Iä! Iä! Cthulhu Fhtagn!

List of Papers

This thesis is based on the following papers, which are referred to in the text by their Roman numerals.

I Orzechowski Westholm J, Nordberg N, Murén E, Ameur A,

Komorowski J, Ronne H (2008) Combinatorial control of gene expression by the three yeast repressors Mig1, Mig2 and Mig3. BMC Genomics 9: 601.

II Orzechowski Westholm J, Tronnersjö S, Nordberg N, Olsson I, Komorowski J, Ronne H (2012) Gis1 and Rph1 regulate glyc-erol and acetate metabolism in glucose depleted yeast cells. PLoS ONE 7: e31577.

III Nordberg N, Olsson I, Hu GZ, Orzechowski Westholm J, Ronne H (2012) The histone demethylase activity of Rph1 af-fects telomeric silencing but is not required for nutrient signal-ing. Manuscript.

IV Nordberg N, Olsson I, Hu GZ, Ronne H (2012) Rph1 functions downstream of TOR and Rim15 in yeast but is not needed for chronological lifespan extension. Manuscript.

Reprints were made with permission from the respective publishers.

Contents

Introduction ................................................................................................... 11

Transcription and chromatin modification .................................................... 13 Transcription factors and transcriptional regulation ................................. 14

C2H2 zinc finger transcription factors .................................................. 15 Chromatin modification ........................................................................... 17

Histone acetylation and deacetylation ................................................. 17 Histone methylation ............................................................................. 18 The Set2/Rpd3(S) pathway .................................................................. 19 Histone demethylation ......................................................................... 22 Histone demethylation in budding yeast .............................................. 23

Nutrient signaling and aging ......................................................................... 27 The growth phases of budding yeast ........................................................ 28 Nutrient signaling pathways ..................................................................... 29

Glucose repression and the Snf1 kinase .............................................. 30 The glucose induction pathway ........................................................... 32 The PKA pathway................................................................................ 32 The TOR pathway................................................................................ 33 The Sch9 kinase ................................................................................... 34 Pho85-Pho80 signaling ........................................................................ 35 Rim15 and downstream transcription factors ...................................... 35

Nutrient signaling as a regulator of aging ................................................ 38

Aims .............................................................................................................. 42

Results and discussion .................................................................................. 43 Paper I ...................................................................................................... 43 Paper II ..................................................................................................... 43 Paper III .................................................................................................... 44 Paper IV ................................................................................................... 46

Future perspectives ....................................................................................... 47

Acknowledgements ....................................................................................... 50

References ..................................................................................................... 53

Abbreviations

AMPK AMP-activated protein kinase cAMP Cyclic adenosine monophosphate CLS Chronological lifespan CR Caloric restriction CTD C-terminal domain CWI Cell wall integrity H2A Histone 2A H2B Histone 2B H3 Histone 3 H4 Histone 4 HAT Histone acetylase HDAC Histone deacetylase JmjC Jumonji C JmjN Jumonji N me1 Monomethylated me2 Dimethylated me3 Trimethylated mRNA Messenger RNA ORF Open reading frame PDS Post diauxic shift PHD Plant homeodomain PIC Preinitiation complex PKA Protein kinase A rDNA Ribosomal DNA Ribi Ribosome biogenesis RLS Replicative lifespan RNA Pol RNA polymerase RP Ribosomal protein Rpd3(L) The large Rpd3 complex Rpd3(S) The small Rpd3 complex rRNA Ribosomal RNA SnRK1 Snf1-related protein kinase STRE Stress response element TBP TATA-binding protein TCA Tricarboxylic acid TORC1 Tor complex 1

TORC2 Tor complex 2 tRNA Transfer RNA UAS Upstream activation sequence URS Upstream repression sequence WT Wild type ZnF Zinc finger

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Introduction

A cell is ultimately defined not only by the DNA it contains, its genome, but also by how this DNA is utilized, a phenomenon known as gene expression. This is evident from the fact that the different cell types constituting a multi-cellular organism all contain the same genome, but can yet be fundamentally different. But besides allowing differentiation into such distinct types, the fact that cells can alter their gene expression also gives them the oftentimes vital ability to adapt to a changing environment. All living cells have there-fore evolved elaborate ways of controlling their gene expression, which at its most basic could be described as deciding whether a gene is turned on (acti-vated) or off (repressed).

One very important environmental condition that cells need to adapt their gene expression in response to is nutrient availability. Basically, cells need to be able to decide if it is time to grow and prosper, or to hunker down and wait for better times. Interestingly, research has revealed that this decision-making is tightly linked to the aging of cells, and even entire multicellular organisms. This finding, that cells in effect have systems controlling their own lifespan, has philosophical ramifications as it alters our long-held view that aging and death are unavoidable processes of deterioration. Adaptation to extracellular nutrients is controlled by intracellular signaling pathways that are strikingly well conserved throughout evolution, the most important end result of which is the activation or repression of transcription, the initial step in gene expression.

The use of model organisms has been of great use in the field of molecu-lar biology. Studies of higher, multicellular organisms can pose many practi-cal problems that may be bypassed by studying the same phenomena in a simpler organism. The rationale behind such studies is of course that many basic functions and pathways have been strongly conserved between organ-isms during evolution, and the information gained from the simpler system can often be extrapolated to higher organisms. The yeast Saccharomyces cerevisiae has long served as such a model organism, and is arguably the simplest eukaryotic model system imaginable. Studies using S. cerevisiae as a model organism have often proven fruitful, two high-impact examples being Leeland Hartwell’s discoveries of cell cycle regulators and Roger Kornberg’s work on transcriptional regulation which garnered them Nobel Prizes in 2001 and 2006, respectively. A bit closer to home as far as this thesis is concerned, S. cerevisiae was used to elucidate the mechanism of

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action of the immunosuppressant rapamycin (Heitman et al., 1991), a drug with remarkable anti-aging properties which is now being discussed as a potential modulator of human lifespan (Bjedov and Partridge, 2011; Kaeberlein and Kennedy, 2009; Sharp and Richardson, 2011).

A small unicellular organism, S. cerevisiae often goes by the more collo-quial name budding yeast, which refers to its manner of division where a small daughter cell buds off from a larger mother cell. It is also referred to as baker’s or brewer’s yeast, owing to the fact that its ability to produce carbon dioxide and ethanol have been used by mankind for thousands of years to manufacture bread and spirits.

The advantages of using budding yeast for research are many. Its genome, which was the first eukaryotic genome to be fully sequenced (Goffeau et al., 1996), is almost entirely free of introns, making the cloning of genes simple. In addition, yeast is capable of very efficient homologous recombination, greatly facilitating genetic manipulations and making gene disruption trivial. There are also a number of well established plasmid systems that can be used for complementation, overexpression and screening studies. Further, bud-ding yeast can be cultivated in both a haploid and diploid state, which offers additional advantages. Conducting experiments in the haploid state facili-tates genetic manipulations and means that the effect of a mutation can be immediately observed, while the diploid state offers the possibility to per-form genetic crosses to produce haploid offspring with, for example, multi-ple knockouts. An obvious advantage is also that budding yeast is very cheap and easy to work with. It can be cultivated much like bacteria, on agar plates or in liquid culture. Nevertheless, yeast is still an eukaryote, and many of its functions are well conserved with those in higher organisms.

This thesis employs budding yeast as a model to study regulation of gene expression in response to nutrient availability. The bulk of the thesis is di-vided into two sections. The first part describes how so called transcription factors contact DNA to activate and repress genes, and deals with aspects of this pertaining to chromatin, the form of DNA present in cells. The second part deals with nutrient signaling, which is controlled by evolutionary con-served pathways that sense environmental cues and ultimately control the activity of transcription factors. These two major sections are followed by an outline of the aims of the thesis, a summary of the four papers included in the thesis, and a discussion about future perspectives.

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Transcription and chromatin modification

As mentioned previously, a very important function in all cells is the ability to control the way it uses its genetic material. Gene expression is defined as the conversion of the information harboured in genes into functional gene products, which are most often proteins. This comes about through a number of sequential processes; transcription of DNA into messenger RNA (mRNA), mRNA splicing and export into the cytoplasm, translation of mRNA into protein and finally post-translational modifications. A cell can control its gene expression by regulating any of these processes. However, as it is the first step, much regulation takes place at the level of transcription. Specialized proteins known as transcription factors are significant players in transcriptional regulation; they bind to specific DNA sequences and affect the activity of RNA polymerases on nearby genes, thereby activating or re-pressing transcription.

There is also an important level of complexity added to this which has emerged in the last decade or so. This is the field of epigenetics; the study of how the output of a gene can be altered without changing the underlying DNA sequence. Epigenetic phenomena were for a long time considered mys-terious, but have now been largely explained by the fact that DNA, or the histone proteins which are intimately associated with DNA, can be covalent-ly modified in a myriad of ways (Goldberg et al., 2007). It is worth pointing out that the term epigenetics in its classical definition described phenotypes that were independent of DNA mutations, but were nonetheless heritable. However, in recent usage the term often does not strictly depend on herita-bility, but instead encompasses all the information contained in chromatin that is not coded by DNA. Indeed, some of the modifications that have been termed epigenetic are extremely dynamic, changing back and forth many times within a single cell cycle.

This chapter begins by giving an overview of transcription regulation, and introduces several budding yeast transcription factors that have been under study in this thesis. This is followed by a discussion of how chromatin, the form of DNA which transcription factors interact with, can be modified, and what the consequences of this may be.

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Transcription factors and transcriptional regulation The nuclear DNA of all eukaryotic cells is transcribed by three RNA poly-merases (Pol I, II and III). RNA Pol I and III together transcribe the riboso-mal RNA (rRNA) and the transfer RNA (tRNA), which only represents a small portion of the genome but can make up more than 90% of the RNA content in a rapidly growing cell. In contrast, RNA Pol II is responsible for producing the mRNAs transcribed from all protein coding genes. While de-tailed knowledge of RNA Pol I and III is emerging (Werner et al., 2009), our deepest understanding of how transcription is regulated comes from studies on RNA Pol II.

Transcriptional regulation begins with transcription factors binding spe-cifically to short stretches of DNA upstream of a gene; these are known as upstream activation sequences (UASs) or upstream repression sequences (URSs). The outcome of this binding is that coactivator and corepressor complexes are recruited, which in turn affects events at another region up-stream of the gene; the core promoter, where the RNA Pol II assembles with a number of general transcription factor complexes to initiate transcription (Hahn, 2004; Hahn and Young, 2011). Importantly, for transcription initia-tion to occur the TATA-binding protein (TBP) needs to be delivered to the core promoter where it binds; this can be achieved by either of the coactivator complexes TFIID and SAGA (Huisinga and Pugh, 2004). At the core promoter, TBP then assembles with the general transcription factor complexes TFIIA and TFIIB into a more stable complex, which in turn re-cruits the RNA Pol II and TFIIF. This is followed by binding of TFIIE and TFIIH, and a structure known as the preinitiation complex (PIC) is formed. A subunit of TFIIH has helicase activity, which unwinds the DNA strands, transforming the PIC into an open complex. This also enables scanning for a downstream transcriptional start site, which is where RNA Pol II initiates transcription and proceeds into elongation. Importantly, one of the subunits of RNA Pol II, Rpb1, has a unique C-terminal domain (CTD) consisting of multiple repeats of seven repeated amino acid residues, which is specifically phosphorylated as the polymerase transitions through these stages (Buratowski, 2009). Thus, the CTD is unphosphorylated when RNA Pol II is initially assembled in PIC, but is then phosphorylated at Ser5 and Ser7 with-in each repeat during initiation by a subunit of TFIIH, and further phosphor-ylated by other kinases at Ser2 during elongation. These phosphorylations regulate many functions associated with RNA Pol II, an example being the interaction with the Set2 methyltransferase, which will be discussed below.

Transcription factors clearly serve an important role in transcriptional regulation as they control the first step leading to transcriptional output; promoting the chain of events outlined above in the case of a transcriptional activator, or counteracting it in the case of a repressor. In general, transcrip-tion factors have a modular structure built up of discrete functional domains;

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these can for example be activating or repressing domains that act by recruit-ing coactivator or corepressor complexes, respectively (Hahn and Young, 2011). The DNA-binding domains are usually very well conserved, and tran-scription factors are therefore grouped based on the type of DNA-binding domain they contain; these general groups are then further subdivided into transcription factor families (Hahn and Young, 2011; Luscombe et al., 2000). The general groups of transcription factors include the helix-turn-helix and the leucine zipper type, but the most abundantly found DNA-binding domain is the zinc-coordinated domain. This DNA-binding domain is characterized by the coordination of zinc ions by conserved cysteine and histidine residues, and transcription factors containing such domains are further subdivided into families depending on how this is accomplished (Krishna et al., 2003). For example, in the case of the C6 domain, also known as the Zn2Cys6 binuclear cluster, two zinc ions are coordinated by six cyste-ine residues. The next section will discuss another type of zinc-coordinated DNA binding domain, the C2H2 zinc finger, as transcription factors contain-ing such domains are of special relevance to this thesis.

C2H2 zinc finger transcription factors The most common type of zinc-coordinated DNA-binding domain in eukar-yotic transcription factors is the C2H2 zinc finger domain; budding yeast, for example, has 53 proteins containing this domain (Böhm et al., 1997). The C2H2 zinc finger is a small domain of around 30 amino acids composed of a two-stranded antiparallel β-sheet and an α-helix. This structure is stabilized by the coordination of a single zinc ion by two conserved cysteines in the β-sheet and two histidines in the α-helix. Zinc fingers bind DNA by inserting the α-helix into the major groove; conserved residues there make specific contacts with three consecutive base pairs of DNA. As it is only a triplet of base pairs that is recognized, a single zinc finger does not bind DNA with a high specificity. However, zinc fingers are usually found in tandem repeats of two or more domains, giving rise to much more precise sequence specific-ity. When this is the case, neighboring fingers are placed so that contact with the major groove is extended in direct succession, with each zinc finger add-ing three consecutive base pairs to the recognition sequence. Therefore, two tandem zinc fingers will recognize six consecutive base pairs of DNA, three zinc fingers will recognize nine base pairs, and so forth (Wolfe et al., 2000). Notably, higher eukaryotes can often have proteins containing ten or more repeated zinc finger domains, but in yeast zinc finger proteins one common-ly finds two or three fingers (Böhm et al., 1997).

This thesis focuses heavily on seven budding yeast transcription factors that all contain two tandem C2H2 zinc fingers; the two homologs Gis1 and Rph1, the two homologs Msn2 and Msn4, and the three homologs Mig1, Mig2 and Mig3 (Figure 1). Notably, the zinc fingers are the most well con-

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served parts between the homologous pairs (or triplet, in the case of Mig1, Mig2 and Mig3). Indeed, the amino acids thought to interact specifically with DNA are completely identical in each of the three cases, and Gis1 and Rph1 are therefore predicted to bind similar DNA sequences, as are Msn2 and Msn4, as well as Mig1, Mig2 and Mig3 (Böhm et al., 1997).

Figure 1. Schematic overview of the seven C2H2 zinc finger transcription factors under study in this thesis. JmjN, Jumonji N; JmjC, Jumonji C; ZnF, zinc finger.

In addition to DNA binding domains, transcription factors also contain other domains needed to perform their function. As has been mentioned, these might be activating or repressing domains that often act by recruiting coactivator or corepressor complexes, respectively (Hahn and Young, 2011). Another example, found for example in the Gis1 and Rph1 transcription factors (Figure 1), are the Jumonji N (JmjN) and Jumonji C (JmjC) domains. These two well conserved domains are present in all eukaryotes and were initially thought to always co-occur (Balciunas and Ronne, 2000). However, this view was widened when it was discovered that there are also many pro-teins containing only a JmjC domain but lacking JmjN, and it was then also hypothesized that JmjC might be an enzymatic domain (Clissold and Ponting, 2001). This hypothesis has been proven correct as JmjC is now known to be a domain capable of catalyzing the removal of methyl groups from histone proteins; this will be discussed in detail in the following sec-tion.

Rph1

Gis1

Msn2

Msn4

Mig1

Mig2

Mig3

JmjN JmjC ZnF

0 100 200 300 400 500 amino acids

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Chromatin modification The DNA which transcription factors and RNA polymerases must interact with is not naked, but is instead found in a densely compact form known as chromatin. The basic unit of chromatin is the nucleosome, composed of an octamer of histone proteins (two each of the core histones H2A, H2B, H3 and H4) around which 147 base pairs of DNA is wrapped (Kornberg and Lorch, 1999). Histones are rich in arginine and lysine residues, giving them a strong positive charge, which means that they bind well to the negatively charged sugar phosphate backbone of DNA. Nucleosomes therefore act as barriers to transcription, as they restrict transcription factors and RNA poly-merases from accessing the DNA (Li et al., 2007a). To overcome this, there are systems that dynamically remodel chromatin and displace nucleosomes; the ATP-dependent SWI/SNF complex, for example, which helps to main-tain a nucleosome-free region in gene promoters (Cairns, 2009; Jiang and Pugh, 2009; Mellor, 2005, 2006; Workman, 2006).

The histone proteins are mainly globular, but their N-terminal tails are unstructured and protrude out from the core. A noteworthy feature of his-tones, and in particular their N-terminal tails, is that they can be modified in a large number of ways. Thus, over 60 individual residues are known to be covalently modified by, for example, acetylation, methylation, phosphoryla-tion, ubiquitination and sumoylation (Kouzarides, 2007). In addition, DNA itself can be methylated, although this is absent in many fungi, including budding yeast. Significantly, there are numerous protein domains that bind specifically to these modified marks; bromodomains bind acetylated histone lysines, for example (Yun et al., 2011). Discovery of these domains led to the proposal of a ‘histone code’, the idea that chromatin contains a vast amount of accessible information, in addition to the underlying genetic code (Jenuwein and Allis, 2001; Strahl and Allis, 2000). However, subsequent work has revealed that there is elaborate crosstalk taking place between dif-ferent histone modifications, and that how, when and where a mark is depos-ited is often of crucial importance to the outcome, suggesting that histone modifications constitute a complex language rather than a strict code (Lee et al., 2010).

Histone acetylation and deacetylation The most well studied type of histone modification is lysine acetylation, and histone acetyltransferases (HATs) as well as histone deacetylases (HDACs) have been well characterized. Acetylation negates the positive charge of the lysine to which it is conjugated, weakening the binding between histone and DNA (Hong et al., 1993). Because nucleosomes make the DNA less accessi-ble, this has lead to the traditional view that acetylation leads to transcrip-tional activation via nucleosome disassembly, while deacetylation counter-

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acts this and leads to repression (Grunstein, 1997; Kornberg and Lorch, 1999; Kouzarides, 2007). For example, the SAGA and TFIID complexes both contain subunits with HAT activity, and this is thought to help them gain access to DNA and recruit TBP to the core promoter during transcrip-tional initiation (Grant et al., 1997; Mizzen et al., 1996).

As it counteracts acetylation, histone deacetylation is in general repressive to transcription, although there are exceptions, as will be seen below. A very well studied and important HDAC is the yeast Sir2 enzyme, which is ex-tremely well-conserved with homologs from bacteria to humans (Blander and Guarente, 2004; Imai et al., 2000; Landry et al., 2000). Sir2 is crucial for maintaining silencing of the yeast mating type loci, telomeres and ribosomal DNA (rDNA), and this also suppresses recombination at the rDNA locus, thereby extending yeast lifespan (Kaeberlein et al., 1999).

Another important HDAC is the budding yeast Rpd3, which, much like Sir2, exhibits striking evolutionary conservation (Rundlett et al., 1996; Yang and Seto, 2008). Rpd3 is a global regulator of gene expression, and even though it is mostly linked to repression, this is not strictly the case; Rpd3 is needed not only for maximum repression, but also for maximum activation of genes (Vidal and Gaber, 1991). Indeed, Rpd3 seems to have a role in op-posing the function of the Sir2 HDAC; deletion of RPD3 causes increased repression at telomeres, rDNA and mating type loci, and also leads to an extended yeast lifespan (Jiang et al., 2002; Kim et al., 1999; Sun and Hampsey, 1999; Zhou et al., 2009). Interestingly, Rpd3 functions in two distinct complexes, both of which have effects on transcription; the large and the small Rpd3 complexes (Rpd3(L) and Rpd3(S), respectively). These two complexes share the subunits Rpd3, Sin3 and Ume1, while Rpd3(S) also contains Eaf3 and Rco1, and Rpd3(L) contains Ash1, Dep1, Cti6, Pho23, Rxt2, Rxt3, Sap30, Sds3 and Ume6 (Carrozza et al., 2005a, 2005b; Keogh et al., 2005). Rpd3(L) has an established role in repression by histone deacetylation in promoter regions; it is recruited to specific promoter motifs by DNA-binding proteins such as Ume6 and Ash1 (Carrozza et al., 2005a; Kadosh and Struhl, 1997). In contrast, Rpd3(S) is recruited to the interior of actively transcribed genes and there works in close conjunction with Set2-mediated histone methylation; this will be discussed in detail further below.

Histone methylation Histone methylation, having received a lot of attention in recent years, has emerged as an important regulator of many biological processes (Martin and Zhang, 2005). Methylation can occur on arginine and lysine residues, and a level of complexity is added because these residues can be modified by more than one methyl group. Thus, arginines can be monomethylated, symmetri-cally dimethylated or asymmetrically dimethylated, while lysines can be monomethylated (me1), dimethylated (me2) or trimethylated (me3)

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(Bannister et al., 2002). In mammals, histone arginine methylation is found on residues 2, 8, 17 and 26 of histone H3 (H3R2, H3R8, H3R17 and H3R26) and on residue 3 of histone H4 (H4R3), while histone lysine methylation is found on residues H3K4, H3K9, H3K27, H3K36, H3K79 and H4K20 (Kouzarides, 2007).

In contrast to acetylation, the charge of the modified residue is not af-fected by methylation, indicating that the mark does not serve to disrupt histone-DNA binding. Instead, it is thought that it serves as a docking point for effector proteins, as a number of domains have now been found that bind to methylated histones. For example, chromodomains, tudor domains and plant homeodomain (PHD) fingers all bind to methylated histone lysines (Martin and Zhang, 2005; Yun et al., 2011). In fact, these domains often bind very specifically to a certain residue of a particular methylation state (a mono-, di- or trimethylated lysine, for example), showing that there is great specificity encoded in the methylation mark. The various effector proteins that bind also produce the result that histone methylation can be associated with both activation and repression of transcription. The general rule for lysine methylation is that H3K4, H3K36 and H3K79 are associated with transcriptional activity, while H3K9, H3K27 and H4K20 are connected to repression, but there are exceptions (Kouzarides, 2007). It should also be noted that being associated with activation or repression does not mean that the mark is the cause of this status. For example, methylation of H3K36, which will be discussed in more detail below, is a modification found in the interior of actively transcribed genes. However, this modification is not itself activating; instead it activates the Rpd3(S) complex, which acts repressively on these genes (see below).

Histone methylation in S. cerevisiae was long thought to be limited to H3K4, H3K36 and H3K79, catalyzed by the Set1, Set2 and Dot1 methyltransferases, respectively (Millar and Grunstein, 2006). It is notable that these modifications have all been found to correlate with transcriptional activity, while the methylated marks that are absent in budding yeast (H3K9, H3K27 and H4K20) are associated with repression (Pokholok et al., 2005). This picture has very recently been slightly altered by the findings of also H2BK37, H3R2, H3K42 and H4K20 as targets of methylation in budding yeast (Edwards et al., 2011; Gardner et al., 2011; Hyland et al., 2011; Kirmizis et al., 2007), as well as indications that low levels of methylated H3K9 might be present (Garcia et al., 2007). However, in all these cases the responsible enzymes are as of yet unknown.

The Set2/Rpd3(S) pathway In budding yeast, methylation of H3K36 is catalyzed by the histone methyltransferase Set2 (Strahl et al., 2002). This modification serves an im-portant role during transcriptional elongation, and is of special relevance to

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this thesis. Set2 interacts with RNA Pol II during transcriptional elongation, as it binds specifically to the hyperphosphorylated CTD (Krogan et al., 2003; Li et al., 2003; Schaft et al., 2003; Xiao et al., 2003). The methylation laid down by Set2 in the wake of the transcribing polymerase is in turn needed for histone deacetylation mediated by the Rpd3(S) complex; this acts repres-sively, thereby preventing erroneous transcription initiation from within open reading frames (Carrozza et al., 2005b; Joshi and Struhl, 2005; Keogh et al., 2005). Notably, Rpd3(S) is, much like Set2, recruited physically to actively transcribed regions by interacting with the phosphorylated CTD of RNA Pol II. However, it is only when H3K36 is methylated that Rpd3(S) actively deacetylates histones (Drouin et al., 2010; Govind et al., 2010). This is most likely due to stabilized binding as two of the subunits of Rpd3(S) have domains that cooperatively bind methylated H3K36; the chromodomain of Eaf3 and the PHD finger of Rco1 (Li et al., 2007c). Taken together, the effect of H3K36 methylation is to ensure that DNA made ac-cessible by transcriptional elongation is swiftly restored to a closed state by histone deacetylation. It is believed that the suppression of cryptic transcrip-tion initiation from within active genes by the Set2/Rpd3(S) pathway occurs genome-wide (Li et al., 2007b; Lickwar et al., 2009).

Interestingly, it has also recently been shown that the Set2/Rpd3(S) path-way enforces strong promoter directionality by suppressing antisense tran-scription, and also represses meiotic recombination (Churchman and Weissman, 2011; Merker et al., 2008). In addition, Set2 and Rpd3(S) has a small but highly significant repressive effect on DNA replication initiation globally, whereas Rpd3(L) has a larger effect on a subset of replication ori-gins (Knott et al., 2009). In support of this, an additional study (Biswas et al., 2008b) has implied that Set2 negatively regulates replication. Intriguing-ly, Set2 has also been reported to have a positive effect on DNA replication; a set2∆ or H3K36R mutation can thus reverse the accelerated S-phase seen in rpd3∆ cells (Pryde et al., 2009). Importantly, this last result shows that methylation of H3K36 by Set2 has a role even in the absence of Rpd3, and therefore demonstrates that the sole purpose of Set2 cannot be to facilitate recruitment of Rpd3(S). Methylation of H3K36 catalyzed by Set2 also has an additional role that is independent of the Rpd3(S) complex in that it pre-vents spreading of silencing from heterochromatin into adjacent euchromatic regions. Thus, deletion of SET2 or mutation of H3K36, but not deletion of the Rpd3(S)-specific subunits EAF3 and RCO1, gives rise to increased si-lencing in regions close to telomeres and mating type loci (Tompa and Madhani, 2007). Intriguingly, it has been shown that Rpd3 and subunits specific to Rpd3(L), but not Rpd3(S), give rise to an anti-silencing function highly reminiscent to that of Set2 (Zhou et al., 2009). This suggests that H3K36 methylation might also be linked to the Rpd3(L) complex by an as of yet unknown mechanism.

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Notably, some recent evidence suggests that the distinction between Rpd3(L) and Rpd3(S) might not be as clear-cut as previously thought. Thus, it has been reported that Rpd3(S) is found only in the interior of genes that also have Rpd3(L) bound to the promoter, which is hard to reconcile with the idea that Rpd3(L) is recruited to specific promoters while Rpd3(S) af-fects transcriptional elongation ubiquitously (Drouin et al., 2010). In addi-tion, subunits of Rpd3(L) have been detected within the coding region (Drouin et al., 2010; Govind et al., 2010), while subunits of Rpd3(S) have been observed to be recruited to gene promoters (Biswas et al., 2008a; Kre-mer and Gross, 2009). It has also been reported that the two complexes sometimes have opposing functions, and that the balance between Rpd3(L) and Rpd3(S) is of crucial importance in certain contexts (Biswas et al., 2008a). Based on some of these observations, it has been suggested that there is a single Rpd3 complex which alters its composition in the transition from transcriptional initiation to elongation, rather than two independent complexes (Drouin et al., 2010).

An additional piece of the puzzle might be that the Rpd3(S) subunit Eaf3, which binds methylated H3K36, is also a subunit of the NuA4 HAT complex (Doyon and Côté, 2004). It has been observed that the Rpd3(S) and NuA4 complexes appear to compete for binding in the ARG3 promoter, which is surprising as H3K36 methylation is not in general found in promoter regions and Rpd3(S) is not thought to function there (Biswas et al., 2008a). Interest-ingly enough, it has also recently been shown that NuA4 promotes transcrip-tional elongation by binding to the CTD of RNA Pol II. This cotranscriptional association is independent of the NuA4 subunits that nor-mally recruit the complex to promoter regions, but is instead strengthened by H3 methylation, although it remains to be determined if it is H3K36 methyl-ation specifically that is important (Ginsburg et al., 2009). Taking this new information into account, the NuA4 HAT complex behaves strikingly similar to the Rpd3(L) and Rpd3(S) HDAC complexes; Rpd3(L) contains subunits that recruit it to promoter regions, while Rpd3(S) is recruited to coding re-gions by association with the CTD of RNA Pol II, and is there activated by Eaf3-mediated binding to methylated H3. Furthermore, the NuA3 HAT complex requires Set2 and H3K36 methylation for its association with chromatin (Martin et al., 2006), and the PHD domain of the NuA3 subunit Nto1 binds specifically to H3K36me3 in vitro (Shi et al., 2007), suggesting that also this complex might be an important downstream target of H3K36 methylation.

An additional complicating factor is that Rpd3(S) in certain settings inter-acts with methylated H3K4, rather than H3K36; this appears to be mediated by the PHD finger of the Rpd3(S)-specific subunit Rco1 (Pinskaya et al., 2009; Pinskaya and Morillon, 2009). In this context it is also interesting to note that Rpd3(L) contains two subunits (Cti6 and Pho23) with PHD fingers that specifically recognize methylated H3K4 (Shi et al., 2007). Additionally,

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H3K36 is a target of acetylation by the SAGA HAT complex, besides meth-ylation by Set2 (Morris et al., 2007). However, it remains to be determined if SAGA and Set2 actively compete for this site, and what the relevance of this may be.

Taken together, methylation of H3K36 catalyzed by Set2 appears to have a role in several important processes, the best studied of which is the interac-tion with the Rpd3(S) complex. Many details remain to be elucidated, how-ever, as has been outlined above. An additional intriguing complication comes from the recent discovery of enzymes capable of opposing Set2; this aspect will be explored in the following two sections.

Histone demethylation For a long time, methylation of histones was believed to be a stable, irre-versible modification, owing to the fact that global turnover of the histone methyl groups occurs at the same speed as turnover of the histones them-selves (Bannister et al., 2002; Byvoet et al., 1972). Therefore, it was thought that for histone methylation to be counteracted, the nucleosome would have to be disassembled. There is no doubt that histone turnover has an important regulatory role (Dion et al., 2007), but it is unlikely that it exists to specifi-cally counteract methylation, as it would also remove all other modifications on the same molecule. As it turns out, however, histone methyl groups can also be enzymatically removed.

The first finding of an enzyme that could counteract histone methylation came when it was discovered that PADI4 could convert methylated arginines to citrulline, thus removing the methyl group (Cuthbert et al., 2004; Wang et al., 2004). Since this reaction leaves behind a citrulline residue rather than an unmodified arginine, it is not a strict removal of the methyl group and has therefore been coined demethylimination rather than demethylation. The first true histone demethylase to be identified was LSD1, which was found to be able to demethylate H3K4me1 and H3K4me2, thereby repressing transcrip-tion (Shi et al., 2004). Intriguingly, when LSD1 is in complex with the an-drogen receptor, it instead demethylates H3K9 and activates transcription, showing that the specificity of the enzyme can be modulated by its interact-ing partners (Metzger et al., 2005).

Because the reaction mechanism utilized by LSD1 requires a protonated nitrogen, it can only demethylate mono- and dimethylated lysines, and has no effect on the trimethylated state. This raised the suspicion that there could also be other types of enzymes with the ability to demethylate histones, and it was proposed that the JmjC domain might be capable of this, via a hy-droxylation reaction using Fe(II) and α-ketoglutarate as cofactors (Trewick et al., 2005). This hypothesis was quickly proven correct when the first JmjC-domain-containing histone demethylase, JHDM1, was discovered (Tsukada et al., 2006). A large number of studies have followed, firmly es-

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tablishing JmjC as a catalytic domain capable of removing mono-, di- and trimethylation from histone lysine residues (Klose et al., 2006a; Klose and Zhang, 2007). In addition, the crystal structure of the JmjC domain of the human JMJD2A protein has been solved, lending insight into which specific residues are responsible for cofactor-interaction (Chen et al., 2006). Notably, JmjC-domain-containing histone demethylases are very substrate specific, both with regard to the lysine residue and the methylated state. To date, JmjC demethylases capable of catalyzing the removal of all methylation states (me1, me2 and me3) from H3K4, H3K9, H3K27 and H3K36 have been characterized. In contrast, an enzyme removing H4K20me1, but not H4K20me2 and H4K20me3, has been identified, and no demethylase revers-ing H3K79 methylation has of yet been discovered (Lan and Shi, 2009; Pedersen and Helin, 2010; Qi et al., 2010). A noteworthy point is that only proteins from the JARID and JHDM3/JMJD2 groups (Klose et al., 2006a), which contain the JmjN domain in addition to JmjC, seem to be capable of demethylating trimethylated lysines, giving rise to the notion that this do-main is needed to affect the trimethylated state. It should be noted that sub-strates other than histone lysines are possible for JmjC-containing proteins, although this remains for the most part unexplored. The exception is the human JMJD6 protein, which has been reported to demethylate histone arginines (Chang et al., 2007) although this finding has been questioned and it has instead been suggested that JMJD6 hydroxylates a protein involved in RNA splicing, or even RNA directly (Hong et al., 2010; Webby et al., 2009).

Although recent years have seen great interest in JmjC proteins, much remains to be discovered concerning their functions and roles. Notably, many of them have connections to human disease, including cancer and neu-rological disorders (Pedersen and Helin, 2010; Shi, 2007), and endeavors to identify specific JmjC inhibitors as potential therapeutic agents are in pro-gress (Lohse et al., 2011).

Histone demethylation in budding yeast In S. cerevisiae, there is no homolog of the LSD1 enzyme, but there are five proteins containing JmjC domains: Jhd1, Jhd2, Ecm5, Rph1 and Gis1. Of the five, Jhd1 is the most distantly related, and is the only one that does not have a JmjN domain. It belongs to the JHDM1 group, while Jhd2 and Ecm5 be-long to the JARID group and Rph1 and Gis1 belong to the JHDM3/JMJD2 group (Klose et al., 2006a). Jhd1 is, like its two human homologues, an ac-tive histone demethylase targeting H3K36me1 and H3K36me2 (Kim and Buratowski, 2007; Tsukada et al., 2006; Fang et al., 2007). The in vivo role of Jhd1 is still unknown, but it has been reported that its PHD finger can interact specifically with H3K4me3, suggesting that Jhd1 might be involved in the interplay between H3K4 and H3K36 methylation (Shi et al., 2007). Furthermore, genome-wide location studies in strains that lack or overex-

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press Jhd1 show subtle shifts in the distribution of H3K36me2 across genes, indicating that its role may be to fine-tune this pattern (Fang et al., 2007).

Of the two JARID group members, Jhd2 and Ecm5, only Jhd2 is known to be an active enzyme, targeting all three methylated states of H3K4 (Huang et al., 2010; Ingvarsdottir et al., 2007; Liang et al., 2007; Seward et al., 2007; Tu et al., 2007). It also seems to be a negative regulator of silencing at telo-meres and mating-type loci, which fits with the fact that Set1, the enzyme that methylates H3K4, has a positive effect on such silencing (Liang et al., 2007; Osborne et al., 2009). Furthermore, Jhd2 has been suggested to have a role in both the activation and attenuation of inducible genes, by altering H3K4 methylation levels (Ingvarsdottir et al., 2007). Ecm5 has a large inser-tion in its JmjC domain and several substitution mutations in important cata-lytic residues (Balciunas and Ronne, 2000; Klose et al., 2006a), making it highly unlikely that it is an active enzyme. Nevertheless, Ecm5 has a PHD finger that can bind specifically to H3K36me3, indicating that it may have a chromatin-related role (Shi et al., 2007).

The remaining two JmjC-containing proteins in budding yeast are Rph1 and Gis1; they belong to the JHDM3/JMJD2 group and also share closely related DNA-binding zinc finger domains, as discussed earlier (Figure 1). Rph1 is a functional histone demethylase, with the ability to reverse H3K36me2 and H3K36me3, both in vitro and in vivo (Chang et al., 2010; Kim and Buratowski, 2007; Klose et al., 2007; Kwon and Ahn, 2011; Tu et al., 2007). Rph1 also has demethylase activity against H3K9, which is sur-prising as this residue is not thought to be methylated in budding yeast (Klose et al., 2007), although low amounts have been reported (Garcia et al., 2007). Interestingly, Rph1 was named after its known role as a repressor of the DNA-damage responsive PHR1 gene (regulator of PHR1), and this re-pression is dependent on the catalytic activity of the JmjC domain (Jang et al., 1999; Liang et al., 2011).

Given its specificity against H3K36, it has been suggested that demethylation by Rph1 promotes transcriptional elongation as it counteracts the Set2-mediated activation of the repressive Rpd3(S) complex which was discussed earlier. Thus, overexpression of RPH1 (or JHD1, which also demethylates H3K36) suppresses the near-lethality imposed by the absence of the positive elongation factor Bur1 (Kim and Buratowski, 2007). The significance of Rph1 in this process is not yet known, but it could be specu-lated that demethylation might counteract the Set2/Rpd3(S) pathway in one of two ways depending on the speed and frequency of the reaction. If the demethylation reaction is fast and frequent, it could occur before Rpd3(S) has been activated by binding the methylated mark, thus directly negating histone deacetylation. This might serve to fine-tune the outcome of the Set2/Rpd3(S) pathway, or to directly oppose it under specific circumstances. Alternatively, if demethylation occurs slowly or infrequently, the reaction might take place after deacetylation by Rpd3(S), thus having more of a

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house-keeping role to ensure that chromatin is eventually returned to the original state after the Set2/Rpd3(S) pathway has played its course.

It is also interesting to speculate about the substrate specificity of Rph1 with regard to the methylation states of H3K36. The Set2 methyltransferase catalyzes all three methylation states (me1, me2 and me3) of H3K36 in bud-ding yeast, but there are differences in how this comes about. Thus, the cata-lytic SET domain of Set2 is in itself sufficient for catalyzing H3K36me1 and -me2, but H3K36me3 is dependent on Set2 interacting with the phosphory-lated CTD of RNA Pol II as well as additional factors such as the Spt6 pro-tein; this suggests that the different methylation states are differentially regu-lated and have distinct functions (Fuchs et al., 2012; Youdell et al., 2008). Indeed, the Rpd3(S) complex binds H3K36me2 and -me3 in vitro, but not H3K36me1 or the unmodified form, and H3K36me2 is sufficient to facilitate recruitment of Rpd3(S) (Li et al., 2009). Furthermore, it has recently been suggested that H3K36 methylation might have both a positive and negative role in transcriptional regulation; H3K36me2 recruits the repressive Rpd3(S) complex while H3K36me3 recruits activating complexes such as NuA3 and NuA4 (Fuchs et al., 2012). Pryde and coworkers (2009) have also reported that H3K36me1 and H3K36me3 have opposing functions in regulating the timing of DNA replication, although H3K36me2 was not investigated in that study. In this context it is interesting to note that Rph1 demethylates H3K36me2 and -me3, while Jhd1 demethylates H3K36me1 and -me2. It is therefore theoretically conceivable that Rph1 and Jhd1 together constitute a system for controlling the state of H3K36 methylation; lack of Rph1 would lead to an enrichment of H3K36me3, while lack of Jhd1 would lead to H3K36me1 accumulation, for example. However, there are no experimental indications that these two JmjC proteins specifically function together and a rph1∆ jhd1∆ double mutant does not display any obvious phenotype (Klose et al., 2007).

Unlike Rph1, Gis1 is likely to be an inactive enzyme due to the fact that it has a histidine-to-tyrosine substitution in one of the iron-coordinating resi-dues of the JmjC domain (Klose et al., 2006a). In support of this, several groups have failed to establish any enzymatic activity when testing Gis1 against a number of methylated substrates; H3K4 (Fang et al., 2007; Huang et al., 2010), H3K36 (Fang et al., 2007; Kim and Buratowski, 2007), H3K79 (Fang et al., 2007; Huang et al., 2010) and H4K20 (Edwards et al., 2011). In one case (Tu et al., 2007), there have been indications that Gis1 is an active demethylase, capable of demethylating H3K36me1 and H3K36me2. How-ever, it should be noted that this conclusion was based on accumulation of methylation states in the knockout strain, which means that it would also be perfectly compatible with a model in which Gis1 somehow negatively regu-lates Rph1, or competes with Rph1 (since Rph1 demethylates H3K36me2 and H3K36me3, one would expect to see an accumulation of H3K36me1 and H3K36me2 in cells where Rph1 activity was enhanced). Apart from

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histone demethylation, Gis1 has a well established role as a transcriptional activator, and it has recently been discovered that the JmjC domain is not required for this function (Yu et al., 2010; Zhang and Oliver, 2010).

When discussing the potential catalytic activity of Gis1, it is noteworthy that the Schizosaccharomyces pombe JmjC protein Epe1 has a substitution mutation identical to the one in Gis1 that renders it enzymatically inactive against histone substrates (Tsukada et al., 2006; Zofall and Grewal, 2006). Despite this, Epe1 has an important role in preventing spreading of silencing from heterochromatic regions and, furthermore, this function relies on the JmjC domain (Ayoub et al., 2003; Trewick et al., 2007; Zofall and Grewal, 2006). This indicates that JmjC domains might also have a function which is independent of its enzymatic activity, or that an alternative enzymatic activi-ty directed against non-histone substrates may exist. It is also interesting that other JmjC proteins have a role similar to that of Epe1; to protect euchromatin from heterochromatic spreading (Tamaru, 2010), a role also ascribed to the Set2 methyltransferase (Tompa and Madhani, 2007).

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Nutrient signaling and aging

The ability to sense environmental conditions and respond accordingly is of vital importance to all living cells. This is as true for the individual cells in a multicellular organism as for a single-celled microorganism, where the cell constitutes the entire organism. Therefore, signaling pathways have evolved that enable cells to sense extracellular cues and adapt their metabolism, growth, proliferation and stress resistance. These pathways are generally very well-conserved in all eukaryotes, although the extracellular cue is al-most always some sort of nutrient in microorganisms, where it might be a growth factor or hormone in a multicellular organism. Because of this con-servation, budding yeast has been an excellent model system to improve our understanding of how these pathways transmit their signal.

An important function controlled by these pathways is that of aging and regulation of lifespan. There is a link between nutrient availability and aging, clearly illustrated by caloric restriction (CR), also known as dietary re-striction, a phenomenon which delays aging and extends lifespan in response to a reduced food intake (Masoro, 2005). As sensing nutrient availability is exactly what the above discussed signaling pathways do, it is not surprising that they have turned out to be important in mediating CR. Strikingly, CR appears to be a universal phenomenon found in both unicellular and multi-cellular organisms. Thus, a yeast cell will have an extended lifespan when starved (Lin et al., 2000), as will a rhesus monkey (Colman et al., 2009). Yeast has therefore turned out to be a surprisingly useful model organism in the endeavor to understand the fundamental question of why we age and die (Kaeberlein, 2010).

Linked to the phenomenon of aging, another very important function con-trolled by these signaling pathways is the decision of when a cell should divide. Indeed, the vast majority of cells in a multicellular eukaryotic organ-ism are in a resting, non-dividing state referred to as quiescence or G0. The entry into and exit from this state are tightly controlled processes, problems with which can have dire consequences in the form of cancer. Yeast can also enter a resting state, often referred to as stationary phase; it is brought about by nutrient limitation and enables the cells to survive for long periods of time. Importantly, some earlier assumptions have recently been called into question by the discovery that a stationary phase yeast culture is actually a heterogeneous mixture made up of two types of cell; quiescent cells, which retain the ability to exit stationary phase and divide, and nonquiescent cells,

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which are alive but rapidly lose the ability to reproduce (Allen et al., 2006; Aragon et al., 2008; Davidson et al., 2011). Undoubtedly, some earlier work will have to be revisited in light of these findings. Nevertheless, many as-pects of yeast cell adaptation for stationary phase are conserved in how cells of higher eukaryotes exit the cell cycle into G0, showcasing the value of this simple model organism. This chapter will start by describing how yeast cells grow and finally enter into stationary phase, followed by an overview of key signaling pathways, and finish off with a discussion of the connections to aging.

The growth phases of budding yeast Basically, yeast cells will enter stationary phase when they run out of one or more nutrients. However, S. cerevisiae is somewhat special with regard to the carbon source as it has adapted to a fermentative lifestyle. This means that it will always anaerobically ferment glucose as long as this preferred carbon and energy source is available, before turning to aerobic respiration and use of other carbon sources. This may seem wasteful, as pyruvate, in-stead of being shunted into the TCA cycle, is metabolized into ethanol which is exported from the cell. However, it is believed that this strategy gives budding yeast an adaptive advantage as it facilitates a more rapid growth combined with the fact that ethanol is inhibitory to many competing micro-organisms. Besides, budding yeast has good capacity to utilize the produced ethanol once glucose has run out. The outcome of this is that yeast cells grown in batch culture, given that no other nutrient is limiting, will go through several distinct growth phases as they shift their carbon metabolism (Figure 2). Notably, yeast that depletes a nutrient other than glucose will not pass through these phases, but instead enter directly into stationary phase (Smets et al., 2010).

When yeast cells are exposed to glucose, they adapt their metabolism to fermentation during what is called the lag phase (Figure 2). This is followed by the exponential phase, or logarithmic phase, where the cells divide rapid-ly, with a doubling time of around 90 minutes. During the exponential phase, as discussed above, the cells ferment glucose into ethanol, which is released into the medium. This is accommodated by induced expression of genes needed for glycolysis and glucose uptake, together with repression of genes involved in the TCA cycle, mitochondrial function, and uptake and metabo-lism of other carbon sources. To facilitate rapid growth, there is also a high expression of genes needed for protein synthesis, such as tRNAs and ribo-somal proteins (DeRisi et al., 1997). When the glucose is finally depleted, a major change known as the diauxic shift occurs, whereby the cell alters its metabolism from fermentation to respiration. This is accomplished by large changes in gene expression; almost 2000 genes, accounting for a third of the

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yeast genome, are significantly up- or downregulated (DeRisi et al., 1997). This basically leads to a reversal of the gene expression patterns described above for the exponential phase; for example, genes coding for proteins in-volved in glycolysis are now repressed, while genes needed for a functioning TCA cycle are activated. Furthermore, genes required for stress resistance and long-term survival are induced. After the diauxic shift follows a period of slower growth during which ethanol is aerobically metabolized, known as the post-diauxic growth phase. Finally, all ethanol will be consumed, and the cells then enter the stationary phase.

Figure 2. Schematic overview of the various growth phases S. cerevisiae goes through when growing in batch culture with glucose as carbon source.

Nutrient signaling pathways The most important nutrient signaling pathways in budding yeast are illus-trated in Figure 3 and will be discussed in this chapter (also recently re-viewed in De Virgilio, 2012 and Smets et al., 2010). These pathways begin with the sensing of an extracellular nutrient; how this is accomplished is known in some cases, in others much remains to be discovered. This is fol-lowed by signal transduction, which has generally been thoroughly investi-gated. Almost all the pathways utilize evolutionary conserved protein kinas-es to transduce the signal by phosphorylation, the downstream outcome be-ing regulation of one or more transcription factors, which in turn alters gene expression. Illustrated in Figure 3, an emerging theme is that there is a vast degree of cross-talk between pathways, with signals often impinging on the same target to yield a more graded response (De Virgilio, 2011). One might therefore be better off imagining a large and elaborate nutrient-sensing net-work, although for simplicity’s sake pathways are often discussed separately.

Exponentialphase

Lagphase

Diauxicshift

Post-diauxicphase

Stationaryphase

Time

Log

cell

num

ber

Fermentation

Respiration

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Figure 3. Nutrient signaling pathways in S. cerevisiae. Arrows and bars represent positive and negative interactions, respectively. Dashed, bidirectional arrows repre-sent proteins that shuttle between cytoplasm and nucleus. For simplicity’s sake, the net outcome of an interaction (positive or negative) is shown, rather than the specific details. For example, PKA and TORC1 are both depicted as negatively regulating Rim15, even though PKA directly inactivates the kinase domain of Rim15 while TORC1 inhibits Rim15 nuclear localization. See text for further details. cAMP, cyclic adenosine monophosphate; HXT, hexose transporter; PDS, post-diauxic shift; PHO, phosphate metabolism; Ribi, ribosome biogenesis; RP, ribosomal protein; STRE, stress response element.

Glucose repression and the Snf1 kinase Glucose is a primary energy source for most cells, and it is therefore crucial that they are able to sense and respond to its presence. This is especially important for an organism such as S. cerevisiae which as described previous-ly has a fermentative lifestyle, meaning that it will preferentially ferment all the available glucose before turning to respiration and alternative carbon sources. Therefore, yeast has several pathways that sense and respond to glucose (Johnston, 1999; Rolland et al., 2002; Santangelo, 2006). One of these is the glucose repression pathway, which ensures that large groups of genes are tightly repressed as long as glucose is present (Ronne, 1995). The-se include genes involved in gluconeogenesis, the TCA cycle, respiration and the uptake and metabolism of other carbon sources.

A key player in the glucose repression pathway is the protein kinase Snf1, which is inactivated in the presence of glucose, resulting in repression of the above mentioned genes (Hedbacker and Carlson, 2008). The Snf1 kinase is evolutionary well-conserved in all eukaryotes and is known as the AMP-

HXTgenes

MIG3 MIG2 PHOgenes

PDSgenes

STREgenes

Glucose-repressed

genes

UV-inducedgenes

Ribi and RPgenes

Nitrogenutilization

genes

Msn2/4

Pho4

Gis1

Gpa2 Gpr1Cyr1Ras1/2

TORC1Rgt2Snf3

Pho81Snf1 PKA

Yck1/2

Pho80Pho85

Std1Mth1

Sch9

Rim15

Mig2 Mig1 Mig3Rgt1 Gln3 Pho4

Rim15

Msn2/4

UVirradiation

Glucose Nitrogen StressPhosphate

Amino acids

cAMP

Mig1Gln3

Cytoplasm

Nucleus

Extracellularmedium

Rph1

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activated protein kinase (AMPK) in animals and as the Snf1-related kinase (SnRK1) in plants (Hardie, 2007; Woods et al., 1994). It should be men-tioned that AMPK is an extremely important metabolic sensor regulating many aspects of cellular function, and that Snf1 likewise is involved in many processes besides glucose repression (Hardie, 2011; Hedbacker and Carlson, 2008). Like all members of the AMPK family, Snf1 functions in a heterotrimeric complex composed of a catalytic α-subunit (Snf1), a regulato-ry β-subunit (Gal83, Sip1 or Sip2), and a regulatory γ-subunit (Snf4) (Ghillebert et al., 2011; Hedbacker and Carlson, 2008).

The most important downstream effector in glucose repression is Mig1 (Figure 1), a DNA-binding zinc finger protein which acts as a transcriptional repressor through its interaction with the Cyc8/Ssn6-Tup1 corepressor com-plex (Klein et al., 1998; Nehlin and Ronne, 1990; Treitel and Carlson, 1995). As mentioned above, Snf1 is active in the absence of glucose, and it then regulates Mig1 by direct phosphorylation, which abolishes the interaction between Mig1 and Cyc8/Ssn6-Tup1 and causes nuclear exclusion of Mig1, leading to derepression of target genes (De Vit et al., 1997; Ostling and Ronne, 1998; Papamichos-Chronakis et al., 2004; Smith et al., 1999; Treitel et al., 1998). Repression by Mig1 is often direct, as it binds physically to a well-characterized promoter motif; (C/G)(C/T)GG(G/A)G (Lundin et al., 1994), but can in addition be indirect as in the case of the GAL genes, where Mig1 represses the transcriptional activator GAL4 (Nehlin et al., 1991).

Yeast also has two other zinc finger proteins closely related to Mig1: Mig2 and Mig3 (Figure 1; Lutfiyya et al., 1998). The residues of the zinc fingers thought to determine DNA binding specificity are conserved among the three proteins (Böhm et al., 1997), and indeed, Mig1, Mig2 and Mig3 have all been shown to physically bind the above described promoter motif in vitro (Badis et al., 2008; Lutfiyya et al., 1998; Lutfiyya and Johnston, 1996; Zhu et al., 2009). Mig2 acts redundantly with Mig1 in the regulation of glucose-repressed genes (Lutfiyya et al., 1998; Lutfiyya and Johnston, 1996; Westergard et al., 2007). However, Mig2 does not seem to be regulat-ed by Snf1, nor is its nuclear localization affected by glucose as is the case for Mig1 (Lutfiyya et al., 1998). Instead, expression of the MIG2 gene is induced by the glucose induction pathway, which will be described below (Kaniak et al., 2004). Mig3 is phosphorylated by Snf1, although this leads to proteolysis rather than nuclear exclusion, as is the case for Mig1 (Dubacq et al., 2004). Intriguingly, MIG3 is, just like MIG2, induced by glucose, alt-hough Mig3 does not affect glucose-repressed genes as do Mig1 and Mig2 (Kaniak et al., 2004; Lutfiyya et al., 1998). Instead, it has been suggested that Mig3 acts as a downstream effector of Snf1 in the UV response (Dubacq et al., 2004; Wade et al., 2009). In summary, (i) MIG2 and MIG3 are in-duced by glucose; (ii) Mig1 and Mig3 are inactivated by Snf1; (iii) Mig1 and Mig2 redundantly repress glucose-repressed genes; and (iv) Mig3 regulates gene expression in response to UV damage.

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The glucose induction pathway To grow well on glucose, yeast needs not only glucose repression, but also the specific induction of genes encoding enzymes involved in uptake and metabolism of glucose. The best studied example is the regulation of the numerous HXT genes, which encode glucose transporters of various affini-ties (Johnston, 1999; Rolland et al., 2002; Santangelo, 2006). These are re-pressed by the transcription factor Rgt1 in the absence of glucose, but can be induced by a signaling cascade when glucose becomes available. Thus, the membrane-bound Snf3 and Rgt2 proteins generate an intracellular signal in response to glucose which activates the Yck1 and Yck2 kinases (Moriya and Johnston, 2004; Özcan et al., 1996). Yck1/2 in turn phosphorylates the regu-latory proteins Mth1 and Std1, which triggers their degradation, leading to inactivation of the Rgt1 repressor (Lakshmanan et al., 2003; Moriya and Johnston, 2004). As mentioned above, Rgt1 also represses MIG2 (and MIG3), setting up an intricate crosstalk between the glucose induction and repression pathways (Kaniak et al., 2004). In addition, Rgt1 is phosphory-lated and thereby inactivated by PKA (see below), further highlighting how interconnected the various signaling pathways are (Kim and Johnston, 2006).

The PKA pathway The protein kinase A (PKA) pathway, also known as the Ras/cyclic AMP (cAMP) pathway, is another evolutionary well-conserved pathway, which in yeast responds to the presence of glucose. Contrary to the Snf1 kinase dis-cussed above, PKA is active in nutrient rich conditions, and has a key inhibi-tory role in the transition from exponential growth to the post-diauxic and stationary phases. Thus, cells with a deficiency in the PKA pathway behave as if starving, even when glucose is abundant (Rolland et al., 2002; Santangelo, 2006). PKA functions as a heterotetramer composed of two catalytic subunits (Tpk1, Tpk2 or Tpk3), and two regulatory subunits (Bcy1) (Toda et al., 1987a, b). PKA is activated when the inhibitory Bcy1 subunits dissociate from the catalytic subunits upon binding of the second messenger cAMP, which is produced in the cell in response to extracellular glucose. Glucose can be sensed by two separate branches of this pathway; the Ras1 and Ras2 proteins are activated by intracellular glucose-6-phosphate, while the G protein Gpa2 directly senses extracellular glucose via the transmembrane receptor Gpr1 (Santangelo, 2006; Smets et al., 2010). Both Ras1/2 and Gpa2 activate the adenylate cyclase Cyr1, which catalyzes the conversion of ATP into cAMP. Thus, glucose stimulates a spike in cAMP production, leading to PKA activation. PKA then phosphorylates a number of proteins, but a key target is the Rim15 protein kinase, which will be dis-cussed below.

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The TOR pathway The target of rapamycin (TOR) protein kinase is another key controller of cell growth responding to environmental cues, in particular nitrogen availa-bility (Jacinto and Hall, 2003; Loewith and Hall, 2011; Rohde et al., 2001; Schmelzle and Hall, 2000; Wullschleger et al., 2006). Originally identified genetically in yeast by mutations that confer dominant resistance to the im-munosuppressant rapamycin, recent years have seen great interest in TOR as rapamycin has striking anti-aging effects in several organisms (Bjedov and Partridge, 2011; Heitman et al., 1991; Sharp and Richardson, 2011). Signifi-cantly, TOR is conserved in all eukaryotes so far examined, although yeast has two kinases, Tor1 and Tor2, whereas higher eukaryotes have only one (Wullschleger et al., 2006).

The TOR kinases at first posed somewhat of an enigma as it was evident they served in two separate roles; Tor1 and Tor2 act redundantly in a path-way that is very specifically inhibited by rapamycin, while Tor2 also has a specific, rapamycin independent, function (Zheng et al., 1995). However, this was settled with the discovery that these separate functions are carried out by two distinct multiprotein complexes; the rapamycin-sensitive TOR Complex 1 (TORC1) and the rapamycin-insensitive TORC2 (Loewith et al., 2002). In yeast, TORC1 can contain either Tor1 or Tor2 as the catalytic sub-unit, while TORC2 always contains Tor2 (Loewith et al., 2002). Higher eu-karyotes, as mentioned, have only a single TOR kinase, but the two distinct complexes, along with their functions and response to rapamycin, are con-served (Loewith et al., 2002). Interestingly, both TOR complexes are local-ized in association with membranes (Wedaman et al., 2003). TORC1 is thus predominantly found at the vacuolar membrane, while TORC2 mainly local-izes in discrete dots at the plasma membrane (Aronova et al., 2007; Sturgill et al., 2008).

TORC1 regulates a multitude of processes in response to nutrients (Jacin-to and Hall, 2003; Loewith and Hall, 2011; Wullschleger et al., 2006). Thus, TORC1 affects protein levels by promoting translation initiation and inhibit-ing autophagy (Barbet et al., 1996; Noda and Ohsumi, 1998). TORC1 also inhibits expression of genes needed for nitrogen source utilization and stress resistance by sequestering the Gln3, Gat1, Msn2 and Msn4 transcription factors in the cytoplasm (Beck and Hall, 1999; Cardenas et al., 1999; Hard-wick et al., 1999). Significantly, localization of these transcription factors is also controlled by Snf1, suggesting a convergence between the TORC1 and Snf1 signaling pathways (Bertram et al., 2002; Mayordomo et al., 2002). Expression of ribosomal protein genes is also controlled by TORC1; when TORC1 is inactivated these genes are strongly repressed and this repression is mediated by the Rpd3(L) HDAC complex (Cardenas et al., 1999; Humph-rey et al., 2004; Powers and Walter, 1999; Rohde and Cardenas, 2003). In addition, TORC1 activates genes involved in glycolysis, while repressing

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genes involved in the TCA cycle (Hardwick et al., 1999). Taken together, it is clear that TORC1 is major regulator of growth; when TORC1 is active, growth is promoted, while inactivation of TORC1, by rapamycin treatment or upon nutrient limitation, rapidly leads to entry into G0.

In contrast to TORC1, less is known about TORC2, but it is an essential complex that controls important processes such as actin cytoskeleton organi-zation and sphingolipid synthesis (Aronova et al., 2008; Cybulski and Hall, 2009; Jacinto et al., 2004; Loewith and Hall, 2011; Schmidt et al., 1996). In yeast, these functions are largely mediated by the TORC2 substrate Ypk2 (Kamada et al., 2005).

As mentioned, rapamycin is a very specific inhibitor of TORC1, and has thus been an extremely useful tool in elucidating TORC1 function. However, there are other TORC1 inhibitors available, synthetically manufactured as well as naturally derived (Zhou et al., 2010). One of these is the small com-pound caffeine; it inhibits TORC1 but is significantly more pleiotropic than rapamycin and has therefore not been as widely used for studies of TORC1 function (Kuranda et al., 2006; Lum et al., 2004; Reinke et al., 2006; Wanke et al., 2008).

The Sch9 kinase Another very important protein kinase in yeast nutrient signaling is Sch9. Originally, Sch9 was thought to be partly redundant with PKA; overexpres-sion of SCH9 suppresses deficiencies in the PKA pathway, while lack of Sch9 can conversely be suppressed by increased PKA activity (Toda et al., 1988). However, it was later established that Sch9 and PKA exert their ef-fects through parallel but distinct signaling pathways, sometimes acting syn-ergistically and sometimes antagonistically (Roosen et al., 2005). Recently, Sch9 has been shown to be an important downstream effector of TORC1; Sch9 is directly phosphorylated by TORC1, and ribosome biogenesis, trans-lation initiation and proper entry into G0, all important TORC1 readouts, depend largely on Sch9 (Urban et al., 2007). Indeed, Sch9 has been termed a central coordinator of protein synthesis as it regulates both RNA Pol I and III, as well as ribosomal protein and biogenesis genes transcribed by RNA Pol II, all in a TORC1-dependent manner (Huber et al., 2009, 2011; Jorgen-sen et al., 2004). Interestingly, this regulation is mediated by recruitment of the Rpd3(L) HDAC complex (Huber et al., 2011). However, Sch9 also has roles that are clearly TORC1-independent, acting in parallel with and some-times opposing TORC1 (Smets et al., 2008). Sch9 was also recently discov-ered to be directly phosphorylated by Snf1, at sites distinct from those used by TORC1 (Lu et al., 2011). This places Sch9 at a convergence point be-tween Snf1 and TORC1 signaling, and suggests that it is a central coordina-tor of cellular aging (see below).

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Lastly, it should be mentioned that Sch9 is homologous to the mammalian protein kinase B (PKB/Akt), an important oncogene (Sobko, 2006). Howev-er, Akt is regulated by TORC2 which does not seem to be the case for Sch9 (Urban et al., 2007). Therefore, it has been proposed that Sch9 might rather be the functional counterpart of mammalian S6K1, which, just like Sch9, is regulated by TORC1 and directly phosphorylates the critically important ribosomal protein S6 (Powers, 2007; Urban et al., 2007).

Pho85-Pho80 signaling In addition to glucose and nitrogen limitation, entry into G0 can also be trig-gered by phosphate starvation. This is regulated by Pho85, a cyclin-dependent protein kinase that can partner up with ten different cyclins, one of which is Pho80, to direct its activity to different substrates (Huang et al., 2007). When phosphate is abundant, Pho80-Pho85 is active and phosphory-lates the transcription factor Pho4, thereby excluding it from the nucleus (O’Neill et al., 1996). Upon phosphate limitation, Pho81 inhibits Pho80-Pho85, which leads to Pho4 localizing to the nucleus where it activates genes involved in phosphate scavenging and metabolism (Mouillon and Persson, 2006). In addition, Pho81 inhibition of Pho80-Pho85 can also promote entry into G0 (Swinnen et al., 2005). This is carried out by controlling the nuclear localization of the Rim15 protein kinase (see below).

Rim15 and downstream transcription factors The protein kinase Rim15 is a downstream master regulator of central im-portance, as it integrates and transmits signals from many of the pathways discussed above (Figure 3; Swinnen et al., 2006). In essence, Rim15 is kept in an inactive state by signals from the upstream pathways. Upon nutrient limitation, however, these signals cease and Rim15 becomes active. Thus, in the cytoplasm, the kinase domain of Rim15 is inactivated by PKA-dependent phosphorylation (Reinders et al., 1998). Working in conjunction with this, Rim15 is prevented from entering the nucleus via phosphorylation at Ser1061 by Sch9 and at Thr1075 by Pho80-Pho85, modifications that facilitate binding to the 14-3-3 proteins Bmh1/2, which thereby anchors Rim15 in the cyto-plasm (Pedruzzi et al., 2003; Wanke et al., 2005, 2008). Notably, the Sch9-mediated phosphorylation at Ser1061 is dependent on TORC1, consistent with Sch9 being a downstream effector of TORC1 as discussed above (Pedruzzi et al., 2003; Urban et al., 2007; Wanke et al., 2008). However, TORC1 also independently affects the phosphorylation status of Thr1075, possibly by in-hibiting a phosphatase (Urban et al., 2007; Wanke et al., 2005). Taken to-gether, four of the nutrient signaling pathways outlined above; PKA, TORC1, Sch9 and Pho80-Pho85, serve to maintain Rim15 in an inactive state in the cytoplasm when nutrients are present.

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The consequence of Rim15 becoming active and translocated to the nu-cleus is that it in turn activates three downstream zinc finger transcription factors; Msn2, Msn4 and Gis1 (Figure 1; Swinnen et al., 2006). Msn2 and Msn4 are functionally redundant and regulate gene expression by binding to the STRE (stress response element; AG4) motif in the promoters of genes induced upon stress and at the diauxic transition (Boy-Marcotte et al., 1998; Martinez-Pastor et al., 1996; Schmitt and McEntee, 1996). Notably, although Rim15 is clearly epistatic over Msn2 and Msn4 in many regards, the activity of these transcription factors is also regulated by other processes (Galdieri et al., 2010; Smets et al., 2010). For example, much like Rim15 itself, Msn2 and Msn4 are kept inactivated by cytoplasmic sequestering, a level of regu-lation exerted by PKA, TORC1 and Snf1, but independent of Rim15 (Beck and Hall, 1999; Gorner et al., 1998; Mayordomo et al., 2002; Roosen et al., 2005).

Gis1 was originally isolated as a multicopy suppressor of a snf1∆ mig1∆ srb8∆ triple mutant (Balciunas and Ronne, 1999), but has later been estab-lished as a Rim15-dependent transcriptional activator at the diauxic transi-tion, acting mainly through the PDS (post-diauxic shift; T(A/T)AG3AT) motif present in promoters of genes needed for stress resistance and metabo-lism (Pedruzzi et al., 2000; Zhang et al., 2009). Subsequent work has shown that there is a substantial degree of functional overlap between Msn2/4 and Gis1, with regards to target genes as well as their preference for STRE and PDS promoter motifs (Cameroni et al., 2004; Roosen et al., 2005). Con-sistent with Msn2/4 and Gis1 cooperatively mediating much of the effect of Rim15 is the finding that the entire set of genes requiring Rim15 for activa-tion at the diauxic transition is included within the combined set of genes that are activated by Msn2/4 and Gis1 (Cameroni et al., 2004). However, Gis1 also has Rim15-independent functions; it both represses and activates genes in glucose-limited cells, but only gene activation seems dependent on Rim15 (Zhang et al., 2009). Noteworthy in this context is that Sch9 seems to modulate Gis1 activity directly, independently of Rim15 (Roosen et al., 2005). Intriguingly, Sch9 can be physically recruited to promoters of stress-responsive genes, opening up the possibility that Gis1 is modulated at its site of action (Pascual-Ahuir and Proft, 2007).

Interestingly, Gis1 is also required for induction of genes involved in spore wall assembly during sporulation (Coluccio et al., 2004; Yu et al., 2010), and it has recently been discovered that Gis1 is regulated through limited proteolysis by the proteasome (Quan et al., 2011; Zhang and Oliver, 2010). The reason for this proteolysis is thought to be to ensure that Gis1 activity is tightly regulated by Rim15, thereby allowing the cell to rapidly resume growth when nutrients become abundant (Bonzanni et al., 2011).

As mentioned earlier, Gis1 also has a homolog, Rph1 (Figure 1). The two proteins are 34% identical overall, but the parts of the zinc finger domains thought to interact with DNA are 100% identical, suggesting they recognize

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the same sequence (Jang et al., 1999). Historically, Rph1 has not been thought to play a part in nutrient signaling as it does not directly affect acti-vation of selected STRE- and PDS-driven reporter genes, which are instead entirely dependent on Msn2/4 and Gis1, respectively (Pedruzzi et al., 2000). However, Gis1 and Rph1 are redundant repressors of the DNA damage-responsive PHR1 gene, and this repression is dependent on a STRE motif in the PHR1 promoter, to which Rph1 is known to physically bind (Jang et al., 1999). In addition, Gis1 and Rph1 both repress DPP1, a gene encoding a diacylglycerol pyrophosphate phosphatase, and this regulation is mediated by three PDS promoter motifs, to which Gis1 binds physically (Oshiro et al., 2003). Furthermore, Rph1 has been identified as a factor binding the STRE motif in several genomic studies (Badis et al., 2008; Treger et al., 1998; Zhu et al., 2009).

Another piece of evidence tying Rph1 to nutrient signaling is that it be-comes phosphorylated at a number of serine residues (Ser425, Ser426, Ser430, Ser459, Ser557 and Ser561) upon treatment with the TORC1 inhibitor rapamy-cin (Huber et al., 2009). Interestingly, it has also been reported that a residue distinct from these (Ser587) is significantly less phosphorylated in rapamycin-treated cells (Soulard et al., 2010). It should also be mentioned that DNA damage, in a Rad53-dependent manner, induces phosphorylation of Rph1 (Chen et al., 2010; Kim et al., 2002), which is thought to lead to dissociation of Rph1 from the PHR1 promoter, followed by derepression of this DNA repair gene (Liang et al., 2011). In this context it is intriguing that Rph1 has been found to be hypophosphorylated in cells lacking the Snf1 kinase (Zhang et al., 2011), as Snf1 in addition to its metabolic function is a regula-tor of the transcriptional response to DNA damage (Wade et al., 2009).

Very recently it was also reported that Rph1 may be involved in cell wall integrity (CWI) signaling (Kuravi et al., 2011; Levin, 2005, 2011). This is especially interesting as CWI signaling is a process in which both TORC1 and TORC2 have been implicated (Ho et al., 2005; Levin, 2005; Loewith and Hall, 2011; Soulard et al., 2010; Torres et al., 2002).

It is unknown if Rph1 is regulated through limited proteolysis by the pro-teasome, as is its homolog Gis1 (Quan et al., 2011; Zhang and Oliver, 2010). However, it is interesting to note that the human JmjC protein which is most closely related to Gis1 and Rph1, KDM4A/JMJD2A, is regulated by ubiqui-tin-mediated degradation by the proteasome (Tan et al., 2011; Van Rechem et al., 2011) In addition, the S. cerevisiae JmjC protein Jhd2, as well as Epe1 in S. pombe, are regulated in this manner (Braun et al., 2011; Mersman et al., 2009).

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Nutrient signaling as a regulator of aging There has been intense interest in the phenomenon of aging since the realiza-tion that aging might be a genetically determined, program-like process, rather than an unavoidable deterioration, which opens up possibilities to modulate it (Blagosklonny, 2006). Much of the work in this field has been done using model organisms; mice, fruit flies, nematodes and yeast. Studies in budding yeast have proven extremely fruitful as there are striking similari-ties between cellular and organismal aging, i.e. many of the same factors affect the lifespans of single celled yeast and entire multicellular organisms (Kaeberlein, 2010). There have been two models of cellular aging estab-lished in yeast: replicative aging and chronological aging. Replicative lifespan (RLS) is measured as the number of daughters a mother cell can produce before senescence, and models aging of mitotically active cells in multicellular organisms (Steinkraus et al., 2008). In contrast, chronological lifespan (CLS), measured as the time a cell can remain viable when kept in a non-dividing state, serves as a model for the aging of post-mitotic cells (Fabrizio and Longo, 2003).

As mentioned at the start of this chapter, there is a strong connection be-tween the aging process and the nutrient signaling pathways outlined above. This connection comes about because these very well conserved pathways mediate caloric restriction (CR), a phenomenon whereby lifespan can be extended by reduction of food intake (Masoro, 2005). The effects of CR exhibit striking evolutionary conservation; CR extends both RLS and CLS in yeast (Lin et al., 2000; Smith et al., 2007), and also prolongs the lifespan of all multicellular organisms that have so far been tested, from nematodes to primates (Fontana et al., 2010).

Since the nutrient signaling pathways mediate CR, one would expect that manipulating them, by genetic mutation or chemical inhibition, would affect aging. And indeed, yeast strains with deficiencies in these pathways are strikingly long-lived; in effect the cells behave as if they are starving, even when nutrients are abundant. Thus, tor1∆ and sch9∆ strains have extended lifespans, in terms of both RLS and CLS (Fabrizio et al., 2001; Kaeberlein et al., 2005; Wei et al., 2008). Likewise, strains deficient in the PKA pathway, such as cyr1∆ and ras2∆, have extended RLS and CLS, and strains with an overly active PKA pathway, such as bcy1∆ or mutants with a constitutively active Ras2, have shortened RLS and CLS (Fabrizio et al., 2001, 2003; Lin et al., 2000; Sun et al., 1994; Wei et al., 2008). In particular for TORC1, there is strong evidence that this connection to aging is very well conserved also in higher eukaryotes (Stanfel et al., 2009).

As has been mentioned, the immunosuppressant rapamycin is a very spe-cific inhibitor of TORC1. Consistent with this, the drug has anti-aging prop-erties; treatment with low doses of rapamycin extends both yeast RLS and CLS (Lu et al., 2011; Powers et al., 2006). Furthermore, in 2009 a widely

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cited study showed that rapamycin significantly extends the lifespan of mice, and strikingly enough, this was seen also when the drug was administered late in life (Harrison et al., 2009). This has prompted discussion of rapamycin as an anti-aging drug for humans, especially since it also has anti-cancer properties (Bjedov and Partridge, 2011; Kaeberlein and Kennedy, 2009; Sharp and Richardson, 2011). Besides rapamycin, there are other in-hibitors of TORC1 being investigated, derivatives of rapamycin, termed rapalogs, as well as naturally occurring compounds (Zhou et al., 2010). One of these natural substances is caffeine, and it extends the CLS of yeast when administered in low doses (Wanke et al., 2008).

As discussed above, Rim15 is an important downstream mediator of sev-eral of the signaling pathways affecting longevity. In line with this, a dele-tion of RIM15 is epistatic over and reverses the lifespan-extending pheno-types of tor1∆, ras2∆, cyr1∆ and sch9∆ strains with regards to CLS, alt-hough the effects on RLS have not been thoroughly investigated (Fabrizio et al., 2001; Wei et al., 2008). Furthermore, a rim15∆ strain does not benefit from the prolonged CLS granted by CR or by treatment with low doses of rapamycin and caffeine (Wanke et al., 2008; Wei et al., 2008).

Downstream of Rim15 are the three transcription factors Msn2, Msn4 and Gis1. Accordingly, deletion of the genes encoding these proteins reverses the lifespan extension gained from treatment with CR or signaling pathway dis-ruption, at least for CLS, which is what has been studied in detail. Thus, a msn2/4∆ gis1∆ strain has its CLS only slightly increased by CR, as opposed to the striking extension seen in a wild type (WT) strain (Wei et al., 2008). Interestingly, this strain nearly, but not completely, resembles the CLS phe-notype of rim15∆, suggesting that some other factor(s) downstream of Rim15 might contribute to CLS extension upon CR. Furthermore, gis1∆ reverses the extended CLS of sch9∆ and ras2∆ strains (Wei et al., 2008), while msn2/4∆, interestingly enough, can reverse the prolonged lifespan of ras2∆ and cyr1∆, but not sch9∆ (Fabrizio et al., 2001, 2003, 2004b). As for RLS, the effects of Gis1 have not been investigated while the effects of Msn2/4 are inconclusive; it has been reported that deletion of MSN2/4 (i) has no effect on the RLS extension of a cdc25-10 mutant (Lin et al., 2000); (ii) completely abolishes RLS extension from CR (Medvedik et al., 2007); and (iii) acts synergistically with a CYR1 mutant to extend RLS (Fabrizio et al., 2004b). The effects of the Gis1 homolog Rph1 on longevity have not been studied at all.

The model that emerges is that shutdown of the PKA pathway or TORC1 (and thereby also Sch9), by treatment with CR or specific inhibitors such as rapamycin or caffeine, leads to activation and nuclear localization of Rim15, which in turn activates Msn2/4 and Gis1 (see Figure 3). Regulation of gene expression by these transcription factor then promotes longevity, although exactly how is unknown (see below). It should be noted that this model in its entirety only applies to CLS; the upstream part of this signaling cascade

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(CR, rapamycin, TORC1, PKA and Sch9) all affect RLS and CLS similarly, but the effects of the downstream part (Rim15, Msn2/4 and Gis1) on RLS have not been investigated or are ambiguous, as discussed above. Also, as mentioned earlier and adding complexity to this model, the nuclear localiza-tion of Msn2/4 is directly regulated by PKA, TORC1 and Snf1 (Beck and Hall, 1999; Gorner et al., 1998; Mayordomo et al., 2002; Roosen et al., 2005), and Gis1 has been suggested to be regulated by Sch9 independently of Rim15 (Roosen et al., 2005).

It is not clear which the target genes downstream of Gis1 and Msn2/4 are that mediate the effects on CLS and RLS. Much evidence points to an im-portant role of induced resistance to oxidative stress in prolonging CLS, and genes involved in this response are therefore candidates, especially the su-peroxide dismutases SOD1 and SOD2 (Cameroni et al., 2004; Fabrizio et al., 2004a; Madia et al., 2009; Mesquita et al., 2010; Weinberger et al., 2010). Thus, deleting SOD2 decreases CLS, and also abolishes the CLS extension seen in sch9∆ and ras2∆ strains (Fabrizio et al., 2003; Weinberger et al., 2010). Conversely, overexpressing SOD1 and SOD2 prolongs CLS, but in-triguingly shortens RLS (Fabrizio et al., 2001, 2003, 2004b). It has also been suggested that chronological aging is mediated specifically by a transient exposure to accumulated acetic acid (Burtner et al., 2009). In support of this, long-lived strains such as sch9∆ and ras2∆ exhibit increased resistance to acetic acid, a phenotype which can be reversed by deletion of RIM15 or GIS1. A further observation is that production of glycerol protects cells against stress, thereby prolonging CLS (Wei et al., 2009). Thus, the glycerol biosynthesis genes GPD1, GPD2 and RHR2 are among the most upregulated genes in sch9∆, tor1∆ and ras2∆ strains, and at least for sch9∆, this effect is reversed by deletion of GIS1. Furthermore, the prolonged CLS of sch9∆ can be reversed by deleting these three glycerol biosynthesis genes (Wei et al., 2009). Finally, it has also been shown that RLS extension by CR is depend-ent on Msn2/4-mediated induction of the nicotinamidase PNC1 gene (Medvedik et al., 2007).

Suggesting a mechanism of aging regulation acting in parallel to the one outlined above, it was very recently shown that the Snf1 kinase regulates RLS in yeast independently of CR (Lu et al., 2011). Thus, when the Snf1 inhibitor Sip2 is acetylated, it binds Snf1 and the aging process is slowed. Interestingly, the acetylation status of Sip2 is controlled by the NuA4 acetyltransferase complex and the Rpd3 deacetylase, both previously thought to mediate their effects primarily through histone acetylation and deacetylation. Downstream of Snf1, and directly phosphorylated by it, is Sch9, known to be a direct target of also TORC1. As lifespan regulation by Snf1 occurs independently of TORC1 and CR, it was proposed that Snf1 and TORC1 define two parallel pathways to control aging that converge on Sch9 (Lu et al., 2011). However, it should be mentioned that there are still unan-swered questions with this model as loss of SnRK1, the plant homolog of

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Snf1, causes accelerated senescence in the moss Physcomitrella (Thelander et al., 2004). Similarly, AMPK, the mammalian homolog of Snf1, seems to have an inhibitory effect on aging in higher eukaryotes; activation of AMPK thus leads to a prolonged lifespan (Lu et al., 2011; Mair et al., 2011, and references therein).

Lastly, something should be said about Sir2, founding member of the sirtuin histone deacetylase family, around which much of the early aging research in yeast revolved. Sir2 has been demonstrated to prolong RLS in yeast by silencing the rDNA locus, thereby repressing the formation of extrachromosomal rDNA circles (ERCs), which are toxic to the cell (Kaeberlein et al., 1999). Notably, ERC formation does not contribute to the aging process in multicellular organisms, but it was nevertheless reported that increased sirtuin activity extends the lifespan of not only yeast, but also the nematode Caenorhabditis elegans and the fruitfly Drosophila melano-gaster (Rogina and Helfand, 2004; Tissenbaum and Guarente, 2001). Fur-thermore, it was claimed that sirtuins are the mediators of the lifespan-extending effect of CR in all three of these organisms; CR alters the NAD+/NADH ratio in the cell thereby activating the sirtuins which in turn slows the aging process (Lin et al., 2000; Rogina and Helfand, 2004; Wang and Tissenbaum, 2006). However, much of this work has proved controver-sial; it has been demonstrated that CR can affect aging independently, and in parallel to the effects of the sirtuins, and Sir2 has been shown to have the opposite effect on yeast CLS to that of RLS (Fabrizio et al., 2005; Garber, 2008; Kaeberlein et al., 2004). Indeed, it was very recently also reported that the previously claimed effects of sirtuins on the lifespan of C. elegans and Drosophila could be explained by second-site mutations and lack of appro-priate controls (Burnett et al., 2011). Therefore, while it is clear that Sir2 affects yeast RLS by preventing ERC formation, it is highly questionable if it acts as a mediator of CR, or how relevant sirtuins are for aging in higher eukaryotes. Nevertheless, sirtuins do appear to have a protective effect against many aging-related diseases in mice, such as diabetes and cancer, making them a relevant field of study even if they do not directly increase longevity in all eukaryotes (Herranz and Serrano, 2010).

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Aims

The general aim of this thesis has been to gain a deeper understanding of how nutrient signaling pathways allow cells to adapt to a changing environ-ment. As these signaling pathways are strikingly conserved throughout eu-karyotic evolution, the simple model organism budding yeast has been used in this investigation. Furthermore, the focus has been on a number of bud-ding yeast transcription factors that directly contact DNA to activate or re-press transcription in response to signals from the upstream pathways, with a particular interest in one transcription factor, Rph1, of which very little was previously known. The specific objectives to achieve the more general aim have thus been:

• To gain a more comprehensive view of how the three homologous

transcription factors Mig1, Mig2 and Mig3 regulate transcription in response to glucose, with a particular focus on combinatorial effects (Paper I).

• To gain a fuller understanding of how the two homologous tran-scription factors Gis1 and Rph1 regulate transcription as the cell transitions through different growth phases (Paper II).

• To investigate to what extent the known histone demethylase ac-tivity of Rph1 is needed for its role in transcriptional regulation (Paper III).

• To elucidate whether or not Rph1 is involved in regulation of CLS alongside the three related transcription factors Gis1, Msn2 and Msn4, all known to be actors in this process (Paper IV).

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Results and discussion

Paper I In paper I, combinatorial gene regulation by the homologous transcription factors Mig1, Mig2 and Mig3 is investigated. Thus, microarrays are used to transcriptionally profile a WT strain, as well as all seven possible combina-tions of deleting the MIG1, MIG2 and MIG3 genes. This is done in cells growing logarithmically in the presence of glucose (2%), as Mig1 is known to be an important mediator of glucose repression (Klein et al., 1998). We find that Mig1 and Mig2 function redundantly in regulating target genes, with Mig1 in general being more important.

Interestingly, an exception to this, with Mig2 having a greater effect than Mig1, is in the repression of HXT6. This gene encodes a high-affinity glu-cose transporter, and follow-up experiments reveal that Mig2 has a large repressive effect of several HXT genes at higher glucose concentrations (10%). For example, the repression of HXT4 is entirely dependent on Mig1 at 2% glucose, but at 10% glucose it is redundantly repressed, with Mig2 having the strongest repressive effect. Taken together, our results suggest that Mig2 is redundant with and of slightly less importance than Mig1 under conditions of normal laboratory growth, but appears to be more important in an environment of high glucose abundance.

In contrast to the findings for Mig1 and Mig2, very few genes are affected by the MIG3 deletion under the conditions investigated; it does not appear to cooperate with Mig1 and Mig2 in regulating gene expression. Interestingly, though, one gene significantly repressed by Mig3 is SIR2, encoding a HDAC involved in silencing and yeast lifespan regulation (Kaeberlein et al., 1999).

Paper II In an approach similar to that in Paper I, microarrays are used to study the effects on gene expression of the two homologous transcription factors Gis1 and Rph1. Prior to this study, Gis1 was an established downstream effector of the highly conserved TORC1, PKA and Sch9 nutrient signaling pathways, acting as an activator of gene expression at the diauxic shift, when the yeast cell switches from a fermentative to a respiratory metabolism (Cameroni et al., 2004; Pedruzzi et al., 2000; Roosen et al., 2005; Zhang et al., 2009). In

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this role, Gis1 functions in parallel with the redundant transcription factors Msn2 and Msn4, and the generally held view was that Gis1 acts through PDS promoter motifs, while Msn2 and Msn4 mediate gene regulation through STRE motifs. In contrast, much less was known about the role of Rph1 in gene regulation, although it had been shown that it acted redundant-ly with Gis1 in repressing the DPP1 and PHR1 genes (Jang et al., 1999; Oshiro et al., 2003), and Rph1 had several times been reported as a factor physically binding the STRE promoter motif (Badis et al., 2008; Jang et al., 1999; Treger et al., 1998; Zhu et al., 2009). As a complete picture of combi-natorial gene regulation by Gis1 and Rph1 was lacking, this study attempts to address this lack of knowledge. We use microarrays to monitor the effects on gene expression of four strains; WT, gis1∆, rph1∆ and gis1∆ rph1∆, at three time-points; logarithmic phase, right after the diauxic shift, and after three days growth.

Our results show that Gis1 and Rph1 are both repressors and activators of gene expression, often acting redundantly or synergistically on the same genes, but also on distinct targets. Significantly, the roles of Gis1 and Rph1 are determined largely by the growth phase; during exponential growth the two transcription factors have a high degree of redundancy and mainly act as repressors of genes containing PDS and STRE motifs in their promoters. In contrast, after the cells have passed the diauxic shift, Gis1 and Rph1 are predominantly activators of gene expression and their target genes are more diverged; Gis1 has a bias towards activating genes with the PDS promoter element, while Rph1 activates mainly genes containing the STRE motif.

We also find that the extracellular levels of acetate and glycerol are af-fected in cultures of yeast cells lacking GIS1 and RPH1. This is interesting as both acetate and glycerol have been reported to be involved in regulation of CLS, a process where Gis1 is known to be involved (Burtner et al., 2009; Wei et al., 2009). Significantly, these discovered phenotypes are consistent with the patterns we see on gene expression; the deletions of GIS1 and RPH1 affect genes involved in glycerol and acetate metabolism. Intriguingly, we also find that several genes involved in acetyl-CoA metabolism are represssed by Gis1, and we hypothesize that there may be a link between this metabolite and the known effects on CLS mediated by Gis1.

Paper III In this paper we investigate the connection between the two known roles of Rph1; an activator and repressor of gene expression (as found in Paper II), and a histone demethylase capable of catalyzing the removal of methyl groups from H3K36 (Chang et al., 2010; Kim and Buratowski, 2007; Klose et al., 2007; Kwon and Ahn, 2011; Tu et al., 2007). We do this by using microarrays to study the effects on gene expression of a yeast strain contain-

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ing an allele of Rph1 with a JmjC domain that has been catalytically inacti-vated by a single point mutation (rph1-H235A). This point mutation has many times over been shown to render Rph1 completely catalytically inac-tive (Chang et al., 2010; Kim and Buratowski, 2007; Klose et al., 2007; Tu et al., 2007). As controls in this experiment we use a WT strain as well as a rph1∆ knockout, and we also study effects on gene expression at two differ-ent growth phases, logarithmic phase and after the diauxic shift, where we have previously (Paper II) seen that Rph1 has rather different functions. In addition, to be able to study the effects of this mutation in the absence of Gis1, we add three more strains to the analysis: gis1∆, gis1∆ rph1-H235A and gis1∆ rph1∆.

The most important finding of this paper is that the histone demethylase activity of Rph1 is largely dispensable for its role in transcriptional regula-tion, as measured under these conditions. Thus, the rph1-H235A strain be-haves much more like the WT than like the rph1∆ strain, and gis1∆ rph1-H235A behaves like gis1∆ rather than like gis1∆ rph1∆, both with regard to whole-genome patterns (principal component analysis and hierarchical clus-tering) and the number of genes affected above a set threshold. However, we do observe more subtle patterns; genes known to be up- or downregulated in the absence of Set2, the enzyme catalyzing methylation of H3K36, are in general affected in the opposite direction by the rph1-H235A mutant, and genes in the subtelomeric regions, known to be downregulated in a set2∆ strain (Tompa and Madhani, 2007), are slightly upregulated in the mutant.

We also find that genes affected in cells lacking the Rpd3 HDAC are dif-ferentially expressed in our strains. Rpd3 is the catalytic subunit of two dis-tinct complexes; Rpd3(L) is recruited to promoters while Rpd3(S) functions within the coding region of genes and depends on H3K36 methylation for its activity (Carrozza et al., 2005b; Keogh et al., 2005). Consistent with Rpd3(S) being a downstream effector of H3K36 methylation, and recapitu-lating the patterns seen for set2∆, we detect a weak trend to the effect that genes up- or downregulated by deletion of subunits specific to Rpd3(S) are affected in the opposite direction by the rph1-H235A mutant. In contrast, genes affected by deletion of RPD3 itself, as well as Rpd3(L)-specific subu-nits, are in general upregulated by deletion of RPH1 and GIS1, but unaffect-ed by the rph1-H235A mutation.

Taken together, the results can best be explained by Rph1 having two separate roles; it binds to the promoters of some genes and thereby activates and represses them (together with Gis1), but can also oppose Set2 activity, which is likely to occur in the interior of active genes, although this is not known for certain. The fact that the effects in the catalytically inactive mu-tant are so modest suggests that this role is one of fine-tuning, though we cannot rule out that it has more dramatic effects under conditions not yet investigated.

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Paper IV In paper IV, the role of Rph1 in determining CLS is studied. An important finding of our previous work (Papers II and III) is that there is a large over-lap in what genes are repressed and activated by the two homologs Gis1 and Rph1, and it is therefore plausible that they are involved in the same pro-cesses. Gis1 is, together with Msn2 and Msn4, known to be an important downstream target of Rim15 in mediating effects on chronological aging (Wei et al., 2008). Surprisingly, the main finding of this paper is that Rph1 does not affect CLS or resistance to acetic acid and other stress factors known to correlate with lifespan. However, we do observe that Rph1 has an effect on growth in the presence of the two TORC1 inhibitors rapamycin and caffeine; overexpression of RPH1 can partially suppress the rapamycin sen-sitivity of a tor1∆ strain, and a gis1∆ rph1∆ double mutant, but not the single deletion strains, is sensitive to caffeine. Furthermore, we find that a deletion of RPH1 is epistatic over a deletion of RIM15 with respect to rapamycin sensitivity. This suggests that Rph1 mediates effects of TORC1 signaling, although these effects are not directly related to lifespan regulation. The finding that GIS1 and RPH1 are redundant with regards to caffeine sensitivi-ty is also interesting in light of the recent discovery that RPH1 is involved in CWI signaling, a process associated with caffeine sensitivity (Kuravi et al., 2011).

This paper also includes an unbiased screen for genes that can, similarly to RPH1, suppress the rapamycin sensitivity of a tor1∆ strain when overex-pressed. We identify eight such suppressors, the strongest of which is URE2. As Ure2 is a well established downstream effector of TORC1 (Beck and Hall, 1999), this finding serves as a proof of concept for our screen. Another of the suppressors is NYV1, encoding a v-SNARE protein involved in ve-sicular fusion (Nichols et al., 1997). Further experiments reveal that NYV1 is a specifically potent suppressor of tco89∆. This is interesting as Tco89 is a subunit of TORC1 which has been shown to serve in a role distinct from the Tor1 kinase in regulating the vacuole import and degradation pathway, a process responsible for quickly degrading gluconeogenic enzymes when the cell encounters glucose (Brown et al., 2010; Reinke et al., 2004). Signifi-cantly, NYV1 has been implicated in this pathway (Brown et al., 2003), and our finding therefore suggests new opportunities to further probe the role of TORC1 in vacuolar function. Other interesting suppressors that we identify include PRE5, a gene encoding a proteasomal subunit (Heinemeyer et al., 1994), and ZDS2, encoding a protein recently described as a regulator of protein phosphatase 2A, an important downstream effector of TORC1 (Loewith and Hall, 2011; Rossio and Yoshida, 2011).

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Future perspectives

Several possible lines of continued research have spawned from the results included in this thesis. The most intriguing and promising of these will be briefly discussed in this chapter.

As has been discussed, Gis1 is a mediator of chronological aging (Wei et al., 2008); we have confirmed this result and added the finding that Rph1 does not affect this process (Paper IV). However, the effects of these tran-scription factors on the replicative lifespan remain unexplored. One of the reasons for this is that the standard assay for RLS is time-consuming and arduous; budding daughter cells are counted and removed by micromanipu-lation after each cell division (Park et al., 2002). One way of overcoming this is the ‘mother enrichment program’, which uses a genetically modified yeast strain where the ability of daughter cells to reproduce can be eliminat-ed through an inducible system (Lindstrom and Gottschling, 2009). Thus, the viability of a yeast culture over time becomes a direct measure of RLS, which can be assessed by simple plating assays. Using this strain, it would be comparably trivial to delete GIS1 and RPH1 and investigate the effect on RLS. As has been mentioned, results concerning the effects of the transcrip-tion factors Msn2 and Msn4 on RLS are also inconclusive; it has been re-ported that deletion of MSN2/4 shortens RLS (Medvedik et al., 2007), as well as extends it (Fabrizio et al., 2004b). It would therefore be interesting to also measure the RLS of strains deleted for MSN2/4, or better yet to investi-gate the roles of all four transcription factors, reminiscent of the analysis we have carried out for CLS (Paper IV).

Another investigative course would be to examine were Gis1 and Rph1 physically bind throughout the genome, using for example the ChIP-on-chip technique. This could potentially shed light on the questions arising from the discrepancy between the notion that Gis1 and Rph1 bind to STRE and PDS motifs in gene promoters, and the observation that H3K36 methylation, the known target of Rph1 demethylation, occurs foremost within the coding region of transcribed genes. However, we have had problems successfully tagging the Gis1 and Rph1 proteins with epitopes, and the fact that Gis1 has recently been discovered to be regulated by proteolysis further complicates any analysis of Gis1 function (Zhang and Oliver, 2010). In addition, using ChIP, several groups have been unsuccessful at detecting yeast JmjC pro-teins, including Rph1, suggesting that their interaction with DNA is weak or transient (Ingvarsdottir et al., 2007; Kim and Buratowski, 2007). An alterna-

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tive approach to elucidate the function of Rph1 would be to assay the amounts of H3K36 methylation using ChIP-on-chip in cells with RPH1 de-leted or mutated. Such data might be used in conjunction with our gene ex-pression profiling of rph1∆ and rph1-H235A strains to gain a clearer picture of the effect of Rph1-mediated histone demethylation on transcription. Be-cause of the epistatic interactions we noticed between gis1∆ and rph1-H235A with regards to for example expression of subtelomeric genes (Paper III) it would also be interesting to include strains with GIS1 deleted in such an analysis.

The recent discovery that Gis1 is regulated by limited proteolysis (Quan et al., 2011; Zhang and Oliver, 2010) is very interesting and raises several questions, one of which is whether or not Rph1 is regulated in the same manner. Especially intriguing is the fact that other JmjC proteins have been found to be regulated by proteasomal degradation; KDM4A/JMJD2A in mammalian cells (Tan et al., 2011; Van Rechem et al., 2011), Jhd2 in bud-ding yeast (Mersman et al., 2009), as well as Epe1 in fission yeast (Braun et al., 2011). It would therefore be interesting to conduct a careful biochemical analysis of Rph1 protein levels before and after treatment with a proteasome inhibitor such as MG132. In the context of proteolysis as a form of regula-tion, it is also interesting to speculate about the two separate roles we have proposed for Rph1; JmjC domain-dependent demethylation of H3K36, most likely occurring in the interior of transcribed genes, and specific regulation of genes containing STRE and PDS promoter elements (Paper II), which appears to be largely independent of the JmjC domain (Paper III), but most likely requires the zinc fingers, as these are known to bind the STRE motif (Badis et al., 2008; Jang et al., 1999; Treger et al., 1998; Zhu et al., 2009). It is theoretically conceivable that a selective processing of Rph1 might be the basis for these two roles; cleavage of the Rph1 protein yielding the N-terminal part containing the JmjC domain and the C-terminal part with the zinc fingers. A way to address this would be to assay if proteasome function has any impact on the demethylase activity of Rph1. For example, one could investigate if proteasome inhibition affects the ability of RPH1 overexpres-sion to suppress the growth defect of a bur1∆ strain, a phenotype that is de-pendent on the ability to demethylate H3K36 (Kim and Buratowski, 2007).

It would also be interesting to examine how and when Rph1 is phos-phorylated. Intriguingly, Rph1 has turned up as a target of phosphorylation in several large-scale screens (Albuquerque et al., 2008; Chen et al., 2010; Chi et al., 2007; Huber et al., 2009; Li et al., 2007d; Smolka et al., 2007; Soulard et al., 2010; Zhang et al., 2011). Since several pieces of our data indicate that Rph1 acts downstream of Rim15 (Papers II and IV), it would be especially interesting to investigate the phosphorylation status of Rph1 in a rim15∆ strain. For example, the hyperphosphorylation of Rph1 in cells treat-ed with rapamycin (Huber et al., 2009) could be tested for Rim15-dependence.

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The patterns we noticed indicating distinct functional overlap between Gis1/Rph1 and the Rpd3(S) and Rpd3(L) complexes (Paper III) are interest-ing and could be followed up experimentally. To summarize briefly, we detected a weak trend that genes affected by deletion of EAF3 and RCO1, encoding Rpd3(S)-specific subunits, were in general affected in the opposite direction by the catalytically inactive rph1-H235A mutant. In contrast, genes affected by deletion of RPD3 itself, as well as Rpd3(L)-specific subunits, were in general always upregulated in our GIS1 and RPH1 deletion strains, with little to no effect of the rph1-H235A mutation. This lead us to hypothe-size that Rph1 functions (together with Gis1) in a demethylation-independent role that involves Rpd3(L), but also has a role in demethylation of H3K36, which is mechanistically linked to Set2 and Rpd3(S). Further support for this notion comes from a recent study that investigated interac-tions between components of the chromatin machinery by analyzing the gene expression profiles of 165 yeast mutants (Lenstra et al., 2011). This work revealed that, as expected, the profile of set2∆ clustered tightly togeth-er with eaf3∆ and rco1∆, strains deleted for subunits specific to the Rpd3(S) complex. In contrast, the gene expression profile of rph1∆ clustered closest to the profiles of strains deleted for RPD3 and Rpd3(L)-specific subunits. One way of further investigating these observations would be to probe for epistatic effects between Gis1/Rph1 and subunits of Rpd3(S)/(L). As readout in such an assay one could use genes whose expression we have found to be differentially expressed in strains where GIS1 and RPH1 have been deleted (Papers II and III) or, alternatively, the rapamycin and caffeine phenotypes we have discovered for gis1∆ and rph1∆ cells (Paper IV).

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Acknowledgements

There are a number of people without which, truth be told, this text would never have seen print.

First of all, I must thank my supervisor, Hans. Your passion for science has been an inspiration, and you oftentimes surprised me with your kindness as a person. You always believed in me and supported me, even when I myself had doubts. The scientific road we travelled turned out to be bumpier than we hoped, but in the end I am glad we took it. I have learned a lot during these years.

My fellow coworkers in the group deserve an honor-heaped mention, as they have directly or indirectly assisted me in getting to this point. Zhen, your work has been so very important, and it has been extremely nice working alongside you during all these years. Thank you! Ida, your work during the-se last years has also been invaluable, and I have really enjoyed all our lunches and scientific discussions. Jacke, as my mind seems to move with glacial speed when it comes to computery stuff, your bioinformatic supervi-sion and assistance was crucial, and you were also an extremely pleasant person to work with. Sanna, you taught me much about science when I was fresh and naïve, and I enjoyed sitting opposite to you in the lab during many a year (you made the trek through the TAP-träsk bearable). Eva, my cosupervisor, you were a solid foundation of our group for many years, and were always a great person to talk to. Tina, your whacked-out sense of hu-mor did a lot to lighten the mood in the lab; never again have I thought of penguins in the same way. Marie, I really enjoyed sitting next to you in the lab for several years; your scientific input was always level-headed and val-uable (even though you laughed mercilessly at my pathetic skills with the tetrad microscope!). Anders, mein doppelganger, I very much enjoyed spending the major part of my PhD alongside you; your cheerfulness cou-pled with the occasional outburst of mortar and pestle-induced rage did much to make our corridor a great working environment. Micke, thanks for many research project-related talks, as well as for sharing my unwholesome passion for loincloth-clad men flexing their biceps under a blistering Dark Sun. Mattias, you are always very friendly and helpful and I continue to enjoy our talks on the most varied of subjects. I am also happy to have worked alongside all the other past and present members of our group;

51

Mattias T, Gunilla, Christine, Andrea, Jenni H, Alvaro, as well as my project students Liwei and Kerstin.

The ‘pombe’ group also deserves a special mention; I was really glad we shared our lab with you for all those years. Pernilla, my second cosupervisor, thanks for all your input on the project; you are always a great person to talk to. Jenny, besides Anders you were my fellow PhD student from day one; it was a fun ride even though you both beat me to the mark with a few too many years! Carolina, talking to you about both science and child rearing strategies lightened up those sometimes very early mornings.

I am also thankful for the collaborations we had with Jan Komorowski and Adam Ameur.

I’d also like to extend my general gratitude to all the nice people at IMBIM who made my years there so pleasant, especially the members of IPhAB and the various PhD students I’ve shared teaching duties with over the years. Also, thanks to all the friendly folks at Mikro who have welcomed me as one of their own.

I am also grateful to friends outside of work that have assisted me mostly in the department of gaining perspective. Helena, Johanna and Pelle, my old study buddies, I really enjoyed hanging with you back in the day and have seen much too little of you lately. Tobias and Michael, thanks for a number of crazy schlager-parties during the early years, followed by pleasant dinners as we all grew older and wiser (who am I kidding?). Mattias and Åsa, for all the relaxed and fun times spent hanging out with kids. Gustav and Sara, for all the dinners and game nights on Stabby Allé. Thanks also to all our new Storvreta neighbors and friends; there is always a place to go if you are desperately out of diapers, or feel the sudden urge to vent about sleepless nights over a glass of wine. I am also grateful to my gaming buddies, the Örebro gang (Peter, Jocke, Jonas, Morgan and Urban), as well as the Uppsala crew (Gustav, Adam, Johan, Mats and Micke) for helping me get some much needed diversion during these years, whether it’s been struggling against elemental evil, kicking nazi übermensch butt, or just enjoying a good old Middenheim carnival. Last but not least, my two oldest friends and brothers-from-other-mothers. Magnus, even though your outward appear-ance is that of a grim Spartan warrior, you are my go-to person if ever I need a male shoulder to cry on. You are, indeed, my best man. Peter, when we were young, you had a great part in helping me find myself and become the person I am today (that can be a compliment or an accusation, depending on perspective!), and the good times we’ve shared are many; late nights at the Terminal, bizarre meals in New Orleans, and all those nice dinner and movie evenings.

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My extended family deserves thanks for always making me feel so very wel-come. Sven and Britt-Marie, you have kept me ridiculously well-fed during all these years. Agneta and Ingvar, it is always fun to see you, and I am especially grateful for the steady supply of ‘mormors smutsiga bullar’. Em-ma, Jonas, Sonja and the newcomer Stellan; your experience with deter-mined young ladies has been invaluable to partake of, and I always look forward to the next time we’ll pack our families together and hang out.

I am also grateful to my parents; I know you are always there for me, and that is a great comfort. My mother, Joany, you are so kind-hearted and thoughtful, and I can only hope that I will display some of these traits to-wards my own children. My father, Leo, you always believed in me, and probably contributed specifically to this thesis when you guided me into studying natural science and math in high school (instead of Latin!). My sister Katie and my brother Jonathan, we have so many shared memories, and I am also very happy to have gotten to know your families; Mattias, little Hugo, and Hanna.

I am also immensely pleased to have been joined on this ride by Max and Molly, small in stature but occupying a significant part of my heart. Think-ing of you two always puts a silly smile on my face.

Last of all I come to Ida, my love; you are my best friend and have taught me more than anyone. I could not have done this without you, and am simp-ly put looking forward to spending the rest of my life with you. Between the laughter and the fun, it’s been tough at times; slogging through daily life in a hallucinatory state induced by sleep-deprivation, but I wouldn’t have wanted to do it with anyone else, and we’ve come out of it with a love stronger than ever. Of that I am both proud and grateful. Besides, du är ju så fin i beige.

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