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Transcriptional and epigenetic control of gene expression in embryo development

Ann Boija

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Doctoral dissertation 2016 Department of Molecular Biosciences, The Wenner-Gren Institute Stockholm University Stockholm, Sweden

©Ann Boija, Stockholm University 2016 ISBN 978-91-7649-537-7 ISBN 978-91-7649-538-4 Printed by Holmbergs, Malmö 2016 Distributor: Department of Molecular Biosciences, The Wenner-Gren Institute

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To my family

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Abstract During cell specification, temporal and spatially restricted gene expression programs are set up, forming different cell types and ultimately a multicellu-lar organism. In this thesis, we have studied the molecular mechanisms by which sequence specific transcription factors and coactivators regulate RNA polymerase II (Pol II) transcription to establish specific gene expression programs and what epigenetic patterns that follows. We found that the transcription factor Dorsal is responsible for establishing discrete epigenetic patterns in the presumptive mesoderm, neuroectoderm and dorsal ectoderm, during early Drosophila embryo development. In addi-tion, these different chromatin states can be linked to distinct modes of Pol II regulation. Our results provide novel insights into how gene regulatory net-works form an epigenetic landscape and how their coordinated actions speci-fy cell identity. CBP/p300 is a widely used co-activator and histone acetyltransferase (HAT) involved in transcriptional activation. We discovered that CBP occupies the genome preferentially together with Dorsal, and has a specific role during development in coordinating the dorsal-ventral axis of the Drosophila em-bryo. While CBP generally correlates with gene activation we also found CBP in H3K27me3 repressed chromatin. Previous studies have shown that CBP has an important role at transcription-al enhancers. We provide evidence that the regulatory role of CBP does not stop at enhancers, but is extended to many genomic regions. CBP binds to insulators and regulates their activity by acetylating histones to prevent spreading of H3K27me3. We further discovered that CBP has a direct regu-latory role at promoters. Using a highly potent CBP inhibitor in combination with ChIP and PRO-seq we found that CBP regulates promoter proximal pausing of Pol II. CBP promotes Pol II recruitment to promoters via a direct interaction with TFIIB, and promotes transcriptional elongation by acetylat-ing the first nucleosome. CBP is regulating Pol II activity of nearly all ex-pressed genes, however, either recruitment or release of Pol II is the rate-limiting step affected by CBP. Taken together, these results reveal mechanistic insights into cell specifica-tion and transcriptional control during development.

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List of Publications I Boija, A. and Mannervik, M. (2016). Initiation of diverse epigenetic

states during nuclear programming of the Drosophila body plan. Proc Natl Acad Sci U S A. 113(31):8735-40

II Holmqvist, P.H., Boija, A., Philip, P., Crona, F., Stenberg, P., & Man-

nervik, M. (2012). Preferential genome targeting of the CBP co-activator by Rel and Smad proteins in early Drosophila melanogaster embryos. PLoS Genetics. 8(6):e1002769

III Philip, P., Boija, A., Vaid, R., Churcher, A.M., Meyers D.J., Cole,

P.A., Mannervik, M. & Stenberg, P. (2015). CBP binding outside of promoters and enhancers in Drosophila melanogaster. Epigenetics & Chromatin, 8:48. doi:10.1186/s13072-015-0042-4

IV Boija, A., Mahat, D.J., Zare, A., Holmqvist, P.H., Philip, P., Meyers

D.J., Cole, P.A., Lis, J.T., Stenberg, P., & Mannervik, M. (2016). CBP regulates promoter-proximal RNA polymerase II. (Submitted).

Related publication

Boija, A. and Mannervik, M. (2015). A time of change: Dynamics of chromatin and transcriptional regulation during nuclear programming in early Drosophila development. Mol. Reprod. Dev., 82: 735–746. doi: 10.1002/mrd.22517

Yeung, K., Boija, A., Karlsson, E., Holmqvist, P.H., Tsatskis, Y., Ni-

soli, I., Yap, D., Lorzadeh, A., Moksa, M., Hirst, M., Aparicio, A., Fanto, M., Stenberg, P., Mannervik, M., & McNeill, H (2016). Whole genome analysis of the transcriptional corepressor, Atrophin, reveals interactions with Trithorax-like in regulation of developmental pattern-ing. (Submitted).

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Abbreviations

Pol II RNA polymerase II TSS Transcription start site PIC Pre-initiation complex GTF General transcription factor TF Transcription factor CTD C-terminal domain CRM Cis regulatory module PTM Post-translational modification HAT Histone acetyl transferase HDAC Histone deacetylase H3K27ac Histone 3 Lysine 27 acetylation H3K27me3 Histone 3 Lysine 27 methylation CBP Creb binding protein P300 300-kDa protein GAF GAGA-factor RTS Rubinstein-Taybi syndrome ChIP Chromatin immunoprecipitation PRO-seq Precision nuclear Run-On and sequencing GRO-seq Global Run-On and sequencing STARR-seq Self-transcribing active regulatory region sequencing FAIRE-seq Formaldehyde-Assisted Isolation of Regulatory Elements PB Pause button DPE Downstream promoter element HMT Histone methyl transferase HDM Histone demethylase PcG Polycomb Group proteins TrxG Trithorax Group proteins PRE Polycomb response element MZT Maternal-to-zygotic transition ZGA Zygotic genome activation Sog short gastrulation Brk brinker

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TableofContents

Introduction..................................................................................11

Chromatinandtranscription.........................................................12Transcription.......................................................................................12EukaryoticDNAiselegantlypackedintochromatin............................13Transcriptionalregulation...................................................................14

KeyfactorsinTranscription...........................................................15Pioneerfactorsopenupthechromatin...............................................15GTFspositionPolIIatthepromoter....................................................16Co-regulatorsintegrateregulatorysignalsduringtranscription...........17

CBP/p300coactivator...................................................................18Conservation.......................................................................................18DomainstructureofCBP/p300............................................................18TheroleofCBP/p300indisease..........................................................19TheroleofCBP/p300inregulatinggeneexpression............................20

KeyregulatoryelementsinTranscription......................................22Promoterelements-determinantsofthemechanismsoftranscriptionalcontrol................................................................................................22Enhancerelements-akeyregulatoryplatformduringtranscriptioncontrol................................................................................................24Insulatorelements..............................................................................27

PromoterproximalpausingofPolII..............................................28PolIICTD............................................................................................28PolIIactivityisregulatedatmultiplestepsduringtranscription.........29KeyplayersofPolIIpausing................................................................30ExperimentalproofofPolIIpausing...................................................30PRO-seq..............................................................................................31TheroleofPolIIpausing.....................................................................32

Histonemodifications...................................................................33Chromatinstates.................................................................................34Theroleofhistonemodificationsintranscription...............................35Histoneacetylation.............................................................................36Histoneacetyltransferases(HATs).......................................................36Histonemethylation...........................................................................37ChIP....................................................................................................38

Epigenetics....................................................................................39Establishmentofcellularidentity........................................................39Stabilityofcellularidentity.................................................................39

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Theepigenomeisplastic.....................................................................40

Development–whenaneggdevelopsintoanorganism...............41Earlyembryodevelopment.................................................................41Zygoticgenomeactivation..................................................................42Patterningoftheembryo....................................................................42Anterior-posterioraxisformation.......................................................43Dorsal-ventralaxisformation..............................................................45Dppsignaling......................................................................................48

Aimofthethesis...........................................................................49

Resultsanddiscussion...................................................................50PaperI................................................................................................50PaperII...............................................................................................52PaperIII..............................................................................................53PaperIV..............................................................................................54

Conclusionandperspectives.........................................................55

Acknowledgements.......................................................................56

Sammanfattning(Swedishsummary):...........................................58

References....................................................................................60

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Introduction

Cell specification is at the core of development, followed by cellular prolif-eration and maturation into a complete organism. Each cell in our body con-tains an identical set up of DNA. Despite this fact some cells will develop into muscle cells and others into neurons. This is a remarkable property of the genome, and the multifaceted processes forming regulated expression are both challenging and amazing to study. Distinct cells express a specific set of genes, i.e. a stretch of DNA that encodes for a function. Regulating which genes that are expressed result in the production of distinct set of proteins and thereby the unique properties of various tissues. These specific gene expression patterns need to be transferred to daughter cells during cell divi-sion to maintain their identical properties. In addition, during life, cells need to adapt to developmental and environmental cues and be able to change the expression status of particular genes. Understanding by what mechanisms gene expression programs are regulated is fundamental for cell identity and development but also for its misregulation in disease.

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Chromatin and transcription

Transcription Transcription is the process where by the genetic information stored in DNA is transferred into RNA. The RNA is then spliced in the nucleus where in-trons are removed to produce messenger RNA (mRNA). The processed mRNA is transported to the cytoplasm and translated by the ribosome into protein. Adding post-translational modifications can in turn regulate the bio-logical activity of the protein. This process of sequence information transfer is called the central dogma of molecular biology. There are also noncoding parts of the DNA that are transcribed into transfer RNA (tRNA) and ribosomal RNA (rRNA) that are involved in the process of translating mRNA into proteins. Recent findings have identified numerous new non-coding RNAs that have been shown to play a significant regulatory role during transcription. Transcription is carried out by three types of RNA polymerases (Pol) (Roeder and Rutter, 1969). The different polymerases synthesize distinct sets of RNAs; Pol I (rRNA), Pol II (proteins, snRNA and miRNA) and Pol III (tRNA, rRNA and small RNA) (Weinmann and Roeder, 1974) (Weinmann et al., 1974). The three polymerases are very similar in subunit composition and structure, but the largest subunit Rpb1 of Pol II has an exclusive C-terminal domain (CTD) that plays a crucial role during tran-scriptional regulation. The central dogma of transcription might seem like a linear and clear-cut process but has been shown to be remarkably complex. Transcription is a tightly regulated process critical for the state of virtually all cells, for cells to take on a specific identity during cellular differentiation and development as well as to avoid diseases.

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Eukaryotic DNA is elegantly packed into chromatin The eukaryotic cell must organize the DNA into a more compact form in order to fit the large amount of DNA into the tiny nucleus. This is achieved by wrapping 147bp of DNA in two super-helical turns around an octamer of histones (two of each H2A, H2B, H3 and H4) forming the basic unit of chromatin, the nucleosome (Kornberg, 1974) (Kornberg and Thomas, 1974) (Luger et al., 1997b). The compaction of DNA into chromatin forming the pattern of beads on a string is not only beautiful like a pearl necklace, it also serves as one important level of control during several essential cellular pro-cesses (Campos and Reinberg, 2009). Chromatin packaging limits the acces-sibility of the DNA and thereby influence key processes in the cell that uti-lizes the genetic information including gene regulation, DNA repair and replication. In addition, the elegant packaging of DNA into the well-structured form of chromatin mediates a highly dynamic structure. Rather than being a static form of packaging, histones arrange the DNA in a way that the sequence information can be stored but also used. The link between chromatin and transcription comes from the discovery that nucleosomes hamper transcription in vitro (Workman and Roeder, 1987). It was later discovered that posttranslational modifications and histone remod-eling including the addition or removal of histones as well as incorporation of histone variants could change the architecture of the chromatin (Li et al., 2007). This explosion in chromatin research provided new means of manipu-lating chromatin and thereby transcription, bringing the field in to the high-light.

Genome organization. DNA is wrapped around an octamer of histones, forming the basic package form, the nucleosomes. Nucleosomes are further organized into chromatin and finally into chromosomes.

chromosome

chromatinnucleosomes

histones

histone tailsDNA

H3 H4

H2A H2B

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Transcriptional regulation Transcription factors (TFs) are DNA binding proteins that bind to regulatory elements of a specific set of genes to control their expression, either acting as activators or repressors. The targeting of a TF to its binding site is influ-enced by protein-protein interactions and the structure of the local chroma-tin. TFs interact with co-regulators that bridge interactions between TFs and the general machinery, composed of a group of general transcription factors (GTFs) and Pol II. Pol II is regulated at the step of recruitment to the pro-moter but also at the step of release from a paused state in order to go into productive elongation. Chromatin modifications recruit a specific set of chromatin regulators but also influence the accessibility of the chromatin to key DNA binding factors.

Transcriptional regulation of a typical eukaryotic gene. Transcription is regulat-ed at several levels; binding of TFs and the recruitment of Coregulators, recruitment and release of Pol II and at the level of chromatin structure.

Activator

Repressor

Coactivator

Corepressor

GTF

PolII PolII PolII

Histone modifications

Pause

release

factor

Pausing

factor

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Key factors in Transcription

Pioneer factors open up the chromatin Chromatin acts as a gatekeeper of regulatory regions by directing the acces-sibility of binding sites for transcription factors. Due to the dynamic nature of chromatin, the accessibility to DNA varies during development and in response to extracellular signaling. A special class of transcription factors, called pioneer factors, has the ability to bind nucleosomal DNA and open up the target region by recruiting chromatin-remodeling enzymes. Cooperative binding of pioneer factors and activators promote an open chromatin con-formation and active transcription. By contrast, cooperative binding between pioneer factors and repressors result in closed chromatin and gene repression (Zaret and Mango, 2016).

Action of pioneer factors. Pioneer factors scan nucleosomal DNA. Upon binding of pioneer factors, they mediate the recruitment of other factors, either an activator or repressor resulting in gene activation or repression, respectively.

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GTFs position Pol II at the promoter Early in vitro studies revealed that isolated purified RNA polymerase to-gether with nucleotides could synthesize RNA, but these RNAs were synthe-sized from random positions. Position specific RNA synthesis was achieved by adding crude nuclear extracts to the reaction (Luse and Roeder, 1980) (Weil et al., 1979). GTFs were later identified in human cells by separating proteins in whole cell extracts based on charge (Matsui et al., 1980). It re-vealed that none of the fractions alone, but the combination of fractions to-gether could drive efficient transcription. The individual transcription factors were then discovered in the different fractions, Transcription Factor for RNA Pol II A (TFIIA) was located in fraction A, TFIID in fraction D and several factors in fraction C. The combined activity as well as the individual roles of the GTFs has been extensively characterized using biochemical studies and crystal structures (Woychik and Hampsey, 2002). The general transcription factors assemble at core promoter sequences (Thomas and Chiang, 2006) (Smale and Kadonaga, 2003) in a stepwise manner to initiate transcription (Orphanides et al., 1996). Promoter recognition is mediated by TATA-binding protein (TBP)-TFIID that recognizes core promoter elements and the interaction between TFIID and DNA is stabilized by TFIIA (Hoiby et al., 2007). TFIIB and TFIIF (Cabart et al., 2011) recruit Pol II and TFIIE and TFIIH mediate promoter opening. The ATP-dependent translocase activity of TFIIH un-winds the DNA to form a transcription bubble. Holoenzyme complexes comprised of only a subset GTFs and Pol II have also been discovered (Chang and Jaehning, 1997) (Myer and Young, 1998). Regardless of the mechanism behind pre-initiation complex (PIC) assembly it will result in transcriptional initiation. However, the initiation may result in several rounds of short nascent transcripts that do not produce full-length transcripts.

Formation of the pre-initiation complex (PIC). Stepwise recruitment of GTFs and Pol II to form the PIC.

TATAA

TBP

TAF

TAF

TAF

TATAA

TBP IIB

TATAA

TBP

IIA

TAF

TAF

IIB

IIF Pol II

TATAA

TBP

IIA

TAF

TAF

IIB

IIF Pol II

IIE IIH TFIIA

TFIIB

TFIIF

Pol II

TFIIE

TFIIH

TAF

TAF IIA

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Co-regulators integrate regulatory signals during transcription While the PIC contains all the minimal factors required for transcription it couldn’t respond to repressors, activators or transcribe genes in chromatin. For this purpose the PIC needs co-regulators and chromatin modifying com-plexes. Biochemical and functional studies have identified a large number of complexes that act as coregulators of specific transcriptional programs. Many coregulators are components of large multisubunit complexes with diverse enzymatic activity that can be summarized into two categories: his-tone modifying enzymes and ATP-dependent remodeling enzymes. Co-regulators associate with DNA binding factors and mediate its regulatory function. They can be divided into coactivators and corepressors with respect to their role during transcription. Mediator One widely used co-activator is the Mediator that acts as a bridge between regulatory factors and the PIC. However, the Mediator has also been shown to stimulate activator-independent transcription and has therefore been sug-gested to categorize as a GTF (Ansari et al., 2009) (Takagi and Kornberg, 2006). The Mediator has a flexible structure and can anatomically be divided into Head, Middle, Tail and Kinase domain. The Head is responsible for the contact with Pol II while the tail domain interacts with transcriptional regula-tors (Borggrefe and Yue, 2011). In contrast to GTFs, the Mediator has a global role in stimulating regulated transcription and exists in different sub-unit compositions (Malik and Roeder, 2010). Thus, in addition to its general activating function, the Mediator may regulate genes in a context specific manner.

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CBP/p300 coactivator

The cyclic-AMP-response element binding (CREB) binding protein (CBP) and its paralog the 300-kDa protein (p300) are two widely used coactivators in metazoan cells. CBP (also called CREBBP and KAT3A) was originally discovered as a nuclear protein that interacted with the phosphorylated form of the transcription factor CREB to promote transcription (Chrivia et al., 1993). p300 (also called EP300 or KAT3B) was found when looking for interaction partners of the adenoviral oncogenic transcription factor E1A. CBP is composed of 2441 amino acids and p300 of 2412 amino acids and the two proteins share high sequence homology but not to other acetyltrans-ferases, and are therefor collectively called CBP/p300 and comprise their own family of acetyltransferases.

Conservation CBP/p300 have been identified in many different species belonging to high-er eukaryotes. There are four CBP/p300 gene orthologs in Arabidopsis (Bordoli et al., 2001) and one copy in flies and roundworms. Rtt109 has been identified as a structural ortholog of CBP/p300 in yeast but lacks func-tional and sequence similarities (Tang et al., 2008). During the evolution of vertebrates, the chromosomal region of CBP/p300 has been duplicated re-sulting in the p300 gene at chromosome 22 and the CBP gene at chromo-some 16 (Giles et al., 1998). The same composition of one CBP gene and one p300 gene can be seen in chicken, opossums, mice and humans, while frogs are missing the p300 gene (Dancy and Cole, 2015).

Domain structure of CBP/p300 CBP is a multidomain protein with a nuclear receptor interaction domain (NRID), several cysteine/histidine regions (CH), a CREB and MYB interac-tion domain (KIX), a bromodomain (binding acetylated lysines), a HAT domain (with an acidic surface that bind lysines and arginines and a nearby loop structure that binds the CoA substrate), a steroid receptor co-activator domain (SID) and an interferon response binding domain (IBiD) (Dancy and Cole, 2015). In addition, the N-terminal part of CBP has an ubiquitin ligase

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function (Grossman et al., 2003) (Shi et al., 2009). The overall sequence similarity between CBP and p300 is 61%, however the acetyltransferase domain and its two flanking domains are especially enriched for sequence conservation (86%) (Chan and La Thangue, 2001). The many overlapping functions of CBP and p300 could be explained by the high sequence homol-ogy. For example, both proteins can bind to E1A and CREB but the two proteins also have distinct functions.

Model of CBP/p300 domain structure. Nuclear receptor interacting domain (NRID), Three cysteine/histidine-rich (C/H) domains which also contain transcrip-tional adaptor zinc fingers (TAZ) or a plant homeo domain (PHD), an interferon binding homology domain (IHD), a CREB and MYB interaction domain (KIX), a bromodomain, a HAT domain, a steroid receptor co-activator domain (SID), an interferon response binding domain (IBiD) and a proline containing PxP domain

The role of CBP/p300 in disease. Altered expression of CBP/p300 results in numerous developmental defects. In humans these are manifested as a developmental disorder named Rubin-stein-Taybi syndrome (RTS) (Petrij et al., 1995). Individuals with RTS ex-hibit short height, learning difficulties, broad thumbs and big toes and char-acteristic facial features including eyes and nose (Rubinstein and Taybi, 1963). The prevalence of the disease is 1 in 100 000 newborns and has been associated with alterations in the CBP gene (Hennekam, 2006). Human stud-ies of CBP have been limited to cell culture models, however, an in vivo system is preferred when studying development. In flies, mice and worms, CBP/p300 is required for cell viability (Goodman and Smolik, 2000). Chromosome translocations that fuse monocytic leukemia zinc-finger pro-tein (MOZ) with CBP or MLL-mixed lineage leukemia (MLL) with CBP have been associated with different types of leukemia. Somatic mutations in HAT- and C/H2-domain as well as truncated p300 are associated with loss of wild-type allele and tumor formation (Iyer et al., 2004). Frequent inacti-vating mutations in CBP and p300 have been found in several types of can-cers, including small-cell lung cancer, bladder cancer, B cell lymphoma etc. (Shen and Laird, 2013).

NR

ID

TA

Z1

/

C/H

1

TA

Z2

/

C/H

3

KIX

bro

mo

PH

D/

C/H

2

HAT CBP

p300

NC

BD

Q-r

ich

E1A

p53

CREB NF B

p53

Nuclear

receptors

PxP

IHD

IBiD

SID

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The role of CBP/p300 in regulating gene expression CBP is a widely used HAT and coactivator with more than 400 interaction partners (Bedford et al., 2010). CBP is a promiscuous HAT in vitro, acetylat-ing many proteins and histone lysines. However, CBP shows much greater substrate selectivity in vivo. In cells, CBP and p300 are required to maintain global levels of H3K18ac and H3K27ac, but dispensable for other histone acetylations (Jin et al., 2011) (Tie et al., 2009). However, a recent study has shown that CBP is also responsible for acetylating H4K8 (Feller et al., 2015). In response to DNA damage, CBP mediates H3K56ac (Das et al., 2009). Furthermore, CBP is known to acetylate a total of 70 proteins in vivo (Wang et al., 2008a). CBP/p300 is well known for its role at enhancers, and genome wide map-ping of CBP has been used to find novel enhancers both in human and flies (Visel et al., 2009) (Negre et al., 2011). Mapping CBP binding in different tissues can be used to predict tissue-specific activity of enhancers (Visel et al., 2009). CBP has been suggested to regulate transcription by three main means, 1) CBP can act as an adaptor protein by bridging activators and GTF that sub-sequently mediate the recruitment of Pol II, 2) CBP can act as scaffolding protein facilitating interaction between proteins and DNA-and-proteins, 3) CBP can act as a HAT, acetylating both histones and non-histone proteins (Holmqvist and Mannervik, 2013). The mechanism by which CBP/p300 is regulating transcription is the topic of paper IV.

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The regulatory roles of CBP. CBP can regulate transcription by acting as (top) a brigde mediating the interaction between pre-initiation complex (PIC) and transcrip-tion factors (TFs), (middle) a scaffolding protein facilitating protein-protein interac-tions e.g. chromatin regulators (CRs) and DNA-protein interactions, (bottom), a HAT acetylating histones and non-histone proteins

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Key regulatory elements in Transcription

Promoter elements- determinants of the mechanisms of transcriptional control The promoter is the central site of action to which the transcriptional ma-chinery binds. Despite the fact that all promoters utilize PIC and that the factors are highly conserved, it is surprising that there is not a DNA element that is shared between all promoters. The core promoter is a region of about 100bp bordering the transcription start site (TSS) that is sufficient to drive correct transcriptional initiation and is composed of several core promoter elements that are recognized by the transcriptional machinery. The majority of promoters contain either of the two motifs TATA-box or Downstream promoter element (DPE) that are recognized by TFIID (Kutach and Kadonaga, 2000). The TATA-box is located -30 base pairs (bp) upstream of TSS and is an A/T rich region bound by TBP subunit of TFIID. DPE is found 30bp downstream of TSS and is bound by the TBP-associated factor (TAF) subunit of TFIID (Burke and Kadonaga, 1996). TFIIB recognition element (BRE) is situated just upstream of TATA-box and mediate an ele-vated affinity of TFIIB for the core promoter (Lagrange et al., 1998). Initia-tor (Inr) is situated right over the TSS and has a role in promoting correct transcriptional initiation (Smale and Baltimore, 1989). The motif 10 element (MTE) is positioned at +18 to +27 bp downstream of TSS and promotes transcription of Pol II (Lim et al., 2004). Pause bottom (PB) is located most commonly at +20 to +30 bp downstream of TSS (Hendrix et al., 2008) and is found at paused genes. The core promoter is extended by the proximal pro-moter, which holds a set of sequence-specific DNA-binding factors that im-pact transcription in different ways. One example of a proximal promoter motif is the GAGA motif commonly found at -100bp to -80bp of TSS, bound by GAGA-factor (GAF) and found at paused genes. A subset of enhancers has a preference for a specific type of core promoter. Studies in the early Drosophila embryo have shown that enhancers in the Antennapedia gene complex and the Bithorax complex have a preference for activating TATA-containing promoters, while the rhomboid enhancer lacks a preference between TATA-containing and TATA-less promoters (Ohtsuki et al., 1998). Furthermore, a subset of enhancers has been found to only acti-vate genes from a DPE containing enhancer and not a TATA containing

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enhancer (Butler and Kadonaga, 2001). This indicates that the composition of the core promoter does not only drive the initiation of transcription but could also regulate enhancer function. One reason for having enhancer speci-ficity to a core promoter could be to enhance accurate association of a distant enhancer and an explicit promoter within a promoter cluster.

Regulatory elements of a typical gene. The core promoter contains a selection of core promoter motifs that mediate its specific regulatory activity. The promoter proximal region holds binding sites for sequence specific DNA binding factors, e.g. GAGA-factor and heat shock factor (HSF). Enhancer elements also exhibit binding sites for sequence specific DNA binding factors and can be located at far distance. Different promoter motifs have been associated with genes that have differ-ent rate-limiting steps during the transcription cycle. Genes that are regulated at the step of recruitment of Pol II to the promoter are rich in TATA-box motif (Chen et al., 2013). By contrast, genes that are or will be regulated at the step of release of the polymerase from the pause site are rich in PB and GAGA-motif (Chen et al., 2013). The Levine lab made a promoter swapping experiment, replacing the highly paused snail promoter with a less-paused promoter and found that it affected the synchrony of gene activation to a more stochastic activation, which resulted in variability of mesoderm invag-ination (Lagha et al., 2013). Thus, the composition of the promoter affects Pol II pausing and mode of gene activation and thereby key developmental processes. Furthermore, it has been shown that promoter motifs bound by Dref and GAGA-factor can separate between ubiquitously expressed house-keeping genes and tissue specific expression of developmental genes, re-spectively (Zabidi et al., 2015). Taken together, the structure of core promoters is thus yet another layer of transcriptional control that contributes to the complexity of organisms.

BRE TATA Inr MTE DPE

-30 bp -37 to -32bp -2bp +18bp +28bp

PB

+25bp

Enhancer 3 GAGA HSE

En

han

cer 2

Core promoter Proximal promoter

-250 to 250bp

GAGA HSF

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Enhancer elements- a key regulatory platform during transcription control While the promoter is sufficient to assemble the RNA pol II machinery and drive low levels of expression of its adjacent gene, high levels of expression often require activity from additional regulatory elements. These elements are called cis- regulatory modules (CRMs) and are located more distant from TSS, sometimes as far as 1Mbp away. CRMs are also called enhancers, due to their role in upregulating, or enhancing transcription of target genes. The term enhancer was first coined after the discovery that the SV40 DNA could ectopically drive the expression of rabbit β-globin gene irrespective of orien-tation (Banerji et al., 1981). Later studies documented endogenous sequenc-es in the immunoglobulin heavy chain locus with similar functions (Neuberger, 1983) (Gillies et al., 1983) (Banerji et al., 1983). Enhancer function Enhancers contain multiple short DNA motifs that act as binding sites for different tissue-specific transcription factors. These factors will in turn re-cruit co-activators and co-repressors and the sum of the combined regulatory activity of all factors bound will determine the transcriptional activity of the enhancer. Looping between enhancer and promoter element has been pro-posed to be critical for enhancer activity and would provide one possible explanation of how enhancers can exert their regulatory activity from far distance (Amano et al., 2009) Enhancer activity has also been associated with DNase I hypersensitivity (Boyle et al., 2008) as well as specific histone modifications of adjacent histones (Heintzman et al., 2009; Heintzman et al., 2007). Active enhancers are dressed with H3K4me1 and H3K27ac while flanking nucleosomes of inactive poised enhancers possess H3K4me1 and H3K27me3 (Shlyueva et al., 2014). Enhancer-derived RNAs (eRNAs) have been isolated but their function is poorly understood (Core et al., 2012) (De Santa et al., 2010). One possibility is that they are involved in keeping the chromatin accessible.

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Enhancers can contact promoters from far distance. Upon binding of a tissue-specific transcription factor, coactivators are recruited and the enhancer and promot-er are brought into close proximity by looping. Flanking nucleosomes of active enhancers are dressed with H3K4me1 and H3K27ac. Global prediction of enhancers The locations of enhancers are hard to predict due to the fact that they can be situated at various distance from their target genes. Advances in DNA se-quencing during the last 10 years have discovered putative enhancers on a global scale. Several strategies in combination with sequencing have been used to predict enhancers on a genome-wide scale (Shlyueva et al., 2014). The chromatin landscape and the binding of regulatory proteins including transcription factors and co-factors has been mapped by ChIP-chip and ChIP-seq. Chromatin accessibility has been mapped using DNase-seq (Boyle et al., 2008), MNase-seq (Yuan et al., 2005) and Formaldehyde-Assisted Isolation of Regulatory elements (FAIRE-seq) (Giresi et al., 2007). The binding of CBP/p300, and the presence of H3K4me1 and H3K27ac have mainly been used to identify active enhancers. Functional test of enhancers Although the above studies have identified a large number of putative en-hancers based on correlation with activity, few have actually been function-ally tested on a global scale. Predicted enhancers have been tested in embry-os by putting the candidate DNA sequence in front of a core promoter fol-lowed by a reporter gene. In situ hybridization can be used to determine the activity of the enhancer by measuring the abundance and localization of the reporter transcript in developing embryos. The enhancer activity could also be measured on the protein level by using enzymatic activity (β -galactosidase), fluorescence (GFP) and antibodies as a read out. These stud-ies require the generation of transgenic animals and are therefore not suitable for enhancer screening genome-wide, but recent efforts have mapped thou-

Pol IIEnhancer

TF

Pol II

EnhancerTF

Coactivator

Coactivator

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sands of sequences for activity in vivo (Tomancak et al., 2007) (Kvon et al., 2014). Advances in functionally testing enhancers have used deep sequencing to be able to test several enhancers in parallel (Shlyueva et al., 2014). Plasmid-based systems have also been used involving placement of the candidate sequence upstream of a minimal promoter followed by a reporter gene con-taining a barcode (Mogno et al., 2013). Self-transcribing active regulatory region sequencing (STARR-seq) brought the functionally testing of enhanc-ers to a genome wide scale (Arnold et al., 2013). Since enhancers can drive the expression of target genes irrespective of orientation, the Stark lab put the enhancer sequence into the reporter gene downstream of the promoter. By directly linking the enhancer activity with its own sequence, and not by barcodes, STARR-seq allows the testing of millions of candidate sequences with variable length to be screened in batches. Systematic analysis of func-tionally characterized enhancer sequences have provided improved under-standing of how regulatory function is encoded in the DNA. The draw back with plasmid-based systems is that you are not assessing the enhancer activi-ty within the genome of a developing animal and might miss important as-pects of developmental gene regulation at the chromatin level (Shlyueva et al., 2014). Recently, new methods to manipulate the activity of enhancers have arisen including transcription activator-like effectors (TALEs) (Crocker and Stern, 2013) and clustered regularly interspaced short palindromic repeat (CRISPR)-Cas9 system (Gilbert et al., 2013). In both systems, transcription factors and cofactors can be targeted to the site of interest and their regulato-ry effect can be monitored. In addition, the two systems can be used to edit the genomic DNA sequence (Reyon et al., 2012) (Jinek et al., 2012) (Mali et al., 2013). These methods will likely give further insights to the functional roles of individual key players.

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Insulator elements Insulator elements are DNA sequence elements that shield genes from inap-propriate regulatory action from the surrounding. Insulators that are located between a promoter and an enhancer can block the ability of the enhancer to activate the promoter (Geyer and Corces, 1992) (Kellum and Schedl, 1991). However, the enhancer is free to activate promoters located on its insulator free side. The blocking ability of insulators has been suggested to be in-volved in structuring chromatin into topological domains with distinct func-tions (West et al., 2002). The insulator can also act as barriers by restricting the spread of condensed chromatin that could shut down the expression of genes (Sun and Elgin, 1999). Insulators are bound by a specific set of insulator proteins for e.g. Su(Hw) (Parkhurst et al., 1988), BEAF (Zhao et al., 1995) and CTCF (Bell et al., 1999) that have been shown to mediate blocking function.

Insulator activity. Insulator elements protect genes from inappropriate regulatory action from the surrounding, (top) by blocking enhancer activity on promoters locat-ed behind the insulator element or (low) by acting as a barrier to prevent the spread of condensed chromatin.

Enhancer Promoter Insu

lato

r

Promoter Promoter

Insu

lato

r

Blocking

Barrier

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Promoter proximal pausing of Pol II

The recruitment of the polymerase to the promoter was long believed to be the major rate-limiting step in transcription. However, more recent studies have shown that the release of a promoter proximal paused polymerase is the rate-limiting step at many genes. The Lis laboratory has contributed with insights on how Pol II pausing occurs by extensive studies on heat shock (hsp) genes in Drosophila (Gilmour and Lis, 1986) (Rougvie and Lis, 1988) (Giardina et al., 1992) (Rasmussen and Lis, 1993). The current view of the transcription cycle is 1. Recruitment of the polymerase to the promoter, 2. Transcriptional initiation. 3.Pausing of the polymerase immediately down-stream of TSS. 4.Release from pausing resulting in productive elongation. 5. Termination of Pol II. 6. Re-initiation of transcription.

Pol II CTD Modifying the activity of the central figure Pol II itself is one important layer of transcriptional regulation and occurs at several checkpoints. Pol II is a multi-subunit enzyme with a C-terminal domain (CTD) on the largest subu-nit, Rpb1, of Pol II. The protruding CTD has multiple heptad repeats and is the key target for regulation of Pol II activity. The number of heptad repeats varies between organisms, budding yeast contain 26-29 repeats, Drosophila 45 repeats and humans 52 repeats (Chapman et al., 2008). Regardless of the length of the CTD, the shared consensus sequence of the heptad repeats is Y1S2P3T4S5P6S7. The CTD is subjected to the action of modifying enzymes that mediate various yet reversible modifications, where phosphorylation of Serines are the best studied. The existence of different forms of the polymer-ase was discovered on SDS-PAGE gels, which revealed two different motili-ties of Rpb1 (Schwartz and Roeder, 1975). One corresponded to a hypo-phosphorylated form of Pol II, which is recruited to the promoter and is a part of the PIC (Lu et al., 1991). The other is a hyperphosphorylated form which is associated with elongation (Cadena and Dahmus, 1987) and needs to be dephosphorylated in order to be recruited to the promoter again for another round of transcription (Cho et al., 1999).

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Pol II activity is regulated at multiple steps during transcription During the transcription cycle, the CTD is extensively modified at distinct steps (Buratowski, 2003, 2009). The three serines of the heptad repeat can be phosphorylated (Ser2, Ser5 and Ser7) (Corden, 2007; Palancade and Bensaude, 2003) which regulate the different steps of the transcription cycle. During PIC formation, Mediator bridges activators and unphosphorylated Pol II resulting in a fully assembled PIC at the promoter, stimulation of the kinase Cdk7 of TFIIH that mediates Pol II CTD phosphorylation at serine 5 and 7 and subsequent release of the Pol II CTD from the Mediator (Sogaard and Svejstrup, 2007) (Murakami et al., 2015). This result in promoter clear-ance, i.e. Pol II is detached from the PIC at the core promoter and starts to initiate transcription. At many genes, Pol II S5P is accumulated 20-50bp downstream of the TSS where it is held in a paused state. Phosphorylation of Pol II CTD Ser2 (Pol II S2P) is required for release into productive elonga-tion. In addition, the CTD has an important regulatory role in serving as a platform for other enzymes participating in RNA maturation including pro-tein complexes that cap, splice and polyadenylate RNA (Bentley, 2014) (Buratowski, 2009) (Egloff et al., 2012). Non-canonical amino-acids are most prevalently found at position number seven of the heptapeptide repeat, where substituting canonical serine 7 to lysine 7 is the most common (Chapman et al., 2008). Lysine 7 is subjected to acetylation by p300 and associated with promoter proximal polymerases at paused genes in mammalian cells. CTD acetylation was further shown to specifically regulate the expression of growth factor induced genes (Schroder et al., 2013). Lysine 7 can also be methylated and associated with the earliest transcription stages before or together with serine 5 and 7 phos-phorylation (Dias et al., 2015). Given that lysine acetylation was critical for only a specific set of genes raises the question: How essential is the CTD for transcription? Early studies in yeast have shown that deleting parts of CTD down to less than 10 repeats are cell lethal, while 10-12 repeats are sufficient for conditional viability (Nonet et al., 1987). Mutation of Serine 7-p shows little effect on the expres-sion of protein coding genes, but has shown to be important for transcription of short non-coding genes (Napolitano et al., 2014). Neither yeast mutants with impaired Ser2p show global effects on gene expression, but selectively affect genes with roles in meiosis (Saberianfar et al., 2011). Taken together, the regulatory function of Pol II CTD is highly complex. Different modifications of the CTD seem to play diverse roles on different sets of genes. Considering all possible CTD modifications of one heptad repeat and then the number of repeats it will give rise to a wide range of variations of different phosphorylation, acetylation and methylation patterns.

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Key players of Pol II pausing Sequence specific transcription factors associate with the promoter and work together with DRB sensitivity-inducing factor (DSIF) and the Negative elongation factor (NELF) to establish a paused polymerase. Upon recruit-ment of positive elongation factor-b (P-TEFb) Ser2 of Pol II CTD becomes phosphorylated as well as NELF (resulting in eviction) and DSIF (trans-formed to a positive elongation factor). Recently, the crystal structure of mammalian Pol II in its transcribing form was resolved and positioned DSIF over the clamp domain of Pol II (Bernecky et al., 2016). P-TEFb is com-posed of cyclin T1 and cyclin-dependent kinase 9 (CDK9). In order for P-TEFb to promote transcription via its phosphorylating activity, it has to be released from an inhibitory complex where it is reversibly associated with a small nuclear ribonucleoprotein (snRNP) complex, 7SK (Zhou et al., 2012).

Experimental proof of Pol II pausing Evidence of a paused polymerase comes from several different experimental techniques including permanganate footprinting that detects hypersensitivity of single stranded thymidines to oxidation. An open transcription bubble could be identified at 20-50 bp downstream of TSS at many genes (Gilmour and Fan, 2009). Recent advances in genome wide studies discovered that this is not just a phenomenon of a few genes but also a widespread signature of 10-40% of genes in human cells and Drosophila (Guenther et al., 2007) (Muse et al., 2007) (Zeitlinger et al., 2007). Chromatin immunoprecipitation sequencing (ChIP-seq) studies detect enrichment of cross-linked polymerase at the pause position. Sequencing of short capped RNAs reveal the presence of transcripts mapping to the region near promoters (Nechaev et al., 2010). Nuclear run on assays, including GRO-seq and PRO-seq, have shown that the paused polymerase is still transcriptionally competent (Core et al., 2008) (Kwak et al., 2013).

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The transcription cycle. Pol II is recruited to the promoter followed by TFIIH me-diated phosphorylation of Pol II CTD on Serine 5, resulting in transcriptional initia-tion. Immediately downstream of TSS, Pol II is paused by NELF and DSIF. Upon recruitment of PTEF-b, Pol II CTD Serine 2 is phosphorylated and Pol II is released from pausing and proceeds into productive elongation. Pol II reaches the end of the gene and is terminated and can then be re-initiated to resume another round of the transcription cycle.

PRO-seq Precision run-on sequencing (PRO-seq) is a technique that was developed in the Lis laboratory to map the levels and orientation of actively transcribing Pol II on a genome wide scale (Kwak et al., 2013). Nuclei are isolated and depletion of ribonucleotide monomers causes Pol II to stop transcribing but is kept in a transcriptional competent state. The run-on is performed by add-ing single biotin-labeled ribonucleotides that are incorporated by transcrip-tionally engaged polymerases and is performed in the presence of Sarkosyl that prevents new initiation of Pol II. A single nucleotide resolution is achieved due to the addition of only one of the four biotin labeled ATP/CTP/GTP/UTP that causes stalling of Pol II and prevents it from fur-ther transcription. Streptavidin beads are used for purifying the nascent RNA followed by deep sequencing The difference between PRO-seq and global run-on sequencing (GRO-seq) is that PRO-seq uses biotin-labeled ribonu-cleotides and GRO-seq BrUTP which results in a longer run-on and thereby decreased resolution (Core et al., 2008). The advantage with the PRO-seq technique is that in contrast to ChIP it does not rely on crosslinking efficien-cy or antibody specificity and detects transcriptionally engaged Pol II irre-spective of phosphorylation status of Pol II CTD.

S5P

S5P

S2P S5P S2P

Pol II

Pol II

Recruitment

Initiation S5P

Pausing

NELF DSIF

TFIIH

Productive elongation

Spt5 Spt5 P

S5P S2P

Termination

P

Spt5 P

Re-initiation

PTEF-b

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The role of Pol II pausing While the existence and importance of Pol II pausing are no longer debated, the role of pausing is still not fully understood. Initially the purpose of paused Pol II was thought to be rapid activation of genes. The reason for this was that the heat shock genes were one of the first genes to be identified to possess a paused Pol II (Rougvie and Lis, 1988). Under high temperature, the heat shock genes are rapidly turned on and the paused polymerase is often believed to prepare the heat shock genes for fast induction. However, recent studies have shown that rapid induction does not always involve paused Pol II (Lin et al., 2011) and that Pol II pausing occurs more frequent-ly in components of signaling cascades rather than inducible target genes (Gilchrist et al., 2012). Furthermore, while Pol II pausing is associated with active genes, its correlation with gene activity is poor (Min et al., 2011; Nechaev et al., 2010). Together these studies indicate that Pol II pausing has a role in mediating the potential of active elongation. At most genes Pol II pausing fine tunes the expression levels in response to environmental cues rather than acting as an on/off switch. The binding of Pol II at the promoter proximal pause site has been suggested to be a mechanism by which the promoter region is kept nucleosome free (Gilchrist et al., 2010). Furthermore, Pol II pausing has been proposed to create a permissive state and is established in a stage specific fashion during development. Ultimate-ly, this permissive state would allow the binding of tissue-specific transcrip-tion factor and when the right combination of transcriptional regulators is binding result in gene activation. Moreover, Pol II pausing is often present at multiple presumptive tissues in early development and is believed to mediate a permissive state that allows the response to morphogens resulting in gene activation in a subset of cells (Gaertner et al., 2012). Pausing could serve as an extra step of transcriptional regulation. Due to the long residence time of Pol II pausing there is time for several interactions with transcriptional regulator that could modify the transcriptional state. Furthermore, the presence of Paused Pol II RNA has been suggested to fur-ther allow interactions with epigenetic modifiers and transcriptional regula-tors.

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Histone modifications

Chromatin modifications influence several nuclear processes including tran-scription. Post-translational modifications (PTMs) of histones can occur both on the globular domain but most prominently on the protruding histone tails. Several different amino acid residues of histones can be modified including lysine (K), arginine (R), serine (S), threonine (T), tyrosine (Y), by small structurally distinct moieties that include acetyl, methyl, phosphate and ubiquitin groups. Modifications of the protruding histone tails have been suggested to affect the inter-nucleosomal interaction and thereby the overall chromatin structure (Luger et al., 1997a). The discovery of acetylated his-tones and its association with high transcription of genes was made over 50 years ago (Allfrey et al., 1964). Since then, Chromatin immunoprecipitation (ChIP) has been used to map a wide range of histone modifications genome wide in various organisms during different developmental stages (Wang et al., 2008b) (Liu et al., 2005) (Negre et al., 2011) (Roudier et al., 2011). Re-cent reports have markedly expanded the list of histone modifications, and their role of action as well as the identification of novel modifications is under intense studies.

Modifications of the four core histones. Histones are dressed with various modifi-cations including acetylation (ac), methylation (me), phosphorylation (Ph) and ubiq-uitination (Ub) on several different amino acids.

H3 H4

H2A H2BS1 K5 K9K13 K36 K119 T120

K20 K120K15

K5 K12S14

R2 T3 K4 R8 K9S10

T11K14

R17K18

K23 R26K27S28K36 K56 K79

S1 R3 K5 K8 K12 K16 K20

ac

acacacacacacac

acac

ac

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acacacacac

acac

me

me

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me meme

me

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Chromatin states Simplified, chromatin can be divided into two states, an open/active eu-chromatic state that is associated with histone acetylation and active tran-scription, and a closed/inactive heterochromatic state associated with histone methylation and repressed transcription. Heterochromatin can further be divided into constitutive heterochromatin marked by H3K9me2/3, which is found at repetitive DNA elements, and facultative heterochromatin marked by H3K27me3 that silences genes in a cell-type specific manner. However, recent studies have divided chromatin into more specialized groups.

Chromatin states. Chromatin can be divided into an acetylated euchromatin that is associated with active transcription, and an inactive heterochromatin state that is methylated. Heterochromatin can further be divided into H3K9me3 constitutive heterochromatin and H3K27me3 facultative heterochromatin. With an attempt to functionally annotate the genome, the Encyclopedia of DNA Elements consortium has generated numerous genome wide data sets containing maps for histone modifications, chromosomal proteins, transcrip-tion factors, transcripts, replication proteins and nucleosome properties (Consortium, 2012). The same approach has been applied for model organ-isms, the model organism Encyclopedia of DNA Elements (modENCODE) project has generated more than 700 genome wide data sets containing maps for different developmental stages and several Drosophila cell lines. Compu-tational approaches are then used to search for recurrent combinations that are grouped together and defined as chromatin states. The combination of 18 chromatin marks has been used to identify 9 broad classes of chromatin states (c1-c9), which can be further subdivided into 30 states (d1-d30), which showed enrichment for specific functions and regulatory elements (mod et al., 2010). Since chromatin not only consist of DNA and histone proteins but also other chromosomal proteins, whole genome maps were generated of 53 chromatin proteins using DamID and led to the distinction of five chromatin states (Filion et al., 2010). Three states represent heterochromatin: GREEN (en-riched in HP1 and H3K9me), BLUE (rich in PcG proteins and H3K27me3) and BLACK (enriched in non-coding elements, Histone H1 and low tran-

H3K27me3 facultative

heterochromatin

Euchromatin Heterochromatin

H3K9me3 constitutive

heterochromatin

Acetylated histones

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scriptional activity). Two states representing euchromatin: RED (enriched for e.g. Brahma and GAF) and YELLOW (rich in Mrg15 and H3K36me3). Taken together, whole genome mapping of chromatin factors and histone modifications can be used to group together recurrent combinations forming different chromatin states. The number of states is arbitrary and dependent on how broad or fine-grained one would like the classification to be. Regard-less of the number of states defined, it is a simplification of tremendous amounts of information that is useful to understanding how the genome is organized. However, identifying the different states seems to be easier than to understand what they mean in the language of biology.

The role of histone modifications in transcription Two models have been suggested to how histone modifications affect tran-scription. One model proposes that the regulatory action lies in the change in charge of histones that result in altered chromatin structure (Zheng and Hayes, 2003). H4K16ac has been shown to regulate higher-order chromatin structure by inhibit the compaction of chromatin into 30nM fibers (Shogren-Knaak et al., 2006). The second model builds upon the existence of a “his-tone code” (Jenuwein and Allis, 2001). The combination of diverse histone modification would form different codes. The code would provide synergis-tic or antagonistic affinities for regulatory proteins that can affect the struc-ture of the chromatin and regulate gene expression. Since information stored in chromatin can also be inherited through cell division, the role of chroma-tin within the field of epigenetics is an area under intense investigation. Different types of histone modifications are localized to a specific position of a gene, together forming a histone landscape with distinct patterns. The precise location of histone modifications is essential for its regulatory output. For example, mistargeting of Set2 that normally methylates H3K36 in the gene body of transcriptionally active genes (Landry et al., 2003) results in gene repression (Strahl et al., 2002). Studies in yeast have revealed a number of hallmarks describing the chromatin landscape of a typical eukaryotic gene (Li et al., 2007). Nucleosomes have a higher density in the gene body than at the promoter, implying that promoter-binding sites of sequence specific tran-scription factors are located in more accessible regions (Bernstein et al., 2004) (Lee et al., 2004) (Sekinger et al., 2005). Active promoters are dressed with H3K4me3 and H3 and H4 acetylation. H3K36me3 and H3K4me2 are present in the coding region, where H3K4me1 is accumulating at the 3’ end. In addition, active or poised metazoan enhancers are enriched for H3K4me1 and H3K27ac (Creyghton et al., 2010). Studies in ES cells have also provid-ed examples of bivalent domains that posses both the repressive mark H3K27me3 and the active mark H3K4me3 (Bernstein et al., 2006). A pro-

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posed role for bivalent domains is to silence developmental genes, but keep them poised for activation at a later stage.

Histone acetylation Histone acetylation has been described as a hallmark of chromatin dressing transcriptionally active genes. How acetylation promotes transcription is not fully understood but has been suggested to involve the weaker interaction between nucleosome and DNA caused by the neutralization of histone tails by acetylation, making them less attractive to the negatively charged DNA. Acetylation of different lysines is often found to coincide, H3K9ac, H3K18ac, H3K27ac and H4ac are found at TSS. In addition H4ac is also found throughout the gene of active genes (Wang et al., 2008b). This fits with the view that histone modifications may act cooperatively and that the cumulative effect of the number acetylated lysines make the gene ready for transcription (Li et al., 2007). The relaxation of the chromatin structure by acetylation mediates increased accessibility and thereby binding of transcrip-tion factors to their target sites. Alternatively, specific patterns of histone modifications may have distinct functions by directing regulatory factors to chromatin and could provide a mechanism for coordinated gene regulation (Kurdistani et al., 2004). Whether acetylation or not is a cause or conse-quence of transcription is still not clear (Roth et al., 2001). The acetylation status of lysines is a highly dynamic process governed by the antagonistic battle between HATs and HDACs. How these widely used proteins regulate specific genes as well as genes on a global level are ongoing questions in the field.

Histone acetyltransferases (HATs) Histone acetyltransferases (HATs) are enzymes that add the reversible acety-lation of specific lysine residues on histones. By contrast histone deacetylas-es (HDACs) removes acetyl groups from lysines resulting in a closed chro-matin conformation. The first protein reported to have HAT activity was the transcriptional adaptor Gcn5 (Brownell et al., 1996). Gcn5 is part of the mul-tisubunit complex SAGA, which can acetylate many lysine residues in vitro (Grant et al., 1999). The Saga complex is highly conserved and promotes transcription via four submodules with individual functions: HAT module acetylates histones, DUB module deubiquitinates H2B which promotes phorphorylation of Pol II CTD (Wyce et al., 2007), SPT module promotes PIC assembly and TAF module is important for the structure of SAGA (Koutelou et al., 2010). SAGA is targeted to chromatin via binding of Sgf29 to H3K4me2/3 resulting in H3K9ac and H3K14ac (Bian et al., 2011).

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Since then, several other HATs have been discovered including CBP/p300, MYST (monocyte leukemia zinc-finger protein (MOZ), Ybf2, Sas2, Tip60) family of transcription factors and nuclear receptor coactivators. HATs are also referred to as KATs (Lysine acetyltransferase) due to their ability to also acetylate non-histone proteins. They are large multidomain and multi-protein complexes that are recruited to DNA by a wide variety of sequence specific transcription factors. In Drososphila, complete loss of CBP is cell lethal and prevents oogenesis, but the hypomorphic allelel nej1 manifest embryonic patterning phenotypes that can be explained by decreased Dpp-signalling caused by reduced ex-pression of tolloid (tld) (Akimaru et al., 1997) (Lilja et al., 2003) (Waltzer and Bienz, 1999). The role of CBP/p300 in pattern formation is addressed in paper II. Many questions still remain about how these enzymes regulate transcription. How are these broadly used HATs recruited to specific target genes? By what mechanism do HATs stimulate transcription? What are the genomic functions of HATs? These questions are addressed in paper II, III and IV.

Histone methylation Histone methylation is both associated with an active and a repressive tran-scriptional state depending on where the methylation mark is located. His-tone methyl transferases (HMTs) mediate the addition of methyl groups from S-adenosylmethionine to lysine that can be both mono- di- and tri-methylated. By contrast, histone demethylases (HDMs) are responsible for the removal of methyl marks. Methylation of H3K9 is mediated by sup-pressor of variegation Su(var)3-9 (but also other HMTs) resulting in re-cruitment of HP1 and heterochromatin formation (Schotta et al., 2002). Polycomb Group (PcG) and Trithorax Group (TrxG) proteins are two key regulatory enzymes that mediate histone methylation but with antagonistic effects. TrxG proteins mediate H3K4 methylation that promotes transcrip-tion through recruitment of HATs and nucleosome remodelling complexes. PcG proteins have a repressive effect on transcription by forming H3K27me3 heterochromatin. 18 and 17 genes, respectively, have been as-signed to PcG and TrxG proteins, and they tend to act in large protein com-plexes (Ringrose and Paro, 2004). Enhancer of zeste E(z) mediate the his-tone methyl transferase activity of Polycomb repressive comlex 2 (PRC2) and specifically methylates H3K27me3 (Czermin et al., 2002). Polycomb (Pc) of Polycomb repressive comlex 1 (PRC1) recognize this mark via its chromodomain (Cao and Zhang, 2004). The Trx subunit of TrxG proteins mediates the methylation of H3K4. The two protein complexes have a con-served core of proteins between Drosophila and human, but also unique accessory proteins. For example, the three sequence specific DNA-binding

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proteins GAF, Pipsquek and Zeste are unique to Drosophila (Ringrose and Paro, 2004). PcG and TrxG target genes hold cis-regulatory elements termed Polycomb response elements (PRE) (Chan et al., 1994). Although target genes are recognized in human, PREs have not been identified. The two regulatory complexes were first identified for their role in Drosophila body patterning by regulating the expression of homeotic (Hox) genes (Ringrose and Paro, 2004). PcG and TrxG are important for maintaining the expression of Hox genes throughout development and adult life, and are therefore im-portant in cellular memory.

ChIP Chromatin immunoprecipitation (ChIP) is a widely used method for identify-ing binding sites of chromatin proteins. It involves crosslinking of your fa-vorite protein to its chromosomal target sites in living cells, chopping up the cross-linked chromatin into smaller pieces to be able to immunoprecipitate the protein with a specific antibody. The crosslinking is reversed and you will isolate DNA that was bound by the protein of interest. The DNA can be analyzed by qPCR, hybridization to an array or more common today, by sequencing. There are several crucial steps in order to achieve a successful ChIP-seq. Crosslinking needs to be done in a concentration and for a time that captures the protein but that does not overcrosslink resulting in artifacts and false peaks. Sonication of your chromatin in a size of about 100-300bp is suitable for sequencing and will give you high resolution. The antibody needs to be highly specific and preferentially verified by western blot and if possible, knockdown of your protein followed by ChIP. It is ideal to use two inde-pendent antibodies that generate consistent results. A recent report has iden-tified non-specific enrichment in ChIP that is not seen in pre-immune serum or input, which they call phantom peaks (Jain et al., 2015). Phantom peaks are enriched in open chromatin of active promoters of highly expressed genes. The authors speculate that phantom peaks could represent regions that are particularly sticky in a combination with low ChIP specificity. When making sense of your genomic data, this is something to have in mind.

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Epigenetics

The term epigenetics means above genetics and was originally coined by C.H. Waddington. His definition of epigenetics was “the causal interactions between genes and their products, which bring the phenotype into being” (Waddington CH. The epigenotype. Endeavour. 1942;1:18–20). Today one common definition of epigenetics is the heritable changes in gene expression that are independent of DNA sequence alterations.

Establishment of cellular identity All cells in the body contain an identical set up of DNA but the expression status of genes differs between cells. Transcription factors have a central role in dictating the expression of a defined set of genes for each cell type. Dur-ing cell differentiation, pluripotent cells receive input from regulatory factors resulting in induction of genes coding for additional transcription factors (Holmberg and Perlmann, 2012). Crossregulation of transcription factors will result in feedforward induction of specific factors that will activate each other and continue to work for an explicit cell fate. Complex transcription factor networks will give rise to a diverse set of gene expression programs, resulting in different cell types. Distinct expression programs of a cell will be accompanied by specific patterns of histone modifications. It might seem peculiar at first that the number of protein-coding genes in humans is only double the amount as that of the Drosophila fly. However, a more complex gene regulatory network could perhaps explain the greater complexity of me versus my flies in the lab. Increasing the number of tran-scription factors involved in regulating target genes would also expand the amount of different gene expression programs, resulting in several different types of cells and increased complexity of an organism.

Stability of cellular identity Cell fusion and transcription factor induced reprogramming experiments have pointed out the importance of continuously active transcription factors for the maintenance of cell identity, a concept suggested already 25 years ago (Blau and Baltimore, 1991). Fusion of somatic cells have shown that

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unknown factors in one cell can reprogram the fusion partner’s genome and induce cell-type specific gene activation (Blau et al., 1983). Forced expres-sion of transcription factors can cause transdifferentiation, i.e. reprogram-ming that result in a switch of cell identity. Misexpression of myoblast de-termination protein 1 (MYOD1) in fibroblasts resulted in a conversion to skeletal muscle cells (Davis et al., 1987). Differentiated states are not always dependent on continuous instructions from key transcription factors. Instead, differential identities may be stabilized by DNA and chromatin modifica-tions. One such example is the body segment identity in Drosophila that is controlled by homeobox (hox) and engrailed genes. Expression of key tran-scription factors in the early embryo set up the hox gene expression patterns, but TrxG and PcG proteins are responsible for maintaining them (Ringrose and Paro, 2004). Interestingly, PcG proteins have been implicated in regula-tion of several genes in the hierarchy that set up hox gene expression (McKeon et al., 1994) (Pelegri and Lehmann, 1994). This implies that tran-scription factor cascades may be supported by chromatin regulatory mecha-nisms and that this begins at an early point of the cascade. The tight link between chromatin structure and regulation of gene expression is illustrated by Position-effect variegation (PEV). This was a phenomena discovered by H.J. Muller that upon the usage of X rays as a mutagen found variegated Drosophila eyes with areas of red and areas of white color (Henikoff, 1990). This phenotype was explained by chromosomal rear-rangement that positioned the white gene, normally located in euchromatin, in close proximity to heterochromatin thus resulting in gene silencing. PEV has been reproduced for many genes when rearranged juxtaposed to hetero-chromatin (Girton and Johansen, 2008).

The epigenome is plastic Numerous studies with diverse strategies have shown that a differentiated state is not irreversible, and that somatic differentiated cells can be repro-grammed all the way back to a pluripotent state. The first evidence comes from John Gurdon, who transferred a somatic differentiated cell nucleus into an enucleated Xenopus frog oocyte. He found that the somatic nucleus could be reprogrammed into pluripotency and had the capacity to develop into a complete animal (Gurdon, 1962). Somatic cells can also be reverted to a pluripotent state by expression of the key transcription factors Oct3/4, Sox2, c-Myc, and Klf4 (Takahashi and Yamanaka, 2006).

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Development – when an egg develops into an organism

The development of no other animal with comparable or superior complexity is more comprehended than that of Drosophila melanogaster. In particular, a lot is known about the genetics of development and homology to Drosophila genes has been used to identify several key developmental genes also in other organisms. Using Drosophila as a model system has several practical advantages including, inexpensive to culture, short generation time of 10 days, use little space, and each female produce hundreds of eggs. In contrast to mouse and human where the embryo develops within the uterus, Drosoph-ila embryo development is visible to study. The genome of Drosophila, which is condensed into 4 chromosomes, is exceptionally well annotated (Hoskins et al., 2015). In fact, 75% of disease causing genes in humans has counterparts also in fly, which makes findings in Drosophila also applicable to human biology and medicine (Reiter et al., 2001) (Chien et al., 2002) (Bier, 2005). The combination of genetic tools and the ChIP technology make the Drososophila embryo a good system to study transcriptional and epigenetic regulation during cell specification in vivo.

Early embryo development During oogenesis, the egg is loaded with maternal mRNA and proteins, which direct the initial development of the early embryo. Upon fertilization, external stimuli activate the egg resulting in a rise of intracellular calcium that triggers the completion of meiosis, remodeling of the male pronucleus, reorganization of the cytoskeleton as well as an alteration in gene regulation. This results in fusion of the male and female pronucleus and is followed by rapid and synchronous cell cycles without gap phases. In Drosophila, the initial cell cycles occur without cytokinesis resulting in a syncytium with 6000 nuclei after 12 divisions, all sharing the same cytoplasm. This property of the syncytium mediates the possibility for key regulatory proteins to dif-fuse across nuclei during early Drosophila development that is of great im-portance during pattern formation. Following nuclear cycle 9, the nuclei migrate to the periphery where they undergo further syncytial divisions to form the syncytial blastoderm (represented by blastula or blastoderm stage in

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other animals). After blastoderm formation, the cell cycles lengthen with inclusion of gap phases in a process termed mid-blastula-transition (MBT). During the Drosophila MBT the peripheral nuclei are enclosed by the plas-ma membrane to form a cellular blastoderm. About 15 nuclei located to the posterior end do not give rise to the cellular blastoderm but develop into pole cells and later germ cells that will form the sperm and egg.

Zygotic genome activation During this initial part of embryogenesis, the mother provides all transcripts and proteins, but as development proceeds, the control is handed over to the zygote that will synthesis all required gene product from its own genome. The zygotic genome will be activated after a species-specific number of mitotic cycles, which is a conserved process in all animals, called the mater-nal-to-zygotic transition (MZT). The MZT can be divided into two distinct chapters, degradation of maternal products and activation of zygotic tran-scription (Tadros and Lipshitz, 2009). In Drosophila, the first part of zygotic genome activation (ZGA) occurs during cycle 8 and involves the activation of only a few tens of genes. The second part occurs at cycle 14 and engages several hundreds of zygotic genes into transcriptional activation. Distinct groups of chromatin marks have been associated with the two dif-ferent parts of ZGA. Acetylation of H4K8, H3K18, and H3K27 accumulate during the first part of ZGA while the chromatin is not dressed with K3K9ac, H3K4me1, H3K4me3, H3K36me3 and H3K27me3 until the se-cond wave of transcription (Chen et al., 2013; Li et al., 2014). This temporal distinction of chromatin marks agrees with analysis of bulk levels of histone modification during Drosophila embryogenesis. High levels of H3K27ac are present during early stages of development while H3K27me3 is not detected until 4 hours of development (Tie et al., 2009). All together, these transcriptional program events result in dramatic morpho-logical changes

Patterning of the embryo The importance of maternally provided RNA and proteins are well illustrat-ed during patterning of the embryo. The principle behind animal embryo pattern formation is that initially equivalent cells in a developing tissue as-sume specific functions by responding to their position along a morphogen gradient. The first pattern is then further refined by short distance regulation resulting in fields of cells with the same properties. The morphogen gradient in the early embryo is composed of maternal gene products that specify the

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body plan along the anterior-posterior and dorsal-ventral axes. Multifaceted regulations of the zygotic genes are used to further specify the embryo bodyplan. The two axes are organized more or less simultaneously but by different regulatory factors.

Anterior-posterior axis formation During the onset of anterior-posterior axis formation, three maternal tran-scription factors Bicoid (Bcd), Hunchback (Hb) and Caudal (Cad) activate its target genes giant (gt), krüppel (kr) and knirps (kni) in spatial domains. Lack of these target genes causes gaps in the body pattern and is therefore collectively called gap genes. The anterior and posterior pole of the embryo is controlled by the activation of the receptor tyorine kinase Torso (Li, 2005). Torso utilizes Ras-MAP signalling pathway to regulate the expression of the two repressors, huckebein (hkb) and tailless (tll). The expression pat-tern of gap genes is further refined by cross-regulation between Tailless, Giant, Krüppel and Knirps. Bicoid, Hunchback and Caudal also activate pair-rule genes whose expres-sion is precisely limited to seven defined stripes perpendicular to the anteri-or-posterior axis by the repressive mechanism of gap genes. This tight regu-lation is mediated by two characteristics, 1. The CRM of pair-rule genes have high complexity with multiple binding sites for many regulatory factors 2. The regulatory function of Gap genes act only on a short range allowing individual CRMs to independently control transcription of the same target genes at the same time. The pair-rule genes in turn encodes for transcription factors that in a combinatorial fashion control the expression of the segment-polarity genes. The 14-stripe expression pattern of segment polarity genes is the precursors of body segments that finally acquire their unique character.

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Patterning of the anterior-posterior axis in early Drosophila embryos. A hierar-chy of factors set up the anterior-posterior axis. Maternally provided Bicoid (Bcd), Hunchback (Hb) and Caudal (Cad) activate gap genes in spatial domains. Bcd, Hb and Cad also activate pair-rule genes whose expression is restricted to seven stripes by the gap genes. The pair-rule genes in turn, regulate the expression of the seg-ment-polarity genes into 14 stripes.

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Dorsal-ventral axis formation Patterning of the dorsal-ventral axis is ultimately controlled by the Rel/NFκB-family transcription factor Dorsal (Hong et al., 2008). The mother deposits Dorsal in the early embryo and is initially cytoplasmic but is trans-located to the nucleus in response to Toll signaling. Pipe is synthesized on the ventral side of the embryo, by cells surrounding the oocyte (follicle cells). Pipe initiates an extracellular proteolytic cascade involving the prote-ases Nudel (Ndl), Gastrulation-defective (Gd), Snake (Snk) and Easter (Ea) resulting in cleaved and active Spätzle (Spz) ligand that binds to Toll recep-tor on the ventral side of the embryo. Toll activates Pelle kinase that in turn phosphorylates Cactus resulting in release of Dorsal protein that migrates into the nucleus and regulates its target genes. Taken together, this result in a gradient of Dorsal with highest concentration on the ventral side. Dorsal regulates about 50 genes along dorsal-ventral axis in order to gener-ate specialized tissues (Levine and Davidson, 2005). Dorsal activates genes in a concentration dependent manner, forming discrete thresholds of gene activity (Chopra and Levine, 2009). High levels of Dorsal at the most ventral side, activates snail (sna) and twist (twi) that possess enhancers with low-affinity Dorsal sites. Expression of these genes mediates the formation of the mesoderm that later becomes muscles and connective tissues. Intermediate to low levels of Dorsal activates short gastrulation (sog) and brinker (brk) that contains more optimal Dorsal sites as well as binding sites for the ubiq-uitously expressed transcription factor Zelda (Zld). Expression of these genes result in formation of the presumptive neuroectoderm and later the ventral nerve chord and ventral epidermis. Other neuroectoderm genes like ventral nervous system defective (vnd), rhomboid (rho) and vein (vn) have enhancers with fixed arrangement of Dorsal and Twist sites, mediating the response to intermediate levels of Dorsal. The Snail repressor prevents the expression of neuroectoderm genes from the ventral most cells. Absence of Dorsal activates decapentaplegic (dpp), tolloid (tll) and zerknüllt (zen) (which are repressed by Dorsal in the mesoderm and neuroectoderm) and results in dorsal ectoderm formation. The repressive function of Dorsal in-volves the interaction with a protein that binds to AT-rich sequences flank-ing the Dorsal site. This result in a conformational change of Dorsal that expose a cryptic peptide motif (Ratnaparkhi et al., 2006) mediating the inter-action with the co-repressor Groucho (Dubnicoff et al., 1997) (Valentine et al., 1998). The concentration dependent response to Dorsal at target genes has been illustrated by using different mutants in the Toll signaling pathway (Anderson et al., 1985). Dominant mutation in the Toll gene (Toll10B) results in a constitutive active form of the receptor (Schneider et al., 1991) and

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thereby the expression of sna and twi and an embryo that entirely consist of mesoderm. Tollrm9/10 mutants have a partially active receptor (Schneider et al., 1991) and ubiquitously express sog and brk leading to neuroectoderm formation in the entire embryo. In pipe mutants and gd7 mutants which lack Toll signaling and are thereby devoid of Dorsal in the nucleus (Moussian and Roth, 2005), expression of genes such as dpp and zen form dorsal ecto-derm in the entire embryo. Taken together, Dorsal controls diverse transcriptional outputs by the com-bined effort of different affinity binding sites, number of binding sites and the co-binding with other transcription factors to mediate the many transcrip-tional outputs (Jiang and Levine, 1993). The molecular mechanisms by which tissue-specific gene expression pat-terns are set up and how they are maintained has been a long-standing ques-tion and is the topic of paper I.

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Patterning of the dorsal-ventral axis in early Drosophila embryos. Maternally provided Dorsal is present in a gradient with highest concentration on the most ven-tral side. Dorsal activates genes in a concentration dependent manner that subdivides the embryo into mesoderm, neuroectoderm and dorsal ectoderm. The Dorsal gradi-ent is established by ventral localized Pipe that triggers a proteolytic cascade involv-ing Nudel, Gastrulation-defective (Gd), Snake and Easter resulting in cleaved and active Spätzle ligand that binds to Toll receptor. Mutants in the Toll signaling path-way that have a homogenous population of cells with respect to Dorsal concentra-tion are depicted to the right. Toll10B mutants have a high Dorsal concentration and will form the mesoderm, Tollrm9/10 mutants have low levels of Dorsal and will form the neuroectoderm and gd7 mutants lack dorsal in the nucleus and will form the dorsal ectoderm in the entire embryo.

Dorsal gradient

mesoderm

neuroectodern

dorsal ectoderm

sna tld

brk twi dpp

sog sna tld

brk twi dpp

sog sna tld

brk twi dpp

Dl

Dl

Dl

Dl

Dl

Dl

Dl

Dl

Dl

Dl

sog

gd7

Tollrm9/10

Toll10B

Pipe

Nudel Gd Snake Easter

Spätzle

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Sna

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Dpp signaling Upon cellularization, transcription factors can no longer diffuse between nuclei and cell signaling is used for communication between cells. Dpp is a homolog of bone morphogenetic protein-4 (BMP-4) that belongs to the TGF-β family of cytokines that are involved in dorsal-ventral patterning also in vertebrates. Dpp acts as a morphogen with peak levels in the most dorsal part of the Drosophila embryo and specifies the dorsal ectoderm and the amnioserosa. Later in development, Dpp is involved in setting up wing im-aginal disc patterning. At the onset of cellularization, dpp is expressed uniformly in cells that lack dorsal in the nucleus but shortly becomes restricted to the most dorsal parts of the embryo. Dpp is inhibited by the two BMP related proteins Sog and Twisted gastrulation (Tsg) that bind Dpp and inhibit it from interacting with its receptor. Sog, a homolog of the vertebrate Chordin, is degraded when bound to Dpp by the metalloproteinase Tld resulting in release of Dpp. Dpp interacts with Screw and the two proteins signal through Thickveins (Tkv) and Saxaphone (Sax) respectively. This result in activation of Smad-signaling where the class I Smad protein Mothers against dpp (Mad) be-comes phosphorylated by Tkv and thereby associated with the class II co-Smad Medea (Whitman, 1998). The two Smads are translocated to the nu-cleus where they bind and activate their target genes. Brk acts as a repressor of Dpp target genes both in the early embryo and in the wing disc (Campbell and Tomlinson, 1999) (Jazwinska et al., 1999a) (Minami et al., 1999). Brk utilizes the long distance co-repressor Groucho (Gro) and the short distance co-repressor CtBP as well as its repressive domain 3R for its repressive ac-tion on target genes (Upadhyai and Campbell, 2013). In the early embryo, Brk together with Sog specify the neuroectoderm and restricts Dpp to more dorsal regions (Jazwinska et al., 1999b).

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Aim of the thesis

The main objective of the thesis is to understand transcriptional and epige-netic regulation of gene expression. More specifically, it aims to describe how epigenetic patterns are first established during cell specification. It also dissects the molecular mechanisms by which the widely used co-activator CBP regulates gene expression and its role in development. The following questions are addressed:

• How is different epigenetic landscapes formed in the dorsal ecto-derm, neuroectoderm and mesoderm in response to the Dorsal mor-phogen? (Addressed in paper I).

• What is the role of CBP at Dorsal target genes during early embryo

development? (Addressed in paper II).

• What is the genomic function of CBP? What function does CBP bindings have outside of promoters and enhancers? (Addressed in paper III).

• How is CBP regulating the different steps in the transcription cycle,

and specifically what is the role of CBP in promoter proximal paus-ing? (Addressed in Paper IV).

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Results and discussion

Paper I In order for a cell to take on a specific fate, transcriptional regulation and epigenetic mechanisms needs to be tightly coordinated. The molecular mechanism for the initiation of specific transcriptional programs has been a subject of considerable interest, including the role of sequence specific tran-scription factor, polymerase and histone modifications in this process. We have performed ChIP-qPCR of key factors in transcriptional control at two neuroectoderm specific genes sog and brk, in naïve embryos and mutant Drosophila embryos that homogenously take on a specific dorsal-ventral fate forming the dorsal ectoderm, neuroectoderm and mesoderm. We find that the pioneer factor Zelda is responsible for establishing a poised polymerase at Dorsal target genes prior to Dorsal activation in naïve cells. Pioneer factors are known for their role in cell differentiation by their unique property in being able to bind condensed chromatin. They endow competen-cy for gene activity or repression by opening the chromatin to transcription factors and histone modifying enzymes. Our results indicate that some pio-neer factors could mediate the establishment of a poised polymerase during the set up of different gene expression patterns. Several outstanding ques-tions remain to be elucidated: By what molecular mechanism does pioneer factors mediate poised Pol II recruitment? What is the regulatory function of the poised polymerase in setting up tissue specific expression patterns? How frequent is this property among pioneer factors and at what types of genes? Given that the process of genome activation is a highly conserved process, it will be interesting to further investigate if this is a general mechanism in animal development. We also found that the key transcription factor Dorsal is responsible for set-ting up diverse epigenetic states. Upon cell specification, Dorsal activates its target genes in the neuroectoderm resulting in recruitment of the co-activator CBP/p300 and a hyperacetylated chromatin. In the mesoderm, sog and brk are not expressed despite the presence of Dorsal in the nucleus. The activity of a transcription factor can be regulated in several ways including competi-tion for binding sites and protein-protein interactions. We find that Dorsal is still bound to these genes but this does not lead to CBP/p300 recruitment or acetylation of the chromatin. We propose that the Snail repressor quenches Dorsal activity leading to a hypoacetylated chromatin. It will be interesting

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to further dissect by what mechanism Snail is modulating Dorsal activity. Furthermore, we find that in the dorsal ectoderm, these genes are held inac-tive by a Polycomb mediated mechanism involving H3K27me3 chromatin. It seems likely that Polycomb is recruited to these genes by a dorsal ectoderm specific factor, but how and what factor remains to be elucidated. Interest-ingly, we find that these two modes of repression result in different RNA polymerase regulation. Whereas a poised polymerase is still bound to the promoter in hypoacetylated chromatin, the polymerase is displaced from H3K27me3-chromatin. Thus, this provides a direct link between different modes of Pol II regulation and chromatin states. It is well appreciated that epigenetic patterns are important for the mainte-nance of tissue-specific gene expression and used as a cellular memory to transfer gene activity programs to daughter cells. However, it is less known how these epigenetic patterns are first established during early development. Our results demonstrate how gene regulatory networks for nuclear pro-gramming initiate diverse epigenetic states to orchestrate embryogenesis.

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Paper II During embryo development, genes must be switched on and off in a highly coordinated and specific manner. Here we investigate the role of the widely used coactivator and histone acetyltransferase CBP/p300 in this process by using ChIP-seq and ChIP-qPCR in wt and mutant embryos lacking Dorsal in the nucleus. Given that CBP interacts and facilitates gene activation by mul-tiple transcription factors from all major families, one might anticipate that CBP should be involved in setting up most transcriptional programs during development. Instead, we find that CBP is specifically regulating gene net-works controlling dorsal-ventral patterning of the embryo. Hundreds of in-teraction partners have been described for CBP but how it is directed to DNA of its target genes is not known. We find that among 40 previously mapped transcription factors, CBP show strongest co-occupancy and also direct interaction with Dorsal. CBP occupancy is selectively Dorsal depend-ent at sites where few factors are bound. In the absence of Dorsal, CBP bind-ing correlates best with Medea, a Smad-protein involved in Dpp-signaling, which is the opposing force that drives patterning of the dorsal-ventral axis. HATs are well known for their role in stimulating transcription, but their broad role in transcriptional regulation of gene expression is not fully under-stood. We find that changes in CBP binding in mutant embryos correlated with the reciprocal change in gene expression levels genome wide. However, we also find that CBP is associated with silent genes and that the presence of H3K27me3 does not prevent the binding of CBP but constrains histone acet-ylation. Given that acetylation and methylation of H3K27 is mutually exclu-sive, this raises the possibility that CBP’s HAT activity could be regulated by substrate availability. Whole-genome mapping of CBP in combination with H3K4me1 has been used to successfully predict enhancers. However, we find that CBP binding to many well-known active enhancers are below our high-confidence cut off, although the binding to many of them are above genome background. While mapping of CBP in different cell-types and developmental stages will in-crease the number of identified enhancers, our data also suggest that in order to get a full picture of regulatory sequences, CBP mapping should be used together with other approaches like mapping of additional HATs. Taken together, CBP is a general coactivator with specific roles in develop-ment by coordinating the dorsal-ventral rather than for example the anterior-posterior axis of the embryo. CBP is also found at hypoacetylated silent genes. Like many other HATs, CBP is a big multidomain protein with sever-al interaction opportunities with multiple regulatory factors, and could per-haps act as a scaffolding protein to regulate gene expression.

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Paper III CBP is a widely used co-activator with HAT activity but its full role in regu-lating transcription and chromatin organisation is not known. We used CBP ChIP-seq in Drosophila S2 cells to map CBP binding genome wide and compared it with modENCODE data profiles of different chromatin proteins. We find that the majority of CBP is binding to active promoters and to both inactive/active enhancers consistent with a regulatory role of CBP in tran-scription. In addition we find that the strongest CBP binding is found at Pol-ycomb regions that contain Polycomb proteins and H3K27me3 marks. At these sites, the binding of CBP does not result in activation or acetylation, indicating that CBP is not antagonizing H3K27me3 as is seen on a global level. Interesting for future studies would be to investigate if the HAT activi-ty of CBP is blocked or if CBP is acetylating chromatin proteins but not histones. We also found that CBP is binding to insulator elements and regu-lates its insulating activity by acetylating histones to prevent spreading of H3K27me3. In summary, CBP is bound to many genomic regions with dif-ferent function including promoters, enhancers, insulators and Polycomb response elements (PREs). A common theme of these regions is that they directly interact with other genomic regions. Interestingly, this indicates that CBP might regulate higher order chromatin structure and the formation of chromosomal domains.

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Paper IV Transcriptional activation requires both the recruitment of the polymerase to the promoter and the subsequent release of Pol II into productive elongation. We used CBP ChIP seq and a highly potent CBP inhibitor in combination with ChIP-qPCR and PRO-seq to map Pol II at single nucleotide resolution to look at the role of CBP in the transcription cycle. We find that CBP is directly interacting with TFIIB and thereby promotes recruitment of Pol II. CBP is also regulating the release of polymerase into productive elongation by acetylating histones. This means that CBP has dual roles during transcrip-tional activation, mediating both of the key steps during the transcription cycle, recruitment and elongation. We further found that CBP is regulating transcription on virtually all ex-pressed promoters but that the rate-limiting step differs. At most promoters, the rate-limiting step is the release of the polymerase from the promoter into elongation. However, we also identified a special class of promoters with high levels of CBP and GAF that have higher levels of paused polymerase than other promoters. These highly paused promoters are marked by a unique set of histone modifications and chromatin factors, are highly ex-pressed through development and largely shared between cell types. At these promoters the rate-limiting step affected by CBP is the recruitment of Pol II to the promoter. Taken together, these results uncover a novel role of CBP at promoters in addition to its well-known role at enhancers. At expressed promoters, CBP has a global role in directly controlling promoter-proximal Pol II to stimulate transcription. It will be interesting to further dissect the role of CBP at pro-moters by investigating if CBP is acetylating key transcriptional regulators and examine how acetylation influence their action.

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Conclusion and perspectives

A few final thoughts on transcription and epigenetic control during devel-opment are as follows. The strategies to regulate transcription are both com-plex and numerous, mediating the amazing diversity of life. While our un-derstanding of how genes are regulated has greatly advanced, it has become evident that chromatin and transcription are two tightly linked key determi-nants in this process. Upon fertilization, the specialized germ cell is un-dressed from chromatin modifications in order for the two pronuclei to come together, forming a totipotent cell. Pioneer factors mediate the transition from a naïve cell to a differentiated cell by opening up the chromatin so oth-er tissue specific factors can bind and recruit cofactors in order to initiate gene expression programs. The commencement of cell specification is ac-companied by distinct histone modifications, first by acetylation and then by methylation that together form chromatin states that maintain proper expres-sion patterns of different tissues. We have shown that the key transcription factor Dorsal is responsible for the establishment of different epigenetic landscapes on target genes in the three developing tissues along the dorsal-ventral axis of the embryo. The next step would be to map these epigenetic landscapes on a genome-wide scale. Given the wide range of players discussed in this thesis that are involved in regulating transcription, understanding the molecular mechanism behind transcriptional control is a big challenge. However, the arrival of high-resolution methods to map chromatin proteins and transcriptionally engaged Pol II on a genome wide scale offer great opportunities to monitor these processes. In addition, the emergence of small molecule inhibitors provides a powerful tool to manipulate the function of specific key factors in transcrip-tion and in combination with genome wide methods will give direct and novel mechanistic insights to their role in gene regulation. Using these methods we discovered an unappreciated role of CBP/p300 at promoters in regulating Pol II pausing. Along with increasing mechanistic insights to tran-scriptional regulation a growing understanding of transcriptional misregula-tion is also being recognized. Small molecule inhibitors targeting key com-ponents in transcriptional control have already shown promising results as potential therapeutic targets in cancer.

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Acknowledgements

There are many people who contributed to this work that I would like to express my gratitude to: First and foremost, I would like to thank my supervisor Mattias Mannervik that with great expertise has provided me with generous guidance and sup-port, but also for challenging me and urged me to do better. It’s been a pleasure to work with you! Per-Henrik for invaluable hands on advises of how to elegantly perform experiments and for being a fantastic co-worker! Olle for being a generous person, giving good advises on cloning and fly-genetics and providing end-less stories about pretty much everything. Filip for teaching me in situ and make excellent RNA preps. Together the four of us shared many years in the lab with endless discussion about science and life and we had a great deal of fun while doing it! Roshan for good collaboration and nice discussions. I would also like to thank all other members in the lab that I had the pleasure of working with Min, Suresh, George, Amanda, Bhumica, Kati, Helin and Olga. Thanks to Christos Samakovlis, Stefan Åström and Qi Dai for their gener-osity in sharing resources and giving valuable advise on my work and career during the years. Monika for taking care of flies and making the department a nice place to work at. Special thanks to Vasilios, for teaching me confocal microscopy and for our discussions about life. Thanks to members of neighboring labs for sharing reagents and contrib-uting to a nice working atmosphere: Ingrid, Naghmeh, Benedetta, Stefanie, Naveena, Emad, Jiang, Chie, Ryo, Georgia, Akkis, Alex, Ana, Dan, Enen, Andreas, Liqun, Katarina, Satish, Fergal, Badrul, Kirsten, Shenqiu, Chun and Taraneh. Thanks are due to Stefan Björklund and Karl Ekwall for introducing me to research during my master studies, and the generous guidance from Nils Elfving, Miriam Larsson and Annelie Strålfors.

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I am grateful for the productive collaboration with Per Stenberg and his team Philge and Aman. Big thanks to Per for taking the time to teach me how to run scripts and for advice on statistics! Many thanks to John Lis and Jay for nice collaboration and long discussions about the role of CBP in pausing! Many thanks to my family, extended family and friends for endless encour-agement during these years. Finally, I want to thank my husband Henric for never-ending support and for sharing happy and difficult times. Thanks to my kids Oscar and Greta for bringing so much love, joy and cheerfulness into my life.

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Sammanfattning (Swedish summary):

Varje cell i vår kropp innehåller en identisk uppsättning av DNA. Trots detta kommer vissa celler att utvecklas till muskelceller, andra till neuroner och vissa till hudceller. Detta är en anmärkningsvärd egenskap hos genomet, styrd av en mångfacetterad process som är både utmanande och fantastisk att studera. Olika celler uttrycker vävnadsspecifika gener, med andra ord en bit av DNA som kodar för en funktion och skrivs av (transkriberas) av ett en-zym som heter Polymeras. Gener kodar för proteiner som ger vävnaden dess unika egenskaper, t.ex. muskelceller uttrycker muskelspecifika proteiner som ger dem förmågan att kontrahera. Dessa specifika genuttrycksmönster måste överföras till dotterceller vid celldelning för att bibehålla dess egen-skaper, vilket sker med hjälp av epigenetiska mekanismer. DNA är rullat runt proteiner som kallas histoner som kan modifieras med små kemiska grupper som t.ex. en acetylgrupp eller en metylgrupp. Dessa modifieringar påverkar uttrycket av gener genom att kontrollera tillgängligheten av DNA och kan också användas för att skapa ett minne av cellens genaktivitet. Cel-ler måste kunna ändra uttrycksstatusen av vissa gener för att anpassa sig till förändringar under utvecklingen samt till olika miljöfaktorer. Att kartlägga mekanismerna som reglerar genuttrycksprogram är avgörande för att förstå hur cellidentitet uppkommer och bibehålls under utvecklingen av en organ-ism men också för att kunna identifiera vad som går fel i denna process vid sjukdom som t.ex. cancer. I denna avhandling har vi studerat de molekylära mekanismerna som an-vänds för att etablera och bibehålla specifika genuttrycksprogram under em-bryoutvecklingen. Många cellulära processer inklusive hur genuttryck kon-trolleras är konserverat mellan bananflugan Drosophila melanogaster och människa. Vi har därför använt oss av de avancerade genetiska verktyg som finns i bananflugan för att följa hur det epigenetiska mönstret förändras på gennivå från naiva celler till specificerade vävnader under embryoutveckl-ing. Vi har identifierat Dorsal som den faktor som både direkt och indirekt sätter upp olika epigenetiska mönster i celler som kommer att bilda muskel celler (mesodermet), nervceller (neuroektodermet) och i celler som kommer att bilda hudceller (dorsala ektodermet). Dorsal rekryterar CBP som acetyle-rar histoner och leder till aktivt genuttryck. Vidare har vi påvisat två mekan-ismer som tystar gener i dessa vävnader, antingen genom Polycomb medie-rad metylering av histone 3 lysin 27 eller genom hypoacetylerade histoner

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etablerade av Snail. Vi kan dessutom länka de olika histone modifierings-mönstren med distinkta sätt att reglera Polymeraset. Våra resultat tillhanda-håller nya insikter till hur genregulatoriska nätverk formar det epigenetiska landskapet och hur dessa koordinerade aktioner specificerar cellidentitet. CBP/p300 har en viktig roll i att reglera uttrycket av gener och mutationer av CBP är associerade med olika typer av cancer. Tidigare studier har visat att CBP har en viktig roll vid regulatoriska element kallade enhancers. Vi till-handahåller bevis för att CBPs regulatoriska roll inte stannar vid enhancers utan sträcker sig till många genomiska regioner. Genom att använda en mycket potent CBP-drog har vi identifierat en ny regulatorisk roll hos CBP i att kontrollera ett nyckelsteg vid genaktivering: Polymerase pausing. CBP stimulerar rekryteringen av Polymeraset, till ett annat regulatoriskt element med namnet promotor, via direkt interaktion med TFIIB. CBP släpper iväg det pausade Polymeraset från promotorn längs genen genom att acetylera första oktameren av histoner. CBP reglerar Polymerasets aktivitet på i prin-cip alla gener, men antingen rekrytering eller frigörandet av Polymeraset är det hastighetsbegränsande steget som påverkas av CBP. Sammantaget bidrar dessa resultat till ny mekanistisk förståelse för tran-skriptionell och epigenetisk kontroll av genuttryck. Att kartlägga de under-liggande mekanismerna som reglerar genuttryck är essentiellt för att förstå hur cancerceller skiljer sig mot vanliga celler.

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References

Akimaru, H., Hou, D.X., and Ishii, S. (1997). Drosophila CBP is required for dorsal-dependent twist gene expression. Nat Genet 17, 211-214. Allfrey, V.G., Faulkner, R., and Mirsky, A.E. (1964). Acetylation and Methylation of Histones and Their Possible Role in the Regulation of Rna Synthesis. Proc Natl Acad Sci U S A 51, 786-794. Amano, T., Sagai, T., Tanabe, H., Mizushina, Y., Nakazawa, H., and Shiroishi, T. (2009). Chromosomal dynamics at the Shh locus: limb bud-specific differential regulation of competence and active transcription. Dev Cell 16, 47-57. Anderson, K.V., Jurgens, G., and Nusslein-Volhard, C. (1985). Establishment of dorsal-ventral polarity in the Drosophila embryo: genetic studies on the role of the Toll gene product. Cell 42, 779-789. Ansari, S.A., He, Q., and Morse, R.H. (2009). Mediator complex association with constitutively transcribed genes in yeast. Proc Natl Acad Sci U S A 106, 16734-16739. Arnold, C.D., Gerlach, D., Stelzer, C., Boryn, L.M., Rath, M., and Stark, A. (2013). Genome-wide quantitative enhancer activity maps identified by STARR-seq. Science 339, 1074-1077. Banerji, J., Olson, L., and Schaffner, W. (1983). A lymphocyte-specific cellular enhancer is located downstream of the joining region in immunoglobulin heavy chain genes. Cell 33, 729-740. Banerji, J., Rusconi, S., and Schaffner, W. (1981). Expression of a beta-globin gene is enhanced by remote SV40 DNA sequences. Cell 27, 299-308. Bedford, D.C., Kasper, L.H., Fukuyama, T., and Brindle, P.K. (2010). Target gene context influences the transcriptional requirement for the KAT3 family of CBP and p300 histone acetyltransferases. Epigenetics 5, 9-15. Bell, A.C., West, A.G., and Felsenfeld, G. (1999). The protein CTCF is required for the enhancer blocking activity of vertebrate insulators. Cell 98, 387-396. Bentley, D.L. (2014). Coupling mRNA processing with transcription in time and space. Nat Rev Genet 15, 163-175. Bernecky, C., Herzog, F., Baumeister, W., Plitzko, J.M., and Cramer, P. (2016). Structure of transcribing mammalian RNA polymerase II. Nature 529, 551-554. Bernstein, B.E., Liu, C.L., Humphrey, E.L., Perlstein, E.O., and Schreiber, S.L. (2004). Global nucleosome occupancy in yeast. Genome Biol 5, R62.

61

Bernstein, B.E., Mikkelsen, T.S., Xie, X., Kamal, M., Huebert, D.J., Cuff, J., Fry, B., Meissner, A., Wernig, M., Plath, K., et al. (2006). A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell 125, 315-326. Bian, C., Xu, C., Ruan, J., Lee, K.K., Burke, T.L., Tempel, W., Barsyte, D., Li, J., Wu, M., Zhou, B.O., et al. (2011). Sgf29 binds histone H3K4me2/3 and is required for SAGA complex recruitment and histone H3 acetylation. EMBO J 30, 2829-2842. Bier, E. (2005). Drosophila, the golden bug, emerges as a tool for human genetics. Nat Rev Genet 6, 9-23. Blau, H.M., and Baltimore, D. (1991). Differentiation requires continuous regulation. J Cell Biol 112, 781-783. Blau, H.M., Chiu, C.P., and Webster, C. (1983). Cytoplasmic activation of human nuclear genes in stable heterocaryons. Cell 32, 1171-1180. Bordoli, L., Netsch, M., Luthi, U., Lutz, W., and Eckner, R. (2001). Plant orthologs of p300/CBP: conservation of a core domain in metazoan p300/CBP acetyltransferase-related proteins. Nucleic Acids Res 29, 589-597. Borggrefe, T., and Yue, X. (2011). Interactions between subunits of the Mediator complex with gene-specific transcription factors. Semin Cell Dev Biol 22, 759-768. Boyle, A.P., Davis, S., Shulha, H.P., Meltzer, P., Margulies, E.H., Weng, Z., Furey, T.S., and Crawford, G.E. (2008). High-resolution mapping and characterization of open chromatin across the genome. Cell 132, 311-322. Brownell, J.E., Zhou, J., Ranalli, T., Kobayashi, R., Edmondson, D.G., Roth, S.Y., and Allis, C.D. (1996). Tetrahymena histone acetyltransferase A: a homolog to yeast Gcn5p linking histone acetylation to gene activation. Cell 84, 843-851. Buratowski, S. (2003). The CTD code. Nat Struct Biol 10, 679-680. Buratowski, S. (2009). Progression through the RNA polymerase II CTD cycle. Mol Cell 36, 541-546. Burke, T.W., and Kadonaga, J.T. (1996). Drosophila TFIID binds to a conserved downstream basal promoter element that is present in many TATA-box-deficient promoters. Genes Dev 10, 711-724. Butler, J.E., and Kadonaga, J.T. (2001). Enhancer-promoter specificity mediated by DPE or TATA core promoter motifs. Genes Dev 15, 2515-2519. Cabart, P., Ujvari, A., Pal, M., and Luse, D.S. (2011). Transcription factor TFIIF is not required for initiation by RNA polymerase II, but it is essential to stabilize transcription factor TFIIB in early elongation complexes. Proc Natl Acad Sci U S A 108, 15786-15791. Cadena, D.L., and Dahmus, M.E. (1987). Messenger RNA synthesis in mammalian cells is catalyzed by the phosphorylated form of RNA polymerase II. J Biol Chem 262, 12468-12474. Campbell, G., and Tomlinson, A. (1999). Transducing the Dpp morphogen gradient in the wing of Drosophila: regulation of Dpp targets by brinker. Cell 96, 553-562.

62

Campos, E.I., and Reinberg, D. (2009). Histones: annotating chromatin. Annu Rev Genet 43, 559-599. Cao, R., and Zhang, Y. (2004). SUZ12 is required for both the histone methyltransferase activity and the silencing function of the EED-EZH2 complex. Mol Cell 15, 57-67. Chan, C.S., Rastelli, L., and Pirrotta, V. (1994). A Polycomb response element in the Ubx gene that determines an epigenetically inherited state of repression. EMBO J 13, 2553-2564. Chan, H.M., and La Thangue, N.B. (2001). p300/CBP proteins: HATs for transcriptional bridges and scaffolds. J Cell Sci 114, 2363-2373. Chang, M., and Jaehning, J.A. (1997). A multiplicity of mediators: alternative forms of transcription complexes communicate with transcriptional regulators. Nucleic Acids Res 25, 4861-4865. Chapman, R.D., Heidemann, M., Hintermair, C., and Eick, D. (2008). Molecular evolution of the RNA polymerase II CTD. Trends Genet 24, 289-296. Chen, K., Johnston, J., Shao, W., Meier, S., Staber, C., and Zeitlinger, J. (2013). A global change in RNA polymerase II pausing during the Drosophila midblastula transition. Elife 2, e00861. Chien, S., Reiter, L.T., Bier, E., and Gribskov, M. (2002). Homophila: human disease gene cognates in Drosophila. Nucleic Acids Res 30, 149-151. Cho, H., Kim, T.K., Mancebo, H., Lane, W.S., Flores, O., and Reinberg, D. (1999). A protein phosphatase functions to recycle RNA polymerase II. Genes Dev 13, 1540-1552. Chopra, V.S., and Levine, M. (2009). Combinatorial patterning mechanisms in the Drosophila embryo. Brief Funct Genomic Proteomic 8, 243-249. Chrivia, J.C., Kwok, R.P., Lamb, N., Hagiwara, M., Montminy, M.R., and Goodman, R.H. (1993). Phosphorylated CREB binds specifically to the nuclear protein CBP. Nature 365, 855-859. Consortium, E.P. (2012). An integrated encyclopedia of DNA elements in the human genome. Nature 489, 57-74. Corden, J.L. (2007). Transcription. Seven ups the code. Science 318, 1735-1736. Core, L.J., Waterfall, J.J., Gilchrist, D.A., Fargo, D.C., Kwak, H., Adelman, K., and Lis, J.T. (2012). Defining the status of RNA polymerase at promoters. Cell Rep 2, 1025-1035. Core, L.J., Waterfall, J.J., and Lis, J.T. (2008). Nascent RNA sequencing reveals widespread pausing and divergent initiation at human promoters. Science 322, 1845-1848. Creyghton, M.P., Cheng, A.W., Welstead, G.G., Kooistra, T., Carey, B.W., Steine, E.J., Hanna, J., Lodato, M.A., Frampton, G.M., Sharp, P.A., et al. (2010). Histone H3K27ac separates active from poised enhancers and predicts developmental state. Proc Natl Acad Sci U S A 107, 21931-21936. Crocker, J., and Stern, D.L. (2013). TALE-mediated modulation of transcriptional enhancers in vivo. Nat Methods 10, 762-767. Czermin, B., Melfi, R., McCabe, D., Seitz, V., Imhof, A., and Pirrotta, V. (2002). Drosophila enhancer of Zeste/ESC complexes have a histone H3

63

methyltransferase activity that marks chromosomal Polycomb sites. Cell 111, 185-196. Dancy, B.M., and Cole, P.A. (2015). Protein lysine acetylation by p300/CBP. Chem Rev 115, 2419-2452. Das, C., Lucia, M.S., Hansen, K.C., and Tyler, J.K. (2009). CBP/p300-mediated acetylation of histone H3 on lysine 56. Nature 459, 113-117. Davis, R.L., Weintraub, H., and Lassar, A.B. (1987). Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell 51, 987-1000. De Santa, F., Barozzi, I., Mietton, F., Ghisletti, S., Polletti, S., Tusi, B.K., Muller, H., Ragoussis, J., Wei, C.L., and Natoli, G. (2010). A large fraction of extragenic RNA pol II transcription sites overlap enhancers. PLoS Biol 8, e1000384. Dias, J.D., Rito, T., Torlai Triglia, E., Kukalev, A., Ferrai, C., Chotalia, M., Brookes, E., Kimura, H., and Pombo, A. (2015). Methylation of RNA polymerase II non-consensus Lysine residues marks early transcription in mammalian cells. Elife 4. Dubnicoff, T., Valentine, S.A., Chen, G., Shi, T., Lengyel, J.A., Paroush, Z., and Courey, A.J. (1997). Conversion of dorsal from an activator to a repressor by the global corepressor Groucho. Genes Dev 11, 2952-2957. Egloff, S., Dienstbier, M., and Murphy, S. (2012). Updating the RNA polymerase CTD code: adding gene-specific layers. Trends Genet 28, 333-341. Feller, C., Forne, I., Imhof, A., and Becker, P.B. (2015). Global and specific responses of the histone acetylome to systematic perturbation. Mol Cell 57, 559-571. Filion, G.J., van Bemmel, J.G., Braunschweig, U., Talhout, W., Kind, J., Ward, L.D., Brugman, W., de Castro, I.J., Kerkhoven, R.M., Bussemaker, H.J., et al. (2010). Systematic protein location mapping reveals five principal chromatin types in Drosophila cells. Cell 143, 212-224. Gaertner, B., Johnston, J., Chen, K., Wallaschek, N., Paulson, A., Garruss, A.S., Gaudenz, K., De Kumar, B., Krumlauf, R., and Zeitlinger, J. (2012). Poised RNA polymerase II changes over developmental time and prepares genes for future expression. Cell Rep 2, 1670-1683. Geyer, P.K., and Corces, V.G. (1992). DNA position-specific repression of transcription by a Drosophila zinc finger protein. Genes Dev 6, 1865-1873. Giardina, C., Perez-Riba, M., and Lis, J.T. (1992). Promoter melting and TFIID complexes on Drosophila genes in vivo. Genes Dev 6, 2190-2200. Gilbert, L.A., Larson, M.H., Morsut, L., Liu, Z., Brar, G.A., Torres, S.E., Stern-Ginossar, N., Brandman, O., Whitehead, E.H., Doudna, J.A., et al. (2013). CRISPR-mediated modular RNA-guided regulation of transcription in eukaryotes. Cell 154, 442-451. Gilchrist, D.A., Dos Santos, G., Fargo, D.C., Xie, B., Gao, Y., Li, L., and Adelman, K. (2010). Pausing of RNA polymerase II disrupts DNA-specified nucleosome organization to enable precise gene regulation. Cell 143, 540-551. Gilchrist, D.A., Fromm, G., dos Santos, G., Pham, L.N., McDaniel, I.E., Burkholder, A., Fargo, D.C., and Adelman, K. (2012). Regulating the

64

regulators: the pervasive effects of Pol II pausing on stimulus-responsive gene networks. Genes Dev 26, 933-944. Giles, R.H., Dauwerse, H.G., van Ommen, G.J., and Breuning, M.H. (1998). Do human chromosomal bands 16p13 and 22q11-13 share ancestral origins? Am J Hum Genet 63, 1240-1242. Gillies, S.D., Morrison, S.L., Oi, V.T., and Tonegawa, S. (1983). A tissue-specific transcription enhancer element is located in the major intron of a rearranged immunoglobulin heavy chain gene. Cell 33, 717-728. Gilmour, D.S., and Fan, R. (2009). Detecting transcriptionally engaged RNA polymerase in eukaryotic cells with permanganate genomic footprinting. Methods 48, 368-374. Gilmour, D.S., and Lis, J.T. (1986). RNA polymerase II interacts with the promoter region of the noninduced hsp70 gene in Drosophila melanogaster cells. Mol Cell Biol 6, 3984-3989. Giresi, P.G., Kim, J., McDaniell, R.M., Iyer, V.R., and Lieb, J.D. (2007). FAIRE (Formaldehyde-Assisted Isolation of Regulatory Elements) isolates active regulatory elements from human chromatin. Genome Res 17, 877-885. Girton, J.R., and Johansen, K.M. (2008). Chromatin structure and the regulation of gene expression: the lessons of PEV in Drosophila. Adv Genet 61, 1-43. Goodman, R.H., and Smolik, S. (2000). CBP/p300 in cell growth, transformation, and development. Genes Dev 14, 1553-1577. Grant, P.A., Eberharter, A., John, S., Cook, R.G., Turner, B.M., and Workman, J.L. (1999). Expanded lysine acetylation specificity of Gcn5 in native complexes. J Biol Chem 274, 5895-5900. Grossman, S.R., Deato, M.E., Brignone, C., Chan, H.M., Kung, A.L., Tagami, H., Nakatani, Y., and Livingston, D.M. (2003). Polyubiquitination of p53 by a ubiquitin ligase activity of p300. Science 300, 342-344. Guenther, M.G., Levine, S.S., Boyer, L.A., Jaenisch, R., and Young, R.A. (2007). A chromatin landmark and transcription initiation at most promoters in human cells. Cell 130, 77-88. Gurdon, J.B. (1962). The developmental capacity of nuclei taken from intestinal epithelium cells of feeding tadpoles. J Embryol Exp Morphol 10, 622-640. Heintzman, N.D., Hon, G.C., Hawkins, R.D., Kheradpour, P., Stark, A., Harp, L.F., Ye, Z., Lee, L.K., Stuart, R.K., Ching, C.W., et al. (2009). Histone modifications at human enhancers reflect global cell-type-specific gene expression. Nature 459, 108-112. Heintzman, N.D., Stuart, R.K., Hon, G., Fu, Y., Ching, C.W., Hawkins, R.D., Barrera, L.O., Van Calcar, S., Qu, C., Ching, K.A., et al. (2007). Distinct and predictive chromatin signatures of transcriptional promoters and enhancers in the human genome. Nat Genet 39, 311-318. Hendrix, D.A., Hong, J.W., Zeitlinger, J., Rokhsar, D.S., and Levine, M.S. (2008). Promoter elements associated with RNA Pol II stalling in the Drosophila embryo. Proc Natl Acad Sci U S A 105, 7762-7767.

65

Henikoff, S. (1990). Position-effect variegation after 60 years. Trends Genet 6, 422-426. Hennekam, R.C. (2006). Rubinstein-Taybi syndrome. Eur J Hum Genet 14, 981-985. Hoiby, T., Zhou, H., Mitsiou, D.J., and Stunnenberg, H.G. (2007). A facelift for the general transcription factor TFIIA. Biochim Biophys Acta 1769, 429-436. Holmberg, J., and Perlmann, T. (2012). Maintaining differentiated cellular identity. Nat Rev Genet 13, 429-439. Holmqvist, P.H., and Mannervik, M. (2013). Genomic occupancy of the transcriptional co-activators p300 and CBP. Transcription 4, 18-23. Hong, J.W., Hendrix, D.A., Papatsenko, D., and Levine, M.S. (2008). How the Dorsal gradient works: insights from postgenome technologies. Proc Natl Acad Sci U S A 105, 20072-20076. Hoskins, R.A., Carlson, J.W., Wan, K.H., Park, S., Mendez, I., Galle, S.E., Booth, B.W., Pfeiffer, B.D., George, R.A., Svirskas, R., et al. (2015). The Release 6 reference sequence of the Drosophila melanogaster genome. Genome Res 25, 445-458. Iyer, N.G., Ozdag, H., and Caldas, C. (2004). p300/CBP and cancer. Oncogene 23, 4225-4231. Jain, D., Baldi, S., Zabel, A., Straub, T., and Becker, P.B. (2015). Active promoters give rise to false positive 'Phantom Peaks' in ChIP-seq experiments. Nucleic Acids Res 43, 6959-6968. Jazwinska, A., Kirov, N., Wieschaus, E., Roth, S., and Rushlow, C. (1999a). The Drosophila gene brinker reveals a novel mechanism of Dpp target gene regulation. Cell 96, 563-573. Jazwinska, A., Rushlow, C., and Roth, S. (1999b). The role of brinker in mediating the graded response to Dpp in early Drosophila embryos. Development 126, 3323-3334. Jenuwein, T., and Allis, C.D. (2001). Translating the histone code. Science 293, 1074-1080. Jiang, J., and Levine, M. (1993). Binding affinities and cooperative interactions with bHLH activators delimit threshold responses to the dorsal gradient morphogen. Cell 72, 741-752. Jin, Q., Yu, L.R., Wang, L., Zhang, Z., Kasper, L.H., Lee, J.E., Wang, C., Brindle, P.K., Dent, S.Y., and Ge, K. (2011). Distinct roles of GCN5/PCAF-mediated H3K9ac and CBP/p300-mediated H3K18/27ac in nuclear receptor transactivation. EMBO J 30, 249-262. Jinek, M., Chylinski, K., Fonfara, I., Hauer, M., Doudna, J.A., and Charpentier, E. (2012). A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337, 816-821. Kellum, R., and Schedl, P. (1991). A position-effect assay for boundaries of higher order chromosomal domains. Cell 64, 941-950. Kornberg, R.D. (1974). Chromatin structure: a repeating unit of histones and DNA. Science 184, 868-871. Kornberg, R.D., and Thomas, J.O. (1974). Chromatin structure; oligomers of the histones. Science 184, 865-868.

66

Koutelou, E., Hirsch, C.L., and Dent, S.Y. (2010). Multiple faces of the SAGA complex. Curr Opin Cell Biol 22, 374-382. Kurdistani, S.K., Tavazoie, S., and Grunstein, M. (2004). Mapping global histone acetylation patterns to gene expression. Cell 117, 721-733. Kutach, A.K., and Kadonaga, J.T. (2000). The downstream promoter element DPE appears to be as widely used as the TATA box in Drosophila core promoters. Mol Cell Biol 20, 4754-4764. Kvon, E.Z., Kazmar, T., Stampfel, G., Yanez-Cuna, J.O., Pagani, M., Schernhuber, K., Dickson, B.J., and Stark, A. (2014). Genome-scale functional characterization of Drosophila developmental enhancers in vivo. Nature 512, 91-95. Kwak, H., Fuda, N.J., Core, L.J., and Lis, J.T. (2013). Precise maps of RNA polymerase reveal how promoters direct initiation and pausing. Science 339, 950-953. Lagha, M., Bothma, J.P., Esposito, E., Ng, S., Stefanik, L., Tsui, C., Johnston, J., Chen, K., Gilmour, D.S., Zeitlinger, J., et al. (2013). Paused Pol II coordinates tissue morphogenesis in the Drosophila embryo. Cell 153, 976-987. Lagrange, T., Kapanidis, A.N., Tang, H., Reinberg, D., and Ebright, R.H. (1998). New core promoter element in RNA polymerase II-dependent transcription: sequence-specific DNA binding by transcription factor IIB. Genes Dev 12, 34-44. Landry, J., Sutton, A., Hesman, T., Min, J., Xu, R.M., Johnston, M., and Sternglanz, R. (2003). Set2-catalyzed methylation of histone H3 represses basal expression of GAL4 in Saccharomyces cerevisiae. Mol Cell Biol 23, 5972-5978. Lee, C.K., Shibata, Y., Rao, B., Strahl, B.D., and Lieb, J.D. (2004). Evidence for nucleosome depletion at active regulatory regions genome-wide. Nat Genet 36, 900-905. Levine, M., and Davidson, E.H. (2005). Gene regulatory networks for development. Proc Natl Acad Sci U S A 102, 4936-4942. Li, B., Carey, M., and Workman, J.L. (2007). The role of chromatin during transcription. Cell 128, 707-719. Li, W.X. (2005). Functions and mechanisms of receptor tyrosine kinase Torso signaling: lessons from Drosophila embryonic terminal development. Dev Dyn 232, 656-672. Li, X.Y., Harrison, M.M., Villalta, J.E., Kaplan, T., and Eisen, M.B. (2014). Establishment of regions of genomic activity during the Drosophila maternal to zygotic transition. Elife 3. Lilja, T., Qi, D., Stabell, M., and Mannervik, M. (2003). The CBP coactivator functions both upstream and downstream of Dpp/Screw signaling in the early Drosophila embryo. Dev Biol 262, 294-302. Lim, C.Y., Santoso, B., Boulay, T., Dong, E., Ohler, U., and Kadonaga, J.T. (2004). The MTE, a new core promoter element for transcription by RNA polymerase II. Genes Dev 18, 1606-1617. Lin, C., Garrett, A.S., De Kumar, B., Smith, E.R., Gogol, M., Seidel, C., Krumlauf, R., and Shilatifard, A. (2011). Dynamic transcriptional events in

67

embryonic stem cells mediated by the super elongation complex (SEC). Genes Dev 25, 1486-1498. Liu, C.L., Kaplan, T., Kim, M., Buratowski, S., Schreiber, S.L., Friedman, N., and Rando, O.J. (2005). Single-nucleosome mapping of histone modifications in S. cerevisiae. PLoS Biol 3, e328. Lu, H., Flores, O., Weinmann, R., and Reinberg, D. (1991). The nonphosphorylated form of RNA polymerase II preferentially associates with the preinitiation complex. Proc Natl Acad Sci U S A 88, 10004-10008. Luger, K., Mader, A.W., Richmond, R.K., Sargent, D.F., and Richmond, T.J. (1997a). Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389, 251-260. Luger, K., Rechsteiner, T.J., Flaus, A.J., Waye, M.M., and Richmond, T.J. (1997b). Characterization of nucleosome core particles containing histone proteins made in bacteria. J Mol Biol 272, 301-311. Luse, D.S., and Roeder, R.G. (1980). Accurate transcription initiation on a purified mouse beta-globin DNA fragment in a cell-free system. Cell 20, 691-699. Mali, P., Yang, L., Esvelt, K.M., Aach, J., Guell, M., DiCarlo, J.E., Norville, J.E., and Church, G.M. (2013). RNA-guided human genome engineering via Cas9. Science 339, 823-826. Malik, S., and Roeder, R.G. (2010). The metazoan Mediator co-activator complex as an integrative hub for transcriptional regulation. Nat Rev Genet 11, 761-772. Matsui, T., Segall, J., Weil, P.A., and Roeder, R.G. (1980). Multiple factors required for accurate initiation of transcription by purified RNA polymerase II. J Biol Chem 255, 11992-11996. McKeon, J., Slade, E., Sinclair, D.A., Cheng, N., Couling, M., and Brock, H.W. (1994). Mutations in some Polycomb group genes of Drosophila interfere with regulation of segmentation genes. Mol Gen Genet 244, 474-483. Min, I.M., Waterfall, J.J., Core, L.J., Munroe, R.J., Schimenti, J., and Lis, J.T. (2011). Regulating RNA polymerase pausing and transcription elongation in embryonic stem cells. Genes Dev 25, 742-754. Minami, M., Kinoshita, N., Kamoshida, Y., Tanimoto, H., and Tabata, T. (1999). brinker is a target of Dpp in Drosophila that negatively regulates Dpp-dependent genes. Nature 398, 242-246. mod, E.C., Roy, S., Ernst, J., Kharchenko, P.V., Kheradpour, P., Negre, N., Eaton, M.L., Landolin, J.M., Bristow, C.A., Ma, L., et al. (2010). Identification of functional elements and regulatory circuits by Drosophila modENCODE. Science 330, 1787-1797. Mogno, I., Kwasnieski, J.C., and Cohen, B.A. (2013). Massively parallel synthetic promoter assays reveal the in vivo effects of binding site variants. Genome Res 23, 1908-1915. Moussian, B., and Roth, S. (2005). Dorsoventral axis formation in the Drosophila embryo--shaping and transducing a morphogen gradient. Curr Biol 15, R887-899.

68

Murakami, K., Mattei, P.J., Davis, R.E., Jin, H., Kaplan, C.D., and Kornberg, R.D. (2015). Uncoupling Promoter Opening from Start-Site Scanning. Mol Cell 59, 133-138. Muse, G.W., Gilchrist, D.A., Nechaev, S., Shah, R., Parker, J.S., Grissom, S.F., Zeitlinger, J., and Adelman, K. (2007). RNA polymerase is poised for activation across the genome. Nat Genet 39, 1507-1511. Myer, V.E., and Young, R.A. (1998). RNA polymerase II holoenzymes and subcomplexes. J Biol Chem 273, 27757-27760. Napolitano, G., Lania, L., and Majello, B. (2014). RNA polymerase II CTD modifications: how many tales from a single tail. J Cell Physiol 229, 538-544. Nechaev, S., Fargo, D.C., dos Santos, G., Liu, L., Gao, Y., and Adelman, K. (2010). Global analysis of short RNAs reveals widespread promoter-proximal stalling and arrest of Pol II in Drosophila. Science 327, 335-338. Negre, N., Brown, C.D., Ma, L., Bristow, C.A., Miller, S.W., Wagner, U., Kheradpour, P., Eaton, M.L., Loriaux, P., Sealfon, R., et al. (2011). A cis-regulatory map of the Drosophila genome. Nature 471, 527-531. Neuberger, M.S. (1983). Expression and regulation of immunoglobulin heavy chain gene transfected into lymphoid cells. EMBO J 2, 1373-1378. Nonet, M., Sweetser, D., and Young, R.A. (1987). Functional redundancy and structural polymorphism in the large subunit of RNA polymerase II. Cell 50, 909-915. Ohtsuki, S., Levine, M., and Cai, H.N. (1998). Different core promoters possess distinct regulatory activities in the Drosophila embryo. Genes Dev 12, 547-556. Orphanides, G., Lagrange, T., and Reinberg, D. (1996). The general transcription factors of RNA polymerase II. Genes Dev 10, 2657-2683. Palancade, B., and Bensaude, O. (2003). Investigating RNA polymerase II carboxyl-terminal domain (CTD) phosphorylation. Eur J Biochem 270, 3859-3870. Parkhurst, S.M., Harrison, D.A., Remington, M.P., Spana, C., Kelley, R.L., Coyne, R.S., and Corces, V.G. (1988). The Drosophila su(Hw) gene, which controls the phenotypic effect of the gypsy transposable element, encodes a putative DNA-binding protein. Genes Dev 2, 1205-1215. Pelegri, F., and Lehmann, R. (1994). A role of polycomb group genes in the regulation of gap gene expression in Drosophila. Genetics 136, 1341-1353. Petrij, F., Giles, R.H., Dauwerse, H.G., Saris, J.J., Hennekam, R.C., Masuno, M., Tommerup, N., van Ommen, G.J., Goodman, R.H., Peters, D.J., et al. (1995). Rubinstein-Taybi syndrome caused by mutations in the transcriptional co-activator CBP. Nature 376, 348-351. Rasmussen, E.B., and Lis, J.T. (1993). In vivo transcriptional pausing and cap formation on three Drosophila heat shock genes. Proc Natl Acad Sci U S A 90, 7923-7927. Ratnaparkhi, G.S., Jia, S., and Courey, A.J. (2006). Uncoupling dorsal-mediated activation from dorsal-mediated repression in the Drosophila embryo. Development 133, 4409-4414.

69

Reiter, L.T., Potocki, L., Chien, S., Gribskov, M., and Bier, E. (2001). A systematic analysis of human disease-associated gene sequences in Drosophila melanogaster. Genome Res 11, 1114-1125. Reyon, D., Tsai, S.Q., Khayter, C., Foden, J.A., Sander, J.D., and Joung, J.K. (2012). FLASH assembly of TALENs for high-throughput genome editing. Nat Biotechnol 30, 460-465. Ringrose, L., and Paro, R. (2004). Epigenetic regulation of cellular memory by the Polycomb and Trithorax group proteins. Annu Rev Genet 38, 413-443. Roeder, R.G., and Rutter, W.J. (1969). Multiple forms of DNA-dependent RNA polymerase in eukaryotic organisms. Nature 224, 234-237. Roth, S.Y., Denu, J.M., and Allis, C.D. (2001). Histone acetyltransferases. Annu Rev Biochem 70, 81-120. Roudier, F., Ahmed, I., Berard, C., Sarazin, A., Mary-Huard, T., Cortijo, S., Bouyer, D., Caillieux, E., Duvernois-Berthet, E., Al-Shikhley, L., et al. (2011). Integrative epigenomic mapping defines four main chromatin states in Arabidopsis. EMBO J 30, 1928-1938. Rougvie, A.E., and Lis, J.T. (1988). The RNA polymerase II molecule at the 5' end of the uninduced hsp70 gene of D. melanogaster is transcriptionally engaged. Cell 54, 795-804. Rubinstein, J.H., and Taybi, H. (1963). Broad thumbs and toes and facial abnormalities. A possible mental retardation syndrome. Am J Dis Child 105, 588-608. Saberianfar, R., Cunningham-Dunlop, S., and Karagiannis, J. (2011). Global gene expression analysis of fission yeast mutants impaired in Ser-2 phosphorylation of the RNA pol II carboxy terminal domain. PLoS One 6, e24694. Schneider, D.S., Hudson, K.L., Lin, T.Y., and Anderson, K.V. (1991). Dominant and recessive mutations define functional domains of Toll, a transmembrane protein required for dorsal-ventral polarity in the Drosophila embryo. Genes Dev 5, 797-807. Schotta, G., Ebert, A., Krauss, V., Fischer, A., Hoffmann, J., Rea, S., Jenuwein, T., Dorn, R., and Reuter, G. (2002). Central role of Drosophila SU(VAR)3-9 in histone H3-K9 methylation and heterochromatic gene silencing. EMBO J 21, 1121-1131. Schroder, S., Herker, E., Itzen, F., He, D., Thomas, S., Gilchrist, D.A., Kaehlcke, K., Cho, S., Pollard, K.S., Capra, J.A., et al. (2013). Acetylation of RNA polymerase II regulates growth-factor-induced gene transcription in mammalian cells. Mol Cell 52, 314-324. Schwartz, L.B., and Roeder, R.G. (1975). Purification and subunit structure of deoxyribonucleic acid-dependent ribonucleic acid polymerase II from the mouse plasmacytoma, MOPC 315. J Biol Chem 250, 3221-3228. Sekinger, E.A., Moqtaderi, Z., and Struhl, K. (2005). Intrinsic histone-DNA interactions and low nucleosome density are important for preferential accessibility of promoter regions in yeast. Mol Cell 18, 735-748. Shen, H., and Laird, P.W. (2013). Interplay between the cancer genome and epigenome. Cell 153, 38-55.

70

Shi, D., Pop, M.S., Kulikov, R., Love, I.M., Kung, A.L., and Grossman, S.R. (2009). CBP and p300 are cytoplasmic E4 polyubiquitin ligases for p53. Proc Natl Acad Sci U S A 106, 16275-16280. Shlyueva, D., Stampfel, G., and Stark, A. (2014). Transcriptional enhancers: from properties to genome-wide predictions. Nat Rev Genet 15, 272-286. Shogren-Knaak, M., Ishii, H., Sun, J.M., Pazin, M.J., Davie, J.R., and Peterson, C.L. (2006). Histone H4-K16 acetylation controls chromatin structure and protein interactions. Science 311, 844-847. Smale, S.T., and Baltimore, D. (1989). The "initiator" as a transcription control element. Cell 57, 103-113. Smale, S.T., and Kadonaga, J.T. (2003). The RNA polymerase II core promoter. Annu Rev Biochem 72, 449-479. Sogaard, T.M., and Svejstrup, J.Q. (2007). Hyperphosphorylation of the C-terminal repeat domain of RNA polymerase II facilitates dissociation of its complex with mediator. J Biol Chem 282, 14113-14120. Strahl, B.D., Grant, P.A., Briggs, S.D., Sun, Z.W., Bone, J.R., Caldwell, J.A., Mollah, S., Cook, R.G., Shabanowitz, J., Hunt, D.F., et al. (2002). Set2 is a nucleosomal histone H3-selective methyltransferase that mediates transcriptional repression. Mol Cell Biol 22, 1298-1306. Sun, F.L., and Elgin, S.C. (1999). Putting boundaries on silence. Cell 99, 459-462. Tadros, W., and Lipshitz, H.D. (2009). The maternal-to-zygotic transition: a play in two acts. Development 136, 3033-3042. Takagi, Y., and Kornberg, R.D. (2006). Mediator as a general transcription factor. J Biol Chem 281, 80-89. Takahashi, K., and Yamanaka, S. (2006). Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126, 663-676. Tang, Y., Holbert, M.A., Wurtele, H., Meeth, K., Rocha, W., Gharib, M., Jiang, E., Thibault, P., Verreault, A., Cole, P.A., et al. (2008). Fungal Rtt109 histone acetyltransferase is an unexpected structural homolog of metazoan p300/CBP. Nat Struct Mol Biol 15, 738-745. Thomas, M.C., and Chiang, C.M. (2006). The general transcription machinery and general cofactors. Crit Rev Biochem Mol Biol 41, 105-178. Tie, F., Banerjee, R., Stratton, C.A., Prasad-Sinha, J., Stepanik, V., Zlobin, A., Diaz, M.O., Scacheri, P.C., and Harte, P.J. (2009). CBP-mediated acetylation of histone H3 lysine 27 antagonizes Drosophila Polycomb silencing. Development 136, 3131-3141. Tomancak, P., Berman, B.P., Beaton, A., Weiszmann, R., Kwan, E., Hartenstein, V., Celniker, S.E., and Rubin, G.M. (2007). Global analysis of patterns of gene expression during Drosophila embryogenesis. Genome Biol 8, R145. Upadhyai, P., and Campbell, G. (2013). Brinker possesses multiple mechanisms for repression because its primary co-repressor, Groucho, may be unavailable in some cell types. Development 140, 4256-4265. Valentine, S.A., Chen, G., Shandala, T., Fernandez, J., Mische, S., Saint, R., and Courey, A.J. (1998). Dorsal-mediated repression requires the formation

71

of a multiprotein repression complex at the ventral silencer. Mol Cell Biol 18, 6584-6594. Visel, A., Blow, M.J., Li, Z., Zhang, T., Akiyama, J.A., Holt, A., Plajzer-Frick, I., Shoukry, M., Wright, C., Chen, F., et al. (2009). ChIP-seq accurately predicts tissue-specific activity of enhancers. Nature 457, 854-858. Waltzer, L., and Bienz, M. (1999). A function of CBP as a transcriptional co-activator during Dpp signalling. EMBO J 18, 1630-1641. Wang, L., Tang, Y., Cole, P.A., and Marmorstein, R. (2008a). Structure and chemistry of the p300/CBP and Rtt109 histone acetyltransferases: implications for histone acetyltransferase evolution and function. Curr Opin Struct Biol 18, 741-747. Wang, Z., Zang, C., Rosenfeld, J.A., Schones, D.E., Barski, A., Cuddapah, S., Cui, K., Roh, T.Y., Peng, W., Zhang, M.Q., et al. (2008b). Combinatorial patterns of histone acetylations and methylations in the human genome. Nat Genet 40, 897-903. Weil, P.A., Luse, D.S., Segall, J., and Roeder, R.G. (1979). Selective and accurate initiation of transcription at the Ad2 major late promotor in a soluble system dependent on purified RNA polymerase II and DNA. Cell 18, 469-484. Weinmann, R., Raskas, H.J., and Roeder, R.G. (1974). Role of DNA-dependent RNA polymerases II and III in transcription of the adenovirus genome late in productive infection. Proc Natl Acad Sci U S A 71, 3426-3439. Weinmann, R., and Roeder, R.G. (1974). Role of DNA-dependent RNA polymerase 3 in the transcription of the tRNA and 5S RNA genes. Proc Natl Acad Sci U S A 71, 1790-1794. West, A.G., Gaszner, M., and Felsenfeld, G. (2002). Insulators: many functions, many mechanisms. Genes Dev 16, 271-288. Whitman, M. (1998). Smads and early developmental signaling by the TGFbeta superfamily. Genes Dev 12, 2445-2462. Workman, J.L., and Roeder, R.G. (1987). Binding of transcription factor TFIID to the major late promoter during in vitro nucleosome assembly potentiates subsequent initiation by RNA polymerase II. Cell 51, 613-622. Woychik, N.A., and Hampsey, M. (2002). The RNA polymerase II machinery: structure illuminates function. Cell 108, 453-463. Wyce, A., Xiao, T., Whelan, K.A., Kosman, C., Walter, W., Eick, D., Hughes, T.R., Krogan, N.J., Strahl, B.D., and Berger, S.L. (2007). H2B ubiquitylation acts as a barrier to Ctk1 nucleosomal recruitment prior to removal by Ubp8 within a SAGA-related complex. Mol Cell 27, 275-288. Yuan, G.C., Liu, Y.J., Dion, M.F., Slack, M.D., Wu, L.F., Altschuler, S.J., and Rando, O.J. (2005). Genome-scale identification of nucleosome positions in S. cerevisiae. Science 309, 626-630. Zabidi, M.A., Arnold, C.D., Schernhuber, K., Pagani, M., Rath, M., Frank, O., and Stark, A. (2015). Enhancer-core-promoter specificity separates developmental and housekeeping gene regulation. Nature 518, 556-559.

72

Zaret, K.S., and Mango, S.E. (2016). Pioneer transcription factors, chromatin dynamics, and cell fate control. Curr Opin Genet Dev 37, 76-81. Zeitlinger, J., Stark, A., Kellis, M., Hong, J.W., Nechaev, S., Adelman, K., Levine, M., and Young, R.A. (2007). RNA polymerase stalling at developmental control genes in the Drosophila melanogaster embryo. Nat Genet 39, 1512-1516. Zhao, K., Hart, C.M., and Laemmli, U.K. (1995). Visualization of chromosomal domains with boundary element-associated factor BEAF-32. Cell 81, 879-889. Zheng, C., and Hayes, J.J. (2003). Structures and interactions of the core histone tail domains. Biopolymers 68, 539-546. Zhou, Q., Li, T., and Price, D.H. (2012). RNA polymerase II elongation control. Annu Rev Biochem 81, 119-143.