Thesis
Reference
The transcription factor Sfp1 regulates growth and division in the yeast Saccharomyces cerevisiae
TOMASSETTI, Susanna
Abstract
The Split Finger Protein 1 (Sfp1) is an yeast nutrient- and stress-sensitive transcription factor that controls for the expression of growth related genes involved in ribosome biogenesis, rRNA processing and translation. Sfp1 was early recognized as an unusual transcription factor that occupies the promoter of a reduced set of its target genes and its mechanism of action have been remained elusive. Sfp1 ChIP-seq experiments performed in this thesis revealed that Sfp1 may play a direct role in the regulation of RNAPII recruitment during PIC formation and this control does not necessarily require Sfp1 promoter occupancy. We have found that SFP1 positively affects the expression of growth related genes whereas it has a negative effect on stress-sensitive genes expression. Finally, we have described that Sfp1 occupies some cell cycle-dependent gene in a cell cycle dependent manner, suggesting a key role of this transcription factor in the intricate process that maintains cell size homeostasis.
TOMASSETTI, Susanna. The transcription factor Sfp1 regulates growth and division in the yeast Saccharomyces cerevisiae. Thèse de doctorat : Univ. Genève, 2016, no. Sc. 4960
URN : urn:nbn:ch:unige-862792
DOI : 10.13097/archive-ouverte/unige:86279
Available at:
http://archive-ouverte.unige.ch/unige:86279
UNIVERSITE DE GENEVE
Département de biologie moléculaire FACULTE DES SCIENCES Professeur David Shore
The transcription factor Sfp1 regulates growth and division in the yeast Saccharomyces cerevisiae
THESE
présentée à la Faculté des sciences de l’Université de Genève pour obtenir le grade de Docteur ès Sciences, mention biologie
par
Susanna Tomassetti
de
Torri in Sabina (Italie)
Thèse n°4960
Atelier d’impression ReproMail 2016
Résumé
La protéine de levure Saccharomyces cerevisiae Sfp1 (Split Finger Protein 1) est un régulateur majeur de la croissance cellulaire s’adaptant aux stress et aux conditions extracellulaires. Dans des conditions favorables, Sfp1 est localisée principalement dans le noyau ou elle peut contrôler l’expression des gènes de croissance (les gènes impliqués dans la biogenèse des ribosomes, la maturation des ARNr ou encore les gènes permettant la traduction). En réponse à un stress ou dans des conditions de croissance devenant défavorable Sfp1 est alors rapidement délocalisée du noyau vers le cytoplasme conjointement à la diminution de l’expression des gènes de croissance. Malgré son rôle clefs, dans la croissance et la division cellulaire, le mécanisme d’action de Sfp1 pour réguler la transcription est encore incomprise et semble différer des facteurs de transcriptions classiques.
Dans cette thèse, des expériences de ChIP-seq ont été menées dans le but d’obtenir une information détaillée sur la manière dont Sfp1 régule le programme transcriptionel. Pour cela, nous avons utilisé deux approches différentes nous permettant à la fois de suivre le recrutement de Sfp1 et la distribution de l’ARN polymerase II (RNAPII) après l’augmentation ou la déplétion de Sfp1. Nous avons ainsi trouvé que Sfp1 joue un rôle direct dans le recrutement de l’ARNPII sur le complexe d’initiation (PIC) sans pour autant nécessiter la présence de Sfp1 sur le promoteur : en effet, Sfp1 est détectée sur le promoteur mais de manière plus surprenante également sur les région codantes (ORF). La surexpression de Sfp1 ou sa déplétion peut à la fois augmenter ou diminuer l’expression de ses gènes cibles, ainsi nos données indiquent que Sfp1 stimulerait les gènes de croissance et réprimerait l’expression des gènes de stress. De manière intéressante, nous avons observé que ces gènes régulés positivement ou négativement par Sfp1 sont enrichies en TBP (TATA binding protein) suggérant que Sfp1 pourrait réguler, au moins en partie, la distribution de la TBP en fonction des conditions extracellulaires et des processus d’assemblage du PIC.
Enfin, nous avons décris pour la première fois que la protéine Sfp1 est recrutée sur certains gènes en fonction du cycle cellulaire. Nous avons notamment identifié les gènes codant pour les cyclines G1, CLN1 et CLN2 qui sont connus pour être des facteurs essentiels à la transition G1/S, une phase du cycle cellulaire connue sous le nom de Start chez la levure.
Finalement, même si Sfp1 n’est pas une protéine essentielle, elle pourrait être un composant déterminant permettant la coordination entre division et croissance permettant de maintenir l’homéostasie cellulaire.
Table of contents
General Introduction ... 9
Studying cell physiology: size under the spotlight ... 9
Cell size regulation ... 9
The cell cycle ... 11
Active size measurement coordinates growth and division to maintain size homeostasis 17 Protein synthesis and ribosome biogenesis: a proxy for cell growth ... 19
General aspects of ribosome production ... 19
Ribosome synthesis and size homeostasis are tightly coupled processes ... 23
Nutrient sensing and signaling pathways controlling cell growth ... 24
Glucose signaling ... 24
Nitrogen regulation of growth and development ... 27
DNA-dependent RNA polymerases mediate genome transcription ... 30
Transcription mediated by RNAPI and III ... 31
The transcription mediated by RNAPII ... 38
Sfp1 is a nutrient- and stress-sensitive transcription factor involved in growth control ... 43
Results ... 75
Chapter I: Sfp1 is a transcriptional activator linked to cell growth ... 75
Introduction ... 75
Sfp1 regulates gene expression by controlling RNAPII recruitment ... 77
Genome-wide analysis reveals Sfp1 binding at both promoters and gene bodies ... 83
Genome-wide analysis of the Sfp1-dependent changes in RNAPII distribution ... 91
Sfp1 target genes display high levels of TBP promoter binding ... 98
Evidence that Sfp1 acts together with gene specific activators at some promoters ... 100
Not only RNAPII: Sfp1 associates with RNAPIII and may be involved in class III gene
expression ... 120
Chapter II: Sfp1 may be involved in “Starting” the cell cycle... 127
Introduction ... 127
Sfp1 is recruited to CLN1 and CLN2 promoters in a cell cycle dependent manner ... 129
Nuclear depletion of Sfp1 delays cell cycle progression ... 133
Chapter III: Sfp1 phosphorylation changes rapidly in response to environmental signals ... 138
Introduction ... 138
Sfp1 phosphorylation level signals extracellular changes ... 141
Discussion ... 151
Promoters that are occupied by Sfp1 have specific structural characteristics ... 153
Sfp1 and gene specific activators may act together to properly control the expression of some genes ... 157
Sfp1 may follow RNAPII during gene transcription ... 159
Sfp1 regulates gene expression by controlling RNAPII recruitment ... 162
Sfp1 can either increase or down regulate specific genes expression ... 163
Sfp1 target genes are enriched for TBP ... 165
TBP is strongly bound on RNAPIII transcribed genes that may also be Sfp1 targets ... 172
RNAPII recruitment on RiBi gene promoters is particularly sensitive to Sfp1 ... 173
Sfp1 transcriptional activity promotes growth and controls the cell cycle ... 176
Cell growth and viability ... 176
Sfp1 is phosphorylated on multiple sites in a highly dynamic manner ... 184
Sfp1: a speculative model for its mechanism of action and the regulation of its subcellular localization ... 192
Tables ... 229
Experimental Procedures ... 259
Yeast strains and growth conditions ... 259
Chromatin immunoprecipitation and mRNA purification ... 259
Chromatin immunoprecipitation ... 259
ChIP-sequencing ... 260
Steady state mRNA measurement ... 261
Protein extraction and western blot analysis ... 262
TCA-Urea protein extraction and Phos-tag method ... 262
Co-immunoprecipitation ... 263
CHX pulse experiment ... 263
Viability Assays ... 263
Cell synchrony experiments ... 263
Cell and colony size measurement ... 264
General Introduction
Studying cell physiology: size under the spotlight
A simple “small room”, the cellula (Latin for cell). This was the original definition of the unifying form of life. Because of its small size, it originally escaped notice of life scientists. It was only after the invention of the microscope and its subsequent technological improvement that the cell became visible. It was soon recognized as the simplest component of both plant and animal tissues, as well as being an independent form of life. Upon the first cell observation by the English scientist Robert Hooke (1635-1703) in cork slices, a large variety of cell types and species started to be described in parallel with microscope technology developments. The field of cell biology prospered consequently, leading to new and exciting discoveries about cell structures and functions. In modern times an enormous cell variability has been described and our starting “small room” appears to be a complex assembly of hidden passages and doors that still have to be opened.
Cell size regulation
Size is one of the most obvious structural characteristics of a cell, and one of the first to be analyzed. Different cell types vary greatly in size, ranging from smallest known eukaryote Ostreococcus tauri, at ~1µm (Palenik et al., 2007) to the ~1mm frog oocyte (Figure 1; (Wallace and Selman, 1981)). Each cell type or unicellular organism maintains a characteristic size required to function efficiently in a given environmental context (Schmoller and Skotheim, 2015). For example, recent observations demonstrated a linear relationship between cell geometry and fitness for bacterial cells growing in a fluctuating environment (Monds et al., 2014). How big a cell can be? And why does it matter? Basic processes of cell physiology are by their nature dependent on cell size (Marshall et al., 2012): for example, nutrients as well as metabolic waste products diffuse or are actively transported across the plasma membrane and as a result, changes in surface-to-volume ratio have a profound effect on the cell
biosynthetic capacity. Therefore, because of functional constraints, cells are selected by the external conditions to maintain a characteristic size. Size control mechanisms are hence essential and they generally rely on the regulation of growth rate and on active size perception to coordinate growth and division.
Figure 1. Cell size comparison of two different eukaryotic cell types
A. Picture of the smallest known eukaryote Ostreococcus tauri (B. Palenik et al 2007). B.
Frog oocytes picture from Cristofori-Armstrong et al 2015.
A B
The cell cycle
During its life a cell duplicates the genetic information encoded by the DNA as well as the essential cellular components (like organelles, membranes, ribosomes etc.) and uses the duplicated material to form a new cell. Therefore every single organism, from prokaryotes to the multicellular mammals, is formed and maintained by repetitive cycles of growth and division. Even though the molecular mechanisms governing cell division are highly variable between cell types, universal characteristics can be described. To produce two genetically identical cells the DNA has to be precisely duplicated and segregated into the daughter cell in such a way that upon division each cell contains a copy of the entire genetic information (Figure 2A). This universal process occurs as a series of highly coordinated events whose sequential execution is achieved by specific controlling mechanisms referred as check points.
Deregulation of cell cycle progression causes genomic instability (Hartwell and Kastan, 1994) and has been extensively correlated with both hereditary and spontaneous cancer (Hall and Peters, 1996; Hartwell and Kastan, 1994; Malumbres and Barbacid, 2009). The eukaryotic cell cycle can be schematically divided into four main phases: two of them, referred as gap-phases (G1 and G2) are required to duplicate cell mass (i.e. to grow) and are the phases where a cell generally spends most of the time of its life cycle (with high time-variability observed between different cell types). Between G1 and G2 DNA synthesis occurs in a highly controlled step called S-phase (for “synthesis phase”). Once the DNA has been appropriately duplicated and organized in complex structures called chromosomes, the physical separation of nuclei and cytoplasm occurs during a phase called Mitosis (Figure 2B).
Figure 2. General aspects of the cell cycle
A. The universal characteristic of cell division is the accurate process of DNA replication and segregation to form two genetically identical cells. B. Schematic representation of the eukaryotic cell cycle as sequential process of four highly controlled and organized phases (G1,S, G2 and M). C. Modified from B. Alberts et al (Molecular biology of the cell 5/e 2008).
Schematic representation of the budding yeast cell cycle.
A B
C
The budding yeast S. cerevisiae is a versatile model system of eukaryotic genetics (Mell, 2003). Mutant screens and segregation analysis are simpler to perform in yeast than in higher eukaryotes. Moreover, upon complete sequencing of the yeast genome (Goffeau et al., 1996) the number of observed homologous genes between yeast and mammals increased (Botstein et al., 1997): ~31% of all the potential protein-encoding genes of yeast were found to have mammalian homologs, revealing a profound evolutionary conservation of essential biological functions. Furthermore, yeast have basic structural elements and chromosome organization that are highly homologous to those of plant and animal cells (Hartwell, 1974). Taken together, these observations explain why S. cerevisiae has been extensively used as a model organism for genetic and biochemical characterization of the eukaryotic cell cycle. S. cerevisiae, because of its asymmetric cell division, gives rise to a smaller daughter cell (the bud) and a bigger mother cell. Following mitosis, the bud needs to growth to a characteristic size (generally referred as the “critical size”) before entering the next division cycle (Figure 2C).
The precise temporal execution of growth, DNA replication and chromosome segregation during the cell cycle is obtained by periodic transcription of nearly 20% of the yeast genome (Guo et al., 2013; Orlando et al., 2008; Spellman et al., 1998). Therefore, progression through the cell cycle is accompanied by a dramatic reorganization of gene expression (Haase and Wittenberg, 2014) that at its most fundamental appears as a continuum of transcriptional activation and deactivation that ultimately controls the cyclin-dependent kinases (CDKs).
CDKs are proline-directed serine and threonine protein kinases whose association with specific activating subunits called cyclins regulates their activity and specificity (Bloom and Cross, 2007; Mendenhall and Hodge, 1998). Budding yeast encodes five different CDKs:
Kin28, Ssn3 and Ctk1 required for direct control of RNAPII transcriptional activity through phosphorylation of its long carboxy-terminal domain (Prelich, 2002), Pho85, controlling cell cycle progression mainly in response to intracellular phosphate levels (Jiménez, 2013), and Cdc28 (the analog of CDK1 in higher eukaryotes), the central coordinator of the yeast cell cycle (Mendenhall and Hodge, 1998). In contrast to higher eukaryotes, where multiple CDKs drive the cell cycle, Cdc28 is the only yeast CDK governing cell cycle progression and
coordinating growth and division (Bloom and Cross, 2007). Control of Cdc28 kinase activity is ensured by the accumulation of several different cyclins. Cyclin-Cdc28 complexes phosphorylate and therefore regulate the activity of specific transcription factors required for the “cell cycle-regulated transcription” (Haase and Wittenberg, 2014). In turn, the transcriptional waves controlling the cell cycle guide the activity of Cdc28 by ensuring appropriately timed expression of the specific cyclin partners, revealing a complex interplay between the transcriptional program, CDK activity and cell cycle progression (Haase and Wittenberg, 2014). The Cdc28 cyclins can be schematically classified into two groups (Mendenhall and Hodge, 1998): the G1 cyclins (from Cln1 to Cln3) primarily regulating the G1- S transition and the B-Type cyclins (from Clb1 to Clb6) expressed from late G1 to M-phase and required for DNA replication. Yeast and animals G1 cyclins, even though not phylogenetically related, carry out very similar functions and they might represent a case of convergent evolution (Bloom and Cross, 2007). Moreover, analysis of yeast B-type cyclin phylogeny reveals that cyclin differentiation occurred following several rounds of gene duplication (Archambault et al., 2005) and is likely to have conferred important fitness advantages compared to the hypothetical “single-cyclin” ancestor (Nasmyth, 1995). Indeed, partial overlap of cyclin function establishes regulatory flexibility: for example, CLB5 overexpression rescues the viability of cells lacking all three G1 cyclins (Bloom and Cross, 2007). The specificity of cyclins, and therefore of Cdc28 activity, is obtained in different means:
by controlling the expression of genes encoding the different cyclins (Bertoli et al., 2013;
Mendenhall and Hodge, 1998), by direct proteasome-mediated degradation of a specific cyclin that is no longer required, or by inhibition of cyclin-Cdc28 complexes using specific inhibitors called Cyclin Kinase Inhibitors (CKIs) (Mendenhall and Hodge, 1998). Moreover, differential subcellular localization has been demonstrated for different cyclin-Cdc28 complexes and is mainly required for the morphogenetic events associated with the G1-S transition (Bailly et al., 2003; Bloom and Cross, 2007; Moffat and Andrews, 2004). Finally, substrate specificity of cyclins allows Cdc28 to control for different processes: for example, experiments where a
cyclin was ectopically express in a cell-cycle phase in which it is not normally present demonstrated that cyclins do possess intrinsic substrate specificity (Erlandsson et al., 2000).
Yeast cells commit to enter a new mitotic cycle during G1 the cell cycle phase were most of the growth program takes place. The activation of cyclin-Cdc28 dependent transcription and therefore cell cycle entrance, is an irreversible decision that is taken by the cell upon achievement of a specific size generally referred as “critical size” (Johnson and Skotheim, 2013). This size-dependent control of cell cycle entrance occurs in daughter cells and requires coordination between growth and division. The all-or-non decision to divide is taken at Start, the G1 checkpoint beyond which the cell will proceed through the cell cycle despite subsequent changes in extracellular conditions (Johnson and Skotheim, 2013). The mechanism controlling Start has been extensively studied and involves a positive feedback loop that ensures the irreversible progression through the cell cycle (Bertoli et al., 2013). The events at Start are highly conserved from yeast to humans (Cobrinik, 2005; Johnson and Skotheim, 2013; Weinberg, 1995). Two sequence-specific heterodimeric transcription factors SBF (SCB-binding factor) and MBF (MCB-binding factor; (Bean et al., 2005; Koch et al., 1996) are required to exploit the G1-S gene expression program and are directly regulated by Cdc28 kinase activity (Lowndes et al., 1991; Primig et al., 1992). They are composed of a shared subunit encoded by SWI6 and required for transcription activation and by the specific DNA- binding subunits encoded by SWI4 and MBP1 respectively (Andrews and Herskowitz, 1989;
Bean et al., 2005; Koch et al., 1996; Primig et al., 1992). SBF-activated genes are predominantly involved in budding and include CLN1 and CLN2, whereas MBF targets mostly regulate transcription of DNA replication genes (Bloom and Cross, 2007; Haase and Wittenberg, 2014; Mendenhall and Hodge, 1998). About 300 genes have been collectively found as SBF and MBF targets (Harris et al., 2013; Iyer et al., 2001). Early in G1, SBF associates at the specific consensus sequence of its target genes. At this time the physical interaction with the negative regulator Whi5 prevents SBF-mediated transcriptional activation (Bertoli et al., 2013). When growth requirements are met, the most upstream G1 cyclin, Cln3,
is released from the endoplasmic reticulum (where it is maintained in an inactive state both as untranslated mRNA and in complex with Cdc28) and accumulates in the nucleus as a complex with Cdc28 (Caudron and Barral, 2013; Gari et al., 2001; Verges et al., 2007; Wang et al., 2004a; Yahya et al., 2014). In the nucleus, the main function of the Cln3-Cdc28 complex is to phosphorylate and partially inactivate the negative regulator Whi5, causing an initial low level of SBF-mediated transcription leading to Cln1 and Cln2 cyclin production (Costanzo et al., 2004). Cln1/2 can then associate with Cdc28, further stimulating their own expression through complete Whi5 inactivation. Once established, the Cln1/2 positive feedback loop boosts SBF target gene expression, thus ensuring irreversible entrance into the cell cycle (Figure 3; (Bean et al., 2006; Skotheim et al., 2008). The cell cycle engine is now activated, and through cyclin specificity and Cdc28 kinase activity, the downstream events of budding and morphogenesis, Spindle Pole Body (SPB) duplication, mitotic spindle formation and chromosomes segregation occur under the control of specific checkpoints that unsure correct completion of one phase before starting the next one (Bloom and Cross, 2007).
Figure 3. The G1/S regulatory network ensures irreversible cell cycle entry
Figure adapted from Schmoller et al 2015. Schematic representation of the main components of the G1/S regulatory network. Blue repressing-arrows refer to negative regulation; Green arrows refer to positive regulation; Black arrows represent transcription of the indicated genes.
Active size measurement coordinates growth and division to maintain size homeostasis
Doubling of cell mass with each division ensures size homeostasis. A cell population of any particular type shows a characteristic size distribution caused by physiological size variation over successive generations (Jorgensen and Tyers, 2004; Turner, 2012). This is particularly evident for budding yeast populations characterized by high size heterogeneity. Mother and daughter cells differ greatly in size, and spontaneous size variability of newborn cells is observed upon mitosis. Moreover, when the size distribution is modified upon changes in the extracellular conditions (for example as a consequence of starvation), the normal distribution is promptly restored after the insult has been removed (Johnston et al., 1977). This strongly suggests that size distribution is under homeostatic control (Mitchison, 1972). Size sensing mechanisms are therefore essential for the cell to perceive its own size and to use this information to coordinate growth and division.
How might cells measure size? Whereas the molecular mechanisms governing cell cycle progression are now well defined, much less is known about how cell size is sensed.
According to cell type different size sensing mechanisms have been observed (Amodeo et al., 2015): a geometric-based mechanism to measure size can be proposed for cell types characterized by little cell-to-cell variability and by a homogeneous shape. For example, the surface-to-volume ratio may represent a good means to geometrically determine size. In multicellular organisms general tissue characteristics like edges, junctions or contact target sites, impose on the single cell a spatial constraint that could determine its size. Finally, a titration-based size sensing mechanism has also been described whereby a “constant- concentration molecule” is measured over an “internal sensor molecule”. S. cerevisiae has been proposed to use such a system, and recent observations have provided new and interesting details about the molecular mechanism controlling yeast cell size. During the pre- replicative G1 phase of a daughter cell, the growth program takes place and therefore the size sensing mechanism is operating (Di Talia et al., 2009). The Start cyclin Cln3 is a highly
unstable protein whose concentration is rate limiting for cell cycle progression. Cln3 protein levels oscillate weakly through the cell cycle and scale linearly with growth (Schmoller et al., 2015; Tyers et al., 1993) causing Cln3 concentration to remain constant during G1. For this reason Cln3 has been proposed as the “constant-concentration molecule” for the yeast size sensing mechanism (Liu et al., 2015; Wang et al., 2004a). Given that Whi5 phosphorylation and therefore inactivation is the main function of Cln3-Cdc28 activity, K. M. Schmoller and coworkers have recently proposed Whi5 as a “sensor molecule” for the size sensing mechanism. By using time lapse microscopy, Cln3 and Whi5 protein concentration was measured at the single cell level, confirming previous observations for Cln3 constant concentration and demonstrating that Whi5 is synthesized during S/G2/M cell cycle phases, in a largely size-independent manner. Therefore, Whi5 concentration decreases as a consequence of the mass accumulation occurring in G1. This mechanism for Whi5 production causes small daughter cells to be born with higher Whi5 concentration that extends G1 phase duration. Thus, differential size dependency of protein synthesis can provide a mechanism to coordinate growth and division and thus maintain size homeostasis of a yeast cell population.
In the work presented in this thesis we discuss some new aspects of the mechanism controlling size and the fundamental balance between growth and division in the yeast Saccharomyces cerevisiae.
Protein synthesis and ribosome biogenesis: a proxy for cell growth
In a proliferating population the growth rate depends on the extracellular conditions and the biosynthetic capacity of the cell. About 40% of the total macromolecules in a steady-state yeast culture are represented by proteins (Lange and Heijnen, 2001; Polymenis, 2015).
Furthermore, about 20% of the proteome is required for the biosynthesis of ribosomes, enormous ribonucleoprotein complexes whose function is to translate the genetic information encoded by a messenger RNA (mRNA) into the appropriate polypeptide (Figure 4A;
(Liebermeister et al., 2014)). Therefore, protein synthesis is generally accepted as a good parameter to evaluate cell growth.
General aspects of ribosome production
Producing ribosomes is an energy intensive process for the cell: it was estimated that during exponential growth ~104 ribosomes per µm3 can be found in a single yeast cell representing
~10% of the total cytoplasmic volume (Albert et al., 2011; Warner, 1999); Figure 4B)) and that every second in a growing yeast cell ~40 ribosomes are produced (Tschochner and Hurt, 2003). During each division, the ribosomal content of the cell needs to be duplicated, to provide daughter cell with starting translational capacity. In order to produce a single ribosome the cell has to coordinate the transcriptional output from the three RNA Polymerases (RNAPs) to ensure balanced production of the different ribosome components as well as their correct assembly. RNAPI is responsible for ~60% of total transcription (Warner, 1999). The 35S precursor (pre-35S) ribosomal RNA (rRNA) is transcribed by ~150 RNAPI complexes that simultaneously transcribe ~50% of the rDNA copies present in the genome (Albert et al., 2011). Indeed, a eukaryotic genome normally contains several hundred copies of rRNA genes:
in S. cerevisiae the 35S rDNA is organized into head-to-tail tandem arrays of rRNA genes (25S, 18S and 5.8S) and is present in ~100-200 copies per cell codified on chromosome XII (Petes, 1979; Schweizer, 1969) whereas in mammalian cells similar head-to-tail units are found dispersed among five different chromosomes. It has been estimated that in yeast cells only ~50% of the repeats are actively transcribed (Dammann et al., 1993; French et al., 2003).
The pre-35S rRNA is processed into 25S, 5.8S and 18S mature rRNAs that will be assembled into the two main ribosomal subunits, the 60S and the 40S respectively (Tschochner and Hurt, 2003; Udem and Warner, 1972). The RNAPII complex, generally involved in mRNAs and Small Nucleolar RNA (snoRNA) production, transcribes the ribosomal protein genes (RPGs).
This transcriptional output represents ~50% of the RNAPII initiation events during growth under optimal nutrient conditions (Warner, 1999). In the yeast genome that are 79 ribosomal proteins (RPs), 59 of which are encoded by a pair of identical or very similar genes. Therefore, a total of 139 RPGs are transcribed in order to generate the full complement of ribosomal proteins. RPG transcription is highly regulated in response to changes in extracellular conditions (DeRisi et al., 1997). This tight control is achieved by a set of specific transcription factors (TFs) that produce an adequate transcriptional output depending on the growth conditions, and therefore on the available energy (Jorgensen and Tyers, 2004; Martin et al., 2004; Rudra and Warner, 2004; Schawalder et al., 2004; Wade et al., 2004). Interestingly, detailed analysis of RPG promoter architecture and the corresponding TF organization was recently obtained (DeRisi et al., 1997; Hall et al., 2006; Jorgensen et al., 2004; Knight et al., 2014; Lavoie et al., 2010; Lee et al., 2002; Lieb et al., 2001; Martin et al., 2004; Reja et al., 2015; Rudra et al., 2005; Schawalder et al., 2004; Wade et al., 2004). Briefly, RPGs promoters can be categorized into two different types both characterized by the binding of the general TF Rap1 and the RPGs-specific regulators Fhl1 and Ifh1. The binding of the HMG-B protein Hmo1 is what distinguishes the two groups, with category I being bound by Hmo1 DNA- interaction and category II not. Equimolar amounts of rRNA and RPs are required for efficient ribosome production: aberrant ribosome biogenesis occurs when any of the 79 RPs is not properly produced (Rudra and Warner, 2004). A molecular mechanism that coordinates the production of ribosomal proteins and rRNA have been recently elucidated (B. Albert under review; (Rudra et al., 2007)). Briefly, the interaction between the activator Ifh1 and the rRNA processing complex (CURI complex composed by CK2, Utp22 and Ifh1) is responsible for the active communication between RNAPI and RNAPII complexes and for the direct control
regulon of ~236 genes generally referred as Ribosome Biogenesis genes (RiBis; (Brown et al., 2008; Jorgensen et al., 2002; Wade et al., 2006). This regulon is heterogeneous in term of function: most of the RiBis encode for accessory factors required for ribosome assembly and rRNA maturation, additional functional categories are composed of genes encoding for RNAPI and RNAPIII subunits, enzymes required for ribonucleotide biosynthesis, transfer-RNA (tRNA) synthetases and translation regulatory factors (Gasch et al., 2000; Jorgensen et al., 2002; Jorgensen et al., 2004; Wade et al., 2006). However, even though this regulon is characterized by functional diversification, RiBi genes have been clustered together based upon their highly similar response to environmental changes, as well as to genetic perturbations (Gasch et al., 2000; Jorgensen et al., 2002; Miyoshi et al., 2003; Wade et al., 2001). Moreover, the presence of two DNA sequence motifs named PAC and RRPE, found upstream of ~72% of the RiBi genes was also used as a parameter to establish this regulon.
The PAC and RRPE motifs can be found alone or in different combinations on RiBi promoters and are responsible, at least in part, for their regulated expression in response to different stimuli (Gasch et al., 2000; Hughes et al., 2000; Jorgensen et al., 2002; Jorgensen et al., 2004). It should be noted that these motifs are not exclusively found on RiBi promoters, but also upstream of other genes that do not belong to this regulon (Jorgensen et al., 2002).
Finally, proper ribosome production and protein synthesis requires the transcriptional activity of the RNAPIII complex. RNAPIII transcribes the 5S rRNA, present in the same rDNA repeat locus as the pre-35S rRNA. In addition, RNAPIII is responsible for the transcription of transfer RNAs (tRNA; required for translation; (Geiduschek and Kassavetis, 2001)) and two non- coding RNAs (ncRNAs), SNR52 and SNR6 (required for rRNA maturation and mRNA splicing, respectively). Therefore, the transcriptional mechanisms coordinating the three RNAP complexes ensure balanced production of the different ribosome components, and, at the same time, of all the auxiliary factors required for ribosome assembly and for translation. The mechanical process of ribosome assembly takes place mostly within the nucleolus, a dedicated nuclear sub-compartment where the rDNA repeats are clustered (Raska et al.,
2006). Upon transcription, the pre-35S rRNA is heavily spliced and modified by methylation and pseudouridylation (Kressler et al., 1999; Melese and Xue, 1995). rRNA maturation occurs in the presence of the ribosomal proteins, which upon translation are imported into the nucleolus. The 90S pre-ribosome complex is formed with rRNA and protein parts of both 40S and 60S subunits. When the pre-35S rRNA is cleaved the 90S complex is split up into the two main subunits and subsequently exported into the nucleoplasm and then into the cytoplasm through the nuclear pore complex (Grandi et al., 2002; Tschochner and Hurt, 2003). During the journey from nucleolus to cytoplasm, pre-ribosomal subunits are associated with non- ribosomal factors required for quality control and transport. The composition of these factors is dramatically altered during ribosome subunit transportation, highlighting the intricate spatial organization of the eukaryotic nucleus and the unidirectional transport of ribosome subunits (Nissan et al., 2002).
Figure 4. The ribosomes
A. Recently published structure of human 80S ribosome (H. Khatter 2015). B. Image of a cryofixed S. cerevisiae cell. Ribosomes can be seen into the cytoplasm, particularly into the zoomed region. No: nucleolus, the electron-dense region of the nucleoplasm (Np; B.
Albert 2012).
A B
Ribosome synthesis and size homeostasis are tightly coupled processes Ribosome biogenesis is the predominant biosynthetic activity of a growing yeast cell. Nutrient availability modulates the ribosome synthesis rate and therefore the number of active ribosomes that finally drive cell growth. Early studies aimed at identifying molecular pathways involved in growth control and cell cycle progression in yeast revealed the previously uncharacterized involvement of ribosome biogenesis factors in the process of critical size setting (Jorgensen et al., 2002). Size measurement and clustering of the complete set of
~5000 S. cerevisiae single gene deletion strains allowed the identification of abnormally small (whi) or large (lge) mutants compared to the wild type (WT) strain size. Large phenotype resulted, for example from mutations in components of the actin cytoskeleton, secretory pathway, translation control or regulatory components of the RNAPII complex. On the other hand whi mutants were obtained upon deletion of genes encoding for ribosomal proteins, respiratory apparatus factors of the mitochondria, glucose signaling proteins, and proteins required for the biogenesis of ribosomes (RiBis). Compared to the deletion of RP genes that causes a large increase in the duplication time, deletion of RiBi genes causes size reduction without a strong parallel effect on the duplication time. This suggests that the identified RiBi genes could have a role in critical cell size setting. To further explore this observation, analysis of pairwise matings of selected whi mutants and known regulators of Start was performed, and a reduced number of epistatic interactions was obtained. This result suggested new and uncharacterized pathways controlling critical cell size, the majority of which appear to involve aspects of ribosome biogenesis. Therefore, early alterations in ribosome production contribute to critical cell size adaptation and cell cycle entry, before any actual change in the protein synthesis rate occurs. In this way, potential energy wastes are prevented under conditions, such as nutrient deprivation, that do not support high levels of protein synthesis and growth.
Nutrient sensing and signaling pathways controlling cell growth
S. cerevisiae is a widespread microorganism in the natural environment. It is commonly found in decaying fruits such as grapes, apples and bananas, and more generally on sugary foods.
Moreover, yeast cells can be also isolated from animal and plant tissues (Jacques and Casaregola, 2008) and they normally compete with other fungal species and bacteria for food supplies. Therefore, in order to adapt to these extremely variable and competitive environments, S. cerevisiae evolved several mechanisms to maximize survival by optimal use of available energy sources. Budding yeast is a heterotroph microorganism being able to oxidize the chemical bonds of a large variety of organic compounds: for instance, carbon is mainly obtained from glucose, fructose, sucrose and maltose. As nitrogen sources, ammonia and urea are mostly used, but other compounds such as amino acids, small peptides and nitrogen bases can be metabolized. Yeast cells also required phosphorus and sulfur in the form of dihydrogen phosphate ion, sulfate ions or as organic sulfur compounds such as methionine and cysteine (Cai and Tu, 2012). The availability of these key nutrients and the abundance of each relative to the other, dictates the growth rate and the developmental decisions of a yeast cell that can undergo quiescence, filamentous growth and meiosis/sporulation, depending upon the particular nutritional circumstances. Multiple interconnected signaling pathways are stimulated by nutrient quality and quantity and are essential to control growth and development in response to extracellular variations (Broach, 2012). Therefore, nutrients have a dual role as metabolites and signaling molecules that orchestrate the metabolic and transcriptional output to optimize survival. Here I briefly describe how yeast cells respond to the two main classes of nutrients: carbon and nitrogen.
Glucose signaling
Yeast cells can metabolize different carbon sources either via fermentation or by oxidative phosphorylation. When glucose (yeast’s preferred carbon source) is available, it is rapidly fermented to extract energy and to generate the building blocks required for growth. CO2 and
rearrangements cause a switch from glycolysis to the aerobic utilization of the ethanol produced by fermentation. Moreover, non-fermentable carbon sources can be converted into glucose to fuel glycolysis and the TCA cycle to produce ATP, CO2 and storage carbohydrates (Futcher, 2006; Zaman et al., 2008). Therefore, profound metabolic alterations occur according to the available carbon source. It has been determined that ~40% of yeast genes shown altered expression upon glucose addition (Wang et al., 2004b). Several different metabolic pathways are required to assess carbohydrate availability and consequently adjust the cellular metabolic output. Most of the glucose signaling in yeast cells is based on the conserved Protein Kinase A (PKA) pathway that plays a critical role in coupling growth and cell cycle progression in response to glucose availability (Zaman et al., 2008). PKA is a heterotetramer composed of two catalytic subunits and two regulatory subunits (encoded by TPK1, 2 and 3, partially redundant genes, and BCY1, respectively; (Broek, 1987; Tamaki, 2007)). In the presence of glucose PKA kinase activity is stimulated by the second messenger molecule cyclic-AMP (cAMP), whose intracellular concentration is strongly regulated by the counteracting action of the Cyr1 protein (adenyly cyclase of yeast) and of the phosphodiesterases Pde1 and Pde2. Glucose signaling impinges directly on Cyr1 activity causing increased cAMP intracellular concentration. Two parallel pathways communicate glucose perception to the adenylyl cyclase: the Cdc25/Ras and Gpr1/Gpa2 pathways. Several experimental observations demonstrated that the Cdc25/Ras pathway plays the major role in mediating glucose effects, whereas Gpr1/Gpa2 has an auxiliary role and both act through modulation of PKA activity (Kraakman et al., 1999; Kubler et al., 1997; Rolland et al., 2000;
Wang et al., 2004b; Xue, 1998; Zaman et al., 2008). cAMP binds to Bcy1, alleviating its inhibitory role on the PKA catalytic subunits (Broach, 2012; Broek, 1987). As a consequence of PKA activation and target protein phosphorylation, genes required for ribosome production and assembly as well as glycolytic genes are strongly induced, whereas genes involved in the stress response, gluconeogenesis and in metabolism of storage carbohydrates, are repressed. Moreover, increased concentration of intracellular cAMP was shown to delay cell cycle progression and increase the critical cell size required for the G1/S transition by negative
regulation of CLN1 and CLN2 expression (Baroni et al., 1992; Tokiwa et al., 1994). Recent data found the SBF-subunit Swi4 to be a direct target for PKA explaining, at least in part, the negative effect of glucose addition on cell cycle progression (Amigoni, 2015). Therefore, activation of the PKA pathway promotes growth through the expression of genes required for mass accumulation and consistently delays cell cycle progression (Wang et al., 2004b). In the absence of glucose, yeast cells can metabolize other fermentable and non-fermentable carbon sources. To do so, metabolic fluxes have to be reorganized and energy expenditure properly controlled. Under these conditions the Snf1-mediated pathway plays an essential role (Hedbacker and Carlson, 2008). The yeast Snf1 kinase is a heterotrimeric complex composed of the Snf1 catalytic subunit (α subunit), the γ-like Snf4 regulatory subunit and the three different β-subunits Gal83, Sip1 and Sip2. The Snf1 catalytic subunit is part of the eukaryotic family of the AMP-activated protein kinases (AMPKs; (Mitchelhill et al., 1994)). In mammals, AMPK is required for energy homeostasis and its activation is obtained upon an increased AMP:ATP ratio through allosteric activation of the γ-subunit by AMP (Townley and Shapiro, 2007). In contrast, AMP does not directly control yeast Snf1 kinase activity (Wilson et al., 1996;
Woods et al., 1994): indeed, glucose starvation causes T120 of Snf1 to be phosphorylated, promoting Snf1 kinase activity (Hong et al., 2003; Nath et al., 2003). This phosphorylation is obtained by the three different kinases Sak1, Elm1 and Tos3 upon glucose depletion or in response to different stresses (Hong and Carlson, 2007). Moreover, under these conditions Snf1 subcellular organization is actively modified: a Gal83-containing complex is imported into the nucleus, Snf1/Sip1 migrates to the vacuolar membrane, whereas the Snf1/Sip2 complex remains in the cytoplasm, suggesting that the functional overlap of Snf1 β-subunits is only partial (Zaman et al., 2008). Unlike in presence of glucose, Snf1 complexes are found in the cytoplasm upon glucose depletion and this localization requires the activity of PKA (Zaman et al., 2008). The downstream effects of Snf1 kinase are obtained through phosphorylation of several different transcription factors that in turn control the expression of genes required for growth in the absence of glucose, stress response and cell cycle progression. For example,
fatty acids oxidation and amino acids transport are activated by the Mig1 and Adr1 transcription factors (TFs); increased gluconeogenesis is achieved by the action of Cat8 and Sip4, and long term adaptation to carbon stress is brought about by Msn2 phosphorylation (Zaman et al., 2008). Moreover, Snf1 was found to directly regulate gene expression by chromatin remodeling obtained through histone H3 phosphorylation on a subset of target genes (Lo et al., 2001; Lo et al., 2005). Interestingly, Snf1 was also demonstrated to play a direct role in the regulation of G1/S transcription by localizing to the SBF/MBF target gene promoters through a direct interaction with the Swi6 subunit (Busnelli et al., 2013; Pessina et al., 2010). As is evident by this brief description, cell physiology is quickly adapted in response to glucose availability prior any actual change of the cell growth potential, avoiding energy waste and controlling size homeostasis.
Nitrogen regulation of growth and development
As observed for carbon sources, yeast cells adjust their growth rate and biosynthetic capacity by assessing quality and quantity of the available nitrogen compounds. In conditions of nitrogen scarcity cells slow down their growth rate by inhibiting ribosome biogenesis and protein translation (Brauer et al., 2008). When nitrogen is completely depleted from the medium, cells enter a quiescent state, which enables survival during long period of starvation (Brauer et al., 2008; De Virgilio, 2012; Klosinska et al., 2011). Compared to cells grown in the complete absence of nitrogen, auxotrophic yeast cells starved for their required amino acids show strongly reduced survival, suggesting that specific adaptations occur when nitrogen sources are limiting. As previously stated, S. cerevisiae is able to metabolize a variety of nitrogen-containing compounds. However, glutamine and ammonia are preferably used as nitrogen sources compared to other amines and amides. The Nitrogen Catabolite Repression pathway (NCR) is responsible for this hierarchical preference, ensuring that the preferred nitrogen compounds are assimilated first (Zaman et al., 2008). When the NCR is active, genes required for alternative nitrogen compound metabolism are repressed and, at the same time, ribosome biogenesis and ribosomal protein gene transcription is promoted to stimulate growth
(Magasanik and Kaiser, 2002). Finally, the general amino acid permeases are post- translationally modified to prevent their membrane localization and the process of autophagy is down regulated under conditions of NCR activation (Magasanik and Kaiser, 2002; Yang and Klionsky, 2009).
During growth on alternative nitrogen sources and upon mitochondrial dysfunction the Retrograde Regulation pathway (RTG) maintains intracellular levels of glutamate and ammonia. Under these conditions, the available nitrogen compounds are converted to ammonia, which is consequently condensed to α-ketoglutarate to give glutamate. The RTG pathway is composed of positive (Rtg1-2 and 3 and Grr1) and negative (Mks1, Bmh1-2 and Lst8) regulators that control the subcellular localization of specific transcription factors required for growth and survival according to the available nitrogen sources (Yang and Klionsky, 2009). The aforementioned nitrogen signaling pathway is mainly controlled and integrated to the other nutrient signaling by the evolutionary conserved TOR (Target Of Rapamycin) kinase (Broach, 2012; Panchaud et al., 2013). This Ser/Thr protein kinase is a member of the phosphatidylinositol kinase-related protein kinase (PIKK; (Keith and Schreiber, 1995; Wullschleger et al., 2006) family and a central regulator of cell physiology.
Saccharomyces cerevisiae, unlike other eukaryotes, has two distinct Tor proteins encoded by TOR1 and TOR2, the first of which was originally identified by mutations that confer resistance to the growth inhibitor rapamycin (Heitman et al., 1991). Tor1 and Tor2 have been shown to be part of two distinct protein complexes found in all eukaryotes examined to date (De Virgilio and Loewith, 2006; Loewith et al., 2002): TORC1, containing Tor1 or Tor2, Kog1 and Lst8, responds primarily to the quality of nitrogen sources, whereas TORC2, containing Tor2, Avo1- 3, Bit61 and Lst8 is mainly controlled by environmental stresses and alteration of plasma membrane tension (Loewith and Hall, 2011; Wullschleger et al., 2006). The TORC1/2 signaling pathways control several different processes required for growth and cell cycle progression. The overlapping functions of Tor1 and Tor2 mediate “temporal control” of cell
nutrient transport (Loewith and Hall, 2011; Schmelzle and Hall, 2000). On the other hand
“spatial control” of cell growth is a Tor2-unique function. Being mainly localized to the plasma membrane, TORC2 controls actin cytoskeleton organization, sphingolipid biosynthesis and endocytosis (Cardenas et al., 1999; Hardwick et al., 1999; Komeili et al., 2000; Loewith and Hall, 2011; Loewith et al., 2002; Rispal et al., 2015; Wullschleger et al., 2006). The macrolide drug rapamycin, in complex with the prolyl isomerase Fkbp12, binds to TORC1 specifically, preventing its interaction with target substrates, whereas TORC2 is insensitive to rapamycin inhibition (Loewith and Hall, 2011). Under optimal growth conditions, yeast cells exploit their maximum growth potential and TORC1 activity is required to coordinate the different processes promoting mass accumulation, such as ribosome biogenesis, amino acids biosynthesis, efficiency of protein translation and nutrient transportation. Moreover, TORC1 prevents the activation of pathways required to metabolize alternative nitrogen sources (Loewith and Hall, 2011). One important mechanism of TORC1 signaling is the regulation of PP2A-type phosphatases that control the expression of genes required for stress responses and for metabolism of alternative nitrogen sources (Duvel et al., 2003). Most of the regulatory activity of TORC1 is obtained by direct phosphorylation and activation of the AGC family kinase Sch9 (Urban et al., 2007), the yeast homolog of the mTORC1 target S6K1 (Isotani, 1999; Sarbassov et al., 2005). Sch9 phosphorylation is strongly controlled in a nutrient- and stress-sensitive manner: rapid dephosphorylation is observed upon rapamycin treatment, carbon or nitrogen starvation, as well as in presence of less favorable nitrogen sources (Zaman et al., 2008). Upon TORC1 mediated activation, Sch9 stimulates transcription of ribosome biogenesis and ribosomal protein genes and also controls cell fate, at least in part by preventing entry into quiescence when nutrients are available (Zaman et al., 2008).
Interestingly, glucose also affects Sch9 function both at the level of protein synthesis and phosphorylation (Jorgensen et al., 2004; Urban et al., 2007), even though the molecular details of this control are not known. Moreover, Sch9 transcriptional targets are partially overlapped with those of PKA, even if Sch9 plays only a minor role in glucose-mediated gene transcription (Jorgensen et al., 2004; Zaman et al., 2008). In sum, analysis of Sch9 phosphorylation and
transcriptional regulation highlights the strong interconnection between nutrient sensing and signaling pathways controlling cell growth.
DNA-dependent RNA polymerases mediate genome transcription
In every cell, the genetic information encoded by the DNA molecule is used to produce proteins, via messenger RNA (mRNA) intermediates, and structural RNAs. Transcription, the first step in gene expression, is a complex series of events that starts with the recruitment of large multi-subunit protein complexes called DNA-dependent RNA polymerases (RNAPs) to promoter regions. In eukaryotic cells, three different RNAP complexes have been isolated over 4 decades ago: the RNAPI, II and III (Roeder and Rutter, 1969, 1970). Their biochemical and physical properties have been studied extensively leading to our current detailed knowledge of their structural composition (Cramer et al., 2008; Cramer et al., 2001; Fernandez-Tornero et al., 2013; Vannini and Cramer, 2012). Moreover, analyzing the different sensitivities of these complexes to specific inhibitors revealed their ability to synthesize different classes of RNA and therefore their specificity in eukaryotic genome transcription (Kedinger, 1970; Lindell, 1970; Seifart and Sekeris, 1969; Weinmann, 1974; Zylber and Penman, 1971). RNAPI synthesizes the 35S ribosomal RNA precursor (pre-35S rRNA), RNAPII transcribes messenger RNAs (mRNA) and small nuclear RNAs (snRNAs), while RNAPIII produces different types of short untranslated RNAs such as transfer RNA (tRNA), small nuclear and nucleolar RNA (snoRNA), the 5S rRNA, the RNA component of nuclear RNase P (RPR1) and the RNA subunit of the signal recognition particle SRP (SCR1). Detailed structural studies, mainly performed with yeast proteins, have shown that the catalytically active central region of these complexes is highly conserved and that strong structural differences on their surfaces account for functional specialization (Cramer et al., 2008). The RNAPs active central core resembles the (α2ββ’ω) subunit composition of the bacterial holoenzyme core (Sentenac, 1992) with 5 extra subunits: Rpb5, Rpb8, Rpb10, and Rpb12 being common between the three complexes (Dumay, 1999; Krapp et al., 1998; Rubbi, 1999) and A12.2, Rpb9 and C11
core complex of eukaryotic RNAPs (Bischler et al., 2002). Associated to this ten-subunit core, 4 specific proteins are found in RNAPI, 2 into RNAPII and 7 in RNAPIII, which together form enormous macromolecular complexes of 589, 514 and 693 KDa, respectively (Cramer et al., 2008). Common transcriptional initiation mechanisms can be described for the three RNAPs.
General transcription factors (GTFs) recognize specific promoter elements, recruit the polymerase and mediate DNA opening and initial RNA synthesis. Moreover, all polymerases require TATA Binding Protein (TBP), which likely interacts with all promoters analyzed to date, inducing a 90° bend in the DNA that is essential to initiate transcription (Blair et al., 2012;
Cormack, 1992; Kim et al., 1993a; Kim et al., 1993b; Rhee and Pugh, 2012; Wu et al., 2001).
The topology and function of the initiation complexes that interact with TBP is what confers gene-class specificity to the three RNAPs. Finally, the mechanism of RNA synthesis during transcription elongation is another common feature of the three RNAPs. During elongation, the RNAPs move along the DNA template producing in its active site a DNA-RNA hybrid between the DNA template strand and the newly synthesized RNA molecule (Cramer 2008).
When a nucleoside triphosphate (NTP) enters the active site it is catalytically added to the growing 3’ end of the nascent RNA, releasing pyrophosphate ion. Further translocation releases the active site for binding to the next NTP (Gnatt, 2001; Poglitsch et al., 1999).
Transcription mediated by RNAPI and III
60% of the total transcription in yeast depends on RNAPI and occurs at the rDNA locus (Warner, 1999). Initially the six-subunit complex UAF (Upstream Activating Factor) associates stably with the rDNA promoter in a sequence-specific manner promoting the association of RNAPI and its specific transcription factors with the core promoter element (Boukhgalter et al., 2002). TBP is also necessary for pre-initiation complex formation (PIC). It was shown in yeast and humans that RNAPI is unable to initiate productive transcription when only UAF is found at the rDNA promoter (Cavanaugh et al., 2008; Yamamoto et al., 1996). To engage active rRNA production, the key regulator Rrn3 (mammalian TIF-IA) is required. Direct interaction between Rrn3 and the subunit Rpa43 mediates RNAPI binding with the specific
transcription factors and therefore transcription. Upon initiation, Rrn3, TBP and the GTFs all leave the promoter, whereas UAF remains stably bound as a scaffold that supports further rounds of RNAPI recruitment and transcription initiation (Figure 5). The described mechanism for RNAPI-mediated transcription suggests that rRNA synthesis may be controlled by a two- step mechanism where the rDNA promoter is first made accessible (open conformation) by UAF binding and consequent active PIC formation is obtained by TBP binding and by direct interaction with the key regulator Rrn3 (Aprikian et al., 2001). Early structural analysis (Aprikian et al., 2001), confirmed by the recently obtained crystal structure of the RNAPI complex (Fernandez-Tornero et al., 2013), revealed one feature that distinguishes RNAPI from the other polymerases: the presence of the heterodimer Rpa49/Rpa34. Curiously, this module was shown to be required for release of Rrn3 during elongation despite being located on the opposite side of the RNAPI complex (Albert et al., 2011; Kuhn et al., 2007). However, the high loading rate of RNAPI on each rDNA gene causes direct interaction between adjacent enzymes, placing the Rpa49/34 module in close proximity to the Rpa43 subunit on an adjacent RNAPI complex. This may explain the ability of the Rpa49/34 module to eject Rrn3, highlighting the cooperative action of adjacent RNAPI complexes to stimulate the elongation step of transcription (Albert et al., 2011). The amount of the initiation-competent form Rrn3- RNAPI is a rate-limiting factor of rRNA production and it was shown to represent an important regulatory point for nutrient signaling pathways. Both yeast and mammals have decreased Rrn3 activity upon inhibition of protein synthesis or under growth-limiting conditions (Cavanaugh et al., 2002; Milkereit, 1998; Zhao J., 2003). For example, when grown into stationary phase, yeast cells decrease the amount of rRNA genes that are actively transcribed and pre-rRNA synthesis is reduced by more than 10 fold (Sandmeier, 2002). Additionally, transcription of rRNA was found to rapidly increase upon glucose addition to starved cells (Kief and Warner, 1981). In contrast, glucose depletion decreases synthesis of Rrn3 (Claypool, 2004; Grummt, 2003; Mayer, 2004). Moreover, TORC1 inactivation upon rapamycin treatment causes a rapid (5-10 minutes) down regulation of rRNA gene transcription and a concomitant
Rudra et al., 2007). It was found that TORC1 inhibition stimulates Rrn3 proteolytic degradation and consequently destabilizes the Rrn3-RNAPI complex (Philippi, 2010). Moreover TORC1 was shown to directly bind the rDNA promoter in a nutrient- and rapamycin-dependent manner (Li et al., 2006) promoting pre-35S synthesis. Sch9 mediates the regulatory effect of TORC1 on RNAPI activity (Huber et al., 2011) by promoting both RNAPI recruitment on rDNA promoters and rRNA processing by as yet undefined mechanisms (Huber et al., 2011;
Jorgensen et al., 2004).
Figure 5. Basal model of RNAPI assembly
Figure from Aprikian et al. 2007. Schematic description of the proposed model for RNAPI transcription cycle.
Together with RNAPI, the transcriptional activity of RNAPIII is responsible for the synthesis of untranslated RNAs required for ribosome production and function as well as for pre-mRNA maturation. In addition to the RNAPIII core, class III gene expression requires three transcription factors: the GTFs TFIIIB and TFIIIC, and the specific transcription factor TFIIIA exclusively found at the RDN5 (5S rRNA) promoter (Acker et al., 2013; Klekamp and Weil, 1982; Ottonello et al., 1987). The transcription complex assembly process starts with TFIIIC binding, which requires the target sequences Box A and Box B (Figure 6A). These sequences can be found in upstream promoter DNA (for RPR1 and SNR52) or within the transcribed region (for all tRNA genes, SNR6 and SCR1). The main role of TFIIIC is to recruit TFIIIB to the DNA during initiation (Figure 6B). However, on long class III genes like SCR1, TFIIIC mediates the RNAPIII reinitiation mechanism, supporting high rates of RNAPIII transcription (Ferrari et al., 2004). Moreover, TFIIIC promotes transcription by competing with nucleosome formation (Marsolier et al., 1995). TFIIIB complex lacks intrinsic DNA-binding activity and is therefore recruited to promoters by direct interaction with TFIIIC (Kassavetis, 1989). TFIIIB can be considered as an assemblage of three transcription factors (Acker et al., 2013): (i) Brf1, which promotes RNAPIII recruitment and works as a scaffold to connect TFIIIB subunits together; (ii) Bdp1, which contains a SANT histone binding module (Aasland, 1996); and TBP.
Once recruited to DNA, Bdp1 and TBP generate a DNA distortion and promote DNA strand separation to initiate transcription (Kassavetis, 1989; Leveillard, 1991). During the last decades, several different signaling pathways have been demonstrated to impinge on the transcriptional activity of RNAPIII (Acker et al., 2013). For example, down regulation of RNAPIII occurs upon entry into stationary phase, as a consequence of DNA damage or oxidative stress, upon TORC1 inhibition and when the secretory pathway is disturbed (Mizuta and Warner, 1994; Willis et al., 2004). In all of these cases, down regulation of RNAPIII transcription requires Maf1 (Desai et al., 2005; Upadhya et al., 2002), the only known negative regulator of RNAPIII. Maf1 physically interacts with the central core subunit C160 (Oficjalska- Pham et al., 2006) in a phosphorylation state-dependent manner. The least phosphorylated form of Maf1 interacts with RNAPIII (Roberts et al., 2006), causing rearrangement of specific
subunits (C82/C34/C31), impairment of RNAPIII recruitment to TFIIIB and inhibition of the transcription initiation step (Vannini et al., 2010). Phosphorylation of Maf1 controls its nuclear localization: experiments performed so far have demonstrated that PKA signaling in response to glucose and TORC1/Sch9 signaling in response to nitrogen cause Maf1 to be phosphorylated on consensus sequences close to its nuclear localization domain. This has been thought to prevent Maf1 nuclear accumulation (Lee et al., 2009; Oficjalska-Pham et al., 2006). On the other hand, PP2A activity counteracts PKA and TORC1-mediated Maf1 phosphorylation. Moreover, even when present in the nucleus, the phosphorylated form of Maf1 does not affect transcription, suggesting that phosphorylation both prevents nuclear accumulation and direct interaction with RNAPIII (Towpik et al., 2008). Finally, it is important to mention that TORC1 and the ubiquitous and highly conserved Casein Kinase 2 (CK2) were found by chromatin immunoprecipitation (ChIP) to be present on class III gene promoters, where they could directly control RNAPIII activity (Acker et al., 2013). For example, CK2 phosphorylates TBP and Bdp1 TFIIIB subunits, transducing DNA damage and cell cycle- specific signals to inactivate RNAPIII (Ghavidel and Schultz, 2001; Hu et al., 2004). Taken together, these observations underlie the coordinated regulation of RNAPI and RNAPIII transcription obtained by several different signaling pathways that allow rapid adjustment of their transcriptional activity under different growth conditions.
Figure 6. RNAPIII dependent transcription
Picture from Acker et al 2013. A. Schematic representation of class III genes and the relative position of downstream and upstream sequences important for transcription. B.
The proposed transcription complex assembly is schematically described. TFIIIa, TFIIIB and TFIIIC are represented as red green and blue oval respectively. The purple oval represent the 17-subunit RNAPIII complex.
A
B
The transcription mediated by RNAPII
The 12-subunit complex RNAPII transcribes eukaryotic protein coding genes. Regulation of RNAPII activity underlies cell responses to environmental changes and, more generally, cellular development. Therefore, detailed investigation of RNAPII’s mechanism of action and regulation will help to elucidate essential aspects of cell physiology. As already described, the activity of RNAPs requires the GTFs that surround the core complex, promote its association with the DNA template and stimulate its enzymatic activity. Nowadays, multiple RNAPII structures are available that help to elucidate the classical model for PIC assembly and transcription initiation (Armache et al., 2003; Armache et al., 2005; Bushnell and Kornberg, 2003; Cramer et al., 2000; Cramer et al., 2001). Initially, TBP is recruited on promoters in a sequence-specific manner on TATA box-containing genes (~30bp upstream of transcription start site (TSS)). However, TBP also binds to the promoters of the majority of yeast protein coding genes that do not contain a canonical TATA box (TATA-less promoters; (Rhee and Pugh, 2012)). At these promoters TBP binds as part of the multi-subunit TFIID complex that is involved in the recognition of different core promoter sequences through multiple TBP- associated factors (TAFs) and modulates TBP activity. Moreover TFIID mediates the interaction with activating transcription factors (Sainsbury et al., 2015). TFIID binding on the promoter induces a TBP-mediated 90° DNA bend required for initiation and assembly of the other components. The TBP-DNA complex is further stabilized by TFIIA and TFIIB that flank TBP on both sites. TFIIB is important for the appropriate orientation of the PIC (Muhlbacher et al., 2014) and upon RNAPII recruitment plays an essential role in positioning the DNA over the enzyme active center and in maintenance of the transcription bubble (Kostrewa et al., 2009). Furthermore, TFIIB functions in setting the TSS (Bangur et al., 1997; Faitar et al., 2001;
Kuehner and Brow, 2006; Li et al., 1994) and stimulates initial RNA synthesis (Sainsbury et al., 2013) before being released from the RNAPII complex once it is engaged in productive elongation (Cabart et al., 2011). RNPII-TFIIF is then recruited to the resulting promoter complex forming the PIC core. Approximately 50% of RNAPII in yeast is associated with TFIIF
(Rani et al., 2004), which prevents unspecific DNA binding and is required for TSS selection and initiation of RNA synthesis (Ghazy et al., 2004; Khaperskyy et al., 2008; Yan et al., 1999).
Finally, TFIIE and TFIIH binding completes the PIC: TFIIE is required to promote the activity of TFIIH, which contains a DNA-dependent ATPase module essential for transcription initiation and a kinase module that phosphorylates the C-terminal domain of Rpb1 (the major central core subunit) to promote elongation ((Conaway and Conaway, 1989, 1993; Kim et al., 1994;
Murakami et al., 2012; Schaeffer et al., 1993) Figure 7).
In addition to the general transcription factors, an increasing body of evidence (Hirst et al., 1999; Malagon et al., 2004; Pan et al., 1997; Prather et al., 2005; Wery et al., 2004) demonstrates that the yeast elongation factor Dst1 (also referred as TFIIS) is involved in the regulation of transcription initiation. TFIIS is a ~30KDa protein belonging to the highly conserved SII family of factors that promote the efficient synthesis of long transcript in vitro (Sekimizu, 1976). It is now well established that TFIIS stimulates the intrinsic RNA cleavage activity of paused RNAPII elongation complexes, allowing backtracking and the consequent generation of a new 3’ transcript end (Wind and Reines, 2000). Yeast TFIIS stimulates PIC formation through direct interaction with RNAPII and independently of its transcript cleavage activity (Kettenberger et al., 2003; Kim et al., 2007). TFIIS recruitment is mediated by co- activators and does not involve RNAPII (Guglielmi et al., 2007; Prather et al., 2005).
Also required for transcription from most RNAPII-dependent promoters is the co-activator multisubunit complex Mediator. As the name suggests, its main function is to bridge the gene specific activators to the general transcription factors (Thomas and Chiang, 2006) promoting both basal and activator-dependent transcription (Hengartner et al., 1995; Holstege et al., 1998; Mittler et al., 2001; Takagi et al., 2006). Furthermore, Mediator stabilizes the PIC and stimulates TFIIH kinase activity (Sogaard and Svejstrup, 2007). Unlike GTFs, Mediator function may be promoter-specific and dependent upon different growth conditions (Fan et al., 2006). The yeast Mediator is a 25 subunit complex organized in four functionally and structurally distinct modules: the “head” and “middle” modules that are essential for viability
