Wesleyan University The Honors College Investigating Adjacent Gene Co-­‐Regulation in the Ribosome Biogenesis Regulons in S. cerevisiae by Linsin Smith Class of 2016 A thesis submitted to the faculty of Wesleyan University in partial fulfillment of the requirements for the Degree of Bachelor of Arts with Departmental Honors in Biology Middletown, Connecticut April, 2016 Acknowledgements I would like to take this opportunity to thank my research advisor, Dr. Michael McAlear, for the hours of support, teaching, constructive criticism, and wonderful conversation we have shared over the past year and a half. Thank you for giving me this opportunity to pursue molecular research and inspiring me to be the best scientist I can be. I’d like to thank my fellow lab mates, both past and present, for giving me the foundation I needed to be successful in the lab, and for being good friends and resources throughout my time here. I’d also like to express my gratitude to the members of the MacQueen lab, especially Karen Voelkel-­‐Meiman and Melanie Parziale for answering my every question and concern with a friendly smile and helpful advice. I’d like to thank the professors who have encouraged me and inspired me to pursue a career in molecular research, namely Dr. Robert Lane and Dr. Scott Holmes. The classes I have taken with you have impacted me beyond the lecture hall, and the advice and care you have given me over the past four years will not be forgotten. A huge thank you is due to my readers: Dr. Amy MacQueen and Dr. Joseph Coolon, for giving me your time and agreeing to be a crucial part of my honors thesis. Finally, I’d like to thank my friends, family, and the brotherhood of Psi Upsilon for being my support system, not just throughout writing this thesis, but throughout my time at Wesleyan. I honestly do not know where I would be without you. *** This thesis is dedicated in the memory of Richard Matthews, my incredible grandfather, who first inspired me to be a scientist. Thank you, I love you, and I wish you could read this. 2 Table of Contents Acknowledgements...........................................................................................................2 Table of Contents...............................................................................................................3 Index of Figures..................................................................................................................5 Index of Tables...................................................................................................................5 Abstract.................................................................................................................................6 Introduction........................................................................................................................7 Section I: Saccharomyces cerevisiae is an ideal model organism for research in molecular biology.......................................................................................................7 Section II: Transcription in S. cerevisiae..................................................................14 Section III: Ribosome Biogenesis in Eukaryotes..................................................20 Section IV: Adjacent Gene Co-­‐Regulation……………..............................................28 Mechanistic Model of Adjacent Gene Co-­‐Regulation....................................39 Questions Addressed in This Study...........................................................................40 Materials and Methods..................................................................................................42 Yeast Strains used in this study...................................................................................42 Primer Sequences used in this study.........................................................................43 Transformation Protocols..............................................................................................44 Protocol for Observing Gene Expression via RT-­‐qPCR......................................46 Protocol for Restriction Digest with SacI................................................................50 Results……………………………………………………………………………………………………..51 Section I: Moving MRX12 next to another RRB gene creates a new co-­‐
regulated gene pair……………………………………………………...….......………...……...……...51 3 Section II: Moving MRX12 next to two non-­‐RRB genes may break AGC….55 Section III: cis-­‐mutations in the MRX12 promoter provide insights into the interaction between paired genes.............................................................................................58 Discussion..........................................................................................................................64 Section I: The modular nature of AGC is evolutionarily significant.............64 Section II: The relationship between transcriptionally paired genes could be explained by promoter co-­‐localization.............................................................................69 References.........................................................................................................................74 4 Index of Figures Figure 1: The delitto perfetto transformation system.....................................................10 Figure 2: The CRISPR/Cas9 genome editing system.......................................................12 Figure 3: The preinitiation complex and associated trans-­‐factors............................15 Figure 4: Chromatin architecture and histone modifications.....................................18 Figure 5: The crystal structure of the yeast ribosome....................................................21 Figure 6: Nutrient signaling pathways regulate ribosome biogenesis....................23 Figure 7: Three distinct gene regulons required for ribosome biogenesis...........26 Figure 8: PAC and RRPE motif logo plots.............................................................................27 Figure 9: The MPP10-­‐MRX12 gene pair and expression profile.................................31 Figure 10: Insertions of active RNA Pol II-­‐transcribed genes can abrogate AGC.........................................................................................................................................................33 Figure 11: Transcriptional regulation of the MPP10-­‐MRX12 gene pair is coordinated by the promoter of MPP10.................................................................................34 Figure 12: Transcriptional regulation of dCFP is coordinated by MPP10..............36 Figure 13: Mutations in trans-­‐factors abrogate AGC.......................................................38 Figure 14: Molecular model of AGC........................................................................................40 Figure 15: Wild-­‐type expression of FUI1 under heat shock conditions.................52 Figure 16: Diagram of transformation to move MRX12 between ECM13 and FUI1........................................................................................................................................................53 Figure 17: MRX12 exhibits a normal heat shock response when positioned between ECM13 and FUI1.............................................................................................................54 Figure 18: Wild-­‐type expression of FIT2 and FIT3 post heat shock.........................56 Figure 19: Diagram of initial transformation to insert pGSHU cassette between FIT2 and FIT3.....................................................................................................................................57 Figure 20: Diagram of the secondary transformation to replace the pGSHU cassette with the MRX12 construct in yMM611..................................................................58 Figure 21: pGSKU insertion sites to mutate cis-­‐sequences in MRX12 promoter.60 Figure 22: Structure of the intermediate strains used to create mutations in the MRX12 promoter..............................................................................................................................61 Figure 23: Healing fragments to replace pGSKU cassettes and generate cis-­‐
mutations in MRX12 promoter...................................................................................................62 Figure 24: Mutation of Abf1p binding site, but not TATA-­‐like element in MRX12 promoter abrogate AGC.................................................................................................................64 Figure 25: RRB gene pairs are conserved across divergent fungal species..........66 Figure 26: Updated molecular model of AGC.....................................................................72 Index of Tables Table 1: Yeast strains used in this study...............................................................................42 Table 2: Primers used in this study.........................................................................................43 5 Abstract The production of a mature functional ribosome requires the coordinated expression of three distinct regulons: the ribosomal protein (RP), the rRNA, and the rRNA and ribosome biogenesis (RRB) regulons. These regulons contain hundreds of genes that need to be regulated in response to changing environmental conditions. We have found a significant enrichment of immediately adjacent co-­‐regulated gene pairs in the RRB and RP regulons. These gene pairs exhibit tighter transcriptional co-­‐regulation to each other than to the rest of the genes of the regulon as a whole, demonstrating a phenomenon we have characterized as adjacent gene co-­‐regulation (AGC). To further understand AGC, we have studied the co-­‐regulated RRB gene pair MPP10 and MRX12, and we have demonstrated that MRX12 is regulated by the MPP10 promoter. Here we show that AGC is modular by creating a new co-­‐regulated gene pair through moving MRX12 to a new locus. To further investigate the role of cis-­‐regulatory sequences in AGC, we have mutated sequences in the promoter of MRX12, and found that the TATA-­‐like element in the MRX12 promoter is not required to maintain AGC. We have also found that mutation of the Abf1p and Med8p binding sites in the MRX12 promoter abrogates AGC, suggesting novel roles for these proteins in the co-­‐regulation of paired genes. Beyond understanding the way large sets of genes are transcriptionally co-­‐regulated, understanding the molecular mechanisms of AGC also has important evolutionary significance. 6 Introduction Section I: Saccharomyces cerevisiae is an ideal model organism for genetic research Biological researchers have used model organisms for centuries to study and understand biological phenomena, and to apply their discoveries to more complex organisms. Studies performed on model organisms have been instrumental in the many biological breakthroughs, such as Gregor Mendel’s work in pea plants, which led to the discovery of the rules of chromosomal heredity. Generally, model organisms are easy to grow and manipulate, have short generation times, and are particularly well suited for specific fields of study. Due to the widespread use of a select few organisms, a great wealth of information is readily available to researchers studying numerous aspects of biology. Humans have used the model organism Saccharomyces cerevisiae for over ten thousand years in the processes of brewing and baking (Liti, 2015). In ancient times, the fermentation reaction carried out in brewing and baking was not understood scientifically, but now scientists know it is a byproduct of yeast metabolism (Lodolo, 2008). Louis Pasteur discovered yeast’s essential role in alcoholic fermentation in 1857, and in the 1930’s a group of brewers in Carlsberg became the first yeast geneticists through breeding experiments aimed to combine desirable brewing traits by crossing different yeast strains (Pasteur, 1858, Barnett, 2007). 7 As a single celled eukaryote, S. cerevisiae is an ideal model organism; it has a quick doubling time, it is inexpensive to cultivate, and is non-­‐pathogenic. S. cerevisiae reproduces through both meiosis and mitosis, and it can exist in one of two haploid mating types: a and α (Botstein et al. 1988). Since budding yeast is a clonal organism, it is easy to grow large, genetically homogenous populations and to isolate single colonies (Botstein and Fink, 2011). Budding yeast can be used as a model organism for a variety of purposes in genetics, molecular biology, and biochemical research. S. cerevisiae’s haploid genome consists of nearly 5,800 protein-­‐coding genes that are arranged into 16 chromosomes (Goffeau et al. 1996). S. cerevisiae was the first eukaryotic organism to have its entire genome sequenced in 1996, an accomplishment that launched a growing data source that is readily available on the Saccharomyces Genome Database (SGD) (Goffeau et al. 1996, Engel et al. 2013). Since it’s sequencing in 1996, we now have some understanding of the biological role of approximately 85% of the S. cerevisiae genome: a higher percentage than any other eukaryote (Bostein and Fink, 2011). This knowledge of the molecular function of the majority of yeast genes makes S. cerevisiae a fantastic model for genetic research. Before the age of molecular biology, classical geneticists made genetic mutations by performing random mutagenesis screens. These experiments often induced DNA damage through X-­‐ray radiation or chemicals to generate unique phenotypes due to random mutations. Geneticists could then work backwards from the novel mutant phenotype to find the mutated gene that was responsible. 8 From there, researchers would try to establish whether the mutations were on the same or different chromosomes through complementation assays (Esposito and Esposito, 1969). These assays also helped to establish gene dominance by assessing whether the resultant offspring displayed a wild type or mutant phenotype. Conditional mutants, such as those found to only affect cell viability at higher temperatures or on media lacking specific nutrients, could be used to link mutations with specific gene functions (Esposito and Esposito, 1969). Today, the process of generating mutations has progressed far beyond random mutagenesis via radiation or chemicals. Current methods harness the endogenous process of homologous recombination to insert and mutate specific sequences in the genome. Homologous recombination is a DNA damage repair process that uses homologous sequences in the diploid cell as the template for repair (Shinohara et al. 1995). The hijacking of the homologous recombination machinery allows for site-­‐directed mutagenesis, resulting in researchers being able to change specific genetic sequences. One such method of site-­‐directed mutagenesis via homologous recombination is called delitto perfetto, which is Italian for “the perfect murder” (Storici et al. 2006). As the name suggests, this method leaves behind no trace of its presence through utilizing a two-­‐step transformation protocol. First, a counterselectable reporter (pCORE) containing specific drug resistance genes for selective growth is inserted into the locus of interest (Storici et al. 2006). The reporter construct also contains an inducible endonuclease that creates a double-­‐strand break to encourage homologous recombination for the second 9 transformation. Once the double strand break has been initiated, the cassette is replaced with a DNA construct containing the desired mutation (Figure 1). The induction of a double-­‐strand break has been found to increase the likelihood of a successful transformation by over 1,000-­‐fold, making this method highly efficient. Figure 1: The delitto perfetto transformation system A diagram of the steps to mutate genetic sequences via the delitto perfetto system. First a pCORE cassette is inserted at a chosen locus by using homologous sequences that match the target locus on either side of the cassette (Step 1). Next, a double strand break is induced in the pCORE to remove the cassette and integrate the desired mutation through an engineered double stranded DNA construct created from oligonucleotides (Step 2). (Stuckey and Storici, 2013) Even though delitto perfetto has proven to be an efficient and reliable transformation method, CRISPR/Cas9 is a relatively new mutagenesis technique that has quickly become one of the most influential methodologies in molecular 10 genetics. Originally discovered as a viral defense system in bacteria, the CRISPR/Cas system utilizes a site-­‐specific Cas9 endonuclease directed by a guide RNA that can been engineered to target any sequence of interest (Cong et al. 2009, Mali et al. 2013). Once targeted to the correct locus, Cas9 initiates a double strand DNA break, stimulating homologous recombination. In order to repair the break, the cell can use an engineered DNA construct as a healing template for homology directed repair (Figure 2). Cells unable to repair the damage will theoretically become senescent or apoptotic due to the double strand break, allowing only the cells that have taken up the engineered DNA construct to survive. The CRISPR plasmid also contains a selectable marker, often the LEU2 gene, in order to isolate those cells that have taken up the plasmid and integrated the mutation through plating on selective media (Ran et al. 2013). Compared to delitto perfetto, CRISPR/Cas is much more efficient, and it can be utilized in cells that do not have the same degree of endogenous homologous recombination as found in budding yeast, including human cells. The ease of CRISPR/Cas9 transformations and the ability for this protocol to be utilized in a multitude of cell types makes it one of the most influential methodologies in molecular biology. 11 Figure 2: The CRISPR/Cas9 genome editing system The CRISPR/Cas9 transformation: The Cas9 protein (orange) creates a double strand break in the DNA after being guided to that locus by a guide RNA (crRNA). This cleavage can be resolved by non-­‐homologous end joining (NHEJ) or with homology directed repair (HDR). HDR utilizes donor DNA with homologous sequences to repair the break. This method allows for researchers to create DNA constructs that can then be taken up in the cell in a site-­‐specific manner. (Figure adapted from NEB.com, article by Alex Reis) The relative ease of making mutations in S. cerevisiae combined with the profound knowledge of the yeast genome makes it an optimal model organism for genetic research. Studies in S. cerevisiae have opened doors for newer genomic sciences and systems biology, including research on genetic and protein interaction networks (Botstein & Fink 2011). Many researchers attribute the creation of genome-­‐wide studies to S. cerevisiae, due to its important role in 12 pioneering genetic research, making it a choice model organism for investigating how the whole cell works together. New genome-­‐focused methodologies can be used to assess genome-­‐wide nucleosome occupancy and transcription factor binding sites, and such studies led to the discovery of a vast array of interlocking transcriptional regulatory networks. This field of functional genomics utilizes methods such as Chip-­‐seq, ChIP-­‐chip (chromatin immunoprecipiatation followed by DNA microarray), and many more methods to map the binding sites of transcription factors across the genome, thereby providing considerable insights into the global regulation of yeast genes (Duina et al, 2014, Lee et al. 2002). As technologies and methodologies progress, the fields of genomics and systems biology continue to expand, resulting in important discoveries that can be applied to many organisms beyond budding yeast. S. cerevisiae also serves as a suitable system for biochemical applications. The yeast two-­‐hybrid system can be used to identify protein-­‐protein interactions in vivo and budding yeast, like E. coli, can also be used to isolate large quantities of proteins for in vitro structural or biochemical analysis (Chien et al 1991, Ito et al. 2001). Immunoprecipitation experiments can be performed in which researchers create fusion proteins between the protein of interest and an affinity tag in order to isolate the protein through the use of antibodies that bind the tag in question. These tagged proteins can be used to discover DNA-­‐protein interactions and to identify transcriptional regulatory networks (Kuo et al. 1999, Lee et al. 2002). Isolated proteins may also be used in X-­‐ray crystallography or 13 nuclear magnetic resonance spectroscopy to obtain and observe their molecular structure. As the first eukaryote to have its genome sequenced and its profound history through its use in brewing and baking, it is clear that budding yeast has had a crucial impact and is an extremely useful model organism in the biological sciences. Section II: Transcription in S. cerevisiae Transcription, the highly regulated process of creating RNA copies from a DNA template, begins with the assembly of the pre-­‐initiation complex (PIC) at the transcription start site (TSS). Many yeast promoters contain a conserved ‘TATAA’ sequence, called a TATA-­‐like element, 35 bp upstream of the TSS (Hahn and Young, 2011). In the first step of transcription initiation, TATA-­‐binding protein (TBP) binds to the TATA-­‐like element, creating a bend in the DNA, which is thought to be essential for the recruitment of other basal transcription factors. TBP is a subunit of the essential basal transcription factor TFIID, which facilitates the recruitment of other basal transcription factors including TFIIA, TFIIB, TFIIE, TFIIF, TFIIH and RNA polymerase II (Conaway and Conaway, 2011). These factors are required for transcription in vivo and constitute the fully assembled preinitiation complex (Grummt, 2003, Smale and Kadonaga, 2003, White, 2011). Regulation of PIC formation and transcriptional initiation is mediated by the presence of the basal transcription factors, but also by other associated proteins including the SAGA complex, Mediator complex, and other transcription factors (Figure 3). The crucial step of recruiting TBP is often facilitated by the 14 SAGA (Spt-­‐Ada-­‐Gcn5-­‐Acetyltransferase) co-­‐activator complex (Huisinga and Pugh, 2004). In S. cerevisiae, the SAGA complex is composed of 21 proteins that comprise distinct functional units, including a recruitment module, acetylation module, a TBP interaction unit, a deubiquitination module, and an architecture unit (Koutelou et al., 2010).These distinct functional modules allow the SAGA complex to interact with the PIC to globally regulate transcription. Similar to the SAGA complex, the Mediator complex is made up of 20 proteins and possesses the ability to interact with transcription factors as well as RNA polymerase, giving it an important job in the regulation of transcription (Conaway and Conaway 2011). The SAGA complex, mediator complex, and general transcription factors define the baseline transcriptional state of the gene, which is then regulated by additional transcription factors (Hahn and Young, 2011). Figure 3: The preinitiation complex and associated trans-­‐factors The RNA polymerase II preinitiation complex and associated trans-­‐factors at the core promoter. The mediator bridges between regulatory DNA element (RE)-­‐
bound activators (ACT), RNA Pol II, and general transcription factors (GTFs). Specific domains of the mediator complex are color coded where blue indicates 15 the head, green is the middle, magenta is the tail and orange is the kinase domain. The SAGA chromatin remodeler complex is similarly associated with upstream regulatory elements. (Adapted from Dr. Martin Seizl, University of Munich) Many genes also contain specific regulatory cis-­‐sequences upstream of the TSS to recruit additional transcription factors (Struhl, 1987). Transcription factors are sequence-­‐specific, DNA binding proteins that ultimately affect PIC formation and activate or repress transcription. These transcription factors may be used both in a hierarchy where one master regulator initiates a recruitment cascade, as well as concurrently where different combinations of factors will result in differential transcription activity (Venters and Pugh, 2009). One such example is Rap1p, which acts as a repressor and activator of transcription depending on the binding context. These secondary trans-­‐acting elements, including the Mediator complex and gene-­‐specific transcription factors, control the rate of transcription through their interaction with PIC assembly, thereby affecting gene expression. Another direct method of transcriptional regulation is the physical organization of the DNA, which can regulate gene expression by restricting or allowing transcription factors access to transcribed regions. The simplest unit of organization of DNA is the nucleosome, a DNA-­‐protein complex that is used to condense linear DNA into chromatin (Jansen and Verstrepen, 2011). Each eukaryotic nucleosome is comprised of approximately 147 base pairs of DNA wrapped twice around eight highly basic histone proteins. Each nucleosome has two copies of each of the four canonical histone proteins: H2A, H2B, H3 and H4. Histone proteins include a structural globular domain that forms the core of the 16 nucleosome, as well as an N-­‐terminal tail that can be modified to remodel the chromatin structure (Jansen and Verstrepen, 2011). In order for a gene to be transcribed the DNA must be in a loose, relaxed state called euchromatin. In this state, transcription factors and the PIC can bind, facilitating transcription of that gene. Conversely, tightly condensed regions of DNA are defined as heterochromatin, which blocks access to of the transcriptional machinery (Grunstein et al. 1998). Nucleosomes are generally absent from the promoters of active genes, allowing for the transcriptional machinery to bind, called a nucleosome-­‐depleted region (NDR)(Lee et al. 2007). In order to regulate transcription, nucleosomes can be covalently modified on histone tails, or physically repositioned by protein complexes (Bannister and Kouzarides, 2011). These modifications directly affect gene regulation by modulating the relationship between nucleosomes and DNA, thereby changing the compaction status of the DNA (Figure 4). 17 Figure 4: Chromatin architecture and histone modifications The relationship between the condensation state of chromatin to resulting transcriptional repression or activation. In the first row, the tightly compacted heterochromatin represses transcription because transcription factors (TF) are unable to bind and promote transcription. Modulation between heterochromatin and euchromatin is done by an assortment of post-­‐
translational histone modifications and by the active movement of nucleosomes via chromatin remodeling complexes. (Ohtani et al, 2011) Posttranslational histone modifications include acetylation, methylation, ubiquitinylation, ribosylation, phosphorylation, and sumoylation often on the amino acids located on the N-­‐terminal tails (Bannister and Kouzarides 2011). Generally speaking, histone acetylation can be associated with transcriptional activation, where methylation usually confers a transcriptionally repressed state, including areas of heterochromatin. However, there are examples of 18 repressive acetylation marks and activating methylation marks, so this association is not always the rule (Bannister and Kouzarides 2011). Histone acetyltransferases (HATs) utilize the cofactor acetyl CoA to transfer an acetyl group to a lysine residue on the histone tail, effectively neutralizing the positive charge of the lysine. This change in charge weakens the interaction between the histone protein and the negatively charged phosphate backbone of DNA, resulting in a local de-­‐compaction of DNA at that locus (Bannister and Kouzarides, 2011; Deckert and Struhl, 2001). Histone de-­‐
acetylases (HDACs) perform the opposite reaction to HATs by removing acetyl marks on histone tails. Lysine residues are often the target of post-­‐translational histone modifications, however other amino acids can be subjected to post-­‐
translational modification as well. Active chromatin remodeling complexes work to regulate and reposition nucleosomes throughout the genome. These multi-­‐protein enzymes utilize ATP in order to reposition nucleosomes in various contexts (Venters and Pugh 2009). Essentially, these complexes use an ATPase domain to break the interaction between the DNA and histones in order to slide nucleosomes into different positions (Clapier and Cairns 2009). Nucleosome remodeling is an important cellular process, as nucleosomes must be cleared from the promoter regions of transcriptionally active genes in order to allow the transcriptional machinery access. Both through post-­‐translational histone modifications and nucleosome repositioning, chromatin architecture plays a significant role in transcriptional 19 regulation. Section III: Ribosome Biogenesis in Eukaryotes The production of mature ribosomes is an essential process for cell growth and proliferation (Warner, 1999). The ribosome is a large protein and ribosomal RNA (rRNA) complex that carries out the process of translation, turning mRNA transcripts into functional proteins. The process of translation is vital to all growing and dividing cells, as they need a constant supply of proteins in order to survive and grow. The importance of ribosomes has been well documented and they are highly conserved across all living organisms (Warner, 1999). Mature eukaryotic ribosomes are composed of a small 40S subunit and a large 60S subunit, which each have different roles in translation. In S. cerevisiae the small subunit contains one rRNA and 33 ribosomal proteins while the large subunit is comprised of 3 rRNAs and 46 ribosomal proteins (Figure 5). These distinct subunits also have distinct roles in the process of translation: the small subunit serves as a “decoding center” by bringing the tRNAs and mRNA together. The large subunit catalyzes the peptidyltransferase reaction, creating the peptide bonds in the growing amino acid chain (Woolford and Baserga, 2013). In a rapidly dividing yeast cell, more than 2,000 ribosomes are produced each minute, creating about 200,000 ribosomes per cell (Warner, 1999). Ribosome assembly is a major undertaking for cells, requiring many resources to create and move the various rRNAs and ribosomal proteins. This process is so complex that all three RNA polymerases play a role; RNA polymerases I and III transcribe the rRNAs and RNA polymerase II transcribes the genes for ribosomal 20 proteins (RP) and ribosome assembly factors (Woolford and Baserga, 2011). Altogether, transcribing the component parts of the ribosome poses a large metabolic cost on the cell. Around 60% of total cellular transcriptional activity is devoted to the production of rRNA, and around 50% of RNA polymerase II activity is devoted to RP mRNA synthesis (Warner 1999). Overall, ribosome biogenesis can account for around 90% of a cell’s metabolic activity (Zaman 2008). Figure 5: The crystal structure of the yeast ribosome The crystal structure of the mature ribosome in S. cerevisiae. The small 40S subunit is shown in blue while the large 60S subunit is yellow and red. Associated ribosomal proteins are shown as ribbons structures on the surface of the rRNAs. (Ben-­‐Shem et al. 2010) In yeast, ribosome biogenesis relies on transcriptional regulation to ensure that equimolar concentrations of the ribosomal components are produced. The large metabolic cost of creating the ribosome makes it essential for cells to be able to sense their extracellular environment and to regulate 21 ribosome production efficiently. Likewise In order for cells to react during times of stress, they must have mechanisms in place to monitor the environment and quickly and efficiently activate the proper survival responses. Favorable environmental conditions stimulate cell proliferation, which requires high amounts of ribosomes. Environmental stressors inhibit proliferation, thereby repressing ribosome biogenesis to conserve resources and keep the cell alive. Two nutrition-­‐sensing pathways, the TOR and ras/PKA cascades, regulate growth and cell division, and have been found to have downstream effects on ribosome biogenesis. The target of the immunosuppressant rapamycin (TOR) nutrient sensing pathway controls progression through the G1 phase (Thomas et al. 1997). The TOR1C kinase complex functions to phosphorylate downstream messengers that activate or repress transcription factors involved in the expression of amino acids, rRNAs and RPs in a nutrient-­‐dependent manner. In the presence of nutrients, Tor1 localizes in the nucleus and directly interacts with the promoter of 35S ribosomal DNA, activating 35S rRNA synthesis, thereby effectively up-­‐
regulating ribosome production (Figure 6) (Li et al., 2006). The ras-­‐cAMP-­‐protein kinase A (PKA) pathway monitors the availability carbon and nitrogen sources, and initiates intracellular signaling cascades promoting or preventing growth and division (Broach and Deschenes 1990). The presence of extracellular glucose activates Ras, which undergoes a conformational change from a GDP bound state to a GTP bound state. Ras, in the active GTP-­‐bound state, activates adenylyl cyclase (cAMP), which in turn 22 activates protein kinase A (PKA). Phosphorylation by PKA on a multitude of downstream targets regulates gene expression at the transcriptional level (Zaman et al, 2009). Active PKA has been found to upregulate transcription of many ribosomal proteins, while inactivated PKA has been found to repress RP transcription, and it confers an inability to stimulate transcription in the presence of glucose (Warner, 1999). Figure 6: Nutrient signaling pathways regulate ribosome biogenesis The TORC1 nutrient regulatory network in regards to ribosomal gene expression in S. cerevisiae. The upper panel depicts the conserved transcriptional regulators of rRNA, RP and RRB genes, and their regulation by the TORC1 pathway during optimal growth conditions. The bottom panel represents the change in transcription factor architecture upon growth inhibition. (Lempiäinen et al. 2009) Both the TOR and Ras/PKA nutrient-­‐sensing pathways directly influence ribosome biogenesis by regulating transcription factors that control activation and repression of genes involved in the production of the ribosome. An example of nutrient dependent ribosome regulation is the inhibition of the Maf1 23 repressor by both the Ras/PKA and TOR pathways (Figure 6) (Zaman et al. 2009). Maf1 activation represses the activity of RNA polymerase III, which in turn represses the expression of multiple rRNAs, tRNAs, and many small nuclear RNAs. As both the Ras/PKA and TOR pathways inhibit Maf1, the result is a de-­‐
repression of RNA polymerase III activity and the up-­‐regulation of ribosome biogenesis. Similarly, both nutrient-­‐sensing pathways regulate the AGC family kinase Sch9 (Figure 6). Sch9 overexpression leads to the expression of ribosome biogenesis genes, and its inhibition leads to transcriptional silencing (Zaman et al. 2008). These examples represent just a few of the many signals used to regulate ribosome biogenesis in an environmental-­‐dependent manner. While these signaling pathways are used to quickly regulate overall production of ribosomes in response to the environment, each gene must also be tightly co-­‐regulated to ensure the production of equimolar amounts of both ribosomal proteins and mature rRNAs. In order to accomplish this, signal transduction pathways regulate ribosome biosynthesis genes at the transcriptional level. Subsets of co-­‐regulated genes are often enriched for promoter-­‐binding motifs that recruit specific transcription factors for the co-­‐
regulation of entire sets of genes. In this way, a few transcription factors can regulate large sets of genes of similar functions, creating a co-­‐regulated set. The production of a fully functional ribosome is not as simple as producing ribosomal proteins and rRNA, as these products must also be processed and modified. A 2001 study of genome-­‐wide microarray data identified a novel set of 65 genes that were regulated similarly to each other and 24 distinctly from other genes during cell-­‐cycle progression, sporulation and diauxic shift (Wade et al. 2001). To confirm the co-­‐regulation of this cluster, quantitative PCR of a selection of five of these genes was performed over a variety of stress conditions, such as release from α-­‐factor inhibition, heat-­‐shock and secretory inhibition with tunicamycin (Wade et al. 2001). Interestingly, this cluster of 65 genes was enriched for genes whose products are integral to the biosynthesis of the ribosome without being a component of the final ribosome product, including transcription factors for RNA polymerases, RNA maturation factors, and RNA endonucleases (Wade et al. 2001). In 2006, this subset of 65 initial genes was expanded to around 200 co-­‐
regulated genes (Wade et al. 2006). Since then, this set of genes has been further expanded to around 280 genes, and the resultant regulon has been defined as the ribosome and rRNA biogenesis regulon (RRB) (Figure 7). The RRB regulon is the largest co-­‐regulated set of genes discovered in S. cerevisiae, making the molecular mechanism behind its co-­‐regulation of important significance (Abu-­‐
Jamous et al, 2014). 25 Figure 7: Three distinct gene regulons required for ribosome biogenesis Production of a mature ribosome requires the coordinated expression of a myriad of ribosomal protein genes, ribosomal RNAs, and ribosomal and rRNA biogenesis genes. (James Arnone) In order to understand how RRB genes are co-­‐regulated, bioinformatic promoter analysis was used to identify common sequences among RRB genes. It was found that RRB genes are enriched for two cis-­‐acting promoter motifs: the polymerase A and C (PAC) and ribosomal RNA processing element (RRPE) motifs, and distinct from those found in the RP and rRNA regulon gene promoters (Figure 8) (Wade et al. 2006, Bailey and Elkan ,1994). These motifs offer a potential mechanism for transcriptional regulation of the regulon, and both PAC and RRPE have been found to be necessary for proper expression of members of this large set of genes (Wade et al. 2001). The PAC motif is present in around 60% of RRB promoters and the RRPE motif is found in around 48% (Wade et al. 2006). 26 RRPE:
PAC:
Figure 8: PAC and RRPE motif logo plots The logo plots for the conserved promoter motifs enriched in the RRB regulon High throughput genome-­‐wide TF binding studies have identified Stb3p as a specific RRPE motif binding protein and Dot6/Tod6 as specific PAC motif binding factors (Liko et al. 2007, Zhu et al. 2009). These factors have been implicated as repressors of the RRB genes through their interaction and recruitment of the Sin3/Rpd3 deacetylase complex, which is a key co-­‐factor in the environmental stress response signaling network (Lempiäinen et al. 2009, Alejandro-­‐Osorio et al. 2009). Stb3 and Dot6/Tod6 are a link between the global nutrient sensing pathways TOR and Ras/PKA and the DNA sequences of RRB gene promoters (Figure 6). Taken altogether, the discovery of the RRB regulon as a set of co-­‐
regulated genes based on microarray data and conserved promoter motifs provides insight on how the gene members of large regulons are transcriptionally coordinated. In the case of the production of the mature ribosome, this involves the coregulated transcription RRB regulon which then interacts with the products of both the rRNA and RP regulons. The conserved RRPE and PAC promoter motifs present in the RRB regulon bind trans-­‐factors which are regulated by downstream effectors of the TOR and Ras/PKA pathways, ensuring that the production of ribosomes is regulated in accordance 27 to changing environmental conditions. Section IV: Adjacent Gene Co-­‐regulation After establishing that the RRB regulon represents a co-­‐regulated gene set with common promoter motifs, researchers looked closer at the organization of RRB genes in the genome. Surprisingly, it was found that approximately 16% (44 out of 282, P = 4.1x10-­‐4) of the RRB genes were located immediately adjacent to each other on the chromosome. Significant levels of immediately adjacent gene pairs were also found in the ribosomal protein (RP) regulon with around 13% (P = 1.1x10-­‐4) of the genes being paired (Wade et al. 2006). Examples of functional gene adjacency with transcriptional significance had already been discovered, including the prokaryotic operon, in which multiple tandemly oriented genes are co-­‐transcribed under the regulation of a single promoter (Cho et al. 1998, Pardee et al. 1959). This phenomenon has also been characterized in eukaryotes as in the divergent gene pair GAL1 and GAL10, which share a 680 base pair bi-­‐directional promoter region that drives the transcription of both genes (West et al. 1984). Unlike these two examples, RRB gene pairs have been found in all three orientations: divergent, tandem, and convergent. Furthermore, it has been shown through computational analysis that adjacently paired genes show correlated expression independent of their orientation, and that these adjacently paired genes tend to have similar functions (Cohen et al, 2000). In order to discover the functional role of RRB and RP gene adjacency, gene members of these two regulons were analyzed and searched for 28 transcriptional regulatory similarities (Arnone et al. 2012). Microarray data was taken following heat-­‐shock, osmotic shock and menadione exposure in order to compare expression levels between unpaired genes and genes from immediately adjacent pairs. Expression values were assessed for how tightly the different gene orientations were regulated through Pearson correlation coefficients (PCCs). The PCC of paired genes was found to be much higher than that of the unpaired genes for both the RRBs (0.91 vs. 0.68) and the RPs (0.77 vs. 0.31). These results that paired genes as a whole are more tightly co-­‐regulated than unpaired genes in the regulon (Arnone et al. 2012). This data suggests that pairing genes on the chromosomes could be a regulatory mechanism to control large sets of genes, including the RRB and RP regulons. To further explore the functional relevance of gene pairing, the evolutionary conservation of gene pairs was investigated across different regulons and species (Arnone et al. 2012). It was found that many other genetic regulons display enrichment for adjacent gene pairing, with the paired genes showing much higher average expression correlations than unpaired genes. These regulons included genes involved in the heat shock response, carbohydrate metabolism, purine base metabolism, and nitrogen metabolism, which all showed a much higher PCC for paired genes as opposed to unpaired (Arnone et al, 2012). As with the RRB and RP gene pairs, the gene pairings found in these diverse functional pathways were found in tandem, divergent, and convergent orientations (Arnone et al. 2012). The prevalence of transcriptionally 29 significant gene pairs in multiple regulons suggests that adjacent gene pairing may be an important regulatory mechanism for controlling large sets of genes. The extent of gene pairing in the RRB and RP regulons was examined across diverse species; including divergent fungal species, flagellates, the fruit fly D. melanogaster, the nematode C. elegans, the flowering mustard plant A. thaliana, the ciliate T. thermophila, the malarial parasite P. falciparum and humans (Arnone et al. 2012). Despite sizeable differences in regulon makeup from species to species, many of them show highly significant preferences toward maintaining paired gene adjacency (Arnone et al. 2012). This suggests that adjacent gene pairing has functional relevance, and may be used across diverse species to tightly co-­‐regulate functionally similar genes. Investigating the Mechanisms of Adjacent Gene Co-­‐Regulation The presence of statistically and transcriptionally significant adjacent gene pairs in large regulons motivated deeper studies into the mechanism of adjacent gene co-­‐regulation (AGC)(Arnone et al. 2011). The case of tightly co-­‐
regulated convergent gene pairs is particularly compelling, as there is currently no described mechanism for interaction between convergent genes. The RRB convergent gene pair MPP10-­‐MRX12(YJR003C) was used as a model for identifying and exploring the molecular mechanism of adjacent gene co-­‐
regulation (AGC) (Figure 9). The essential MPP10 gene encodes for a component of the U3 snoRNP, which functions in maturation of the 18S pre-­‐rRNA. The gene MRX12 (previously known as YJR003C) does not yet have a known function but is predicted to be involved in ribosome biogenesis and its protein associates with 30 the mitochondrial ribosome (Reinders et al. 2006, Kehrein et al 2015). This gene pair displays an interesting promoter motif makeup, where the promoter of MPP10 contains both the conserved RRPE and PAC motifs (located 63 and 94 bases upstream of the start ATG respectively) while the promoter of MRX12 doesn’t contain a stringent match to the RRPE or PAC consensus motifs (Arnone et al. 2011). Rela.ve"Expression"
120"
100"
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!!
Chromosome"X:!
RP
!! !! !!MPP10
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EBP2%
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Figure 9: The MPP10-­‐MRX12 gene pair and expression profile 30"
The convergent gene pair MPP10 and MRX12 located on chromosome X and separated by a 348 bp intergenic region as well as RRB singleton EBP2 on chromosome XI. Graph shows relative expression of MPP10, MRX12, and EBP2 post heat shock (This study) One of the questions addressed with the MPP10-­‐MRX12 gene pair was what the cis-­‐requirements were for adjacent gene co-­‐regulation to occur. To test this, a strain was designed to break the immediate adjacency between MPP10 and MRX12 (Arnone et al. 2011). A pCORE genetic reporter cassette was inserted 31 in the intergenic region between the two genes, increasing the distance between the two promoters by approximately 3.2 kb. After insertion of the cassette, the transcriptional regulation of MPP10 remained unchanged, however, the cassette disrupted the normal regulation of MRX12. This data indicates that immediate adjacency may be required in order for MRX12 to be properly co-­‐regulated (Arnone et al. 2011). To better understand the degree to which gene adjacency is required, various other sequences were inserted into the region between MPP10 and MRX12, including the coding sequence for LEU2 (Arnone et al. 2014). The insertion of LEU2 was found to have no effect on the regulation of MPP10, however uncoupled the co-­‐regulation phenomenon for MRX12, resulting in the de-­‐repression MRX12 over a heat shock time course (Arnone et al. 2014). However, when the inserted LEU2 was transcriptionally repressed, the co-­‐
regulation between MPP10 and MRX12 was unaffected, despite the greater than 2,000 bp increase in the intergenic region (Figure 10). These findings were similar for both orientations of the integrated LEU2 gene. Adjacent gene co-­‐
regulation was also found to be undisturbed by the insertion of a 0.7 RNA polymerase III-­‐transcribed tRNA Thr gene inserted in the intergenic region. These experiments suggest that the distance between the co-­‐regulated genes is less important for abrogating AGC than the presence of an actively transcribing RNA polymerase II complex (Arnone et al. 2014). 32 Figure 10: Insertions of active RNA Pol II-­‐transcribed genes can abrogate AGC Strains were grown in either SC-­‐leucine (A and B), SC media (C and D), or YPD (E and F). Strains were then subjected to a 37°C heat shock and monitored for their expression profiles at the EBP2, MPP10, and YJR003C (MRX12) genes by RT-­‐PCR. The profiles of the leftward-­‐oriented MPP10.LEU2 insert (YMM554) are represented in panels A and C, and the profiles of the rightward-­‐oriented MPP10.LEU2 insert (YMM559) are presented in panels B and D. (E) YMM555 MPP10. tRNA (Thr); (F) YMMM556 MPP10.Ty tRNA (Thr). (Arnone et al. 2014) In order to elucidate the function of the cis-­‐ acting binding motifs in the 33 promoter of MPP10, the RRPE and PAC motifs were mutated through site-­‐
directed mutagenesis (Arnone et al. 2011). These mutations yielded viable strains, meaning that neither motif is critical for basal transcription of the essential gene MPP10. The single and double knockout strains were subjected to either heat shock or glucose replenishment, and the resultant change in expression was monitored via quantitative PCR. The mutations in the promoter of MPP10 prevented proper regulation of MPP10 in response to changing environmental conditions. Surprisingly the effect of these mutations extended to MRX12, whose promoter is in the opposite direction and lays approximately 3.8 kilobases away (Arnone et al. 2011). This result suggests that the MPP10 promoter controls the transcription of both MPP10 and MRX12. Figure 11: Transcriptional regulation of the MPP10-­‐MRX12 (YJR003C) gene pair is coordinated by the promoter of MPP10 Gene expression levels were monitored for MPP10, MRX12 (YJR003C), and the unpaired RRB gene EBP2 in wild-­‐type (A) and ∆RRPE ∆PAC (B) strain (yMM514) (Arnone and McAlear, 2011) In order to address whether adjacent gene co-­‐regulation is gene-­‐specific, 34 a strain was constructed replacing both the ORF and promoter of MRX12 with that of the antibiotic resistance gene KanMX (Arnone et al. 2014). Quantitative PCR expression profiles over a heat shock time course show that substitution of MRX12 with KanMX uncoupled the gene pair. Transcription of MPP10 displayed normal RRB regulation while KanMX remained unchanged throughout the heat shock. This result suggests that adjacent gene co-­‐regulation is gene-­‐specific between MPP10 and MRX12, and that the promoter of MPP10 is not able to assume control of the transcriptional regulation of KanMX. However, this result does not indicate what specific part of the MRX12 locus allows co-­‐regulation to occur: whether it is the 3’ UTR, coding sequence, or promoter region. In order to elucidate the degree of gene-­‐specificity of AGC, a strain was created in which just the open reading frame of MRX12 was replaced with the open reading frame of LEU2 under the endogenous MRX12 promoter. Transcription of these genes in both the wild type and the LEU2 ORF replacement mutant follow the characteristic heat shock expression pattern for RRB genes: a quick repression followed by a gradual recovery. These data indicate that AGC is ORF independent. These results have been corroborated through the replacement of the MRX12 ORF with a destabilized cyan fluorescent protein (dCFP) to create a fluorescent reporter of AGC. The replacement of MRX12 with dCFP maintained AGC, suggesting that either the MRX12 promoter or 3’ UTR sequence may have functional significance in maintaining gene co-­‐
regulation. 35 Figure 12: Transcriptional regulation of dCFP is coordinated by MPP10 The MPP10-­‐dCFP construct and expression profile depicting the relative change in transcription of MPP10, dCFP and EBP2 in the YJR003C::dCFP mutant tested with one biological replicate and three technical replicates (Michael McAlear). Trans-­‐acting factors mediate adjacent gene co-­‐regulation In order to identify possible trans-­‐acting players in AGC, a bioinformatics approach was taken to survey an expression profile data set for a library of 165 deletion mutants of genes implicated in gene regulation and chromatin architecture (Arnone et al. 2014). This data set was screened for factors that significantly (P<0.005) disrupted expression of RRB or RP genes. Mutants associated with the SWI/SNF, SAGA/ADA, RSC, CCR4/NOT, NuA4 and Mediator complexes were all found to have large effect on RRB regulation. The PAC motif-­‐
binding factor Tod6 was also found in this initial screen, as would be expected for an RRB-­‐specific binding factor. This analysis was then repeated to look for factors that preferentially (P<0.05) disrupted paired RRB or RP genes 36 specifically. This screen identified many of the same transcriptional regulators, including members of the SAGA complex in RRB pairs, and Mediator, SWI/SNF, INO80, and CCR4/NOT complexes in RP pairs (Arnone et al. 2014). These results suggest that there may be multiple classes of transcriptional regulators involved in AGC, and some of these factors may be specific for the transcriptional regulation of paired genes. In order to confirm the roles of these trans-­‐factors, deletion strains with mutations in SNF2, CHD1, SPT20, ISW1, ASF1, and SWR1 were subjected to heat shock in order to monitor the resultant change in expression for the model RRB pair MPP10 and MRX12 (Arnone et al. 2014). Interestingly, an uncoupling defect was observed for the spt20Δ, chd1Δ and snf2Δ mutants, which maintained normal repression of MPP10 but lost repression of MRX12. The deletion mutants isw1Δ, swr2Δ and asf1Δ were found to not have an effect on adjacent gene co-­‐
regulation, indicating that the coregulated repression of MPP10 and YJR003C is mediated in part by the trans-­‐factors Snf2p, Chd1p and Spt20p (Arnone et al. 2014). 37 Figure 13: Mutations in trans-­‐factors abrogate AGC The indicated yeast strains were grown in YPD media to early log phase, subjected to a 37°C heat shock, and monitored for their expression profiles by RT-­‐PCR. (A) YMM593 snf2Δ; (B) YMM565 chd1Δ; (C) YMM562 spt20Δ; (D) YMM566 isw1Δ; (E) YMM595 asf1Δ; (F) YMM596 swr1Δ. (Arnone et al. 2014) In addition to trans-­‐acting factors, it is possible that changes in nucleosome conformation may also play a role in AGC, as nucleosomes often have to be cleared from the transcription start sites of genes in order to facilitate the recruitment of the transcription machinery. Studies were performed to assess nucleosome occupancy through micrococcal nuclease assay and ChIP analysis. The MNase assay showed that the positions of the nucleosomes present in the MRX12 and MPP10 promoter did not change a significant amount upon 38 heat shock (Arnone et al, 2014). The ChIP experiments found changes in the H4K12 acetylation levels at the promoter of MPP10 upon heat shock, but not at the promoter of MRX12. It was also found that histone H3 occupancy changes at the MPP10 promoter upon heat shock, but not at the promoter of MRX12 (Arnone et al, 2011). Although it is not clear exactly what the roles of histone modifications nor nucleosome positioning are in AGC, it is clear that there must be an interaction between the promoter regions of paired genes that involves conserved cis-­‐sequences and the trans-­‐factors Snf2p, Chd1p, and Spt20p. Mechanistic Model of Adjacent Gene Co-­‐Regulation One model for adjacent gene co-­‐regulation of RRB genes is a DNA looping mechanism in which the promoter of MPP10 and MRX12 are physically brought together (Figure 14). This model proposes that trans-­‐acting factors that bind the promoter of one gene may influence the promoter of the adjacent gene. The physical co-­‐localization of the paired promoter regions may allow trans-­‐factors, including Stb3p and Dot6p, to associate with both promoters in order to co-­‐
regulate both genes. Should looping occur, the transcriptional uncoupling of the MPP10-­‐YJR003C gene pair due to mutations of spt20Δ, chd1Δ and snf2Δ indicates that these trans-­‐factors are involved in forming the DNA loop, effectively promoting AGC. Deletion of these binding factors may prevent DNA looping and abrogate the cross talk between the two promoters, allowing only MPP10 to be properly regulated. In this model, deletion of the RRPE and/or PAC motifs in the MPP10 promoter would then result in abnormal regulation of both genes, reflecting the experimental data collected. The DNA looping hypothesis would 39 also allow promoter co-­‐localization for all adjacent gene pair orientations, as the DNA could bend in 3 dimensions to bring the promoter regions together. Rpd3%
Dot6/Tod6%
Stb3%
R P
!! !!
!! MPP10
SAGA%
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Figure 14: Molecular model of AGC The promoters of MPP10 and MRX12 may be brought together by DNA looping mediated by the SAGA (via Spt20) and Swi/Snf complexes. The physical co-­‐
localization of both promoters allows simultaneous transcriptional regulation of both genes by the PAC and RRPE binding factors targeting the promoter of MPP10. Stb3p, Dot6p/Tod6p, and Rpd3p all repress transcription of RRB genes, and are localized to the MPP10 promoter by the RRPE and PAC motifs (green, pink). Questions Addressed in this Study This study aims to investigate the molecular mechanisms of adjacent gene co-­‐regulation. This study tests two important components of AGC: the requisite of immediate adjacency to induce co-­‐regulation and the importance of trans-­‐
factor binding sites in the promoter regions of paired genes. This study tests whether AGC is a modular phenomenon in which genes can be swapped in and out of pairs in order to create differential regulatory schemes. The importance of trans-­‐factors and cis-­‐sequences in AGC has been demonstrated through 40 experiments mutating the conserved PAC and RRPE sequences in the MPP10 promoter. In this study mutations are induced in the promoter of MRX12. These mutations assess what sequences are required in the MRX12 promoter in order to maintain its interaction with MPP10. This study aims to elucidate the cis-­‐
sequences involved in the co-­‐regulation of adjacent paired genes and to understand the degree to which AGC is modular. 41 Materials and Methods Table 1. Yeast Strains used in this study Strain Genotype Source yMM13 yMM591 MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 pGSHU cassette inserted between ECM13 and FUI1, MRX12::Hyg MATa his3-­‐1 ura3-­‐0 leu2-­‐0 met15-­‐0 MRX12::KanMX Wade et al. 2001 Teryn Citino yMM557 yMM597 yMM602 yMM603 yMM604 yMM605 yMM606 yMM607 yMM608 yMM611 yMM612 yMM613 MATa leu2Δ1 trp1Δ63 ura3-­‐52 MRX12::LEU2 MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 MRX12 inserted between ECM13 and FUI1 MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 MRX12::LEU2, pGSKU cassette inserted in MRX12 promoter 67 bp upstream of the ATG MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 pGSKU cassette inserted in MRX12 promoter 67 bp upstream of the ATG MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 MRX12::LEU2, pGSKU cassette inserted in MRX12 promoter 175 bp upstream of the ATG MATa leu2-­‐1 trp1-­‐63 ura3-­‐52, pGSKU cassette inserted in MRX12 promoter 175 bp upstream of the ATG MATa leu2-­‐1 trp1-­‐63 ura3-­‐52, MRX12::LEU2, pGSKU cassette inserted in MRX12 promoter 310 bp upstream of the ATG MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 pGSKU cassette inserted in MRX12 promoter 310 bp upstream of the ATG MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 pGSHU inserted between FIT2 and FIT3 MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 MRX12 TATA-­‐like element at 175 bp upstream of ATG replaced with SacI cut site. Generated from intermediate strain yMM606. MATa leu2-­‐1 trp1-­‐63 ura3-­‐52 MRX12 Abf1p/Med8p binding site at 310 bp upstream of ATG replaced with SacI cut site. Generated from intermediate strain yMM608. 42 MacQueen/Holmes deletion collection (Open Biosystems) Teryn Citino This Study This Study This Study This Study This Study This Study This Study Natalie Favret This Study This Study Table 2. Primers Used in this Study Name ΔABF1mut3w ACT1qRTW Sequence from 5’ to 3’ TCCATTTAACTATGGTTGAAGAAAAAAAGAGCCAGGA
TGAAAATTATGAACAATAAGAGCTCGCGTTTCG TGTCGTACCTGCTATTCTAAAACGGGTACTGTACAGTT
AGTACATTGAGTCGAAACGCGAGCTCTTATTG TAGCTGTTAAGCATTTGAAAAAGTAGCTCTCATCGAA
CTCTTTCATGGTGATGAAATTGAATTCTCTACT CCGCTCGTTTCATTACCCGAAGGCTGTTTCAGTAGACC
ACTGATTAAGTAAGTAGAGAATTCAATTTCAT CTTCGGGTAATGAAACGAGCGGTAACGCTCACAAATT
TTTCTTTTTACGAACAAAA GAGCTC TTTAATAC ATACTTTTCTAGTTCTTATCCTTTTCCGTCTCACCGCA
GATTTTATCATAGTATTAAA GAGCTCTTTTGT ATCGTTATGTCCGGTGGTACC ACT1qRTC TGGAAGATGGAGCCAAAGC ARS1015pGSKUw1 EBP2qRTW TGGCGGTTAGCTGTTAAGCATTTGAAAAAGTAGCTCT
CATCGAACTCTTTCATGGTGATGTTCGTACGCTGCAGG
TCGAC TTTCATTACCCGAAGGCTGTTTCAGTAGACCACTGATT
AAGTAAGTAGATGAAAAAATTTCCGCGCGTTGGCCGA
TTCAT CGGGTAATGAAACGAGCGGTAACGCTCACAAATTTTT
CTTTTTACGAACAAAATATAAATTTCGTACGCTGCAG
GTCGAC ATAATACTTTTCTAGTTCTTATCCTTTTCCGTCTCACC
GCAGATTTTATCATAGTATTAACCGCGCGTTGGCCGAT
TCAT ATGAACTGTCCATTTAACTATGGTTGAAGAAAAAAAG
AGCCAGGATGAAAATTATGAACATTCGTACGCTGCAG
GTCGAC CTGCTATTCTAAAACGGGTACTGTACAGTTAGTACAT
TGAGTCGAAATATACGAAATTATCCGCGCGTTGGCCG
ATTCAT AACGCTACCTTACAGAAACG EBP2qRTC TCCGTTAGGCCTGCCTCTATCGAA ECM13YJRwp1 TACTTTGCGAATAAAACTACGAATTCAAACGTAAAAT
AAGACTACAGAGACTAAGGTAATGTCATTAACCCCTT
AAAGG AAGGTTTGGTTCTCAAGAGAGCTCACGCAAAATTTTT
TCTTACGTTCCCGTAGAACTGATTTCAACCTATGAAGG
TAAAG ACGAAGATCCAGATGCTGACAC GGCAAGTTCATCCGTCTGCTCG GGTCCTTCGGTGACAACCTTTG ΔABF1mut3c ΔRRPEmut1w ΔRRPEmut1c ΔTATAmut2w ΔTATAmut2c ARS1015pGSKUc1 ARS1015pGSKUw2 ARS1015pGSKUc2 ARS1015pGSKUw3 ARS1015pGSKUc3 ECM13YJRcp1 ECM13qRTW1 ECM13qRTC1 FIT2c1 43 Source This Study This Study This Study This Study This Study This Study James Arnone James Arnone This Study This Study This Study This Study This Study This Study James Arnone James Arnone Anand Soorneedi Anand Soorneedi This Study This Study This Study FIT3w1 FIT3c1 FIT2pGSHUw1 FUI1qRTW FUI1qRTC KANcp5 TTTCGGCGGCAGAAGATTCAGC TGCTGAGAGTATCACCACCACC TCGTATTATTATTATTGTTATTATTATTATTATCATT
ACTTTTATTAATATTCGTACGCTGCAGGTCGAC GTATCTAATTAAAATAAATATACCCGAAATCATACTA
AAAAAATAGTTAATAGGATAACAGGGTAATCCGCGCG
TTGGCCGATTCAT TCGTATTATTATTATTGTTATTATTATTATTATCATT
ACTTTTATTAATAGTAATGTCATTAACCCCTTAAAGG GTATCTAATTAAAATAAATATACCCGAAATCATACTA
AAAAAATAGTTAATGATTTCAACCTATGAAGGTAAAG TACTGCTTCCATGTCCACGTTC AAGGTGTCTTGGCTGCAAAGAG CGCGGCCTCGAAACGTGAGTC KLUwp1 KLUcp1 LEU2qRTW CTTGACGTTCGTTCGACTGATGAGC GAGCAATGAACCCAATAACGAAATC GTTACATGGTCTTAAGTTGGCG LEU2qRTC TGTGGGTGGTCCTAAATGGGG LEU-­‐YJR-­‐TESTwp1 LEU-­‐YJR-­‐TESTcp1 MPP10qRTW TGTGCTGAATAATTTTTTGTTG CAATTCGTAAACTTTTTCATC CGAGGAGGAGGAGGCTATTTAT MPP10qRTC CCTCCTCATCCGCAAATAAGTC SAG-­‐NSA-­‐FP1 URA3TEST1 GTTTCGGCACACGAGTTTCATTGG CCTTGGAAGACAATTCAGCAAGC YJRCHIPwp01 YJRqRTW CCTCTTGTTAGATAACGTAGCC ACCACCATTGACCCATACTCTC YJRqRTC GACCACTTCCATCAGTTCATCA FIT2pGSHUc1 FIT2MRX12wp1 FIT2MRX12cp1 This Study This Study Natalie Favret Natalie Favret This Study This Study This Study This Study James Arnone Q. Sun Q. Sun Teryn Citino Teryn Citino This Study This Study Anand Soorneedi Anand Soorneedi J. Arace Teryn Citino J. Arace Anand Soorneedi Anand Soorneedi Transformation Protocols Generation of Mutations via delitto perfetto A 1.5 ml aliquot of an overnight yeast culture was added to 50 ml YPD media and incubated at 30°C for 4 hours. Cells were then pelleted and 5 ml of a solution of 0.1 M LiAc and 1X TE was added. Cells were pelleted, washed with 50 ml ddH2O and resuspended in 250 µl of 0.1 LiAc and 1X TE. From this, 50 µl 44 samples were aliquoted to eppendorf tubes along with 300 µl of a solution of 0.1 LiAc and 50% PEG 4000 and 10 µl of concentrated PCR construct. The cells were incubated at 30°C for 30 minutes, then submitted to a heat shock at 42°C for 15 minutes. These cells were then pelleted and grown in YPD media for 3-­‐4 hours before being plated onto desired agar media. Single colony isolates emerging after 4 days were then plated onto various selective media to determine phenotype, as well as tested via PCR to determine genotype. Generation of Mutations via CRISPR/Cas9 A 1.5 ml aliquot of an overnight yeast culture was added to 50 ml YPD media and incubated at 30°C for 3-­‐4 hours. Cells were pelleted, washed with 50 ml ddH20 and resuspended in 250 µl of 0.1 LiAc and 1X TE. From this, 50 µl samples were aliquoted to eppendorf tubes along with 300 uL of a solution of 0.1 LiAc and 50% PEG 4000, 5 µl of denatured carrier DNA, 500 ng of CRISPR/Cas9 plasmid, and 10 µl concentrated PCR product. The cells were then incubated at 30°C for 30 minutes, and then submitted to a heat shock at 42°C for 15 minutes. The cells were pelleted and resuspended in 100 µl ddH20, then plated onto selective media. Single colony isolates emerging after 4 days were then plated onto various selective media to determine phenotype, as well as tested via PCR to confirm genotype. Alternate CRISPR/Cas9 Protocol A 1 mL aliquot of an overnight yeast culture was added to 20 ml YPD media and incubated at 30°C for 4-­‐5 hours. 10 ml aliquots were pelleted, washed 45 with 10 ml ddH20 and resuspended in 100 µl of 0.1 LiOAc. 100 µl of this solution was transferred to eppendorf tubes along with 240 µl 50% PEG (mw 3500), 36 µl 1M LiOAc, and 25 µl denatured carrier DNA (2mg/mL salmon sperm). The cell mixture was then vortexed lightly, in order to mix and allow the carrier DNA to coat the cells. Then 500-­‐800 ng of CRISPR/Cas9 plasmid was added along with 45 µl PCR product. The cells were then incubated at 30°C for 45 minutes, and then submitted to a heat shock at 42°C for 25 minutes. The cells were pelleted and resuspended in 1 ml ddH20 and 200 µl of that solution was plated onto selective media. Single colony isolates emerging after 4-­‐5 days were then plated onto various selective media to determine phenotype, as well as tested via PCR to confirm genotype. (This protocol used courtesy of the MacQueen Lab, Wesleyan University, 2016) Protocol for Observing Gene Expression via RT-­‐qPCR: 1. Heat-­‐Shock: The desired yeast strain was grown overnight at 30°C with shaking to an OD of 0.4 to 0.8 in 100 ml of YPD media. 15 ml aliquots of the culture were then transferred to 5 individual falcon tubes labeled 0, 5, 10, 20, and 30 respectively. The 5, 10, 20, and 30 time points were then set in a 37°C water bath and removed at the indicated time. 2. RNA Extraction: After heat shock, the cells were pelleted and resuspended in 1 ml of diethyl pyrocarbonate (DEPC) treated water. Cells were then treated with 450 µl 46 of DEPC treated TES and 400 µl phenol and heated to 65°C for 1 hour. Every 15 minutes the cells were vortexed in order to break open the cell membranes. The samples were then chilled on ice for 5 minutes. 400 µl phenol was added, and the cells were vortexed and spun down for 5 minutes. The aqueous phase was removed to a new eppendorf tube, and 400 µl chloroform was added. After the cells were vortexed and spun down, the aqueous phase was removed to a new eppendorf and 400 µl chloroform was added. After vortexing and spinning down the cells again, the aqueous phase was removed and transferred to a new eppendorf. 1 ml 100% ice-­‐cold ethanol was then added to the samples, as well as 40 µl (10% of total volume) 3M sodium acetate, in order to precipitate out the RNA. The samples were mixed and spun down at max speed for 5 minutes. The supernatant was removed and the RNA pellet was resuspended in 1 ml 70% ethanol to wash the RNA. The samples were then spun down again and the supernatant removed. The RNA pellet was resuspended in 50 µl DEPC treated water. 2 µl of the resuspended RNA was run on a DEPC agarose gel for 45 minutes at 80 volts to visualize the RNA. 3. DNase Treatment: In order to measure mRNA expression, all gDNA was degraded via DNase treatment. 10 µg of RNA for each time point was pipetted to an eppendorf tube, along with 2 µl of DNase enzyme and 5 µl DNase buffer. The final volume was brought to 50 µl with water (Ambion DNA-­‐free™ DNA Removal Kit AM1906). The samples were incubated at 37°C for one hour before adding 5 µl inactivation 47 enzyme. The samples were incubated at room temperature for 2 minutes and then centrifuged at 10,000 xG for 2 minutes in order to separate the sample from the DNase inactivation enzyme. 38 µl of the aqueous layer was transferred to new eppendorf tubes. 1 µl of each sample was then used in PCR amplification to confirm degradation of any contaminating DNA. In the case of contaminating DNA being present, the DNase treated samples were subjected to a secondary DNase treatment by adding 2 µl DNase enzyme to the samples. The samples were then incubated at 37°C for one hour before adding 5 µl inactivation enzyme. The samples were incubated at room temperature for 2 minutes and then centrifuged at 10,000 xG for 2 minutes in order to separate the sample from the DNase inactivation enzyme. 32 µl of the aqueous layer was transferred to new eppendorf tubes. 1 µl of each sample was then used in PCR amplification to confirm degradation of any contaminating DNA. 4. Generation of cDNA: In order to quantify mRNA expression, the RNA must first be converted to cDNA in order to be measured by RT-­‐qPCR. 10 µl of DNase-­‐treated RNA was combined with 2 µl oligo dT primer (Ambion RETROscript® Reverse Transcription Kit AM1710) and heated at 85°C for 3 minutes. Then the samples were transferred to ice for 5 minutes, and 2 µl 10x RT buffer, 4 µl dNTPs, 1 µl RNase inhibitor and 1 µl MMLV-­‐RT reverse transcriptase enzyme were added. Each tube was vortexed and spun before being heated at 44°C for 1 hour and 48 then 92°C for 10 minutes. 1 µl of each sample was then used in PCR amplification to confirm cDNA generation. 5. RT-­‐qPCR and Ct value analysis: In order to have enough sample volume for RT-­‐qPCR, the cDNA samples were diluted 1:5 in sterile ddH20. The primers being used were optimally designed for use in cDNA quantification via RT-­‐qPCR. The primers did not exceed 200 bp in length and must not anneal to an intron. The melting temperature of RT-­‐qPCR primers was between 57°C and 63°C with an ideal melting temperature of 60°C. These primers ended with either a G or C, and had a 50%-­‐60% GC composition. The primers being used did not have any self-­‐
complementarity or tendency to form inhibitory secondary structures in order to be used for RTqPCR. Primers were designed to anneal to the 5’ end of mRNA transcripts, as mRNA is degraded in a 3’-­‐5’ direction. A master-­‐mix was made for each gene whose expression is being analyzed. The master-­‐mix contained 200 µl Sybr Green master mix (Life Technologies Power SYBR® Green PCR Master Mix 4368708), 140 µl ddH20, and 20 µl of each primer. 18 µl aliquots of the mixture were then transferred to an optically clear 96 well plate for use in RT-­‐qPCR (Axygen, PCR-­‐96-­‐AB-­‐C 321-­‐65-­‐
051, or Thermo Scientific ABgene SuperPlate, AB-­‐2100). 2µl of cDNA was then added to each well, bringing the final volume of each well to 20 µl. The plate was then covered in optically clear film, sealing each individual well, and centrifuged at 2000rpm. To quantify expression levels, the plate was then placed in the 7300 49 Real-­‐Time PCR System (Applied Biosystems) and absolute quantification Ct values were obtained for each sample. In order to calculate percent-­‐change in expression, gene expression was normalized to the expression of ACT1 by subtracting the Ct of ACT1 from the Ct of the gene of interest to obtain a ΔCt value. The 0 minute ΔCt value was subtracted from each time point in order to obtain a ΔΔCt value, as baseline expression of each gene is defined as 100% expression. Ct values were on a Log2 scale that is inversely proportional to gene expression. ΔΔCt values were multiplied by a factor of -­‐1 in order to make ΔΔCt directly proportional to gene expression. These values were converted to a linear scale by raising 2 to the power of the ΔΔCt and multiplying the resulting value by 100. Technical replicates were then averaged in order to plot the data as well as calculate standard error. All data shown is generated from 2 biological replicates and 6 technical replicates unless otherwise stated. Protocol for Restriction Digest with SacI 2 µl PCR product was added to a mixture of 2 µl NEBuffer 1.1 10x, 1 µL SacI enzyme, and 15 µL ddH2O per sample. Samples were heated at 37°C for 1.5 hours, then tested for successful digest via polyacrylamide gel electrophoresis. (New England BioLabs, NEBuffer 1.1 10x concentration B7201S, and SacI Restriction Enzyme R0156L) 50 Results Section I: Moving MRX12 next to another RRB gene creates a new co-­‐
regulated gene pair Previous studies have demonstrated that mutations in the conserved RRPE and PAC motifs present in the MPP10 promoter affect the expression of both MPP10 and MRX12. Studies have also shown that the insertion of an actively transcribing gene between MPP10 and MRX12 uncouples the expression of MRX12 from MPP10 (Arnone et al. 2014). This result suggests that without the presence of MPP10 in an immediately adjacent position, MRX12 would not be regulated in typical RRB fashion (Arnone et al. 2011). Since gene pairs can be readily uncoupled, as demonstrated through the insertion of LEU2, we wondered whether it would also be possible to create novel co-­‐regulated gene pairs through genetic manipulation. To test whether co-­‐regulated gene pairs can be created through genetic manipulation, the MRX12 gene was moved next to another RRB gene, FUI1, on chromosome II. FUI1 codes for a high affinity uridine permease and exhibits canonical RRB repression after heat shock (Figure 15). FUI1 also has two RRPE motifs in its promoter region, located 194 bp and 223 bp upstream of the ATG. The role of FUI1 in ribosome biogenesis is through the uridine metabolism pathway, and the presence of two RRPE motifs as well as its response to stress made FUI1 an optimal candidate to induce adjacent gene pairing with MRX12. 51 Chromosome(X:
Chromosome(II:
RR
EBP2 150 Relative Expression !
! ! !!
! MPP10
MRX12
!
FUI1
! ECM13
RP
FUI1 MRX12 100 50 0 0 5 10 15 Time 20 25 30 Figure 15: Wild-­‐type expression of FUI1 under heat shock conditions The relative organization of ECM13, FUI1, MRX12, and MPP10. The RRPE motifs are notated in green and PAC motifs in pink. The relative expression of EBP2 (RRB singleton), FUI1, and MRX12 under heat shock conditions, quantified by RTqPCR. In order to move the MRX12 gene, we first had to obtain a deletion strain in which the endogenous copy of MRX12 was replaced with the KanMX gene (yMM557, Open Biosystems). This was to ensure that upon insertion of MRX12 next to FUI1 there would not be two copies of MRX12 present in the genome. From this deletion strain we inserted a pGSHU cassette in between ECM13 and FUI1 through the delitto perfetto system (yMM591, Teryn Citino). The pGSHU cassette was then replaced with a DNA construct containing 256 bp of the 3’ UTR, the MRX12 coding sequence, and 447 bp of the 5’ UTR region using the delitto perfetto transformation protocol (Figure 16). In order to select for transformants that had incorporated the MRX12 construct in place of the pCORE cassette, cells were plated on 5FOA media and tested for MRX12 integration via 52 PCR. This transformation created strain yMM602: in which the promoter region, coding sequence, and 3’ UTR of MRX12 is located next to FUI1 in a tandem orientation. MRX12%
Chromosome II: ECM13%
pGSHU"
FUI1%
yMM591 R"R"
ECM13%
MRX12%
FUI1%
yMM602 Figure 16: Diagram of transformation to move MRX12 between ECM13 and FUI1. Transformation strategy to insert MRX12 between ECM13 and FUI1. The MRX12 construct was amplified from a wild type strain using primers with 20 bp of homology to sequences around MRX12 (light orange rectangles) and 50 bp of homology to the regions between ECM13 and FUI1 (light green rectangles). Using the delitto perfetto transformation protocol, MRX12 was inserted between ECM13 and FUI1, creating the resultant strain yMM602. To test the change in expression of MRX12 in this new locus, a heat shock was performed and resultant expression profile was created through RTqPCR (Figure 17). We found that when adjacent to FUI1, MRX12 displays a typical RRB expression profile. This result implies that FUI1 may be able to influence the expression of MRX12 the same way that MPP10 does on chromosome X, suggesting that AGC is a modular phenomenon. This result is a direct contrast to the experiment performed by Arnone et al. in which the LEU2 construct was inserted between MPP10 and MRX12, resulting in the loss of typical repression of 53 MRX12. This exciting result suggests that AGC is a flexible, modular phenomenon that allows genes to be swapped in and out of pairs in order to change gene expression. !! ECM13
!! !!
200(
EBP2%
FUI1%
MRX12%
FIT3%
150(
100(
50(
0(
B: 100(
50(
0(
5(
10(
15(
Time(
20(
25(
Chromosome(II:!
200(
150(
0(
0(
10(
15(
20(
25(
30(
25(
30(
Chromosome(X:!
R" P"
FUI1&
MPP10&
100(
50(
5(
Time(
R"R"
MRX12&
ECM13&
0(
30(
120(
FUI1%
100(
MPP10%
Relative(Expression(
MRX12
FIT3%
FUI1%
MPP10%
150(
0(
Relative(Expression(
Relative(Expression(
Relative(Expression(
200(
FUI1
!!
A: Chromosome(X:!
RP
!! !! !!MPP10
RR
!!
Chromosome(II:!
MRX12::KanMX&
MRX12%
80(
60(
40(
20(
0(
5(
10(
15(
Time(
20(
25(
30(
0(
5(
10(
15(
Time(
20(
Figure 17: MRX12 exhibits a normal heat shock response when positioned between ECM13 and FUI1. A: Schematic showing the relative organization of genes in strain yMM13 and subsequent graphs depicting gene expression post-­‐heat shock for EBP2 (RRB singleton), FIT3 (non RRB control), FUI1, MRX12, and MPP10. B: Schematic showing the relative organization of genes in strain yMM602 and subsequent graphs depicting gene expression post heat-­‐shock. 54 Section II: Moving MRX12 next to two non-­‐RRB genes may break AGC Previous data demonstrated that when MRX12 is separated from its position next to MPP10 it loses its canonical expression pattern under heat-­‐
shock, and is in effect transcriptionally un-­‐coupled from the RRB regulon. In order to further test the extent to which MRX12 can function autonomously without input from MPP10, we sought to move it to a new locus in the genome between two non-­‐RRB genes. This experiment also serves as a control to confirm the creation of a new co-­‐regulated pair between MRX12 and FUI1. We searched the genome for possible insertion sites removed from RRB genes, such that the locus in which MRX12 was to be inserted would not influence its expression, and we found the convergent gene pair FIT2 and FIT3 on chromosome XV. Both of these genes are involved in iron transport and are upregulated under stress conditions, making the intergenic region between them a good candidate for MRX12 insertion (Figure 18). Moreover, since FIT2 and FIT3 have similar expression profiles and are involved in the same biological process, it is possible that they themselves are a co-­‐regulated pair, meaning that the insertion of MRX12 between them would allow us to test AGC in another genetic regulon. 55 !!FIT2
450"
MRX12
FIT2%
FIT3%
MRX12%
EBP2%
400"
Relative(Expression(
FIT3
!!
Chromosome"X:!
RP
!! !! !!MPP10
!!
Chromosome"XV:!
350"
300"
250"
200"
150"
100"
50"
0"
0"
5"
10"
15"
Time"
20"
25"
30"
Figure 18: Wild-­‐type expression of FIT2 and FIT3 post heat shock The relative organization of the FIT2 and FIT3 genes on the XV chromosome and their relative expression levels after heat shock. The RRPE motifs are notated in green and PAC motifs in pink. The relative expression of EBP2 (RRB singleton), FIT2, FIT3 and MRX12 under heat shock conditions. To move MRX12 to a neutral place in the genome, we utilized a strain from the deletion library that had the MRX12 coding region replaced by a KanMX gene (yMM557, Open Biosystems). The pGSHU cassette was inserted in the intergenic region between FIT2 and FIT3 through the delitto perfetto transformation protocol (Figure 19). In order to select for transformant colonies that had taken up the pGSHU cassette, cells were plated on SC-­‐Ura and then tested genetically through PCR. 56 pGSHU&
FIT2%
FIT3%
yMM557 FIT2%
pGSHU&
FIT3%
yMM611 Figure 19: Diagram of initial transformation to insert pGSHU cassette between FIT2 and FIT3. Transformation strategy to integrate the pGSHU cassette in between FIT2 and FIT3. The cassette was amplified using primers containing 20 bp of homology to the pGSHU region and 50 bp of homology to the intergenic region between FIT2 and FIT3. After the insertion of the pGSHU cassette, a CRISPR/Cas9 transformation was performed to replace the pGSHU cassette with the MRX12 gene. The CRISPR/Cas9 transformation protocol was used with a CRISPR plasmid containing a guide RNA targeting the Hyg gene in the pGSHU cassette in order to facilitate a double strand break. This is designed to induce homology directed repair with a DNA construct containing 256 bp of the 3’ UTR, the MRX12 coding sequence, and 447 bp of the 5’ UTR region, along with 50 bp of homology with the intergenic FIT2-­‐FIT3 region on either side (Figure 20). Transformants are currently being selected on SC-­‐Leu media and once obtained will be tested for the integration of MRX12 through PCR. 57 FIT2%
pGSHU&
FIT3%
yMM611 LEU2%
MRX12%
%
s9
Ca
FIT2%
MRX12%
Hyg%
gRNA%
FIT3%
in progress) (currently Figure 20: Diagram of the secondary transformation to replace the pGSHU cassette with the MRX12 construct in yMM611. Transformation strategy to replace the pGSHU cassette with the MRX12 gene in between FIT2 and FIT3. The MRX12 construct was amplified using primers containing 20 bp of homology to the 3’ and 5’ UTRs around MRX12 (yellow rectangles) and 50 bp of homology to the intergenic region between FIT2 and FIT3 (light blue rectangles). Section III: cis-­‐mutations in the MRX12 promoter provide insights into the interaction between paired genes Previous experiments have shown that RRPE and PAC motifs within the MPP10 promoter are essential for appropriate transcriptional regulation of both MPP10 and MRX12. Mutation of these sequences resulted in the transcriptional up-­‐regulation of both MPP10 and MRX12 after heat shock, suggesting that the expression of MRX12 is regulated by the MPP10 promoter. To fully understand the nature of the interaction between the two genes, we also wanted to understand which sequences in the MRX12 promoter are responding to the regulatory information given by MPP10. Previous studies which replaced the ORF of MRX12 with LEU2 show that the regulation of MRX12 by MPP10 is ORF 58 independent, suggesting that the promoter region of MRX12 may be responding to MPP10. Several possible transcription factor binding sites were annotated in the MRX12 promoter through the computational analysis program PROMO, which analyzes promoter regions for transcription factor binding sites generated from TRANSFAC (Messeguer et al. 2002, Farré et al. 2003). This program found several putative TF binding sites, including Med8 binding sites, Abf1 binding sites, and TATA-­‐like elements. In addition to this, we identified a less stringent RRPE-­‐like sequence in the MRX12 promoter that was not found in earlier studies due to the use of the MAST program, which searches for the best combined match to a specific motif and used an E value of 50 (Bailey and Gribskov, 1998, Wade et al, 2006). In this study we were able to identify a less stringent RRPE motif using the FIMO program, which searches for individual matches to a motif without using E values (Grant et al, 2011). The less stringent RRPE motif found through FIMO has the sequence ‘TGAAAAAATTTC’ and a p-­‐value of 2.69x10-­‐5, a substitution of 2 bases when compared to the RRPE consensus motif. In order to assess whether any of these sequences were responsible for MRX12’s response to MPP10, mutations were created in the less stringent RRPE motif (ls RRPE), the TATA-­‐like element, and the Abf1/Med8 binding site through the insertion of a pGSKU cassette through delitto perfetto (Figure 21). 59 "
3’" MRX12&
ATTGTAATAACTTACCGCCAATCGACAATTCGTAAACTTTTTCATCGAGAGTAGCTTGAG
ls#RRPE#motif#
AAAGTACCACTAC
*TTTAAAAAAGTAGATGAATGAATTAGTCACCAGATGACTTTGTCGG
1"
CTTAAG
TATA.like#element#
AAGCCCATTACTTTGCTCGCCATTGCGAGTGTTTAAAAAGAAAAATGCTTGTTTTATATT
CTCGA
TA*AATTATGATACTATTTTAGACGCCACTCTGCCTTTTCCTATTCTTGATCTTTTCATAA
2
"
G
TAATCTCAATTCTATACTTGACAGGTAAATTGATACCAACTTCTTTTTTTCTCGGTCCTA
Abf1/Med8p#binding#sites#
CTTTTAATACTTGTTAT
*TAAAGCATATAAAGCTGAGTTACATGATTGACATGTCATGGG
3" CTCGAGCGC
SAG1&"
CAAAATCTTATCGTCCATGCTGTTTTCGTC 5’"
Figure 21: pGSKU insertion sites to mutate cis-­‐sequences in MRX12 promoter The promoter region of MRX12 with the proposed mutations to the ls RRPE motif, TATA-­‐like element, and Abf1/Med8 binding site. The three purple stars represent the pGSKU insertion sites at 67 bp upstream of the ATG, 175 bp upstream of the ATG, and 310 bp upstream of the ATG respectively. Each insertion site is targeted to mutate a specific consensus sequence, which are notated in green. The restriction enzyme sites that replace the consensus sequences are notated in red. The start and stop codons of MRX12 and SAG1 respectively are marked with a thick underline. The pGSKU cassettes were inserted through the delitto perfetto transformation protocol on two different background strains: wild-­‐type (yMM13) and a strain in which the coding region of MRX12 had been replaced with LEU2 (yMM597). The yMM597 strain was used so that a construct with the promoter mutations could then be moved to other locations, through the use of LEU2 as a counterselectable marker. This would allow us to investigate the affects of MRX12 promoter mutations in other genetic loci. This transformation created 6 intermediate strains (yMM603-­‐yMM608) that have the pGSKU cassette 60 inserted in 3 different mutation sites in two strain backgrounds (Figure 22). pGSKU$
pGSKU$
yMM604$
MRX12&
yMM606$
67$bp$
175$bp$
310$bp$
LEU2&
67$bp$
pGSKU$
MRX12&
67$bp$
175$bp$
175$bp$
310$bp$
yMM603$
pGSKU$
310$bp$
LEU2&
67$bp$
175$bp$
pGSKU$
310$bp$
yMM605$
pGSKU$
yMM608$
MRX12&
67$bp$
175$bp$
310$bp$
yMM13
LEU2&
67$bp$
175$bp$
310$bp$
yMM607$
yMM591
Figure 22: Structure of the intermediate strains used to create mutations in the MRX12 promoter The relative structure of the six intermediate strains. The three pGSKU insertion sites are represented in green (less stringent RRPE motif, 67 bp upstream of the ATG), red (TATA-­‐like element, 175 bp upstream of the ATG), and the purple/blue rectangle (putative Abf1p binding site and Med8p binding site, 310 bp upstream of the ATG). The cassettes are inserted in two background strains: yMM13 and yMM591. After the insertion of pGSKU cassettes into the three distinct mutation sites in two different background strains, a CRISPR/Cas9 transformation was performed on the wild type based strains: yMM606 and yMM608 to replace the cassettes with healing DNA fragments containing the desired mutations. The transformation on yMM604 was not performed due to contamination of the intermediate strain. On the strains in which the coding region for MRX12 has been replaced with LEU2 (yMM603, yMM605, and yMM607), CRISPR/Cas9 cannot be performed due to the presence of LEU2, so the mutations are to be inserted through delitto perfetto. The healing fragments produced to replace the pGSKU cassettes were created from oligonucleotides that were turned into 61 double stranded DNA through PCR. The healing fragments contain a restriction enzyme cut site replacing the sequence of interest, resulting in a 120 bp DNA fragment containing the mutation in place of the mutation target site (Figure 23). The restriction enzyme cut site allows for a secondary test to ensure that the cell has taken up the mutation, as a digest can be performed and the resultant bands can be visualized through gel electrophoresis. Mutation site 1: Original(Sequence:(
5’& TAGCTGTTAAGCATTTGAAAAAGTAGCTCTCATCGAACTCTTTCATGGTGATGAAATTTTTTCATCTACTTACTTAATCAGTGGTCTACTGAAACAGCCTTCGGGTAATGAAACGAGCGG 3’&
3’& ATCGACAATTCGTAAACTTTTTCATCGAGAGTAGCTTGAGAAAGTACCACTACTTTAAAAAAGTAGATGAATGAATTAGTCACCAGATGACTTTGTCGGAAGCCCATTACTTTGCTCGCC 5’&
!
ΔRRPEmut1w(and(ΔRRPEmut1c:(
! 5’&
TAGCTGTTAAGCATTTGAAAAAGTAGCTCTCATCGAACTCTTTCATGGTGATGAAATTGAATTCTCTACT
TACTTTAACTTAAGAGATGAATGAATTAGTCACCAGATGACTTTGTCGGAAGCCCATTACTTTGCTCGCC 3’&
Mutation site 2: Original(Sequence:(
5’& CTTCGGGTAATGAAACGAGCGGTAACGCTCACAAATTTTTCTTTTTACGAACAAAATATAAATTTAATACTATGATAAAATCTGCGGTGAGACGGAAAAGGATAAGAACTAGAAAAGTAT 3’&
3’& GAAGCCCATTACTTTGCTCGCCATTGCGAGTGTTTAAAAAGAAAAATGCTTGTTTTATATTTAAATTATGATACTATTTTAGACGCCACTCTGCCTTTTCCTATTCTTGATCTTTTCATA 5’&
!
ΔTATAmut2w(and(ΔTATAmut2c:(
! 5’&
CTTCGGGTAATGAAACGAGCGGTAACGCTCACAAATTTTTCTTTTTACGAACAAAAGAGCTCTTTAATAC
TGTTTTCTCGAGAAATTATGATACTATTTTAGACGCCACTCTGCCTTTTCCTATTCTTGATCTTTTCATA 3’&
Mutation site 3: Original(Sequence:(
5’& TCCATTTAACTATGGTTGAAGAAAAAAAGAGCCAGGATGAAAATTATGAACAATAATTTCGTATATTTCGACTCAATGTACTAACTGTACAGTACCCGTTTTAGAATAGCAGGTACGACA 3’&
3’& AGGTAAATTGATACCAACTTCTTTTTTTCTCGGTCCTACTTTTAATACTTGTTATTAAAGCATATAAAGCTGAGTTACATGATTGACATGTCATGGGCAAAATCTTATCGTCCATGCTGT 5’&
!
ΔABFmut3w(and(ΔABFmut3c:(
! 5’&
TCCATTTAACTATGGTTGAAGAAAAAAAGAGCCAGGATGAAAATTATGAACAATAAGAGCTCGCGTTTCG
GTTATTCTCGAGCGCAAAGCTGAGTTACATGATTGACATGTCATGGGCAAAATCTTATCGTCCATGCTGT 3’&
Figure 23: Schematic of healing fragments to replace pGSKU cassettes and generate cis-­‐mutations in MRX12 promoter The healing fragments created to replace the pGSKU cassettes in intermediate strains yMM603 through yMM608. Each figure shows the original dsDNA sequence and healing oligonucleotide created to insert a restriction enzyme cut site in place of a cis-­‐sequence. The blue nucleotides represent the consensus sequence of the motif to be mutated, the green nucleotides represent the overlap of the oligonucleotides, and the red nucleotides represent the restriction enzyme cutsite sequence. For the third mutation site, the pink bases are additional bases changed to ensure a full deletion of the consensus motif. In order to select for transformant colonies that had taken up the desired mutation, transformants were plated onto selective media and tested via PCR 62 and subsequent restriction digest. CRISPR/Cas9 tranformations were performed on srains yMM606 and yMM608, replacing the respective TATA-­‐like element and Abf1/Med8 binding sites with SacI restriction enzyme cut sites (creating strains yMM612 and yMM613 respectively). After confirmation of the integration of the mutation through a SacI restriction digest, the cells were treated with heat shock and subsequent RTqPCR. In strain yMM612, upon heat shock, MRX12 appears to maintain typical RRB expression, implying that the mutation in the ‘TATAA’ sequence has no effect on the regulation of MRX12 (Figure 24). For yMM613, the deletion of the Abf1p and Med8p binding motifs seems to abrogate AGC, resulting in the loss of repression of MRX12 after heat shock. These results helped to define the regions of the MRX12 promoter that are responsible for its regulation. In particular, the phenotype of the ΔAbf1/Med8 mutant suggests new roles for Abf1p and the Mediator complex in the transcriptional co-­‐regulation of the MPP10-­‐MRX12 gene pair. 63 RP
!! !! !!MPP10
120!
Rela%ve'Expression'
EBP2(
MPP10(
MRX12(
60!
20!
0!
0!
!! !! !!MPP10
175bp!
80!
40!
RP
B:&
yMM612&(ΔTATA)&
100!
MRX12(
120!
MRX12(
310bp!
yMM613&(ΔAbf1/Med8)&&
100!
Rela%ve'Expression'
A:&
80!
60!
40!
20!
0!
5!
10!
Time'
20!
30!
0!
5!
10!
Time''
20!
Figure 24: Mutation of Abf1p binding site, but not TATA-­‐like element in MRX12 promoter abrogate AGC The schematic and relative expression of EBP2, MPP10, and MRX12 in the ΔTATA mutant (yMM612) (A) and the ΔAbf1p/Med8p mutant (yMM613) (B). In both strains the consensus motif has been replaced with a SacI restriction enzyme cut site (light red X). Data generated from 1 biological and 3 technical replicates. Discussion The modular nature of AGC is evolutionarily significant In order to understand the importance of adjacent gene co-­‐regulation, we must first try to imagine its evolutionary history by asking questions regarding how and why AGC evolved. Transcriptionally significant gene adjacency is not an isolated phenomenon, in fact, it was described in the pioneering work on the prokaryotic operon by François Jacob and Jacques Monod (Jacob and Monod, 1961). Even though AGC and the prokaryotic operon are both mechanisms used to achieve transcriptional co-­‐regulation, there are many differences between the two. In the prokaryotic operon, multiple genes are arranged in a tandem 64 30!
orientation and they lack individual promoter regions, instead they are regulated by a single promoter and operator region. The genes in the operon are co-­‐transcribed, creating a single mRNA, which then goes through translation to create distinct proteins (Pardee et al. 1959). Due to this, the group of genes is regulated as a set, creating a common state of transcriptional co-­‐regulation. In AGC, co-­‐regulated genes are oriented in pairs, but can be present on either strand in any orientation. Each gene retains its endogenous genomic promoter and is transcribed on its own. However the pair is still transcriptionally co-­‐
regulated, possibly due to a DNA looping mechanism that allows the promoter regions to interact. The differences between the two co-­‐regulatory systems are striking, as AGC allows for much more fluidity regarding which genes are co-­‐regulated and under which circumstances. Since AGC does not require the loss of individual promoter regions, genes can simply be swapped into pairs to create novel co-­‐
regulated states. In contrast, in order to add a new gene to an operon, that gene must lose its promoter region and be inserted on the same strand in the same direction. In addition, once a gene is added to an operon it can never be removed, as it has lost its endogenous promoter and would not be able to function autonomously. Due to this, and the co-­‐transcription of operon gene members, the operon is an extremely stable system that does not allow for fluidity in gene members. The apparent flexibility in the gene members of AGC has been 65 documented through the comparison of paired RRB genes across divergent fungal lineages. Of the 22 RRB pairs found in S. cerevisiae, 20 are conserved in at least one closely related fungal species, however multiple gene pairs have been broken, either existing as single genes or in pairs with other RRBs (Figure 25). The MPP10-­‐MRX12 gene pair is conserved in S. mikatae and S. bayanus, however both genes are found unpaired in S. paradoxus, C. glabrata, C. albicans, and C. dubliniensis. In K. waltii, S. kluyveri, K. lactis, and S. pombe, MRX12 and MPP10 are found paired to different RRB genes. Both the presence of conserved gene pairs and the observation that multiple RRB gene pairs have been swapped in S.#cerevisiae#
#
S.#paradoxus#
#
S.#mikatae#
#
S.#bayanus#
#
C.#glabrata#
#
K.#waltii#
#
S.#kluyveri#
#
K.#lactis#
#
C.#albicans#
#
C.#dublinienus#
#
S.#pombe##
#
divergent fungal species suggests that there is fluidity in the relative organization of RRB genes. Schizosaccharomyces#pombe#
NSA2>LCP5#
#
RRP4>TRM5#
Candida#dubliniensis#
#
NIP7>PUS1#
#
Candida#albicans#
NOP2>GCD10#
#
Candida#glabrata#
RSA3>UTP13#
#
Saccharomyces#bayanus#
FAP7>NRP1#
#
RIX7>GRC3#
Saccharomyces#mikatae#
#
RRP3>SSF1#
Saccharomyces#cerevisiae#
#
DBP2>RPC19#
Saccharomyces#paradoxus#
#
NOG2>ESF2#
Kluyveromyces#waltii#
#
DBP8>NMD3#
#
Saccharomyces#kluyveri#
RRP15>NOC4#
#
#
MPP10>MRX12#
Kluyveromyces#lactis#
#
RPF1>GAR1#
#
NOC2>RET1#
#
NIP1>YMR310C#
#
MTR3>NSR1#
#
Same%adjacent%pairing%
PWP1>NOP56#
#
Both%genes%paired%to%another%partner%
BFR2>PRO1#
One%gene%paired%with%another%partner%
#
YNL247W>RPA49#
Both%genes%are%unpaired%
#
NOC3>CMS1#
Homologue%not%present%
#
DIS3>TSR4#
#
66 Figure 25: S. cerevisiae RRB gene pairs are conserved across divergent fungal species. The phylogenetic relationship of divergent fungal lineages and heat map depicting the degree of conservation of RRB pairs across fungal species. (Figure adapted from Arnone et al. 2012) The hypothesis that AGC is a modular phenomenon is supported both by evolutionary trends and the creation of a co-­‐regulated pair between FUI and MRX12. If MRX12 were not co-­‐regulated by FUI1 in this position, we would expect MRX12 to maintain expression upon heat shock, as it is when the MPP10-­‐
MRX12 gene pair is broken through the insertion of a transcriptionally active LEU2 gene. Further analysis on the expression level of MRX12 when it is positioned between FIT2 and FIT3 should provide insights into the regulation of MRX12 when it is separated from other members of the RRB regulon. In order to explain why AGC evolved in eukaryotes and the operon system evolved in prokaryotes, it is important to think of the differences in DNA structure between the two life forms. In prokaryotes, DNA is not condensed into chromatin and therefore transcriptional regulation through chromatin modification does not exist. Since we have found that chromatin interacting proteins such as Spt20p, Chd1p and Snf2p are required to maintain co-­‐
regulation of MPP10 and MRX12, it is possible that AGC evolved after the evolution of higher order DNA structures (Arnone et al, 2014). The proposed model of AGC involves DNA looping, which requires chromatin remodelers and other trans-­‐acting factors in order to hold the DNA in the correct conformation. It is possible that AGC represents an evolutionarily older method of controlling 67 transcription solely through genomic organization, where genes are swapped in in and out of pairs to control expression levels easily (as opposed to creating new transcription factors or regulatory networks). While adjacent gene pairing is present in higher-­‐order eukaryotes, it is not nearly as prevalent as it is in yeast. It is possible that AGC is a progenitor of enhancer regions, which are not present in yeast but are in higher-­‐order eukaryotes. Enhancers are regions of distal DNA sequences that function in cis to increase transcription of their target gene (Mora et al, 2015). Enhancers can exist anywhere upstream or downstream of their target gene, and can be as far as 100 Mbp away. Enhancers interact with the core promoter of genes through DNA looping to physically localize trans-­‐acting factors and enhance transcription. AGC could represent an evolutionarily earlier form of enhancers, where gene regulatory regions are physically co-­‐localized in order to regulate transcription. In higher-­‐order eukaryotes, AGC may have evolved to be independent of distance, such that genes no longer needed to be immediately adjacent to each other, but instead could interact from distal regions. We can speculate on the possible evolutionary history of AGC, however this study has concretely shown that adjacent gene co-­‐regulation is a modular phenomenon, in which genes can be swapped in and out of pairs to create novel regulatory states. These experiments show that immediately adjacent co-­‐
regulated pairs can be uncoupled by the presence of a transcriptionally active gene between them, and also created by matching genes with similar molecular 68 functions and conserved promoter motifs. The modular nature of AGC has also been demonstrated through studies on diverse eukaryotic species, which exhibit a conservation of gene pairing in many regulons. Supplemented with the experimental data presented in this study, we have characterized a novel, flexible system for the co-­‐regulation of large regulons. The relationship between paired genes could be explained by promoter co-­‐localization The DNA looping model holds significant explanatory power for the co-­‐
regulation of immediately adjacent paired genes. In yeast, short-­‐range DNA loops have been found to mediate individual gene expression and recent studies have shown DNA loops can localize regions containing 1-­‐5 genes (O’Sullivan et al. 2004, Hseih et al. 2015). In the case of AGC, the proposed looping mechanism is not dependent on cis-­‐sequences in the open reading frame, as AGC has been shown to be ORF-­‐independent (Arnone et al, 2014). ORF-­‐independent looping supports the idea that paired, adjacent promoter sequences are co-­‐localized to mediate transcriptional coupling. A looping strategy would allow cis-­‐mutations made in the promoter of one gene to influence the juxtaposed promoter of the adjacent gene, resulting in the co-­‐regulation of the two genes. This could account for the relationship between MPP10 and MRX12, in which mutations in the conserved PAC and RRPE motifs affect the expression level of both genes. Mutations made in the promoter of MRX12 to assess the relationship 69 between co-­‐regulated paired genes provide insights into the nature of the interaction between MPP10 and MRX12. The ΔTATA mutant (yMM612) suggests that the TATA-­‐like element located 175 bp upstream of the ATG in the promoter of MRX12 is not required for appropriate repression of MRX12. This result is consistent with the DNA-­‐looping hypothesis, as since the TATA-­‐like element in the MPP10 promoter remains intact, it could be enough to recruit the basal transcriptional machinery to both promoter regions, resulting in the transcription of both genes. It seems clear from this result that the TATA sequence in the MRX12 promoter is not responsible for the interaction between the two promoter regions, nor is it required for typical RRB repression of MRX12. The ΔAbf1/Med8 mutant (yMM613) phenotype suggests that the predicted Abf1 and Med8 binding sites located 310 bp upstream of the ATG in the MRX12 promoter are required for the co-­‐regulation of MRX12. This result implies that these cis-­‐sequences may be responding to the regulatory information sent from MPP10. In yeast, Abf1p has been found to bind to many RP and rRNA gene regulatory regions, and can act as a repressor or activator depending on the binding context (Planta, 1997). In this case, is possible that Abf1 acts as a repressor of MRX12, such that the removal of the Abf1p binding site results in the loss of repression of MRX12. Abf1p has also been identified as having a role in rendering adjacent cis-­‐sequences available to other trans-­‐factors through influencing the chromatin structure (Planta et al, 1995). Functioning as a facilitator of other trans-­‐factors, Abf1p could allow for other proteins to 70 associate with the coupled promoters, suggesting a role for Abf1p in AGC. This finding supports our current model of AGC, as it is clear that Abf1p is involved in maintaining the co-­‐regulation between the MPP10-­‐MRX12 gene pair. Med8p, an essential component of the Mediator complex, is required to form the RNA polymerase II holoenzyme in order to initiate transcription (Kornberg, 2005). It is possible that Med8 may be required to physically co-­‐
localize the promoters of MRX12 and MPP10, as the Mediator complex has been shown to play a role in DNA looping (Hsieh et al, 2015). In the case of MRX12 expression, it is possible that mutation of the Med8p binding site impedes the interaction of part of the Mediator complex with the MRX12 promoter, resulting in the uncoupling of MRX12 from the transcriptional repression of MPP10. The results from the ΔTATA and ΔAbf1/Med8 mutants are both generated from one biological replicate and need to be verified by additional replicates in order to make definitive statements about the affect of these promoter mutations on the expression levels of MRX12 and MPP10. However, we can infer new insights into the possible mechanism of AGC based on the data already collected, namely the large role of Abf1p in achieving transcriptional co-­‐
regulation (Figure 26). In this new model, Abf1p represses MRX12 and may also play a role in facilitating additional trans-­‐factors in the DNA loop. This model also adds a role for the Mediator complex in its interaction with the Med8p binding site in the MRX12 promoter, and may be responsible for transcriptional initiation of the paired genes. From this study, we have also characterized that 71 the TBP present in the MPP10 promoter may facilitate transcription of MRX12, as deletion of the TATA-­‐like element in the MRX12 promoter does not affect its expression. Rpd3%
Dot6/Tod6%
Stb3%
R P
!! MPP10
!! !! !!
Swi2/Snf2%
TBP%
Mediator%
SAGA%
Chd1%
Med8% Abf1%
!!
MRX12&
!!
Figure 26: Updated molecular model of AGC The promoters of MPP10 and MRX12 may be brought together by DNA looping mediated by the SAGA complex, Swi/Snf, Mediator, and Abf1p. The physical co-­‐
localization of both promoters allows simultaneous transcriptional regulation of both genes by the PAC and RRPE binding factors, as well as Abf1p, targeting the promoter of MPP10. The TBP recruited to the MPP10 promoter may also be responsible for transcriptional initiation of MRX12. Further mutation of cis-­‐sequences in the promoter of MRX12 may identify additional essential cis-­‐sequences involved in AGC. These mutations may also identify key DNA binding factors that may play a role in DNA looping or paired transcriptional co-­‐regulation. Additionally, mutations in the promoter of MRX12 may also affect the regulation of MPP10, suggesting that the co-­‐regulatory state is not one sided, but instead involves input from both promoter regions. This result would be consistent with the DNA looping model, as the physical co-­‐
72 localization of the promoter regions would allow for interactions between trans-­‐
factors, which could be recruited to cis-­‐sequences present in either promoter. In order to confirm that DNA looping is in fact the mechanism of AGC, more experiments would need to be performed assessing whether the promoter regions of paired genes are physically brought together. Due to the proximity of the two promoter regions, experiments such as 3C (chromosome conformation capture) cannot be performed with high confidence. However, recent innovations such as Micro-­‐C, which utilizes a MNase treatment instead of restriction enzyme digest to fragment the DNA, may be able to provide the resolution necessary to image the interaction between the promoter regions of paired genes (Mozziconacci and Koszul, 2015). Hopefully future experiments such as Micro-­‐C or additional cis-­‐ and trans-­‐factor mutants could definitively prove DNA looping as the mechanism of AGC. Taken altogether, this study has clarified the nature of the interaction between paired genes through site-­‐directed mutagenesis in the promoter of MRX12. Our findings have identified novel roles for cis-­‐ and trans-­‐acting sequences in AGC that are consistent with the DNA looping model of AGC. More experiments need to be performed in order to clarify the modular nature of AGC as well as define the mechanism by with adjacent genes are co-­‐regulated. However, this study has identified novel players in the co-­‐regulation of adjacent genes and proven that AGC is a modular system in which genes can be swapped in and out of pairs to change transcriptional regulatory states. 73 References Abu-­‐Jamous B, Fa R, Roberts DJ, and Nandi AK. Comprehensive analysis of forty yeast microarray datasets reveals a novel subset of genes (APha-­‐RiB) consistently negatively associated with ribosome biogenesis. BMC Bioinformatics. 2014 Sep 29;15:322. doi: 10.1186/1471-­‐2105-­‐15-­‐322. 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