Chromatin Remodelling by BRG1 and SNF2H190933/FULLTEXT01.pdf · Chromatin remodeling by BRG1 and SNF2H Biochemistry and Function 4 This thesis is based on the following papers: Paper

Chromatin Remodeling by BRG1 and SNF2H Biochemistry and Function

Patrik Asp

Department of Cell Biology Wenner‐Gren Institute Stockholm University

Stockholm 2004

Chromatin remodeling by BRG1 and SNF2H Biochemistry and Function

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ʺThe nature of reality excites me far more than any of the many alternatives contrived by ourselvesʺ KMPA. 2004

© Patrik Asp ISBN 91‐7265‐914‐9

Akademitryck, Edsbruk 2004

Introduction 3

SUMMARY Chromatin is a highly dynamic, regulatory component in the process of transcription, repair, recombination and replication. The BRG1 and SNF2H proteins are ATP‐dependent chromatin remodeling proteins that modulate chromatin structure to regulate DNA accessibility for DNA‐binding proteins involved in these processes. The BRG1 protein is a central ATPase of the SWI/SNF complexes involved in chromatin remodeling associated with regulation of transcription. SWI/SNF complexes are biochemically hetero‐geneous but little is known about the unique functional characteristics of the various forms. We have shown that SWI/SNF activity in SW13 cells affects actin filament organization dependent on the RhoA signaling pathway. We have further shown that the biochemical composition of SWI/SNF complexes qualitatively affects the remodeling activity and that the composition of biochemically purified SWI/SNF complexes does not reflect the patterns of chromatin binding of individual subunits. Chromatin binding assays (ChIP) reveal variations among subunits believed to be constitutive, suggesting that the plasticity in SWI/SNF complex composition is greater than suspected. We have also discovered an interaction between BRG1 and the splicing factor Prp8, linking SWI/SNF activity to mRNA processing. We propose a model whereby parts of the biochemical heterogeneity is a result of function and that the local chromatin environment to which the complex is recruited affect SWI/SNF composition. We have also isolated the novel B‐WICH complex that contains WSTF, SNF2H, the splicing factor SAP155, the RNA helicase II/Guα, the transcription factor Myb‐binding protein 1a, the transcription factor/DNA repair protein CSB and the RNA processing factor DEK. The formation of this complex is dependent on active transcription and links chromatin remodeling by SNF2H to RNA processing. By linking chromatin remodeling complexes with RNA processing proteins our work has begun to build a bridge between chromatin and RNA, suggesting that factors in chromatin associated assemblies translocate onto the growing nascent RNA.


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This thesis is based on the following papers:

Paper I Patrik Asp, Margareta Wihlborg, Mattias Karlén and Ann‐

Kristin Östlund Farrants (2002) BRG1, a human SWI/SNF subunit, affects actin filament organization through the RhoA signaling pathway, Journal of Cell Science 115, 2735‐2746.

Paper II Patrik Asp, Erica Cavellán, Jessica Tångefjord and Ann‐Kristin Östlund Farrants (2004) Variations in biochemical composition and

chromatin association of mammalian SWI/SNF complexes. Manuscript

Paper III Erica Cavellán, Patrik Asp and Ann‐Kristin Östlund Farrants

(2004) WSTF‐SNF2h interacts with several nuclear proteins in a transcription dependent manner, to form a functional unit B‐WICH Manuscript

Introduction 5

Table of contents INTRODUCTION 6 Structure‐function relationships in the eukaryotic nucleus 6 Nucleosomes and chromatin formation 7 Chromatin and DNA accessibility 10 Chromatin remodeling 11 Covalent remodeling 13 ATP‐dependent remodeling 14

Chromatin Remodeling ATPases 15 The SWI2/SNF2 protein family 15 Table I 16 The mammalian SNF2 proteins BRG1 and BRM 17 The SWI/SNF complex 19 Table II 20 SWI/SNF in vitro activity 25 The biology of SWI/SNF complexes 26 The mammalian ISWI proteins SNF2H and SNF2L 32 The ISWI complex 33 ISWI in vitro activity 35 The biology of ISWI complexes 36

PRESENT INVESTIGATION 39 REFERENCES 51 ACKNOWLEDGEMENTS 59 PAPERS 61


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INTRODUCTION

Structure‐function relationships in the eukaryotic nucleus The very large molecules of DNA that make up our chromosomes are located within the envelope of the eukaryotic nucleus and are covered by a plethora of proteins to form a complex nucleo‐protein structure known as chromatin. Chromatin was initially regarded as a static storage structure with the sole purpose of protecting and stabilizing the macromolecules of DNA. However, research in the last 15 years has shown, in increasing detail that the nucleus and its chromatin content are highly dynamic and capable of responding rapidly to cues from the extranuclear environment with processes such as transcription, replication and the import/export of molecules. We know today that chromatin itself is a fundamental component in the regulation of nuclear processes. Advances in microscopy and immunocytochemistry and the ability to express fluorescently tagged proteins in vivo has revealed multiple internal chromatin structures in the nucleus and details of their organization (Dundr and Misteli, 2001; Lamond and Earnshaw, 1998; Misteli, 2001). It is clear that there is a close connection between these structures and the functional state of the chromatin, even though the precise relationship is not yet understood in molecular detail. Some of the structural changes are functional consequences of activities taking place on the chromatin while others precede and regulate chromatin activities. These changes can be global, such as whole chromatin domains shifting position within the nucleus, while others act on only small, local stretches of chromatin. All processes, such as transcription, replication, repair and re‐combination, that take place on chromatin are highly context‐dependent, and it is essential to focus both on the small and well defined chromatin areas involved in each specific event, as well as on the global context wherein they take place in order to fully understand them (Spector, 2003).

Introduction 7

Nucleosomes and chromatin formation Chromatin is a complex structure of DNA and DNA associated proteins. A large part of the protein constituent is made up of the histone proteins that together with DNA form nucleosomes, the smallest organizing unit of chromatin (Figure 1).

Figure 1. Strua) Side viewterminal histmark the DN Two copies eoctameric coleft‐handed h2000; Luger along the Ddetermined stretch of DN

a b

cture of the nucleosome at 2.5 Å resolution (Harp et al., 2000) . The dotted line marks the dyad axis. b) Front view with the N‐one tails extending out through the DNA strand. The arrows A entry and exit points.

ach of the four histone proteins, H2A, H2B, H3 and H4 form an re structure with about 146 bp of DNA wrapped around it in a elix of approximately 1.7 turns (Davey et al., 2002; Harp et al.,

et al., 1997). Nucleosomes are positioned side by side in an array NA strand, and the distance between them is dynamically by the functional state of the chromatin. The nucleosome‐free A separating them is referred to as linker DNA. Nucleosomes are


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assembled mainly during DNA replication and this assembly take place by two parallel processes. One is called ʺparental nucleosome transferʺ, and is the distribution of pre‐existing nucleosomes between the two new molecules of DNA. The other process is known as ʺdenovo nucleosome assemblyʺ, and involves a series of protein factors that target newly synthesized histone proteins to the site of DNA replication. During this process, the H3/H4 tetramer is the first histone component that binds to DNA, partially wrapping the double helix around itself. Subsequent incorporation of the two H2A‐H2B dimers completes the 1.7 turns of DNA around the outside of the core particle. The finished nucleosome is pseudosymmetric and can be divided into two halves along an axis, the dyad (Figure 1a). The assembly process is highly regulated and orchestrated in concert with the cell‐cycle machinery and it involves many co‐factors in addition to histone proteins and DNA (Akey and Luger, 2003; Krude and Keller, 2001; Mello and Almouzni, 2001). There is also a continuous dynamic exchange of histones and a rearrangement of nucleo‐somes that is uncoupled from replication, known as ʺreplication independentʺ (RI) assembly. As an example, Drosophila express the H3 variants H3.3 and Cid and they are deposited throughout the cell cycle and mark transcriptionally active euchromatin and centromeric heterochromatin respectively (Ahmad and Henikoff, 2002). Histone 1 (H1) binds to the outside of the nucleosome core where the DNA enters and exits the structure and is not a constitutive part of the nucleosome. The H1 protein has a more complex pattern of association than the core histones. The entry and exit points of the DNA strand in the nucleosome particle flank the dyad axis and are not necessarily aligned because the DNA does not always wrap all around the core histones in two complete turns (Figure 1b). Complete wrap‐around can occur when H1, the linker histone, binds to the dyad center. H1 has a strong stabilizing effect on higher order chromatin structures, even though it is not essential for the structure to form (Carruthers and Hansen, 2000; Wolffe, 1998; Wolffe and Hayes, 1999), and reports suggest that the positively charged C‐terminus of H1 neutralizes electrostatic forces that may otherwise destabilize higher order chromatin compaction (Vila et al., 2000). H1 does not have a global inhibiting effect on transcription, but it can act in concert with transcription factors as a gene‐ specific transcriptional repressor (Dou and Gorovsky, 2000).

Introduction 9

Nucleosomes are not randomly distributed on DNA but are deposited in a specific pattern that depend both on inherent qualities of the nucleotide sequence and on the functional state of the particular stretch of DNA. During replication, the genetic information in the actual DNA sequence must be faithfully preserved, but equally important is the preservation of the epigenetic information that is present in the chromatin. Information such as the pattern of DNA methylation, nucleosome positions and modifications of histone N‐termini must be passed down to the newly synthesized chromatin to maintain correct gene activity, preserve cell identity and ensure genomic integrity (Lowary and Widom, 1997; McNairn and Gilbert, 2003). The DNA around a nucleosome is aligned so that the minor grooves of the double helix form channels through which the tails of H3 and H2B pass, while the tails of H4 and H2A seem to pass over the DNA (Figure 1b). The N‐termini of H3 and H4 do not have a detectable structure in solution but adopt a helical structure when incorporated into a nucleosome, and this also seems to be the case for H2A and H2B (Baneres et al., 1997; Harp et al., 2000; Luger et al., 1997). The histone N‐termini contains several arginine and lysine residues, giving the tails an overall positive charge. They readily interact, therefore, with the negative phosphate backbone of DNA, which significantly adds to the stability of nucleosomes. The positively charged N‐termini of the core histone proteins also extend beyond the nucleosome particle itself, allowing histone N‐termini to bind histones and DNA in neighboring nucleosomes. This contributes to the stabilization of nucleosome arrays and to the formation of higher order chromatin structures (Carruthers and Hansen, 2000). The large number of histone‐DNA interactions within and between nucleosome particles gives them a high degree of stability. It allows them to be dynamic and to facilitate significant structural changes without becoming unstable and falling apart. This combination of stability and plasticity is crucial for their role as the smallest organizing unit of chromatin.


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Chromatin and DNA accessibility A fundamental functional consequence of nucleosome and chromatin formation is its effect on DNA accessibility. The strong interaction between DNA and histones and the steric block that this interaction creates have profound effects on the ability of DNA‐binding proteins to gain access to and bind to their cognate binding sites. As a consequence, regulation of DNA‐ associated processes, such as transcription, replication, recombination and repair, will to a large extent depend on local chromatin topology. There are several layers to chromatin topology, but from a genome‐wide perspective it is generally divided into two classes: heterochromatin and euchromatin. Nucleosomes in heterochromatin are organized in a closed configuration, contain specific histone‐DNA modifications, and are packaged together with an array of silencing proteins that help to condense the chromatin fiber. This structure is transcriptionally inert. Euchromatin, on the other hand, is mostly devoid of silencing factors and is therefore accessible and transcriptionally active. Importantly, cells can utilize the mechanisms for heterochromatin formation to form highly localized domains of heterochromatin within eu‐chromatin and thereby specifically regulate the activity of individual genes (Henikoff, 2000; Moazed, 2001). Within euchromatin, nucleosomes are commonly positioned to block transcription factor binding sites in promoter regions, and this repressive positioning of nucleosomes often takes place during replication. Interestingly, repressive nucleosome positioning can also be initiated by transcription factors. Nucleosomes on the un‐induced MMTV promoter are randomly positioned, but when GR (the glucocorticoid receptor) binds they become precisely positioned and block several important transcription factor binding sites (Belikov et al., 2000; Hayes and Wolffe, 1992). In contrast, promoters of genes with a continuous steady‐state level of transcription, housekeeping genes, are directly assembled into transcriptionally competent configurations during replication and are thereby largely insensitive to nucleosome repression. The regulatory use of nucleosome positioning is usually limited to genes that are required to alternate between an induced and a repressed state. Nucleosome‐ and chromatin formation have far‐reaching consequences for all biological processes that utilize DNA. On a global scale, nucleosomes facilitate the organized compaction of chromatin into higher order structures and, on a

Introduction 11

local scale, they serve as highly dynamic regulatory units of specific processes. This dual effect on both the large‐scale and on the short‐scale organization of chromatin does not come about through two separate processes because these processes are to a large extent functionally inter‐dependent. The dynamic range and local specificity in DNA packing and chromatin organization depend on changes in protein‐DNA and protein‐protein interactions that are strictly regulated in response to signals that regulate transcription, replication, repair and recombination. Cells have therefore evolved a wide range of protein modifications and energy‐dependent mechanisms to modulate chromatin structure in order to control and regulate the complex network of chromatin‐dependent biological processes. Some of these chromatin‐ modifying mechanisms, such as methylation of cytosines, target DNA itself but most of them directly manipulate the structure of the nucleosome in a process known as chromatin remodeling. Chromatin remodeling The term ʺchromatin remodelingʺ is used to describe a wide variety of changes in chromatin structure and is generally defined as any activity that generates detectable changes in histone‐DNA interactions. Such changes are often mapped as alterations in the nuclease digestion pattern of a particular stretch of DNA assembled into nucleosomes. Certain nucleases are unable to cut DNA assembled into a nucleosome and can therefore be used to monitor chromatin structure. Consequently, chromatin domains can be altered from a nuclease‐ insensitive to a nuclease‐hypersensitive conformation, or vice versa, which in vivo can be a highly localized event that acts on a single nucleosome, or an event that involves several kilobases of chromatin. Nucleosome positioning is commonly described as translational positioning or rotational positioning (Li et al., 1997). Translational positioning (Figure 2a) defines where on a particular stretch of DNA a nucleosome is positioned, and it determines if a protein‐binding site is incorporated into a nucleosome and where it is located within the nucleosome. In most cases, such a location causes a steric block in DNA accessibility but it is possible that a site is still accessible even inside a nucleosome which depends on its rotational position. Rotational positioning (Figure 2b) defines the internal orientation of a binding site relative to the nucleosome surface.


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A binding sequence may face inwards, towards the histone proteins, making it inaccessible or it may face outwards, which grants certain DNA‐binding proteins access to their sites. The mechanistic consequences in terms of chromatin remodeling of translational versus rotational positioning are important because in some cases it would be necessary to move or remove entire nucleosomes, while in other instances a twist of the DNA strand within a single nucleosome could open up a site. All of these scenarios have been observed in vivo where both sliding of nucleosomes, nucleosome eviction and nucleosome remodeling occur and, consequently, different groups of re‐modeling machines have mechanistically distinct modes of action.

Figure 2 Translational and Rotational positioning. A. Translational positioning: localization of a DNA‐binding site between nucleosomes makes it accessible while incorporation into a nucleosome makes it inaccessible. B. Rotational positioning: A DNA‐binding site facing the histone surface is inaccessible. Twisting the DNA makes the DNA‐binding site face away from the histone surface, making it accessible.

Introduction 13

Covalent remodeling Chromatin remodeling is usually separated into two main categories: covalent remodeling and ATP‐dependent remodeling. Covalent chromatin remodeling is the addition or removal of various molecular moieties on histone proteins, mostly involving the N‐terminal tails. Acetylation (Carrozza et al., 2003; Thiagalingam et al., 2003), methylation (Sims et al., 2003), phosphorylation (Iizuka and Smith, 2003), and ubiquitination (Bach and Ostendorff, 2003) are the most well‐studied covalent histone modifications, while histone ADP‐ ribosylation is still relatively unexplored in the context of chromatin remodeling, but has a role in DNA repair (Virag and Szabo, 2002). There is a complex network of functional interactions among these modifications and the effect of modifying histone N‐termini can be very local, acting on nucleosomes in individual promoters, as well as affecting the organization of whole chromosomes (Gregory et al., 2001; Zhang and Reinberg, 2001). As mentioned earlier, the histone N‐terminal tails are highly positive due to the presence of several arginine and lysine residues and these charges mediate interactions with a large array of proteins and with the negative phosphate backbone of DNA. Modifying the lysine residues by adding or removing acetyl groups alters the positive charge and the specific charge distribution of the tails which leads to changes in the pattern of association with DNA and proteins and to modifications of the strengths of these interactions. In a similar manner, the addition or removal of phosphate groups on serine residues leads to changes in charge distribution. The added molecules can also serve as markers and/or docking sites for proteins that in turn affect chromatin structure. For instance, methylation of lysine 9 on H3 (H3metK9) can recruit HP1, leading to heterochromatin formation, while methylation of certain residues on H4 is instead a marker for open and transcriptionally active euchromatin (Sims et al., 2003). The rapidly expanding field of covalent remodeling has involved the isolation and identification of a multitude of proteins responsible for such activities and work in this field has begun to unravel their complex patterns of biological function. A major advance in this field was the formulation of the histone code hypothesis, which explains how these modifications regulate one another and result in activation or repression of transcription. In general, acetylation of histones increases DNA accessibility, while deacetylation leads to a decrease (Jenuwein and Allis, 2001; Strahl and Allis, 2000). The number and variety of


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covalent histone modifications are steadily increasing and translating the histone code will be challenging. Especially since the influence of histone modifications on chromatin structure and corresponding biological response is intimately connected with ATP dependent chromatin remodeling.

ATP‐dependent remodeling ATP‐dependent chromatin remodeling is a process in which the energy of ATP hydrolysis is used to open up or close protein binding sites in chromatin by altering the rotational and/or translational positions of nucleosomes. ATP‐dependent remodeling is performed by the SWI2/SNF2 family of nucleic acid stimulated ATPases (Havas et al., 2001; Lusser and Kadonaga, 2003; Tsukiyama and Wu, 1997; Varga‐Weisz, 2001) and several models have been proposed for this type of remodeling. The exact mechanism, however, has not yet been fully resolved (Langst and Becker, 2004; Lusser and Kadonaga, 2003). The Twist‐Diffusion model suggests that SWI2/SNF2 ATPases uses the energy of ATP hydrolysis to twist DNA and to generate negative superhelical torsion that diffuses through the nucleosome. This results in structural changes in DNA‐histone interactions and, subsequently, modifies DNA accessibility. The Loop‐Recapture model proposes that DNA is detached from the nucleosome in a 30‐35 bp loop that propagates through the nucleosome as a “wave” of altered accessibility. Yet another model, the Cross‐Transfer model, suggests that the DNA helix undergoes cross‐transfer across the surface of the nucleo‐some, and that parts of the helix are actually rolling off the histone octamer, thereby escaping the repressive nucleosome environment altogether. There are distinct differences in the way the various subgroups within the SWI2/SNF2 family remodel chromatin substrates in vitro, so it is probable that several mechanisms are used. However, the results presented so far are still unclear and depend strongly on the type off assay used. It is important to remember that these investigations have for the most part used single nucleosomes and poly‐nucleosomal arrays assembled on defined short stretches of DNA and these substrates may not correctly reflect the situation in vivo with its much more complex and variable chromatin environment (Havas et al., 2000; Langst and Becker, 2001; Narlikar et al., 2001). Whatever the specific details in remodeling mechanisms will turn out to be, all of them change the trans‐lational and/or rotational positioning of nucleosomes and cause changes in DNA accessibility.

Introduction 15

Chromatin Remodeling ATPases All known ATP‐dependent chromatin remodeling is performed by protein complexes that contain a central ATPase from the SWI2/SNF2 family. The founding member of this family is the yeast SWI2/SNF2 protein, which was identified as a chromatin remodeling protein in the early 1990s. The members of this family are evolutionary conserved from yeast to mammals, and several of them are essential for viability. The number of SWI2/SNF2 proteins is still growing, adding to the increasing complexity of the function and regulation of this important family of proteins.

The SWI2/SNF2 protein family Genetic screens in yeast in the 1980s identified a series of mutations in genes affecting mating type switching and sucrose metabolism. One of these genes was designated swi2/snf2 for switch deficient 2/sucrose non‐fermenting 2 and it became the founding member of the ATP‐dependent chromatin remodeling family SWI2/SNF2 (Table I) (Abrams et al., 1986; Egel et al., 1984; Neigeborn and Carlson, 1984). The discovery that the SWI2/SNF2 protein was involved in the modification of chromatin structure in association with transcription (Hirschhorn et al., 1992; Peterson and Herskowitz, 1992; Peterson et al., 1991) marked the beginning of a rapid expansion in the field of chromatin remodeling research. The number of SWI2/SNF2 family members has grown considerably in the last ten years, and proteins belonging to this family have been found in species from yeast to mammals (Table 1). The common denominator is the bipartite ATPase domain with an N‐terminal half called the SNF2‐N domain that is unique to the SNF2 family, and a C‐terminal helicase C domain with a high degree of similarity to the helicase C family. However, helicase activity is thought to possibly be possessed by only one of the member proteins, INO80. All SWI2/SNF2 proteins have DNA stimulated and/or nucleosome stimulated ATPase activity and remodel chromatin in vitro. The defining ATPase domain is highly conserved across species, but outside this domain the SWI2/SNF2 proteins have little sequence identity. Another characteristic feature of this protein family is the combination of the ATPase domain with additional structures such as BROMO, SANT or CHROMO domains. This feature has allowed the members of the family to be classified into 7 groups: 1 SNF2, 2 ISWI, 3 CHD, 4 INO80, 5 CSB, 6 RAD54 and 7 DDM1 (Table 1) (Lusser and Kadonaga, 2003).

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Table I

The SWI2/SNF2 Family

Characteristic Group Species‐Protein Domain Function

1. SNF2

S.cerevisiae: Drosophila: Mammalia:

Snf2, STH1 Brahma BRG1, BRM

BROMO

Transcription DNA Repair

2. ISWI


Iswi1p, Iswi2p ISWI SNF2H SNF2L

SANT

Transcription Replication Global chromosome structure

3. CHD1


CHD1 Hrp1,3 Mi2‐α/CHD3 Mi2‐β/CHD4

CHROMO PHD

Transcription

4. INO80

S.cerevisiae:

Ino80

Divided SNF2 domain

Transcription

5. CSB

S.cerevisiae Mammalia

Rad26 CSB/ERCC6

Acidic N‐

terminal domain critical for function.

Patterning of genome‐ wide DNA methylation

DNA Repair

6. RAD54

S.cerevisiae Mammalia

Rad54 ATRX, ARIP4

Recombination Transcription

7. DDM1

A.thaliana

DDM1/Lah1

DNA methylation

Introduction 17

The work presented in this thesis is focused on the mammalian SWI2/SNF2 homologs BRG1 and SNF2H that belong to the first and second group respectively.

The mammalian SNF2 proteins BRG1 and BRM The SNF2 group contains the archetypical SWI2/SNF2 proteins. This group has two members in yeast: the founding member ySWI2/SNF2 and ySTH1 (Snf‐two‐homolog 1) (Abrams et al., 1986; Laurent et al., 1992). Swi2/snf2 deletion strains are viable but grow more slowly than the wild type, while Sth1 is essential. Two mammalian homologs of ySWI2/SNF2 have been identified: BRM and BRG1, while no mammalian protein has been designated as being a ySTH1 equivalent. BRM was named after the Drosophila SNF2 protein Brahma (Tamkun et al., 1992), while BRG1 stands for Brahma related gene 1 (Chiba et al., 1994; Khavari et al., 1993; Muchardt and Yaniv, 1993). BRG1 and BRM have a high degree of similarity in peptide sequence and in vitro enzymatic activity but have both overlapping and unique functions in vivo. This difference is strikingly reflected in the fact that Brg1 is essential for viability while Brm is not (Bultman et al., 2000; Reyes et al., 1998).

Figure 3. Domain analysis of BRG1 and BRM: I. Charged, P/Q rich domain, II. Charged domain, III. Bipartite ATPase domain, IV. E7 box (LXCXE motif), V. AT‐hook, VI. BROMO domain

Domain analysis of the BRM and BRG1 peptides (Figure 3) shows that the N‐terminal part contains two highly charged regions, where region 1 has a high proline content (~25%). The central region is the bipartite ATPase domain unique for the SNF2 family. The actual nucleotide binding site is highly conserved with a lysine residue critical for function that can be substituted for an arginine (K783R in hBRG1) to create an ATPase‐deficient form that is

I II III IV V VI 1648 aaBRG1

1591 aa BRM


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biochemically indistinguishable from the wild type protein (Chiba et al., 1994; Khavari et al., 1993; Muchardt and Yaniv, 1993; Wong et al., 2000). The C‐terminal region contains a BROMO domain (Figure 3) which is found in many chromatin‐associated proteins (Tamkun et al., 1992). These domains bind to histone N‐termini and recent results show that the affinity is higher for acetylated histone N‐termini (Dhalluin et al., 1999; Jacobson et al., 2000). This affinity suggests that the BROMO domains in BRG1 and BRM facilitate substrate recognition and binding, leading to stabilization of these proteins on chromatin in response to the acetylation status of histones (Marmorstein and Berger, 2001). BRM and BRG1 also contain a region that is similar to the AT‐hook motif found in HMG I/Y proteins (Figure 3). This region is a DNA binding domain essential for BRM function in vivo, and in vitro experiments show that BRM can bind to DNA harboring A⋅T bp repeats in a gel shift assay. Interestingly, the BROMO domain is not absolutely essential for BRM mediated growth repression (Bourachot et al., 1999). BRG1 and BRM function in large protein complexes, and some of the associated proteins contain an HMG domain or an ARID (AT‐rich interaction domain) (Wilsker et al., 2004) which, together with the presence of the AT‐hook motif, suggests that these ATP‐dependent remodeling complexes have intrinsic DNA‐binding activity. However, it is unlikely that this activity is strong enough for autonomous and undirected chromatin binding that results in random chromatin remodeling, because such binding could result in improper gene regulation and have negative repercussions for the cell. An alternative scenario is that the BROMO, AT‐hook, ARID and HMG domains present in BRG1, BRM, and their associated proteins, give SWI/SNF complexes an ʺaffinityʺ towards DNA. Once recruited to a specific chromatin site this affinity would help stabilize substrate binding and facilitate remodeling which would be in agreement with data showing that BRG1 and BRM are recruited to chromatin by sequence‐ specific transcription factors (Sudarsanam and Winston, 2000). BRG1 and BRM also contain an Rb binding LXCXE motif (Figure 3) indicative of a role in cell cycle control via the Rb/E2F network, and both proteins have growth‐ suppressing properties in transformed cells, that identify them as tumor suppressors. Interestingly, despite their high degree of similarity, the growth‐ suppressive property is clearly stronger for BRG1 than for BRM (Strober et al., 1996).

Introduction 19

The SWI/SNF complex All proteins within the SNF2 group so far investigated function as ATPases in large protein complexes. The original yeast SWI2/SNF2 protein gave its name to the yeast SWI/SNF complex, which contains approximately 11 different polypeptides (Cairns et al., 1994; Cote et al., 1994; Peterson et al., 1994). The essential ySTH1 protein functions within a complex called RSC (remodels the structures of chromatin), which contains 15 polypeptides (Cairns et al., 1996). The compositions of the yeast SWI/SNF and RSC complexes are fairly constant and have been well characterized. The situation, however, is much more complex when it comes to the mammalian SNF2 proteins BRM and BRG1. These ATPases also function in large protein assemblies but the composition of these mammalian SWI/SNF complexes can vary from between 8‐20 subunits. Variants of mammalian SWI/SNF complexes have been purified from various sources and the complexity in subunit composition and in vivo function is staggering and not yet fully resolved. Below follows a discussion of SWI/SNF complexes isolated from mammalian sources, where the reference for each complex can be found in Table II together with the designated name and a description of the subunit composition. Two mammalian SWI/SNF complexes were initially isolated by conventional chromatography: SWI/SNF‐A and SWI/SNF‐B. Both of these complexes had in vitro remodeling activity and stimulated activator binding to DNA (Kwon et al., 1994). In a large scale purification of mammalian SWI/SNF complexes from various human, mouse and rat cell lines, combining chromatography with immunoaffinity purification, it was shown that BRG1 and BRM are mutually exclusive subunits, that SWI/SNF complexes have overall similar polypeptide composition but that they also display a significant degree of variation. Proteins associated with the complexes were called BAFs (for ʺBRG1/BRM associated factorsʺ), and a number designating the apparent MW of the polypeptide. The cloning and identification of BAF proteins were also initiated during these purifications (Wang et al., 1998; Wang et al., 1996a; Wang et al., 1996b; Zhao et al., 1998). SWI/SNF complexes isolated from erythroid cells revealed an association with the Ikaros family of transcription factors and a role in β‐globin expression (Armstrong et al., 1998; Kim et al., 1999; OʹNeill et al., 1999; OʹNeill et al., 2000). The characterization of these purified complexes revealed a link between the SNF2 and Mi2 groups by identifying components of both the SWI/SNF and the NuRD complexes (not shown in the table) within

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Table II

21


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these novel SWI/SNF variants (OʹNeill et al., 1999; OʹNeill et al., 2000; Tong et al., 1998; Xue et al., 1998). This discovery coupled SWI/SNF‐mediated chromatin remodeling not only to the activation but also to the repression of transcription, since the NuRD complex contains HDACs, which have a repressive effect on transcription. A role for SWI/SNF in repression of transcription was further supported by the isolation of the N‐Cor 1 and N‐Cor 2 complexes, which also combine ATP‐dependent remodeling with the presence of HDAC proteins. In addition, N‐Cor2 contains mSin3A, a mammalian homolog of the yeast co‐repressor Sin3. Interestingly, components of the splicing machinery (SAP130 and SF3a120) were also found in these complexes (Underhill et al., 2000). The BAF180 protein has been suggested to define a specific form of mammalian SWI/SNF complexes called PBAF. By combining chromatography with immunoaffinity, two types of SWI/SNF complexes were isolated that differed with respect to the presence of BAF250 and BAF180. Cloning and analysis of BAF180 revealed that it is highly similar to the chicken Polybromo protein (Goodwin and Nicolas, 2001; Nicolas and Goodwin, 1996) and that it contains multiple BROMO domains, an unusual and characteristic feature. To distinguish the BAF180‐SWI/SNF from BAF250‐SWI/SNF they were designated PBAF (Polybromo‐BAF) and BAF respectively. Database searches revealed that three yRSC subunits: Rsc1, 2 and 4, have a combined domain structure highly similar to BAF180, suggesting that PBAF is a mammalian functional equivalent to the yeast RSC complex. Yeast RSC plays a role in cell cycle progression, and the subsequent immunolocalization of BAF180 to kinetochores and spindle poles during prometaphase is in agreement with the hypothesis that PBAF is a mammalian RSC equivalent (Angus‐Hill et al., 2001; Xue et al., 2000). The division of SWI/SNF complexes into BAF and PBAF has been confirmed by others using conventional chromatography purification strategies, which further showed a specificity for PBAF in mediating hormone receptor‐activated in vitro transcription (Lemon et al., 2001). However, the division into BAF and PBAF is far from clear, since a similar purification strategy as applied by Xue etal. (2000), using FLAG‐tagged BRG1, BRM and SNF5 proteins, did not yield any BAF180‐containing SWI/SNF complexes. Instead it revealed an association with the HDAC/mSin3A repressor complex, in agreement with previous findings (Sif et al., 2001; Sif et al., 1998). Interestingly, the only difference in the purification strategies used in these

Introduction 23

reports is that for the isolation of the PBAF complex, the nuclear extract was first fractionated over a P11 cation exchange column before αFLAG immunopurification. A neuronal SWI/SNF complex designated ʺbBAFʺ was isolated using a neuro‐ specific BAF53 homolog, BAF53b, as a marker. This complex contains two subunits, p180 and p160, that either are special variants of BAF180 and BAF155 or novel proteins (Olave et al., 2002). By an identical approach as that described earlier (Sif et al., 2001), the previously uncharacterized p66 poly‐peptide was identified as PRMT5, a type II arginine methyl transferase. This SWI/SNF complex methylates H3 and H4 in vitro and is recruited to the cad promoter by Myc/Mad heterodimers resulting in transcriptional repression (Pal et al., 2003). A strong connection between SWI/SNF and leukemia was established when a complex called EBAF was isolated. EBAF is a SWI/SNF variant that contains the ENL protein, and the enl gene is commonly fused to the mll gene in leukemia. MLL‐ENL fusion proteins could subsequently recruit SWI/SNF to genes normally regulated by MLL alone, resulting in aberrant gene expression and the induction of leukemia. Reporter assays using the HoxA7 promoter, an endogenous target gene for MLL, supported this hypothesis by showing a synergistic effect between BRG1 and an MLL‐ENL fusion protein in the transcriptional activation of the reporter (Nie et al., 2003). The SWI/SNF variant WINAC combines the WSTF (Williams‐syndrome‐transcription‐factor) protein with SWI/SNF components, with components of the replication and nucleosome assembly machinery and with the VDR (vitamin D receptor). Interestingly, the WSTF protein had previously been found only in complex with ISWI proteins, the second group of the SNF2 family. The WINAC has in vitro nucleosome assembly activity, in vivo promoter targeting and, interestingly, vitamin D3‐independent complex assembly and promoter binding, while it has, on the other hand, ligand‐dependent transactivation (Kitagawa et al., 2003). The latest addition to this multifaceted family is the NUMAC which contains CARM1, an arginine specific histone methyl transferase protein (H3metR17). Its association with SWI/SNF dramatically increases its specificity towards nucleosomal histones compared to its activity towards free histones. The NUMAC is involved in hormone receptor‐activated transcription, and activates ER responsive genes in vivo (Xu et al., 2004).


24

The diversity discovered by the purifications of SWI/SNF complexes has important implications for their biological function. The power inherent in ATP‐dependent remodeling seems to have been harnessed and refined by evolution to meet specific demands that arose when the genome expanded, since the general trend is that the larger the genome of an organism, the more complex is the biochemical make‐up of SWI/SNF. This model fits well with the isolation of cell type specific and tissue specific forms capable of regulating genes necessary for phenotypic identity, since an increase in the number of genes would require a more sophisticated network of regulation to ensure a properly executed and maintained differentiation and homeostasis. As a result, the actual in vivo function of these complexes is likely to be highly gene‐ specific and linked to local chromatin topology. It is, therefore, necessary to focus on each individual gene and determine its specific relationship to SWI/SNF, in order to dissect and fully understand the biology of SWI/SNF complexes Table II shows that many of the isolated SWI/SNF variants contain proteins that are involved in repression of transcription, such as HDACs. This is difficult to reconcile with the fact that identification of SWI/SNF regulated genes have for the most part found genes to be up regulated in response to SWI/SNF and not repressed (Hendricks et al., 2004; Liu et al., 2001; Sudarsanam et al., 2000). An unknown factor with the many forms of SWI/SNF that have been isolated is that there is very limited information about the relative amounts of these complexes in cells. It is possible, therefore, that the population containing repressive components is a minor one, explaining why activation of transcription is predominant. Another possibility is that our understanding of SWI/SNF activity on promoters is incomplete and that there are mechanisms we have not yet discovered. From a technical point of view, another important issue is that the biochemical conditions used to purify SWI/SNF may have a profound effect on the forms of SWI/SNF complexes that the procedure will yield. Some examples of this are the presence of actin in SWI/SNF and the division into BAF and PBAF, both of which vary depending on biochemical conditions during purification (See Present Investigation for an analysis and discussion of the SWI/SNF‐actin connection and BAF/PBAF division).

Introduction 25

SWI/SNF in vitro activity It is well‐established that BRG1‐containing SWI/SNF complexes remodel mononucleosomes in an ATP‐dependent manner in vitro, and that they also change the nuclease digestion pattern of an array of nucleosomes. Interestingly, BRM‐containing SWI/SNF may only be active on a nucleosome array and not on mononucleosomes (Sif et al., 2001). The process is partially independent of histone N‐termini, although it is less efficient on trypsin treated nucleosomes (Guyon et al., 1999). Remodeling usually takes place without evicting histone octamers from the DNA strand unless a high molar ratio (1:3) of SWI/SNF to nucleosomes is used (Phelan et al., 2000). It has been estimated that there are between 100‐200 SWI/SNF complexes in yeast nuclei (Cairns et al., 1996) and if this is the case also in mammals, the in vivo molar ratio of SWI/SNF to nucleosomes is very much lower than in these in vitro assays. As a consequence, trans‐displacement of histones may not be a physiological mode of action in a natural chromatin environment. On the other hand, recruitment mechanisms of SWI/SNF to promoters may increase the local concentration enough to promote trans‐displacement. SWI/SNF complexes also change the topology of DNA assembled into nucleosomes by reducing the amount of negative supercoiling (Havas et al., 2000) and SWI/SNF complexes can shift the translational position of nucleosomes in a process known as sliding (Jaskelioff et al., 2000; Whitehouse et al., 1999). Recombinant BRG1 and BRM can remodel nucleosomes on their own, albeit less efficiently than a complete SWI/SNF complex. A unit as active as a whole SWI/SNF complex can be formed by combining BRG1 with only three additional subunits: INI1, BAF155 and BAF170 (Phelan et al., 1999). This suggests that the complex is organized around a central remodelling core and that BAF proteins function as regulators or modulators of activity and/or to mediate interactions with components of the transcription machinery. Such a model agrees well with the biochemical results that always show INI1, BAF155 and BAF170 as a part of the complex, while many other components vary (Table II). This type of modular structure around a central functional core allows for great flexibility, in concordance with the compositional diversity seen among SWI/SNF complexes. However, it has not been tested if other combinations of BAFs together with BRG1 or BRM result in equally high remodeling activity in vitro.


26

The biology of SWI/SNF complexes Development It is a challenge to investigate the biological aspects of SWI/SNF function given the high complexity in subunit composition. Studies show that both BRG1 and hBRM are expressed ubiquitously in all adult tissues (Khavari et al., 1993; Muchardt and Yaniv, 1993) but have distinctly different spatial and temporal patterns of expression during development. F9 murine embryonal carcinoma cells have an absolute requirement for BRG1 (Sumi‐Ichinose et al., 1997) and mouse zygotes with a homozygous deletion of BRG1 cannot grow into viable embryos (Bultman et al., 2000). Developing blastulas die before implantation in the uterus, and neither the inner cell mass nor the trophectoderm are viable in ex vivo cultures. Heterozygous BRG1+/‐ mice are viable, but the number of offspring is significantly lower than it is for wild type animals. These mice also display an increased predisposition for exencephaly and tumors. Baf47/INI1‐/‐ mice are non‐viable and have an identical phenotype as BRG1‐/‐ embryos. Heterozygous individuals survive, but develop tumors in the nervous system and soft tissue sarcomas (Klochendler‐Yeivin et al., 2000). In stark contrast, BRM is not essential for survival, and homozygous knockout mice are viable and fertile. The only identifiable phenotype is that overall body size is larger than normal, as are certain internal organs. The protein level of BRG1 is elevated in these mice, suggesting that BRG1 can functionally compensate for the loss of BRM (Reyes et al., 1998). Differentiation The expression patterns of BRG1 and BRM during embryo differentiation have spatial and temporal tissue‐specific distribution in mice, in which BRM is specifically expressed as soon as the blastula starts to differentiate (Dauvillier et al., 2001; LeGouy et al., 1998; Randazzo et al., 1994). Similar patterns are seen in developing chicken embryos (Schofield et al., 1999). SWI/SNF activity has also been coupled to the differentiation of murine neural precursor cells, where BRG1 is present at a constant level throughout this process while the level of BRM increases dramatically. Identical results have been obtained in murine P19 cells during their differentiation to muscle, neural, and endodermal and mesodermal cell types (Machida et al., 2001). Other reports have shown that differentiation of NIH3T3 fibroblasts into muscle cells depends on both BRG1 and BRM in cooperation with the transcription factor MyoD. Expression of dominant negative ATPase‐deficient forms of BRG1 and

Introduction 27

BRM severely inhibits this process and specifically represses remodeling of promoters of MyoD‐activated genes in vivo (de la Serna et al., 2001a; de la Serna et al., 2001b). These results clearly show that chromatin remodeling by SWI/SNF complexes is important in development and differentiation and they show that BRG1 and BRM are likely to have both shared and unique biological functions. SWI/SNF and cell cycle regulation The SWI/SNF complex is intimately involved in cell cycle regulation. The BRG1 and BRM proteins are both phosphorylated and excluded from condensed chromosomes during the M‐phase, but the outcome of the phosphorylation is different. The level of BRG1 remains constant throughout the cell cycle, but the level of the BRM protein drops considerably during the M‐phase when it is degraded in response to phosphorylation. BRG1 is reactivated by de‐phosphorylation in late M/early G1 and, at the same time, de novo synthesis of BRM rapidly brings the protein back up to normal levels (Muchardt et al., 1996; Stukenberg et al., 1997). The SWI/SNF subunit BAF155 is also phosphorylated in a cell‐cycle dependent pattern similar to BRG1 and BRM, and SWI/SNF complexes isolated from M‐phase cells are inactive in remodeling assays (Sif et al., 1998). Data from yeast show that genes that must be activated in the boundary between M and G1 in the cell cycle, where chromatin is still very condensed, depend on SWI/SNF for transcriptional activation (Krebs et al., 2000). Clearly, highly condensed chromatin is still somewhat susceptible to SWI/SNF remodeling and it is, therefore, possible that remodeling during the M‐phase interferes with proper chromosome structure and separation, explaining why the complex must be inactivated. The SWI/SNF complex interacts with a number of regulatory components in the cell cycle machinery. Over‐expression of BRG1 or BRM in human SW13 cells, which are deficient in these proteins, causes cell cycle arrest and cell senescence. This arrest depends on an interaction between BRG1 and the cell cycle repressor protein pRb , which is essential for BRG1‐mediated cell cycle arrest in SW13 cells (Dunaief et al., 1994; Shanahan et al., 1999). Co‐expression of BRG1 and a constitutively active pRb allele in C33A cells that are normally deficient in these proteins induces cell cycle arrest (Strobeck et al., 2000a; Strobeck et al., 2000b). SWI/SNF has been found in a functional unit together with pRb and HDACs in a complex that represses the cyclin E and A genes.


28

Phosphorylation of pRb disrupts the interaction with HDACs, but the pRb‐ SWI/SNF complex persists and instead maintains cyclin A expression (Zhang et al., 2000). BRG1 and BAF155 also directly interact with cyclin E (Shanahan et al., 1999). The involvement of SWI/SNF in cyclin A expression and the interactions with pRb and cyclin E define a role for SWI/SNF in the exit from the G1 and S phases of the cell cycle. The SWI/SNF subunit Ini1/BAF47 binds to the early response protein c‐myc in vivo and facilitates c‐myc transactivation of target genes (Cheng et al., 1999). In SW13 cells, SWI/SNF represses endogenous c‐fos expression in an Rb‐dependent manner that is independent of E2F (Murphy et al., 1999), while transactivation by Fos/Jun dimers is partially regulated by the SWI/SNF subunit BAF 60a (Ito et al., 2001). The data above clearly show that SWI/SNF activity is important in cell cycle regulation, and in agreement with these data the SWI/SNF complex has been identified as a tumor suppressor. Many human cancer cell lines show a down‐regulation of expression or lack expression altogether of several SWI/SNF components and a number of mutations in genes coding for SWI/SNF components have been identified (Decristofaro et al., 2001; DeCristofaro et al., 1999; Reisman et al., 2003; Reisman et al., 2002; Wong et al., 2000). The SWI/SNF subunit Ini1 is strongly connected to cancer development and is mutated or undetectable in several forms of cancer, in particular in pediatric rhabdoid tumors (Biegel et al., 1999; Versteege et al., 1998). Strikingly, inactivation of INI1 expression in mice leads to CD8+ T‐cell lymphoma with 100% penetrance (Roberts et al., 2002). Expression of a functional Ini1 protein in Ini1‐deficient tumor cells restores cell cycle control and leads to cell cycle arrest and apoptosis (Ae et al., 2002; Betz et al., 2002). Surprisingly, despite having such severe biological effects, loss of Ini1 does not impair the expression of several known SWI/SNF‐ dependent genes, nor is the protein required for the structural integrity of the SWI/SNF complex (Doan et al., 2004). Similarly, the Drosophila INI1 homolog SNR1 is not required for all functional aspects of the Brahma complex (Zraly et al., 2003). BRG1 also binds to BRCA1, a tumor suppressor that is mutated in familial breast cancer, and BRCA1‐stimulation of p53‐mediated transcription depends on BRG1 (Bochar et al., 2000b). These data clearly show that SWI/SNF complexes have a fundamental role in tumor suppression.

Introduction 29

Transcription SWI/SNF complexes in yeast and mammalian cells are involved in the regulation of transcription and are recruited to promoters by sequence‐specific transcription factors (Kadam and Emerson, 2003; Prochasson et al., 2003). The chromatin remodeling activity then facilitates binding of both specific and general transcription factors, and it facilitates the binding of factors involved in repression, such as HDACs. It is important to recognize that chromatin remodeling per se does not determine whether transcription will be activated or repressed, although SWI/SNF activity has so far mostly been associated with activation. A well‐studied aspect of SWI/SNF‐mediated transcriptional activation is the interaction between SWI/SNF and nuclear hormone receptors (Jeon et al., 1997). The GR (glucocorticoid receptor) recruits SWI/SNF to the MMTV promoter, resulting in increased DNA accessibility that is essential for transcriptional activation (Fryer and Archer, 1998; Ostlund Farrants et al., 1997). Chromatin remodeling of the MMTV promoter cannot be substituted with other ATP‐dependent chromatin remodeling complexes besides SWI/SNF and, in addition, the MMTV promoter specifically requires the BAF variant of SWI/SNF complexes for transcriptional activation, and not PBAF (Trotter and Archer, 2004). Selectivity in the recruitment of specific forms of SWI/SNF complexes is a general pattern for nuclear hormone receptors. The VDR (vitamin D receptor) selectively recruits the PBAF form of SWI/SNF (Lemon et al., 2001) and the AR (androgen receptor) has a preference for BRM‐containing SWI/SNF complexes with important implications in prostate cancer development (Marshall et al., 2003). The ER (estrogen receptor) recruits the SWI/SNF complex to estrogen‐responsive genes, where it activates transcription in co‐operation with HATs (Belandia et al., 2002; DiRenzo et al., 2000; Ichinose et al., 1997), and this ER‐SWI/SNF interaction is essential for breast cancer treatment with estrogen antagonists (Wang et al., 2004). In the SWI/SNF‐hormone receptor interaction, another layer of complexity is that it has been mapped to specific subunits besides BRG1 and BRM, which greatly increases the number of potential combinations in the interaction between SWI/SNF complexes and hormone receptors (Belandia et al., 2002; Debril et al., 2004; Hsiao et al., 2003; Inoue et al., 2002; Koszewski et al., 2003; Lemon et al., 2001).


30

Detailed studies in mammalian cells have begun to unravel the complex dynamics between SWI/SNF recruitment to promoters, PIC (pre‐initiation complex) formation, promoter firing and other chromatin remodeling activities and have revealed variations in the order of events. Histone acetylation and partial PIC formation precedes SWI/SNF recruitment on the mammalian IFN‐β (interferon‐beta) promoter (Agalioti et al., 2000). The mammalian PPARγ2 promoter is regulated in a different order in which TBP binding and partial PIC assembly happen before SWI/SNF recruitment. Full PIC assembly takes place concomitant with SWI/SNF recruitment, followed by H3 acetylation and subsequent promoter firing (Salma et al., 2004). The human α1‐AT gene is induced by differentiation signals, but TBP, TFIIB and the activator protein HNF‐1 are constitutively bound to the promoter even in the absence of such signals. Differentiation triggers rapid recruitment of RNApolII and GTFs (general transcription factors), leading to full PIC formation but not to activation of transcription. Promoter firing requires the subsequent recruitment of SWI/SNF and two HATs (CBP and P/CAF), resulting in remodeling and acetylation of proximal nucleosomes (Soutoglou and Talianidis, 2002). These examples of variations in SWI/SNF involvement in transcriptional regulation reveal significant differences in promoter‐specific SWI/SNF requirement and function. However, the situation may be even more complex, as almost all work on promoter recruitment of SWI/SNF has been done with ChIP (chromatin immunoprecipitation) directed towards BRG1 or BRM, and very little work has been done in elucidating the pattern of promoter recruitment of specific BAFs. It is of significant interest to expand promoter analyses to also encompass the BAFs, in order to fully understand the biology of mammalian SWI/SNF in transcription in relation to the biochemical hetero‐geneity and the apparent promoter‐specific variations in SWI/SNF activity, SWI/SNF‐regulated genes Since most aspects of the biology of SWI/SNF complexes are coupled to regulation of transcription, several large‐scale attempts have been made to identify SWI/SNF‐regulated genes. Expression profiling in yeast comparing wild type and swi2/snf2 strains revealed ~300 genes to be up‐ or down‐ regulated but no discernible pattern could be seen among them (Holstege et al., 1998; Sudarsanam et al., 2000). Similar approaches have been used in mammalian cells, using the BRG1‐deficient and BRM‐deficient cell line SW13 (Leibovitz et al., 1973; Muchardt and Yaniv, 1993).

Introduction 31

Twenty‐two thousand genes were screened using mRNA from FACS‐sorted SW13 cells transiently transfected with BRG1 together with GFP as a marker, and this screen yielded 80 up‐regulated and two down‐regulated genes (Liu et al., 2001). Interestingly, there was a significant degree of variation among these genes, where some were absolutely dependent on SWI/SNF for expression while others seemed only to require SWI/SNF for maximal induction. The identification of SWI/SNF‐dependent genes is further complicated by the fact that there is a cell type‐specific interaction between SWI/SNF and promoters. A microarray screen of 40,000 human cDNA sequences using transient expression of BRG1 in the BRG1‐deficient human breast cancer cell line ALAB identified 70 up‐regulated and 65 down‐regulated genes. Several novel targets were confirmed to be SWI/SNF‐regulated but, interestingly, some genes displayed a cell type‐specific dependence on SWI/SNF for activation. BRG1 expression in SW13 cells strongly induced cd44, osteonectin and csf1, while BRG1 expression in ALAB cells only induced osteonectin. Brm was also identified and verified as a BRG1 target promoter, a connection not seen in several other cell types (Hendricks et al., 2004). It is highly interesting that remodeling by SWI/SNF is not only promoter‐ specific, but also varies depending on cell type. One explanation is that additional transcription factors besides SWI/SNF are simply not expressed in ALAB cells, preventing SWI/SNF‐mediated promoter activation/stimulation. Another possibility is that cells in culture are epigenetically unstable, so that the regulatory chromatin topology over promoters changes over time so that it no longer reflects the correct physiological situation. Such a scenario raises questions about the physiological relevance of cell culture experiments. Alternatively, epigenetic patterns established during development could result in the same promoter having tissue‐specific chromatin topology and, consequently, to require different promoter activities for transcriptional activation or repression. Depending on tissue origin, the same gene could subsequently show variations in SWI/SNF dependency for its expression in different cell types.


32

The mammalian ISWI proteins SNF2H and SNF2L The second group in the SNF2 family contains the ISWI (imitation switch) proteins. The founding member of this group is the Drosophila ISWI protein, which was identified based on its sequence homology to the yeast Swi2/Snf2 protein. These proteins appeared, in early experiments, to have the same in vitro remodeling activity as the SNF2 proteins, imitating them (Elfring et al., 1994). Yeast contains two ISWI homologs, Iswi1 and Iswi2 (Table I), which are not essential for viability although null mutant strains are sensitive to a wide range of stress factors (Tsukiyama et al., 1999). Mammals contain two highly similar homologs of ISWI, designated SNF2L (Snf2‐like) and SNF2H (Snf2 homolog). The SNF2H protein is essential for viability. (Table I & Figure 4) (Aihara et al., 1998; Okabe et al., 1992; Stopka and Skoultchi, 2003).

Figure 4. Domain analysis of SNF2H and SNF2L I. ATPase domain, II SANT domain, III SLIDE domain.

SNF2H and SNF2L share the same bipartite ATPase domain as the SNF2 group, but have a SANT (SWI3‐Ada2‐N‐Cor‐TFIIB DNA‐binding) domain instead of a BROMO domain (Figure 4). The SANT domain is related to the DNA‐binding domain found in MYB transcription factors (Aasland et al., 1996). Results suggest that this domain binds to histone N‐terminal tails in a manner that depends on the acetylation status of the tails. Interestingly, the SWI/SNF components BAF155 and BAF170 also contain a SANT domain (Boyer et al., 2002; Boyer et al., 2004). The SNF2L and SNF2H proteins also contain a SANT related SLIDE domain, which plays a role in nucleosome binding and chromatin remodeling (Figure 4) (Grune et al., 2003). There is no sequence conservation between the ISWI proteins and the SNF2 proteins, with the exception of the conserved ATPase domain, which suggests that they have different biological functions.

I II III SNF2H 1053 aa

SNF2L 1034 aa

Introduction 33

This is further illustrated by the facts that the ATPase activity of ISWI proteins is stimulated by DNA but requires nucleosomes for maximum activation, and that ISWI activity depends on histone N‐termini, a clear difference compared to the SNF2 proteins, see above and (Boyer et al., 2000; Corona et al., 1999).

The ISWI complex The ISWI proteins function in complexes in vivo and these complexes were first identified in Drosophila and later in mammals. The different ISWI‐ containing complexes that have been isolated to date have not revealed the large biochemical heterogeneity seen in the SWI/SNF family of protein complexes, but have a more limited repertoire in subunit composition. Yeast contains three ISWI complexes: ISWI1a, which contains Iswi1 and Ioc3; ISWI1b, which contains Iswi1, Ioc2 and Ioc4; and ISWI2, which contains Iswi2 and Itc1. Yeast also has the unique feature of containing free Iswi1 protein (Gelbart et al., 2001; Vary et al., 2003) The NURF (nucleosome remodeling factor) was the first ISWI‐containing complex isolated, and it was found in Drosophila as a factor that disrupts a regularly spaced nucleosome array in co‐operation with the GAGA transcription factor (Tsukiyama and Wu, 1995). NURF is made up of 4 polypeptides: Nurf301, ISWI, Nurf55 and Nurf38. A mammalian equivalent to NURF has recently been isolated, and is the only remodeling factor that contains the SNF2L protein (Barak et al., 2003). Another factor, ACF (ATP dependent chromatin remodeling and assembly factor), was also originally isolated from Drosophila embryo extracts and is made up of two subunits: Acf1 and ISWI. ACF was identified as a factor that facilitates the ordered spacing of nucleosomes in chromatin assembly reactions in vitro (Ito et al., 1997; Ito et al., 1999). A homologous complex has been identified in human cells with an identical subunit composition and identical in vitro activity. It is called hACF or WCRF after the human Acf1 homolog WCRF180, and contains the SNF2H ATPase (Bochar et al., 2000a; LeRoy et al., 2000). The chromatin accessibility complex, CHRAC, was also found in Drosophila and is highly similar to ACF. It contains 4 polypeptides: Acf1, ISWI and two novel, histone fold proteins: p17 and p15 (Varga‐Weisz et al., 1997). Topoisomerase II was reported as a part of the complex in the original purification, but additional purification revealed that this interaction was non‐specific and that the in vitro activity of CHRAC is unchanged with or without topo II. CHRAC assists in the creation of ordered nucleosome arrays, an activity it shares with


34

the ACF, which sets them apart from the NURF that mediates the opposite reaction. Human CHRAC has the exact same polypeptide composition as its Drosophila counterpart and contains the mammalian ISWI homolog SNF2H (Poot et al., 2000). The WICH complex combines SNF2H with the WSTF (Williams syndrome transcription factor) protein and this complex mediates the formation of an ordered nucleosome array in vitro. WHICH is present in NIH3T3 cells, in HeLa cells and in Xenopus oocyte extracts (Bozhenok et al., 2002). The NoRC (nucleolar remodeling complex) was isolated from mice and contains Tip5 and murine SNF2H and has the same in vitro activity as CHRAC (Strohner et al., 2001). The SNF2H‐cohesin complex combines SNF2H with Mi2/NURD components and cohesin proteins and is probably involved in loading cohesin onto chromosomes (Hakimi et al., 2002). RSF (remodeling and spacing factor) was isolated as a factor that facilitates activator‐mediated transcription initiation in vitro and facilitates both the spacing and remodeling of nucleosomal arrays. It was isolated from HeLa cells and is made up of two components: Rsf‐1 and SNF2H. No Rsf‐1‐related proteins have been identified (LeRoy et al., 1998; Loyola et al., 2003; Loyola et al., 2001). Interestingly, most of the proteins associated with ISWI, SNF2H and SNF2L, such as yItc1, Acf1, Nurf301, Tip5 and WSTF, belong to a novel protein family called the WAL/BAZ/BAH (wstf‐acf‐like/ bromo‐adjacent zinc finger/ bromo‐ adjacent homology) family (Jones et al., 2000). These proteins are not very similar on the level of amino acid sequence, except for a number of small and defined domains, and ordinary BLAST (basic local alignment search tool) searches do not reveal significant relationships. However, it becomes evident when comparing the overall domain structure of these proteins that they have a high degree of structural similarity. Database searches using the NCBI CDART (conserved domain architecture retrieval tool) search engine indicate that there are uncharacterized WAL/BAZ/BAH proteins within the human genome, suggesting that there might be novel SNF2H‐containing or SNF2L‐ containing complexes that have not yet been identified. The functional significance of the WAL/BAZ/BAH proteins in chromatin remodeling has not yet been fully elucidated, but in vitro studies show that they have a profound effect on the chromatin remodeling activity of ISWI proteins.

Introduction 35

ISWI in vitro activity Analysis of ISWI in vitro activity has revealed that these ATPases change the translational position of nucleosomes without perturbing the actual nucleosome structure. This mechanism is known as ʺslidingʺ but differs from the sliding activity of the SNF2 proteins in that these proteins actually change the nucleosome structure during the sliding process. On reconstituted mono‐nucleosomes, ISWI ATPases alter the translational position of the octamer from a central position to the extreme end of the DNA strand, and can actually push it slightly off the end of the DNA. A proposed mechanism for this is that ISWI proteins contact both linker DNA and the nucleosome, creating a position of leverage, and then ʺpushʺ the DNA against the histone octamer, in agreement with the Loop‐Recapture model (Langst and Becker, 2001; Langst et al., 1999). Chromatin remodelling by ISWI proteins depends on the presence of histone tails and on the acetylation state of the histone N‐termini. Results show that a 20 amino acid epitope on the H4 tail is essential, together with DNA, for ISWI binding, and that acetylation of nearby residues abolishes this interaction. These observations are consistent with the histone binding proper‐ties of the SANT domain (Clapier et al., 2001; Clapier et al., 2002). Remarkably, the activity of ISWI‐containing complexes differs from that of the free ISWI protein. Proteins associated with ISWI ATPases not only increase the specific activity of the enzyme but reverse the directionality of the sliding reaction so that nucleosomes are moved from the end of DNA fragments to a central position. This indicates that the geometry of the native complex is important for the specific properties of the catalytic core (Eberharter et al., 2001; Langst et al., 1999). ISWI‐containing complexes also modulate the spacing of nucleosomal arrays; some of them produce ordered arrays while others produce disordered arrays (Ito et al., 1997; Tsukiyama and Wu, 1995).


36

The biology of ISWI complexes Development In Drosophila, homozygous ISWI null mutants are not viable, but survive to the late larval or early pupal stage due to a large maternal contribution of ISWI protein. The entire X chromosome in male larvae is severely distorted, directly linking chromatin remodeling by ISWI to chromatin structure in vivo (Deuring et al., 2000). The structural effect of ISWI deficiency on the male X chromosome depends on the acetylation state of H4, where H4K16 acetylation counteracts substrate binding and chromatin compaction by ISWI (Corona et al., 2002). It was shown, using tissue‐specific expression of dominant negative forms of the ISWI protein, that the expression of the en and Ubx genes is severely reduced without ISWI activity. ISWI does not strongly co‐localize with RNApol II on polytene chromosomes, in contrast to these results, which is indicative of a role in repression rather than activation. It was suggested that ISWI was involved in both activation and repression due to the presence of tissue‐specific ISWI complexes, or that the effect on En and Ubx transcription was indirect (Deuring et al., 2000). Heterozygous SNF2H+/‐ mice develop normally and have a normal phenotype, while homozygous SNF2H‐/‐ embryos die during the peri‐implantation stage. Neither the inner cell mass nor the trophectoderm are viable in ex vivo cultures, and antisense depletion of SNF2H drastically inhibits the differentiation of CD34+GlyA‐ hematopoetic stem cells into erythrocytes (Stopka and Skoultchi, 2003). Both of the mammalian ISWI proteins are present in ES cells and incorporated into chromatin remodeling complexes (Bozhenok et al., 2002). Nevertheless, the closely related SNF2L protein cannot compensate for the loss of SNF2H, which is in stark contrast to the situation with BRG1 and BRM, where BRG1 is able to compensate for the loss of BRM. The mammalian ISWI proteins have distinctly different patterns of expression in mouse embryos. SNF2L is highly expressed in terminally differentiated cells in the brain, testis, ovaries and lymphocytes; while SNF2H is highly expressed in proliferating cells in the same organs. This correlation between SNF2H and proliferation is consistent with the phenotype seen in homozygous null embryos (Lazzaro and Picketts, 2001).

Introduction 37

Transcription The role of ISWI proteins in transcription has been studied most extensively in yeast, which contains the ISWI1a, ISWI1b and ISWI2 complexes. The ISWI1a and ISWI1b complexes have both shared and unique target genes as revealed by expression profiling, while ISWI2 regulates a separate set of genes (Vary et al., 2003). The Iswi2 complex represses early meiotic genes and is recruited by the sequence‐specific, DNA binding protein Ume6 (Goldmark et al., 2000). In vivo analysis of promoter structure combined with expression analysis revealed that both Iswi1 and Iswi2 mostly mediate repression of transcription, an observation consistent with the low degree of co‐localization of ISWI and RNApol II on Drosophila polytene chromosomes. Iswi1 and Iswi2 deletion strains have an altered chromatin structure on certain promoters associated with an up‐regulation of transcription from these de‐repressed genes (Kent et al., 2001). Interestingly, yIswi proteins take part not only in initiation of transcription but also in elongation and termination (Mellor and Morillon, 2004). The mammalian NoRC is also involved in the repression of transcription, specifically of rRNA genes. Repression occurs at a step preceding PIC formation, and NoRC is unable to repress rRNA genes that are already active. NoRC facilitates the recruitment of HDACs and histone methylases, leading to HP1 binding and the formation of heterochromatin‐like structures on rDNA (Santoro et al., 2002; Strohner et al., 2001; Strohner et al., 2004; Zhou et al., 2002). In contrast, mammalian NURF plays a role in the activation of transcription. Both SNF2L and the mammalian Nurf301 homolog BPTF are associated with the promoter regions of the human En‐1 and En‐2 genes, regulators of mid‐hindbrain development. RNAi mediated knock‐down of SNF2L expression and BPTF expression reduced the levels of En‐1 and En‐2 mRNA levels (Barak et al., 2003). Replication Most studies of ISWI complexes in Drosophila and mammals have focused on their role in replication. In Drosophila embryos, loss of Acf1, a subunit of both ACF and CHRAC, decreases the density of nucleosome spacing and induces a faster rate of transition through the S‐phase. It has been suggested that ACF and/or CHRAC represents the major nucleosome spacing activities in these embryos, and that these factors are therefore responsible for the formation of repressive chromatin structures. The replication machinery is therefore able to


38

replicate the genome more rapidly due to the general decrease in the amount of repressive chromatin following the loss of Acf1 (Fyodorov et al., 2004). The mammalian ACF and CHRAC have also been linked to replication. However, in contrast to the situation in Drosophila embryos, RNAi depletion of Acf1 in mammalian cells leads to a delay in the replication of pericentromeric heterochromatin. It has been suggested that without the assistance of ACF and/or CHRAC, the replication machinery requires a longer time to replicate heterochromatin because it has difficulties in traversing and replicating these dense structures (Collins et al., 2002). The apparent disparity between Drosophila and mammals in the role of Acf1 in replication is not clearly understood, but it has been suggested that this disparity is related to the difference in species or that it is a result of the difference between somatic and embryonic chromosomes. Interestingly, replication and histone deposition onto DNA in cell‐free systems using Xenopus egg extracts is unaffected by ISWI depletion, but the overall spacing of nucleosomes is greatly disturbed (MacCallum et al., 2002). Mammalian WICH has also been indirectly linked to replication, since it accumulates in pericentromeric heterochromatin during replication (Bozhenok et al., 2002).

39 Present Investigation

PRESENT INVESTIGATION

Paper I:

Expression of BRG1, a human SWI/SNF component, affects the organization of actin filaments through the RhoA signaling pathway

Summary

1. BRG1 expression induces a dramatic rearrangement in the organization of the actin filament system in SW13 cells. Untreated cells have a very weak organization of filamentous actin but form thick stress fiber like structures in response to BRG1 (Paper I: Figure 1).

2. The effect is absolutely dependent on a functional ATPase domain in

BRG1 since the BRG1K298R ATPase deficient mutant protein has no detectable effect on actin filament organization (Paper I: Figure 1).

3. The reorganization is mediated through the RhoA signaling pathway at

a point downstream of the RhoA protein itself since no change in the activation status of RhoA is detected. This suggests that one or more components in this signaling pathway are affected by BRG1‐SWI/SNF (Paper I: Figure 5 and 6).

4. The level of the RhoA downstream effector protein ROCK 1 is elevated

in response to BRG1. Expression of both wt and constitutively active forms of this protein in SW13 cells induce a reorganization of actin filaments that is morphologically highly similar to that induced by BRG1 (Paper I: Figure 7 and 8).

5. The increase in ROCK 1 protein levels is posttranscriptional since no

change in mRNA levels is detected (data not shown).


40

Supplementary information The discovery of the connection between BRG1 and actin filament organization was made during our efforts to isolate stable BRG1 expressing clones of SW13 cells. It is well established that BRG1 expression induces cell cycle arrest, flat cell formation, senescence and cell death in these cells. Changes in cell morphology are intimately connected to the microfilament system which made us investigate the organization of F‐actin in BRG1 positive SW13 cells. Interestingly, we have discovered that filament reorganization is strongly dependent on the level of BRG1, suggesting a dose‐response relationship. We used a strong expression vector that rapidly caused flat cell formation and cell death, in agreement with published data. We were, however, able to do some analyses before these clones senesced which lead to the discovery of the microfilament system reorganization. Expressing BRG1 from a slightly weaker expression vector enabled us to isolate viable stable clones. These BRG1 positive SW13 cells are larger than the parental cells and display the same actin filament reorganization as the unstable clones obtained with the stronger expression vector, although the changes are not as dramatic. The cell cycle profile of the BRG1 positive clone revealed less cells in the G1‐ phase and in the M‐phase and an increase in the number of cells with a DNA‐content larger than 4N, indicative of problems with chromosome separation (Figure 5: compare b and c). These cells have a longer generation time than the original SW13 cells. However, we have not detected signs of senescence associated β‐gal activity which, in combination with the fact that only a low level of BRG1 expression allowed the isolation of stable clones, suggests that BRG1 mediated effects on both actin filament organization and the cell cycle are closely related to the level of BRG1 expression. Over time the clones slowly loose their BRG1 expression, indicating that there is a continuous negative selective pressure against BRG1, and the cell cycle profile becomes similar to that for SW13 cells (Figure 5f and compare c and d with b). In parallel with these lower BRG1 levels, the slightly larger size of the clones and the reorganization of the actin filament system slowly revert back to a point where the cells become morphologically indistinguishable to the parental SW13 cells. The ATPase deficient BRG1 protein does not have a detectable effect on the cell cycle profile of SW13 cells (Figure 5e).

Present Investigation 41

A HeLa B SW13

C SW13BRG1 6p D SW13BRG1 57p

E SW13BRG1/K783R 53p F

Figure 5. Cell cycle profile of SW13, BRG1wt+ and BRG1K783R+ cells. a‐e. Cells were stained with propidiumiodide and DNA Content analyzed using a Becton Dickinson FACS. f. BRG1 expression in FACS analyzed cells in b‐e. p6 and p57 refer to the number of passages of the BRG1 positive clone.

Interestingly, we have also been able to link BRG1 and microfilament organization in the BRG1 deficient, human breast cancer cell line ALAB. However, antisense depletion of BRG1 expression in HeLa cells causes cell death, and dominant negative interference using the ATPase mutant form of BRG1, does not interfere with actin filament organization in HeLa cells or in human 1523 fibroblasts. Considering the fact that there is an apparent cell type specific SWI/SNF regulation of genes (Hendricks et al., 2004), it is not unlikely that different cell types will vary in their response to BRG1.


42

The ATPase deficient BRG1 protein also has an effect on SW13 cells, although it is subtle. Clones with a stable expression of BRG1K783R have an increase in the number of multi‐nucleated cells compared to the parental SW13 cells, something also seen in the BRG1wt expressing clone, and the cells are slightly larger. These clones are so far completely stable in culture and show no decrease in the expression of the mutant BRG1 protein. This indicates that the SWI/SNF complex as a whole has more functions in transcription than just chromatin remodeling, maybe in mediating interactions between components in the transcription machinery. This ATPase independent effect has also been seen in the activation of the MMTV promoter (Trotter and Archer, 2004). Our experiences from creating the stable BRG1 positive clones suggests that there is a threshold above which BRG1 expression induces rapid cell cycle arrest, flat cell formation, senescence and cell death but if BRG1 expression is kept below this threshold, the cells are able to survive and develop a characteristic phenotype. It is possible that the effects observed when over‐expressing BRG1 in SW13 cells may not be physiologically relevant and that high levels of BRG1 protein are toxic and not a physiological outcome of SWI/SNF activity. This would be analogous to the effects seen when over‐expressing Rb‐E2F proteins and may explain why it is difficult to get stable BRG1 expressing clones of cells that already have endogenous BRG1 expression. Consequently, care must be taken when interpreting the outcome of transient over‐expression of BRG1.


Paper II:

Variations in biochemical composition and chromatin

association of mammalian SWI/SNF complexes Summary

1. The composition of biochemically purified SWI/SNF complexes from mammalian cells differs from the interactions observed in immuno‐precipitations and from the patterns in chromatin binding of individual subunits. The biochemically purified complexes show differences in remodeling activity on reconstituted mononucleosomes, suggesting that small differences in complex composition can have a significant impact on function (Paper II: Figure 1‐4).

2. Most of the Baf180 protein interacts strongly with Baf250 in an immuno‐

precipitation but have both separate and overlapping binding to chromatin. Consequently, the PBAF form of SWI/SNF constitutes a minor fraction of the total SWI/SNF population in HeLa cells. This is also reflected in the low amount of complex 5, the only purified complex that contains detectable amounts of Baf180 (Paper II: Figure 1 & 4).

3. The Baf155 and Baf170 proteins have been regarded as constitutive

subunits of SWI/SNF but show both overlapping and separate binding to chromatin. This shows that the plasticity of SWI/SNF complex composition is larger than previously believed (Paper II: Figure 6).

4. We do not detect an interaction between SWI/SNF and actin using

biochemical methods but actin is found associated to the same chromatin locations as SWI/SNF components (Paper II: Figure 2,4 &6).

5. BRG1 interacts with the splicing factor Prp8, linking SWI/SNF to mRNA

processing (Paper II: Figure 5).

6. We propose a model where the biochemical heterogeneity among SWI/SNF complexes is partially a result of function dependent on the local chromatin environment to where the complexes are recruited.


44

Supplementary information ChIP cloning: In order to investigate the full range of possible chromatin targets for WI/SNF complexes we designed a ChIP cloning procedure using the BRG1 protein as bait and HeLa chromatin. The use of chromatin immunoprecipitation has become extensive in the last years and is usually combined with PCR detection of precipitated DNA or slot‐blot hybridization which is a way of avoiding unspecific signals. In a ChIP cloning procedure, however, all fragments will be picked up in the cloning procedure, even unspecific DNA. To eliminate such sequences we performed a ChIP from SW13 cells that have no detectable expression of BRG1. To increase the efficiency of this approach, we scaled‐up this ChIP ten times compared to the one performed in HeLa cells but. Note that this SW13 ChIP included αBRG1 antibodies.

.

>500bp

< 500bp

Figure 6 Dot‐blot detection of unspecific DNA in αBRG1 ChIP clones from HeLa cells Genomic DNA from SW13 cells and HeLa cells as positive controls. Negative control: pORSVI cloning vector DNA. Notice the weak but distinct signal from the negative control.


The cloned fragments from the HeLa ChIP were dot‐blotted onto a nylon membrane, and the fragments from the SW13 ChIP were labeled with p32 and used to probe the clones for unspecific DNA (Figure 6). We obtained 200 colonies from the HeLa ChIP but only 120 of these contained detectable inserts. These 120 clones were screened for unspecific sequences as seen in figure 6. Using the signal from the negative vector control as background, we estimated that 75 clones remained as possible candidates. We have so far sequenced 20 of these clones and 8 of them have been positive BRG1 targets in an analytical ChIP assay. Using this as an estimate, our screen will have generated approximately 30 true BRG1 targets out of 75 relatively likely candidates. This example clearly illustrates the potential for unspecific results in ChIP cloning and the need to verify their validity. The next step in our cloning will be to elucidate the functional aspects of these sequences. Those that are located in proximal promoter regions will be relatively easy to investigate by analyzing mRNA levels of these genes when expressing BRG1 in BRG1 negative cell lines such as SW13, ALAB or C33A. The same is true for those targets that are located deep within coding regions, especially since we have identified a possible function for BRG1 in mRNA processing (Paper II: Figure 5). Those that are located in intergenic regions, however, will require more elaborate approaches to determine their function, for example as distal enhancer elements. SWI/SNF and actin: The interaction between SWI/SNF and βactin or, more precisely, the proposed role of βactin as a true subunit of SWI/SNF complexes has proven to be difficult to verify. In some cases this interaction is strong and unambiguous while in other, βactin is not detected at all. We therefore asked ourselves two questions: 1. Can we detect nuclear βactin? 2. Can we find any evidence that an interaction between βactin and BRG1 is even possible? To answer the first question, we carefully fractionated HeLa cells and separated the nuclei from the cytoplasmic compartment. The nuclei were then extracted with increasing concentrations of salt, 0‐1M KCl. As markers for the degree of cytoplasmic contamination of the nuclei, we probed for the presence of tubulin and vinculin. As seen in figure 7a, β‐actin is located in the nuclear compartment and is even more strongly associated with nuclear structures than BRG1 since it requires slightly more salt to be efficiently extracted.


46

As judged by the absence of tubulin and vinculin in the nucleoplasmic fractions, we believe that the nuclei are mostly free from cytoplasmic contamination, and that the βactin signal is a sign of a true nuclear localization (Figure 7a). In figure 7b, we have stained HeLa cells with the same βactin antibody as in a, and co‐stained with DAPI. There is a clear nuclear localization behind the strong cytoplasmic signal. The fact that nucleoli are visible supports the validity of the immunofluorescence localization (Figure 7b).

Figure 7. Nuclear localization of βActin in HeLa cells A. Cellular fractionation of HeLa cells where nuclei were extracted with increasing KCl concentrations. B. Immuno‐fluorescence images of HeLa cells stained with monoclonal anti‐ bodies against βActin and co‐stained with DAPI.

Even though a pool of nuclear actin was detected in HeLa cells it did not necessarily mean that it interacts with SWI/SNF. We therefore performed Superose 6 HR gelfiltration analyses of crude HeLa nuclear extract to see if there was any high Mr actin present in the same fractions as BRG1.


Figure 8. Gelfiltration analysis on the effect of ATP on BRG1 and βactin. 0.5 mg crude HeLa nuclear extract was separated on a Superose 6 HR column using the same buffers as described in Paper II, with or without ATP. Flow‐rate: 0.2 ml/min, Fraction size: 0.6 ml. 100μl of each fraction was precipitated with TCA and separated in a 7% SDS‐PAGE. NE: 30μg crude HeLa nuclear extract.

From the elution profiles in figure 8 (‐ATP) it is clear that no βactin is present in high Mr weight fractions together with BRG1 under normal conditions, which is an indication that there is no significant interaction between BRG1 and βactin. However, actin and actin dynamics is influenced by ATP and the addition of 0.1 mM ATP during extract preparation and column run induced a change in βactin elution so that it was detected in fractions together with BRG1. This should not necessarily be interpreted as an ATP‐induced stabilization of a protein‐protein interaction between βactin and BRG1. It could mean that the ATP drives the βactin in the nuclear extract to polymerize which, in fact, could destroy an interaction with BRG1 by recruiting those molecules away from SWI/SNF and into filaments. Despite this possibility, I decided to include ATP in the purification of SWI/SNF, because without it, there was no high Mr actin and, therefore, no possibility of an interaction. In addition, ATP does not have a significant effect on the SWI/SNF complex as seen by the elution profiles of BRG1. The slightly wider peak of BRG1 from the column with ATP than from that without ATP is an effect of the fact that there is more material loaded onto the column in the ATP sample. Together with the fact that the BRG1 distribution varies slightly between experiments, with or without ATP, made me believe that ATP did not have any significant effect on the SWI/SNF complex itself.


48

Paper III:

WSTF‐SNF2h interacts with several nuclear proteins in a transcription dependent manner, to form a functional unit B‐WICH.

Summary

1. The previously identified WSTF‐SNF2H complex, WICH, interacts with several nuclear proteins to form a novel unit designated B‐WICH with an estimated mass of 2‐3 MD (Paper III: Figure 1).

2. Components of B‐WICH so far identified are: WSTF, SNF2H, the

splicing factor SAP155, the RNA helicase RNAhelicase II/Guα, the transcription factor Myb‐binding protein 1a, the transcription factor/DNA repair protein CSB and the RNA processing factor DEK (Paper III: Figure 2).

3. The interactions between the WSTF‐SNF2H core and the other

components are mediated by RNA and are sensitive to RNAse digestion (Paper III: Figure 3).

4. The formation of the B‐WICH complex is dependent on active

transcription since the RNApol II inhibitors actinomycin D and DRB prevent the formation of the complex. Interestingly, the smaller, B‐WICH like complex found in these extracts is still larger than the original WICH complex (Paper III: Figure 4).

5. We suggest that the WSTF protein functions as a platform capable of

mediating the assembly of several large complexes in vivo involved in processes such as replication, transcription and RNA processing. This is another example of the growing connection found between ATP‐dependent chromatin remodeling and RNA processing.


Supplementary information Biochemical properties of the WSTF protein: The discovery of the interaction between WSTF and SNF2H was made based on the identification of the WSTF protein as a member of the WAL/BAZ family. During the initial stages of our characterization of this interaction, the isolation of the WICH complex was reported, containing WSTF and SNF2H, and this complex was suggested to have a role in DNA replication. However, we had discovered that the majority of the WSTF protein was not efficiently extracted from HeLa nuclei below 0.7 M KCl, and that these extracts contained two forms of WSTF containing complexes. In our efforts to find the optimal biochemical conditions to purify these complexes, we had also discovered that passing the extract over a Heparin column efficiently disrupted the B‐WICH complex, leaving only the WICH complex, as seen in Superose 6 HR gelfiltrations. These two discoveries made us to conclude that the high Mr complex seen in our extracts was a physiologically relevant find and this eventually led to the identification of the B‐WICH complex. The cellular localization of the WSTF protein: We have also investigated the cellular localization of the WSTF protein. This was brought about by the difference in staining obtained by the commercial C‐terminal αWSTF antibody used by us and the N‐terminal antibody developed by Varga‐Weisz and co‐workers. Interestingly, we saw a clear variation in staining between NIH3T3 cells, used in the identification of WICH, and HeLa cells. As shown in figure 9, the WSTF protein in human cells, A‐D, is localized to the nucleus in a granular pattern that is evenly spread throughout the nucleoplasm. Mouse, rat and porcine cells (Figure 9 E‐H) have a similar localization but, in addition, WSTF is found in a rim‐like structure around the entire nucleus. This could indicate that a portion of the total WSTF population in these cells is associated with the nuclear membrane, either directly or indirectly. It is possible that the WSTF protein in humans and other species have slightly different properties and differs in regulation and/or function. This suggests that care must be taken when interpreting data about WSTF and when applying them to other species.


50

Figure 9. Immunofluorescence localization of the WSTF protein. A‐D human cells, E and F mouse cells, G rat cells and H porcine cells.


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ACKNOWLEDGEMENTS So here I am at last, after all these pages and all these years, and I canʹt quite convince myself that it is all over, or maybe I should limit myself to saying that this part is over. There are so many people that I am indebted to for making it all possible and I hope I donʹt forget anyone: To Anki I hope you realize what a perfect supervisor you have been. Always knowing the exact moment when I needed to be slowed down, and being able to do so without making me feel restrained or locked into a corner. I can only imagine how exhausting it must have been at times to try to keep up with me running around and doing experiments close to light speed. Bit you did it, you survived, and now you can finally rest. Ha Ha Thank you so much for you patience, for your ability to teach, for your sharp eye that always spotted my mistakes, for all the laughs and for having the same weird humor as myself. And maybe the most important thing of all: Thank you, for letting me be me. To Erica I donʹt think that I ever again will I have the privilege and joy of working with a colleague such as you. Your sense of humor perfectly matches my own weird humor and your critical eye always spotted every attempt I made at cutting corners (Ha Ha). Your ability to teach and explain science to me and the students we had (except for the unfortunate incident with the matrix explanation). Thank you for being a huge part in making these years some of the most interesting, most rewarding, funniest and stimulating I have ever experienced. Good luck with your own thesis next year, I will be there! To Margareta Your humor, your knowledge and your generosity and patience when teaching me all those things when I was a fresh, overambitious and unstoppable PhD student. I must have been so annoying and exhausting! Thank you for all your help and for being a big and important part of my time at the department.


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To the people at the Dept. of Cellbiology I would like to thank Prof. Uno Lindberg and Prof. Roger Karlsson for always making time for me and my questions and for their helpful and challenging discussions. Ingegärd, how you always know where to find things is still a mystery to me but thank you for all your help and support. To Staffan, the myth, the man the legend: Your sharp mind, your support, your wit and humor has meant a lot to me: Thank you! To Li‐Sophie, Louise, Thomas and Yu: thank you for all the help and all the fun over the years and good luck with your own thesis work. (Thomas, without your help I would still be lost in cyberspace). Thank you Ingrid and Ingela for being helpful colleagues and thanks for all the fun. Former colleagues I would like to thank Herwig, Kartik, Mats, Petra, Thomas and Åsa for rewarding discussions, help and a lot of fun. I would especially like to thank Maria for all the discussions, help and support and for all the laughs and strange conversations. You are one of a kind, don’t ever stop! My family Mom, dad, ʺsyster‐ysterʺ Madde and my nephew Dennis: Thank you for your support, your encouragement and unstoppable belief in my abilities. Thank you to the rest of my family for being supportive and for being who you are. You know I really like my extended family with grandparents, aunts and uncles, cousins and so on. To Christian I know you don’t really understand what Iʹve been doing or why Iʹve been doing it but you have never failed to support me anyway. Thank you for being an amazing friend despite the fact that Iʹve regularly disappeared into science for months at an end in the last five years. Big hug to you! I am grateful for the financial support: Stipend in memory of Wanda Pokora‐Kulinska, The JA Ekströms foundation, The E & L Åqvist foundation, The Liljewals foundation and for financial support granted to Dr. A‐K Östlund Farrants from The Swedish Natural Science Research Council, The Carl Trygger Foundation, Magnus Bergvall’s foundation, the Nilsson‐Ehle Foundation, Foundation for Lars Hiertas Minne and Marcus Borgström’s Foundation


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Chromatin Remodelling by BRG1 and SNF2H190933/FULLTEXT01.pdf · Chromatin remodeling by BRG1 and SNF2H Biochemistry and Function 4 This thesis is based on the following papers: Paper