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Helsinki University Biomedical Dissertations No. 155

Mammalian Mat1 subunit of Transcription Factor IIH in gene-specific and general transcription

Katja Helenius

Institute of Biotechnology

Genome-Scale Biology Program and Institute of Biomedicine,

Faculty of Medicine

Helsinki Biomedical Graduate School

University of Helsinki, Finland

Academic dissertation

To be publicly discussed with the permission of the Faculty of Medicine of the University of Helsinki, in Biomedicum Helsinki, Lecture Hall 2, Haartmaninkatu 8, Helsinki

on September 9th, 2011, at 12 noon.

HELSINKI 2011

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Thesis supervisor Tomi P. Mäkelä, M.D., Ph.D. Professor Institute of Biotechnology and Genome-Scale Biology Program University of Helsinki Helsinki, Finland Thesis follow-up committee Olli A. Jänne, M.D., Ph.D. Mikko Frilander, Ph.D.

Professor Institute of Biotechnology Institute of Biomedicine University of Helsinki

Faculty of Medicine Helsinki, Finland University of Helsinki Helsinki, Finland

Reviewers appointed by the Faculty Lea Sistonen, Ph.D. Olli A. Jänne, M.D., PhD. Professor Professor Department of Biology Institute of Biomedicine Åbo Akademi University Faculty of Medicine Turku Center for Biotechnology University of Helsinki Turku, Finland Helsinki, Finland Opponent appointed by the Faculty John T. Lis Professor Department of Molecular Biology and Genetics Cornell University Ithaca, New York, USA

ISBN 978-952-10-7141-6 (paperback) ISBN 978-952-10-7142-3 (PDF) ISSN: 1457-8433 http://ethesis.helsinki.fi Yliopistopaino Helsinki 2011

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“Harder, Better, Faster, Stronger” -daft punk-

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TABLE OF CONTENTS

ORIGINAL PUBLICATIONS………………………………………………………………....5

ABBREVIATIONS……………………………………………………………………………...6

ABSTRACT……………………………………………………………………………………...9

REVIEW OF THE LITERATURE……………………………………………………………10

1. RNA Polymerase II-mediated transcription…………………………………...................10 The chromatin template……………………………………………………........................11 Phosphorylation of the C-terminal domain of RNAPII……………………..................13 TFIIH and Mat1……………………….…………………………………….......................16 Cotranscriptional mRNA processin……………………………………….......................19 Regulation of mRNA stability and decay..…………………………………………….…..21 Chromatin modifications…………………..………………………………………………..23

2. Transcriptional regulation of energy metabolism……………………………..…………26

The transcriptional cascade of adipocyte differentiation….…………….....................26 The PPAR coactivator-1 family members and fatty acid oxidation……….……….....29 Transcriptional control of hepatic lipogenesis……...…………………………………...30 TFIIH: linking general and gene-specific transcription.…...………………................33

AIMS OF THE STUDY………………………………………………………………………...35

MATERIALS AND METHODS…………………………………………………….................36

1. Materials……………………….………………………………………………...............36 2. Methods……………………………….………………………………………………….37

RESULTS ……………………………………………………………………………………….44

1. Mat1 is required for phosphorylation of RNA Polymerase II C-terminal domain,

transcription and efficient mRNA turnover ………………...….……………….……….44 2. The Cdk7 kinase submodule of TFIIH acts as a physiological roadblock to adipogenesis

through inhibitory phosporylation of PPARγ....................................................................46 3. Mat1 is critical for cardiac energy metabolism in vivo via regulation of PGC1…………48 4. Hepatic-specific deletion of Mat1 in mice results in fatty liver through activation of

SREBP-1c target genes via loss of DMAP1 and Dnmt1………………………………...49

DISCUSSION & PERSPECTIVES……………………………………………………………52

1. Mat1, Cdk7 and RNA Polymerase II CTD phosphorylation……………………………52 2. Tissue-specific functions of Mat1……………………………………………………….54 3. mRNA stabilization: linking transcription progression to mRNA turnover…………….55

ACKNOWLEDGEMENTS……………………………………………………………………58 REFERENCES…………………………………………………………………………………60

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ORIGINAL PUBLICATIONS The thesis is based on the following original publications, which have been assigned the following roman numerals: I. Helenius K, Yang Y, Tselykh T.V, Pessa H.K.J., Frilander M., Mäkelä T.P. Requirement of TFIIH kinase subunit Mat1 for RNA Pol II C-terminal domain Ser5 phosphorylation, transcription and mRNA turnover. Nucleic Acids Research. Vol 3 (2011). II. Helenius K, Yang Y, Alasaari J, Mäkelä TP. Mat1 inhibits peroxisome proliferator-activated receptor gamma-mediated adipocyte differentiation. Molecular and Cellular Biology. Vol. 20, No. 2. 315-23 (2009). III. Sano M*, Izumi Y*, Helenius K, Asakura M, Rossi DJ, Xie M, Taffet G, Hu L, Pautler RG, Wilson CR, Boudina S, Abel ED, Taegtmeyer H, Scaglia F, Graham BH, Kralli A, Shimizu N, Tanaka H, Mäkelä TP, Schneider MD. Ménage-à-trois 1 is critical for the transcriptional function of PPARgamma coactivator 1. Cell Metabolism. Vol. 5, Issue 2:129-42 (2007). IV. Helenius K, Mäkelä TP. Hepatosteatosis and derepression of SREBP-1c target genes following ablation of the TFIIH kinase subunit Mat1. Manuscript. The articles are reproduced with the permission of copyright holders. ∗=Equal contribution Publication I was also used in the thesis of Timofey Tselykh in the Faculty of Biosciences.

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ABBREVIATIONS Ab antibody ACC acetyl CoA carboxylase ACL acetyl CoA lyase ACS acetyl CoA synthase ADP adenosine diphosphate AF-2 ligand-dependent transactivation domain AR androgen receptor ARE adenylate-uridylate rich element as analog-sensitive ATP adenosine-5-triphosphate bp base pairs BAT brown adipose tissue BRE TFIIB recognition element c-myc cellular myelocytomatosis viral oncogene homolog CAK Cdk-activating kinase CBP CREB-binding protein Cdk cyclin-dependent kinase C/EBP CCAAT/enhacer binding protein ChIP chromatin immunoprecipitation ChREBP carbohydrate response element binding protein CKO conditional knockout CPSF cleavage polyadenylation specificity factor CPT1 carnitine palmytoyl transferase 1 CstF cleavage stimulation factor CTD C-terminal domain DBD DNA binding domain DCE downstream core element DGAT diacylglycerol acyltransferase DNA deoxyribonucleic acid Dnmt DNA methyltransferase DPE downstream promoter element Elovl6 elongation of long chain fatty acids family member 6 ER estrogen receptor ERR estrogen-related receptor FACT facilitates chromatin transcription FAS fatty acid synthase FAO fatty acid oxidation FCP1 TFIIF-associating CTD phosphatase 1 GAPDH glyceraldehyde 3-phosphate dehydrogenase Gcn5 general control of amino acid synthesis protein 5 G6PD glucose-6-phosphate GTF general transcription factor GPAT glycerol phosphate acyltansferase GR glucocorticoid receptor

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GST glutathionine-S transferase H histone HAT histone acetylase HDAC histone deacetylase HDM histone demethylase HMT histone methyltransferase HNF hepatocyte nuclear factor Hsp70 heat shock protein 70 HuR Hu antigen R ICM inner cell mass Inr initiator element IRE iron-response element JNK c-Jun N-terminal kinase LBD ligand-binding domain MAPK mitogen-activated protein kinase Mat1 Menage-a-trois 1 ME1 malic enzyme 1 MEF mouse embryonic fibroblast MEF2 muscle enhancer factor 2 MTE motif ten element mRNA messenger RNA NER nucleotide excision repair NHR nuclear hormone receptor NMD nonsense-mediated decay NR nuclear receptor NRF nuclear respiratory factor Oct-1 octamer binding transcription factor 1 PCR polymerase chain reaction PIC preinitiation complex PARN poly(A) ribonuclease PARP1 poly-ADP-ribosylate protein 1 PGC1 PPARγ co-activator 1 PGDH phosphogluconate dehydrogenase PKA protein kinase A PKC protein kinase C PPAR peroxisome proliferating antigen receptor PPRE PPAR response element PR progesterone receptor P-TEFb positive transcription elongation factor b qRT-PCR quantitative real-time polymerase chain reaction RAR retinoic acid receptor RNA ribonucleic acid RNAP II ribonucleic acid polymerase II RXR retinoid X receptor SAM S-adenosyl-L-methionine SCAP SREBP cleavage-activating protein

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SCD1 stearoyl CoA desaturase 1 SCP1 small c-terminal domain phosphatase 1 SET Su(var)3-9, Enhancer of zeste E(z), Trithorax Ser serine siRNA small inhibitory RNA snRNA small nuclear RNA Spt suppressor of Ty SRE sterol regulatory element SREBP sterol regulatory element binding protein SRC-1 steroid receptor coactivator 1 Swi/Snf SWItch/Sucrose NonFermentable TAF transcription-associated factor TBP TATA-binding protein TFII transcription factor II ts temperature-sensitive TSS transcription start site TZD thiazolidinediones UCP-1 uncoupling protein 1 UTR untranslated region VDR vitamin D receptor VEGF vascular endothelial growth factor WAT white adipose tissue XCPE1 X core promoter element 1 XPB/D/F/G xeroderma pigmentosum complementation group B/D/F/G

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ABSTRACT

All protein-encoding genes in eukaryotes are transcribed into messenger RNA (mRNA) by RNA

Polymerase II (RNAP II), whose activity therefore needs to be tightly controlled. An important

and only partially understood level of regulation is the multiple phosphorylations of RNAP II

large subunit C-terminal domain (CTD). Sequential phosphorylations regulate transcription

initiation and elongation, and recruit factors involved in co-transcriptional processing of mRNA.

Based largely on studies in yeast models and in vitro, the kinase activity responsible for the

phosphorylation of the serine-5 (Ser5) residues of RNAP II CTD has been attributed to the

Mat1/Cdk7/CycH trimer as part of Transcription Factor IIH. However, due to the lack of good

mammalian genetic models, the roles of both RNAP II Ser5 phosphorylation as well as TFIIH

kinase in transcription have provided ambiguous results and the in vivo kinase of Ser5 has

remained elusive.

The primary objective of this study was to elucidate the role of mammalian TFIIH, and

specifically the Mat1 subunit in CTD phosphorylation and general RNAP II-mediated

transcription. The approach utilized the Cre-LoxP system to conditionally delete murine Mat1 in

cardiomyocytes and hepatocytes in vivo and and in cell culture models. The results identify the

TFIIH kinase as the major mammalian Ser5 kinase and demonstrate its requirement for general

transcription, noted by the use of nascent mRNA labeling. Also a role for Mat1 in regulating

general mRNA turnover was identified, providing a possible rationale for earlier negative

findings.

A secondary objective was to identify potential gene- and tissue-specific roles of Mat1 and the

TFIIH kinase through the use of tissue-specific Mat1 deletion. Mat1 was found to be required for

the transcriptional function of PGC-1 in cardiomyocytes. Transriptional activation of lipogenic

SREBP1 target genes following Mat1 deletion in hepatocytes revealed a repressive role for

Mat1apparently mediated via co-repressor DMAP1 and the DNA methyltransferase Dnmt1.

Finally, Mat1 and Cdk7 were also identified as a negative regulators of adipocyte differentiation

through the inhibitory phosphorylation of Peroxisome proliferator-activated receptor (PPAR) γ.

Together, these results demonstrate gene- and tissue-specific roles for the Mat1 subunit of TFIIH

and open up new therapeutic possibilities in the treatment of diseases such as type II diabetes,

hepatosteatosis and obesity.

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REVIEW OF THE LITERATURE

I. RNA Polymerase II-mediated transcription

The genetic blueprint for the building blocks of all living organisms on Earth is contained

within their DNA. Transcription by RNA polymerase II (RNAP II) is the process of

copying a defined sequence of DNA into a complementary messenger RNA (mRNA)

sequence, which is then translated into proteins by ribosomes. In addition to RNAP II,

eukaryotic cells contain two other polymerases: RNAP I which transcribes the majority

of ribosomal RNA (rRNA) and RNAP III, which is responsible for producing transfer

RNA (tRNA), 5S rRNA and other small RNAs. RNAP II-mediated transcription is

divided into four stages: initiation, promoter escape, elongation and termination.

Regulation of transcription by RNAP II is the key event in determining the fate of any

given cell, and thus it is not surprising that this regulation is complex. Classically,

regulation of transcription initiation is thought to occur via binding of specific

transcription factors to promoters and enhancers of a given gene, which recruit the

general transcription factors (GTFs) and the polymerase machinery to the scene.

Following formation of this pre-initiation complex (PIC), RNAP II escapes the PIC and

progresses into elongation to produce the nascent transcript. Once the polymerase

encounters the termination signal at the end of the gene, it exits the DNA and is ready to

begin a new transcription cycle.

Recent genome-wide maps of RNAP II distribution in human and Drosophila have

demonstrated that in addition to RNAP II recruitment to promoters, transcription is also

often regulated at the stage of progression into elongation: many genes contain a

promoter-proximal paused polymerase (Guenther et al., 2007). These genes are often

ones that require rapid activation in response to external stimuli (Muse et al., 2007) as

well as genes involved in pattern formation during development (Wang et al., 2007;

Zeitlinger et al., 2007). Hence the paused polymerase is thought to provide a swift means

of temporal and special regulation for transcription. However, regardless of the status of

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RNAP II, all nuclear protein-encoding genes in eukaryotes exist as chromatin and their

mRNAs are mostly processed in the same way to produce mature transcripts.

The chromatin template

In eukaryotes, DNA is packaged into chromatin, the main constituents of which are

nucleosomes: 147 base pairs of DNA wrapped around highly basic histone octamers

(Figure 1). The histone octamers consist of two H2A-H2B dimers and an H3-H4

tetramer. Between the core nucleosomes, the DNA is occupied by a linker histone, H1,

which can additionally compact the chromatin. The chromatin structure and nucleosome

positioning relate directly to transcription: tightly packed chromatin is often silent

whereas highly transcribed regions and, in particular promoters, associate with lower

nucleosome occupancy (Bernstein et al., 2004; Lee et al., 2004; Lee et al., 2007; Sekinger

et al., 2005). In order for RNAP II to progress along the DNA, the tightly packed

chromatin template must be altered; this involves partial disassembly of nucleosomes by

chromatin remodellers such as Swi/SNF, FACT and Spt6 (Belotserkovskaya et al., 2003;

Ivanovska et al., 2010 ; Takahata et al., 2009) as well as histone modifications. The major

histone modifications are acetylation, methylation, phosphorylation and ubiquitination;

the combination of these modifications is termed the ‘histone code’ (reviewed in

Jenuwein and Allis, 2001). These modifications directly influence the physical structure

of the chromatin to enable or alternatively block the passage of RNAP II and additionally

recruit other co-regulatory proteins that control transcription progression. These

modifications are discussed in more detail in ‘Chromatin modifications’.

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Figure 1. The chromatin template and the core promoter. The genetic information of eukaryotes in chromosomes is packaged into chromatin, which consists of DNA and histone proteins. 147 bp of DNA wrapped around histone octamers forms the core component of chromatin: the nucleosome. Transcription of DNA into mRNA begins from the transcription start site (TSS), which is located in the core promoter region. The main constituents of the core promoter are the TFIIB recognition element (BRE), TATA box, Initiator element (Inr) and the downstream promoter element (DPE), which together dictate where RNA Polymerase II and the transcription machinery assemble.

Although the structure of the entire chromatin template affects transcription and vice

versa, it is specific sequences in the DNA that dictate where the PIC is assembled. The

core promoter is the nucleotide sequence in DNA to which the transcription machinery

binds in order to initiate transcription. Most promoters exist in nucleosome-free regions

(NRFs), regardless of their status of transcriptional activity (Albert et al., 2007; Lee et al.,

2007; Mavrich et al., 2008; Shivaswamy et al., 2008). Two types of core promoters exist:

focused and dispersed. The majority of vertebrate promoters are dispersed: they contain

numerous transcription start sites (TSS) diffused along 100 bp (Carninci et al., 2005;

Carninci et al., 2006). A typical focused core promoter is composed of several

nucleotides and includes one TSS or a cluster of start sites. Focused promoters are often

found on tissue-specific genes and typically contain the consensus sequence TATAA

termed the TATA box located 28-34 bp upstream of the TSS, which recruits the TATA-

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binding protein (TBP) (Carninci et al., 2006). Dispersed promoters are common in

ubiquitously expressed genes and typically have CpG islands, genomic stretches enriched

with GC dinucleotides, which can be methylated to repress transcription (Carninci et al.,

2006). In addition to the TATA box, the paramount core promoter motifs are the TFIIB

recognition element BRE, the initiator element (Inr), and the downstream promoter

element, DPE (reviewed in Smale and Kadonaga, 2003) (Figure 1). All, some or none of

these elements may be included in a particular core promoter. The Inr comprises the start

site for transcription and is functionally similar to the TATA box in that they are the only

known promoter elements that can alone recruit PIC (Smale and Baltimore, 1989). The

DPE is typically found in TATA-less promoters and acts in conjunction with the Inr as a

TFIID binding site (Zhou and Chiang, 2001). In classic PIC assembly, TFIID binds to

these core elements either through TBP or TAFs (transcription associated factors),

followed by sequential recruitment of the other GTFs: TFIIA, TFIIB, TFIIF, TFIIE and

TFIIH (reviewed in Baumann et al., 2010). To initiate transcription, accessory factors

such as the Mediator, chromatin modifiers, and co-activators are often required in the

vicinity of the promoter (reviewed in Malik and Roeder, 2010). The basic structure of the

core promoter and chromatin is depicted as a schematic in Figure 1.

In addition to the promoter elements described above, core promoters may contain the

more rare sequence motifs Downstream core element (DCE) (Lewis et al., 2000), X core

promoter element 1 (XCPE1) (Tokusumi et al., 2007) and Motif ten element (MTE) (Lim

et al., 2004). Although the role of core promoter elements in PIC assembly and TSS

definition has been widely accepted, it should be noted that recent advances in

nucleosome profiling have led to speculations that nucleosome positioning may

determine the location of the TSS and dictate PIC assembly (reviewed in Jiang and Pugh,

2009).

Phosphorylation of the C-terminal domain of RNA PII

Comprising at least 12 distinct subunits and with a molecular mass of over 500 kD,

eukaryotic RNA polymerase II is the protagonist in transcription of mRNA. The C-

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terminal domain (CTD) of the largest subunit of RNAP II (Rpb1) holds a unique, highly

conserved feature: a tandem repeat of the heptapeptide consensus sequence YSPTSPS.

The repeat length varies: yeast has 26-27 repeats, C. elegans contains 34 repeats,

Drosophila carries 43 repeats, and the human and mouse have 52 (reviewed in Egloff and

Murphy, 2008). The CTD is essential for viability: deletion analyses and truncation

mutants show that the minimal length of 11 repeats in yeast and 29 repeats in human cells

is necessary for survival (Bartolomei et al., 1988; Nonet et al., 1987). The CTD

heptapeptide repeat can be phosphorylated at the three serine residues of the consensus

sequence, Ser2, Ser5 and Ser7 (Akhtar et al., 2009; Chapman et al., 2007; Glover-Cutter

et al., 2009; Kim et al., 2009; Zhang and Corden, 1991), and to some extent, tyrosine and

threonine residues (Baskaran et al., 1993; Zhang and Corden, 1991).

The Ser5 phosphorylated form of RNAP II is mainly localized to 5’-ends of genes where

transcription initiation occurs, whereas RNAP II containing phosphorylations of the Ser2

residue has been detected further in gene bodies and therefore associated with elongation

(Figure 2) (Cheng and Sharp, 2003; Donner et al., 2010; Glover-Cutter et al., 2009;

Morris et al., 2005). Ser7 has been demonstrated to be crucial for the transcription of

snRNAs, but not for protein-encoding genes (Egloff et al., 2007). After RNAP II has

finished transcribing, it is dephosphorylated by CTD phosphatases FCP1 (Lin et al.,

2002b) and SCP1 (Yeo et al., 2003), and the unphosphorylated polymerase can re-enter a

PIC to initiate another cycle of transcription (Figure 2). As RNAP II progresses along the

gene, the sequential phosphorylations of the CTD create a dynamic scaffold for

chromatin modifiers and mRNA processing enzymes, temporally coordinating the length

and maturity of the nascent transcript. This coupling has often been termed the ‘CTD

code’ (reviewed in Egloff and Murphy, 2008), and its implications are discussed in detail

in the secion entitled ‘mRNA processing’).

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Figure 2. Schematic representation of the CTD phosporylation cycle. (1) General transcription factors (GTFs) and RNA Polymerase II (RNAPII) assemble at the promoter (here depicted by TATA) to form the pre-initiation complex (PIC). (2) TFIIE recruits TFIIH to the PIC, which co-incides with the phosphorylation of the C-terminal domain (CTD) of RNAPII on Serine 5 residues. This results in promoter escape, and the transcription of ∼30 nucleotides of mRNA. At this stage, RNAPII can either pause or proceed onto elongation. (3) At the elongation stage, P-TEFb is recruited to the transcription site and RNAPII CTD is further phosphorylated on Serine 2 residues. This results in progression of elongation and transcription of the rest of the gene. (4) After RNAPII has finished transcribing, the CTD is dephosphorylated by phosphatases, and it can be recycled to begin a new transcription cycle (5).

Transcriptional cell-cycle dependent kinases (Cdks) are the major encoders responsible

for molding the CTD. To date, the majority of in vitro and in vivo data indicate that the

Ser2 residues are phosphorylated by Cdk9 as part of Positive transcription elongation

factor b (P-TEFb) (reviewed in Peterlin and Price, 2006). Recently, Cdk7 has been

identified as the kinase responsible for Ser7 phosphorylation (Akhtar et al., 2009; Glover-

Cutter et al., 2009; Kim et al., 2009). In vitro, Ser5 of RNAP II can be phosphorylated by

Cdk7 but in addition, a number of other kinases, such as Cdk8, Cdk9, and MAPK

(Gomes et al., 2006; Ramanathan et al., 2001; Rickert et al., 1999; Trigon et al., 1998;

Zhou et al., 2000). Since Cdk7 as part of TFIIH is recruited to promoters upon

transcription induction and coincides with Ser5 phosphorylation, Cdk7 has been one of

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the best candidates for the in vivo kinase (Cheng and Sharp, 2003; Komarnitsky et al.,

2000; Schwartz et al., 2003; Spilianakis et al., 2003). Figure 2 illustrates the RNAP II

CTD phosphorylation cycle during transcription

TFIIH and Mat1

The general transcription factor IIH (TFIIH) is a conserved protein complex required for

RNAP II-mediated transcription and for nucleotide excision repair (Drapkin et al., 1994;

Schaeffer et al., 1993; van Vuuren et al., 1994; Wang et al., 1994). TFIIH consists of ten

subunits divided into a core (p5, p34, p44, p52, p66, XPB, XPD) (Giglia-Mari et al.,

2004) and a kinase (Mat1, Cdk7, Cyclin H) submodule apparently not required for DNA

repair (Coin et al., 2008) (Figure 3). The helicase activity of the XPB core subunit is

required for general transcription at the stage of transcription initiation to melt promoter

DNA (reviewed in Saunders et al., 2006).

In mammalian cells, the kinase submodule consists of the catalytic cyclin-dependent

kinase Cdk7 (Roy et al., 1994), its cognate cyclin CycH (Serizawa et al., 1995;

Shiekhattar et al., 1995), and a third subunit Mat1 (Adamczewski et al., 1996) required

for stability of the complex in vitro (Devault et al., 1995; Tassan et al., 1995) and in vivo

(Korsisaari et al., 2002; Rossi et al., 2001) (Figure 3). This submodule has been

implicated in many functions not related to general transcription - and indeed partly

independently of core TFIIH (Li et al., 2010): cell cycle regulation as the Cdk-activating

kinase CAK (Larochelle et al., 2007; Li et al., 2010), differentiation (Patel and Simon,

2010; Wang et al., 2006), and regulation of specific transcriptional activators (Compe et

al., 2005; Keriel et al., 2002; Rochette-Egly et al., 1997).

The Mat1 subunit is a 36-37 kD protein with a hydrophobic C-terminal domain, a central

coiled-coil domain and an N-terminal RING-finger motif which contains a zinc-binding

domain (Busso et al., 2000). The RING finger has the typical ββαβ topology of the

RING family, but additionally, it accommodates a short α-helix and an extended basic

surface, which may be required for the characteristic activities and regulation of Mat1

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(Gervais et al., 2001). The RING-finger motif is associated with TFIIH-mediated

transcriptional activation; its deletion inhibits basal transcription and phosphorylation of

the CTD of RNAP II (Busso et al., 2000; Warren, 2002). Deletion mutant analysis shows

that the hydrophobic and coiled-coil domains of Mat1 join the CAK complex to the N-

terminal region of XPD (Sandrock and Egly, 2001) and perhaps also to XPB (Busso et

al., 2000), which links it to the core TFIIH through interactions with p44. The C-terminal

hydrophobic domain interacts with the Cdk7-Cyclin H dimer; according to

coimmunoprecipitation studies, Mat1 is able to bind to both subunits independently and

that this domain appears to be sufficient for Cdk7 kinase activity (Busso et al., 2000).

XPD

XPB

Mat1cycH

p34

p52

p62

p44

Cdk7

core TFIIH

Kinase submodule(CAK)

The Cdk7 submodule of TFIIH was first suggested to be involved in regulation of general

transcription when it was found to phosphorylate the C-terminal domain (CTD) of

mammalian RNAP II (Mäkelä et al., 1995; Serizawa et al., 1995; Shiekhattar et al.,

1995). The TFIIH kinase can phosphorylate Ser5 of the CTD (Gebara et al., 1997; Roy et

al., 1994; Trigon et al., 1998), and on mammalian genes localization of Ser5

phosphorylated RNAP II is consistent with promoter clearance (Cheng and Sharp, 2003;

Donner et al., 2010; Gomes et al., 2006; Morris et al., 2005). However, genetic studies on

the role of Cdk7 and the TFIIH kinase in Ser5 phosphorylation and general transcription

do not provide a unified picture even in the most extensively studied budding yeast

system, where temperature-sensitive alleles of the TFIIH kinase demonstrate dramatically

Figure 3. Cartoon of TFIIH structure. TFIIH consists of 7 subunits forming the core TFIIH (XPD, XPB, p62, p52, p44, p34, p5 (not depicted here)) and three subunits which make up the kinase submodule (Mat1, Cdk7, Cyclin H), also termed the Cdk-activating kinase (CAK). The cartoon is based on documented interactions between Mat1-XPD (Sandrock and Egly, 2001), Mat1- Cdk7-CycH(Busso et al., 2000), p44-XPB (Iyer et al., 1996), p52-XPB (Coin et al., 2007), p34-p44 (Fribourg et al., 2001) and p44-XPB (Coin et al., 1999) as well as the molecular structure of TFIIH resolved by electron microscopy (Muse et al., 2007; Schultz et al., 2000).

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reduced mRNA levels (Faye et al., 1997; Holstege et al., 1998; Valay et al., 1995). In one

study, RNAP II occupancy was dramatically decreased (Schroeder et al., 2000) whereas

another reported only a modest decrease in RNAP II occupancy despite a general loss of

Ser5 phosphorylation (Ser5-P) (Komarnitsky et al., 2000). Subsequent studies using

analogue-sensitive KIN28 (Cdk7 homolog) alleles either show no significant global

change in mRNA levels, or in occupancy, while Ser5 phosphorylation and capping are

decreased (Kanin et al., 2007) or show a significant decrease in mRNA levels without

major changes in occupancy or Ser5 phosphorylation (Hong et al., 2009).

In fission yeast Mcs6 (Cdk7 homolog) ts mutants demonstrate a moderate decrease in

total Ser5-P and a 35% reduction in median mRNA abundance (Lee et al., 2005) whereas

,inhibition of an analog-sensitive Msc6 allele leads to decreased global Ser5-P but not a

global deregulation of mRNAs (Viladevall et al., 2009). In C. elegans, partial loss of

Cdk7 resulted in decreased global Ser5-P as well as defective transcription (Wallenfang

and Seydoux, 2002). Cdk7 temperature-sensitive mutants in Drosophila in some studies

do not display transcriptional defects (Larochelle et al., 1998), whereas in others defects

have been noted in early zygotic transcription (Leclerc et al., 2000) or on the widely

studied hsp70 locus (Schwartz et al., 2003). In the latter study, a decrease in Ser5-P was

not associated with the transcriptional defects.

In mammalian systems two genetic approaches have been used to investigate the role of

the endogenous TFIIH kinase in Ser5 phosphorylation and general transcription. Despite

decreased Ser5 and Ser2 phosphorylation, no defect was noted in murine Mat1-/- embryo

outgrowths in their ability to express GFP from microinjected plasmids (Rossi et al.,

2001), and loss of Mat1 from Schwann cells did not lead to phenotypes consistent with

general transcriptional defects (Korsisaari et al., 2002). In HCT116 human cancer cells

with an integrated ATP-analogue sensitive Cdk7 allele, addition of the inhibitor (3-MB-

PP1) did not detectably block general transcription or result in Ser5-P changes at global

levels or on the studied genes c-myc and gapdh (Glover-Cutter et al., 2009; Larochelle et

al., 2007). Therefore, the evidence for TFIIH in Ser5 phosphorylation and general

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transcription across species has been controversial and the mammalian in vivo Ser5

kinase remained elusive.

Co-transcriptional mRNA processing

The transcription of a gene is not enough to produce a mature transcript that can be

translated into protein: the nascent transcript must additionally be processed into

messenger RNA (mRNA). This includes co-transcriptional capping of the protruding

5’end of the nascent mRNA, splicing of introns and polyadenylation of the pre-mRNA.

All these processes have been linked to phosphorylation of RNAP II CTD as described in

detail below.

Capping, the first step in mRNA maturation, is the process through which a guanine

nucleotide at the 5’end of the pre-mRNA is methylated on the 7-position by cap

methyltransferase (Figure 4). This 7-methylguanosine (m7G) cap is thought to provide

stability to the newly transcribed mRNA by protecting it from 5-exonucleases. In vitro,

phosphorylated RNAP II CTD stimulates capping enzyme activity (Ho and Shuman,

1999). Furthermore, capping enzymes directly associate with Ser5-phosphorylated RNAP

II CTD (Cho et al., 1997; McCracken et al., 1997a). Studies in both budding and fission

yeast demonstrate that capping enzymes are recruited by Ser5 phosphorylated RNAP II

and in turn, recruit Cdk9 to phosphorylate Ser2 (Guiguen et al., 2007; Pei et al., 2003;

Qiu et al., 2009; Viladevall et al., 2009). Similarly, in Drosophila resumption of

elongation of a paused RNAP II is correlated with Ser5 CTD phosphorylation (O'Brien et

al., 1994) and capping (Rasmussen and Lis, 1993), whereas P-TEFb and Ser2

phosphorylation are noted on genes after the pause site (Boehm et al., 2003). Therefore,

current data shows that capping occurs simultaneously to RNAP II pausing

approximately 30 bp downstream from the TSS and may therefore provide a potential

checkpoint for progression into elongation.

The majority of eukaryotic pre-mRNA is composed of non-coding sequences, introns,

which must be spliced out to form the mature mRNA (Figure 4). Eukaryotic cells contain

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two types of introns: the more common U2-type introns and the highly conserved, less

common U12-type introns, which are present in a small subset of “information

processing genes” (reviewed in Patel and Steitz, 2003). This removal of introns is

achieved by the spliceosome, which is made up of uridine-rich small nuclear RNAs (U

snRNAs), working together with over 200 auxiliary proteins, often termed splicing

factors (reviewed in Valadkhan and Jaladat, 2010). The spliceosomal snRNAs recognize

the intron-exon boundaries through base-pairing to a consensus sequence and the intron is

removed by two consecutive transesterification reactions. Phosphorylated CTD has been

shown in vitro to stimulate splicing activity (Hirose et al., 1999; Millhouse and Manley,

2005), and in vivo hypophosphorylated RNAP II co-localizes and associates with splicing

factors (Mortillaro et al., 1996). In Xenopus oocytes, blocking CTD phosphorylation with

kinase inhibitors dimishes transcription-coupled splicing (Bird et al., 2004). Spt6 mutants

that are unable to bind Ser2 phosphorylated CTD display splicing defects, suggesting one

possible mediator between splicing and CTD phosphorylation (Yoh et al., 2007).

The final step in producing a mature mRNA is polyadenylation, where the 3’end of the

transcript is first cleaved endonucleolytically after an AAUAAA sequence followed by

the addition of a poly(A) tail consisting of multiple adenosine monophosphates (Figure

4). This is catalyzed by the multisubunit enzyme complexes CstF and CPSF in mammals

(Perez Canadillas and Varani, 2003). The poly(A) tail stabilizes the mRNA and targets it

for translation and nuclear export. Budding yeast ctk1 (Ser2 kinase) mutant strains that

lack Ser2 phosphorylation are unable to recruit polyadenylation factors (Ahn et al., 2004)

and although both nonphosphorylated and phosphorylated RNAP II bind CstF and CPSF

similarly in vitro, the phosphorylated form is more active in an endonuclease cleavage

assay (Hirose and Manley, 1998).

Interestingly, comparison of truncation mutants has revealed that different segments of

the CTD independently stimulate either capping, splicing or 3’-end processing (Fong and

Bentley, 2001). This suggests that specific CTD repeats may be differentially

phosphorylated during the transcription cycle to recruit the necessary factors, providing

yet higher specificity to the CTD code.

21

Figure 4. Transcription-coupled mRNA processing. (1) A 7-methylguanosine cap is added to the protruding nascent pre-mRNA by the capping machinery to protect it from nuclease activity. The capping machinery is recruited to the site of transcription by Ser5 phosphorylated RNAP II. (2) Capping enzymes recruit P-TEFb, which phosphorylates RNAP II CTD on Ser2. This provides a potential ‘checkpoint’ whether or not to proceed onto transcript elongation. (3) Non-coding introns are removed during transcription elongation by spliceosomes. (4) A poly-A tail is added to the pre-mRNA, the mRNA is cleaved and ready to be transported to the cytosol for translation into proteins by ribosomes.

Regulation of mRNA stability and decay

In addition to transcription rate, the abundance of mRNAs is also regulated by their

stability and decay, which in turn can be reflected on the amount of protein produced

from the transcript. Typically, mRNAs with short half-lives encode proteins that need to

be produced quickly in response to external stimuli, whereas stable mRNAs with long

half-lives represent constitutively and highly expressed genes such as house-keeping

genes or maternal mRNAs (reviewed in Guhaniyogi and Brewer, 2001).

Several different mRNA decay pathways exist in mammalian cells, and they may all

work in concert or separately from one another. In the deadenylation pathway, the

protective poly(A) tail is chewed off by at least one poly(A) ribonuclease (PARN)

(Korner and Wahle, 1997; Korner et al., 1998). To hasten the process, the stabilizing

5’cap is also removed, and the mRNA is bidirectionally degraded by exonucleases. The

process of mRNA decay once again highlights the importance of proper mRNA

processing to protect the nascent transcript. The complex responsible for mRNA

22

degradation has been termed the exosome and consists almost exclusively of

exoribonucleic proteins. mRNAs can also be degraded by a deadenylation-independent

pathway. In this process, RNA endonucleases recognize specific sequences within the

mRNA and begin to chew up the transcript from within (reviewed in Guhaniyogi and

Brewer, 2001). In addition to normal mRNA turnover, the exosome functions as a quality

controller of pre-mRNA: if any of the mRNA processing steps are faulty, the mRNA is

degraded (reviewed in Raijmakers et al., 2004). Lastly, the nonsense-mediated decay

pathway (NMD) targets aberrant transcripts formed during transcription, serving as an

mRNA surveillance mechanism and preventing the formation of truncated proteins

(reviewed in Chang et al., 2007).

mRNA turnover can be regulated at individual transcript level by specific sequence

elements, the best studied to date being the adenylate-uridylate rich elements (AREs)

located in the 3’ untranslated region (3’UTR). These elements have been found in a

number of proto-oncogenes, growth factors and cytokines (Bakheet et al., 2001). External

cues signal ARE-binding proteins to localize to these elements, hence either stabilizing

them or hastening their turnover by promoting decay. For example, mRNA levels of

vascular endothelial growth factor (VEGF) rise in response to hypoxia due to binding of

the HuR protein to AREs in VEGF 3’UTR, resulting in increased stability and longer

half-life (Levy et al., 1998). In addition to AREs, other decay determinants that have been

identified include iron-response element (IRE), C-rich element and JNK-response

element (Guhaniyogi and Brewer, 2001).

In addition to transcript-specific stabilization, widespread changes in mRNA turnover

have been noted in budding yeast, where 36.2% of mRNAs show altered stability

following oxidative stress (Molina-Navarro et al., 2008). In mammalian cells, studies

comparing mRNA steady-state levels and nascent transcription following transcriptional

inhibition have suggested general stabilization of mRNAs (Dolken et al., 2008; Friedel et

al., 2009; Meininghaus et al., 2000), although this phenomenon has yet to be widely

recognized. Nevertheless, recent advances in studying nascent transcription by labeling

23

newly transcribed mRNA have highlighted the importance of mRNA turnover in the

regulation of transcript levels.

Chromatin modifications

To provide a yet higher level of regulation and specificity to gene expression, the

chromatin template itself can be modified through enzymatic activities. This comprises

DNA methylation and covalent post-translational modifications of histones: acetylation,

methylation, phosphorylation, ADP-ribosylation, ubiquitination and biotinylation

(reviewed in Vaquero et al., 2003). Histone tail modifications are typically dynamic and

transient, whereas DNA methylation can be passed on through cell divisions and

generations, and forms the core of epigenetic inheritance.

Histones are modified on their N-terminal tails, and the various modifications change

their affinity towards different proteins, determining for e.g. which transcription factors,

chromatin modifiers, and enhancers are recruited to the DNA. Acetylation is the most

extensively studied histone modification and is mostly associated with transcriptional

activation (Bernstein et al., 2005; Pokholok et al., 2005; Schubeler et al., 2004).

Therefore, it is not surprising that many transcriptional co-activators such as Gcn5

(Imoberdorf et al., 2006), p300/CBP (Ogryzko et al., 1996) and TAF250 (Mizzen et al.,

1996) contain histone acetyltransferase (HAT) activity. HATs catalyze the transfer of an

acetyl group from acetyl coenzyme A to lysine residues of H2B, H3 and H4 (reviewed in

Vaquero et al., 2003) and it is hypothesized that this modification partially neutralizes the

positively charged histones, hence loosening their grip on the DNA. The removal histone

acetylations is catalyzed by histone deacetylases (HDACs), which are therefore involved

in transcriptional repression (Hu et al., 2000).

Methylation can take place on arginine and lysine residues of H3 and H4. The transfer of

methyl groups from S-adenosyl-L-methionine (SAM) to these residues is catalyzed by

histone methyltransferases (HMTs) and conversely removed by histone demethylases

(HDMs) (reviewed in Zhang and Reinberg, 2001). Interestingly, HMTs Set1 and Set2,

24

which catalyze H3 lysine 4 (H3K4) methylation and H3 lysine 36 (H3K36) methylation

respectively, interact with phosphorylated RNAP II CTD (Krogan et al., 2003; Li et al.,

2003; Xiao et al., 2003), providing a direct link between sequence-specific regulation and

the general transcription machinery. In particular, H3K4 trimethylation appears to require

Cdk7 kinase activity in yeast (Ng et al., 2003). Genome-wide studies have shown that

H3K4 methylation is associated with active transcription and occurs in the vicinity of

transcription start sites, whereas H3K36 methylation occurs in the bodies of actively

transcribed genes (Bernstein et al., 2005; Guenther et al., 2007; Pokholok et al., 2005;

Santos-Rosa et al., 2002). Methylated H3K9 on the other hand, has been shown to

localize to sites of repressed chromatin (Ebert et al., 2006).

Additionally, histones can undergo phosphorylation, ubiquitination and ADP-

ribosylation, which to date have not been as extensively studied. All histones have been

shown to be phosphorylated, and the phosphorylation of H1 and H3 has been linked to

chromosome condensation and segregation (Guo et al., 1995). Additionally, in mammals,

phosphorylated H3 correlates with transcriptional activation (Thomson et al., 1999).

Ubiquitination occurs on lysine residues of at least H2A, H2B, H3 and H1 (Belz et al.,

2002). Active transcription has been associated with monoubiquitinated histones (Davie

et al., 1991). ADP-ribosylation is achieved on all histones by poly-ADP-ribosylate

protein 1 (PARP1) (de Murcia et al., 1988) and has been linked to aging, as the amount of

ribosylated histones has been shown to decrease with age (Mishra and Das, 1992).

DNA methylation occurs on cytosine residues on CpG islands and is catalyzed in

mammals by DNA methyltransferases (DNMTs) Dnmt1, Dnmt1, Dnmt3a and Dnmt3b

(reviewed in Svedruzic, 2008). The latter two DNMTs are involved in de novo

methylation whereas Dnmt1, the most abundant of the three in mammalian cells,

maintains methylation patterns on CpG dinucleotides during DNA replication (reviewed

in Dhe-Paganon et al., 2011). Genome-scale DNA methylation maps and in silico

prediction studies suggest that the majority of the mammalian genome is in fact

methylated, with only 1-2% unmethylated domains consisting mostly of CpG islands,

1000 bp long CG-rich stretches often occurring at promoters (Bird et al., 1985; Eckhardt

25

et al., 2006; Illingworth et al., 2008; Rabinowicz et al., 2003; Rakyan et al., 2004; Suzuki

et al., 2007). Although the role of intragenic methylation is still unresolved, methylation

at CpG islands of promoters is typically associated with transcriptional silencing

(reviewed in Suzuki and Bird, 2008). Although typically DNA methylation has been

thought of as a relatively stable, inherited form of repression, recent evidence

demonstrated that certain promoters are methylated in a transient cyclical fashion with

rapid kinetics, which directly influences RNAP II occupancy and transcription

(Kangaspeska et al., 2008; Kim et al., 2009; Metivier et al., 2008). Dnmt1 deregulation as

well as DNA hyper- and hypomethylation states have been associated with cellular

differentiation as well as numerous diseases such as cancer and diabetes (reviewed in

Dhe-Paganon et al., 2011).

26

II. Transcriptional regulation of energy metabolism

Although almost all cells in the human body contains the same information within its

DNA, over 250 different cell lineages exist with distinct morphologies, varying protein

compositions and highly divergent functions. This fine-tuning of gene expression is

achieved largely through RNAP II regulation by cell and stimuli-specific transcription

factors as well as transcriptional co-activators, many of which contain enzymatic

chromatin modifying activities described previously. One of the major challenges is the

maintenance of energy balance, requiring constant attention of energy intake and energy

expenditure in order to sustain homeostasis both in cells and tissues to avoid conditions

such as obesity or the metabolic syndrome. Different cell types react in very diverse ways

to alterations in metabolic state through regulating catabolic (breaking down) and

anabolic (building up) metabolic pathways. Since the energy requirements of specific

tissues within the body as well as in response to different external and nutritional stimuli

differ greatly, energy metabolism is an excellent example of transcriptional regulation by

specific transcription factors, co-activators and chromatin modifiers.

The transcriptional cascade of adipocyte differentiation

Two types of adipose tissue exist in mammals: brown adipose tissue (BAT) and white

adipose tissue (WAT). WAT forms the main fat depots in the body, the subcutaneous and

visceral fat, and serves as the major energy store for situations such as fasting, whereas

BAT contains high amounts of mitochondria and less fat droplets and is largely absent

from humans. For decades, adipose tissue was disregarded by scientists due to its

apparent function mainly as an insulator, a storage of energy and cushion system of the

body. However, it is now recognized as an endocrine organ, secreting peptide hormones

such as leptin and resistin along with fatty acids that, in addition to regulating appetite

and energy metabolism, affect bone mass, heamatopoesis, and blood pressure (reviewed

in Kershaw and Flier, 2004). With the explosion of obesity and related metabolic diseases

such as type II diabetes within the last 30 years in the Western world, numerous scientists

27

have concentrated their efforts into understanding the adipose tissue and deciphering how

cells differentiate into adipocytes.

Differentiation of mesenchymal preadipocytes to mature adipocytes (adipogenesis)

involves contact inhibition followed by hormonal stimuli such as insulin/IGF-1, which

leads to the expression of key transcription factors, namely PPARγ, Krox-20 and

CCAAT/enhanced binding proteins (C/EBPs) (Chen et al., 2005). Their expression

coincides with two rounds of mitosis termed clonal expansion (Fajas et al., 2002; Lane et

al., 1999; Tang et al., 2003). Following clonal expansion, the cells are committed to

differentiate into adipocytes through the expression of adipogenic genes such as

adiponectin, adipsin and aP-2, which result in the accumulation of lipids and eventually,

in the fully differentiated adipocyte phenotype (reviewed in Rosen and MacDougald,

2006). Relatively little is known about the differentiation of brown adipocytes, but

PPARγ-coactivator-α (PGC1-α) appears to play a role in it. When PGC-1a is introduced

in white adipocytes, they begin to resemble their brown counterparts; expression and

activity of UCP-1, an enzyme specific for brown adipocytes, is increased and

mitochondrial biogenesis initiated (Puigserver et al., 1998; Tiraby et al., 2003). The

traditional view of the process of adipocyte differentiation is illustrated in Figure 5.

The primary transcript of the PPARγ gene is alternatively spliced to encode three

different protein isoforms: PPARγ1 which is ubiquitously expressed, PPARγ3 which

dominates in the large intestine and PPARγ2 which is predominant in white adipose

tissue (Braissant et al., 1996). PPARγ2 has been coined the master regulator of

adipogenesis as it is both sufficient and essential for this process (Rosen et al., 1999;

Tontonoz et al., 1994). Upon activation through binding of endogenous metabolites

acting as ligands, it heterodimerizes with retinoid X receptor (RXR) and binds to PPAR

response elements (PPREs) on target genes, thus initiating transcription (Chandra et al.,

2008). PPARγ2 is a phosphoprotein that can be phosphorylated on S112 by mitogen-

activated protein kinase (MAPK) (Adams et al., 1997; Camp and Tafuri, 1997) and Cdk7

in vitro (Compe et al., 2005). However, in contrast to activating phosphorylation

observed in the majority of nuclear hormone receptors, the phosphorylation of PPARγ on

28

S112 inhibits its activity (Adams et al., 1997; Camp and Tafuri, 1997; Hu et al., 1996).

Furthermore, mutated PPARγ-S112A in preadipocyes results in increased adipogenesis

(Hu et al., 1996). In addition to ligand binding and phosphorylation, PPARγ activity is

further regulated in some tissues by co-activators such as PGC-1 and SRC-1 (Puigserver

et al., 1999).

Figure 5. The process of adipocyte differentiation. Undifferentiated cells that have undergone growth arrest can differentiate into adipocytes upon induction by hormones or in response to external stimuli. The induced cells undergo clonal expansion, which commits them to the adipogenic lineage. Expression of adipogenic transcription factors results in the fully differentiated adipocyte characterized by large lipid vacuoles and increased mitochondria. PGC-1α expression directs the differetiation process towards brown adipocytes, which contain a higher amount of mitochondria and many small lipid droplets. The required adipogenic transcription factors are highlighted in red and placed at the differentiation stage when their expression peaks.

PPARγ has attracted much clinical interest since it was identified to be the target of

synthetic antidiabetic drugs called thiazolidinediones (TZDs) (Lehmann et al., 1995).

TZDs exert their effect by reducing hyperlipidemia and insulin resistance (reviewed in

Olefsky, 2000). Paradoxially, they activate PPARγ and thus induce adipogenesis

(Lehmann et al., 1995). Although the amount of adipocytes increases in response to TZD

treatment, the size of the new adipocytes is smaller, they exhibit characertistics of brown

adipocytes and appear to be metabolically more active, thereby potentially restoring the

insulin sensitivity of fat (Wilson-Fritch et al., 2004; Yamauchi et al., 2001). In support of

this, although PPARγ-S112A mice demonstrate an increased potential for adipogenesis,

29

they do not gain weight and show increased insulin sensitivity (Rangwala et al., 2003).

Hence, it is now widely recognized that the regulation of PPARγ in vivo is more complex

than previously thought and that it may be clinically beneficial to activate PPARγ more

selectively through modulating its phosphorylation on S112 (reviewed in Cock et al.,

2004).

The PPARγ co-activator-1 family members and fatty acid oxidation

The PPARγ co-activator-1 (PGC-1) family members are transcriptional co-activators that

bind to a myriad of transcription factors, increasing their ability to stimulate transcription

of genes involved in glucose, lipid and energy homeostasis. The two major isoforms

present in tissues with high energy requirements (heart, liver, skeletal muscle and brown

fat) are PGC-1α and PGC-1β. Originally, PGC-1α was discovered as a thermogenic

switch in brown fat, where it is required to stimulate fuel intake, fatty acid oxidation and

heat production (Puigserver et al., 1998), but it is now recognized that dysregulation of

these coactivators plays an important part in the development of pathophysiologies such

as heart failure and diabetes (reviewed in Lin et al., 2005).

PGC-1 co-activators facilitate the transcription of mitochondrial fatty acid oxidation

(FAO) genes. These co-activators do not contain a DNA-binding domain, but are

mobilized to target genes by tissue-specific transcription factors like PPARγ (brown fat)

(Tiraby et al., 2003), muscle enhancer factor 2 (MEF2) (Handschin et al., 2003) and

hepatocyte nuclear factor 4α (HNF4α) (Rhee et al., 2006) or ubiquitous factors such as

nuclear respiratory factors (NRFs) (Wu et al., 1999). In turn, they recruit chromatin-

modifying enzymes, for example HATs p300 and steroid receptor co-activator (SRC-1)

(Puigserver et al., 1999) and the Mediator complex to initiate transcription (Wallberg et

al., 2003). PGC-1 co-activators are themselves tightly regulated by environmental cues

and nutritional stimuli. For example, strenuous exercise increases PGC-1α mRNA levels

in skeletal muscle (Baar et al., 2002; Goto et al., 2000), and in brown fat PGC-1α mRNA

is induced in response to cold exposure (Puigserver et al., 1998).

30

The heart is the largest consumer of energy in the body taking into account its size. The

majority of ATP produced in the heart is a result of mitochondrial β-oxidation of fatty

acids, with 5-10% coming from glycolysis. It is not surprising that due to its large

consumption of ATP, both PGC-1α and PGC-1β are very highly expressed in the heart

(Lin et al., 2002a; Puigserver et al., 1998). PGC-1α is especially important in the

neonatal developing heart when the workload and energy requirement of the heart

dramatically increase and a metabolic switch from glycolysis to fatty acid oxidation

occurs (Lehman et al., 2000). In the heart, PGC-1α promotes the transcription of genes

required for mitochondrial biogenesis through direct interaction with NRFs and ERRs

and of genes involved in fatty acid import and utilization through PPARα (reviewed in

Rowe et al., 2010). In addition, PGC-1α activates a large array of angiogenic factors,

including VEGF through ERRα (Arany et al., 2008), providing a link between high

energy expenditure and the vasculature which supplies the heart with nutrients.

The role of mitochondrial dysfunction in cardiomyopathy and heart failure is evident

from genetic human diseases: over 50% of mitochondrial DNA mutations and most

mutations of the nuclear-encoded genes for fatty acid oxidation and transport result in

cardiomyopathy (reviewed in Rowe et al., 2010). Mouse models of cardiac dysfunction

have suggested that PGC-1α would be at least one common culprit, as a number of these

mice exhibit decreased expression of PGC-1α (Garnier et al., 2003; Palomer et al., 2009;

Sano et al., 2004). Additionally, heart-specific PGC-1α knockout mice show decreased

fatty acid oxidation, abnormal mitochondria, reduced ATP synthesis and eventually, at

two months of age, cardiac failure (Arany et al., 2005; Arany et al., 2006; Lehman et al.,

2008).

Transcriptional control of hepatic lipogenesis

Mammalian lipid homeostasis is largely controlled by a family of membrane-bound

transcription factors, the sterol regulatory binding-element proteins (SREBPs). SREBPs

exist as three isoforms generated from two genes; SREBP-1a and SREBP-1c encoded by

the same gene but through utilization of a different transcription start site, and SREBP-2

31

(Sato). SREBP-1a is mainly expressed in cultured cells, SREBP-1c in hepatocytes and

muscle and SREBP-2 ubiquitously in mammals (Shimomura et al., 1997). Transgenic and

knockout mouse models of SREBPs have demonstrated that SREBP-1c is the chief

regulator of lipogenesis in the liver (Shimano et al., 1997a; Shimomura et al., 1999;

Shimomura et al., 1998; Tobe et al., 2001), whereas SREBP-2 governs cholesterol

synthesis (Horton et al., 1998; Shimano et al., 1997b).

Figure 6. Regulation of lipid and cholesterol metabolism by SREBPs. Acetyl CoA is produced from glycolysis of glucose in the mitochondria and used for synthesis of fatty acids or cholesterol. SREBP-1c regulates the transcription of the genes coding for the main enzymes required for the synthesis of triglycerides and phospholipids (depicted in grey). SREBP-2 regulates the expression of the genes involved in cholesterol synthesis. In addition, the main enzymes that catalyze the production of NADPH are encoded by SREBP-1c target genes. ACS; acetyl CoA synthase, ACL; acetyl CoA lyase, ACC; acetyl CoA carboxylase, FAS; fatty acid synthase, Elovl6: elongation of long chain fatty acids family member 6, SCD1; stearoyl CoA desaturase, GPAT; glycerol phosphate acyltransferase, DGAT; diacylglycerol acyltransferase, ME1; malic enzyme, G6PD; glucose-6-phosphate dehydrodenase, PGDH; phosphogluconate dehydrogenase.

32

As inactive precursors, SREBPs are bound to the endoplasmic reticulum. Upon stimulus

such as increased cholesterol levels, the SREBP cleavage-activating protein SCAP

transfers SREBP to the Golgi apparatus, where it is cleaved by Site-1 and Site-2

proteases (Matsuda et al., 2001). The processed and active SREBPs are transported to

the nucleus where they can bind to sterol regulatory elements (SREs) in promoters of

target genes. These genes include all the main enzymes of the fatty acid biosynthetic

pathways depicted in Figure 6, including ACC1, ACLY, FAS, G6PD, GPAT, and SCD1

(Shimomura et al., 1998).

Hepatosteatosis is the accumulation of triglycerides in the liver. It can exist as a benign,

non-inflammatory form or progress on to steatohepatitis with strong inflammation, cell

death and fibrosis. Insulin resistance, a characteristic of type II diabetic patients, is the

most common metabolic problem associated with hepatosteatosis. In the classic model,

insulin-resistant adipocytes increase their secretion of fatty acids, which are then taken up

by hepatocytes, resulting in accumulation of fat in the liver (reviewed in Browning and

Horton, 2004). Findings in the last decade also point towards a de novo pathway of

hepatosteatosis formation, where increased synthesis of fatty acids in the liver takes place

due to transcriptional activation of lipogenic SREBP-1c target genes (Shimomura et al.,

1999). For example, transgenic mice that overexpress SREBP-1c in the liver develop a

classic fatty liver due to activation of lipogenic genes (Shimano et al., 1997a). Likewise,

absence of SREBP-1c ameliorates the fatty liver phenotype of diabetic leptin receptor

knockout mice (Yahagi et al., 2002). In addition to the well-documented role of SREBP1

in hepatosteatosis, the carbohydrate response element binding protein (ChREBP) and

PPARγ have been shown to participate in the formation of fatty liver (Iizuka et al., 2004;

Kim et al., 1998), although these molecular pathways have not been fully defined.

Importantly, PPARγ has been suggested to function downstream of SREBP-1c, as

SREBP-1c can transcriptionally activate PPARγ (Fajas et al., 1999; Kim et al., 1998)

Although PPARγ, PGC-1 and SREBP1 have been discussed here in a rather tissue-

specific context for structural purposes of the thesis, it is important to note that in some

tissues as well as cell culture and disease models, they in fact co-operate to promote the

33

transcription of lipogenic, adipogenic and FAO genes. In addition, in some models they

induce the expression of one another and also regulate each other via

feedback/feedforward loops. Novel concepts as well as potential caveats that have risen

during the thesis work will be elaborated on in ‘Discussion and Perspectives’.

TFIIH: Linking general and gene-specific transcription

In addition to its role as a general transcription factor, TFIIH has been implicated as a

regulator of specific transcription factors, in particular the nuclear hormone receptors.

Nuclear hormone receptors (NHRs) include androgen receptor (AR), progesterone

receptor (PR), glucocorticoid receptor (GR), estrogen receptors α and β (ER) as well as

the non-steroid receptors retinoid acid receptors (RARs), peroxisome proliferator-

activated receptors (PPARs) and vitamin D receptor (VDR). All NHRs are structurally

similar within the C-terminal domain: they all contain a ligand-binding domain (LBD),

which encompasses a ligand-dependent transactivation domain (AF-2) followed by a

DNA binding domain (DBD). The A/B domain of the N-terminal region on the other

hand is highly variable both in length and sequence (reviewed in Rochette-Egly, 2003).

In the absence of a ligand, nuclear hormone receptors are inactive and typically bound by

chaperone proteins. Upon ligand binding, they dissociate from the chaperones, dimerize

and translocate to the nucleus where they bind response elements on the DNA of their

target genes. Binding to the response elements brings the receptors to the vicinity of

TFIIH: appropriately, Cdk7 has been identified to phosphorylate a number of these

receptors on their A/B domain. In addition to the previously described PPARγ, these

include PPARα, retinoic acid receptors α and γ and estrogen receptor α (Bastien et al.,

2000; Chen et al., 2000; Compe et al., 2005; Keriel et al., 2002; Rochette-Egly et al.,

1997). Many of the nuclear hormone receptors have additionally been shown to be

phosphorylated by other kinases such as MAPK, Akt, PKC, PKA and JNKs (reviewed in

Rochette-Egly, 2003).

34

In addition to regulating nuclear hormone receptors, TFIIH has been implicated in the

phosphorylation of various other transcription factors. TFIIH enhances the binding of

tumor suppressor p53 to DNA through phosphorylation (Ko et al., 1997; Lu et al., 1997).

Cdk7 is able to directly phosphorylate elongation factors Cdk11 and Spt5 (Larochelle et

al., 2006), Cdk5 (Rosales et al., 2003) and the octamer binding transcription factor Oct-1

(Inamoto et al., 1997). These data imply that TFIIH may have more specific, regulatory

roles in different tissues, cell-types or in response to certain stimuli and that this protein

complex could serve as a link between general and gene-specific transcription initiation.

35

AIMS OF THE STUDY This study was undertaken to study the role of the mammalian TFIIH kinase complex and in particular the Mat1 subunit in the regulation of general and gene-specific transcription of RNA Polymerase II transcribed genes. The work utilizes the Cre-LoxP system to conditionally delete Mat1 in various tissues in vivo and as well as in cell culture models. Specific aims of the study were to characterize: I. Investigate the possible general role of the mammalian TFIIH kinase in RNA Polymerase II C-terminal domain phosphorylation, transcription and co-transcriptional processes. II. Idenfity and characterize possible gene-specific and tissue-specific functions for the TFIIH kinase using conditional deletion of the Mat1 subunit during adipocyte differentiation in vitro, and in cardiomyocytes and hepatocytes in vivo.

36

MATERIALS AND METHODS The materials and methods used in this study are listed below and are described in detail in the original publications, which are here referred to using Roman numerals. 1.Materials Mouse lines Description Source or reference Used in αMHC-Cre cardiac-specific Cre, expressed

under myosin heavy chain promoter Gaussin et al, 2002 III

C57BL/6J-Tg(Mx1-cre) liver-specific Cre, expressed under Mx1 promoter JAX Laboratories IV

Mat1 flox Mat1 allele with loxP sites, deletion inducible by Cre Korsisaari et al, 2002 I-IV

Cells Description Source or reference Used in Primary MEFs mouse embryonic fibrobasts prepared by author I-III Immortalized MEFs MEF cell line immortalized

with Retro-p53 prepared by author I-II HepG2 human hepatocarcinoma ATCC

cell line IV 3T3-L1 mouse preadipocyte cell line ATCC II Phoenix 293T human embryonic kidney ATCC

cell line II Plasmids/constructs Description Source or reference Used in GST-CTD triple repeat of RNAPII CTD

fused to GST provided by Dr. Young II GST-Cdk2 Cdk2 fused to GST provided by Dr. Young II PPRE3-TK-Luc 3 x PPRE fused to luciferase provided by Dr.

Chatterjee II pCMV-SPORT6- PPARγ2 PPARγ2 expressed under

CMV promoter MGC Gene collection II pSV-SPORT- PPARγS112A non-phosphorylated PPARγ2 mutant provided by Dr.

Spiegelman II Antigens (Name) Description Source or reference Used in Actin (AC-40) rabbit polyclonal ab Sigma I-IV BrdU (BU33) mouse monoclonal ab Dako II, III Cdc2-T161-P (9114) rabbit polyclonal ab Cell Signaling II Cdk2-T160-P (2561) rabbit polyclonal ab Cell Signaling II Cdk2 (sc-163) rabbit polyclonal ab Santa Cruz II

37

Cdk7 (C-4) mouse monoclonal ab Santa Cruz I-IV C/EBPβ (ab15049) mouse monoclonal ab Abcam II DMAP1 (ab2848) rabbit polyclonal ab Abcam IV Dnmt1 (ab13537) mouse monoclonal Abcam IV Mat1 (FL-309) rabbit polyclonal ab Santa Cruz I-IV m7G-cap (K121) mouse monoclonal ab Calbiochem I catalase (ab1877) rabbit polyclonal ab Abcam I, IV p44 rabbit polyclonal ab Genway I p62 rabbit polyclonal ab Genway I PPARg (ab41928) mouse monoclonal ab Abcam II PPARg-S112-P (IF5) mouse monoclonal ab Euromedex II PGC-1 rabbit polyclonal ab US Biologicals III RNAPII (N20) rabbit polyclonal ab Santa Cruz I,III,IV RNAPII-S2 (H5) mouse monoclonal ab Nordic Biosite I, III RNAPII-S5 (H14) mouse monoclonal ab Nordic Biosite I, III RNAPII-S5 (ab5131) rabbit polyclonal ab Abcam I SREBP-1 (ab28481) rabbit polyclonal ab Abcam IV TAF10 mouse monoclonal ab provided by Dr. Tora II XPD mouse monoclonal ab provided by Dr. Egly I Viruses Description Source or reference Used in AdCre encodes Cre recombinase provided by Dr. Parada I, II, III AdGFP encodes GFP provided by Dr. Parada II AdLacZ encodes LacZ provided by Dr. Zhu I, II, III Retro-p53 encodes CTD of p53 provided by Dr. Klefstrom II Retro-Mat1 encodes Mat1 Biomedicum Virus Core II siRNA/shRNA Description Source or reference Used in shRNA-K7/pENTR-H1 short hairpin RNA towards

human Cdk7 prepared in Mäkelä lab II shControl/pENTR-H1 short nontargeting RNA prepared in Mäkelä lab II siMat1 (mouse) Mat1 targeting oligos:

L-047470-00-0005 Dharmacon III siMat1 (human) Mat1 targeting oligos:

L-003281-00-0005 Dharmacon IV siDMAP1 (human) DMAP1 targeting oligos: D-001810-10-05 Dharmacon IV siSREBP-1 (human) SREBP-1 targeting oligos:

L-006891-00-0005 Dharmacon IV siNONTARGETTING nontargeting oligos: D-001810-10-20 Dharmacon I, II, IV 2. Methods Method Used in Adenoviral infection I-III Adenovirus production I-III Adipocyte differentiation II Affymetrix expression profiling I, III

38

BrdU labeling II Cell culture I-IV Chromatin immunoprecipitation (ChIP) I, IV Flow cytometry analysis III Generation of immortalized MEFs II Glucose and fatty acid oxidation assay I GST-pulldown II, III Immunofluorescence analysis (IF) I-III Immunohistochemistry (IHC) I, III, IV Immunoprecipitation I-III Isolation of primary MEFs I-III Luciferase reporter assay I-III In vitro kinase assay I, II Microscopy I-IV Mitochondrial enzyme and respiration assays III Methyl-cap mRNA IP I Mouse breeding /crossing II-IV Nascent mRNA analysis with FU labeling I Nascent mRNA analysis with EU labeling I Real-time quantitative PCR I-IV Retroviral transduction II RNA extraction I-IV siRNA knockdown I, II, IV Splicing analysis I Oil RedO staining II Poly(A) mRNA IP I pI-pC mouse injections IV Transfection of cells I-IV Triglyceride assay IV Western blot analysis I-IV

39

Methods

The following section describes in detail the main methods used by the author.

Adipocyte differentiation of MEFs and 3T3-L1 cells (II)

Two-day post-confluent MEFs or 3T3-L1 preadipocytes were treated with differentiation

medium (Dulbecco’s modified Eagle medium with 10% calf serum supplemented with 10

µg/insulin, 1 µM dexamethasone, 0.25 mM 3-isobutyl-1-methylxanthine) for 2 days,

followed by 2 days of insulin-supplemented medium, followed by 4 days in normal

growth medium. For analysis of lipids, cells were fixed after the differentiation protocol

with 10% formalin for 30 min at +37°C and stained with Oil Red O (Sigma).

Gene expression profiling and data analysis (I, III)

mRNAs were collected from tissues or cells using RNEasy Kit (Qiagen). cDNA synthesis

using oligo dT primers, target labeling, and hybridization were done using Affymetrix

GeneChip Reagents according to manufacturer’s protocol using Affymetrix high-density

Mouse Genome 430 2.0 Arrays, and scanned with an Agilent microarray laser scanner.

Data from all arrays has been deposited at ArrayExpress (www.ebi.ac.uk/microarray-

as/ae/). Data analysis and comparison of the data set to other published microarray

studies were performed using GeneSpring GX (Agilent Technologies), Microsoft Excel,

and Filemaker Pro software. For standard comparison of differentially regulated genes at

a given time point samples arranged by genotype were filtered for expression (at least 1/6

raw values between 20-100% percentile for inclusion), for significance (paired T-test,

p<0.05) and fold change (>1.5). For analysis of general changes in signal intensities in

MEFs (I), a normalization between genotypes at the 72 h time point was performed using

a normalization factor generated through pairwise comparisons of triplicate averages of

45 Probe Sets: AFFX labeling controls (28 probes), AFFX prelabeled hybridization

controls (excluding AFFX-Cre due to the Adeno-CRE infection; 14 probes) and AFFX

mouse non-PolII/PolI controls (3 probes) leading to a normalization factor of 1.0374 for

Mat1-/- samples.

40

Analysis of capped, FU labeled and EU labeled transcripts (I)

Capped and methylated mRNAs were immunoprecipitated from total RNA using α-m7G-

cap antibody (H20; K121 Calbiochem) in cap binding buffer (150 mM NaCl, 0.1% NP-

40, 10 mM Tris, pH 8.0) and kept on ice for 1 h. After this, the samples were incubated

overnight at + 4°C with 100 µl of Protein A Sepharose beads + 5 µl DTT + 1.25 µl

RNAseOUT. The following day, the samples were washed five times with ice-cold

binding buffer containing 2.5 mM DTT, beads were resuspended in 200 µl of ProtK

solution and nutated at + 37°C for 30 min. The precipitated RNA was purified, measured

and calculated as % of input RNA. Immunoprecipiated material was additionally

analyzed with qRT-PCR.

For 5-Flourouridine (FU) experiments, cells were pulsed with 1 mM FU in KH buffer (30

mM KCl, 10 mM HEPES pH 7.4) for 10 min and pulse-chased for various times.

Immunoprecipiattion was performed using α-BrdU (DAKO), otherwise following the

same protocol as for the capping analysis described above. Immunoprecipitated material

was additionally analyzed with qRT-PCR.

5-Ethinyl uridine (EU) labeling experiments were done according to the protocol of

Click-iT Nascent RNA Capture kit (Invitrogen). Briefly, cells were pulsed with 0.5 mM

EU, total RNA was isolated, and used in a copper catalyzed click reaction with azide-

modified biotin. The nascent transcripts were captured using streptavidin magnetic beads.

cDNA synthesis was performed using the immunoprecipitated material directly on the

beads using Superscript VILO cDNA synthesis kit (Invitrogen) followed by analysis with

qRT-PCR.

Chromatin immunoprecipitation (I, IV)

For chromatin immunoprecipitation of cells (ChIP) 5-10 x 106 cells were cross-linked

with 1% paraformaldehyde (PFA) for 10 minutes at room temperature. The cross-linking

reaction was terminated with glycine and cross-linked cells washed 3 x with phosphate

buffered saline (PBS). For ChIPs from tissues, 15-30 mg liver pieces were cut into 1 mm2

pieces and fixed with 1% PFA for 15 minutes at room temperature, terminated with

glycine and samples washed 3 x 5 minutes with PBS. Fixed tissues were lysed with

41

Lysing Matrix D beads using Precellys-24 and sonicated. Immunoprecipitation was

performed using 5-10 µg of antibodies described in Materials in rotation overnight in +

4°C. Binding was performed for 3 hours at + 4°C with 50% slurry protein-A sepharose

(PAS) followed by washes once with low salt buffer (20 mM Tris-HCl pH8.0, 150 mM

NaCl, 2 mM EDTA, 1% Triton-X100, 0.1% SDS), 3 times in high salt buffer (20 mM

Tris-HCl pH8.0, 500 mM NaCl, 2 mM EDTA, 1% Triton-X100, 0.1% SDS), once in

LiCl containing buffer (10 mM Tris-HCl pH8.0, 250 mM LiCl, 1 mM EDTA, 1% DOC,

1% NP-40) and twice with TE, pH 8.0. Elution of precipitated proteins was performed

using elution buffer (0.1 M NaHCO3, 1% SDS) containing 50 µg of proteinase K

(Fermentas) at + 42°C for 45 min. Reverse cross-linking of precipitated chromatin was

done at + 67°C overnight with NaCl solution to 200 mM final concentration. Controls:

antibodies: rabbit IgG and peroxisomal α−catalase (ab1877), negative genomic region:

SBiosciences intergenic negative control primers. The ChIP samples were purified using

QIAquick PCR purification kit spin columns (Qiagen) and eluted twice in 10 mM Tris-

HCl pH8.5 buffer. Immunprecipitated material was analyzed with qRT-PCR, normalized

to IgG/control antibody and calculated as % input.

Generation of immortal Mat1-/flox MEFs (I-III)

MEFs were isolated from E12.5 Mat1-/flox embryos and cultured in DMEM (Gibco)

supplemented with 10% fetal calf serum and antibiotics. Immortal Mat1-/flox MEFs were

generated by infecting cells from passage 3 with a retrovirus encoding residues 302 to

309 of p53 (Klefstrom, J et al, 1997) and selected with hygromycin (Invitrogen). For

deletion of Mat1, MEFs were infected with adenoviruses encoding CRE-recombinase

(AdCre) or LacZ (AdLacZ) or GFP (AdGFP) using multiplicity of infection of 1500

overnight at 37 °C.

Immunostaining of murine tissue and cultured cells (I-IV)

For paraffin and frozen sections, mice were fixed using intracardiac perfusion with 1%

PFA under anesthesia. Tissue sections were washed with PBS, permeabilized using 0.3%

PBS-Triton X (Fluka Biochemicals) and blocked with PBS containing 5% donkey serum,

0.05% sodium azide, 0.2% bovine albumin serum, and 0.3% Triton X. The tissue slides

42

were incubated with antibodies described in Materials diluted in blocking solution

overnight, followed by incubation with secondary antibodies. Nuclei were stained with

DAPI. Immunofluorescence samples were mounted with Vectashield mounting medium

(H-1200, Vector).

For immunostaining of cells, cells were grown on coverslips, fixed with 3.5% PFA 15

min and washed with PBS. Fixed cells were permeabilized with PBS-0.25% Triton X for

10 min and blocked with PBS-0.25% Triton X-5% normal goat serum for 20 min. Cells

were incubated with primary antibodies listed in Materials diluted in blocking solution

for 1 h at room temperature, and with secondary antibodies diluted in blocking buffer for

30 min at room temperature. Nuclei were stained with Hoechst and coverslips mounted

with Elvanol.

Tissue samples were analyzed with a confocal microscope (Zeiss LSM 510; Carl Zeiss)

and cell samples using Zeiss Axioplan 2 microscope and Axiovision software.

Quantitative real time PCR (qRT-PCR) analysis of mRNA levels (I-IV)

RNA was isolated using RNEasy isolation kit (Qiagen) according to the manufacturer’s

protocol. Double-stranded complementary DNA was amplified using Power SYBR

Green PCR Master Mix (Applied Biostystems). Relative mRNA amounts were assayed

by normalizing PCR cycles to GAPDH for each sample unless otherwise indicated using

7500 Fast Real-Time PCR System Software or BioStep1 plus RT-PCR Software

(Applied Biosystems). Technical triplicates were run from 3-7 independent experiments

in all cases.

Transfections and cell treatments (I-IV)

For siRNA knockdown experiments, Mat1-/flox MEFs were transfected with 100 nmol

siRNA diluted in OptiMEM (Gibco) and incubated with Lipofectamine 2000 (Invitrogen)

following the manufacturer's instructions. Transfections were repeated 24 h later. RNA

was collected 72 h from first transfection. siRNAs used are listed in the Materials chart.

For reporter gene assays, the Luciferase constructs desrcribed in Materials were

transfected transiently using Superfect transfection reagent (Qiagen) or Effectene

(Qiagen) according to manufacturer’s instructions. Troglitazone treatment of MEFs

43

(Cayman Chemical Company) was done at 10 µM from 4-24 h. 5-Aza-cytidine (Sigma)

treatment of HepG2 cells was with 10 µΜ for 48 h.

Triglyceride analysis

Homogenization of 15-30 mg of liver was done in 10% Tween-H2O with Precellys-24

machine (Bertin Technologies) at settings 5000, 2 x 20 sec, after which the samples were

incubated in +70C for 5 min. After centrifugation at 5000 rpm for 1 min, the supernatant

was collected into a new tube and centrifuged again at 14,000 rpm for 3 minutes.

Triglycerides were measured from the supernatant using Triglyceride kit

(ThermoScientific) at OD520 and calculated as ratio to protein from the same supernatant.

Western blotting and kinase assays

For Western blotting analysis of proteins MEFs were lysed in Laemmli sample buffer and

centrifuged at 13 000 rpm for 15 min. Cleared samples were run in SDS-polyacrylamide

gels and blotted onto nitrocellulose membranes (Protran) using either wet blotting

overnight (Applied Biosystems) or semi-dry blotting for 1.5 h (Owl Scientific).

Membranes were incubated with antibodies from Materials overnight in 5% milk-PBS,

detected with secondary antibodies visualized with ECL reagent (Thermo Scientific).

For in vitro kinase assays, freshly prepared cell lysates were immunoprecipitated with

α−Cdk7 and 50% PAS slurry for 1.5 h in + 4°C. Kinase activity of immunoprecipitates

was measured against in vitro GST substrates described in Materials in the presence of 20

µCi γ-32PATP in 20 mM Tris-HCL, pH 7.5, 50 mM KCl, 5 mM MgCl2 with 2.5 mM

MnCl2 and 1 mM DTT. Samples were resolved in SDS-polyacrylamide gels and detected

with autoradiography.

44

RESULTS

1. Mat1 is required for phosphorylation of RNA Polymerase II C-terminal domain,

transcription and efficient mRNA turnover (I)

The contradictory evidence from different model organisms concerning the in vivo RNAP

II CTD Ser5 kinase and the role of TFIIH in transcription prompted us to undertake a

study of global transcriptional effects following temporal deletion of the Mat1 subunit of

TFIIH in mouse embryonic fibroblasts (MEFs). This was achieved by expressing Cre

recombinase through adenoviral infection in MEFs carrying a floxed Mat1 allele

(Korsisaari et al., 2002). Within 48 h following AdCre infection, Mat1 as well as Cdk7

protein levels were undetectable by Western blot, and at 72 h, phosphorylation of both

Ser5 and Ser2 of RNAP II CTD were dramatically reduced at global levels (down to 17%

and 21%, respectively). In addition, Ser5-P was decreased to 10-25% both at the TSS and

in gene bodies of all analyzed genes (Bact, Rab2b, Rpl30, Tuba4). These results

demonstrate that Mat1 is required for phosphorylation of RNAP II CTD and strongly

suggest that Cdk7 is the mammalian kinase responsible for this activity.

Global gene expression profiling of steady-state mRNAs revealed that although Mat1-/-

MEFs showed a small but significant general decrease in mRNA expression, the majority

of mRNAs remained unchanged. However, chromatin immunoprecipitation (ChIP)

experiments demonstrated that RNAP II levels were decreased in the bodies of all genes

analyzed independent of whether their steady-state levels were decreased, increased or

unchanged. This suggested a global defect in the progression of RNAP II into elongation,

which would not be reflected on the steady-state levels of most mRNAs. Defective

transcription was also supported by analysis of c-Fos mRNA levels after serum

stimulation as well as Hsp70 mRNA levels following heat shock: in both induction

models, Mat1-/- MEFs demonstrated deficient and delayed accumulation of the analyzed

mRNAs. Analysis of nascent transcripts using two alternative methods of in vivo RNA

labeling revealed a global defect in transcription following Mat1 deletion: a pulse-chase

of 1 h demonstrated a global decrease in labeled RNA as well as a decrease of all

45

analyzed mRNAs in Mat1-/- MEFs. A time-course labeling experiment analyzing three

genes with unchanged steady-state mRNAs (c-Myc, Pim1 and Gapdh) showed that the

labeled mRNA persisted for a much longer time in the Mat1-deleted cells, conveying

altered mRNA turnover and increased mRNA half-life following the transcriptional

attenuation in these cells.

In addition to the global defect in RNAP II progression into elongation, analysis of 5’end

capping and splicing of both U2- and U12-type introns demonstrated reduced levels of

both capped and spliced transcripts. These findings are in concert with those from other

models (Bird et al., 2004; Cho et al., 1997; Glover-Cutter et al., 2008; McCracken et al.,

1997a; McCracken et al., 1997b; Mortillaro et al., 1996; Moteki and Price, 2002;

Viladevall et al., 2009) and suggest that transcription and mRNA processing events in

mammalian cells would progress much in the same way as in yeast: TFIIH

phosphorylates Ser5 needed for the recruitement of capping enzymes, which in turn

recruit Cdk9, thus resulting in Ser2 phosphorylation and transcription elongation

(Guiguen et al., 2007; Pei et al., 2003; Qiu et al., 2009). The results demonstrate that

mammalian Mat1 is required for global transcription and efficient capping and suggest

that this requirement is mediated via phosphorylation of the Pol II CTD on Ser5.

Furthermore, the results reveal an unprecedented stabilization of mRNAs following

transcriptional attenuation.

46

2. The Cdk7 kinase submodule of TFIIH acts as a physiological roadblock to

adipogenesis through inhibitory phosphorylation of PPARγ (II)

As part of characterizing the functions of the kinase submodule of TFIIH, different

tissues were assessed for the protein levels of Cdk7 and Mat1. Unexpectedly, we

discovered that white adipose tissue exhibited very low levels of these subunits of TFIIH.

This was an unexpected finding, as Mat1 and Cdk7 had been postulated as ubiquitous

proteins. Since both subunits are expressed in undifferentiated fibroblasts, this suggested

that these proteins may be downregulated during the process of adipocyte differentiation.

Differentiation of 3T3-L1 fibroblasts into adipocytes demonstrated that indeed, both

Mat1 and Cdk7 protein levels decreased progressively during adipogenesis. To further

characterize the role of this kinase complex in adipogenesis, both a transient knockdown

approach (siRNA and shRNA targeting Cdk7) and a conditional genetic deletion model

(Mat1-/- MEFs) were used to assess the adipogenic role of Mat1 and Cdk7. Interestingly,

fibroblasts with a deficient kinase submodule demonstrated an increased capacity for

adipogenesis, increased activation of PPARγ as assessed by luciferase reporter assays and

induced transcription of endogenous PPARγ target genes. These phenotypes were rescued

by re-expression of Mat1, demonstrating that the intact kinase complex blocked PPARγ

activity.

As Cdk7 had been shown to phosphorylate PPARγ in vitro on Serine-112 (Ser112)

(Compe et al., 2005), and this phosphorylation had been demonstrated to inhibit the

activity of PPARγ (Hu et al., 1996), Ser112 phosphorylation was the most likely

hypothesis for the mode of action of the Cdk7 kinase. To test this model, Mat1-/- MEFs

and shCdk7 U2Os cells were transfected with PPARγ and stained for both total PPARγ as

well as the Ser112 phosphorylated form. Although the levels of total PPARγ were

identical to wild-type cells, both cell types with deficient Cdk7 kinase were unable to

generate the Ser112 phosphorylated form of PPARγ, confirming that intact Cdk7 kinase

activity is crucial for Ser112 phosphorylation of PPARγ. To directly assess the role of

Ser112 phosphorylation in the acquired responsiveness to PPARγ in Cdk7 knockdown

47

models, MEFs were transfected with a nonphosphorylatable PPARγ mutant, PPARγ-

Ser112A. This mutant did not result in a similar increase in luciferase reporter activity as

the wild-type PPARγ, demonstrating that the increased adipogenic potential noted in

Cdk7-kinase deficient cells is mediated via inhibitory phosphorylation of PPARγ on

Ser112. These results show that both Mat1 and Cdk7 proteins are decreased during

adipogenesis and undetectable in mature, fully differentiated adipocytes, and that this

kinase complex is required for the phosphorylation of PPARγ on Ser112. Thus they

support a model where the Cdk7 kinase complex maintains cells in an undifferentiated

state through the inhibitory Ser112 phopshorylation of PPARγ and suggest that in some

circumstances, for e.g. adipocyte differentiation, the downregulation of this complex is

required to surpass a roadblock to differentiation.

48

3. Mat1 is critical for cardiac energy metabolism in vivo via regulation of PGC1 (III)

Previous findings had demonstrated that the Cdk7/CycH/Mat1 complex becomes

activated in human heart failure and in mice with chronic cardiac hypertrophy (Sano et

al., 2002). In order to further characterize the potential role of this complex in the heart,

cardiac-specific Mat1 knockout mice were generated by crossing Mat1 flox mice

(Korsisaari et al., 2002) with mice expressing the Cre recombinase under cardiomyocyte-

specific α–myosin heavy chain (αMHC) promoter (Gaussin et al., 2002), (referred to in

text as CKO). The hearts of Mat1 CKO mice developed normally and did not show

defects in phosphorylation of Cdk2, Cdc2 or RNAP II despite reduced levels of Mat1,

Cdk7 and CycH.

Within 5-6 weeks, Mat1 CKO mice began to show signs of cardiomyopathy as evidenced

by increased apoptosis of cardiomyocytes, systemic edema, ventricular dilation and

fibrosis. All Mat1-/- mice died within 8 weeks as a result of cardiac failure. The fact that

Mat1-deficient hearts developed normally for the first month suggested that the lethal

cardiac phenotype was due to specific transcriptional changes. To investigate this, global

gene expression profiling (Affymetrix) was performed from hearts of 4 week old mice.

mRNAs involved in mitochondrial fatty acid β-oxidation, the tricarboxylic acid cycle and

the respiratory chain were found to be downregulated in Mat1 CKO hearts compared to

control littermates. Many of these mRNAs are regulated by the transcriptional co-

activator PGC-1α, which suggested that Mat1 deletion may be required for PGC-1

mediated transcriptional activation. Indeed, PGC-1α failed to activate two well-

characterized target genes Atp5b and Cycs in Mat1-/- MEFs and deletion of Mat1

suppressed the co-activation of PPARα and ERRα by PGC-1 in MEFs. Finally, a direct

interaction between PGC-1α and both Mat1 and Cdk7 was detected, suggesting that the

Cdk7/CycH/Mat1 complex may be involved in recruiting PGC-1 to target gene

promoters, and hence be required for transcription of PGC-1 downstreatm targets.

49

4. Hepatic-specific deletion of Mat1 in mice results in fatty liver through induction

of SREBP-1c target genes via loss of DMAP1 and Dnmt1 (IV)

This study was undertaken to study the role of the Cdk7 kinase complex in hepatic lipid

metabolism. Liver-specific conditional Mat1 knockout mice were generated by breeding

Mat1 flox mice (Korsisaari et al., 2002) with interferon-response sensitive Mx1-Cre

deletor mice (Kuhn, R. et al, 1995) which can be strongly induced in the liver following

polyinosinic-polycytidylic acid (pI-pC) injections. Induction of Mx1-Cre was achieved

using 3 consecutive injections of (pI-pC) every third day, and depletion of Mat1 protein

levels was observed at day 14 from the first injection. At this timepoint, livers of Mx1-

Cre;Mat1-/- mice displayed significant accumulation of triglycerides characteristic of

hepatosteatosis. As Mat1 deletion in MEFs results in adipogenesis through activation of

the PPARγ pathway (II), PPARγ as well as PPARα (the major PPAR isoform in liver),

target gene expression was analyzed using qRT-PCR but the results did not indicate

detectable alterations. Furthermore, levels of mRNAs that were identified to be

dysregulated at the steady-state level following Mat1 deletion in MEFs were also

quantitated but no significant differences were found, demonstrating that in different cell

types, Mat1 regulates the steady-state levels of different mRNAs.

SREBP-1c is the key transcription factor in regulating hepatic lipid metabolism

(Shimomura et al., 1998) and as been shown to be involved in the development of fatty

liver in response to insulin (reviewed in Foufelle and Ferre, 2002). Therefore, the levels

of SREBP-1c target gene mRNAs were quantitated using qRT-PCR, and found to be

paradoxically upregulated in livers of Mx1-Cre;Mat1-/- mice compared to healthy

littermates. This indicated that deletion of Mat1 in mouse hepatocytes results in

triglyceride synthesis and accumulation due to induction of the lipogenic SREBP-1c

downstream genes. A similar response was noted in cultured HepG2 hepatocarcinoma

cells, where acute depletion of Mat1 using siRNA knockdown led to the increase in the

levels of SREBP-1c target gene mRNAs. This induction requires SREBP-1, as following

SREBP-11 knockdown no increase was noted. However, the induction of SREBP-1c

target mRNAs was apparently not due to increased levels of SREBP1 based on

50

unchanged mRNA levels, suggesting an alternative transcriptional regulatory mechanism

involving Mat1 and yet specific for SREBP-1c downstream geens.

The finding that Mat1 deletion resulted in upregulation of specific mRNAs was

surprising, since the majority of Mat1 functions have pointed to an activating role in

transcription. However, Mat1 has been shown to bind to DMAP1 (Kang et al., 2007), a

transcriptional co-repressor, which acts partially by recruiting DNA methyltransferase 1

(Dnmt1) to DNA (Lee et al., 2010). To investigate whether this mechanism might

underlie the increased SREBP-1c target mRNAs following Mat1 deletion, DMAP1 levels

were downregulated using siRNA, and Dnmt1 activity inhibited with 5-Azacytidine in

HepG2 cells. Remarkably, both of these treatments resulted in specific upregulation of

SREBP-1c target gene mRNAs. Furthermore, double knockdowns of Mat1 and DMAP1

together or in combination with Dnmt1 inhibition did not have additive effects on the

increased levels of SREBP-1c target gene mRNAs. This data indicated that Mat1,

DMAP1 and Dnmt1 collaborate to maintain appropriate levels of SREBP-1c lipogenic

mRNAs in healthy hepatocytes by suppressing transcription through the DNA

methyltransferase activity of Dnmt1.

To investigate whether DMAP1 and Dnmt1 occupy the promoters of repressed SREBP-

1c target genes and whether their presence is dependent on Mat1, chromatin

immunoprecipitations (ChIP) with Mat1, DMAP1 and Dnmt1 were performed from

livers of Mx1-Cre;Mat1-/- mice along with control littermates. The ChIP experiments

demonstrated that the promoters of SREBP-1c target genes ME1, Scd1 and Gapt are

indeed co-occupied by DMAP1 and Dnmt1 in wild-type livers, consistant with a

repressive role on these SREBP-1c target genes. Furthermore, Mat1 deletion in livers

resulted in decreased occupancy of both DMAP1 and Dnmt1 on all the analyzed SREBP-

1c target gene promoters. Mat1 deletion also resulted in increased occupancy of SREBP-

1 on these genes. These data suggest that the induction of SREBP-1c dependent mRNAs

following Mat1 depletion in hepatocytes is due to loss of co-repressor DMAP1 and

Dnmt1 from and recruitment of SREBP-1c to promoters of ME1, Gpat1 and Scd1.

51

These findings reveal a novel transcriptional repressor function for mammalian Mat1 in

the regulation of hepatic lipid metabolism through SREBP-1c in vivo. The results

demonstrate that Mat1 is required for the maintenance of appropriate levels of lipogenic

SREBP-1c target genes and promoter occupancy of both DMAP1 and Dnmt1 on SREBP-

1c target genes in hepatocytes, indicating that in a healthy liver SREBP-1c target genes

are repressed through the recruitment of co-repressor DMAP1 and the DNA

methyltransferase Dnmt1 by Mat1. The discovery that Mat1 and Cdk7 are significantly

decreased both at mRNA and protein level in fatty livers of diabetic db/db mice

implicates that downregulation of the Cdk7 kinase complex may play a role in the

development of obesity-related hepatosteatosis.

52

DISCUSSION AND PERSPECTIVES The use of Cre-mediated Mat1 deletion in cell culture and tissue-specific mouse models

used in this study has for the first time clearly established a general role for Mat1 and the

TFIIH kinase in RNAP II CTD Ser5 phosphorylation and transcription in mammalian

cells. Furthermore, the approach has revealed novel gene- and tissue-specific functions

for Mat1 and the TFIIH kinase both as an activator (PGC-1 mediated transcriptional

activation in cardiomyocytes) and as a repressor (PPARγ inhibitory phosphorylation in

preadipocytes; recruitment of DMAP1 and Dnmt1 to SREBP-1c target genes in

hepatocytes). The results attained during this thesis work provide important novel

discoveries in mechanisms of basic science but also offer potential new therapeutic

windows for diseases such as type II diabetes, hepatosteatosis and the metabolic

syndrome.

Mat1, Cdk7 and RNA Polymerase II CTD phosphorylation

The data presented here provides compelling evidence that in mouse embryonic

fibroblasts, Cdk7 is the main Ser5 RNAP II CTD kinase (I). Although it is formally

possible that the depletory effect on Ser5 following Mat1 deletion in MEFs would be

mediated independently of Cdk7, this appears unlikely as in all model systems used in

this thesis as well as in other studies, Cdk7 levels and kinase activity decrease following

Mat1 ablation. Furthermore, no Cdk7-independent functions of Mat1 have been

described. As TFIIH has been mainly associated with PIC formation at the promoter

region and not associated further along genes (Cheng and Sharp, 2003; Donner et al.,

2010; Gomes et al., 2006; Morris et al., 2005), the Ser5 phosphorylation of RNAP II

during elongation has been attributed to some other CTD kinase. However, our results

show that Ser5 phosphorylation decreases also in gene bodies (I). This suggests that

either a) Cdk7 is in fact the kinase responsible for the majority of all RNAP II CTD Ser5

phosphorylation events independent of where RNAP II is located on a gene b) Mat1

additionally regulates some other downstream Ser5 kinase or c) TSS Ser5

53

phosphorylation is necessary for the action or recruitment of a second, downstream Ser5

kinase. Mat1-/- MEFs also demonstrated globally reduced Ser2 phosphorylation (I). As

Cdk7 has not been shown to phosphorylate Ser2, the observed decrease in Ser2

phosphorylation is most probably due to the lack of recruitment of a Ser2 kinase, for e.g.

Cdk9 (Guiguen et al., 2007; Viladevall et al., 2009), as a result of decreased Ser5-P

levels.

In tissues, the role of Mat1 and Cdk7 in RNAP II CTD phosphorylation appears to be

more complicated. Deletion of Mat1 in myocardium did not reveal defects in global Ser5

levels (III), although the possibility –and probability- of reduced levels of the Ser5-P

form of RNAP II on specific genes exists. On the other hand, different genes may have a

differential requirement for the amount of Ser5 phosphorylated RNAP II. This is

supported by the ChIP analysis of Ser5-P levels from Mat1-deficient livers: although

SREBP-1c target genes have 30-50% less Ser5 phosphorylated RNAP II in Mat1- deleted

livers compared to heterozygous littermates, the mRNA levels of these genes are

increased. Another interpretation of the results is that some tissues may contain another

(or other) RNAP II Ser5 kinase(s). Alternatively, although Cdk7 may be the major CTD

Ser5 kinase in all mammalian cells under unperturbed circumstances, certain tissues may

be able to re-direct the substrate specificity of another kinase towards Ser5 when Cdk7

activity is depleted. For example, the HIV Tat protein is able to re-target Cdk9 to

phosphorylate Ser5 (Zhou et al., 2000). In support of this, the timescale between the cell

culture and in vivo mouse studies must be considered: in the MEF model, immediate

effects of Mat1 deletion are investigated, whereas the Mat1-deficient tissues are analyzed

weeks and months later, giving the cells time to possibly compensate for the lack of Cdk7

kinase activity. The conditional Mat1 mouse models generated during these studies

would therefore provide an excellent tool for the discovery of novel RNAP II CTD Ser5

kinases.

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Tissue-specific roles of Mat1

Three of the studies presented here highlight a role for Mat1, and in the case of

adipogenesis also for Cdk7, in metabolism (II-IV). PGC-1α has been shown to act as a

co-activator of PPARα and PPARγ (Puigserver et al., 1998; Vega et al., 2000), and

SREBP-1c as an upstream regulator of PPARγ (Fajas et al., 1999; Kim et al., 1998).

Therefore, one might contemplate the possibility that as these pathways are

interconnected, all the observed tissue-specific defects could be in fact mediated by the

same mechanism. This remains highly unlikely based on tissue-specific expression

patterns of PPARs, PGC-1 and SREBP-1c. Furthermore, our data demonstrated that Mat1

deletion in MEFs resulted in differentiation towards white adipocytes (II), whereas PGC-

1α is implicated in the brown adipocyte phenotype (Puigserver et al., 1998). In the Mx1-

Cre;Mat1flox mouse model, the primary culprit of triglyceride accumulation was PPARγ

based on previous in vitro findings (Compe et al., 2005) and our own earlier data from

MEFs, but neither PPARγ nor PPARα demonstrated defective activation (IV).

Furthermore, the mode of action in each case was demonstrated to be different. In

adipocyte differentiatation, the Cdk7 subunit phosphorylates and inhibits PPARγ thereby

indirectly influencing transcription of downstream targets, whereas in hepatocytes, the

mode of transcriptional repression occurs directly on promoters of SREBP-1c target

genes via recruitment of the co-repressor DMAP1 and the DNA methyltransferase

Dnmt1.

The unexpectedly minor role of the mammalian TFIIH kinase in global transcription in

the analyzed tissues and differentiating cells is interesting in regard of described

physiological settings with active mRNA transcription regardless of reduced TFIIH

kinase activity or Ser5 phosphorylation. The cell cycle in budding yeast is one such case:

quiescent cells lack detectable levels of Ser5-P and Ser2-P, yet 460 stationary-phase

specific genes are actively transcribed (Radonjic et al., 2005). Cdk7 kinase activity and

Ser5-P are repressed also during mitosis in mammalian cells, yet at least β-actin and

cyclin B1 mRNAs are actively synthesized (Long et al., 1998; Sciortino et al., 2001; Xu

and Manley, 2007). The work in this thesis demonstrates that mouse adipose tissue lacks

55

detectable levels of Mat1 and Cdk7 (II); nevertheless, adipocytes are transcriptionally

active, demonstrating that TFIIH kinase-independent transcription occurs in certain

tissues in vivo. Both Mat1 and Cdk7 are downregulated during adipocyte differentiation

to permit activation of PPARγ (ΙΙ), suggesting a model where in certain circumstances

the general TFIIH kinase elongation function identified in MEFs can be compromised

when inhibition of TFIIH kinase is beneficial or critical for other functions (Figure 7).

Furthermore, in the liver, Mat1 has a repressive modulatory function in transcription,

which appears to be crucial for the maintenance of appropriate levels of SREBP1 target

mRNAs (IV).

mRNA stabilization: linking transcription progression to mRNA turnover

Perhaps the most novel and intriguing discovery of this thesis work was the finding that

the perturbation of transcription by Mat1 deletion in MEFs resulted in prolonged mRNA

half-lives (I). A similar phenomenon has been previously observed in budding yeast

following oxidative stress (Molina-Navarro et al., 2008) and in mammalian cells treated

with transcriptional inhibitors (Meininghaus et al., 2000), but this has not been widely

acknowledged.

How might mRNAs be stabilized at a global level? An obvious mechanism that emerges

is the retention of mRNAs in the nucleus by blocking nuclear export. However, if this

were the case, Mat1-/- MEFs would have shown decreased protein levels (unless a

simultaneous stabilization of proteins occurred), which were not observed. A second

possible mechanism is via capping-dependent regulation of translation The majority of

mRNAs are translated in a cap-dependent manner (Van Der Kelen et al., 2009). As the

Mat1-/- MEFs demonstrated defective capping of most analyzed mRNAs, this could result

in slower translation which would naturally increase the half-lives of the mRNAs. If the

translation rate would not form a bottle-neck, such a defect would not be detected in total

protein levels, similar to DRB-treated HeLa cells, where inhibition of Cdk7 activity slows

down capping without having an effect on the amount of capped end product (Moteki and

Price, 2002). A third possibility worth considering is the binding of stabilizing, protective

56

proteins to mRNAs. These proteins would have to be general rather then the sequence-

specific ARE-binding proteins. For example, poly(A) binding proteins (PABPs) prevent

deadenylation of mRNAs by binding to the poly(A) tail and hence protect the mRNA

from degradation (Bernstein et al., 1989). PABPs have been shown to interact with the

5’cap structure, thereby providing a possible communication link between transcription

and translation.

This discovery of altered mRNA turnover raises the question whether this phenomenon

takes place in vivo. For example, might the cardiomyopathy observed in αMHC-

Mx1;Mat1-/- mice simply reflect a timing issue: Mat1-deficient cardiomyocytes are able

to maintain protein levels by stabilizing mRNAs but only for a certain period of time,

eventually resulting in lethality. What does the nascent mRNA profile of white

adipocytes look like: do they exhibit altered mRNA turnover compared to

undifferentiated cells? All in all, these findings warrant the necessity of utilizing nascent

mRNAs rather then mRNA steady-state levels for reliable conclusions concerning

transcriptional activity in any given system.

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Figure 7. Summary of the roles of mammalin Mat1 in transcriptional activation and repression. (a) In undifferentiated and proliferating cells, such as preadipocytes and mouse embryonic fibroblasts (MEFs), Mat1 is required for the stability of the Cdk7 kinase module to TFIIH and for phosphorylation of Ser5 of RNAP II CTD (1). Ser5 phosphorylation is crucial for the recruitment of the cap complex (2), which in turn is required for recruitment of P-TEFb to phosphorylate RNAP II CTD on Ser2 residues (3). These consecutive steps are required for efficient transcription elongation to produce the nascent mRNA reflected in the levels of total mRNA. Furthermore, thep preadipocyte state is maintained through the inhibitory phosphorylation of Ser112 of PPARγ by Mat1/Cdk7 (4). (b) Upon Mat1 deletion, Cdk7 is unable to bind to the rest of TFIIH, Ser5 phosphorylation of RNAP II CTD does not occur and the cap complex and P-TEFb are not recruited to the site of transcription (1). This results in transcriptional attenuation and decreased nascent mRNA amounts. This defect is not reflected in the majority of total mRNA, suggesting the existence of a global mRNA stabilizing mechanism (2). Due to loss of Cdk7 activity, PPARγ is no longer maintained in a phosphorylated state, and in the presence of appropriate hormonal cues, differentiation into adipocytes can take place (3). (c) In some terminally differentiated tissues (e.g. WAT) and during the differentiation process as well as pathophysiologies like hepatosteatosis, Mat1 and Cdk7 protein levels are downregulated (1). This leads to activation of specific transcriptional pathways, depending on tissue and cell type (2), suggesting that the general transcriptional function demonstrated in a-b can be compromised in circumstances where downregulation of Mat1/Cdk7 is required. Whether specific tissues and differentiation processes require an alternate RNAP II CTD Ser5 kinase (3), display differences in recruitment of cap complex, P-TEfB and synthesis of nascent mRNA (4) and possible stabilization of mRNA remains to be studied.

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ACKNOWLEDGEMENTS This work was carried out in the Institute of Biotechnology, Viikki and Genome-Scale Biology Program (Biomedicum Helsinki), University of Helsinki during 2004-2011 under the supervision of Professor Tomi Mäkelä. I thank the staff of Biomedicum Helsinki and the Institute of Biotechnology for the pleasant working environments and excellent research facilites, with particual thanks to Olli Jänne, Tomi Mäkelä, Maart Saarma and Esa Korpi. First and foremost, I would like to express my gratitude to my thesis supervisor Tomi Mäkelä. You have provided me with a lab with an excellent social and productive atmosphere whose close-knit members are always keen on both science and fun. Your enthusiasm towards science is inspiring and despite your busy schedule, you always found time to discuss experiments/data/manuscripts with me. I appreciate you letting me in many cases go my own way and make my own mistakes – after all, that is the best way to learn. Thank you Tomi. I am extremely grateful to Mikko Frilander and Olli Jänne for acting as my Thesis Committee, without whom this thesis may have taken another 5 years. Thank you for your valuable insights on the projects throughout the PhD, which have in many cases changed the course of experiments. I express my gratitude to Olli Jänne (again) and Lea Sistonen for reviewing my thesis on a tight schedule and providing me with numerous valuable comments that further strengthened the thesis. I would additionally like to thank Kari Alitalo for scientific advice and support. The Helsinki Biomedical Graduate School is sincerely acknowledged for the provided opportunities as well as financial support. I thank all my co-authors in and out of Finland without whom this work would not have been possible. Particular thanks go to Heli Pessa and Mikko Frilander, who provided novel insight and analyses to my research. Motoaki Sano and Michael D. Schneider are acknowledged for the fruitful collaboration. I want to express my warmest gratitude to the entire Mäkelä lab, both past and present members – it has truly been a pleasure! In particular I want to thank my coauthors: Ying Yang for fruitful scientific discussions and the cleanest pipetting hands I have witnessed, as well as Timofei Tselykh for patiently providing the data which significantly impacted our results. I want to thank Saana Ruusulampi and Birgitta Tjäder for taking care of basically everything in the lab and knowing where everything is – you both managed the lab in your own exceptional ways. Outi Kokkonen, Kirsi Mänttäri and Susanna Räsänen – thank you for excellent help with basically anything I could think to ask. Special thanks go to Jenny Bärlund and Saana Ruusulampi (again) for superb hands-on help with experiments, without which I surely wouldn’t have been able to meet my deadlines. In addition to the long work hours spent together, I want to thank all the Makelab members for an absolutely brilliant time doing things not involving science. Special thanks to Bianca, Kaisa, Tea, Kari, Pekka, Thomas, Saana, Lina and Outi for friendship. In particular I want to thank Emilia Kuuluvainen, who has been terrific company in the office, as my benchmate, at transcription meetings, at Faculty club, at the numerous parties – and afterparties. I also want to acknowledge the past and present members of Klefalab (in particular Johanna Partanen, Anni Nieminen, Annika Hau) and Ojalalab (in particular Annika Järviluoma, Grzegorz Sarek and Sonja Koopal) for so many countless occasions of fun – both science related and not so much…

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Additionally, I have received technical and administrative help outside the lab, with special thanks to Satu Sankkila, Ritva Lautala, Aija Kaitera, Tapio Tainola and Jukka Rahkonen. Thank you also to Carl Kyrklund for valuable help with Swedish grant applications. I wish to extend my thanks to the excellent party crowd of HBGS/Biomedicum: Katja P, Joao D, Mira H, Sanna V, Maurice J, Can H, Tea B, Alex G, Alex T, Roxana O, Iulia D, Rahkonen and of course all the members of HBGS student council – it was a blast! Cheers! Outside of Biomedicum, I owe a huge thanks to the people who kept me half-sane throughout all this. I cannot even express how thankful I am to all my many wonderful friends, especially during this last year, which would have been hell without you guys! Katja Kurhela, Jenny Miettinen, Niina Puustinen, Paula Kemppainen, Paula Koivumäki, Tuuli Lappalainen, Martta Viitaniemi, Laura Kalliokoski, Essi Lind, Emilia Gaal, Sanna Vuoristo, Marika Kontio, Eeva Niini, Sini Koivisto, Laura Helenius, Laura Hirvensalo – whether it was girlie movie nights, wine tastings, bird watching excursions, pole evenings or trips to Ibiza, you kept me going throughout all this. My very special thanks go to Johanna Partanen for always being there for me, for always understanding me. You kick ass!! I wish to express my gratitude to my parents, Tarja and Matti Helenius. Although I guess you never really understood why I chose to do this as a career, you have always let me make my own choices and supported me in whatever road I took. In the end, I am sure none of this would have been possible without the opportunities you provided me when growing up. Dearest Laura and Janne, you are the best possible sis and bro anyone could have. Thanks for all your support and putting up with me! Tuomas, my dear husband: thank you for endlessly supporting me, thank you for understanding me, thank you for being patient with me, thank you for making me smile (occasionally laugh as well), thank you for listening to me, thank you for taking care of me, thank you for believing in me (especially when I didn’t), thank you for being you. I love you – always, forever. I have been financially supported by the Helsinki Biomedical Graduace School, the Biomedicum Helsinki Foundation, the Paulo Foundation, the Finnish Cultural Foundation, the Orion Farmos Research Foundation, the Emil Aaltonen Foundation, the Diabetes Research Foundation, and Nylands Nation. All are sincerely acknowledged.

Helsinki, September 9th, 2011

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REFERENCES Adamczewski, J.P., Rossignol, M., Tassan, J.P., Nigg, E.A., Moncollin, V., and Egly, J.M. (1996). MAT1, cdk7 and cyclin H form a kinase complex which is UV light- sensitive upon association with TFIIH. The EMBO journal 15, 1877-1884. Adams, M., Reginato, M.J., Shao, D., Lazar, M.A., and Chatterjee, V.K. (1997). Transcriptional activation by peroxisome proliferator-activated receptor gamma is inhibited by phosphorylation at a consensus mitogen-activated protein kinase site. The Journal of biological chemistry 272, 5128-5132. Ahn, S.H., Kim, M., and Buratowski, S. (2004). Phosphorylation of serine 2 within the RNA polymerase II C-terminal domain couples transcription and 3' end processing. Molecular cell 13, 67-76. Akhtar, M.S., Heidemann, M., Tietjen, J.R., Zhang, D.W., Chapman, R.D., Eick, D., and Ansari, A.Z. (2009). TFIIH kinase places bivalent marks on the carboxy-terminal domain of RNA polymerase II. Molecular cell 34, 387-393. Albert, I., Mavrich, T.N., Tomsho, L.P., Qi, J., Zanton, S.J., Schuster, S.C., and Pugh, B.F. (2007). Translational and rotational settings of H2A.Z nucleosomes across the Saccharomyces cerevisiae genome. Nature 446, 572-576. Arany, Z., Foo, S.Y., Ma, Y., Ruas, J.L., Bommi-Reddy, A., Girnun, G., Cooper, M., Laznik, D., Chinsomboon, J., Rangwala, S.M., et al. (2008). HIF-independent regulation of VEGF and angiogenesis by the transcriptional coactivator PGC-1alpha. Nature 451, 1008-1012. Arany, Z., He, H., Lin, J., Hoyer, K., Handschin, C., Toka, O., Ahmad, F., Matsui, T., Chin, S., Wu, P.H., et al. (2005). Transcriptional coactivator PGC-1 alpha controls the energy state and contractile function of cardiac muscle. Cell Metab 1, 259-271. Arany, Z., Novikov, M., Chin, S., Ma, Y., Rosenzweig, A., and Spiegelman, B.M. (2006). Transverse aortic constriction leads to accelerated heart failure in mice lacking PPAR-gamma coactivator 1alpha. Proc Natl Acad Sci U S A103, 10086-10091. Baar, K., Wende, A.R., Jones, T.E., Marison, M., Nolte, L.A., Chen, M., Kelly, D.P., and Holloszy, J.O. (2002). Adaptations of skeletal muscle to exercise: rapid increase in the transcriptional coactivator PGC-1. Faseb J 16, 1879-1886. Bakheet, T., Frevel, M., Williams, B.R., Greer, W., and Khabar, K.S. (2001). ARED: human AU-rich element-containing mRNA database reveals an unexpectedly diverse functional repertoire of encoded proteins. Nucleic acids research 29, 246-254.

61

Bartolomei, M.S., Halden, N.F., Cullen, C.R., and Corden, J.L. (1988). Genetic analysis of the repetitive carboxyl-terminal domain of the largest subunit of mouse RNA polymerase II. Molecular and cellular biology 8, 330-339. Baskaran, R., Dahmus, M.E., and Wang, J.Y. (1993). Tyrosine phosphorylation of mammalian RNA polymerase II carboxyl-terminal domain [see comments]. Proc Natl Acad Sci U S A90, 11167-11171. Bastien, J., Adam-Stitah, S., Riedl, T., Egly, J.M., Chambon, P., and Rochette-Egly, C. (2000). TFIIH interacts with the retinoic acid receptor gamma and phosphorylates its AF-1-activating domain through cdk7. The Journal of biological chemistry 275, 21896-21904. Baumann, M., Pontiller, J., and Ernst, W. (2010) Structure and basal transcription complex of RNA polymerase II core promoters in the mammalian genome: an overview. Molecular biotechnology 45, 241-247. Belotserkovskaya, R., Oh, S., Bondarenko, V.A., Orphanides, G., Studitsky, V.M., and Reinberg, D. (2003). FACT facilitates transcription-dependent nucleosome alteration. Science 301, 1090-1093. Belz, T., Pham, A.D., Beisel, C., Anders, N., Bogin, J., Kwozynski, S., and Sauer, F. (2002). In vitro assays to study protein ubiquitination in transcription. Methods 26, 233-244. Bernstein, B.E., Kamal, M., Lindblad-Toh, K., Bekiranov, S., Bailey, D.K., Huebert, D.J., McMahon, S., Karlsson, E.K., Kulbokas, E.J., 3rd, Gingeras, T.R., et al. (2005). Genomic maps and comparative analysis of histone modifications in human and mouse. Cell 120, 169-181. Bernstein, B.E., Liu, C.L., Humphrey, E.L., Perlstein, E.O., and Schreiber, S.L. (2004). Global nucleosome occupancy in yeast. Genome biology 5, R62. Bernstein, P., Peltz, S.W., and Ross, J. (1989). The poly(A)-poly(A)-binding protein complex is a major determinant of mRNA stability in vitro. Molecular and cellular biology 9, 659-670. Bird, A., Taggart, M., Frommer, M., Miller, O.J., and Macleod, D. (1985). A fraction of the mouse genome that is derived from islands of nonmethylated, CpG-rich DNA. Cell 40, 91-99. Bird, G., Zorio, D.A., and Bentley, D.L. (2004). RNA polymerase II carboxy-terminal domain phosphorylation is required for cotranscriptional pre-mRNA splicing and 3'-end formation. Molecular and cellular biology 24, 8963-8969.

62

Boehm, A.K., Saunders, A., Werner, J., and Lis, J.T. (2003). Transcription factor and polymerase recruitment, modification, and movement on dhsp70 in vivo in the minutes following heat shock. Molecular and cellular biology 23, 7628-7637. Braissant, O., Foufelle, F., Scotto, C., Dauca, M., and Wahli, W. (1996). Differential expression of peroxisome proliferator-activated receptors (PPARs): tissue distribution of PPAR-alpha, -beta, and -gamma in the adult rat. Endocrinology 137, 354-366. Browning, J.D., and Horton, J.D. (2004). Molecular mediators of hepatic steatosis and liver injury. J Clin Invest 114, 147-152. Busso, D., Keriel, A., Sandrock, B., Poterszman, A., Gileadi, O., and Egly, J.M. (2000). Distinct regions of MAT1 regulate cdk7 kinase and TFIIH transcription activities. The Journal of biological chemistry 275, 22815-22823. Camp, H.S., and Tafuri, S.R. (1997). Regulation of peroxisome proliferator-activated receptor gamma activity by mitogen-activated protein kinase. The Journal of biological chemistry 272, 10811-10816. Carninci, P., Kasukawa, T., Katayama, S., Gough, J., Frith, M.C., Maeda, N., Oyama, R., Ravasi, T., Lenhard, B., Wells, C., et al. (2005). The transcriptional landscape of the mammalian genome. Science 309, 1559-1563. Carninci, P., Sandelin, A., Lenhard, B., Katayama, S., Shimokawa, K., Ponjavic, J., Semple, C.A., Taylor, M.S., Engstrom, P.G., Frith, M.C., et al. (2006). Genome-wide analysis of mammalian promoter architecture and evolution. Nature genetics 38, 626-635. Chandra, V., Huang, P., Hamuro, Y., Raghuram, S., Wang, Y., Burris, T.P., and Rastinejad, F. (2008). Structure of the intact PPAR-gamma-RXR-alpha nuclear receptor complex on DNA. Nature, 350-356. Chang, Y.F., Imam, J.S., and Wilkinson, M.F. (2007). The nonsense-mediated decay RNA surveillance pathway. Annu Rev Biochem 76, 51-74. Chapman, R.D., Heidemann, M., Albert, T.K., Mailhammer, R., Flatley, A., Meisterernst, M., Kremmer, E., and Eick, D. (2007). Transcribing RNA polymerase II is phosphorylated at CTD residue serine-7. Science 318, 1780-1782. Chen, D., Riedl, T., Washbrook, E., Pace, P.E., Coombes, R.C., Egly, J.M., and Ali, S. (2000). Activation of estrogen receptor alpha by S118 phosphorylation involves a ligand-dependent interaction with TFIIH and participation of CDK7. Molecular cell 6, 127-137. Chen, Z., Torrens, J.I., Anand, A., Spiegelman, B.M., and Friedman, J.M. (2005). Krox20 stimulates adipogenesis via C/EBPbeta-dependent and -independent mechanisms. Cell Metab 1, 93-106.

63

Cheng, C., and Sharp, P.A. (2003). RNA polymerase II accumulation in the promoter-proximal region of the dihydrofolate reductase and gamma-actin genes. Molecular and cellular biology 23, 1961-1967. Cho, E.J., Takagi, T., Moore, C.R., and Buratowski, S. (1997). mRNA capping enzyme is recruited to the transcription complex by phosphorylation of the RNA polymerase II carboxy-terminal domain. Genes & development 11, 3319-3326. Cock, T.A., Houten, S.M., and Auwerx, J. (2004). Peroxisome proliferator-activated receptor-gamma: too much of a good thing causes harm. EMBO reports 5, 142-147. Coin, F., Bergmann, E., Tremeau-Bravard, A., and Egly, J.M. (1999). Mutations in XPB and XPD helicases found in xeroderma pigmentosum patients impair the transcription function of TFIIH. The EMBO journal 18, 1357-1366. Coin, F., Oksenych, V., and Egly, J.M. (2007). Distinct roles for the XPB/p52 and XPD/p44 subcomplexes of TFIIH in damaged DNA opening during nucleotide excision repair. Molecular cell 26, 245-256. Coin, F., Oksenych, V., Mocquet, V., Groh, S., Blattner, C., and Egly, J.M. (2008). Nucleotide excision repair driven by the dissociation of CAK from TFIIH. Molecular cell 31, 9-20. Compe, E., Drane, P., Laurent, C., Diderich, K., Braun, C., Hoeijmakers, J.H., and Egly, J.M. (2005). Dysregulation of the peroxisome proliferator-activated receptor target genes by XPD mutations. Molecular and cellular biology 25, 6065-6076. Davie, J.R., Lin, R., and Allis, C.D. (1991). Timing of the appearance of ubiquitinated histones in developing new macronuclei of Tetrahymena thermophila. Biochemistry and cell biology = Biochimie et biologie cellulaire 69, 66-71. de Murcia, G., Huletsky, A., and Poirier, G.G. (1988). Modulation of chromatin structure by poly(ADP-ribosyl)ation. Biochemistry and cell biology = Biochimie et biologie cellulaire 66, 626-635. Devault, A., Martinez, A.M., Fesquet, D., Labbe, J.C., Morin, N., Tassan, J.P., Nigg, E.A., Cavadore, J.C., and Doree, M. (1995). MAT1 ('menage a trois') a new RING finger protein subunit stabilizing cyclin H-cdk7 complexes in starfish and Xenopus CAK. The EMBO journal 14, 5027-5036. Dhe-Paganon, S., Syeda, F., and Park, L. (2011). DNA methyltransferase 1: regulatory mechanisms and implications in health and disease. Int J Biochem Mol Biol 2, 58-66. Dolken, L., Ruzsics, Z., Radle, B., Friedel, C.C., Zimmer, R., Mages, J., Hoffmann, R., Dickinson, P., Forster, T., Ghazal, P., et al. (2008). High-resolution gene expression

64

profiling for simultaneous kinetic parameter analysis of RNA synthesis and decay. RNA (New York, NY 14, 1959-1972. Donner, A.J., Ebmeier, C.C., Taatjes, D.J., and Espinosa, J.M. (2010). CDK8 is a positive regulator of transcriptional elongation within the serum response network. Nature structural & molecular biology 17, 194-201. Drapkin, R., Reardon, J.T., Ansari, A., Huang, J.C., Zawel, L., Ahn, K., Sancar, A., and Reinberg, D. (1994). Dual role of TFIIH in DNA excision repair and in transcription by RNA polymerase II. Nature 368, 769-772. Ebert, A., Lein, S., Schotta, G., and Reuter, G. (2006). Histone modification and the control of heterochromatic gene silencing in Drosophila. Chromosome Res 14, 377-392. Eckhardt, F., Lewin, J., Cortese, R., Rakyan, V.K., Attwood, J., Burger, M., Burton, J., Cox, T.V., Davies, R., Down, T.A., et al. (2006). DNA methylation profiling of human chromosomes 6, 20 and 22. Nature genetics 38, 1378-1385. Egloff, S., and Murphy, S. (2008). Cracking the RNA polymerase II CTD code. Trends Genet 24, 280-288. Egloff, S., O'Reilly, D., Chapman, R.D., Taylor, A., Tanzhaus, K., Pitts, L., Eick, D., and Murphy, S. (2007). Serine-7 of the RNA polymerase II CTD is specifically required for snRNA gene expression. Science 318, 1777-1779. Fajas, L., Landsberg, R.L., Huss-Garcia, Y., Sardet, C., Lees, J.A., and Auwerx, J. (2002). E2Fs regulate adipocyte differentiation. Developmental cell 3, 39-49. Fajas, L., Schoonjans, K., Gelman, L., Kim, J.B., Najib, J., Martin, G., Fruchart, J.C., Briggs, M., Spiegelman, B.M., and Auwerx, J. (1999). Regulation of peroxisome proliferator-activated receptor gamma expression by adipocyte differentiation and determination factor 1/sterol regulatory element binding protein 1: implications for adipocyte differentiation and metabolism. Molecular and cellular biology 19, 5495-5503. Faye, G., Simon, M., Valay, J.G., Fesquet, D., and Facca, C. (1997). Rig2, a RING finger protein that interacts with the Kin28/Ccl1 CTD kinase in yeast. Mol Gen Genet 255, 460-466. Fong, N., and Bentley, D.L. (2001). Capping, splicing, and 3' processing are independently stimulated by RNA polymerase II: different functions for different segments of the CTD. Genes & development 15, 1783-1795. Foufelle, F., Ferre, P. (2002). New perspectives in the regulation of hepatic glycolytic and lipogenic genes by insulin and glucose: a role for the transcription factor sterol regulatory element binding protein-1c. Biochem J 366, 377-91.

65

Fribourg, S., Romier, C., Werten, S., Gangloff, Y.G., Poterszman, A., and Moras, D. (2001). Dissecting the interaction network of multiprotein complexes by pairwise coexpression of subunits in E. coli. Journal of molecular biology 306, 363-373. Friedel, C.C., Dolken, L., Ruzsics, Z., Koszinowski, U.H., and Zimmer, R. (2009). Conserved principles of mammalian transcriptional regulation revealed by RNA half-life. Nucleic acids research 37, e115. Gaussin, V., Van d Putte, T., Mishina, T., Hanks, M.C., Zwijsen, A., Huylebroeck, D., Behringer, R.R., Schneider. M.D. (2002). Endocardial cushion and myocardial defects after cardiac myocyte-specific conditional deletion of the bone morphogenetic protein receptor ALK3. Proc Natl Acad Sci U S A99, 2878-83. Garnier, A., Fortin, D., Delomenie, C., Momken, I., Veksler, V., and Ventura-Clapier, R. (2003). Depressed mitochondrial transcription factors and oxidative capacity in rat failing cardiac and skeletal muscles. The Journal of physiology 551, 491-501. Gebara, M.M., Sayre, M.H., and Corden, J.L. (1997). Phosphorylation of the carboxy-terminal repeat domain in RNA polymerase II by cyclin-dependent kinases is sufficient to inhibit transcription. Journal of cellular biochemistry 64, 390-402. Gervais, V., Busso, D., Wasielewski, E., Poterszman, A., Egly, J.M., Thierry, J.C., and Kieffer, B. (2001). Solution structure of the N-terminal domain of the human TFIIH MAT1 subunit: new insights into the RING finger family. The Journal of biological chemistry 276, 7457-7464. Giglia-Mari, G., Coin, F., Ranish, J.A., Hoogstraten, D., Theil, A., Wijgers, N., Jaspers, N.G., Raams, A., Argentini, M., van der Spek, P.J., et al. (2004). A new, tenth subunit of TFIIH is responsible for the DNA repair syndrome trichothiodystrophy group A. Nature genetics 36, 714-719. Glover-Cutter, K., Kim, S., Espinosa, J., and Bentley, D.L. (2008). RNA polymerase II pauses and associates with pre-mRNA processing factors at both ends of genes. Nature structural & molecular biology 15, 71-78. Glover-Cutter, K., Larochelle, S., Erickson, B., Zhang, C., Shokat, K., Fisher, R.P., and Bentley, D.L. (2009). TFIIH-associated Cdk7 kinase functions in phosphorylation of C-terminal domain Ser7 residues, promoter-proximal pausing, and termination by RNA polymerase II. Molecular and cellular biology 29, 5455-5464. Gomes, N.P., Bjerke, G., Llorente, B., Szostek, S.A., Emerson, B.M., and Espinosa, J.M. (2006). Gene-specific requirement for P-TEFb activity and RNA polymerase II phosphorylation within the p53 transcriptional program. Genes & development 20, 601-612.

66

Goto, M., Terada, S., Kato, M., Katoh, M., Yokozeki, T., Tabata, I., and Shimokawa, T. (2000). cDNA Cloning and mRNA analysis of PGC-1 in epitrochlearis muscle in swimming-exercised rats. Biochem Biophys Res Commun 274, 350-354. Guenther, M.G., Levine, S.S., Boyer, L.A., Jaenisch, R., and Young, R.A. (2007). A chromatin landmark and transcription initiation at most promoters in human cells. Cell 130, 77-88. Guhaniyogi, J., and Brewer, G. (2001). Regulation of mRNA stability in mammalian cells. Gene 265, 11-23. Guiguen, A., Soutourina, J., Dewez, M., Tafforeau, L., Dieu, M., Raes, M., Vandenhaute, J., Werner, M., and Hermand, D. (2007). Recruitment of P-TEFb (Cdk9-Pch1) to chromatin by the cap-methyl transferase Pcm1 in fission yeast. The EMBO journal 26, 1552-1559. Guo, X.W., Th'ng, J.P., Swank, R.A., Anderson, H.J., Tudan, C., Bradbury, E.M., and Roberge, M. (1995). Chromosome condensation induced by fostriecin does not require p34cdc2 kinase activity and histone H1 hyperphosphorylation, but is associated with enhanced histone H2A and H3 phosphorylation. The EMBO journal 14, 976-985. Handschin, C., Rhee, J., Lin, J., Tarr, P.T., and Spiegelman, B.M. (2003). An autoregulatory loop controls peroxisome proliferator-activated receptor gamma coactivator 1alpha expression in muscle. Proc Natl Acad Sci U S A100, 7111-7116. Hirose, Y., and Manley, J.L. (1998). RNA polymerase II is an essential mRNA polyadenylation factor. Nature 395, 93-96. Hirose, Y., Tacke, R., and Manley, J.L. (1999). Phosphorylated RNA polymerase II stimulates pre-mRNA splicing. Genes & development 13, 1234-1239. Ho, C.K., and Shuman, S. (1999). Distinct roles for CTD Ser-2 and Ser-5 phosphorylation in the recruitment and allosteric activation of mammalian mRNA capping enzyme. Molecular cell 3, 405-411. Holstege, F.C., Jennings, E.G., Wyrick, J.J., Lee, T.I., Hengartner, C.J., Green, M.R., Golub, T.R., Lander, E.S., and Young, R.A. (1998). Dissecting the regulatory circuitry of a eukaryotic genome. Cell 95, 717-728. Hong, S.W., Hong, S.M., Yoo, J.W., Lee, Y.C., Kim, S., Lis, J.T. and Lee, D.K. (2009). Phosphorylation of the RNA polymerase II C-terminal domain by TFIIH kinase is not essential for transcription of Saccharomyces cerevisiae genome. Proc Natl Acad Sci U S A106, 14276-80. Horton, J.D., Shimomura, I., Brown, M.S., Hammer, R.E., Goldstein, J.L., and Shimano, H. (1998). Activation of cholesterol synthesis in preference to fatty acid synthesis in liver

67

and adipose tissue of transgenic mice overproducing sterol regulatory element-binding protein-2. J Clin Invest 101, 2331-2339. Hu, E., Chen, Z., Fredrickson, T., Zhu, Y., Kirkpatrick, R., Zhang, G.F., Johanson, K., Sung, C.M., Liu, R., and Winkler, J. (2000). Cloning and characterization of a novel human class I histone deacetylase that functions as a transcription repressor. The Journal of biological chemistry 275, 15254-15264. Hu, E., Kim, J.B., Sarraf, P., and Spiegelman, B.M. (1996). Inhibition of adipogenesis through MAP kinase-mediated phosphorylation of PPARgamma. Science 274, 2100-2103. Iizuka, K., Bruick, R.K., Liang, G., Horton, J.D., and Uyeda, K. (2004). Deficiency of carbohydrate response element-binding protein (ChREBP) reduces lipogenesis as well as glycolysis. Proc Natl Acad Sci U S A101, 7281-7286. Illingworth, R., Kerr, A., Desousa, D., Jorgensen, H., Ellis, P., Stalker, J., Jackson, D., Clee, C., Plumb, R., Rogers, J., et al. (2008). A novel CpG island set identifies tissue-specific methylation at developmental gene loci. PLoS biology 6, e22. Imoberdorf, R.M., Topalidou, I., and Strubin, M. (2006). A role for gcn5-mediated global histone acetylation in transcriptional regulation. Molecular and cellular biology 26, 1610-1616. Inamoto, S., Segil, N., Pan, Z.Q., Kimura, M., and Roeder, R.G. (1997). The cyclin-dependent kinase-activating kinase (CAK) assembly factor, MAT1, targets and enhances CAK activity on the POU domains of octamer transcription factors. The Journal of biological chemistry 272, 29852-29858. Ivanovska, I., Jacques, P.E., Rando, O.J., Robert, F., and Winston, F. (2010). Control of chromatin structure by spt6: different consequences in coding and regulatory regions. Molecular and cellular biology 31, 531-541. Iyer, N., Reagan, M.S., Wu, K.J., Canagarajah, B., and Friedberg, E.C. (1996). Interactions involving the human RNA polymerase II transcription/nucleotide excision repair complex TFIIH, the nucleotide excision repair protein XPG, and Cockayne syndrome group B (CSB) protein. Biochemistry 35, 2157-2167. Jenuwein, T., and Allis, C.D. (2001). Translating the histone code. Science 293, 1074-1080. Jiang, C., and Pugh, B.F. (2009). Nucleosome positioning and gene regulation: advances through genomics. Nat Rev Genet 10, 161-172. Kang, B.G., Shin, J.H., Yi, J.K., Kang, H.C., Lee, J.J., Heo, H.S., Chae, J.H., Shin, I., and Kim, C.G. (2007). Corepressor MMTR/DMAP1 is involved in both histone deacetylase

68

1- and TFIIH-mediated transcriptional repression. Molecular and cellular biology 27, 3578-3588. Kangaspeska, S., B. Stride, R. Metivier, M. Polycarpou-Schwarz, D. Ibberson, R.P. Carmouche, V. Benes, F. Gannon, and G. Reid. 2008. Transient cyclical methylation of promoter DNA. Nature. 452:112-5. Kanin, E.I., Kipp, R.T., Kung, C., Slattery, M., Viale, A., Hahn, S., Shokat, K.M., and Ansari, A.Z. (2007). Chemical inhibition of the TFIIH-associated kinase Cdk7/Kin28 does not impair global mRNA synthesis. Proc Natl Acad Sci U S A 104, 5812-5817. Keriel, A., Stary, A., Sarasin, A., Rochette-Egly, C., and Egly, J.M. (2002). XPD mutations prevent TFIIH-dependent transactivation by nuclear receptors and phosphorylation of RARalpha. Cell 109, 125-135. Kershaw, E.E., and Flier, J.S. (2004). Adipose tissue as an endocrine organ. The Journal of clinical endocrinology and metabolism 89, 2548-2556. Kim, J.B., Wright, H.M., Wright, M., and Spiegelman, B.M. (1998). ADD1/SREBP1 activates PPARgamma through the production of endogenous ligand. Proc Natl Acad Sci U S A95, 4333-4337. Kim, M., Suh, H., Cho, E.J., and Buratowski, S. (2009). Phosphorylation of the yeast Rpb1 C-terminal domain at serines 2, 5, and 7. The Journal of biological chemistry 284, 26421-26426. Kim, M.S., T. Kondo, I. Takada, M.Y. Youn, Y. Yamamoto, S. Takahashi, T. Matsumoto, S. Fujiyama, Y. Shirode, I. Yamaoka, H. Kitagawa, K. Takeyama, H. Shibuya, F. Ohtake, and S. Kato. 2009. DNA demethylation in hormone-induced transcriptional derepression. Nature. 461:1007-12. Ko, L.J., Shieh, S.Y., Chen, X., Jayaraman, L., Tamai, K., Taya, Y., Prives, C., and Pan, Z.Q. (1997). p53 is phosphorylated by CDK7-cyclin H in a p36MAT1-dependent manner. Molecular and cellular biology 17, 7220-7229. Komarnitsky, P., Cho, E.J., and Buratowski, S. (2000). Different phosphorylated forms of RNA polymerase II and associated mRNA processing factors during transcription. Genes & development 14, 2452-2460. Korner, C.G., and Wahle, E. (1997). Poly(A) tail shortening by a mammalian poly(A)-specific 3'-exoribonuclease. The Journal of biological chemistry 272, 10448-10456. Korner, C.G., Wormington, M., Muckenthaler, M., Schneider, S., Dehlin, E., and Wahle, E. (1998). The deadenylating nuclease (DAN) is involved in poly(A) tail removal during the meiotic maturation of Xenopus oocytes. The EMBO journal 17, 5427-5437.

69

Korsisaari, N., Rossi, D.J., Paetau, A., Charnay, P., Henkemeyer, M., and Makela, T.P. (2002). Conditional ablation of the Mat1 subunit of TFIIH in Schwann cells provides evidence that Mat1 is not required for general transcription. Journal of cell science 115, 4275-4284. Krogan, N.J., Kim, M., Tong, A., Golshani, A., Cagney, G., Canadien, V., Richards, D.P., Beattie, B.K., Emili, A., Boone, C., et al. (2003). Methylation of histone H3 by Set2 in Saccharomyces cerevisiae is linked to transcriptional elongation by RNA polymerase II. Molecular and cellular biology 23, 4207-4218. Kuhn, R., Schwenk, F., Aguet, F., and Rajewsky, K. (1995). Inducible gene targeting in mice. Science 269, 1427-9. Lane, M.D., Tang, Q.Q., and Jiang, M.S. (1999). Role of the CCAAT enhancer binding proteins (C/EBPs) in adipocyte differentiation. Biochem Biophys Res Commun 266, 677-683. Larochelle, S., Batliner, J., Gamble, M.J., Barboza, N.M., Kraybill, B.C., Blethrow, J.D., Shokat, K.M., and Fisher, R.P. (2006). Dichotomous but stringent substrate selection by the dual-function Cdk7 complex revealed by chemical genetics. Nature structural & molecular biology 13, 55-62. Larochelle, S., Merrick, K., Terret, M., Wohbold, L., Barboza, N.M., Zhang, C., Shokat, K.M., Jallepalli, P.V., and Fisher, R.P. (2007). Requirements for Cdk7 in the Assembly of Cdk1/Cyclin B and Activation of Cdk2 Revealed by Chemical Genetics in Human Cells. Molecular cell 25, 839-850. Larochelle, S., Pandur, J., Fisher, R.P., Salz, H.K., and Suter, B. (1998). Cdk7 is essential for mitosis and for in vivo Cdk-activating kinase activity. Genes & development 12, 370-381. Leclerc, V., Raisin, S., and Leopold, P. (2000). Dominant-negative mutants reveal a role for the Cdk7 kinase at the mid-blastula transition in Drosophila embryos. The EMBO journal 19, 1567-1575. Lee, C.K., Shibata, Y., Rao, B., Strahl, B.D., and Lieb, J.D. (2004). Evidence for nucleosome depletion at active regulatory regions genome-wide. Nature genetics 36, 900-905. Lee, G.E., Kim, J.H., Taylor, M., and Muller, M.T. (2010). DNA methyltransferase 1-associated protein (DMAP1) is a co-repressor that stimulates DNA methylation globally and locally at sites of double strand break repair. The Journal of biological chemistry 285, 37630-37640. Lee, K.M., Miklos, I., Du, H., Watt, S., Szilagyi, Z., Saiz, J.E., Madabhushi, R., Penkett, C.J., Sipiczki, M., Bahler, J., et al. (2005). Impairment of the TFIIH-associated CDK-

70

activating kinase selectively affects cell cycle-regulated gene expression in fission yeast. Molecular biology of the cell 16, 2734-2745. Lee, W., Tillo, D., Bray, N., Morse, R.H., Davis, R.W., Hughes, T.R., and Nislow, C. (2007). A high-resolution atlas of nucleosome occupancy in yeast. Nature genetics 39, 1235-1244. Lehman, J.J., Barger, P.M., Kovacs, A., Saffitz, J.E., Medeiros, D.M., and Kelly, D.P. (2000). Peroxisome proliferator-activated receptor gamma coactivator-1 promotes cardiac mitochondrial biogenesis. J Clin Invest 106, 847-856. Lehman, J.J., Boudina, S., Banke, N.H., Sambandam, N., Han, X., Young, D.M., Leone, T.C., Gross, R.W., Lewandowski, E.D., Abel, E.D., et al. (2008). The transcriptional coactivator PGC-1alpha is essential for maximal and efficient cardiac mitochondrial fatty acid oxidation and lipid homeostasis. American journal of physiology 295, H185-196. Lehmann, J.M., Moore, L.B., Smith-Oliver, T.A., Wilkison, W.O., Willson, T.M., and Kliewer, S.A. (1995). An antidiabetic thiazolidinedione is a high affinity ligand for peroxisome proliferator-activated receptor gamma (PPAR gamma). The Journal of biological chemistry 270, 12953-12956. Levy, N.S., Chung, S., Furneaux, H., and Levy, A.P. (1998). Hypoxic stabilization of vascular endothelial growth factor mRNA by the RNA-binding protein HuR. The Journal of biological chemistry 273, 6417-6423. Lewis, B.A., Kim, T.K., and Orkin, S.H. (2000). A downstream element in the human beta-globin promoter: evidence of extended sequence-specific transcription factor IID contacts. Proc Natl Acad Sci U S A97, 7172-7177. Li, B., Howe, L., Anderson, S., Yates, J.R., 3rd, and Workman, J.L. (2003). The Set2 histone methyltransferase functions through the phosphorylated carboxyl-terminal domain of RNA polymerase II. The Journal of biological chemistry 278, 8897-8903. Li, X., Urwyler, O., and Suter, B. (2010). Drosophila Xpd regulates Cdk7 localization, mitotic kinase activity, spindle dynamics, and chromosome segregation. PLoS genetics 6, e1000876. Lim, C.Y., Santoso, B., Boulay, T., Dong, E., Ohler, U., and Kadonaga, J.T. (2004). The MTE, a new core promoter element for transcription by RNA polymerase II. Genes & development 18, 1606-1617. Lin, J., Handschin, C., and Spiegelman, B.M. (2005). Metabolic control through the PGC-1 family of transcription coactivators. Cell Metab 1, 361-370. Lin, J., Puigserver, P., Donovan, J., Tarr, P., and Spiegelman, B.M. (2002a). Peroxisome proliferator-activated receptor gamma coactivator 1beta (PGC-1beta ), a novel PGC-1-

71

related transcription coactivator associated with host cell factor. The Journal of biological chemistry 277, 1645-1648. Lin, P.S., Dubois, M.F., and Dahmus, M.E. (2002b). TFIIF-associating carboxyl-terminal domain phosphatase dephosphorylates phosphoserines 2 and 5 of RNA polymerase II. The Journal of biological chemistry 277, 45949-45956. Long, J.J., Leresche, A., Kriwacki, R.W., and Gottesfeld, J.M. (1998). Repression of TFIIH transcriptional activity and TFIIH-associated cdk7 kinase activity at mitosis. Molecular and cellular biology 18, 1467-1476. Lu, H., Fisher, R.P., Bailey, P., and Levine, A.J. (1997). The CDK7-cycH-p36 complex of transcription factor IIH phosphorylates p53, enhancing its sequence-specific DNA binding activity in vitro. Molecular and cellular biology 17, 5923-5934. Mäkelä, T.P., Parvin, J.D., Kim, J., Huber, L.J., Sharp, P.A., and Weinberg, R.A. (1995). A kinase-deficient transcription factor IIH is functional in basal and activated transcription. Proc Natl Acad Sci USA 92, 5174-5178. Malik, S., and Roeder, R.G. (2010). The metazoan Mediator co-activator complex as an integrative hub for transcriptional regulation. Nat Rev Genet 11, 761-772. Matsuda, M., Korn, B.S., Hammer, R.E., Moon, Y.A., Komuro, R., Horton, J.D., Goldstein, J.L., Brown, M.S., and Shimomura, I. (2001). SREBP cleavage-activating protein (SCAP) is required for increased lipid synthesis in liver induced by cholesterol deprivation and insulin elevation. Genes & development 15, 1206-1216. Mavrich, T.N., Jiang, C., Ioshikhes, I.P., Li, X., Venters, B.J., Zanton, S.J., Tomsho, L.P., Qi, J., Glaser, R.L., Schuster, S.C., et al. (2008). Nucleosome organization in the Drosophila genome. Nature 453, 358-362. McCracken, S., Fong, N., Rosonina, E., Yankulov, K., Brothers, G., Siderovski, D., Hessel, A., Foster, S., Shuman, S., and Bentley, D.L. (1997a). 5'-Capping enzymes are targeted to pre-mRNA by binding to the phosphorylated carboxy-terminal domain of RNA polymerase II. Genes & development 11, 3306-3318. McCracken, S., Fong, N., Yankulov, K., Ballantyne, S., Pan, G., Greenblatt, J., Patterson, S.D., Wickens, M., and Bentley, D.L. (1997b). The C-terminal domain of RNA polymerase II couples mRNA processing to transcription. Nature 385, 357-361. Meininghaus, M., Chapman, R.D., Horndasch, M., and Eick, D. (2000). Conditional expression of RNA polymerase II in mammalian cells. Deletion of the carboxyl-terminal domain of the large subunit affects early steps in transcription. The Journal of biological chemistry 275, 24375-24382.

72

Metivier, R., R. Gallais, C. Tiffoche, C. Le Peron, R.Z. Jurkowska, R.P. Carmouche, D. Ibberson, P. Barath, F. Demay, G. Reid, V. Benes, A. Jeltsch, F. Gannon, and G. Salbert. 2008. Cyclical DNA methylation of a transcriptionally active promoter. Nature. 452:45-50. Millhouse, S., and Manley, J.L. (2005). The C-terminal domain of RNA polymerase II functions as a phosphorylation-dependent splicing activator in a heterologous protein. Molecular and cellular biology 25, 533-544. Mishra, S.K., and Das, B.R. (1992). (ADP-ribosyl)ation pattern of chromosomal proteins during ageing. Cellular and molecular biology 38, 457-462. Mizzen, C.A., Yang, X.J., Kokubo, T., Brownell, J.E., Bannister, A.J., Owen-Hughes, T., Workman, J., Wang, L., Berger, S.L., Kouzarides, T., et al. (1996). The TAF(II)250 subunit of TFIID has histone acetyltransferase activity. Cell 87, 1261-1270. Molina-Navarro, M.M., Castells-Roca, L., Belli, G., Garcia-Martinez, J., Marin-Navarro, J., Moreno, J., Perez-Ortin, J.E., and Herrero, E. (2008). Comprehensive transcriptional analysis of the oxidative response in yeast. The Journal of biological chemistry 283, 17908-17918. Morris, D.P., Michelotti, G.A., and Schwinn, D.A. (2005). Evidence that phosphorylation of the RNA polymerase II carboxyl-terminal repeats is similar in yeast and humans. The Journal of biological chemistry 280, 31368-31377. Mortillaro, M.J., Blencowe, B.J., Wei, X., Nakayasu, H., Du, L., Warren, S.L., Sharp, P.A., and Berezney, R. (1996). A hyperphosphorylated form of the large subunit of RNA polymerase II is associated with splicing complexes and the nuclear matrix. Proc Natl Acad Sci U S A93, 8253-8257. Moteki, S., and Price, D. (2002). Functional coupling of capping and transcription of mRNA. Molecular cell 10, 599-609. Muse, G.W., Gilchrist, D.A., Nechaev, S., Shah, R., Parker, J.S., Grissom, S.F., Zeitlinger, J., and Adelman, K. (2007). RNA polymerase is poised for activation across the genome. Nature genetics 39, 1507-1511. Ng, H.H., Robert, F., Young, R.A., and Struhl, K. (2003). Targeted recruitment of Set1 histone methylase by elongating Pol II provides a localized mark and memory of recent transcriptional activity. Molecular cell 11, 709-719. Nonet, M., Sweetser, D., and Young, R.A. (1987). Functional redundancy and structural polymorphism in the large subunit of RNA polymerase II. Cell 50, 909-915.

73

O'Brien, T., Hardin, S., Greenleaf, A., and Lis, J.T. (1994). Phosphorylation of RNA polymerase II C-terminal domain and transcriptional elongation. Nature 370, 75-77. Ogryzko, V.V., Schiltz, R.L., Russanova, V., Howard, B.H., and Nakatani, Y. (1996). The transcriptional coactivators p300 and CBP are histone acetyltransferases. Cell 87, 953-959. Olefsky, J.M. (2000). Treatment of insulin resistance with peroxisome proliferator-activated receptor gamma agonists. J Clin Invest 106, 467-472. Palomer, X., Alvarez-Guardia, D., Rodriguez-Calvo, R., Coll, T., Laguna, J.C., Davidson, M.M., Chan, T.O., Feldman, A.M., and Vazquez-Carrera, M. (2009). TNF-alpha reduces PGC-1alpha expression through NF-kappaB and p38 MAPK leading to increased glucose oxidation in a human cardiac cell model. Cardiovascular research 81, 703-712. Patel, A.A., and Steitz, J.A. (2003). Splicing double: insights from the second spliceosome. Nature reviews 4, 960-970. Patel, S.A., and Simon, M.C. (2010). Functional analysis of the Cdk7/cyclin H/Mat1 complex in mouse embryonic stem cells and embryos. The Journal of biological chemistry 285, 15587-98. Pei, Y., Schwer, B., and Shuman, S. (2003). Interactions between fission yeast Cdk9, its cyclin partner Pch1, and mRNA capping enzyme Pct1 suggest an elongation checkpoint for mRNA quality control. The Journal of biological chemistry 278, 7180-7188. Perez Canadillas, J.M., and Varani, G. (2003). Recognition of GU-rich polyadenylation regulatory elements by human CstF-64 protein. The EMBO journal 22, 2821-2830. Peterlin, B.M., and Price, D.H. (2006). Controlling the elongation phase of transcription with P-TEFb. Molecular cell 23, 297-305. Pokholok, D.K., Harbison, C.T., Levine, S., Cole, M., Hannett, N.M., Lee, T.I., Bell, G.W., Walker, K., Rolfe, P.A., Herbolsheimer, E., et al. (2005). Genome-wide map of nucleosome acetylation and methylation in yeast. Cell 122, 517-527. Puigserver, P., Adelmant, G., Wu, Z., Fan, M., Xu, J., O'Malley, B., and Spiegelman, B.M. (1999). Activation of PPARgamma coactivator-1 through transcription factor docking. Science 286, 1368-1371. Puigserver, P., Wu, Z., Park, C.W., Graves, R., Wright, M., and Spiegelman, B.M. (1998). A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell 92, 829-839.

74

Qiu, H., Hu, C., and Hinnebusch, A.G. (2009). Phosphorylation of the Pol II CTD by KIN28 enhances BUR1/BUR2 recruitment and Ser2 CTD phosphorylation near promoters. Molecular cell 33, 752-762. Rabinowicz, P.D., Palmer, L.E., May, B.P., Hemann, M.T., Lowe, S.W., McCombie, W.R., and Martienssen, R.A. (2003). Genes and transposons are differentially methylated in plants, but not in mammals. Genome research 13, 2658-2664. Radonjic, M., Andrau, J.C., Lijnzaad, P., Kemmeren, P., Kockelkorn, T.T., van Leenen, D., van Berkum, N.L., and Holstege, F.C. (2005). Genome-wide analyses reveal RNA polymerase II located upstream of genes poised for rapid response upon S. cerevisiae stationary phase exit. Molecular cell 18, 171-183. Raijmakers, R., Schilders, G., and Pruijn, G.J. (2004). The exosome, a molecular machine for controlled RNA degradation in both nucleus and cytoplasm. Eur J Cell Biol 83, 175-183. Rakyan, V.K., Hildmann, T., Novik, K.L., Lewin, J., Tost, J., Cox, A.V., Andrews, T.D., Howe, K.L., Otto, T., Olek, A., et al. (2004). DNA methylation profiling of the human major histocompatibility complex: a pilot study for the human epigenome project. PLoS biology 2, e405. Ramanathan, Y., Rajpara, S.M., Reza, S.M., Lees, E., Shuman, S., Mathews, M.B., and Pe'ery, T. (2001). Three RNA polymerase II carboxyl-terminal domain kinases display distinct substrate preferences. The Journal of biological chemistry 276, 10913-10920. Rangwala, S.M., Rhoades, B., Shapiro, J.S., Rich, A.S., Kim, J.K., Shulman, G.I., Kaestner, K.H., and Lazar, M.A. (2003). Genetic modulation of PPARgamma phosphorylation regulates insulin sensitivity. Developmental cell 5, 657-663. Rasmussen, E.B., and Lis, J.T. (1993). In vivo transcriptional pausing and cap formation on three Drosophila heat shock genes. Proc Natl Acad Sci U S A90, 7923-7927. Rhee, J., Ge, H., Yang, W., Fan, M., Handschin, C., Cooper, M., Lin, J., Li, C., and Spiegelman, B.M. (2006). Partnership of PGC-1alpha and HNF4alpha in the regulation of lipoprotein metabolism. The Journal of biological chemistry 281, 14683-14690. Rickert, P., Corden, J.L., and Lees, E. (1999). Cyclin C/CDK8 and cyclin H/CDK7/p36 are biochemically distinct CTD kinases. Oncogene 18, 1093-1102. Rochette-Egly, C. (2003). Nuclear receptors: integration of multiple signalling pathways through phosphorylation. Cell Signal 15, 355-366. Rochette-Egly, C., Adam, S., Rossignol, M., Egly, J.M., and Chambon, P. (1997). Stimulation of RAR alpha activation function AF-1 through binding to the general transcription factor TFIIH and phosphorylation by CDK7. Cell 90, 97-107.

75

Rosales, J., Han, B., and Lee, K.Y. (2003). Cdk7 functions as a cdk5 activating kinase in brain. Cell Physiol Biochem 13, 285-296. Rosen, E.D., and MacDougald, O.A. (2006). Adipocyte differentiation from the inside out. Nature reviews 7, 885-896. Rosen, E.D., Sarraf, P., Troy, A.E., Bradwin, G., Moore, K., Milstone, D.S., Spiegelman, B.M., and Mortensen, R.M. (1999). PPAR gamma is required for the differentiation of adipose tissue in vivo and in vitro. Molecular cell 4, 611-617. Rossi, D.J., Londesborough, A., Korsisaari, N., Pihlak, A., Lehtonen, E., Henkemeyer, M., and Makela, T.P. (2001). Inability to enter S phase and defective RNA polymerase II CTD phosphorylation in mice lacking Mat1. The EMBO journal 20, 2844-2856. Rowe, G.C., Jiang, A., and Arany, Z (2010). PGC-1 coactivators in cardiac development and disease. Circulation research 107, 825-838. Roy, R., Adamczewski, J.P., Seroz, T., Vermeulen, W., Tassan, J.P., Schaeffer, L., Nigg, E.A., Hoeijmakers, J.H., and Egly, J.M. (1994). The MO15 cell cycle kinase is associated with the TFIIH transcription- DNA repair factor. Cell 79, 1093-1101. Sandrock, B., and Egly, J.M. (2001). A yeast four-hybrid system identifies Cdk-activating kinase as a regulator of the XPD helicase, a subunit of transcription factor IIH. The Journal of biological chemistry 276, 35328-35333. Sano, M., Abdellatif, M., Oh, H., Xie, M., Bagella, L., Giordano, A., Michael, L.H., DeMayo, F.J., and Schneider, M.D. (2002). Activation and function of cyclin T-Cdk9 (positive transcription elongation factor-b) in cardiac muscle-cell hypertrophy. Nat Med 8, 1310-1317. Sano, M., Wang, S.C., Shirai, M., Scaglia, F., Xie, M., Sakai, S., Tanaka, T., Kulkarni, P.A., Barger, P.M., Youker, K.A., et al. (2004). Activation of cardiac Cdk9 represses PGC-1 and confers a predisposition to heart failure. The EMBO journal 23, 3559-3569. Santos-Rosa, H., Schneider, R., Bannister, A.J., Sherriff, J., Bernstein, B.E., Emre, N.C., Schreiber, S.L., Mellor, J., and Kouzarides, T. (2002). Active genes are tri-methylated at K4 of histone H3. Nature 419, 407-411. Sato, R. (2010). Sterol metabolism and SREBP activation. Archives of biochemistry and biophysics 501, 177-181. Saunders, A., Core, L.J., and Lis, J.T. (2006). Breaking barriers to transcription elongation. Nature reviews 7, 557-567.

76

Schaeffer, L., Roy, R., Humbert, S., Moncollin, V., Vermeulen, W., Hoeijmakers, J.H., Chambon, P., and Egly, J.M. (1993). DNA repair helicase: a component of BTF2 (TFIIH) basic transcription factor [see comments]. Science 260, 58-63. Schroeder, S.C., Schwer, B., Shuman, S., and Bentley, D. (2000). Dynamic association of capping enzymes with transcribing RNA polymerase II. Genes & development 14, 2435-2440. Schubeler, D., MacAlpine, D.M., Scalzo, D., Wirbelauer, C., Kooperberg, C., van Leeuwen, F., Gottschling, D.E., O'Neill, L.P., Turner, B.M., Delrow, J., et al. (2004). The histone modification pattern of active genes revealed through genome-wide chromatin analysis of a higher eukaryote. Genes & development 18, 1263-1271. Schultz, P., Fribourg, S., Poterszman, A., Mallouh, V., Moras, D., and Egly, J.M. (2000). Molecular structure of human TFIIH. Cell 102, 599-607. Schwartz, B.E., Larochelle, S., Suter, B., and Lis, J.T. (2003). Cdk7 is required for full activation of Drosophila heat shock genes and RNA polymerase II phosphorylation in vivo. Molecular and cellular biology 23, 6876-6886. Sciortino, S., Gurtner, A., Manni, I., Fontemaggi, G., Dey, A., Sacchi, A., Ozato, K., and Piaggio, G. (2001). The cyclin B1 gene is actively transcribed during mitosis in HeLa cells. EMBO reports 2, 1018-1023. Sekinger, E.A., Moqtaderi, Z., and Struhl, K. (2005). Intrinsic histone-DNA interactions and low nucleosome density are important for preferential accessibility of promoter regions in yeast. Molecular cell 18, 735-748. Serizawa, H., Makela, T.P., Conaway, J.W., Conaway, R.C., Weinberg, R.A., and Young, R.A. (1995). Association of Cdk-activating kinase subunits with transcription factor TFIIH. Nature 374, 280-282. Shiekhattar, R., Mermelstein, F., Fisher, R.P., Drapkin, R., Dynlacht, B., Wessling, H.C., Morgan, D.O., and Reinberg, D. (1995). Cdk-activating kinase complex is a component of human transcription factor TFIIH. Nature 374, 283-287. Shimano, H., Horton, J.D., Shimomura, I., Hammer, R.E., Brown, M.S., and Goldstein, J.L. (1997a). Isoform 1c of sterol regulatory element binding protein is less active than isoform 1a in livers of transgenic mice and in cultured cells. J Clin Invest 99, 846-854. Shimano, H., Shimomura, I., Hammer, R.E., Herz, J., Goldstein, J.L., Brown, M.S., and Horton, J.D. (1997b). Elevated levels of SREBP-2 and cholesterol synthesis in livers of mice homozygous for a targeted disruption of the SREBP-1 gene. J Clin Invest 100, 2115-2124.

77

Shimomura, I., Bashmakov, Y., and Horton, J.D. (1999). Increased levels of nuclear SREBP-1c associated with fatty livers in two mouse models of diabetes mellitus. The Journal of biological chemistry 274, 30028-30032. Shimomura, I., Shimano, H., Horton, J.D., Goldstein, J.L., and Brown, M.S. (1997). Differential expression of exons 1a and 1c in mRNAs for sterol regulatory element binding protein-1 in human and mouse organs and cultured cells. J Clin Invest 99, 838-845. Shimomura, I., Shimano, H., Korn, B.S., Bashmakov, Y., and Horton, J.D. (1998). Nuclear sterol regulatory element-binding proteins activate genes responsible for the entire program of unsaturated fatty acid biosynthesis in transgenic mouse liver. The Journal of biological chemistry 273, 35299-35306. Shivaswamy, S., Bhinge, A., Zhao, Y., Jones, S., Hirst, M., and Iyer, V.R. (2008). Dynamic remodeling of individual nucleosomes across a eukaryotic genome in response to transcriptional perturbation. PLoS biology 6, e65. Smale, S.T., and Baltimore, D. (1989). The "initiator" as a transcription control element. Cell 57, 103-113. Smale, S.T., and Kadonaga, J.T. (2003). The RNA polymerase II core promoter. Annu Rev Biochem 72, 449-479. Spilianakis, C., Kretsovali, A., Agalioti, T., Makatounakis, T., Thanos, D., and Papamatheakis, J. (2003). CIITA regulates transcription onset viaSer5-phosphorylation of RNA Pol II. The EMBO journal 22, 5125-5136. Suzuki, M.M., and Bird, A. (2008). DNA methylation landscapes: provocative insights from epigenomics. Nat Rev Genet 9, 465-476. Suzuki, M.M., Kerr, A.R., De Sousa, D., and Bird, A. (2007). CpG methylation is targeted to transcription units in an invertebrate genome. Genome research 17, 625-631. Takahata, S., Yu, Y., and Stillman, D.J. (2009). FACT and Asf1 regulate nucleosome dynamics and coactivator binding at the HO promoter. Molecular cell 34, 405-415. Svedruzic, Z.M. (2008). Mammalian cytosine DNA methyltransferase Dnmt1: enzymatic mechanism, novel mechanism-based inhibitors, and RNA-directed DNA methylation. Curr Med Chem 15, 92-106. Tang, Q.Q., Otto, T.C., and Lane, M.D. (2003). CCAAT/enhancer-binding protein beta is required for mitotic clonal expansion during adipogenesis. Proc Natl Acad Sci U S A 100, 850-855.

78

Tassan, J.P., Jaquenoud, M., Fry, A.M., Frutiger, S., Hughes, G.J., and Nigg, E.A. (1995). In vitro assembly of a functional human CDK7-cyclin H complex requires MAT1, a novel 36 kDa RING finger protein. The EMBO journal 14, 5608-5617. Thomson, S., Mahadevan, L.C., and Clayton, A.L. (1999). MAP kinase-mediated signalling to nucleosomes and immediate-early gene induction. Seminars in cell & developmental biology 10, 205-214. Tiraby, C., Tavernier, G., Lefort, C., Larrouy, D., Bouillaud, F., Ricquier, D., and Langin, D. (2003). Acquirement of brown fat cell features by human white adipocytes. The Journal of biological chemistry 278, 33370-33376. Tobe, K., Suzuki, R., Aoyama, M., Yamauchi, T., Kamon, J., Kubota, N., Terauchi, Y., Matsui, J., Akanuma, Y., Kimura, S., et al. (2001). Increased expression of the sterol regulatory element-binding protein-1 gene in insulin receptor substrate-2(-/-) mouse liver. The Journal of biological chemistry 276, 38337-38340. Tokusumi, Y., Ma, Y., Song, X., Jacobson, R.H., and Takada, S. (2007). The new core promoter element XCPE1 (X Core Promoter Element 1) directs activator-, mediator-, and TATA-binding protein-dependent but TFIID-independent RNA polymerase II transcription from TATA-less promoters. Molecular and cellular biology 27, 1844-1858. Tontonoz, P., Hu, E., and Spiegelman, B.M. (1994). Stimulation of adipogenesis in fibroblasts by PPAR gamma 2, a lipid-activated transcription factor. Cell 79, 1147-1156. Trigon, S., Serizawa, H., Conaway, J.W., Conaway, R.C., Jackson, S.P., and Morange, M. (1998). Characterization of the residues phosphorylated in vitro by different C- terminal domain kinases. The Journal of biological chemistry 273, 6769-6775. Valadkhan, S., and Jaladat, Y. (2010). The spliceosomal proteome: At the heart of the largest cellular ribonucleoprotein machine. Proteomics. Valay, J.G., Simon, M., Dubois, M.F., Bensaude, O., Facca, C., and Faye, G. (1995). The KIN28 gene is required both for RNA polymerase II mediated transcription and phosphorylation of the Rpb1p CTD. Journal of molecular biology 249, 535-544. Van Der Kelen, K., Beyaert, R., Inze, D., and De Veylder, L. (2009). Translational control of eukaryotic gene expression. Critical reviews in biochemistry and molecular biology 44, 143-168. van Vuuren, A.J., Vermeulen, W., Ma, L., Weeda, G., Appeldoorn, E., Jaspers, N.G., van der Eb, A.J., Bootsma, D., Hoeijmakers, J.H., Humbert, S., et al. (1994). Correction of xeroderma pigmentosum repair defect by basal transcription factor BTF2 (TFIIH). EMBO J 13, 1645-1653.

79

Vaquero, A., Loyola, A., and Reinberg, D. (2003). The constantly changing face of chromatin. Sci Aging Knowledge Environ 2003, RE4. Vega, R.B., Huss, J.M., and Kelly, D.P. (2000). The coactivator PGC-1 cooperates with peroxisome proliferator-activated receptor alpha in transcriptional control of nuclear genes encoding mitochondrial fatty acid oxidation enzymes. Molecular and cellular biology 20, 1868-1876. Viladevall, L., St Amour, C.V., Rosebrock, A., Schneider, S., Zhang, C., Allen, J.J., Shokat, K.M., Schwer, B., Leatherwood, J.K., and Fisher, R.P. (2009). TFIIH and P-TEFb coordinate transcription with capping enzyme recruitment at specific genes in fission yeast. Molecular cell 33, 738-751. Wallberg, A.E., Yamamura, S., Malik, S., Spiegelman, B.M., and Roeder, R.G. (2003). Coordination of p300-mediated chromatin remodeling and TRAP/mediator function through coactivator PGC-1alpha. Molecular cell 12, 1137-1149. Wallenfang, M.R., and Seydoux, G. (2002). cdk-7 Is required for mRNA transcription and cell cycle progression in Caenorhabditis elegans embryos. Proc Natl Acad Sci U S A99, 5527-5532. Wang, J.G., Barsky, L.W., Davicioni, E., Weinberg, K.I., Triche, T.J., Zhang, X.K., and Wu, L. (2006). Retinoic acid induces leukemia cell G1 arrest and transition into differentiation by inhibiting cyclin-dependent kinase-activating kinase binding and phosphorylation of PML/RARalpha. Faseb J 20, 2142-2144. Wang, X., Lee, C., Gilmour, D.S., and Gergen, J.P. (2007). Transcription elongation controls cell fate specification in the Drosophila embryo. Genes & development 21, 1031-1036. Wang, Z., Svejstrup, J.Q., Feaver, W.J., Wu, X., Kornberg, R.D., and Friedberg, E.C. (1994). Transcription factor b (TFIIH) is required during nucleotide-excision repair in yeast. Nature 368, 74-76. Warren, A.J. (2002). Eukaryotic transcription factors. Curr Opin Struct Biol 12, 107-114. Wilson-Fritch, L., Nicoloro, S., Chouinard, M., Lazar, M.A., Chui, P.C., Leszyk, J., Straubhaar, J., Czech, M.P., and Corvera, S. (2004). Mitochondrial remodeling in adipose tissue associated with obesity and treatment with rosiglitazone. J Clin Invest 114, 1281-1289. Wu, Z., Puigserver, P., Andersson, U., Zhang, C., Adelmant, G., Mootha, V., Troy, A., Cinti, S., Lowell, B., Scarpulla, R.C., et al. (1999). Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell 98, 115-124.

80

Xiao, T., Hall, H., Kizer, K.O., Shibata, Y., Hall, M.C., Borchers, C.H., and Strahl, B.D. (2003). Phosphorylation of RNA polymerase II CTD regulates H3 methylation in yeast. Genes & development 17, 654-663. Xu, Y.X., and Manley, J.L. (2007). The prolyl isomerase Pin1 functions in mitotic chromosome condensation. Molecular cell 26, 287-300. Yahagi, N., Shimano, H., Hasty, A.H., Matsuzaka, T., Ide, T., Yoshikawa, T., Amemiya-Kudo, M., Tomita, S., Okazaki, H., Tamura, Y., et al. (2002). Absence of sterol regulatory element-binding protein-1 (SREBP-1) ameliorates fatty livers but not obesity or insulin resistance in Lep(ob)/Lep(ob) mice. The Journal of biological chemistry 277, 19353-19357. Yamauchi, T., Kamon, J., Waki, H., Murakami, K., Motojima, K., Komeda, K., Ide, T., Kubota, N., Terauchi, Y., Tobe, K., et al. (2001). The mechanisms by which both heterozygous peroxisome proliferator-activated receptor gamma (PPARgamma) deficiency and PPARgamma agonist improve insulin resistance. The Journal of biological chemistry 276, 41245-41254. Yeo, M., Lin, P.S., Dahmus, M.E., and Gill, G.N. (2003). A novel RNA polymerase II C-terminal domain phosphatase that preferentially dephosphorylates serine 5. The Journal of biological chemistry 278, 26078-26085. Yoh, S.M., Cho, H., Pickle, L., Evans, R.M., and Jones, K.A. (2007). The Spt6 SH2 domain binds Ser2-P RNAPII to direct Iws1-dependent mRNA splicing and export. Genes & development 21, 160-174. Zeitlinger, J., Stark, A., Kellis, M., Hong, J.W., Nechaev, S., Adelman, K., Levine, M., and Young, R.A. (2007). RNA polymerase stalling at developmental control genes in the Drosophila melanogaster embryo. Nature genetics 39, 1512-1516. Zhang, J., and Corden, J.L. (1991). Identification of phosphorylation sites in the repetitive carboxyl-terminal domain of the mouse RNA polymerase II largest subunit. The Journal of biological chemistry 266, 2290-2296. Zhang, Y., and Reinberg, D. (2001). Transcription regulation by histone methylation: interplay between different covalent modifications of the core histone tails. Genes & development 15, 2343-2360. Zhou, M., Halanski, M.A., Radonovich, M.F., Kashanchi, F., Peng, J., Price, D.H., and Brady, J.N. (2000). Tat modifies the activity of CDK9 to phosphorylate serine 5 of the RNA polymerase II carboxyl-terminal domain during human immunodeficiency virus type 1 transcription. Molecular and cellular biology 20, 5077-5086.

81

Zhou, T., and Chiang, C.M. (2001). The intronless and TATA-less human TAF(II)55 gene contains a functional initiator and a downstream promoter element. The Journal of biological chemistry 276, 25503-25511.