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 Dro- sophila 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 Snail- dependent 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 Adaptor protein 2 AP axis Anterior-posterior axis ATM Ataxia-telangiectasia mutated ATR ATM and Rad-3 related Atro Atrophin Bks Brakeless CRM Cis regulatory module CtBP C-terminal binding protein DV axis Dorsal-ventral axis Gro Groucho GTF General transcription factor H3K9 Histone H3 lysine 9 HAT Histone acetyl transferase Hb Hunchback HDAC Histone deacetylase HMT Histone methyl transferase HP1 Heterochromatin protein 1 Knirps Kni Kruppel Kr NCoR Nuclear receptor corepressor PcG Polycomb group PEV Position effect variegation PRC1 Polycomb repressive complex 1 Rho Rhomboid Sim Single-minded SMRT Silencing mediator for retinoic acid and thyroid hormone receptors Sog Short-gastrulation TF Transcription factor Tip60 Tat interaction protein 60kD Tll Tailless TrxG 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 RNA pol II A D F TATA E H

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). Tran- scription 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 poly-

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).

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 chemical- induced 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 so- matic 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 Dro- sophila 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

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

amnioserosa B A

zen Dorsal

Dorsal ectoderm Acron

s Telson

o Dorsal gradient h r Posterior Ventral ectoderm Anterior Sna Ventral Head Tail Mesoderm Thorax 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 mi- gration 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-

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 Dor- sal 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)

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 combina- tory role with other factors is also involved in regulating Dorsal target genes. Twist protein regulates nearly half of the known Dorsal target genes syner- gistically 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, cell- cycle 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

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. An- other 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

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 Gi- ant 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 uni- formly 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.,

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 pro- ducing the seven stripes. This enhancer autonomy is partially due to short- range 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 Bitho- rax 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).

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 Ac Ac Ac Ac U H2A S K K K K K K 1 5 9 13 15 36 119 M Ac Ac P Ac Ac M Ac U H2B K K S K K K K K 5 12 14 15 20 23 24 120 M M M P M AcAc P Ac Ac M Ac Ac M M P Is M M Is Ac H3 R T K R K S T K R K K R K S P K K P K 2 3 4 8 9 10 11 14 17 18 23 26 2728 30 36 37 38 56 M M P M Ac Ac Ac Ac Ac H4 S R K K K K K 1 3 5 8 12 16 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.

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). Het- erochromatin 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).

HP1 Eaf3 Pc Rsc4 14-3-3 CHD1

Bdf1 Brd2 Taf1 Bromo Chromo Chromo JMJD2A Chromo Chromo CRB2 Ac P Me Me Me T H3 PH B Bromo Me D M H4 Bromo Bromo Tudor K K S K K K Ac 27 14 10 9 L(3)MBTL Ac Ac Me 36 BPTF 4 TAF3 K K K ING2 8 12 16 K 20 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.

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 uni- versal.

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. How- ever, 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).

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 is- lands via proteins such as methyl-CpG binding proteins and methyl-CpG- binding-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

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 P H2Av

Sensors Rad17, Rad9, Rad1, Hus1, Mei-41(ATR) P MRN complex P dATM

Chk2 P Mus304 P Transducers ATM, ATR, DNA-PK

P p53 Grp(Chk1) P

CDC25 Effectors 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

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 higher- order 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 CBP Type II: HDAC4, 5, 7, 9 SRC-1 HDAC6, 10 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.

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 chromatin- assembly-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 TBP- associated 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

Ubp8 and Sgf11 (Kohler et al., 2006). Both normal and polyglutamine- ex- panded 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 GCN5- null 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 c- Myc, 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). Sub- sequently, 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

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 C- terminal 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).

Tip60Tip60 E(Pc)E(Pc) ING3ING3 TRRAPTRRAP DominoDomino DMAP1Reptin Pontin BAF53BAF53DMAP1Reptin Pontin Eaf6 AActinctin Eaf6 Brd8Brd8 GAS41GAS41 MRG15MRG15 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) (En- hancer of Polycomb)) and ING3 (yeast Yng2, Drosophila dIng3) form a core

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. Dom- ino/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 Tip60- coactivated 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

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). Interest- ingly, Reptin and Pontin may play similar or antagonistic roles in a context- dependent 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 sumoy- lated 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.

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 zinc- catalyzed 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 protein- protein 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; Kawa- guchi 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).

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 deacetylase- activating domain (DAD) that is essential for HDAC3 activation. Addition- ally, 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 well- studied 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 receptor- regulated 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

(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 ab- sentia) 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. Heterochro- matin 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

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 de- methylase (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 com-

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 (Pleio- homeotic) 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).

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 De- Pinho, 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 long-

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 activator- coactivator 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

Co- Repressor Family Range Reference repressor 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)

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 Droso- phila 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 CtBP- interacting 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 eluci- dated. 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). Further- more, 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

(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 WRPW- like 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 Bus- carlet 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 con- verted 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 expan- sion 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 malforma- tion 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

(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 N- terminal 929 amino acids, but BksB is longer and has a 1373-residue C- terminal 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 (recombination- activating gene1) and rag2 expression (Zhang et al., 2006b).

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.

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. Contradic- tory 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

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. Fur- ther, 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 Droso- phila (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 Dom- ino. 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

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. How- ever, 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; Me- neghini et al., 2003). However, no change was found in binding of H2Av to polytene chromosomes from domino mutant larvae. It is possible that PRC1- mediated 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 Snail- target genes requires both dCtBP and Ebi, and that Snail recruits both CtBP

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 a.a. 245 Zinc fingers a.a. 390 Sna P-DLS-K P-DLS-R DNA-binding CtBP CtBP

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-DLS- R. 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 pro-

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 simulta- neously 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.

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 co- factors, 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 interest- ing 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 influ- ence 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 Droso- phila development.

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 sup- portive 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 occa- sional 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 orga- nizing the nice boat trip; Satish, for being a good party-mood maker; Vasil- ios, for showing us how devoted a scientist should be; Nicole, Naumi, Olivi- ano, Chie and Ryo, for creating a good working atmosphere.

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 mov- ing, 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.

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