Genome wide targeting of the epigenetic regulatory protein CTCF to gene promoters by the transcription factor TFII-­‐I. by Rodrigo A. Peña-­‐Hernández Principal Investigator Dr. Michael Witcher Submitted December 2015 Department of Experimental Medicine McGill University Montreal, Quebec Canada A thesis submitted to the Faculty of Graduate Studies and Research in partial fulfillment of the requirements of the degree of MASTER OF SCIENCE ©Rodrigo A. Peña-­‐Hernández
TABLE OF CONTENTS____________________________________________________________ Abstract……………………………………………………………………………………………………………………………...... iii Résumé........................................................................................................................................ iv Acknowledgments……………………………………………………………………………………………………………….… v 1. Introduction……………………………………………………………………………………………………………….…..... 1 DNA TRANSCRIPTIONAL REGULATION…………………………………………………………………………..…… 2 Transcription Preinitiation…………………………………………………………………………………….……. 2 Transcription Initiation………………………………………………………………………………………….……. 5 Transcriptional Pausing………………………………………………………………………………………….…… 7 Transcriptional Elongation………………………………………………………………………………………….. 9 Transcriptional Termination……………………………………………………………………………………….. 12 Transcription Reinitiation……………………………………………………………………………………………. 13 EPIGENETIC REGULATION………………………………………………………………………………………………….. 13 DNA Methylation………………………………………………………………………………………………………... 14 Histone Modifications…………………………………………………………………………………………………. 15 Histone Acetylation…………………………………………………………………………………………. 16 Histone Methylation………………………………………………………………………………………… 17 GENERAL TRANSCRIPTION FACTOR II-­‐I (TFII-­‐I) …………………………………………………………………… 20 Genomic distribution of TFII-­‐I and epigenetic role………………………………………………….…… 24 CCCTC-­‐BINDING FACTOR (CTCF)………………………………………………………………..………………………. 26 CTCF Role in Genome Organization…………………………………………………………………………….. 29 CTCF and Insulator activity………………………………………………………………………………. 29 Role of CTCF in Chromatin Boundaries ……………………………………………………………. 30 Role of CTCF in Chromatin loop formation………………………………………………..…….. 31 CTCF Post-­‐Translational Modifications……………………………………………………………………….. 32 Relationship between CTCF functions and PARylation..………………………….……….. 34 CTCF and Epigenetic Regulation of Transcription……………………………………………………….. 35 CTCF and Transcription…………………………………………………………………………………... 37 2. Aims...………………………………….……………………………………………………………................................. 39 3. Materials and Methods Cell Culture and growth curves………………………………………………………………………………..... 40 Generation of CTCF Knockdowns……………………………………………………………………………….. 40 i 4.
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Antibodies………………………………………………………………………………………………………………….. 40 Cell Cycle and Flow cytometry Analysis………………………………………………………………………. 41 Mass Spectrometry and Protein sequencing………………………………………………………………. 41 Column Chromatography……………………………………………………………………………………………. 42 Confocal microscopy…………………………………………………………………………………………………… 42 Western Blot………………………………………………………………………………………………………………. 43 Gene Expression Analysis……………………………………………………………………………………………. 43 Co-­‐Immunoprecipitation…………………………………………………………………………………….………. 42 Chromatin Immunoprecipitation (ChIP)………………………………………………………………………. 44 ChIP-­‐seq and bioinformatics……………………………………………………………………………………….. 45 Gene Ontology and KEGG analysis………………………………………………………………………………. 46 Results ……………………………………………………………………………………………………………………………... 47 Identification of CTCF Interacting Partners…………………………………………………………………. 47 CTCF interaction with TFII-­‐I.……………………………………………………………………………………….. 47 TFII-­‐I and CTCF cooperate to regulate of gene expression…………………………………..……… 48 CTCF and TFII-­‐I cooperation regulates cell survival…………………………………………………….. 51 Figures.……………………………..……………………………………………………………………………………………… 53 Discussion..………………………………………………………………………………………………………………………. 64 Identification of CTCF Interacting Partners...……………………………………………………………… 64 CTCF interaction with TFII-­‐I..………………………………………………………………………………………. 65 TFII-­‐I and CTCF cooperate to regulate of gene expression………………………………………….. 66 CTCF and TFII-­‐I cooperation regulates cell survival……………………………………………………… 69 Conclusion...……………………………………………………………………………………………………………………... 75 References.…………………………………………………………………………………………………………………..…... 76 Appendices…………………………………………………………………………………………………………….…………. 96 ii Abstract_______________________________________________________________________ CCCTC binding factor CTCF is a key regulator of nuclear chromatin structure and gene regulation. The impact of CTCF on transcriptional output is quite varied, ranging from repression, to transcriptional pausing and transactivation. The multifunctional nature of CTCF may be directed solely through remodelling chromatin architecture. However, another hypothesis is that the multifunctional nature of CTCF is mediated, in part, through differential association with protein partners having unique properties. Consistent with this hypothesis, our Mass-­‐Spec analyses of CTCF interacting partners reveal an association with four functional groups; chromatin remodelers, transcription factors, DNA damage repair proteins and mRNA processing factors. Biochemical fractionation of CTCF indicates that distinct CTCF complexes are indeed assembled on DNA. Unexpectedly, we found that the interaction between CTCF and one of our newly identified cofactors, TFII-­‐I is important for directing binding of CTCF to the p14ARF gene. Using Chip-­‐seq analysis we found TFII-­‐I essential for directing CTCF to the promoter proximal regulatory regions of target genes across the genome, particularly at genes involved in metabolism. At genes co-­‐regulated by CTCF and TFII-­‐I, we find knockdown (KD) of TFII-­‐I results in diminished CTCF binding, lack of CDK8 recruitment, and an attenuation of RNA Pol II phosphorylation at Serine 5. Phenotypically, KD of TFII-­‐I alters the cellular response to metabolic stress. Our data indicate TFII-­‐I directs CTCF binding to target genes where the two proteins cooperate to recruit CDK8 and enhance transcription initiation. iii Résumé_______________________________________________________________________ CCCTC Binding factor (CTCF) est un facteur de transcription qui joue un rôle primordial dans la structure de la chromatine et la régulation de l'expression génique. Il régule un large éventail de fonctions transcriptionnelles telles que l'activation, la répression, et la pause de la transcription. Le caractère multifonctionnel de CTCF pourrait être dicté par sa capacité à façonner l'architecture de la chromatine. Cependant, une autre hypothèse stipule que c'est l'association avec différents partenaires protéiques qui serait à la base du caractère multifonctionnel de CTCF. Nos résultats de spectroscopie de masse sont en accord avec cette hypothèse et révèlent que CTCF interagit avec une multitude de protéines. De par leurs fonctions, les partenaires de CTCF se catégorisent au sein de quatre groupes; le remodelage de la chromatine, les facteurs de transcription, la réparation des dommages à l'ADN et les facteurs de maturation de l'ARN. En utilisant des méthodes de fractionnement biochimique, nous avons pu confirmer que certains complexes de CTCF s'assemblent sur l'ADN. Nous avons identifié le facteur général de transcription TFII-­‐I comme nouveau partenaire de CTCF. Par immunoprécipitation de la chromatine suivie de séquençage (ChIP-­‐seq) nous avons montré que TFII-­‐I est essentiel pour le ciblage de CTCF aux promoteurs proximaux des gènes cibles à l'échelle du génome particulièrement ceux des gènes impliqués dans le métabolisme. Au niveau des gènes régulés par CTCF et TFII-­‐I, la déplétion de TFII-­‐I conduit à une diminution de la liaison de CTCF et CDK8 et à une atténuation de la phosphorylation de l’ARN Pol II à la sérine 5. L'absence de TFII-­‐I altère la réponse cellulaire au stress métabolique. Nos résultats indiquent que TFII.I joue un rôle dans la liaison de CTCF aux promoteurs des gènes cibles et les deux facteurs coopèrent pour le recrutement de CDK8 pour amorcer l'initiation de la transcription. iv Acknowledgements_____________ ______________________________________________ First, I would like to express my sincere appreciation and gratitude to Dr. Michael Witcher for his continuous support and guidance throughout the duration of this project. During this time, he was as great mentor and helped me to improve my technical and scientific rational skills. He was very accessible in the moments that I need to. I appreciate all the encouragement to pursue all of my ideas and future career plans. I am also grateful for Dr. Witcher’s lab members, in particular with Dr. Maud Marques, for all her insights and advices in the development of this project, as well as with the rest of the members, Khalid Hilmi for his invaluable advices and Tiejun Zhao for all his assistance I want to extent my gratitude to my thesis committee members for all their assistance, suggestions and advices. Finally, I am very thankful for the support of my family and friends, their kind words of encouragement and support that they always provided me when needed. v 1. Introduction_________________________________________________________________ Cancers are often associated with both an environmental and genetic component. Recent knowledge on the impact of the genetic component in promoting human cancer has grown considerably and we have begun to understand that genetics contributes to a fraction of disease pathology, but other factors are also equally as important. Soon it came clear that epigenetics may contribute in a potent manner, similar to genetic mechanisms, to the development of cancer (Virani, Colacino et al. 2012). Epigenetic alteration refers to heritable changes in gene expression that occur without changes in DNA sequence, and influence gene expression (Berger, Kouzarides et al. 2009). It remains to be seen what portion of epigenetic marks are meiotically stable, however epigenetic marks are at least mitotically stable. One of the first epigenetic marks discovered was DNA methylation in 1948 (Hotchkiss 1948). By 1969 it was implied that DNA demethylation might have a biological role in long term memory in the brain (Griffith and Mahler 1969) and later in gene expression, leading to cellular differentiation (Taylor and Jones 1979, Beck and Rakyan 2008). Besides DNA methylation, histone modification and regulation of gene expression by microRNAs are known as key components of epigenetic regulation (Dawson and Kouzarides 2012). The influence of epigenetic modification is most obviously observed on the process of transcription. Because transcription is fundamental for homeostasis and survival, it is of great importance to understand how the dynamic of epigenetic modifications modulates the transcriptional output. In this study it will be explored how cooperation between transcription factors regulates the expression of a gene cohort, primarily involved in metabolism. We propose a model where cooperation of the epigenetic regulator CTCF and transcriptional factor TFII-­‐I links epigenetic processes directly to gene regulation. 1 DNA TRANSCRIPTIONAL REGULATION DNA stores the genetic information necessary for all cells to function normally. This includes programs for processes such as growth and differentiation. This information is inscribed into mRNA by a large molecular machine called the RNA polymerase (RNA Pol). Particularly, RNA Pol II is responsible for transcribing all the protein-­‐coding genes (Fuda, Ardehali et al. 2009), whereas RNA Pol I transcribes ribosomal RNAs (rRNAs) and RNA Pol III transcribes small RNAs such as transfer RNAs (tRNAs) (Clancy 2008). Transcription is a complex process that consists of multiple stages that direct the recruitment and processing activity of RNA Pol II. The six steps of transcription are; 1. Preinitiation; 2. Initiation; 3. Pausing; 4. Elongation; 5. Termination and 6. Reinitiation. (Danko, Hah et al. 2013). Transcription Preinitiation Gene expression is a dynamic process and is regulated by the sequences around a specific gene promoter and these sequences help to dictate, when, where and at what level specific genes are to be transcribed. The three most important DNA regulatory regions are the core promoter, promoter proximal regions (often bound by transcription factors such as Sp1), and distal enhancer sequences (Fuda, Ardehali et al. 2009). Formation of the pre-­‐initiation complex (PIC), is the earliest stage of transcription, and this complex is composed of general transcription factors (GTF, which are regulatory proteins whose function is to activate transcription of DNA by binding to specific DNA sequences) and RNA Pol II. Studies in vitro have determined that there are five basal GTF needed for the initiation of transcription (TFIIB, TFIID, TFIIE, TFIIF and TFIIH) (Reese 2003), and their primary role is to position RNA Pol II on the promoter region, facilitating initial access of RNA pol II catalytic subunit to target sites (Barrero and Malik 2013). The first step in transcription initiation is the recognition of the core promoter region. Core promoter elements are comprised by TATA boxes, Inr and DPE DNA motifs, among others. TATA box is an A/T rich sequence located around -­‐25 to -­‐30 nucleotides upstream of the transcription start site and is recognized by the TATA-­‐binding protein (TBP), present in the TFIID 2 complex (Thomas and Chiang 2006). Inr (initiator) contains a pyrimidine-­‐rich sequence and it is recognized by the transcription associated factors 1 and 2 (TAF1/TAF2) subunits of TFIID; DPE (downstream promoter element), on the other hand, is located +28 to +34 nucleotides downstream transcription start site and its sequence can be identified by TAF6/9 subunits of TFIID (Thomas and Chiang 2006). Usually core promoter elements are recognized by TFIID, which is formed by at least 14 TAF (Burley and Roeder 1996, Green 2000). TBP, subunit of TFIID, was initially identified as an exclusive TATA binding protein. However, in mammalians, the majority of promoters lack of TATA-­‐box and it has been demonstrated that TBP can also bind to this TATA-­‐less promoters through a different combinations of TAFs (Burke and Kadonaga 1997, Thomas and Chiang 2006, Tora and Timmers 2010). TFIID has also been implicated as a “writer” and “reader” of the chromatin epigenetic program through the presence of an Histone Acetyl Transferase (HAT) domain and double bromodomain of TAF1 (Mizzen, Yang et al. 1996, Jacobson, Ladurner et al. 2000), as well as the presence of a PHD domain in TAF3 that is capable of binding methylated lysine 4 of histone 3 (van Ingen, van Schaik et al. 2008), suggesting a link in the recognition of promoter DNA sequences and chromatin modifications (Papai, Weil et al. 2011). Binding of TFIID to promoter region, causes a bend in the DNA and acts as anchor for the association of the remaining factors (Reese 2003), such as TFIIB, that stabilizes TFIID at the promoter region by contacting TFIID and sequences flanking the TATA box (Barrero and Malik 2013). TFIIB can also bind to an upstream and downstream recognition element (BRE), that is usually located close to TATA-­‐box and in TATA-­‐less promoter regions, providing additional contact points, enhancing TFIID binding to core promoter regions (Thomas and Chiang 2006). TFIIB, is involved in the recruitment of RNA Pol II-­‐TFIIF complex, that stabilizes the DNA-­‐
TBP-­‐TFIID complex and it also mediates the incorporation of TFIIE and TFIIH into the PIC (Fig. i) (Woychik and Hampsey 2002, Reese 2003). 3 Fig. i. Elements of the Pre-­‐Initiation Complex. Formation of the pre-­‐initiation complex is a multistep process that requires the sequential binding of different transcriptional factor. TFIID is involved in the initiation of this complex, where it recognizes the promoter region and binds to specific sequences, such TATA box through TBP subunit. Later on, other transcriptional factors binds such as IIF and IIB allowing a platform for the binding of RNA Pol II, thus forming and active transcriptional complex that depending on the context will be post-­‐translational modified allowing the transcriptional elongation. Promoter clearance is a process that depends on ATP energy, where the helicase from TFIIH unwinds the DNA and expanding the transcriptional bubble. The addition of nucleoside triphosphate (NTPs) allows the elongation complex to extent the RNA (Weaver 2004). The general transcription factor TFIIE comprises two different subunits (TFIIEα/β) and it’s primarily function is related to the formation of an early open complex structure associated to basal transcription states. It has been described that TFIIE has the ability to bind single stranded DNA, therefore it its speculated that it binds to the non-­‐template DNA strand, helping to stabilize TFIIB and to recruit TFIIH (Grünberg and Hahn 2013). On the other hand, TFIIH contains ten subunits, from which three of them have ATP-­‐
dependent enzymatic activities. Among these, CDK7 mediates phosphorylation of RNA Pol II carboxyl-­‐terminal-­‐domain (CTD); Rad3/XPD, and Ssl2/XPB have DNA helicase activity with opposite polarity, involved in the unwinding of the promoter region from the transcriptional bubble at the active centre of RNA Pol II, once engaged in transcription (Kim, Ebright et al. 2000, Woychik and Hampsey 2002, Gibbons, Brignole et al. 2012). Besides of its role in 4 transcription TFIIH, is also implicated in the DNA excision repair (NER) mechanism, where unwinds DNA in order to excise the damaged DNA strand (Grünberg and Hahn 2013). TFIIE and TFIIH play an important role in the transition from initiation to elongation of transcripts (Okamoto, Yamamoto et al. 1998), where they are required for ATP-­‐dependent formation of the open promoter complex and cooperate to supress promoter-­‐proximal stalling of RNA Pol II (Dvir, Conaway et al. 2001). Transcription Initiation After the PIC has been assembled, RNA Pol II needs to escape the promoter region. Promoter escape can be described as the process in which RNA Pol II breaks its contacts with promoter-­‐sequence elements and promoter bound factors and simultaneously tightens its grip on the nascent RNA (Saunders, Core et al. 2006). It has been shown that TFIIB plays a role regulating RNA Pol II escape by stabilizing the a short 4-­‐5 nucleotides of nascent RNA with RNA Pol II (Bushnell, Westover et al. 2004), however it is suggested that upon elongation, newly synthesised RNA starts to collide with TFIIB, provoking the dissociation between TFIIB and RNA Pol II, thus, favouring the escape from promoter region (Saunders, Core et al. 2006). One of the key factors that regulate transcription is the phosphorylation status of RNA Pol II. RNA Pol II, contains on its CTD tandem heptapeptide repeats of the consensus amino acid sequence Tyr-­‐Ser-­‐Pro-­‐Thr-­‐Ser-­‐Pro-­‐Ser (YSPTSPS) (Prelich 2002), which is target for different protein complexes during transcription. Mainly, RNA Pol II can be phosphorylated on Serine 2 (Ser2) or Serine 5 (Ser5) of the heptapeptide sequence, and the dynamic of these events will determine the status of transcription. During transcription initiation, the levels of Ser5 phosphorylation on RNA Pol II increase, but as transcription progress, there is a decrease of Ser5 phosphorylation and this mark is replaced by Ser2 phosphorylation on RNA Pol II, associating Ser2 levels with productive elongation (Komarnitsky, Cho et al. 2000, Boehm, Saunders et al. 2003). TFIIH has been involved in the RNA Pol II CTD phosphorylation. It has been shown that a subunit of TFIIH, Cyclin-­‐dependent kinase 7 (CDK7) is able to phosphorylate Ser5 (Prelich 2002), 5 as well as Cyclin-­‐dependent kinase 8 (CDK8), found in the general cofactor Mediator (Donner, Ebmeier et al. 2010). Mediator complex is an evolutionary conserved regulatory complex and it is composed by about of 30 distinct subunits (Takahashi, Parmely et al. 2011). Mediator is a global regulator of gene expression, and it can be considered as a general transcription factor, however it differs from other transcription factors, by the high degree of structural flexibility and variable subunit composition (Malik and Roeder 2010, Poss, Ebmeier et al. 2013). In humans, CDK8 seems to be associated with various forms of Mediator, but it has been proposed that up to a 30% of CDK8 is independent of mediator (Knuesel, Meyer et al. 2009). Biochemical studies have shown that CDK8 requires MED12 for its kinase activity towards RNA Pol II CTD and other targets, and it depends on MED13 to mediate the interaction between CDK8 and the rest of components of the Mediator complex (Galbraith, Donner et al. 2010, Nemet, Jelicic et al. 2014). Similarly, it is proposed that CDK8 can also phosphorylate H3 on chromatin, which is associated with transcriptional activation. H3 phosphorylation facilitates the access of trans-­‐
acting factors to their target DNA sequences, causing the de-­‐condensation or relaxation of chromatin (Thomson, Mahadevan et al. 1999, Hans and Dimitrov 2001). Initially CDK8 was associated with a negative role in transcriptional control by phosphorylation and inactivation of the cyclin H subunit of TFIIH (Akoulitchev, Chuikov et al. 2000), and also by disrupting Mediator-­‐RNA Pol II interactions (Knuesel, Meyer et al. 2009). Recently, evidence shows that CDK8 can directly regulate several transcriptional programs, including the p53 network, Wnt/β-­‐catenin pathway and serum response network (Galbraith, Donner et al. 2010). Particularly, in the p53 network it was shown that upon UV-­‐induced DNA damage, activation of p53 target genes such as p21, undergo a rapid activation of transcription of these genes, by an increased recruitment of CDK8, Cyclin C and MED12, during conditions of sustained promoter activity (Donner, Szostek et al. 2007, Donner, Ebmeier et al. 2010). In a similar way, it was shown on HCT116 colon cancer cells, that upon knockdown of CDK8, there is a downregulation in the expression of approximately 26 genes out of 29 that are 6 usually activated upon serum stimulation (Donner, Ebmeier et al. 2010). Knockdown of CDK8, does not have an effect on recruitment of total RNA Pol II, however, is associated with a decrease in Ser5 phosphorylation, a mark commonly associate with transcriptional initiation. Assays done to study genes that are activated upon serum stimulation, such as FOS (a transcriptional factor implicated as regulator of cell proliferation, differentiation, and transformation) and EGR-­‐1 (Early growth response protein 1), have determined that depletion of CDK8, shows a decrease on the production of pre-­‐mRNA of these genes, suggesting that it has an impact in RNA Pol II elongation efficiency (Donner, Ebmeier et al. 2010). CDK8 is also required for the regulation of gene expression by HIF1A, a transcription factor involved in gene activation upon hypoxic stress (Galbraith, Allen et al. 2013), whereas approximately 200 genes showed reduce expression upon CDK8 knockdown in colon cancer cell line HCT116 having a positive impact in RNA Pol II elongation, via the CDK8 dependant recruitment of the Super Elongation Complex (SEC) and positive transcriptional elongation factor-­‐b (pTEF-­‐b), through the interaction with HIF1A (Galbraith, Allen et al. 2013). Besides its role in transcription, there is also evidence that in colon cancer, CDK8 is associated with the regulation of Wnt/β-­‐catenin pathway (Nemet, Jelicic et al. 2014). CDK8 was found to be amplified in many of colon cancer (Firestein, Bass et al. 2008), therefore it was shown that overexpression of CDK8 has the ability of transform NIH3T3 cells, allowing to growth in an anchorage-­‐independent manner as well as tumor formation in animals, but when expression of CDK8 is suppressed, it has an effect in inhibition of proliferation in colon cancer cells that are characterized by β-­‐catenin hyperactivity (Firestein, Bass et al. 2008). CDK8 can overcome the repression of β-­‐catenin by the transcription factor E2F1 in Drosophila and mammalian cells, where CDK8 interacts and phosphorylates E2F1, affecting the ability of E2F1 to activate or repress transcriptional activity (Morris, Ji et al. 2008, Zhao, Ramos et al. 2013). Transcriptional Pausing Transcriptional pausing is a process that is marked by a short transcriptional activity of RNA Pol II. Initial observation of this process was described in Drosophila heat shock genes 7 (Hsp), where transcriptionally engaged polymerase accumulates just downstream of the Hsp promoters and is associated with a 20-­‐60 nucleotide long nascent RNA (Rasmussen and Lis 1993). Early work in 1990, demonstrated that RNA Pol II elongates inefficiently through promoter-­‐proximal region, displaying a strong tendency to terminate elongation within the first 100 nucleotides (Kephart, Marshall et al. 1992, Marshall and Price 1992). Experiments done using an inhibitor of transcription known as DRB (Sehgal, Darnell et al. 1976), showed that treatment with this inhibitor causes a production of short transcripts, suggesting that elongation by RNA Pol II is affected. From this studies it was shown that RNA Pol II paused state is mainly coordinated by association of DRB-­‐sensitivity-­‐inducing factor (DSIF; also known as SPT5–SPT4) and Negative Elongation Factor (NELF). DSIF binds directly to RNA Pol II through its Spt5 subunit (Yamaguchi, Wada et al. 1999, Narita, Yamaguchi et al. 2003), and it can have positive or negative effects during transcription (Gilchrist, Nechaev et al. 2008). On the other hand, it seems that NELF does not bind substantially to DSIF or RNA Pol II alone, but it does bind to the complex DSIF and RNA Pol II (Yamaguchi, Inukai et al. 2002), and it is suggested that the formation of this complex triggers RNA Pol II Pausing (Price 2000, Narita, Yamaguchi et al. 2003). It is also speculated that one mechanism by which these factors mediate the pausing state in the promoter proximal region is throughout the affinity that NELF has with nascent RNA, therefore, interaction between short transcripts and NELF can also stimulate this paused state (Fujinaga, Irwin et al. 2004). Analysis done with mammalian cell cultures provide evidence that once transcription is initiated, it does not necessarily produce a full length transcript (Fraser, Sehgal et al. 1978), indicating that RNA Pol II faces a barrier that does not allow it to extend fully. Recent studies highlight that distribution of RNA Pol II genome wide is not uniform. In this sense, new evidence shows that in mammalian promoter regions, a large fraction of genes display a RNA Pol II signal that is concentrated near to TSS, indicating that the polymerases recruited to these promoters are not efficiently released downstream into the gene body, suggesting a default paused state of RNA Pol II, where only short transcripts can be produced 8 and final elongation might take place only after the right signalling event (Guenther, Levine et al. 2007). Similarly, global nuclear run-­‐on sequencing (GRO-­‐seq) assays, that maps the position, amount, and orientation of transcriptionally engaged RNA polymerases genome-­‐wide; confirmed that in human primary lung fibroblasts, promoter associated RNA Pol II molecules are paused in most of the coding genes (approximately 30% of all genes) (Core, Waterfall et al. 2008). This paused estate can be resumed by the treatment with Sarkosyl, a detergent that is thought to remove pause-­‐inducing factors, confirming paused state of these promoters (Core, Waterfall et al. 2008). Pausing represents an additional regulatory step in the transcription cycle, where it has the ability to regulate gene expression. Distribution of RNA Pol II in a poised state has been associated with constitutively expressed genes, such as signalling and transcription factors. In this genes, paused stated can be linked to an extremely rapid and synchronous activation in the expression of this genes, where can provide a mechanism to tune these key genes in response to cellular and external regulatory cues (Adelman and Lis 2012). Transcriptional Elongation One of the key aspects of transcriptional elongation is the recruitment of the positive transcription elongation factor b (P-­‐TEFb), which is associated with the phosphorylation of DSIF-­‐
NELF complex (Fujinaga, Irwin et al. 2004), as well as phosphorylation of RNA Pol II Ser2 residue (Ramanathan, Rajpara et al. 2001). P-­‐TEFb is a heterodimer complex of Cyclin T (CYC-­‐T) and Cyclin Dependent Kinase (CDK9) (Price 2000), and it can be recruited by the Mediator complex in a subset of genes. It has been shown that the subunit of Mediator complex, MED26 can serve as a binding platform for super-­‐
elongation complexes (SEC) that contains P-­‐TEFb, thus, stimulating the active elongation process (Takahashi, Parmely et al. 2011). Recruitment of P-­‐TEFb can be achieved by other mechanisms that involved activators such as c-­‐MYC, that can interact in the CYC-­‐T subunit of P-­‐TEFb during transcriptional activation and can induce transcriptional elongation (Eberhardy and Farnham 2002). In a similar way, 9 there is evidence that REL-­‐A, a subunit of NF-­‐κB, also binds to CYC-­‐T, allowing the recruitment of P-­‐TEFb to TNF-­‐α target genes (Barboric, Nissen et al. 2001). P-­‐TEFb can also be targeted to most of the genes by the interaction with the coactivator BRD4, which has a bromodomain that recognize acetylated histone tails, such as H4K16. Upon recognition of acetylated histones, BRD4 can recognize the CYC-­‐T subunit of P-­‐TEFb (Zippo, Serafini et al. 2009), showing that P-­‐TEFb is necessary for an active transcription elongation. Escape from transcriptional pausing requires the phosphorylation of DSIF-­‐NELF complex. It has been suggested that P-­‐TEFb initially phosphorylates the SPT5 subunit of DSIF, stimulating its function as positive elongation factor (Kim and Sharp 2001, Yamada, Yamaguchi et al. 2006) and then phosphorylates NELF having the consequence of releasing NELF from the double stranded RNA-­‐DNA hybrid, favouring RNA Pol II elongation, however the exact order of these events remains to be proven (Fujinaga, Irwin et al. 2004, Peterlin and Price 2006). RNA Pol II phosphorylation of Ser2 is carried out by CDK9 subunit of P-­‐TEFb that creates a platform for binding of additional RNA-­‐processing factors that facilitates productive RNA synthesis (Peterlin and Price 2006, Adelman and Lis 2012, Hsin and Manley 2012). RNA Pol II phosphorylation and dephosphorylation is a dynamic process that is not only essential for the recruitment and assembly of transcriptional complexes but also controls transcription and mRNA processing during the cell cycle (Zhang, Kim et al. 2006). As transcriptional elongation takes place, RNA Pol II dephosphorylation is carried out by the action of two phosphatases, Fcp1 and Scp1. The kinetics of these two phosphatases is dependent on its target. It has been shown that Fcp1 has a higher affinity towards Ser2, whereas Scp1 has a higher affinity towards Ser5 (Hausmann and Shuman 2002, Yeo, Lin et al. 2003). The importance of this catalytic sensitivity is likely to play a role in timing of transcription and mRNA processing, as Ser5 dephosphorylation is associated with transcriptional elongation and Ser2 dephosphorylation is a mark of transcriptional termination and RNA Pol II recycling for the next round of transcription (Hsin and Manley 2012). Apart from P-­‐TEFb, there are other factors that are necessary for an efficient elongation. There is evidence that Elongin-­‐related proteins, such as ELL1, is necessary to increase the net 10 catalytic rate of RNA Pol II, by reducing the transient pauses in gene bodies (Luo, Lin et al. 2012, Kwak and Lis 2013). ELL1 is part of the SEC complex that is form by multiple subunits that can be changed dependent on the genomic context, giving different specificity to the sequences at which they regulate (Kwak and Lis 2013). As transcription is a dynamic process, its regulation will also depend in the presence of nucleosomes during active elongation. It has been previously shown that FACT (Facilitates Chromatin Transcription) helps to overcome nucleosome barriers (Orphanides, LeRoy et al. 1998) by disassembling an H2A-­‐H2B dimer from nucleosomes, allowing RNA Pol II to carry out transcription (Belotserkovskaya, Oh et al. 2003). FACT is form by two subunits, SPT16 and SSRP1. SPT16 can interact with histones H3 and H4 (Bortvin and Winston 1996) and can also help to increase the elongation rate of RNA Pol II (Endoh, Zhu et al. 2004). Polymerase-­‐associated factor complex (PAF) has a role in elongation, although it seems that does not contain any enzymatic activity, it is thought that it serves as a platform for recruitment of elongation factor complexes such as SEC and FACT, as well as histone modifying enzymes (Sims, Belotserkovskaya et al. 2004). Besides all of these factors, it is widely known that transcriptional elongation is tightly regulated through the physical barrier imposed by nucleosomes. Importantly, histone tail modification such as acetylation of lysine side chains have an impact “loosening” the DNA wrapped into the nucleosomes, increasing the accessibility of transcription machinery (Bintu, Ishibashi et al. 2012), and it has been suggested that histone modifications play an import role in transcription, as these modifications can cause changes to the kinetics of RNA Pol II as it goes though the nucleosomes (Kwak and Lis 2013). Recent studies suggest a model of transcription in which Pol II cannot mechanically detach the DNA from the histones. Instead, when faced with the nucleosomal barrier, the enzyme stops, backtracks, and advances only when the DNA spontaneously unwraps from the surface of the core nucleosome. Thus, the polymerase behaves as a ratchet that rectifies the spontaneous DNA wrapping/unwrapping fluctuations of the nucleosome (Hodges, Bintu et al. 2009). 11 Through use of optical tweezers to follow real-­‐time trajectories of individual Pol II complexes as they transcribe through nucleosomes containing modifications at their N-­‐terminal histone tails or at specific histone-­‐DNA contacts, it was found that dynamics of the polymerase (its pause density and durations) is uniquely controlled to different extents by the histone tails, histone-­‐DNA contacts, and the DNA sequence (Bintu, Ishibashi et al. 2012). Particularly, it was shown that histone tails act mainly as a gate for RNA Pol II to access the nucleosomal region and upon acetylation of key lysine residues of the histone tails, there is a decrease in the transcriptional barrier by allowing the binding of chromatin remodelling factors to the nucleosomes, having an impact in transcriptional elongation (Bintu, Ishibashi et al. 2012). Transcriptional Termination Transcription termination occurs when RNA Pol II ceases RNA synthesis and both RNA Pol II and the nascent RNA are released from the DNA template (Kuehner, Pearson et al. 2011). Termination of transcription in mammals can be achieved by different pathways depending on the RNA 3’-­‐end processing signals and termination factors that are present at the end of the gene, among these the most studied is the poly(A)-­‐dependent pathway (Kuehner, Pearson et al. 2011). The ploy(A)-­‐dependent termination process can be described as two distinct stages. The first stage is characterized by the transcription of the poly(A) site, followed by pausing of RNA Pol II transcription downstream of this site and cleavage of the nascent transcript. The second stage is associated with the cleavage of the polyadenylated site and degradation of downstream product from polyadenylated site (Kuehner, Pearson et al. 2011). It has been discovered that the Cleavage and Polyadenylation Specificity factor (CPSF) is recruited to the elongation complex by the interaction with RNA Pol II body. At the same time, there is a Cleavage Stimulatory Factor (CstF) that interacts with the RNA Pol II CTD. When the AAUAA sequence that CPSF recognizes is transcribed, CPSF induces pausing. After this, CstF binds to downstream GU-­‐rich processing signal and dislodges CPSF, leading to the cleavage of polyadenylated nascent RNA (Kuehner, Pearson et al. 2011). 12 Apart from this, it has been shown that nuclease XRN2 is recruited to the elongation complex by p54/PSF (Kaneko, Rozenblatt-­‐Rosen et al. 2007), a multifunctional protein dimer that binds to the RNA Pol II CTD (Rosonina and Blencowe 2004). XRN2 will degrade the downstream RNA that is tether to the RNA exit channel from RNA Pol II. Collision of XRN2 with Pol II would then promote termination (Kuehner, Pearson et al. 2011). Transcription Reinitiation Reinitiation of transcription is a process that varies enormously from one gene to the other, and even for the same gene under different conditions (Dieci and Sentenac 2003). Transcription initiation rates depend on how fast the PIC is assembled and is also influenced by the combination of activators and repressors of transcription. One important mechanism by which activators stimulate transcription is by the efficiency of template reutilization for multiple transcription cycles through the stabilization of preinitiation intermediates (Hahn 1998, Dieci and Sentenac 2003). It is suggested that at least some components of the PIC can persist at the promoter during multiple transcription cycles (Dieci and Sentenac 2003). In this sense, there is evidence that RNA Pol II reinitiation intermediate includes transcription factors such as TFIID, TFIIA, TFIIH, TFIIE and Mediator, that can be recognized by RNA Pol II and whose stabilization by transcriptional activators could be a key mechanism in gene activation (Dieci, Fermi et al. 2014). Reinitiation, can bypass some rate-­‐limiting transition steps such as the binding of TFIID, having a higher rate than transcription from initiation, thus facilitating high levels of transcription (Liu, Yuan et al. 2014). EPIGENETIC REGULATION Transcription programs are different in every cell type, dictating cell identity, but these programs are highly regulated by interplay between signal transduction pathways, transcription factors and chromatin packaging. However, transcription programs of differentiated cells retain certain plasticity to ensure appropriate responses to the cellular environment. 13 The role of the epigenetic landscape in transcriptional regulation is multi-­‐faceted and encompasses nucleosome density, positioning, and recruitment of chromatin remodelling complexes and transcription factors (Lee and Ge 2014). Here we will discuss the two most commonly studied epigenetic modifications, post-­‐translational covalent modification of the N-­‐
terminal tail of core histones and DNA methylation. DNA Methylation DNA methylation refers to the addition of a methyl group (-­‐CH3) covalently attached to the base cytosine in the dinucleotide 5’-­‐CpG-­‐3’ (Lim and Maher 2010). Methylation patterns in normal cells occurs predominantly in repetitive genomic regions, including satellite DNA, long interspersed transposable elements (LINES) and short interspersed transposable elements (SINES) (Robertson 2005). In contrast, unmethylated CpGs are not randomly distributed, but are clustered in CpG islands, which are regions near promoter areas of many genes (Bird 1986). Mechanistically, the methylation machinery has two distinct components, the DNA Methyltransferases (DNMTs) responsible for the methylation, and the readers of the methylated marks known as Methyl CpG binding proteins (MBDs) (Robertson 2005). DNMTs mediate the transfer of a methyl group from the S-­‐adenosylmethionine (SAM) to cytosines. There are three enzymatically active enzymes DNMT1, DNMT3a and DNMT3b. DNMT1 is known as a maintenance methyltransferase that preserves methylation patterns during cell division and it preferentially methylates hemimethylated CG dinucleotides. On the other hand DNMT3 enzymes are responsible for de novo methylation during embryonic development (Denis, Ndlovu et al. 2011). Catalytic activity between DNMT3a and DNMT3b seems to be influenced by sequences next to the target CG sites, suggesting a sequence preference by de novo DNMTs in human genomic methylation patterns (Handa and Jeltsch 2005). Typically, DNA methylation represses transcription directly, by inhibiting the binding of specific transcriptional factors, and indirectly, due to the recruitment of MBDs and their associated repressive chromatin remodelling factors (Robertson 2005). 14 The importance of DNA methylation is emphasized by the growing number of human diseases that are known to occurs when the DNA methylation pattern is not properly established or maintained (Robertson 2005). Particularly in cancer, the genome of cancer cells become hypomethylated, due to the loss of methylation in repetitive regions (Yoder, Walsh et al. 1997), resulting in genomic instability (Rizwana and Hahn 1999). Alternatively, aberrant hypermethylation in cancer cells usually occurs at CpG islands near promoter regions of genes involved in cell-­‐cycle regulation, tumor cell invasion, DNA repair and chromatin remodelling, among others. The overall consequence of the hypermethylated phenotype will be changes in chromatin structure, such as histone hypoacetylation, that will produce transcriptional silencing (Robertson 2005). Aberrant DNA methylation patterns are associated with an overexpression of DNMT enzymes in cancer; particularly it was shown that in breast cancer DNMT3b is highly overexpressed in comparison with DNMT1 and DNMT3a (Girault, Tozlu et al. 2003, Veeck and Esteller 2010). However, there is evidence that supports another layer of regulation by controlling the expression levels through microRNAs, where in lung cancer cell lines, miR-­‐29 can target DNMT3a/b and enforced expression of miR-­‐29s restores the normal methylation patterns and inhibits tumorigenicity in vitro and in vivo (Fabbri, Garzon et al. 2007), evidencing the importance of proper DNA methylation profiles for cellular homeostasis. Histone Modifications It is widely known that DNA is compacted in the nucleus through wrapping around nucleosomes. Nucleosomes are comprised of 146 bp DNA segments that are wrapped around 8 core histones. Nucleosomes represent a major barrier to transcription (Petesch and Lis 2012). Typically histone tails are covalently modified on their N-­‐terminal (Kouzarides 2007) tail by the addition of a plethora of post-­‐translational modifications such as methyl and acetyl groups, phosphorylation, ubiquitylation, sumoylation among others (Berger 2007, Kouzarides 2007). Such diverse modifications could be expected to have diverse effects, and indeed this is the case. Modification may impact the affinity of DNA towards the histones, allowing either tighter or loosen association with the histones, hence influencing the availability of DNA for 15 transcription activation or repression. N-­‐terminal modification may also serve as docking sites for many factors that modulate transcription. Among the most studied modifications, methylation and acetylation have been shown to robustly regulate transcription. Histone Acetylation Histone acetylation is a mark that is commonly found at active genes and the level of acetylation tends to be higher in the 5’ regulatory regions (Liang, Lin et al. 2004, Pokholok, Harbison et al. 2005). Acetylation is carried out by histone acetyltransferases (HATs) enzymes that can be grouped in three principal families, GNAT, MYST and CBP/p300 (Sterner and Berger 2000). HATs catalyzes the acetylation of core histones through the addition of an acetyl group from the pseudo-­‐substrate acetyl coenzyme A to the lysine residues on the N-­‐terminal of histones (Schneider, Chatterjee et al. 2013). Amongst the acetylation marks that are associated with transcriptional activation is H3K9ac, which is catalyzed by the Gcn5 enzyme, a member of the GNAT family. Gcn5 contains a bromodomain that binds acetyl-­‐lysine and it has been shown that is a subunit of the chromatin-­‐
associated transcriptional complex SAGA (Spt-­‐Ada-­‐Gcn5 acetyltransferase) (Lee, Sardiu et al. 2011, Cieniewicz, Moreland et al. 2014). Studies have shown that the bromodomain of Gcn5, a functional motif that is involved in the recognition of acetylated lysines (Zeng and Zhou 2001), is essential for the HAT activity, as point mutations in this region decrease histone acetylation at promoter regions in vivo, affecting transcriptional rates, as well as a global decrease in acetylation level of nucleosomes (Syntichaki, Topalidou et al. 2000). Besides H3K9ac, acetylation of H3K18 and H3K27 is enriched in areas close to the TSS, suggesting their role in transcription activation (Wang, Zang et al. 2008). H3K9ac and H3K27 have an antagonistic role towards methylation of this sites, where when methylated these two residues have a potent role in transcription repression (Kouzarides 2007). Histone acetylation can also function as a platform for different transcriptional factors and chromatin remodelers (Fischle, Wang et al. 2003). In this context, chromatin remodelers such as SWI/SNF complex, possess bromodomains, that in vivo are important for the maintenance of transcriptional active chromatin regions (Hassan, Prochasson et al. 2002). 16 The SWI/SNF complex is evolutionary conserved and is composed for at least nine proteins including an invariant core complex (SNF5) and variable subunits that contribute to transcriptional activation or repression (Roberts and Orkin 2004). When this complex is recruited to chromatin, it hydrolyzes ATP and uses this energy to remodel nucleosomes (Reisman, Glaros et al. 2009). The energy for the SWI/SNF chromatin remodelling function is carried out by the catalytic subunit, BRG1 or BRM, where both have DNA-­‐dependant ATPase activity, allowing the hydrolysis of ATP (Vignali, Hassan et al. 2000). It is believed that nucleosome remodelling can occur by sliding of histone octamer along the same strand of DNA or by transferring complete histone octamers to adjacent regions of exposed DNA (Roberts and Orkin 2004). Antagonistic to HAT, there are enzymes associated with the removal of acetylated residues, known as histone deaceytalases (HDACs). Many HDACs form a part of multi-­‐protein complexes such as the transcriptional co-­‐repressor Sin3, N-­‐COR and SMRT (Glass and Rosenfeld 2000), which are target to specific genomic regions by interactions with DNA associated proteins such as transcriptional factors and other epigenetic modifiers (Ropero and Esteller 2007). HDACs repress transcription by various mechanisms (Kouzarides 2007). Deacetylation of histones causes a change in the net charge of nucleosomes, strengthening histone tail-­‐DNA interactions and blocking the access for the transcriptional machinery to the DNA template (Gallinari, Marco et al. 2007). Besides this, it has been observed that deacetylation allows further modification by repressive histone methyltransferases (Martin and Zhang 2005). Further, the methyl binding protein MeCP2 is able to recruit HDAC-­‐containing complexes to methylated gene promoters as a mechanism of repression (Jones, Jan Veenstra et al. 1998, Ropero and Esteller 2007). Histone Methylation Histone methylation is another mark that has in impact in transcription. N-­‐terminal residues of histones can be mono-­‐, di-­‐, or tri-­‐methylated; and depending on the precise lysine being modified, can show either a transcriptional activation or repression (Saunders, Core et al. 17 2006). Among histone methylation marks, modification on Histone 3 has been extensively studied, particularly on lysine residues 4, 9, 27, 36 and 79. H4 lysine 20 has also been well investigated and these modifications can impact chromatin structure, and transcriptional regulation as well as DNA damage response (Zhang, Eugeni et al. 2003, Shi and Whetstine 2007). Histone methylation is carried out by Histone Methyltransferases (HMTs) and this enzyme utilizes S-­‐adenosyl-­‐methionine as the methyl donor group (Bannister, Schneider et al. 2002). Lysine methy-­‐transferases are specific compared to acetyltransfereases and they usually modified one single lysine on a single histone (Bannister and Kouzarides 2005). Transcriptional activation through methylation can be mediated by the modification of H3K4, H3K36 and H3K79. H3K4me3 is a mark whose abundance is greater in the 5’ end of genes and correlates strongly with activation of transcription (Schneider, Bannister et al. 2004, Pokholok, Harbison et al. 2005). Importantly, it has been shown that H3K4me3, can be recognized by TAF3, a submit of TFIID, promoting transcriptional activation by stimulating PIC formation (van Ingen, van Schaik et al. 2008). H3K36me3 is found to accumulate at the 3’end of active genes and is associated with RNA Pol II Ser2 phosphorylation and transcriptional elongation (Joshi and Struhl 2005, Kouzarides 2007). The function of methylation at H3K79 is not well known, however it has been suggested that can be involved in the activation and elongation of the HOXA9 gene (Kouzarides 2007), and that H3K79me3 is enriched near the transcription start site (Vakoc, Sachdeva et al. 2006). There is evidence that several complexes have the ability to bind H3K4me3 and participate in gene induction. Among these, SAGA complex, that contains Gnc5, associates with the chromodomain protein and chromatin remodeler CHD1. CHD1 binds to methylated lysine 4 residue of histone 3, and allows the interaction with SAGA complex leading to histone acetylation, however it remains to be seen which is the exact mechanism CHD1 mediates histone acetylation (Pray-­‐Grant, Daniel et al. 2005, Sims, Chen et al. 2005, Berger 2007). More recently, it has been demonstrated that the SGF29 (a subunit of the SAGA complex) binds to H3K4me3 and plays a role in the ER stress response. Its been suggested that SGF29 coordinates H3K4me3 levels and maintains a ‘poised’ chromatin state on ER stress target 18 gene promoters. Following ER stress induction, SGF29 is required for increased H3K14 acetylation on these genes, which then results in full transcriptional activation, thereby promoting cell survival (Schram, Baas et al. 2013). Besides this complex, documented evidence shows that other proteins can interact with the H3K4me3 mark, such as the plant homeodomain (PHD) finger that helps to recruit positive active enzymatic complexes. H3K4me3 stabilizes binding of the nucleosome remodelling factor (NURF) to target genes and allows the binding of the bromodomain PHD transcription factor (BPTF), regulating the transcriptional status of target HOX locus (Wysocka, Swigut et al. 2006). Interestingly, in some contexts, H3K4me3 may also be associated with transcriptional repression. In response to DNA damage, the SIN3-­‐HDAC1 complex binds to the H3K4me3 mark through the PHD domain of the inhibitor of growth family member 2 (Ing2), which stimulates the HDAC1 activity leading to repression of genes involved in proliferation to modulate cellular response to genotoxic damage (Shi, Hong et al. 2006), showing a dual function of this mark, and the flexibility of the histone code. However, there are marks such as H3K9me, H3K27me and H4K20me that are strictly associated with transcriptional repression. Particularly, H3K9me is associated with heterochromatin formation. There is evidence that repression involves recruitment of methylating enzymes SUV39H1 and the heterochromatin protein 1 (HP1) that binds specifically to H3 N-­‐terminal tails when lysine residue 9 is methylated. HP1 is a protein capable of heterodimerization and associates with many other proteins such as HDACs (Muchardt, Guillemé et al. 2002, Yamamoto and Sonoda 2003). H3K27me3 has been implicated in the silencing of Hox gene expression and X chromosome inactivation during imprinting (Plath, Fang et al. 2003, Kouzarides 2007), but it has also been shown that can have a bivalent function, as in a mono-­‐methylated state this mark is enriched at actively transcribed promoters, whereas the tri-­‐methylated mark is associated with silenced promoters (Barski, Cuddapah et al. 2007). H3K27me3 is a mark that has is established by the Polycomb Repressive Complex 2 (PRC2), which is formed by EZH2, involved in the methylation of H3K27. Once this mark has 19 been placed the Polycomb Repressive Complex 1 (PRC1) can recognize it and forms a repressive chromatin domain (Aoto, Saitoh et al. 2008) All of these modifications are important as regulators of transcription, however, as has been described earlier, transcriptional factors help to determine in which context a gene or a subset of genes are going to be expressed. GENERAL TRANSCRIPTION FACTOR II-­‐I (TFII-­‐I) TFII-­‐I was discovered in 1991 as a basal transcriptional factor that has the ability to bind to core promoter elements such as pyrimidine-­‐rich initiator (Inr) element sequences, as well as upstream regulatory elements (E-­‐box) in the promoters of the adenovirus major late (AdML) and human immunodeficiency virus-­‐1 (HIV-­‐1) in vitro. The capacity to bind to these two elements, gives to TFII-­‐I a dual characteristic to function as basal factor and as activator that can interact with complexes assembled at upstream regulatory sites (Roy, Meisterernst et al. 1991). Recent data from Chromatin Immunoprecipitation (ChIP) suggest that TFII-­‐I genomic localization is mostly intergenic rather than intragenic, and TFII-­‐I binds upstream of the transcription start site of expressed genes, whereas in silenced genes it can bind both upstream and downstream of the transcription start site in some stress responsive genes, such as ATAF3 in human erythroleukemia cell line K562 (Fan, Papadopoulos et al. 2014). It has been shown that GTF2i is frequently deleted in the Williams-­‐Beuren syndrome, a neurodevelopmental disorder characterized by a complex phenotype that includes mental deficiency, premature aging of skin, supravalvular aortic stenosis, dental and craniofacial malformations (Bayés, Magano et al. 2003), showing the importance of TFII-­‐I during developmental stages. TFII-­‐I has several alternative spliced isoforms in human and mice. The original sequence identified for TFII-­‐I codes for a 957 amino acid form known as TFII-­‐IΔ. TFII-­‐Iα (977 amino acids) is encoded by exon A, TFII-­‐Iβ (978 amino acids) is encoded by exon B and TFII-­‐Iɣ (998 amino acids) arises by the combination of exon A and B (Cheriyath and Roy 2001). Isoforms of TFII-­‐I are differently expressed in tissues. TFII-­‐Iβ is expressed at a much higher rate in murine cells 20 than in human cells, on the other hand, TFII-­‐Iα isoforms appears to be lacking in murine cells and there is evidence that TFII-­‐Iɣ is expressed predominantly in the neuronal cells (Pérez Jurado, Wang et al. 1998). Each of these isoforms contains the same structural characteristics as TFII-­‐IΔ (here after referred as TFII-­‐I). TFII-­‐I is characterized by the presence of six direct I-­‐repeats (R1-­‐R6), comprised by a highly conserved 90 residues repeats, each containing a helix-­‐loop-­‐helix domain, implicated in distinct protein-­‐protein interactions. TFII-­‐I also possess a basic region (BR) sequence, before R2 repeat, that is necessary for DNA binding and a Leucine Zipper, for homo-­‐ or hetero-­‐dimerization (Fig. ii) (Roy, Du et al. 1997, Roy 2012). Fig. ii. TFII-­‐I structure and isoforms. TFII-­‐I is characterized by the presence of 4 distinct isoforms product of alternative splicing of exon A and B. All isoforms of TFII-­‐I are characterized by the presence of sis I-­‐
Repeats (R1-­‐6) with a helix-­‐loop-­‐helix domain, implicated in protein interactions, a nuclear localization signal (NLS) and a basic region (BR) involved in DNA binding and a Leucine Zipper (LZ) that is used for dimerization (Roy 2012). So far, it is unclear if there is another region of TFII-­‐I that is necessary or implicated to DNA binding. Experiments done by deleting 90 amino acids from the N-­‐terminal, which contains a leucine zipper motif, led to a loss of binding of TFII-­‐I to the Vβ and c-­‐fos promoter region, even though this mutant has an intact BR region. Interestingly, deletion of the N-­‐terminal region also leads to a failure in homomeric dimerization with the wild-­‐type protein, suggesting 21 that the leucine zipper present in this area mediates the homo-­‐ or heteromeric interaction required for DNA binding (Cheriyath and Roy 2001). The ability of TFII-­‐I to works as a transcriptional factor depends on the cellular signalling pathways. The role of TFII-­‐I in cellular signalling cascades is mainly accomplished by the phosphorylation of serine/threonine and tyrosine residues (Novina 1998, Cheriyath, Desgranges et al. 2002). Under basal conditions TFII-­‐I is phosphorylated in these residues, but upon activation of signalling pathways, such as growth factors, TGFβ signalling and ER stress, TFII-­‐I becomes hyper-­‐phosphorylated (Novina 1998). Conversely, when tyrosine residues are dephosphorylated, the capacity of TFII-­‐I to bind regulatory elements, remains unchanged, but transcriptional activity becomes affected (Novina 1998). It has been shown that a variety of growth-­‐promoting and mitogenic signals such as epidermal growth factor (EGF), platelet-­‐derived growth factor (PDGF) and serum can enhance TFII-­‐I tyrosine phosphorylation. Studies have determine that transcriptional activity of TFII-­‐I requires phosphorylation of tyrosine residues, particularly in the residue 248 (Y248) (Cheriyath, Desgranges et al. 2002). Upon growth factor stimulation, it has been shown that TFII-­‐I can be phosphorylated by the Src kinase family (Roy 2012). In B cells, there is evidence that TFII-­‐I phosphorylation is associated with Bruton’s tyrosine kinase (BTK), here TFII-­‐I is tyrosine-­‐phosphorylated by Bruton’s tyrosine kinase in vitro, and upon immunoglobulin receptor cross-­‐linking in B cells it is released from Bruton’s tyrosine kinase and is translocated to the nucleus, showing that TFII-­‐I is a downstream effector of several signal transduction pathways mediating signal-­‐responsive activator complexes to general transcriptional machinery (Cheriyath and Roy 2000). There is also evidence that supports the role of TFII-­‐I as a downstream effector in Ras and Rho pathways. Interestingly, TFII-­‐I seems to interact with p190RhoGAP protein, a potent Rho inhibitor. Upon stimulation with PDGF, TFII-­‐I becomes phosphorylated and interaction with p190RhoGAP is disrupted, leading to an enhance transcription of serum-­‐inducible genes including c-­‐fos (Jiang, Sordella et al. 2005). Although tyrosine phosphorylation of TFII-­‐I is required for its basal transcription function, it still remains unknown the precise mechanism by which phosphorylation exerts its 22 functions. Evidence suggest that DNA binding seems to be unaffected by phosphorylation, thus it is likely that the protein-­‐protein interactions of TFII-­‐I with the basal machinery may be dependent upon its phosphorylation status. It has also been considered that phosphorylation of TFII-­‐I is required for its nuclear translocation, however TFII-­‐I mutants that do not phosphorylate are translocated to the nucleus upon ectopic over-­‐expression (Novina 1998). Ectopic over-­‐expression of TFII-­‐I isoforms provide evidence that all of them localize preferentially in the nucleus and exhibit similar transcriptional activity (Cheriyath and Roy 2000). However, TFII-­‐IΔ shows a cytoplasmic localization, whereas TFII-­‐Iβ is localized in the nucleus, but upon positive extracellular signals, TFII-­‐IΔ, is imported to the nucleus and TFII-­‐Iβ is exported from nucleus to the cytoplasm, leading to activation of transcription in the c-­‐fos promoter (Hakre 2006). This differential distribution of endogenous isoforms in various cell suggest distinct and non-­‐redundant functional roles in vivo (Cheriyath and Roy 2000, Hakre 2006). Besides relaying signal transduction signals into a transcriptional output, TFII-­‐I Δ has also been related with endoplasmic reticulum stress (ER-­‐stress) regulation (Parker, Phan et al. 2001). Upon induction of ER-­‐stress, TFII-­‐I is recruited to the promoter of the response elements Grp78, which is part of a family of stress-­‐induced chaperones (Parker, Phan et al. 2001). ER-­‐stress triggers TFII-­‐I phosphorylation and as result is translocated to the nucleus and subsequently recruited to the promoter site. Activation of ER chaperones, such as Grp78 activates a general pathway that triggers unfolded protein response, inducing cell death as well as pro-­‐survival mechanism (Hong, Lin et al. 2005). In addition to TFII-­‐I transcriptional activity, TFII-­‐I might be implicated in the regulation of Ca2+ influx (Caraveo, Rossum et al. 2006). Ca2+ is an important ion for the maintenance of homeostasis and it affects processes such as transcription and cell proliferation, among others (Clapham 2007). Activation of growth factor and immune receptors result in activation of PLC-­‐ɣ (phospholipase C) and temporary intracellular influx of Ca2+ via transient receptor potential channel 3 (TRPC3). TFII-­‐I interacts directly with PLC-­‐ɣ thorough the interaction of Src-­‐homology domains of PLC-­‐ɣ, which is necessary for the binding with TRPC3 and calcium influx. Upon PLC-­‐ɣ 23 and TFII-­‐I association, PLC-­‐ɣ is unable to bind to the TRCP3 receptors reducing the influx of calcium (Caraveo, Rossum et al. 2006). Apart from the interactions described above, there is evidence that support the role of TFII-­‐I in cell cycle progression. Particularly, it has been shown by Desgranges et al, that TFII-­‐I is recruited to the Cyclin D1 promoter in vivo and activates the transcription of this gene, resulting in accelerated entry and exit into S phase. Upon genotoxic stress TFII-­‐I is marked for proteosomal degradation mediated by activation of p53 ATM (Ataxia telangiectasia mutated), leading to cell cycle arrest (Desgranges, Ahn et al. 2005). Based on these examples, TFII-­‐I functions serve as link that connects extracellular signals with intracellular environment. Genomic distribution of TFII-­‐I and epigenetic role The TFII-­‐I DNA binding domain is located directly preceding repeat domain 2, where there are some basic residues (amino acids 301–306), which function as a DNA binding domain. Deletion of these residues abrogates the binding to Inr/Inr-­‐like elements at the Vβ and c-­‐fos promoters (Roy 2012). Several studies have also tried to determine the consensus DNA binding sequence of TFII-­‐I, however the precise DNA binding motif remains controversial and seems to be highly variable. It is believed that TFII-­‐I functions as a transcriptional activator as well as a repressor, likely in a promoter context dependent and partner interaction dependent, stressing the versatility of this protein (Roy 2012). Gemonic studies, particulary ChIP-­‐seq analysis on mouse embryonic stem (ES) cells, have identified that binding of TFII-­‐I and BEN (member of the helix-­‐loop-­‐helix protein family) are predominantly located at either promoter or enhancer regions. The prominence of TFII-­‐I at enhancer regions indicate the possibility of long-­‐range enhancer-­‐promoter interactions as regulatory mechanism whereby TFII-­‐I controls transcription (Makeyev, Enkhmandakh et al. 2012). It was shown in another study that TFII-­‐I could regulate transcription by itself or through the interaction with BEN. This two proteins have the capacity to bind to the same site or to different sites, adding a differential regulatory mechanism that can have different transcriptional outcomes (Bayarsaihan, Makeyev et al. 2012). 24 By ChIP in mouse stem cells, TFII-­‐I and BEN bind to a significant amount of genes related to highly cooperative cellular functions including signal transduction (TFII-­‐I), cell-­‐cell signalling and cytoskeleton organization (BEN) as well as cell fate commitment (TFII-­‐I and BEN). However, it was also found that TFII-­‐I binding to promoter or regulatory regions has a different pattern in embryonic stem cells (ESC) compared to embryonic craniofacial tissues (ET) (Makeyev, Enkhmandakh et al. 2012). TFII-­‐I binding to promoter regions in ESC depends also on the presence of BEN. In ET, promoter-­‐bound BEN allows the de novo recruitment of TFII-­‐I, showing the cooperativity of these two factors in the regulation of transcription and underscores a role in cell fate commitment (Makeyev, Enkhmandakh et al. 2012). Besides TFII-­‐I role in cellular differentiation, it has been implicated in epigenetic regulatory mechanisms. Expression analysis of TFII-­‐I in null mice embryos, show that TFII-­‐I is responsible for maintaining the expression of histone-­‐modifying enzymes including EZH2 and EED, both are core components of the PRC2. In a similar way, the H3K4me3 demethylation gene Aof2 and H3K36me3 methyltransferase gene Nsd, are regulated by TFII-­‐I. Altered expression of these genes, has as consequence the downregulation of cellular homeostasis, potentially explaining the embryonic lethality of TFII-­‐I knockout embryos (Enkhmandakh, Makeyev et al. 2009). Recent studies show that TFII-­‐I binds a multitude of silenced genes across the genome enriched with the repressive histone mark H3K27me3. Conversely, it is also highly enriched at euchromatic regions marked with H3K4me3, often at proximal transcription start sites (Fan, Papadopoulos et al. 2014). It was also reported that it can interact with components of the SWI/SNF chromatin remodelling complex, such as Brg-­‐1, showing a potential role in epigenetic regulation (Fan, Papadopoulos et al. 2014). The presence of TFII-­‐I at both active and silenced genes may suggest an unknown role of TFII-­‐I as a platform that can recruit different elements that may influence gene activation and repression. Importantly, the association of TFII-­‐I and Brg-­‐1 suggest that is able to enhance gene activation by remodelling chromatin states and conversely, association of TFII-­‐I in regions 25 enriched with H3K27me3, may indicate that has an influence in the recruitment of HDAC that will enhance gene repression (Fan, Papadopoulos et al. 2014). It its suggested that TFII-­‐I can recruit HDAC3 to target promoters, helping to modulate the transcriptional activity of TFII-­‐I, however is not fully understood how this interaction plays a role in transcription and what are the signals that trigger the recruitment of HDACs by TFII-­‐I, and the impact that TFII-­‐I has as positive/negative regulator of transcription (Wen, Cress et al. 2003). TFII-­‐I may also play a role in RNA Pol II paused state. ChIP-­‐seq experiments shown that TFII-­‐I binding is present in genes where RNA Pol II is paused, such as genes related to stress response elements (Adelman and Lis 2012). Particularly, in the promoter of the EFR3A gene, it is suggested that TFII-­‐I interacts with Elongin A, a positive activator of transcriptional elongation (Conaway and Conaway 1999), and these two proteins could recruit Elonging B and C, enhancing ubiquitination signal in RNA Pol II, producing the degradation and removal of RNA Pol II, thus in EFR3A gene, TFII-­‐I could have a negative impact in the regulation of this gene (Fan, Papadopoulos et al. 2014). Overall, TFII-­‐I functions both as a transcriptional activator as well as a repressor, where cytoplasmic signals can play an important role determining the outcome in transcription as well as the interaction with partner proteins. CCCTC-­‐BINDING FACTOR (CTCF) CTCF was first described by Baniahmad et al. in 1990 as a transcriptional repressor-­‐
silencer of Myc and lysozyme genes in chicken (Baniahmad, Steiner et al. 1990). A few years later, CTCF gene was localized to a region that is frequently deleted in breast and prostate cancer (Filippova, Lindblom et al. 1998). CTCF is a highly conserved multifunctional protein and it is considered to be a master weaver present in all multicellular organism involved in diverse processes such as gene activation (Klenova, Nicolas et al. 1993, Filippova, Fagerlie et al. 1996), gene repression (Lobanenkov, Nicolas et al. 1990), enhancer blocking (Bell, West et al. 1999, Hark, Schoenherr 26 et al. 2000) RNA Pol II pausing and recently in chromatin domain formation and maintenance as well as in 3D organization of the genome (Recillas-­‐Targa, de la Rosa-­‐Velázquez et al. 2011). One of the most remarkable features of CTCF is its structure, characterized by the presence of eleven zinc-­‐finger domains (Fig. iii) (Ohlsson, Renkawitz et al. 2001). Given the wide range of CTCF functions, it is thought that CTCF uses a combination of zinc-­‐fingers that allows it to bind to a great variety of sequences and at the same time allows it to interact with many proteins (Filippova, Fagerlie et al. 1996). Genome-­‐wide data sets have enabled the identification of an 11-­‐15bp core consensus binding sequence that is consistent in all cell types assed by independent studies (Phillips and Corces 2009). Early studies have reported the association of many more fingers with an extended 50–60 bp sequence (Ohlsson, Renkawitz et al. 2001). By ChIP-­‐seq, analyses it was showed CTCF uses distinct groups of contiguous zinc fingers creating different binding subdomains that are able to recognize a core DNA binding sequence. Interestingly, it was also shown that mutations in peripheral zinc fingers decrease CTCF binding, thus indicating that adjacent zinc fingers modulate CTCF binding (Nakahashi, Kwon et al. 2013). CTCF consensus binding sequence contains CpG and can be subject to DNA methylation. It has been shown that CTCF binds preferentially unmethylated DNA sequences as it has been reported in the H19 insulator of the Igf2 locus (Bell and Felsenfeld 2000, Witcher and Emerson 2009, Holwerda and de Laat 2013). Initially CTCF was discovered by its ability to bind in the promoter-­‐proximal regulatory region of the chicken c-­‐Myc gene (Klenova, Nicolas et al. 1993), as well as to the upstream region from the Amyloid β-­‐Protein precursor transcription start site (Vostrov and Quitschke 1997). Since these initial observations, it has been shown that CTCF binds across all the genome. Genome wide studies have revealed that CTCF is able to bind in approximately at 15000 sites through the genome in IMR90 human fibroblast, where its global distribution pattern was reported in intergenic regions (46%), intronic (22%), exonic (12%) and in whithin 2.5Kb of promoters (20%) (Barski, Cuddapah et al. 2007, Kim, Abdullaev et al. 2007). In another study it 27 has been shown by ChIP-­‐seq that CTCF can be found in approximately 20000 sites in CD4+ T cells (Barski, Cuddapah et al. 2007). More recently, by ChIP-­‐seq it has been shown that in mouse embryonic stem cells, CTCF can be found in 40000 target sites genome wide(Chen, Xu et al. 2008). However, is not clear whether these cell type-­‐specific differences in occupancy are functionally significant or merely due different experimental approaches (Phillips and Corces 2009), regardless, is evident that CTCF has the ability to bind to thousands of sites, highlighting the importance in chromatin organization. Fig. iii. Representation of CTCF structure. CTCF is a conserved protein that is characterized by presence of 11 Zinc-­‐fingers (ZF) in its core structure. Binding to DNA sequences can be achieved by a different combination of ZFs, originating a diversity of target sequences (Klenova, Morse et al. 2002). To better understand CTCF functions, it is important to describe how chromatin is organized in the nucleus. Genomes of higher eukaryotes are packaged into several hierarchical levels of organization. DNA is wrapped around histones to form the 10 nm nucleosomal fiber, which is folded and looped into sophisticated higher-­‐order structures (Phillips and Corces 2009). 28 The arrangement of chromatin within the nucleus is not random; instead de-­‐condensed chromosomes occupy areas defined as chromosome territories, which are dynamic compartments where chromosomes are able to interact with each other (Ohlsson, Lobanenkov et al. 2010). Chromosomes are organized in radial position in the nucleus, where gene-­‐dense chromosomes are located towards the centre of the nucleus (Ohlsson, Lobanenkov et al. 2010). In these regions, enhancers (regulatory DNA sequences that, when bound by specific transcription factors, enhance the transcription of an associated gene), insulators (DNA sequence elements that prevent inappropriate interactions between adjacent chromatin domains) and promoters are in contact with one another when genes are being transcribed (Wallace and Felsenfeld 2007). Active genes are transcribed while associated with RNA Pol II transcription factories containing many Pol II molecules. A given factory is visited by many genes, with the effect of bringing individual genes that are widely separated on a chromosome, or even situated on different chromosomes, into proximity. The formation of loop domains appears to be a necessary process associated with transcription in eukaryotes (Wallace and Felsenfeld 2007). It has been shown that CTCF can interact with itself and with other proteins that may allow it to form clusters, which, as in the case of Gypsy insulator from Drosophila would lead to the formation of discrete domains (Yusufzai, Tagami et al. 2004). Recent data confirm that CTCF molecules bound to distant sites, even on different chromosomes, can interact with one another in vivo (Ling, Li et al. 2006). CTCF Role in Genome Organization CTCF has been implicated in diverse roles in gene regulation, including context-­‐
dependent promoter activation/repression, enhancer blocking or barrier insulation, as well as genomic imprinting and long-­‐range chromosomal interactions (Phillips and Corces 2009). CTCF and Insulator activity Chromatin insulators are commonly defined as DNA elements that prevent inappropriate interactions between neighbouring genes. Insulators are classified into enhancer 29 blockers, that prevent enhancer promoter interactions if placed between these elements, thus repressing transcription; and barrier elements, that prevent spreading of heterochromatin (a repressive chromatin environment) into neighbouring active domains (Ohlsson R. 2010). The ability of CTCF to act as an insulator has been one of the most studied and intriguing characteristics of this protein to date, due to the fact that CTCF is the only known protein in vertebrates to mediate this function (Dunn and Davie 2003). The role of CTCF as insulator suggests that acts to prevent the action of enhancers or repressors on distal gene promoters. This role is consistent with reports that CTCF binding can be detected near regions of genes that are transcriptionally coregulated (Kim, Abdullaev et al. 2007). One of the first evidences of CTCF as an enhancer blocker comes from the imprinted control region (ICR) H19 of the Igf2 locus. Regulation at this site requires the binding of CTCF in a differentially methylated region of H19. CTCF binds to the unmethylated site in the maternal allele, hence acting as a barrier between Igf2 and its enhancer, preventing the transcription of Igf2 (Bell and Felsenfeld 2000). Methylation present in the paternally inherited ICR disrupts CTCF binding, allowing the enhancer to potentiate Igf2 transcription. Interestingly, mutation on the H19 region in the maternal allele led to loss of CTCF binding and an increase in DNA methylation in mice, having as consequence the biallelic expression of Igf2 (Engel, Thorvaldsen et al. 2006). Various studies where CTCF binding sites have been mutated reveal that CTCF is critical to prevent the spread of DNA methylation. Particularly, in the promoter region of the Retinoblastoma gene, absence of CTCF binding is able to contribute to a more rapid and extensive DNA methylation pattern and a robust epigenetic silencing in transgenes in glioma cell lines (Dávalos-­‐Salas, Furlan-­‐Magaril et al. 2011), highlighting the role of CTCF in chromatin boundary functions. Role of CTCF in Chromatin Boundaries By computational approaches using 12 mammalian genomes and ChIP experiments, it was shown that CTCF localization is not randomly distributed throughout the genome, instead, 30 it tends to be distributed close to the regions where genes are located across the whole genome (Kim, Abdullaev et al. 2007, Xie, Mikkelsen et al. 2007). Mapping of CTCF binding sites shows that a majority of DNA fragments where CTCF binds are located in within intergenic regions, far from gene promoters (Kim, Abdullaev et al. 2007, Soshnikova, Montavon et al. 2010). Such target sites were proposed to function as insulators, preventing the undesirable activation of genes by tissue specific enhancers located at their vicinity, and evidencing CTCF role in the maintenance of chromatin boundaries (Soshnikova, Montavon et al. 2010). However, It has been shown that approximately 7% of CTCF binding sites are located within a one kb window from the TSS (Soshnikova, Montavon et al. 2010), suggesting a role of CTCF in gene activation. CTCF participates in gene regulation by maintaining chromatin boundaries (Cuddapah, Jothi et al. 2009, Witcher and Emerson 2009). Chromatin boundaries are region across the genome that marks active and repressive chromatin states. Repressive chromatin is the default state and if it is not contained it will spread passively throughout a chromosome (Talbert and Henikoff 2006). Genomic analysis evidence that there is proportion of CTCF binding sites that demarcate boundaries between repressive an active chromatin, where it has been shown that CTCF binding correlates with enrichment or repressive marks such as H3K27me3 and activating mark H2AK5Ac (Cuddapah, Jothi et al. 2009). Similarly it has also been shown that there is a correlation between the presence of euchromatic regions with CTCF binding sites, stressing the role of CTCF in the maintenance of chromatin boundaries (Wen, Wu et al. 2012). Role of CTCF in Chromatin loop formation It is believed that CTCF can perform most of its functions by the formation of dimers or through the interaction with other proteins and partners that facilitate the formation of chromatin loops (Pant, Kurukuti et al. 2004, Yusufzai, Tagami et al. 2004). Through the interaction of CTCF with nucleophosmin, Yusufzai et al, was able to provide one of the first evidences that show the capacity of CTCF to interact with nuclear matrix, where it was demonstrated that CTCF-­‐nucleophosmin complexes co-­‐localize at the nucleolar surface, 31 creating a loop domain structure in the chicken β-­‐globin locus. The formation of this structure has an impact in chromatin structure that prevents enhancer-­‐promoter interactions, blocking the action of the distal enhancer (Magdinier, Yusufzai et al. 2004, Yusufzai, Tagami et al. 2004). This is in keeping with the insulator function of CTCF (Bell and Felsenfeld 2000). It remains to be seen if all CTCF insulator sites are tethered to the nucleolus. Other studies have shown that CTCF homodimerization may be another possible mechanism to organize chromatin domains creating active loops that may facilitate the activation of genes (Pant, Kurukuti et al. 2004). CTCF has also been implicated in mediation of long range chromosomal interactions, where it has proposed that CTCF forms a complex with cohesin, which will create a contact between chromatin fibers in cis, as seen at the Ifng locus. It is believed that cohesion brings proximal CTCF binding sites together, providing sufficient affinity between these sites to stabilize and immobilize chromatin fibres (Hadjur, Williams et al. 2009). CTCF Post-­‐Translational Modifications Perhaps, one of the most important aspects of CTCF in the regulation of gene expression is its post-­‐translational modifications. CTCF functions are modulated by both phosphorylation and by poly-­‐ADP-­‐ribosylation (PARylation), as well as SUMOlyation, of these, PARylation is the most studied modification due to its wide range of functions. CTCF phosphorylation occurs in the carboxyl terminus through the action of the enzyme casein kinase II (CK2). Phosphorylation is associated with a change from a transcriptionally repressive function for CTCF to an activating one in a chicken c-­‐myc reporter gene (El-­‐Kady and Klenova 2005). There is evidence that CTCF phosphorylation is related to control of cell growth. When serine residues (Serine-­‐612) undergoing phosphorylation are mutated, a negative effect on cell growth is observed, even though binding of CTCF and nuclear localization is not affected by phosphorylation (Klenova, Chernukhin et al. 2001). SUMOylation, is the addition of small ubiquitin-­‐like modifications, carried out by SUMO proteins. SUMO modification of a protein does not target it for degradation, instead, it can play roles in transcriptional regulation, chromatin structure and DNA repair; but at the same time it is usually associated with repression of gene expression, genomic imprinting and insulator 32 function (MacPherson, Beatty et al. 2009). SUMOylation of CTCF occurs in the C-­‐terminal and N-­‐
terminal domain and is effect is associated with organization of repressive chromatin domains on the c-­‐myc promoter (MacPherson, Beatty et al. 2009, Kitchen and Schoenherr 2010). PARylation is a covalent modification catalyzed by the poly(ADP-­‐ribose) polymerase enzyme (PARP) (Kim, Zhang et al. 2005). This modification consist in the addition of ADP-­‐ribose units that form a chain that can containing up to 200 ADP ribose units, that are preferentially incorporated in glutamic acid, aspartic acid and lysine residues (Boulikas 1989). One of the important features of this modification is that can be reversed by the action of the poly(ADP-­‐
ribose) glycohydrolase enzyme (PARG) (Hassa, Haenni et al. 2006). Particularly, it has been demonstrated that there are six distinct residues of CTCF N-­‐terminal domain that can be PARylated (Farrar, Rai et al. 2010, Zhang, Wang et al. 2013). PARP-­‐1 is the founding member of the PARP family, which contains about 18 distinct proteins in humans, and all of its members share a highly conserved catalytic domain. In addition, PARP enzymes contain different domains that could include zinc fingers, BRCA1 C-­‐
terminus like, and macro domains (Kim, Zhang et al. 2005). Addition of PAR to proteins may alter protein activity by functioning as a site-­‐specific covalent modification, a protein-­‐binding matrix or a steric block (Kim, Zhang et al. 2005). In vivo, PAR is involved in specific interactions with a variety of effector proteins, and by proteomic analysis it was identified a macro domain that is composed by a 190 amino acid domain found in a wide variety of proteins, that functions as an ADP-­‐ribose-­‐binding module (Karras, Kustatscher et al. 2005). Recent studies have demonstrated that macro domains have the potential to orchestrate various chromatin-­‐based biological tasks, including DNA repair, chromatin remodelling, transcription, cell death and insulator functions among others (Kim, Zhang et al. 2005, Han, Li et al. 2011). Particularly it has been shown that CTCF can be PARylated in the N-­‐
terminal domain (Yu, Ginjala et al. 2004), and it has been shown that many of the processes that involve CTCF (Wallace and Felsenfeld 2007), can be modulated by this mark. 33 Relationship between CTCF functions and PARylation The effect that PARylation has on CTCF it has been the most studied. CTCF PARylation was first associated in the mouse H19 imprinting control region, where it can function as an insulator (Yu, Ginjala et al. 2004). In this sense, one of the first evidence of CTCF and enhancer blocking insulation was proposed on the 5’HAS insulator sequence upstream of the chicken β-­‐globin locus (Bell, West et al. 1999). Later it was shown that CTCF has also a role in the imprinted control region H19 of the Igf2 locus (Bell and Felsenfeld 2000). Importantly, in the H19 region, it has been shown that by using 3-­‐aminobenzamine, an inhibitor of PARP activity, compromised CTCF function as insulator in this locus (Yu, Ginjala et al. 2004). In a similar way, it has been shown that by using another PARP1 inhibitor (PJ34) and a CTCF mutant that does not undergo PARylation, it compromises CTCF activity in the p19ARF promoter-­‐driven reporter gene, suggesting that CTCF function as transcriptional regulator is dependent on PARylation (Farrar, Rai et al. 2010). It has been hypothesised that PARylation of CTCF plays an important role in breast cancer. Analysis of breast cancer cell lines, tumors and normal breast tissues, suggest that a majority of breast tumors harbour low levels of CTCF in a PARylated form. It has been suggested a link with breast cancer progression, proposing that loss of CTCF PARylation contributes to the progression of this disease (Docquier, Kita et al. 2009). It can be speculated that in these tumors PARP activity is low or that there might be a hyper activity of PARG, thus influencing low levels of PARylation that could affect different protein-­‐protein interactions. Studies done in vitro and in vivo indicate that CTCF can induce PARP-­‐1 activity and can lead to its PARylation causing the inhibition of DNA methyltrasnferase (DNMT1) and contributing to an increase in the hypomethylation profile genome wide (Guastafierro, Cecchinelli et al. 2008). It is believed that in cancer, loss of PARylation of CTCF can perturb DMNT1 inhibition and cause an aberrant DNA hypermethylation over a particular sub-­‐set of the CTCF target genes (Recillas-­‐Targa, de la Rosa-­‐Velázquez et al. 2011). PARylation of CTCF can function as a dock that may allow interaction of CTCF with other protein partners. 34 CTCF and Epigenetic Regulation Of Transcription Besides playing a role in the maintenance of chromatin boundaries, one of the most important functions of CTCF is maintaining activation of gene transcription. By genome wide analysis, it has been shown that between 7% to 15% of the CTCF binding sites in the human genome are located within two Kbp from transcription start sites (Barski, Cuddapah et al. 2007, Soshnikova, Montavon et al. 2010). The association of CTCF with regulatory elements, such as enhancer and insulators, suggest its role in transcriptional regulation (Barski, Cuddapah et al. 2007). Several studies have demonstrated the ability of CTCF to regulate the expression of genes. Among these, it has been shown that CTCF can modulate the gene expression of tumor suppressor genes such as BRAC1, RB, p53, CDKN2A (Butcher and Rodenhiser 2007, De La Rosa-­‐
Velázquez, Rincón-­‐Arano et al. 2007, Witcher and Emerson 2009, Soto-­‐Reyes and Recillas-­‐Targa 2010). CTCF is located in a genomic region that is frequently deleted in breast and prostate tumors causing loss of heterozygosity (LOH) affecting CTCF integrity and as previously described (Filippova, Lindblom et al. 1998), however it seems that the role of CTCF, at least in breast cancer is not related to somatic or germ line mutations. In breast cancer, studies showed that the role of CTCF as positive activator of transcription might be accomplished by its ability to act as regulator of boundary or barrier elements. Particularly, in the BRCA1 locus, there are two CTCF binding sites described and it has been demonstrated that along with the interaction of SP1 (special protein 1) can create a boundary between methylated and unmethylated DNA upstream of the transcriptional start site (TSS) of BRCA1, preventing the spread of methylated DNA in the core promoter of this gene (Butcher, Mancini-­‐DiNardo et al. 2004). By analyzing sporadic breast tumor samples, it was found that CTCF localization in a subset of samples shifted from nuclear to cytoplasmic and as consequence the promoter region of BRCA1 becomes hypermethylated (Butcher and Rodenhiser 2007). Apart from the role of CTCF in the regulation of BRCA1, CTCF has the ability to bind to the promoter region of the RB gene. Interestingly, it was shown by De La Rosa-­‐Velázquez, that 35 upon altering the CTCF binding sequence in vitro in the promoter region of RB by point mutations, causes an epigenetic silencing of a reporter gene randomly integrated in the genome, however when the binding sequence remains unchanged there is not a decrease in the expression of the reporter gene. At the same time, it was also described that the binding of CTCF is sensitive to the methylation status of its target sequence (De La Rosa-­‐Velázquez, Rincón-­‐Arano et al. 2007). It was also shown that in the CTCF binding region there is an overlapping site for the methyl-­‐CpG-­‐
binding protein Kaiso and upon loss of CTCF and binding of Kaiso, the latter can recruit co-­‐
repressors like N-­‐CoR that has the ability to attract chromatin enzymes like histone methylases and deacetylases having as end result the silencing of this gene (Yoon, Chan et al. 2003, Recillas-­‐Targa, de la Rosa-­‐Velázquez et al. 2011). An alternative explanation for the silencing of RB has been proposed based on the fact that Kasiso can interact in vitro with the C-­‐terminal portion of CTCF (Defossez, Kelly et al. 2005), suggesting that such association might stimulate or block chromatin loop formation or induce alternative loops that could act as a protective mechanism against abnormal DNA methylation spread over regulatory elements (De La Rosa-­‐Velázquez, Rincón-­‐Arano et al. 2007). On the other hand, CTCF has also been implicated in the regulation of p53, whereas in the upstream region from the TSS, it was shown enrichment in the incorporation of repressive chromatin marks such as H3K9me3 and H3K27me3 upon reduced levels of CTCF, but not an increase of DNA methylation, supporting the role of CTCF in gene expression regulation at different levels, protecting not only from DNA methylation but also from the incorporation of repressive marks (Soto-­‐Reyes and Recillas-­‐Targa 2010). In the same way, it was also found that CTCF is involved in the regulation of the p16INK4a gene and loss of CTCF binding correlates with silencing of p16. Particularly, the loss of CTCF is correlated with a defective state of CTCF PARylation that has as consequence a destabilization of chromosomal boundaries and changing the epigenetic landscape by the loss of the histone variant H2A.Z associated with active transcriptional genes, thus promoting the silencing of this gene as well as RASSF1A and CDH1 (Witcher and Emerson 2009). 36 CTCF and Transcription Besides its epigenetic role and regulation of transcription, there is evidence that suggests that CTCF can interact with RNA Pol II, thus affecting in a direct way the transcriptional process (Chernukhin, Shamsuddin et al. 2007). Particularly, it was shown that a subpopulation of CTCF is able to interact with the large subunit of RNA Pol II in vivo and in vitro. This interaction is sufficient to activate the transcription, where transfection of a vector containing a CTCF binding site fused with a luciferase reporter gene and its mutant version incapable to bind CTCF, demonstrated that CTCF and RNA Pol II cooperate to activate transcription of the reporter gene, as luciferase levels were significantly higher in wild-­‐type binding site, compared to mutant. In the same report, it was also shown that CTCF has the ability to recruit RNA Pol II to a particular subset of CTCF target genes (Chernukhin, Shamsuddin et al. 2007). CTCF can also promote inclusion of exons by mediating local RNA Pol II pausing in a mammalian model system for alternative splicing of CD, here CTCF binding to CD45 exon 5 is inhibited by DNA methylation, leading to reciprocal effects on exon 5 inclusion (Shukla, Kavak et al. 2011). Other evidence that supports the role of CTCF with direct regulation of transcription is its association with transcription factors. Among these binding partners is the zinc finger transcription factor Yy1, that has been implicated in X-­‐chromosome inactivation process (Donohoe, Zhang et al. 2007). Studies suggest that there is a strong pattern of colocalization between these two factors at predicted boundary elements, suggesting that they could act synergistically in delimiting chromatin domains (Wang, Lunyak et al. 2011, Schwalie, Ward et al. 2013). YB-­‐1 is another transcription factor that is associated in regulation of DNA replication, transcription and RNA processing (Chernukhin, Shamsuddin et al. 2007). YB-­‐1 and CTCF are able to modulate the differential expression in the serotonin transporter gene. CTCF will interact with YB-­‐1 to form a protein complex throughout the DNA-­‐binding domain of YB-­‐1. CTCF interactions abolish the ability of YB-­‐1 to bind to its consensus sequence, having an impact in the expression of the gene (Klenova, Scott et al. 2004). 37 It has been reported that the Upstream Binding Factor (UBF), interacts with CTCF. UBF is an abundant nucleolar protein that is involved in DNA binding. UBF binds dynamically throughout the rDNA repeats (regions that codify for rRNAs clustered in long tandem repeats on one or a few chromosomes) (Jantzen, Admon et al. 1990), and not only plays a role as a transcriptional activator of RNA Pol I, but also in transcription elongation (Stefanovsky, Langlois et al. 2006). CTCF binds immediately upstream of the ribosomal spacer promoter in a methylation sensitive manner, and activates spacer promoter transcription. CTCF binding controls the loading of UBF onto rDNA, and the binding of RNA polymerase I and H2A.Z near the spacer promoter, hence influencing transcription (van de Nobelen, Rosa-­‐Garrido et al. 2010). Altogether, CTCF is multifunctional protein that is involved in a diverse range of transcriptional and epigenetic functions. Here we will discuss a novel example of how CTCF can be targeted to promoter regions, suggesting a new role of How CTCF can influence transcription in eukaryotes organism. 38 2. Aim________________________________________________________________________ Transcription is a dynamic process that is tightly regulated by diverse mechanisms. Genome wide alterations to transcription are ubiquitous in cancer and transcription is commonly disrupted in other diseases as well. In cancer, disruption of the epigenetic landscape contributes to the aberrant transcription profile observed in cancer. CTCF is a key epigenetic regulator that has diverse functions. It is speculated that the accessory proteins bound to CTCF allow it carry out such disparate roles. However this has not been proven. We hypothesize that CTCF cooperates with transcription factors to regulate target genes. Genome wide analysis of CTCF binding revealed that flanking sequences influence CTCF binding, again supporting the notion that CTCF partners dictate its function and even its binding. There is little knowledge about the identity of proteins that can target CTCF to distinct sequences. Therefore, the objective of this study is to answer questions such as what factors cooperate with CTCF to direct its binding? Does CTCF cooperate with transcription factors to activate transcription? To achieve these goals, proteomic and genomic tools will be used to define CTCF interacting partners, promoter regions bound by CTCF, and the mechanisms whereby CTCF can directly modulate gene expression. 39 3. Materials and Methods___ __________________________________________________ Cell Culture and growth curves. WEHI-­‐231 control and WEHI-­‐231 TFII-­‐I KD cell lines were a generous gift of Ananda Roy, and were cultured as previously described (Ashworth and Roy 2007). MDA-­‐MB-­‐435 and HCT-­‐116 cells were cultured in RPMI and DMEM respectively, supplemented with 10% FBS. (Balakrishnan, Witcher et al. 2012). For proliferation assays, 2x105 WEHI-­‐231 control and KD cells were cultured in 2.5ml of DMEM with 25mM (high) or 5mM (low) glucose, +/-­‐ glutamine and 5% FBS. Cells were counted at 24h and 48h and cell viability was evaluated by Trypan blue exclusion assay. Generation of CTCF KDs. shRNA against mouse CTCF was purchased from Sigma (SHCLNG-­‐NM_181322). Briefly, virus was packaged in Hek293T cells. 5.5x106 cultured cells were transfected with 5µg of packaging vector Pax2, 2µg of envelope vector MD2G and 7µg of shRNA against CTCF using 42µg of PEI (1mg/ml). The mix of PEI and plasmid DNA was prepared in DMEM without FBS and incubated at room temperature for 15 minutes. After this time, the transfection mix was added to the cells and viruses were collected 72h after transfection. For infection of CTCF shRNA viral supernatants into WEHI-­‐231 cells, 0.3 x106 cells/ml were seeded in 6 well dishes and 500µL of virus were added along with 8µg of hexadimethrine bromide (Polybrene 4mg/ml). 24h post initial infection, a second round of infection was carried out. After 72h of initial infection, cells were selected with 0.5µg of Puromycin for 48h and then collected. Antibodies. Primary antibodies for Co-­‐IPs and ChIPs used were rabbit polyclonal anti-­‐
CTCF (Millipore), mouse anti-­‐TFII-­‐I (BD), rabbit polyclonal anti-­‐H2A.Z (Millipore), rabbit polyclonal anti-­‐RNA Poll II (Santa Cruz), mouse monoclonal anti-­‐RNA Pol II phosphor-­‐Ser5 (H14, Covance), rabbit polyclonal anti-­‐H3K24me3, rabbit polyclonal anti-­‐H3K27ac and goat polyclonal anti-­‐CDK8 (Santa Cruz). For Western Blot, rabbit monoclonal anti-­‐NBS1 (Millipore). Mouse monoclonal anti-­‐Brg1 (G7), mouse monoclonal anti-­‐MRE11 (18), goat polyclonal anti-­‐hnRNPD, rabbit polyclonal anti-­‐Sp1, mouse monoclonal nucleolin (c23) and mouse monoclonal anti B23 40 Nucleophosmin were from Santa Cruz biotechnology. Rabbit polyclonal parp1 and parp 2 were from Cell Signalling technology. Cell Cycle and Flow Cytometry Analysis. Briefly, cells cultured under high and low glucose conditions were washed in 1X PBS with 3%FBS and spun down for 5 min at 1750 rpm. Cells were fixed in ethanol at 75% in 1X PBS and stored at 4°C. Before propidium iodide staining, cells were centrifuged to remove fixation media and washed in 1X PBS with 3%FBS followed by overnight staining. Before FACs analysis cells were resuspended in 300µL of fresh propidium iodide and analyzed in BD FACS Calibur. For apoptosis analysis, Annexin V-­‐CF633 (Biotium) staining protocol was followed as per the manufacturer’s instructions and analysis was carried out using FlowJo software. Mass Spectrometry and Protein Sequencing. MDA-­‐MB-­‐435 whole cell extracts spiked with 1µg recombinant CTCF were subject to immunoprecipitation with rabbit polyclonal anti-­‐
CTCF. Immunoprecipitated proteins were resolved by SDS-­‐PAGE and visualized with coomassie blue staining. Coomassie blue stained protein bands were excised and subjected to disulfide reduction and alkylation. Trypsin-­‐digested samples were injected onto nano-­‐HPLC system with a C18 capillary column. Peptides were eluted with a linear gradient from 5 to 45% acetonitrile and effluent was electro-­‐sprayed into the LTQ mass spectrometer. The mass spectrometer was operated in data dependent mode to automatically switch between survey MS and MS/MS acquisitions. The conventional MS spectra (survey scan) were acquired in the Orbitrap at a resolution of 60,000 followed by three MS/MS scanning, peptide charged over 10,000 were kept for sequencing. MS/MS spectra were searched against a IPI human database (IPI version 3.24 november 2008) using Mascot (version 2.3.2, Matrix Science); the threshold of parental and ionic fragment were respectively set at +/-­‐ 0.4 and 0.1 Da. Manual inspection of all MS/MS spectra for modified peptides was performed to validate assignments. Peptides with a Mascot inferior to 100 were excluded from further analysis 41 Column Chromatography. All procedures were performed at 4 °C and all chromatography matrices were purchased from (GE Healthcare Bio-­‐Sciences). Flow rate was 0.5 ml/minute except for DNA Cellulose, which was set at 0.1 ml/minute. HeLa whole cell extracts were sonicated and diluted 10 fold in binding buffer (25 mM HEPES, pH 7.6, 50 mM KCl, 10% Glycerol, 12.5 mM MgCl2, and 1 mM DTT). Phospho-­‐cellulose p11 column (cation exchange) was prepared according to the manufacturer's instructions and equilibrated with 2X column volume with binding buffer. The diluted cell extract was applied to column and washes were performed with 2X column volumes of binding buffer. The retained proteins were eluted with increasing concentrations of KCL. The eluted fractions were combined and salt concentration and pH were adjusted before loading to Q-­‐Sepharose (anion exchange). After washes, a gradient of linearly increasing salt concentration was applied to elute the sample components from the column. The eluted protein fractions were pooled and loaded on DNA cellulose column (native double stranded calf -­‐thymus DNA cellulose). DNA cellulose column was prewashed before use in 20 mM Tris-­‐HCl ph8.0, 50 mM KCl, 10% glycerol, 1mM MgCl2, 1μM ZnCl2 and 1mM DTT. After washing the column extensively, the bound proteins were eluted with the above buffer containing increasing concentrations of KCL. Confocal microscopy. MDA-­‐MB-­‐435 cell monolayers grown on glass coverslips in 6-­‐well plates were washed twice with 1X PBS and fixed at room temperature (RT) in 1% PFA containing 0.25% Triton X-­‐100 for 15 min. The cells were then washed twice with PBS and blocked in blocking buffer (1X PBS, 1% BSA, 2% normal goat serum) for 60min. Antibodies targeting CTCF (BD, 1:50), TFII-­‐I (cell signaling, 1:200) were then added directly to the blocking buffer and incubated overnight. The cells were then washed three times in 1X PBS and incubated in blocking buffer containing respective Alexa-­‐conjugated secondary antibodies (Invitrogen, 1:1000) and incubated for 1 hour at RT. The cells were then washed three times in 1X PBS and mounted on slides with vectashield containing DAPI. Images were captured using a 60X oil immersion objective on the Wave FX spinning disk confocal microscopy system (Quorum technologies) and analyzed using Volocity®. Intensity scatter plot and corresponding Pearson correlation coefficient for CTCF and TFII-­‐I double-­‐stained cells were generated using Volocity®. 42 Western Blot. Cell were lysed with Whole Cell Lysis buffer (20mM Tris pH 7.5, 420 mM NaCl, 2mM MgCl2, 1mM EDTA, 10% glycerol , 0.5% NP40, 0.5% Triton) supplemented with fresh 1mM DTT and PSMF and protease and phosphatase inhibitor cocktail, BGP, P8340 and NAF. Whole cell lysis buffer was added 2 times the volume of the cell pellet and left on ice for 25 min. Lysates were centrifuge at top speed at 4°C and supernatant was collected and transferred to a new tube. Protein concentration was measured by Bradford assay. 35µg of protein were loaded into 8% acrylamide gels and Transfer was done overnight at 30V at 4°C. Membranes were washed 3 times with TBST buffer (Tris base 20mM, NaCl 137mM, 0.1% Tween 20) for 5, 10 and 15 minutes. Membrane was blocked with 5% milk in TBST and incubated with primary antibody overnight at 4°C. After primary antibody incubation, membrane was washed as previously described and incubated with secondary antibody for 1h at room temperature. Western blot was revealed using Clarity Western Enhanced chemiluminescence kit (bio-­‐Rad). Gene Expression Analysis. Gene Elute Mammalian Total RNA kit (Sigma) was used for RNA extraction following manufactured instructions. 1µg at 50ng/µL of total WEHI-­‐231 cell RNA were send to Innovation Génome Québec for microarray analysis. Illumina Beads technology was used (MouseWG-­‐6_V2). Microarray analysis was performed using Bioconductor package “limma”(Smyth 2005). Validation of target genes identified was realized as followed: total RNA was extracted with Gene Elute Mamalian Total RNA kit following manufacturer's instructions. cDNA was generated by reverse transcription using 1µg of RNA. EasyScript kit from ABM was used following manufacturer's protocol. qPCR reactions were carried out using SYBERgreen dye from Promega with specific primers at 50µM concentration. Nascent mRNA was examined for CDKN2B/Arf after on column DNAse digestion and using primers spanning the first exon and intron. qPCR primer sequences are listed in Table S1. Microarray data sets are available at the Gene Expression Omnibus (GEO) website, www.ncbi.nlm.nih.gov/geo, under accession ID: GSE60915. 43 Co-­‐Immunoprecipitation. Cells were collected, washed with 1X PBS and lysed as described above. Protein concentration was quantified by Bradford assay and 1mg of protein was used to do the immunoprecipitation. Protein lysates were diluted 5X in IP buffer (20mM Tris pH7.5, 50mM NaCl, 10mM MgCl2, 2mM EDTA, 0.5% Triton X100) and PSMF was added to avoid protein degradation. Protocol for samples treated with Nucleases (DNAse and RNAse) was carried out as previously described (Zhang, Kuznetsov et al. 2008). Lysates were pre-­‐
cleared for 2h with protein G agarose beads. After the pre-­‐clearing stage, beads were pelleted, supernatant was collected and transferred to a new tube where antibodies were added and nutated overnight at 4°C. Antibodies were collected by adding fresh Protein G agarose beads and nutated at 4° for 2h. Beads were pelleted and the supernatant was discarded. The collected beads were washed 3 times by adding 1mL with IP buffer (100 mM NaCl) and centrifuged at 4000rpm to pellet the beads. The final wash was done with IP buffer containing 0.1% Triton-­‐X and spun down at 5000rpm. Proteins were eluted from the beads by adding 25µL of 2X SDS loading buffer (3M Tris pH 6.8, 20% SDS, 100% glycerol, H2O and bromophenol blue) and heated at 100°C for 10 minutes. Beads were again pelleted and the resulting supernatant was loaded to acrylamide gel and blotted as described above. Chromatin Immunoprecipitation (ChIP). WEHI-­‐231 Ctl and TFII-­‐I KD cells were collected and crosslinked with 1% folmaldehyde in PBS for 10 min at room temperature. The crosslinking reaction was stopped with 125 mM Glycine, the cells washed with 1X PBS, and stored at -­‐80°C until the assay was carried out. Cells were lysed and DNA sheered by sonication in cell lysis/ChIP buffer (0.25% NP-­‐40, 0.25% Trinton-­‐X100, 0.25% Sodium deoxycholate, 0.1% SDS, 50mM Tris pH 8.0, 50mM NaCl, 5mM EDTA), 15 times for 15 seconds each. Lysates were centrifuged for 15min at 14000 rpm at 4°C and supernatant was collected. 1mg of protein was precleared for 2h with Protein G agarose beads (50% slurry blocked with salmon sperm DNA) at 4°C. Immunoprecipitation was carried out by adding 2µg of antibody and 30µl of agarose G beads, nutating overnight at 4°C. After immunoprecipitation, beads were pelleted by centrifugation, followed by 4 washes to remove unspecific binding using a variety of buffers with varying concentrations of salt. Buffers 1 to 3 contained 0.1% SDS, 1% Triton-­‐X, 2mM EDTA, 20mM Tris 44 pH 8.0 and 150mM NaCl, 300mM NaCl, 500mM NaCl respectively. Buffer 4 contained 0.25M LiCl, 1% NP-­‐40, 1% Sodium deoxycholate, 1mM EDTA and 10mM Tris pH 8.0. Two additional washes with TE were done to remove any residual buffers from the beads. Complexes bound to the beads were eluted with 500µl of elution buffer (1% SDS, 1mM EDTA, 50mM Tris pH 8.0) at 65°C for 25min. Beads were pelleted by centrifugation and supernatant was collected. Reverse crosslinking was done by adding 0.2mM NaCl at 65°C overnight followed by treatment with Proteinase K at 45°C for 1h, and a second incubation of 15min at 65°C. DNA recovery was carried out with separation using 500µl of Phenol-­‐Chloroform. The aqueous layer was recovered and a second recovery was done with Chloroform alone to ensure the complete removal of phenol. DNA precipitation was done overnight by adding 2µL of ytRNA as carrier, 17uL of sodium acetate and 900µL of 95% ethanol. DNA was pelleted by centrifugation at top speed for 15min at 4°C and pellets were washed with 70% ethanol and dried by vacuum centrifugation. DNA was resuspended in 100µl of H2O and store at -­‐20°C. ChIP-­‐seq and bioinformatics. The ChIP protocol described above was used for ChIP-­‐seq with the exception of the DNA recovery step; in this case, DNA was retrieved using a PCR purification kit (Qiagen) following manufacturer’s instruction. CTCF ChIP was performed in duplicate using lysates from WEHI-­‐231-­‐shCtl and WEHI-­‐231-­‐TFII-­‐I KD cells. Recovered DNA was sent to the IRIC (Institut de Recherche en Immunologie et Cancérologie) sequencing facility where library construction and sequencing (100 bases, paired-­‐end, HiSeq, Illumina) were performed. DNA sequences obtained were trimmed to 45 bases (quality score > 30) and were aligned to the mouse genome (U.S. National Center for Biotechnology Information (NCBI) Build 37, July 2007, mm9) using the BWA algorithm (Li and Durbin 2009). After alignment, duplicates were removed and only the sequences with MAPQ score ≥ 30 were kept for further analysis. The model based analysis of ChIP-­‐Seq (Zhang, Liu et al. 2008) peak-­‐finding algorithm (MACS) was used to identify peaks in Ctl and TFII-­‐I KD conditions using the default settings. The loss of CTCF binding sites in TFII-­‐I KD cells was quantified using MACS with TFII-­‐I KD data set as background; during this process MACS software normalized the total tags count between the two samples. The intersect function of Bedtools with an overlap window of 50bp was used to 45 identify region of co-­‐localization between TFII-­‐I and CTCF in K562 cells in Fig. 10B (GSE51065). HOMER was used to annotate CTCF peaks and determine their genomic distribution (Heinz, Benner et al. 2010). seqMiner software was used to generate all the cluster and heat map data (Ye, Krebs et al. 2011). Data sets are available at the Gene Expression Omnibus (GEO) website, www.ncbi.nlm.nih.gov/geo, under accession ID: GSE60917. Gene Ontology and KEGG analysis. R (version 3.1.0) and Bioconductor (version 2.14) package “biomaRt”(Durinck, Moreau et al. 2005), “GOstat” (Falcon and Gentleman 2007) and “KEGGprofile” (Zhao and Shyr 2012) were used to perform all the Gene Ontology and pathway analysis on expression and ChIP-­‐seq data. 46 4. Results_____________________________________________________________________ Identification of CTCF interacting partners We previously observed strong binding between CTCF and known protein partners in MDA-­‐MB-­‐435 cells by CoIP (Witcher and Emerson 2009). Therefore, we decided to define CTCF interacting proteins in these cells using an affinity purification approach, followed by Mass-­‐
Spectrometry. For these experiments, conditions were sufficiently stringent that little background binding was observed in IgG controls (Fig. 1A). LC/MS of the purified proteins revealed several previously identified proteins (including YY1, Ybx1, TopoII, Npm and Ubtf) and novel interactors (Mascot score cut-­‐off of 100), primarily falling into four functional categories; SWI/SNF chromatin remodelers, transcription factors, splicing factors and DNA damage repair. (Fig. 1B, complete list with Mascot scores Appendix 1). Using Ingenuity Pathway Analysis, the top predicted network for CTCF interacting partners integrated members of the DNA replication, recombination, DNA repair, gene expression and protein synthesis pathways, attesting to the multi-­‐functional nature of CTCF (Appendix 2). Binding to newly identified partner candidates from each category was verified using Co-­‐IP (Fig. 2A). It has been shown that one of the CTCF binding partner TopoIIβ co-­‐localizes only to a subset of CTCF targets, suggesting the existence of distinct CTCF complexes (Witcher and Emerson 2009). Using biochemical purification, it was found that CTCF can be purified in two distinct complexes (as shown by size exclusion chromatography Appendix 3), by fractionated CTCF from cell extracts using column chromatography, with the final step in our purification scheme utilizing DNA cellulose as a binding matrix (Fig. 2B). CTCF interaction with TFII-­‐I Based on our MS analysis (Appendix 1) and Co-­‐IPs (Fig. 2A), one of the strongest CTCF binding partners was the transcription factor TFII-­‐I. Binding to TFII-­‐I was validated by Co-­‐IP and reverse Co-­‐IPs (Fig.3 A-­‐C) using lysates from cells of three tissue types (MDA-­‐MB-­‐435, HCT116 and WEHI-­‐231). Confocal microscopy indicated that co-­‐localization of CTCF and TFII-­‐I was not cell type dependent (Fig. 3). 47 Colocalization of CTCF and TFII-­‐I was assessed by confocal microscopy (Fig 4). To do this we employed Volocity software, which quantifies pixel colocalization within the two channels used to detect CTCF (Green, Alexa Fluor 488) and TFII-­‐I (Red, Alexa Fluor 594) for Z-­‐series confocal data (Dunn, Kamocka et al. 2011). The Volocity-­‐calculated Pearson correlation coefficient of 0.85 indicates a significant degree of three-­‐dimensional colocalization (1.0 = 100% overlap of signal). It has been proposed that interaction between CTCF and other protein partners is related to CTCF post-­‐translational modifications (Witcher and Emerson 2009). In MDA-­‐MB-­‐435 cell line by using 3-­‐amino-­‐benzamide (3-­‐ABA), it was shown that CTCF interaction between CTCF and TFII-­‐I is disrupted (Fig. 5A). To further test this, a mutant of CTCF that does not become poly-­‐ADP-­‐ribosyl-­‐ated (PARylated) by mutating the amino acids that are PARylated (Farrar, Rai et al. 2010) was used to check if other interaction becomes disrupted. Indeed, it was found by Co-­‐IP, that interaction with BRG-­‐1 is altered in Hek293T cell line (Fig. 5B). However it seems that the PARylation effect, is cell type dependent as in WHEI-­‐231 cell line, upon treatment with 3-­‐ABA is not enough to disrupt the interaction between CTCF and TFII-­‐I (Fig. 5C). Based on the fact that post-­‐translational modifications might play a role in the CTCF interaction with protein partners in a cell type dependant manner, interaction between CTCF and TFII-­‐I was mapped by Co-­‐IP, using mutated forms of CTCF that do not contain the C-­‐
terminal domain, N-­‐terminal domain and another one that contains the Zinc-­‐finger domain (Fig. 6A). In Hek-­‐293T cells, CTCF and TFII-­‐I interact with C-­‐terminal and N-­‐Terminal domains (Fig. 6B) however it was also shown that CTCF Zinc finger alone does not have the ability to form an association with TFII-­‐I (Fig. 6B). TFII-­‐I and CTCF cooperate to regulate gene expression KD studies show CTCF primarily acts as a positive regulator of target genes (Soshnikova, Montavon et al. 2010) and we surmised that CTCF may work together with TFII-­‐I to regulate the production of target transcripts. To test the possibility that CTCF and TFII-­‐I may cooperatively regulate expression of common target genes, we first identified genes whose expression is modulated by TFII-­‐I through microarray analysis of mRNA from TFII-­‐I KD cells (Ashworth and 48 Roy 2007) (Fig. 7A). A panel of the top hits from our microarray data were validated by conventional RT-­‐qPCR (Fig. 7B). We identified 500 genes differentially regulated between the control and the TFII-­‐I KD cells (Top500: fold change KD vs CT > 1.6 and p<0.05). Attesting to the specificity of this data, the TFII-­‐I coding gene (Gtf2i) was the top gene down regulated. To functionally categorize these genes, we performed functional annotation, Gene Ontology (GO) and pathway (KEGG) analysis. Of the genes regulated by TFII-­‐I, over 75% are predicted to be involved in metabolism (Fig, 7C). KEGG pathway analysis also showed a significant enrichment for transcripts involved in metabolic processes, with 46 metabolic genes regulated by TFII-­‐I being predicted to function in the same pathway (Fig. 6D, p=0.0002). One of the genes identified by microarray and down regulated in TFII-­‐I depleted cells was Cdkn2aArf (Fig. 7B). This tumor suppressor gene was previously shown to be transcriptionally regulated by CTCF (Witcher and Emerson 2009) and therefore might prove a suitable model to study cooperativity between TFII-­‐I and CTCF. Chromatin immunoprecipitation (ChIP) shows both proteins co-­‐localize to the Cdkn2aArf promoter region (Fig. 8A). As expected, TFII-­‐I binding was lost in KD cells (Fig. 8A), but surprisingly, CTCF binding was significantly diminished as well (p<0.05) (Fig. 8A). This loss was not generalized to all CTCF binding sites in tumor suppressor genes, as binding at Zmynd10 was undisturbed (Fig. 8B). To ensure that depletion of TFII-­‐I was directly responsible for the loss CTCF binding at Cdkn2aArf, we reconstituted our TFII-­‐I KD cells with exogenous TFII-­‐I (β and Δ isoforms (Hakre 2006)) and probed for CTCF binding by ChIP. CTCF binding was indeed restored at the Cdkn2aArf promoter in the TFII-­‐I KD complemented with exogenous TFII-­‐I (Fig. 8C). By depleting CTCF (Fig. 8D), there is also a downregulation of Cdkn2aArf and other TFII-­‐I target genes, including Khk, Pdhb and Glud1, in a similar manner to what is observed post-­‐TFII-­‐I KD (Fig. 7B). Our data indicate CTCF binding is targeted by TFII-­‐I and further support a cooperative role between CTCF and TFII-­‐I to transcriptionally regulate a subset of genes. Knowing that depletion of TFII-­‐I has an impact in gene regulation and that CTCF plays a role in the maintenance of epigenetic boundaries (Witcher and Emerson 2009), the mechanism by which epigenetic regulation of transcription plays a role was assessed. To do this, ChIP experiments were carried out using different epigenetic marks associated with active 49 transcription (H3K4ac, H2A.Z) and repressive marks such as (H3K27me3) in WEHI-­‐231 control and TFII-­‐I KD cells, finding no significant change among them (Fig. 8E). There is evidence that related CTCF with the recruitment of RNA Pol II to target genes (Chernukhin, Shamsuddin et al. 2007) and since TFII-­‐I is known as an initiation factor (Roy, Meisterernst et al. 1991), we wanted to look at Pol II recruitment to the Cdkn2aArf promoter. Total RNA polymerase II association with the Cdkn2aArf promoter did not shown difference between CT and TFII-­‐I KD cells (Fig. 8E), but we found a decrease in serine 5 phosphorylation of Pol II a mark associated with transcription initiation (Fig. 8E). Next, we wanted to identify the kinase responsible for Pol II modification when Cdkn2aArf is bound by TFII-­‐I and CTCF. Serine 5 of the Pol II CTD hepta-­‐repeat is primarily targeted by CDK7 and CDK8 (Rickert, Corden et al. 1999, Ramanathan, Rajpara et al. 2001, Galbraith, Donner et al. 2010, Helenius, Yang et al. 2011, Egloff, Dienstbier et al. 2012). While we did not observe any change in association of CDK7 at the Cdkn2aArf proximal promoter after TFII-­‐I KD, CDK8 binding was clearly disrupted (Fig. 8E), suggesting that CTCF and TFII-­‐I cooperate to form a scaffolding complex required for the efficient recruitment of the CDK8 complex to the Cdkn2aArf promoter. To explore whether TFII-­‐I might be involved in directing CTCF to binding sites genome wide we carried out ChIP-­‐Seq experiments to evaluate CTCF binding to genomic DNA in CT and TFII-­‐I KD cells. Of the 24,169 CTCF peaks identified in these experiments, 6978 were lost in absence of TFII-­‐I (p=0.03), consistent with the data we collected using the Cdkn2aArf promoter as a model (Fig. 9A-­‐B). Distribution analysis revealed that CTCF was primarily displaced from promoter and proximal upstream regulatory regions (Fig. 9C-­‐D). Of the 3986 CTCF sites located within +/-­‐ 3kb of a transcription start site, 777 were lost, using a stringent cut-­‐off of 3.7 fold loss of sequence tags at a given site (Fig. 9E). Visualization of our ChIP-­‐seq data at the Znf219 locus using the UCSC genome browser highlighted the specificity of CTCF at promoter regions. Here, CTCF sites were found at the 5’ regulatory region, the proximal promoter and within multiple exons. Of these, only CTCF binding at the proximal promoter was dependent on TFII-­‐I (Fig. 10A). 50 Recently, a genome-­‐wide screen of TFII-­‐I binding sites was carried out using the K562 cell line as a model (Fan, Papadopoulos et al. 2014). We aligned these sites with CTCF binding sites from the same cell line using publically available Encode data. We find 20% of TFII-­‐I sites localized near transcription start sites are co-­‐occupied by CTCF (Fig. 10B). Whereas CTCF can be found at less than 10% of TFII-­‐I sites bound outside promoter proximal regions. Again, this overlap underscores the potential importance of cooperativity between CTCF and TFII-­‐I. By comparing the overlap between genes regulated by TFII-­‐I from our microarray data, with gene promoters where CTCF binding was occluded after TFII-­‐I KD, our analysis revealed that of the 519 genes significantly changed upon TFII-­‐I KD (fold change ≥1.6, p < 0.05), approximately 219 genes have a CTCF binding site and out of these, 58 genes have a lost on CTCF binding in TFII-­‐I KD cells (Fig. 10C). Gene ontology analysis showed that of the 777 genes where CTCF binding was displaced after TFII-­‐I KD showed enrichment for genes involved in metabolic processes (Fig. 10D). This is highly consistent with our GO and KEGG pathway analysis of genes regulated by TFII-­‐I (Fig. 7C, D). As a model, we showed that CTCF binding to the Cdkn2aArf locus is lost after TFII-­‐I KD, and our ChIP-­‐seq data reveals that this model is extended to other CTCF targets, including the metabolic genes Khk and PdhB (Fig 11A). At the Khk gene, where TFII-­‐I targets CTCF to the proximal promoter (Fig. 11A, B), we see that TFII-­‐I KD has little impact on total Pol II recruitment or epigenetic modification. But similar to our data at the Cdkn2aArf gene, both CDK8 recruitment and serine 5 phosphorylation of Pol II are compromised in TFII-­‐I KD cells (Fig. 11B). CTCF and TFII-­‐I cooperation regulates cell survival Our data suggest a relation between TFII-­‐I and genes involved in metabolism. Therefore, we test TFII-­‐I KD cells for an altered response to nutrient deprivation, such as glucose. It was found that TFII-­‐I KD cells display a survival advantage over control cells, when grown in low glucose (Fig. 12A). This was associated with less apoptosis in this nutrient depleted environment as evidence by a strikingly lower sub-­‐G1 population (Fig. 12B, C). Apoptosis analysis by Annexin V and PI staining, confirm our findings that CT cells are more sensitive to low glucose conditions, 51 showing higher density of cells in late apoptotic stages (Fig. 12D), while TFII-­‐I KD cells show an early apoptotic stage after 2 days of growth in low glucose media. Overall CT cells show a higher frequency of Annexin V positive cells than TFII-­‐I KD cells (Fig. 12E). Another key energy source is the amino acid glutamine. There are multiple pathways through which glutamine can be metabolized for energy production. We see TFII-­‐I KD cells can adapt to these conditions and show much less sub-­‐G1 content (Fig. 13A) Even under conditions of low glucose and glutamine deprivation, TFII-­‐I cells are resistant to apoptosis (Fig. 13B). 52 5. Figures_____________________________________________________________________ Fig. 1. CTCF interactome reveals incorporation of CTCF into four functional groups. (A) Affinity purification of CTCF and interacting proteins using an anti-­‐CTCF antibody from MDA-­‐MB-­‐435 extract spiked with recombinant CTCF. Result shown is a representative coomassie stained gel of immunoprecipitated material and IgG control. (B) Schematic of LC/MS results showing that CTCF interacting partners can be classified in 4 distinct categories; transcription factors, DNA damage repair proteins, organizers of chromatin and mRNA processing. 53 Fig. 2. Interaction of CTCF with protein partners reveals integration of CTCF into multiple complexes. (A) Validation of mass spectrometry results by co-­‐IP with MDA-­‐MB-­‐435 extracts. Interaction was assessed by immunoblotting. (B) Biochemical purification of CTCF-­‐containing complexes by column chromatography. Immunoblotting for CTCF in eluents demonstrate integration of CTCF into multiple protein complexes on DNA. The purification scheme and a representative silver stained gel of the DNA cellulose purified fraction 9 is shown on the left. Western blotting for proteins co-­‐purifying with CTCF after separation on a DNA cellulose matrix is shown on the right. Proteins interacting with CTCF were eluted in fractions 9 and 17. 54 Fig. 3. CTCF interacts with transcription factor TFII-­‐I. (A) Forward and reverse co-­‐
immunoprecipitation (co-­‐IP) of MDA-­‐MB-­‐435 extracts using CTCF and TFII-­‐I antibodies. Inputs represent 2% of immunoprecipitated material. Treatment of extracts with nucleases (DNAse and RNAse) showed the interaction between the two proteins is independent of nucleic acids. (B) Interaction between CTCF and TFII-­‐I was assessed by co-­‐IP (forward and reverse) using HCT116 lysates. DDX5 interacts with CTCF in a RNAse-­‐dependent manner. (C) Interaction between CTCF and TFII-­‐I was assessed by co-­‐IP (forward and reverse) using WEHI-­‐231 lysates. 55 Fig. 4. CTCF and TFII-­‐I nuclear Colocalization by confocal microscopy. (a-­‐d) MDA-­‐435 cells were grown on coverslips, fixed and stained for confocal microscopy. Antibodies against TFII-­‐I (secondary, Alexa 594, red) and CTCF (secondary, Alexa 488, green) were used to bind endogenous proteins, followed by visualization using secondary antibodies. Scale bar 10μm. (e-­‐
g) Top-­‐down and rotated view of stained cells. Large panel (y-­‐x) = Top-­‐down, small panels (y-­‐z, left panel) and (z-­‐x, top panel) = view rotated along designated axis. The intersecting lines point to an example of colocalizing clusters. Scale bar: Axis 3μm. (h) Volocity-­‐generated Scatter plot of CTCF and TFI-­‐II intensities with Calculated Pearson correlation coefficient of 0.855. (i-­‐l) Same experiment as in (a-­‐d) except with the omission of the primary antibodies. Scale bar 10μm. 56 Fig. 5. CTCF interaction is associated with post-­‐translational modification. (A) Co-­‐
immunoprecipitation (Co-­‐IP) of MDA-­‐MB-­‐435 cell line treated with 3-­‐amino-­‐benzamide (3-­‐ABA, 3µM) shows that interaction between CTCF and TFII-­‐I is dependant of CTCF PARylation. (B) CTCF mutated form (M2) was overexpressed in Hek293T cells along with CTCF wild-­‐type (WT), revealing that interaction with Brg-­‐1, another CTCF interaction partner is disrupted once CTCF loss its ability to become PARylated. (C) Interactions dependent on CTCF PARylation status seem to be cell type dependant as Co-­‐IP of CTCF in WEHI-­‐231 cells treated with 3-­‐ABA, shows that this interaction is not dependant on PARylation. 57 Fig. 6. N-­‐terminal domain and C-­‐terminal domain of CTCF are needed to interact with TFII-­‐I. (A) Schematic representation of CTCF constructs generated to map CTCF interaction with TFII-­‐I. FL: full length, ΔC: deletion of C-­‐terminal; ΔN: deletion of the N-­‐terminal; ZF: Zinc finger domain (N and C terminal deleted). Lower panel shows Western Blot of the expression of CTCF constructs in Hek293T cell line. (B) CTCF constructs were ectopically expressed in the Hek293T cell line and Co-­‐IP of HA-­‐tagged (FL, ΔC) or Myc-­‐tagged (FL, ΔN, ZF) proteins were carried out. Interaction between CTCF and TFII-­‐I is mediated by regions within the C-­‐terminal and N-­‐
terminal domains. The CTCF ZF domain alone is incapable of mediating an interaction between the two proteins. 58 Fig. 7. Microarray profile of genes regulated by TFII-­‐I. (A) A heat map of three independent WEHI-­‐231 RNA samples from scrambled control (Ctl) and TFII-­‐I KD cells is shown on the left. Only those genes showing minimally a 1.6 fold change between the control and the TFII-­‐I KD samples and p< 0.05 are represented. On the right is a Western blot of TFII-­‐I levels from Ctl and TFII-­‐I KD. (B) RT-­‐qPCR validation of a subset of the genes identified differentially expressed between CT and TFII-­‐I depleted cells. Relative mRNA levels were calculated by 2-­‐ΔCT method and normalized to β-­‐Actin. Error bars represent SEM from at least 3 independent experiments (two-­‐
tailed Student t-­‐test, *** p≤0.05, **** p≤0.01). (C) Pie chart of Gene Ontology analysis performed for the Top500 genes (fold change > 1.6 and p < 0.05). (D) Pathway analysis using KEGG was performed on the Top500 genes differentially expressed. List of 46 genes found in a single metabolic pathway (p= 0,00017). 59 Fig. 8. TFII-­‐I KD impairs CTCF binding at Cdkn2a/Arf promoter. ChIP analysis of TFII-­‐I and CTCF at Cdkn2a/Arf-­‐p16 locus (n=3, two-­‐tailed Student t test, ***p≤0.05) (A) or Zmynd10 gene (n=3) (B) in Control and TFII-­‐I KD cells. (C) ChIP of CTCF at the Cdkn2a/ARF locus in CT, TFII-­‐I KD, and TFII-­‐I KD + exogenous TFII-­‐I. Western Blot of TFII-­‐I in the control, TFII-­‐I KD and TFII-­‐I complemented cells (n=3). (D) RT-­‐qPCR of TFII-­‐I identified target genes in Control and CTCF KD 60 cells from at least three independent experiments. Relative mRNA levels were calculated by 2-­‐
ΔCT
method and normalized to Ftl1 (two-­‐tailed Student t test, ****p≤0.01). Western Blot of CTCF in control and KD cells with and without exogenous TFII-­‐I. (E) ChIP-­‐qPCR of H2A.Z, H3K27me3, H3K27Ac, total RNA polymerase, Phospho-­‐Ser5-­‐RNA polymerase II, CDK7 and CDK8 (n=3), two-­‐
tailed Student t test, ***p≤0.05) at the Cdkn2a/ARF locus in Control and TFII-­‐I KD cells. Relative enrichment was calculated fold Ab over no Ab. For all graphs error bars represent SEM. Fig. 9. CTCF binding is disrupted at promoter regions genome wide upon TFII-­‐I KD. (A) Histogram of the number of CTCF binding sites identified in Control and TFII-­‐I KD cells by MACS (*p=0.03; ChIP-­‐seq data represent results from 2 (n=2) independent experiments). (B) Heatmap of CTCF binding sites in Control and TFII-­‐I KD cells within a +/-­‐ 5kb window surrounding the binding sites. (C) Pie chart of CTCF peak distribution at various genomic loci. (D) Histogram of the log2 fold change in CTCF peak distribution between Control and TFII-­‐I KD cells. (E) Smooth 61 lined scatter of CTCF tag density in +/-­‐ 5kb window around TSS and its coverage represented by number of tags per bp. Fig. 10. CTCF binding sites in promoter regions is related to genes associated with metabolic processes. (A) Visual representation of CTCF binding in gene Znf219 in control and KD cells using USC Genome Browser. UCSC CTCF reference track was used for comparison. (B) Colocalization of TFII-­‐I and CTCF ChIP-­‐seq Peaks from K562 cell line. Bedtools intersect function was used to calculate the number of overlapping peaks in the whole genome or at promoter region between TFII-­‐I and CTCF in K562 cell line. (C) Of the 519 genes revealed by microarray analysis to undergo significant change upon TFII-­‐I KD, 219 harbour CTCF binding sites. Of these 59 show a loss of CTCF binding upon TFII-­‐I KD. (D) Gene Onthology analysis of the 777 genes where CTCF binding is lost in promoter regions, shows that approximately 55% of these genes are related to metabolic processes such as RNA, nucleic acid, phosphorum and protein metabolism. 62 Fig. 11. TFII-­‐I targets CTCF and CDK8 to target genes. (A) Visual representation of CTCF binding in select genes in Ctrl and TFII-­‐I KD using the UCSC genome browser. A UCSC CTCF reference track was added for comparison. Aldh3B1 was use as a negative control as it does not show a 63 CTCF binding site. (C) ChIP analysis of the promoter region of Khk. A loss of CTCF and TFII-­‐I binding upon TFII-­‐I knock down, as well loss of phosphor-­‐serine 5 and CDK8 (n=3) are observed. Relative enrichment was calculated fold Ab over no Ab. For all graphs error bars represent SEM. ***p≤0.05 (two-­‐tailed Student t test). Fig. 12. TFII-­‐I KD cells show a survival advantage in response to nutrient deprivation. (A) CT and TFII-­‐I KD cells were cultured in normal media conditions with 5% FBS and low glucose (5mM) for two days. TFII-­‐I KD show a significant survival advantage under low glucose 64 conditions compared to Ctl cells. (B) Representative cell cycle analysis by propidium iodide staining after two days of culture in regular media or with low glucose. Ctl cells display a higher Sub-­‐G1 population than TFII-­‐IKD in low glucose. (C) Histogram shows the frequency of cells in Sub-­‐G1 population culture under low glucose conditions. For all graphs error bars indicate SEM (****p<0.01, two-­‐tailed Student t test). Fig. 13. TFII-­‐I KD cells adapt to survival under depletion of glutamine. (A) Cells were cultured in 5% FBS with either normal levels of glucose and glutamine or low glucose and depleted of glutamine. Under low glucose/no glutamine conditions, TFII-­‐II KD cells are more resistant to cell death as indicated by a low SubG1 population. (B) Representative histogram shows the frequency of SubG1 cells cultured under media without glutamine. 65 6. Discussion__________________________________________ _____________________ Identification of CTCF interacting partners Due the wide range of functions in which CTCF has been implicated, we decided to identify novel protein partners of CTCF. By mass spectrometry we were able to identify both novel interactors and those that have been shown previously to bind CTCF such as YB1, Yy1, TopoII, Npm and Ubtf. The isolation of these known binding partners reassured us that our data was of high quality, and consistent with previous studies. Based on our analysis and bioinformatics approaches, were able to group protein partners in four functional categories: SWI/SNF chromatin remodelers, transcription factors, splicing factors and DNA damage repair, highlighting the variation of functions of CTCF. The interaction between CTCF and SWI/SNF subunits, as well as splicing factors provides insights into previous work from other groups. CTCF colocalizes with SWI/SNF complexes on chromatin throughout the genome (Euskirchen, Auerbach et al. 2011). Our data suggests this overlap might be due to a direct interaction between the CTCF and core SWI/SNF subunits, and not based solely on functional cooperation. Since SWI/SNF subunits have no intrinsic DNA binding capacity of themselves, it is tempting to speculate that CTCF may recruit subunits of the SWI/SNF complex to target sites. Likewise, our data complement a recent report implicating CTCF as a regulator of RNA splicing (Shukla, Kavak et al. 2011). This study suggested CTCF regulates exon incorporation indirectly through promotion of RNA Pol II pausing (Shukla, Kavak et al. 2011). However, our data indicate direct recruitment of spliceosome proteins by CTCF, which provides an additional level of splicing control and offers insight into the functional relevance of CTCF binding within introns and exons, where it is commonly found globally (Chen, Tian et al. 2012). We previously showed the CTCF binding partner TopoIIβ localized to only a subset of CTCF targets, suggesting the existence of distinct CTCF complexes (Witcher and Emerson 2009). Consistent with a previous report (van de Nobelen, Rosa-­‐Garrido et al. 2010), we find size exclusion chromatography cannot separate CTCF into distinct complexes as CTCF containing elutions are constrained to sizes above one megadalton (Appendix 3). This suggests that CTCF is 66 integrated into at least one, large multi-­‐protein complex. To probe for biochemically distinct CTCF complexes, we fractionated CTCF from cell extracts using column chromatography, with the final step in our purification scheme utilizing DNA cellulose as a binding matrix. Eluents from DNA cellulose were relatively uncomplicated, with approximately 45 proteins being visible by silver staining in the first CTCF-­‐positive fraction. This approach showed CTCF is indeed integrated into biochemically distinct complexes that have unique affinities for DNA. This biochemical separation also served to validate our Mass Spec data, as many of the same proteins, from each of the major functional groups, co-­‐purified. CTCF interaction with TFII-­‐I We observed a robust interaction between CTCF and TFII-­‐I in all cell types tested by forward and reverse Co-­‐IPs (Fig. 3). We carried out Co-­‐IPs to detect the interaction between CTCF and TFII-­‐I in the presence of both DNAse and RNAse in multiple cell lines (Fig. 3 A-­‐C). The use of nucleases allow us to rule out the possibility that CTCF and TFII-­‐I interaction was mediated by nucleic acids, as recent evidence suggests that CTCF interacts with some protein cofactors in a RNA-­‐dependent manner (Yao, Brick et al. 2010). Here, we show that the interaction between CTCF and TFII-­‐I is not dependent on either DNA or RNA. The association of DDX5 with CTCF was found to be dependent on RNA as previously reported (Yao, Brick et al. 2010), but it was not the case for TFII-­‐I (Fig. 3B). The interaction of CTCF with TFII-­‐I was also supported using confocal microscopy to show colocalization of TFII-­‐I and CTCF within MDA-­‐MB-­‐435 cells (Fig. 4). Volocity software allow us to correlate the colocatization of CTCF and TFII-­‐I within the nucleus, were it was shown that indeed, both proteins are located in the same region with a correlation coefficient of 0.85, strengthen our hypothesis that CTCF and TFII-­‐I act in cooperative manner. CTCF interactions could be dependent on CTCF post-­‐translational modifications (Witcher and Emerson 2009). CTCF can be phosphorylated on C-­‐terminal by the casein kinase II (CKII) and this modification has been implicated in regulation of cellular growth (Klenova, Chernukhin et al. 2001). CTCF can also be modified in the N-­‐terminal by addition of chains of ADP-­‐ribose residues by the enzyme PARP1 (Yusufzai, Tagami et al. 2004), and it has been shown 67 that inhibition of PARP1 enzyme by 3-­‐aminobenzamide, impacts chromatin insulator function of CTCF, having an implication in gene expression (Yu, Ginjala et al. 2004). Based on the impact that PARylation has in CTCF activity, we decided to test if interaction between novel CTCF partners are dependent on PARylation status of CTCF. We were able to identify that CTCF interaction with TFII-­‐I is indeed disrupted upon treatment with PARP inhibitor 3-­‐ABA in MDA-­‐435 cell line (Fig. 5a). Based on this observation, we decided to take another approach and test if a mutant of CTCF that does not become PARylated can have an effect on its interaction. By Co-­‐immunoprecipitation assays, it was shown that interaction with Brg1-­‐1, a component of the SWI/SNF machinery (Hohmann and Vakoc 2014), is also dependent on this modification. In WEHI-­‐231 cells, treatment with 3-­‐ABA did not produced an ablation of the interaction between CTCF and TFII-­‐I, suggesting that PARylation might not always be necessary for this interaction. It is possible that other proteins bind CTCF in a PARylation-­‐
dependent manner and these interactions might stabilize TFII-­‐I and CTCF contact. Regardless, the capacity of PARylation to mediate the interaction between the two proteins seems to be cell-­‐type specific. By Co-­‐IP, we were able to determine the CTCF domains that interact with TFII-­‐I. We found that TFII-­‐I is able to interact with multiple domains, as deletion of either C-­‐terminal or N-­‐
terminal does not affect the interaction with CTCF. When only overexpressing the Zinc Finger domain of CTCF, interaction with TFII-­‐I becomes disrupted. The zinc finger domain of CTCF does not interact with TFII-­‐I, indicating this interaction is not solely dependent binding to DNA. Similarly, a multi-­‐domain interaction profile was observed between CTCF and the transcription factor YY1 (Donohoe, Zhang et al. 2007). One possible explanation for the multi-­‐domain interactions is that N-­‐terminal and C-­‐terminal domain might contain similar sequences that could from similar tertiary structures that could be recognized by TFII-­‐I, hence facilitating the binding. It could also be speculated that different molecules of TFII-­‐I can interact with CTCF, thus stabilizing the interaction with CTCF. 68 TFII-­‐I and CTCF cooperate to regulate of gene expression Out of all the proteins identified to interact with CTCF, one of the strongest interactions was the transcriptional factor TFII-­‐I. Since KD studies show CTCF primarily acts as a positive regulator of target genes (Soshnikova, Montavon et al. 2010), we supposed that CTCF may cooperate with TFII-­‐I to regulate the production of target transcripts. TFII-­‐I was originally found as a component of the basal transcription machinery (Roy, Meisterernst et al. 1991), but its binding at target genes may also be mediated by growth signals (Kim and Cochran 2000, Hakre 2006), as is commonly seen with other trans-­‐activators. Thus, the interaction between TFII-­‐I and CTCF represents a novel link between the extracellular environment and epigenetic organization. From our gene expression profile study, we were able to identify that upon KD of TFII-­‐I, most of the genes that had a significant change in expression were associated with metabolic functions. It has been shown that cancer cells display an altered metabolism (Hsu and Sabatini 2008). One of the key aspect in the metabolic change of cancer cells is the switch between oxidative phosphorylation and the glycolytic pathway, termed as Warburg effect (Koppenol, Bounds et al. 2011). Based on this assumption, we decided to test if there is a difference in proliferation between CT and TFII-­‐I KD cells on nutrient deprivation that will be addressed further on, since some of the genes regulated by CTCF and TFII-­‐I are repressed in cancer, such as Khk, we speculate that disruption of CTCF binding might lead to this repression as is seen with other silenced genes. Perhaps loss of CTCF binding at tumor suppressor genes commonly silenced in cancer could be due to disruption of the interaction between CTCF and TFII-­‐I that would lead to a loss of CTCF binding. TFII-­‐I ablation leads to changes on gene expression of other genes separate from the metabolic pathways. In our case, we found that expression of Cdkn2aArf, a known target of CTCF (Witcher and Emerson 2009) is downregulated upon knockdown of TFII-­‐I. When reintroducing TFII-­‐I (Δ and β isoforms) to TFII-­‐I KD cells, binding of CTCF was restored, showing that indeed TFII-­‐I cooperated with CTCF and targets it promoter regions, regulating the transcription of this gene. We also tried to determine, if this cooperation between CTCF and TFII-­‐I is extended to other genes. We indeed found that upon knockdown of CTCF, expression of target genes 69 downregulated by TFII-­‐I, are also affected by CTCF, showing that this cooperation extends to a set of genes. CTCF has epigenetic insulator activity, being capable of preventing the spread of repressive histone modifications such as H3K27me3 (Cuddapah, Jothi et al. 2009, Soto-­‐Reyes and Recillas-­‐Targa 2010) and enabling the local accumulation of activating marks such as histone acetylation and the histone variant H2A.Z (Splinter, Heath et al. 2007, Witcher and Emerson 2009, Soto-­‐Reyes and Recillas-­‐Targa 2010). Therefore, we investigated whether the diminished Cdkn2aArf expression and decreased CTCF binding we observed after TFII-­‐I KD were concomitant with changes to the epigenetic landscape. No significant changes of the repressive mark H3K27me3 or the activating marks H2A.Z, and H3K27Ac were observed. Eukaryotic transcription initiation is a dynamic and complex process requiring the association of general transcription factors to form a pre-­‐initiation complex (PIC) at promoter regions as an initial step (Li, Virbasius et al. 1999). Once this complex has been established, and RNA Pol II is recruited, Pol II is able to clear the proximal promoter and initiate mRNA synthesis after it is phosphorylated at Ser5 or Ser7 of the C-­‐terminal domain on the largest RNA Pol II subunit (CTD) (Egloff and Murphy 2008, Napolitano, Lania et al. 2014). CTCF has been implicated in the recruitment of RNA Pol II to target genes (Chernukhin, Shamsuddin et al. 2007) and TFII-­‐I is known as an initiation factor (Roy, Meisterernst et al. 1991), therefore, we thought that an obvious target for gene regulation by either protein might be RNA Pol II recruitment to the Cdkn2aArf promoter. Surprisingly, total RNA Pol II association with the Cdkn2aArf promoter was consistent between the control and KD cells, suggesting the transcriptional regulation lies downstream of PIC formation and RNA Pol II recruitment. Phosphorylation of Ser5 of RNA Pol II CTD hepta-­‐repeat is primarily targeted by CDK7 and CDK8 (Rickert, Corden et al. 1999, Ramanathan, Rajpara et al. 2001, Galbraith, Donner et al. 2010, Helenius, Yang et al. 2011, Egloff, Dienstbier et al. 2012). Analysis of Ser5 phosphorylation revealed a substantial decrease of this modification upon TFII-­‐I KD. RNA Pol II phosphorylation on Ser5 is required for transcription initiation and its loss can explain the reduction in transcriptional output of the Cdkn2aArf transcript. . 70 As CDK8 belong to the Mediator complex, is important to note that this complex has been associated with transcription activation and repression (Conaway and Conaway 2013). In human cells, Mediator complex containing a kinase module contributes to repression of RNA Pol II transcription by the DNA binding transcription factor C/EBPβ, whereas Mediator lacking the kinase module but containing MED26 contributes to activation by the same transcription factor (Mo, Kowenz-­‐Leutz et al. 2004). Besides CDK8, mediator subunits MED12 and MED13 have been shown to interact with or to be required for transcriptional activation by a variety of DNA binding transcriptional factors, such as β-­‐catenin. Direct interactions between the β-­‐catenin activation domain and MED12 have been shown to contribute to Mediator recruitment to genes (Kim, Xu et al. 2006), stressing the importance of this complex in transcriptional regulation. It is important to consider that Mediator has also been implicated in post-­‐initiation transcriptional activities. In Drosophila it has been shown that Mediator can be recruited to heat shock genes only upon heat shock and its recruitment is coincident with the release of paused RNA Pol II in these genes (Park, Werner et al. 2001). In mouse ES cells it was found that deletion of MED23 interfere with the activation of expression of the serum response gene Egr1 and to result both in the loss of Mediator recruitment to the Egr1 promoter and in a failure to release paused RNA Pol II (Wang, Balamotis et al. 2005). Evidence suggests that one way in which mediator might contribute to elongation control is by overcoming or bypassing the activities of factors that negatively regulate elongation. In this sense, it has been shown that Mediator complexes could overcome a block to transcription imposed by DSIF, in a reconstituted transcription system that is apparently devoid of P-­‐TEFb (Malik, Barrero et al. 2007), raising the possibility that, at least under some circumstances, Mediator may function independently or in parallel with, P-­‐TEFb to allow productive elongation by Pol II (Conaway and Conaway 2013). This evidence suggested us that the Mediator complex may be recruited to TFII-­‐I target genes. We carried out ChIP experiments towards the Mediator complex (MED1), but no significant differences were found between TFII-­‐I CT and KD cells (Appendix 5). As CTCF binds other core transcription factors, such as TAF3 (Liu, Scannell et al. 2011), it is possible that CTCF 71 and TFII-­‐I integrate into a larger scaffolding complex at core promoter regions enabling the recruitment of CDK8. This may be similar to the scaffolding complex previously shown to promote reinitiation (Liu, Kung et al. 2004). However, we do not rule out the possibility that other subunits of Mediator complex might be involved in this scaffolding complex. By analysing CTCF binding across the genome, our data predict that TFII-­‐I directs CTCF binding primarily to promoter regions, concordant with its own role as a general transcription factor. CTCF has previously been shown to facilitate the binding of UBTF to ribosomal DNA (van de Nobelen, Rosa-­‐Garrido et al. 2010), but to our knowledge TFII-­‐I is the first mammalian factor shown to enhance the association of CTCF to chromatin. In Drosophila, the introduction of deletion mutations to the insulator protein Cp190 virtually abolished CTCF binding to polytene chromosomes (Mohan, Bartkuhn et al. 2007). Thus, there is a precedent for zinc finger proteins directing CTCF to target sites. Recently it was demonstrated that sequences flanking core CTCF binding elements influence CTCF binding in vivo (Nakahashi, Kwon et al. 2013). Cooperative loading of CTCF and protein binding partners onto chromatin may explain this phenomenon. This data also suggests that the position of CTCF binding relative to the transcription start site may greatly influence its role in the transcriptional process. Our data indicates proximal promoter bound CTCF acts as an activator of initiation, whereas other reports have demonstrated CTCF bound within exonic regions acts as a negative regulator of transcription through enhanced pausing (Gomes and Espinosa 2010, Shukla, Kavak et al. 2011). CTCF and TFII-­‐I cooperation regulates cell survival Because of the link between TFII-­‐I and genes involved in metabolism, we wanted to test TFII-­‐I KD cells for an altered response to nutrient deprivation. Glucose represents one of the principle sources of cellular energy and we see the altered expression of several genes that might alter the utilization of glucose as a fuel source after TFII-­‐I KD. These include PdhB, a key mediator of glycolytic flux (Koike, Urata et al. 1990), and other genes that may impact TCA cycle progression such as Me2 (Raimundo, Baysal et al. 2011). This prompted us to examine growth of control and TFII-­‐I KD cells under conditions of low glucose. TFII-­‐I KD cells display a survival advantage over control cells, when grown in low glucose. This was associated with less cell 72 death in this nutrient depleted environment as evidence by a strikingly lower sub-­‐G1 population and lower Annexin V positive cells stained. But how do these cells have the ability to survive at lower glucose levels? A recent study suggest that the capacity of these cells to survive to glucose stress, is due to the production of lactic acidosis, which confers a protective advantage by reducing glucose consumption and increase the efficiency of glucose utilization (Wu, Ding et al. 2012). In this sense, it has been shown in gastric cancer, lactate production is necessary for cells to proliferate, and upon PdhB overexpression, the production of lactate and proliferation is decreased (Cai, Zhao et al. 2010), which fits to our proliferation assays where TFII-­‐I KD cells are more resistant than control cells with normal levels of PDHB. Another key energy source is the amino acid glutamine. There are multiple pathways through which glutamine can be metabolized for energy production. One key pathway is the conversion of glutamine to α-­‐ketoglutarate by Glud1, which provides α-­‐ketoglutarate for subsequent utilization by the TCA cycle. Because we see lowered levels of the Glud1 transcript in TFII-­‐I KD cells we examined the response to glutamine deprivation. Again, we see TFII-­‐I KD cells can adapt to these conditions and show much less sub-­‐G1 content. Even under conditions of low glucose and glutamine deprivation, TFII-­‐I KD cells are resistant to cell death. TFII-­‐I has been involved in the regulation of genes that are associated to Endoplasic Reticulum stress (ER stress) (Parker, Phan et al. 2001). Regulation of gene expression by nutrients is an important mechanism that confers and advantage under adverse conditions. A decrease in cellular carbohydrate levels causes the transcriptional regulation of a number of genes that encode proteins associated with ER stress, such as the glucose-­‐regulated-­‐proteins (GRP) (Chang, Barbosa-­‐Tessmann et al. 2002). There is evidence that TFII-­‐I is associated in ER stress, through the regulation of gene expression of GRPs (Parker, Phan et al. 2001). One of the consequences of ER stress is the activation of the unfolded protein response. In response to ER stress GRP78 binds to unfolded proteins to facilitate refolding, helping the cell to cope with ER stress, but if ER is persistent, then it can also contribute to cell death throughout apoptosis (Badiola, Penas et al. 2011). Therefore it possible, that in control cells, there is a higher rate of death due to ER stress response mechanism, but upon TFII-­‐I deregulation, cells might not be able to elicit an efficient ER response. 73 Recently, it has been shown that TFII-­‐I is associated with a proliferative capacity of the cells (Desgranges, Ahn et al. 2005, Ashworth and Roy 2009, Petrini, Meltzer et al. 2014), suggesting that TFII-­‐I function needs to be tightly regulated. There is evidence that mutations in GTF2i increase the proliferative potential of cells (Petrini, Meltzer et al. 2014). While it remains to be proven, it is also possible that loss of function GTF2i mutations, may tip of the metabolic balance of the cell to provide a survival advantage. Based on our results, it is probable, that upon loss of expression of TFII-­‐I, downregulation of tumor suppressor genes, such as Cdkn2aARF can contribute to an enhance proliferative potential (Brown, Harwood et al. 2004), and depending on the context and the level of expression or activity, TFII-­‐I can have both pro-­‐
proliferative and anti-­‐proliferative roles. Cancer cells undergo substantial metabolic reprogramming to enable high rates of proliferation and survival in a less than ideal environment. Amongst the changes already characterized is decreased expression of Khk in renal carcinomas (Hwa, Kim et al. 2006), a gene which we show here, is dependent on TFII-­‐I for proper expression. In the future, we will explore whether TFII-­‐I or CTCF binding to such genes is compromised leading to diminished expression in a different subset of cancers. 74 7. Conclusion_________________________ _______________________________________ Herein we show TFII-­‐I is a key regulator of metabolic gene expression, and we propose that this effect is mediated, at least in part, by promoting CTCF binding to the promoter region of a subset of these genes. Our data indicate TFII-­‐I and CTCF cooperate to promote CDK8 recruitment and Pol II phosphorylation on serine 5. Intriguingly, CDK8 recruitment to early response genes has been documented in response to various extracellular signals including hypoxia, serum and hormones (Belakavadi and Fondell 2010, Donner, Ebmeier et al. 2010, Galbraith, Allen et al. 2013). It will be important for future studies to probe the possibility that extracellular signals known to modify TFII-­‐I activity, impact the transcription of metabolic regulatory genes through the TFII-­‐I/CTCF/CDK8 axis that we have describe in this manuscript. 75 8. References_________________ ________________________________________________ Adelman, K. and J. T. Lis (2012). "Promoter-­‐proximal pausing of RNA polymerase II: emerging roles in metazoans." Nat Rev Genet 13(10): 720-­‐731. Akoulitchev, S., S. Chuikov and D. Reinberg (2000). "TFIIH is negatively regulated by cdk8-­‐containing mediator complexes." Nature 407(6800): 102-­‐106. Aoto, T., N. Saitoh, Y. Sakamoto, S. Watanabe and M. Nakao (2008). "Polycomb Group Protein-­‐
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435 proteins. 11 mg extract were loaded onto a sephacryl p300 column and size fractionated using an AKTA HPLC (GE Healthcare). Coloured standards were pre-­‐run to establish molecular weight of fractions. CTCF is seen to elute as a single high molecular weight fraction larger than 669 Daltons. Sp1 elutes separately from CTCF at 300-­‐400 Daltons, and b-­‐Actin elutes in multiple, non-­‐concurrent fractions. Thus the experimental approach enabled us to efficiently fractionate cellular proteins based complex size. 99 Appendix 4. 25 Relafve Enrichment α-­‐MED1 20 15 10 5 0 α-­‐MED1 -­‐ + Ctl -­‐ + TFII-­‐I KD Binding of Mediator complex to Cdkn2a/Arf promoter. ChIP against MED1 in the promoter region of Cdkn2a/Arf does not show a significant difference between control and TFII-­‐I KD cells. Error bars represent SEM. 100