Download Transcriptional and epigenetic control of gene expression in embryo

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 multicellular 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 addition, these different chromatin states can be linked to distinct modes of Pol II
regulation. Our results provide novel insights into how gene regulatory networks form an epigenetic landscape and how their coordinated actions specify 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 embryo. 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 transcriptional 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 regulatory 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 acetylating the first nucleosome. CBP is regulating Pol II activity of nearly all expressed genes, however, either recruitment or release of Pol II is the ratelimiting step affected by CBP.
Taken together, these results reveal mechanistic insights into cell specification 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., & Mannervik, M. (2012). Preferential genome targeting of the CBP coactivator 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., Nisoli, 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 patterning. (Submitted).
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Abbreviations
Pol II
TSS
PIC
GTF
TF
CTD
CRM
PTM
HAT
HDAC
H3K27ac
H3K27me3
CBP
P300
GAF
RTS
ChIP
PRO-seq
GRO-seq
STARR-seq
FAIRE-seq
PB
DPE
HMT
HDM
PcG
TrxG
PRE
MZT
ZGA
Sog
Brk
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RNA polymerase II
Transcription start site
Pre-initiation complex
General transcription factor
Transcription factor
C-terminal domain
Cis regulatory module
Post-translational modification
Histone acetyl transferase
Histone deacetylase
Histone 3 Lysine 27 acetylation
Histone 3 Lysine 27 methylation
Creb binding protein
300-kDa protein
GAGA-factor
Rubinstein-Taybi syndrome
Chromatin immunoprecipitation
Precision nuclear Run-On and sequencing
Global Run-On and sequencing
Self-transcribing active regulatory region sequencing
Formaldehyde-Assisted Isolation of Regulatory Elements
Pause button
Downstream promoter element
Histone methyl transferase
Histone demethylase
Polycomb Group proteins
Trithorax Group proteins
Polycomb response element
Maternal-to-zygotic transition
Zygotic genome activation
short gastrulation
brinker
TableofContents
Introduction..................................................................................11
Chromatinandtranscription.........................................................12
Transcription.......................................................................................12
EukaryoticDNAiselegantlypackedintochromatin............................13
Transcriptionalregulation...................................................................14
KeyfactorsinTranscription...........................................................15
Pioneerfactorsopenupthechromatin...............................................15
GTFspositionPolIIatthepromoter....................................................16
Co-regulatorsintegrateregulatorysignalsduringtranscription...........17
CBP/p300coactivator...................................................................18
Conservation.......................................................................................18
DomainstructureofCBP/p300............................................................18
TheroleofCBP/p300indisease..........................................................19
TheroleofCBP/p300inregulatinggeneexpression............................20
KeyregulatoryelementsinTranscription......................................22
Promoterelements-determinantsofthemechanismsoftranscriptional
control................................................................................................22
Enhancerelements-akeyregulatoryplatformduringtranscription
control................................................................................................24
Insulatorelements..............................................................................27
PromoterproximalpausingofPolII..............................................28
PolIICTD............................................................................................28
PolIIactivityisregulatedatmultiplestepsduringtranscription.........29
KeyplayersofPolIIpausing................................................................30
ExperimentalproofofPolIIpausing...................................................30
PRO-seq..............................................................................................31
TheroleofPolIIpausing.....................................................................32
Histonemodifications...................................................................33
Chromatinstates.................................................................................34
Theroleofhistonemodificationsintranscription...............................35
Histoneacetylation.............................................................................36
Histoneacetyltransferases(HATs).......................................................36
Histonemethylation...........................................................................37
ChIP....................................................................................................38
Epigenetics....................................................................................39
Establishmentofcellularidentity........................................................39
Stabilityofcellularidentity.................................................................39
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Theepigenomeisplastic.....................................................................40
Development–whenaneggdevelopsintoanorganism...............41
Earlyembryodevelopment.................................................................41
Zygoticgenomeactivation..................................................................42
Patterningoftheembryo....................................................................42
Anterior-posterioraxisformation.......................................................43
Dorsal-ventralaxisformation..............................................................45
Dppsignaling......................................................................................48
Aimofthethesis...........................................................................49
Resultsanddiscussion...................................................................50
PaperI................................................................................................50
PaperII...............................................................................................52
PaperIII..............................................................................................53
PaperIV..............................................................................................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 proliferation and maturation into a complete organism. Each cell in our body contains 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 division 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 introns 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 biological 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 transcriptional 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 processes (Campos and Reinberg, 2009). Chromatin packaging limits the accessibility of the DNA and thereby influence key processes in the cell that utilizes the genetic information including gene regulation, DNA repair and
replication. In addition, the elegant packaging of DNA into the wellstructured 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 remodeling 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 manipulating chromatin and thereby transcription, bringing the field in to the highlight.
chromosome
DNA
histone tails
H3
H4
nucleosomes
chromatin
H2A H2B
histones
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.
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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 influenced by protein-protein interactions and the structure of the local chromatin. 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 promoter 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.
Pause
release
factor
Pausing
factor
Coactivator
Activator
Repressor
PolII
GTF
PolII
PolII
Histone modifications
Corepressor
Transcriptional regulation of a typical eukaryotic gene. Transcription is regulated at several levels; binding of TFs and the recruitment of Coregulators, recruitment
and release of Pol II and at the level of chromatin structure.
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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 accessibility 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 conformation 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 together with nucleotides could synthesize RNA, but these RNAs were synthesized 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 revealed that none of the fractions alone, but the combination of fractions together 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 unwinds 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.
TAF
TAF
TFIIA
TFIIB
TAF
IIA
TBP
TAF
TBP IIB
TATAA
TATAA
TAF
TFIIF
Pol II
TAF
TAF
IIA
IIF
TBP IIB
TATAA
Pol II
TFIIE
TFIIH
IIE
TAF
TAF
IIA
IIF
IIH
Pol II
TBP IIB
TATAA
Formation of the pre-initiation complex (PIC). Stepwise recruitment of GTFs and
Pol II to form the PIC.
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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 complexes. 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: histone modifying enzymes and ATP-dependent remodeling enzymes. Coregulators 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 suggested 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 regulators (Borggrefe and Yue, 2011). In contrast to GTFs, the Mediator has a
global role in stimulating regulated transcription and exists in different subunit 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 acetyltransferases, 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 higher 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 functional and sequence similarities (Tang et al., 2008). During the evolution of
vertebrates, the chromosomal region of CBP/p300 has been duplicated resulting in the p300 gene at chromosome 22 and the CBP gene at chromosome 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 interaction 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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PxP
IBiD
SID
HAT
E1A
p53
NCBD
Q-rich
PHD/
C/H2
bromo
KIX
CREB
IHD
NF B
p53
TAZ1/
C/H1
NRID
Nuclear
receptors
TAZ2/
C/H3
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 homology. For example, both proteins can bind to E1A and CREB but the two
proteins also have distinct functions.
CBP
p300
Model of CBP/p300 domain structure. Nuclear receptor interacting domain
(NRID), Three cysteine/histidine-rich (C/H) domains which also contain transcriptional 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 Rubinstein-Taybi syndrome (RTS) (Petrij et al., 1995). Individuals with RTS exhibit short height, learning difficulties, broad thumbs and big toes and characteristic 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 studies 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 protein (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 inactivating mutations in CBP and p300 have been found in several types of cancers, including small-cell lung cancer, bladder cancer, B cell lymphoma etc.
(Shen and Laird, 2013).
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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, acetylating 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 mapping 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 subsequently 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 transcription factors (TFs), (middle) a scaffolding protein facilitating protein-protein interactions 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 machinery 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 elevated affinity of TFIIB for the core promoter (Lagrange et al., 1998). Initiator (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 promoter, which holds a set of sequence-specific DNA-binding factors that impact 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 activate genes from a DPE containing enhancer and not a TATA containing
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Enhancer 2
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 specificity to a core promoter could be to enhance accurate association of a distant
enhancer and an explicit promoter within a promoter cluster.
HSF
HSE
GAGA
GAGA
Proximal promoter
-250 to 250bp
BRE
TATA
Inr
-37 to -32bp
-30 bp
-2bp
MTE
+18bp
PB
DPE
+25bp
+28bp
Enhancer 3
Core promoter
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 different 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 invagination (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 housekeeping genes and tissue specific expression of developmental genes, respectively (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.
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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 orientation (Banerji et al., 1981). Later studies documented endogenous sequences 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 recruit 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 proposed 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.
24
Coactivator
TF
Enhancer
Pol II
Enhancer
TF
Coactivator
Pol II
Enhancers can contact promoters from far distance. Upon binding of a tissuespecific transcription factor, coactivators are recruited and the enhancer and promoter 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 sequencing 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 enhancers based on correlation with activity, few have actually been functionally tested on a global scale. Predicted enhancers have been tested in embryos by putting the candidate DNA sequence in front of a core promoter followed 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 studies require the generation of transgenic animals and are therefore not suitable
for enhancer screening genome-wide, but recent efforts have mapped thou25
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). Plasmidbased systems have also been used involving placement of the candidate
sequence upstream of a minimal promoter followed by a reporter gene containing a barcode (Mogno et al., 2013). Self-transcribing active regulatory
region sequencing (STARR-seq) brought the functionally testing of enhancers 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 functionally characterized enhancer sequences have provided improved understanding of how regulatory function is encoded in the DNA. The draw back
with plasmid-based systems is that you are not assessing the enhancer activity within the genome of a developing animal and might miss important aspects 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 regulatory 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.
26
Insulator elements
Insulator elements are DNA sequence elements that shield genes from inappropriate 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 involved in structuring chromatin into topological domains with distinct functions (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.
Promoter
Insulator
Blocking
Enhancer
Promoter
Promoter
Insulator
Barrier
Insulator activity. Insulator elements protect genes from inappropriate regulatory
action from the surrounding, (top) by blocking enhancer activity on promoters located behind the insulator element or (low) by acting as a barrier to prevent the spread
of condensed chromatin.
27
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 downstream 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 subunit, 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 polymerase was discovered on SDS-PAGE gels, which revealed two different motilities of Rpb1 (Schwartz and Roeder, 1975). One corresponded to a hypophosphorylated 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).
28
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 clearance, 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 elongation. In addition, the CTD has an important regulatory role in serving as a
platform for other enzymes participating in RNA maturation including protein 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 phosphorylation (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 expression 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.
29
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 recruitment of positive elongation factor-b (P-TEFb) Ser2 of Pol II CTD becomes
phosphorylated as well as NELF (resulting in eviction) and DSIF (transformed 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 composed of cyclin T1 and cyclin-dependent kinase 9 (CDK9). In order for PTEFb 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).
30
Recruitment
Pol II
Re-initiation
Termination
Initiation
S5P
S2P
S5P
Spt5
TFIIH
P
Pol II
Productive
elongation
Pausing
S5P
S2P
PTEF-b
S5P
Spt5
P
S2P
S5P
Spt5
DSIF
NELF
P
The transcription cycle. Pol II is recruited to the promoter followed by TFIIH mediated phosphorylation of Pol II CTD on Serine 5, resulting in transcriptional initiation. 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 adding single biotin-labeled ribonucleotides that are incorporated by transcriptionally 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 further 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 ribonucleotides 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 efficiency or antibody specificity and detects transcriptionally engaged Pol II irrespective of phosphorylation status of Pol II CTD.
31
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 frequently 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. Ultimately, this permissive state would allow the binding of tissue-specific transcription 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 further allow interactions with epigenetic modifiers and transcriptional regulators.
32
Histone modifications
Chromatin modifications influence several nuclear processes including transcription. 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 histones 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). Recent 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.
ac
K36
ac
ac
ac
K13
Ph
K9
S1 K5
me
me
ac
ac Ph ac
me
ac
S28 K36 K56
ac K23 R26 K27
me
ac R17 K18
Ph K14
me Ph
ac S10T11
me
me
K9
me Ph
R8
R2 T3 K4
ac ac
Ph
K20
ac ac S14 K15
K5 K12
Ub
Ph
K119 T120
H2A H2B
me
K79
H3
H4
Ub
K120
ac me
ac
ac ac K12 K16 K20
Ph me
K8
K5
S1 R3
Modifications of the four core histones. Histones are dressed with various modifications including acetylation (ac), methylation (me), phosphorylation (Ph) and ubiquitination (Ub) on several different amino acids.
33
Chromatin states
Simplified, chromatin can be divided into two states, an open/active euchromatic state that is associated with histone acetylation and active transcription, 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.
Euchromatin
Acetylated histones
Heterochromatin
H3K9me3 constitutive
heterochromatin
H3K27me3 facultative
heterochromatin
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, transcription factors, transcripts, replication proteins and nucleosome properties
(Consortium, 2012). The same approach has been applied for model organisms, 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. Computational 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 (enriched in HP1 and H3K9me), BLUE (rich in PcG proteins and H3K27me3)
and BLACK (enriched in non-coding elements, Histone H1 and low tran34
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. Regardless 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 transcription. 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 (ShogrenKnaak et al., 2006). The second model builds upon the existence of a “histone code” (Jenuwein and Allis, 2001). The combination of diverse histone
modification would form different codes. The code would provide synergistic or antagonistic affinities for regulatory proteins that can affect the structure of the chromatin and regulate gene expression. Since information stored
in chromatin can also be inherited through cell division, the role of chromatin 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 transcription 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 provided examples of bivalent domains that posses both the repressive mark
H3K27me3 and the active mark H3K4me3 (Bernstein et al., 2006). A pro-
35
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 transcription 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 consequence 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 acetylation of specific lysine residues on histones. By contrast histone deacetylases (HDACs) removes acetyl groups from lysines resulting in a closed chromatin conformation. The first protein reported to have HAT activity was the
transcriptional adaptor Gcn5 (Brownell et al., 1996). Gcn5 is part of the multisubunit 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).
36
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 multiprotein 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 expression 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 transcriptional state depending on where the methylation mark is located. Histone methyl transferases (HMTs) mediate the addition of methyl groups
from S-adenosylmethionine to lysine that can be both mono- di- and trimethylated. By contrast, histone demethylases (HDMs) are responsible for
the removal of methyl marks. Methylation of H3K9 is mediated by suppressor of variegation Su(var)3-9 (but also other HMTs) resulting in recruitment 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 transcription 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 assigned to PcG and TrxG proteins, and they tend to act in large protein complexes (Ringrose and Paro, 2004). Enhancer of zeste E(z) mediate the histone 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 conserved core of proteins between Drosophila and human, but also unique
accessory proteins. For example, the three sequence specific DNA-binding
37
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 important in cellular memory.
ChIP
Chromatin immunoprecipitation (ChIP) is a widely used method for identifying binding sites of chromatin proteins. It involves crosslinking of your favorite 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 independent antibodies that generate consistent results. A recent report has identified 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.
38
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. During 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 transcription 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
39
unknown factors in one cell can reprogram the fusion partner’s genome and
induce cell-type specific gene activation (Blau et al., 1983). Forced expression of transcription factors can cause transdifferentiation, i.e. reprogramming that result in a switch of cell identity. Misexpression of myoblast determination 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 modifications. One such example is the body segment identity in Drosophila that is
controlled by homeobox (hox) and engrailed genes. Expression of key transcription 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 regulation of several genes in the hierarchy that set up hox gene expression
(McKeon et al., 1994) (Pelegri and Lehmann, 1994). This implies that transcription factor cascades may be supported by chromatin regulatory mechanisms 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 rearrangement 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 heterochromatin (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 reprogrammed 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).
40
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, Drosophila 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 diffuse across nuclei during early Drosophila development that is of great importance 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
41
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 plasma 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 maternal-to-zygotic transition (MZT). The MZT can be divided into two distinct
chapters, degradation of maternal products and activation of zygotic transcription (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 different 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 second 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 morphological changes
Patterning of the embryo
The importance of maternally provided RNA and proteins are well illustrated during patterning of the embryo. The principle behind animal embryo
pattern formation is that initially equivalent cells in a developing tissue assume 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
42
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 transcription 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 pattern 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 expression is precisely limited to seven defined stripes perpendicular to the anterior-posterior axis by the repressive mechanism of gap genes. This tight regulation 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 segmentpolarity genes. The 14-stripe expression pattern of segment polarity genes is
the precursors of body segments that finally acquire their unique character.
43
Patterning of the anterior-posterior axis in early Drosophila embryos. A hierarchy 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 segment-polarity genes into 14 stripes.
44
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 translocated 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 proteases Nudel (Ndl), Gastrulation-defective (Gd), Snake (Snk) and Easter (Ea)
resulting in cleaved and active Spätzle (Spz) ligand that binds to Toll receptor 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 generate 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 lowaffinity 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 ubiquitously 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 involves the interaction with a protein that binds to AT-rich sequences flanking the Dorsal site. This result in a conformational change of Dorsal that
expose a cryptic peptide motif (Ratnaparkhi et al., 2006) mediating the interaction 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
45
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 ectoderm in the entire embryo.
Taken together, Dorsal controls diverse transcriptional outputs by the combined effort of different affinity binding sites, number of binding sites and
the co-binding with other transcription factors to mediate the many transcriptional outputs (Jiang and Levine, 1993).
The molecular mechanisms by which tissue-specific gene expression patterns are set up and how they are maintained has been a long-standing question and is the topic of paper I.
46
gd7
Dorsal
gradient
sna
sog
twi
brk
Dl
tld
Dl
brk
Dl
neuroectodern
dpp
Dl
Sna
Dl
Sna
sog
Dl
tld mesoderm
brk
Dl
dpp
Dl
twi
sna
Dl
twi
dorsal
ectoderm
dpp
sog
sna
Dl
tld
Tollrm9/10
Toll10B
Toll
Spätzle
Easter
Snake
Gd
Nudel
Pipe
Patterning of the dorsal-ventral axis in early Drosophila embryos. Maternally
provided Dorsal is present in a gradient with highest concentration on the most ventral side. Dorsal activates genes in a concentration dependent manner that subdivides
the embryo into mesoderm, neuroectoderm and dorsal ectoderm. The Dorsal gradient is established by ventral localized Pipe that triggers a proteolytic cascade involving Nudel, Gastrulation-defective (Gd), Snake and Easter resulting in cleaved and
active Spätzle ligand that binds to Toll receptor. Mutants in the Toll signaling pathway that have a homogenous population of cells with respect to Dorsal concentration 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.
47
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 imaginal 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 Smadsignaling where the class I Smad protein Mothers against dpp (Mad) becomes phosphorylated by Tkv and thereby associated with the class II coSmad Medea (Whitman, 1998). The two Smads are translocated to the nucleus 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 action 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).
48
Aim of the thesis
The main objective of the thesis is to understand transcriptional and epigenetic 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 ectoderm, neuroectoderm and mesoderm in response to the Dorsal morphogen? (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 pausing? (Addressed in Paper IV).
49
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 transcription 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 competency for gene activity or repression by opening the chromatin to transcription
factors and histone modifying enzymes. Our results indicate that some pioneer factors could mediate the establishment of a poised polymerase during
the set up of different gene expression patterns. Several outstanding questions 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 setting 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 competition 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
50
to further dissect by what mechanism Snail is modulating Dorsal activity.
Furthermore, we find that in the dorsal ectoderm, these genes are held inactive 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. Interestingly, 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 maintenance 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 programming initiate diverse epigenetic states to orchestrate embryogenesis.
51
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 multiple 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 networks controlling dorsal-ventral patterning of the embryo. Hundreds of interaction 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 dependent at sites where few factors are bound. In the absence of Dorsal, CBP binding 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 understood. 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 acetylation. Given that acetylation and methylation of H3K27 is mutually exclusive, 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 increase 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 development by coordinating the dorsal-ventral rather than for example the anteriorposterior axis of the embryo. CBP is also found at hypoacetylated silent
genes. Like many other HATs, CBP is a big multidomain protein with several interaction opportunities with multiple regulatory factors, and could perhaps act as a scaffolding protein to regulate gene expression.
52
Paper III
CBP is a widely used co-activator with HAT activity but its full role in regulating 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 transcription. In addition we find that the strongest CBP binding is found at Polycomb 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 activity 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 regulates its insulating activity by acetylating histones to prevent spreading of
H3K27me3. In summary, CBP is bound to many genomic regions with different 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.
53
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 transcriptional 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 expressed 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 expressed 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 promoters by investigating if CBP is acetylating key transcriptional regulators
and examine how acetylation influence their action.
54
Conclusion and perspectives
A few final thoughts on transcription and epigenetic control during development are as follows. The strategies to regulate transcription are both complex and numerous, mediating the amazing diversity of life. While our understanding of how genes are regulated has greatly advanced, it has become
evident that chromatin and transcription are two tightly linked key determinants in this process. Upon fertilization, the specialized germ cell is undressed 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 other tissue specific factors can bind and recruit cofactors in order to initiate
gene expression programs. The commencement of cell specification is accompanied by distinct histone modifications, first by acetylation and then by
methylation that together form chromatin states that maintain proper expression 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 dorsalventral 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 highresolution 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 transcription 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 transcriptional regulation a growing understanding of transcriptional misregulation is also being recognized. Small molecule inhibitors targeting key components in transcriptional control have already shown promising results as
potential therapeutic targets in cancer.
55
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 support, 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 endless 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 generosity 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 contributing 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.
56
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 encouragement 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.
57
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 enzym 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 egenskaper, 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. Celler 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 organism 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 används för att etablera och bibehålla specifika genuttrycksprogram under embryoutvecklingen. Många cellulära processer inklusive hur genuttryck kontrolleras 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 embryoutveckling. 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 acetylerar histoner och leder till aktivt genuttryck. Vidare har vi påvisat två mekanismer som tystar gener i dessa vävnader, antingen genom Polycomb medierad metylering av histone 3 lysin 27 eller genom hypoacetylerade histoner
58
etablerade av Snail. Vi kan dessutom länka de olika histone modifieringsmönstren med distinkta sätt att reglera Polymeraset. Våra resultat tillhandahåller 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 tillhandahåller 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 princip 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 transkriptionell och epigenetisk kontroll av genuttryck. Att kartlägga de underliggande mekanismerna som reglerar genuttryck är essentiellt för att förstå
hur cancerceller skiljer sig mot vanliga celler.
59
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