Iowa State University Capstones, Theses and Graduate Theses and Dissertations Dissertations

2012 Understanding Higher Order Heterochromatin Structure And Its Determinants Abdelhamid Azzaz Iowa State University

Follow this and additional works at: https://lib.dr.iastate.edu/etd Part of the Biochemistry Commons

Recommended Citation Azzaz, Abdelhamid, "Understanding Higher Order Heterochromatin Structure And Its Determinants" (2012). Graduate Theses and Dissertations. 12266. https://lib.dr.iastate.edu/etd/12266

This Dissertation is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Graduate Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. Understanding Higher Order Heterochromatin Structure And Its Determinants

By

Abdelhamid Azzaz

A dissertation submitted to graduate faculty in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSPHY

Major: Biochemistry

Program of Study Committee: Michael Shogren-Knaak, Major Professor Amy Andreotti Kristen Johansen Yanhai Yin Louisa Tabatabai

Iowa State University

Ames, Iowa 2012

Copyright © Abdelhamid Azzaz, 2011. All rights reserved ii

To Rania and Omar

iii

Table of Contents ABSTRACT ...... v CHAPTER 1: GENERAL INTRODUCTION ...... 1 1.1 Histone methylation and the “histone code” ...... 1 1.2 Heterochromatin formation ...... 2 1.2.1 The Chromo domain ...... 4 1.2.2 The Chromo shadow domain ...... 5 1.2.3 The hinge region ...... 5 1.2.4 Localization ...... 6 1.2.5 Functions ...... 6 1.2.6 Transcription and RNA interface in the formation of Heterochromatin ...... 8 1.3 Dynamic nature of heterochromatin protein 1 binding ...... 9 1.4 HP1 and higher-order chromatin structure ...... 10 1.5 Generation of in vitro heterochromatin model using native chemical ligation of histone proteins ...... 11 Dissertation Organization ...... 12 References ...... 12 CHAPTER 2. MULTIPLE INTERACTIONS BETWEEN HETEROCHROMATIN PROTEIN 1 (HP1) AND NUCLEOSOMES...... 16 Abstract ...... 17 Introduction ...... 18 Experimental procedures ...... 19 RESULTS ...... 27 Discussion ...... 39 References ...... 42 Supplemental Figures ...... 46 CHAPTER 3. HP1-MEDIATED CONDENSATION OF METHYLATED CHROMATIN ...... 50 Abstract ...... 51 Introduction ...... 51 Experimental procedures ...... 52 Results...... 55 Discussion ...... 64 References ...... 66 iv

Supplementary figures ...... 68 CHAPTER 4. GENERAL CONCLUSIONS...... 74 ACKNOWLEGMENT ...... 76

v

ABSTRACT

Heterochromatin is a higher order unique compacted chromatin structure present at the centromeres and the telomeres of chromosomes. It is characterized by the methylation of histone H3 on lysine 9 and the binding of a key structural and functional protein called heterochromatin protein 1 (HP1) to that histone mark. The structural determinants for such binding were not clear. In this study, we show that in vitro HP1 can bind to uniformlyH3-K9 methylated nucleosomal arrays in a methylation dependent fashion, and this binding promotes the formation of a more compacted structure. This binding is shown to be dependent on the ability of HP1 to form a dimeric structure through its chromoshadow domain and also upon its ability to bind methylated lysine through its chromo domain. Also,

HP1 has shown some ability for non-specific DNA-dependent binding to unmethylated arrays under certain conditions. The more specific methylation dependent binding is shown to promote intra-molecular and inter-molecular compaction of methylated arrays. In vivo, we show that the ability of HP1 to bind methylated lysine and to dimerize is essential for its in vivo functionality. Using molecular modeling we show that HP1 can adopt several binding positions to methylated arrays without steric hindrance. 1

CHAPTER 1: GENERAL INTRODUCTION

In eukaryotes, the nuclear DNA associates tightly with basic histone proteins and nonhistone proteins to form chromatin fibers, which make up chromosomes. The chromatin fibers are composed of nucleosome arrays, with each nucleosome consisting of 147 bp of

DNA coiled roughly 1.7 times around an octamer of histone proteins (2 copies of each of the core histones, H2A, H2B, H3 &H4)[1]. Each histone protein consists of a core part folded in what is known as the histone fold (helix-turn-helix), and an N-terminal “tail” of 15-38 amino acids (H2A has also a C-terminal tail), which extends past the DNA and is unstructured.

Extended chromatin appears as an array of nucleosomes, but in the nucleus, the chromatin fibers forming chromosomes undergo several levels of folding, resulting in increasing degrees of condensation[2]. Histone tails play an important role in this folding process[3].

1.1 Histone methylation and the “histone code”

An important characteristic of histones, and especially of their tails, is the large number and the type of modified residues they possess. There are at least eight distinct types of modifications found on histones (acetylation, lysine methylation, arginine methylation, phosphorylation, ubiquitylation, sumoylation, ADP ribosylation, deimination and proline isomerization)[4]. Histone methylation occurs on arginine and lysine residues and is catalyzed by enzymes belonging to three distinct families of proteins, 1) the PRMT1 family,

2) the SET-DOMAIN containing protein family and, 3) the non-SET-domain proteins 2

DOT1/DOT1L. Lysine methylation is better understood and has several important functions[5].

Considering the importance of histone tails in forming higher order chromatin structures and the fact that they are the target of methylation, it is not surprising that histone methylation is used to modify chromatin structure. This could affect chromatin-based processes such as transcription, heterochromatin formation and DNA repair, assuming that folding alters the accessibility of DNA to the proteins that mediate these processes. There is no direct evidence that lysine methylation directly affects chromatin dynamics. As lysine methylation does not change the charge, any direct effect of lysine methylation on chromatin folding would have to occur through a non-electrostatic mechanism (for example, through hydrophobic interactions)[5]. An alternative hypothesis proposes that specific histone modifications, including lysine methylation, are binding sites for different proteins that mediate downstream effects[6]. Methylated lysines are recognized by the chromo-like domains of the Royal family (chromo, tudor, MBT) and non-related PHD domains[4].

1.2 Heterochromatin formation

Histone modifications help partition the genome into distinct domains such as euchromatin, where DNA is kept “accessible” for transcription and heterochromatin, where chromatin is “inaccessible” for transcription. Heterochromatin is an important structure with diverse functions, including silencing of transcription[7], protection of chromosome ends[4], also heterochromatin formation is required for the proper segregation of chromosomes during mitosis[5, 8] and it plays a crucial role in recombination events that are associated with 3

mating-type switching in yeast[9]. A link between heterochromatin formation and gene silencing has been inferred from the loss of most of the gene activity on the inactive X chromosome, which is visibly condensed in female mammals, and from the loss of gene expression, correlated with condensed packaging, in position-effect variegation (PEV) in

Drosophila and other organism. PEV occurs when a gene that is normally euchromatic is juxtaposed with heterochromatin, through rearrangement or transposition; the resultant variegating phenotype indicates that the gene has been silenced in a proportion of the cells in which it is normally active[10]. Studies of PEV in Drosophila melanogaster have resulted in the identification of a number of PEV suppressors including Su(var)3-9[11], and its human homologue Suv39H1 that were later shown to be histone methyltransferases with a specificity for histone H3 lysine9 (H3-K9)[12]. A role for Suv39H1 and its associated H3-K9 methyltransferase activity in heterochromatin function was indicated by the demonstration that it associates with the heterochromatin protein HP1[13]. Subsequently, it was shown that methylation of H3-K9 provides a binding site for the chromo domain of the HP1 protiens[14]. These molecular events were conserved during evolution. For example, in fission yeast, H3-K9 methylation is catalyzed by the SUV39H1 homologue Clr4[15]. Clr4 mediated H3-K9 methylation serves to recruit Swi6, the fission yeast homologue of mammalian HP1a[14, 15].

The sequence and structure of HP1 proteins can be divided into three regions (figure

1). First, the chromo domain (CD) at the amino terminus that is responsible for HP1

binding to di- and trimethylated H3-K9[14]. Second, the carboxy-terminal chromo shadow

domain (CSD) is involved in homo- and/or heterodimerization and interaction with other 4

proteins[16, 17]. Third, the chromo domain is separated from the CSD by a variable linker

or hinge region containing a nuclear localization sequence[18].

Figure 1. HP1 domains and their interacting partners.

1.2.1 The Chromo domain

The structure of the CD has been analyzed by nuclear magnetic resonance spectrometry[19] and X-ray crystallography[20]. The domain folds into a globular conformation approximately 30 Å in diameter, consisting of an antiparallel three-stranded β sheet packed against an  helix in the carboxy-terminal segment of the domain[19]. Methyl lysine recognition involves a conserved aromatic pocket, where the aromatic residues Tyr24,

Trp45 and Tyr48 form a 3-walled cage into which the methyl ammonium group inserts[20].

Additionally, the CD specificity for H3-K9 methylation is accomplished by binding interactions with residues flanking H3-K9me. Residues Q5, T6, A7, and R8 of H3 tail form β sheet interactions with residues E23, Y24, V26, N60, D62 in the CD. T6 and A7 form complementary Van der Waals contacts, whereas H3-Ser 10 forms H-bond with the CD Glu

56[20].

5

1.2.2 The Chromo shadow domain

The overall structure of the CSD is very similar to that of the CD. Like the CD, the

CSD is composed of 3 antiparallel β strands. Unlike the CD, which has a subsequent single α helix that folds against the sheet, the CSD has 2 carboxy-terminal α helices. Additionally, unlike the CD, the CSD can dimerize[21]. The dimerization through the CSD results in the simultaneous formation of a putative protein-protein interaction pit at the dimer interface with a non-polar base that can accommodate HP1-interacting proteins containing the consensus sequence PXVXL[16]. The CSD was shown to be responsible for the targeting of

Drosophila HP1a to heterochromatin[18].

1.2.3 The hinge region

The hinge region connects the CD and the CSD and is less well conserved. This region contains the most variable amino-acid sequence between HP1 proteins, between proteins both from the same species and from different species[22]. The linker is highly susceptible to post-transcriptional modifications, especially phosphorylation[23, 24].

Additionally, modifications within this region have been shown to affect localization, interactions and function[23]. It was shown that the hinge region has a bipartite nuclear localization sequence (NLS) that is conserved among eukaryotic species, and it is responsible for the distinct localization patterns observed among different Drosophila melanogaster HP1 homologs[18]. Therefore, the linker could be a central region in the regulation of HP1 proteins.

6

1.2.4 Localization

HP1 proteins localize not only to heterochromatin as their name suggests, but also to euchromatic sites in the eukaryotic nuclei[18, 23, 25]. The localization seems to be isoform- specific: in mammalian cells, HP1α and HB1β are mainly heterochromatic, whereas HP1γ is observed in both heterochromatin and euchromatin[25]. In Drosophila, HP1a is heterochromatic, HP1b is distributed in both heterochromatin and euchromatin, while HP1c is exclusively euchromatic[18].

Repetitive DNA segments, found at centromeres (pericentric heterochromatin) and telomeres, are enriched in HP1α (HP1a in Drosophila).26 HP1 proteins have been localized to the nuclear periphery and this may be associated with their interaction with the lamin B receptor and/or with the localization of centromeric heterochromatin. 26 HP1α has also been shown to co-localize with transcriptionally active domains of polytene chromosomes and, in both mouse and human, HP1 proteins, in particular HP1γ, has been associated with transcriptional elongation[23]. 28 Thus, despite its name and its predominant localization at heterochromatin, HP1 seems to have different roles in different nuclear environments[22].

1.2.5 Functions

Heterochromatin formation and gene silencing are two of the primary functions of

HP1. Targeting HP1 to chromatin in vivo was shown to require not only histone H3 lysine 9 methylation but also a direct protein-protein interaction between HP1 on one side and

SUV39h1 and other auxiliary factors on the other side (figure 2)[7, 26]. Thus through such interaction with the H3K9 methyltransferase, HP1 forms the basis for its spreading and for an autoregulatory loop, helping to maintain the methylated state of heterochromatin[26]. In 7

vitro, HP1 showed selective recognition and specificity in binding to methylated lysine 9 on histone H3[14, 27].

Figure 2. HP1 and its interactions. HP1 interacts with H3K9me2 and H3K9me3 through its chromodomain, and with SU(VAR)3-9 through its chromoshadow domain. By interacting with both the modified histone and the enzyme responsible for the histone modification, HP1 provides a foundation for heterochromatin spreading and epigenetic inheritance.

HP1 was reported to interact directly through its chromo shadow domain or indirectly with various non-histone proteins with varying functions. These interacting proteins are involved in cellular processes ranging from transcriptional regulation, chromatin modification, and replication to DNA repair, nuclear architecture and chromosomal maintenance[22]. The chromo shadow domain binding proteins have the motif PXVXL which is sufficient for interaction with dimerized chromo shadow domains[28]. Interaction occurs through binding of the peptide across the HP1 dimer interface, so that it forms a parallel β sheet with the carboxy-terminal tail of one monomer and an antiparallel β sheet with the tail of the other monomer. Stable retention of HP1 in heterochromatin was shown to require this interaction with PXVXL-containing motifs, besides the simultaneous interaction of the chromo domain with methylated H3K9 required for efficient localization. 8

1.2.6 Transcription and RNA interference in the formation of Heterochromatin

The RNA interference (RNAi) machinery was found to be essential for the establishment and maintenance of heterochromatin domains[22]. The RNAi machinery components were shown to be important for the proper localization of HP1, as the loss of or mutations in these components in

Schizosaccharomyces pombe (fission yeast), Drosophila and mouse resulted in abnormal localization of HP1[29-31].

The model for RNAi-mediated heterochromatin assembly and silencing in S.pombe is shown in figure 3[10]. Centromeric repeat (dg and dh) transcripts produced by Pol II are processed by the

RNAi machinery, including the complexes RITS and RDRC (which interact with each other and localize across heterochromatic regions). The slicer activity of Ago1 (a component of RITS) and the

RNA-directed RNA polymerase activity of Rdp1 (a component of RDRC) are required for processing the repeat transcripts into siRNAs. The siRNA-guided cleavage of nascent transcripts by Ago1 might make these transcripts preferential substrates for Rdp1 to generate double-stranded RNA, which in turn is processed into siRNAs by Dcr1. The targeting of histone-modifying effectors, including the

Clr4-containing complex, is thought to be mediated by siRNAs. This process most probably involves the base-pairing of siRNAs with nascent transcripts, but the precise mechanism remains undefined.

Methylation of H3K9 in Schizosaccharomyces pombe (fission yeast) by Clr4 is necessary for the stable association of RITS with heterochromatic loci, apparently through binding to the chromo domain of Chp1. This methylation event also recruits Swi6, which, together with other factors, mediates the spreading of various effectors, such as SHREC. SHREC might facilitate the proper positioning of nucleosomes to organize the higher-order chromatin structure that is essential for the diverse functions of heterochromatin, including transcriptional gene silencing. Swi6 also recruits an antisilencing protein, Epe1, that modulates heterochromatin to facilitate the transcription of repeat 9

elements, in addition to other functions. A dynamic balance between silencing and antisilencing activities determines the expression state of a locus within a heterochromatic domain[10].

Figure 3. Model showing RNAi-mediated heterochromatin assembly and silencing in S. pombe.

1.3 Dynamic nature of heterochromatin protein 1 binding

HP1 binding to protein was shown to be transient and dynamically exchanged from chromatin, with a half-life of 2.5 seconds as determined by fluorescence recovery after photobleaching (FRAP) on CHO cells expressing the GFP-HP1 isoforms[32]. In addition, most HP1 typically dissociates from chromosomes during mitosis[33], which was shown to result from histone H3 Ser 10 phosphorylation by Aurora b, and which is a part of a

“methyl/phos switch” mechanism that displaces HP1 and perhaps other proteins from mitotic heterochromatin[34]. Recently, it was shown that the H3K9 trimethyl mark, which was 10

considered a stable modification[35], can be removed from modified H3K9 by the enzyme

JHDM3A (jumonji c (JmjC)-domain containing histone demethylase 3A, also known as

JMJD2A), which abrogates recruitment of HP1 to heterochromatin, indicating a role of

JHDM3A in antagonizing methylated H3K9 nucleated events[36]. Furthermore, HP1a was shown to interact with the linker histone H1 in vivo and in vitro through their hinge and C- terminal domains, respectively [37, 38]. The phosphorylation of H1 by CDK2, which is required for efficient cell cycle progression, disrupts this interaction[38]. This raised the assumption that phosphorylation of H1 provides a signal for the disassembly of higher order chromatin structures during interphase, independent of histone H3-lysine 9 (H3-K9) methylation, by reducing the affinity of HP1a for heterochromatin[38].

1.4 HP1 and higher-order chromatin structure

Previous studies have shown that HP1a is not a chromatin fiber-bridging architectural protein[39]. It was shown that it facilitates the intramolecular folding of a nucleosomal array to generate compacted secondary chromatin structures without affecting intermolecular fiber- fiber interactions. And using nucleosomal arrays constructed with the histone variant

H2A.Z, it was proposed that HP1a is intrinsically attracted to condensed chromatin fibers, and upon binding, further local compaction occurs to generate distinct secondary structures with H2A.Z[39]. Other studies suggested an efficient binding of HP1 to chromatin can be achieved only when several binding sites are present within the chromatin substrate such as,

ACF1, which is a major chromatin assembly factor in Drosophila, and the histone 11

methyltransferase SU(VAR)3-9[26]. However, these studies lacked the basic requirement of uniformly methylated H3-K9 nucleosomal arrays that are characteristic to heterochromatic regions[4, 5].

1.5 Generation of in vitro heterochromatin model using native chemical ligation of histone proteins

A major problem with any previous in vitro study for heterochromatin models is the lack of a uniform histone H3 lysine 9 methylation. Although some studies attempted to generate such a uniformly methylated model using the Drosophila H3 methyltransferase

SU(VAR)3-9[26], but this enzymatic approach has some drawbacks represented in the possible partial activity of the methyltransferase and/or the non-uniformity of the methyl mark produced. In another study, methyllysine analogues (MLA) were used to generate such uniformly methylated arrays[40]. However, these arrays, in addition to its unnatural chemical structure, did not show significant specificity for Swi6 (fission yeast HP1 analogue) binding to the methylated arrays compared to the unmethylated ones.

In our laboratory, we made use of our previous experience of native chemical ligation[41] in generating uniformly modified histone proteins[42]. In this approach the tail fragments are synthesized by standard Fmoc-based solid phase peptide chemistry, where commercially available trimethylated lysine residue is directly incorporated into the desired peptide. Following chemical manipulations, the peptides are ligated to the recombinantly expressed carboxy-terminal fragment of H3, generating full length histone H3 with the 12

desired histone modification. Purified ligation product can routinely be generated in milligram quantities, which is sufficient for our biochemical and biophysical studies.

Modified histones can then be incorporated into more complicated chromatin structures using standard techniques. Using salt dialysis, histones can be incorporated into histone octamers[43], using rapid dilution and step-wise salt dialysis, histone octamers in conjunction with DNA templates can be incorporated into mononucleosomes and nucleosomal arrays respectively[44, 45]. Standard nucleosomal arrays have been generated using 601-177-12 templates[46].

Dissertation Organization

This dissertation is organized into 4 chapters. Chapter 1 covers the background literature related to the research project. Chapters 2 and 3 are the papers prepared for submission for publication. Chapter 2 is multiple interactions between heterochromatin protein 1 (HP1) and nucleosomes. Chapter 3 is HP1-mediated condensation of methylated chromatin. Chapter 5 is the conclusion of our study.

References 1. Crystal structure of the nucleosome core particle at 2.8 A resolution. Luger K, Mäder AW, Richmond RK, Sargent DF, Richmond TJ. 1997, Nature, pp. 251-260. 2. Conformational dynamics of the chromatin fiber in solution: determinants, mechanisms, and functions. Hansen, J C. 2002, Annual review of biophysics and biomolecular structure, pp. 361-392. 3. The core histone N termini function independently of linker histones during chromatin condensation. Carruthers LM, Hansen JC. 2000, The journal of biological chemistry, pp. 37285-37290. 13

4. Chromatin modifications and their function. Kouzarides, T. 2007, Cell, pp. 693-705. 5. The diverse functions of histone lysine methylation. Martin C, Zhang Y. 2005, Nature reviews. Molecular cell biology, pp. 838-849. 6. Translating the histone code. Jenuwein T, Allis CD. 2001, Science, pp. 1074-1080. 7. Relationship between histone H3 lysine 9 methylation, transcription repression, and heterochromatin protein 1 recruitment. Stewart MD, Li J, Wong J. 2005, Molecular and cellular biology, pp. 2525-2538. 8. The role of heterochromatin in centromere function. Pidoux AL, Allshire RC. 2005, Philosophical transactions of the royal society of London. Series B, Biological sciences, pp. 569-579. 9. Heterochromatin regulates cell type-specific long-range chromatin interactions essential for directed recombination. Jia S, Yamada T, Grewal SI. 2004, Cell, pp. 469-480. 10. Transcription and RNA interference in the formation of heterochromatin. Grewal SI, Elgin SC. 2007, Nature, pp. 399-406. 11. The protein encoded by the Drosophila position-effect variegation suppressor gene Su(var)3-9 combines domains of antagonistic regulators of homeotic gene complexes. Tschiersch B, Hofmann A, Krauss V, Dorn R, Korge G, Reuter. 1994, The EMBO Journal, pp. 3822-3831. 12. Regulation of chromatin structure by site-specific histone H3 methyltransferases. . Rea S, Eisenhaber F, O'Carroll D, Strahl BD, Sun ZW, Schmid M, Opravil S, Mechtler K, Ponting CP, Allis CD, Jenuwein T. 2000, Nature, pp. 593-599. 13. Functional mammalian homologues of the Drosophila PEV-modifier Su(var)3-9 encode centromere-associated proteins which complex with the heterochromatin component M31. . Aagaard L, Laible G, Selenko P, Schmid M, Dorn R, Schotta G, Kuhfittig S, Wolf A, Lebersorger A, Singh PB, Reuter G, Jenuwein T. 1999, pp. 1923-1938. 14. Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Bannister AJ, Zegerman P, Partridge JF, Miska EA, Thomas JO, Allshire RC, Kouzarides T. 2001, Nature, Vol. 410, pp. 120-124. 15. Role of histone H3 lysine 9 methylation in epigenetic control of heterochromatin assembly. Nakayama J, Rice JC, Strahl BD, Allis CD, Grewal SI. 2001, Science, Vol. 292, pp. 110-113. 16. Cowieson NP, Partridge JF, Allshire RC, McLaughlin PJ. Cowieson NP, Partridge JF, Allshire RC, McLaughlin PJ. 2000, Current Biology, Vol. 10, pp. 517--525. 17. Structural basis of HP1/PXVXL motif peptide interactions and HP1 localisation to heterochromatin. Thiru A, Nietlispach D, Mott HR, Okuwaki M, Lyon D, Nielsen PR, Hirshberg M, Verreault A, Murzina NV, Laue ED. 2004, The EMBO Journal, Vol. 23, pp. 489-499. 18. The hinge and chromo shadow domain impart distinct targeting of HP1-like proteins. Smothers JF, Henikoff S. 2001, Molecualr and cellular biology, Vol. 21, pp. 2555-2569. 19. Structure of the chromatin binding (chromo) domain from mouse modifier protein 1. Ball LJ, Murzina NV, Broadhurst RW, Raine AR, Archer SJ, Stott FJ, Murzin AG, Singh PB, Domaille PJ, Laue ED. 1997, The Embo Journal, Vol. 16, pp. 2473-2481. 20. Structure of HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Jacobs SA, Khorasanizadeh S. 2002, Science, Vol. 295, pp. 2080-2083. 14

21. The structure of mouse HP1 suggests a unique mode of single peptide recognition by the shadow chromo domain dimer. Brasher SV, Smith BO, Fogh RH, Nietlispach D, Thiru A, Nielsen PR, Broadhurst RW, Ball LJ, Murzina NV, Laue ED. 2000, The EMBO Journal, Vol. 19, pp. 1587-1597. 22. The Heterochromatin Protein 1 family. . Lomberk G, Wallrath L, Urrutia R. 2006, Genome Biology, Vol. 7, p. 228. 23. Evidence for the existence of an HP1-mediated subcode within the histone code. Lomberk G, Bensi D, Fernandez-Zapico ME, Urrutia R. 2006, Nature cell biology, Vol. 8, pp. 407-415. 24. Mutations in the heterochromatin protein 1 (HP1) hinge domain affect HP1 protein interactions and chromosomal distribution. Badugu R, Yoo Y, Singh PB, Kellum R. 2005, Chromosoma, Vol. 113, pp. 371-384. 25. HP1gamma associates with euchromatin and heterochromatin in mammalian nuclei and chromosomes. Minc E, Courvalin JC, Buendia B. 2000, Cytogenetics and cell genetics, Vol. 90, pp. 279-284. 26. HP1 binding to chromatin methylated at H3K9 is enhanced by auxiliary factors. Eskeland R, Eberharter A, Imhof A. 2007, Molecular and cellular biology, Vol. 27, pp. 453-465. 27. Specificity of the HP1 chromo domain for the methylated N-terminus of histone H3. Jacobs SA, Taverna SD, Zhang Y, Briggs SD, Li J, Eissenberg JC, Allis CD, Khorasanizadeh S. 2001, The EMBO Journal, Vol. 20, pp. 5232-5241. 28. The HP1 chromo shadow domain binds a consensus peptide pentamer. Smothers JF, Henikoff S. 2000, Current biology, Vol. 10, pp. 27-30. 29. Regulation of heterochromatic silencing and histone H3 lysine-9 methylation by RNAi. Volpe TA, Kidner C, Hall IM, Teng G, Grewal SI, Martienssen RA. 2002, Science, Vol. 297, pp. 1833-1837. 30. Heterochromatic silencing and HP1 localization in Drosophila are dependent on the RNAi machinery. Pal-Bhadra M, Leibovitch BA, Gandhi SG, Rao M, Bhadra U, Birchler JA, Elgin SC. 2004, Science, Vol. 303, pp. 669-672. 31. Dicer-deficient mouse embryonic stem cells are defective in differentiation and centromeric silencing. Kanellopoulou C, Muljo SA, Kung AL, Ganesan S, Drapkin R, Jenuwein T, Livingston DM, Rajewsky K. 2004, Genes & development, Vol. 19, pp. 489- 501. 32. Maintenance of stable heterochromatin domains by dynamic HP1 binding. Cheutin T, McNairn AJ, Jenuwein T, Gilbert DM, Singh PB, Misteli T. 2003, Science, Vol. 299, pp. 721-725. 33. Molecular behavior in living mitotic cells of human centromere heterochromatin protein HPLalpha ectopically expressed as a fusion to red fluorescent protein. Sugimoto K, Tasaka H, Dotsu M. Source. 2001, Cell structure and funtion, pp. 705-718. 34. Histone H3 serine 10 phosphorylation by Aurora B causes HP1 dissociation from heterochromatin. Hirota T, Lipp JJ, Toh BH, Peters JM. 2005, Nature, Vol. 438, p. 22. 35. Histone methylation: dynamic or static? . Bannister AJ, Schneider R, Kouzarides T. 2002, Cell, Vol. 109, pp. 801-806. 15

36. The transcriptional repressor JHDM3A demethylates trimethyl histone H3 lysine 9 and lysine 36. Klose RJ, Yamane K, Bae Y, Zhang D, Erdjument-Bromage H, Tempst P, Wong J, Zhang Y. 2006, Nature, Vol. 442, pp. 132-316. 37. Heterochromatin formation in mammalian cells: interaction between histones and HP1 proteins. Nielsen AL, Oulad-Abdelghani M, Ortiz JA, Remboutsika E, Chambon P, Losson R. 2001, Molecualr Cell, Vol. 7, pp. 729-739. 38. Phosphorylation of the linker histone H1 by CDK regulates its binding to HP1alpha. Hale TK, Contreras A, Morrison AJ, Herrera RE. 2006, Molecular cell, Vol. 22, pp. 693- 699. 39. H2A.Z alters the nucleosome surface to promote HP1alpha-mediated chromatin fiber folding. Fan JY, Rangasamy D, Luger K, Tremethick DJ. 2004, Molecular cell, Vol. 16, pp. 656-661. 40. Chromodomain-mediated oligomerization of HP1 suggests a nucleosome-bridging mechanism for heterochromatin assembly. Canzio D, Chang EY, Shankar S, Kuchenbecker KM, Simon MD, Madhani HD, Narlikar GJ, Al-Sady B. 2011, Molecular cell, Vol. 41, pp. 67-81. 41. Synthesis of proteins by native chemical ligation. Dawson PE, Muir TW, Clark-Lewis I, Kent SB. 1994, Science, Vol. 266, pp. 776-779. 42. Creating designer histones by native chemical ligation. Shogren-Knaak MA, Peterson CL. 2004, Methods in enzymology, Vol. 375, pp. 62-76. 43. Preparation of nucleosome core particle from recombinant histones. Luger K, Rechsteiner TJ, Richmond TJ. 1999, Methods in enzymology, Vol. 304, pp. 3-19. 44. Nucleosome interactions and stability in an ordered nucleosome array model system. Blacketer MJ, Feely SJ, Shogren-Knaak MA. 2010, Jornal of biological chemistry, Vol. 285, pp. 34597-34607. 45. Assembly of defined nucleosomal and chromatin arrays from pure components. Carruthers LM, Tse C, Walker KP 3rd, Hansen JC. 1999, Methods in enzymolgy, Vol. 304, pp. 19-35. 46. Chromatin fiber folding: requirement for the histone H4 N-terminal tail. Dorigo B, Schalch T, Bystricky K, Richmond TJ. 2003, Journal of molecular biology, Vol. 327, pp. 85-96.

16

CHAPTER 2. MULTIPLE INTERACTIONS BETWEEN HETEROCHROMATIN PROTEIN 1 (HP1) AND NUCLEOSOMES

A paper in preparation for submission

Abdelhamid M. Azzaz1*, Michael W. Vitalini2*, Melissa J. Blacketer1, Andrew S.

Thomas2, Diane E. Cryderman2, Adrian H. Elcock2, Michael A. Shogren-Knaak1 and

Lori L. Wallrath2

*equal contribution

From The Department of Biochemistry1, Biophysics and Molecular Biology

Iowa State University1, Ames, Iowa, 50011, and the Department of Biochemistry2,

University of Iowa2, Iowa City, Iowa 52242

To whom correspondence should be addressed: Lori L. Wallrath, Ph.D., Department of

Biochemistry, 3135 MERF, University of Iowa, Iowa City, IA 52242; E-mail: lori-

[email protected]

Keywords: chromatin, heterochromatin, HP1, nucleosomes

17

Abstract

The Heterochromatin Protein 1 (HP1) family is comprised of conserved nonhistone chromosomal proteins that package chromatin, regulate transcription and function in DNA repair. HP1 proteins contain an N-terminal chromo domain and C- terminal chromo shadow domain, separated by a hinge. HP1 dimerizes and interacts with histone H3 di- and tri-methylated at lysine 9. However, it is not known how HP1 interacts with nucleosomes to form heterochromatin. To address this question, a multi- disciplinary approach using in silico modeling, in vitro biochemistry and in vivo assays was undertaken. Molecular dynamic simulations and atomic resolution modeling revealed that human HP1Hs is a flexible protein that possesses the ability to bind both adjacent and non-adjacent methylated nucleosomes within an array. Models generated in silico were used to guide in vitro experiments showing that HP1Hs promotes chromatin compaction by bridging nucleosomes within an array and enhancing interactions between nucleosome arrays. These in vitro findings were supported by in vivo studies in which Drosophila HP1a was found to promote interactions between distant chromosome sites; such interactions were reduced in the presence of HP1a that lacked the hinge or the ability to dimerize. HP1a mutants that cannot interact with methylated histones or dimerize failed to support viability of an HP1a null.

Surprisingly, flies expressing only the hinge deletion were viable and possessed nucleosome arrays characteristic of heterochromatin. Taken together, dimerization and the flexibility of HP1 allow for multiple interactions with nucleosomes and the hinge is dispensable for heterochromatic nucleosome formation and viability in Drosophila. 18

Introduction

Heterochromatin Protein 1 (HP1) proteins comprise a group of evolutionarily conserved, non-histone chromosomal proteins that function in DNA replication, DNA repair, telomere capping, gene regulation and heterochromatin formation [1-5]. The human genome possesses three HP1 proteins, HP1Hsα, HP1Hsβ and HP1Hsγ, encoded by different genes [6-9].

They exhibit partial overlap on chromosomes and have non-redundant functions [8, 10]. One or more of the HP1 proteins are commonly mis-regulated in different types of cancer [11-13].

HP1 proteins contain an N-terminal chromatin modifier (chromo) domain (CD) that specifically interacts with di- and tri-methylated lysine 9 of the histone H3 N-terminal tail

(H3K9me2/3) [14-16]. A C-terminal chromo shadow domain (CSD) of similar sequence and structure mediates dimerization of HP1 proteins and interactions with partner proteins containing a PxVxL motif (‘x’ is any amino acid) [17-19]. A less conserved hinge region separates the CD and CSD. The hinge contains sites of post-translational modification and has been implicated in protein- and nucleic acid interactions [19-21].

Association of HP1 with chromatin is sufficient to induce an ordered arrangement of nucleosomes characteristic of heterochromatin and silence gene expression [22, 23]. The mechanism by which this is accomplished is unknown. Models to explain heterochromatin formation depict HP1 bridging adjacent nucleosomes [1, 23, 24]. Support for such a model comes from studies of the HP1-related protein Swi6 in S. pombe [25, 26]. However, data on interactions between metazoan HP1 proteins and nucleosome arrays are lacking.

A unique multi-disciplinary approach has been undertaken to discern HP1-nucleosome interactions in metazoans using human HP1Hsα and Drosophila HP1a as models. These 19

studies combine in silico, in vitro and in vivo methods. In silico atomic-scale modeling and molecular dynamics simulations suggest that HP1Hsα is flexible and participates in multiple interaction scenarios with nucleosome arrays. The in silico-generated models were tested by in vitro biochemical experiments, leading to the discovery that HP1Hsα promotes chromatin compaction by both intra- and inter-strand nucleosome interactions. Drosophila in vivo data demonstrated that HP1a promotes interactions between distant chromosome sites, supporting in vitro findings. In addition, our data demonstrate that methyl histone binding and dimerization are essential for viability; functions provided by the hinge are not. These data support a model whereby HP1 joins non-adjacent nucleosomes, expanding the repertoire of

HP1-nucleosome interactions, and demonstrate the hinge is dispensable for heterochromatic nucleosome organization and organismal viability.

Experimental procedures

Molecular dynamics simulations of HP1 conformational dynamics - Molecular dynamics simulations of the human HP1Hsα dimer were performed to assess the range of relative motion available to the CDs. Explicit-solvent MD simulations were carried out using

Gromacs 4 software (Hess, 2008) with a total of eight different simulations being performed: four independent simulations of human wild-type HP1Hs dimers and four independent simulations of the hinge HP1Hs dimer (in which amino acid residues P81-G116 were deleted).

Each of the independent simulations was performed with a different starting geometry of the CDs relative to the dimerized CSDs (Fig. 1S). In the MD simulations the HP1Hsα protein 20

was described by the Amber forcefield ff99SB [27]; the thousands of explicitly modeled water molecules were described by the SPC/E parameters [28]. To approximately mimic the physiological environment, 150 mM KCl was added to the system with a number of additional K+ ions included to neutralize the net charge of the HP1Hsα dimers (6 K+ were added for the wild-type HP1 simulations and 16 K+ for the hinge simulations); in the simulations, the K+ and Cl– ions were described by the SPC/E specific parameters developed by Joung & Cheatham [29].

Simulations of the wild type HP1Hs dimer and hinge dimer were carried out in truncated dodecahedron boxes with initial box diameters of 211.542 and 188.621 Å length, respectively: the wild type and hinge simulations contained approximately 668,000 and

476,000 atoms, respectively. All bonds were constrained with the LINCS algorithm [30] and a 2.5 fs time step was used. All non-bonded interactions were calculated directly up to a 10 Å cutoff, and all longer-range electrostatic interactions were calculated with the Particle Mesh

Ewald method [31]. Simulations were performed at 298 K and 1 atm using the Nose-Hoover thermostat [32, 33] and Parrinello-Rahman barostat [34], respectively. Each of the independent simulations was first energy minimized and then subjected to a 1 ns equilibration period in which the temperature was gradually raised to 298K. Following this point, production simulations were performed for 100 ns with coordinates saved every 1 ps for analysis. To assess the distance between the CDs within a dimer, we measured the distances between the centers of mass of the aromatic rings of the three residues (Y19, W40

& F43) that form each H3K9me binding site. Each 100 ns MD simulation required ~55,000 hours of cpu time on 256 cores using the TACC supercomputer Ranger. Therefore, the total computer time invested in the simulations amounts to ~440,000 hours. 21

In silico modeling of HP1-chromatin interactions - The exact arrangement of nucleosomes within the 30 nm chromatin fiber remains controversial [35, 36]. Although the high-resolution crystal structure of a tetranucleosome [37] provided a clear suggestion of the possible arrangement of nucleosomes within an oligonucleosome array, it did not contain fully connected DNA strands. Therefore, it was unclear how to use this model to build a complete, i.e. fully connected, atomic model of a 12-mer nucleosome array. To overcome this problem, we constructed a complete atomic model using a high-resolution mononucleosome crystal structure as a starting point [38]; pdb code 1KX5. In this structure,

146 nucleotides of DNA are wrapped around the histone octamer. Since the ‘612’ sequence of our 12-mer arrays has a repeat of 177 nucleotides, the length of the ‘linker’ DNA connecting successive nucleosomes must be 31 nucleotides. In the 1KX5 crystal structure the

DNA at each end is essentially B-form. This made it possible to append additional B-form

DNA to both ends to construct a repeating mononucleosomal unit, which in turn made it possible to construct a polymeric model. The procedure used to build the array is outlined in

Fig. 2S. Satisfyingly, the resulting structure of the 12-mer nucleosome array is visually consistent with the tetranucleosome crystal structure [37], but has the advantage of being a complete and fully-connected atomic model.

To determine how HP1Hs might bind chromatin, we constructed different models in which HP1Hs dimers were docked to the 12-mer nucleosome array. Three models were constructed in which: (a) a HP1Hs dimer was bound to adjacent nucleosomes, (b) a HP1Hs dimer was bound to two every other nucleosome and (c) a HP1Hs dimer was bound to every third nucleosome. To model HP1Hs, we took as our starting point the atomic structure of the 22

human HP1Hs CD bound to a N-terminal fragment of the H3 histone tail (H3K9me peptide)[36]. We manually positioned two copies of the CD close to where the H3 N- terminal tails exit from the nucleosome core. With the CDs positioned, the atomic structure of the mouse HP1 CSD dimer [17] was added and positioned in a way that made it feasible for the unstructured hinge domain to complete the connection between the C-terminus of a

CD with the N-terminus of a CSD in each HP1Hs monomer. The feasibility of performing this connection was explicitly demonstrated by using the program Loopy [39] to build an atomic model of the hinge domain sequence consistent with the spatial restrictions imposed by placement of the CD and the CSD. The final step of the modeling process was to verify that the modeled HP1Hsdimers were also capable of binding to the nucleosome array. This was again explicitly demonstrated by using Loopy to build atomic models of the histone H3 tails that correctly connected the C-terminal residue of the H3K9me N-terminal peptide bound in the co-crystal structure of the CD (S10) with the most N-terminal residue of the H3 tail (K36) resolved in the nucleosome core crystal structure. Additional modeling of the unresolved N- and C-terminal tails of HP1Hs was accomplished using in-house software and final adjustments of the residue identities in the CSD domain so that they were consistent with the human HP1Hs CSD sequence was accomplished with the homology-modeling program DeepView [40]. Once a complete structure of a HP1Hs dimer bound to the nucleosome array had been constructed, additional HP1Hs dimers were added by applying the symmetry transformations used to generate the original 12-mer nucleosome array. We visually verified that the additional copies of HP1Hsα constructed in this way did not engage in unfavorable steric clashes with each other. 23

Generation and characterization of HP1Hs protein - BL21 (DE3) pLys S cells containing either wild-type human 6His-HP1 plasmid (a gift from Professor Christian

Muchardt, Pasteur Institute in Paris), W71A 6His-hHP1 plasmid, or I165E 6His-hHP1 plasmid were grown to an OD600 of 0.5-0.6 and then induced with IPTG (0.2 mM final concentration) for 4 hours. Pelleted cells were resuspended in 20 ml lysis buffer (50 mM

NaH2PO4 pH8, 300 mM NaCl, 10 mM Imidazole, 1 mM 2-mercaptoethanol, 10% glycerol,

1M PMSF, 10 g/ml leupeptin, 10 g/l aprotinin and 50 g/ml DNase I) per 500 ml of culture media, flash frozen, and stored at -80C. Cells were lysed cold using a French press, and the lysate was mixed with Ni-NTA agarose resin (Qiagen). The resin was washed twice

(lysis buffer + 20 mM imidazole), and then the protein was eluted from the resin (lysis buffer

+ 250 mM imidazole). Aggregated HP1Hs was removed from the eluate by gel filtration chromatography (Sephacryl S-300 resin with lysis buffer as the mobile phase). The desired

HP1Hs fractions were pooled and concentrated to about 30 M concentration, as determined by UV absorbance.

The extent of dimerization was determined by sedimentation equilibrium analysis. The protein was dialyzed in a buffer containing 100 mM NaCl, 50 mM NaH2PO4 pH 8, 0.2 mM

EDTA, 0.05 mM TCEP. 120 ml of protein samples were prepared with OD280 of 0.3, 0.5 and

0.7, centrifuged at 30000 rpm in a Beckman XL-A ProteomeLab ultracentrifuge and the data analyzed using the Ultrascan software.

The binding of HP1Hs to H3K9me3 peptide (fragments 2-23) was analyzed using fluorescence anisotropy (Fig. 3S). H3 peptide (1-23) with K9 trimethylation and an amino- terminal fluorescein label was generated by standard Fmoc-based solid-phase peptide 24

synthesis (University of Wisconsin Biotechnology Center), where fluorescein-5-EX succinimidyl ester, was coupled to the amino-terminus prior to peptide to resin cleavage/side- chain protection and purification according to standard peptide synthesis protocols. For fluorescence anisotropy experiments, HP1 was dialyzed in a buffer containing 50 mM K2PO4 pH 8, 25 mM NaCl and 2 mM DTT as reported in [41]. 100 nM fluorescein peptide were titrated with increasing concentrations of HP1, ranging from 0.1 to 100 M, left at room temperature for 30 minutes and the anisotropy vales determined using a Varian Fluorescence spectrophotometer fitted with a polarizer.

In vitro chromatin assembly - Recombinant, unmodified, Xenopus laevis H2A, H2B, H3, and H4 histones were prepared according to standard protocols [42]. H3 histone uniformly trimethylated at lysine 9 was generated by native chemical ligation of a synthesized H3 1-23 peptide (University of Wisconsin Biotechnology Center) and a recombinant H3 C-terminal fragment, according to previously described methods [43]. Methylated and unmethylated histone octamers were assembled according to standard techniques [42]. 601-177-12 and

601-177-1 DNA templates were liberated from a 601-177-12-containing plasmid using

EcoRV and ScaI, respectively. Carrier DNA was prepared by PCR amplification as previously described [42]. Following phenol-chloroform extraction, DNA fragments were further purified by preparative gel electrophoresis, electroelution, butanol extraction, and ethanol precipitation. 601-177-1 mononucleosomes were prepared by rapid dilution deposition with molar ratios of octamer to DNA between 0.8 and 1.2 [42]. Assembled mononucleosomes were dialyzed against 2.5 mM NaCl, 10 mM Tris, pH 8.0, 0.25 mM

EDTA with three buffer exchanges and concentrated. Saturation was determined by native 25

gel analysis. 601-177-12 arrays were assembled by salt step dialysis as previously described

[42], with ratios of template to carrier to octamer of 1.0:0.3:1.1-1.3.

EMSA Analysis - HP1Hs was dialyzed in 50 mM HEPES pH 8, 150 mM NaCl, 1 mM

EDTA and 10% glycerol. Nucleosome arrays or mononucleosomes were mixed with HP1Hs to a final volume of 20 l containing 50 mM NaCl, 23.33 mM HEPES pH 8, 0.067% TCEP, and 0.33 mM EDTA. The HP1Hs arrays or mononucleosome mixture was mixed gently on the vortexer for 30 minutes, and then applied on 0.5% agarose (type IV) midi-gel made in 50 mM HEPES pH 8 and 150 mM NaCl buffer, and run in the same buffer at 70 volts and 0.4 amp for 2 hours. Gels were visualized with either ethidium bromide or Sybr Gold.

Differential sedimentation analyses- Reversible self-association of nucleosomal arrays in the presence of HP1Hs was characterized using a differential sedimentation assay previously described [44]. Briefly, HP1 in HP1 buffer (150mM NaCl, 50mM HEPES, 0.2mM EDTA, pH=8.0) was diluted to a stock concentration of 13.2µM and final buffer concentrations of

150mM NaCl, 12.5mM HEPES, and .05mM EDTA. Just prior to mixing with arrays,

Hs HP1 was added to varying concentrations of MgCl2 in array buffer (2.5mM NaCl, 10mM

Hs Tris pH8.0, 0.25mM EDTA, 0.1mM TCEP) to create 2X MgCl2 and HP1 solutions, which were then mixed in equal volume with nucleosomal arrays (9.1 ng/µl final concentration template DNA). After incubation for 15 minutes at room temperature, samples were centrifuged at 14,000g for 10 minutes. The percentage of arrays remaining in the supernatant was determined by comparing the OD260 of each sample with the OD260 of a sample with no

MgCl2 added. 26

Drosophila cultures – Drosophila stocks were maintained at room temperature on standard sucrose/cornmeal medium. The generation of the LacI-HP1a and lacO repeat transgenic stocks were described previously [23]. For the genetic complementation studies, genetic crosses were set up to introduce a single copy of a given transgene into a trans- heterozygous null background and the resulting larvae were heat shocked daily at 37° C for 1 hour throughout development to maintain transgenic expression of the LacI-HP1a protein.

Rescued trans-heterozygous adult flies containing two null alleles of Su(var)2-5 were scored based on the wing phenotype for loss of the CyO balancer chromosome.

Polytene chromosome staining – Third instar larvae were subjected to a 1 hour heat- shock at 37° C. After a one-hour recovery period at room temperature, salivary glands were dissected, fixed, squashed and stained with mouse antibodies to LacI (Upstate Biotechnology no. 05-501, clone 9A5, 1:300 dilution) followed by donkey-anti-mouse Alexa Fluor 488

(Invitrogen, 1:300). Images were obtained using a Biorad confocal miscroscope.

MNase digestion of chromatin – Nuclear preparations, MNase treatments and Southern procedures were performed as described previously (23). Adult ‘rescued’ flies were obtained as described above, control flies (y,w) underwent the same heat-shock regimen as transgene- containing stocks. Flies were harvested by snap-freezing in liquid N2 1-3 days post-eclosion and ~6 hours post heat-shock, and stored at -80° C. Between 0.5 and 0.75 ml of adults

(measured in a microfuge tube) were used for each MNase experiment.

27

RESULTS

In silico modeling of HP1 conformational dynamics. HP1 proteins associate with heterochromatin through an interaction between the CD and H3K9me2/3 [14]. However, the geometry in which this interaction occurs is unknown. It is possible that each HP1 dimer interacts with both histone H3 tails on a single nucleosome, with one histone H3 tail from each of two adjacent nucleosomes, or with one histone H3 tail from each of two non-adjacent nucleosomes. As a means of distinguishing among these possibilities, we determined the range of relative motion between the two CDs in a dimer. A limited range of relative motion, for example, might severely restrict the possible binding geometries of HP1 that would need to be considered. To this end, we performed a series of computationally intensive in silico molecular dynamics (MD) simulations of complete atomic models of both wild type HP1Hsα and a hinge mutant in which residues 81–116 have been removed (Fig. 1). Four independent simulations of each construct were performed with the initial positions of the two CDs being altered in each in order to explore a wide range of initial relative orientations.

The distance between the H3K9me binding sites on the two CDs was then followed over the course of each simulation and plotted as a histogram (Fig. 1B and C). 28

29

Figure 1. The hinge region imparts flexibility to HP1Hsa. An atomic-scale model of an HP1Hsα homodimer was generated using the known structures for the CD (blue) and the CSD (purple) (A). Regions of the protein for which no structure has been determined were modeled de novo (grey, see Experimental Procedures). The model was used in molecular dynamics simulations to assess the range of motion for the two CDs over 100 ns (B). At the start of each simulation the two CDs were positioned at different distances from one another (Supplemental Figure 1). The probability of each distance sampled was plotted and color coded with respect to the starting position: blue, 51 Å; red, 108 Å; green, 138 Å; purple, 72 Å for wild type (B) and blue, 125 Å; red, 58 Å; green, 98 Å; purple, 34 Å for hinge (C).

The histograms obtained from each of the four independent simulations of the wild type

HP1Hs show a range of different behaviors. In simulation ‘1’, for example, the two CDs begin in close proximity (Fig. 1S) and remain in contact throughout the entire 100 ns simulation (in Fig. 1B, blue line). By way of contrast, in simulation ‘3’, the two CDs explore a very wide range of relative orientations: the distance between the two H3K9me binding sites varies from 70 to 155 Å over the course of 100 ns (Fig. 1B, green line). The simulations demonstrate that wild type HP1Hs has a very wide range of conformational capabilities.

Surprisingly, deletion of the hinge did not alter this flexibility (Fig. 1C). The range of distances sampled between the two H3K9me binding sites is somewhat narrower, varying from 40 to 120 Å (compared with 35 to 160 Å for the wild type HP1Hs) consistent with the loss of the 36 residues connecting each CD to the CSD. Nevertheless, the four independent simulations again show widely differing behaviors, indicating that the hinge HP1Hs is likely to have a high degree of conformational flexibility similar to that of wild type.

In silico modeling of HP1-nucleosome interactions. To more directly assess the possible geometry in which HP1Hs dimers might bind to nucleosomal structures, we constructed complete, albeit putative, atomic models for the interaction of HP1Hs dimers with (a) a mononucleosome, (b) a dinucleosome, and (c) a model of a 12-mer nucleosome array. We 30

determined that a single HP1Hs dimer could readily bind to both H3K9me binding sites within a mononucleosome (Fig. 2A) and could successfully bridge between the two nucleosomes of a dinucleosome model (Fig. 2B). Importantly, the distances between the

H3K9me binding sites in these two models (~53 Å and 78 Å, respectively) are well within the range of possibilities for both the wild type and the hinge HP1Hs seen in our MD simulations (Fig. 1).

Figure 2. Atomic resolution models of an HP1Hs dimer bound to a mono- and di- nucleosome. An all atom atomic resolution model was generated for HP1Hs (see Experimental Procedures) and docked onto the histone H3 tails emerging from a modeled nucleosome (see Experimental Procedures) (A) or a modeled di-nucleosome (B). CD, blue; CSD, purple; nucleosome, hinge region and unstructured regions of HP1, gray; DNA green and yellow

To address how HP1Hs might bind to the chromatin fiber, we constructed atomic models of HP1Hs bound to a nucleosome array (Fig. 3). Based on attempts to manually construct complete atomic models of the HP1Hs-oligonucleosome complex that are free of any steric clashes, three basic binding configurations appear possible. First, the HP1Hs dimer might bridge adjacent nucleosomes (Fig 3A). In such a binding orientation the two H3K9me binding sites on the HP1Hs molecule would be separated by ~65 Å. Although this distance 31

must be considered only a rough estimate – the flexibility inherent in both the HP1Hs hinge domain and the H3 N-terminal tail allow for a great deal of latitude in the manual positioning of the HP1Hs dimers – this distance is well within the range of possible conformations sampled by the HP1Hs dimer in MD simulations (Fig. 1). Second, the HP1Hs dimer might bridge every other nucleosome. In order to bind in this manner we find that the two H3K9me binding sites on the HP1 molecule would need to be separated by ~62 Å, also possible according to the MD simulations (Fig. 1) Third, the HP1 dimer might bridge every fourth nucleosome; to achieve this the H3K9me binding sites would need to be separated by ~51 Å, also feasible according to the MD simulations. Taken together, the MD simulations and modeling strongly suggest that all the conformations tested are plausible from a structural perspective. 32

33

Figure 3. Atomic resolution model of HP1Hs bound to a nucleosome array. A full atom model of an HP1Hs dimer was docked onto the histone H3 tail emanating from adjacent nucleosomes (A), every other nucleosome (B) or every third nucleosome (C) within a 12- nucleosome array. A schematic representation is shown below each model.

In vitro EMSA tests of predicted HP1-nucleosome interactions. The in silico modeling predicted that HP1Hs dimers can associate with mononucleosomes. In vitro EMSAs were performed using purified HP1Hsα to test for that interaction. Mononucleosomes consisting of purified histones H2A, H2B, H3, H4 and one copy of the 177 bp 601 nucleosome positioning sequence were incubated with purified HP1Hs. Addition of 0.2 M HP1Hsα to mononucleosomes containing histone H3K9me, resulted in the formation of a slower- migrating species, with a complete shift to the slower band occurring by 6 M, suggesting

HP1Hsα –mononucleosome association (Fig. 4A). This interaction was H3K9me3-dependent, as the slower-migrating band was not observed when up to 6 M HP1Hsα was added to unmethylated mononucleosomes (Fig. 4A). Incubation of a mutant version of HP1Hsα

(W17A) that disrupts the interaction with histone H3K9me3 [45] resulted in a weak band at the highest concentration of HP1Hsα used (6 M), compared to results obtained with wild type HP1Hsα (Fig. 4A). Interestingly, a mutant form of HP1Hsα that cannot dimerize (I195E)

[17] also failed to produce a slower migrating band in the presence of methylated mononucleosomes. These data indicate that dimerization is required for the HP1- mononucleosome interaction.

34

Figure 4. Associations of HP1Hsα with mononucleosomes or 12-nucleosome arrays in vitro. EMSAs were performed using purified wild type HP1Hsa, a mutant HP1Hsa that lacked H3K9me2/3 interaction (CD mutant, W71A HP1) and a CSD dimerization (CSD mutant, I195E HP1). The HP1Hsa proteins were incubated with mono-nucleosomes (A) or 12- nucleosome array (B) containing unmodified or H3K9 histones. Differential sedimentation analysis was performed using the HP1Hsa proteins and 12-mer methylated and unmethylated arrays. The percent of material in the supernatant is plotted as a function of MgCl2 concentration.

EMSA analysis was performed using 12-nucleosome arrays as binding substrates. Similar to the results obtained with mononucleosomes, wild type HP1Hsα association with the nucleosome arrays was dependent on H3K9me (Fig. 4B). Addition of ~0.1 M HP1Hsα was sufficient to completely shift the arrays to the slower migrating species, suggesting a higher affinity of HP1Hsα for the nucleosome arrays as compared to the mononucleosomes (compare

Fig. 4A and 4B). Interestingly, binding was also apparent in the absence of histone 35

H3K9me3, as a faint band of the slower-migrating species was present in experiments using unmethylated nucleosomes (Fig. 4B). Likewise, incubation with 2M or higher concentrations W71A resulted in a shift to the slower migrating species. The dimerization mutant, I165E, displayed a reduced ability to shift the nucleosome arrays. Thus, HP1Hsα possesses the ability to interact with unmethylated nucleosomes, but shows a clear preference for histone H3K9me3 nucleosomes, consistent with prior publications [14, 45].

Differential sedimentation analysis of HP1-nucleosome interactions. An in vitro approach was used to determine whether HP1Hsα interactions are confined to a single nucleosome array, or cross-link independent arrays. We hypothesized that addition of HP1Hsα to nucleosome arrays would facilitate intra- and/or inter-array interactions, resulting in higher levels of sedimentation [46] at lower concentrations of MgCl2 as compared to reactions lacking HP1Hsα. Incubation of unmethylated 12-nucleosome arrays with 1 m HP1Hsα had no effect on salt-dependent condensation (Fig. 4C). However, a dose-dependent effect of HP1Hsα was observed when incubated with 12-nucleosome arrays containing H3K9me; increasing amounts of HP1 increased the amount of condensation observed for a given concentration of

Hsα MgCl2 (Fig. 4C). The ability of HP1 to enhance salt-dependent condensation required dimerization, as the I165E mutant had no effect (Fig. 4C), consistent with the reduced ability of this form of the protein to bind the nucleosome arrays (Fig. 4C). These results suggest that in vitro, dimerized HP1Hsα interacts with non-adjacent nucleosomes to promote inter-array interactions.

In vivo analysis of HP1-nucleosome interactions. Both the in silico and in vitro data suggested that HP1Hsα facilitates interactions between non-adjacent nucleosomes to induce a condensed chromatin structure. To test for this property in vivo, we performed studies on 36

Drosophila HP1a. HP1a shares amino acid sequence identity with HP1Hsα [47]. In addition,

HP1Hsα can rescue the lethality associated with HP1a nulls, demonstrating conserved functions [8]. Drosophila offers large salivary gland polytene chromosomes to visualize

HP1a localization and in vivo functional tests.

Previously we developed a system to tether a LacI-HP1a fusion to genomic sites possessing integrated copies of the lacO repeat [23]. Tethered HP1a caused the formation of a less accessible chromatin structure, ‘chromosome loops’, and gene silencing [23]. Here, we tethered HP1a and assayed for chromosome loops after fixation, as evidence of non- adjacent nucleosome interactions (Fig. 5A). Over 36% of the chromosomes bound by HP1a exhibited loops, as compared to 8.9% control LacI-GFP bound chromosomes (Fig. 5B).

Interestingly, the HP1a dimerization mutant I191E (analogous to HP1Hsα I195E) displayed a reduced level of self-association, 15.9%, as compared to LacI-HP1a (Fig. 5A). This decreased level of association is consistent with the decreased HP1Hs-nucleosome association observed for the dimerization mutant in vitro.

A mutant form of HP1 lacking the hinge region (LacI-Δhinge) was also tested for the ability to induce intra-chromosomal associations. If flexibility provided by the hinge is important for association with non-adjacent nucleosomes, then LacI-Δhinge should show reduced levels of chromosome self-association as compared to LacI-HP1. Indeed, targeted binding of LacI-Δhinge induced intra-chromosomal associations at an intermediate level

(24.5%) as compared to the LacI-HP1a and LacI-I191E proteins (Fig. 5B). These results suggest that HP1a is capable of associating with non-adjacent nucleosomes in vivo; this association is heavily dependent on dimerization, but may not require the hinge region. 37

Figure 5. Targeted binding of HP1a induces intra-chromosomal associations. Polytene chromosomes from larvae expressing LacI-HP1 fusion proteins, or a LacI-GFP control, and possessing lacO repeats were stained with anti-LacI antibodies and scored for self- association (A). The total number and percentage of chromosomes that showed an extended or associated phenotype for each genetic background are shown (B). The value for each construct was significantly different from that of each other construct at p<0.001 (chi square analysis). 38

HP1 dimerization and interaction with H3K9me2/3, is essential for viability, the hinge region is not. Both the in vitro and in vivo data suggested that dimerization of HP1 influences

HP1-nucleosome interactions. In order to determine whether dimerization of HP1a is essential for viability, genetic complementation assays were performed. The Δhinge form of

HP1a was also tested for viability to determine whether the flexibility imparted to HP1 by the hinge is essential to the function of the protein. Flies trans-heterozygous for null or loss of function alleles of Su(var)2-5 (the Drosophila gene encoding HP1) die at the third larval instar stage. Consistent with previous studies, the transgene encoding LacI-HP1a (wtHP1) was capable of rescuing transheterozygous animals to adulthood (Table 1). A transgene encoding a mutant form of HP1 containing the amino acid substitution V26M (analogous to the V22M substitution in HP1Hsα), which is defective in H3K9me3 binding, failed to rescue animals to adulthood, consistent with the defect in nucleosome association observed in vitro and previous in vivo results [Su(var)2-502 is an EMS-induced allele that encodes the V26M substitution and that is lethal in combination with null alleles of Su(var)2-5]. In addition, the

I191E dimerization mutant also failed to rescue, suggesting that dimerization is necessary for the essential functions of HP1 in vivo. Surprisingly, the transgene encoding LacI-Δhinge

(Δhinge) was capable of rescuing flies to adulthood, albeit at a lower rate than the wtHP1 transgene (18% of expected class compared to 42% for wtHP1, Table 1). All fusion proteins were expressed at similar levels (<1.8 fold difference) as measured by western blot (Fig. S4).

Together, these data suggest that the CD and CSD both perform functions essential for the ability of HP1 to support viability in Drosophila (H3K9me3 binding and dimerization, respectively). In contrast, the hinge region does not appear to be essential for viability. 39

Parents Progeny % of Expected Straight- Class Female Male Curly-Wings Wings (rescued) wtHP1 Su(var)2-5 Su(var)2-5 ; 446 64 42 wtHP1 CyO CyO Su(var)2-5 V26M Su(var)2-5 ; 426 0 0 CyO V26M CyO

I191E, Su(var)2-5 Su(var)2-5 413 0 0 CyO CyO Δhinge Su(var)2-5 Su(var)2-5 ; 302 19 18 Δhinge CyO CyO

Table 1. Genetic complementation analysis

Discussion

Binding modes. Our biophysical experiments show that HP1 protein can mediate methylation-dependent binding with individual nucleosomes, as well as with arrays of nucleosomes (both within and between such arrays). To what extent each of these bound states occurs in vivo is unclear. Based on the relative concentrations of HP1 necessary to induce these bound states, the greatest affinity state appears to be for binding of arrays with multiple nucleosomes, potentially making it the most favorable state. However, these states are not necessarily mutually exclusive. For example, it is possible that at relatively high densities of bound HP1, some HP1 molecules mediate binding within an array, while others mediate interactions between arrays. Additionally, different nuclear chromatin contexts may favor one interaction over another. For example, if there are nucleosome regions with few or only a single methylated nucleosome, HP1-mediated binding may only 40

be possible within a nucleosome. Moreover, other heterochromatin-associated protein and

RNA factors may stabilize such a binding state.

Structures of bound states. HP1-mediated interactions with individual nucleosomes require both H3K9 methylation and dimerization of HP1, suggesting that the two chromodomains of a HP1 dimer could bridge the two K9 methylated H3 histone tails within a nucleus. Indeed, our in silico dynamic and modeling studies of HP1 indicate that the hinge domain provides sufficient flexibility and length to adopt such a state. Because of the apparent increased affinity for HP1 with methylated nucleosomal arrays, as well as the requirement for HP1 dimerization, the simplest model for such an interaction is that the

HP1 dimer bridges different nucleosomes within a strand. Again, our in silico dynamic and modeling studies show that such bridging is feasible. However, these studies indicate that multiple types of nucleosome bridging are possible, and further in vitro studies will be necessary to better characterize the nature of the bridged state.

Interchromatin contacts. Visualization of heterochromatin in vivo shows that it is highly dark stained, potentially due to a highly condensed nature. The ability of HP1 to mediate binding within methylated nucleosomal arrays might account for this condensation.

However, this condensation may also occur through strand-to-strand interactions between

HP1-associated chromatin. Our in vitro and in vivo studies indicate that HP1 and its fly homolog can facilitate such long-range contacts between chromatin, and may underscore an important structural feature of in vivo heterochromatin.

Importance of HP1 dimerization. In vitro, HP1 dimerization is very tight, and in our computational studies this dimerization is assumed. In vitro validation of the importance 41

of this dimerization in nucleosome binding comes from the fact that all of the forms of

HP1-mediated nucleosome interactions we observe are strongly disrupted by the loss of

HP1 dimerization. The importance of this dimerization is further underscored by our in vivo results. Loss of HP1 dimerization in a mutant fly homolog of HP1introduced into a background of wild-type HP1 results in severely reduced localization of the mutant HP1 isoform to induced heterochromatic sites. Moreover, the lethality that results from the total loss of the fly HP1 isoform cannot be rescued by HP1 proteins that do not dimerize.

Importance of the HP1 hinge domain. While a flexible hinge domain is the basis of our in silico modeling of potential HP1-mediated binding structures, our in vivo studies indicate that, while important, this hinge region may not be essential. Deletion of the hinge region of fly HP1 in the background of wild-type HP1 result in reduced localization of the mutant isoform. However, some localization occurs above background levels. Similarly, some degree of rescue of the lethality occurs in strains in which the hinge deletion isoform complements the wild-type delete. These results suggest that the hinge is not essential. One possibility is that dimers of HP1 lacking the hinge domain can form the same or similar types of nucleosome bound species as wild-type HP1 dimers. Alternatively, because heterodimerization of HP1 isoforms is possible, it is possible that a heterodimer of fly HP1 with another HP1 isoform that also can localize to heterochromatin, such as the fly homolog of HP1, might be able to bind methylated nucleosome, albeit in a less efficient manner.

HP1 binding in other species. Recently, Canzio and coworkers have shown that an

HP1 homolog in fission yeast, Swi6, can be bound to nucleosomes and within nucleosomal arrays containing a mimic of H3 K9 methylation [48]. While in agreement with some aspects 42

our studies, there are also a number of differences. In their system, they observe mononucleosome binding that is tighter than we observed and that is dimerization independent. Further, in all of their studies, they observe a relatively high level of binding to non-methylated nucleosomes. In addition to potential differences in the way the chromodomain binds native trimethylated lysines, versus a trimethylation mimic, the observed differences could be due to differences in the nature of the HP1 molecules. The hinge domain of HP1 is believed to contain the non-specific DNA binding activity, and this domain is not highly conserved between HP1 isoforms within and between species. Thus, these differences could be the basis of the differences in affinity and specificity for methylated mononucleosomes. Additionally, there appears to be difference in the nature of the chromoshadow domain that mediates dimerization. The deletion of Swi6 cannot be complemented by mouse HP1. Moreover, a swap containing the mouse CD can rescue the Swi6 delete, but a CSD swap cannot. Thus, how CSD function in heterochromatin structure may also be different [49].

References

1. Bannister, A.J., et al., Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature, 2001. 410(6824): p. 120-4. 2. Cheutin, T., et al., Maintenance of stable heterochromatin domains by dynamic HP1 binding. Science, 2003. 299(5607): p. 721-5. 3. Danzer, J.R. and L.L. Wallrath, Mechanisms of HP1-mediated gene silencing in Drosophila. Development, 2004. 131(15): p. 3571-80. 4. Singh, P.B., HP1 proteins--what is the essential interaction? Genetika, 2010. 46(10): p. 1424-9. 5. Zeng, W., A.R. Ball, Jr., and K. Yokomori, HP1: heterochromatin binding proteins working the genome. Epigenetics, 2010. 5(4): p. 287-92. 43

6. James, T.C., et al., Distribution patterns of HP1, a heterochromatin-associated nonhistone chromosomal protein of Drosophila. Eur J Cell Biol, 1989. 50(1): p. 170- 80. 7. Sharma, G.G., et al., Human heterochromatin protein 1 isoforms HP1(Hsalpha) and HP1(Hsbeta) interfere with hTERT-telomere interactions and correlate with changes in cell growth and response to ionizing radiation. Mol Cell Biol, 2003. 23(22): p. 8363-76. 8. Ma, J., et al., Expression and functional analysis of three isoforms of human heterochromatin-associated protein HP1 in Drosophila. Chromosoma, 2001. 109(8): p. 536-44. 9. Kato, M., et al., Functional domain analysis of human HP1 isoforms in Drosophila. Cell Struct Funct, 2007. 32(1): p. 57-67. 10. Szakal, B., et al., Cloning, characterization and localization of Chinese hamster HP1 isoforms. Chromosome Res, 2004. 12(5): p. 483-93. 11. Kirschmann, D.A., et al., Down-regulation of HP1Hsalpha expression is associated with the metastatic phenotype in breast cancer. Cancer Res, 2000. 60(13): p. 3359- 63. 12. Norwood, L.E., et al., A requirement for dimerization of HP1Hsalpha in suppression of breast cancer invasion. J Biol Chem, 2006. 281(27): p. 18668-76. 13. Dialynas, G.K., M.W. Vitalini, and L.L. Wallrath, Linking Heterochromatin Protein 1 (HP1) to cancer progression. Mutat Res, 2008. 647(1-2): p. 13-20. 14. Jacobs, S.A. and S. Khorasanizadeh, Structure of HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Science, 2002. 295(5562): p. 2080-3. 15. Nielsen, P.R., et al., Structure of the HP1 chromodomain bound to histone H3 methylated at lysine 9. Nature, 2002. 416(6876): p. 103-7. 16. Fischle, W., et al., Molecular basis for the discrimination of repressive methyl-lysine marks in histone H3 by Polycomb and HP1 chromodomains. Genes Dev, 2003. 17(15): p. 1870-81. 17. Thiru, A., et al., Structural basis of HP1/PXVXL motif peptide interactions and HP1 localisation to heterochromatin. EMBO J, 2004. 23(3): p. 489-99. 18. Cowieson, N.P., et al., Dimerisation of a chromo shadow domain and distinctions from the chromodomain as revealed by structural analysis. Curr Biol, 2000. 10(9): p. 517-25. 19. Smothers, J.F. and S. Henikoff, The hinge and chromo shadow domain impart distinct targeting of HP1-like proteins. Mol Cell Biol, 2001. 21(7): p. 2555-69. 20. Badugu, R., et al., Mutations in the heterochromatin protein 1 (HP1) hinge domain affect HP1 protein interactions and chromosomal distribution. Chromosoma, 2005. 113(7): p. 370-84. 21. Meehan, R.R., C.F. Kao, and S. Pennings, HP1 binding to native chromatin in vitro is determined by the hinge region and not by the chromodomain. Embo J, 2003. 22(12): p. 3164-74. 22. Lechner, M.S., et al., Molecular determinants for targeting heterochromatin protein 1-mediated gene silencing: direct chromoshadow domain-KAP-1 corepressor interaction is essential. Mol Cell Biol, 2000. 20(17): p. 6449-65. 44

23. Li, Y., et al., Effects of tethering HP1 to euchromatic regions of the Drosophila genome. Development, 2003. 130(9): p. 1817-24. 24. Nakayama, J., et al., Role of histone H3 lysine 9 methylation in epigenetic control of heterochromatin assembly. Science, 2001. 292(5514): p. 110-3. 25. Aygun, O. and S.I. Grewal, Assembly and functions of heterochromatin in the fission yeast genome. Cold Spring Harb Symp Quant Biol, 2010. 75: p. 259-67. 26. Grewal, S.I., RNAi-dependent formation of heterochromatin and its diverse functions. Curr Opin Genet Dev, 2010. 20(2): p. 134-41. 27. Song, K., et al., Computational analysis of the mode of binding of 8-oxoguanine to formamidopyrimidine-DNA glycosylase. Biochemistry, 2006. 45(36): p. 10886-94. 28. van Gunsteren, W.F. and H.J. Berendsen, Thermodynamic cycle integration by computer simulation as a tool for obtaining free energy differences in molecular chemistry. J Comput Aided Mol Des, 1987. 1(2): p. 171-6. 29. Joung, I.S. and T.E. Cheatham, 3rd, Determination of alkali and halide monovalent ion parameters for use in explicitly solvated biomolecular simulations. J Phys Chem B, 2008. 112(30): p. 9020-41. 30. Hess, C.P., et al., A software system for interactive MR signal processing. Magn Reson Imaging, 1997. 15(1): p. 127-30. 31. Sciortino, F., et al., Crystal stability limits at positive and negative pressures, and crystal-to-glass transitions. Phys Rev E Stat Phys Plasmas Fluids Relat Interdiscip Topics, 1995. 52(6): p. 6484-6491. 32. Nosé, S., A unified formulation of the constant temperature molecular dynamics methods J. Chem. Phys., 1984. 81(1): p. 9. 33. Hoover, W.G., Canonical dynamics: Equilibrium phase-space distributions. Phys Rev A, 1985. 31(3): p. 1695-1697. 34. Parrinello, M.R., A., Polymorphic transitions in single crystals: A new molecular dynamics method Journal of Applied Physics, 1981. 52(12): p. 9. 35. Kruithof, M., et al., Single-molecule force spectroscopy reveals a highly compliant helical folding for the 30-nm chromatin fiber. Nat Struct Mol Biol, 2009. 16(5): p. 534-40. 36. Wu, C., A. Bassett, and A. Travers, A variable topology for the 30-nm chromatin fibre. EMBO Rep, 2007. 8(12): p. 1129-34. 37. Schalch, T., et al., X-ray structure of a tetranucleosome and its implications for the chromatin fibre. Nature, 2005. 436(7047): p. 138-41. 38. Davey, C.A., et al., Solvent mediated interactions in the structure of the nucleosome core particle at 1.9 a resolution. J Mol Biol, 2002. 319(5): p. 1097-113. 39. Xiang, Z., C.S. Soto, and B. Honig, Evaluating conformational free energies: the colony energy and its application to the problem of loop prediction. Proc Natl Acad Sci U S A, 2002. 99(11): p. 7432-7. 40. Guex, N. and M.C. Peitsch, SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis, 1997. 18(15): p. 2714-23. 41. Jacobs, S.A., W. Fischle, and S. Khorasanizadeh, Assays for the determination of structure and dynamics of the interaction of the chromodomain with histone peptides. Methods Enzymol, 2004. 376: p. 131-48. 45

42. Luger, K., T.J. Rechsteiner, and T.J. Richmond, Preparation of nucleosome core particle from recombinant histones. Methods Enzymol, 1999. 304: p. 3-19. 43. Shogren-Knaak, M.A. and C.L. Peterson, Creating designer histones by native chemical ligation. Methods Enzymol, 2004. 375: p. 62-76. 44. Schwarz, P.M. and J.C. Hansen, Formation and stability of higher order chromatin structures. Contributions of the histone octamer. J Biol Chem, 1994. 269(23): p. 16284-9. 45. Jacobs, S.A., et al., Specificity of the HP1 chromo domain for the methylated N- terminus of histone H3. EMBO J, 2001. 20(18): p. 5232-41. 46. Blacketer, M.J., S.J. Feely, and M.A. Shogren-Knaak, Nucleosome interactions and stability in an ordered nucleosome array model system. J Biol Chem, 2010. 285(45): p. 34597-607. 47. Wreggett, K.A., et al., A mammalian homologue of Drosophila heterochromatin protein 1 (HP1) is a component of constitutive heterochromatin. Cytogenet Cell Genet, 1994. 66(2): p. 99-103. 48. Canzio, D., et al., Chromodomain-mediated oligomerization of HP1 suggests a nucleosome-bridging mechanism for heterochromatin assembly. Mol Cell, 2011. 41(1): p. 67-81. 49. Wang, G., et al., Conservation of heterochromatin protein 1 function. Mol Cell Biol, 2000. 20(18): p. 6970-83.

46

Supplemental Figures

(A) (B)

(C) (D)

(E) (F)

(G) (H)

47

Figure S 1. Initial positions for the MD simulations shown in Fig 1. Initial structures for the independent simulations of both wild-type HP1 dimers and hinge HP1 dimers.

Figure S 2. Diagram of the construction of the atomic model for a 12 nucleosome array. We started with two copies of the mononucleosome unit, with short stretches of canonical B- DNA attached at either end (Fig. 1Sa). Structural superposition of three base pairs of the B- DNA at the ‘beginning’ of one copy (green) with three basepairs of the B-DNA at the ‘end’ of a second copy (red), makes it possible to construct a dinucleosome (Fig. 1b; the superimposed region of the DNA is indicated by green and red stripes). Repeating the procedure with a third copy of the mononucleosome produces the trinucleosome structure shown in Fig. 1c; repetition of the procedure with a fourth copy of the mononucleosome produces a tetranucleosome structure (Fig. 1d) and so on. It should be noted that while this procedure is guaranteed to produce a fully connected oligonucleosomal structure, it is not guaranteed to produce a structure that is free of steric clashes. In fact, when we assume that the twist angle between adjacent nucleotides in the linker DNA is 36° the procedure produces a polynucleosome structure that has serious clashes between the i:i+2 nucleosomes. To solve this problem we found it necessary to increase slightly the twist angle between adjacent nucleotides of the linker DNA to 37°.

48

49

Figure S 3. Molecular Weights (determined by sedimentation equilibrium) and fluorescence anisotropy binding results of purified HP1Hsa proteins.

Figure S 4. The transgene-encoded HP1 proteins are expressed at similar levels. Western analysis of transgene-encoded LacI-HP1 protein levels is shown. Genotypes are given at top, molecular weight positions at left, antibodies at right, and relative fold difference in LacI- HP1 levels is given at bottom. y,w is a wild-type fly stock that does not contain a transgene.

50

CHAPTER 3. HP1-MEDIATED CONDENSATION OF METHYLATED CHROMATIN

A paper in preparation for submission

Abdelhamid M. Azzaz1, Melissa J. Blacketer1, J. P. Price2, Lori L. Wallrath2,

Christopher L. Woodcock3, Michael A. Shogren-Knaak1

From The Department of Biochemistry1, Biophysics and Molecular Biology

Iowa State University1, Ames, Iowa, 50011, The Department of Biochemistry2,

University of Iowa2, Iowa City, Iowa 52242, the Biology Department3, University of

Massachusetts, Amherst, Massachusetts 01003

To whom correspondence should be addressed: Michael A. Shogren-Knaak, Ph.D.,

Department of Biochemistry, Biophysics and Molecular Biology, 4214 MBB, Iowa State

University, Ames Iowa 50011; Email: [email protected].

51

Abstract

In eukaryotic organisms heterochromatin is a prevalent and biologically critical form of genomic DNA. As a model of constitutive heterochromatin, we show that heterochromatin protein HP1 is sufficient to induce significant chromatin compaction in nucleosomal arrays fully methylated on histone H3 lysine 9. In addition to requiring histone methylation, this transition to the compacted state is favored by high HP1 concentrations, extensive nucleosome saturation, and pre-existing chromatin condensation. Chromatin compaction by nonspecific DNA interactions can also occur, but is significantly weaker in magnitude.

Introduction

Heterochromatin is a cytologically distinct form of DNA found in eukaryotic organisms, where approximately 10% of human genomic DNA exists in this form.

Heterochromatin directly and indirectly impacts a wide range of DNA-mediated processes, including regulation of gene transcription and maintenance of genomic stability [1, 2].

Because it stains darker than other genomic DNA, heterochromatin has long thought to be an especially condensed form of chromatin [3], but the actual higher-order structure of heterochromatin has remained elusive.

Heterochromatin exists in multiple forms in terms of molecular composition and genomic localization, suggesting that heterochromatin structure is heterologous [3].

However, the composition and localization of one form of heterochromatin, constitutive heterochromatin, is well maintained across species and cell types [4]. Found as large domains 52

proximal to centromeres and telomeres, constitutive heterochromatin generally contains chromatin that is di- or trimethylated on lysine nine of histone H3 as well as the presence of

HP1, heterochromatin protein 1 [5]. HP1 proteins contain three domains, a chromodomain that binds H3K9 methylation, a flexible hinge domain, and a chromoshadow domain that mediates HP1 dimerization [6-9]. Thus, one of the original models of constitutive heterochromatin structure is that HP1 dimers mediate compaction of methylated chromatin by bridging neighboring nucleosomes [10]. In support of this model, in vitro and in vivo binding of HP1 to methylated chromatin is well established [6, 11, 12]. However, binding is not necessarily synonymous with compaction of chromatin. Moreover, studies have revealed additional or potentially alternative structural roles for HP1 [13-15]. It has been shown that

HP1 can associate with heterochromatin in a chromodomain-independent manner, potentially through non-specific binding to DNA and RNA[16]. HP1 has also been shown to localize to non-heterochromatin regions [15, 17]. Finally, HP1 has been shown to directly bind to numerous proteins, serving as a recruitment platform for heterochromatin and non- heterochromatin factors [15]. Thus, to what extent HP1 directly mediates chromatin compaction by binding methylated nucleosomes or whether HP1 mediates chromatin compaction at all, has been unclear.

Experimental procedures

Generation And Characterization of HP1Hs Protein - BL21 (DE3) pLys S cells containing either wild-type human 6His-HP1 plasmid (a gift from Professor Christian

Muchardt, Pasteur Institute in Paris), W71A 6His-hHP1 plasmid, or I165E 6His-hHP1 53

plasmid were grown to an OD600 of 0.5-0.6 and then induced with IPTG (0.2 mM final concentration) for 4 hours. Pelleted cells were resuspended in 20 ml lysis buffer (50 mM

NaH2PO4 pH8, 300 mM NaCl, 10 mM Imidazole, 1 mM 2-mercaptoethanol, 10% glycerol,

1M PMSF, 10 g/ml leupeptin, 10 g/l aprotinin and 50 g/ml DNase I) per 500 ml of culture media, flash frozen, and stored at -80C. Cells were lysed cold using a French press, and the lysate was mixed with Ni-NTA agarose resin (Qiagen). The resin was washed twice

(lysis buffer + 20 mM imidazole), and then the protein was eluted from the resin (lysis buffer

+ 250 mM imidazole). Aggregated HP1Hs was removed from the eluate by gel filtration chromatography (Sephacryl S-300 resin with lysis buffer as the mobile phase). The desired

HP1Hs fractions were pooled and concentrated to about 30 M concentration, as determined by UV absorbance.

In Vitro Chromatin Assembly - Recombinant, unmodified, Xenopus laevis H2A, H2B,

H3, and H4 histones were prepared according to standard protocols [18]. H3 histone uniformly trimethylated at lysine 9 was generated by native chemical ligation of a synthesized H3 1-23 peptide (University of Wisconsin Biotechnology Center) and a recombinant H3 C-terminal fragment, according to previously described methods [19].

Methylated and unmethylated histone octamers were assembled according to standard techniques [18]. 601-177-12 and 601-177-1 DNA templates were liberated from a 601-177-

12-containing plasmid using EcoRV and ScaI, respectively. Carrier DNA was prepared by

PCR amplification as previously described [18]. Following phenol-chloroform extraction,

DNA fragments were further purified by preparative gel electrophoresis, electroelution, butanol extraction, and ethanol precipitation. 601-177-1 mononucleosomes were prepared by rapid dilution deposition with molar ratios of octamer to DNA between 0.8 and 1.2 [18]. 54

Assembled mononucleosomes were dialyzed against 2.5 mM NaCl, 10 mM Tris, pH 8.0,

0.25 mM EDTA with three buffer exchanges and concentrated. Saturation was determined by native gel analysis. 601-177-12 arrays were assembled by salt step dialysis as previously described [18], with ratios of template to carrier to octamer of 1.0:0.3:1.1-1.3.

EMSA Analysis - HP1Hs was dialyzed in 50 mM HEPES pH 8, 150 mM NaCl, 1 mM

EDTA and 10% glycerol. Nucleosome arrays or mononucleosomes were mixed with HP1Hs to a final volume of 20 l containing 50 mM NaCl, 23.33 mM HEPES pH 8, 0.067% TCEP, and 0.33 mM EDTA. The HP1Hs arrays or mononucleosome mixture was mixed gently on the vortexer for 30 minutes, and then applied on 0.5% agarose (type IV) midi-gel made in 50 mM HEPES pH 8 and 150 mM NaCl buffer, and run in the same buffer at 70 volts and 0.4 amp for 2 hours. Gels were visualized with Sybr Gold.

Preparation of Electron Microscopy Samples. Samples were fixed with 160 ml 0.1% glutaraldehyde in the appropriate salt buffer (8 mM or 50 mM NaCl, 23.33 mM HEPES pH

8, 0.33mM EDTA) containing 3.24% glycerol for 4 h at 4oC, and then dialyzed overnight in

2.5 mM NaCl at 4oC. Fixed arrays were diluted 5-10-fold with 50 mM NaCl, applied to glow-discharged thin carbon films for 5 min, then washed with 5mm magnesium acetate, and stained for 10 sec with 0.04% aqueous uranyl acetate, washed with H2O and air dried. Grids were examined with a Tecnai 12 TEM (FEI co., Hillsboro, OR) operated at 100KV in tilted darkfield mode, and images recorded using a 2048x2048 CCD camera (TVIPS, Gauting,

Germany) with 2x2 binning. The final pixel size was 0.908 nm.

55

Results

A key technical difficulty to demonstrating that HP1 directly mediates chromatin compaction by binding methylated nucleosomes has been generating a chromatin model system homogenously methylated on histone H3K9. Recently we have adapted native chemical ligation to uniformly install histone H3 K9 trimethylation in 601-177 12-mer nucleosomal arrays [19, 20]. Using an electrophoretic mobility shift assay (EMSA) we have demonstrated that addition of HP1 can induce the formation of a chromatin species with distinct electrophoretic mobility in a manner dependent on H3K9 methylation ([21] and Fig.

1a). This species is consistent with a more condensed form of chromatin. However, because

EMSA does not directly demonstrate the structure of this species, and because of the complex conditions that molecules experience during gel electrophoresis, we turned to electron microscopy (EM) to more directly visualize the HP1-associated species in solution.

By visual inspection of EM images, we observe that the addition of HP1 to methylated arrays both induces overall array compaction and increases the number of foci containing multiple nucleosomes (Fig 1b, left panels). Each effect can be quantified and shown to be highly statistically significant. By measuring the radius of the smallest circle that encloses the methylated arrays (Supplementary Fig. 1, left panels), we find that the addition of HP1reduces the average minimal bounding radius from 15.3 ± 0.46 a.u. to 12.9 ± 0.39 a.u, with a t-test p value of 0.000067, where lengths are measured in arbitrary units, a.u., and deviation about the population mean is indicated by the standard error. For a given array we can also count the total number of visible particles corresponding to either individual nucleosomes, or clusters of nucleosomes that cannot be differentiated. (Supplementary Fig.

2). Using this technique, the distribution of particles per array is visibly different, and an 56

analysis using a non-parametric Kolmogorov-Smrnov test indicated a p-value of

0.000000043. 57

Figure 6. Effect of HP1 on nucleosomal arrays containing histone H3 K9 methylation. (a) EMSA results, where arrays were visualized by SYBR Gold staining. (b) Electron microscopy analysis at 8 mM NaCl. EM samples for each condition were analyzed to determine the distribution of radii for the smallest circle that contains a given array. The 58

average minimal bounding radius with the associated standard error and sample size is indicated. Like our EMSA results, this HP1-mediated compaction effect appears to have a strong methylation-specific component. Unmethylated arrays were prepared with the same initial ratio of histone octamer to DNA as the methylated arrays and gave a similar distribution of nucleosome occupancy, where an average number of nucleosomes observed for the unmethylated of 11.2 ± 0.21 nucleosomes was similar to the 11.1 ± 0.20 nucleosomes for the methylated (Supplementary Fig. 3a). The methylated and unmethylated arrays were also similar in their compaction in the absence of HP1, with EM and sedimentation velocity analysis indicating that the methylated arrays adopt a slightly more extended conformation than the unmethylated arrays (Supplementary Fig. 1 top panels and 3b), potentially due to the increased negative charge character of the H3 histone tail from the A25C substitution used to install H3K9 methylation by native chemical ligation. However, despite the similarity of these two arrays, the unmethylated array is significantly less sensitive to the addition of the HP1(Fig 1b, right panels). Quantitative measurements of the average minimal bounding radius before and after HP1 addition show a slight decrease in radius (13.7 ± 0.45 a.u. and 13.2 ± 0.31 a.u., respectively) that is not statistically significant (p=0.3)

(Supplementary Fig. 1 right panels). With respect to total visible particle distribution, addition of HP1 to the unmethylated arrays does induce a statistically significant change

(p=0.014) (Supplementary Fig. 2). However, this p-value is more than 10,000-fold less than for the methylated array. Altogether, the EM analysis indicates HP1induces chromatin compaction in a manner highly sensitive to histone H3 K9 methylation and suggests that the gel-shifted species in the EMSA analysis is the compacted state. 59

Although methylation-independent compaction was a relatively small effect in our

EM results, others have reported relatively large effects for such interactions [22]. To better understand the contribution of such binding EMSA HP1 titration experiments were performed on unmethylated arrays. We observed that at concentrations of HP1 higher than our EM studies, generation of a gel-shifted species could be observed (Fig 2a). A contribution to this methylation-independent binding could be due to the previously identified non-specific DNA binding of HP1. Because such interactions are often strongly electrostatic in nature, we performed EMSA HP1 titration experiments under a range of ionic strengths. With unmethyated arrays we observed that increasing ionic strength did indeed decrease the affinity for this methylation-independent binding (Fig 2b), consistent with DNA-mediated binding. To confirm that this interaction was DNA mediated; titrations were performed with the nucleosomal array in the absence of histones. These experiments show that free DNA shows similar, if not enhanced HP1-mediated gel shift, suggesting that our minor methylation-independent nucleosomal array compaction is due to non-specific

DNA binding. Interestingly, this DNA-mediated binding to DNA is not universal to all human HP1 isoforms. Like HP1, HP1 demonstrates DNA binding. However, HP1 does not exhibit DNA binding under similar conditions (Supplementary Fig. 4).

60

Figure 7. Methylation-independent binding. (a) Effect of increasing concentrations of HP1 dimer on unmethylated arrays as determined by EMSA analysis under our standard conditions. Arrays were visualized by SYBR Gold staining. EMSA analysis of HP1 addition to (b) unmethylated arrays or (c) free DNA under various NaCl concentrations.

To dissect other factors responsible for this compaction, we first investigated how changes to array saturation influenced generation of the condensed species. In our studies described so far, nucleosomal arrays with a high occupancy of methylated histone octamers were used (59% with 12 nucleosomes). To determine how array saturation affects compaction, arrays with greater and lesser nucleosome occupancy were investigated (Fig 3). 61

For arrays with a greater nucleosome saturation (79% with 12 nucleosomes), the transition to the gel shifted species occurs at a lower concentration of HP1, while a greater concentration was required for arrays with less nucleosome saturated (13% with 12 nucleosomes). This trend indicates that lower degrees of array occupancy are not an absolute impediment toward array compaction, as greater amounts of HP1 dimer can still induce compaction. However, increasing occupancy of arrays promotes HP1 -mediated compactions. One potential explanation for this trend is that saturated arrays are known to more readily adopt compacted structures.

Figure 8. Effects of varying concentrations of HP1 dimer. EMSA results for arrays with a nucleosome concentration of 8 nM. R indicates the molar ratio of histone octamer to array template nucleosome positioning sequence used in array assembly.

To more explicitly test the effect of pre-existing array compaction on HP1-mediated compaction, EM studies similar to those shown in figure 1 were repeated at a 50 mM NaCl

(Fig 4). Those studies were performed at relatively low salt conditions (8 mM NaCl), and it is known that increased cation concentrations can drive array compaction. By raising the salt concentration from 8 to 50 mM, we observed that in the absence of HP1, the average 62

bounding radius did indeed decrease for both the methylated (15.3 ± 0.46 a.u. to 12.8 ± 0.49 a.u.) and unmethylated arrays (13.7 ± 0.45 a.u. to 11.2 ± 0.48 a.u) (Figs. 1 and 4,

Supplementary Figs. 2 and 5, note – we tested salt concentration above 50 mM NaCl, however, beyond 50 mM NaCl, arrays showed very extensive compaction in the absence of

HP1, making HP1induced increase in compaction less obvious. Additionally, at higher salt concentrations for EM arrays conditions, significant array-to-array interaction was induced). Addition of HP1 to methylated arrays at 50 mM NaCl results in a statistically significant further reduction of average minimal bounding radius (12.8 ± 0.49 a.u. to 10.6 ±

0.47 a.u., p=0.0012). Additionally, unmethylated arrays did not show compaction in the presence of HP1, increasing slightly in average minimal bounding radius (11.2 ± 0.48 a.u. to 12.9 ± 0.45 a.u., p=0.012). Thus our results for methylated arrays suggest that driving chromatin to a more compacted state allows HP1 to more efficiently condense the chromatin. 63

Figure 9. Electron microscopy analysis at 50 mM NaCl. EM samples for each condition were analyzed to determine the distribution of radii for the smallest circle that contains a given array. The average minimal bounding radius with the associated standard error and sample size is indicated.

64

Discussion

Altogether, our data shows that the interaction of HP1 with methylated nucleosomes is sufficient to induce the formation of compacted chromatin. Thus, additional protein and RNA factors associated with heterochromatin are not required for heterochromatin compaction [23, 24]. Nonetheless, our experiments show that a number of factors can help facilitate HP1mediated compaction, including HP1 concentration, nucleosome saturation, and pre-existing chromatin compaction. These factors are likely to be promoted by additional proteins and RNAs associated with heterochromatin. For example, the effective concentration of HP1 may be enhanced by HP1-binding partners that recruit

HP1 to heterochromatic sites. Similarly, proteins complexes that facilitate nucleosome assembly and spacing, as well as nucleosome methylation (and deacetylation and dephosphorylation), are likely to generate an optimal methylated chromatin substrate for

HP1 association. Further, because HP1 concentration is limiting in the nucleus, by establishing optimal nucleosome domain, these regions of the genome will be able to outcompete other regions of the genome for HP1.

What the role of non-specific DNA binding is in heterochromatin binding is less clear. For HP1, nonspecific DNA binding appears significantly reduced at monovalent salt concentration (150 mM NaCl in our EMSA with arrays) approaching physiological conditions (a normal saline solution is 0.9% NaCl which is equal to about 155 mM NaCl).

Thus, one would expect that because of limiting HP1 concentration, nonspecific 65

localization of HP1 to unmethylated nucleosome regions would be limited. Nonetheless, because such non-specific binding can occur at high HP1 concentration, it is still possible that targeted recruitment of HP1 to non-methylated regions might suffice to stabilize HP1 binding. Additionally, our panel of human HP1 isoforms indicates that different forms of

HP1 have differing DNA affinities and these differences within and across species may account for some of the observed targeting of HP1 molecules to unmethylated regions of the genome. These isoform differences, as well as the utilization of highly variable salt conditions and HP1 concentrations, may also help to explain why in vitro experiments have traditionally shown such variability with respect to binding to non-methylated arrays. It should also be pointed out that the ability of HP1 to bind DNA may be used in conjunction with methylation-specific binding to increase the overall affinity of HP1 molecules to methylated nucleosomes.

66

References

1. Dillon, N., Heterochromatin structure and function. Biol Cell, 2004. 96(8): p. 631-7. 2. Grewal, S.I. and D. Moazed, Heterochromatin and epigenetic control of gene expression. Science, 2003. 301(5634): p. 798-802. 3. Silver, L.M. and S.C. Elgin, Immunofluorescent analysis of chromatin structure in relation to gene activity: a speculative essay. Curr Top Dev Biol, 1979. 13 Pt 1: p. 71-88. 4. Dimitri, P., et al., Constitutive heterochromatin: a surprising variety of expressed sequences. Chromosoma, 2009. 118(4): p. 419-35. 5. Stewart MD, L.J., Wong J., Relationship between histone H3 lysine 9 methylation, transcription repression, and heterochromatin protein 1 recruitment. Molecular and cellular biology, 2005. 25(7). 6. Bannister, A.J., et al., Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature, 2001. 410(6824): p. 120-4. 7. Cowieson, N.P., et al., Dimerisation of a chromo shadow domain and distinctions from the chromodomain as revealed by structural analysis. Curr Biol, 2000. 10(9): p. 517-25. 8. Thiru, A., et al., Structural basis of HP1/PXVXL motif peptide interactions and HP1 localisation to heterochromatin. EMBO J, 2004. 23(3): p. 489-99. 9. Smothers, J.F. and S. Henikoff, The hinge and chromo shadow domain impart distinct targeting of HP1-like proteins. Mol Cell Biol, 2001. 21(7): p. 2555-69. 10. Grewal, S.I., RNAi-dependent formation of heterochromatin and its diverse functions. Curr Opin Genet Dev, 2010. 20(2): p. 134-41. 11. Ball, L.J., et al., Structure of the chromatin binding (chromo) domain from mouse modifier protein 1. EMBO J, 1997. 16(9): p. 2473-81. 12. Jacobs, S.A. and S. Khorasanizadeh, Structure of HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Science, 2002. 295(5562): p. 2080-3. 13. Yahi, H., et al., Differential cooperation between heterochromatin protein HP1 isoforms and MyoD in myoblasts. J Biol Chem, 2008. 283(35): p. 23692-700. 14. Singh, P.B., HP1 proteins--what is the essential interaction? Genetika, 2010. 46(10): p. 1424-9. 15. Zeng, W., A.R. Ball, Jr., and K. Yokomori, HP1: heterochromatin binding proteins working the genome. Epigenetics, 2010. 5(4): p. 287-92. 16. Meehan, R.R., C.F. Kao, and S. Pennings, HP1 binding to native chromatin in vitro is determined by the hinge region and not by the chromodomain. Embo J, 2003. 22(12): p. 3164-74. 17. Badugu, R., et al., Mutations in the heterochromatin protein 1 (HP1) hinge domain affect HP1 protein interactions and chromosomal distribution. Chromosoma, 2005. 113(7): p. 370-84. 18. Luger, K., T.J. Rechsteiner, and T.J. Richmond, Preparation of nucleosome core particle from recombinant histones. Methods Enzymol, 1999. 304: p. 3-19. 67

19. Shogren-Knaak, M.A. and C.L. Peterson, Creating designer histones by native chemical ligation. Methods Enzymol, 2004. 375: p. 62-76. 20. Shogren-Knaak, M.A., C.J. Fry, and C.L. Peterson, A native peptide ligation strategy for deciphering nucleosomal histone modifications. J Biol Chem, 2003(In Press). 21. Azzaz, A.M., et al., Multiple interactions between heterochromatin protein 1 (HP1) and nucleosomes. Journal of Biological Chemistry, 2011. 22. Canzio, D., et al., Chromodomain-mediated oligomerization of HP1 suggests a nucleosome-bridging mechanism for heterochromatin assembly. Mol Cell, 2011. 41(1): p. 67-81. 23. Eskeland, R., A. Eberharter, and A. Imhof, HP1 binding to chromatin methylated at H3K9 is enhanced by auxiliary factors. Mol Cell Biol, 2007. 27(2): p. 453-65. 24. Stewart, M.D., J. Li, and J. Wong, Relationship between histone H3 lysine 9 methylation, transcription repression, and heterochromatin protein 1 recruitment. Mol Cell Biol, 2005. 25(7): p. 2525-38.

68

Supplementary figures

Figure S 5. Effect of HP1 on nucleosomal arrays containing histone H3 K9 methylation at 8 mM NaCl. Shown are the EM derived distribution of the minimal bounding radii for the methylated and unmethylated array in the presence and absence of HP1. The average minimal bounding radius and its standard error for each condition are as follows: Methylated arrays in the absence of HP1, 15.3 ± 0.46 a.u. (arbitrary length units), n = 74; Unmethylated arrays in the absence of HP1, 13.7 ± 0.45 a.u., n = 67; Methylated arrays in the presence of HP1, 12.9 ± 0.36 a.u., n = 120; Unmethylated arrays in the presence of HP1, 13.2 ± 0.31 a.u., n = 116.

69

Figure S 6. Distribution of number of particles per methylated array with and without addition of HP1at 8 mM NaCl. Particles were counted from EM images and represent either individual nucleosomes or clusters of nucleosomes that could not be resolved.

70

Figure S 7. Characterization of methylated and unmethylated arrays. (a) Distribution of the number of nucleosomes per array for methylated and non-methylated arrays from EM images. For the methylated arrays the average number of particles per array and their standard errors were 11.1 ± 0.20, n = 23. For the unmethylated arrays, the average was 11.2 ± 0.21, n = 23. Arrays were characterized at a low salt concentration (8 mM NaCl) so that the arrays could be as extended as possible. (b) Distribution of the minimal bounding radii for the methylated and unmethylated arrays from the same images in a. For the methylated arrays, the average minimal bounding radius and its standard error were 15.3 ± 0.46 a.u. (arbitrary length units), n = 74. For the unmethylated arrays, the average minimal bounding radius was 13.7 ± 0.45 a.u, n = 67. (c) Integrated sedimentation velocity distribution for the methylated and unmethylated arrays at 2.5 mM NaCl. Distributions were generated by van Holde-Weischet analysis and normalized to standard conditions (water solution at 20° C). The methylated arrays had a median Sw,20 of 24.5 S. The unmethylated arrays median Sw,20 of 26.0 S. Because the distribution of nucleosomes per array is so similar (part a), comparisons of the sedimentation velocity values are expected to directly report on differences in the 71

friction coefficients between the arrays, where friction coefficients are directly proportional to the Stokes radius of the array.

72

Hsα HP1 0 1 8 16 32X

Hsβ HP1 0 1 3 10 30X

Hs HP1 γ 0 1 3 10 30X

Figure S 8. Binding of different human HP1 isoforms to DNA as determined by EMSA analysis. Arrays were visualized by radiography using 32P-labled DNA.

73

Figure S 9. Effect of HP1 on nucleosomal arrays containing histone H3 K9 methylation at 50 mM NaCl. Shown are the EM derived distribution of the minimal bounding radii for the methylated and unmethylated arrays in the presence or absence of HP1. The average minimal bounding radius and its standard error for each condition are as follows: Methylated arrays in the absence of HP1, 12.8 ± 0.49 a.u. (arbitrary length units), n = 53; Unmethylated arrays in the absence of HP1, 11.2 ± 0.48 a.u., n = 41; Methylated arrays in the presence of HP1, 10.6 ± 0.47 a.u., n = 46; Unmethylated arrays in the presence of HP1, 12.9 ± 0.45 a.u., n = 62.

74

CHAPTER 4. GENERAL CONCLUSIONS

This study reveals several new aspects in the mechanism of formation and structural requirements of a unique form of chromatin; heterochromatin. For the first time in the field of chromatin biology, we were able to generate a uniformly methylated nucleosomal array on lysine 9 of H3 and successfully use these arrays with recombinantly expressed HP1to show very specific and strong modes of binding without the use of any additional proteins or cell lysates, and under almost physiological conditions. This binding was shown to be methylation specific and independent of any previously reported ability for HP1 to bind to naked DNA or unmethylated arrays.

This unique mode of binding was shown to induce a significant compaction of the methylated arrays in comparison to the unmethylated ones. It was also shown to be capable of crosslinking the methylated arrays in an intra- and an inter-molecular fashion as revealed by the EM experiments and the differential sedimentation experiments. The great flexibility of the HP1 dimer can allow it to cover wide range of distance from 40 to 160 Å, giving the chance for different possible modes of binding within the nucleosomal array, which is a point that remains to be investigated more.

The tighter more specific binding that was shown to occur at higher salt concentrations and higher nucleosomal density sheds light on how HP1 might be functional in vivo, as the limiting concentrations of HP1 inside the nucleus will be poised to bind to areas of high methylation density, but will also retain the ability of binding to less densely methylated loci in the presence of the proper stabilizing and recruiting factors. 75

On one hand, the ability of HP1 to form such tight binding was dependent on its ability to form a dimer structure. As when this ability was disrupted in the chromoshadow domain I195E mutant, it lost its ability to cross link the H3-K9 methylated arrays in vitro and could not support chromosomal looping or rescue the deletion of wild type HP1a in vivo.

On the other hand, the heterochromatin forming capacity of HP1a (the Drosophila

HP1 homologue) was not significantly compromised by the deletion of the hinge domain as evidenced by the fact that the hinge mutant was able to support chromosomal looping and rescued the deletion of wild type HP1 in vivo. Additionally, this mutant showed a good deal of flexibility in the molecular dynamics simulations. Both of these facts can explain the evolutionary preservation of both the chromodomain and the chromoshadow domain in contrast to the hinge which is the least conserved domain, as it is the least important in the establishment of a proper heterochromatic structure.

We also had the chance to explore some of the DNA binding aspects of the other two

HP1 isoforms; the - and the -isoforms. Surprisingly, the -isoform showed no DNA binding compared to the  and - isoforms. That might be due to intrinsic differences between the three isoforms in their hinge region which is thought to be responsible for DNA binding. The individual affinities and specificities of the - and -isoforms to methylated chromatin system is something that requires more investigation.

76

ACKNOWLEGMENT

I would like to thank my advisor and major professor Dr. Michael Shogren-Knaak for his guidance and support throughout this tough project. I would like to thank him for his encouragement and continuous help as a great mentor. It has been six wonderful years that I got to know him and work under his supervision, which seeded what I wish to be a lifelong friendship. I would like also to thank the amazing Dr. Melissa Blacketer for her huge help in different stages of the project. My gratitude extends to my lab mates, Divya Sinha, Shanshan

Li, Chitvan Mittal, Haitham Alabsi, Sarah Feely and Isaac Young for their advice and support. Special thanks for my committee members for their time and support along the years of my Graduate school.

Finally, I thank my wife Rania, my son Omar and my parents for their patience, understanding and continuous help.