care of our flies, making embryo injections and sharing
your Chinese mushrooms; Kostas, for making fly food and taking a lot of
boring tasks in the department.
Magdalena, for keeping track of us all and giving me a lot of personal help.
All the rest of the members at Devbio, for making it such a nice place to
work, especially, Stefan, for providing yeast SAGA strains and suggestions
on my projects; other yeast guys Emad, Sid and Jiang, for the nice occasional communications and the great pub time; Dana, for celebrating our
common birthday and the tasty Slovakia wine; Nafiseh, for helping with cell
culture work; Kirsten, for the protocols and suggestions; Katarina, for organizing the nice boat trip; Satish, for being a good party-mood maker; Vasilios, for showing us how devoted a scientist should be; Nicole, Naumi, Oliviano, Chie and Ryo, for creating a good working atmosphere.
48
The former PH.D students, Peggy, Andreas, Johanna, Nikos and Marco, for
always being so kind and helpful to the juniors and exemplifying a good
student.
My friends in Sweden, Bing Tang, Wei Zhao, Liqun He, Ying Sun, Wei
Jiao, Zhi Wang, Guiling Zhao and Dongbo Zhao, for helping us with moving, parties and communications; all my friends in China, especially,
Yongqun Zhang, for sending me good wishes and my favorite food from
8000 kms away.
Last but not least, I am deeply grateful to my family for the support: my
parents, thanks for your loving and spoiling; my brother Tao, thanks for
following me to Sweden and making my life here easier and happier; Qiu ¯s
family, thanks for believing in me; my dear husband Qiu, for sharing all the
happiness and difficulties with me and giving me endless encouragement
and love.
49
References
Aronson, B. D., Fisher, A. L., Blechman, K., Caudy, M. and Gergen, J. P.
(1997). Groucho-dependent and -independent repression activities of Runt
domain proteins. Mol Cell Biol 17, 5581-7.
Asahara, H., Dutta, S., Kao, H. Y., Evans, R. M. and Montminy, M.
(1999). Pbx-Hox heterodimers recruit coactivator-corepressor complexes in
an isoform-specific manner. Mol Cell Biol 19, 8219-25.
Asai, Y., Chan, D. K., Starr, C. J., Kappler, J. A., Kollmar, R. and
Hudspeth, A. J. (2006). Mutation of the atrophin2 gene in the zebrafish
disrupts signaling by fibroblast growth factor during development of the
inner ear. Proc Natl Acad Sci U S A 103, 9069-74.
Ayer, D. E. (1999). Histone deacetylases: transcriptional repression with
SINers and NuRDs. Trends Cell Biol 9, 193-8.
Babiarz, J. E., Halley, J. E. and Rine, J. (2006). Telomeric heterochromatin boundaries require NuA4-dependent acetylation of histone variant
H2A.Z in Saccharomyces cerevisiae. Genes Dev 20, 700-10.
Baek, S. H., Ohgi, K. A., Rose, D. W., Koo, E. H., Glass, C. K. and
Rosenfeld, M. G. (2002). Exchange of N-CoR corepressor and Tip60 coactivator complexes links gene expression by NF-kappaB and beta-amyloid
precursor protein. Cell 110, 55-67.
Banin, S., Moyal, L., Shieh, S., Taya, Y., Anderson, C. W., Chessa, L.,
Smorodinsky, N. I., Prives, C., Reiss, Y., Shiloh, Y. et al. (1998). Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science 281, 1674-7.
Bantignies, F., Goodman, R. H. and Smolik, S. M. (2000). Functional
interaction between the coactivator Drosophila CREB-binding protein and
ASH1, a member of the trithorax group of chromatin modifiers. Mol Cell
Biol 20, 9317-30.
Barbaric, S., Reinke, H. and Horz, W. (2003). Multiple mechanistically
distinct functions of SAGA at the PHO5 promoter. Mol Cell Biol 23, 346876.
Barlev, N. A., Emelyanov, A. V., Castagnino, P., Zegerman, P.,
Bannister, A. J., Sepulveda, M. A., Robert, F., Tora, L., Kouzarides, T.,
Birshtein, B. K. et al. (2003). A novel human Ada2 homologue functions
with Gcn5 or Brg1 to coactivate transcription. Mol Cell Biol 23, 6944-57.
Barolo, S. and Levine, M. (1997). hairy mediates dominant repression in
the Drosophila embryo. Embo J 16, 2883-91.
Barolo, S., Stone, T., Bang, A. G. and Posakony, J. W. (2002). Default
repression and Notch signaling: Hairless acts as an adaptor to recruit the
50
corepressors Groucho and dCtBP to Suppressor of Hairless. Genes Dev 16,
1964-76.
Bassi, M. T., Ramesar, R. S., Caciotti, B., Winship, I. M., De Grandi, A.,
Riboni, M., Townes, P. L., Beighton, P., Ballabio, A. and Borsani, G.
(1999). X-linked late-onset sensorineural deafness caused by a deletion involving OA1 and a novel gene containing WD-40 repeats. Am J Hum Genet
64, 1604-16.
Bauer, A., Chauvet, S., Huber, O., Usseglio, F., Rothbacher, U., Aragnol,
D., Kemler, R. and Pradel, J. (2000). Pontin52 and reptin52 function as
antagonistic regulators of beta-catenin signalling activity. Embo J 19, 612130.
Bauer, A., Huber, O. and Kemler, R. (1998). Pontin52, an interaction
partner of beta-catenin, binds to the TATA box binding protein. Proc Natl
Acad Sci U S A 95, 14787-92.
Bellosta, P., Hulf, T., Balla Diop, S., Usseglio, F., Pradel, J., Aragnol, D.
and Gallant, P. (2005). Myc interacts genetically with Tip48/Reptin and
Tip49/Pontin to control growth and proliferation during Drosophila development. Proc Natl Acad Sci U S A 102, 11799-804.
Berger, S. L. (2007). The complex language of chromatin regulation during
transcription. Nature 447, 407-12.
Berger, S. L., Pina, B., Silverman, N., Marcus, G. A., Agapite, J., Regier,
J. L., Triezenberg, S. J. and Guarente, L. (1992). Genetic isolation of
ADA2: a potential transcriptional adaptor required for function of certain
acidic activation domains. Cell 70, 251-65.
Bergmann, A., Stein, D., Geisler, R., Hagenmaier, S., Schmid, B., Fernandez, N., Schnell, B. and Nusslein-Volhard, C. (1996). A gradient of
cytoplasmic Cactus degradation establishes the nuclear localization gradient
of the dorsal morphogen in Drosophila. Mech Dev 60, 109-23.
Bernstein, C., Bernstein, H., Payne, C. M. and Garewal, H. (2002). DNA
repair/pro-apoptotic dual-role proteins in five major DNA repair pathways:
fail-safe protection against carcinogenesis. Mutat Res 511, 145-78.
Beuchle, D., Struhl, G. and Muller, J. (2001). Polycomb group proteins
and heritable silencing of Drosophila Hox genes. Development 128, 9931004.
Bird, A. W., Yu, D. Y., Pray-Grant, M. G., Qiu, Q., Harmon, K. E.,
Megee, P. C., Grant, P. A., Smith, M. M. and Christman, M. F. (2002).
Acetylation of histone H4 by Esa1 is required for DNA double-strand break
repair. Nature 419, 411-5.
Blander, G. and Guarente, L. (2004). The Sir2 family of protein deacetylases. Annu Rev Biochem 73, 417-35.
Boudreault, A. A., Cronier, D., Selleck, W., Lacoste, N., Utley, R. T.,
Allard, S., Savard, J., Lane, W. S., Tan, S. and Cote, J. (2003). Yeast
enhancer of polycomb defines global Esa1-dependent acetylation of chromatin. Genes Dev 17, 1415-28.
Boulay, J. L., Dennefeld, C. and Alberga, A. (1987). The Drosophila developmental gene snail encodes a protein with nucleic acid binding fingers.
Nature 330, 395-8.
51
Boulton, S. J., Brook, A., Staehling-Hampton, K., Heitzler, P. and Dyson,
N. (2000). A role for Ebi in neuronal cell cycle control. Embo J 19, 5376-86.
Boyd, J. M., Subramanian, T., Schaeper, U., La Regina, M., Bayley, S.
and Chinnadurai, G. (1993). A region in the C-terminus of adenovirus 2/5
E1a protein is required for association with a cellular phosphoprotein and
important for the negative modulation of T24-ras mediated transformation,
tumorigenesis and metastasis. Embo J 12, 469-78.
Boyer, L. A., Langer, M. R., Crowley, K. A., Tan, S., Denu, J. M. and
Peterson, C. L. (2002). Essential role for the SANT domain in the functioning of multiple chromatin remodeling enzymes. Mol Cell 10, 935-42.
Brand, M., Moggs, J. G., Oulad-Abdelghani, M., Lejeune, F., Dilworth,
F. J., Stevenin, J., Almouzni, G. and Tora, L. (2001). UV-damaged DNAbinding protein in the TFTC complex links DNA damage recognition to
nucleosome acetylation. Embo J 20, 3187-96.
Braunstein, M., Sobel, R. E., Allis, C. D., Turner, B. M. and Broach, J.
R. (1996). Efficient transcriptional silencing in Saccharomyces cerevisiae
requires a heterochromatin histone acetylation pattern. Mol Cell Biol 16,
4349-56.
Breen, T. R. and Harte, P. J. (1991). Molecular characterization of the
trithorax gene, a positive regulator of homeotic gene expression in Drosophila. Mech Dev 35, 113-27.
Brodsky, M. H., Weinert, B. T., Tsang, G., Rong, Y. S., McGinnis, N. M.,
Golic, K. G., Rio, D. C. and Rubin, G. M. (2004). Drosophila
melanogaster MNK/Chk2 and p53 regulate multiple DNA repair and apoptotic pathways following DNA damage. Mol Cell Biol 24, 1219-31.
Brownell, J. E., Zhou, J., Ranalli, T., Kobayashi, R., Edmondson, D. G.,
Roth, S. Y. and Allis, C. D. (1996). Tetrahymena histone acetyltransferase
A: a homolog to yeast Gcn5p linking histone acetylation to gene activation.
Cell 84, 843-51.
Bu, P., Evrard, Y. A., Lozano, G. and Dent, S. Y. (2007). Loss of Gcn5
acetyltransferase activity leads to neural tube closure defects and exencephaly in mouse embryos. Mol Cell Biol 27, 3405-16.
Buggy, J. J., Sideris, M. L., Mak, P., Lorimer, D. D., McIntosh, B. and
Clark, J. M. (2000). Cloning and characterization of a novel human histone
deacetylase, HDAC8. Biochem J 350 Pt 1, 199-205.
Buscarlet, M. and Stifani, S. (2007). The 'Marx' of Groucho on development and disease. Trends Cell Biol 17, 353-61.
Byrd, K. N. and Shearn, A. (2003). ASH1, a Drosophila trithorax group
protein, is required for methylation of lysine 4 residues on histone H3. Proc
Natl Acad Sci U S A 100, 11535-40.
Candau, R. and Berger, S. L. (1996). Structural and functional analysis of
yeast putative adaptors. Evidence for an adaptor complex in vivo. J Biol
Chem 271, 5237-45.
Candau, R., Scolnick, D. M., Darpino, P., Ying, C. Y., Halazonetis, T. D.
and Berger, S. L. (1997). Two tandem and independent sub-activation domains in the amino terminus of p53 require the adaptor complex for activity.
Oncogene 15, 807-16.
52
Carre, C., Szymczak, D., Pidoux, J. and Antoniewski, C. (2005). The
histone H3 acetylase dGcn5 is a key player in Drosophila melanogaster
metamorphosis. Mol Cell Biol 25, 8228-38.
Carrozza, M. J., Li, B., Florens, L., Suganuma, T., Swanson, S. K., Lee,
K. K., Shia, W. J., Anderson, S., Yates, J., Washburn, M. P. et al. (2005).
Histone H3 methylation by Set2 directs deacetylation of coding regions by
Rpd3S to suppress spurious intragenic transcription. Cell 123, 581-92.
Carrozza, M. J., Utley, R. T., Workman, J. L. and Cote, J. (2003). The
diverse functions of histone acetyltransferase complexes. Trends Genet 19,
321-9.
Cavallo, R. A., Cox, R. T., Moline, M. M., Roose, J., Polevoy, G. A.,
Clevers, H., Peifer, M. and Bejsovec, A. (1998). Drosophila Tcf and
Groucho interact to repress Wingless signalling activity. Nature 395, 604-8.
Chen, G. and Courey, A. J. (2000). Groucho/TLE family proteins and transcriptional repression. Gene 249, 1-16.
Chen, G., Fernandez, J., Mische, S. and Courey, A. J. (1999). A functional interaction between the histone deacetylase Rpd3 and the corepressor
groucho in Drosophila development. Genes Dev 13, 2218-30.
Chen, G., Nguyen, P. H. and Courey, A. J. (1998). A role for Groucho
tetramerization in transcriptional repression. Mol Cell Biol 18, 7259-68.
Chen, J. D. and Evans, R. M. (1995). A transcriptional co-repressor that
interacts with nuclear hormone receptors. Nature 377, 454-7.
Chinnadurai, G. (2002). CtBP, an unconventional transcriptional corepressor in development and oncogenesis. Mol Cell 9, 213-24.
Cho, S. G., Bhoumik, A., Broday, L., Ivanov, V., Rosenstein, B. and
Ronai, Z. (2001). TIP49b, a regulator of activating transcription factor 2
response to stress and DNA damage. Mol Cell Biol 21, 8398-413.
Clements, A., Poux, A. N., Lo, W. S., Pillus, L., Berger, S. L. and Marmorstein, R. (2003). Structural basis for histone and phosphohistone binding by the GCN5 histone acetyltransferase. Mol Cell 12, 461-73.
Cortez, D., Wang, Y., Qin, J. and Elledge, S. J. (1999). Requirement of
ATM-dependent phosphorylation of brca1 in the DNA damage response to
double-strand breaks. Science 286, 1162-6.
Costa, F. F. (2005). Non-coding RNAs: new players in eukaryotic biology.
Gene 357, 83-94.
Courey, A. J. and Jia, S. (2001). Transcriptional repression: the long and
the short of it. Genes Dev 15, 2786-96.
Czermin, B., Melfi, R., McCabe, D., Seitz, V., Imhof, A. and Pirrotta, V.
(2002). Drosophila enhancer of Zeste/ESC complexes have a histone H3
methyltransferase activity that marks chromosomal Polycomb sites. Cell 111,
185-96.
Daniel, J. A. and Grant, P. A. (2007). Multi-tasking on chromatin with the
SAGA coactivator complexes. Mutat Res 618, 135-48.
de Napoles, M., Mermoud, J. E., Wakao, R., Tang, Y. A., Endoh, M.,
Appanah, R., Nesterova, T. B., Silva, J., Otte, A. P., Vidal, M. et al.
(2004). Polycomb group proteins Ring1A/B link ubiquitylation of histone
H2A to heritable gene silencing and X inactivation. Dev Cell 7, 663-76.
53
de Ruijter, A. J., van Gennip, A. H., Caron, H. N., Kemp, S. and van
Kuilenburg, A. B. (2003). Histone deacetylases (HDACs): characterization
of the classical HDAC family. Biochem J 370, 737-49.
Dong, X., Tsuda, L., Zavitz, K. H., Lin, M., Li, S., Carthew, R. W. and
Zipursky, S. L. (1999). ebi regulates epidermal growth factor receptor signaling pathways in Drosophila. Genes Dev 13, 954-65.
Downs, J. A., Allard, S., Jobin-Robitaille, O., Javaheri, A., Auger, A.,
Bouchard, N., Kron, S. J., Jackson, S. P. and Cote, J. (2004). Binding of
chromatin-modifying activities to phosphorylated histone H2A at DNA
damage sites. Mol Cell 16, 979-90.
Downs, J. A., Nussenzweig, M. C. and Nussenzweig, A. (2007). Chromatin dynamics and the preservation of genetic information. Nature 447, 951-8.
Doyon, Y., Selleck, W., Lane, W. S., Tan, S. and Cote, J. (2004). Structural and functional conservation of the NuA4 histone acetyltransferase
complex from yeast to humans. Mol Cell Biol 24, 1884-96.
Dubnicoff, T., Valentine, S. A., Chen, G., Shi, T., Lengyel, J. A., Paroush,
Z. and Courey, A. J. (1997). Conversion of dorsal from an activator to a
repressor by the global corepressor Groucho. Genes Dev 11, 2952-7.
Dugan, K. A., Wood, M. A. and Cole, M. D. (2002). TIP49, but not
TRRAP, modulates c-Myc and E2F1 dependent apoptosis. Oncogene 21,
5835-43.
Ebert, A., Lein, S., Schotta, G. and Reuter, G. (2006). Histone modification and the control of heterochromatic gene silencing in Drosophila. Chromosome Res 14, 377-92.
Eissenberg, J. C., Wong, M. and Chrivia, J. C. (2005). Human SRCAP
and Drosophila melanogaster DOM are homologs that function in the notch
signaling pathway. Mol Cell Biol 25, 6559-69.
Erkner, A., Roure, A., Charroux, B., Delaage, M., Holway, N., Core, N.,
Vola, C., Angelats, C., Pages, F., Fasano, L. et al. (2002). Grunge, related
to human Atrophin-like proteins, has multiple functions in Drosophila development. Development 129, 1119-29.
Etard, C., Gradl, D., Kunz, M., Eilers, M. and Wedlich, D. (2005). Pontin
and Reptin regulate cell proliferation in early Xenopus embryos in collaboration with c-Myc and Miz-1. Mech Dev 122, 545-56.
Fang, J., Chen, T., Chadwick, B., Li, E. and Zhang, Y. (2004). Ring1bmediated H2A ubiquitination associates with inactive X chromosomes and is
involved in initiation of X inactivation. J Biol Chem 279, 52812-5.
Fang, M., Li, J., Blauwkamp, T., Bhambhani, C., Campbell, N. and
Cadigan, K. M. (2006). C-terminal-binding protein directly activates and
represses Wnt transcriptional targets in Drosophila. Embo J 25, 2735-45.
Fanto, M., Clayton, L., Meredith, J., Hardiman, K., Charroux, B., Kerridge, S. and McNeill, H. (2003). The tumor-suppressor and cell adhesion
molecule Fat controls planar polarity via physical interactions with Atrophin,
a transcriptional co-repressor. Development 130, 763-74.
Fischle, W., Tseng, B. S., Dormann, H. L., Ueberheide, B. M., Garcia, B.
A., Shabanowitz, J., Hunt, D. F., Funabiki, H. and Allis, C. D. (2005).
54
Regulation of HP1-chromatin binding by histone H3 methylation and phosphorylation. Nature 438, 1116-22.
Fisher, A. L. and Caudy, M. (1998). Groucho proteins: transcriptional
corepressors for specific subsets of DNA-binding transcription factors in
vertebrates and invertebrates. Genes Dev 12, 1931-40.
Flores-Saaib, R. D., Jia, S. and Courey, A. J. (2001). Activation and repression by the C-terminal domain of Dorsal. Development 128, 1869-79.
Frank, S. R., Parisi, T., Taubert, S., Fernandez, P., Fuchs, M., Chan, H.
M., Livingston, D. M. and Amati, B. (2003). MYC recruits the TIP60 histone acetyltransferase complex to chromatin. EMBO Rep 4, 575-80.
Fulco, M., Schiltz, R. L., Iezzi, S., King, M. T., Zhao, P., Kashiwaya, Y.,
Hoffman, E., Veech, R. L. and Sartorelli, V. (2003). Sir2 regulates skeletal
muscle differentiation as a potential sensor of the redox state. Mol Cell 12,
51-62.
Funakoshi, Y., Minami, M. and Tabata, T. (2001). mtv shapes the activity
gradient of the Dpp morphogen through regulation of thickveins. Development 128, 67-74.
Galasinski, S. C., Resing, K. A., Goodrich, J. A. and Ahn, N. G. (2002).
Phosphatase inhibition leads to histone deacetylases 1 and 2 phosphorylation
and disruption of corepressor interactions. J Biol Chem 277, 19618-26.
Gallant, P. (2007). Control of transcription by Pontin and Reptin. Trends
Cell Biol 17, 187-92.
Gao, L., Cueto, M. A., Asselbergs, F. and Atadja, P. (2002). Cloning and
functional characterization of HDAC11, a novel member of the human histone deacetylase family. J Biol Chem 277, 25748-55.
Gaughan, L., Logan, I. R., Cook, S., Neal, D. E. and Robson, C. N.
(2002). Tip60 and histone deacetylase 1 regulate androgen receptor activity
through changes to the acetylation status of the receptor. J Biol Chem 277,
25904-13.
Gavaravarapu, S. and Kamine, J. (2000). Tip60 inhibits activation of
CREB protein by protein kinase A. Biochem Biophys Res Commun 269, 75866.
Glozak, M. A., Sengupta, N., Zhang, X. and Seto, E. (2005). Acetylation
and deacetylation of non-histone proteins. Gene 363, 15-23.
Goldstein, R. E., Cook, O., Dinur, T., Pisante, A., Karandikar, U. C.,
Bidwai, A. and Paroush, Z. (2005). An eh1-like motif in odd-skipped mediates recruitment of Groucho and repression in vivo. Mol Cell Biol 25,
10711-20.
Goldstein, R. E., Jimenez, G., Cook, O., Gur, D. and Paroush, Z. (1999).
Huckebein repressor activity in Drosophila terminal patterning is mediated
by Groucho. Development 126, 3747-55.
Govind, C. K., Zhang, F., Qiu, H., Hofmeyer, K. and Hinnebusch, A. G.
(2007). Gcn5 promotes acetylation, eviction, and methylation of nucleosomes in transcribed coding regions. Mol Cell 25, 31-42.
Grant, P. A., Berger, S. L. and Workman, J. L. (1999). Identification and
analysis of native nucleosomal histone acetyltransferase complexes. Methods
Mol Biol 119, 311-7.
55
Greenblatt, M. S., Bennett, W. P., Hollstein, M. and Harris, C. C. (1994).
Mutations in the p53 tumor suppressor gene: clues to cancer etiology and
molecular pathogenesis. Cancer Res 54, 4855-78.
Gregoire, S., Xiao, L., Nie, J., Zhang, X., Xu, M., Li, J., Wong, J., Seto,
E. and Yang, X. J. (2007). Histone deacetylase 3 interacts with and deacetylates myocyte enhancer factor 2. Mol Cell Biol 27, 1280-95.
Grewal, S. I. and Jia, S. (2007). Heterochromatin revisited. Nat Rev Genet
8, 35-46.
Guarente, L. (2001). SIR2 and aging--the exception that proves the rule.
Trends Genet 17, 391-2.
Guelman, S., Suganuma, T., Florens, L., Swanson, S. K., Kiesecker, C.
L., Kusch, T., Anderson, S., Yates, J. R., 3rd, Washburn, M. P., Abmayr,
S. M. et al. (2006). Host cell factor and an uncharacterized SANT domain
protein are stable components of ATAC, a novel dAda2A/dGcn5-containing
histone acetyltransferase complex in Drosophila. Mol Cell Biol 26, 871-82.
Guenther, M. G., Lane, W. S., Fischle, W., Verdin, E., Lazar, M. A. and
Shiekhattar, R. (2000). A core SMRT corepressor complex containing
HDAC3 and TBL1, a WD40-repeat protein linked to deafness. Genes Dev
14, 1048-57.
Hahn, S. (2004). Structure and mechanism of the RNA polymerase II transcription machinery. Nat Struct Mol Biol 11, 394-403.
Hall, I. M., Shankaranarayana, G. D., Noma, K., Ayoub, N., Cohen, A.
and Grewal, S. I. (2002). Establishment and maintenance of a heterochromatin domain. Science 297, 2232-7.
Harding, K. and Levine, M. (1988). Gap genes define the limits of antennapedia and bithorax gene expression during early development in Drosophila. Embo J 7, 205-14.
Hasson, P., Muller, B., Basler, K. and Paroush, Z. (2001). Brinker requires two corepressors for maximal and versatile repression in Dpp signalling. Embo J 20, 5725-36.
Helmlinger, D., Hardy, S., Eberlin, A., Devys, D. and Tora, L. (2006).
Both normal and polyglutamine- expanded ataxin-7 are components of
TFTC-type GCN5 histone acetyltransferase- containing complexes. Biochem
Soc Symp, 155-63.
Hemavathy, K., Meng, X. and Ip, Y. T. (1997). Differential regulation of
gastrulation and neuroectodermal gene expression by Snail in the Drosophila
embryo. Development 124, 3683-91.
Hirao, A., Kong, Y. Y., Matsuoka, S., Wakeham, A., Ruland, J., Yoshida,
H., Liu, D., Elledge, S. J. and Mak, T. W. (2000). DNA damage-induced
activation of p53 by the checkpoint kinase Chk2. Science 287, 1824-7.
Hoch, M., Gerwin, N., Taubert, H. and Jackle, H. (1992). Competition for
overlapping sites in the regulatory region of the Drosophila gene Kruppel.
Science 256, 94-7.
Holliday, R. (1994). Epigenetics: an overview. Dev Genet 15, 453-7.
Horlein, A. J., Naar, A. M., Heinzel, T., Torchia, J., Gloss, B., Kurokawa,
R., Ryan, A., Kamei, Y., Soderstrom, M., Glass, C. K. et al. (1995).
56
Ligand-independent repression by the thyroid hormone receptor mediated by
a nuclear receptor co-repressor. Nature 377, 397-404.
Hubbert, C., Guardiola, A., Shao, R., Kawaguchi, Y., Ito, A., Nixon, A.,
Yoshida, M., Wang, X. F. and Yao, T. P. (2002). HDAC6 is a microtubule-associated deacetylase. Nature 417, 455-8.
Ikura, T., Ogryzko, V. V., Grigoriev, M., Groisman, R., Wang, J.,
Horikoshi, M., Scully, R., Qin, J. and Nakatani, Y. (2000). Involvement
of the TIP60 histone acetylase complex in DNA repair and apoptosis. Cell
102, 463-73.
Ip, Y. T. and Gridley, T. (2002). Cell movements during gastrulation: snail
dependent and independent pathways. Curr Opin Genet Dev 12, 423-9.
Ip, Y. T., Levine, M. and Small, S. J. (1992). The bicoid and dorsal
morphogens use a similar strategy to make stripes in the Drosophila embryo.
J Cell Sci Suppl 16, 33-8.
Ip, Y. T., Maggert, K. and Levine, M. (1994). Uncoupling gastrulation and
mesoderm differentiation in the Drosophila embryo. Embo J 13, 5826-34.
Jackle, H., Hoch, M., Pankratz, M. J., Gerwin, N., Sauer, F. and Bronner, G. (1992). Transcriptional control by Drosophila gap genes. J Cell Sci
Suppl 16, 39-51.
Jaquet, Y., Delattre, M., Spierer, A. and Spierer, P. (2002). Functional
dissection of the Drosophila modifier of variegation Su(var)3-7. Development 129, 3975-82.
Jiang, J., Cai, H., Zhou, Q. and Levine, M. (1993). Conversion of a dorsal-dependent silencer into an enhancer: evidence for dorsal corepressors.
Embo J 12, 3201-9.
Jimenez, G., Guichet, A., Ephrussi, A. and Casanova, J. (2000). Relief of
gene repression by torso RTK signaling: role of capicua in Drosophila terminal and dorsoventral patterning. Genes Dev 14, 224-31.
Jimenez, G., Paroush, Z. and Ish-Horowicz, D. (1997). Groucho acts as a
corepressor for a subset of negative regulators, including Hairy and Engrailed. Genes Dev 11, 3072-82.
Juan, L. J., Shia, W. J., Chen, M. H., Yang, W. M., Seto, E., Lin, Y. S.
and Wu, C. W. (2000). Histone deacetylases specifically down-regulate
p53-dependent gene activation. J Biol Chem 275, 20436-43.
Kaminker, J. S., Canon, J., Salecker, I. and Banerjee, U. (2002). Control
of photoreceptor axon target choice by transcriptional repression of Runt.
Nat Neurosci 5, 746-50.
Kanazawa, I. (1999). Molecular pathology of dentatorubral-pallidoluysian
atrophy. Philos Trans R Soc Lond B Biol Sci 354, 1069-74.
Kankel, M. W., Duncan, D. M. and Duncan, I. (2004). A screen for genes
that interact with the Drosophila pair-rule segmentation gene fushi tarazu.
Genetics 168, 161-80.
Kao, H. Y., Ordentlich, P., Koyano-Nakagawa, N., Tang, Z., Downes, M.,
Kintner, C. R., Evans, R. M. and Kadesch, T. (1998). A histone deacetylase corepressor complex regulates the Notch signal transduction pathway.
Genes Dev 12, 2269-77.
57
Kasten, M. M., Dorland, S. and Stillman, D. J. (1997). A large protein
complex containing the yeast Sin3p and Rpd3p transcriptional regulators.
Mol Cell Biol 17, 4852-8.
Kawaguchi, Y., Kovacs, J. J., McLaurin, A., Vance, J. M., Ito, A. and
Yao, T. P. (2003). The deacetylase HDAC6 regulates aggresome formation
and cell viability in response to misfolded protein stress. Cell 115, 727-38.
Keller, S. A., Mao, Y., Struffi, P., Margulies, C., Yurk, C. E., Anderson,
A. R., Amey, R. L., Moore, S., Ebels, J. M., Foley, K. et al. (2000).
dCtBP-dependent and -independent repression activities of the Drosophila
Knirps protein. Mol Cell Biol 20, 7247-58.
Khanna, K. K. and Jackson, S. P. (2001). DNA double-strand breaks: signaling, repair and the cancer connection. Nat Genet 27, 247-54.
Kim, J. H., Choi, H. J., Kim, B., Kim, M. H., Lee, J. M., Kim, I. S., Lee,
M. H., Choi, S. J., Kim, K. I., Kim, S. I. et al. (2006). Roles of sumoylation of a reptin chromatin-remodelling complex in cancer metastasis. Nat
Cell Biol 8, 631-9.
Kim, J. H., Kim, B., Cai, L., Choi, H. J., Ohgi, K. A., Tran, C., Chen, C.,
Chung, C. H., Huber, O., Rose, D. W. et al. (2005). Transcriptional regulation of a metastasis suppressor gene by Tip60 and beta-catenin complexes.
Nature 434, 921-6.
Kimura, A. and Horikoshi, M. (1998). Tip60 acetylates six lysines of a
specific class in core histones in vitro. Genes Cells 3, 789-800.
Klymenko, T., Papp, B., Fischle, W., Kocher, T., Schelder, M., Fritsch,
C., Wild, B., Wilm, M. and Muller, J. (2006). A Polycomb group protein
complex with sequence-specific DNA-binding and selective methyl-lysinebinding activities. Genes Dev 20, 1110-22.
Kobayashi, M., Goldstein, R. E., Fujioka, M., Paroush, Z. and Jaynes, J.
B. (2001). Groucho augments the repression of multiple Even skipped target
genes in establishing parasegment boundaries. Development 128, 1805-15.
Kobor, M. S., Venkatasubrahmanyam, S., Meneghini, M. D., Gin, J. W.,
Jennings, J. L., Link, A. J., Madhani, H. D. and Rine, J. (2004). A protein complex containing the conserved Swi2/Snf2-related ATPase Swr1p
deposits histone variant H2A.Z into euchromatin. PLoS Biol 2, E131.
Kohler, A., Pascual-Garcia, P., Llopis, A., Zapater, M., Posas, F., Hurt,
E. and Rodriguez-Navarro, S. (2006). The mRNA export factor Sus1 is
involved in Spt/Ada/Gcn5 acetyltransferase-mediated H2B deubiquitinylation through its interaction with Ubp8 and Sgf11. Mol Biol Cell 17, 4228-36.
Kouzarides, T. (2007). Chromatin modifications and their function. Cell
128, 693-705.
Krogan, N. J., Keogh, M. C., Datta, N., Sawa, C., Ryan, O. W., Ding, H.,
Haw, R. A., Pootoolal, J., Tong, A., Canadien, V. et al. (2003). A Snf2
family ATPase complex required for recruitment of the histone H2A variant
Htz1. Mol Cell 12, 1565-76.
Kumar, V., Carlson, J. E., Ohgi, K. A., Edwards, T. A., Rose, D. W.,
Escalante, C. R., Rosenfeld, M. G. and Aggarwal, A. K. (2002). Transcription corepressor CtBP is an NAD(+)-regulated dehydrogenase. Mol Cell
10, 857-69.
58
Kuo, M. H., vom Baur, E., Struhl, K. and Allis, C. D. (2000). Gcn4 activator targets Gcn5 histone acetyltransferase to specific promoters independently of transcription. Mol Cell 6, 1309-20.
Kurdistani, S. K., Tavazoie, S. and Grunstein, M. (2004). Mapping global
histone acetylation patterns to gene expression. Cell 117, 721-33.
Kusch, T., Florens, L., Macdonald, W. H., Swanson, S. K., Glaser, R. L.,
Yates, J. R., 3rd, Abmayr, S. M., Washburn, M. P. and Workman, J. L.
(2004). Acetylation by Tip60 is required for selective histone variant exchange at DNA lesions. Science 306, 2084-7.
Kusch, T., Guelman, S., Abmayr, S. M. and Workman, J. L. (2003). Two
Drosophila Ada2 homologues function in different multiprotein complexes.
Mol Cell Biol 23, 3305-19.
La Rosee-Borggreve, A., Hader, T., Wainwright, D., Sauer, F. and
Jackle, H. (1999). hairy stripe 7 element mediates activation and repression
in response to different domains and levels of Kruppel in the Drosophila
embryo. Mech Dev 89, 133-40.
LaJeunesse, D. and Shearn, A. (1995). Trans-regulation of thoracic homeotic selector genes of the Antennapedia and bithorax complexes by the
trithorax group genes: absent, small, and homeotic discs 1 and 2. Mech Dev
53, 123-39.
Landry, J., Sutton, A., Hesman, T., Min, J., Xu, R. M., Johnston, M. and
Sternglanz, R. (2003). Set2-catalyzed methylation of histone H3 represses
basal expression of GAL4 in Saccharomyces cerevisiae. Mol Cell Biol 23,
5972-8.
Larschan, E. and Winston, F. (2001). The S. cerevisiae SAGA complex
functions in vivo as a coactivator for transcriptional activation by Gal4.
Genes Dev 15, 1946-56.
Lebrecht, D., Foehr, M., Smith, E., Lopes, F. J., Vanario-Alonso, C. E.,
Reinitz, J., Burz, D. S. and Hanes, S. D. (2005). Bicoid cooperative DNA
binding is critical for embryonic patterning in Drosophila. Proc Natl Acad
Sci U S A 102, 13176-81.
Lee, J. H., Lee, E., Park, J., Kim, E., Kim, J. and Chung, J. (2003). In
vivo p53 function is indispensable for DNA damage-induced apoptotic signaling in Drosophila. FEBS Lett 550, 5-10.
Leptin, M. (1991). twist and snail as positive and negative regulators during
Drosophila mesoderm development. Genes Dev 5, 1568-76.
Leptin, M. (1999). Gastrulation in Drosophila: the logic and the cellular
mechanisms. Embo J 18, 3187-92.
Leptin, M., Casal, J., Grunewald, B. and Reuter, R. (1992). Mechanisms
of early Drosophila mesoderm formation. Dev Suppl, 23-31.
Levanon, D., Goldstein, R. E., Bernstein, Y., Tang, H., Goldenberg, D.,
Stifani, S., Paroush, Z. and Groner, Y. (1998). Transcriptional repression
by AML1 and LEF-1 is mediated by the TLE/Groucho corepressors. Proc
Natl Acad Sci U S A 95, 11590-5.
Li, B., Carey, M. and Workman, J. L. (2007a). The role of chromatin during transcription. Cell 128, 707-19.
59
Li, B., Samanta, A., Song, X., Iacono, K. T., Bembas, K., Tao, R., Basu,
S., Riley, J. L., Hancock, W. W., Shen, Y. et al. (2007b). FOXP3 interactions with histone acetyltransferase and class II histone deacetylases are required for repression. Proc Natl Acad Sci U S A 104, 4571-6.
Li, J., Wang, J., Wang, J., Nawaz, Z., Liu, J. M., Qin, J. and Wong, J.
(2000). Both corepressor proteins SMRT and N-CoR exist in large protein
complexes containing HDAC3. Embo J 19, 4342-50.
Li, S., Xu, C. and Carthew, R. W. (2002). Phyllopod acts as an adaptor
protein to link the sina ubiquitin ligase to the substrate protein tramtrack.
Mol Cell Biol 22, 6854-65.
Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F. and Richmond, T. J. (1997). Crystal structure of the nucleosome core particle at 2.8
A resolution. Nature 389, 251-60.
Luo, J., Nikolaev, A. Y., Imai, S., Chen, D., Su, F., Shiloh, A., Guarente,
L. and Gu, W. (2001). Negative control of p53 by Sir2alpha promotes cell
survival under stress. Cell 107, 137-48.
Makino, Y., Kanemaki, M., Kurokawa, Y., Koji, T. and Tamura, T.
(1999). A rat RuvB-like protein, TIP49a, is a germ cell-enriched novel DNA
helicase. J Biol Chem 274, 15329-35.
Malave, T. M. and Dent, S. Y. (2006). Transcriptional repression by Tup1Ssn6. Biochem Cell Biol 84, 437-43.
Mannervik, M. and Levine, M. (1999). The Rpd3 histone deacetylase is
required for segmentation of the Drosophila embryo. Proc Natl Acad Sci U S
A 96, 6797-801.
Mannervik, M., Nibu, Y., Zhang, H. and Levine, M. (1999). Transcriptional coregulators in development. Science 284, 606-9.
Margueron, R., Trojer, P. and Reinberg, D. (2005). The key to development: interpreting the histone code? Curr Opin Genet Dev 15, 163-76.
Matsuoka, S., Huang, M. and Elledge, S. J. (1998). Linkage of ATM to
cell cycle regulation by the Chk2 protein kinase. Science 282, 1893-7.
Matsuyama, A., Shimazu, T., Sumida, Y., Saito, A., Yoshimatsu, Y.,
Seigneurin-Berny, D., Osada, H., Komatsu, Y., Nishino, N., Khochbin, S.
et al. (2002). In vivo destabilization of dynamic microtubules by HDAC6mediated deacetylation. Embo J 21, 6820-31.
Matsuzawa, S. I. and Reed, J. C. (2001). Siah-1, SIP, and Ebi collaborate
in a novel pathway for beta-catenin degradation linked to p53 responses. Mol
Cell 7, 915-26.
Meneghini, M. D., Wu, M. and Madhani, H. D. (2003). Conserved histone
variant H2A.Z protects euchromatin from the ectopic spread of silent heterochromatin. Cell 112, 725-36.
Mizuguchi, G., Shen, X., Landry, J., Wu, W. H., Sen, S. and Wu, C.
(2004). ATP-driven exchange of histone H2AZ variant catalyzed by SWR1
chromatin remodeling complex. Science 303, 343-8.
Moran, E. and Jimenez, G. (2006). The tailless nuclear receptor acts as a
dedicated repressor in the early Drosophila embryo. Mol Cell Biol 26, 344654.
60
Morel, V., Lecourtois, M., Massiani, O., Maier, D., Preiss, A. and
Schweisguth, F. (2001). Transcriptional repression by suppressor of hairless
involves the binding of a hairless-dCtBP complex in Drosophila. Curr Biol
11, 789-92.
Morisato, D. (2001). Spatzle regulates the shape of the Dorsal gradient in
the Drosophila embryo. Development 128, 2309-19.
Moussian, B. and Roth, S. (2005). Dorsoventral axis formation in the Drosophila embryo--shaping and transducing a morphogen gradient. Curr Biol
15, R887-99.
Munshi, N., Agalioti, T., Lomvardas, S., Merika, M., Chen, G. and
Thanos, D. (2001). Coordination of a transcriptional switch by HMGI(Y)
acetylation. Science 293, 1133-6.
Muratoglu, S., Georgieva, S., Papai, G., Scheer, E., Enunlu, I., Komonyi,
O., Cserpan, I., Lebedeva, L., Nabirochkina, E., Udvardy, A. et al.
(2003). Two different Drosophila ADA2 homologues are present in distinct
GCN5 histone acetyltransferase-containing complexes. Mol Cell Biol 23,
306-21.
Murr, R., Loizou, J. I., Yang, Y. G., Cuenin, C., Li, H., Wang, Z. Q. and
Herceg, Z. (2006). Histone acetylation by Trrap-Tip60 modulates loading of
repair proteins and repair of DNA double-strand breaks. Nat Cell Biol 8, 919.
Nasiadka, A., Grill, A. and Krause, H. M. (2000). Mechanisms regulating
target gene selection by the homeodomain-containing protein Fushi tarazu.
Development 127, 2965-76.
Nasiadka, A. and Krause, H. M. (1999). Kinetic analysis of segmentation
gene interactions in Drosophila embryos. Development 126, 1515-26.
Nibu, Y. and Levine, M. S. (2001). CtBP-dependent activities of the shortrange Giant repressor in the Drosophila embryo. Proc Natl Acad Sci U S A
98, 6204-8.
Nibu, Y., Zhang, H., Bajor, E., Barolo, S., Small, S. and Levine, M.
(1998a). dCtBP mediates transcriptional repression by Knirps, Kruppel and
Snail in the Drosophila embryo. Embo J 17, 7009-20.
Nibu, Y., Zhang, H. and Levine, M. (1998b). Interaction of short-range
repressors with Drosophila CtBP in the embryo. Science 280, 101-4.
Niessing, D., Rivera-Pomar, R., La Rosee, A., Hader, T., Schock, F.,
Purnell, B. A. and Jackle, H. (1997). A cascade of transcriptional control
leading to axis determination in Drosophila. J Cell Physiol 173, 162-7.
Nieto, M. A. (2002). The snail superfamily of zinc-finger transcription factors. Nat Rev Mol Cell Biol 3, 155-66.
Ochoa-Espinosa, A., Yucel, G., Kaplan, L., Pare, A., Pura, N., Oberstein,
A., Papatsenko, D. and Small, S. (2005). The role of binding site cluster
strength in Bicoid-dependent patterning in Drosophila. Proc Natl Acad Sci U
S A 102, 4960-5.
Ogryzko, V. (2000). Mammalian histone acetyltransferase complexes.
Medicina (B Aires) 60 Suppl 2, 21-6.
Ogura, T. and Wilkinson, A. J. (2001). AAA+ superfamily ATPases:
common structure--diverse function. Genes Cells 6, 575-97.
61
Ohdate, H., Lim, C. R., Kokubo, T., Matsubara, K., Kimata, Y. and
Kohno, K. (2003). Impairment of the DNA binding activity of the TATAbinding protein renders the transcriptional function of Rvb2p/Tih2p, the
yeast RuvB-like protein, essential for cell growth. J Biol Chem 278, 1464756.
Oren, M. (2003). Decision making by p53: life, death and cancer. Cell
Death Differ 10, 431-42.
Pal-Bhadra, M., Leibovitch, B. A., Gandhi, S. G., Rao, M., Bhadra, U.,
Birchler, J. A. and Elgin, S. C. (2004). Heterochromatic silencing and HP1
localization in Drosophila are dependent on the RNAi machinery. Science
303, 669-72.
Pan, D. and Courey, A. J. (1992). The same dorsal binding site mediates
both activation and repression in a context-dependent manner. Embo J 11,
1837-42.
Pankotai, T., Komonyi, O., Bodai, L., Ujfaludi, Z., Muratoglu, S., Ciurciu, A., Tora, L., Szabad, J. and Boros, I. (2005). The homologous Drosophila transcriptional adaptors ADA2a and ADA2b are both required for
normal development but have different functions. Mol Cell Biol 25, 8215-27.
Papoulas, O., Beek, S. J., Moseley, S. L., McCallum, C. M., Sarte, M.,
Shearn, A. and Tamkun, J. W. (1998). The Drosophila trithorax group
proteins BRM, ASH1 and ASH2 are subunits of distinct protein complexes.
Development 125, 3955-66.
Paroush, Z., Finley, R. L., Jr., Kidd, T., Wainwright, S. M., Ingham, P.
W., Brent, R. and Ish-Horowicz, D. (1994). Groucho is required for Drosophila neurogenesis, segmentation, and sex determination and interacts
directly with hairy-related bHLH proteins. Cell 79, 805-15.
Patel, J. H., Du, Y., Ard, P. G., Phillips, C., Carella, B., Chen, C. J., Rakowski, C., Chatterjee, C., Lieberman, P. M., Lane, W. S. et al. (2004).
The c-MYC oncoprotein is a substrate of the acetyltransferases
hGCN5/PCAF and TIP60. Mol Cell Biol 24, 10826-34.
Perissi, V., Aggarwal, A., Glass, C. K., Rose, D. W. and Rosenfeld, M. G.
(2004). A corepressor/coactivator exchange complex required for transcriptional activation by nuclear receptors and other regulated transcription factors. Cell 116, 511-26.
Peterson, C. L. (2002). Chromatin remodeling: nucleosomes bulging at the
seams. Curr Biol 12, R245-7.
Peterson, C. L. and Cote, J. (2004). Cellular machineries for chromosomal
DNA repair. Genes Dev 18, 602-16.
Petruk, S., Sedkov, Y., Smith, S., Tillib, S., Kraevski, V., Nakamura, T.,
Canaani, E., Croce, C. M. and Mazo, A. (2001). Trithorax and dCBP acting in a complex to maintain expression of a homeotic gene. Science 294,
1331-4.
Plaster, N., Sonntag, C., Schilling, T. F. and Hammerschmidt, M. (2007).
REREa/Atrophin-2 interacts with histone deacetylase and Fgf8 signaling to
regulate multiple processes of zebrafish development. Dev Dyn 236, 1891904.
62
Poortinga, G., Watanabe, M. and Parkhurst, S. M. (1998). Drosophila
CtBP: a Hairy-interacting protein required for embryonic segmentation and
hairy-mediated transcriptional repression. Embo J 17, 2067-78.
Puri, T., Wendler, P., Sigala, B., Saibil, H. and Tsaneva, I. R. (2007).
Dodecameric structure and ATPase activity of the human TIP48/TIP49
complex. J Mol Biol 366, 179-92.
Qi, D., Larsson, J. and Mannervik, M. (2004). Drosophila Ada2b is required for viability and normal histone H3 acetylation. Mol Cell Biol 24,
8080-9.
Qin, S. and Parthun, M. R. (2006). Recruitment of the type B histone acetyltransferase Hat1p to chromatin is linked to DNA double-strand breaks.
Mol Cell Biol 26, 3649-58.
Rao, Y., Pang, P., Ruan, W., Gunning, D. and Zipursky, S. L. (2000).
brakeless is required for photoreceptor growth-cone targeting in Drosophila.
Proc Natl Acad Sci U S A 97, 5966-71.
Regulski, M., Harding, K., Kostriken, R., Karch, F., Levine, M. and
McGinnis, W. (1985). Homeo box genes of the Antennapedia and bithorax
complexes of Drosophila. Cell 43, 71-80.
Ricci, A. R., Genereaux, J. and Brandl, C. J. (2002). Components of the
SAGA histone acetyltransferase complex are required for repressed transcription of ARG1 in rich medium. Mol Cell Biol 22, 4033-42.
Ringrose, L. and Paro, R. (2004). Epigenetic regulation of cellular memory
by the Polycomb and Trithorax group proteins. Annu Rev Genet 38, 413-43.
Rivera-Pomar, R., Lu, X., Perrimon, N., Taubert, H. and Jackle, H.
(1995). Activation of posterior gap gene expression in the Drosophila blastoderm. Nature 376, 253-6.
Rogakou, E. P., Pilch, D. R., Orr, A. H., Ivanova, V. S. and Bonner, W.
M. (1998). DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273, 5858-68.
Rosenfeld, M. G., Lunyak, V. V. and Glass, C. K. (2006). Sensors and
signals: a coactivator/corepressor/epigenetic code for integrating signaldependent programs of transcriptional response. Genes Dev 20, 1405-28.
Roth, S. (2003). The origin of dorsoventral polarity in Drosophila. Philos
Trans R Soc Lond B Biol Sci 358, 1317-29; discussion 1329.
Roth, S. Y., Denu, J. M. and Allis, C. D. (2001). Histone acetyltransferases.
Annu Rev Biochem 70, 81-120.
Rottbauer, W., Saurin, A. J., Lickert, H., Shen, X., Burns, C. G., Wo, Z.
G., Kemler, R., Kingston, R., Wu, C. and Fishman, M. (2002). Reptin and
pontin antagonistically regulate heart growth in zebrafish embryos. Cell 111,
661-72.
Rudolph, T., Yonezawa, M., Lein, S., Heidrich, K., Kubicek, S., Schafer,
C., Phalke, S., Walther, M., Schmidt, A., Jenuwein, T. et al. (2007). Heterochromatin formation in Drosophila is initiated through active removal of
H3K4 methylation by the LSD1 homolog SU(VAR)3-3. Mol Cell 26, 103-15.
Rusch, J. and Levine, M. (1996). Threshold responses to the dorsal regulatory gradient and the subdivision of primary tissue territories in the Drosophila embryo. Curr Opin Genet Dev 6, 416-23.
63
Ryu, J. R. and Arnosti, D. N. (2003). Functional similarity of Knirps CtBPdependent and CtBP-independent transcriptional repressor activities. Nucleic
Acids Res 31, 4654-62.
Sapountzi, V., Logan, I. R. and Robson, C. N. (2006). Cellular functions
of TIP60. Int J Biochem Cell Biol 38, 1496-509.
Saurin, A. J., Shao, Z., Erdjument-Bromage, H., Tempst, P. and Kingston, R. E. (2001). A Drosophila Polycomb group complex includes Zeste
and dTAFII proteins. Nature 412, 655-60.
Schreiber-Agus, N. and DePinho, R. A. (1998). Repression by the
Mad(Mxi1)-Sin3 complex. Bioessays 20, 808-18.
Schuettengruber, B., Chourrout, D., Vervoort, M., Leblanc, B. and
Cavalli, G. (2007). Genome regulation by polycomb and trithorax proteins.
Cell 128, 735-45.
Seligson, D. B., Horvath, S., Shi, T., Yu, H., Tze, S., Grunstein, M. and
Kurdistani, S. K. (2005). Global histone modification patterns predict risk
of prostate cancer recurrence. Nature 435, 1262-6.
Senti, K., Keleman, K., Eisenhaber, F. and Dickson, B. J. (2000). brakeless is required for lamina targeting of R1-R6 axons in the Drosophila visual
system. Development 127, 2291-301.
Shen, X., Mizuguchi, G., Hamiche, A. and Wu, C. (2000). A chromatin
remodelling complex involved in transcription and DNA processing. Nature
406, 541-4.
Shen, Y., Lee, G., Choe, Y., Zoltewicz, J. S. and Peterson, A. S. (2007).
Functional architecture of atrophins. J Biol Chem 282, 5037-44.
Shukla, A., Bajwa, P. and Bhaumik, S. R. (2006). SAGA-associated
Sgf73p facilitates formation of the preinitiation complex assembly at the
promoters either in a HAT-dependent or independent manner in vivo. Nucleic Acids Res 34, 6225-32.
Silverstein, R. A. and Ekwall, K. (2005). Sin3: a flexible regulator of
global gene expression and genome stability. Curr Genet 47, 1-17.
Simon, J., Chiang, A. and Bender, W. (1992). Ten different Polycomb
group genes are required for spatial control of the abdA and AbdB homeotic
products. Development 114, 493-505.
Small, S., Blair, A. and Levine, M. (1992). Regulation of even-skipped
stripe 2 in the Drosophila embryo. Embo J 11, 4047-57.
Smith, M. M. (2002). Histone variants and nucleosome deposition pathways.
Mol Cell 9, 1158-60.
Song, H., Hasson, P., Paroush, Z. and Courey, A. J. (2004). Groucho
oligomerization is required for repression in vivo. Mol Cell Biol 24, 4341-50.
St Johnston, D. and Nusslein-Volhard, C. (1992). The origin of pattern
and polarity in the Drosophila embryo. Cell 68, 201-19.
Stathopoulos, A. and Levine, M. (2002). Dorsal gradient networks in the
Drosophila embryo. Dev Biol 246, 57-67.
Stathopoulos, A., Van Drenth, M., Erives, A., Markstein, M. and Levine,
M. (2002). Whole-genome analysis of dorsal-ventral patterning in the Drosophila embryo. Cell 111, 687-701.
64
Stewart, M. D., Li, J. and Wong, J. (2005). Relationship between histone
H3 lysine 9 methylation, transcription repression, and heterochromatin protein 1 recruitment. Mol Cell Biol 25, 2525-38.
Stifani, S., Blaumueller, C. M., Redhead, N. J., Hill, R. E. and Artavanis-Tsakonas, S. (1992). Human homologs of a Drosophila Enhancer of
split gene product define a novel family of nuclear proteins. Nat Genet 2,
343.
Stockinger, E. J., Mao, Y., Regier, M. K., Triezenberg, S. J. and
Thomashow, M. F. (2001). Transcriptional adaptor and histone acetyltransferase proteins in Arabidopsis and their interactions with CBF1, a transcriptional activator involved in cold-regulated gene expression. Nucleic Acids
Res 29, 1524-33.
Strahl, B. D. and Allis, C. D. (2000). The language of covalent histone
modifications. Nature 403, 41-5.
Strahl, B. D., Grant, P. A., Briggs, S. D., Sun, Z. W., Bone, J. R., Caldwell, J. A., Mollah, S., Cook, R. G., Shabanowitz, J., Hunt, D. F. et al.
(2002). Set2 is a nucleosomal histone H3-selective methyltransferase that
mediates transcriptional repression. Mol Cell Biol 22, 1298-306.
Strunk, B., Struffi, P., Wright, K., Pabst, B., Thomas, J., Qin, L. and
Arnosti, D. N. (2001). Role of CtBP in transcriptional repression by the
Drosophila giant protein. Dev Biol 239, 229-40.
Sundqvist, A., Sollerbrant, K. and Svensson, C. (1998). The carboxyterminal region of adenovirus E1A activates transcription through targeting
of a C-terminal binding protein-histone deacetylase complex. FEBS Lett 429,
183-8.
Swaminathan, J., Baxter, E. M. and Corces, V. G. (2005). The role of
histone H2Av variant replacement and histone H4 acetylation in the establishment of Drosophila heterochromatin. Genes Dev 19, 65-76.
Tamburini, B. A. and Tyler, J. K. (2005). Localized histone acetylation
and deacetylation triggered by the homologous recombination pathway of
double-strand DNA repair. Mol Cell Biol 25, 4903-13.
Taubert, S., Gorrini, C., Frank, S. R., Parisi, T., Fuchs, M., Chan, H. M.,
Livingston, D. M. and Amati, B. (2004). E2F-dependent histone acetylation and recruitment of the Tip60 acetyltransferase complex to chromatin in
late G1. Mol Cell Biol 24, 4546-56.
Teng, Y., Yu, Y. and Waters, R. (2002). The Saccharomyces cerevisiae
histone acetyltransferase Gcn5 has a role in the photoreactivation and nucleotide excision repair of UV-induced cyclobutane pyrimidine dimers in the
MFA2 gene. J Mol Biol 316, 489-99.
Towb, P., Galindo, R. L. and Wasserman, S. A. (1998). Recruitment of
Tube and Pelle to signaling sites at the surface of the Drosophila embryo.
Development 125, 2443-50.
Tsai, C. C., Kao, H. Y., Yao, T. P., McKeown, M. and Evans, R. M.
(1999). SMRTER, a Drosophila nuclear receptor coregulator, reveals that
EcR-mediated repression is critical for development. Mol Cell 4, 175-86.
65
Tsuda, L., Kaido, M., Lim, Y. M., Kato, K., Aigaki, T. and Hayashi, S.
(2006). An NRSF/REST-like repressor downstream of Ebi/SMRTER/Su(H)
regulates eye development in Drosophila. Embo J 25, 3191-202.
Tsuda, L., Nagaraj, R., Zipursky, S. L. and Banerjee, U. (2002). An
EGFR/Ebi/Sno pathway promotes delta expression by inactivating
Su(H)/SMRTER repression during inductive notch signaling. Cell 110, 62537.
Utley, R. T. and Cote, J. (2003). The MYST family of histone acetyltransferases. Curr Top Microbiol Immunol 274, 203-36.
Utley, R. T., Ikeda, K., Grant, P. A., Cote, J., Steger, D. J., Eberharter,
A., John, S. and Workman, J. L. (1998). Transcriptional activators direct
histone acetyltransferase complexes to nucleosomes. Nature 394, 498-502.
Valentine, S. A., Chen, G., Shandala, T., Fernandez, J., Mische, S., Saint,
R. and Courey, A. J. (1998). Dorsal-mediated repression requires the formation of a multiprotein repression complex at the ventral silencer. Mol Cell
Biol 18, 6584-94.
van Attikum, H. and Gasser, S. M. (2005). ATP-dependent chromatin
remodeling and DNA double-strand break repair. Cell Cycle 4, 1011-4.
Vega, R. B., Matsuda, K., Oh, J., Barbosa, A. C., Yang, X., Meadows, E.,
McAnally, J., Pomajzl, C., Shelton, J. M., Richardson, J. A. et al. (2004).
Histone deacetylase 4 controls chondrocyte hypertrophy during skeletogenesis. Cell 119, 555-66.
Verdel, A. and Moazed, D. (2005). RNAi-directed assembly of heterochromatin in fission yeast. FEBS Lett 579, 5872-8.
Verdin, E., Dequiedt, F. and Kasler, H. G. (2003). Class II histone deacetylases: versatile regulators. Trends Genet 19, 286-93.
Vlachonasios, K. E., Thomashow, M. F. and Triezenberg, S. J. (2003).
Disruption mutations of ADA2b and GCN5 transcriptional adaptor genes
dramatically affect Arabidopsis growth, development, and gene expression.
Plant Cell 15, 626-38.
Wahi, M., Komachi, K. and Johnson, A. D. (1998). Gene regulation by the
yeast Ssn6-Tup1 corepressor. Cold Spring Harb Symp Quant Biol 63, 44757.
Wang, H., Wang, L., Erdjument-Bromage, H., Vidal, M., Tempst, P.,
Jones, R. S. and Zhang, Y. (2004). Role of histone H2A ubiquitination in
Polycomb silencing. Nature 431, 873-8.
Wang, L., Rajan, H., Pitman, J. L., McKeown, M. and Tsai, C. C. (2006).
Histone deacetylase-associating Atrophin proteins are nuclear receptor
corepressors. Genes Dev 20, 525-30.
Wedeen, C., Harding, K. and Levine, M. (1986). Spatial regulation of
Antennapedia and bithorax gene expression by the Polycomb locus in Drosophila. Cell 44, 739-48.
Wehn, A. and Campbell, G. (2006). Genetic interactions among scribbler,
Atrophin and groucho in Drosophila uncover links in transcriptional repression. Genetics 173, 849-61.
Wiren, M., Silverstein, R. A., Sinha, I., Walfridsson, J., Lee, H. M.,
Laurenson, P., Pillus, L., Robyr, D., Grunstein, M. and Ekwall, K.
66
(2005). Genomewide analysis of nucleosome density histone acetylation and
HDAC function in fission yeast. Embo J 24, 2906-18.
Wood, M. A., McMahon, S. B. and Cole, M. D. (2000). An ATPase/helicase complex is an essential cofactor for oncogenic transformation
by c-Myc. Mol Cell 5, 321-30.
Wurtele, H. and Verreault, A. (2006). Histone post-translational modifications and the response to DNA double-strand breaks. Curr Opin Cell Biol 18,
137-44.
Xiao, H., Chung, J., Kao, H. Y. and Yang, Y. C. (2003). Tip60 is a corepressor for STAT3. J Biol Chem 278, 11197-204.
Xu, L., Lavinsky, R. M., Dasen, J. S., Flynn, S. E., McInerney, E. M.,
Mullen, T. M., Heinzel, T., Szeto, D., Korzus, E., Kurokawa, R. et al.
(1998). Signal-specific co-activator domain requirements for Pit-1 activation.
Nature 395, 301-6.
Xu, W., Edmondson, D. G., Evrard, Y. A., Wakamiya, M., Behringer, R.
R. and Roth, S. Y. (2000). Loss of Gcn5l2 leads to increased apoptosis and
mesodermal defects during mouse development. Nat Genet 26, 229-32.
Yamamoto, T. and Horikoshi, M. (1997). Novel substrate specificity of the
histone acetyltransferase activity of HIV-1-Tat interactive protein Tip60. J
Biol Chem 272, 30595-8.
Yamauchi, T., Yamauchi, J., Kuwata, T., Tamura, T., Yamashita, T.,
Bae, N., Westphal, H., Ozato, K. and Nakatani, Y. (2000). Distinct but
overlapping roles of histone acetylase PCAF and of the closely related
PCAF-B/GCN5 in mouse embryogenesis. Proc Natl Acad Sci U S A 97,
11303-6.
Yang, J. and Steward, R. (1997). A multimeric complex and the nuclear
targeting of the Drosophila Rel protein Dorsal. Proc Natl Acad Sci U S A 94,
14524-9.
Yang, P., Shaver, S. A., Hilliker, A. J. and Sokolowski, M. B. (2000).
Abnormal turning behavior in Drosophila larvae. Identification and molecular analysis of scribbler (sbb). Genetics 155, 1161-74.
Yoon, H. G., Chan, D. W., Huang, Z. Q., Li, J., Fondell, J. D., Qin, J.
and Wong, J. (2003). Purification and functional characterization of the
human N-CoR complex: the roles of HDAC3, TBL1 and TBLR1. Embo J 22,
1336-46.
Yoon, H. G., Choi, Y., Cole, P. A. and Wong, J. (2005). Reading and function of a histone code involved in targeting corepressor complexes for repression. Mol Cell Biol 25, 324-35.
You, A., Tong, J. K., Grozinger, C. M. and Schreiber, S. L. (2001).
CoREST is an integral component of the CoREST- human histone deacetylase complex. Proc Natl Acad Sci U S A 98, 1454-8.
Zhang, C. L., McKinsey, T. A., Chang, S., Antos, C. L., Hill, J. A. and
Olson, E. N. (2002a). Class II histone deacetylases act as signal-responsive
repressors of cardiac hypertrophy. Cell 110, 479-88.
Zhang, C. L., Zou, Y., Yu, R. T., Gage, F. H. and Evans, R. M. (2006a).
Nuclear receptor TLX prevents retinal dystrophy and recruits the corepressor
atrophin1. Genes Dev 20, 1308-20.
67
Zhang, F., Thomas, L. R., Oltz, E. M. and Aune, T. M. (2006b). Control
of thymocyte development and recombination-activating gene expression by
the zinc finger protein Zfp608. Nat Immunol 7, 1309-16.
Zhang, H. and Levine, M. (1999). Groucho and dCtBP mediate separate
pathways of transcriptional repression in the Drosophila embryo. Proc Natl
Acad Sci U S A 96, 535-40.
Zhang, S., Xu, L., Lee, J. and Xu, T. (2002b). Drosophila atrophin homolog functions as a transcriptional corepressor in multiple developmental
processes. Cell 108, 45-56.
Zhang, W., Deng, H., Bao, X., Lerach, S., Girton, J., Johansen, J. and
Johansen, K. M. (2006c). The JIL-1 histone H3S10 kinase regulates dimethyl H3K9 modifications and heterochromatic spreading in Drosophila.
Development 133, 229-35.
Zhang, Y., Ng, H. H., Erdjument-Bromage, H., Tempst, P., Bird, A. and
Reinberg, D. (1999). Analysis of the NuRD subunits reveals a histone
deacetylase core complex and a connection with DNA methylation. Genes
Dev 13, 1924-35.
Zoltewicz, J. S., Stewart, N. J., Leung, R. and Peterson, A. S. (2004).
Atrophin 2 recruits histone deacetylase and is required for the function of
multiple signaling centers during mouse embryogenesis. Development 131,
3-14.
68
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