ATPase dependent and independent roles of Brahma in

ATPASE DEPENDENT AND INDEPENDENT ROLES
OF BRAHMA IN TRANSCRIPTION AND PRE-MRNA
PROCESSING
Simei Yu
ATPase dependent and independent
roles of Brahma in transcription and
pre-mRNA processing
Simei Yu
©Simei Yu, Stockholm University 2015
ISBN 978-91-7649-245-1
Printed in Sweden by Holmbergs, Malmö 2015
Department of Molecular Biosciences, The Wenner-Gren Institute
To my family
Abstract
SWI/SNF is a chromatin-remodeling complex and Brahma (BRM) is the ATPase
subunit of SWI/SNF. BRM regulates transcription by remodeling the nucleosomes
at promoter regions. BRM is also associated with RNA and affects pre-mRNA processing together with other SWI/SNF subunits. In this thesis, I will discuss the roles
of BRM in both transcription and pre-mRNA processing. In Paper I, we showed
that BRM, as well as other SWI/SNF subunits SNR1 and MOR, affects the alternative processing of a subset of pre-mRNAs, as shown by microarray analysis. This
observation was validated by RNAi experiments both in Drosophila S2 cells and in
vivo. In Paper II, we characterized the trans-splicing of transcripts derived from the
mod(mdg4) gene. RNA interference (RNAi) and overexpression experiments revealed that BRM regulates the trans-splicing of mod(mdg4)-RX in an ATPase independent manner. In Paper III, we analyzed the expression of two BRM-target genes
identified in Paper I, CG44250 and CG44251. RNAi and overexpression experiments showed that the expression levels of these two genes were affected by BRM
in a manner that is independent of its ATPase activity. Transcriptome analysis further proved that BRM affects gene expression both in ATPase dependent and independent manners. In Paper IV, we showed that BRM is present at the 3’-end of two
analyzed genes, CG5174 and CG2051. BRM facilitates the recruitment of the cleavage and polyadenylation machinery to the cleavage sites through protein-protein
interactions that do not require the ATPase activity of BRM. Morevoer, BRM promotes the cleavage of the CG5174 and CG2051 pre-mRNAs. To sum up, SWI/SNF
plays important roles not only in transcription but also in pre-mRNA processing. To
regulate transcription, BRM can either act as an ATPase-dependent chromatin remodeler or in a manner that does not involve ATPase activity. Additionally, BRM
interacts with RNA-binding proteins to regulate the processing of a subset of premRNAs, and this function of BRM is independent of its chromatin remodeling activity.
List of articles included
Waldholm J, Wang Z, Brodin D, Tyagi A, Yu S, Theopold U, Östlund Farrants AK, Visa N. (2011). SWI/SNF regulates the alternative processing of a
specific subset of pre-mRNAs in Drosophila melanogaster. BMC Mol Biol
12:46.
Yu S, Waldholm J, Böhm S, Visa N. (2014). Brahma regulates a specific
trans-splicing event at the mod(mdg4) locus of Drosophila melanogaster.
RNA Biol 11(2):134-45.
Yu S, Waldholm J, Jordán-Pla A, Källman T, Östlund Farrants AK, Visa N.
The ATPase-dependent and ATPase-independent functions of Brahma in
transcription regulation.
Manuscript.
Yu S, Gàñez-Zapater A, Jordán-Pla A, Rolicka A, Östlund Farrants AK,
Visa N. A role for SWI/SNF in pre-mRNA 3’-end processing.
Manuscript.
Contents
Abbreviations ................................................................................................. 9 Introduction .................................................................................................. 11 Nucleosome ................................................................................................................. 11 Chromatin remodeling ................................................................................................. 12 Histone modification .................................................................................................... 12 Histone acetylation ................................................................................................. 13 Histone methylation ................................................................................................ 14 Other histone modifications .................................................................................... 14 ATP-dependent chromatin remodeling complex.......................................................... 15 SWI/SNF chromatin remodeling complexes........................................................... 15 ATPase dependent and ATPase independent roles of BRM ................................. 17 Other chromatin remodeling enzymes ................................................................... 18 Non-coding RNAs and chromatin ........................................................................... 19 Transcription ................................................................................................................ 20 Core promoter elements......................................................................................... 20 Transcription initiation ............................................................................................ 21 Transcription elongation and termination ............................................................... 22 Pre-mRNA Splicing ...................................................................................................... 23 Regulation of splicing and alternative splicing ............................................................. 24 Cis- and trans-splicing ................................................................................................. 27 The Mod(mdg4) gene .................................................................................................. 29 Pre-mRNA 3’-end processing ...................................................................................... 29 Alternative cleavage and polyadenylation .............................................................. 30 Cleavage factor Im ................................................................................................. 31 The present investigation ............................................................................. 33 Aim of the studies ........................................................................................................ 33 Model System .............................................................................................................. 33 High throughput methods ............................................................................................ 34 Project summary .......................................................................................... 36 Paper I ......................................................................................................................... 36 Paper II ........................................................................................................................ 38 Paper III ....................................................................................................................... 40 Paper IV ....................................................................................................................... 44 Conclusions ................................................................................................. 47 Acknowledgements ...................................................................................... 48 References ................................................................................................... 49 Abbreviations
BRG1– Brahma-related gene 1
BRE – TFIIB recognition element
BRM – Brahma
CA – Cleavage site
CFIm – Cleavage factor I
CFIIm – Cleavage factor II
CHD – Chromodomain helicase DNA-binding
CPSF – Cleavage and polyadenylation specificity factor
CstF – Cleavage stimulation factor
CTCF – CCTC-binding factor
CTD – C-terminal domain of the largest subunit of RNA polymerase II
DPE – Downstream promoter element
Dscam – Down syndrome cell adhesion molecule
Gpdh – Glycerol 3 phosphate dehydrogenase
HAT – Histone acetyltransferase
HDAC – Histone deacetylase
HSA – Helicase-SANT–associated domain
hnRNP – Heterogeneous nuclear ribonucleoprotein
Inr – Initiator
ISWI – Imitation switch
lola – longitudinals lacking
microRNA – miRNA
mod(mdg4) – modifier of mdg4
Mor – Moira
MRG15 – Mortality factor on chromosome 4-related gene on chromosome 15
NGS – Next generation sequencing
ncRNA – non-coding RNA
PAS – Polyadenylation site
PIC – Preinitiation complex
Pol-II – RNA Polymerase II
9
pre-mRNA – Precursor messenger RNA
PTB – Polypyrimidine tract-binding protein
QLQ – Glutamine-Leucine-Glutamine domain
SAM68 – Src-associated protein in mitosis 68 kDa
Small interfering RNA – siRNA
Snr1 – Snf5-related 1
snRNP – Small nuclear ribonucleoprotein
SR protein – Serine/Arginine rich protein
SRp20 (SRSF3) – Serine/Arginine splicing factor 3
SWI/SNF – SWItch/Sucrose NonFermentable
TF – Transcription factors
TSS – Transcription start site
10
Introduction
Gene expression is the process by which the information from a gene is used to
synthesize a functional gene product such as a protein or a functional RNA. There
are a series of steps in the gene expression process: transcription, RNA processing,
RNA export, translation and post-translational events. The transcription of a proteincoding gene results in the synthesis of a pre-mRNA. The pre-mRNA is then processed to mRNA. Splicing is a step in the pre-mRNA processing by which introns
are removed and exons joined. The cleavage and polyadenylation machinery catalyzes the pre-mRNA cleavage and polyA tail synthesis at the 3’end of nascent transcripts. The components of these machineries interact with each other and coordinate the different steps of gene expression.
Chromatin structure also contributes to the mechanism of transcription regulation.
Chromatin remodeling complexes with ATPase activity change histone-DNA interactions and affect transcription factor binding through the change of nucleosome
structure (Luo and Dean, 1999). Recent studies showed that chromatin remodeling
has an impact not only on transcription, but also on the processing of the premRNAs. For example, it has been revealed that SWItch/Sucrose NonFermentable
(SWI/SNF) chromatin remodeling complex regulates the abundance of alternative
splicing transcripts in Drosophila (Tyagi et al., 2009). In the present study, I aim at
understanding the links between chromatin remodelling, transcription and premRNA processing, and how these steps are regulated by SWI/SNF.
Nucleosome
In the cell nucleus, DNA is highly compacted into chromatin to fit the small size of
the nucleus. Chromatin is composed of nucleosomes. The nucleosomes consist of a
core of histones that is wrapped around twice by approximate 147 bp of DNA. The
histone core is an octamer that contains two copies of each H2A, H2B, H3 and H4
(Luger et al., 1997). Nucleosomes are linked by 10-70 bp of linker DNA and assembled into chromatin. Nucleosome structure is highly conserved among eukaryotic
species (McGhee et al., 1980). Nucleosome structure is dynamic and this process is
11
important for DNA replication, repair and transcription to occur. Nucleosomes transiently unwrap and naked DNA is exposed for DNA binding factors (Ngo et al.,
2015). Work from Ngo and coworkers indicated that the unwrapping of DNA was
not symmetrical and that the direction of the unwrapping was decided by the DNA
sequence.
Chromatin remodeling
In eukaryotes, RNA polymerases and transcription factors are needed for transcription to occur. However, the highly condensed structure of chromatin limits the access of these proteins to the DNA. Thus, chromatin remodeling is a necessary process during which DNA segments are exposed and can interact with DNA binding
proteins. The complexes that regulate chromatin structure can be divided into two
major groups: one is ATP-dependent chromatin remodellers and the other is histonemodifying enzymes, for example histone acetyltransferases and histone deacetylases
(Vignali et al., 2000). The ATP-dependent factors use the energy from ATP hydrolysis to modify the association of histones with DNA, and the second group regulates
expression by determining the properties of the nucleosomal histones (Mohrmann
and Verrijzer, 2005). In addition to these two major groups, recent studies reported
that small RNAs together with RNA binding protein Argonaute (Ago) also function
in chromatin remodeling and transcription regulation. The RNA binding protein
Ago can be recruited to promoter regions guided by microRNAs (miRNAs) and
promote the binding of other chromatin modifying proteins, and remodel chromatin
structure through heterochromatin formation or histone modification (Li, 2014).
Histone modification
According to the different packing states, chromatin can be divided into euchromatin and heterochromatin. Euchromatin is a lightly packed form of chromatin and
often under active transcription, whereas heterochromatin is more condensed and
transcriptionally inactive. Histone tails protruding from the octamer could be modified in many ways post translationally, such as acetylation, methylation, phosphorylation and ubiquitination (Figure 1). These modifications perform as a platform to
12
recruit transcription factor or chromatin remodeling complexes, thereby regulating
nucleosome stability and dynamics (Tessarz and Kouzarides, 2014).
Figure 1. Post-translationally
modified histone tails. DNA
is wrapped around core histones and is represented as a
thick brown line. Histone
tails are represented as thin
black lines. Different modifications at histone residues
are represented as purple
squares, yellow stars and
blue circles.
Histone acetylation
Histone acetylation is the best characterized modification. It is mediated by histone
acetyltransferases (HATs) and occurs on lysine residues. Histone acetylation takes
place both on tails and internal histones. Histone 3 is commonly modified. Acetylation of histones is often enriched in euchromatin and participates in the regulation of
transcribed genes. For instance, acetylation of the lysine 27 in histone H3
(H3K27ac) distinguishes active from inactive enhancers and acts as a marker of
active transcription (Creyghton et al., 2010). H3K56ac within DNA Entry-Exit region of nucleosome might facilitate the unwrapping of DNA from histone, increase
DNA breathing and regulate chromatin structure accordingly (Simon et al., 2011;
Neumann et al., 2009). H3K122ac and H3K64ac have a role in active transcription
by decreasing nucleosome stability. H3K64ac was also shown to change DNAhistone octamer interaction and evict histone at promoter regions (Simon et al.,
2011; Cerbo et al., 2014). Acetylation is not only modified at lysines of histone H3
13
but also found in other histones such as H4, H2A and H2B (Berger, 2007). Histone
acetylation is not a permanent state. These acetylation marks can be removed by
histone deacetylases (HDACs), which usually leads to transcriptional repression
(Xhemalce et al., 2011).
Histone methylation
Histone methylation occurs in lysine and arginine residues, functioning as a mark of
either active or repressed transcription. There are three possibilities of each methylation: mono, di or tri-methylation. Methylation occurs at different regions. H3K4me
is enriched at enhancers and is correlated with transcriptional activation. Genome
wide studies indicated H3K4me3 marks are enriched at transcription initiation sites,
while H3K9me3 and H3K27me3 are detected mainly at transposons and repetitive
sequences (Yin et al., 2011). H3K9me3 is the heterochromatin protein 1 binding
target and contributes to heterochromatin formation (Hublitz et al., 2009). Histone
methylation marks are also involved in euchromatin-heterochromatin structure
maintenance and distinction. H3K9me2 was not found at 5’-end regions but enriched at euchromatin-heterochrmatin transition regions (Yasuhara and Wakimoto,
2008). Apart from lysine methylation, arginine methylation also affects nucleosome
stability. For examples, the asymmetric form of H3R42me2 disrupts histone and
DNA interactions and makes chromatin more accessible to the transcription machinery accordingly (Casadio et al., 2013).
Other histone modifications
There are also other histone modifications such as phosphorylation, ubiquitination
and isomerization. Phosphorylation of H3T118 is involved in nucleosome disassembly and increases DNA accessibility (North et al., 2011). H3S10ph has an impact on Gcn5 mediated H3 acetylation and promotes Gcn5-dependent gene transcription (Lo et al., 2000). H2A ubiquitination plays a negative role in transcription
initiation and represses the formation of H3K4 di and tri-methylation (Nakagawa et
al., 2008). However, H2B ubquitination is associated with RNA polymerase and
facilitates transcription elongation (Xiao et al., 2005).
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Many histone modifications contribute to regulate the gene expression levels
through the recruitment of chromatin-associated factors (Bannister and Kouzarides,
2011). These histone modifications are not individual marks but compose histone
codes in a complex manner (Jenuwein and Allis, 2001). Some of the histone modification considered as a negative mark can also define activated chromatin depending
on the chromatin context. H3K4me3 is a positive mark for transcription. The combination of H3K4me3 and negative mark H3K9me3 at open reading frames indicated a dynamic status of transcription. The combination of H3K4me3 and negative
mark H3K27me3 in embryonic stem cells defined a region that is poised for transcription (Berger, 2007). All these examples above suggest histone marks are no
longer single “on” or “off” codes but a complex “language” depended on the chromatin context.
ATP-dependent chromatin remodeling complex
Currently, four different families of ATP-dependent chromatin remodeling enzymes
are identified: SWI/SNF, ISWI, CHD and Ino80 (Swygert and Peterson, 2014).
Each family is defined by the presence of a unique ATPase subunit.
SWI/SNF chromatin remodeling complexes
SWI/SNF (switch/ Sucrose nonfermentable) is a 2MDa complex that was initially
identified by the isolation of mutants in yeast (Carlson and Laurent, 1994).
SWI/SNF is an ATP-dependent chromatin-remodeling complex that contains 8-10
subunits (Figure 2) (Dingwall et al., 1995; Mohrmann and Verrijzer, 2005). In yeast,
insects and human cells, there are at least two different SWI/SNF complexes
(Mohrmann and Verrijzer, 2005). Each complex contains one distinct ATPase subunit and several common or specific subunits. In Drosophila, there is only one
ATPase subunit Brahma (BRM). In human cells, there are two ATPase subunits:
BRM and Brahma-related gene 1 (BRG1). Previous work indicated that BRM is
found in two different SWI/SNF complexes: BAP and PBAP. They share 7 subunits, and in addition BAP contains OSA, whereas PBAP contains Polybromo and
BAP170 and SYAP (Kal et al., 2000; Mohrmann et al., 2004; Papoulas et al., 1998;
Chalkley et al., 2008). BRM contains several domains: ATPase domain was first
15
identified in yeast (Laurent et al., 1993) Bromo domain was shown to recognize and
bind to the acetylated lysines from histone H3 and H4 (Singh et al., 2007). Apart
from these two domains, other domains of BRM are also involved in protein interaction or DNA binding (Figure 3) (Marmorstein, 2001). For instance, the helicaseSANT–associated (HSA) domain acts as a platform and facilitates the binding of
nuclear actin-related proteins. Through this interaction, HSA regulates the ATPase
activity of the chromatin-remodeling complex (Szerlong et al., 2008). There was no
effect on BRM function and chromatin binding when deleting its bromodomain.
However, the HSA domain instead of bromodomain was found to be essential for
BRM function and for the stability of the BRM complex in Drosophila (Elfring et
al.,1998).
BAP
OSA
PBAP
BAP
170
BRM
SAYP
SNR1
MOR
BRM
SNR1
PB
MOR
Figure 2. The BAP and PBAP complexes of Drosophila melanogaster. These two
complexes have the same core subunits: BRM, Moira and SNR1. Signature subunit
OSA is found in BAP, whereas BAP170, Polybromo and SAYP are characteristic
of the PBAP complex.
Figure 3. The Brahma domains annotated by Uniprot. There are five domains
described in the protein: Glutamine-Leucine-Glutamine domain(QLQ), HelicaseSANT–associated domain (HSA), ATPase domain, helicase C-terminal domain
and Bromo domain.
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Previous findings provide direct evidence that SWI/SNF complexes regulate many
genes by remodeling nucleosomes at promoter regions (Bouazoune and Brehm,
2006). Genome wide studies suggest that human SWI/SNF core subunits co-localize
with RNA polymerases extensively and have higher occupancy at promoters, enhancers and CTCF motifs (Euskirchen et al., 2011). SWI/SNF complexes have dual
roles either in transcription activation or repression. For example, BRM has been
proved to associate with active RNA polymerase II (Armstrong et al., 2002), and
regulate the homeotic genes (Kal et al., 2000). BRM has been reported associated
with transcription initiation factor TFIID and involved in active transcription (Vorobyeva et al., 2009). BRM was also observed to act as a negative regulator of transcription. Recent studies revealed that BRM increases nucleosome occupancy and
limits transcription output (Kwok et al., 2015). There are a few genes that are not
affected by BRM, for example the induced heat shock genes. BRM is not associated
with these genes and their expression is not influenced by BRM loss function (Armstrong et al., 2002). Other components of SWI/SNF have an effect on BRM function. Snf5-related 1 (SNR1) was found to inhibit BRM activity in a cell type specific
manner and therefore repress transcription (Marenda et al, 2004). Collins and
Treisman revealed that OSA overexpression repressed wingless regulated genes
through SWI/SNF complex (Collins and Treisman, 2000). Apart from transcription,
SWI/SNF has other functions. Co-purification of SWI/SNF subunits has led to the
identification of proteins associated with other aspects of gene expression, such as
mRNA processing, DNA replication and DNA repair (Euskirchen et al., 2011).
BRG1 is recruited to DNA double strand breaks and promotes homologous recombination in mammalian cells (Qi et al., 2015). BRM is known to be involved in cell
viability and development. For example, expression of dominant-negative BRM
caused nervous system defects (Elfring et al., 1998). Loss of BRM impairs intestinal
stem cell proliferation and differentiation through Hippo pathway that regulates cell
proliferation and apoptosis in Drosophila (Jin et al., 2013).
ATPase dependent and ATPase independent roles of BRM
SWI/SNF is known to destabilize the DNA-histone interaction using energy from
ATP hydrolysis. However, studies from both our and other groups indicated that
17
BRM could regulate transcription and pre-mRNA processing in an ATPase independent manner. Studies from Kwok et al. (2015) suggested that BRM increased
nucleosome density in an ATPase dependent manner. They also observed that BRM
influences gene expression through a mechanism that does not involve the ATPase
activity of BRM. This conclusion was based on experiments using a catalytically
inactive mutant, BRM-K804R. The BRM ATPase mutant may facilitate the binding
of repressors and pausing factors to promoters and lead to a pausing of RNA polymerase without changing the nucleosome occupancy in these regions.
Our studies further proved the conclusion that BRM can regulate transcription
through an ATPase-independent mechanism (see Paper III). We found that BRM
regulates the transcription of the CG44250/44251 genes in an ATPase independent
manner. RNA sequencing analysis data showed that 1031 genes were affected significantly when we overexpressed BRM wild-type, and 770 of them were affected in
an ATPase independent manner.
Other chromatin remodeling enzymes
The ISWI family contains three complexes: NURF, ACF and CHRAC. All these
complexes contain an ISWI subunit, an ATP-dependent motor that alters nucleosome structure. ISWI has a role in transcription repression and chromosome structure maintenance. ISWI mutations alter the structure of the male X chromosome in
Drosophila (Deuring et al., 2000).
The CHD family in Drosophila includes four complexes: dCHD1, dCHD3, dNuRD
and Kismet (Murawska and Brehm, 2011). CHD chromatin remodelers are involved
in either transcription activation or repression. Genome wide studies revealed that
CHD1 plays a role in nucleosome disassembly at promoters (Walfridsson et al.,
2007). NuRD facilitates histone H3K27 deacetylation and acts as a transcription
repressor (Reynolds et al., 2012). Moreover, similar to SWI/SNF complex, the
CHD chromatin remodelers are also involved in pre-mRNA processing. The human
CHD1 can be co-purified with the U2snRNP spliceosomal complex, and depletion
of CHD1 decreases splicing efficiency in human cells (Sims et al., 2007).
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The INO80 complex has multiple functions in transcription regulation, DNA repair
and DNA replication. Microarray studies revealed that the transcription of up to
20% of genes in yeast was either up or down regulated by INO80 (Jónsson et al.,
2004). INO80 has also been found to localize to DNA double strand breaks, and
depletion of INO80 caused a DNA repair defect (Gospodinov et al., 2011).
Non-coding RNAs and chromatin
Apart from messenger RNA, there are RNAs transcribed from DNA but not translated into proteins. These RNAs are functional and known as non-coding RNAs
(ncRNAs). The functions of tRNA and rRNA in protein synthesis have been known
since long. Small nuclear RNA (snRNA) plays an important role in splicing and
were identified long ago (Rogers and Wall, 1980). Other ncRNAs have been discovered more recently and their functions are intensely studied at present. These include
small interfering RNA (siRNA), microRNA (miRNA), piwi-interacting RNA (piRNA) and different types of long ncRNAs (lncRNA). Many of these ncRNAs play a
role in the regulation of chromatin structure (Mattick and Makunin, 2006; Böhmdorfer and Wierzbicki, 2015).
miR-320 is a conserved miRNA transcribed from POLR3D antisense strand in
mammalian cells. Recent evidence suggests that miR-320 guides the RNA-binding
protein Ago to the POLR3D promoter region, which further recuits the Polycomb
group component EZH2. As a result, H3K27 becomes methylated (H3K27me3) in
this region and transcription is silenced (Kim et al., 2008). In another study, siRNA
and Ago1 were found to be targeted to CCR5 promoters and caused gene silencing
through heterochromatin formation in human cells (Kim et al., 2006). The long noncoding RNA HOTAIR interacts with Polycomb group component PRC2, induces
H3K27 methylation, and silence sthe transcription of the HOXD locus (Rinn et al.,
2007). On the contrary, miRNAs have also been reported as activators of gene expression. For example, miRNA-373 triggered E-cadherin and CSDC2 expression,
and Pol-II enrichment was observed at promoters of targeted genes (Place et al.,
2008). miRNA-744 induced cyclin B1 expression and increased polymerase as well
as H3K4me3 occupancy at the transcription start region (Huang et al., 2012). miRNA-589 mediates COX-2 gene activation together with the RNAi factors Ago2 and
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GW182 (Matsui et al., 2013). In summary, an increasing number of evidences suggested that ncRNAs are involved in chromatin modification and gene expression
through a large variety of mechanisms.
Transcription
Transcription of protein-coding gene in eukaryotes is catalyzed by RNA polymerase
II (Pol-II). Pol-II uses one strand of the DNA as the template and synthesizes a
complementary strand of RNA. It is a 550KDa complex that contains 12 subunits
(Liu et al., 2013). The carboxy-terminal domain of the largest subunit of Pol-II
(CTD) contains a repeat heptapeptide sequence: Y1-S2-P3-T4-S5-P6-S7. The number of repeats correlates to the genomic complexity of the organism. There are 26
repeats in yeast, 32 in C.elegans and 45 in Drosophila (Hsin and Manley, 2012).
The phosphorylation state of S2 and S5 regulates progression through the different
phases of transcription (Srivastava and Ahn, 2015).
Core promoter elements
Genes transcribed by RNA Pol-II contain sequences that are recognized by transcription machinery. Four major promoter elements functioning in transcription
regulation have been reported: TATA box, TFIIB recognition element (BRE), initiator (Inr) and downstream promoter element (DPE) (Kadonaga, 2012).
The TATA box, one of the most characterized core promoter elements, contains a
T/A-rich sequence that is located approximately 20-30 nucleotides upstream of the
transcription start site (TSS). In Drosophila, the TATA box appears in around 43%
of the genes (Kutach and Kadonaga, 2000). The BRE motif is located either upstream or downstream of the TATA box. This motif was reported to activate or
repress transcription (Deng and Roberts, 2005). The Inr is located at the +1 of TSS.
According to the genome sequence analysis, the Inr contains consensus sequence
TCAGTY in Drosophila (FitzGerald et al., 2006). In the TATA-less promoters, a
DPE serving as a binding site of transcription factors has been characterized. The
DPE consensus sequence RGWCGTG (where R is A or G, and W is A or T) is conserved from Drosophila to human. It could be located from TSS to around 35nt
20
downstream, and can compensate for defects in transcription when the upstream
TATA box is mutated (Burke and Kadonaga, 1996).
Transcription initiation
Pol-II is recruited to the promoters to form a preinitiation complex (PIC) and initiate
transcription with the help of general transcription factors (TFs). Transcription factor TFIID binds to the TATA box, and then recruits TFIIA and TFIIB. After the
proper assembly, Pol-II can recognize the protein complex and binds to it. Together
with other transcription factors (TFIIF, TFIIE, TFIIH), transcription is initiated.
When transcription initiates, the Kin28 subunit of TFIIH phosphorylates the Ser5
residues of the CTD (Kim et al., 2010). Then the Pol-II elongation complex is
paused at a checkpoint to ensure pre-mRNA capping. Pausing is regulated in different ways. First, the amount of pausing factors NELF and DRB-sensitivity-inducing
factor (DSIF) is responsible to stabilize the paused Pol-II. The balance between
these factors and p-TEFb, which facilitates the release of Pol-II, regulates the stages
between pausing and productive elongation. (Wu et al., 2003; Jonkers and Lis,
2015). Previous studies indicated that GAGA factor GAF also promotes pausing
through aiding NELF recruitment both before and after transcription initiation (Li et
al., 2013). Mediator also participates in transcription initiation and pausing regulation. Studies indicated that the human mediator subunit MED26 interacts with transcription factors and thereby mediates transition of Pol-II from pausing to productive elongation (Takahashi et al., 2011). The SR protein SRSF2, which is known to
function in splicing, has been revealed to promote the release of paused Pol-II (Ji et
al., 2013). Apart from these factors, Ser2 phosphorylation catalyzed by P-TEFb
facilitates the release from pausing (Ahn et al., 2004). Finally, RNAs also play a
role in this process. For example, ncRNAs together with mediators promote chromatin looping structure, which releases the paused polymerase (Lai et al., 2013). Another study suggested that enhancer RNA had a similar role through extruding
NELF from immediate early genes in neurons (Schaukowitch et al., 2014).
During transcription, Pol-II pausing is a common feature of genes and contributes to
active transcription. Studies suggest that Pol-II pausing and P-TEFb dependent re21
lease of Pol-II happens at all active transcribed genes, which indicates pausing is a
regulatory step of transcription (Jonkers et al., 2014). Knocking down of pausing
factor NELF leads to nucleosome occupation at promoter regions and impaired
transcription factor binding (Gilchrist et al., 2010).
Transcription elongation and termination
After the release of Pol-II form the promoter-proximal region, Pol-II will start the
productive elongation phase of transcription. During the elongation, Ser5 is
dephosphorylated by protein phosphatases such as Ssu72 and FCP1 (Werner-Allen
et al., 2011; Fuda et al., 2012; Hsin and Manley, 2012). The elongation rates are not
constant, and several factors may modulate transcription elongation rates. One such
factor is exon composition (Jonkers et al., 2014). Exons usually have a higher GC
content than the surrounding introns, and the relatively high GC content may delay
transcription. This deceleration might be associated with the outcome of cotranscriptional splicing.
Nucleosome density affects polymerase elongation rate and may cause a transient
stalling, while chromatin remodelers helps the polymerase to overcome the nucleosome barriers (Subtil-Rodríguez and Reyes, 2011). Histone marks can also affect
Pol-II elongation rate by changing the interaction between nucleosomes and DNA,
or facilitating histone chaperone recruitment (Venkatesh and Workman, 2015).
Moreover, other accessory factors are required for efficient transcription. Extensive
studies indicated that histone chaperones FACT, Spt6 and ASF1 are involved in
effective elongation via exchanging or removing histones (Venkatesh and Workman, 2015). Polymerase associated factor (PAF) is known to recruit super elongation complex (SEC) to elongating polymerase to regulate transcription (He et al.,
2011).
Transcription termination mechanisms are complex because Pol-II genes do not
have conserved termination site. The distance between termination site and the 3’end of the mature RNA is variant between a hundred to several thousand base pairs
(Proudfoot, 1989). The end of the pre-mRNA is not determined by the site of transcription termination but by the cleavage of the pre-mRNA at a fixed position 25-30
22
nt downstream of the consensus poly(A) signal (AAUAAA, see page 30). The distances between the 3’-end of the pre-mRNA and termination are variant. Pol-II
continues transcribing up to 2 kilobases downstream of the poly(A) signal (Sims et
al., 2004). Transcription termination is triggered by the polyadenylation signal.
After cleavage of the nascent transcript, the exoribonuclease XRN2 will recognize
the 5’-end of the downstream cleavage product, which will lead to exonucleolytic
degradation from the free 5’-end. The free 3’-end of cleaved RNA will be degraded
by the exosome when released from Pol-II (West et al., 2004). Transcriptional termination is affected in multiple aspects. In vitro transcriptional termination experiments indicate that cleavage of the pre-mRNA is associated with termination efficiency. Polymerase release will be delayed when there is no cleavage (Porrua and
Libri, 2015). Moreover, Pol-II pausing plays a role in transcription termination
(Gromak et al., 2006). One possible mechanism could be the formation of
RNA/DNA hybrids (R-loops) at pausing site downstream of poly(A) signal facilitates XRN2 entry and transcription termination (Skourti-Stathaki et al., 2011).
Pre-mRNA Splicing
Splicing is a process by which introns are removed from the pre-mRNA and exons
are ligated together. Splicing is catalyzed by an RNA-protein complex that contains
five small nuclear ribonucleoproteins (snRNPs): U1, U2, U4, U5 and U6, and also a
large number of associated protein factors (Matera and Wang, 2014). In the premRNA, each intron has a GU sequence at its 5’-end as well as a branch point A, a
poly-pyrimidine tract and an AG sequence at its 3’-end. During the splicing process,
the U1 snRNP binds to the 5’GU of intron by base pairing between snRNA and
mRNA. Then U2 snRNP binds to the branch site with the help of U2 auxiliary factors (U2AFs). At this stage, an interaction between U1 and U2 snRNPs brings the 5’
splice site close to the branch point. Then other spliceosome components, including
the tri-snRNP U4-U5-U6, bind the complex and bring the 3’ splice site also close to
the branch point. Finally the intron is spliced out through two transesterification
reactions catalyzed by U6-snRNP (Matlin et al., 2005).
23
In some cases, exons join together in different combinations through a process
called alternative splicing. Differential inclusion and exclusion of exons is responsible for encoding multiple proteins by a single gene, which increases the diversity of
proteins encoded in the genome to a great extent (Nilsen and Graveley, 2010). In
Drosophila, 20% of the genome encodes exons or mature transcripts. Compared to
the human genome, the genome of Drosophila is more compact and contains much
shorter introns (Graveley et al., 2011). There were 13918 protein-coding genes and
3384 non-coding genes annotated by the year of 2014 (dos Santos et al., 2014).
RNA sequencing data further showed that alternative splicing was observed in 31%
of Drosophila genes (Daines et al., 2011).
Regulation of splicing and alternative splicing
Alternative splicing is the result of a competition among potential splice sites, such
as alternative 3’splice sites or alternative 5’splice sites. Both cis-acting and transacting factors participate in the regulation of the splice-site choice. There are cisregulatory sequences in the pre-mRNA that act as either splicing enhancers (SEs) or
splicing silencers (SSs). Depending on their location, these sequences can be divided into exonic SEs, exonic SSs, intronic SEs and intronic SSs (Figure 4). SEs and
SSs recruit different proteins, for instance Serine/Arginine rich proteins (SR proteins) and heterogeneous nuclear ribonucleoproteins (hnRNPs), that bind to distinct
sequences to recruit splicing factors or prevent their interaction with the pre-mRNA,
and in this way regulate the splicing process (Lopez, 1998; Smith and Valcárcel,
2000). The binding affinity of SEs and SSs is correlated to copy numbers of splicing
elements (Zhang and Chasin, 2004). SR proteins often bind to exonic SEs and promote splicing. SR proteins contain a C-terminal protein domain with long repeats of
serine and arginine amino acid residues. The C-terminal RS domains are conserved
and phosphorylated, and the phosphorylation status affects protein–protein interactions (Graveley and Maniatis, 1998; Xiao and Manley, 1997). The N-terminal RNArecognition motifs (RRMs) mediate the binding to SEs (Graveley and Maniatis,
1998). HnRNP proteins often act as repressors and bind to exonic SSs (Pozzoli and
Sironi, 2005). For example, hnRNP I represses splicing by blocking interactions
between U1 and U2 snRNPs (Izquierdo et al., 2005). HnRNP A1 inhibits internal 5'
splice site though loop structure formation (Nasim et al., 2002). Apart from the
24
sequence of the pre-mRNA, secondary structures are also shown to regulate splicing. For example, docking site:selector sequence interactions are necessary for mutually exclusive splicing in Drosophila down syndrome cell adhesion molecule
(Dscam) gene (Graveley, 2005).
Figure 4. The locations of exonic SEs, exonic SSs, intronic SEs and intronic SSs.
SR proteins bind preferentially to splicing enhancers and facilitate splicing while
hnRNPs bind to splicing silencers and block splicing.
Splicing occurs not only post-transcriptionally but also co-transcriptionally (Baurén
and Wieslander, 1994). Previous studies showed that the CTD constitutes a platform
for the recruitment to the nascent transcript of factors involved in 3’end processing,
splicing and chromatin remodeling (Phatnani and Greenleaf, 2006). The CTD is
needed for splicing and the length of the CTD affects the outcome of splicing
(Rosonina and Blencowe, 2004). Das and co-workers purified SR proteins together
with the Pol-II holoenzyme (Das et al., 2007). The CTD is crucial for the recruitment of the SR protein SRp20, and this recruitment inhibits the alternative exon
inclusion independently of Pol-II elongation rate (de la Mata M and Kornblihtt AR,
2006).
There is also a kinetic coupling model that indicates that Pol-II elongation rate affects splice-site choice. Compared to distal 3’ splice sites, the proximal 3’ splice
sites are weaker and therefore a low Pol-II elongation rate favors the recognition of
proximal 3’ splice sites, which results in the inclusion of alternative exons (Ip et al.,
2011). The study of the fibronectin extra domain I (EDI) supported the kinetic model. A Pol-II with intrinsically low processivity resulted in more inclusion of EDI
exon (de la Mata et al., 2003). Different mechanisms can modulate the elonga-
tion rate of Pol-II. It has been shown that the splicing factor SKIP interacts with
PTEF-b and affects transcription elongation (Brès et al., 2005). Apart from splicing
25
factors, Pol-II elongation rate is also regulated by DNA binding proteins such as the
CCCTC-binding factor (CTCF). CTCF promotes the inclusion of weak upstream
exons of the CD45 gene through Pol-II pausing (Shukla et al., 2011). Another example of alternative splicing regulated through changes in Pol-II elongation is provided by the chromatin remodeling factor SWI/SNF (Batsché et al., 2006). The
SWI/SNF catalytic subunit BRM associates with splicing factor Src-associated protein in mitosis 68 kDa (SAM68) and with other spliceosome components, and promotes exon inclusion by decreasing Pol-II elongation rate at the alternative exons
(Batsché et al., 2006). Cases of siRNA-mediated Pol-II pausing have also been reported. Transfection with siRNAs, which target the downstream intron of an alternative exon, leads to a condensed chromatin structure with higher H3K9me2 and
H3K27me3 marks. Ago1 and heterochromatin-associated protein HP1α are involved
in this process. Highly compacted chromatin decreases Pol-II elongation rate, leading to an inclusion of alternative exons (Alló et al., 2009).
In addition, it is becoming evident that chromatin structure is a key factor of exon
recognition and splicing regulation. High-throughput sequencing data suggested that
nucleosome density was higher in exons with weak splice sites, and positively correlated with exon inclusion rates (Tilgner et al., 2009). This occupancy pattern was
also found in silent genes, indicating that the nucleosome positioning occurred independently of transcription. Histone modifications are also involved in alternative
splicing regulation. Different histone marks affect alternative splicing by mediating
the recruitment of different splicing factors or change Pol-II elongation rate. For
example, H3K9ac facilitates Pol-II elongation rate and thereby causes skipping of
an alternative exon (Schor et al., 2009). On the contrary, H3K9me3 favors Pol-II
accumulation and alternative exons inclusion of CD44 (Saint-André et al., 2011).
Last but not least, it has been studied that small RNA-associated protein Argonaute
promotes spliceosome recruitment, facilitates histone H3K9 methylation and modulates Pol-II elongation rate, which suggests that Argonaute proteins regulate alternative splicing through coupling histone modification to polymerase elongation
(Ameyar-Zazoua et al., 2012).
26
Some non-coding RNAs also have an impact on alternative splicing. The long nuclear-retained regulatory RNA (nrRNA) MALAT1 regulated alternative splicing
through interacting with SR proteins. MALAT1 could regulate the distribution of
SR proteins and other splicing factors such as U2AF-65 at nuclear speckles. The
phosphorylation level of SR proteins was also mediated by MALAT1 (Tripathi et
al., 2010). Another mechanism of splicing regulation is masking the splice sites
from the spliceosome. Expression of a natural antisense ncRNA complementary to
the 5’ splice site of the human mRNA Zeb2 prevented splicing. The retained intron,
containing an internal ribosome entry site recognized by translation machinery, was
essential for Zeb2 translation (Beltran et al., 2008).
It has also been demonstrated that the SWI/SNF complex can regulate the abundance of alternatively spliced transcripts in both Drosophila and mammalian cells
(Batsché et al., 2006, Tyagi et al., 2009). The human BRM (hBRM) was connected
to the inclusion of alternative exons through a mechanism that involves modifications of the phosphorylation state of the CTD and that modulate the Pol-II elongation rate (Batsché et al., 2006). Similarly, Drosophila BRM (dBRM) also acts as a
regulator of pre-mRNA processing. A fraction of dBRM was not associated with
chromatin but with nascent pre-mRNPs, suggesting that SWI/SNF affects premRNA processing by acting at the RNA level (Tyagi et al., 2009). In agreement
with this idea, SNR1, one of the SWI/SNF core subunits, plays a negative role in
splicing regulation of Eig71Eh pre-mRNA through a mechanism that is independent
of Pol-II elongation rate (Zraly and Dingwall, 2012).
Cis- and trans-splicing
There are two kinds of splicing: cis-splicing and trans-splicing. Cis-splicing is the
common form of splicing that occurs within the same pre-mRNA, whereas transsplicing joins exons from two different pre-mRNAs. Trans-splicing usually occurs
in Caenorbabditis elegans, whose genes are clustered and transcribed polycistronically (Horiuchi and Aigaki, 2006). Trans-splicing also happens in mammalian cells
(Caudevilla et al., 1998). Previous evidence indicated that trans-splicing might be
due to an experimental artifact during RT-PCR, when reverse transcriptase switches
from one template to another homologous template (Cocquet et al., 2006). However,
27
high-throughput sequencing of Drosophila hybrid mRNA identified 80 genes that
undergo trans-splicing and firmly demonstrated that the trans-splicing of modifier of
mdg4 (mod(mdg4)) and longitudinals lacking (lola) in Drosophila was not an artifact of template switching (McManus et al., 2010). Evidence demonstrated that the
trans-splicing at the mod(mdg4) locus is independent of the chromosomal position
of common exons and specific exons (Dorn et al., 2001). What triggers initiation of
trans-splicing instead of cis-splicing is not known.
There are two main catagories of spliceosomal trans-splicing events: spliced leader
(SL) trans-splicing and genic trans-splicing. When SL trans-splicing happens, a
common short leader exon attaches to the 5’-ends of different mRNAs, and produces mRNAs containing the same 5’-end sequence (Lasda and Blumenthal, 2011).
Genic trans-splicing involves two transcripts produced from the same gene or from
two different genes, and the exons that join together can be encoded on the same or
opposite DNA strands. In the mod(mdg4) locus of Drosophila, common exons located close to the 5’end of the gene are spliced to several alternative exons that are
transcribed from the same locus. Some alternative exons are transcribed from the
other DNA strand in opposite direction. Therefore exons encoded from opposite
DNA strands join together (Figure 5). We have shown that the alternative splicing of
mod(mdg4) is regulated by SWI/SNF (Waldholm et al., 2011).
Figure 5. Trans-splicing in the mod(mdg4) locus. The arrow indicates a splicing
event that involves the 3’-end of an exon encoded in the sense strand joined
together with the 5’-end of an exon encoded in the antisense strand.
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The Mod(mdg4) gene
The mod(mdg4) gene produces 31 splicing isoforms and encodes for insulator proteins that are involved in the establishment and maintenance of chromatin structure
(Dorn and Krauss, 2003). All MOD(MDG4) proteins have same N-terminal domain
which contains 402 amino acids and includes a BTB/POZ domain. Different protein
isoforms have different C-terminal domains that vary from 28 to 208 amino acids in
length (Büchner et al., 2000; Krauss and Dorn, 2004). Most C-terminal domains
contain a FLYWCH motif (Dorn and Krauss Büchner, 2003). The MOD(MDG4)
proteins are encoded by both strands of the mod(mdg4) gene. In the mod(mdg4)
gene, one promoter drives the transcription of the major (sense) mod(mdg4) premRNA. Several additional promoters drive the expression of anti-sense transcripts.
These antisense transcripts contain several alternative exons that can be spliced
together by a conventional cis-splicing reaction, but some of the exons in the antisense pre-mRNA can also be spliced with the constant sense exons through transsplicing. The common N-terminal part of the MOD(MDG4) proteins is encoded in
four common exons that are located in the 5’-end of locus and are included in all the
mod(mdg4) transcripts. Nine exons encoded in the complementary strand are also
joined together with these four common exons, which indicates that at least nine
transcripts are generated by trans-splicing. Moreover, global analysis of transsplicing indicated that the exons encoded in the same strand (sense strand) also
undergo trans-splicing since they are from different pre-mRNA molecules
(McManus et al., 2010). Each MOD(MDG4) protein isoform has a DNA-binding
domain that confers sequence specificity and that is presumably associated with a
specific function, and the variation in DNA-binding preference plays an important
role in regulating chromatin structure of different genes (Büchner et al., 2000).
Pre-mRNA 3’-end processing
The majority of eukaryotic pre-mRNAs will undergo 3’-end processing which consists of cleavage and polyadenylation to form a mature mRNA. This process is catalyzed by multiple factors: Cleavage and Polyadenylation Specificity Factor (CPSF),
Cleavage Stimulation Factor (CstF), Cleavage Factor I (CFIm), Cleavage Factor II
(CFIIm), polyA polymerase and polyA binding protein (Figure 6). CPSF consists of
29
six polypeptides: CPSF1 (160kDa), CPSF2 (100kDa), CPSF3 (73kDa), CPSF4
(30kDa), FIP1L1 and WDR33. CstF is composed of CSTF1 (55kDa), CSTF2
(64kDa) and CSTF3 (77kDa). CFIm is a heterodimer containing two 25kDa subunits (CPSF5) and two larger subunits either 59 (CPSF7) or 68 kDa (CPSF6). These
factors have different RNA binding sites: CPSF1 recognizes the polyA signal sequence AAUAAA. CstF binds to a GU-rich region downstream of the CPSF binding
site (downstream sequence element). CFIm binds to a UGUAA sequences (upstream
sequence elements) and facilitates the recruitment of CPSF even if the AAUAAA
sequence is missing (Zhao et al., 1999). Once the CPSF complex is assembled,
CPSF3 acts as the processing endonuclease and cuts RNA at cleavage site (Mandel
et al., 2006). Then PolyA tail is added to the 3’end of the cleaved transcript by PolyA polymerase. The CPSF complex and polyA binding protein II are involved in the
control of the length of the polyA tail (Kuhn et al., 2009).
Figure 6. Factors involved in cleavage and polyadenylation. CFIm recognizes the
upstream sequence elements (USE) located approximately 50 nt upstream of the
polyadenylation site (PAS). CPSF recognizes polyA signal (AAUAAA).
Alternative cleavage and polyadenylation
More than 70% of mammalian genes and around half of Drosophila genes have
more than one polyadenylation sites (PAS) (Tian and Manley, 2013). Alternative
polyadenylation is regulated at different levels. Firstly, the abundance of 3’-end
processing factors affects polyA sites usage. For instance, co-depletion of the
CSTF2 and its paralog CstF64τ leads to an increase of distal PolyA site usage (Yao
et al., 2012). Secondly, transcription regulates cleavage efficiency and polyA sites
usage. Highly expressed genes tend to use the proximal PAS whereas lowly ex30
pressed genes tend to use the distal PA sites (Ji et al., 2011). RNA Pol-II kinetics
has an influence on alternative polyadenylation as well. It has been found that lower
polymerase elongation rate favors proximal PA site (Pinto et al., 2011). Loading of
transcription elongation factor ELL2 to RNA polymerase is linked to CstF64 and
promotes proximal polyA site usage, which suggests that transcription factors recruit 3’ processing factors during transcription (Martincic et al., 2009). Transcription activators are also shown to promote 3’end processing and recruit CPSF and
CstF through RNA polymerase associated factors (Nagaike et al., 2011). In addition,
alternative polyadenylation is coupled to splicing. One of the best studied mechanisms of coupleing between splicing and 3’end processing is a mechanism that involves U1 snRNP. U1 snRNP has a negative role in PAS usage by hindering cleavage and polyadenylation in introns (Berg et al., 2012). Finally, regulation of alternative polyadenylation correlates with nucleosome occupancy around the polyA
site. Genome-wide studies of nucleosome occupancy revealed nucleosomes depletion around polyA sites, whereas in regions downstream of the PAS nucleosomes
are more enriched (Spies et al., 2009). The fact that nucleosome landscape affects
polyA site selection suggests a possible role for chromatin remodeling factors in the
regulation of alternative polyadenylation.
Cleavage factor Im
In Drosophila, CFIm only consists of two CPSF5 subunits (approx. 25kDa) and two
CPSF6 subunits (approx. 70 kDa), but not CPSF7. CPSF6 contains three domains:
RRM, Proline-rich and RS like domain (Ruepp et al., 2011). CPSF5 was revealed as
the major RNA binding subunit of CFIm, but CPSF6 facilitates the binding of
CPSF5 and via its RRM domain (Dettwiler et al., 2004; Yang et al., 2011). The RS
domain functions as a protein interaction domain and interacts with spliceosomal SR
proteins: hTra2β, SRp20 and 9G8 (Ruegsegger et al., 1998). CFIm is recruited during the initial transcription stage and elongates with the transcription machinery
(Venkataraman et al., 2005). The binding of CFIm to the pre-mRNA is the earliest
step in the assembly of the cleavage and polyadenylation complex, and CFIm recruits other 3’-end processing factors (Dettwiler et al., 2004).
31
CFIm has been shown to regulate alternative polyadenylation. Depletion of CPSF6
caused an upstream shift in polyA site selection (Kim et al., 2010; Martin et al.,
2012). Depletion of CPSF5 also affects alternative PAS selection of a subset of
genes (Kubo et al., 2006). A model that involves RNA looping has been proposed to
explain this polyA site shift. According to this model, CFIm bind to both proximal
and distal USEs and form a loop. The proximal PAS is then looped out and can not
be accessed by CPSFs, which facilitates the usage of distal PAS (Yang et al., 2011).
Moreover, CFIm is also involved in splicing regulation. Apart from interaction between CPSF6 and SR proteins, CPSF7 has been identified to interact with U2AF65
(Millevoi et al., 2006).
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The present investigation
Aim of the studies
The main focus of my studies is the ATPase dependent chromatin remodeling complex SWI/SNF and how it functions in transcription and pre-mRNA processing.
Previous studies by Batsché et al. (2006) suggested hBRM, one of the ATPase subunits of the human SWI/SNF, is involved in alternative splicing through a mechanism that decreases Pol-II elongation rate. In this Thesis, we intend to address how
BRM regulates alternative splicing and 3’end processing in Drosophila. An important question that I have analyzed is whether the ATPase activity of BRM is
necessary for these processes. Apart from pre-mRNA processing, we also want to
understand the mechanisms by which BRM regulates transcription.
Model System
The Drosophila melanogaster is one of the most intensively studied organisms in
biology. It is a model system to investigate developmental, genetic and cellular
processes. It is easy to create mutants and carry out experiments, and the functions
of many genes have been established (Adams et al., 2000). There are fly strains
constructed to make RNAi experiments and these strains are available to order from
Drosophila resource centers. The mammalian SWI/SNF chromatin remodeling
complex contains one of two highly homologous ATPases, BRG1 and BRM, while
the Drosophila SWI/SNF complex only contains one type of ATPase: BRM. For
this reason, it is more straightforward to study the function and regulation of the
ATPase subunit of SWI/SNF in Drosophila than in mammalian systems.
Schneider 2 cells (S2 cells) are commonly used Drosophila melanogaster cell lines.
They were derived from a primary culture of late stage (20–24 hours old) Drosophila melanogaster embryos. They are derived from a hemocyte lineage. Approaches
of knocking down proteins by RNAi and overexpression by transfection of the recombinant plasmid are easy to carry out (Clemens et al., 2000).
33
High throughput methods
Apart from conventional methodology in molecular biology, two high-throughput
methods have been essential in my projects. One is Next Generation Sequencing
(NGS) and the other one is High Throughput Mass Spectrometry.
I have applied NGS to understand the function of BRM on a genome-wide scale. I
have used ChIP sequencing (ChIP-seq) to map BRM recruitment sites. For this
purpose, I have expressed epitope-tagged BRM in S2 cells and carried out immunoprecipitation experiments using an antibody against the tag. I have then purified the
immunoprecipitated DNA, and sequenced it. The preparation of DNA libraries for
sequencing includes several steps. The purified DNA undergoes end repair to ensure
blunt ended DNA fragments. Adapters with specific sequences are ligated to 3’ and
5’-ends of these fragments. Then Illumina technology is carried out for sequencing.
All the sequencing steps occur in a flow cell with 8 separate lanes (HiSeq 2500).
First, double stranded DNA is denatured into single stranded DNA that can hybridize to the oligos on the flow cell surface. These attached fragments are copied and
used to generate a cluster of identical fragments. After bridge amplification steps,
these clusters are ready to be sequenced. The nucleotide sequence of each cluster is
read by fluorescently labelled dNTPs and one nucleotide is read in each cycle in a
process called “sequencing by synthesis” or SBS. SBS implies that the sequencing
of the nucleotides is based on a DNA polymerase reaction. In NGS, the nucleotides
that are used for SBS emit a fluorescent pulse when they are incorporated into a
nascent DNA strand during sequencing, and each distinct fluorescent signal is detected (Buermans and Dunnen, 2014).
I have carried out RNA sequencing (RNA-seq) to identify transcripts that are affected by BRM depletion or overexpression. Compared to microarrays, RNA sequencing can detect novel transcripts without using pre-designed probes. In addition,
RNA sequencing is more specific and sensitive. It detects transcripts that are relatively lowly expressed (Zhao et al., 2014). In the RNA-seq experiments that are
included in my projects, the purified total RNA was polyA selected and converted to
cDNA fragments with adaptors. Then these cDNAs were amplified and 125 bp were
34
sequenced from each end. The sequencing technology for RNA-seq is basically the
same as described above for DNA. After sequencing, all the reads were aligned to
the reference genome, the reads aligned to transcribed genes were quantified, and
the transcripts that showed changed levels in treated cells (BRM depleted or overexpressed) were identified following standard bioinformatics procedures (Wang et al.,
2009).
Mass spectrometry is an analytical technique that measures the mass-to-charge ratio
of charged particles. It is widely used in molecular cell biology for the identification
of biomolecules, in particular proteins. In the most simple setup, the proteins are
cleaved to smaller peptides by trypsin, the peptides are separated by chromatography, and then introduced into a mass spectrometer. The spectrometer measures the
mass/charge ratio of each peptide. By measuring the masses of each peptide and
comparing them to the masses predicted for each protein encoded in the genome,
different proteins can be identified. In our study, we used high pressure liquid chromatography (HPLC) coupled with tandem MS to analyze the protein-protein interactions. First, the proteins immunoprecipitated by the anti-BRM antibody were
digested, separated by HPLC, and then loaded onto two mass spectrometers linked
by a collision cell. Tandem mass spectrometers provide sequence information. After
comparing MS/MS data to peptide sequences on a database, the protein candidates
associated with BRM were identified. To assess the specificity of our results, we
also identified the unspecific binding of BRM by analysing negative control samples.
35
Project summary
Paper I
SWI/SNF regulates the alternative processing of a specific subset of premRNAs in Drosophila melanogaster
Aims
Previous studies proved that BRM is not only associated with chromatin but also
with pre-mRNP fractions, and that depletion of BRM affects alternative splicing
and/or alternative polyA site selection (Tyagi et al., 2009). The aims of this study
are:
1) Analyze the microarray data in a more stringent way to understand the role of
SWI/SNF in pre-mRNA processing.
2) Apart from BRM, do other SWI/SNF subunits also modulate pre-mRNA processing?
3) Study if this regulation also takes place in vivo
Results
We analyzed the microarray data from Moshkin and coworkers (Moshkin et al.,
2007) and selected genes with more than one probe-set. We calculated signal ratios
for each pair of probe-sets, and selected the ones with more than 2-fold change.
Among these 243 probe-sets corresponding to 149 genes, we found 45 genes that
were affected in terms of alternative pre-mRNA processing. Among them, 15 genes
were affected when knocking down BRM, 14 genes were affected when knocking
down SNR1, and 12 genes were affected by knocking down of Moria. Next we
validated four genes: Gpdh, CG3884, mod(mdg4) and lola derived from microarray
data. The transcripts that we validated from these four genes were affected not only
by BRM alone but also the other SWI/SNF core subunits.
36
To ensure this was a direct effect, we carried out chromatin immunoprecipitation
(ChIP) experiments to see whether BRM, SNR1 and Moira were associated with
these four genes. Our ChIP results confirmed the presence of SWI/SNF core subunits at the target genes, which suggested that the observed effects were direct.
Finally we asked whether we could see the level of SWI/SNF core subunits modulates pre-mRNA processing in vivo. Knocking down BRM in Drosophila larvae
resulted in changes in the relative abundances of alternative transcripts, which suggested that BRM modulates pre-mRNA processing both in vitro and in vivo.
Discussion
Precious studies suggested that BRM affects alternative splicing in human (Batsché
et al, 2006). But how BRM modulates alternative pre-mRNA processing in Drosophila was not clear yet. By microarray and RT-qPCR analysis, we found that
BRM influenced the pre-mRNA processing of a subset of genes in S2 cells and
larvae. Knocking down of SNR1 and Moira suggested that BRM incorporated with
SWI/SNF complex to regulate alternative pre-mRNA processing instead of working
alone. We also examined the decay of the transcripts affected by SWI/SNF subunits.
Knocking down of SWI/SNF subunits did not alter half-life time of these transcripts,
which excluded differences in RNA stability. The differences of transcripts relative
abundances were due to differential regulation of pre-mRNA processing.
Two models regarding mechanisms of pre-mRNA processing by SWI/SNF can be
envisioned. First, SWI/SNF could affect pre-mRNA processing by altering transcription elongation rate, which is consistent with the mechanism of human CD44
gene. The other model is a mechanism that is more direct and independent of transcription kinetics. We observed in a previous study that BRM was associated with
pre-mRNP fractions (Tyagi et al., 2009). We propose that SWI/SNF might play a
role in pre-mRNA processing by facilitating splicing factors recruitment and therefore regulate alternative processing.
37
Paper II
Brahma regulates a specific trans-splicing event at the mod(mdg4) locus of Drosophila melanogaster
Aims
In Paper I, Waldholm and coworkers proved that the patterns of splicing of Gpdh,
CG3884, lola and mod(mdg4) were affected when knocking down BRM. We had
evidence that BRM affects splicing of exons encoded in the same strand, but transsplicing was not studied. To study trans-splicing, we chose the mod(mdg4) gene in
which exons are encoded by both DNA strands. The aims of this paper were:
1) Characterize the expression of one of the mod(mdg4) anti-sense regions.
2) Investigate whether BRM is involved in the trans-splicing of mod(mdg4) transcripts encoded in this anti-sense region.
3) Study whether BRM regulates mod(mdg4) trans-splicing through a mechanism
that is dependent on the chromatin remodeling function of SWI/SNF.
Results
We investigated mod(mdg4) genetic locus and characterized the expression of one
of the mod(mdg4) anti-sense regions (region 1) which contain the following exons:
UE, RY, RX, RAE, RAD, RW, RV, RU (see Figure 1 in the manuscript). We identified several trans-spliced mod(mdg4) transcripts in S2 cells that contained these
exons. In these transcripts, exons from sense and anti-sense strands combine in
highly complex ways. By RT-PCR, we demonstrated that all the exons except RAE
in region 1 were trans-spliced in vivo. After that, we characterized the anti-sense
region 1 using RACE experiments. We mapped the TSSs and PASs for the transcripts in region 1. We found two TSSs and 12 alternative PASs.
In another series of experiments, we asked whether the abundance of antisense transcripts was a limiting factor for the trans-splicing. Un-spliced exons were used as
substrate for trans-splicing, but trans-splicing was not facilitated when we overexpressed part of the anti-sense transcript. This result revealed that trans-splicing is not
merely regulated by the abundance of the substrates but needs other factors. Howev38
er, we cannot exclude the possibility that trans-splicing requires sequences that were
not present in the recombinant cDNA that was used in this experiment.
Knocking down BRM in vitro and in vivo caused a significant decrease of transspliced RX mRNA without affecting the transcription level. On the contrary, transspliced RX increased significantly when we overexpressed BRM in S2 cells. A
BRM mutant that lacked ATPase activity and had a dominant negative effect on
transcription had been previously characterized (Khavari et al., 1993; Elfring et al.,
1998). Expression of this BRM mutant, K804R, resulted in a similar increase of RX
mRNA levels. This result supported the conclusion that BRM affects trans-splicing
of some mod(mdg4) transcripts through a mechanism that does not require its
ATPase activity and is independent of the chromatin remodeling function of
SWI/SNF. ChIP-qPCR experiments showed that both Pol-II and phosphorylated
Ser5-CTD density increased in the mod(mdg4) anti-sense region 1 in cells that overexpressed BRM, which could be a consequence of a reduced elongation rate.
Discussion
We have shown that BRM regulates mod(mdg4) trans-splicing in vitro and in vivo,
but the mechanism of this affection is not known. Previous work in the group
demonstrated that other SWI/SNF core subunits also contribute to the regulation of
mod(mdg4) splicing, which means that BRM might not affect splicing alone. Moreover, a BRM ATPase mutant that lacks the function of chromatin remodeling can
still affect RX trans-splicing. Therefore we concluded that SWI/SNF complex has
independent functions in splicing. ChIP-qPCR data showed that the density of Pol-II
in the antisense region 1 increased in cells that overexpressed BRM, and the increase was also found when using an antibody against the Ser5-CTD. We also
showed that the overexpression of BRM did not increase the abundance of the premRNAs transcribed from the antisense region 1. Therefore we concluded that BRM
does not activate transcription of the antisense region 1 even though there is a local
accumulation of Pol-II. Instead the increased Pol-II density can reflect a reduced
elongation rate. The same tendency for Pol-II and Ser5-CTD was observed at the
human CD44 gene (Batsché et al., 2006). However, the explanation that Pol-II elongation rate decides the use of alternative exons may not be applicable for
mod(mdg4) gene where the exons are transcribed in opposite strands.
39
Paper III
The ATPase-dependent and ATPase-independent functions of Brahma in transcription regulation
Aims
From the analysis in Paper I, we showed that the levels of BRM affect the alternative pre-mRNA processing of CG3884 transcripts. In a more recent release of FlyBase, CG3884 was re-annotated into two overlapping genes: CG44250 and
CG44251. We re-analyzed the regulation by BRM in these two genes and found that
BRM affected the transcription of CG44250 instead of pre-mRNA processing. In
this paper, we aimed to:
1) Establish the role of BRM in regulating the transcription of the CG44250/51
genes.
2) Investigate whether BRM regulates the expression of CG44250/51 gene in an
ATPase dependent manner.
3) Is BRM alone or the whole SWI/SNF involved in CG44250/51 transcription
regulation?
4) Study the catalytic and non-catalytic roles of BRM in transcription at a genomewide scale.
Results
CG3884 was re-annotated as CG44250 and CG44251 in a complex locus that contains three different promoters. First we re-characterized the CG44250/51 locus. By
RT-PCR, we detected six different mRNAs generated by alternative splicing and
alternative polyadenylation. RNAs transcribed from promoter 2 (P2) were much
more abundant compared to promoter 1 (P1) and promoter 3 (P3). ChIP-qPCR experiments suggested that BRM was recruited to the entire CG44250/51 region, but
P2 had slightly more occupancy than P1 and P3.
To investigate whether BRM affects CG44250/51 pre-mRNA processing, we depleted BRM by RNAi in S2 cells and quantified the abundance of each transcript
40
derived from CG44250/51 locus using exon-exon junction primers designed to detect the alternatively spliced isoforms. The abundance of every transcript was decreased to approximately the same extent when we depleted BRM. We also measured exon abundance, transcribing from P2, which was also down regulated to the
same extent. Exon abundance transcribed from P1 was not affected. This evidence
suggested that BRM regulated the transcription instead of pre-mRNA processing of
CG44250/51. Similar effect was also observed in larvae, which revealed that BRM
modulated CG44250/51 transcription both in vitro and in vivo. This finding was
further confirmed by ChIP-qPCR experiments where we measured RNA polymerase
density at the CG44250/51 locus. We saw a decrease of Pol-II CTD occupancy at P2
and downstream regions in BRM-depleted cells, whereas P1 and P3 were not influenced.
Next we wondered whether the ATPase activity of BRM was essential for the transcriptional regulation described above. By overexpressing either BRM wild-type
(BRM-WT) or an ATPase mutant form of BRM (BRM-K804R), we found that both
BRM-WT and BRM-K804R upregulated CG44250/51 transcription. Intriguingly,
BRM-K804R caused more upregulation than BRM-WT. ChIP-qPCR analysis indicated that overexpression of BRM resulted in increased BRM, Pol-II and Ser5 phosphorylated Pol-II occupancy. We also detected more occupancy of H3K9ac and
H3K27ac, suggesting that in the CG44250/51 locus BRM promotes a chromatin
state compatible with more active transcription. The observations above showed that
the role of BRM in transcription regulation at CG44250/51 gene is independent of
its ATPase activity.
The bromodomain of BRM functions in protein-protein interaction so we hypothesized that the bromodomain might be required for the transcription of the
CG44250/51 locus. Overexpressing a bromodomain mutant form of BRM (BRMYN1501AA) reproduced the effect of BRM depletion, which confirmed our hypothesis.
Then we asked whether the regulation of CG44250/51 transcription was carried out
by BRM alone or as part of SWI/SNF. To answer this question, we analyzed public
41
microarray expression data from experiments in which the effect of depleting different subunits of SWI/SNF on the transcriptome of S2 cells had been studied (Moshkin et al., 2007). Microarray hybridization revealed that the PBAP-specific subunits
PB and BAP170 were essential for proper CG44250/51 transcription while the
BAP-specific subunit OSA was not necessary, which showed PBAP but not BAP
functions in CG44250/51 transcription regulation.
Finally we analyzed the ATPase dependent and independent roles of BRM in transcription at a genome-wide scale. RNA-sequencing results indicated that 770 genes
were affected by BRM in an ATPase independent manner, and 261 genes were
affected in an ATPase dependent manner. TATA box frequency analysis suggested
that many of these affected genes were TATA-less genes. We further carried out
motif enrichment analysis and showed a non-catalytic role of BRM as a negative
regulator of genes involved in Hox/TGF-beta signaling pathways. In addition, genes
affected by BRM were characterized by lower nucleosome density than the average
of genome.
Discussion
We have shown that BRM acts as an activator to regulate CG44250/51 transcription
at promoter 2. Apart from BRM, other subunits of the PBAP complex are also involved in this regulation. Most importantly, our studies pointed out an ATPase independent role of BRM in transcription regulation at CG44250/51, which is consistent with previous work (Zraly et al., 2006). Genome-wide studies further confirmed this observation and suggested that the non-catalytic role of BRM in transcription regulation is not an exception but is much more widespread than initially
anticipated. Motif analysis identified genes regulated by BRM contain binding sequences for transcription factors, suggesting BRM may modulate the recruitment of
transcription factors and thereby regulate the transcription of the target genes. Based
on all these observations, we conclude that BRM regulates transcription at two different levels: 1) in some genes, it remodels chromatin structure using the energy
from ATP hydrolysis. 2) in others, BRM modulates the recruitment of transcription
activators or repressors through protein-protein interactions. Some of the results
presented here might be secondary effects of BRM overexpression, because BRM
42
regulates the expression of many transcription factors. In order to rule out indirect
effects, we have carried out ChIP-seq experiments that will allow us to distinguish
the false positive hits (genes are affected in the RNA-seq but BRM is not recruited
to these genes) from those where BRM is present and therefore can play a direct,
local role.
43
Paper IV
A role for SWI/SNF in pre-mRNA 3’-end processing
Aims
Batsché et al. (2006) suggested that hBRM associates with the mRNA-binding protein Sam68 to decrease the elongation rate of Pol-II and to promote the accumulation of Pol-II at specific sites. This accumulation promotes splicing at weak, proximal splice sites. We suspected that in Drosophila, BRM could also regulate splicing
or polyadenylation through mRNA binding proteins. To investigate this possibility,
we immunoprecipitated BRM from chromosomal RNP fractions prepared from S2
cells. We pulled down BRM by immunoprecitation with anti-BRM-specific antibodies, and identified BRM-interacting proteins by high-throughput mass spectrometry.
We found several RNA-binding proteins that are involved in pre-mRNA processing
among the identified proteins. We also carried out a similar proteomics analysis in
human cells, in collaboration with the Östlund Farrants group. We detected interactions between the SWI/SNF ATPases and components of the 3’-end processing
machinery in both human and fly cells. Based on this observation, we aimed to:
1) Validate the interaction between 3’-end processing factors and dBRM in S2 cells
and hBRG1 in human C33A cells.
2) Elucidate to what extent dBRM affects pre-mRNA processing on a global scale.
3) Analyze whether the regulation of 3’-end processing by dBRM and hBRG1 is
ATPase dependent.
Results
We immunoprecipitatated dBRM, hBRM and hBRG1 to identify proteins that interacted with the SWI/SNF complexes of fly and human cells. The experiments were
carried out in parallel using nuclear fractions enriched in chromatin-bound RNP
complexes. The immunoprecipitated proteins were analyzed by mass-spectrometry.
We identified 271 proteins binding to hBRG1, 128 to hBRM and 130 to dBRM.
Gene ontology term analysis revealed that the term “poly(A) RNA binding” was
significantly enriched in all three interactomes. hBRG1 was associated with CPSF1
44
and CPSF2, while dBRM was associated with CPSF6. We further validated these
interactions by co-immunoprecipitation and Western blotting experiments.
We next aimed at investigating the functional significance of the interactions, and
we analyzed two transcripts that were identified as BRM targets in Paper I: CG5174
and CG2051. We depleted BRM and measured the abundance of uncleaved premRNAs by RT-qPCR. Approximately 20% more uncleaved CG5174 and CG2051
pre-mRNAs were observed in BRM-depleted cells compared to control samples.
Next we used the ATPase mutant form of BRM, recBRM-K804R, and overexpressed either wild-type recBRM or recBRM-K804R in S2 cells. We measured 3’end cleavage in either recBRM or recBRM-K804R overexpressed samples. Overexpression of recBRM caused a significant reduction of uncleaved pre-mRNAs in
CG5174 and CG2051. However, overexpression of recBRM-K804R resulted in less
reduction of uncleaved CG5174 and an increase of uncleaved CG2051. The data
above indicated BRM regulated CG5174 cleavage in an ATPase independent manner. For CG2051, the ATPase activity was needed to promote cleavage. We also
quantified GPRC5A and CD44 pre-mRNA cleavage in human C33A cells, and the
results also suggested a role for hBRG1 in the 3’-end processing of the analyzed
genes. Overexpression of recombinant CPSF6 resulted in a decrease of uncleaved
pre-mRNA and this effect was undermined when dBRM was depleted, suggesting
that the effect of CPSF6 was mediated by dBRM.
We further addressed whether dBRM facilitates the interaction of CPSF6 with the
cleavage site. We carried out ChIP-qPCR experiments in S2 cells expressing recCPSF6. First we depleted dBRM and measured CPSF6 occupancy by ChIP-qPCR.
Less CPSF6 was recruited to the cleavage sites of the analyzed genes in dBRMdepleted cells. We also carried out ChIP-qPCR experiments expressing both recBRM and recCPSF6, and we observed increased CPSF6 levels both in BRM wildtype and ATPase mutant. The data presented above revealed that dBRM facilitates
the interaction of CPSF6 with the cleavage site through a mechanism that does not
need the ATPase activity of BRM.
45
Finally, we carried out RNA-sequencing experiments either knocking down or overexpressing BRM to understand the effect of BRM on cleavage on a genome-wide
scale. Metagene analysis showed that depletion of BRM caused a pileup of reads
downstream of the PAS. This effect was more significant in highly expressed genes.
Cluster analysis of the highly expressed genes according to their effects on premRNA processing suggested different structural features such as poly(A) signal,
USE and DSE. In addition, gene ontology studies revealed that the different gene
clusters were associated with specific biological functions.
Discussion
Co-immunoprecipitation experiments proved a physical link between SWI/SNF and
the 3’-end processing machinery, and our results provided direct evidence that
SWI/SNF is involved in 3’-end processing of a subset of pre-mRNAs in S2 cells.
Our results suggest that SWI/SNF might regulate cleavage through different mechanisms. We have ruled out indirect effects due to a possible influence of dBRM on
the expression of genes that code for 3’-end processing factors. We have also ruled
out possible effects of dBRM levels on Pol-II elongation rate. In CG5174 and
CG2051, overexpression of BRM wild-type and ATPase mutant suggested that
BRM is involved in the recruitment of CPSF6 to the PAS through a mechanism that
is independent of its catalytic activity. However, the regulation of the cleavage efficiency, at least in the CG2051 gene, was ATPase dependent. One possible explanation is that the recruitment of CPSF6 is sufficient for efficient cleavage in a subset
of genes, whereas other genes may need chromatin remodeling to facilitate cleavage.
RNA-sequencing analysis revealed that BRM favors the usage of proximal cleavage
sites in genes with alternative PASs. This is consistent with the fact that in CG5174
and CG2051, depletion of BRM causes a defect of cleavage. Depletion of BRM
might cause a shift from proximal to distal PAS. Gene cluster studies indicated that
the genes affected by dBRM are involved in biological processed that are consistent
with the function of SWI/SNF in embryonic development, gametogenesis and cell
proliferation.
46
Conclusions
SWI/SNF, a master regulator of cell differentiation and development, regulates
many genes by remodeling nucleosomes at promoter regions. My studies have revealed additional roles of SWI/SNF and have provided new knowledge about the
mechanisms by which SWI/SNF regulates gene expression. We have found that
BRM, the ATPase subunit of SWI/SNF, regulates the transcription of a subset of
genes in an ATPase dependent manner as expected. Intriguingly, we also established
that the transcription of many genes in Drosophila S2 cells is regulated by BRM in
an ATPase independent manner, and we have identified the genes that are regulated
by BRM through non-catalytic mechanisms in S2 cells. Apart from the role in transcription regulation, BRM plays roles in pre-mRNA processing, both alternative
splicing and alternative 3’-end processing. We have shown that SWI/SNF regulates
the alternative processing of a subset of pre-mRNAs both in vitro and in vivo. In the
context of splicing regulation, we have shown that BRM has an effect on transsplicing. The studies of the mod(mdg4) gene indicated that the regulation of transsplicing of mod(mdg4)-RX by BRM is independent of the ATPase activity of BRM.
BRM promotes the recruitment of 3’-end processing factors to the PAS, as shown
for the CG5174 and CG2051 pre-mRNAs, without involvement of its catalytic activity. And the increased recruitment of CPSF6 to the PASs correlates in some cases
with more efficient 3’-end cleavage. In summary, SWI/SNF plays important roles in
transcription and pre-mRNA processing and acts through two different mechanisms.
One of the mechanisms needs the catalytic activity of BRM, either for nucleosome
remodeling or for other reactions that require ATP hydrolysis. The other mechanism
does not require the ATPase activity of BRM and is therefore uncoupled from
BRM’s activity in chromatin remodeling. In the latter case, BRM might function as
a platform to modulate the interactions of transcription factors, splicing factors or
3’-end processing factors with their target binding sites through protein-protein
interactions.
47
Acknowledgements
It would not have been possible to write this thesis without the help and support
from people around me. I would like to sincerely thank everyone who has assisted
me in these years:
First I take this opportunity to express my profound gratitude to my supervisor Neus
Visa for accepting me as a PhD student, and for your constant guidance and encouragement throughout the projects. I learned a lot during these five years, both the
basic knowledge and technical skills. Thanks for revising my manuscripts and thesis
as well. My thesis would not have been possible without your help and useful suggestions. To my co-supervisor Ann-Kristin Östlund Farrants, thank you for support and professional advice on my project, especially the helpful discussions and
comments on my manuscripts.
To current and past members in our group: Johan Waldholm, thanks for teaching
me many lab skills. It has been a pleasure working with you. Antonio Jordán Pla,
thanks for all the help on my projects and RNA-sequencing analysis. Andrea
Eberle, thanks for helping me in the lab with common stuffs and leaving the heater
for us. Judit Domingo Prim, we had a great time in the lab and also other PhD
activities. Noemí Sánchez Hernández, thanks for cheering me up in the lab. I feel
really motivated when we were all busy and working hard. Franziska Bonath,
thanks for assistance on programming and feeding us with super delicious cakes.
Consuelo Marin-Vicente, thanks for always being encouraging me and keeping a
positive mood all the time. Anne von Euler, thank you for keeping happiness in the
lab. Magnus Hansson, it has been nice time to share the office and the lab with you.
I would also thank part of the Anki’s group for collaborations and nice meetings:
Antoni Gàñez Zapater, thank you for hard working on CPSF6 project and sharing
ham with us. Anna Rolicka, thanks for your qPCR and ChIP experiments for
CPSF6 story.
Thanks to every one in our old Molbio and MBW:) It is a pleasure working with all
of you!
Finally, I would particularly like to thank my family: 谢谢我的父母对我的爱和一
如既往的支持与照顾. Thanks Zhe Yuan for being a witty guy and standing by me
all the timeJ
48
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