Evidence for Coupling Transcription and Splicing in Vivo in Saccharomyces Cerevisiae

EVIDENCE FOR COUPLING TRANSCRIPTION AND SPLICING IN VIVO IN SACCHAROMYCES CEREVISIAE

DISSERTATION

Presented in Partial Fulfillment of the Requirement for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Luh Tung, M.S.

* * * * *

The Ohio State University 2007

Dissertation Committee: Dr. Tien-Hsien Chang, Advisor Dr. Venkat Gopalan Dr. Paul K. Herman Dr. Amanda Simcox

ABSTRACT

This dissertation describes the study of two DExD/H-box proteins in the budding yeast Saccharomyces cerevisiae: The first part (Chapter 1 to 4) outlines the discovery that specific alterations can bypass the requirement of Sub2p, an essential DExD/H-box protein that functions in precursor messenger RNA

(pre-mRNA) splicing and in coupling transcription to mRNA export. However, its precise mode of action remains to be defined. Here, I show that the otherwise essential Sub2p can be made dispensable by specific alterations of the intron branch-site-binding protein (BBP) within its conserved branch-site recognition domain. This result suggests that Sub2p acts as a Ribonucleoprotein ATPase

(RNPase) to remodel the splicing complex by removing BBP from the branch site, thereby allowing subsequent binding of the U2 snRNP. Unexpectedly, specific alterations of several transcription factors, as well as perturbing transcription elongation by 6-arauracil (6-AU), can also eliminate the requirement of Sub2p.

Chromatin-immunoprecipitation (ChIP) experiments revealed that these perturbations significantly reduce the co-transcriptional recruitment of BBP, thus offering a satisfactory explanation as to how Sub2p is bypassed. Most

significantly, these results provide compelling evidence that transcription and splicing in yeast are coupled and that this strategy may be conserved.

The second part (Chapter 5) documents the genetic characterization of

Ded1p, an evolutionarily conserved DExH/D-box protein in the budding yeast.

Ded1p is indispensable for translation, but it is also functionally linked to pre-mRNA splicing and virus propagation. In this Chapter, I report a novel aspect of Ded1p’s functions. I first showed that combinations of mutant ded1 alleles with a deletion allele of TIF4631, which encodes one of the two eIF4G translation initiation factors, resulted in a synthetic-lethal growth phenotype.

Unexpectedly, an open-ended search led to the identification of RTG3, which encodes a component involved in the retrograde (RTG) signaling pathway employed in yeast for responding to mitochondrial dysfunction. Further analyses revealed that this synthetic-lethal phenotype is related to RTG1, RTG2, and RTG3, which are required for turning on the glyoxylate cycle. However, deletion of other genes involved in the glyoxylate cycle did not result in the same lethal phenotype.

Consistent with the rapamycin-resistant phenotype exhibited by rtg mutants, ded1 mutants are also rapamycin resistant, thereby suggesting a relationship of

Ded1p to the TOR (target of rapamycin) signaling pathway. Since the TOR pathway was reported to control the protein stability of eIF4G, Ded1p may also be involved in regulating eIF4G level via the TOR signaling pathway.

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Dedicated to my parents

ACKNOWLEDGMENTS

I thank my advisor Dr. Tien-Hsien Chang for providing me with an excellent training environment with intellectual support and guidance during my graduate school career. I also thank Dr. Venkat Gopalan, Dr. Paul K. Herman, Dr. Amanda

Simcox, and Dr. Lee F. Johnson for their helpful discussions, suggestions and encouragements as my committee members. I wish to thank all members of the

Chang laboratory especially Rosemary Hage and Dr. Jean-Leon Chong for their friendship, support and assistance. I am grateful to Hsin-yue, Liz Oakley,

Marianne, and Dr. Michael Chan for their encouragement and support during these years. Thanks also go to Dr. Christine Guthrie, Dr. Michael Hampsey, Dr.

Michael Rosbash, Dr. Bertrend Séraphin, Dr. Ed Hurt and Dr. Grant A. Hartzog for providing reagents. Finally, I am deeply indebted to my dear parents, brother and Li-chi Chang for their support, encouragement and understanding during my graduate school years.

VITA

Oct 1975...... Born – Taipei, Taiwan

July 1997...... B.S. in Department of Zoology, National Taiwan University, Taiwan

July 1999...... M.S. in Institute of Molecular Medicine, National Taiwan University, Taiwan

1999-present...... Graduate Research and Teaching Associate, Graduate Program in Molecular, Cellular and Development Biology (MCDB), The Ohio State University, Columbus, OH, United States

PUBLICATIONS

Pryor, A., Tung, L., Yang, Z., Kapadia, F., Chang, T.-H. and Johnson, L.F. (2004). Growth-regulated expression and G0-specific turnover of the mRNA that encodes URH49, a mammalian DExH/D box protein that is highly related to the mRNA export protein UAP56. Nucleic Acids Research 32, 1857-1865.

Chong, J.-L., Chuang, R.-Y., Tung, L. and Chang, T.-H. (2004). Ded1p, a conserved DExD/H-box translation factor, can promotoe yeast L-A virus negative-strand RNA synthesis in vitro. Nucleic Acids Research 32, 2031-2038.

Huang, C.F., Liu, Y.W., Tung, L., Lin, C.H. and Lee FJ. (2003) Role for Arf3p in development of polarity, but not endocytosis, in Saccharomyces cerevisiae. Mol. Biol. Cell 14, 3834-3847.

Huang, C.F., Chen, C.C., Tung, L., Buu, L.M., and Lee, F.J. (2002) The yeast ADP-ribosylation factor GAP, Gcs1p, is involved in maintenance of mitochondrial morphology. J. Cell Sci. 115(Pt 2), 275-282.

FIELDS OF STUDY

Major Field: Molecular, Cellular, and Developmental Biology

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TABLE OF CONTENTS

Page

Abstract ...... ii Dedication ...... iv Acknowledgments...... v Vita ...... vi

List of Tables ...... xiii List of Figures ...... xv

Chapters:

1. INTRODUCTION ...... 1

1.1 Pre-mRNA Splicing ...... 2 1.2 The Spliceosome ...... 4 1.3 The Dynamic Nature of the Spliceosome ...... 5 1.3.1 Assembly of the Spliceosome ...... 5 1.3.2 Recognition of the Intron-Branch Site by BBP and Mud2p ...... 7 1.3.2.1 Branch Site Binding Protein (BBP) ...... 7 1.3.2.2 U2 Auxiliary Factor (U2AF) and Mud2p ...... 10 1.3.3 DExD/H-box Proteins Function as RNPase in Splicing ...... 13 1.3.4 UAP56 and Sub2p ...... 14 1.3.4.1 Sub2p Functions in pre-mRNA Splicing ...... 15 1.3.4.2 Sub2p Functions in mRNA Export ...... 16 1.4 Gene Expression Pathways Are Tightly Coupled ...... 16 1.4.1 The RNA polymerase II (Pol II) C-terminal domain (CTD) Serves as a Recruitment Platform ...... 18

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1.4.2 Transcription and Splicing Are Coupled...... 19 1.4.3 Chromatin Immunoprecipitation (ChIP) ...... 21 1.5 Goal of This Work...... 22

2. MATERIALS AND METHODS ...... 27

2.1 Yeast Strains ...... 27 2.2 Plasmids...... 27 2.3 Oligos ...... 27 2.4 Analysis of msl5 Alleles Capable of Bypassing sub2 ...... 27 2.5 Genetic Screening of the sub2 Bypass Mutants ...... 29 2.6 Southern blot analysis of SUB2 alleles of the sub2 Bypass Mutants ...... 29 2.7 Genetic characterization of the sub2 Bypass Mutants ...... 30 2.8 Identification of the sub2 Bypass Mutants ...... 31 2.9 Perturbations of the Transcription Machinery ...... 31 2.10 Examination of the capability of prp40-1 to bypass sub2 ...... 33 2.11 Examination of the Requirement of SUB2 in Medium Containing Different Carbon Sources ...... 33 2.12 Examination of the Genetic Interaction between SUA7 and MUD2 ...... 34 2.13 Examination of the Genetic Interaction Between MSL5 and MUD2 ...... 34 2.14 Examination of the Genetic Interaction Between MSL5 and RPB2 ...... 35 2.15 Preparation of Splicing Extract ...... 35 2.16 Preparation of SUA7-depleted Splicing Extract ...... 36 2.17 Preparation of the [32P]-labeled transcript in vitro ...... 37 2.18 In vitro Pre-mRNA Splicing Assay ...... 38 2.19 Commitment Complex and Spliceosome Formation Assay of SUB2 Bypass Extract ...... 39 2.20 Chromatin Immunoprecipitation ...... 40 2.21 Immunoprecipitation of Chromatin-RNA Complexes ...... 41 2.22 Real-Time PCR ...... 42

2.23 Plasmid Constructs in URH49 Project ...... 43 2.24 Yeast Methods in URH49 Project ...... 44

3. RESULTS ...... 68

3.1 Specific alterations of BBP can eliminate the requirement of Sub2p in vivo ...... 68 3.2 Alterations of transcription factor TFIIB can bypass SUB2 ...... 73 3.3 Perturbations of transcription also bypass Sub2p requirement ...... 76 3.4 prp40-1 Failed to Bypass sub2 ...... 78 3.5 Reduced BBP recruitment to intron-containing gene by sub2 bypass suppressor mutations ...... 79 3.6 Impact of BBP recruitment to a pre-mRNA intron by mud2 and msl5-S194P mutations ...... 81 3.7 UAP56 and URH49 both complement a yeast sub2 ...... 82

4. DISCUSSION ...... 125

4.1 The In vivo Targets of DExH/D-box Proteins ...... 125 4.2 BBP is an in vivo Target of Sub2p ...... 127 4.3 Only Specific and Subtle Alteration Can Bypass sub2 ...... 130 4.4 Transcription and Splicing are Coupled in Yeast ...... 133 4.5 Prospectus ...... 136

5. GENETIC ANALYSIS OF Ded1p, AN ESSENTIAL DEXH/D-BOX TRANSLATION FACTOR ...... 139

5.1 INTRODUCTION ...... 139 5.1.1 Ded1p, an Evolutionarily Conserved Essential Translation Initiation Factor ...... 139

5.1.2 The Role of eIF4G in Translation Initiation ...... 141 5.1.3 Retrograde Signal Transduction Pathways ...... 143 5.1.4 TOR Signal Transduction Pathway ...... 144 5.2 MATERIALS AND METHODS ...... 149 5.2.1 Yeast Strains ...... 149 5.2.2 Plasmids ...... 149 5.2.3 Oligos ...... 149 5.2.4 Serial Deletion of the C-terminal Region of Ded1p ...... 149 5.2.5 Examination of C-terminal Deleted Ded1p by Immuno Blot Analysis ...... 150 5.2.6 Genetic Screening and Identification of Mutants Which Are Synthetic Lethal to ded1-120 ...... 151 5.2.7 Examination of Genetic Interaction between DED1 and Other Non-essential Genes...... 152 5.2.8 Rapamycin Sensitivity/Resistance Test ...... 152 5.3 RESULTS ...... 159 5.3.1 C-terminal Truncation of Ded1p is Lethal ...... 159 5.3.2 Truncated Ded1p Is Over-expressed but Unstable in vivo ...... 160 5.3.3 Deletion of TIF4631 but not TIF4632 Caused a Lethal Growth Phenotype with ded1-120 and ded1-199 ...... 160 5.3.4 Deletion of RTG1, RTG2, or RTG3 is Synthetic Lethal with ded1-120 and ded1-199 ...... 161 5.3.5 Deletion of Genes Related to Glyoxylate Cycle Do Not Result in Synthetic Lethality in Combination of ded1-120 or ded1-199 ...... 162 5.3.6 ded1 Mutant Strains Are Rapamycin Resistant ...... 163 5.4 DISCUSSION ...... 168 5.4.1 Ded1p’s C-terminal Region is Important for Its Stability in vivo ..... 168 5.4.2 RTG signaling pathway is genetically linked to Ded1p’s function .. 169 5.5 PROSPECTUS ...... 171

BIBLIOGRAPHY ...... 173

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LIST OF TABLES

Table Page

2.1 Yeast Strains ...... 46

2.2 MSL5 Plasmids ...... 55

2.3 MUD2 Plasmids ...... 58

2.4 PRP40 Plasmids ...... 59

2.5 RPB2 Plasmids ...... 60

2.6 SUA7 Plasmids ...... 61

2.7 SUB2 Plasmids ...... 64

2.8 Oligos ...... 65

3.1 Examination of the Lethal Phenotype Caused by Various Alleles in sub2-CEN.PK2 Strain Carrying SUB2/URA3/CEN ...... 84

3.2 Summary of SUA7 alleles tested of bypassing sub2 ...... 85

3.3 Summary of mutant alleles capable of bypassing sub2 ...... 86

5.1 Yeast Strains ...... 153

5.2 DED1 and RTG3 Plasmids ...... 156

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5.3 Oligos ...... 158

5.4 Growth Phenotype of Strains Containing C-terminal Serially Deleted Ded1p at Different Temperature...... 166

5.5 Alleles Used for the Synthetic Lethality Test in Relation to ded1-120 and ded1-199 ...... 167

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LIST OF FIGURES

Figure Page

1.1 DExD/H-box Proteins Participate in pre-mRNA Splicing Regulation...... 23

1.2 Commitment Complexes and Pre-spliceosome Formation...... 24

1.3 Conserved Sequence Motifs of Several DExD/H-box Proteins...... 25

1.4 A Simplified Model of Co-transcriptional Recruitment of RNA Processing Factors...... 26

3.1 Amino acid sequence comparisons of the conserved RNA-binding domain (RH domain) of Msl5p and position of mutations in msl5-S194P and msl5-V195D...... 87

3.2 The cold-sensitive chromosomal msl5-S194P and msl5-V195D alleles can bypass sub2...... 88

3.3 Schematic overview of the interactions between RNA-binding domain of human BBP (Splicing Factor 1 [SF1]) and a BPS-containing RNA...... 90

3.4 Cell growth of strains carried msl5 mutations...... 92

3.5 Cell growth of strains carried msl5 mutations at 37°C...... 93

3.6 Cell growth of strains carried msl5 mutations at 16°C...... 94

3.7 Synthetic lethality between msl5 alleles and mud2...... 95

3.8 Scheme of sub2-bypass suppressor screen...... 98

3.9 Southern blot analysis of the sub2-bypass suppressors...... 100

3.10 sua7-L214S can bypass sub2...... 102

3.11 Examination of the sub2-bypass capability of sua7 alleles...... 103

3.12 Sua7p depletion from the wild-type splicing extract...... 105

3.13 Depletion of Sua7p From Splicing Extracts Prepared From mud2 and msl5-S194P Strains...... 106

3.14 Examination of the depletion of Sua7p from mud2 and msl5-S194P splicing extract by anti-Ded1p antibodies...... 107

3.15 In vitro Splicing Assay...... 108

3.16 Commitment Complexes Formation Assay...... 110

3.17 Addition of 6- azauracil (6-AU) Can Bypass sub2...... 112

3.18 rpb2-7 But Not rpb2-10 Can Bypass sub2...... 113

3.19 rpb2 Alleles Are Not Synthetic Lethal With msl5-S194P...... 114

3.20 Examination of BBP Recruitment to an Actively Transcribed Intron- containing Reporter Gene By Chromatin-Immunoprecipitation (ChIP). .... 115

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3.21 Comparison of the in vivo ChIP Profile of BBP with the Recruitment of U1, U2 and U5...... 116

3.22 Examination of BBP Recruitment to An Actively Transcribed Intron- containing Reporter Gene at wild-type and mud2 Background By ChIP.117

3.23 Examination of BBP Recruitment to an Actively Transcribed Intron- containing Reporter Gene at Wild-type and sua7-L214S Background By ChIP...... 118

3.24 Examination of BBP Recruitment to an Actively Transcribed Intron- containing Reporter Gene Under 6-AU Treatment By ChIP...... 119

3.25 Examination of BBP Recruitment to an Actively Transcribed Intron- containing Reporter Gene at Wild-type and msl5-S194P Background By ChIP...... 120

3.26 Examination of BBP Recruitment to an Actively Transcribed Intron- containing Reporter Gene at Wild-type and msl5-V195D Background By ChIP...... 121

3.27 The Level of BBP Recruitment to an Actively Transcribed Intron- containing Reporter at Different Genetic Backgrounds and Under 6-AU Treatment...... 122

3.28 Examination of the recruitment of BBP to the YRA1 intron region by RNA-IP Assay...... 123

3.29 URH49 rescues the lethal sub2 deletion in yeast ...... 124

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4.1 Structure of Rpb2p...... 138

5.1 Glyoxylate Cycle ...... 147

5.2 Regulation of the RTG Pathway...... 148

5.3 Protein Expression of C-terminal truncate Ded1p in yeast...... 164

5.4 Cells carrying ded1-120 and ded1-100 display rapamycin resistant phenotype...... 165

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CHAPTER 1

INTRODUCTION

Recent advances in molecular and genome-wide analyses have revealed that cells need to be studied in a holistic manner that integrates the activities of numerous cellular processes. For example, the genetic information transfer pathway, which includes transcription, pre-mRNA processing, mRNA export and translation were recently found to function in a highly integrated manner (Maniatis and Reed, 2002). Organization of these cellular machines, which consist of multiple ribonucleoprotein (RNP) complexes, can be thought to function akin to a tightly regulated factory assembly line. Furthermore, these RNP machines and their interactions are continuously being remodeled by DExD/H-box proteins

(Schwer, 2001), enzymes that harness the energy from ATP hydrolysis to drive conformational changes within, and among, RNPs (Tanner and Linder, 2001). In this thesis, I present a study in which Sub2p, an essential and evolutionarily conserved DExD/H-box protein that is involved in both pre-mRNA splicing and mRNA export, can be made dispensable by several specific genetic alterations of

transcription and splicing apparatus. Subsequent in vivo examinations of the co-transcriptionally recruited splicing apparatus demonstrate that the interaction between the nascent pre-mRNA and an essential splicing factor, BBP (or Msl5p), is reduced when the transcription machinery is altered, which in turn results in bypassing the requirement of Sub2p.

In this chapter, I will start by introducing the pre-mRNA splicing mechanism

(1.1), the splicing machinery (1.2), and the participation of DExD/H-box proteins in pre-mRNA splicing (1.3). In the end, I will summarize recent experimental findings supporting coupling within the multi-stage gene expression pathway

(1.4).

1.1 Pre-mRNA Splicing

The majority of eukaryotic genes are discontinuous, in that the coding sequences are often interrupted by non-coding intervening sequences called introns. During gene expression, the intron sequences on pre-mRNAs have to be removed and the coding sequences, i.e. exons, joined accurately in a process called pre-mRNA splicing. Because introns often contain stop codons in-frame with the upstream protein-coding sequence, the precision and fidelity of intron removal is of paramount importance for cell viability.

During splicing, conserved sequence elements within the pre-mRNA transcript are used to provide the cis-acting signals required for accurate intron removal. These conserved sequences include 5’ and 3’ splice sites, an intron

branch point sequence (BPS) containing a strictly conserved adenosine (A) residue, and a stretch of pyrimidines that is located between the BPS and the 3’ splice site. Although these sequence elements are similar in all eukaryotes, they are much more conserved in the yeast Saccharomyces cerevisiae than in metazoa, especially mammals. The 5’ splice site sequence signal in yeast is almost always GUAUGU, while in mammals only the first two positions (i.e., GU) are highly conserved. Similarly, the intron BPS found in yeast is almost invariable UACUAAC, while the mammalian intron BPS sequences are degenerate, with a consensus sequence of YNCURAY (Y = pyrimidine; R= purine;

N = any nucleotide) (Moore et al., 1993). In both yeast and mammals, the 3’ splice site signal is short, consisting of a pyrimidine (U or C) followed by AG.

These differences in splicing signal sequence conservation probably reflect the relative complexity of the splicing systems in each living organism.

Most metazoan genes contain multiple introns and their transcripts can undergo alternative splicing to produce transcript variants containing or leaving out intron(s). Thus, by virtue of this alternative splicing mechanism, a single gene can produce multiple forms of proteins expressed at different levels in a temporal and tissue-specific manner (Horowitz and Krainer, 1994; Maniatis, 1991).

In contrast, being a simple eukaryote, yeast is known to have far fewer intron-containing genes (~250 out of a total of ~5,700 annotated genes), and fewer than 10 contain more than one intron (Spingola et al., 1999). So far, native

alternative splicing appears not to exist in yeast, although there are examples in which alternative splicing can be induced to occur upon alterations of splicing signals within the introns (Howe and Ares, 1997; Howe et al., 2003).

1.2 The Spliceosome

Splicing is accomplished in a highly dynamic macromolecular machine, termed the spliceosome, which is generally conserved in evolution (Brody and

Abelson, 1985; Brow, 2002). The spliceosome is made up of five functionally distinct small nuclear ribonucleoprotein (snRNPs) particles each of which consists of one of the five conserved small nuclear RNAs (snRNAs; U1, U2, U4, U5 and

U6) and spliceosome-associated proteins. More than 145 different proteins were found in the human spliceosome (Zhou et al., 2002) and among these, over 90 homologues have been identified in the yeast spliceosome (Burge et al., 1995;

Stevens et al., 2002).

Consistent with the fact that splicing regulation is more complex in mammalian systems, many metazoa-specific proteins that are present in human spliceosome are entirely lacking in yeast. One such example is the essential SR protein family which contains at least 10 proteins in human spliceosome (Fu, 1995;

Graveley, 2000; Tacke and Manley, 1999). SR proteins are thought to play a role in splice-site selection and regulation of alternative splicing through interactions of their serine/arginine (SR) or arginine/serine (RS) dipeptide repeats with RNA and other splicing factors (Bourgeois et al., 2004; Fu, 1995; Graveley, 2000; Zahler et

al., 1992). Although no obvious SR family protein homologues were found in yeast, several snRNP-associated yeast splicing factors do contain SR-like domains, implying a similar activity may exist in yeast (Hurt et al., 2004).

1.3 The Dynamic Nature of the Spliceosome

1.3.1 Assembly of the Spliceosome

The spliceosome assembles in an ordered pathway with the five snRNPs sequentially joining the pre-mRNA (Figure 1.1) (Cheng and Abelson, 1987;

Konarska and Sharp, 1986; Konarska and Sharp, 1987). The first step is to commit pre-mRNA to the splicing pathway by binding U1 snRNP to the 5’ splice site (5’ SS) in an ATP-independent manner. This 5’ SS recognition is mediated by a combination of base-pairing of the U1 snRNA to the 5’ SS and the sequence-specific binding activity of some of the U1-snRNP proteins, such as

U1C protein (Du and Rosbash, 2002). Next, U2 snRNP joins the complex and forms the pre-spliceosome in an ATP-dependent manner (Seraphin and Rosbash,

1989). Similar to the 5’-SS recognition, the association of U2 snRNP to the pre-mRNA is via sequence-specific interaction between U2 snRNP/snRNA and the conserved BPS. The recruitment of U2 snRNP is mediated in part by the U1 snRNP as well as additional non-snRNP factors that bridge the two components

(Ares, 1986; Parker et al., 1987). Subsequently, the U4/U6/U5 tri-snRNP and other splicing factors are added to complete the spliceosome assembly. Within the tri-snRNP particle, there is extensive base pairing between the U4 and U6

snRNAs, and the U5 snRNP appears to associate with U4/6 snRNP via protein-protein interactions (Moore et al., 1993; Staley and Guthrie, 1998). Once all five snRNPs are present in the spliceosome, the complex undergoes a series of rearrangements. First, the interaction between the U1 snRNA and the 5’ SS is disrupted, releasing the U1 snRNP from the complex. Simultaneously or at least about the same time, the base pairing between the U4 and U6 snRNAs is disrupted as well, allowing the release of the U4 snRNP and the annealing of the now free U6 snRNA to U2 snRNA. The activated spliceosome, now containing only U2, U6 and U5 snRNPs, then catalyzes the two sequential trans-esterification reactions, which result in the formation of intron lariat. After both 5’ and 3’ splice sites are cleaved and exons ligated, the lariat intron is released from the spliceosome and degraded (Moore et al., 1993; Staley and

Guthrie, 1998).

Despite the prevailing view that the spliceosome assembles in a stepwise fashion, an apparently pre-assembled 45S particle, or called penta-snRNP, containing all of the spliceosomal snRNPs has been described in yeast (Stevens et al., 2002). This 45S complex is capable of catalyzing splicing in vitro (Stevens et al., 2002), suggesting an alternative route of splicing in vivo. In addition, a

200S supraspliceosome complex which contains perhaps multiple copies of penta-snRNPs, most non-snRNP splicing factors, and pre-mRNA was isolated from HeLa nuclear extracts (Azubel et al., 2004; Miriami et al., 1995; Raitskin et al., 2002). Recent ChIP experiments performed both in yeast and in the

mammalian cells, however, argue strongly for the stepwise model of spliceosome assembly co-transcriptionally in vivo (Gornemann et al., 2005). Thus, penta-snRNP may represent a storage complex of the excessive snRNPs to prevent interference of splicing or degradation.

1.3.2 Recognition of the Intron-Branch Site by BBP and Mud2p

During early spliceosome assembly, U1 and other non-snRNP proteins first bind to the pre-mRNA to form a biochemically detectable complex termed commitment complex (CC) in yeast (Seraphin and Rosbash, 1989) and early (E) complex in the mammalian system (Michaud and Reed, 1991). In yeast extracts, two forms of commitment complex termed commitment complex 1 (CC1) and 2

(CC2) can be resolved by native gel electrophoresis (Seraphin and Rosbash,

1989). CC1 is assembled by the binding of U1 snRNP to the 5’ SS, and CC2 is built by the subsequent recruitment of BBP and Mud2p to the intron BPS in CC1 in an ATP-independent manner (Figure 1.2) (Abovich et al., 1994; Rosbash and

Seraphin, 1991; Rutz and Seraphin, 1999).

1.3.2.1 Branch Site Binding Protein (BBP)

The gene encoding the yeast BBP, MSL5 (MUD2 Synthetic Lethal), was first identified as a mutant allele (msl5-G230S) that caused a synthetic-lethal phenotype when placed together with a mud2 allele that produces a truncated

Mud2p (Abovich and Rosbash, 1997). BBP may bridge the 3’ and 5’ splice-site

ends of the intron during pre-mRNA splicing because of its genetic and physical interaction with Mud2p and Prp40p, an U1 snRNP protein (Abovich and Rosbash,

1997; Berglund et al., 1997). The N-terminal region of BBP was found to physically interact with Prp40p by the yeast two-hybrid assay. Because Prp40p also interacts with Prp8p, an essential U5 snRNP component, Prp40p may form a bridge to connect U1 and U5 snRNPs (Abovich and Rosbash, 1997). The interaction between BBP and Prp40p is thought to be mediated by BBP’s proline-rich region and the two tandem WW motifs within Prp40p (Bork and Sudol,

1994). A similar interaction has been proposed for the mammalian SF1 and two proteins that share a conserved WW motif with Prp40p (Abovich and Rosbash,

1997; Bedford et al., 1997; Bedford et al., 1998).

The primary structure of yeast and mammalian BBP is essentially conserved.

Both contain a KH domain, an RNA-binding motif found in many RNA-binding proteins (Musco et al., 1996; Siomi et al., 1993), and two retroviral Zn-knuckle domains (Darlix et al., 1995). UV cross-linking experiments showed that BBP binds to or proximal to the branch-site during splicing commitment complex formation (Berglund et al., 1997). In addition, a recombinant form of BBP binds with a relatively low affinity to a short branch-site containing RNA. Therefore, it was speculated that its interaction with the branch-site region may be facilitated by other splicing factors (Berglund et al., 1997).

The interaction between BBP and the branch-site region has been studied in detail by nuclear magnetic resonance (NMR) spectroscopy. This study

employed a fragment of the mammalian BBP (or SF1) containing the KH domain and the QUA2 motif to bind to a short synthetic RNA containing the UACUAAC branch-site sequence (Kramer and Utans, 1991). The specific binding is mediated by specific contacts between amino acid residues located in both the

KH and QUA2 domains and both the sugar-phosphate backbone and all the nitrogen bases in the RNA. As a result, this interaction is stabilized by hydrogen bonding, hydrophobic, and electrostatic interactions.

BBP is found only in CC2 (Abovich and Rosbash, 1997; Rutz and Seraphin,

1999), but not in CC1 and in the pre-spliceosome, which already contains U2 snRNP (Rutz and Seraphin, 1999). Furthermore, BBP is required for the transition from CC1 to CC2, formation of which was blocked by msl5 mutations

(Rutz and Seraphin, 2000). Thus, it is generally believed that the interaction of

BBP with the conserved UACUAAC sequence during early spliceosome assembly

(Abovich and Rosbash, 1997) must be disrupted to allow a mutually exclusive binding of the U2 snRNP to the branch site (Berglund et al., 1998; Rutz and

Seraphin, 1999). However, Abovich and Rosbash (1997) have shown that depletion of BBP to >99% from the splicing extract does not block pre-spliceosome formation nor affect the kinetics of spliceosome formation in vitro, suggesting that the trace amounts of BBP are catalytically recycled for CC2 formation (Rutz and Seraphin, 1999).

1.3.2.2 U2 Auxiliary Factor (U2AF) and Mud2p

U2AF in metazoa (Zamore and Green, 1989) or Mud2p in yeast (Abovich et al., 1994) is another non-snRNP splicing factor crucial for assembly of the proper metazoan E complex and the yeast CC2 splicing complex. U2AF was first identified and purified from HeLa cell extracts as a heterodimer, composed of

U2AF65 (U2AF 65-kDa subunit) and U2AF35 (U2AF 35-kDa subunit) (Zamore and Green, 1989). The essential U2AF splicing factor is evolutionarily conserved and required for the binding of the U2 snRNP to the intron branch site in metazoa (Kanaar et al., 1993; Potashkin et al., 1993; Zamore et al., 1992; Zorio and Blumenthal, 1999). U2AF65 contains an arginine-serine-rich (RS) domain and three RNA recognition motifs (RRMs), while U2AF35 has a degenerate RRM and a carboxyl-terminal RS domain (Zamore and Green, 1989). The U2AF65 subunit is sufficient for the essential U2AF splicing activity for most of the tested genes in vitro (Zamore and Green, 1989), and the U2AF35 subunit is required for splicing process of only a subset of genes (Merendino et al., 1999). The yeast

U2AF65 homolog, Mud2p (Mutant-U-Die), was later identified from a genetic screen that searched for mutations synthetic lethal to a U1 snRNA partial deletion allele (Abovich et al., 1994). Unlike metazoa, yeast does not have the U2AF35 homolog, reflecting the different nature of splicing processes in two systems

(Abovich et al., 1994).

In the E complex, U2AF65’s C-terminal RNA binding domains directly bind to the polypyrimidine tract (Singh et al., 1995) and U2AF35 recognizes the AG at 3’

SS in an ATP-independent manner (Merendino et al., 1999; Wu et al., 1999a;

Zorio and Blumenthal, 1999). In contrast to its yeast homolog, U2AF65 was also detected in the A complex (the pre-spliceosome) (Bennett et al., 1992;

Champion-Arnaud and Reed, 1994), suggesting a possible additional function.

Similar to U2AF65, the yeast Mud2p has been suggested to bind to the non-conserved polypyrimidine region that immediately follows the conserved branch site sequence (Rain and Legrain, 1997). One of the major distinctions between mammalian and yeast introns is that mammalian introns often possess a degenerate branch site, which, in sharp contrast, is nearly absolutely conserved in yeast. In addition, mammalian introns possess a highly conserved stretch of polypyrimidine, which is often missing in most yeast introns (Burge et al., 1998).

This contrasting nature is consistent with U2AF being an essential splicing factor in mammals, and MUD2 being non-essential in yeast.

Like BBP, Mud2p is found in CC2, but not in CC1 and in pre-spliceosome

(Abovich et al., 1994; Rutz and Seraphin, 1999). The binding of Mud2p to the

CC1 requires the binding of U1 snRNP to the 5’ SS and a proper branch point sequence (Seraphin et al., 1991). BBP and Mud2p are both important for

CC1-to-CC2 transition in yeast (Rutz and Seraphin, 1999). However, BBP is considered to contribute more to the proper CC2 formation, because there is CC2 formation in mud2 extract (Abovich et al., 1994), but no detectable CC2 in BBP

mutant extract (Abovich and Rosbash, 1997; Rutz and Seraphin, 2000). This observation is also consistent with the perfect conservation of the branch site sequence in nearly all the yeast introns.

BBP and Mud2p are thought to join the CC1 as a complex because of the absence of detectable intermediates, containing either BBP or Mud2p, during the transition from CC1 to CC2 (Rutz and Seraphin, 1999). This proposal is also supported by the fact that the yeast two-hybrid assay showed that BBP and

Mud2p interact (Fromont-Racine et al., 1997) and the N-terminal region of BBP co-immunoprecipitates with Mud2p (Abovich and Rosbash, 1997). Unlike

U2AF65, Mud2p does not have the RS domain which has been speculated to facilitate the base pairing of U2 snRNA with the branch site (Valcarcel et al., 1996).

This suggests that Mud2p interacts less strongly with the splicing machinery and the pre-mRNA than its metazoan counterpart.

Although MUD2 is not an essential gene in yeast, mud2 deletion is synthetic lethal with alterations of components of the U1 and U2 snRNPs (Abovich et al.,

1994; Tang et al., 1996), thus suggesting a role in bridging the U1 and U2 snRNPs. The physical and genetic interaction between Mud2p and Sto1p, the nuclear pre-mRNA cap binding protein, provides further support for this idea

(Fortes et al., 1999). Another critical function of Mud2p might be to stabilize the interaction between BBP and intron BPS. Interestingly, mud2 deletion was found to bypass the requirement of an essential DExD/H-box protein Sub2p (Kistler and

Guthrie, 2001), which will be discussed below.

1.3.3 DExD/H-box Proteins Function as RNPase in Splicing

DExD/H-box (D=Asp; E=Glu; H=His; x can be any amino acid) proteins are ubiquitous, present in organisms ranging from viruses to humans (de la Cruz et al.,

1999; Tanner and Linder, 2001). DExD/H-box proteins belong to ATPase superfamily II and possess six conserved motifs (Figure 1.3) that are important for nucleoside triphosphate (NTP) binding (motifs I, II, V and VI), RNA interaction

(motifs Ia, Ib, IV and V), NTP hydrolysis (motifs II, i.e. DExD/H, III and VI), and

RNA unwinding (motif III) (Tanner and Linder, 2001). DExD/H-box proteins are known to be involved in almost all aspects of RNA-related events, including ribosomal biogenesis, mRNA synthesis, pre-mRNA splicing, mRNA export, translation, mRNA turnover and mitochondrial gene expression in eukaryotic cells

(Linder et al., 1989; Tanner and Linder, 2001). In budding yeast, eight out of thirty-four DExD/H-box protein proteins have been reported to participate in pre-mRNA splicing, including Sub2p, Brr2p, Prp2p, Prp5p, Prp16p, Prp22p,

Prp28p, and Prp43p. (Figure 1.1). Notably, most DExD/H-box proteins are essential for cell viability, implying that they are functionally distinct and are likely to act on non-overlapping set of substrate.

Owing to their NTP binding/hydrolysis and RNA unwinding activities,

DExD/H-box proteins are often considered as RNA helicases that employ the energy derived from NTP hydrolysis to drive the unwinding of RNA duplexes.

However, recent discoveries from our laboratory and other research groups have led to a shift in this paradigm. For example, we showed that the function of the

normally essential Prp28p can be bypassed by a series of mutations mapped to genes encoding several U1 snRNP components (Chen et al., 2001). This discovery argues strongly that Prp28p may act as a ribonucleoprotein ATPase

(RNPase) to remodel the early splicing complex, so that U1 snRNP can be removed from the 5’ SS, allowing U6 snRNP binding to the same site.

Biochemical experiments by Jankowsky and co-workers also revealed that

DExD/H-box proteins can enforce protein removal from RNA in vitro (Bowers et al.,

2006; Jankowsky and Bowers, 2006; Yang and Jankowsky, 2006), consistent with the RNPase hypothesis. These advances notwithstanding, the modes of action of these DExD/H-box proteins and their cellular substrates remain poorly understood.

1.3.4 UAP56 and Sub2p

UAP56 (U2AF associated protein of 56 kDa) was initially identified and purified from HeLa cell extracts via its association with U2AF65 and found to be an essential DExD/H-box protein that catalyzes the first ATP-dependent reaction in spliceosome assembly (Fleckner et al., 1997). Because UAP56 is a

DExD/H-box protein, it was speculated that UAP56 facilitates the recruitment of

U2 snRNP to the intron BPS by its putative RNA helicase activity (Fleckner et al.,

1997). Since BBP and U2 snRNP bind to intron branch point in a mutually exclusive manner, UAP56 was proposed to modulate the accessibility of the intron

BPS or the U2 snRNA for base-pairing (Fleckner et al., 1997). The yeast

ortholog of the mammalian UAP56 is Sub2p, which shares 63% identities with

UAP56 (Kistler and Guthrie, 2001). SUB2 (suppressors-screening of brr1-1), an essential gene in yeast, was first identified as a high-copy suppressor (Kistler and

Guthrie, 2001) of a snRNP biogenesis mutant (Noble and Guthrie, 1996a; Noble and Guthrie, 1996b). Independently, Sub2p was also uncovered from a genetic screen (Libri et al., 2001) searching for factors interacting with Prp40p and Nam8p, proteins implicated in stabilization of the U1 snRNP-pre-mRNA interaction (Kao and Siliciano, 1996; Puig et al., 1999), consistent with Sub2p’s role in splicing.

1.3.4.1 Sub2p Functions in pre-mRNA Splicing

Both UAP56 and Sub2p are essential for splicing (Fleckner et al., 1997;

Kistler and Guthrie, 2001; Libri et al., 2001). Consistent with the order of spliceosome assembly, the recruitment of UAP56 to the pre-mRNA requires

U2AF65 (Fleckner et al., 1997). Sub2p’s role at early spliceosome assembly is also supported by the finding that splicing was blocked before the formation of pre-spliceosome in a Sub2p-inactivated or depleted splicing extract (Kistler and

Guthrie, 2001; Zhang and Green, 2001). Addition of purified Sub2p restored the splicing (Zhang and Green, 2001). Upon inactivation of Sub2p, altered forms of

CC1 and CC2 were observed (Kistler and Guthrie, 2001; Libri et al., 2001; Zhang and Green, 2001), indicating that Sub2p impacts the architecture of CCs. Finally,

Kistler and Guthrie (2000) observed that mud2 deletion appears to bypass the requirement of SUB2, suggesting that Mud2p may very well be a target of Sub2p.

1.3.4.2 Sub2p Functions in mRNA Export

In addition to its role in splicing, Sub2p also plays a key role in mRNA export.

First, a temperature-sensitive sub2-85 allele is synthetic lethal with a mutant allele of YRA1, which encodes a protein essential for mRNA export (Strasser and Hurt,

2001). Second, Sub2p depletion results in significant nuclear accumulation of mRNAs derived from both intron-containing and intronless genes (Strasser and

Hurt, 2001). Third, Sub2p overexpression results in blocking mRNA export, possibly by sequestering mRNA export factors (Strasser and Hurt, 2001). Fourth,

Sub2p and Mex67p/Mtr2p, a complex required for mRNA export, interact with

Yra1p in a mutually exclusive and RNA-independent manner (Strasser and Hurt,

2001). Nearly identical data were also reported for UAP56 in metazoa (Luo et al.,

2001). Finally, Sub2p and Yra1p are present in the TREX (transcription and export) complex, which also contains Tho2p, Hpr1p, Mft1p and Thp2p that can form a distinct THO complex involved in transcription elongation. Because TREX complex is co-transcriptionally recruited to the intron-containing and intronless genes by splicing machinery (Abruzzi et al., 2004), it has been proposed that

TREX complex couples transcription with mRNA export by means of recruiting

Yra1p to the mature mRNP.

1.4 Gene Expression Pathways Are Tightly Coupled

The eukaryotic gene expression pathway encompasses transcription, mRNA processing, and export of the mature mRNAs to the cytoplasm, where they are

translated and ultimately degraded. Not knowing a priori that splicing and mRNA export are coupled, studies over the past 20 years, however, have consistently observed that the presence of an intron can greatly enhance gene expression in mammalian cells (Buchman and Berg, 1988; Hamer and Leder, 1979). As a result, many commercial expression vectors were designed to include an engineered intron. In 1985, for the first time, electron-microscopy (EM) visualization of the Drosophila chorion transcripts revealed that RNP particles are found at the splice junction sequences (Osheim et al., 1985). It was interpreted at that time to reflect the natural kinetic order of transcription and splicing. It was not until recent years that different steps of the gene expression pathway were found to be extensively coupled in a temporal, spatial, and functional mannar (Ares and

Proudfoot, 2005; Guthrie and Steitz, 2005; Maniatis and Reed, 2002). One of the key observations is that splicing in vivo is more rapid than that of the reactions performed in vitro. For example, the in vitro splicing reaction takes more than 20 min to complete. Yet, splicing in vivo can be completed within 30 sec

(Wetterberg et al., 2001). To fully understand this difference, one must then take a holistic view to re-consider the interaction among all the involved processes as a single entity, in which individual process may influence other processes in either a synergistic or a negative manner.

1.4.1 The RNA polymerase II (Pol II) C-terminal domain (CTD) Serves as a

Recruitment Platform

The concept of coupling gradually took its root from the finding that the truncated CTD of RNA Pol II inhibits splicing, 3’ site cleavage, and polyadenylation due to the failure of the recruitment of factors required for RNA processing (Cho et al., 1997; McCracken et al., 1997a; McCracken et al., 1997b). The CTD is an essential domain of the largest subunit (Rpb1p) of RNA Pol II, however this domain is not present in RNA Pol I and III (Archambault and Friesen, 1993). This suggests a specific role of the CTD in producing mRNAs, and yet this specific function is either not needed for tRNA and rRNA synthesis. The CTD domain is made up of tandem repeats of a hepta-peptide, Y1S2P3T4S5P6S7 that is evolutionarily conserved. In the repeats, the Ser5 residues are phosphorylated by TFIIH-associated kinase during transcription initiation (Stiller and Cook, 2004).

In the later stage of transcription in yeast, Ser2 residues are phosphorylated by kinases CTK1 and PTEFb (positive transcription elongation factor b) (Phatnani and Greenleaf, 2006), resulting in a higher Ser2 phosphorylation ratio at the 3’ end (Phatnani and Greenleaf, 2006).

Many proteins involved in RNA processing were found to associate with the phosphorylated CTD (Phatnani and Greenleaf, 2004). These include capping enzymes (Cho et al., 1997; McCracken et al., 1997a; Yue et al., 1997), splicing factors (Bird et al., 2004; Chabot et al., 1995; Kim et al., 1997; Mortillaro et al.,

1996; Yuryev et al., 1996), 3’ cleavage and polyadenylation machinery (Hirose et

al., 1999; McCracken et al., 1997a; McCracken et al., 1997b). Furthermore, overexpression of phosphorylated CTD peptides in vivo inhibits splicing in mammalian cells (Du and Warren, 1997). These results were taken to suggest that CTD plays a role in regulating RNA processing in a co-transcriptional manner.

It was hypothesized that CTD serves as a major platform to increase local concentrations of processing factors so as to promote efficient RNA processing, kinetic coupling of RNA processing, and regulating processing machinery possibly via the dynamic phosphorylation state of CTD (Aguilera, 2005; Bentley, 2005).

1.4.2 Transcription and Splicing Are Coupled

Several lines of evidence have revealed a reciprocal relationship between transcription and splicing in metazoa, in that Pol II facilitates the recruitment of splicing factors to nascent introns that in turn promote transcription (Bentley, 2005;

Kornblihtt et al., 2004). First, splicing factors were found to directly bind to the phosphorylated CTD upon transcription activation (Morris and Greenleaf, 2000;

Mortillaro et al., 1996; Phatnani and Greenleaf, 2004). Second, splicing is blocked under the conditions that CTD is truncated, depleted, or titrated (Bird et al., 2004; Du and Warren, 1997; McCracken et al., 1997b; Yuryev et al., 1996), indicating that CTD is essential for splicing. Third, purified phosphorylated Pol II can activate splicing and facilitate spliceosome formation (Hirose et al., 1999), possibly by providing associated splicing factors. Fourth, a promoter proximal intron was shown to stimulate transcription, probably through the interaction

between U snRNPs and the elongation factor TAT-SF1 (Fong and Zhou, 2001).

Finally, switching promoters or altering the transcription elongation rate resulted in altering the pattern of alternative splicing (Blencowe, 2006; de la Mata et al., 2003;

Smith, 2005).

In the budding yeast, though, the supporting data for coupling transcription and splicing are not as robust as in mammalian systems. A crucial difference is that the yeast CTD, despite its ability to improve the 3’ end formation and polyadenylation (Licatalosi et al., 2002), is not required at all for splicing and even the 3’ end formation (Licatalosi et al., 2002). This observation thus paints a scenario that the CTD recruitment of RNA processing factors may not play a major role in yeast, in contrast to the situation found in the metazoa. However, a growing body of evidence continues to support the notion of coupling of transcription to splicing. First, Prp40p, a U1-snRNP protein, was found to bind to yeast CTD (Morris and Greenleaf, 2000; Phatnani and Greenleaf, 2004).

Second, the TREX complex (Abruzzi et al., 2004) and the five snRNPs

(Gornemann et al., 2005; Kotovic et al., 2003; Lacadie and Rosbash, 2005) were found to be co-transcriptionally recruited to nascent transcripts. Importantly, the latter study also demonstrated that the spliceosome assembles in a stepwise manner co-transcriptionally (Gornemann et al., 2005; Lacadie and Rosbash,

2005). Third, deletion of genes encoding transcription elongation factors, such

as TFIIS, resulted in different co-transcriptional splicing phenotypes, suggesting altering the transcription machinery may impact on the efficiency of splicing

(Lacadie and Rosbash, 2005; Lacadie et al., 2006).

1.4.3 Chromatin Immunoprecipitation (ChIP)

ChIP is a standard approach that has been used widely for examining the physical interactions between DNA-binding proteins and their corresponding binding sites on the chromatin in the living cells (Orlando, 2000). This approach employs formaldehyde to produce both protein-nucleic acid and protein-protein crosslinks, provided that these targets are located within a short distance of 2Å.

After shearing chromatin to appropriate sizes, the protein-nucleic-acid complex can be immunoprecipitated by specific antibodies against the protein of interest.

After reversing the crosslinks, the abundance of the DNA (or RNA) targets can then be quantified by real-time PCR. Most recently, the ChIP approach has been successfully exploited to study yeast spliceosome assembly in vivo. The rationale of developing this application rests upon the assumption that splicing in yeast is also a co-transcriptional event. As a result, one would anticipate to crosslink the splicing actors, upon their recruitment to the nascent pre-mRNA, to the nearby relevant chromatin region. In this dissertation, I have employed this technique to study the co-transcriptional recruitment of BBP in details.

1.5 Goal of This Work

A major focus of our laboratory has been to illuminate the functions of

DExD/H-box proteins. In recent years, our research on Prp28p has helped to usher in a major paradigm shift in the field of DExD/H-box proteins. It is now generally believed that DExD/H-box proteins are likely to function as RNPases to remodel RNP complexes, so as to govern their itineraries and fates from their birth to their ultimate demise along the genetic information transfer pathway.

This dissertation work was inspired by our earlier work on Prp28p, in which mutations bypassing the otherwise essential Prp28p eventually led us to a deeper understanding of Prp28p’s role in splicing and its potential in vivo targets.

Applying the same logic, I set out with a goal to look for in vivo targets of Sub2p, another essential DExD/H-box splicing factor. This pursuit, though, took an unexpected, but fruitful, turn, which led me to provide a compelling evidence for the functional coupling of transcription and splicing.

Figure 1.1 DExD/H-box Proteins Participate in pre-mRNA Splicing Regulation. Eight DExD/H-box proteins are labeled in yellow circles; U snRNPs are labeled in green circles. (Adapted from de la Cruz et al., 1999)

Figure 1.2 Commitment Complexes and Pre-spliceosome Formation. U1 snRNP first binds to pre-mRNA at 5’ SS to form commitment complex 1 (CC1). BBP and Mud2p next recognize intron branch point sequence (BPS) and form commitment complex 2 (CC2). After the removal of BBP and Mud2p in an ATP-dependent manner, U2 snRNP binds to the intron BPS to form the pre-spliceosome.

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CHAPTER 2

MATERIALS AND METHODS

2.1 Yeast Strains

All yeast strains used in this work are listed in Table 2.1.

2.2 Plasmids

All plasmids used in this study are listed in Table 2.2-2.7.

2.3 Oligos

All the oligos used in this study are listed in Table 2.8.

2.4 Analysis of msl5 Alleles Capable of Bypassing sub2

All yeast procedures were performed using standard protocols (Guthrie and

Fink, 1991). The mutant msl5 allele (msl5-S194P) in strain CEN.PK2 (or

YTC876; a gift from M. C. López Cuesta, Universidad de Salamanca) (Lopez et al., 1998) was amplified by PCR and cloned into pRS415 and pRS406 (Sikorski

and Hieter, 1989) to yield pMSL5005 and pMSL5006, respectively. The msl5-V195D allele was constructed via a PCR-based site-directed mutagenesis procedure (QuickChangeTM; Stratagene) by using pMSL5002 (MSL5/LEU2/CEN) as the template. Briefly, pMSL5002 was amplified by Pfu Turbo DNA

Polymerase (Stratagene) to extend the two opposing primers containing desired mutation at the center. After amplification, the methylated template strands were digested by DpnI and the resulting unmethylated mutant plasmid was transformed into E. coli XL1-Blue strain for identification of mutant clones, which were validated by DNA sequencing. The msl5-V195D fragment was next sub-cloned into pRS406 to yield pMSL5008 for the following experiment. I employed a standard “pop-in-pop-out” method (Guthrie and Fink, 1991) to replace the wild-type MSL5 gene with either msl5 alleles. BglII-linearized pMSL5006 and pMSL5008 were separately integrated into strain YTC803 [MATa sub2::HIS3 ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 can1-100 pSUB2002

(=SUB2/LEU2/CEN)]. The resulting Ura+ tansformants were cultured non-selectively in YPD liquid medium and plated onto 5-FOA plates (Guthrie and

Fink, 1991). Colonies that appeared on the 5-FOA plates were replica-plated onto YPD plates, which were incubated at 16˚C to identify cold-sensitive colonies.

Previous studies (L. Tung and T.-H. Chang, unpublished) have shown that a cold-sensitive phenotype is linked to msl5 alleles. We validated the correct gene replacements in two ways. First, we crossed the presumed mutant strains with strain YTC811 [MATa sub2::HIS3 ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112

can1-100 pSUB2001 (=SUB2/URA3/CEN)]. Tetrad analysis invariably yielded

2+:2- segregation of the predicted cold-sensitive phenotype. Second, DNA sequencing of the PCR-amplified msl5 alleles from the constructed mutant strains confirmed the designed point mutations. Finally, we swapped the original pSUB2002 in the mutant strains with pSUB2001 (Table 2.2) for testing the bypass activities of the mutant msl5 alleles. Unless stated otherwise, all sub2 strains used in this study were derivatives from strain yCG470 (kindly provided by C.

Guthrie, UCSF) (Kistler and Guthrie, 2001).

2.5 Genetic Screening of the sub2 Bypass Mutants

Approximately ~3x107 cells of the yeast strain YTC802 [MATa sub2::HIS3 ade2-1 his3-11 leu2-3,112 trp1-1 ura3-1 can1-100 pSUB2003

(=SUB2/ADE2/URA3/CEN)] was irradiated by UV for 25 sec on plates to achieve

80% killing. UV-irradiated plates were wrapped in aluminum foil to prevent the activation of the light-inducible DNA repair system in yeasts and incubated at

30ºC for 3 days. The survival colonies were replica-plated onto 5-FOA (1 mg/ml) plates and incubated at 30˚C for 5 days. A total of 137 Ade- (red)/5-FOA+ were identified for further analyses.

2.6 Southern blot analysis of SUB2 alleles of the sub2 Bypass Mutants

To determine if the collected sub2 bypass candidates have truly lost the

SUB2 allele, the genomic DNA of 16 better-growing candidates was extracted by

the standard glass-bead method (Guthrie and Fink, 1991) followed by phenol/chloroform/isoamyl alcohol purification and ethanol precipitation. The isolated genomic DNAs were next digested by EcoRI, separated by agarose gel electrophoresis, transferred to nylon membrane, and then probed with a 2.1-kb

[32P]-labeled SUB2 probe. The SUB2 probe was prepared by PCR amplification with UAP56-11 and UAP56-12 oligos (Table 2.3) from pSUB2003 as the template

(see Figure 3.9). After EcoRI digestion, total 25 ng of the amplified SUB2 fragment was used to label with [a-32P] dATP by using the RediprimeTM II Random

Prime Labelling System (Amersham Biosciences) kit. The standard procedure

(Sambrook et al, 1989) was used for hybridization and wash of the membrane which was next exposed to an X-ray film at -80ºC for 40 hours. None of the 16 strains tested showed any residual SUB2 presence.

2.7 Genetic characterization of the sub2 Bypass Mutants

To determine whether the sub2 bypass mutants carried by the candidates are dominant or recessive, diploids were then made by mating four mutant strains to an isogenic wild-type strain YTC854 (MAT sub2::HIS3 ade2-1 his3-11 leu2-3,112 trp1-1 ura3-1 can1-100 pSUB2003) and tested for their 5-FOA sensitivity. None grew on the 5-FOA plate, indicating that all four bypass mutations are recessive. Subsequent tetrad analysis revealed 2+ : 2- segregation of the 5-FOA phenotype, suggesting that in each case only one mutation is responsible for the bypass activity.

2.8 Identification of the sub2 Bypass Mutants

To clone the wild-type alleles corresponding to the bypass mutations, two sub2 bypass strains containing pSUB2003 were transformed with a YCp50

(LEU2) genomic library to yield ~11,400 transformants in each case. 5-FOA- colonies were identified upon replica-plating transformants onto 5-FOA plates.

Two YCp50 plasmids that reproducibly yielded the 5-FOA- phenotype were selected for DNA sequencing from both ends of their inserts by oligos YCP50-1 and YCP50-2 (Table 2.3), which showed that they are identical. Database search revealed that the insert covered a region of chromosome XVI containing five ORFs, YPR084w, YPR085c, SUA7, VPS69, and SRP54. Re-cloning and re-testing of these five ORFs individually identified SUA7 as the gene responsible for the 5-FOA- phenotype. The two mutant sua7 alleles were cloned by PCR using genomic DNAs prepared from the two sub2 bypass mutants as templates.

2.9 Perturbations of the Transcription Machinery

5-FOA plates containing 150 µg/ml of 6-Azauracil (or 6-AU; Sigma A-1757) were employed to achieve perturbation of transcription elongation for cells grown on the solid medium. Eleven haploid deletion strains (MATa xxx::kanMX4 his31 leu2 met15 ura3; where xxx is a transcription-elongation-factor gene) were obtained from Open Biosystems and crossed to YTC811 (see above).

Sporulation and tetrad dissection of the resulting diploid strains allowed the identification of the desired recombinant (His+/G418R/ Ura+) spores, which were

streaked out on the 5-FOA plates to test for their abilities to grow. To assess the impact by polII mutations on the requirement of SUB2, a tester strain was first constructed by crossing YTC1147 (MAT sub2::kanMX4 his31 leu2 lys20 ura3 pSUB2001) with FY1649 [MATa rpb2297::HIS3 his3200 leu21 lys2-128 ura3-52 pRP212 (=RPB2/URA3/CEN)] (or termed YTC868). The two endogenous URA3 plasmids were then cured by 5-FOA counterselection and another plasmid carrying both SUB2 and RPB2 was introduced into the diploid strain. Tetrad dissection led to the isolation of the tester strain, YTC960 [MATa rpb2297::HIS3 sub2::kanMX4 his3 lys2 leu2 ura3 (RPB2/SUB2/URA3/CEN)].

Plasmids were recovered from strains FY1650 and FY1651 (Hartzog et al., 1998) and the respective rpb2-7 and rpb2-10 alleles were recloned into the pRS315 backbone for introducing into the YTC960. FY strains were gifts from G. Hartzog

(UC Santa Cruz). To assess the impact by sub1 on the requirement of SUB2, a sub1 strain YMH476 (a gift from M. Hampsey) (MATa his3200 ura3-52 leu21 lys2202 trp163 sub1::hisG) was crossed to YTC1147 (see above) and the resulting diploid strain was analyzed after tetrad dissection. None of the 36 sets of tetrads dissected yielded more than two 5-FOA+ spores per tetrad, suggesting that sub1 cannot bypass the requirement of SUB2. For the test of spt4,

GHY368 (YTC892) strain was crossed to YTC1146 to generate a diploid strain.

After tetrad analysis, the original pSUB2001 (=SUB2/LEU2/CEN) plasmid was replaced by pSUB2002 (=SUB2/URA3/CEN) plasmid from G148+His+ spores.

Subsequent examination was done on 5-FOA plates to check if spt4 can bypass

sub2. For the test of spt5-242, GHY92 (YTC891) strain was crossed to

YTC1148 followed by tetrad dissection. G148+Ura+ and cold-sensitive (i.e. spt5-242 allele) spores were collected and test for the lethality on 5-FOA plates.

2.10 Examination of the capability of prp40-1 to bypass sub2

To generate the dominant prp40-1 allele, oligos PRP40-3 and PRP40-4

(Table 2.3) were used to conduct site-directed mutagenesis on pPRP4001 for the

S240F substitution, resulting the pPRP4003 (prp40-1/LEU2/CEN) clone. Since the prp40-1 is a dominant allele, pPRP4003 was first introduced into YTC725

[sub2::HIS3 pSUB2001 (=SUB2/URA3/CEN)] strain and the cells were spotted on 5-FOA plates to test if pSUB2001 plasmid can be lost. Strain YTC1139

[sub2:::kanMX4 prp40::kanMX4 pPRP4004 (=SUB2/PRP40/URA3/CEN)] was also generated and spotted on 5-FOA plates to test if prp40-1 can bypass sub2 and hence lose the pPRP4004 plasmid.

2.11 Examination of the Requirement of SUB2 in Medium Containing

Different Carbon Sources

To examine if carbon sources other than glucose can eliminate the requirement of Sub2p, 2% (w/v) galactose, raffinose and glycerol were used in preparation of 5-FOA plates containing different carbon sources. Yeast strain

YTC802 (see above) and controls (YTC803, YTC1151, and YTC1152, see Table

2.1) grown to mid-log phase were spotted onto each plate and incubated at 30ºC.

2.12 Examination of the Genetic Interaction between SUA7 and MUD2

Strain YTC878 (MATa ura3 leu2 his3 met15 mud2::kanMX4) was crossed to YTC1152 [MATa his3D1 leu2D0 ura3D0 LYS2(?) MET15(?) sua7::kanMX4 SUA7/URA3/CEN (=pSUA7004)] followed by tetrad dissection to obtain spores which carry both mud2::kanMX4 and sua7::kanMX4 as well as the SUA7 allele harbored on an URA3-marked plasmid. Next, various sua7 alleles carried on pRS315 (=LEU2/CEN) were introduced into this strain to test if they can grow on 5-FOA plate.

2.13 Examination of the Genetic Interaction Between MSL5 and MUD2

First, strain YTC878 (see above) was crossed to YTC797 [MATa msl5::kanMX4 ura3 his3 leu2 met15(?*) lys2(?) MSL5/LEU2/CEN

(=pMSL5002)] followed by tetrad dissection to obtain spores (=YTC1187) which carry both mud2::kanMX4 and msl5::kanMX4 as well as the MSL5 plasmid.

Plasmid pMSL5011 (=MSL5/URA3/CEN) was next introduced into YTC1187 to replaced pMSL5002 and the strain was named YTC1189. Finally, plasmids which carry different msl5 alleles were individually transformed into YTC1189 and transformants were streaked on the 5-FOA plate for viability test.

* Genotypes labeled with (?) are not determined.

2.14 Examination of the Genetic Interaction Between MSL5 and RPB2

First, plasmid pRPB2004 (=RPB2/LEU2/CEN) was introduced into strain

YTC868 to replace pRPB2001 (=RPB2/URA3/CEN) followed by transformation of the BglII-digested pMSL4006. After growing in YPD medium for two days to pop-out the integrated pMSL4006, the cold-sensitive colonies were isolated and their MSL5 alleles were sequenced to confirm the presence of msl5-S194P on the chromosome. Next, pRPB2001 was introduced back into this strain and replaced the pRPB2004 plasmid. Finally, the resulting strain was used to introduce pRPB2005 (=rpb2-7/LEU2/CEN) and pRPB2006 (=rpb2-10/LEU2/CEN) to test if they can grow on the 5-FOA plate.

2.15 Preparation of Splicing Extract

Yeast cells were cultured in 1 L of YPD medium overnight, and collected at late log phase (OD600 ~2-3) by centrifugation in a cold GS3 rotor at 3,500 X g. at

4°C for 15 min. Cells were then washed by 200 ml of cold AGK buffer (10mM

HEPES pH7.9, 1.5 mM MgCl2, 200 mM KCl, 10% glycerol and 0.5 mM DTT).

After centrifugation, cell pellets were resuspended in 20 ml of AGK buffer and transferred to 50-ml Falcon tubes followed by centrifugation in a GS3 rotor at

3,500 X g at 4°C for 15 min. The volume of cell pellet was estimated and 0.4-cell volumes of AGK buffer with protease inhibitors (Roche Complete Mini, EDTA-free) was used to resuspend cells. The cell suspension was directly drizzled into a mortar filled half way with liquid nitrogen followed by pounding and grinding to

break up cells to a very fine powder. After grinding, the power was collected and thawed in a 25°C water bath with gentle swirling. Once thawed, cell extract was spun in a SS34 rotor at 35,000 X g at 4°C for 30 min. The clarified cell extract was immediately transferred into a 60 Ti ultracentrifuge tube and spun in a Ti 70.1 rotor (Beckman) at 100,000 x g at 4°C for one hour. After ultracentrifugation, two thirds of the clear pale yellow liquid in the middle of the tube was transferred into a dialysis tubing for dialysis against 2 liters of buffer D (20 mM HEPES pH 7.9, 2 mM EDTA, 50 mM KCl, 20% glycerol and 0.5 mM DTT) for 1.5 hr twice. The dialyzed splicing extract was removed into 1.5 ml tubes, spun to remove sediments, frozen in liquid nitrogen, and stored at -80°C.

2.16 Preparation of SUA7-depleted Splicing Extract

To study whether Sua7p plays a role in splicing, yeast strain YTC920 was used to prepare the splicing extract (see above) for Sua7p depletion. Briefly, IgG

SepharoseTM 6 Fast Flow (17-0969-01 Amersham Biosciences) was activated by

0.5 M CH3COOH (pH 3.4) and then equilibrated by the high salt buffer D (110 mM

KCl). The KCl concentration of the splicing extract was also adjusted to 500 mM by addition of 2.7 M of KCl. To deplete Sua7p from the splicing extract, 70 l of splicing extract was used to incubate with 10 l of IgG Sepharose beads in a 1.6 ml of eppendorf tube on a rotating table at 4°C for 1.5 hr. Next, the splicing extract was dialyzed against 1.5 L of buffer D in a Slide-A-Lyzer¡ Mini Dialysis

Unit (10,000 MWCO dialysis tubing; No. 69572 Pierce) at 4°C for 1.5 hr twice to

adjust the concentration of KCl back to 50 mM for the subsequent in vitro splicing assay. Western blotting examination revealed that > 95% of Sua7p was depleted under this condition.

2.17 Preparation of the [32P]-labeled transcript in vitro

A plasmid with an actin intron-containing actin allele was linerized by EcoRI near its 3’ end. The in vitro transcription reaction was set up with 1.5 g of linerized intron-containing DNA, 1X T7 Transcription Buffer (NEB), 10 mM DTT,

10 mM NTP, 15 units of RNasin (Promega), 3 l of -[32P]-UTP (3000Ci/mmol), and 1.5 l of T7 RNA Polymerase (Gibco). After 2 h incubation at 37°C, 15 l of formamide dye was added to stop the reaction. The reacting tube was next moved to a 65°C water bath for 3 min to denature newly synthesized transcripts and then transferred to ice. The in vitro transcription mixture was loaded into a

5% polyacrylamide gel with 8M urea and electrophoresed in 1X TBE buffer at 500

V until the xylene cyanol dye was 3-4 cm away from the bottom of the gel. The positions of the radiolabeled transcripts were identified by autoradiography. After excising the transcript-containing part from the gel, the gel slices were transferred into two 1.5-ml eppendorf tubes with 500 l elution buffer (500 mM NaOAc pH 5.0 and 1 mM EDTA pH8.0) and stored at 4°C overnight. The next day, the tubes containing the labeled transcripts were inverted into a poly-prep chromatography column (BioRad) in a 50 ml falcon tube and spun to remove the gel debris. The spun liquid was extracted by phenol/chloroform/isoamyl alcohol (pH 5.2) and

precipitated by ethanol at -80°C for 1 hr. The [32P]-labeled transcripts were then resuspended in water and standardized to 60,000 cpm/l for an in vitro splicing assay or to 200,000 cpm/l for commitment complexes formation analysis by native gel electrophoresis.

2.18 In vitro Pre-mRNA Splicing Assay

The in vitro splicing reaction was set up in a 10-l reaction (60 mM K2PO4

32 pH7.0, 3% PEG 8000, 2.5 mM MgCl2, 2 mM ATP, 1 mM spermidine, [ P]-labeled actin pre-mRNA transcript [60,000 cpm]) with 4 l of splicing extract (~15-30 mg/ml) and incubate at 25°C water bath for 20 min or at 16°C water bath for one hour. To terminate the reaction, 200 l splicing extraction buffer (50 mM

Na-Acetate, 1 mM EDTA pH 8.0 and 0.1% SDS) and 200 l of phenol/chloroform/isoamyl alcohol (pH 5.2) were added for RNAs extraction.

RNAs were then precipitated by ethanol and dissolved in 3 l of ddH2O and 7 l of sequencing gel loading dye [98% formamide, 10mM EDTA pH 8.0, 0.025% (w/v) xylene cyanol FF, 0.025% (w/v) bromophenol blue]. After incubation at 65°C for

2 min and then on ice for 2 min, an aliquot of 5-l RNA suspension was loaded onto an 8% polyacrylamide gel with 8 M urea and run in 1X TBE buffer at 500V till the xylene cyanol dye to approximately 1-2 cm from the bottom of the gel. The gel was next transferred onto 3 MM paper for autoradiography.

2.19 Commitment Complex and Spliceosome Formation Assay of SUB2

Bypass Extract

The commitment complex formation reaction was assembled in a 5-l reaction containing 2 l of splicing extract, 2 l of splicing salts [150 mM

K-phosphate pH7.0, 6.25 mM MgCl2 and 7.5% (w/v) PEG 6000], 2 mM ATP,

[32P]-labeled actin pre-mRNA (100,000 cpm) and 125 ng of oligo U2 (RB60).

After incubating at 25°C for 20 min, 2.5 g of yeast transfer-RNA (Sigma) in 5 l of

R buffer (2 mM magnesium acetate, 20 mM EDTA and 50 mM HEPES pH 7.5) was added to dissociate nonspecific binding proteins from spliceosome for 10 min at 4°C. Then 2.5 l of the loading buffer (50% glycerol, 2.5X TBE and 0.1% bromophenol blue) was added and the whole reaction mixture was analyzed in a native gel [3% acrylamide (50:1), 0.5% agarose. 0.5X TBE and 4.15% glycerol] and run in 0.5X TBE at 70V for 20 h at 4°C. Spliceosome formation was analyzed in the same native gel, except the reaction was assembled in a 5-l reaction containing 2 l of splicing extract, 2 l of splicing salts, and [32P]-labeled pre-mRNA (100,000 cpm). After incubating at 25°C for 20 min, 2.5 g of yeast transfer-RNA (Sigma) in 5l of R buffer was added to dissociate nonspecific binding proteins from spliceosome for 10 min at 4°C followed by electrophoresing native gel at 70V for 20 h at 4°C.

2.20 Chromatin Immunoprecipitation

Strains carrying pHZ18 (Lacadie and Rosbash, 2005) were grown in 50-ml synthetic medium containing 2% raffinose but without uracil until reaching 0.8-1.0

OD600. Galactose was added to 2% to induce the reporter gene for 1 h. To crosslink proteins to nucleic acids, formaldehyde (Sigma F-1268) was added to a final concentration of 1% (v/v) for 10 min at room temperature (RT) with occasional swirling of the cultures. Crosslinking was terminated by the addition of 2.5 M glycine to 360 mM followed by incubation at RT for 5 min. Cells were then collected and washed twice with 30-ml cold TBS (20 mM Tris-HCl pH 7.5,

150 mM NaCl), and resuspended in 400-l ChIP buffer [50 mM HEPES pH 7.5,

140 mM NaCl, 1% Triton X-100, 0.1% deoxycholic acid (DOC), 0.1% SDS, protease inhibitor (Roche Complete Mini, EDTA-free)]. Cells were broken by glass beads with 16 cycles of vortexing, each for 30 sec. After separating the cell lysate from glass beads, chromatin pellets was washed twice with ChIP buffer, resuspended in 400-l ChIP buffer, and sonicated 6 times on ice, each for 1 min, at 30% power output (Branson Sonifier 250, 0.5-cm-diameter probe). From each of the 400-l supernatant after sonication, 20 l was saved as “Input”, 5 l for

Western blotting, and the remaining 375 l was subjected to IP. Fifty-l IgG agarose beads (in 100-l slurry) was activated, balanced by ChIP buffer containing 500 g of BSA and 400 g of yeast tRNA followed by an overnight incubation with the IP lysate in a total volume of 1 ml on a nutator at 4˚C. Next,

IgG beads were washed twice with 1-ml ChIP buffer, twice with 1-ml high-salt (500

mM NaCl) ChIP buffer, twice with 1-ml DOC buffer (10 mM Tris-Cl pH 8.0, 250 mM

LiCl, 0.5% NP-40, 0.5% DOC, 1 mM EDTA), and twice with 1-ml TE buffer (50 mM

Tris-Cl pH 8.0, 10 mM EDTA). To elute the protein-DNA complex and reverse the cross-linking, IgG beads were resuspended with 200-l TES (50 mM Tris-Cl pH 8.0, 10 mM EDTA, 1% SDS) and incubated at 65˚C overnight. The 20-l

Input samples were treated identically. The next day, 200-l TE and 25-l proteinase K (10 mg/ml) were added to all samples and incubated at 42˚C for 3 hr.

Finally, DNA fragments were purified through Qiagen PCR columns and eluted in

75-l EB buffer. IP DNA was then concentrated to a volume of 18 l. The amounts of DNA in 1 l of the samples were then determined by real-time PCR.

2.21 Immunoprecipitation of Chromatin-RNA Complexes

For Chromatin-RNA IP, cells were cross-linked and collected as described in

ChIP (see above), except the pellets were resuspended in RNA-IP buffer [50 mM

Tris-HCl pH 7.5, 1% (v/v) NP-40, 0.5% sodium deoxycholate, 0.05% SDS, 1 mM

EDTA, 150 mM NaCl] containing complete protease inhibitors Complete Mini

(Kanaar et al.) and RNasin (Promega). Cells were broken and the cross-linked complexes were sonicated as described in ChIP. To remove DNAs, 12 l of RQ1

DNase (Promega) was added to each of the 400-l supernatant after sonication followed by 1-hr incubation at 37̓C, and the insoluble material was removed by centrifugation. From each of the 400-l supernatant, 20 l was saved as “Input”, and the remaining 380 l was subjected to IP. 50-l IgG agarose beads (in

100-l slurry) was activated, balanced by RNA-IP buffer containing 500 g of BSA and 400 g of yeast tRNA followed by incubation with the IP lysate in a total volume of 1 ml on a nutator at 4˚C overnight. Next, IgG beads were washed twice with 1-ml RNA-IP buffer, twice with 1-ml high-salt (500 mM NaCl) RNA-IP buffer, twice with 1-ml DOC buffer, and twice with 1-ml TE buffer. To elute the protein-RNA complex and reverse the cross-linking, IgG beads were resuspended with 200-l TES (50 mM Tris-Cl [pH 8.0], 10 mM EDTA, 1% SDS) and incubated at 65˚C overnight. The 20-l Input samples were treated identically. The next day, 200-l TE and 25-l proteinase K (10 mg/ml) were added to all samples and incubated at 42˚C for 3 hr. Finally, RNA fragments were purified by standard phenol/chloroform extraction method. The Input and IP RNA were resuspended in 40 l and 12 l of water, respectively. Complementary DNA was prepared from 1 l of each sample in a 20-l reverse transcription reaction, consisting of steps of 95˚C/30 sec, 60˚C/30 sec, and 37˚C/45 min. The cDNAs were analyzed by quantitative PCR with primers spanning YRA1 intron (oligo YRA1-10, table 2.3) and exon 2 (oligo YRA1-12, table 2.3). The relative proportions of RNA fragments were determined on the basis of the threshold cycle (Ct) for each PCR product in the real-time PCR method described below.

2.22 Real-Time PCR

PCR reactions were performed on 96-well plates with each 20-µl reaction consisting of 1-l IP DNA (input 1:80), 10-l iQ SYBR Green Supermix (BioRad),

and a primer pair (0.2 µM each). The reaction was initiated by a hot start (95˚C/3 min), which was followed by 40 cycles of amplification, each consisting of steps of

95˚C/30 sec, 60˚C/30 sec, and 72˚C/30 sec. The temperature cycling was done on an iCycler iQ Real-Time Detection System (BioRad) run by the OSU

Plant-Microbe Genomics Facility (PMGF). Six sets of primer pairs (Lacadie and

Rosbash, 2005) were used for all the experiments. All samples in a single PCR run were assayed in triplicates. All data represent the average of at least three independent experiments, with the error bars displaying the average deviation.

The enrichment of the BBP above background was calculated using the following

(Input Ct – IP Ct) (Input Ct – IP Ct) formula: [2 ]MSL5-TAP strain - [2 ]MSL5-nonTAP strain, where Ct is the threshold cycle reported by the instrument for each PCR reaction. Data were normalized to the signal of the third primer pair (Figure 3.20) of wild-type before plotting.

2.23 Plasmid Constructs in URH49 Project

To examine the ability of UAP56 and URH49 to complement the sub2 in yeast, HA-tagged versions of UAP56 (UAP56-HA) and URH49 (URH49-HA) were

PCR-amplified and placed under the yeast GAL1 promoter control in p415GAL1 vector (Mumberg et al., 1995). The UAP56-HA ORF was amplified using primers UAP56-1 and UAP56-2. Primer UAP56-1 (cccggtctagaaaaaATGgcaga- gaacgat) contains an optimal yeast translation start sequence (bold) upstream of the initiation codon (ATG) of UAP56; primer UAP56-2 (acatgactcgag-[ctaagcgtag-

tctgggacgtcgtatgggta]-ccgtgtctgttcaatgtaggaggagatgtc; HA-epitope, in brackets) is complementary to the last 30 nucleotides of the UAP56 open reading frame.

The ~1.7-kb PCR product was digested with XbaI and XhoI (sites underlined in primer sequences) and cloned into p415GAL1 to yield plasmid

UAP56-HA/p415GAL1. Plasmid URH49-HA/p415GAL1 was constructed in a similar manner, using primer URH49-1 (cccggtctagaaaaaATGgcagaacaggat) and primer URH49-2 (aagccgctcgag-[ctaagcgtagtctgggacgtcgtatgggta]-ccggctctgctcg- atgtatgtggagatgtc; HA-epitope, in brackets) as PCR primers. The XbaI- and

XhoI-digested PCR product was then cloned into p415GAL1 to yield plasmid

URH49-HA/p415GAL1. Plasmid SUB2/p415GAL1, a positive control, was constructed by isolating from pCG788 (=SUB2/CEN/URA3) a 2-kb SalI

(blunted)-XbaI fragment, which was cloned into p415GAL1. Plasmid pCG788 was a gift from Christine Guthrie (Kistler and Guthrie, 2001).

Small-scale plasmid preparations were performed using a QIAprep Spin

Miniprep kit (Qiagen). Large-scale plasmid preparations were performed using a

Plasmid Maxi Kit (BioRad). The sequences of all plasmid inserts were confirmed by DNA sequence analysis.

2.24 Yeast Methods in URH49 Project

Plasmids URH49-HA/p415GAL1, UAP56-HA/p415GAL1, as well as

SUB2/p415GAL1 were transformed into yeast strain yCG470 (Kistler and Guthrie,

2001), which contains the sub2::HIS3 chromosomal deletion and pCG788.

Yeast strain yCG470 was provided by Christine Guthrie. Growth of these strains was examined on synthetic medium lacking leucine and containing galactose and

5-fluoroorotic acid (5-FOA) for counter-selecting pCG788 (Sikorski and Boeke,

1991).

Name Genotype Source YTC725 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 C. Guthrie1 can1-100 sub2::HIS3 SUB2/URA3/CEN (=pSUB2001)

YTC726 MATa/a his3/his3 leu2/leu2 ura3/ura3 Open met15/MET15 lys2/LYS2 msl5::kanMX4/MSL5 Biosystems

YTC795 MATa ura3 his3 leu2 met15(?) lys2(?) This Study msl5::kanMX4 MSL5/LEU2/CEN (=pMSL5002)

YTC797 MATa ura3 his3 leu2 met15(?) lys2(?) This Study msl5::kanMX4 MSL5/LEU2/CEN (pMSL5002)

YTC802 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 SUB2/ADE2/URA3/CEN (=pSUB2003)

YTC803 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 SUB2/LEU2/CEN (=pSUB2002)

YTC810 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 SUB2/LEU2/CEN (=pSUB2002)

YTC811 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 SUB2/URA3/CEN (=pSUB2001)

YTC828 MATa leu2 ura3 his3 trp1 sub2::kanMX4 E. Hurt2 SUB2/TRP1/CEN (=SUB2/pRS314)

YTC829 MATa leu2 ura3 his3 trp1 sub2::kanMX4 E. Hurt sub2-85/TRP1/CEN (=sub2-85/pRS314)

Table 2.1 Yeast Strains

(continued)

Table 2.1 (continued)

Name Genotype Source YTC837 MATa trp1 leu2 ura3 his3 ade2 lys2(?) met15(?) This Study sub2::HIS3 msl5::kanMX4 MSL5/SUB2/URA3/CEN (=pMSL5010)

YTC838 MATa trp1 leu2 ura3 his3 ade2 lys2(?) met15(?) This Study sub2::HIS3 msl5::kanMX4 MSL5/SUB2/URA3/CEN (=pMSL5010)

YTC854 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 SUB2/ADE2/URA3/CEN (=pSUB2003)

YTC865 MATa/a his31/his31 leu2/leu2 met15/MET15 Open ura3/ura3 lys2/LYS2 sub2::kanMX4/SUB2 Biosystems

YTC868 MATa his3200 lys2-128Γ leu21 ura3-52 G. Hartzog3 rpb2297::HIS3 RPB2/URA3/CEN (=pRP212)

YTC869 MATa his3200 lys2-128Γ leu21 ura3-52 G. Hartzog rpb2297::HIS3 rpb2-10/URA3/CEN (=pRP2-10[U])

YTC870 MATa his3200 lys2-128Γ leu21 ura3-52 G. Hartzog rpb2297::HIS3 rpb2-7/URA3/CEN (=pRP2-7[U])

YTC871 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::kanMX4 SUA7/SUB2/ADE2/URA3/CEN (=pSUA7006)

YTC872 MATa his31 leu2 ura3 lys2(?) met15(?) This Study sub2::kanMX4 SUA7/SUB2/ADE2/URA3/CEN (=pSUA7006)

YTC873 MATa his31 leu2 ura3 lys2(?) met15(?) This Study sua7::kanMX4 SUA7/SUB2/ADE2/URA3/CEN (=pSUA7006)

YTC874 MATa his31 leu2 ura3 lys2(?) met15(?) This Study sua7::kanMX4 SUA7/SUB2/ADE2/URA3/CEN (=pSUA7006) (continued)

Table 2.1 (continued)

Name Genotype Source YTC878 MATa ura3 leu2 his3 met15 mud2::kanMX4 Open Biosystems

YTC879 MATa ura3 ade2 his3 leu2 trp1 E. Hurt SUB2-CBP-TEV-ProtA::TRP1-KL

YTC888 MATa his4-912Γ lys2-128Γ trp163 ura3-52 G. Hartzog leu21 spt5202::LEU2 SPT5/URA3/CEN (=pMS4)

YTC889 MATa his4-912Γ lys2-128Γ leu21 ura3-52 G. Hartzog rpb1-244

YTC890 MATa his4-912Γ lys2-128Γ leu21 ura3-52 G. Hartzog rpb1-221

YTC891 MATa his4-912Γ lys2-128Γ leu21 ura3-52 G. Hartzog spt5-242

YTC892 MATa his4-912Γ lys2-128Γ leu21 ura3-52 G. Hartzog spt42::HIS3

YTC900 MATa ura3-52 leu2-3,112 his3-1 trp163 M. Hampsey4 ssu72::LEU2 SSU72/TRP1/CEN

YTC901 MATa ura3-52 leu2-3,112 his3-1 trp163 M. Hampsey ssu72::LEU2 ssu72-3/TRP1/CEN

YTC902 MATa ura3-52 leu2-3,112 his3-1 trp163 M. Hampsey ssu72::LEU2 ssu72-7/TRP1/CEN

YTC903 MATa ura3-52 leu2-3,112 trp1::ura3 sua7-1 M. Hampsey ssu72-1

YTC904 MATa his3200 ura3-52 leu21 lys2202 trp163 M. Hampsey sub1::hisG

YTC920 MATa his31 leu20 met150 ura3 Open SUA7-TAP-HIS3 Biosystems

(continued)

Table 2.1 (continued)

Name Genotype Source YTC947 MATa CYC1 his3-1 leu2-3,112 trp1-289 ura3-52 M. Hampsey ade1-100 (can1-100?) sua7::LEU2 SUA7/URA3/CEN (=pDW11)

YTC948 MATa ade2-1 ade322 his3-11,15 leu2-3,112 trp1-1 This Study ura3-1 can1-100 sua7::kanMX4 SUA7/ADE3/URA3/CEN (=pSUA7034)

YTC949 MATa ade2-1 ade322 his3-11,15 leu2-3,112 trp1-1 This Study ura3-1 can1-100 sua7::kanMX4 SUA7/ADE3/URA3/CEN (=pSUA7034)

YTC959 MATa his3 leu2 ura3-52(or ) lys2(?) met15(?) This Study sub2::kanMX4 rpb2297::HIS3 SUB2/RPB2/URA3/CEN (pRPB2007)

YTC960 MATa his3 leu2 ura3-52(or ) lys2(?) met15(?) This Study sub2::kanMX4 rpb2297::HIS3 SUB2/RPB2/URA3/CEN (pRPB2007)

YTC963 MATa his31 leu20 met150 ura3 msl5-S194P This Study SUA7-TAP

YTC969 MATa/a ura3/ura3 leu2/leu2 MSL5-HA/MSL5 Open Biosystems

YTC970 MATa ura2 his3 leu2 met15(?) lys2(?) This Study msl5::kanMX4 MSL5/pRS316 (=pMSL5011)

YTC971 MATa his3 leu2 met15(?) lys2(?) This Study msl5::kanMX4 msl5-V195D/LEU2/CEN (=pMSL5007)

YTC988 MATa ura3 leu2 his3 met15 rad26::kanMX4 Open Biosystems

YTC989 MATa ura3 leu2 his3 met15 ppr2::kanMX4 Open Biosystems

(continued)

Table 2.1 (continued)

Name Genotype Source YTC990 MATa ura3 leu2 his3 met15 snf11::kanMX4 Open Biosystems

YTC991 MATa ura3 leu2 his3 met15 isw1::kanMX4 Open Biosystems

YTC992 MATa ura3 leu2 his3 met15 ioc4::kanMX4 Open Biosystems

YTC993 MATa ura3 leu2 his3 met15 chd1::kanMX4 Open Biosystems

YTC994 MATa ura3 leu2 his3 met15 rtf1::kanMX4 Open Biosystems

YTC995 MATa ura3 leu2 his3 met15 hpr1::kanMX4 Open Biosystems

YTC996 MATa ura3 leu2 his3 met15 thp2::kanMX4 Open Biosystems

YTC997 MATa ura3 leu2 his3 met15 taf14::kanMX4 Open Biosystems

YTC998 MATa ura3 leu2 his3 met15 ctk1::kanMX4 Open Biosystems

YTC1026 MATa ade2 arg4 leu2-3,112 trp1-289 ura3-2 B. Séraphin5 MSL5-TAP-TRP1-KL

YTC1030 MATa his31 leu20 met150 ura3 Open MSL5-TAP-HIS3 Biosystems

YTC1032 MATa his31 leu20 met150 ura3 This Study MSL5-TAP-HIS3

YTC1033 MATa his3 leu2 ura3 lys2(?) met15(?) This Study MSL5-TAP-HIS3 rpb2297::HIS3 RPB2/URA3/CEN (=pRP212 )

(continued)

Table 2.1 (continued)

Name Genotype Source YTC1034 MATa his3 leu2 ura3 lys2(?) met15(?) This Study MSL5-TAP-HIS3 rpb2297::HIS3 RPB2/URA3/CEN (=pRP212)

YTC1035 MATa his3 leu2 ura3 lys2(?) met15(?) This Study sua7::kanMX4 MSL5-TAP-HIS3 SUA7/SUB2/ADE2/URA3/CEN (=pSUA7006)

YTC1037 MATa his3 leu2 ura3 met15 BBP-TAP-His3 This Study mud2::kanMX4

YTC1038 MATa his3 leu2 ura3 met15 BBP-TAP-His3 This Study ppr2::kanMX4

YTC1039 MATa his3 leu2 ura3 met15 BBP-TAP-His3 This Study taf14::kanMX4

YTC1040 MATa his3 leu2 ura3 met15 BBP-TAP-His3 This Study taf14::kanMX4

YTC1120 MATa ura3 leu2 his3 met15 mud2::kanMX4 This Study msl5-S194P MUD2/URA3/CEN (=pMUD2004)

YTC1122 MATa his31 leu20 met150 ura3 This Study msl5-S194P-TAP-HIS3

YTC1125 MATa/a his3D1/his3D1leu2D0/leu2D0 Open met15D0/MET15 ura3D0/ura3D0 lys2D0/LYS2 Biosystems prp40::kanMX4/PRP40

YTC1131 MATa his3D1 leu2D0 ura3D0 lys2D met15D0(?) This Study prp40::kanMX4 PRP40/URA3/CEN (=pPRP4002)

YTC1132 MATa his3D1 leu2D0 ura3D0 lys2D met15D0(?) This Study prp40::kanMX4 PRP40/URA3/CEN (=pPRP4002)

YTC1139 MATa his3 leu2 ura0 lys2(?) met15D0(?) ade2(?) This Study trp1(?) can1(?) prp40::kanMX4 sub2::HIS3 PRP40/SUB2/ADE2/URA3/CEN (=pPRP4004) (continued)

Table 2.1 (continued)

Name Genotype Source YTC1139 MATa his3 leu2 ura0 lys2(?) met15D0(?) ade2(?) This Study trp1(?) can1(?) prp40::kanMX4 sub2::HIS3 PRP40/SUB2/ADE2/URA3/CEN (=pPRP4004)

YTC1146 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::kanMX4 SUB2/LEU2/CEN (=pSUB2002)

YTC1147 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::kanMX4 SUB2/URA3/CEN (=pSUB2001)

YTC1150 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::kanMX4 SUB2/URA3/CEN (=pSUB2001)

YTC1151 MATa his3D1 leu2D0 ura3D0 lys2(?) met15(?) This Study sua7::kanMX4 SUA7/LEU2/CEN (=pSUA7002)

YTC1152 MATa his3D1 leu2D0 ura3D0 lys2(?) met15(?) This Study sua7::kanMX4 SUA7/URA3/CEN (=pSUA7004)

YTC1153 MATa his3D1 leu2D0 ura3D0 lys2(?) met15(?) This Study sub2::kanMX4 SUB2/LEU2/CEN (=pSUB2002)

YTC1154 MATa his3200 lys2-128∂ leu21 ura3-52 This Study rpb2297::HIS3 rpb2-7/LEU2/CEN (=pRPB2005)

YTC1155 MATa/a msl5::kanMX4/MSL5 This Study mud2::KanMX4/MUD2 ura3/ura3 his3/his3 leu2/leu2 met15(?)/met15 lys2(?)/LYS2

YTC1156 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::kanMX4 msl5-S194P-URA3 (=pMSL5006) SUB2/LEU2/CEN (=pSUB2002)

YTC1157 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::kanMX4 msl5-V195D-URA3 (=pMSL5008) SUB2/LEU2/CEN (=pSUB2002)

YTC1158 MATa MSL5-TAP-HIS3 his31 leu20 met150 This Study ura3 msl5-V195D-URA3 (=pMSL5008) (continued)

Table 2.1 (continued)

Name Genotype Source YTC1159 MATa his31 leu20 ura30 lys2(?) met15(?) This Study sub2::KanMX4 msl5-S194P SUB2/LEU2/CEN (=pSUB2002)

YTC1168 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-S194P-URA3 (=pMSL5006) SUB2/LEU2/CEN (=pSUB2002)

YTC1169 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-V195D-URA3 (=pMSL5008) SUB2/LEU2/CEN (=pSUB2002)

YTC1170 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-S194P SUB2/LEU2/CEN (=pSUB2002)

YTC1171 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-S194P SUB2/LEU2/CEN (=pSUB2002)

YTC1181 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-S194P SUB2/ADE2/URA3/CEN (=pSUB2003)

YTC1182 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-S194P SUB2/ADE2/URA3/CEN (=pSUB2003)

YTC1183 MATa ura3 his3leu2 met15(?) lys2(?) This Study msl5::kanMX4 mud2::kanMX4 MSL5/URA3/2m (=pMSL5016)

YTC1184 MATa ura3 his3 leu2 met15(?) lys2(?) This Study msl5::kanMX4 mud2::kanMX4 MSL5/URA3/2m (=pMSL5016)

(continued)

Table 2.1 (continued)

Name Genotype Source YTC1187 MATa ura3 his3leu2 met15(?) lys2(?) This Study msl5::kanMX4 mud2::kanMX4 MSL5/LEU2/CEN (=pMSL5002)

YTC1188 MATa his31 leu20 ura30 met15(?) lys2(?) This Study msl5-S194P sub2::kanMX4 SUB2/ADE2/URA3/CEN (=pSUB2003)

YTC1189 MATa ura3 his3leu2 met15(?) lys2(?) This Study msl5::kanMX4 mud2::kanMX4 MSL5/URA3/CEN (=pMSL5011)

YTC1190 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-V195D SUB2/LEU2/CEN (=pSUB2002)

YTC1192 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-V195D SUB2/URA3/CEN (=pSUB2001)

YTC1194 MATa ura3 his3 leu2 can1-100 met15(?) lys2(?) This Study ade2(?) trp1-1(?) msl5-V195D

YTC1195 MATa ura3 his3 leu2 can1-100 met15(?) lys2(?) This Study ade2(?) trp1(?) msl5-V195D

YTC1196 MATa ura3 his3 leu2 can1-100 met15(?) lys2(?) This Study ade2(?) trp1-1(?) msl5-V195D-TAP

YTC1197 MATa ade2-1 ura3-1 trp1-1 his3-11 leu2-3,112 This Study can1-100 sub2::HIS3 msl5-V195D SUB2/ADE2/pRS316 (=pSUB2003)

1. C. Guthrie: Christine Guthrie (University of California, San Francisco) 2. E. Hurt: Ed Hurt (Universitat Heidelberg, Heidelberg, Germany) 3. G. Hartzog: Grant A. Hartzog (University of California, Santa Cruz) 4. M. Hampsey: Michael Hampsey (Robert Wood Johnson Medical School, New Jersey) 5. B. Séraphin: Bertrand Séraphin (Centre de Génétique Moléculaire, Avenue de la Terrasse, Gif sur Yvette, France)

Name Description pMSL5001 A ~2.2 kb PCR fragment amplified by oligos MSL5-1 and MSL5-4 (Table 2.8) was cloned into T-Vector. pMSL5002 A ~2.2 kb MSL5 fragment from pMSL5001 was cut by XhoI/BamHI and cloned into pRS315. pMSL5003 A ~2.2 kb MSL5 fragment from pMSL5001 was cut by SacI/SacII and cloned into pSUB2001. pMSL5004 A ~2.2 kb MSL5 fragment from pMSL5002 was cut by XhoI/BamHI and cloned into pRS414. pMSL5005 The msl5-S194P fragment was subcloned from pMSL5009 onto pRS315 after XhoI/BamHI digestion. pMSL5006 The msl5-S194P fragment was cut from pMSL5005 by XhoI/BamHI and subcloned into pRS406. pMSL5007 Oligos MSL5-6 and MSL5-7 (Table 2.8) were used to conduct site-directed mutagenesis of msl5-V195D on pMSL5002. pMSL5008 The msl5-V195D fragment was subcloned into pRS406. pMSL5009 A ~2.2 kb msl5-S194P fragment was amplified by oligos MSL5-1(XhoI) and MSL5-4(BamHI) (Table 2.8) from the genomic DNA isolated from CEN.PK2 strain followed by cloning onto T-V. pMSL5010 A ~2.2 kb MSL5 fragment from pMSL5001 was cut and cloned into pSUB2003 to generate MSL5/SUB2/pRS316. pMSL5011 A ~2.2 kb MSL5 fragment from pMSL5001 was cut by XhoI/BamHI and cloned into pRS316. pMSL5012 Oligos MSL5NAFw and MSL5NARev (Table 2.8) were used to conduct site-directed mutagenesis of msl5-N163A on pMSL5002. pMSL5013 Oligos MSL5REFw and MSL5RERev (Table 2.8) were used to conduct site-directed mutagenesis of msl5-R172E on pMSL5002.

Table 2.2 MSL5 Plasmids (continued)

(Table 2.2 continued)

Name Description pMSL5013 Oligos MSL5REFw and MSL5RERev (Table 2.8) were used to conduct site-directed mutagenesis of msl5-R172E on pMSL5002. pMSL5014 Oligos MSL5KAFw and MSL5KARev (Table 2.8) were used to conduct site-directed mutagenesis of msl5-K196A on pMSL5002. pMSL5015 Oligos MSL5LAFw and MSL5LARev (Table 2.8) were used to conduct site-directed mutagenesis of msl5-L266A on pMSL5002. pMSL5016 A ~2.2 kb MSL5 fragment from pMSL5001 was cut by XhoI/BamHI and cloned into pRS426. pMSL5017 Site-directed mutagenesis was conducted on pMSL5002 to generate S194A mutant. pMSL5018 Site-directed mutagenesis was conducted on pMSL5002 to generate S194N mutant. pMSL5019 Site-directed mutagenesis was conducted on pMSL5002 to generate S194D mutant. pMSL5020 Site-directed mutagenesis was conducted on pMSL5002 to generate S194R mutant. pMSL5021 Site-directed mutagenesis was conducted on pMSL5002 to generate S194C mutant. pMSL5022 Site-directed mutagenesis was conducted on pMSL5002 to generate S194E mutant. pMSL5023 Site-directed mutagenesis was conducted on pMSL5002 to generate S194Q mutant. pMSL5024 Site-directed mutagenesis was conducted on pMSL5002 to generate S194G mutant. pMSL5025 Site-directed mutagenesis was conducted on pMSL5002 to generate S194H mutant.

(continued)

(Table 2.2 continued)

Name Description pMSL5026 Site-directed mutagenesis was conducted on pMSL5002 to generate S194I mutant. pMSL5027 Site-directed mutagenesis was conducted on pMSL5002 to generate S194L mutant. pMSL5028 Site-directed mutagenesis was conducted on pMSL5002 to generate S194K mutant. pMSL5029 Site-directed mutagenesis was conducted on pMSL5002 to generate S194M mutant. pMSL5030 Site-directed mutagenesis was conducted on pMSL5002 to generate S194F mutant. pMSL5031 Site-directed mutagenesis was conducted on pMSL5002 to generate S194Y mutant. pMSL5032 Site-directed mutagenesis was conducted on pMSL5002 to generate S194W mutant. pMSL5033 Site-directed mutagenesis was conducted on pMSL5002 to generate S194T mutant. pMSL5034 Site-directed mutagenesis was conducted on pMSL5002 to generate S194V mutant.

Name Description pMUD2001 A ~2.4 kb PCR fragment amplified by oligos MUD2-1 and MUD2-4 (Table 2.8) was cloned into T-Vector. pMUD2002 A ~2.4 kb MUD2 fragment from pMUD2001 was cut by XhoI/BamHI and subcloned into pRS315. pMUD2003 A ~2.4 kb MUD2 fragment from pMUD2002 was cut by XhoI/BamHI and cloned into pRS314. pMUD2004 A ~2.4 kb MUD2 fragment from pMUD2001 was cut by XhoI/BamHI and cloned into pRS316.

Table 2.3 MUD2 Plasmids

Name Description pPRP4001 A ~2.4 kb PCR fragment amplified by oligos PRP40-1(promoter included) (Table 2.8) and PRP40-2 was digested by XbaI/BamHI and then cloned into pRS415. pPRP4002 A ~2.4 kb PRP40 fragment from pPRP4001 was digested by NotI/BamHI and then subcloned into pRS416. pPRP4003 pPRP4001 was used as the template for site-directed mutagenesis of prp40-1(prp40-S240F). pPRP4004 SmaI/NotI digested PRP40 (from pPRP4001) fragment was ligated to SacII-fillin SUB2/ADE2/pRS316 plasmid to form SUB2/PRP40/ADE2/pRS316.

Table 2.4 PRP40 Plasmids

Name Description pRPB2001 RPB2/URA3/CEN (=pRP212), recovered from strain YTC868 pRPB2002 rpb2-7/URA3/CEN (=pRP2-7[U]), recovered from strain YTC870 pRPB2003 rpb2-10/URA3/CEN (=pRP2-10[U]), recovered from strain YTC869 pRPB2004 The ~5.5 kb RPB2 fragment was digested from pRPB2001 by XbaI/SalI and then cloned into pRS315. pRPB2005 The ~5.5 kb rpb2-7 fragment was digested from pRPB2002 by XbaI/SalI and then cloned into pRS315. pRPB2006 The ~5.5 kb rpb2-10 fragment was digested from pRPB2003 by XbaI/SalI and then cloned into pRS315. pRPB2007 The ~5.5 kb RPB2 fragment was digested from pRPB2001 by SalI-fill-in and XbaI and then cloned into NotI-fill-in/XbaI treated pSUB2001.

Table 2.5 RPB2 Plasmids

Name Description pSUA7001 A ~2.2 kb PCR fragment amplified by oligos SUA7-1(XhoI) and SUA7-2(SalI) (Table 2.8) was ligated to T-vector. Orientation of SUA7 is undetermined. pSUA7002 A ~2.2 kb SUA7 fragment was digested by NotI from pSUA7001 and subcloned into pRS315. Orientation undetermined pSUA7003-1 A ~2.2 kb sua7-L214S fragment was amplified from the sub2 bypass strain (#34) by oligos SUA7-1 and SUA7-2 (Table 2.8) and then cloned into T-vector. Orientation of sua7-L214S is undetermined. pSUA7003-2 A ~2.2 kb sua7-L214S fragment was amplified from the sub2 bypass strain by oligos SUA7-1 and SUA7-2 (Table 2.8) and then cloned into T-vector. Orientation of sua7-L214S is undetermined. pSUA7003-3 A ~2.2 kb sua7-L214S fragment was digested by NotI from pSUA7003-1 and subcloned into pRS315. Orientation of sua7-L214S is undetermined. pSUA7004 A ~2.2 kb SUA7 fragment was digested by NotI from pSUA7002 and subcloned to pRS316. Orientation of SUA7 is undetermined. pSUA7005 A ~2.2 kb SUA7 fragment was digested by NotI from pSUA7002 and subcloned to pSUB2001 (pCG788=SUB2/pRS316). Orientation of SUA7 is undetermined. pSUA7006 A ~2.2 kb SUA7 fragment was digested by NotI from pSUA7002 and subcloned to pSUB2003 (SUB2/ADE2/pRS316). Orientation of SUA7 is undetermined. pSUA7007 Wild-type SUA7/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM299) pSUA7008 sua7-1(E62K)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM326)

Table 2.6 SUA7 Plasmids (continued)

(Table 2.6 continued)

Name Description pSUA7009 sua7-3(R78C)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM325) pSUA7010 sua7-5(E62K-R78C)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM342) pSUA7011 sua7-18(L323P)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM392) pSUA7012 sua7-36(S53P)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM507) pSUA7013 sua7-8(E62R)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM365) pSUA7014 sua7-11(L136P)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM376) pSUA7015 sua7-12(V146M)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM377) pSUA7016 sua7-13(C149Y)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM378) pSUA7017 sua7-17(F289S)/CEN/HIS3 (probably pRS313); a gift from Mike Hampsey (pM391) pSUA7018 sua7-19(E62R-R78E)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM395) pSUA7019 sua7-26(N208S)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM402) pSUA7020 sua7-28(T101P)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM404) pSUA7021 sua7-20(P25L)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM415)

(continued)

(Table 2.6 continued)

Name Description pSUA7022 sua7-21(L56P)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM416) pSUA7023 sua7-29(K201I)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM500) pSUA7024 sua7-31(L284Q)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM502) pSUA7025 sua7-33(K166E)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM504) pSUA7026 sua7-34(L52P)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM505) pSUA7027 sua7-35(K166Q)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM506) pSUA7028 sua7-40(V51A)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM511) pSUA7029 sua7-41(K310I)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM512) pSUA7030 sua7-43(S273A)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM514) pSUA7031 sua7-44(N19D)/CEN/HIS3 (pRS313); a gift from Mike Hampsey (pM515) pSUA7032 sua7-L214S fragment was subcloned to pRS313 from pSUA7003-3 (NotI). Orientation undetermined pSUA7033 A ~1.26 kb SUA7 fragment was amplified by oligos SUA7-6(XhoI) and SUA7-2 (Table 2.8) and then cloned into pRSET-A (XhoI/NcoI) to yield a bacterial expression (His)6-Sua7p. pSUA7034 A ~2.2 kb SUA7 fragment digested by NotI from pSUA7001 was subcloned to pHT4467(ADE3/URA3/CEN). The orientation of SUA7 was not determined.

Name Description pSUB2001 pCG788 (a gift from C. Guthrie lab, UCSF). Both (SalI/XbaI SUB2 fragment) and the (~181 bp KpnI/SalI fragment) from KS vector were cloned into KpnI/XbaI digested pRS316. However, XhoI site which locates in between KpnI and SalI sites was not found on this clone. pSUB2002 pCG817 (a gift from C. Guthrie lab, UCSF). The ApaI/NotI digested SUB2 fragment was cloned into pRS315 with both XhoI and XbaI sites. pSUB2003 The ~2.5 kb BglII digested ADE2 fragment from pASZ11 was first filled-in by Klenow enzyme and then cloned into XbaI/fill-in treated pSUB2001. The internal NdeI site of ADE2 is more close to SUB2 on this plasmid.

Table 2.7 SUB2 Plasmids

Name Sequence MSL5-1 CCCCCTCGAGGGTACGCCTTCCGAACTTTT MSL5-2 CCCCCTCGAGAAGCTTTATAGAAGATATTT MSL5-3 CCAATTAGCAAAAGGATCCAACACCGTGCT MSL5-4 ACGGTACTGAACTCGATCAAGGATCCGTCA MSL5-5 TATCCCGGTGGATCAGTATC MSL5-6 GAGGAAGAGGTTCAGACAAAGAAGGTAAGAA MSL5-7 TTCTTACCTTCTTTGTCTGAACCTCTTCCTC MSL5-8 CCCCCGGATCCATGAGTTTTAGAAGGATTAA MSL5-9 CCCCCCTCGAGTGTATTATAATCCGGGAGGC MSL5AlaFor TTAGAGGAAGAGGTGCAGTCAAAGAAGGTAAG MSL5AlaRev CTTACCTTCTTTGACTGCACCTCTTCCTCTAA MSL5ArgFor TTAGAGGAAGAGGTCGAGTCAAAGAAGGTAAG MSL5ArgRev CTTACCTTCTTTGACTCGACCTCTTCCTCTAA MSL5AsnFor TTAGAGGAAGAGGTAACGTCAAAGAAGGTAAG MSL5AsnRev CTTACCTTCTTTGACGTTACCTCTTCCTCTAA MSL5AspFor TTAGAGGAAGAGGTGACGTCAAAGAAGGTAAG MSL5AspRev CTTACCTTCTTTGACGTCACCTCTTCCTCTAA MSL5CysFor TTAGAGGAAGAGGTTGTGTCAAAGAAGGTAAG MSL5CysRev CTTACCTTCTTTGACACAACCTCTTCCTCTAA MSL5GlnFor TTAGAGGAAGAGGTCAAGTCAAAGAAGGTAAG MSL5GlnRev CTTACCTTCTTTGACTTGACCTCTTCCTCTAA MSL5GluFor TTAGAGGAAGAGGTGAGGTCAAAGAAGGTAAG MSL5GluRev CTTACCTTCTTTGACCTCACCTCTTCCTCTAA MSL5GlyFor TTAGAGGAAGAGGTGGTGTCAAAGAAGGTAAG MSL5GlyRev CTTACCTTCTTTGACACCACCTCTTCCTCTAA MSL5HisFor TTAGAGGAAGAGGTCACGTCAAAGAAGGTAAG MSL5HisRev CTTACCTTCTTTGACGTGACCTCTTCCTCTAA MSL5IleFor TTAGAGGAAGAGGTATAGTCAAAGAAGGTAAG MSL5IleRev CTTACCTTCTTTGACTATACCTCTTCCTCTAA MSL5LeuFor TTAGAGGAAGAGGTTTAGTCAAAGAAGGTAAG MSL5LeuRev CTTACCTTCTTTGACTAAACCTCTTCCTCTAA MSL5LysFor TTAGAGGAAGAGGTAAGGTCAAAGAAGGTAAG MSL5LysRev CTTACCTTCTTTGACCTTACCTCTTCCTCTAA MSL5LysRev CTTACCTTCTTTGACCTTACCTCTTCCTCTAA MSL5MetFor TTAGAGGAAGAGGTATGGTCAAAGAAGGTAAG MSL5MetRev CTTACCTTCTTTGACCATACCTCTTCCTCTAA MSL5PheFor TTAGAGGAAGAGGTTTCGTCAAAGAAGGTAAG

Table 2.8 Oligos (continued)

(Table 2.8 continued)

Name Sequence MSL5PheRev CTTACCTTCTTTGACGAAACCTCTTCCTCTAA MSL5ThrFor TTAGAGGAAGAGGTACCGTCAAAGAAGGTAAG MSL5ThrRev CTTACCTTCTTTGACGGTACCTCTTCCTCTAA MSL5TrpFor TTAGAGGAAGAGGTTGGGTCAAAGAAGGTAAG MSL5TrpRev CTTACCTTCTTTGACCCAACCTCTTCCTCTAA MSL5TyrFor TTAGAGGAAGAGGTTATGTCAAAGAAGGTAAG MSL5TyrRev CTTACCTTCTTTGACATAACCTCTTCCTCTAA MSL5ValFor TTAGAGGAAGAGGTGTTGTCAAAGAAGGTAAG MSL5ValRev CTTACCTTCTTTGACAACACCTCTTCCTCTAA MSL5NAFw GTATCCTGATGTCGCTTTTGTTGGTTTA MSL5NARev TAAACCAACAAAAGCGACATCAGGATAC MSL5REFw ATTATTAGGTCCTGAGGGACGTACGTTAA MSL5RERev TTAACGTACGTCCCTCAGGACCTAATAAT MSL5KAFw GAAGAGGTTCAGTCGCGGAAGGTAAGAATGC MSL5KARev GCATTCTTACCTTCCGCTGAACCTCTTC MSL5LAFw GAACTTAATGGTACTGCAAGAGAAGATAACAG MSL5LARev CTGTTATCTTCTCTTGCAGTACCATTAAGTTC MUD2-1 GATTCTCTCGAGCCCATTCGGATCATCCTT MUD2-2 TCCATCTCGAGTTTCTCTTTTCTCCTCACC MUD2-3 CCCAAGTAGCGGATCCAAATGCAACAATGT MUD2-4 AGTCCTTTAAGGATCCCTCGTTGGCCGCTT MUD2-5 AGTAGTCACAATACTGGGCC PRP40-1 CACGTCGGATCCAGTGTTAGGGACTTCAAC PRP40-2 ACACTGGTCTAGACACAGTTCGGACAGGTG PRP40-3 CAATATACAAACACTTCGTGGTCAATGAAA PRP40-4 TTTCATTGACCACGAAGTGTTTGTATATTG PRP40-5 CTGGGGACCAGAGATCCAAG SUA7-1 GCAAATCTCGAGATCATACCGAATTTGGAG SUA7-2 CGAATAACTTGTCGACCTTTCATCCTTTCG SUA7-3 CTCATGTGTTCTACGCCCAT SUA7-4 GAACTGAATAAGGCACAAGG SUA7-5 TTCGCTGCGGTAATAGGGAT SUA7-6 GTAGATACTCGAGATGATGACTAGGGAGAGCA SUA7-7 GAGGAATGGGCGGAGGAATGCCTGATATGA SUA7-8 ATATACCGCAAAGGCATGTAAAGAGATC SUA7-9 GATCTCTTTACATGCCTTTGCGGTATAT SUA7-10 TGAATATACCGCAGCGAAATGTAAAGAG SUA7-11 CTCTTTACATTTCGCTGCGGTATATTCA pHZ18-1 CGCTTGACGGTCTTGGTTCT

(continued)

(Table 2.8 continued)

Name Sequence pHZ18-2 CAACTTGTTAGTATACTGAT pHZ18-3 GTATGTTAATATGGACTA AA pHZ18-4 ATGAAATTAGGTATTAATCA DT44 GTCAAGCGTGCTTCTAAGGC SALO83 ATGACCGGATCCCAAG SALO84 CGGTATCGCAGTTCCATATCGTCTGAAAATATCGTC SALO96 ATCTTCCTGAGGCCGATACTGTCGTCGTCC SALO97 TAGATGGGCGCATCGTAACCGTGCATCTGC SALO110 GGAGGCTGAAGTTCAGATGTGCGGCGAGTT SALO111 ACCCTGCCATAAAGAAACTGTTACCCGTAG SALO120 ACGAGCATCATCCTCTGCATGGTCAGGTCA SALO121 TTCATCAGCAGGATATCCTGCACCATCGTC SALO254 TGAAAGTTCCAAAGAGAAGGTTTTTTTAGGCTA SALO255 AGTTGCCTGGCCATCCACGCTATATA CACGCCT SALO257 TCTTTTCACCAGTGAGACGGGCAACAGCCA RTG3-1 GCCCAGCTCTAGAACAATGTCTTCGCAGATTC RTG3-2 TACATAGGATCCTGCTTGCCTATCTCTTCCAC RTG3-3 TCCGAATTTCTGTCT RTG3-4 CAGCATGGCACCGAT RTG1-1 AGAGAGGTCGACTCATCCTCATAGCGAGGG RTG1-2 GGTCAGGGATCCAACATGAAAGTATCGCCC RTG2-1 CCCTCCTCAAGCTTGCCAGTATCAAAACCT RTG2-2 TGAAGGTGGATCCGTTGGTATCATCGGTGC YCP50-1 CCATACCCACGCCGAAACAAGCGCTC YCP50-2 ATATGCGTTGATGCAATTTCTATGCGCAC YRA1-9 CATTAAGCAGGATGCTGTAA YRA1-10 GCGTATCATCCAATACTGAA YRA1-11 TTCAGTATTGGATGATACGC YRA1-12 ACACCACCTACTTGAGATGC

CHAPTER 3

RESULTS

3.1 Specific alterations of BBP can eliminate the requirement of Sub2p in vivo

My initial interest in Sub2p stemmed from its role in splicing and mRNA export and was further heightened by new data which suggested that DExD/H-box proteins are enzymes that can remodel RNP complexes (Chen et al., 2001; Kistler and Guthrie, 2001; Schwer, 2001). In this vein, Kistler and Guthrie (2001) have proposed that one role of Sub2p is to remove Mud2p from the branch point sequence (BPS), because the requirement for the otherwise essential SUB2 gene can be eliminated by mud2 deletion (mud2). If such is the case, one would predict that Sub2p should also dissociate BBP from BPS, because U2AF65, the mammalian homologue of Mud2p, interacts with BBP to promote their cooperative binding to the BPS region (Abovich and Rosbash, 1997; Berglund et al., 1997) and both appear to depart from the pre-spliceosome around the same time (Rutz and Seraphin, 1999). To test the hypothesis that BBP is also a target of Sub2p, I

searched for mutant alleles of MSL5 that can render SUB2 dispensable, reasoning that, by altering BBP’s affinity, U2 snRNP may still accomplish binding to BPS without being aided by Sub2p.

To that end, I was at once intrigued by a report that SUB2 is not an essential gene in a yeast strain named CEN.PK2 (Lopez et al., 1998). I surmised that the

CEN.PK2 strain may have already acquired mutations in MSL5 and/or MUD2, thus making SUB2 redundant. As an initial test of this hypothesis, I constructed a tester strain in the CEN.PK2 background, in which the chromosomal SUB2 was deleted and replaced by a copy of SUB2 carried on a URA3-marked plasmid. As expected, this strain grew on plates containing 5-fluoroorotic acid (5-FOA), which can be converted into toxic compounds by Ura3p. This result thus indicated that the URA3/SUB2 plasmid can be freely lost. However, upon acquisition of another plasmid carrying MSL5, but not MUD2, the resulting strain failed to grow on the 5-FOA plate (Table 3.1). These results strongly suggest that a recessive msl5 allele is present in the CEN.PK2 genetic background to render SUB2 dispensable. Indeed, sequence analysis of the PCR-amplified MSL5 region from

CEN.PK2 revealed a transition (TCA-to-CCA) within the MSL5 open reading frame (ORF), predicting a serine-to-proline change at position 194 (S194P)

(Figure 3.1). Interestingly, S194 is located within the KH domain, a noted

RNA-binding motif, conserved among all BBPs (Figure 3.1). To confirm that the newly discovered msl5-S194P allele is truly a bypass suppressor for the sub2 deletion (sub2), I examined its ability to relieve the requirement of SUB2 in a

commonly used laboratory strain, in which SUB2 is known to be essential. Using this lab strain, I first constructed a tester strain in which the chromosomal SUB2 is deleted, so that the cell viability is dependent upon a wild-type SUB2 gene harbored on a URA3-marked plasmid. This tester strain cannot grow on plates containing 5-fluoroorotic acid (5-FOA), because of the toxic compounds converted from 5-FOA by Ura3p (Figure 3.2). I then replaced the chromosomal MSL5 in this tester strain with the msl5-S194P allele using a standard procedure (see

Materials and Methods). Remarkably, the resulting strain, despite growing slower than the tester strain on the YPD plate at 16˚C, grew on the 5-FOA plate, suggesting that the SUB2/URA3 plasmid can now be freely lost (Figure 3.2).

Subsequent transformation of a plasmid-born wild-type MSL5 gene abolished growth on the 5-FOA plate, indicating a recessive nature for the msl5-S194P allele

(data not shown). These results thus suggest that the BBP-S194P variant is likely to assume an altered binding affinity to BPS and, as a result, does not require Sub2p for its removal during splicing.

Consistent with this view, structural analysis of human BBP (or splicing factor 1

[SF1]) revealed that S182, which is equivalent to S194 in the yeast BBP, interacts with the last adenine base of a BPS-containing RNA (5’-U1AUACUAACAA11) via both electrostatic interactions and hydrogen bonding (Figure 3.3) (Liu et al.,

2001a). This observation led me to speculate that other msl5 alleles with similar properties may also bypass SUB2. After surveying a panel of 12 conditional msl5 mutants previously reported by Rutz and Séraphin (2000), I singled out

msl5-5 as one such potential allele. Sequencing analysis of the msl5-5 allele predicted a total of 13 amino-acid alterations in the resulting BBP variant (Rutz and Seraphin, 2000). However, only two, V195D and E258V, were shown to be essential for msl5-5’s cold-sensitive phenotype (Rutz and Seraphin, 2000).

Interestingly, the V195-equivalent in the human SF1, V183, forms hydrophobic interactions with the bases of A8, C9, and A10 in the BPS-containing RNA (Figure

3.3) (Liu et al., 2001a). In contrast, the E258-equivalent in SF1, E246, does not interact with the RNA (Figure 3.3) (Liu et al., 2001a). I therefore engineered a novel msl5-V195D allele and tested for its ability to bypass the sub2. As expected, replacing the wild-type MSL5 gene with this msl5-V195D allele allowed the tester strain to grow on the 5-FOA plate and yielded a slightly slow growth phenotype at 16˚C (Figure 3.2).

Liu and colleagues (Liu et al., 2001a) have shown that several BBP variants containing alterations in their RNA binding domain interact less stably with the

BPS-containing RNA in vitro. These human BBP variants include N151A,

R160E, K184A, and R255A, which are equivalent to yeast msl5-N163A, msl5-R172A, msl5-K196A and msl5-L266A, respectively. Introduction of these msl5 alleles harbored on a LEU2-marked plasmid into a tester strain (YTC970) which contains a chromosomal msl5-deletion and a wild-type MSL5/URA3/CEN plasmid revealed that msl5-R172A is a lethal mutation due to the growth defect on a 5-FOA plate (Figure 3.4). To test if the rest of yeast BBP mutations can bypass sub2, I constructed a tester strain (YTC837) in which the chromosomal SUB2

and MSL5 are both deleted, so that the cell viability is dependent upon a

URA3-marked plasmid carrying both wild-type SUB2 and MSL5 genes.

Introduction of these msl5 alleles harbored on a LEU2-marked plasmid into the tester strain could not relieve its growth defect on 5-FOA plates, indicating that none of these alleles are able to bypass sub2 (data not shown). Furthermore, none of the resulting strains are temperature sensitive at 37˚C or 16˚C (Figure

3.5-3.6).

Because msl5-S194P and msl5-V195D alleles are predicted to alter the

RNA-binding activity of BBP, one would expect that a further compromise of

Mud2p’s activity may altogether prevent the cellular recognition of the BPS region and consequently result in cell death. This indeed is the case, in that cells

(YTC1120) containing a mud2 together with msl5-S194P or msl5-V195D mutation cannot grow without the presence of a plasmid-born wild-type MSL5 gene at 30̓C (Figure 3.7). An examination of other amino-acid substitutions of serine 194 of MSL5 showed that msl5-S194N, msl5-S194R, msl5-S194G, msl5-S194K, msl5-S194F and msl5-S194Y also produced a lethal phenotype in the mud2 background (Figure 3.7). Taken together, the data argue strongly that one key function of Sub2p in the cell is to remove both BBP and Mud2p after their initial binding to the intron BPS region.

3.2 Alterations of transcription factor TFIIB can bypass SUB2

Previous studies by Rosbash and colleagues (Abovich and Rosbash, 1997;

Berglund et al., 1997; Legrain et al., 1988) suggested that factors in addition to

BBP and Mud2p are needed for optimal recognition of the intron BPS. To pursue this idea further, I carried out a genetic screen to search for mutations that can eliminate the requirement of Sub2p. Briefly, UV irradiation was used to mutagenize a starting strain that contains a chromosomal deletion of SUB2

(sub2::HIS3) and a SUB2 allele harbored on a URA3/ADE2-marked plasmid

(Figure 3.8). Candidate mutants acquiring the desired Ade- (red color)/Ura-

(5-FOA+) phenotype were then examined for loss of the SUB2 plasmid by

Southern. DNA isolated from these candidates were digested with EcoRI and probed for the presence of the chromosomal sub2::HIS3 allele and the plasmid-borne SUB2 allele. This probe (Figure 3.9) hybridized to a 2.1-kb DNA band corresponding to the chromosomal SUB2 (Figure 3.9, lane 5 WT). As expected, in the Ade+/Ura+ starting strain, the 2.1-kb band is replaced by an

5.5-kb band corresponding to the sub2::HIS3 allele and by an additional 9.5-kb band (Figure 3.9, lane 4 SS) corresponding to the plasmid-borne SUB2 allele

(Figure 3.9). Strikingly, the 9.5-kb band is undetectable in the Ade-/Ura- suppressor strains (Figure 3.9, lanes 1-3), indicating that the plasmid-born SUB2 allele has been lost. It is unlikely that a portion of the SUB2 gene not covered by the probe is still present and functional in the Ade-/Ura- strains. Such a truncated

Sub2p would have lost the first 409 amino acids (out of a total of 429 amino acids)

encompassing the conserved motifs I—VI (Koonin and Gorbalenya, 1992) essential for ATP-binding, ATP-hydrolysis, and RNA unwinding (Gross and

Shuman, 1995; Pause et al., 1993; Pause and Sonenberg, 1992). I concluded that extragenic suppressor mutations allow bypass of Sub2p function in the

Ade-/Ura- strains. Two independent bypass suppressor mutants were subjected to further genetic analysis, which showed that the bypass phenotype was caused by a single recessive mutation (data not shown).

To identify genes corresponding to the two suppressor mutations, I screened a genomic library for clones that can inhibit the mutant cell growth on the 5-FOA plate (see Materials and Methods). To my surprise, in both cases, this led to the identification of SUA7, which encodes the transcription factor TFIIB (Pinto et al.,

1992). Sequence analysis of the mutant sua7 alleles revealed that both harbor a transition at codon 214 (TTA to TCA), predicting an L214S amino-acid substitution

(Figure 3.10). I introduced the sua7-L214S allele on the plasmid into a tester strain in which the chromosomal SUB2 and SUA7 were simultaneously deleted and complemented by a URA3/ADE2-marked plasmid carrying both SUB2 and

SUA7 genes. As expected, when grown non-selectively, the resulting strain could readily yield Ura-/Ade-/5-FOA+ progenies, indicating that the sua7-L214S mutation permitted the plasmid loss of SUB2 (Figure 3.10).

The structure of TFIIB includes a zinc-binding domain near the N-terminus and two imperfect repeats in the C-terminal two-thirds of the protein (Pinto et al., 1992)

(Figure 3.11). Hampsey and colleagues (Chen and Hampsey, 2004; Wu et al.,

1999b) have previously isolated a library of sua7 conditional mutant alleles and found that the amino-acid changes are distributed throughout the TFIIB protein.

The sua7-L214S allele which I uncovered turned out to be identical to one of the

27 characterized sua7 alleles (Table 3.2) and L214 is located within a segment bridging the two imperfect repeats (Figure 3.11). To examine whether alterations in a specific region of TFIIB can bypass sub2 deletion, I transformed a collection of plasmids carrying various sua7 mutant alleles into the tester strain described above (Figure 3.11). Remarkably, among the 24 alleles tested, 14 were capable of bypassing sub2 (Table 3.2) (Figure 3.11). Among the 14 sub2-bypass alleles, only sua7-C149Y was reported to produce weak alteration in transcription start site selection, while none of the other 13 alleles were reported to produce detectable defect in transcription start site selection (Wu et al., 1999b). In contrast, five of the 10 alleles that failed to bypass sub2 yielded strong defects on transcriptional start site selection (Chen and Hampsey, 2004). These data thus suggest that the bypass phenotype is unlikely due to inactivation of a specific domain in TFIIB. Rather, a subtle compromise of general TFIIB activity results in the bypass phenotype. Consistent with this argument, I found that only the non-bypass sua7 alleles, when placed in conjunction with mud2, resulted in a lethal phenotype (Table 3.2). This most likely reflects a synergistically deleterious effect caused by a robust transcriptional defect and a moderate splicing deficiency (see below).

3.3 Perturbations of transcription also bypass Sub2p requirement

One way to explain the bypassing activity of certain sua7 alleles is to assume that TFIIB has a novel role in splicing similar to that of the BBP. However, despite repeated attempts using splicing extracts prepared from several temperature-sensitive sua7 mutants, no splicing defects were detected. In these experiments, a TAP-tagged version of Sua7p (Sua7p-TAP) was extensively depleted (>95%) by IgG-Sepharose beads in the splicing extract prepared from a

SUA7-TAP strain containing either chromosomal mud2 or msl5-S194P mutation

(Figure 3.12-14). The Sua7p-depleted splicing extracts were used to conduct the in vitro splicing assays for examining the RNA splicing products (Figure 3.15) and the formation of the splicing commitment complexes (Figure 3.16). I was unable to detect any splicing defects on the basis of these two independent assays (Figure 3.15-16). These data thus suggest that TFIIB is less likely to participate in splicing. I therefore turned to an alternative hypothesis, on the basis of functional coupling of transcription and splicing (Bentley, 2005; Kornblihtt,

2006; Kornblihtt et al., 2004; Maniatis and Reed, 2002; Neugebauer, 2002), that subtle alterations of transcription, such as by certain sua7 mutations, may allow

BBP and/or Mud2p to “fall off” their binding sites more frequently. As a result, the need for Sub2p is substantially diminished.

To test this hypothesis, I used both chemical and genetic means to perturb the yeast transcription process and asked whether a plasmid-born SUB2 can be made dispensable in the presence of a chromosomal sub2. Strikingly, when

treated with 6-azauracil (6-AU), a drug known to increase the arrest of elongating

RNA polymerase II (Pol II) by reducing cellular UTP and GTP levels (Powell and

Reines, 1996), the tester strain could grow on plates containing both 6-AU and

5-FOA (Figure 3.17). If the 6-AU-induced bypass phenotype were derived from slowing down transcription elongation, one would anticipate that elimination of genes encoding transcription elongation factors could achieve the same effect. I chose 14 nonessential genes encoding components of transcription elongation factors (Table 3.2) and recombined their deletion alleles into a tester strain.

Deletions of five genes that encode CHD1, IOC4, HPR1, TFIIF, and TFIIS were capable of bypassing sub2 (Table 3.2). In the second genetic test, I introduced into the same tester strain two mutant alleles of RPB2 (see Materials and

Methods), which encodes the second largest subunit of pol II for the same purpose (Powell and Reines, 1996). Although rpb2-7 and rpb2-10 mutants are both sensitive to 6-AU, rpb2-7 is substantially more so than rpb2-10 and displays a measurable sensitivity to mycophenolic acid, a drug that depresses the cellular

GTP pool (Powell and Reines, 1996). I found that rpb2-7, instead of rpb2-10, could bypass SUB2 (see Discussion) (Figure 3.18). Unlike mud2, none of these two rpb2 alleles are synthetic lethal to msl5-S194P (Figure 3.19), suggesting that Rpb2p may not play a role in splicing directly like Msl5p and

Mud2p, despite the capability to bypass sub2. In summary, my data provided strong evidence that SUB2 can be made dispensable by altering the transcription elongation rate and that transcription and splicing are kinetically coupled in vivo.

3.4 prp40-1 Failed to Bypass sub2

Rosbash and colleagues (Berglund et al., 1997) have shown that the RNA binding domain of yeast BBP has a relatively lower binding affinity to the RNA

BPS in comparison to other RNA-binding proteins in vitro. The RNA binding domain of yeast BBP displays a KD of 500 nM whereas other RNA-binding proteins exhibit KD ≤ 1 nM (Daly and Wu, 1989; Hall and Stump, 1992). They speculated that this weak binding reflects the requirement of other splicing factors that may tether BBP to the intron BPS, obviating the need for a single protein with high-affinity binding. In a multi-step pathway, such as spliceosome assembly, a weak binding is perhaps preferrable to avoid a rate-limiting step requiring the subsequent removal of a high-affinity-binding protein (Abovich and Rosbash,

1997; Chiara et al., 1996). In this regard, Prp40p (Kao and Siliciano, 1996), an

U1-snRNP protein, is also considered to play such a role in tethering BBP to the intron BPS. For example, truncation in Prp40p was reported to cause a lethal growth phenotype in combination with mud2 (Abovich and Rosbash, 1997).

Prp40p was reported to interact with BBP in a yeast two-hybrid screen (Abovich and Rosbash, 1997), and Prp40p in the splicing extracts was found to interact with purified GST-BBP in vitro (Abovich and Rosbash, 1997). To test if Prp40p also stabilizes BBP's interaction with the intron BPS similar to Mud2p, I examined if the reported dominant prp40-1 allele (Kao and Siliciano, 1996) can bypass sub2. No sub2-bypass growth was observed in this test (data not shown).

Nonetheless, this negative result cannot eliminate the possibility that Prp40p is also a component that involves in the recruitment of BBP to the intron BPS in splicing.

3.5 Reduced BBP recruitment to intron-containing gene by sub2 bypass suppressor mutations

A reasonable hypothesis to explain the data presented above is to assume that less BBP and/or Mud2p are present in the coupled transcription/splicing complexes under a less favorable transcription elongation condition when comparing with a normal transcription condition. To test this, I adopted a chromatin immunoprecipitation (ChIP) protocol developed by Rosbash and coworkers (Abruzzi et al., 2004; Lacadie and Rosbash, 2005) for examining co-transcriptional spliceosome assembly. In this method, a well-studied 2 µm plasmid-encoded chimeric reporter gene, including the 5’ end of RP51A with the complete intron and the lacZ ORF, driven by GAL1 promoter was characterized for its recruitment of BBP in various genetic backgrounds. Upon galactose-induced gene expression, TAP-tagged BBP was immunoprecipitated with IgG-Sepharose beads from formaldehyde-crosslinked chromatin and the

DNA was analyzed by quantitative real-time PCR using six sets of primers directed to different regions of the reporter gene (see Materials and methods)

(Figure 3.20). I found that BBP was poorly detected in promoter region, but peaked just past the intron and then decreased substantially into exon 2 and

continued to decrease towards the end of exon 2 (Figure 3.20). This pattern is consistent with the fact that splicing trails kinetically behind transcription and is highly similar, if not identical to, the pattern of U1 snRNP previously reported by others using the same system (Kotovic et al., 2003; Lacadie and Rosbash, 2005).

When the BBP recruitment pattern was superimposed (Figure 3.21) onto the data obtained by Lacadie and Rosbash (2005), in which the recruitment of U1, U2 and

U5 snRNPs were studied (Figure 1C in Lacadie and Rosbash, 2005), the BBP signal appears to peak slightly behind that of the U1 snRNP. This pattern is again consistent with the order of the well-studied spliceosomal assembly process

(Abovich and Rosbash, 1997; Berglund et al., 1997; Rutz and Seraphin, 1999;

Rutz and Seraphin, 2000), in that U1 snRNP first recognizes and interacts with the 5’ splice site followed by the interaction of BBP and the intron branch site.

Notably, the BBP peak preceded the reported U2 and U5 snRNP patterns that peak ~500 bp after the intron, which is again consistent with the established order of spliceosome assembly (Lacadie and Rosbash, 2005). The BBP signal was reduced to background levels when the strain was grown in raffinose (Figure 3.20), indicating that the BBP recruitment to the reporter gene is dependent upon transcription. I then examined the impact on BBP recruitment by three mutations, i.e. mud2, sua7-L214S and msl5-V195D, and 6-AU treatment. As expected, these mutants both displayed significantly reduced levels of BBP recruited to chromatin, especially at the peak region (the third oligo pair) but not the other five regions (Figure 3.22-24). This result suggests that the initial co-transcriptional

recruitment of BBP at the two 5’ testing regions was not heavily influenced; however, the further recruitment of BBP to the BPS was not fully supported, probably due to the suboptimal microenvironment or BBP’s lower RNA binding affinity. Interestingly, the levels of BBP recruitment by all these perturbations are reasonably similar, about ~40% of that in the wild-type background (Figure 3.27).

In contrast, the non-bypass sua7-T101P mutation did not significantly alter the

BBP recruitment (~75% of the wild-type level) (Figure 3.27). This perhaps reflects a subtle balance between maintaining adequate amount of BBP for correct recognition of BPS and the upper threshold of BBP presence, above which

Sub2p would be in demand to perform the BBP clearance off the BPS.

Unexpectedly, the recruitment level of BBP-S104P variant detected was 3-fold higher than wild-type BBP (Figure 3.25), which likely results from an unusual conformation caused by the proline substitution.

3.6 Impact of BBP recruitment to a pre-mRNA intron by mud2 and msl5-S194P mutations

The ChIP approach permits assessment of the recruitment of BBP to an actively transcribed reporter gene in vivo. To directly follow BBP’s recruitment to the intron region in the pre-mRNA produced from YRA1, an endogenous intron-containing gene, I used a procedure modified from the established ChIP assay. In this RNA-IP assay, chromatin was first removed by DNase digestion prior to immunoprecipitation, so that the immunoprecipitated nucleic acid

represents the BBP-bound RNA alone. After BBP-bound RNAs were extracted, they were used as template for synthesizing cDNA corresponding to YRA1 pre-mRNA in a reverse transcription reaction. Finally, the BPS-containing region was analyzed by the real-time PCR amplification. As shown in Figure 3.23, mud2 resulted in only 30% of the RNA cross-linked to BBP in comparison to that of the wild-type background. This pattern is consistent with the ChIP result

(Figure 3.21), in which the BBP recruitment is reduced by 70% as well in mud2 background. This result strongly supports the hypothesis that Mud2p is a component to stabilize the interaction of BBP and intron RNA. Unexpectedly, more (25%) intron-containing RNA was recovered from the msl5-S194P mutant background than that in the wild-type background. This seemingly conflicting result may be due to a dramatic alteration of the BBP variant’s protein structure by the mutation (S194P). As a result, this altered BBP binds instead with a higher affinity to the chromatin or other chromatin-binding proteins and hence it has higher chance to be cross linked to the pre-mRNA. This speculation was validated, at least in part, by the ChIP data obtained from msl5-S194P mutant background (Figure 3.25), which again showed that this altered BBP was recovered in far greater abundance than that in the wild-type background.

3.7 UAP56 and URH49 both complement a yeast sub2

In the budding yeast, SUB2 is the only gene encodes the DExD/H-box protein orthologous to the mammalian UAP56. Previous studies showed that the

human UAP56 can rescue the lethal deletion of sub2 in yeast (Zhang and Green,

2001). Recently, Johnson and colleagues (Pryor et al., 2004), however, has identified a novel protein, URH49 (UAP56-related helicase, 49kDa), which is 90% identical to UAP56, in mouse and human cells. To determine if URH49 could also complement the sub2 deletion, I placed the HA-epitope-tagged UAP56

(UAP56-HA) and URH49 (URH49-HA) under the control of the yeast GAL1 promoter and introduced them into a yeast tester strain containing a chromosomal sub2 complemented by SUB2 carried on a URA3-marked plasmid. A

LEU2-marked plasmid that expresses SUB2 from the GAL1 promoter was used as a control. As expected, after 5-FOA counter-selection of the URA3-marked

SUB2 plasmid, galactose-induced SUB2 expression readily rescued the chromosomal sub2 (Figure 3.27). Consistent with the previous report (Zhang and Green, 2001), UAP56-HA complemented the sub2, although to a much lesser extent than SUB2. Likewise, URH49-HA allowed cells to grow in the absence of SUB2 in a manner nearly identical to that of the UAP56-HA. I therefore conclude that both URH49 and UAP56 are capable of replacing Sub2p’s function in yeast.

Plasmid 5-FOA-Leu Viability sub2-bypass Allele LEU2/CEN +++ No MUD2/LEU2/CEN +++ No MSL5/LEU2/CEN - Yes RAD3/LEU2/CEN +++ No SUA7/LEU2/CEN +++ No PRP5/LEU2/CEN +++ No

Table 3.1 Examination of the Lethal Phenotype Caused by Various Alleles in sub2-CEN.PK2 Strain Carrying SUB2/URA3/CEN

SUA7 allele Bypass SUB2 Synthetic Interaction with mud2 wild-type No No N19D No No P25L No No V51A Yes No L52P Yes No S53P Yes No L56P Yes No E62K No Synthetic Lethal E62R No Synthetic Lethal E62K-R78C No Synthetic Lethal E62R-R78E No Synthetic Lethal R78C No Very sick T101P No No L136P Yes No V146M Yes No C149Y Yes No K166E Yes No K166Q Yes No K201I Yes No N208S No No L214S Yes No S273A Yes No L284Q Yes No F289S No No K310I Yes No L323P Yes No

Table 3.2 Summary of SUA7 alleles tested of bypassing sub2

Factor Gene Alleles Tested sub2 Bypassa TFIIB SUA7 See Fig. 2B; Table 3.2 + PolII RPB2 rbp2-7 + rpb2-10 - TFIIF TAF14 taf14::KanMX4 + CSB RAD26 rad26::KanMX4 - TFIIS PPR2/DST1 ppr2::KanMX4 + P-TEFb CTK1 ctk1::KanMX4 - SWI/SNF SNF11 snf11::KanMX4 - Isw1p ISW1/SGN2 isw1::KanMX4 - IOC4 ioc4::KanMX4 + Chd1p CHD1 chd1::KanMX4 + PAF RTF1 rtf1::KanMX4 - THO HPR1 hpr1::KanMX4 + THP2 thp2::KanMX4 - DSIF SPT4 spt4::HIS3 - SPT5 spt5-242 - Sub1p SUB1 sub1::hisG - BBP MSL5 msl5-S194P + msl5-V195D + Prp40p PRP40 prp40-1 - a (+): able to bypass sub2; (-): unable to bypass

Table 3.3 Summary of mutant alleles capable of bypassing sub2

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Figure 3.2 The cold-sensitive chromosomal msl5-S194P and msl5-V195D alleles can bypass sub2. Top: shown are the growth phenotypes of strains which carry a chromosomal MSL5, msl5-S194P or msl5-V195D allele at 30°C and 16°C. Bottom right: shown are the sub2-bypass phenotypes of the sub2-deletion (sub2::HIS3) strains containing a SUB2/ADE2/URA3/CEN plasmid and either a chromosomal MSL5, msl5-S194P or msl5-V195D allele on a 5-FOA plate. Bottom left: shown are the growth phenotypes of the sub2-deletion (sub2::HIS3) strain containing a chromosomal MSL5 allele and a wild-type SUB2 plasmid (SUB2/URA3/CEN) and sub2-deletion strains containing a chromosomal msl5-S194P or msl5-V195D allele on a YPD plate. For all spot tests, cells were grown to mid-log phase, 10-fold serially diluted, and spotted on YPD or 5-FOA plates and incubated at 30°C for 5 days or 16°C for 10 days.

Figure 3.2

Figure 3.3 Schematic overview of the interactions between RNA-binding domain of human BBP (Splicing Factor 1 [SF1]) and a BPS-containing RNA.

The BPS-containing RNA (5’-U1AUACUAACAA11) nucleotides (single-letter code) are labeled in pink: A, adenine; C, cytosine; and U, uracil. Protein side chains (single-letter code) and their positions are indicated in black: A, Ala; C, Cys; D, Asp; E, Glu; F, Phe; G, Gly; H, His; I, Ile; K, Lys; L, Leu; M, Met; N, Asn; P, Pro; Q, Gln; R, Arg; S, Ser; T, Thr; V, Val; W, Trp; and Y, Tyr. Hydrophobic interactions (red dotted lines), hydrogen bonds and electrostatic interactions (dashed blue lines) stabilizing the protein/RNA complex from the results of the nuclear magnetic resonance (NMR) titration experiments (Liu et al., 2001) are shown here. The human S182 and V183 (equivalent to S194 and V195 of yeast BBP respectively) are circled and the equivalent sub2-bypass mutations indicated. Both S182 and V183 interact with the last adenine (A11) base of the BPS-containing RNA via hydrogen bonding, electrostatic interactions and hydrophobic interaction, respectively (adapted from Liu et. al, 2001)

Figure 3.3

Figure 3.4 Cell growth of strains carried msl5 mutations. Yeast strain YTC970 containing a chromosomal msl5 deletion (msl5::KanMX4) and MSL5/URA3/CEN plasmid was introduced with six different msl5 alleles harbored on pRS315 (LEU2/CEN) plasmids. The pRS315 vector and wild-type MSL5/LEU2/CEN plasmid were also introduced into YTC970 as controls. The resulting strains were streaked out on a 5-FOA plate lacking leucine (5-FOA-Leu) subsequently and incubated at 30°C for comparison of their growth. Cells which obtained msl5-R172A failed to grow on the 5-FOA-Leu plate, indicating that msl5-R172A is a null allele and it cannot complement the function of wild-type MSL5 allele. The locations of the different strains on the plate are indicated in a cartoon on the left of the image.

Figure 3.5 Cell growth of strains carried msl5 mutations at 37°C. Yeast strain YTC970 containing a chromosomal msl5 deletion (msl5::kanMX4) and MSL5/URA3/CEN plasmid was introduced with five different msl5 alleles harbored on pRS315 (LEU2/CEN) plasmids. The wild-type MSL5/LEU2/CEN plasmid was also introduced into YTC970 as a control. The resulting strains were streaked out on a 5-FOA plate lacking leucine (5-FOA-Leu) subsequently and incubated at 37°C for comparison of their growth. None of the tested msl5 alleles are temperature-sensitive at 37°C. The locations of the different strains on the plate are indicated in a cartoon on the left of the image.

Figure 3.6 Cell growth of strains carried msl5 mutations at 16°C. Yeast strain YTC970 containing a chromosomal msl5 deletion (msl5::kanMX4) and MSL5/URA3/CEN plasmid was introduced with five different msl5 alleles harbored on pRS315 (LEU2/CEN) plasmids. The wild-type MSL5/LEU2/CEN plasmid was also introduced into YTC970 as a control. The resulting strains were streaked out on a 5-FOA plate lacking leucine (5-FOA-Leu) subsequently and incubated at 16°C for comparison of their growth. Cells carried the sub2-bypass msl5-S194P plasmid are extremely sick at 16°C. Surprisingly, the sub2-bypass msl5-V195D allele did not yield the cold-sensitive growth phenotype when carried on a plasmid. The locations of the different strains on the plate are indicated in a cartoon on the left of the image.

Figure 3.7 Synthetic lethality between msl5 alleles and mud2. Shown are the growth phenotypes of cells containing different msl5 alleles and mud2. Cells which carry chromosomal mud2 and msl5 double deletions (mud2 msl5) and a MSL5/URA3/CEN plasmid and cells which carry chromosomal msl5 deletion (msl5) and a MSL5/URA3/CEN plasmid were introduced with different msl5 alleles carried on LEU2-marked plasmids. The resulting transformants were purified and streaked out on 5-FOA plates. Figure 3.7.1 shows msl5 alleles including S194P, V195D, S194N, S194A, S194N, S194D, S194R, S194C, S194E, S194Q, S194G, S194H and S194I. Figure 3.7.2 shows msl5 alleles including S194L, S194K, S194M, S194F, S194Y, S194W, S194T and S194V.

Figure 3.7

(continued)

(Figure 3.7 continued)

Figure 3.7 (continued)

Figure 3.8 Scheme of sub2-bypass suppressor screen. UV irradiation was used to mutagenize a starting strain which contains a chromosomal deletion of SUB2 (sub2∆::HIS3) and a SUB2 allele harbored on a ADE2/URA3-marked plasmid. Survival colonies were tested on 5-FOA plates, and 5-FOA (+) cells were collected for examination of the existence of SUB2 alleles by Southern blotting analysis. Cells which have truly lost SUB2 allele were next crossed with the starting strain to form diploids. After sporulation, tetrads were examined to determine how many sub2∆-bypass suppressor alleles are there in the isolated candidates. Only those which have only one sub2∆-bypass suppressor allele were collected for verifying whether the sub2∆-bypass suppressor allele is dominant or recessive. In this study, all the isolated candidates carry recessive sub2∆-bypass suppressor alleles. Finally, a wild-type library carried on a LEU2-marked plasmid was used to clone the sub2∆-bypass suppressors from the isolates.

Figure 3.8

Figure 3.9 Southern blot analysis of the sub2-bypass suppressors. Left panel: restriction maps of the chromosomal SUB2, sub2::HIS3 (labeled as HIS3), and the plasmid-borne SUB2 alleles. The starting strain used in this screen has a chromosomal sub2 deletion (sub2::HIS3) and a SUB2 allele carried on an ADE2/URA3-marked plasmid. The probe used in the Southern blot analysis (right) is a ~2 kb EcoRI DNA fragment. The corresponding sizes of EcoRI-digested DNA fragments hybridized by the probe are indicated: the chromosomal SUB2 allele, 2.1 kb; the chromosomal sub2∆::HIS3 allele, 5.5 kb; and the plasmid-borne SUB2 allele, 9.5 kb. Right panel: Southern blot analysis of the sub2-bypass suppressor strains. Genomic DNAs isolated from the wild-type SUB2 strain (WT), starting strain (SS), and three independent isolates which carry sub2-bypass suppressor alleles (Lanes 1-3) were digested by EcoRI and analyzed by Southern blotting using the probe shown in the right panel.

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Figure 3.9

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Figure 3.10 sua7-L214S can bypass sub2. Cells containing a chromosomal sua7 deletion (sua7::KanMX4), and either a sua7-L214S or wild-type SUA7 allele carried on LEU2-marked plasmids were grown to mid-log phase, 10-fold serially diluted, and spotted on a YPD plate for comparison of their growth phenotype. Cells containing sua7-L214S allele [sua7(L214S), row one] grew slightly more slowly than wild-type cells (WT, row three). Cells containing chromosomal deletions of both alleles of sua7 (sua7::KanMX4) and sub2 (sub2::KanMX4) as well as a sua7-L214S allele carried on a LEU2-marked plasmid were also grown to mid-log phase, 10-fold serially diluted, and spotted on the YPD plate for comparison of the growth phenotype. The sub2-bypass cells were not lethal but grew extremely slowly (row two).

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Figure 3.11 Examination of the sub2-bypass capability of sua7 alleles. Top, schematic summary of the sub2-bypass capability of sua7 alleles. (Adapted from Figure 1 in Wu et al., 1999) The single amino acid replacements are indicated by dotted and thick lines at their approximate positions within Sua7p. Thick lines indicate positions of amino acid replacements which can bypass sub2, and dotted lines indicate positions of amino acid replacements which failed to bypass sub2. The position of L214S is labeled by an arrow. Ball-and-stick figures indicate positions of amino acid substitution that affect the accuracy of transcription start site selection and failed to bypass sub2. Zn near the N-terminus denotes the zinc ribbon domain; the two arrows in the C-terminal two-thirds of the protein denote the two imperfect repeats of the core domain (Pinto et al., 1994). Bottom, allele specificity of sub2-bypass capability of sua7 alleles. Cells which contain chromosomal deletions of both sua7 (sua7::KanMX4) and sub2 (sub2::KanMX4) alleles and a SUA7/URA3/CEN plasmid were transformed with different sua7 mutant alleles carried on a HIS3-marked plasmid and spotted on a 5-FOA plate (bottom left). Bottom right, sua7 alleles which can bypass sub2 were shaded with gray background. Mutations that result in altered transcription start sites were labeled in black background.

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Figure 3.11

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Figure 3.12 Sua7p depletion from the wild-type splicing extract. Sua7p was depleted from the splicing extract prepared from a wild-type strain which has a chromosomal TAP-tagged SUA7 allele. TAP moiety contributes ~22kDa to the size of Sua7p (~38kDa) and hence resulted in a ~60kDa fusion protein (Sua7p-TAP, indicated by an arrow on the right). Lanes 1-8, different amount (40 mg, 30 mg, 20 mg, 10 mg, 8 mg, 6 mg, 4 mg, and 2 mg) of the Sua7p-TAP extract before depletion. Lane 9, 80 mg of Sua7p-depleted extract. Western blot was conducted by using the 1:5000 diluted pre-immune serum as the primary antibody, which can bind to the protein A moiety on the TAP tag.

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Figure 3.13 Depletion of Sua7p From Splicing Extracts Prepared From mud2 and msl5-S194P Strains. 80 mg of the splicing extract prepared from mud2 (lane 1) and msl5-S194P (lane 2) strains which have a chromosomal TAP-tagged SUA7 allele after Sua7p depletion. Lanes 3-4, 80 mg of the splicing extract prepared from mud2 SUA7-TAP (lane 3) and msl5-S194P SUA7-TAP (lane 4) strains after mock depletion by sepharose beads. Lanes 5-10, different amount (50 mg, 25 mg, 12 mg, 6 mg, 3 mg, and 1.5 mg) of the Sua7p-TAP extract before depletion. Western blot was conducted by using the 1:5000 diluted pre-immune serum as the primary antibody, which can recognize the protein A moiety on the TAP tag.

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Figure 3.14 Examination of the depletion of Sua7p from mud2 and msl5-S194P splicing extract by anti-Ded1p antibodies. Sua7p was depleted from the splicing extract prepared from mud2 SUA7-TAP and msl5-S194P SUA7-TAP strains. IgG, splicing extracts were incubated with IgG-Sepharose beads for Sua7p depletion. Mock, splicing extracts were incubated with Sepharose beads for mock depletion. Western blot was conducted by using 1:5000 diluted poly-clonal anti-Ded1p antibodies so the amount of Ded1p can serve as an internal control of each sample.

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Figure 3.15 In vitro Splicing Assay. Sua7p was depleted from splicing extracts prepared from wild-type (SUA7-TAP), mud2 and msl5-S194P strains. U, undepleted; M, Mock (Sepharose beads) depletion, G, IgG-Sepharose beads depletion. Equal amounts of splicing extracts were used to perform the in vitro splicing assay at 25̓C.

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Figure 3.15

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Figure 3.16 Commitment Complexes Formation Assay. In vitro commitment complexes formation was conducted by using the splicing extracts prepared from the following strains: mud2, msl5-S194P, wild-type (SUA7), sua7-L136P (a sub2 bypass suppressor), sua7-E62K-R78C (a non-sub2 bypass suppressor), and wild-type (DED1) strains.

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Figure 3.16

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Figure 3.17 Addition of 6-azauracil (6-AU) Can Bypass sub2. Wild-type cells (SUB2) and cells which carry chromosomal sub2 deletion (sub2::HIS3 or sub2::kanMX4) and a SUB2/ADE2/URA3/CEN plasmid were streaked out on a 5-FOA plate and a 6-AU 5-FOA plate (which contains 50 mg/ml of 6-AU and 20 mg/ml of 5-FOA).

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Figure 3.18 rpb2-7 But Not rpb2-10 Can Bypass sub2. Cells carry chromosomal deletions of both rpb2 (rpb2∆297::HIS3) and sub2 (sub2::KanMX4) alleles together with a SUB2/RPB2/pRS316 plasmid were introduced with either wild-type RPB2, rpb2-7 or rpb2-10 allele harbored on pRS315 and spotted on a 5-FOA plate.

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Figure 3.19 rpb2 Alleles Are Not Synthetic Lethal With msl5-S194P. Cells carry chromosomal rpb2 deletion, msl5-S194P and a wild-type RPB2 carried on an URA3-marked plasmid were transformed with rpb2 alleles harbored on pRS315. The purified transformants were streaked on a 5-FOA-Leu plate.

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Figure 3.20 Examination of BBP Recruitment to an Actively Transcribed Intron-containing Reporter Gene By Chromatin-Immunoprecipitation (ChIP). Six pairs of oligos were used to amplify different regions (marked by short lines on top) of the galactose-inducible reporter. The reporter plasmid (pHZ18) contains an artificial construct which has two RP51B exons (shown in gray boxes), intron (the line between two gray boxes), and a fused LacZ (shown in white box). Cells cultured in galactose and raffinose were collected and used for the ChIP assay. All the data from real-time PCR were first normalized to the INPUT, subtracted the background, and finally normalized to the signal of the third oligo pair from wild-type. The mean and standard deviation from these independent experiments are indicated.

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Figure 3.21 Comparison of the in vivo ChIP Profile of BBP with the Recruitment of U1, U2 and U5. The ChIP profile of BBP to the pHZ18 reporter from Figure 3.20 was superimposed with U1, U2 and U5’s profile done in the same system by Lacadie and Rosbash (Molecular Cell 19, 65-75, 2005; Figure 1C).

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Figure 3.22 Examination of BBP Recruitment to An Actively Transcribed Intron-containing Reporter Gene at wild-type and mud2 Background By ChIP. Both mud2 and its isogenic wild-type strains were introduced with the reporter plasmid (pHZ18) and the transcription was induced by galactose for 1 hr. BBP was pulled-down by ChIP method and the bound DNA was analyzed by real-time PCR.

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Figure 3.23 Examination of BBP Recruitment to an Actively Transcribed Intron-containing Reporter Gene at Wild-type and sua7-L214S Background By ChIP. Both sua7-L214S and its isogenic wild-type strains were introduced with the reporter plasmid (pHZ18) and the transcription was induced by galactose for 1 hr. BBP was pulled-down by ChIP method and the bound DNA was analyzed by real-time PCR.

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Figure 3.24 Examination of BBP Recruitment to an Actively Transcribed Intron-containing Reporter Gene Under 6-AU Treatment By ChIP. Wild-type cells containing the pHZ18 reporter was treated with or without 100 mg/ml of 6-AU prior to addition of galactose. After 1 h induction, BBP was pulled-down by the ChIP method and the bound DNA was analyzed by real-time PCR.

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Figure 3.25 Examination of BBP Recruitment to an Actively Transcribed Intron-containing Reporter Gene at Wild-type and msl5-S194P Background By ChIP. Both msl5-S194P and its isogenic wild-type strains were introduced with the reporter plasmid (pHZ18) and the transcription was induced by galactose for 1 hr. BBP was pulled-down by ChIP method and the bound DNA was analyzed by real-time PCR.

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Figure 3.26 Examination of BBP Recruitment to an Actively Transcribed Intron-containing Reporter Gene at Wild-type and msl5-V195D Background By ChIP. Both msl5-V195D and its isogenic wild-type strains were introduced with the reporter plasmid (pHZ18) and the transcription was induced by galactose for 1 hr. BBP was pulled-down by ChIP method and the bound DNA was analyzed by real-time PCR.

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Figure 3.27 The Level of BBP Recruitment to an Actively Transcribed Intron-containing Reporter at Different Genetic Backgrounds and Under 6-AU Treatment. The control, sua7-T101P, is a non-bypass mutation. The peak heights of the BBP recruitment (data from the third oligo pair) under various experimental conditions (Figures 3.20,22-24,26) are normalized to that of the wild-type strain (WT).

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Figure 3.28 Examination of the recruitment of BBP to the YRA1 intron region by RNA-IP Assay. Cells were cross-linked by formaldehyde and the chromatin complexes were sheared into 300-500 bp fragments. After immunoprecipitation of BBP and IgG-Sepharose beads, the RNA crosslinks were reversed and extracted. The YRA1 intron region was reverse transcribed to generate the cDNA and amplify by real-time PCR subsequently. The data from real-time PCR was first normalized to input, subtracted the background, and finally normalized to the wild-type (WT) signal.

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Figure 3.29 URH49 rescues the lethal sub2 deletion in yeast. Shown are the growth phenotypes of the sub2-deletion strains containing plasmids expressing SUB2, UAP56-HA and URH49-HA, which were placed under the control of GAL1 promoter. Transformation with vector alone was used as a negative control. Cells were grown to mid-log phase, 10-fold serially diluted, and spotted on a leucine drop-out plate containing 5-FOA and galactose. The plate was incubated at 30°C for 5 days.

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CHAPTER 4

DISCUSSION

4.1 The In vivo Targets of DExH/D-box Proteins

The ubiquitous DExD/H-box proteins are often referred to as RNA helicases, because, in vitro, they can harness the energy released from NTP hydrolysis to unwind RNA duplexes (de la Cruz et al., 1999; Fuller-Pace, 1994; Jankowsky et al., 2005; Jankowsky et al., 2000). However, whether or not the observed RNA unwinding activity truly represents their actions in the cell is unclear. This is mainly because RNAs are unlikely to exist in naked forms without bound proteins

(Daneholt, 2001; Linder, 2004). Furthermore, RNA duplexes are rarely longer than ten contiguous base pairs and are stabilized by RNA-binding proteins (Staley and Guthrie, 1998). Therefore, it seems more plausible that the physiological targets of DExD/H-box proteins are RNA-protein complexes rather than RNA duplexes alone. Recent studies on several DExD/H-box proteins, including

Prp28p (Chen et al., 2001), Sub2p (Kistler and Guthrie, 2001), Prp5p (Perriman et al., 2003), and Dbp5p (Lund and Guthrie, 2005) appear to support the hypothesis

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that DExD/H-box proteins act as ribonucleoprotein ATPases (RNPases) to remodel RNA-protein complexes in vivo (Linder, 2004; Schwer, 2001).

One of the key findings came from our lab while studying Prp28p (Chen et al.,

2001), an essential DEAD-box splicing factor known to play a role in switching U1 snRNP for U6 snRNP at the 5’ SS (Staley and Guthrie, 1998). It was discovered that specific alterations of the U1C protein, an U1 snRNP component known to stabilize the U1/5’ SS duplex, bypassed the requirement of Prp28p (Chen et al.,

2001). This result suggests that Prp28p counteracts the stabilizing function of

U1C protein by disrupting the interaction of U1C and the 5’ SS (Chen et al., 2001).

Subsequent experiments further indentified two U1-snRNP proteins, Snu71p and

Prp42p, when specifically altered, also bypass the requirement of Prp28p (R.

Hage et al., unpublished). Taken together, it appears that Prp28p may achieve its function at the 5’ SS by remodeling the U1 snRNP via U1C, Snu71p, and/or

Prp42p.

A related observation came from the study of Sub2p (Kistler and Guthrie,

2001). Sub2p is the yeast homolog of the mammalian UAP56, which was implicated in spliceosome assembly via its interaction with U2AF65, presumably by promoting U2 snRNP recruitment to the spliceosome (Zhang and Green, 2001).

Kistler and Guthrie (2001) observed that deletion of the non-essential MUD2 gene rescued the lethal phenotype of sub2. This finding was interpreted as Sub2p playing a role in counteracting Mud2p’s function.

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These two independent lines of research thus suggest an evolutionarily conserved strategy employed at the 5’ SS and the branch site for counteracting the stabilizing effect imposed by sequence-specific binding proteins.

Presumably, in both cases, DExD/H-box proteins remodel the RNP complexes by displacing one or more RNA-bound proteins from their binding sites to permit spliceosome assembly. However, it remains unclear how this is accomplished.

For example, we do not know how individual DExD/H-box proteins locate their targets, how they discriminate non-target sites, and how they interact and exert their effects at the target sties. To gain further insights into these fascinating questions, I set out to identify the in vivo targets of Sub2p by applying the logic used in our previous study on Prp28p.

4.2 BBP is an in vivo Target of Sub2p

A steady and precise RNA-protein interaction is contributed by a set of at least two different forces, both of which can be modulated by DExD/H-box proteins to achieve RNA-protein complex remodeling purposes. In the case of the BPS recognition by BBP, the first supporting force may come from other neighboring components, such as Mud2p, and they could be targeted by Sub2p in spliceosome assembly. BPS-BBP interaction has been proposed to be mediated by multiple proteins in a manner similar to 5’ SS recognition (Zhang and Rosbash,

1999). Zhang and Rosbash (1999) showed that eight proteins can be cross-linked to the 5’ SS in the commitment complex (Zhang and Rosbash, 1999),

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indicating that these protein directly bind to 5’ SS and perhaps can mediate the recruitment of U1 snRNP to form CC1. In addition, Berglund et al. (1997) found that the recombinant RNA binding domain of BBP binds to the BPS with a relatively low affinity with a KD of 500 nM, while other RNA-binding proteins have

KD of 1 nM or even lower (Daly and Wu, 1989; Hall and Stump, 1992). It was proposed that cells preferred utilizing this kind of protein in a multi-component-assembly pathway to avoid a potential problematic rate-limiting step caused by a high affinity binding of a particular protein. Doing so also allows cells to fine-tune the interaction between RNA and protein by regulating the availability of those proteins. In higher eukaryotes, mammalian homolog SF1 shows very limited sequence specificity, with the preference binding of poly(G) and Poly(U) (Arning et al., 1996). SF1’s BPS binding affinity is even lower than that of BBP (Berglund et al., 1997), probably reflecting the nature of a highly degenerated mammalian BPS. It is hence likely that there are more proteins involved in stabilizing the interaction between SF1 and BPS, and alternative splicing can be regulated perhaps by adjusting the recruitment/interaction of assisting proteins in response to different condition. Consequently, it is crucial to identify these assisting proteins to gain more insight of how BPS-BBP interaction is regulated.

The second fundamental force that contributes to the RNA-protein stabilization is the intrinsic binding affinity between BBP and BPS. BBP recognizes a very specific and highly conserved sequence UACUAAC (Berglund

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et al., 1997), which is carried in 209 (91.67%) out of 228 annotated yeast introns

(Spingola et al., 1999). RNA containing alterations of any one of the UACUAAC nucleotides caused significant reduction of BBP’s binding by 5-20 fold or even completely abolished (Berglund et al., 1997). This sequence-specific binding activity of BBP is presumably achieved by the conserved RNA-binding regions

(RH and QUA domains). Consistent with this view, Liu et al. (2001) employed

NMR spectroscopy to show that multiple amino acids in the mammalian SF1’s

RH-QUA fragment interact with a BPS-containing RNA fragment,

U1A2U3A4C5U6A7A8C9A10 A11 (branch point A underlined; conserved BPS in bold).

Alterations of SF1’s BPS-interacting residues abolished its BPS-RNA binding activity in a band-shift experiment (Liu et al., 2001a), suggesting that the KH domain is critical for BPS binding in vitro. In this study, I have isolated two sub2-bypassed BBP variants, encoded by msl5-S194P and msl5-V195D, both of which have substitutions of a conserved KH amino acid residue. The SF1 structure (Liu et al., 2001a) shows that the corresponding yeast Ser194 interacts with the nitrogen base of A11, a neighboring residue which is conserved in ~50% of the yeast introns (Spingola et al., 1999), through hydrogen bonding and electrostatic interaction. A substitution by proline, a rigid cyclic amino acid, at this position may disrupt the secondary structure of the RH domain hence reduce the RNA binding affinity. The corresponding yeast Val195 in SF1 interacts with the nitrogen bases of C9, A10, conserved in ~66% of the yeast introns, and A11 via hydrophobic interaction. A substitution of a very hydrophobic Valine by a

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charged amino acid, such as Aspartic Acid, may abolish the hydrophobic interaction. Based on these observations and assumptions, it is very likely that these two isolated BBP variants have a lower BPS binding affinity in vivo because of the diminished interaction by the original residues. Consistent with this speculation, when the interaction between BPS-BBP variants is reduced to a certain degree, the enzymatic activity required for removing BBP from BPS is dispensable. In this case, msl5-S194P and msl5-V195D both bypass the requirement of Sub2p, strongly arguing that Sub2p is the DExD/H-box protein which functions in the removal of BBP and Mud2p (Kistler and Guthrie, 2001) in spliceosome assembly.

4.3 Only Specific and Subtle Alteration Can Bypass sub2

Another intriguing finding in this study is that only specific and subtle alterations can bypass sub2. The two isolated BBP variants, which carry

Ser-to-Pro or Val-to-Asp substitutions, mainly lose the interaction with the partially conserved (50—65%) neighboring sequence of BPS, i.e. the last three bases (C9,

A10, and A11) of the short RNA, U1A2U3A4C5U6A7A8C9A10A11, used in previously cited structural studies (Liu et al., 2001a). This then raises a hypothesis that only delicate alterations that do not significantly impact on the respective cellular processes can bypass the requirement of Sub2p. Indeed, alterations of other corresponding BPS-interacting residues of BBP, such as Asn163, Arg172 and

Lys196, which may dramatically disrupt the interaction between KH and BPS,

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failed to bypass sub2 (data not shown). In agreement with this proposal,

Sua7p variants shown to elicit a biochemically detected impact (Wu et al., 1999b) on transcription start site selection cannot bypass sub2 in my study.

Another line of evidence that shows sub2 can be bypassed only by a subtle perturbation is from the study of Pol II. In Saccharomyces cerevisiae, RNA polymerase II (RNAP II) consists of 12 subunits encoded by genes termed

RPB1-RPB12 (Woychik, 1998; Woychik and Hampsey, 2002). The catalytic core of RNAP II comprises four subunits, Rpb1p, Rpb2p, Rpb3p and Rpb11p, which are the counterparts of the bacterial a2bb’ RNAP core enzyme. Two mutant alleles, rpb2-7 and rpb2-10, of the RPB2 gene, encode for the second largest subunit of RNAP II, have been isolated and shown to yield 6-AU sensitive phenotype (Powell and Reines, 1996), suggesting they result in defecting elongation. However, only rpb2-10 yielded a slower transcription elongation rate in vitro (Powell and Reines, 1996), suggesting rpb2-10 might result in a severer elongation defect than rpb2-7. In a genetic study of SPT5, which encodes an elongation factor, (Hartzog et al., 1998) showed that only rpb2-10 but not rpb2-7 can suppress a cold-sensitive spt5-242 allele, presumably by reducing the elongation rate. Their finding also indicates the elongation defect caused by rpb2-7 is perhaps milder. In this study, I found that only rpb2-7 can bypass sub2, whereas rpb2-10 cannot (Figure 3.18), again consistent with the proposal that the sub2-bypass alterations must be subtle and specific.

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A high-resolution image of a transcribing Pol II complex reported by Cramer and colleagues (2001) provides a chance to view these two rpb2 mutations in detail. The Rpb2p variant encoded by rpb2-7 has an R122C substitution located within the conserved N' -terminal region (Powell and Reines, 1996), which is thought to interact with other elongation factors such as Rpb12p (Sweetser et al.,

1987) (Figure 4.1). The structure of Rpb2p shows that this N-terminal conserved region forms a protrusion and is located at the peripheral area of Rpb2p (Figure

4.1) (Cramer et al., 2001). In sharp contrast, the Rpb2p variant encoded by rpb2-10 has a P1018S substitution located at the conserved catalytic center of

Rpb2p (Powell and Reines, 1996), which is known to interact with DNA/RNA hybrid (Figure 4.2) and with the largest subunit of RNA polymerase, Rpb1p

(Cramer et al., 2001). Thus, the latter rpb2-10 mutation is expected to exert a much severe negative impact on transcription. Indeed, (Mason and Struhl, 2005) showed that rpb2-10 causes a reduced elongation rate and processivity in vivo.

These findings are again consistent with the model that rpb2-10 impaired transcription elongation activity in a more significant manner than rpb2-7.

Judging from the structural data, the rpb2-7 mutation may only impact on

Rpb2-7p’s interaction with other transcription factors, but not on the intrinsic elongation activity of Pol II, and therefore it can bypass sub2.

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4.4 Transcription and Splicing are Coupled in Yeast

In metazoa, data are abundant to support the functional coupling between transcription and splicing (Aguilera, 2005; Bentley, 2005). Yet this conclusion has been harder to draw in yeast, mainly due to the apparent functional variation of the RNAP II CTD tail (see Introduction). In yeast, the supporting data mainly came from direct interactions of splicing factors with transcription machinery, but how these interactions contribute to the coupling remain largely unclear. For example, Prp40p, an U1-snRNP component, was found to bind to the phosphorylated Pol II CTD (Morris and Greenleaf, 2000) and Sub2p was found to associate with transcription elongation THO complex (Strasser et al., 2002).

Apart from these physical interactions, perhaps a more compelling line of evidence came from the observation that spliceosome assembly occurs in a co-transcriptional and stepwise manner in yeast (Gornemann et al., 2005;

Lacadie and Rosbash, 2005). Nonetheless, whether the observed physical interactions and snRNP recruitment simply reflect the consequence of two connecting processes or they imply a functional coupling remain unclear.

Furthermore, Lacadie et al. (2006) showed that the actual chemistry of splicing, i.e., the cleavage and formation of phosphodiester may occur ~1 kb after transcription past the 3’ splice site of a reporter gene in vivo. In fact, for >90% of yeast genes having second exons shorter than 1 kb, their splicing processes are thought to have taken place post-transcriptionally in vivo (Tardiff et al., 2006).

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In this study, I have presented in vivo evidence demonstrating that specific perturbations of transcription significantly impact on the recruitment of BBP, suggesting that a properly setup of transcription machinery is required for efficient recruitment of splicing factors in yeast. More interestingly, these specific perturbations allow the otherwise essential Sub2p to be dispensable.

Two hypotheses can be advanced to explain the reduced BBP recruitment resulting from the identified perturbations. The first is a kinetic model. In this view, cells carrying these alterations may have a lower Pol II elongation rate and/or processivity, two tightly connected properties of Pol II (Mason and Struhl,

2005). Data have been accumulating to show that that elongation rates could impact on splicing in terms of exon selection both in metazoa (de la Mata et al.,

2003) and in yeast (de la Mata et al., 2003; Howe et al., 2003). These studies identified a similar trend that a fast elongation favors exon skipping while a slower elongation favors exon inclusion (de la Mata et al., 2003; Howe et al., 2003).

They proposed that a slower elongation allows splicing factors to recognize the first intron in a first-come-first-serve fashion, thereby committing splicing of the first intron and hence inclusion of the second exon. In contrast, a faster elongation rate is predicted to increase the probability of the co-existence two introns flanking an exon in a single pre-mRNA. As a result, the likelihood for the distal 5’ SS to pair with the far-end 3’ SS would have been enhanced, leading to the skipping of the exon flanked by the two introns. One possible interpretation of my data can therefore be entertained along this line of thought. It is plausible

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that a slower transcription elongation resulting from chemical or mutational perturbations may allow a wider window of time for U2 snRNP to compete for binding to the intron BPS with BBP and hence bypass the requirement of Sub2p.

Also, a lowering of the Pol II processivity, if that also occurs upon perturbations, is expected to reduce the density of transcription machinery per transcribing gene, and thus results an overall reduced recruitment of BBP. Indeed, one of the sub2-bypass reagent, 6-AU, has been shown to inhibit Pol II elongation rate and processivity to one third of that of the wild-type rate (Mason and Struhl, 2005).

This is consistent with the finding that 6-AU treatment reduces the BBP recruitment to 35% of that in untreated cells shown in Figure 3.24.

Several perturbations that led to the bypass of Sub2p do not seem to yield detectable transcriptional elongation defects. These include rpb2-7 and deletions of genes encoding several elongation factors, including TFIIS, TFIIF,

Chd1p, Hpr1p and Isw1p. An in vivo study showed that the elongation rate and/or Pol II processivity were not detectably altered in mutant strains lacking

TFIIS, Chd1p, or Hpr1p (Mason and Struhl, 2005). These considerations have led me to a second hypothesis that the identified elongation factors may help to recruit key splicing factors, such as BBP, directly or indirectly in a co-transcriptional manner. This hypothesis is supported by the following observations. First, Lacadie et al. (2006) showed that a loss of TFIIS yielded a different splicing phenotype of a reporter and concluded that TFIIS impacts on splicing at a stage later than U1 snRNP recruitment. Second, Howe et al. (2003)

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observed that overexpression of TFIIS stimulates exon skipping in yeast.

Because TFIIS does not appear to play a major role in transcription elongation

(Mason and Struhl, 2005), this exon skipping might have been caused by an increase recruitment of BBP, rather than by enhancement of elongation rate.

Third, elongation factor Hpr1p, a component of THO complex, has been shown to associate with Sub2p together with other THO components, including Tho2p,

Mft1p and Thp2p (Strasser et al., 2002). Thus, co-transcriptional recruitment of splicing factors may raise the local concentration of splicing factors in the vicinity of the nascent transcript to promote splicing.

4.5 Prospectus

Observations reported here provide in vivo evidence that transcription and splicing are functionally coupled in yeast. Several future directions can be pursued to gain deeper insights of the coupling mechanisms. First, it would be interesting to probe whether the exon inclusion and skipping is a consequence of the differentials of the local concentration of co-transcriptionally recruited BBP and other splicing factors under different elongation rates. Second, besides a role in transcription start site selection, Sua7p apparently has a different activity in transcription elongation due to its 6-AU sensitive phenotype (Chen and Hampsey,

2004). It would be interesting to know whether Sua7p and other elongation factors form a complex to recruit critical splicing factors, such as BBP. Third, if the recruiting model is valid, it would be useful to investigate the physical

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interactions between participating elongation factors and splicing factors. Fourth, biochemical studies of Sub2p’s RNPase function can help to elucidate if Sub2p indeed removes both BBP and Mud2p from intron BPS. Finally, even though splicing occurs predominantly post-transcriptionally in yeast, it would be important to know if coupling of transcription and splicing benefits transcription (perhaps at the elongation stage).

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CHAPTER 5

GENETIC ANALYSIS OF Ded1p, AN ESSENTIAL DEXH/D-BOX

TRANSLATION FACTOR

5.1 INTRODUCTION

5.1.1 Ded1p, an Evolutionarily Conserved Essential Translation Initiation

Factor

Ded1p is an essential ATP-dependent DExH/D-box RNA helicase that is conserved in all eukaryotes (Linder, 2003). Like some other DEAD-box proteins,

Ded1p is able to unwind RNA duplexes (Iost et al., 1999), displace proteins from

RNA in an ATP-dependent manner (Fairman et al., 2004), and displays RNA strand-annealing activity in the absence of ATP (Yang and Jankowsky, 2005).

Despite all these biochemical characterization of Ded1p, its role in the cell remains to be fully defined.

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Historically, Ded1p was first thought to participate in pre-mRNA splicing because a ded1 mutant allele could rescue the growth and splicing defects caused by the prp8-1 mutation (Jamieson et al., 1991). In addition, Ded1p may also have a role in RNA polymerase III (Pol III) transcription, because its overexpression could suppress the growth defect of a Pol III mutant (Thuillier et al., 1995). Later, Dr. Ray-Yuan Chuang, a former graduate student in the Chang laboratory, has convincingly demonstrated that Ded1p plays a key role in translation (Chuang et al., 1997). It was observed that protein synthesis was completely shut off in cold-sensitive ded1 mutants at 16̓C with concurrent accumulation of 80S complexes (Chuang et al., 1997). Supporting this observation, it was found that a combination of mutant alleles of DED1 and TIF1, which encodes the translation initiation factor eIF4A, results in a lethal phenotype

(Chuang et al., 1997). Critically, depletion of Ded1p from cell extract abolishes protein synthesis in vitro, and addition of purified recombinant GST-Ded1p restores the activity (Chuang et al., 1997). Notably, polysome profile analysis revealed that ded1 mutants accumulate inactive forms of 80S ribosomes at 16̓C in vivo and detailed in vitro translation analysis showed that messenger ribonucleoprotein complexes (mRNPs) accumulated upon Ded1p depletion

(Chuang et al., 1997). Taken together, these data suggest that Ded1p is likely to function at an early stage of translation initiation prior to the recruitment of the 40S ribosomal subunit to mRNPs (see also Chuang, 1997).

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Ded1p may have an additional role in serving as a host factor to promote viral replication in several evolutionarily related RNA viruses. First, Ded1p is required for translating mRNAs produced by brome mosaic virus (BMV) in yeast (Noueiry et al., 2000). Second, Dr. Jean-Leon Chong, a former graduate student in the

Chang laboratory, showed that Ded1p promotes negative-strand synthesis of the yeast L-A virus in vitro (Chong et al., 2004). Third, the human ortholog of Ded1p was found to interact with the hepatitis C virus (HCV) core protein (Mamiya and

Worman, 1999; Owsianka and Patel, 1999; You et al., 1999).

Besides its essential function in translation initiation, several lines of unpublished data from our laboratory suggest that Ded1p may also play a role in mRNA splicing. First, genome-wide analysis using splicing-sensitive microarrays revealed that intron-containing transcripts of the ribosomal protein

(RP) genes preferentially accumulated in ded1 mutants (Burckin et al., 2005;

Chang, T.H. et al., unpublished data). Second, the spliceosomal small nuclear

RNAs (snRNAs; U1, 2, 4, 5, and 6) can be co-immunoprecipitated with Ded1p

(Burckin et al., 2005); Chong, 2005). Third, alteration of the splicing commitment complexes was detected in splicing reactions assembled using extracts prepared from ded1 mutants (Chong, 2005).

5.1.2 The Role of eIF4G in Translation Initiation

To expand our understanding of Ded1p’s function, Dr. Jean-Leon Chong initiated an open-ended genetic screen for mutations that, in combination with the

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ded1-120 mutation, result in a synthetic-lethal phenotype (J.-L. Chong, unpublished). This led us to the identification of TIF4631, which encodes one of the two eIF4G translation initiation factors, again supporting Ded1p’s role in translation initiation. In yeast, there are two eIF4G homologs encoded by

TIF4631 and TIF4632 and the two proteins share 53% identity (Goyer et al., 1993).

Deletion of TIF4631 yielded a slow-growth and cold-sensitive phenotype, whereas

TIF4632-deleted cells exhibited no detectable growth phenotype (Goyer et al.,

1993). However, simultaneous deletion of TIF4631 and TIF4632 is lethal, indicating that eIF4G is functionally essential (Goyer et al., 1993).

The role of eIF4G in translation is to serve as a dynamic scaffold for assembling translation initiation factors for effective translation initiation. eIF4G, together with eIF4E and eIF4A, forms a holoenzyme termed eIF4F to promote the recruitment of the 40S ribosomal subunit to mRNAs (Prevot et al., 2003). This recruitment is mediated by eIF4G’s interaction with eIF3, which in turns interacts with the 40S ribosomal subunit, thus leading to the formation of the 43S initiation complex (Prevot et al., 2003). Because eIF4G can simultaneously interacts with the poly (A)-binding protein (Pab1p) and eIF4E, the cytoplasmic cap-binding protein, it presumably facilitates the circularization of the mRNA into a form efficient for subsequent rounds of translation using the same ribosome (Imataka et al., 1998; Tarun and Sachs, 1996; Wells et al., 1998). Surprisingly, genetic and biochemical studies showed that eIF4G also interacts with Cbp80p, the nuclear cap-binding protein (Fortes et al., 2000; Fortes et al., 1999). Notably, the

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Cbp80p-interacting region in eIF4G overlaps with the eIF4E/eIF4A-binding site

(Fortes et al., 2000). It was therefore suggested that the mRNP-bound nuclear cap-binding complex, including Cbp80p and Cbp20p, is replaced by the cytoplasmic eIF4E during or soon after mRNP’s exit from the nucleus (Fortes et al., 2000). In mammalian cells, eIF4G was also observed to associate with nuclear cap-binding complex in nucleus (McKendrick et al., 2001), and eIF4G was speculated to participate in the pioneer round of nuclear translation and facilitate mRNAs export (Ishigaki et al., 2001).

5.1.3 Retrograde Signal Transduction Pathways

In another effort to identify novel synthetic-lethal mutations, an allele of RTG3 was identified (L. Tung and J. Yang, unpublished). RTG3 encodes a basic helix-loop-helix-leucine zipper (bHLH/Zip) transcription factor (Crespo et al., 2002;

Jia et al., 1997; Rothermel et al., 1997) and forms a complex with another bHLH/Zip protein, Rtg1p (Liao and Butow, 1993). In response to changes of the functional state of mitochondria, Rtg3p/Rtg1p activates the retrograde (RTG) signaling pathway by altering the expression of a large set of nuclear genes

(Parikh et al., 1987; Shyjan and Butow, 1993). RTG signaling is turned on when mitochondrial respiratory function is compromised, or when the key metabolic pathways, such as TCA cycle, are disrupted (Epstein et al., 2001; Liao et al., 1991;

Liu and Butow, 1999; McCammon et al., 2003). This retrograde mitochondrial signaling pathway is also involved in regulating apoptosis, aging, and many

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pathophysiological states (Butow and Avadhani, 2004). RTG signaling relies on translocation of Rtg1p/Rtg3p complex from cytoplasm into nucleus to activate

RTG target genes, which includes CIT2 (Jia et al., 1997; Liao et al., 1991; Sekito et al., 2000), encoding the isoform of citrate synthase that functions in the glyoxylate cycle (Figure 5.1) (Lewin et al., 1990; McCammon et al., 1990). The subcellular translocation of Rtg1p/Rtg3p complex is facilitated by Rtg2p (Liao and

Butow, 1993; Liu et al., 2003), probably via sensing the cellular glutamate concentration, and negatively regulated by Mks1p and Bmh1p/Bmh2p complex

(Liu et al., 2003; Sekito et al., 2002) (Figure 5.2).

5.1.4 TOR Signal Transduction Pathway

The nuclear accumulation of Rtg1p/Rtg3p complex and the resulting alteration of nuclear gene expression can also be induced by rapamycin, a specific inhibitor of the target of rapamycin (TOR) kinases (Komeili et al., 2000;

Shamji et al., 2000). Rapamycin was originally used as an immunosuppressive drug that blocks signal transduction pathways required for the activation of helper

T cell (Heitman et al., 1991; Schreiber, 1991). Because of its specificity, rapamycin is now widely used to inhibit the TOR signaling pathway. Rapamycin acts by forming a complex with Fpr1p (a peptidyl-prolyl isomerase) in yeast or with

FKBP (FK506-binding protein, a proline rotamase) in mammalian cells (Heitman et al., 1991). The rapamycin/Fpr1p or rapamycin/FKBP complex then inhibits the yeast Tor1p and Tor2p and their mammalian homolog mTOR/FRAP/RAFT

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(Heitman et al., 1991; Stan et al., 1994). Both Tor1p and Tor2p, which are highly conserved from yeast to human, contain a C-terminal kinase domain that is responsible for phosphorylating the serine and threonine residues of their target proteins (Martin and Hall, 2005).

Treatment of yeast cells with rapamycin or inactivation of both TOR genes induces a starvation-like response (Barbet et al., 1996), including shutdown of translation initiation, cell-cycle arrest at the G1 phase, and expression of

G0-specific genes (Barbet et al., 1996; Chan et al., 2000; Kunz et al., 1993). As a result, TOR pathway is believed to play a critical role in regulating cell growth in response to the availability of nutrients. Importantly, TOR pathway is reported to coordinate and integrate cell physiology and environmental cues for proper cell growth (Hardwick et al., 1999; Mayer and Grummt, 2006) by activating translation initiation, organization of the actin cytoskeleton, membrane traffic, protein degradation, PKC (Protein Kinase C) signaling, ribosome biogenesis, and transcription (Cardenas et al., 1999; Rohde et al., 2001; Schmelzle and Hall,

2000). When nutrients are limiting, the TOR pathway responds by negatively regulating the production of ribosomes (Hannan et al., 2003), which consumes most of the energy in the cell (Warner, 1999). TOR signaling achieves this aim by modulating the transcription activities of Pol I, II, and III, translation of mRNAs encoding ribosomal proteins, and pre-rRNA processing (Mayer and Grummt,

2006).

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To modulate translation initiation, TOR signaling pathway appears to act by influencing the stability of eIF4G in yeast (Berset et al., 1998). Addition of rapamycin was found to cause degradation of eIF4G over a period of 2—3 hours, while other translation initiation factors remained unaffected (Berset et al., 1998).

In mammalian cells, this degradation of eIF4G is speculated to be a consequence of dissociating eIF4E from eIF4G (Beretta et al., 1996; von Manteuffel et al., 1996).

However, the nature of this degradation pathway of eIF4G remains unknown.

In this work, I will present preliminary genetic and biochemical data linking Ded1p to the RTG- and TOR-signaling pathways.

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Figure 5.1 Glyoxylate Cycle (Adapted and modified from Lehninger-Principles of Biochemistry by Nelson/Cox, 4th ed. W. H. Freeman Publishing, 2005)

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Figure 5.2 Regulation of the RTG Pathway. When the mitochondrial function is compromised, rapamycin is present, or the cellular glutamate level is low, the RTG signaling pathway is turned on by association of Rtg2p and partially phosphorylated Mks1p. This event releases the inhibition of Mks1p/Bmh1/2p complex and allows translocation of the Rtg1p/Rtg3p complex from cytoplasm to nucleus. Nuclear Rtg1p/Rtg3p complex binds to the promoter of RTG target genes such as CIT2, which encodes a citrate synthase functions in glyoxylate cycle to provide intermediates for TCA cycle. When glutamate is available, TOR proteins are functional, or mitochondria are healthy, RTG pathway is turned off by keeping hyperphosphorylated Rtg3p as well as Rtg1p in the cytoplasm. Adapted and modified from (Liu et al., 2003).

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5.2 MATERIALS AND METHODS

5.2.1 Yeast Strains

All yeast strains used in this work are listed in Table 5.1.

5.2.2 Plasmids

All plasmids used in this study are listed in Tables 5.2-5.3.

5.2.3 Oligos

All the oligos used in this study are listed in Table 5.4.

5.2.4 Serial Deletion of the C-terminal Region of Ded1p

Six oligos (DED1-53 to DED1-58) were designed for amplifying a serial

C-terminal deleted DED1 fragments by PCR. All the truncated ded1 alleles were cloned onto the LEU2-marked vector pRS315 resulting in pDED1107-to-pDED1112 plasmids. Each plasmid was next transformed into yeast strain YTC74 [MATaʳura3-52 lys2-801 ade2-101 trp1-1 his3200 leu2-1 ded1::TRP1 DED1/URA3/CEN (=pDED1008)]. The resulting transformants were purified and streaked on 5-FOA plates to test if the acquired truncated ded1 allele can replace the existing wild-type DED1/URA3/CEN plasmid in YTC74.

5-FOA+ cells were collected, spotted on YPD plates, and incubated at 16̓C, 30̓C and 37̓C for temperature sensitivity examination.

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5.2.5 Examination of C-terminal Deleted Ded1p by Immuno Blot Analysis

Yeast strains carrying different C-terminal truncated DED1 alleles were grown in 50 ml of YPD until OD600=1. Cells were collected and broken by glass beads method in lysis buffer [50 mM Tris-HCl (pH 7.5), 100 mM NaCl, 1 mM DTT, 1 mM

EDTA, and 1 mM PMSF]. The debris was removed from cell extract by high speed centrifugation at 4̓C. Protein concentration was determined by Bradford assay (BioRad). About 45 mg of total protein of each cell extract was separated on 8% SDS-PAGE, and then transferred to a nitrocellulose membrane by electroblotting. The membrane was blocked in 100 ml of blocking buffer containing 5% nonfat dry milk (Meijer) in TBST [20 mM Tris-HCl (pH 8.0), 150 mM

NaCl, 0.05% Tween-20] for 1 hour at room temperature. The primary polyclonal rabbit anti-Ded1p or anti-Dbp5p antibodies were diluted 5000-fold in the blocking buffer and incubated with the membrane at 4̓C overnight. The membrane was washed three times with TBST followed by a 1-hour incubation with 5000-fold diluted secondary HRP (horseradish peroxidase)-conjugated goat anti-rabbit antibodies in the blocking buffer at room temperature. The membrane was washed three times with TBST and then developed using ECL chemiluminescence detection system (Amersham).

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5.2.6 Genetic Screening and Identification of Mutants Which Are Synthetic

Lethal to ded1-120

All yeast procedures were performed using standard protocols (Guthrie and

Fink, 1991). The yeast strain YTC896 (MATa ded1::TRP1 ade2 ade3 his3 leu2 lys2 ura3 pDED1090 [=ded1-120/LYS2/CEN] pDED1120

[=DED1/ADE3/URA3/CEN]) was mutagenized by UV irradiation to achieve 80% killing rate. Survival colonies were replica-plated onto 5-FOA plates lacking lysine (5-FOA-Lys) for maintenance of ded1-120 plasmid (pDED1090) and identification of 5-FOA sensitive colonies, candidates which may carry synthetic lethal mutations. To identify the wild-type allele corresponding to the mutant from the 5-FOA-sensitive isolates, each candidate was transformed with a

LEU2-marked (YCp50) genomic library to yield ~8000 transformants on

SD-Leu-Lys double drop-out plates. 5-FOA+ colonies were subsequently identified upon replica-plating the transformants onto 5-FOA-Leu-Lys plates.

YCp50 plasmids from all 5-FOA+ isolates were extracted and re-transformed back to the 5-FOA-sensitive candidate to confirm the rescue of each YCp50 clone.

YCp50 plasmids that reproducibly yielded the 5-FOA+ phenotype were selected for DNA sequencing from both ends of the insert by oligos YCP50-1 and YCP50-2

(Table 2.3). Finally, each ORF carried on the YCp50 plasmid was individually cloned and transformed into the synthetic lethal candidate to examine which one is the corresponding allele. TIF4631 was identified from screening of yeast strain YTC982 and RTG3 was identified from YTC1109.

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5.2.7 Examination of Genetic Interaction between DED1 and Other

Non-essential Genes

To investigate the genetic interaction between DED1 and other non-essential genes, double-deletion strains containing chromosomal deletion of each gene of interest (e.g. tif4631::kanMX4), ded1, and the complementary DED1 allele on a URA3-marked plasmid (pDED1008) were constructed. Next, four

LEU2-marked plasmids carrying DED1 (pDED1009), ded1-120 (pDED1009), ded1-199 (pDED1009), and ded1-95 (pDED1009) alleles were transformed into each double-deletion strain. Finally, the resulting transformants were purified and streaked on 5-FOA plates to examine the synthetic lethality between each

DED1 allele and deletion of each gene of interest.

5.2.8 Rapamycin Sensitivity/Resistance Test

To test the rapamycin sensitivity/resistance of each testing strain, equal amount of cells carrying either wild-type DED1 (strain YTC1085), ded1-120 (strain

YTC1086), ded1-199 (strain YTC1087), or ded1-95 (strain YTC1088) allele were serially diluted and spotted on YPD plates containing different concentrations

(25-200 mg/ml) of rapamycin and incubated at 30̓C for six days.

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Name Genotype Source YTC74 MATaʳura3-52 lys2-801 ade2-101 trp1-1 his3200 Chang Lab leu2-1 ded1::TRP1 DED1/URA3/CEN (=pDED1008)

YTC896 MATa ade2 ade3 his3 leu2 lys2 ura3 Chang Lab ded1::TRP1 ded1-120/LYS2/CEN (=pDED1090) DED1/ADE3/URA3/CEN (=pDED1120)

YTC982 MATa ade2 ade3 his3 leu2 lys2 ura3 Chang Lab ded1::TRP1 sl49-1 ded1-120/LYS2/CEN (=pDED1090) DED1/ADE3/URA3/CEN (=pDED1120)

YTC1004 MATa tif4631::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1005 MATa tif4632::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1009 MATa ade2 ade3 his3 leu2 lys2 ura3 Chang Lab ded1::TRP1 sl11 ded1-120/LYS2/CEN (=pDED1090) DED1/ADE3/URA3/CEN (=pDED1120)

YTC1025 MATa rtg3::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1056 MATa rtg1::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1057 MATa rtg2::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1058 MATa cit1::kanMX4 ura3 leu2 his3 met15 Open Biosystems

Table 5.1 Yeast Strains

(continued)

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Table 5.1 (continued)

Name Genotype Source YTC1059 MATa cit2::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1060 MATa mdh1::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1061 MATa mdh2::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1064 MATa msl2::kanMX4 ura3 leu2 his3 met15 Open Biosystems

YTC1068 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 cit1::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1070 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 mdh1::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1072 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 mdh2::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1074 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 rtg1::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1077 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 rtg2::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1078 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 msl2::kanMX4 DED1/URA3/CEN (=pDED1008)

(continued)

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Table 5.1 (continued)

Name Genotype Source YTC1085 MATaʳura3-52 lys2-801 ade2-101 trp1-1 his3200 This Study leu2-1 ded1::TRP1 DED1/LEU2/CEN (=pDED1009)

YTC1086 MATaʳura3-52 lys2-801 ade2-101 trp1-1 his3200 This Study leu2-1 ded1::TRP1 ded1-120/LEU2/CEN (=pDED1018)

YTC1087 MATaʳura3-52 lys2-801 ade2-101 trp1-1 his3200 This Study leu2-1 ded1::TRP1 ded1-199/LEU2/CEN (=pDED1020)

YTC1088 MATaʳura3-52 lys2-801 ade2-101 trp1-1 his3200 This Study leu2-1 ded1::TRP1 ded1-95/LEU2/CEN (=pDED1022)

YTC1100 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 cit2::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1107 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 rtg3::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1022 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 tif4631::kanMX4 DED1/URA3/CEN (=pDED1008)

YTC1023 MATa ade2-101 ura3 leu2 his3 met15(?) lys2(?) This Study trp1(?) ded1::TRP1 tif4632::kanMX4 DED1/URA3/CEN (=pDED1008)

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! Name Description pDED1008 A 2.9 kb XhoI-SalI fragment containing the entire length of the DED1 gene isolated from pUC-Sc2605 (gift from K. Struhl) was cloned into pRS316 (URA3/CEN). pDED1009 A 2.9 kb XhoI-SalI fragment containing the entire length of the DED1 gene isolated from pUC-Sc2605 (gift from K. Struhl) was cloned into pRS315 (LEU2/CEN). pDED1018 Derived from pDED1009 by hydroxylamine mutagenesis, carrying ded1-120 mutant allele (G108D & G494D) on pRS315 (LEU2/CEN). pDED1020 Derived from pDED1009 by hydroxylamine mutagenesis, carrying ded1-199 mutant allele (G368D) on pRS315 (LEU2/CEN). pDED1022 Derived from pDED1009 by hydroxylamine mutagenesis, carrying ded1-95 mutant allele (T408I) on pRS315 (LEU2/CEN). pDED1090 A ~2.9 kb ded1-120 fragment isolated from pDED1018 was cloned onto pRS317 (LYS2/CEN) vector. pDED1107 PCR product of DED1 fragment containing 15 amino acids deleted at the C-terminal generated by using oligos DED1-53/T3 and pDED1001 as the template was cloned to pRS315 (LEU2/CEN). pDED1108 PCR product of DED1 fragment containing 30 amino acids deleted at the C-terminal generated by using oligos DED1-54/T3 and pDED1001 as the template was cloned to pRS315 (LEU2/CEN). pDED1109 PCR product of DED1 fragment containing 45 amino acids deleted at the C-terminal generated by using oligos DED1-55/T3 and pDED1001 as the template was cloned to pRS315 (LEU2/CEN).

Table 5.2 DED1 and RTG3 Plasmids

(continued)

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Table 5.2 (continued)

Name Description pDED1110 PCR product of DED1 fragment containing 60 amino acids deleted at the C-terminal generated by using oligos DED1-56/T3 and pDED1001 as the template was cloned to pRS315 (LEU2/CEN). pDED1111 PCR product of DED1 fragment containing 75 amino acids deleted at the C-terminal generated by using oligos DED1-57/T3 and pDED1001 as the template was cloned to pRS315 (LEU2/CEN). pDED1112 PCR product of DED1 fragment containing 90 amino acids deleted at the C-terminal generated by using oligos DED1-58/T3 and pDED1001 as the template was cloned to pRS315 (LEU2/CEN). pDED1120 A ~2.9 kb DED1 fragment isolated from pDED1008 was cloned onto pHT4467 (ADE3/URA3/CEN) vector. pRTG3001 A 2.4~ kb RTG fragment was amplified by oligo RTG3-1 and RTG3-2 and cloned onto pRS415 (LEU2/CEN).

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Name Sequence DED1-53 TTTCTCGAGTCAAGACTTGGAATCGCTACC DED1-54 TTTCTCGAGTCAATCTCTGCTTCTTGAAGA DED1-55 TTTCTCGAGTCAGGCCTTACGGTAATCTCT DED1-56 TTTCTCGAGTCAACGGCTGTTGCTTCTGCT DED1-57 TTTCTCGAGTCACAAGAATGATGGGACTTC DED1-58 TTTCTCGAGTCACAAACCTTTAACAATGTT RTG3-1 GCCCAGCTCTAGAACAATGTCTTCGCAGATTC RTG3-2 TACATAGGATCCTGCTTGCCTATCTCTTCCAC RTG3-3 TCCGAATTTCTGTC RTG3-4 CAGCATGGCACCGAT T3 ATTAACCCTCACTAAAG

Table 5.3 Oligos

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5.3 RESULTS

5.3.1 C-terminal Truncation of Ded1p is Lethal

Previously Ded1p was shown to physically interact with the yeast L-A virus

(Chong et al., 2004). To define a minimum region in Ded1p that is responsible for this interaction, I engineered a set of deletions in which 15 amino-acid segments were incrementally removed from Ded1p C-terminus. A total six ded1-truncate alleles, i.e. ded1-15-a.a. to ded1-90-a.a., were constructed and cloned on a LEU2-marked plasmid. To examine the impact of these deletions on cell growth in the absence of the wild-type Ded1p, I introduced these clones into a tester strain (YTC74), in which the chromosomal DED1 was deleted and complemented by a DED1 allele carried on a URA3-marked plasmid (pDED1008).

The resulting transformants were spotted on 5-FOA plates to inspect if the ded1-truncate allele in question can replace the function of the plasmid-borne

DED1 at various temperatures. As shown in Table 5.3, deletions up to 45 amino acids permit the cells to grow equally well as the wild-type strains, indicating that the remaining 559 amino acids are sufficient to support wild-type Ded1p’s function.

Cells carrying ded1-60-a.a. and ded1-75-a.a. alleles are viable but sick at all tested temperatures, suggesting that the remaining 529 amino acids still provide some essential but compromised Ded1p activity. Deletion of 90 amino acids, though, resulted in a lethal growth phenotype, implying that the deleted region is critical for Ded1p’s function.

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5.3.2 Truncated Ded1p Is Over-expressed but Unstable in vivo

The protein expression levels of the truncated Ded1p variants were examined by immunoblots using anti-Ded1p antibody (Chuang et al., 1997). As shown in

Figure 5.3, each truncated Ded1p was expressed in a size as expected.

However, truncated Ded1p variants that lacks 60 and 75 amino acids (lanes 4 and 5) from the C-terminus is over-expressed and appears unstable in vivo.

Cells carrying the ded1-60-a.a. truncation exhibited the severest growth phenotype (Table 5.3), but express the highest level of the truncated Ded1p together with its degradation products (Figure 5.3). This overexpressed but unstable pattern is similar to that of the production of the mutant Ded1p in ded1-120 and ded1-199 mutants (J.-L. Chong, unpublished data).

5.3.3 Deletion of TIF4631 but not TIF4632 Caused a Lethal Growth

Phenotype with ded1-120 and ded1-199

In an open-ended genetic screen for mutations that act synergistically with the ded1-120 mutation, a candidate recessive mutation was isolated (J.-L. Chong, unpublished). Subsequent screening of a genomic library led to a plasmid clone harboring the TIF4632 gene that can rescue the synthetic lethality. Because neither TIF4631 nor TIF4632 is an essential gene, one would expect that their deletion alleles might be synthetic lethal to ded1-120 as well. To test this, I constructed two tester strains each containing either a tif4631 (strain YTC1022) or tif4632 (strain YTC1023) in conjunction with a chromosomal deletion of ded1,

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which was complemented by a URA3-marked DED1 plasmid. These two strains were transformed with plasmids containing either ded1-120 and ded1-199 alleles and the resulting strains were examined on the 5-FOA plates. Interestingly, only the combination tif4631 with either ded1-120 or ded1-199 failed to permit cell growth on the 5-FOA plates. This result indicates that tif4631, but not tif4632, is synthetic lethal with the ded1 mutant alleles (Table 5.4), consistent with a previous finding by de la Cruz et al. (1997). It is likely that the observed rescue of the synthetic-lethal phenotype by TIF4632 is caused by a dosage-dependent effect due to the high-degree similarity between Tif4361p and Tif4362p.

5.3.4 Deletion of RTG1, RTG2, or RTG3 is Synthetic Lethal with ded1-120 and ded1-199

A second mutation that is synthetic lethal to ded1-120 was uncovered from another screen (C.-H. J. Yang and L. Tung, unpublished). Screening a genomic library, I found the synthetic-lethal phenotype can be rescued by a plasmid clone containing two intact ORFs, SFT2 and RTG3. Subcloning analysis demonstrated that RTG3 is responsible for the rescue. The mutant rtg3 allele was found to contain an insertion within codon 328 (AAT to AAAT), resulting in a frame shift and predicting a truncated RTG3 gene product of 335, instead of the original 486, amino acid residues. Because RTG3 is a non-essential gene and the isolated rtg3 allele is recessive, I tested the synthetic lethality between rtg3 and ded1 mutant alleles using a similar approach described earlier (see 5.4.3)

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and found that they are indeed synthetic lethal to each other (Table 5.4). Since

Rtg3p functionally interacts with Rtg1p and Rtg2p, I then tested if the same synthetic lethality could be observed between ded1 alleles and rtg1 or rtg2.

This proved to be the case (Table 5.4).

5.3.5 Deletion of Genes Related to Glyoxylate Cycle Do Not Result in

Synthetic Lethality in Combination of ded1-120 or ded1-199

Because RTG1, RTG2 and RTG3 are involved in the glyoxylate cycle by activating the CIT2 gene transcription, it is tempting to speculate that the synthetic-lethal phenotype is a result of the glyoxylate cycle failure. I tested this hypothesis using the approach described previously (see 5.4.3) by recombining deletion alleles of genes encoding enzymes related to glyoxylate cycle, cit1, cit2, mdh1, mdh2, and mls2, individually into the ded1 mutant background.

CIT1 and CIT2 encode two isoforms of citrate synthase involved in TCA and glyoxylate cycles, respectively. MDH1 and MDH2 encode two of the three isoforms of malate dehydrogenase involved in TCA cycle and gluconeogenesis, respectively. MLS2 encodes malate synthase that is required to convert glyoxylate into malate. None resulted in a synthetic-lethal phenotype (Table 5.4), suggesting that synthetic-lethal phenotype is not caused by general glyoxylate cycle breakdown.

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5.3.6 ded1 Mutant Strains Are Rapamycin Resistant

It has been suggested that the RTG signaling pathway is linked to the TOR signaling pathway, because treatment of rapamycin results in nuclear translocation of Rtg1p/Rtg3p as well as the expression of the CIT2 gene (Dilova et al., 2004). Because deletion of RTG genes causes a rapamycin resistant phenotype (Xie et al., 2005), I examined if cells carrying ded1-120 or ded1-199 alleles also respond to rapamycin. As expected, ded1-120 and ded1-199 strains grew better than the wild-type cells in the presence of rapamycin with a concentrating ranging 25 to 200 mg/ml (Figure 5.4). This result suggests that

TOR signaling is blocked in the ded1 mutant strains.

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Figure 5.3 Protein Expression of C-terminal truncate Ded1p in yeast. Truncate forms of Ded1p were examined by Western blot from cell extracts prepared from yeast strains carrying different C-terminal deleted ded1 alleles. 45 mg of total cell extracts was loaded per lane. 1: ded1-15-a.a.; 2: ded1-30-a.a.; 3: ded1-45-a.a.; 4: ded1-60-a.a.; 5: ded1-75-a.a.. Top panel, polyclonal rabbit anti-Ded1p antibodies diluted 5000 fold was used. Bottom panel, 5000-fold diluted polyclonal rabbit anti-Dbp5p antibodies were used as an internal loading control.

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Figure 5.4 Cells carrying ded1-120 and ded1-100 display rapamycin resistant phenotype. Yeast cells containing either wild-type DED1, ded1-120, ded1-199 and ded1-95 alleles were serially diluted and spotted on YPD plates with (right panel) and without (left panel) rapamycin at a concentration of 75 mg/ml).

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DED1 Allele 16̓C 30̓C 37̓C DED1 ++ +++ +++ ded1-15-a.a. ++ê ++ê ++ ded1-30-a.a. +++ +++ ++ê ded1-45-a.a. +++ +++ ++ ded1-60-a.a. + ++ +ê ded1-75-a.a. ++ê ++ê + ded1-90-a.a. - - -

Table 5.4 Growth Phenotype of Strains Containing C-terminal Serially Deleted Ded1p at Different Temperature

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Allele Function ded1-120 ded1-199 rtg1 transcription activator - - rtg2 RTG regulator - - rtg3 transcription activator - - cit1 citrate synthase +++ +++ cit2 citrate synthase +++ +++ mdh1 malate dehydrogenase +++ +++ mdh2 malate dehydrogenase +++ +++ mls2 malate synthase +++ +++ tif4631 eIF4G - - tif4632 eIF4G +++ +++ sub2-85 mRNA splicing/export factor +++ +++ mex67-6 mRNA export factor +++ +++

Table 5.5 Alleles Used for the Synthetic Lethality Test in Relation to ded1-120 and ded1-199

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5.4 DISCUSSION

In this chapter, I describe a series of preliminary analysis aiming to understand better Ded1p’s role in the cell. Unexpectedly, this analysis led to the finding that the RTG signaling pathway is genetically linked to Ded1p’s function, which in turn is related to the TOR signaling pathway.

5.4.1 Ded1p’s C-terminal Region is Important for Its Stability in vivo

I showed that deletion of the C-terminal 90 amino acids from Ded1p is lethal

(Table 5.3), despite the fact that the truncated Ded1p would still maintain all the essential functional domains (Figure 1.2) present in all DExH/D-box proteins. I observed that the truncated forms of Ded1p lacking the C-terminal 60 or 75 amino acid residues were expressed twice as much as the wild-type Ded1p in vivo

(Figure 5.3). A similar pattern was previously documented for two mutant forms of Ded1p encoded by ded1-120 and ded1-199 alleles (J.-L. Chong, unpublished).

It is therefore tempting to speculate that cells compensate the partial loss of function of Ded1p resulted from either mutations or truncations by overexpressing

Ded1p. In the cases of C-terminal deleted Ded1p, the partial loss of function is probably due to protein instability in vivo, because more degraded products can be detected in the corresponding extracts (Figure 5.3).

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5.4.2 RTG signaling pathway is genetically linked to Ded1p’s function

When yeast cells sense mitochondrial defects and/or low level of glutamate, a precursor critical for synthesizing other amino acids and nucleotides, the Rtg transcription factors are mobilized to activate the glyoxylate cycle, thereby compensating the loss of metabolic intermediates required for TCA cycle (Crespo et al., 2002; Liao and Butow, 1993; Liu and Butow, 1999; Rothermel et al., 1997).

In respiratory competent cells or when the glutamate is supplemented in the growth medium, Rtg3p is hyperphosphorylated and is sequestered in an inactive complex with Rtg1p in cytoplasm (Sekito et al., 2000). When the respiration is repressed or glutamate is low in the medium, Rtg3p is partially dephosphorylated and translocates into the nucleus with to activate RTG target genes such as CIT2

(Sekito et al., 2000). This translocation is regulated by the dynamic interaction between Rtg2p and Mks1p (Liu et al., 2003) as well as by 14-3-3 proteins (Bmh1p and Bmh2p in yeast) (van Heusden and Steensma, 2001) and Lst8p (Liu et al.,

2001b; Roberg et al., 1997).

There are at least three hypotheses to explain the genetic interaction between DED1 and RTG genes observed in this study. First, Ded1p is involved in the TCA cycle, because a simultaneous compromise of both TCA and glyoxylate cycle results in a lethal phenotype, as evidenced by the fact that a cit1 rtg1 double mutant is inviable (Small et al., 1995). This seems unlikely, though, because ded1 cit2 and ded1 mls2 double mutants are viable. Both CIT2 and

MSL2 genes are involved in glyoxylate cycle, but not in TCA cycle.

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The second hypothesis is that Ded1p plays a role in the glyoxylate cycle.

However, the fact that ded1 cit1 and ded1 mdh1 double mutants are also viable does not provide support for this hypothesis. . Both CIT1 and MDH1 genes are involved in TCA cycle, but not in glyoxylate cycle.

The third hypothesis is that the Rtg proteins have a novel role related to one of the documented functions of Ded1p, such as translation initiation and pre-mRNA splicing. Additional work is required to specifically investigate this possibility in the future.

5.4.3 Ded1p is a Possible Regulator of eIF4G Degradation in TOR Pathway

Existing data suggest that the RTG and the TOR signaling pathways are linked to each other. For example, rapamycin treatment induces nuclear translocation of Rtg1p and Rtg3p (Komeili et al., 2000) and Lst8p, a negative regulator of the RTG pathway (Liu et al., 2001b), is present in the TOR complexes in yeast (Chen and Kaiser, 2003; Loewith et al., 2002; Wedaman et al., 2003).

The finding that Ded1p is linked to the RTG pathway prompted an investigation of a possible linkage of Ded1p to the TOR pathway. This speculation was borne true by the fact that ded1-120 and ded1-199 mutants are rapamycin resistant

(Figure 5.4). Inactivation of the TOR genes, rapamycin treatment, and starvation have all been reported to cause translation initiation arrest (Barbet et al., 1996).

Several reports have documented that eIF4G is rapidly degraded upon rapamycin treatment or at diauxic growth (starvation), whereas eIF4E and eIF4A remain

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stable (Berset et al., 1998; Kuruvilla et al., 2001; Powers and Walter, 1999), thus providing a possible explanation of the translation initiation arrest caused by inactivation of the TOR pathway. This reasoning is also supported by observation that the amount of eIF4G is significantly reduced in cells that are deficient of Ypk1p, a Ser/Thr protein kinase that regulates translation initiation through TOR pathway (Gelperin et al., 2002). It has been reported that in yeast and mammalian cells, the free form of eIF4G is more susceptible to degradation than the eIF4E-bound form (Berset et al., 1998; De Benedetti et al., 1991). The fact that tif4631 is synthetic lethal with ded1 mutant alleles (Table 5.4; de la Cruz et al., 1997) and that ded1 mutants are resistant to rapamycin thus raises a possibility that the rapamycin-induced degradation of eIF4G might be inhibited in strains carrying ded1 mutant alleles.

5.5 PROSPECTUS

Despite much effort over the years, the precise role of Ded1p remains enigmatic, validating, unfortunately, a pronouncement that “Nevertheless, it is clear that Ded1p is a very mysterious protein involved in various pathways” by

(Hayashi et al., 1996). Observations reported here provide additional links of

Ded1p’s function to RTG and TOR signaling pathways. Several future directions merit consideration to gain a deeper understanding of Ded1p’s cellular role. First, to investigate whether Rtg proteins indeed participate in translation, polysome profiles in the rtg strains should be investigated as a starting point. Second,

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because Ded1p apparently has a role in several biological processes, such as translation, splicing, and Pol III transcription, an open-ended genetic search of

Rtg1p-, Rtg2p- or Rtg3p-interacting proteins is likely to be fruitful. Third, it would be useful to examine the stability of eIF4G in ded1 mutant background under rapamycin treatment to probe the possibility that altered Ded1p helps to stabilize eIF4G upon rapamycin challenge. Finally, since the nuclear translocation of

Rtg1p/Rtg3p complex can be induced by rapamycin, it is also important to study the nuclear interaction of Ded1p and Rtg1p/Rtg3p complex in the presence of rapamycin.

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