STRUCTURAL AND FUNCTIONAL

ANALYSIS OF CRITICAL PROTEINS

INVOLVED IN mRNA DECAY

CHENG ZHIHONG

NATIONAL UNIVERSITY OF SINGAPORE

2006

STRUCTURAL AND FUNCTIONAL

ANALYSIS OF CRITICAL PROTEINS

INVOLVED IN mRNA DECAY

CHENG ZHIHONG

(B.Sc)

Ease China University of Science and Technology

A THESIS SUBMITTED

FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

DEPARTMENT OF BIOLOGICAL SCIENCES

NATIONAL UNIVERSITY OF SINGAPORE

2006

To My Wife

i

Index

Index………………………………………………………………………………………i Acknowledgements …………………………………………………………….………..ii Table of Contents………………………………………………………………………..iii Abstract ……………………………………………………………………………...... viii Lists of Figures……………………………………………………………………...…....x Lists of Tables ……………………………………………………………………….…xii Lists of Abbreviations ………………………………………………………………...xiii

References…………………………………………………………………………...…147 Appendix I MOPS Minimal Medium ……………………………………………..168 Appendix II Publication list…………………………………………………………170

ii

Acknowledgements

I would like to thank sincerely my supervisor, Dr Song Haiwei, for offering me an

opportunity being a postgraduate. Without his great guidance, patience and advice, I

could not finish my Ph.D program successfully.

Many thanks also go to our collaborator from Howard Hughes Medical Institute,

The University of Arizona, Professor Roy Parker, Drs. Jeff Coller and Denise Muhlrad,

for their great experiment data and precious discussions.

I would like to give my appreciation to the members of my thesis committee:

Associate Professor Kunchithapadam Swaminathan, Associate Professor Wang Yue from

Institute of Molecular and Cell (IMCB) and Assistant Professor J Sivaraman from

Department of Biological Sciences (NUS), for guidance and discussions throughout my

studies.

I also thank all my colleagues in my laboratory for their kind help. Thanks

especially go to Dr. Kong Chunguang, Dr. Wu Mousheng, Miss She Meipei, Miss Chen

Nan and Dr Zhou Zhihong for all the valuable discussions in all the experiments. I want

to thank Dr. Christian, Dr. Rohini, Miss Portia and Sharon, for their critical reading of the

manuscript of this thesis, and Lim Mengkiat for his great help in the experiment.

I also thank all administrative staffs in Department of Biological Sciences (NUS) and IMCB for their supports. Thanks also go to the shared facilities in IMCB including

DNA sequencing facility and Mass Spectrometry facility for experimental supports.

Finally, I would like to thank my family, especially my wife for her full support, patience, encouragement and inspiration all the time. I could not image the situation without her precious support.

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Table of contents

Chapter 1 Introduction

1.1. Biological significance of mRNA decay………………………………………..1

1.2. General mRNA decay pathway…………………………………………………1

1.2.1. Deadenylation……………………………………………………………..4

1.2.2. Decapping………………………………………………………………....6

1.2.2.1. The Dcp1-Dcp2 Decapping enzyme complex………………………6

1.2.2.2. Regulation of the decapping activity……………………………...... 7

1.2.2.3. Scavenger decapping enzyme DcpS………………………………...9

1.2.3. Enzymes involved in mRNA body degradation…………………………10

1.2.3.1. 5’ to 3’ degradation by Xrn1……………………………………….10

1.2.3.2. 3’ to 5’ degradation by the exosome complex…………………...... 11

1.3. mRNA quality control mechanisms targeting aberrant mRNAs………………13

1.3.1. Nonsense-mediated mRNA decay…………………………………….....13

1.3.1.1. NMD factors……………………………………………………….15

1.3.1.2. Translation and NMD……………………………………………...16

1.3.1.3. Definition of premature termination codons……………………….18

1.3.1.4. Recognition of PTC in mammals………………………………...... 20

1.3.2. Nonstop mRNA decay …………………………………………………..22

1.4. Specialized mRNA decay pathways…………………………………………...23

1.5. Project I: Structural and functional studies of Ski8…………………………...27

1.5.1. Brief introduction of Ski8………………………………………………..27

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1.5.2. Structural characteristics of WD-repeat proteins………………………...29

1.5.3. Aims of this project………………………………………………………33

1.6. Project II: Structural and functional studies of Dhh1…………………………34

1.6.1. Brief review of Dhh1…………………………………………………….34

1.6.2. Structural characterization of Superfamily 2 helicases……………...... 35

1.6.3. Aims of this project………………………………………………………40

1.7. Project III: Structural and functional analysis of hUpf1……………………….41

1.7.1. Previous functional and biochemical studies of Upf1………………...... 41

1.7.2. Structural studies of Superfamily 1 helicases……………………………43

1.7.3. Aims of this project………………………………………………………44

Chapter 2 Cloning, Protein Purification, Crystallization and Structure Determination 2.1. Gene cloning and protein expression strain construction……………………...45

2.1.1. Yeast genomic DNA isolation…………………………………………...45

2.1.2. Polymerase chain reaction (PCR) ……………………………………….45

2.1.3. Agarose gel electrophoresis……………………………………………...46

2.1.4. Purification of PCR products…………………………………………….46

2.1.5. Enzyme digestion, dephosphorylation and purification……………...... 46

2.1.6. Ligation and transformation…………………………………………...... 47

2.1.7. Plasmid preparation and positive clone screening……………………….47

2.1.8. DNA sequencing………………………………………………………....47

2.1.9. E. coli expression strain transfomation ………………………………….48

2.1.10. Protein expression test and expression strain storage……………………48

2.1.11. SDS-PAGE………………………………………………………………48

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2.2. Protein purification………………………………………………………….....49

2.2.1. Large-scale cell culture for protein expression…………………………..49

2.2.2. Purification procedures………………………………………………...... 49

2.2.2.1. Cell lysis…………………………………………………………...51

2.2.2.2. First GST affinity column chromatography……………………….51

2.2.2.3. Second GST affinity column chromatography…………………….51

2.2.2.4. Ion exchange chromatography…………………………………...... 51

2.2.2.5. Gel filtration chromatography…………………………………...... 52

2.3. Crystallization……………………………………………………………….....53

2.4. Structure determination……………………………………………………...... 55

2.4.1. Heavy atom derivative preparation…………………………………...... 55

2.4.2. Data collection and processing…………………………………………..56

2.4.2.1. Data collection and processing of SeMet Ski8…………………….56

2.4.2.2. Data collection and processing of the Br derivative of Dhh1……..57

2.4.2.3. Data collection and processing of SeMet hUpf1………………...... 58

2.4.3. Structure determination…………………………………………………..59

2.4.3.1. Phasing, modeling and refinement of Ski8……………………...... 59

2.4.3.2. Phasing, modeling and refinement of Dhh1……………………….62

2.4.3.3. Structure determination of hUpf1………………………………….65

2.5. Biochemical and molecular biological experiments………………………...... 72

2.5.1. Experiments for Ski8………………………………………………….....72

2.5.1.1. Site-directed mutagenesis and yeast two-hybrid assay……………72

2.5.1.2. GST pull-down assay……………………………………………...72

vi

2.5.2. Experiments for Dhh1……………………………………………………73

2.5.2.1. CD-spectroscopy…………………………………………………...73

2.5.2.2. Mutagenesis and in vivo mRNA turnover assays………………….73

2.5.2.3. Limited Proteolysis………………………………………………...74

2.5.2.4. In vitro RNA binding assay………………………………………..74

2.5.3. Experiments for hUpf1…………………………………………………..75

2.5.3.1. Mutagenesis and In vitro ATPase activity…………………………75

2.5.3.2 In vitro ATP binding assay………………………………………...75

2.5.3.3. In vitro RNA binding assay………………………………………..76

2.5.3.4. In vivo NMD analysis and P-body formation…………………...... 76

2.5.3.5. Surface plasmon resonance (SPR) ………………………………...77

Chapter 3 Crystal Structure and Mutagenesis Studies of Ski8

3.1. Results…………………………………………………………………………78

3.1.1. Overall structure determination………………………………………….78

3.1.2. Comparison with other WD repeat proteins……………………………..80

3.1.3. Location of protein-protein interaction sites on the β propeller…………83

3.1.4. Mutational Analysis of Ski8……………………………………………..87

3.2. Discussion…………………………………………………………………..….90

Chapter 4 Structural and Functional Analysis of Dhh1

4.1. Results ………………………………………………………………………....94

4.1.1. Structural overview of the Dhh1…………………………………………94

4.1.2. Structural Comparison ………………………………………………...... 96

4.1.3. Location of the conserved sequence motifs……………………………...98

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4.1.4. Interactions of the conserved Motifs……………………………………103

4.1.5. Identification of residues required for RNA binding …………………..107

4.1.6. Conformational changes in Dhh1………………………………………114

4.2. Discussion…………..………………………………………………………..116

Chapter 5 Structural and Functional Insights into hUpf1

5.1. Results………………………………………………………………………..120

5.1.1. Overall structure ………..………………………………………………120

5.1.2. Nucleotide Binding site and ATP hydrolysis…………………………...124

5.1.3. Conformational change during the Upf1 ATPase cycle………………..128

5.1.4. Allosteric effect of ATP binding coupled with RNA binding .………...133

5.1.5. Differential effects of Upf1 mutants on P-body formation……………..141

5.2. Discussion………………………………………………………………….....143

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Abstract

The control of mRNA translation and degradation is important for gene

expression in eukaryotic cells. mRNA decay, including mRNA quality control, is a multi-

step processing event composed of deadenylation, decapping and degradation of the

mRNA body. Three proteins, hUpf1, Dhh1 and Ski8 involved in eukaryotic mRNA decay

were structurally and functionally studied in this thesis.

Ski8 is a WD-repeat protein with an essential role for the Ski complex assembly

in an exosome-dependent 3'-to-5' mRNA decay. Additionally, Ski8 is involved in meiotic

recombination by interacting with Spo11. Crystal structure of Ski8 from Saccharomyces

cerevisiae was determined at 2.2Å resolution. It reveals that Ski8 folds into a seven- bladed beta propeller. Mapping sequence conservation and hydrophobicities of amino

acids on the molecular surface of Ski8 reveals a prominent site on the top surface of the

beta propeller. It was proposed that this top surface mediates interactions of Ski8 with

Ski3 and Spo11, which was confirmed by mutagenesis combined with yeast two-hybrid

and GST pull-down assays. The functional implications for Ski8 function in both mRNA

decay and meiotic recombination was also discussed.

Dhh1, a DEAD-box protein, functions both to repress translation and enhance

decapping. The crystal structure of the N- and C-terminal truncated Dhh1 from budding

yeast was determined at 2.1Å resolution. The structure reveals that truncated Dhh1 is

composed of two RecA-like domains with a unique arrangement. In contrast to the

structures of eIF4A and mjDEAD, in which no motif interactions exist, motif V in Dhh1

interacts with motif I and the Q-motif, thereby linking the two domains together.

ix

Electrostatic potential mapping combined with mutagenesis reveals that motifs I, V, and

VI are involved in RNA binding. In addition, trypsin digestion of the truncated Dhh1 in

the absence or presence of RNA or ligand suggests that ATP binding enhances an RNA-

induced conformational change. Interestingly, some mutations located in the conserved

motifs and at the interface between the two RecA-like domains confer dominant negative

phenotypes in vivo and disrupt the conformational switch in vitro, suggesting that this

conformational change is required for Dhh1 function.

Upf1 is a critical protein involved in triggering nonsense-mediated mRNA decay,

an mRNA surveillance pathway that recognizes and degrades aberrant mRNAs

containing premature stop codons. Upf1 belongs to the helicase Superfamily 1 (SF1), and

is thought to utilize the energy of ATP hydrolysis to promote transitions in the structure of RNA or RNA-protein complexes. The crystal structure of the catalytic core of human

Upf1 determined in three states (phosphate-, AMPPNP- and ADP-bound forms) reveals an overall structure composed of two RecA-like domains with two additional domains protruding from the N-terminal RecA-like domain. Structural comparison combined with mutagenesis studies identified a likely ssRNA binding channel, and a cycle of

conformational change coupled to ATP binding and hydrolysis. These conformational

changes alter the likely ssRNA-binding channel in a manner that can explain how ATP

binding destabilizes ssRNA binding to Upf1.

Keywords: mRNA decay, nonsense-mediated mRNA decay, WD-repeat protein, Ski8,

DEAD-box protein, Dhh1, RNA helicase, Upf1, X-ray crystallography

x

Lists of Figures

Figure 1-1 General mRNA decay pathway…………………………………………...3 Figure 1-2 General models for nonsense-mediated mRNA decay (NMD) in S. cerevisiae and mammalian cells……………………………………14 Figure 1-3 A proposed model for mammalian NMD ……………………………….21 Figure 1-4 The general model for nonstop-mediated mRNA decay (NSD) in S. cerevisiae…………………………………………………………...23 Figure 1-5 Architecture of the β-propeller fold…………………………………...... 30 Figure 1-6 A diagrammatic representation of one blade of the WD-repeat protein…………………………………………………………………....31 Figure 1-7 Architecture of WD-repeat protein in protein complex……………...... 33 Figure 1-8 Crystal structures of Superfamily 2 helicases………………………...... 40 Figure 1-9 Crystal structures of Superfamily 1 helicases………………………...... 44 Figure 2-1 Purification of Ski8……………………………………………………....52 Figure 2-2 Purification of Dhh1……………………………………………………..53 Figure 2-3 Purification of hUpf1………………………………………………….....53 Figure 2-4 Crystals of Ski8, Dhh1 and hUpf1, as well as hUpf1 in complex with AMPPNP, ATPγS and ADP………………………………………..55 Figure 2-5 A partial electron density map of Ski8 shown in the program O………..60 Figure 2-6 Flow chart of structure determination of Ski8…………………………...61 Figure 2-7 A partial electron density map of Dhh1 shown in the program O……….63 Figure 2-8 Flow chart of structure determination of Dhh1…………………………..64 Figure 2-9 Flow chart of structure determination of hUpf1……………………...... 67 Figure 2-10 Evaluation of 1000 trials by SnB program………………………………68 Figure 2-11 A partial electron density map of hUpf1-AMPPNP shown in the program ………………………………………………………………….69 Figure 3-1 Overall structure of Ski8…………………………………………………79 Figure 3-2 Sequence alignment of Ski8 homologs………………………………...... 82 Figure 3-3 Comparison of Ski8p with Gβ and Tup1c……………………………….83 Figure 3-4 Molecular surface views of Ski8…………………………………………85

xi

Figure 3-5 Location of the protein-protein interactions site on the top face of the β propeller…………………………………………………………86 Figure 3-6 Mutational analysis of Ski8 mutants……………………………………..89 Figure 4-1 Orthogonal view of tDhh1…………………………………………….....96 Figure 4-2 Sequence alignment of S. cerevisiae Dhh1, H. sapiens Rck/p54, S. cerevisiae eIF4A and M. janaschii mjDEAD…………………………99 Figure 4-3 Comparison of tDhh1 with eIF4A and mjDEAD………………………102 Figure 4-4 Stereo view of interactions of the conserved motifs……………………106 Figure 4-5 CD spectroscopy of various single mutants, double mutants and wild-type Dhh1…………………………………………………………108 Figure 4-6 In vitro RNA binding assay of Dhh1…………………………………...109 Figure 4-7 In vivo mutagenesis analysis of Dhh1………………………………...... 113 Figure 4-8 Analysis of limited trypsin digestion of Dhh1………………………….115 Figure 4-9 Comparison of Dhh1 to eIF-4A………………………………………...119 Figure 5-1 Domain structure of hUpf1 and sequence alignment of the helicase core domain……………………………………………………122 Figure 5-2 Structure of hUpf1hd and its comparison with PcrA and Vasa helicases………………………………………………………………...123 Figure 5-3 Nucleotide binding and hydrolysis of hUpf1hd……………………...... 127 Figure 5-4 Conformational changes of hUpf1hd upon nucleotide binding and hydrolysis…………………………………………………………..132 Figure 5-5 Protein gel filtration profile of hUpf1 WT and mutants 1B∆ and 1C∆………………………………………………………………....133 Figure 5-6 The channel between domains 1B and 1C involved in ssRNA binding………………………………………………………………….135 Figure 5-7 Allosteric effects of ATP binding/hydrolysis on RNA binding………..137 Figure 5-8 Surface plasma resonance analysis of RNA binding to hUpf1…………139 Figure 5-9 Structure comparison of hUpf1hd-AMPPNP with hUpf1hd-ATPγS………………………………………………………...141 Figure 5-10 Visualization of Dcp2-GFP in live yeast strains expressing Upf1 mutants…………………………………………………………....143

xii

Lists of Tables

Table 1-1 Critical proteins involved in general mRNA decay pathway……………..3 Table 1-2 Regulators of mRNA decapping………………………………………...... 9 Table 1-3 Components of the exosome……………………………………………..12 Table 1-4 NMD factors in eukaryotic cells…………………………………………16 Table 1-5 Biochemical properties of the conserved motifs of RNA helicases……..36 Table 1-6 Crystal structures of helicases in the Protein Data Bank………………...37 Table 1-7 Summaries of biochemical properties of Upf1 mutants……………...... 43 Table 2-1 Purification procedures of Ski8, Dhh1 and hUpf1……………………….50 Table 2-2 Recipes of buffers used for purification………………………………….50 Table 2-3 Crystallization conditions and cryo-protectants used for Ski8 and Dhh1………………………………………………………...….54 Table 2-4 Crystallization conditions and cryo-protectants used for hUpf1………...54 Table 2-5 Statistics of the data collection and processing of Ski8………………….57 Table 2-6 Statistics of the data collection and processing of Dhh1………………...58 Table 2-7 Statistics of the data collection and processing of hUpf1………………..59 Table 2-8 Se atom sites of Ski8 found by the program SOLVE………………...... 60 Table 2-9 Phase determination and refinement statistics of Ski8 structure………...62 Table 2-10 Br atom sites found by the program SOLVE…………………………….63 Table 2-11 Phase determination and refinement statistics of Dhh1 structure………..65 Table 2-12 Se atom sites found by the program SnB……………………………...... 66 Table 2-13 Result of rotation function by Molrep using domain 1A as a searching model………………………………………………………..70 Table 2-14 Result of rotation function by Molrep using domain 2A as a searching model and fixing two copies of domain 1A…………………70 Table 2-15 Phase determination and refinement statistics of hUpf1…………………71

xiii

Lists of Abbreviations

AMPPNP Adenosine 5′-(β,γ-imido)triphosphate tetralithium salt hydrate

ADP adenosine diphosphate

AMD ARE-mediated mRNA decay

ARE AU-rich element

ATP adenosine triphosphate

ATPγS adenosine 5 -[γ-thio]triphosphate

ATR ATM and Rad3-related

Arf ADP-ribosylation factor

CCP4 collaborative computational project No.4

CNS crystallography and NMR system

DSE Downstream sequence elements

DTT dithiothreitol

EDTA ethylenediaminetetraacetic acid

EJC Exon-junction complex

Gal4-BD Gal4 DNA binding domain

Gal4-AD Gal4 DNA activating domain

GST Glutathione-S-transferase

GTPase GTP hydrolase

IPTG isopropyl β-D-thiogalactopyranoside kD kilo Dalton

M molar

xiv

MAD Multiple-wavelength anomalous dispersion ml milliliter mM millimolar mg milligram

NCS non-crystallographic symmetry

NGD No-go mRNA decay

NSD Non-stop mRNA decay nm nanometer

NMD Nonsense-mediated decay

OD optical density

PAGE poly-acrylamide gel electrophoresis

PCR polymerase chain reaction

PEG polyethylene glycol

PI3K phosphoinositide 3-kinase

PMSF phenyl-methyl-sulfonyl fluoride

PTC Premature termination codon

RCC1 regulator of chromosome condensation 1

RISC RNA-induced silencing complex

SAD Single-wavelength anomalous dispersion

SDS sodium dodecyl sulfate siRNA Small interference RNAs

SLBP Stem-loop binding protein

SMD Staufen1-mediated decay

xv

SURF SMG1, Upf1, release factors eRF1 and eRF3

Tris Tris(hydroxymethyl)aminomethane

WT wide type

µl microliter

µM micromolar

UV ultraviolet

1

Chapter 1

Introduction

1.1 Biological significance of mRNA decay

During the lifetime of a cell, eukaryotic mRNAs are bound by a large number of

proteins, which change in coordination with various processing events. This requires

gene expression to be tightly regulated and permits a cell to alter its pattern of protein

synthesis in response to changing physiological conditions. Eukaryotic gene expression

can be controlled at several different levels including transcription, splicing and export,

translation, post-translational modification and protein localization and degradation. In

addition to these processing events, mRNA turnover becomes a critical control point for

regulating gene expression. mRNA decay targets not only endogenous mRNA molecules,

but also viral double-stranded RNAs (dsRNAs) for antiviral defense in a specialized

pathway termed RNA interference (RNAi) (van Hoof et al., 2002; Waterhouse et al.,

2001). Moreover, eukaryotic cells have evolved specialized mRNA decay mechanisms to

recognize and degrade the aberrant mRNAs generated during transcription due to

frequent mutations or faulty splicing, thereby increasing the quality control of mRNA

biogenesis and protein synthesis (Maquat et al., 2001).

1.2 General mRNA decay pathway

Various mRNAs in eukaryotes have different half-lives. For instance, in yeast the most unstable RNAs have a half-life of about 2-3 minutes while stable RNAs can survive 2

more than 90 minutes. In higher eukaryotes unstable mRNAs have a half-life of 15

minutes while stable mRNAs may exist for more than 24 hours (Herrick et al., 1990;

Wang et al., 2002; Shyu et al., 1989). Two general mRNA decay pathways exist in

eukaryotes and both are deadenylation-dependent and initiated by the removal of the

poly(A) tail at the 3’ end (Figure 1-1; Meyer et al., 2004; Coller et al., 2004; Parker et

al., 2004). Subsequently, deadenylated mRNAs are degraded by cleavage of the 5’-cap

structure by the Dcp1-Dcp2 decapping complex (Decapping proteins) , followed by 5’ to

3’ degradation by the Xrn1 exonuclease (Exoribonuclease 1; Decker et al., 1993; Steiger et al., 2003; Larimer et al., 1992). Alternatively, poly(A)-shortened mRNAs can be degraded in a 3’ to 5’ direction by the exosome, a large protein complex consisting of 10 exonucleases with the aid of Ski7 and the Ski2/3/8 complex (Jacobs et al., 1998;

Mukherjee et al., 2002). The residue 5’-cap structure is then cleaved by a scavenger enzyme DcpS (van Dijk et al., 2003).

In addition to deadenylation-dependent 5’ to 3’ and 3’ to 5’ degradation, some eukaryotic mRNAs are degraded by endonucleolytic cleavage without deadenylation

(Figure 1-1). Examples include Transferrin receptor, Vitellogenin and Xenopus β-globin mRNAs (Bremer et al., 2003; Binder et al., 1994; Cunningham et al., 2000), which have a wide variety of endonucleolytic cleavage sites. The general pathways of eukaryotic mRNA decay are illustrated in Figure 1-1 and critical enzymes and regulators are summarized in Table 1-1.

3

Processing step Enzymes Regulators Yeast Mammals Deadenylation Pan2-Pan3 Pan2-Pan3 PABPC CCR4-NOT CCR4-NOT complex complex Cap PARN Decapping Dcp1-Dcp2 Dcp1-Dcp2 Edc1, Edc2,Edc3 Pat1,Lsm1-7, Dhh1, PABPC Cap hydrolysis Dcs1 DcpS 5’-3’ degradation Xrn1 Xrn1 3’-5’ degradation Exosome Exosome Ski2/3/8 complex, Ski7 Table 1-1 Critical proteins involved in general mRNA decay pathway

Figure 1-1 General mRNA decay pathway. The first step in decay is shortening of the poly(A) tail, which can be catalyzed by several different enzymes. Following deadenylation, the body of the mRNA is attacked from either the 5’ or 3’ ends. The 5’ to 3’ decay pathway is initiated by cleavage of the cap structure by the decapping complex Dcp1-Dcp2, followed by 5’→3’ exonucleolytic degradation by Xrn1. 3’ to 5’ decay is catalyzed by a large complex of exonucleases termed the exosome, leading to production of 5’ cap structure, which can be broken down by the scavenger decapping protein DcpS. Decay of some mRNAs are initiated by endonucleolytic digestion.

4

1.2.1 Deadenylation

Deadenylation is the first and rate-limiting step in the degradation of regular mRNA from yeast to higher eukaryotes. To date, three different proteins or protein complexes have been identified as mRNA deadenylases involved in mRNA degradation

(Parker et al., 2004).

The Ccr4-Not complex is the predominant enzyme controlling the mRNA poly(A) tail length in Saccharomyces cerevisiae. In addition to the two nucleases, Ccr4 and Pop2, this complex also contains several accessory proteins, Not1 to Not5, Caf4, Caf16, Caf40 and Caf130 (Tucker et al., 2001; Denis et al, 2003; Martine A. Collart 2003). The most important nuclease in the Ccr4-Not complex is Ccr4 (carbon catabolite repressor 4 factor), which was first identified as a gene expression regulator (Denis et al., 1984), then later found to be involved in mRNA deadenylation (Tucker et al., 2001). Sequence analysis shows that Ccr4 belongs to the ExoIII family of nucleases (Dlakic M. 2000). It has been shown that the activity of Ccr4 is inhibited by the poly(A)-binding protein

(Pab1), but not affected by the cap structure of the mRNA, suggesting its preference for mRNA substrates with a shorter poly(A) tail (Tucker et al., 2001; Viswanathan et al.,

2003). The second protein in the Ccr4-Not complex with deadenylase activity is

Pop2/Caf1 (PGK-promoter directed overproduction/Ccr4-associated factor 1; Daugeron et al., 2001). Pop2 belongs to the DEDD nuclease superfamily composed of RNases and

DNases (Daugeron et al., 2001; Zuo et al., 2001). A recently reported crystal structure of the RNase D domain from yeast Pop2 shows that the nuclease domain adopts a similar fold to DNA exonucleases. These enzymes are proposed to coordinate two divalent metal ions in the active site to catalyze hydrolysis of the phosphodiester bond (Thore et al., 5

2003; Joyce et al., 1995). The situation of two adenylases existing in the Ccr4-Not

complex involved in mRNA deadenylation remains puzzling. It could be possible that

Pop2 facilitates poly(A) targeting to the Ccr4-Not complex.

Another protein complex involved in cytoplasmic mRNA deadenylation is the

Pan2-Pan3 complex (Pab1-stimulated poly(A) ribonuclease), which encodes the

predominant alternative deadenylase (Boeck et al., 1996; Brown et al., 1996; Yamashita

et al., 2005; Tucker et al., 2001). Sequence analysis shows that Pan2 also belongs to the

RNase D superfamily and it is proposed to use a similar hydrolysis mechanism to Pop2

(Moser et al., 1997). The activity of the Pan2-Pan3 complex is also involved in trimming

mRNAs that initially have longer poly(A) tails to the message-specific length of 60 to 80

nucleotides (Brown et al., 1998). Recently, Yamashita et al. (2005) found that biphasic

deadenylation exists in mammalian mRNA decay, which requires the deadenylases Pan2-

Pan3 and the Ccr4-Not complex, but not PARN (see below). The Pan2-Pan3 complex

carries out the first phase of deadenylation to shorten the poly(A) tails to ~110A

nucleotides, followed by the second phase of deadenylation by the Ccr4-Not complex

(Yamashita et al., 2005).

Mammals have a third deadenlase, the poly(A)-specific ribonuclease (PARN),

which is the predominant deadenylase in these organisms (Astrom et al., 1992; Korner et

al., 1997). Sequence analysis shows that PARN also belongs to the RNase D superfamily along with Pop2 and Pan2 (Moser et al., 1997). In addition to the nuclease domain,

PARN also contains an R3H domain, which binds to single-stranded RNA (ssRNA). A

recently determined crystal structure of human PARN both with and without RNA shows

that PARN functions as a dimer with a similar catalytic site to those of Pop2 and ε186 6

(Wu et al., 2005). Unlike Ccr4 and Pop2, the deadenylase activity of PARN is inhibited by Pab1 and stimulated by a 5’ cap structure, suggesting that mRNAs with both an exposed cap and poly(A) tail are the preferred substrates for PARN (Korner et al., 1997;

Dehlin et al., 2000).

1.2.2 Decapping

There are two distinct decapping events required for mRNA decay pathway: one is initiated at the beginning of mRNA degradation in a deadenylation-dependent regular mRNA decay pathway by the decapping enzymes Dcp1-Dcp2; the other targets the remaining 5’-cap structure by DcpS after the degradation of bulk mRNA by the exosome.

1.2.2.1 The Dcp1-Dcp2 Decapping Enzyme Complex

In the regular mRNA decay pathway, decapping is triggered by the shortening of the poly(A) tail by the deadenylases Ccr4-Pop2, Pan2-Pan3 or PARN. Dcp1 and Dcp2 form a decapping holoenzyme, in which Dcp2 is the catalytic subunit and Dcp1 enhances the decapping activity (Dunckley et al., 1999; Steiger et al., 2003; van Dijk et al., 2002;

Wang et al., 2002). Dcp2 contains a Nudix domain, which is found in many proteins that cleave nucleotide diphosphates (Bessman et al., 1996). In addition to the Nudix domain,

Dcp2 also contains two highly conserved regions, termed Box A, which confers the cleavage specificity, and Box B, which has been implicated in RNA binding (Piccirillo et al., 2003). Dcp2 requires divalent cations for activity and prefers capped RNA more than

20 bases in length (Steiger et al., 2003; Stevens, 1988; van Dijk et al., 2002). The recently solved crystal structure of the N-terminal domain of Schizosaccharomyces pombe Dcp2 reveals a two-domain architecture with a helical N-terminal domain and a typical Nudix- 7

fold C-terminal domain (She et al., 2006). The authors identified a conserved surface on the N-terminal domain that mediates the Dcp1-Dcp2 interaction and is essential for decapping in vivo.

Dcp1 is conserved in eukaryotes and is required for decapping in vivo (Beelman et al., 1996; Hatfield et al., 1996). In vitro assays have shown that Dcp1 can stimulate the activity of Dcp2 (Steiger et al., 2003). The crystal structure of yeast Dcp1 shows that it belongs to a novel class of EVH1 domains (She et al., 2004). Two conserved sites have been identified, namely patch 1, which is thought to bind to proline-rich sequences, is proposed to be involved in binding to decapping regulatory proteins, and patch 2, which is required for the function of the holoenzyme. Recent results by Fenger-Grφn et al.

(2006) show that hDcp1a and hDcp2 are components of a larger decapping complex, which contains some decapping regulators, such as Hedls, Edc3 and Rck/p54, suggesting that decapping is more complicated in higher eukaryotes and subject to control by additional regulators.

1.2.2.2 Regulation of the decapping activity

Due to the importance of the decapping event in mRNA decay, decapping activity is regulated by numerous factors (Table 1-2; Coller et al., 2004; Meyer et al., 2004).

Negative regulators include the poly(A)-binding protein 1 (Pab1) and the cap-binding

protein (eIF4E). Pab1 couples decapping with deadenylation and inhibits decapping by

promoting the formation of the translation initiation complex (Caponigro et al., 1995;

Morrissey et al., 1999). In vivo and in vitro assays have shown that eIF4E can inhibit

decapping activity by competitively binding to the 5’-cap structure (Schwartz et al., 1999;

Schwartz et al., 2003). 8

It has been shown that several decapping regulators can promote decapping efficiency. The RNA-binding proteins Edc1, Edc2 and Edc3, can enhance decapping activity. Edc1 and Edc2 are capable of stimulating decapping activity in vitro, although mutations of these two genes show little phenotypic effect in vivo (Dunckley et al., 2001;

Schwartz et al., 2003; Steiger et al., 2003). Edc3 can specifically promote the decapping activity of the RPS28B mRNA by interacting with the RPS28 protein and the decapping complex (Badis et al., 2004).

Mutation in pat1 gene shows a decapping defect in vivo, but not in vitro, suggesting that Pat1 could be a decapping regulator (Bonnerot et al., 2000; Bouveret et al., 2000). It has been shown that Pat1 also functions in translation repression (Coller et al., 2005). Pat1 has been shown to associate with the cytoplasmic Lsm complex. There are two types of Lsm complexes in the cell: a nuclear Lsm complex (Lsm2-8), which associates with the U6 spliceosomal snRNA, and a cytoplasmic Lsm complex (Lsm1-7), which functions in mRNA decay (Boeck et al., 1998; Bouveret et al., 2000; He et al.,

2000; Tharun et al., 2000). The Lsm-Pat1 complex either promotes rearrangement of the mRNP structure or facilitates the recruitment of the decapping complex (He et al., 2000).

The DEAD-box protein Dhh1, the focus of Project II (see section 1.6), has been identified to be involved in mRNA decapping (Coller et al., 2001; Fischer et al., 2002).

Dhh1, like Pat1, may promote the transition from the translating state to the translation repression state and stimulate decapping (Coller et al., 2005).

Interestingly, all of the regulating factors required for regular mRNA decapping are dispensable for decapping in Nonsense-mediated mRNA decay (NMD; see section 9

1.3.1). The NMD factors may help the decapping enzymes circumvent the requirement

for additional factors to stimulate decapping activity.

Regulators Properties Functions Interactions Pab1 RRM domain and Blocks mRNA decapping eIF4G, eRF3, a proline-rich C Stimulates translation Pan2-pan3 terminus eIF4E Cap-binding Translational initiation Lsm7, eIF4G, protein complex, blocks mRNA eIF4A, eIF4B, decapping Pab1 Edc1, Small basic Stimulate mRNA decapping Dcp1, Dcp2, Edc2, proteins

Edc3 Contains five Edc3 regulates decapping of Dcp1, Dcp2, conserved domains RPS28B mRNA Lsm1-7, Ccr4- Pof2, Dhh1, Rps28B, Lsm8 Pat1 No recognizable Stimulates mRNA decapping Dcp1, Dcp2, motifs and formation of P-bodies in Lsm1-7, Dhh1, vivo Xrn1 Lsm1-7 Sm-like proteins Stimulate mRNA decapping Dcp1, Dcp2, Dhh1, Pat1, Xrn1, Upf1 Dhh1 DEAD-box proteinStimulates mRNA decapping Dcp1, Dcp2, and associates with Lsm1-7, Ccr4, deadenylases Pop2, pab1, Edc3 Table 1-2 Regulators of mRNA decapping (Coller et al., 2004).

1.2.2.3 Scavenger decapping enzyme DcpS

The second decapping activity occurs at a later phase of mRNA decay. After

mRNA is degraded by the exosome in the 3’ to 5’ direction, the remaining 5’-cap

structure is cleaved by the DcpS protein via a mechanism that is different from that of

Dcp2 (Cougot et al., 2004). DcpS belongs to the histidine triad (HIT) family, which is involved in the cleavage of nucleotide pyrophosphate bonds (Brenner et al., 2002). In 10 contrast to Dcp2, which has efficient decapping activity with long RNA, DcpS prefers free m7GpppG or a Cap structure with fewer than ten nucleotides. This is explained by the recently-solved crystal structure of human DcpS (Liu et al., 2002; Gu et al., 2004).

The structure shows that the DcpS dimer performs decapping in a closed cavity. This cavity is located between the two HIT domains and a swinging ‘cap’ composed of nuclear transport factor 2 (NTF2)-like domains and will only accommodate a short capped RNA (Gu et al., 2004). In additon to the decapping activity in the 3’ to 5’ degradation pathway, recent analyses show that hDcpS also cleaves m7GDP, the product of the decapping enzyme Dcp2, to m7GMP and a phosphate moiety (Wang et al., 2001; van Dijk et al., 2003). The mechanism of m7GDP cleavage has been characterized in the crystal sructures of apo hDcpS or of hDcpS in complex with m7GDP (Chen et al., 2005).

Since m7GMP is the final product of decapping, it suggests that the cell may monitor mRNA decay by detecting the level of m7GMP or other specific by-products.

1.2.3 Enzymes involved in mRNA body degradation

1.2.3.1 5’ to 3’ degradation by Xrn1

Xrn1, a divalent cation-dependent exonuclease, is responsible for rapid mRNA degradation from the 5’ to 3’ direction after the cap structure is removed by the Dcp1-

Dcp2 complex. The Xrn1 enzyme is highly conserved from yeast to higher eukaryotes

(Bashkirov et al., 1997; Till et al., 1998; Newbury et al., 2004). Mutation of Xrn1 in yeast leads to the accumulation of full-length mRNAs without a 5’ cap structure (Hsu et al.,

1993; Muhlrad et al., 1994). Activity assays have shown that Xrn1 prefers a 5’- monophosphated RNA rather than RNA with a 5’-hydroxyl terminus and that Xrn1 has 11

no activity on capped RNA (Stevens, 1980). In addition to functioning in regular mRNA

decay, NMD (see section 1.3.1) and degradation of specific mRNAs (see section 1.4),

Xrn1 has been reported to be involved in suppressing viral RNA recombination by rapidly removing 5'-truncated RNAs (Cheng et al., 2006).

1.2.3.2 3’ to 5’ degradation by the exosome complex

Two versions of the exosome complex exist in the cell: one in the nucleus and one in the cytoplasm. The functions for the nuclear exosome have been found in many aspects of RNA processing, such as 5.8 S rRNA synthesis, rRNA trimming, snRNA and

snoRNA processing, and degradation of incorrectly-processed mRNAs or pre-mRNAs

(van Hoof et al., 2000; Butler, 2002). The cytoplasmic exosome plays a critical role in 3’

to 5’ degradation of mRNA in various decay pathways, such as the regular mRNA decay,

NMD, NSD (see section 1.3.2) etc. (Anderson et al., 1998; Chen et al., 2001; van Hoof et

al., 2002; Takahashi et al., 2003; Gatfield et al., 2004).

The core exosome contains six exonucleases (Rrp41, Rrp42, Rrp43, Rrp45, Rrp46

and Mtr3), three hydrolytic RNases (Rrp4, Rrp40 and Rrp44) and one RNA-binding

protein (Csl4), all of which are well-conserved from yeast to mammals (Table 1-3, van

Hoof et al., 1999; Mitchell et al., 2000). In the nucleus, there are two proteins associated

with the core exosome, the hydrolytic exonuclease Rrp6 and the DEAD-box RNA

helicase Mtr4. In the cytoplasm, several co-factors have been identified, namely Ski7,

Ski2, Ski3 and Ski8, whose mutations result in the death of yeast cell caused by

overexpression of a toxin produced by the L-A double-stranded RNA virus (Araki et al.,

2001; Maquat 2002; Takahashi et al., 2003). In the wild-type yeast cells, these genes are

required for degradation of the toxin-encoding transcripts from the double-stranded RNA 12

virus. The Ski8 protein is the topic of Project I in this thesis (see section 1.5). Together,

these co-factors stimulate mRNA degradation by recruiting the exosome complex and/or

disrupting the specific secondary structure of RNA substrates.

Subunit Similarity In vitro activity Core subunits Rrp4 S1 RNA BP 3’ exo hydrolase Rrp40 S1 RNA BP Rrp41 RNase PH 3’ exo phosphorolase Rrp42 RNase PH Rrp43 RNase PH Rrp44 RNase II 3’ exo hydrolase Rrp45 RNase PH Rrp46 RNase PH Mtr3 RNase PH Csl4 S1 RNA BP Nuclear subunits Rrp6 RNase D 3’ exo hydrolase Mtr4 RNA helicase Co-factors Ski2 RNA helicase Ski3 TPR domains Ski complex Ski8 WD domains Ski7 G protein Table 1-3. Components of the exosome. Subunits highlighted in grey form the ring structure of the core exosome. The other three core subunits associate tightly with this ring. The co-factors Ski2, Ski3 and Ski8 form the Ski complex, which associates with the exosome.

Recently, two groups have reported the structures of the archaeal exosome: one

from Sulfolobus solfataricus (Lorentzen et al., 2005a and 2005b) and the other from

Archaeoglobus fulgidus (Büttner et al., 2005). The structure of the S. solfataricus

exosome consists of two subunits Rrp41 and Rrp42, which form a trimer of Rrp41-Rrp42 heterodimers in a hexameric ring. Additionally, Büttner et al. (2005) have determined two nine-subunit exosome isoforms from A. fulgidus, consisting of a hexameric ring of 13

RNase PH subunits with three phosphorolytic active sites and an RNA entry pore

composed of S1-domain subunits.

1.3 mRNA quality control mechanisms targeting aberrant mRNAs

Eukaryotic cells have evolved numerous elaborate mRNA quality-control

mechanisms to recognize and degrade mRNAs that have not been properly spliced or that

contain deleterious mutations, ensuring that error-free mRNAs are used for protein

synthesis. There are two surveillance pathways in cells. Nonsense-mediated decay

(NMD) targets mRNAs containing premature stop codons, while nonstop-mediated

mRNA decay (NSD) degrades those mRNAs without translation termination codons

(Conti et al., 2005; Maquat, 2005; Behm-Ansmant et al., 2006).

1.3.1 Nonsense-mediated mRNA decay

Nonsense-mediated mRNA decay is an mRNA surveillance pathway that is highly conserved from yeast to higher eukaryotes. NMD has been recently implicated in regulating the mRNA level of wild-type transcripts (Hillman et al., 2004). Initiation of

NMD in yeast and mammals requires the “pioneer round” of translation. Downstream sequence elements (DSE) in yeast or the exon-exon junction complex (EJC) in C. elegans

and mammalian cells, as well as NMD factors are essential for the recognition of the

premature termination codon (PTC; Gonzalez et al., 2001; Lynne Maquat, 2004). After

PTC recognition, nonsense-containing mRNAs are degraded faster than regular

transcripts by either deadenylation-independent or accelerated deadenylation pathways

(Figure 1-2A; Muhlrad et al., 1994; Mitchell et al., 2003; Lejeune et al., 2003; Couttet et 14

al., 2004). Enzymes involved in regular mRNA decapping, 5’ to 3’ or 3’ to 5’ mRNA

body degradation are also required for NMD.

Surprisingly, degradation of PTC-containing mRNAs in Drosophila melanogaster

is initiated with endonucleolytic cleavage in the vicinity of the PTC (Gatfield et al., 2003;

Gatfield et al., 2004), although the identity of the endonuclease involved in this process remains unclear (Figure 1-2B). A

B

Figure 1-2 General models for nonsense-mediated mRNA decay (NMD) in S. cerevisiae and mammalian cells (A), and in D. melanogaster (B). mRNA decay requires a “pioneer round” of translation. The premature-termination codon (PTC) is recognized by a translating ribosome and surveillance complex Upf1-3. After PTC recognition, mRNA undergoes 5’ to 3’ or 3’ to 5’ decay in yeast and mammals (A), while mRNA decay initiates with endonucleolytic cleavage in D. melanogaster (B). 15

1.3.1.1 NMD factors

The key molecules involved in the NMD pathway were initially identified by

genetic screens in S. cerevisiae (three genes: Upf1-3) , C. elegans (seven genes: SMG1-

7), and then in H. sapiens (seven genes: hSMG-1, hUpf1, -2, -3, hSMG5-7) (Table 1-4;

Cui et al., 1995; Hodgkin et al., 1989; leeds et al., 1991; Perlick et al., 1996; Mendell et al., 2000; Serin et al., 2001; Chiu et al., 2003; Cali et al., 1998; Cali et al., 1999; Denning et al., 2001; Anders et al., 2003;). These factors are highly conserved from yeast to mammals (Maquat, 2004). Recruitment of one of these three Upf proteins to a site more than fifty nucleotides (nt) downstream of a termination codon triggers mRNA degradation by the NMD pathway (Lykke-Andersen et al., 2000). The Upf1, Upf2 and

Upf3 proteins (known as SMG-2, SMG-3 and SMG-4 in C. elegans) are core components of the surveillance complex (He et al., 1997). Upf3 is a shuttling protein with two paralogs in human cells (Upf3 and Upf3X, or Upf3a and Upf3b) and is also one of the components of the exon-exon junction complex (EJC; Le Hir et al., 2000; Kim et al.,

2001). Upf2 is perinuclear and might attach to exporting mRNAs by interacting with

Upf3 (Lykke-Andersen et al., 2000). The crystal structure of the interacting domains of hUpf2 and hUpf3b shows that the RNA-binding domain (RBD) of Upf3 is involved in the interaction with one MIF4G domain of Upf2, rather than RNA binding (Kadlec et al.,

2004).

Of these three Upf proteins, Upf1, a cytoplasmic ATP-dependent RNA helicase, is the central player of the surveillance complex in NMD (Czaplinski et al., 1995; Weng et al., 1996a, 1996b, 1998). Upf1 is the topic of Project III of this thesis (see secion 1.7).

Briefly, Upf1 forms a surveillance complex with Upf2 and Upf3 and associates with the 16 translation release factors eRF1/eRF3, providing a link between the surveillance complex and the translation machinery (He et al., 1997; Czaplinski et al., 1998). The function of

Upf1 in NMD is regulated through phosphorylation/dephosphorylation by SMG-1 and

SMG5/6/7 in higher eukaryotes (Pages et al., 1999; Pal et al., 2001; Cali et al., 1999;

Denning et al., 2001; Yamashita et al., 2001; Grimson et al., 2004; Anders et al., 2003).

The crystal structure of the N-terminal domain of SMG7 reveals that this protein contains a 14-3-3-like phosphoserine-binding domain, which is involved in the association with phosphorylated Upf1 (Fukuhara et al., 2005). Conservation of this 14-3-3-like domain among SMG5, SMG6 and SMG7 suggests that these proteins act as similar adaptors in mediating dephosphorylation of Upf1 in NMD.

Name Function Yeast Mammal Worm Fly Upf1p Upf1 SMG-2 Upf1-PA RNA-dependent ATPase and 5’ to 3’ helicase Upf2p Upf2 SMG-3 Upf2-PA Bridging Upf1 and Upf3 Upf3p Upf3 and SMG-4 Upf3-PB and mRNA binding, associates with the Upf3X Upf3-PC EJC complex Smg1 SMG-1 Smg1-PA Phosphatidyl kinase-related kinase Smg5 SMG-5 Smg5-PA Phosphoprotein phosphatase/PP2A complex, TPR/PIN domain Smg6 SMG-6 Smg6-PA TPR/PIN domains Smg7 SMG-7 Phosphoprotein phosphatase/PP2A complex, TPR/PIN domain, Upf1 targeting to mRNA decay bodies Table 1-4 NMD factors in eukaryotic cells.

1.3.1.2 Translation and NMD

In yeast and higher eukaryotes, it is well known that initiation of the NMD pathway depends on the recognition of a premature stop codon by the translation machinery. Several lines of evidence support the fact that NMD requires translation. For 17

example, NMD is inhibited by blocking translation by adding antibiotics or by mutating

components of the translation machinery (Qian et al., 1993; Menon et al., 1994; Carter et

al., 1995; Herrick et al., 1990; Peltz et al., 1992; Zhang et al., 1997; Welch and Jacobson,

1999; Zuk and Jacobson, 1998). Also, NMD can be prevented by suppressor transfer

RNAs, which guide the incorporation of an amino acid at premature stop codons instead of initiating NMD (Belgrader et al., 1993; Li et al., 1997; Losson et al., 1979; gozalbo et al., 1990). NMD can also be inhibited by a specific secondary structure in the 5’ untranslated region that directly or indirectly via a bound protein, impedes the scanning of 40S ribosomal subunits from binding the initiation codon (Belgrader et al., 1993;

Thermann et al., 1998). It has been shown that NMD is inhibited when components of the translation machinery, such as eukaryotic initiation factors eIF4GI and eIF4GII and poly(A)-binding protein, are inactivated and/or cleaved by the polio virus (Gradi et al.,

1998; Kuyumcu-Martinez et al., 2002).

The most direct evidence showing the relationship between NMD and translation is the finding that the essential factors of NMD interact with two release factors eRF1 and eRF3 that mediate translation termination (Czaplinski et al., 1998; Wang et al., 2001). eRF1 is a multi-functional release factor that detects all three stop codons and induces the release of the nascent peptide synthesized by the ribosome (Frolova et al., 1994). eRF1 provides a dramatic example of molecular mimicry. The three-dimensional structure of eRF1 resembles to the shape and charge distribution of a tRNA molecule. This mimicry is thought to allow the release factor eRF1 to enter the A-site on the ribosome and trigger translation termination (Song et al., 2000). eRF3 has been found to recycle eRF1 in a similar way as its prokaryotic counterpart RF3 (Frolova et al., 1996; Zavialov et al., 18

2001). Sequence analysis indicates that eRF3 contains at least two functional regions: an

N-terminal region, which is not necessary for translation termination but is involved in binding to poly(A)-binding protein, suggesting a link between the termination event and

the initiation process in protein biosynthesis, and a conserved C-terminal eEF1α-like region, which is essential for translation termination and viability (Kisselev et al., 2000;

Zhouravleva et al., 1995; Ter-Avanesyan et al., 1993; Hoshino et al., 1999; Inge-

Vechtomov et al., 2003; Uchida et al., 2002). The crystal structure of the eEF1α-like region of eRF3 from S. pombe reveals an overall structure similar to EF-Tu, but with different domain arrangements (Kong et al., 2004). Based on structural analysis and mutagenesis studies, the eRF1 binding region has been identified. In the free form of eRF3, the eRF1 binding site is occupied by an N-terminal extension, which is rich in acidic amino acids. Therefore it is thought that eRF1 competes with the N-terminal

extension to bind to eRF3. In addition to its interaction with eRF1, eRF3 also interacts

via its eEF1α-like region with the components of the NMD surveillance complex to

initiate mRNA decay (Czaplinski et al., 1999; Wang et al., 2001).

1.3.1.3 Definition of premature termination codon

Although NMD has been found to be conserved in all the eukaryotic organisms

that have been studied so far (Wagner et al., 2002; Hentze et al., 1999), different species

have developed distinct mechanisms by which premature termination codons (PTCs) are

recognized (Lejeune et al., 2005; Conti et al., 2005). In S. cerevisiae, some mRNAs

contain loosely defined downstream sequence elements (DSEs), which associate with

their binding partner Hrp1, an hnRNA-like factor that functions in discriminating PTCs

from normal stop codons (Ruiz-Echevarria et al., 1998; Gonzalez et al., 2000; Zhang et 19 al., 1995). Due to the fact that DSEs are not highly conserved among yeast transcripts, an alternative mechanism has been proposed, whereby the 3’ UTR combined with a specific set of proteins may confer the appropriate context that is required for recognition of normal stop codons, and this is thought to fail in the case of NMD (Hilleren et al., 1999;

Amrani et al., 2004). Surprisingly, unlike in mammals, homologs of human EJC components in Drosophila are not involved in PTC recognition (see below), because

PTC-containing transcripts from intron-free genes are degraded by NMD in yeast and

Drosophila (Gatfield et al., 2003).

In mammals, the NMD pathway is dependent on pre-mRNA splicing, because transcripts from intron-free genes are immune to NMD (Maquat et al., 2001; Brocke et al., 2002). The exon-junction complex (EJC) is deposited 20-24 nucleotides upstream of exon-exon junctions after RNA splicing (Maquat L.E. 2004). Experiments using reconstituted NMD in which components of the EJC are tethered downstream of a nonsense codon show that the prerequisite of pre-mRNA splicing for NMD is based on the positioning of the EJC (Bono et al., 2004; Lykke-Andersen et al., 2001; Palacios et al., 2004; Shibuya et al., 2004). Additionally, the relative position of the EJC to the PTC is important for PTC recognition: a stop codon either less than 50-55 nucleotides upstream of the 3’-most exon-exon junction or downstream of this junction are not recognized to initiate NMD. It has been proposed that when the ribosome stalls at the

PTC, it does not displace the EJC which is located more than 55 nucleotides downstream, by which the NMD is triggered.

The EJC is a highly dynamic protein complex (Tange et al., 2004). The minimal core of EJC is a stable ternary complex containing Y14, Mago, eIF4AIII and Btz (Jurica 20

et al., 2003; reichert et al., 2002; Palacios et al., 2004; Shibuya et al., 2004; Tange et al.,

2005; Ballut et al., 2005). Y14 and Mago form a heterodimer, which is also a component

of the spliceosome. Crystal structures of Y14-Mago show that the RBD domain of Y14 mediates the interaction with Mago (Bono et al., 2004; Fribourg et al., 2003; Shi et al.,

2003). This complex adopts a similar structure to the Upf2:Upf3 complex (Fribourg et al.,

2003), suggesting that Y14 does not bind directly to spliced mRNA. eIF4AIII, a member of the eIF4A family of RNA helicases, has been identified as an EJC-anchoring factor.

This finding is supported by both in vivo and in vitro data (Shibuya et al., 2004;

Ferraiuolo et al., 2004; Palacios et al., 2004; Chan et al., 2004). In addition, the EJC

complex also contains the splicing co-activators SRm160 and RNPS1, the splicing factor

Pinin and the export factors UAP56, REF/Aly and TAP/NFX1:p15 (Tange et al., 2004;

Lejeune et al., 2005). It has been shown that Upf3, a component of the surveillance

complex, also associates with the EJC, suggesting a functional link between the

surveillance complex and the EJC (Kim et al., 2001; Gehring et al., 2003).

1.3.1.4 Recognition of PTC in mammals

As mentioned above, NMD initiation involves at least three sets of protein

complexes: peptide release factors, the surveillance complex and the EJC, which together

form an elaborate molecular interacting network that bind sequentially to initiate NMD.

Based on previous studies and a recent report by Kashima et al. (2006), a stepwise assembly model for NMD has been proposed (Figure 1-3).

During pre-mRNA splicing, EJCs are deposited at every exon-exon junction on the mRNA. Upf3 then associates with the EJC. After nuclear export of the mRNA, the perinuclear-localized Upf2 binds to the complex via its interaction with Upf3. During the 21

“pioneer round” of translation, the ribosome stalls at the PTC, and then the release factors

eRF1 and eRF3 bind to it. Unphosphorylated Upf1 and SMG1 then transiently join this

complex forming the SURF complex (SMG1, Upf1, release factors eRF1 and eRF3).

Interaction between Upf1 and Upf2 induces the formation of a second transient complex formed between SURF and the EJC, named the decay-inducing complex (DECID).

DECID stimulates Upf1 phosphorylation by SMG1, resulting in the release of eRF1 and

eRF3 from this complex. Decapping or accelerated deadenylation are then triggered by

an unknown mechanism (Figure 1-3). Phosphorylated Upf1 is recycled by SMG5/6/7

with the aid of PP2A (Behm-Ansmant et al., 2006).

Figure 1-3 A proposed model for mammalian NMD. Translation termination at the premature stop codon (PTC) leads to the assembly of the SURF complex, which consists of SMG1, Upf1, eRF1 22

and eRF3. The SURF complex interacts with Upf2 and Upf3 in concert with the EJC complex resulting in the formation of the DECID complex, which triggers Upf1 phosphorylation and the dissociation of eRF1 and eRF3. Phosphorylated Upf1 triggers the 5’ or 3’ decay events by an unknown mechanism. SMG7 and PP2A are involved in the dephosphorylation of Upf1 recycling these proteins for a new round of NMD.

1.3.2 Nonstop mRNA decay

In addition to mRNAs containing a premature stop codon, there are also mRNAs

that lack a termination codon, such as those with a 3’ truncation. In eubacteria, translation

of these aberrant mRNAs results in ribosomes stalling at the 3’ end. The ribosomes are

recycled by a tmRNA, a molecule with properties of both transfer and mRNA (Keiler et

al., 1996; himeno et al., 1997). The tmRNA-guided trans-translation results in protein products being tagged by a specific peptide, which is recognized and degraded by

specific proteases. The ribosome-free truncated mRNAs are then degraded by 3’ to 5’

exonucleases (Yamamoto et al., 2003). In eukaryotic cells, nonstop mRNA decay has

evolved to cope with ribosomes stalling at the 3’ end of aberrant mRNAs (Frischmeyer et

al., 2002; van Hoof et al., 2002; Inada et al., 2005). It has been proposed that Ski7

recognizes the stalled ribosome at the 3’ end of a nonstop mRNA by its GTPase domain

and recruits the exosome complex with the aid of the Ski complex via its amino-terminal

domain (van Hoof et al., 2002). Nonstop mRNAs can also be decapped and degraded 5’

to 3’ by Xrn1 in the absence of Ski7 (Inada et al., 2005). A proposed model for the

nonstop mRNA decay pathway is shown in Figure 1-4. 23

Figure 1-4 The general model for nonstop-mediated mRNA decay (NSD) in S. cerevisiae. During the pioneer round of translation, the ribosome stalls at the poly(A) tail. This displaces Pab1 and stimulates translation repression. The stalled ribosome on the mRNA may be recognized by Ski7, which in turn recruits the exosome along with the Ski complex for 3’ to 5’ degradation. Alternatively, the displacement of Pab1 from the 3’ end of the mRNA results in accelerated decapping and 5’ to 3’ decay.

1.4 Specialized mRNA decay pathways

ARE-mediated mRNA decay:

The degradation rates of specific mRNAs are regulated by sequence elements

located throughout the transcript. The AU-rich element (ARE), which is in the 3’ UTR of

transcripts, including those from proto-oncogenes, interleukins and cytokines (Chen et al., 1995; Shim et al., 2002), is the most studied sequence element in yeast and mammals

(Vasudevan et al., 2001). Several RNA-binding proteins that interact with ARE have been identified, including the destabilizing factors AUF1, TTP and KSRP and the stabilizing factors HuR and NF90 (Wagner et al., 1998; Blackshear et al., 2002; Gherzi et al., 2004; Brennan et al., 2001; Shim et al., 2002). It has been shown that ARE-mediated 24

mRNA decay is achieved by independently activating several distinct mRNA decay

steps, including deadenylation, decapping, 3’ to 5’ degradation by the exosome and 5’ to

3’ degradation by Xrn1 (Chen et al., 2001; Lykke-Andersen et al., 2005; Lai et al., 2003;

Gao et al., 2001; Mukher-jee et al., 2002; Stoecklin et al., 2006; Wang et al., 2001).

Staufen1-mediated mRNA decay:

Recently, Kim et al. (2005) identified a new mRNA decay mechanism, so called

Staufen1-mediated mRNA decay (SMD), because it requires Staufen1, as well as Upf1

and a termination codon. Mammalian staufen1 is an RNA-binding protein that binds to

extensive RNA secondary structures by a dsRNA-binding domain (Marion et al., 1999;

Wickham et al., 1999). The function of Staufen1 is well-studied in Drosophila, where it

is involved in the specific transport, localization and translational control of bicoid and

oskar mRNAs (Broadus et al., 1998; Matsuzaki et al., 1998; Shen et al., 1998). In

humans, SMD requires only one component of the surveillance complex, human Upf1

(hUpf1), because SMD is not affected by downregulating Upf2 or Upf3. Mammalian

Staufen1 can bind Upf1 and the 3’ UTR of Arf1 mRNA (ADP-ribosylation factor),

thereby reducing the Arf1 mRNA abundance independently of the EJC. A model has

been proposed for EJC-independent SMD, whereby Staufen1, bound downstream of the

normal termination codon, mimics the role of the EJC to elicit mRNA decay.

Histone mRNA decay:

In addition to its roles in NMD and SMD, Upf1 is also required for the

degradation of histone mRNAs (Kaygun et al., 2005). It has been shown that the degradation of histone mRNAs, the only non-adenylated mRNAs in metazoans, is 25

replication-dependent (Dominski et al., 199). The 3’ end of histone mRNAs contain a

conserved stem-loop structure, which interacts with stem-loop binding protein (SLBP).

These cis and trans elements have been shown to be required for the degradation of histone mRNAs in a DNA replication-dependent manner, but the underlying mechanism remains unclear. Kaygun et al. (2005) have shown that Upf1, the central player of the surveillance complex, and ATR, a critical regulator of the DNA-damage-checkpoint pathway, are required for the degradation of histone mRNAs. A proposed model suggests that inhibition of DNA synthesis triggers the ATR pathway, which leads to phosphorylation of Upf1 and/or SLBP at their S/T-Q motifs. Phosphorylation by the

ATR pathway might result in tight binding of Upf1 to SLBP, allowing Upf1 to be efficiently recruited to the 3’ end of the histone mRNA and to activate the subsequent mRNA degradation pathway.

Autoregulation of RPS28B mRNA decay:

Autoregulation of RPS28B mRNA by Edc3 in yeast has been identified by Badis el al. (2004) as another case of deadenylation-independent decay. It has been shown that

Edc3 is an enhancer for decapping of both unstable and stable mRNAs in yeast

(Kshirsagar et al., 2004). edc3 deletion shows a synergistic effect on the decapping defects of dcp1 or dcp2 temperature sensitive (ts) mutants (kshirsagar et al., 2004). Edc3 also associates with some decapping regulators, such as Dhh1, Lsm1 and Xrn1 in decay processing bodies (P-bodies), suggesting its role in decapping regulation. Badis et al.

(2004) compared transcript profiles of wild-type and edc3∆ cells using microarrays, and identified a single mRNA, RPS28B mRNA encoding a ribosomal protein, as the target of 26

Edc3 function. Rps28 interacts with a conserved hairpin structure in the 3’ UTR of its own mRNA and Edc3 binding to Rps28 is critical for autoregulation. A model for the

Edc3-mediated autoregulation of the RPS28B mRNA has been proposed, whereby excess

Rps28 binds to the conserved hairpin in the 3’ UTR of RPS28B mRNA, which results in deadenylation-independent decapping and subsequent degradation of RPS28B mRNA.

No-go mRNA decay:

Very recently, Doma et al. (2006) identified a new decay mechanism, referred to as “no-go decay” (NGD), whereby translation elongation is impeded by specific secondary structures in mRNA transcripts. NGD is mediated by two conserved proteins:

Dom34 and Hbs1, which are similar to the translation-termination factors eRF1 and eRF3 respectively (Davis et al., 1998; Inagaki et al., 2003). The “no-go decay” recruits an unknown endonuclease to initiate mRNA degradation, completed by the 5’ to 3’ exonuclease Xrn1p and the 3’ to 5’ exosome. A model for the initiation of NGD has been proposed, whereby Dom34 recognizes stalled ribosomes and Hbs1 mimics eRF3 to release the stalled ribosomes and trigger endonucleolytic cleavage of the mRNA

(Clement et al., 2006).

27

1.5 Project I: Structural and functional studies of Ski8

1.5.1 Brief introduction of Ski8

The function of the exosome in cytoplasmic mRNA degradation requires several

cofactors including the putative GTPase Ski7, and the Ski complex consisting of Ski2,

Ski3 and Ski8 (Araki et al. 2001; Maquat 2002; Takahashi et al. 2003). These superkiller

(SKI) genes were initially identified from mutations that cause overexpression of a killer

toxin encoded by the latent double-stranded RNA virus (Toh et al. 1978). Subsequent

work demonstrated that the products of the ski2, ski3 and ski8 genes are necessary for 3’ to 5’ mRNA degradation and for repression of translation from nonpolyadenylated RNA

(Jacobs Anderson and Parker 1998; Araki et al. 2001; Masison et al. 1995). Ski2 and Ski3 are a putative RNA helicase and a tetratricopeptide repeat (TPR) protein, respectively, while Ski8 contains WD-repeats (Rhee et al. 1989; Matsumoto et al. 1993; Widner and

Wickner 1993). The three Ski proteins form a stable complex with 1:1:1 stoichiometry and the complex is localized to the cytoplasm (Brown et al. 2000). The Ski complex has

been suggested to be an mRNA decay specific cofactor for the exosome because

mutations in the Ski genes inhibit 3’ to 5’ mRNA decay, but have no effect on the

functions of exosome in nuclear RNA processing (Brown et al. 2000; Jacobs Anderson

and Parker 1998; van Hoof et al. 2000). However, the nature of the interactions between the subunits of the Ski complex and with the exosome, and the specific role of the Ski

complex in 3’ mRNA decay remain elusive. It has been shown that different regions of

the N-terminal domain of Ski7 interact with the exosome and the Ski complex, thereby

recruiting both complexes to the 3’ end of mRNA (Araki et al. 2001). 28

In addition to the role of Ski8 in mRNA decay, yeast strains lacking ski8 have

reduced meiotic recombination, but have no detectable effects on mitotic recombination

or DNA repair (Malone et al. 1991; Gardiner et al. 1997; Pecina et al. 2002; Evans et al.

1997; Fox and Smith 1998). Moreover, this meiotic role of Ski8 appears to be conserved

(Arora et al. 2004; Tesse et al. 2003; Fox and Smith 1998; Evans et al. 1997). However,

the meiotic role of Ski8 is unclear and because of its direct role in mRNA decay, it could

be an indirect effect of defects in 3’ to 5’ mRNA decay.

Several observations indicate that Ski8 has a distinct role in meiosis that is separate from its function in mRNA decay. Firstly, although ski8∆ yeast strains show defects in meiosis, ski2∆ or ski3∆ strains, which equally affect mRNA decay, do not affect meiotic recombination (Arora et al. 2004). Secondly, Ski8 binds tightly to Spo11,

which is homologous to the archaeal topoisomerases. Spo11 acts in concert with at least

nine other proteins (including Ski8) to create DNA double-strand breaks (DSBs), whose

repair leads to meiotic recombination (Arora et al. 2004; Tesse et al. 2003). Thirdly, the

level of ski8 mRNA increases about fifteen fold upon entry to meiosis (Gardiner et al.

1997), and the Ski8 protein redistributes from the cytoplasm to the nucleus and localizes to chromosomes specifically during meiosis (Arora et al. 2004). These results indicate that Ski8 plays distinct roles in meiotic recombination and mRNA decay by changing its localization and interacting with different protein partners via as yet undetermined interactions.

29

1.5.2 Structural characteristics of WD-repeat proteins

Possible insight into how Ski8 interacts with other proteins comes from analysis of the Ski8 sequence. The Ski8 protein contains multiple repeats of the “WD” motif, originally identified in β-transducin (Matsumoto et al. 1993; Evans et al. 1997). These motifs are found in many proteins with diverse functions and are thought to mediate protein-protein interactions (Smith et al. 1999). WD-repeat proteins belong to a large family of β-propeller proteins, which have diverse sequences and functions. The fold of

β-propeller proteins adopts a highly symmetrical structure with a 4-8 repeats of a four- stranded antiparallel β-sheet motif (Figure 1-5). The twisted blades are packed around a central tunnel and the entire structure is maintained predominantly by hydrophobic interactions between the blades. The last blade contains four antiparallel β strands from both termini of the propeller domain, which seems to be required to stabilize the circular structure. 30

Figure 1-5 Architecture of the β-propeller fold. (A) The four-bladed haemopexin domain, pdb entry code 1hxn. (B) The five-bladed Tachylectin, pdb entry code 1tl2. (C) The six-bladed Sialidase, pdb entry code 2sil. (D) The seven-bladed Transducin G β subunit, pdb entry code 1got. (E) The eight-bladed Nitrite reductase, pdb entry code 1aof.

In addition to the WD motif, β-propeller proteins also contain many conserved sequence motifs, such as the kelch motif, tryptophan docking motif, aspartate box, tolB repeat and YWTD repeat. It is of interest to note that no WD-repeat motif has been identified in archaebacterial sequences, suggesting that the WD motif may be specific to eukaryotic β-propeller-containing proteins. For example, nearly half of the 212 β- 31

propeller proteins in yeast are likely to be WD-repeat proteins (Smith et al., 1999), which

function in signal transduction, cell division, cytoskeleton changes, chemotaxis and RNA

processing as adapters mediating protein-protein interactions.

To date, several crystal structures of WD-repeat proteins have been determined

including the G protein β subunit (Wall et al. 1995; Gaudet et al. 1996; Lambright et al.

1996; Sondek et al. 1996), the C-terminal β-propeller domain of Tup1 (Tup1c) and its

human homolog Groucho/TLE1 protein (hTle1-C) (Pickles et al. 2002; Sprague et al.

2000), the Aip1 protein involved in actin depolymerization (Voegtli et al. 2003), the human regulator of chromosome condensation (RCC1) and its complex with nuclear GTP binding protein Ran (Renault et al., 1998; Renault et al., 2001), the F box protein β-

TrCP1 in complex with Skp1 and β-catenin peptide (Wu et al., 2003), the p40 subunit of

Arp2/3 complex (Robinson et al., 2001). Interestingly, a comparison of the available crystal structures of WD-repeat proteins shows that each WD motif corresponds to the first three strands of one blade and the last strand of the previous blade (Figure 1-6).

Figure 1-6 A diagrammatic representation of one blade of the WD-repeat protein. The sequence repeat is in a different phase from the structural repeat (blade), whereby the first strand (designated D) of the sequence repeat forms the last strand of the previous propeller blade. The conserved residues in WD motif are shown.

32

The crystal structures of WD-repeat proteins show that these proteins adopt an

antiparallel β-propeller architecture and form a thick circular plate with three potential flat surface regions (top, bottom and side) for protein-protein interactions (Figure 1-7 A

and B), which have been observed in several complexes containing WD-repeat proteins

or domains (Figure 1-7). For example, in the crystal structure of the trimeric G protein,

the ‘top’ surface of G β-subunit mainly contacts the α-subunit, while the side region

shows a close interaction with the helix of the γ-subunit. The ‘bottom’ surface of the G β-

subunit is also involved in interaction with the γ-subunit (Figure 1-7C). In the structure

of human RCC1 with Ran, the ‘top’ surface of β-propeller protein RCC1 is absolutely

required for interaction with the Switch II region of the GTP-binding protein (Figure 1-

7D). In the structure of the β-TrCP1-Skp1-β-catenin complex, the majority of the β-

TrCP1 residues that contact β-catenin are located on the ‘top’ surface of the WD-repeat

domain (Figure 1-7E). This manner of protein-protein interaction has also been observed

in the crystal structure of the Arp2/3 complex (Figure 1-7F).

33

Figure 1-7 Architecture of a WD-repeat protein in a complex. WD repeat proteins are shown in cyan. (A) Top view of WD-repeat protein. (B) Side view of WD-repeat protein. (C) WD-repeat G β-subunit in the G trimeric complex, pdb entry code 1got. (D) β-propeller protein RCC1 complexed with Ran, pdb entry code 1i2m. (E) WD-repeat containing β-TrCP1 complexed with Skp1 and β-Catenin, pdb entry code 1p22. (F) ARPC1 p40 subunit in Arp2/3 complex, pdb entry code 1k8k.

1.5.3 Aims of this project

To gain insight into how the adapter protein Ski8 interacts with Ski2 and Ski3 as well as Spo11 by structural and mutagenesis studies of Ski8 from S. cerevisiae.

34

1.6 Project II: Structural and functional studies of Dhh1

1.6.1 Brief review of Dhh1

The critical step in the decapping of eukaryotic mRNAs is the exiting of

translation and the assembly of an mRNP state that targets the mRNA for decapping. It

has been shown that Dhh1, a DEAD-box protein of Superfamily 2 RNA helicases,

functions in moving mRNAs from translation to the non-translating pool of mRNAs, which are concentrated in P-bodies, the specific subcellular sites of mRNA decapping and degradation (Coller & Parker, 2005). Dhh1 is also known to increase the rate of

decapping of mRNAs (Bonnerot et al, 2000; Bouveret et al, 2000; Coller et al, 2001;

Fischer & Weis, 2002; Tharun et al, 2000). Thus, Dhh1 functions as both activators of decapping and repressors of translation, and are critical proteins for modulating the transition from translation to mRNA degradation.

The Dhh1 protein is very important for translation repression and mRNA decapping and has been implicated in a wide range of biological phenomena. For example, and consistent with the findings that Dhh1 functions as a translation repressor,

Dhh1 homologs in Drosophila (Me31b), C. elegans (cgh-1), and Xenopus (Xp54) are involved in the storage of maternal mRNAs in a translationally repressed state (Ladomery

et al, 1997; Nakamura et al, 2001; Navarro et al, 2001). In addition, the Dhh1 homolog in

Schizosaccharomyces pombe, Ste13, is required for meiosis (Maekawa et al, 1994).

Finally, the Dhh1 homolog in mammals (RCK/p54) has been implicated in certain tumor

types (Akao et al, 1995; Nakagawa et al, 1999). Interestingly, over-expression of

RCK/p54 or Xp54 can rescue the loss of Dhh1 in yeast, demonstrating that the 35 underlying biochemical function of Dhh1 is conserved (Tseng-Rogenski et al, 2003;

Westmoreland et al, 2003).

1.6.2 Structural characterization of Superfamily 2 RNA helicases

RNA helicases are enzymes that are thought to catalyze the unwinding of double- stranded nucleic acids using energy from the hydrolysis of dNTPs, generally ATP.

Because of the direction of double-stranded RNA-unwinding activity, helicases can be divided into two types: 5’ to 3’ helicases and 3’ to 5’ helicases. RNA helicases are critical for all living organisms from bacteria and viruses to human, because they are involved in all processes involving RNA, such as transcription, editing, splicing, ribosome biogenesis, RNA export, translation, RNA turnover, and organellar gene expression. A general classification of helicases into five superfamilies has been made by Gorbalenya and Koonin (1993) according to the occurrence and characteristics of conserved motifs

(designated I, Ia, Ib, II, III, IV, V, VI) in the protein sequence. The biochemical functions of these conserved motifs are summarized in Table 1-5. Among these families,

Superfamily 1 and 2 show a high sequence similarity in at least seven of their conserved motifs and a preference for functioning as monomers or dimers (Tuteja et al., 2004), whilst Superfamily 3, 4 and 5 helicases, which are found in bacteria and viruses, are mostly hexameric (Patel and Picha, 2000).

36

Domain Motif Biochemical function I P-loop, Walker A NTP-binding motif; binds to phosphates of NTP Ia Binds substrate through sugar-phosphate backbone Ib Substrate binding 1 II Walker B NTP-binding and hydrolysis motif; binds β and γ phosphate through Mg2+; mediates NTP hydrolysis through water molecule III Binds γ phosphate; couple NTP hydrolysis with unwinding activity IV Substrate binding 2 V Binds substrate through sugar-phosphate backbone VI Binds γ phosphate; couple NTP binding/hydrolysis with conformational change Table 1-5 Biochemical properties of the conserved motifs of RNA helicases.

The largest subgroup of Superfamily 2, referred to as the DExD/H helicase family, contains the DEAD-box proteins and the related DEAH, DExH and DExD families, sharing eight conserved motifs (Tanner et al., 2001). A unique motif Q has been identified recently in DEAD-box families (Cordin et al., 2004). Many of the DEAD-box proteins studied in vitro have been shown to be RNA-dependent ATPases, suggesting that these proteins function to rearrange RNA or RNP structure (Rocak & Linder, 2004).

Although most DEAD-box proteins are thought to possess ATP-dependent RNA helicase activity, it is difficult to demonstrate helicase activity for many of the DEAD-box proteins in vitro presumably due to stringent substrate specificities or missing cofactors/helper proteins. Many structures of helicases in various superfamilies have been determined (summarized in Table 1-6). The structures of Superfamily 2 will be reviewed in the following paragraphs, while the structures of Superfamily 1 will be reviewed in section 1.7.2.

37

Family Helicase PDB code Co-crystallization PcrA 1PJR SF1 1QHG,1QHH ADPNP, Mg2+ 2- 2PJR,3PJR ss/dsDNA,ADPNP,SO4 Rep 1UAA ssDNA eIF4A 1FUK,1FUU,1QDE,1QVA SF2 MjDEAD 1HV8 DEAD Rck/p54 1VEC -box UAP56 1T5I,1T6N Vasa 2DB3 AMPNP, Mg2+ HCV NS3 1CU1,1HEI,1JR6,1ONB,8OHM 1A1V DNA BRCH1 1T15 BRCA1 RecG 1GM5, 3-way DNA junction RecQ 1D8B,1OYW SF2 1OYY ATPγS

SecA 1M6N, ,1NKT,1NL3, ,1TF5 2+ 1M74,1TF2 ADP, Mg SV40 1N25 UvrB 1C4O,1D2M,1D9X 2+ 1D9Z ATP, Mg DnaB 1B79,1JWE(*) Rep40 1S9H 1U0J ADP

RepA 1G8Y, 1OLO

SO 3- 1NLF 4 SF3 Rho 1A62,1A63(*), 1A8V 1PV4 ssDNA ssRNA, ADPNP 1PVO RNA 2A8V T7 gene4 1CR0,1NUI,1Q57 1CR1,1CR2,1CR4 dTTP, dATP, dTDP ANPNP, Mg2+ 1E0J,1E0K Table 1-6 Structures of helicases in the Protein Data Bank. All structures were determined by X- ray crystallography except NMR structures indicated by asterisk.

The first crystal structure of a full length DEAD-box protein was eIF4A, that of which was solved by Caruthers et al. (2000), followed by the DEAD-box protein

MjDEAD (Story et al., 2001), and UAP56 (Shi et al., 2004; Zhao et al., 2004). All of the solved crystal structures of Superfamily 2, as well as Superfamily 1 (see below), contain two covalently linked RecA-fold core domains, spanning about 350-400 residues (Figure

1-8; Story et al., 1992). The N-terminal domain (named domain 1 in Superfamily 2 and 38

domain 1A in Superfamily 1) contains motifs Q (only applicable for DEAD-box protein),

I, Ia, Ib, II and III, whilst the other three motifs IV, V and VI are located on the C-

terminal domain (referred to as domain 2 in Superfamily 2 and domain 2A in

Superfamily 1) . It is interesting to note that the distance and orientation of the two core

domains in these crystal structures of Superfamily 2 helicases are quite different as shown in Figure 1-8, suggesting that the linker region is flexible and that the domains can move with respect to one another. This structural flexibility could be important for their

functions in RNA unwinding or RNP rearrangement. A structural comparison of various

Superfamily 2 helicases or of different conformations observed within a single protein in

the presence or absence of a bound nucleotide or oligonucleotide gives insight into the

domain rearrangement upon ATP binding/hydrolysis and substrate binding/unwinding

(Caruthers et al., 2002). Although no structural data on conformational changes within

the Superfamily 2 helicases are available to date, it has been proposed that

conformational changes do occur in Superfamily 2 helicases upon ATP

binding/hydrolysis and RNA unwinding (Tanner et al., 2003; Shi et al., 2004; Sengoku et

al., 2006). In addition to the conserved core domains, the variable flanking sequences in

the DEAD-box proteins or additional domains (such as domain 3 in HCV NS3, see

below) appear to provide an additional interaction platform for specific RNA substrates

or regulators.

The details of the interaction between RNA and Superfamily 2 RNA helicases

have been observed in two reported crystal structures: Hepatitis C virus NS3 helicase

domain complexed with single-stranded DNA (Kim et al., 1998) and Vasa in complex

with single-stranded RNA (ssRNA) and AMPPNP, a non-hydrolysable ATP analogue 39

(Sengoku et al., 2006). The crystal structure of the NS3 helicase domain with single-

stranded DNA shows that the protein contains two conserved RecA-folds, domains 1 and

2, as well as an additional domain 3 consisting of predominantly α helices (Kim et al.,

1998). The bound single-stranded DNA lies in a channel between domain 3 and domains

1 and 2, with its 5’ end residing at the interface of domains 2 and 3, and its 3’ end

positioned at the interface of domains 1 and 3 (Kim et al., 1998). In the crystal structure

of Vasa in complex with ssRNA and AMPPNP, the ATP analogue interacts with both the

two core domains, which may bring these two domains together. The closed

conformation forms a continuous RNA-binding surface, which coordinates a 10mer

poly(U) that is bent (Sengoku et al., 2006). It has been proposed that the bending of the

bound single-stranded RNA weakens the RNA duplex interactions and facilitates the

RNA helicase activity.

40

Domain 2 Domain 1

Figure 1-8 Crystal structures of Superfamily 2 helicases. (A) Common structural features of DEAD-box protein. Motifs Q, I, Ia, Ib, II, III, IV, V, VI and the ATP-binding pocket are labeled. Domain 1 is shown in pink and domain 2 in blue. (B) Yeast initiation factor 4a (eIF4A), pdb entry code 1fuu. (C) A DEAD-box protein from Methanococcus Jannaschii, pdb entry code 1hv8. (D) Human ATP-dependent splicing and export factor UAP56, pdb entry code 1xtj. (E) Hepatitis C virus NS3 RNA helicase domain, pdb entry code 1a1v. (F) Drosophila Vasa DEAD-box protein, pdb entry code 2db3.

1.6.3 Aims of this project

(1) Solve the crystal structure of Dhh1 using X-ray crystallography.

(2) Based on the structural analysis, mutagenesis studies combined with in

vitro and in vivo assay will be carried out to investigate the conformational

changes of Dhh1 upon ligand binding, as well as its biological implication in

mRNA decay. 41

1.7 Project III: Structural and functional analysis of hUpf1

1.7.1 Previous functional and biochemical studies of Upf1

It has been shown that Upf1 is associated with polysomes (Atkin et al., 1995;

Atkin et al., 1997; Pal et al., 2001), and forms a conserved surveillance complex with

Upf2 and Upf3 (He et al., 1997; Maderazo et al., 2000). Moreover, Mendell et al. (2002) demonstrated that hUpf1 plays separable roles in NMD and NAS (nonsense-mediated altered splicing). Recent studies show that hUpf1-mediated nonsense surveillance is a crucial post-transcriptional regulatory event that controls the expression of a broad range of physiological transcripts (Mendell et al., 2004). Based on these findings, nonsense surveillance can be regarded as a critical molecular system to manipulate individual gene activities and suppress genomic mutations (Alonso et al., 2005). Failure of the mRNA surveillance pathway will lead to the accumulation of nonsense mutations and/or some pivotal physiological transcripts that cause one-third of human genetic diseases and many types of cancer. Hence this pathway could be manipulated for potentially beneficial pharmacological effects (Frischmeyer et al., 1999; Wilusz et al., 2001; Holbrook et al.,

2004).

Upf1 has been reported to be associated with the Dcp1-Dcp2 decapping complex, suggesting a functional link between the surveillance complex and mRNA decapping

(Dunckley et al., 1999; Jens et al., 2002; Sheth et al., 2003). In addition to their role in the

NMD pathway, Upf proteins are also involved in translation termination by interacting with the translation release factors eRF1 and eRF3 (Leeds et al., 1992; Weng et al.,

1996a, 1996b; Czaplinski et al., 1998; Kobayashi et al., 2004). And the role of Upf1 in translation termination is modulated by ATP (Weng et al., 1998). hUpf1 has been shown 42

to be involved in SMD. In SMD, hUpf1, rather than hUpf2 or hUpf3, is recruited by

Staufen1 for degradation of some natural mRNA targets (Kim et al., 2005). hUpf1 is also required for the regulated degradation of histone mRNAs (Kaygun et al., 2005).

Sequence analysis indicates that Upf1 contains two conserved functional regions, an N-terminal Cys/His-rich domain required for binding to Upf2 and a C-terminal helicase domain (Applequist et al, 1997; Culbertson & Leeds, 2003), which belongs to the Superfamily 1 RNA helicases (see below), and contains all seven conserved motifs common to the Superfamily 1 and 2 helicases (Applequist et al, 1997). Purified Upf1 protein demonstrates a nucleic-acid-dependent ATPase activity and 5’ to 3’ DNA/RNA helicase activity (Czaplinski et al., 1995; Bhattacharya et al., 2000). The mutational analysis of its ATPase, RNA binding and RNA unwinding activities have been investigated. The results of mutagenesis studies are summarized in Table 1-7.

Interestingly, the RNA-binding activity of Upf1 is decreased by adding ATP or its analogue ATPγS.

N-terminal Cys/His-rich C-terminal helicase domain domain C72S C65S, H94R, K436Q DE572AA RR793AA TR800AA H110R C84S, H98R (Motif I) (Motif II) (Motif VI) (Motif VI) C148S C122S, C125S ATP binding ND ND ND - + - + ATPase activity + + + - - - - RNA binding + + + + + - - RNA unwinding + + - - - - - Table 1-7 Summaries of biochemical properties of Upf1 mutants. ND means not determined. “+” indicates binding, while “–” indicates no binding detected.

43

1.7.2 Structural studies of Superfamily 1 helicases

To date, only two crystal structures of Superfamily 1 DNA helicases have been

reported: DNA helicase PcrA and E. coli Rep protein in complex with DNA substrates

and nucleotide (Figure 1-9; Korolev et al, 1997; Velankar et al, 1999). These two DNA

helicases contain two RecA-like domains (domain 1A and 2A), as observed in crystal

structures of Superfamily 2 helicases. The structures show that they also contain two

additional domains (domain 1B and 2B) as insertions in domain 1A and 2A, respectively.

The crystal structure of Rep helicase complexed with single-stranded DNA gave

us a first indication of the interaction between helicase and DNA. It is intriguing to note

that two molecules with different conformations (referred to “open” and “closed”) sit

together and interact with one single-stranded 15mer poly(T) oligonucleotide. The fact

that the dimer exists in the crystal structure is consistent with experimental data showing

Rep dimerization upon DNA binding. The difference between these two molecules is the

rigid body rotation of the domain 2B by about 130°, suggesting a translocation

mechanism of Rep dimer along the single-stranded DNA. The crystal structure of PcrA in

complex with dsDNA containing a single-stranded tail has been solved in two states: the

3- substrate complex (AMPPNP-bound) and the product complex (PO4 -bound). The ligand

binds to the cleft between domain 1A and 2A by a similar manner to the Superfamily 2

helicases. The double-stranded part of the DNA substrate contacts mainly the additional

domains 1B and 2B, while the single-stranded part of the DNA substrate locates on the

interface between domain 1A and 2A (Figure 1-9). The binding manner of single-

stranded DNA to the core domains 1A and 2A is very similar to that observed in the

structures of NS3 and Vasa (Figure 1-8). 44

Figure 1-9 Crystal structures of Superfamily 1 helicases. The labeled colors for domain 1A, 2A, 1B and 2B are pink, blue, green and yellow, respectively. The backbone of single-stranded or dsDNA is shown in orange. (A) Crystal structure of PcrA helicase bound to dsDNA and ADPNP, pdb entry code 3pjr. (B) Two copies (“open” and “closed”) of E.coli Rep helicase bound to single- stranded DNA, pdb entry code 1uaa.

1.7.3 Aims of this project

(1) Due to the critical role of the Upf1 protein in translation termination and

NMD pathway, the crystal structure of Upf1 in complex with ATP analog or ADP

will be determined by X-ray crystallography.

(2) The structural information gained will be used to generate mutants for in

vitro biochemical studies and in vivo functional studies to investigate the

biological function of Upf1. 45

Chapter 2

Cloning, Protein Purification, Crystallization and Structure

Determination

2.1 Gene cloning and protein expression strain construction

2.1.1 Yeast genomic DNA isolation

Yeast genomic DNA was used as the template to clone the genes ski8 and dhh1.

The yeast genomic DNA was prepared according to the yeast Smash & Grab DNA

miniprep protocol provided by the Fangman/Brewer Lab (modified from M.D. Rose, F.

Winston, and P. Hieter. 1990. Methods in Yeast Genetics: A Laboratory Course Manual.

Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York).

2.1.2 Polymerase chain reaction (PCR)

Gene fragments were amplified by PCR using yeast genomic DNA (for ski8 and dhh1) or plasmids (for hUpf1) as the template. PCR reactions were performed on an iCycler Thermal Cycler (Bio-Rad) with the parameters shown in the following table:

Reaction mixture components Volume (μl) Template 0.5 10× Pfu buffer 5 Turbo Pfu DNA polymerase 1 10 μM 5’ primer 1 10 μM 3’ primer 1 100 mM MgSO4 0.5 25 mM dNTPs 0.5 H2O 40.5 Total 50

46

Cycle parameters 1 cycle 95°C 5 min 95°C 30 seconds 30 cycles 50-55°C 30 seconds 72°C 2-3 min 1 cycle 72°C 7 min 15°C ∞

2.1.3 Agarose gel electrophoresis

5 μl of the PCR products were run on a 1.5% agarose gel mixed with 1× TAE buffer (40 mM Tris, 1.14% acetic acid, 1 mM EDTA) as running buffer.

2.1.4 Purification of PCR products

The PCR products were purified using a QIAquick PCR purification kit

(QIAGEN) according to the manufacturer’s instructions.

2.1.5 Enzyme digestion, dephosphorylation and purification

Restriction enzyme digestion of plasmid vectors and the purified PCR fragments was performed according to the following protocol:

Volumn (μl) Reagent Vector Purified PCR fragment DNA 20 20 10x Buffer 5 5 Enzyme1 2 2 Enzyme2 2 2 100x BSA 0.5 0.5 Distilled water 20.5 20.5 Total 50 50

The digestions were incubated in 37˚C for 3-5 hours. The vector mixture was further treated with calf intestinal alkaline phosphatase at 37˚C for 1 hour. The alkaline phosphatase was then inactivated at 75˚C for 10 minutes. The digested products were separated on an agarose gel as described above. The digested DNA fragments were 47

excised from the gel and purified using a QIAquick Gel Extraction kit (QIAGEN) according to the manufacturer’s instructions.

2.1.6 Ligation and transformation

Ligation was performed at 15°C overnight. 5μl of the ligation product was then

transformed into 100μl of DH5α competent cells for plasmid amplication according to

standard transformation protocols. The reaction mixtures are as follows:

Reaction mixture components Volume (μl) Double digested vector 3 Double digested PCR fragment 5 10x Ligation buffer 1 T4 ligase 1 Total 10

2.1.7 Plasmid preparation and positive clone screening

Several clones were inoculated in 3ml of LB medium containing the appropriate

antibiotic at 37°C overnight. Positive clones were screened by colony PCR. Plasmids

from positive clones were then obtained using the QIAprep Spin Miniprep kit and submitted for sequencing.

2.1.8 DNA sequencing

The sequences of positive clones determined by colony PCR were confirmed by

DNA sequencing. The reaction mixtures (20μl) are as follows:

Reaction mixture components Volume (μl) BigDye 3.1 sequencing mixture 8 Positive clone 5 10μM Primer 1 DI water 6 Total 20

The PCR sequencing reaction was performed as follows: 48

1 cycle 95°C 1 min 25 cycles 95°C 10 seconds 50°C 5 seconds 60°C 4 min 1 cycle 15°C ∞

Purification of the PCR products and running of the DNA sequencing gel were

carried out in an ABI Prism 377 DNA sequencer (Perkin Elmer) by staff from the DNA

sequencing unit at Institute of Molecular and Cell Biology (IMCB).

2.1.9 E. coli expression strain transfomation

Following sequencing, the recombinant plasmids were transformed into BL21

Star or Ril expression strains for overexpression in E. coli. Transformation was done as

described above except that 0.5μl of plasmid and 25μl of competent cells were used instead.

2.1.10 Protein expression test and expression strain storage

The expression strains containing the appropriate plasmids were inoculated into 2 ml of LB medium with the appropriate specific antibiotics and grew overnight at 37˚C.

600μl of cells were stored in 40% glycerol at -80˚C. For trial expression, 100μl of cells was transferred into 10 ml of fresh LB medium and cultured at 37˚C. When the optical density at 600nm (OD600) reached about 0.5-0.6, 0.5 mM of Isopropyl-ß-D-

thiogalactopyranosid (IPTG) was added to stimulate gene expression for an additional 3

hours at 37˚C. 100μl of cells were pelleted and boiled in SDS-PAGE loading buffer. The

samples were separated by SDS-PAGE using a 12% polyacrylamide gel

2.1.11 SDS-PAGE

SDS polyacrylamide gel electrophoresis (SDS-PAGE) was performed according

to Laemmli’s protocol (Laemmli, 1970). The gel was stained in staining buffer (45% 49

methanol, 10% acetic acid and 0.25% Coomassie Brilliant Blue R-250), and then de-

stained in 5% methanol and 7.5% acetic acid.

2.2 Protein purification

2.2.1 Large-scale cell culture for protein expression

The expression strains were inoculated into 150 ml of LB medium containing the

appropriate antibiotics at 37˚C overnight. 120 ml of cell culture was added into 6 L of LB

medium and shake-incubated (220rpm) at 37˚C. At a turbidity of OD600 = 0.5-0.6, protein

expression was induced by addition of IPTG to a final concentration of 0.1 mM and

further cell growth was performed overnight at 18˚C. The cells were harvested by

centrifugation, resuspended in buffer (20 mM Tris, 200 mM NaCl, pH7.4) and stored at -

80˚C for protein purification.

2.2.2 Purification procedures

The purification procedures for all three proteins are very similar. The differences

lie in the buffer compositions, ion exchange columns and gel filtration columns used. The

entire purification procedures and recipes of buffers are shown in Tables 2-1 and 2-2. The details of purification procedures are described in subsequent sections.

50

Purification steps Ski8 Dhh1 hUpf1 1 Cell lysis in lysis buffer for 30min 2 Sonication using Soniprep 150 for eight cycles 3 Spin down cell pellets with 18000rpm 1hr 4°C 4 Supernatant pass through Glutathione Sepharose 4B column 5 Wash with lysis buffer and elute with elution buffer 6 Protein quantity determination by Coomassie Protei Assay kit, add PreScission protease, 4°C overnight 7 Desalting by HiPrep Desalting column with ion exchange buffer A (described below) 8 Supernatant pass through Glutathione Sepharose 4B column 9 MonoQ ion-exchange MonoS ion-exchange 10 Superdex-75 gel filtration Superdex-200 gel chromatography filtration chromatography Table 2-1 Purification procedures of Ski8, Dhh1 and hUpf1. All of these steps were performed at 4°C.

Buffers Ski8 Dhh1 hUpf1 Lysis buffer 20 mM Tris, 500 20 mM MES, 500 20 mM Tris, 500 mM mM NaCl, 2 mM mM NaCl, 2 mM NaCl, 2 mM DTT, 2 DTT, 2 mM DTT, 2 mM mM benzamidine, 0.1 Bendazole, 0.1mM Bendazole, 0.1 mM PMSF, pH 6.0, 1 PMSF, pH 7.6, 1 mM PMSF, pH mg/ml lysozyme mg/ml lysozyme 6.0, 1 mg/ml lysozyme Elution buffer Lysis buffer with Lysis buffer with Lysis buffer with 20 20 mM GSH 20 mM GSH mM GSH Desalting buffer Corresponding ion exchange buffer A Buffer 20 mM Tris, 100 20mM HEPES, 20 mM HEPES, 100 Ion A mM NaCl, 2 mM 100mM NaCl, mM NaCl, 2 mM exchange DTT, pH 7.6 2mM DTT, pH 7.0 DTT, pH 7.0 buffer Buffer 20 mM Tris, 100 20mM HEPES, 20 mM HEPES, 1000 B mM NaCl, 2 mM 1000mM NaCl, mM NaCl, 2 mM DTT, pH 7.6 2mM DTT, pH 7.0 DTT, pH 7.0 Gel filtration buffer 20 mM Tris, 200 20 mM HEPES, 20 mM HEPES, 200 mM NaCl, 2 mM 200 mM NaCl, 2 mM NaCl, 2 mM DTT, pH 7.6, 10% mM DTT, pH 7.0 DTT, pH 7.0 glycerol Table 2-2 Recipes of buffers used for purification. All buffers are filtered using 0.45 μl filter.

51

2.2.2.1 Cell lysis

Cells were re-suspended in lysis buffer (Table 2-2, supplemented with 1mg/ml

Hen egg lysozyme) for 30 minutes and disrupted by sonication with 8 bursts of 15 seconds using a Soniprep 150 (SANYO). The resulting crude extract was cleared by centrifugation for 1 hour using the J6-HC Centrifuge (Beckman Coulter). The supernatant was filtered (0.45 μm) and loaded onto a self-packed Glutathione Sepharose 4B column

(Amersham), equilibrated in the above lysis buffer.

2.2.2.2 First GST affinity column chromatography

After binding of supernatant to the Glutathione Sepharose 4B beads, the column was washed with about 50 ml of the same buffer and the GST-fusion protein was eluted with 15 ml of elution buffer (Table 2-2). The protein concentrations were determined by the Bradford method using the Coomassie Protein Assay kit (Pierce) with BSA as a standard. PreScission protease (Amersham) was added at a ratio of 50:1 (w/w) to cleave off the GST tag. The reaction mixture was then left to incubate at 4°C overnight.

2.2.2.3 Second GST affinity column chromatography

The protease-treated protein sample was loaded onto a HiPrep Desalting column

(Amersham) equilibrated in desalting buffer (Table 2-2). After desalting, the protein sample was passed through a regenerated Glutathione Sepharose 4B column to remove the cleaved GST fragment. The flowthrough was further purified by ion exchange chromatography.

2.2.2.4 Ion exchange chromatography

The flowthrough from the second GST column was loaded onto a FPLC MonoQ

HR 10/10 column (Amersham) or MonoS HR 10/10 column (Amersham) ion-exchange 52 column (Table 2-1), equilibrated with ion-exchange buffer B (Table 2-2) and subsequently buffer A (Table 2-2). The proteins were eluted with a 100 ml linear gradient of NaCl (0.1 to 1.0 M) using the same buffers (A and B). The peak fractions were checked by SDS-PAGE and pooled together to be further purified by gel filtration chromatography.

2.2.2.5 Gel filtration chromatography

The pooled peak fractions were applied to a FPLC gel filtration column Superdex-

75 or Superdex-200 (Amersham) pre-equilibrated in gel filtration buffers (Table 2-2).

The peaks were verified by SDS-PAGE and the protein-containing peak fractions were pooled in vivaspin 20 concentrators (10,000 MWCO, Vivascience), concentrated to a final concentration of 10-15 mg/ml and stored at -80°C. The gel filtration chromatograms and SDS-PAGE gels of Ski8, Dhh1 and hUpf1 are shown in Figures 2-1, 2-2 and 2-3, respectively.

Figure 2-1 Purification of Ski8 (residues 1-397, MW: 45kD) (A) Superdex-75 gel filtration column Chromatogram (B) 12% SDS-PAGE gel of peak fractions

53

Figure 2-2 Purification of Dhh1 (residues 30-425, MW: 49kD ) (A) Superdex-75 gel filtration column Chromatogram (B) 12% SDS-PAGE gel of peak fractions

Figure 2-3 Purification of hUpf1 (residues 295-914, MW: 70kD) (A) Superdex-200 gel filtration column Chromatogram (B) 10% SDS-PAGE gel of fractions from the peak

2.3 Crystallization

Crystallization was carried out at 20°C by the hanging drop vapour diffusion method using Hampton Research 24-well plastic plates. The crystallization screen kits were purchased from Hampton Research. 1 μl of protein solution was mixed with an equal volume of reservoir solution for screening. Initial crystallization conditions were 54

obtained and used for further optimization. The dimension and quality of single crystals were further improved by microseeding and/or macroseeding techniques. Diffraction- quality crystals were cryoprotected in specific cryo-protectants and flash-frozen in liquid nitrogen for data collection. The optimized crystallization conditions and cryo-protectants

for each of the three proteins are shown in Tables 2-3 and 2-4. Pictures of crystals are shown in Figure 2-4.

Ski8 Dhh1 Crystallization 25-28%PEG4000, 50 mM 18-20%PEG400, 300 mM condition sodium citrate, KBr, 50 mM MES, pH 6.0 20% ethylene, pH 5.6 Cryo-protectant Same as crystallization 35% PEG400, 300 mM condition KBr, 50 mM MES, pH 6.0 Table 2-3 Crystallization conditions and cryo-protectants used for Ski8 and Dhh1

Free hUpf1 or co-crystallized with ligands PO4 AMPPNP AMPγS ADP Crystallization 100 mM 50 mM MES, 100 mM 100 mM Tris, condition Sodium/Potasium pH6.0, 1- MES, pH 6.0, pH7.5, 8-13% Phosphate, pH 3%PEG4000, 8-10% PEG3350, 140 5.8, 6-8% 0.005 mM PEG400, 5 mM PEG3350, 10 MgSO4, 10 mM MgCl2, (NH4)3PO4, mM DTT mM DTT 50 mM KCl 10% EG, 10 mM DTT Cryo- Crystallization Crystallization Crystallization Crystallization protectant condition with conditions with conditions conditions with 25%PEG400 25%PEG400 with 10%MPD 25%PEG400 Table 2-4 Crystallization conditions and cryo-protectants used for hUpf1 55

Figure 2-4 Crystals of Ski8, Dhh1 and hUpf1, as well as hUpf1 in complex with AMPPNP, ATPγS and ADP.

2.4 Structure determination

2.4.1 Heavy atom derivative preparation

In this thesis study, structure determinations of Ski8 and hUpf1 were based on Se-

Met derivatives, while the structure of Dhh1 came from a Br derivative because a high concentration of Br ion was contained in the crystallization condition for Dhh1. 56

Expression, purification and crystallization of Se-Met proteins were performed by the

same methods as their native proteins except that: (a) LB media for large cell culture was

replaced by MOPS media (see appendix); (b) IPTG-induced protein expression was

performed at 28°C for about 5 hours: (c) All buffers for purification and crystallization

contained 10 mM DTT to reduce Selenium oxidation.

2.4.2 Data collection and processing

Diffraction data were collected using flash-frozen Se-Met crystals of Ski8 and

hUpf1 and native protein crystals of Dhh1 grown from conditions containing 300 mM

NaBr. For MAD data collection, an appropriate x-ray energy was first determined by

fluorescence detection. The peak wavelength was chosen at the peak of absorption to give

a maximal ƒ” value and large Bijvoet differences. The edge wavelength (inflection point)

was selected at the ascending inflection point to give a minimal ƒ’ value. The remote

wavelength was chosen on the high-energy side of the absorption edge, which gave a

large difference in ƒ’ from the inflection point. After wavelength detection, the strategy

of data collection was determined by processing several diffraction images separated by

90 degrees. Usually, diffraction data at the peak wavelength were collected first, followed

by data collection at the edge and high-remote wavelengths. In the case of SAD data

collection, the strategy of inverse-beam geometry was used to collect Friedel related

reflections for a redundant set of anomalous signals. All data were collected with an

oscillation angle of 0.5 degrees.

2.4.2.1 Data collection and processing of SeMet Ski8

A MAD dataset for Ski8 was collected at outstation BW7A, DESY (Hamburg,

Germany). The three-wavelength MAD data for Ski8 were processed using DENZO 57

(Otwinowski and Minor, 1997) and intensities were reduced and scaled using

SCALEPACK (Otwinowski and Minor, 1997). The Se-Met derivative crystals contained one molecule per asymmetric unit and belonged to the space group P212121. The data collection and processing statistics are summarized in Table 2-5.

Se-Met MAD data

λ1 (peak) λ2 (edge) λ3 (remote)

Wavelength (Å) 0.9789 0.9791 0.9724 Resolution (Å) 2.2 2.3 2.3 a/b/c(Å) 65.5/66.8/81.5 α/β/γ (°) 90/90/90 Completeness (%) 98.8(100) 98.9(99.9) 99.8(99.9) Redundancy 3.5 3.5 3.5 I/σ (I) 10.3(2.0) 8.7(2.1) 7.5(2.0) a Rmerge (%) 6.3 7.3 8.6 Table 2-5 Statistics for the data collection and processing of Ski8. Values in parentheses indicate a the specific values in the highest resolution shell. Rmerge = ∑|Ij-|/∑Ij, where Ij is the intensity of an individual reflection, and is the average intensity of that reflection.

2.4.2.2 Data collection and processing of the Br derivative of Dhh1

Data collection of the Br derivative of Dhh1 was performed at ID29, ESRF

(Grenoble, France). The set of MAD data was collected and processed using MOSFLM

(Leslie et al., 1992) and the CCP4 package (CCP4, 1994). Crystals of Dhh1 belonged to the space group P21, with one molecule per asymmetric unit. Data processing statistics are summarized in Table 2-6.

58

Br MAD data

λ1 (peak) λ2 (edge) λ3 (remote) Wavelength (Å) 0.9195 0.9198 0.9134 Resolution (Å) 2.1 2.1 2.1 a/b/c(Å) 48.0/80.4/54.8 α/β/γ (°) 90/100.6/90 Completeness (%)a 99.5(98.7) 99.5(99.2) 99.2(98.5) Redundancy 3.6 3.7 3.6 I/σ (I) 12.8(6.8) 12.8(7.2) 11.6(5.5) a Rmerge (%) 4.0(10.6) 3.8(9.3) 4.4(12.8) Table 2-6 Statistics for the data collection and processing of Dhh1. Values in parentheses a indicatethe specific values in the highest resolution shell. Rmerge = ∑|Ij-|/∑Ij, where Ij is the intensity of an individual reflection, and is the average intensity of that reflection.

2.4.2.3 Data collection and processing of SeMet hUpf1

Data collection of the Se-Met derivative of hUpf1 was performed at ID29, ESRF

(Grenoble, France). One set of SAD data was collected for hUpf1-PO4, hUpf1 in complex with AMPPNP and hUpf1 in complex with ADP at the peak wavelength. The

SAD data were processed using MOSFLM (Leslie et al., 1992) and the CCP4 package

(CCP4, 1994). Data collection for hUpf1-ATPγS was carried out on a Rigaku x-ray generator (λ = 1.5418Å; Institute of Molecular and Cell Biology) and the data were processed using d*trek (Rigaku Corporation, 1991). Crystals of hUpf1-PO4, hUpf1-

AMPPNP and hUpf1-ATPγS belonged to the space group P43212 with one molecule per asymmetric unit, while hUpf1-ADP crystals belonged to the space group P1 with two molecules per asymmetric unit. Data processing statistics are summarized in Table 2-7.

59

hUpf1hd- hUpf1hd- hUpf1hd- hUpf1hd- Data collection AMPPNP ATPγS PO4 ADP Derivative SeMet - - - Wavelength(Å) 0.9793 1.5418 0.9793 0.9793 Resolution limit (Å) 2.6 3.2 2.8 2.4 Space group P43212 P43212 P43212 P1 Cell parameters 189.0/189.0 189.6/189.6/ 195.2/195.2/ 63.3/67.8 a/b/c(Å) /44.9 45.0 45.5 /87.3 114.4/90.1 α/β/γ (°) 90/90/90 90/90/90 90/90/90 /110.2 Unique reflections (N) 27959 17023 24896 43423 I/σ 6.0 (2.1) 3.4(2.0) 4.5(2.3) 8.8(3.1) Completeness(%) 97.2(98.9) 100(100) 100(100) 90.8(90.8) a Rmerge 0.094(0.379) 0.154(0.415) 0.102(0.395) 0.059(0.24) Table 2-7 Statistics for the data collection and processing of hUpf1. Values in parentheses a indicatethe specific values in the highest resolution shell. Rmerge = ∑|Ij-|/∑Ij, where Ij is the intensity of an individual reflection, and is the average intensity of that reflection.

2.4.3 Structure determination

2.4.3.1 Phasing, modeling and refinement of Ski8

For phase determination, the multiple-wavelength anomalous dispersion (MAD)

method was used. Four selenium sites were located using the automated Patterson search

routine implemented in the program SOLVE (Terwilliger and Berendzen 1999). The

searched Se heavy atom sites are shown in Table 2-8. Superimposition of the predicted vectors calculated from the heavy atom sites on the real patterson maps was performed to evaluate the heavy atom position using the program VECTORS (CCP4, 1994). Phases calculated with SOLVE were further improved by density modifications implemented in the program RESOLVE (Terwilliger 2002). The electron density map calculated from the improved phase was traceable (Figure 2-5). A partial model containing nearly 80% of the amino acids in the polypeptide chain was built automatically with RESOLVE 60

(Terwilliger 2002). The rest of the model was built manually using the program O (Jones

et al. 1991). Refinement was performed using the program CNS (Brunger et al.1998). A

test dataset (Rfree) was introduced into the refinement to monitor and validate the model refinement. After multiple rounds of model building and CNS refinement, water

molecules were added to the model automatically using CNS (Brunger et al, 1998). The

final round of the refinement was carried out with the program REFMAC5 (Murshudov

et al. 1997). The quality of the model was assessed with the program PROCHECK

(Laskowski et al. 1993), showing that 85.7 % of the residues lie in the most favored

region with no residues in the disallowed regions in a Ramachandran plot. The flow chart

of structure determination of Ski8 is shown in Figure 2-6. Crystallographic statistics are

summarized in Table 2-9.

TOP SOLUTION FOUND BY SOLVE ( = 0.37; score = 17.38) X Y Z OCCUP B HEIGHT/SIGMA 1 0.726 0.264 0.162 0.773 23.5 20.2 2 0.568 0.172 0.234 0.861 32.6 19.7 3 0.110 0.092 0.185 0.538 43.4 15.3 4 0.012 0.395 0.177 0.515 33.2 13.6 Table 2-8 Se atom sites of Ski8 found by the program SOLVE.

Figure 2-5 A partial electron density map of Ski8 shown in the program O. Several strands were clearly identified. Contour level: 1σ.

61

Peak data Edge data Remote data

Integration Integration Integration with DENZO with DENZO with DENZO

Scaling with Scaling with Scaling with SCALEPACK SCALEPACK SCALEPACK

Phasing with SOLVE

Model building with RESOLVE Model refinement with CNS Manual model building with O

Model building with REFMAC5

Figure 2-6 Flow chart of structure determination of Ski8.

62

Phase determination Number of sites 4 Figure of merit Before density modification 0.37 After density modification 0.59 Refinement Statistics Resolution range (Å) 2.2 Reflection used 17717 a Rcryst 24.6 b Rfree 27.9 Nonhydrogen atoms Protein (N) 2816 Waters (N) 159 R.m.s deviations Bond length (Å) 0.007 Bond angle (°) 1.06 Ramachandran plot Most favored region 93.3 Number of outliers 1 Table 2-9 Phase determination and refinement statistics of Ski8 structure. Values in a parenthesesindicate the specific values in the highest resolution shell. Rcryst = ∑||Fo| - |Fc||/∑|Fc|, where Fo denotes the observed structure factor amplitude, and Fc denotes the structure factor b amplitudecalculated from the model. Rfree is as for Rcryst but calculated with 5.2% of randomly chosen reflections omitted from the refinement.

2.4.3.2 Phasing, modeling and refinement of Dhh1

The structure of Dhh1 was also solved by MAD phasing. Four Br sites were

located using the program SOLVE (Terwilliger & Berendzen, 1999) (Table 2-10). The heavy atom sites were evaluated by superimposition of the predicted vectors on the real patterson maps using the program VECTORS (CCP4, 1994). Starting with the input of the four heavy atom sites found by SOLVE, the SHARP program further found another five heavy atom sites. Solvent flattening was performed by the program SOLOMON, which is implemented in the user interface of the SHARP program (De la Fortelle & 63

Bricogne, 1997). The electron density calculated using the improved phase from SHARP

was of excellent quality (Figure 2-7). 98% of the final model was built automatically by

the ARP/wARP program (Perrakis et al, 1999). The rest of the model was built manually

using the program O (Jones et al, 1991). After multiple rounds of model building and

CNS refinement, water molecules were added to the model automatically using CNS

(Brunger et al, 1998). The quality of the model was assessed with the program

PROCHECK (Laskowski et al. 1993) at every round of refinement. The final model from

CNS refinement was refined further by REFMAC5 (Murshudov et al, 1997). The flow chart of structure determination of Dhh1 is shown in Figure 2-8. Statistics for structure refinement are summarized in Table 2-11.

TOP SOLUTION FOUND BY SOLVE ( = 0.43; score = 27.11) X Y Z OCCUP B HEIGHT/SIGMA 1 0.837 0.501 0.135 0.652 26.4 9.9 2 0.886 0.556 0.409 0.910 19.2 11.8 3 0.059 0.160 0.131 0.807 44.9 10.8 4 0.864 0.155 0.340 0.556 19.2 9.8 Table 2-10 Br atom sites found by the program SOLVE

Figure 2-7 A partial electron density map of Dhh1 shown in the program O. Several strands were clearly identified. Contour level: 1σ. 64

Peak data Edge data Remote data

Data integration Data integration Data integration with MOSFLM with MOSFLM with MOSFLM

Data scaling with Data scaling with Data scaling with SCALA SCALA SCALA

Data truncation Data truncation Data truncation with TRUNCATE with TRUNCATE with TRUNCATE

Data combination with CAD

Three wavelength scaling with SCALEIT

Phasing with SOLVE

Heavy atom refinement, search, density modification with SHARP

Automatic model building with ARP/wARP Model refinement with CNS Manual model building with O

Model refinement with REFMAC5

Figure 2-8 Flow chart of structure determination of Dhh1.

65

Phase determination Number of sites 9 Figure of merit Before density modification 0.45 After density modification 0.82 Refinement Statistics Resolution range (Å) 20-2.1 Reflection used 22719 a Rcryst (%) 20.0(20.3) b Rfree (%) 23.4(23.6) Nonhydrogen atoms Protein (N) 3009 Waters (N) 253 R.m.s deviations Bond length (Å) 0.009 Bond angle (°) 1.09 Ramachandran plot Most favored region 88.2 Number of outliers 0 Table 2-11 Phase determination and refinement statistics for the Dhh1 structure. Values a inparentheses indicate the specific values in the highest resolution shell (2.25-2.1Å). Rcryst = ∑||Fo| - |Fc||/∑|Fc|, where Fo denotes the observed structure factor amplitude, and Fc denotes the b structure factor amplitude calculated from the model. Rfree is as for Rcryst but calculated with 5.0%of randomly chosen reflections omitted from the refinement.

2.4.3.3 Structure determination of hUpf1

The flow chart of structure determination of hUpf1 is shown in Figure 2-9. The structure of hUpf1-AMPPNP was solved by SAD phasing. Eleven Se sites were located using the program SnB (Miller R et al, 1994) from random atoms with 1000 trials. The sites of Se are shown in Table 2-12. Evaluations of 1000 trials are shown in Figure 2-10.

The heavy atom sites found by the SnB program were evaluated by superimposition of the predicted vectors on the real patterson maps using the program VECTORS (CCP4,

1994). Another two heavy atom sites were found by the SHARP program (De la Fortelle

& Bricogne, 1997). After refinement of these sites, the phases were improved by density 66

modifications implemented in SHARP. The electron density calculated using the improved phase from SHARP was of excellent quality (Figure 2-11), enabling 60% of

the final model to be built automatically by RESOLVE (Terwilliger, 2002). The rest of

the model was built manually using the program O (Jones et al, 1991). Crystallographic

refinement was performed using the programs CNS (Brunger et al, 1998) and REFMAC5

(Murshudov et al, 1997).

X Y Z Height 1 0.775382 0.544653 0.898875 18.41 2 0.956117 0.636535 0.900388 15.99 3 0.773498 0.041413 0.868174 14.45 4 0.808364 0.465599 0.898285 11.48 5 0.360180 0.113186 0.965427 9.85 6 0.827206 0.439998 0.927763 9.30 7 0.410556 0.083373 0.944319 8.97 8 0.115252 0.401748 0.911050 7.85 9 0.345389 0.179757 0.990991 7.82 10 0.648406 0.726533 0.976268 7.47 11 0.979000 0.644062 0.962606 7.01 Table 2-12 Se atom sites found by the program SnB.

67

Peak data Peak data hUpf1-AMPPNP hUpf1-ATPγS hUpf1-ADP, hUpf1-PO4

Data integration Data integration with MOSFLM with MOSFLM

Data scaling with Data processing Data scaling with SCALA with d*trek, SCALA

Heavy atom search with Data truncation with SnB TRUNCATE

Heavy atom refinement, density modification with SHARP

Molecular Automatic model replacement with building with Molrep/Phaser RESOLVE Model refinement with CNS

Manual model Manual model building with O building with O

Model refinement with Model refinement with REFMAC5 REFMAC5

Figure 2-9 Flow chart of structure determination of hUpf1.

68

Figure 2-10 Evaluation of 1000 trials by SnB program. (A) Histogram of Rmin function. It indicates which trial has the current best (smallest) value of the minimal function (Rmin). R(true) and R(random) denote the expected values of the minimal function for sets of true and random phases, respectively. (B) The Rmin trace. It shows a cycle-by-cycle trace of the value of Rmin for the best trial (lowest Rmin).

69

Figure 2-11 A partial electron density map of hUpf1-AMPPNP shown in the program O. Several strands were clearly identified. Contour level: 1σ.

The ADP-bound structure was solved by molecular replacement using Molrep

(CCP4, 1994). The structure of domain 1A (amino acids 295-700, excluding residues

325-415 and residue 555-610; Chapter 5) of hUpf1-AMPPNP was used as the search model . A solution containing two copies of the N-terminal domain was found (Table 2-

13). Fixing the solution of domain 1A, two copies of domain 2A (residues 705-914) were located (Table 2-14). Subsequently, rigid body refinement of two copies of domains 1A and 2A was carried out using Refmac5 (Murshudov et al, 1997). The difference Fourier map showed clear electron density for the remaining domains 1B and 1C. These two domains were placed manually into the electron density map and subjected to several rounds of manual building and refinement using the program O (Jones et al, 1991).

Crystallographic refinement was performed using CNS with restrained Non- crystallography symmetry (NCS) being used throught the refinement (Brunger et al,

1998). The final refinement of the model was carried out with REFMAC5 (Murshudov et al, 1997). 70

theta phi Chi Alpha beta Gamma Rf Rf/Sigma Sol_RF 1 159.69 -94.99 82.99 135.33 26.59 145.32 45.63 11.09 Sol_RF 2 102.92 40.17 177.25 226.31 154.02 325.97 43.49 10.57 Sol_RF 3 11.12 76.20 44.68 8.16 8.41 35.77 19.28 4.69 Sol_RF 4 93.96 158.34 177.50 355.87 171.70 219.18 18.77 4.56 Sol_RF 5 11.94 58.80 39.73 348.26 8.06 50.67 18.49 4.49 Table 2-13 Result of rotation function by Molrep using domain 1A as a search model. The two best solutions clearly give a markedly high value than the rest.

theta phi Chi Alpha Beta Gamma Rf Rf/Sigma Sol_RF 1 155.11 -67.51 69.34 170.38 27.71 125.41 30.18 7.32 Sol_RF 2 77.85 -146.64 171.78 194.51 154.37 307.78 28.19 6.84 Sol_RF 3 40.62 89.80 175.56 86.88 81.17 87.28 18.75 4.55 Sol_RF 4 130.65 93.84 177.89 275.46 98.68 267.77 17.88 4.34 Sol_RF 5 79.55 -171.31 96.54 110.18 94.43 272.80 17.60 4.27 Table 2-14 Result of rotation function by Molrep using domain 2A as a search model and fixing two copies of domain 1A. The two best solutions clearly give a markedly high value than the rest.

The structure of hUpf1-PO4 was solved by a combination of SAD phasing and molecular replacement method. The structure of hUpf1-ATPγS was solved by molecular replacement using the structure of hUpf1-AMPPNP without ligand as the search model.

Manual model building and refinement were carried out with the programs O and CNS

(Jones et al., 1991; Brunger et al., 1998). The final crystallographic refinement was performed with the program REFMAC5 (Murshudov et al., 1997). Statistics for refinement are summarized in Table 2-15.

71

hUpf1- hUpf1- hUpf1- hUpf1- AMPPNP ATPγS PO4 ADP Number of sites 13 Figure of merit Before density modification 0.284 After density modification 0.847 Refinement Statistics Data range(Å) 20-2.6 20-3.2 20-2.8 20-2.4 Reflections used 24065 13391 21121 41163 a Rcryst 24.5 25.4 25.8 23.5 b Rfree (%) 27.1 30.6 29.8 27.2 Nonhydrogen atoms Protein (N) 4872 4877 4715 9545 Waters (N) 110 0 63 243 R.m.s.deviations Bond length (Å) 0.006 0.006 0.012 0.008 Bond angle (°) 1.048 0.955 1.294 1.011 Ramachandran plot Most favored region 83.9% 86.7% 85.2% 87.6% Number of outliers 1 0 1 0

a Table 2-15 Phase determination and refinement statistics of hUpf1. Rcryst = ∑||Fo| - |Fc||/∑|Fc|, where Fo denotes the observed structure factor amplitude, and Fc denotes the structure b factoramplitude calculated from the model. Rfree is as for Rcryst but calculated with 5.0% of randomly chosen reflections omitted from the refinement.

72

2.5 Biochemical and molecular biological experiments

2.5.1 Experiments for Ski8

2.5.1.1 Site-directed mutagenesis and yeast two-hybrid assay

To check the interactions of Ski8 with Ski2 or Ski3 or Spo11, yeast two-hybrid assay was performed using the Matchmaker3 system (Clontech). Site-directed mutagenesis was performed using the Quick-Change system according to the manufacture’s instructions (Stratagene). Gal4-BD domain fusion of wild type Ski3, Ski2 and Spo11 were constructed by inserting double enzyme-digested fragments into pGBKT7 vector, using restriction endonucleases EcoRI & BamHI, SmaI & PstI, respectively. Gal4-AD domain fusion of wild type and mutant Ski8 were prepared by inserting restriction endonucleases NdeI & XhoI digested fragments into pGADT7 vector. Haploid yeast strains AH109 carrying Gal4-AD fusion protein (strains carrying only vector pGADT7 as negative control) and Y187 containing Gal4-BD fusion protein were mated in appropriate pairwise combinations and the resulting diploids were grown on synthetic dropout medium without leucine and tryptophan. All vectors and yeast strains used here are from Clontech. For yeast two-hybrid assays between variant Ski8 and Ski2/Ski3, corresponding diploid colonies were spread on the synthetic dropout medium without tryptophan, leucine, histidine and adenine. For β-galactosidase activity assay, overnight diploid cultures were diluted with fresh medium and grown to mid-log phase (3-4hrs). Cells were broken by freeze/thaw method and activity assay were carried out according to standard protocols (Clontech). One unit of β-galactosidase hydrolyzes 1

μmol of o-nitrophenyl β-D-galactopyranoside per min per cell.

2.5.1.2 GST pull-down assay

73

Ski3 and Spo11 were translated in vitro using the recombinant plasmids pGBKT7

described above as templates in the presence of 35S-methionine with The TNT® T7

Quick Coupled Transcription/Translation System (Promega). For GST-pull-down assays,

500 μg of GST or GST fusion Ski8 variants (wild type, Ski8-F59A and Ski8-Topmutant)

were immobilized on glutathione-Sepharose. Bound fusion proteins were incubated with

5μl of the in vitro translated Ski3 and Spo11 at 4°C for 1-2 hour. The beads were washed

five times with binding buffer (20 mM HEPES, 1 mM EDTA, 10 mM MgCl2, 4 mM

DTT, 10% glycerol, 0.1 mM PMSF, 0.5% Triton X-100, pH 7.9). Bound proteins were

eluted in SDS loading buffer and resolved by SDS/PAGE (0.5 µl of in vitro translated

protein as input), and visualized by autoradiography.

2.5.2 Experiments for Dhh1

2.5.2.1 CD-spectroscopy

CD spectra were measured at 20˚C on a Jasco J-810 spectrapolarimeter at a

resolution of 0.1 nm, with a bandwidth of 2 nm. Five spectra were averaged for each

derivative. For far UV spectroscopy (200-250 nm), various protein samples were diluted

in 50 mM NaCl, pH6.0, at a concentration of 4 µM (derived from UV spectroscopy)

using a cell of 0.1 cm path-length. The mean residue ellipticity was calculated according

to the CD measurement manual.

2.5.2.2 Mutagenesis and in vivo mRNA turnover assays

These experiments were performed by Dr. Jeff Coller and Dr. Roy Parker from

University of Arizona. Alleles of Dhh1 used in both the in vitro and in vivo analyses were

created using Quicksite-mutagenesis (Stratagene). Specific point mutants were generated 74

and verified by sequence analysis. Complementation and dominant negative growth

assays were performed by transforming each plasmid into either a wild-type (yRP841), or

dhh1∆ deletion (yRP1561) (Coller et al., 2001) strain. Cells were then patched onto

selective media and grown at either 30°C or 37°C.

RNA analysis was performed as previously described (Coller et al., 2001).

Briefly, cells were transformed with plasmids expressing the various Dhh1p alleles or

controls, and then grown in selective media until reaching mid-log phase. At this point,

the cells were harvested, RNA was extracted and the EDC1 mRNA was visualized by

northern analysis using radiolabeled DNA oligonucleotide oRP1236 as a probe. The

SCR1 RNA was probed as a loading control using radiolabeled DNA oligonucleotide

oRP100.

2.5.2.3 Limited Proteolysis

The reactions were performed in a buffer containing 20 mM Tris, 25 mM NaCl, 2

mM MgCl2, 2 mM CaCl2, pH 7.0. 20 μM of the wild-type and mutant tDhh1 (truncated

Dhh1) proteins and 10 μM trypsin were used for each reaction. The concentrations of ligands used were 40 μM poly(U), 2 mM ATP, AMP-PNP or ADP. Reactions were started by the addition of protease and stopped at various times from 5 to 60 minutes by the addition of standard protein sample buffer containing 100 mM NaOH, and the

reaction solutions were then immediately boiled for 10 min. After neutralization by the addition of 1N HCl, the samples were then analyzed by SDS-PAGE.

2.5.2.4 In vitro RNA binding assay

Equal amounts (15 μg) of BSA, wild type and mutant tDhh1 proteins were dot-

blotted on a PVDF membrane. The membrane was blocked in buffer A (25 mM NaCl, 10 75

mM MgCl2, 10 mM HEPES, 0.1 mM EDTA, 1 mM DTT, 3% BSA, pH 7.0) for 1 hour at

room temperature. 12mer poly (U), dsRNA-I and ss/dsRNA-II were purchased from

Dharmacon. The later two RNA substrates were annealed by boiling at 95°C 10min and

then slowly cooling down to room temperature. RNA substrates were labeled with T4

kinase (Invitrogen) in the present of [γ-32P]ATP (Amersham). 10 pmol of 32P-labelled

RNA substrates were added for binding assay in buffer B (50 mM NaCl, 10 mM MgCl2,

10 mM HEPES, 0.1 mM EDTA, 1 mM DTT, 1.5% BSA, pH 7.0). After incubation in buffer B for two hours, the membrane was washed with 20 ml of buffer B three times and quantified using a phosphoimager (Molecular Dynamics). BSA protein served as a negative control.

2.5.3 Experiments for hUpf1

2.5.3.1 Mutagenesis and In vitro ATPase activity

Alleles of hUpf1hd (helicase domain) used in the in vitro analysis were created using Quicksite-mutagenesis (Stratagene). Specific point mutants were generated and verified by sequence analysis. ATPase assay was performed in a 20 µl reaction containing the purified 200nM wild-type or mutant hUpf1 proteins in 50 mM MES (pH

6.0), 50 mM KAc, 2.5 mM Mg(Ac)2, 1 mM DTT, 500 nM 15mer poly(C), 0.1 mg/ml

BSA, 10 µM cold ATP and 1 µCi [γ-32P]ATP. After incubation for 1 h at 37 °C, the reaction was terminated by adding 1 µl 500mM EDTA. For TLC analysis, 1 µl of the reaction mixture was spotted on a PEI-cellulose plate and developed by 0.3M K2HPO4

(pH 7.6).

2.5.3.2 In vitro ATP binding assay 76

Equal amounts (15µg) of BSA, wild-type and mutant hUpf1hd proteins were dot-

blotted on a PVDF membrane. The membrane was blocked in buffer A (20 mM HEPES,

50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 3% BSA, pH 7.0) for 1 hour at room temperature. 50 μCi of 5000Ci/mmol [γ-32P] ATP (Amersham) was added for the binding assay in buffer B (20 mM HEPES, 50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 1.5%

BSA, pH 7.0). After incubation in buffer B for 30 minutes, the membrane was washed

with 20 ml of buffer B twice and quantified using a phosphorimager (Molecular

Dynamics). BSA protein served as a negative control.

2.5.3.3 In vitro RNA binding assay

Equal amounts (15µg) of BSA, wild-type and mutant hUpf1hd proteins were dot-

blotted on PVDF membrane. The membrane was blocked in buffer A (20 mM HEPES,

50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 3% BSA, pH 7.0) for 1 hour at room temperature. Single-stranded RNA (5’CGCCCGAGGCTGTGCCGT3’) was purchased from Dharmacon. The RNA substrates were labeled with T4 kinase (Invitrogen) in the presence of [γ-32P] ATP (Amersham). 1 pmol of 32P-labled RNA substrate was added for

the binding assay in buffer B (20 mM HEPES, 50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM

DTT, 1.5% BSA, pH 7.0). After incubation in buffer B for 1 hour, the membrane was washed with 20 ml of buffer B three times and quantified using a phosphorimager

(Molecular Dynamics). BSA protein served as a negative control.

2.5.3.4 In vivo NMD analysis and P-body formation

These experiments were performed by Dr. Denise Muhlrad and Dr. Roy Parker

from University of Arizona. The UPF1 plasmids used were on a CEN LEU2 vector.

Wild-type, pRP910; DE572AA, pRP912. Additional UPF1 point mutants were created 77 using the Quick Change method (Stratagene) on a UPF1 BglII fragment cloned into pBluescript (pRP1318). Mutations were verified by sequencing and then put back into pRP910. Plasmids created were: pRP1319 (∆261-351), pRP1320 (∆494-546), pRP1321

(K436A), pRP1324 (Q601A), pRP1325 (R639A), pRP1329 (R801A). These mutants were analyzed in the yeast strain yRP1864: MATa, ura3, his4, leu2, lys2, trp1,

UPF1::URA3, DCP2GFPNEO. Strains containing the UPF1 plasmids were grown in minimal selective media to mid log and either harvested for RNA, or visualized microscopically as previously described (Sheth and Parker, 2006). Northern blots were probed either for the 7s RNA (oRP100) or the CYH2 mRNA (oRP1299 and/or 1300).

2.5.3.5 Surface plasmon resonance (SPR)

SPR was carried out on a Biacore 2000 instrument (Biacore). The 5’ end biotin- labeled ssRNA (TGTCATTCGAGTACAGTCTGTTCAGCTAGTCTCC) purchased from CureVAc was attached to a streptavidin-coated sensor chip (Biacore). A buffer of

10 mM HEPES, pH7.0, 150 mM NaCl, 0.005% (v/v) Tween-20 was flowed across the chip until the trace leveled off. The biotin-labeled RNA was then attached to the flow cell

2 by injecting 20 μl of 100nM RNA in 0.3 M NaCl at flow rate 5 μl/min. After immobilization, flow cell 2 and reference flow cell 1 were blocked with 100 μl of 1 mg/ml biotin at 5 μl/min. A binding buffer of 20 mM HEPES, pH7.0, 100 mM NaCl, 2 mM MgCl2, 2 mM DTT, 0.002% (v/v) Tween-20 was flowed across flow cell 1 and 2.

Typically, 150 μl of 100 nM protein sample was injected into the same buffer across the chip at 50 μl/min. The data were fitted to a 1:1 binding mode with mass transfer using

BIAevaluation 3.1.

78

Chapter 3

Crystal Structure and Mutagenesis Studies of Ski8

3.1 Results

3.1.1 Overall structure

The structure of full length Ski8 from S. cerevisiae was solved by MAD phasing, which yielded an easily interpretable electron density map at 2.2 Å resolution, allowing

80 % of the model to be built automatically. The current model has been refined with working and free R factors of 24.6 % and 27.9 %, respectively. Five regions of the polypeptide chain are not visible in the electron density map and are assumed to be disordered, namely, residues 101-104, residues 133-136, residues 225-233, residues 277-

287, and residues 370-375. The details of structure determination are shown in Chapter 2.

As shown in Figure 3-1, the polypeptide chain of Ski8 folds into a seven-bladed β propeller similar to that observed in the β subunits of heterotrimeric G proteins (Gβ) and in the C terminus of the yeast transcriptional repressor Tup1 (Tup1c). The propeller fold is characterized by seven blades that are pseudo symmetrically arranged around a central axis. Each blade consists of a four-stranded antiparallel β sheet, with the strands in each blade (labeled A-D) running approximately parallel to the pseudo-seven-fold axis and ranging from the inside to the outside of the propeller. The center of the propeller is formed by the edges of the seven “A” strands which delineate an axial channel running through the center of the propeller, which is filled with a large number of very well- ordered solvent molecules. The eponymous Trp-Asp motif is only present in blades 1 and 79

6 in Ski8 (Figure 3-2), being replaced by other amino acids in blades 2, 3, 4, 5 and 7.

Blades 1-6 are formed by contiguous segments of the polypeptide chain. In blade 7, the innermost three strands come from the extreme C-terminus while the outmost strand is provided by a strand from N-terminus.

A 5 B 6D-7A 6 Top N 4

C 7 A B C D 3 3C-3D 1 H3 2 2A-2B Bottom

C

N N

C C

Figure 3-1 Overall structure of Ski8. (A) Ribbon diagram of Ski8p showing the seven-bladed β propeller structure. Each blade consists of a four-stranded β sheet (labeled A-D). The view is looking down the central axis of the propeller onto the top surface. The “top” surface is defined by the presence of the D-A loops connecting sequential blades. (B) View rotated 90° about the horizontal axis in (A), looking at the propeller from the side. The top face and bottom face are marked according to the convention for WD-repeat protein structure. (C) Stereoview of a Cα trace of Ski8p with every tenth residue labeled and marked with a closed circle. Figures 3-1, 3-3, 3-4(c) and 3-5 were generated using MOLSCRIPT (Kraulis 1991).

80

3.1.2 Comparison with other WD repeat proteins

Although the sequence of Ski8 shows only 15.5 % of identity with Gβ and 18.5%

of identity with Tup1c, the β propeller of Ski8 superimposes strikingly well with Gβ with

an r.m.s.d of 1.5 Å over 238 Cα atoms, and with Tup1, with an r.m.s.d of 1.3 Å over 233

Cα atoms (Figures 3-3A and 3-3B). The structure of a single blade is strikingly conserved both within the Ski8p structure (Figure 3-3C), and between Ski8 blades and individual blades in Gβ and Tup1c. For instance, blade 1 of Ski8 superimposes with other

Ski8p blades with Cα-Cα r.m.s.d. of 0.9-1.4 Å, and with a typical blade of Gβ, blade 4 or with a typical blade of Tup1c, blade 3, with the same r.s.m.d. of 1.2 Å. The 'structural tetrad' or hydrogen-bonding network, as described in Gβ and Tup1c (Wall et al. 1995;

Lambright et al. 1996; Sprague et al. 2000), is also observed, but only in blade 1 and 6 of

Ski8 propeller (Figure 3-3D). This tetrad is formed between Trp in strand C, Ser/Thr in strand B, His in the DA loop and the nearly invariant Asp in the tight turn between strands B and C.

Despite the similarities, some structural differences exist between the propeller of

Ski8 and those of Gβ and Tup1c. The most notable differences occur in N-terminus and the loop regions. Ski8 lacks the N-terminal extension Gβ and Tup1c have. In Gβ, the N-

terminal extension forms a α helix, which participates in a coil-coil with the N-terminal

segment of the Gγ subunit in the trimeric G protein complex (Wall et al. 1995; Lambright

et al. 1996). The N-terminal 50 amino acids in Tup1c forms a subdomain that is joined to

the propeller by β sheet interactions that extend blade 6 into a six-stranded sheet (Figure

3-3A) while the similar region in hTle1-C forms a β hairpin that extends blade 5 into a

six-stranded sheet (Pickles et al. 2002; Sprague et al. 2000). In Ski8, 6D-7A loop which 81 is about 26 residues long protrudes from the top face of the propeller (Figures 3-1B and

3-3B). Since this loop is only present in Ski8p and not in other Ski8p homologs (Figure

3-2), it is not likely to be central to Ski8p function and is probably involved in yeast- specific functions. Loop 2A-2B connects strands 2A and 2B, which are substantially longer than their corresponding strands in both Gβ and Tup1c (Figure 3-3B). The 3C-3D loop stretches out from the bottom face and contains a short helix. Such loop structure is not observed in both Gβ and Tup1c (Figure 3-3B), but is conserved in Ski8 across species (Figure 3-2). Interestingly, strand 3D seems only present in Ski8 from S. cerevisiae (ScSki8) and S. macrospora (SmSki8) but not in the rest of the Ski8 homologs

(Figure 3-2). Since strand 3D is essential for formation of blade 3 and structurally conserved in Ski8p, Gβ and Tup1c, it is likely that Ski8 homologs other than ScSki8 and

SmSki8 may use part of the 3C-3D loop to form strand 3D. 82

Figure 3-2 Sequence alignment of Ski8 homologs. The secondary structures of S. cerevisiae Ski8 are shown. Invariant residues are white letters, similar residues are red and others are black. Residues speculated for interactions with Ski3 and Spo11 are indicated by *.

83

A B 6D-7A Blade 6 Gβ

Tup1c

3C-3D H3 Ski8p 2A-2B

C D

Ser A B C D Trp His

Asp

Figure 3-3 Comparison of Ski8p with Gβ and Tup1c. (A) Superposition of Ski8, Gβ and Tup1c. Ski8p is shown in red, Gβ in blue and Tup1c in green. The view of Ski8p is as in Figure 1A (B) Same as (A) but with the view looking at the side of the propeller. (C) Superposition of all seven blades of Ski8p. The Cα backbone for each of the seven blades was aligned with respect to blade 1. Four strands of each blade are indicated by A, B, C and D. blade 1, green; blade 2, red; blade 3, yellow; blade 4, cyan; blade 5, blue; blade 6, maroon; blade 7, magenta. (D) Superposition of two typical blades, blade 1 (green) and blade 6 (maroon) containing the eponymous Trp-Asp motif. Four conserved residues, which involved in the structural tetrad are showed in stick models and hydrogen bonds are indicated with dashed lines.

3.1.3 Location of protein-protein interaction sites on the β propeller

Previous analyses indicated that Ski8 interacts with Ski3 and/or Ski2 in 3’ to 5’

mRNA decay, and Spo11 during meiotic recombination (Brown et al. 2000; Arora et al. 84

2004). To identify the key regions on the surface of Ski8, which are likely to be involved

in interaction with Ski3, Ski2 and Spo11, the sequence conservation shared by eukaryotic

Ski8 proteins were mapped on the molecular surface of the budding yeast Ski8 structure.

This analysis revealed a prominent conserved patch that is situated on the top face of the

β propeller and encompasses nearly all of the DA and BC loops (Figure 3-4A).

Moreover, mapping of the side chains of hydrophobic residues on the molecular surface of Ski8 reveals that a large hydrophobic patch consisting of residues F20, F89, W125,

F188, W293, W311 and F358 is located on the top face of the β propeller, overlapping with the conserved patch identified by conservation mapping (Figures 3-4B & 3-4C).

The presence of such a conserved patch of hydrophobic residues suggests this region is a site of protein-protein interaction. Inspection of both the side and bottom surfaces of the propeller shows that there is no obvious conserved or hydrophobic patch, which is large enough for potential protein-protein interactions.

Consistent with the top face of Ski8 being a site of protein interaction, in the Ski8 crystal lattice, loops 3C-3D and 2A-2B from symmetry-related molecules bind to the top surface area, burying a pairwise accessible surface area of 1260 Å2. The interactions of

loops 3C-3D and 2A-2B with the top face are predominantly hydrophobic in nature with

some additional hydrogen bonds. The top surface that interacts with loops 3C-3D and

2A-2B is composed mainly of hydrophobic amino acids (F20, F89, V146, K147, F188,

N205, R237, W293, M295, W311, F358; Figure 3-5A). Among them, F20, F188, R237,

W293, M295 and F358 are located in the DA loop of blade 1, 4, 5, 6 and 7, respectively,

and F89, V146, K147, N205 and W311 lie in BC loop of blade 2, 3, 4 and 6, respectively.

Most of these residues are highly conserved in Ski8 homologs across species, except for 85

V146, N205 and W311 (Figure 3-2). The surface area of loops 3C-3D and 2A-2B that interacts with the top face consists of hydrophobic residues A78, L165, L167, and the

additional polar and charged residues R76, D77, D160, E161, S162 and T166 (Figure 3-

5A). Among them, only E161 and T166 are highly conserved across species (Figure 3-

2). These results suggest that the top face of the β propeller of Ski8 is a hydrophobic

patch and is likely involved in binding Ski3p, Ski2 and Spo11 or other unidentified

interacting proteins.

A B

W293 W293 W311 R237 F358 F188 F188 W125 F20 W125 F89

C

W293

W311 R237

F188 F358 W125 F20 F89

Figure 3-4 Molecular surface views of Ski8. (A) Surface representation of Ski8p showing the regions of high to low sequence conservation shared by the eukaryotic Ski8 proteins, corresponding to a color ramp from red to blue, respectively. Invariant residues are labeled. The view is as in Figure 1A. (B) Molecular surface of Ski8p colored according to residue property, with hydrophobic residues green and other residues gray. The hydrophobic residues are labeled. The view is as in (A). (C) The worm model showing the Cα backbones of Ski8p. Residues located either in hydrophobic patch or in conserved patch are shown in stick models. The view is as in (A). Figures (A) and (B) were produced using GRASP (Nicholls et al. 1991). 86

A W293 R237 M295 W311 S162 T166 N205 Helix H3 in the D160 L167 F188 3C-3D loop L165 V146 F358 E161 W125 A78 K147 F20 2A-2B loop F89 R76 D77

B

3C-3D 2A-2B β-catenin peptide The helix from Gα

Figure 3-5 Location of the protein-protein interactions site on the top face of the β propeller. (A) Close-up view of the interface between the top face of the β propeller in Ski8p (yellow) and the loops of 2A-2B and 3C-3D from the symmetry related molecule (purple). Residues involved in the interface are shown in stick models. (B) Comparison of the protein-protein interactions on the top surface for three WD-repeat proteins. Left: the top face of Ski8p (yellow) interacting with its symmetry related loop regions 3C-3D and 2A-2B (purple). Medium: the top face of β-TrCP1 WD- 40 domain (sky blue) with bound doubly phosphorylated β-catenin peptide (red). pdb code: 1p22. Right: the top face of Gβ (light green) interacting with the helix of Gα (blue). pdb code: 1gp2. All residues involved in interaction on the top face of the β propeller are shown in CPK model. 87

3.1.4 Mutational Analysis of Ski8

Evidence suggests that Ski8 interacts directly with the Ski3 and Spo11 proteins,

and may interact directly, or in combination with Ski3 to Ski2. Specifically, co-

immunoprecipitation showed that Ski3 interacts with Ski8 without the involvement of

Ski2 (Brown et al. 2000). However, Ski2 did not associate with Ski3 in the absence of

Ski8, nor did Ski2 associate with Ski8 in the absence of Ski3, suggesting that the binary complex formed by Ski3 and Ski8 is required for stable binding of Ski2 to Ski8 and/or

Ski3. Moreover, Ski8 also interacts directly with Spo11 and participates in meiotic recombination (Arora et al. 2004). However, the structural mechanism of these interactions has not been elucidated, and the amino acids residues critical for Ski8p interaction with Ski3, Spo11, or possibly Ski2, have not been identified.

Conservation and hydrophobicity mapping suggested that the top face of the β propeller of Ski8 is likely involved in the interaction of Ski8 with its binding partners. To examine the role of the amino acids located on the top face of Ski8 play in mediating interactions with Ski3, Ski2 and Spo11, a Ski8 variant where the top surface was altered was created by site-directed mutagenesis. The resulting variant Ski8 protein was examined for its ability to bind to Ski3, Ski2 and Spo11 by yeast two-hybrid and GST pull-down assays. In the latter assay, wild type and mutant Ski8 were immobilized on

glutathione-Sepharose and examined for their binding to Ski3 and Spo11 translated in

vitro in the presence of 35S-methionine. The specific mutant created (referred to as “top”

mutant) contains alanine substitutions of six residues (F20A, F89A, W125A, W293A,

W311A, F358A) located at the hydrophobic patch plus R237, a well-conserved residue in

Ski8 family found with the conserved top surface. An important result was the alteration 88 of the top surface of Ski8 substantially reduced binding of Ski8 to either Ski3 or Spo11

(Figures 3-6A and 3-6B). These results indicate that the top surface of Ski8 is required for binding to Ski3 and Spo1l, although the possibility of whether there are additional contacts between Ski3 and Spo11 to the side surface of Ski8 can not be excluded (see below).

Interestingly, mutation of the Ski8 top surface appears to have little effect on the binding of Ski8 to Ski2 (Figure 3-6C). This suggests that interactions between Ski2p and

Ski8p do not require the top surface and therefore a Ski8-Ski2 interaction is likely to involve a different surface of the Ski8. In addition, because the “top” mutant strongly inhibits Ski8-Ski3 interaction, this result raises the possibility that Ski8 and Ski2 directly interact even in the absence of Ski3. The fact that the “top” mutant still interacts with

Ski2 implies that these mutations present in the Ski8 “top’ mutant have not grossly altered the overall structure of the Ski8, consistent with the changes simply being changes in surface structure. Finally, our results suggest that the binding of Spo11 to Ski8 may also involve interactions with the side of the β propeller structure. Specifically, mutation of residue F59 located at the side surface to alanine (F59A) caused a substantial reduction of binding to Spo11, but did not affect the binding of Ski8 to Ski3 (Figures 3-6A and 3-

6B).

89

Figure 3-6 Mutational analysis of Ski8 mutants. (A) Effects of mutations at the top surface of the β propeller on the interactions of Ski8 with Ski3 and Spo11 respectively. β-galactosidase activity from various transformants estimated as described in Chapter 2. The Ski8-topmutant refers to a mutant Ski8 protein where seven residues (F20A, F89A, W125A, R237A, W293A, W311A, F358A) were mutated to alanine. Left: Ski8 vs. Ski3; Right: Ski8 vs. Spo11. (B) Ski3 and Spo11 were translated in vitro in the presence of 35S-methionine with the TNT T7 Quick Coupled Transcription/Translation system (Promega). 500 μg of GST or GST fusion Ski8p variants (wild type, Ski8-F59A and Ski8-Topmutant) were immobilized on glutathione-sepharose. Pulldown assay was performed at 4˚C for 1-2 h. (C) Effects of mutations at the top surface of the β propeller on the interactions of Ski8 with Ski2, Ski3 respectively. Alanine substitutions for seven top residues of Ski8 (Ski8-topmutant) abolish the two-hybrid interaction between Ski3 and Ski8, whereas such mutations have no effect on the interactions between Ski2 and Ski8. Plus and minus signs indicate positive and negative interactions, respectively. F59A serves as negative control. 90

3.2 Discussion

Solving the structure of Ski8 has revealed that this protein folds into a classic β

propeller structure similar to the known structures of other WD motif containing proteins.

In addition, the Ski8 structure and experimental analysis presents evidence that the top

surface of the Ski8 β propeller structure functions as a site of protein-protein interactions

and is required for interactions between Ski8 and both Ski3 and Spo11. This conclusion

is based on the presence of a conserved patch of largely hydrophobic residues on the top

surface (Figure 3-4A), and on the observation that mutation of this surface disrupts two

hybrid interactions between Ski8 and both Ski3 and Spo11 (Figure 3-6).

Similar to Ski8, several other WD-repeat proteins use their top faces for

interacting with their protein partners. For example, in the structure of heterotrimeric G

protein, the top face of Gβ mediates its interaction with Gα-GDP (Wall et al. 1995;

Lambright et al. 1996; Figure 3-5B). Morevoer, the structure of the β-TrCp1-SkP1-β-

catenin ternary complex shows that the β-catenin peptide binds the top face of the β

propeller of β-TrCp1 (Wu et al. 2003; Figure 3-5B). Similarly, the structure of Gβ complexed with phosducin showed that the N-terminal domain of phosducin interacts with all of the top loops of the β propeller of Gβ (Gaudet et al. 1996). Finally, eleven point mutations in the yeast Tup1p that specifically affect its interaction with Matα, a promoter-specific DNA-binding protein, have been mapped on the top face of the Tup1c propeller (Sprague et al. 2000; Komachi and Johnson 1997). Taken together, these results suggest that the top face of the propeller is a site for protein-protein interactions that may be conserved in WD repeat domains in general, although the specific residues involved in protein-protein interaction in the individual WD repeat domains may vary. 91

These results suggest a model for the assembly of the Ski complex involved in

mRNA decay wherein the Ski8 plays an important role in bringing Ski3 and Ski2

together. Mutation of the top surface of Ski8 affects Ski3 binding, but does not appear to

affect Ski8-Ski2 interaction, which suggests that the ski complex is formed according to

the following interactions. First, interactions between the top surface of Ski8 and Ski3

would nucleate the complex, and based on the co-immunoprecipitation experiments and

our GST pull-down assay are stable enough to persist even in the absence of Ski2 (Brown

et al. 2000). Moreover, since TPR and WD-repeat proteins are often found in association

with each other (Goebl and Yanagida 1991; Neer et al. 1994; van der Voorn and Ploegh

1992; Smith et al. 1999), a reasonable hypothesis is that the TPR domain of Ski3 interacts

with Ski8. Second, Ski2 has interactions with either the side or bottom of Ski8, which are stabilized by additional interactions between Ski2 and Ski3. The findings presented here is in consistent with the studies reported by Wang et al. (2005).

Ski8 is an interesting protein since it has been shown to be essential in two distinct cellular processes: mRNA metabolism and meiotic recombination. Moreover, emerging information shows that Ski8 plays fundamentally different roles in RNA metabolism and DSB formation during meiotic recombination. In 3’ to 5’ mRNA decay,

Ski8 localizes predominantly to the cytoplasm and interacts with Ski2 and Ski3 to form the Ski complex, thereby mediating the exosome-dependent mRNA decay. During meiotic recombination, Ski8 re-localizes from the cytoplasm to the nucleus and interacts with Spo11, thus either affecting the ability of Spo11 to bind DNA or acting in concert with Spo11 to recruit Rec102 and Rec104 to the chromosomes. WD-repeat proteins have been demonstrated to act as scaffolding or adaptor proteins to interact with multiple 92 protein partners and to carry out different roles. However, of the WD proteins characterized so far, only Ski8 shows extremely different functional roles in terms of non-overlapping protein partners, sub-cellular localization, and even different target substrates (RNA vs. DNA; Arora et al. 2004).

Related to the role of Ski8 in meiosis, Ski8 has been shown to interact with Spo11 in a two-hybrid system (Uetz et al. 2000; Arora et al. 2004). As discussed above, our results suggest that the interaction between Ski8 and Spo11 requires the top surface of the

Ski8 structure. Moreover, the results from Arora et al. (2004) showed that alanine substitutions for residues Gln-376, Arg-377 and Glu-378 in Spo11 both abolished the

Spo11-Ski8 two-hybrid interaction and disrupted meiotic DSB formation. Based on these observations, the two surfaces of Spo11 and Ski8 required for interaction have begun to be identified. Moreover, since the F59A mutation in Ski8 also affects Spo11 interaction, it seems likely that Spo11 also interacts with the side of the β propeller structure to some extent.

An intriguing issue is why Ski8 is involved in both mRNA decay and meiosis.

One possibility is that this bifunctional nature is simply an example of a protein being co- opted for an additional use through evolution. Alternatively, the use of Ski8 in meiosis could be a way of coordinating changes in mRNA turnover with the meiotic program.

This is potentially relevant because there are clear changes in mRNA decay rates during meiosis (Surosky et al. 1994). In addition, two observations raise the possibility that Ski8 function in mRNA decay might be inhibited during meiosis. First, because the Spo11 and

Ski3 binding sites both require the top surface of Ski8, the induction of Spo11 during meiosis might inhibit assembly of the Ski complex by titrating the available Ski8. In 93 addition, the translocation of Ski8 into the nucleus during meiosis would also be expected to limit its function in cytoplasmic mRNA decay. However, despite this intriguing connection, additional experiments will be required to test if there is a functional connection between the Ski8 role in mRNA decay and meiosis.

94

Chapter 4

Structural and Functional Analysis of Dhh1

4.1 Results

4.1.1 Structural overview of the Dhh1

Full length Dhh1 from Saccharomyces cerevisiae was initially expressed and

purified for crystallization. However, purified full length Dhh1 underwent substantial

proteolysis. Compared to its homologs in other organisms, and related DEAD-box

proteins, Dhh1 contains extensions of ~50 amino acids at both its N- and C-terminals,

which flank the conserved core domain (Figure 4-2). Given these extensions, an N- and

C-terminal truncated yeast Dhh1 (residues 30-425; tDhh1), which contains the entire helicase domain, has been expressed and purified to homogeneity. The structure of tDhh1 was determined by MAD phasing with Br derivative, which yielded an easily interpretable electron density map at a resolution of 2.1Å. The final model contains two disordered regions: residues 30-45 and residues 423-425. The details of structure determination and refinement have been described in Chapter 2.

The polypeptide chain of tDhh1 is folded into two α/β domains, each with a

RecA-like topology as observed in other DEAD-box proteins (Figure 4-1). The N- terminal domain (residues 46-247) is composed of a parallel seven-stranded β sheet flanked by four helices (α1-4) on one side and five helices on the other (α5-9). The N-

terminal domain is connected to the C-terminal domain by a short stretch of peptide

(residues 247-253). The C-terminal domain (residues 253-422) is also a parallel α/β 95 structure but with three helices (α10-12) on one side and two helices (α13-14) on the other, surrounding the central seven-stranded β sheet. Consistent with their basic RecA- like topology, the N- and C-terminal domains are similar to each other. Superposition of these two domains gives an r.m.s.d for 81 equivalent Cα atoms of 1.5Å. The N-terminal domain contains sequences motifs associated with ATP binding and hydrolysis while the

C-terminal domain contains motifs associated with RNA binding. The N-terminal domain interacts with the C-terminal domain through the previously identified conserved motifs and helix α12 from the C-terminal domain, thereby creating a large channel between two domains (Figure 4-1). Such a marked structural feature has not been seen in other

DEAD-box structure and may have important functional implications for Dhh1 (see below).

96

A B

IV a12 C- d o m ai n VI V Q I III II

Ia Ib N-domain

Figure 4-1 Orthogonal view of tDhh1. Ribbon diagram of tDhh1 with the N- and C-terminal domains colored in paleblue and lightgrey, respectively. The coloring scheme for conserved motifs are as follows: the Q-motif in red, I in green, Ia in yellow, Ib in purple, II in orange, III in blue, IV in cyan, the α-12 helix in greenyellow, V in navy and VI in magenta. The secondary structural elements are labeled. Figures 4-2, 4-3 and 4-4 were generated using MOLSCRIPT (Kraulis, 1991).

4.1.2 Structural Comparison

Comparison of the tDhh1 structure with the structures of eIF4A, mjDEAD, and

UAP56 reveals three significant points. First, as pointed out earlier, the common feature of the DEAD-box proteins is that the conserved region of these proteins is composed of only two α/β domains (Caruthers et al, 2000; Story et al, 2001). Comparison of tDhh1 with eIF4A and mjDEAD shows that the individual domain structures are similar among these proteins. Superposition of the N-terminal domain of tDhh1 with those of eIF4A, mjDEAD and UAP56 gives an r.m.s.d of 1.0Å, 1.3Å and 1.1 Å, respectively. When the

C-terminal domain is superimposed, the r.m.s.d values for tDhh1p vs. eIF4A, tDhh1 vs. 97

mjDEAD and tDhh1 vs. UAP56 are 1.3Å, 1.2Å and 1.1 Å, respectively. Thus, the

overall structure of each domain is conserved between different DEAD-box proteins.

A second point is that the relative orientation of the N- and C-terminal domains in these DEAD-box proteins is significantly different. Yeast eIF4A is a ‘dumbbell’

structure, having two domains connected to each other by an extended linker. mjDEAD is a compact molecule with two domains connected by a short linker. tDhh1p and UAP56 are more compact molecules with the two domains connected to each other by extensive interactions (see below). The orientation of the N and C terminal domains of Dhh1p is strikingly different in comparison to the other Superfamily 2 helicase structures. When the two N-terminal domains are superimposed, the C-terminal domain of eIF4A is rotated

50° with respect to the C-terminal domain of tDhh1 (Figure 4-3A), whereas the corresponding rotation involving mjDEAD is 30° (Figure 4-3B). Similarly, when the N- terminal domain of tDhh1 is superimposed with that of UAP56, the orientation of the C- terminal domain differs by 40°.

Finally, in addition to the differences in their relative domain orientations, there are local structural changes as well. In eIF4A, the N terminus forms a short antiparallel β strand which interacts with β1 and extends the seven-stranded β sheet to eight strands.

The corresponding region (residues 30-46) in tDhh1 is disordered. mjDEAD lacks a C- terminal region (residues 410-419 in Dhh1), which has different conformations in tDhh1 and eIF4A. Helix α5 and residues 199-205, which is a region immediately downstream of

DEAD motif, showed large positional shifts between tDhh1 and eIF4A and mjDEAD.

The conformation of the loop connecting the α10 helix and β-9 strand in the C-terminal domain of tDhh1 has changed dramatically compared to that of the corresponding regions 98

in both eIF4A and mjDEAD. Residues 90-95, 340-352 and 371-382, which respectively

correspond to motifs I, V and VI, showed larger conformational changes that may have

functional implications (see below).

4.1.3 Location of the conserved sequence motifs

Dhh1 contains nine conserved sequence motifs as observed in other DEAD-box proteins (Figures 4-1 and 4-2). Structural and functional studies of several DNA and

RNA helicases have suggested the general roles for some of these motifs (Caruthers &

McKay, 2002; Tanner & Linder, 2001). Motif I, or the Walker A motif (Walker et al,

1982), forms a loop structure (P loop) that accommodates the phosphate groups of ATP.

In tDhh1 structure, the P loop adopts an “open” conformation (Figure 4-3C) similar to

that observed in mjDEAD, but opposed to the “closed conformation” in the structures of

eIF4A and BstDEAD without bound ligand (Carmel & Matthews, 2004; Johnson &

McKay, 1999; Story et al, 2001). Such an open conformation of motif I would facilitate

ATP binding. Motifs Ia and Ib have been suggested to be involved in binding of RNA substrates (Caruthers & McKay, 2002), but their precise functional roles are not fully understood. The conformations of these two motifs in Dhh1p are very similar to those observed in eIF4A and mjDEAD (Figure 4-3C). 99

** Q I

Ia Ib

** II III

IV

** ** V VI

Figure 4-2 Sequence alignment of S. cerevisiae Dhh1, H. sapiens Rck/p54, S. cerevisiae eIF4A and M. janaschii mjDEAD. The secondary structures of tDhh1 are shown. Invariant residues are shown in white letters, similar residues in red and others in black. Residues mutated for in vivo and in vitro assays are indicated by *.

100

Motif II is also called the Walker B, or DEAD motif (Walker et al, 1982).

Structural, mutational and biochemical studies indicated that motif II forms interactions with the β- and γ-phosphates of ATP through Mg2+ and is required for ATP hydrolysis

(Benz et al, 1999; Caruthers & McKay, 2002; Pause & Sonenberg, 1992; Pause &

Sonenberg, 1993; Schmid & Linder, 1991). In tDhh1p, the conformation of the DEAD

motif is very similar to that in eIF4A or mjDEAD. However, a region immediately

downstream of the DEAD motif, residues 199-205, shows large conformational changes

with a positional shift of up to 8Å for equivalent Cα atom between tDhh1, and eIF4A and

mjDEAD (Figure 4-3C). Whether this difference is idiosyncratic to Dhh1, or reflects a

different basic conformation of the DEAD-box protein fold remains to be seen.

Motifs III and IV in tDhh1 are very similar structurally to those in mjDEAD and

eiF4A (Figure 4-3C & 4-3D). Motifs III and VI are proposed to link ATP binding and

hydrolysis with conformational changes required for helicase activity (Caruthers &

McKay, 2002). Similar to mjDEAD and eIF4A, the SAT motif in tDhh1p interacts with

the final aspartic acid of the DEAD motif providing a conserved connection between

motifs II and III. Motif IV is involved in single strand DNA (ssDNA) binding in other

helicases including PcrA and Rep, HCV (Caruthers & McKay, 2002; Kim et al, 1998). It

is suggested that two Arg residues in motif IV of eIF4A might interact with single strand

RNA (Caruthers & McKay, 2002).

Motif V in tDhh1 adopts a strikingly different conformation from those in mjDEAD

and eIF4A (Figure 4-3D). Superposition of motif V (residues 340-352) with those of

mjDEAD and eIF4A gives an r.m.s.d for 13 Cα atoms of 3.0Å and 3.2Å, respectively.

The conformation of motif V is similar to that of UAP56 without bound ADP, although 101

motif V is disordered in the structure of UAP56 with bound ADP (Shi et al., 2004). The

conformation of motif V in tDhh1 allows it to interact with motif I and the Q-motif, thus

linking the N- and C-terminal domains together (see below). This observation is similar

to the findings that motif V in the DNA helicases PcrA and Rep bridges the contacts

between domains 1A and 2A through interactions with motif II (Korolev et al, 1997;

Velankar et al, 1999). This raises the possibility that motif V might play a general role in

this class of proteins in bringing the two domains together. Interestingly, motif V also

interacts with the DNA substrate in PcrA and Rep, suggesting this motif can have

multiple roles and might couple substrate binding to domain interactions.

Structural studies on DNA helicases indicated that motif VI is involved in contacts with

the γ phosphate of ATP (Caruthers & McKay, 2002). Mutational and biochemical studies

suggested that motif VI in eIF4A is required for RNA binding and ATP hydrolysis (Pause

et al, 1993). In tDhh1, the conformation of motif VI is dramatically different from that in both mjDEAD and UAP56, while the same region is disordered in eIF4A (Figure 4-3D).

In mjDEAD, the Q-motif, motifs I and II in the N-domain and motif VI in C-domain, form part of the ATP-binding site between these domains (Story et al, 2001). The second

Arg residue in motif VI (HRxGRxGR) of DEAD-box proteins is proposed to perform a similar function as the “arginine finger” in G-proteins by sensing the γ-phosphate of the

ATP (Caruthers & McKay, 2002). Although it seems unlikely that motif VI of tDhh1p could make contact with the γ phosphate of ATP, it does interact with motif V (see below). Such interaction may push motif V forward for contacting motifs in the N- terminal domain. 102

A B

C-d o m ai n

N-domain

C

N-domain

D

C-d o m ai n

a12 a12

Figure 4-3 Comparison of tDhh1 with eIF4A and mjDEAD. (A) Superposition of the N-terminal domain of tDhh1 with that of eIF4A. tDhh1 is shown in lightgrey and eIF4A in green. The view of tDhh1 is as in Figure 4-1A. (B) Superposition of the N-terminal domain of tDhh1 with that of mjDEAD (magenta). (C) Stereo view of the superposition of the N-terminal domain of tDhh1 with those of eIF4A and mjDEAD. tDhh1p is shown in lightgrey, eIF4A in lightgreen and mjDEAD in lightpink. (D) Stereo view of the superposition of the C-terminal domain of tDhh1 with those of eIF4A and mjDEAD.

103

The Q-motif is a newly identified motif and has been suggested to be involved in

regulation of ATP binding by modulating the state of the P loop (Tanner et al, 2003).

More recently, it has been shown that the Q-motif regulated RNA-binding and helicase

activity (Cordin et al, 2004). In tDhh1, the Q-motif interacts extensively with the α12 helix and motif V from the C-terminal, thus contributing part of the interactions between the N- and C-terminal domains (see below). Furthermore, structural comparison shows that the conformation of this motif in tDhh1p is similar to that in UAP56, but different from those in mjDEAD and eIF4A with deviations of the equivalent Cα atoms being as large as 2.0 Å (Figure 4-3C).

4.1.4 Interactions of the conserved Motifs

An important issue in understanding the DEAD-box proteins is to understand how the conserved motifs function to couple ATP hydrolysis to perform work. Thus, a goal in gaining insight into helicase function is to understand the structural and dynamic interactions of the conserved motifs, particularly between the two different domains of the proteins. The possibility of dynamic interactions is suggested by multiple lines of evidence indicating that at least some of the DEAD-box proteins undergo conformational changes in response to substrate and/or ATP binding. For example, ATP and RNA binding have been found to change the protease digestion pattern in eIF4A and Dhh1 (see below) (Lorsch & Herschlag, 1998b). Moreover, the structures of PcrA and Rep (Korolev et al, 1997; Velankar et al, 1999) showed that these helicases contain four domains including two α/β domains (domains 1A and 2A), which are tightly packed against each other and correspond to the N- and C-terminal domains observed in DEAD-box proteins. 104

Furthermore, the cleft between the two α/β domains closes upon ATP/DNA binding.

Interestingly, the canonical helicase motifs in PcrA and Rep are close to each other and are clustered in the cleft between the two α/β domains with motif V of domain 2A interacting with motif II in domain 1A, suggesting that the conserved motifs modulate the interaction between the two domains (Korolev et al, 1997; Velankar et al, 1999).

Consistent with this possibility, in the structure of the related hepatitis C virus (HCV) helicase, all eight conserved motifs are found in a cleft between the two α/β domains

(Kim et al, 1998).

In the eIF4A and mjDEAD structures there are few interactions between the conserved motifs within different protein domains. The distant locations of the N- and C- terminal domains in the crystal structure of eIF4A prevent interactions between the two domains (Caruthers et al, 2000). However, it has been suggested that ATP and/or RNA binding would induce a dramatic conformational changes in eIF4A, bringing the two domains as well as the conserved motifs closer together (Caruthers et al, 2000).

Although mjDEAD has a compact two-domain structure, the two domains are in an open conformation and no motif interactions have been observed (Story et al, 2001).

In tDhh1p, the N- and C-terminal domains share an extensive interface similar to the two α/β domains in PcrA and Rep (Korolev et al, 1997; Velankar et al, 1999). The interaction between the N- and C-terminal domains in tDhh1 buries a pairwise accessible surface area of 1601.5Å2. Close inspection of the tDhh1 structure shows several of the conserved motifs are clustered in the cleft between the N- and C-terminal domains with motif V and the α12 helix in the C-terminal domain and motif I and the Q-motif in the N- terminal domain mediating the inter-domain interactions (Figure 4-1). Specifically, as 105 shown in Figure 4-4, the backbone carbonyl oxygen atoms of both Leu342 and Thr344 in motif V are hydrogen-bonded to the NZ atom of Lys91 in motif I while the side-chain of

Arg345 in motif V stacks against that of Arg89 in motif I. The side-chain and the main- chain carbonyl oxygen of Leu343 in motif V make Van der Waals (VDW) contacts with the main-chain carbonyl oxygen of Gly93 and the side-chain of Thr94 in motif I, respectively. Furthermore, the side-chains of Leu343 and Thr344 make extensive hydrophobic interactions with Pro71 in the Q-motif. Helix α12 is situated in close proximity with the Q-motif and the putative ATP-binding pocket, thereby interacting with both the Q-motif and motif I via multiple contacts. Specifically, residues Arg330 and Phe326 in the α12 helix form a salt bridge and stack against with Glu74 and Pro71 in the Q-motif, respectively while His327 in the α12 helix stacks against the methylene group of Lys68 in the Q-motif. Moreover, residues Gln319, Gln320, Asn323 and His327 in helix α12 contact residues Gly95, Phe66, Ser70 and Pro69 in the Q-motif, respectively through multiple VDW interactions. Interestingly, helix α12 also contacts the Q-motif in the structure of UAP56, suggesting this interaction will be common to multiple members of the Superfamily 2 (Shi et al., 2004).

Apart from the interactions with motifs I and Q, motif V also interacts with motif

VI and the α12 helix. For example, the main-chain carbonyl oxygen of Ile347 is hydrogen-bonded to the NH2 group of Arg376 in motif VI while Gln350 interacts with

R376 in motif VI. Inspection of the interaction between motif V and the α12 helix revealed that Ile349 makes extensive hydrophobic interactions with Phe326 and Phe329 of the α12 helix, and Asp341 forms a salt bridge with residue Arg322 in the α12 helix. 106

An additional contact is made by the side-chain of residue Leu343 of motif V stacking

against the methylene group of Arg322 in helix α12.

The interactions between the conserved motifs are unique features of tDhh1 and

correlate with the novel domain arrangement and conformational changes in motifs I, V,

and VI in the tDhh1 structure. This observation is similar to the findings that motif V in

the DNA helicases PcrA and Rep bridges the contacts between domains 1A and 2A

through interactions with motif II (Velenkar et al., 1999; Korolev et al., 1997). Similar

but weak interactions between motifs V and I have been observed in UAP56 (Shi et al.,

2004). This raises the possibility that motif V of the Superfamily 2 protein family might play a general role in bringing the ATPase domain and the RNA-binding domain into close proximity. This would facilitate the coupling of ATP-binding and hydrolysis with the RNA specific functions. However, it is currently unclear if the unique features of the tDhh1p structure represent specific aspects of Dhh1p function, or represent a more general conformation that many DEAD-box proteins adopt at some stage of their conformational changes. This will be resolved as more structures of other DEAD-box proteins, and Dhh1p in different conformations are reported.

a12 a12

Figure 4-4 Stereo view of interactions of the conserved motifs. Motifs Q, I, V, VI and the α12 helix are colored as in Figure 2. Residues involved in the interactions are labeled and shown in stick models. Hydrogen bonds are shown in broken lines.

107

4.1.5 Identification of residues required for RNA binding

DEAD-box proteins are generally capable of RNA binding. To identify potential surfaces for RNA binding on Dhh1, the electrostatic potential was mapped on the molecular surface of tDhh1. This analysis revealed a prominent positively charged patch on the molecular surface of tDhh1 (Figure 4-6A). Part of the patch occupies the cleft between the N- and C-terminal domains with the rest of the patch running across the C- terminal domain. Motifs I, V and VI contribute amino acids side chains forming this patch. Strikingly, the residues that form this positive surface are highly conserved among

Dhh1 homologs, suggesting that this region might represent a conserved RNA binding pocket in these family members. To test whether this region is involved in RNA binding and is functionally important, mutations of several of the amino acids in this region to alanines (Figure 4-6B) has been generated and their effect on RNA binding in vitro and protein function in vivo has been performed.

To examine the RNA binding in vitro, the mutant tDhh1 proteins were expressed in E. coli and purified. The yield for each mutant protein was comparable to that of the wild type protein. Moreover, CD spectroscopy gave the same spectrum for wild type each mutant protein (Figure 4-5). These observations indicate that these mutations had no significant effect on the general folding or stability of the Dhh1 protein. Wild-type and mutant tDhh1 proteins were examined for their binding, in the presence or absence of

ATP, to a single-stranded poly(U) RNA oligo (12mer), a hairpin dsRNA (dsRNA-I), and a hairpin dsRNA with the 11-bases 5’-single-stranded tail (ss/dsRNA-II) (Figure 4-6C).

In the absence of ATP, wild-type tDhh1 binds to all three RNA substrates and the addition of ATP to the RNA binding buffer had no effect on RNA binding. This 108 observation implies that RNA binding to Dhh1 is not stimulated by ATP binding, at least under these biochemical conditions. The ability of Dhh1 to bind RNA independent of

ATP is similar to the interaction of eIF4A with RNA, which can also occurs independent of ATP, although ATP can enhance the eIF4A interaction with RNA (Lorsch and

Herschlag, 1998a).

Figure 4-5 CD spectroscopy of various single muants, double mutants and wild-type Dhh1. Protein samples were diluted in 50 mM NaCl, pH 6.0 at a concentration of 4 µM. Data were collected at a resolution of 0.1 nm and a bandwidth of 2 nm, and five spectra were averaged. The spectra of mutation derivatives are completely same as that of wild-type Dhh1.

109

A B

A 6 A 4 1 3 K302 9 ,G ,K A A A A A A A R2 9 5 A A A 5 6 5 5 6 9 0 9 9 1 9 9 4 4 4 6 7 8 8 9 1 1 3 3 3 3 3 R R K D E R R G H R K362 R3 7 0 H369 K91 Q IIaIbIIIII IVVVI R32 2 R3 45 R89 N-Domain C-Domain

E196 D195

C G 5' GGAACCUAGCAGC U dsRNA-I 3' CCUUGGAUCGUCG A A

G U 3' CGCUGUCGGAGC ss/ dsRNA-II 5' GGUCAUUAAGAGCGACAGCCUCG A A

D 12mer polyU

dsRNA-I

ss/ dsRNA-II

A T A A A A A A A A A A S 1 9 1 5 6 5 6 9 0 W 6 8 9 9 9 4 6 B 9 4 4 7 K R K 1 1 3 3 3 3 , 3 D E R G H R A ,G 9 A 8 5 R 4 3 R

E 120 12mer polyU dsRNA-I 100 ss/ dsRNA-II

80

60

40 % of RNA binding of RNA % 20

0 T A A A A A A A A A A 1 6 6 9 5 5 6 9 0 W 1 8 9 9 9 4 4 6 7 9 4 K 1 3 R 1 3 3 3 3 ,K D E R G H R A ,G 9 A 8 5 R 4 3 R

Figure 4-6 In vitro RNA binding assay of Dhh1. (A) Solvent accessible and electrostatic potential of tDhh1 viewed as in Figure 4-2A showing a prominent RNA binding channel. Residues located in the putative RNA-binding channel, and Asp195 and Glu196 are labeled. (B) Schematic diagram showing the position of the various point mutants in Dhh1. (C) Chart of the two RNA substrates: dsRNA-I and ss/dsRNA-II. (D) Effects of mutations of tDhh1 on RNA-binding in vitro. BSA serves as a negative control. (E) Quantitation of RNA binding to wild-type and mutant tDhh1 proteins using three different RNA substrates. The percentage of RNA binding activity of each mutant relative to that of wild type Dhh1 is shown. 110

An important result is that several changes in the putative RNA binding channel

significantly reduced RNA binding compared to the wild-type protein. Specifically, Ala

mutations of Arg370 in motif VI, and Arg89 or Lys91 or both in motif I drastically reduced binding to all three RNA substrates (Figures 4-6D and 4-6E). Mutation of these

residues to Ala would weaken the positively-charged potential on the molecular surface

of tDhh1, thereby potentially reducing the binding of RNA. It was also observed that

mutations of Asp195 and Glu196 in the DEAD motif to Ala (D195A and E196A)

increased RNA binding compared to the wild-type protein (Figures 4-6D and 4-6E).

These residues are located in the vicinity of the RNA binding channel, and mutation of

these negatively-charged residues to Ala might thereby facilitate the RNA binding by

reducing charge-charge repulsion. These results demonstrate that this region of Dhh1 is

required for RNA binding. The simplest interpretation is that this positive charged region

directly interacts with the RNA substrates. However, until a co-crystal of Dhh1 with

RNA bound is analyzed, it is possible that these mutations indirectly affect RNA binding by influencing conformational changes in the protein.

The effects of mutations in motif V on RNA binding have also been examined.

Individual substitutions of Arg345 and Gly346 by Ala (R345A and G346A) did not lead to substantial defects in RNA binding (Figures 4-6D and 4-6E). However, altering

Arg345 and Gly346 together (R345A G346A) dramatically decreased RNA binding

(Figures 4-6D and 4-6E). These results indicate that defects in motif V can affect RNA binding. Because multiple changes are required, it was supposed that this effect may be caused by the combination of changes in the RNA binding channel and conformational changes in Dhh1. 111

To determine how these biochemical defects in tDhh1 are related to in vivo

function, the tDhh1 variant proteins were examined in yeast for their effects on cell

growth and mRNA turnover in both a wild-type and dhh1∆ strain. In these experiments, the variant Dhh1 are expressed in yeast under the control of their own promoter from single copy plasmids. It has been observed that single mutants of D195A, E196A, and

R370A, and double mutants of R345A and G346A (R345A G346A), and R89A and

K91A (R89A K91A), which all altered RNA binding in vitro (Figures 4-6D and 4-6E), failed to complement the temperature sensitivity of the dhh1∆ strain (Figure 4-7A). For

R345A G346A double mutant, the loss of the ability to complement a dhh1∆ mutant was not specific to either Arg345 or Gly346, as similar results were seen with both R345A and G346A (Figure 4-7A).

The growth phenotypes of the Dhh1 mutant in vivo directly correlated with defects in mRNA turnover, which were determined by examining the levels of the EDC1 mRNA, which is sensitive to the loss of Dhh1 (Muhlrad and Parker, 2005). In vivo studies show that mutants of D195A, E196A, R370A, and double mutants of R345A and

G346A (R345A G346A), and R89A and K91A (R89A K91A) led to the accumulation of

EDC1 mRNA in a dhh1∆ strain (Figures 4-7B and 4-7C). Interestingly, individual substitution of Arg89 and Lys91 to Ala seems to be partially defective in both RNA binding and growth phenotype, and double mutation of these two residues (R89A K91A) is more extreme due to the additive effects of the singles (Figures 4-6 and 4-7). In combination, these results indicated a strong correlation between the in vitro defects in

RNA binding and function of Dhh1 in vivo. The simplest interpretation of these observations is that RNA binding is required for Dhh1p function. 112

It has also been observed that the double mutants of R345A and G346A (R345A

G346A), and R89A and K91A (R89A K91A) caused a dominant negative phenotype and

caused death of wild-type cells at 37°C (Figure 4-7A). For the combination of R345A

and G346A, the dominant negative effects are solely due to G346A, as R345A did not by

itself show a dominant negative phenotype. The dominant negative phenotype of R89A

and K91A requires both mutations as single mutants of both R89A and K91A have no

effect on Dhh1 activity in wild type cells both by growth and RNA decay analysis

(Figure 4-7). Consistent with the dominant negative growth phenotype, double mutants of R345A and G346A (R345A G346A), and R89A and K91A (R89A K91A), and G346A also led to an increase in EDC1 mRNA levels even in a wild-type strain (Figures 4-7B and 4-7C). Since G346A confers a dominant negative phenotype, yet still binds RNA, and mutants R89A, K91A and R370A fail to bind RNA and are not dominant negative, the dominant negative phenotype is not simply due to the inability to bind RNA. Thus, it has been suggested that the alterations seen in G346A and R89A K91A double mutant

alter the function of Dhh1 by locking the protein in a dominant negative conformation.

Both of these changes would affect the structure of the interface between the two

domains in tDhh1. For example, the presence of an alanine at position 346 would be

expected to interfere with motif VI moving close to and interacting with motif V. This

steric interference would be expected to prevent Dhh1 from adopting the closed

conformation seen in the structure. Similarly, substitutions of both Arg89 and Lys91 for

Ala would reduce the specific contacts between the ATPase and RNA-binding domains

of Dhh1. 113

Figure 4-7 In vivo mutagenesis analysis of Dhh1. (A) Shown are the growth of yeast cells expressing various Dhh1 mutants in either a dhh1∆ strain or wild-type (WT) background at 30°C or 37°C. (B) Shown is the effect of the various Dhh1 mutants on the steady-state accumulation of the EDC1 mRNA in a dhh1∆ strain or wild-type (WT) background. (C) Relative abundance of the EDC1 mRNA for the various Dhh1p mutants to that of the wild type Dhh1p in a dhh1∆ strain or wild-type (WT) background.

114

4.1.6 Conformational changes in Dhh1

DEAD-box proteins are proposed to use the energy from ATP hydrolysis to rearrange inter- or intra-molecular RNA structures or disrupt RNA-protein interactions

(Rocak & Linder, 2004; Tanner & Linder, 2001). In tDhh1, the interactions of motif V

with motif I, the Q-motif, links the C-terminal domain to the N-terminal domain, thereby

providing a mechanism for coupling ATP binding and hydrolysis with RNA-specific

activities, probably through conformational changes. To test whether Dhh1 undergoes

conformational changes upon ATP and/or RNA binding, the trypsin cleavage pattern of tDhh1 in the presence of ATP and /or RNA has been analyzed.

Evidence that Dhh1 undergoes a conformational change was that the cleavage of tDhh1 by trypsin was slowed in the presence of ATP, whereas RNA alone had little, if any, effect on the cleavage of tDhh1 (Figure 4-8). This implies that tDhh1 probably undergoes a significant conformational change upon ATP binding rather than RNA binding. In addition, ADP and AMP-PNP failed to confer trypsin resistance (Figure 4-8).

In the presence of both ATP and RNA, Dhh1 conferred strong trypsin resistance to such an extent that very little tDhh1 was cleaved even after one hour of incubation with trypsin

(Figure 4-8). These observations provide evidence for conformational changes in tDhh1p and indicate that the most protease resistance conformation forms in the presence of both

ATP and RNA. A similar conformational change of eIF4A has also been revealed by trypsin digestion (Lorsch and Hershlag, 1998b); while no biochemical studies have been performed on mjDEAD protein.

A reasonable hypothesis is that the protease resistant form of Dhh1 is most likely a compact structure similar to the observed structure. This predicts that alterations to 115

Dhh1 that would prevent the closed conformation might prevent protease resistance in the presence of ATP and RNA. In order to examine this prediction, the protease sensitivity of the double mutant with the R345A and G346A mutations has been investigated. This double mutant would disrupt the close packing of the two domains due to the substitution of G346 with the larger alanine residue. Trypsin digestion showed that this mutant protein did not change its protease sensitivity with added RNA and ATP (Figure 4-8).

This suggests that this protein is unable to efficiently form the protected conformation of

Dhh1 and has implications for the function of Dhh1 (see below).

tDhh1p R345A G346A

0' 5' 10' 15' 20' 30' 60' 0' 5' 10' 15' 20' 30' 60'

No ligand No ligand

+ATP +ATP +RNA

+AMP-PNP

+ADP

+RNA

+ATP +RNA

Figure 4-8 Analysis of limited trypsin digestion of Dhh1. SDS-PAGE analysis of time courses for trypsin cleavage of wild type tDhh1 (A) or the R345A, G346A mutant (B) in the absence of ligands or presence of saturating concentrations of ATP, AMP-PNP, ADP, and 12mer poly(U) or their combinations. The position of the undigested tDhh1p is indicated by an arrow. 116

4.2 Discussion

The crystal structure of tDhh1 revealed that tDhh1 is generally similar to eIF4A,

mjDEAD and UAP56. However, in Dhh1 the two domains are arranged in a unique

manner and some of the conserved motifs show different structures than seen in earlier

structures. One striking structural feature of tDhh1 is that motif V interacts with motif I and the Q-motif, thereby connecting the two domains. Since at least some mutations altering this interaction (R89A K91A and G346A), cause both loss of function phenotypes and confer dominant negative phenotypes, this interaction appears to be functionally important. Moreover, because motif V also comprises and interacts with motif VI to create the RNA binding site, motif V therefore may function in connecting the ATP binding site and the RNA binding site together. This is analogous, although the molecular details differ, to the finding that motif V “bridges” the ATP-binding site and the oligonucleotide-binding site in DNA helicases, suggesting that motif V may play a general role in this class of proteins in coordinating domain interactions affected by ATP and substrate binding. Consistent with this view, in the structure of UAP56, the two helicase domains also contact each other through interactions between motifs V and I, and helix α12 and the Q-motif (Shi et al., 2004), but these interactions are not as extensive as those observed in tDhh1.

Analysis of the electrostatic potential on the molecular surface of tDhh1combined with mutagenesis identified a likely RNA binding region, which is primarily composed of motifs I, V and VI. An interesting feature of this binding surface is that it is comprised of residues from both the N- and C-terminal domains of Dhh1. This fact dictates that the

RNA binding surface will be different dependent on the orientation of the two domains. 117

Given this, it has been hypothesized that the nucleotide effects on RNA binding in

DEAD-box proteins might generally be that ATP binding generates a close spatial

arrangement of the two domains that forms an RNA binding surface from residues in each domain. Future experiments to examine a Dhh1-RNA co-crystal should shed light

on this issue.

Evidence that Dhh1 undergoes a conformational change has come from

examination of the trypsin cleavage pattern in the presence of nucleotide and/or RNA.

This process is likely to occur in two steps, but probably without an obligate order in a

manner similar to what has been seen with eIF4A (Lorsch and Hershlag, 1998b). In one

step, ATP binding would induce a conformational change of tDhh1. In a second step,

RNA binding, coupled with ATP binding, would induce a further conformational change.

Such conformational changes are facilitated by the inter-domain flexibility of DEAD-box

proteins. In tDhh1, the open conformation of motif I may facilitate ATP binding. This

would lead to large scale movements in the protein aided by specific interactions between

motif V, and motif I and the Q-motif which bring the ATP binding and RNA binding

domains into close proximity. Such domain rearrangement may also create the optimal

RNA binding surface by driving conformational transitions in the motif VI region,

thereby exposing the Arg residues in motif VI to the RNA binding channel. In support of

this notion, mutations of any one of the three Arg residues in motif VI drastically reduce

eIF4A binding to RNA (Pause et al, 1993). Taken together, these results suggest a model

of Dhh1 function involving conformational changes, ATP binding and hydrolysis, which

facilitate a cycle of interactions between the conserved motifs and alter the nature of the

RNA binding surface. 118

Additional evidence that a conformational change in Dhh1 is required for function

has come from the analysis of mutations at the interface between the two domains. Most

strikingly, mutations of either G346 or K91A and R89A, lead to loss of function and

dominant negative phenotypes. These residues are located at the junction between the two

domains (Figure 4-9A) and would be predicted to block the formation of the closed

structure due to steric hindrance (in the case of G346A), or to weaken the closed

conformation by loss of interactions between the two domains (in the case of the K91A

R89A mutant). Consistent with this, a Dhh1 variant containing the G346A proteins is defective in entering the protease resistant conformation. Interestingly, this G in motif V is highly conserved in Superfamily 2 helicases. In contrast, the residues analogous to

Lys89 and Arg91 are not generally conserved in Superfamily 2 members and, for example, have been replaced by glutamine at these positions in eIF-4A (Figure 4-9B).

Given this, it seems that the G346 residue may be a critical residue in allowing the

adoption of the closed conformation due to its small size in Superfamily 2 members in

general, although the molecular details of the interface will differ between different

proteins.

119

A B

G322

G346 R3 2 1 R345 K91 Q66

R8 9 Q64

Dhh1p eIF4A

Figure 4-9 Comparison of Dhh1 to eIF-4A. (A) Shown is the position of Lys 89, Arg 91, and Gly 346. These residues span the junction between the N-terminal and C-terminal domains and appear to mediate the interaction between these two domains. (B) Also shown for comparison is the relative position of these residues in the eIF-4A structure. Here Dhh1 R89 and K91 are replaced by Q64 and Q66 in eIF-4A, while G346 is conserved in eIF-4A at position 322.

120

Chapter 5

Structural and Functional Insights into hUpf1

5.1 Results

5.1.1 Overall structure

The helicase domain of hUpf1 (residues 295-914, designated as hUpf1hd, Figure

5-1A) was expressed in E. coli and purified to homogeneity. The structure determination

of hUpf1hd-PO4 and in complex with AMPPNP, ATPγS and ADP has been described in

Chapter 2. Residues 584-587 are disordered in the structure of hUpf1hd-AMPPNP. The

ADP-bound complex (hUpf1hd-ADP) contains three disordered regions: residues 349-

355, 583-586 and 844-849 for both molecules in the asymmetric unit. Since no substantial differences are observed between two subunits (r.m.s.d of 0.6Å for all the equivalent Cα atoms), all subsequent analyses are based on subunit A. The structure of phosphate-bound hUpf1hd (hUpf1hd-PO4) contains four disordered regions: residues

337-338, 367-373, 401-405 and 584-587.

In the AMPPNP, ADP, and PO4 form, hUpf1hd is composed of two domains

(domains 1 and 2) (Figure 5-2A). Both of these two domains contain a “RecA-like” α/β domain designated as 1A and 2A, respectively. In addition to the subdomain 1A, domain

1 also contains two insertions, each of which forms a distinct subdomain, denoted as domains 1B and 1C, respectively. Domain 1A consists of residues 295-324, 415-555 and

610-700 and forms a parallel seven-stranded β-sheet surrounded by seven helices on one

side and three helices on the other. Domain 2A contains a parallel six-stranded β-sheet

flanked by three helices on both sides. The seven classical sequence motifs of 121

Superfamily 1/2 helicases are located in these two domains with motifs I, Ia, II and III in domain 1A and motifs IV, V and VI in domain 2A (Figure 5-1B). The nucleotide binding site is located in a deep cleft separating domains 1A and 2A. The core RecA-like architecture of hUpf1hd resembles that found in Superfamily 1 DNA helicase PcrA

(Velankar et al, 1999) and Superfamily 2 RNA helicase Vasa (Sengoku et al, 2006).

Superposition of these two domains (1A and 2A) with those of PcrA (Velankar et al,

1999) with bound ADPNP and dsDNA and Vasa with bound ssRNA and AMPPNP, gives an r.m.s.d of 1.5Å and 1.9Å, for 212 and 149 equivalent Cα atoms, respectively

(Figures 5-2B and 5-2C). This suggests that the relative orientations of domains 1A and

2A are largely conserved among these helicases with hUpf1 being more similar to PcrA.

Subdomains 1B and 1C are structurally unique domains for hUpf1 and not closely related to equivalent regions in any of the previously determined helicase structures.

Domain 1B, residues 325-414, is a β-barrel consisting of six anti-paralleled β strands that is located above the interface between domains 1A and 2A (Figure 5-2A). Domain 1C contains three helices formed by residues 556-609 and is situated on domain 1A, making a few contacts with domain 1B (Figure 5-2A).

122

Figure 5-1 Domain structure of hUpf1 and sequence alignment of the helicase core domain. (A) Schematic diagram showing the domain organization of hUpf1. (B) Sequence alignment of the helicase core domains of human and yeast Upf1 proteins. Conserved helicase motifs (I, II, III, IV, V, VI and VII) are marked. Mutated residues are marked with “*”. Secondary structural elements of hUpf1hd are indicated. The loop 349-355 is shown in red. The coloring scheme for domains as in (A).

123

Figure 5-2 Structure of hUpf1hd and its comparison with PcrA and Vasa helicases. The ribbon diagrams are drawn with domains 1A (pink) and 2A (wheat) in the same orientation. Bound nucleotide is shown in stick model and Mg2+ ion in purple sphere. (A) hUpf1hd-AMPPNP. Domains 1B and 1C are colored in lime and slate, respectively. (B) The PcrA/ADPNP/dsDNA complex. Domains 1B and 2B are shown in lime and yellow respectively. Bound DNA is shown in orange cartoon tube. (C) The Vasa/AMPPNP/ssRNA complex. Bound ssRNA is shown in purple cartoon tube.

124

5.1.2 Nucleotide Binding site and ATP hydrolysis

AMPPNP is bound in the pocket between domains 1A and 2A (Figure 5-2A).

The interactions of AMPPNP with the protein residues are similar to those observed in the PcrA/DNA/ADPNP complex (Velankar et al, 1999) with some notable differences.

As shown in Figure 5-3A, the adenine base is held in a hydrophobic pocket between

Tyr702 and Pro469. Tyr702 is structurally equivalent to Tyr286 of PcrA while Pro469 has no structural counterpart in PcrA. Val500 contacts the ribose ring via hydrophobic interactions. In contrast, Arg39 of PcrA, which corresponds to Val500 in hUpf1, interacts with the ribose ring of ATP through its methylene group. Four residues in hUpf1hd make direct contacts with the γ-phosphate of AMPPNP, namely Lys498, Gln665, Arg703,

Arg865 from conserved motifs I, III, IV and VI, respectively. In contrast, in PcrA, only three residues Gln254, Arg287, and Arg610 interact directly with the γ-phosphate of the nucleotide with Lys37 (Lys498 in hUpf1) contacting the β-phosphate. In PcrA, Arg610 acts as “arginine finger” which has been proposed to stabilize the transition state, thus facilitating ATP hydrolysis (Caruthers & McKay, 2002; Velankar et al, 1999). Residue

Arg610 in PcrA corresponds to Arg865 in Upf1p, which has a similar interaction with the

γ-phosphate and might fulfill the same role. In addition, Gln665 in hUpf1hd appears to act as “γ-phosphate sensor”, similar to Glu194 in RecA, which has been proposed to detect the presence or absence of the γ-phosphate of ATP and relay the information to other sites within the molecules through conformational changes (Story & Steitz, 1992).

Full-length hUpf1 (hUpf1-FL) has been shown to have RNA-dependent ATPase activity (Bhattacharya et al, 2000). In order to find out which residues are critical for

ATP hydrolysis, several residues in the AMPPNP binding pocket were mutated to alanine 125

and their effects on RNA-dependent ATPase activity were examined. Similar to hUpf1-

FL, hUpf1hd has RNA-dependent ATPase activity. However, the best ATPase activity

of hUpf1hd was obtained in the presence of poly(C) while the optimal ATPase activity

was obtained in the presence of poly(dT) for hUpf1-FL (Figure 5-3B; Bhattacharya et al,

2000). Consistent with previous results of both yeast and human Upf1p, the mutants

K498A and DE636AA were strongly defective for ATPase activity (Weng et al., 1996;

Bhattacharya et al., 2000) (Figures 5-3C). In addition, mutation of Gln665, Arg865, or

Arg703 to Ala nearly eliminated the ATPase activity of hUpf1hd (Figures 5-3C). In

order to identify whether the defects of ATPase are due to the incapability of ATP

binding to these mutants, the ATP binding assay was also performed. It showed that the mutants Q665A and DE636AA show the similar ATP binding activity as the wild-type

protein while mutations of Lys498, Arg703 and Arg865 were strongly defective for ATP

binding (Figure 5-3D), suggesting that residues K498, R703 and R865 are involved

mainly in ATP binding while the remaining residues play a critical role in ATP

hydrolysis. This result is similar to the observation that mutation of these three residues

in PcrA (Gln254, Arg287, and Arg610; Figure 5-3A) to Ala impaired ATPase activity

(Velankar et al, 1999). In line with the observation that mutation R865A abolished the

ATPase activity of hUpf1hd, the TR800AA form of yeast Upf1p in which both Thr800 and Arg801 (Arg865 in hUpf1) were mutated to Ala, was defective in RNA-dependent

ATPase activity (Weng et al., 1996). These data suggest that the Superfamily 1 helicases share a similar ATP binding and hydrolysis mechanism regardless of their DNA or RNA substrates. 126

To test if these residues were required for Upf1p function in vivo, the corresponding amino acids of Upf1 in S. cereviase Upf1p were mutated to alanine and their effect on NMD in yeast cells were examined by investigating the steady state levels of the CYH2 pre-mRNA, which is normally rapidly degraded and thereby reduced in level by NMD (He et al., 1993). Replacement with alanine of Gln601, Arg639, and

Arg801, which are analogous to Gln665, Arg703, and Arg865 in hUpf1 respectively, prevented the function of Upf1p in NMD and led to the accumulation of CYH2 pre- mRNA (Figure 5-3E). This observation indicates that these residues are also required for Upf1p function in vivo.

127

Figure 5-3 Nucleotide binding and hydrolysis of hUpf1hd. (A) Stereo view of superposition of the nucleotide binding sites of hUpf1hd-AMPPNP and PcrA. hUpf1hd-AMPPNP and PcrA in the PcrA/ADPNP/dsDNA complex are colored in pink and orange, respectively. (B) ATPase activities of wild-type proteins (200nM) in the presence of 500nM 15mer poly(A), poly(G), poly(C) and poly(U). (C) ATPase activities of wild-type and mutant hUpf1hd proteins (200nM) in the presence of 500nM 15mer poly(C). (D) ATP binding activities of wild-type and mutants hUpf1hd proteins (15ug) (B). (E) Northern analysis of the yeast CYH2 mRNA from strains expressing UPF1 mutant plasmids. mRNA from a UPF1 deletion strain containing plasmids expressing deletions (1B∆ or 1C∆), empty vector (UPF1∆), wild-type (WT), or point mutations as listed above each lane. The upper panel shows the unspliced precursor (upper band) and spliced CYH2 mRNA (lower band). The % of unspliced mRNA for each strain is given under each lane. The lower panel shows the 7S RNA used for standardization. 128

5.1.3 Conformational change during the Upf1 ATPase cycle

The coupling of conformational changes to NTP binding and hydrolysis is a general mechanism by which helicases and motor proteins catalyze physical transitions.

In order to figure out the conformational changes during the ATPase cycle of hUpf1, the structural comparison of the hUpf1hd-PO4, hUpf1-AMPPNP and hUpf1-ADP was performed. The comparison shows that all four individual domains are similar to one another among the three crystal structures (PO4, ADP- and AMPPNP-bound form) with mean pair-wise Cα r.m.s.d. about 0.5Å. Despite these similarities of the individual

domains, the relative orientation of domain 2A with respect to domain 1A differs

significantly among all three states of hUpf1hd. When domain 1A of hUpf1hd-ADP is

superimposed with that of hUpf1hd-PO4, the orientations of domain 2A differ by 14°

(Figure 5-4A). Similarly, when the corresponding domains of hUpf1hd-PO4 and

hUpf1hd-AMPPNP are compared to each other, the orientations of domain 2A differ by

10° (Figure 5-4B).

Inspection of the structures shows that the large-scale rigid-body movement of

domain 2A relative to domain 1A is caused by closure of the cleft between domains 1A

and 2A induced by nucleotide binding. In the PO4 structure, a phosphate ion in the

nucleotide binding pocket occupies the position equivalent to the γ-phosphate of

AMPPNP and interacts with Gln665, Arg703 and Arg865 (Figure 5-4C). These

interactions thereby contribute to the phosphate bound structure being more similar to the

AMPPNP bound structure.

Comparison of the phosphate and AMPPNP bound states illustrates that the presence of the nucleotide and/or γ-phosphate stabilizes a network of interactions 129 between domains 1A and 2A. Upon AMPPNP binding, motif V (residues 827-835) in domain 2A is positioned ~2Å closer to domain 1A, thus narrowing the nucleotide binding cleft between domains 1A and 2A (Figure 5-4C). Specifically, residue Glu833 undergoes displacement of 1.7Å to interact with the O3 group of the ribose ring of the bound nucleotide. Moreover, Arg703 and Arg865 are displaced 1.6Å and 1.5Å respectively to interact with the γ-phosphate (Figure 5-4C). The position of Gln665, a residue acting as

γ-phosphate sensor, is unchanged compared to its corresponding position in the PO4 state

(Figure 5-4C). The closure of the cleft between domains 1A and 2A (Figures 5-4A and

5-4C), and the direct interactions between residues acting as “arginine finger” or “γ- phosphate sensor” with the γ-phosphate (Figure 5-3A), suggests that the AMPPNP- bound complex represents the substrate-bound state. Thus, the presence of AMPPNP in the nucleotide binding pocket stabilizes a network of interactions between domains 1A and 2A. In this manner, AMPPNP appears to act as a cross-linking bridge inducing cleft closure, thus bringing domains 1A and 2A into close proximity. A similar role for

AMPPNP in cleft closure between domains 1A and 2A has been observed in PcrA

(Velankar et al, 1999) and Vasa (Sengoku et al, 2006).

Comparison of the AMPPNP and ADP structures of hUpf1hd reveals that conformational changes would occur with nucleotide hydrolysis. In the ADP-bound structure, the interactions of the adenine base, ribose ring and the β-phosphate group with the protein residues are the same as those observed in the AMPPNP bound structure

(Figure 5-4D). Despite these similarities, motif V in the ADP structure is displaced 4Å from its corresponding position in the AMPPNP structure (Figure 5-4D). Consequently, residues Gln665, Arg703 and Arg865 are displaced 3.1Å, 2.6Å and 4.5Å respectively 130

from their respective positions in the AMPPNP structure. Such structural changes are

most likely induced by the absence of the γ-phosphate group as these residues make

direct contacts with the γ-phosphate group in the AMPPNP bound structure (Figure 5-

4D). In agreement with this view, extensive analysis of crystal packing showed that the closure and opening of the cleft between domains 1A and 2A are not caused by crystal packing as the interface of domains 1A and 2A is distant from any symmetry-related molecules in all three structures. Consistent with the importance of the γ-phosphate group, the presence of a phosphate in the same position in the phosphate bound structure is sufficient, even in the absence of a nucleotide, to partially organize the cleft between the two RecA-like domains, thereby narrowing down the cleft between them. In addition to the bound ADP, a free phosphate is located 4.8Å from the β-phosphate of ADP, but the position of this phosphate does not correspond to the γ-phosphate group of AMPPNP

(Figure 5-4D). The opening of the nucleotide binding pocket and the coexistence of ADP with a free phosphate in the nucleotide binding pocket, suggest that the ADP-bound form represents the product-bound state after ATP hydrolysis.

In addition to the closure and opening of the cleft between domains 1A and 2A in response to nucleotide binding, there are significant movements of domains 1B and 1C that alter their positions relative to each other. In the AMPPNP state, the movement of domain 2A towards domain 1A results in extensive interactions between domain 2A and domain 1B with a buried solvent accessible surface of 1215.8Å2. These interactions also

displace domain 1B towards domain 1C, thereby narrowing the large channel between

them by 2.0Å (Figure 5-4A). Upon ATP hydrolysis (the ADP state), the opening of the

cleft between domains 1A and 2A leads to domain 2A making fewer contacts with 131 domain 1B with a buried solvent accessible surface of 310.5Å2, thereby widening the channel between domains 1B and 2A. Consequently, the channel between domains 1B and 1C in the ADP state is widened by 4.3Å compared to that in the AMPPNP state

(Figure 5-4B). Such conformational changes are the consequences of the nucleotide binding, and may have important mechanistic implications in the catalytic mechanism of hUpf1 (see below). 132

Figure 5-4 Conformational changes of hUpf1hd upon nucleotide binding and hydrolysis. (A) Stereo view of superposition of domain 1A of hUpf1hd-AMPPNP with that of hUpf1hd-PO4. (B) Stereo view of superposition of domain 1A of hUpf1hd-ADP with that of hUpf1hd-PO4. (C) Stereo view of superposition of the nucleotide binding sites of hUpf1hd-PO4 and hUpf1hd- AMPPNP. (D) Stereo view of superposition of the nucleotide binding sites of hUpf1hd-AMPPNP and hUpf1hd-ADP. The PO4, AMPPNP- and ADP-bound forms of hUpf1hd are shown in limon, pink and cyan, respectively. AMPPNP, ADP, phosphate ion and the residues involved in interaction with ligands or Mg2+ coordination are shown in stick models. Mg2+ ion is shown as a purple sphere.

133

5.1.4 Allosteric effect of ATP binding coupled with RNA binding

The Upf1 protein has been shown to bind ssRNA in a manner modulated by ATP

(Czaplinski et al, 1995; Weng et al, 1996; Weng et al, 1998). Mutagenesis combined with biochemical characterization demonstrated that ATP binding rather than hydrolysis promoted dissociation of the Upf1-RNA complex (Weng et al, 1996). However, the mechanism by which ATP modulates the RNA binding activity remains unclear.

In order to identify the ssRNA binding site and examine how RNA binding relates to the ATPase cycle of Upf1, the electrostatic potential was mapped on the molecular surface of hUpf1hd in the three structures. The electrostatic potential mapping showed that in both PO4 and ADP-bound states, a positively charged patch consisting of Arg549 of domain 1A, and Lys599, Arg600, Arg604 of domain 1C, is in close proximity to the outlet of the large channel between domains 1B and 1C (Figure 5-6A). In the AMPPNP state, this patch is extended to domain 1B due to nearly complete closure of the channel between domains 1B and 1C (Figure 5-6A). Notably, the channel between domains 1B and 1C is rich in conserved basic residues (Figures 5-6B and 5-6C), suggesting that this channel may be involved in RNA binding.

Figure 5-5 Protein gel filtration profile of hUpf1 WT and mutants 1B∆ and 1C∆. Superdex 200 column is equilibrated with buffer 20 mM HEPES, pH7.0, 200 mM NaCl, 2 mM DTT. 200 μl of 30 μM purified wild type and two mutated proteins were injected into column. Column was developed by the same buffer. The WT protein, as well as these two mutants are in monomeric form. 134

To examine biochemically the importance of domains 1B and 1C in RNA

binding, two variants in which domains 1B (1B∆) and 1C (1C∆) were deleted respectively, were expressed in E. coli and purified. Gel filtration showed that 1B∆ and

1C∆ behaved similarly to the wild type protein in solution (Figure 5-5), indicating that

removal of these domains had no significant effect on the general folding or stability of

the hUpf1 protein. ssRNA binding assays showed that deletion of domain 1C abolished

the RNA binding activity of hUpf1hd and removal of domain 1B substantially decreased

the RNA binding (Figures 5-6D). Surface plasmon resonance (SPR) assay showed that

1C∆ had no ssRNA binding activity either in the presence or the absence of ATP, while the 1B deletion not only reduced the binding of ssRNA, but also slowed down

dramatically the dissociation of the hUpf1hd-RNA complex, and rendered hUpf1hd

insensitive to the effect of the ATP binding (Figure 5-8). This latter observation argues

that the destabilization of RNA binding by ATP requires domain 1B (see below). Further

evidence that domain 1C is involved in ssRNA binding comes from the observation that

substitution of both Lys599 and Arg600 to Ala reduced the RNA binding to hUpf1hd

substantially, while mutant R604A showed a moderate effect on the RNA binding

(Figures 5-6D). In line with the observations that domain 1C is essential for RNA binding, the RNA-dependent ATPase activity of mutant 1C∆ is substantially reduced

(Figures 5-3C). Moreover, consistent with domains 1B and 1C having an important functional role in binding RNA, deletion of the 1B and 1C domains from yeast Upf1p abolished Upf1p function in vivo (Figure 5-3E).

135

Figure 5-6 The channel between domains 1B and 1C involved in ssRNA binding. (A) Solvent accessible surface and electrostatic potentials of hUpf1hd-AMPPNP, hUpf1hd-PO4 and hUpf1hd- ADP. Residues involved in RNA binding are labeled. (B) Distribution of the basic residues in the channel between domains 1B and 1C in hUpf1hd-AMPPNP. Superimposed ssDNA from the PcrA/ADPNP/dsDNA complex and ssRNA from the Vasa/AMPPNP/ssRNA complex are shown in orange and magenta cartoon tubes, respectively based on the superposition of domains 1A and 2A among hUpf1hd, PcrA and Vasa. (C) Distribution of the basic residues in the channel between domains 1B and 1C in hUpf1hd-ADP. Nucleic acids are generated and shown as in (B). (D) ssRNA binding activity of wild-type and mutant hUpf1hd proteins. 136

An additional approach to identifying the RNA binding channel in Upf1p takes

advantage of the fact that the two RecA-like domains are very similar among hUpf1hd,

PcrA, and Vasa (see above; Figure 5-2). The ssDNA of PcrA overlays very well with the ssRNA of Vasa, which share the same sequence polarity: 5’-to-3’ direction from domains

2A to 1A. Based on these observations, it has bee proposed that the ssRNA would bind

the analogous site in hUpf1 given the high structural similarity shared among these

enzymes. Superposition of domain 1A in either PcrA or Vasa with that of hUPf1hd

results in a model in which the binding of ssRNA to hUpf1hd involves domains 1A, 2A

and 1B with the 3’ end pointing to the channel between domains 1B and 1C (Figures 5-

6B and 5-6C), which electrostatic mapping and mutagenesis had identified as being

important in RNA binding (see above).

An interesting aspect of this likely RNA binding channel is how the

conformational changes induced by nucleotide binding affects the possible RNA binding

channel. In this model, only one loop (residues 349-355) from domain 1B in the AMPNP

state clashes with the ssRNA (Figure 5-7A) whereas this loop is disordered in the ADP

state (Figure 5-7B). Similarly, in the PO4 state, although most part of this loop is

ordered, the tip of this loop (residues 352-354), which would be involved in steric clashes

with the ssRNA in the AMPPNP state, is disordered. This suggests that the ordering of

the 349-355 loop by ATP binding may interfere with RNA binding. Consistent with this

view, SPR experiments shows that two deletion mutants of the Loop 349-355 (Loop 351-

355∆ and Loop 352-354∆) have a similar RNA-binding level to the wild-type protein and

their RNA-binding is ATP-independent (Figure 5-8E and F). Furthmore, these two

deletion mutants have the same ATPase activity as the wild type protein (Figure 5-3C). 137

Another conformational change altered by nucleotide binding that may affect RNA binding is the status of the channel formed between domains 1B and 1C. In both PO4 and

ADP states, the channel is wide open, which would be accessible for the 3’ end of the ssRNA (Figures 5-6A and 5-6C). In contrast, in the AMPPNP state, the channel is nearly closed, thus potentially blocking the ssRNA binding (Figures 5-6A and 5-6B).

Figure 5-7 Allosteric effects of ATP binding/hydrolysis on RNA binding. (A) The figure depicts the potential location of the ssRNA in hUpf1hd-AMPPNP. Left: hUpf1hd-AMPPNP is shown in ribbon diagram (skyblue) covered with transparency surface. Nucleic acids are generated and shown as in Figure 5-6B. The loop (349-355) region in domain 1B is shown in red. Right: Close up view of the zoomed region in left. The loop (758-763) and helix α15 are shown in green and yellow, respectively. (B) The potential location of the ssRNA in hUpf1hd-ADP. Left: Ribbon diagram of ADP-hUpf1hd (slate) covered with transparency surface. Nucleic acids are generated and shown as in Figure 5-6B. Right: Close up view of the zoomed region in left. The disordered loop (349-355) region is shown in dotted red line. The color coding for loop (758-763) and helix α15 are as in (C). 138

Close inspection of the structures of hUpf1hd in three states shows that both the

closure of the channel between domains 1B and 1C, and the potential blocking of the

ssRNA binding by the loop (residues 349-355) are caused by the allosteric effects of

nucleotide binding. Specifically, conformational changes induced by ATP binding and

hydrolysis are relayed to the RNA binding site through residues in motif VI. Upon

AMPPNP binding, the presence of the γ-phosphate in the nucleotide binding pocket

sensed by the residues in motif VI (Figure 5-4C), leads to the rigid-body movement of

domain 2A toward domain 1A (Figure 5-4A), thereby narrowing the nucleotide binding

cleft. This large domain movement also leads to helix α15 and the loop (residues 758-

763) linking β17 and β18 in domain 2A to make direct contacts with the loop 349-355 of

domain 1B (Figure 5-7A), thereby stabilizing the 349-355 loop and pushing domain 1B

toward domain 1C to close the channel between them. In the ADP state, as the

consequence of ATP hydrolysis, domain 2A moves away from domain 1A (Figure 5-

4B). Consequently, domain 2A makes very few contacts with domain 1B, resulting in the

disorder of loop 349-355 and opening of the channel between domains 1B and 1C

(Figure 5-7B). The disorder of loop 349-355 in domain 1B and the opening of the channel between domains 1B and 1C in the ADP form of hUpf1 possibly would enhance the ssRNA binding to hUpf1 as compared to the structure formed in the presence of

AMPPNP. Consistent with this view, SPR assay showed that the binding of ssRNA to hUpf1hd is stronger in the presence of ADP than in the presence of ATP (Figure 5-8).

139

Figure 5-8 Surface plasma resonance analysis of RNA binding to hUpf1. Sensorgrams of the single-stranded RNA binding at the absence or presense of 2 mM Ligands to 100 nM of the wild- type protein (A), mutated proteins 1B∆ (B), 1C∆ (C), R865A (D), Loop351-355∆ (E) and Loop352-354∆ (F).

140

The structural changes induced by AMPPNP binding provide a good explanation

for the dissociation of the Upf1-RNA complex by ATP binding rather than its hydrolysis,

and correlate well with the previous biochemical data (Weng et al, 1996; Weng et al,

1998). In this regard, the AMPPNP-bound state observed likely mimics the ATP binding state. Consistent with this view, SPR assay showed that AMPPNP, ATPγS and ATP

affected the ssRNA binding to hUpf1hd to a similar extent (Figure 5-8). This finding is

in contrast to the previous studies carried out by Weng et al (1998) in which AMPPNP

did not promote the dissociation of the Upf1p-ssRNA complex, whereas ATP and

ATPγS, at higher concentration, reduced the ssRNA binding to Upf1p. In order to check

whether the structural conformation is different upon AMPPNP or ATPγS binding, the

crystal structure of hUpf1hd in complex with ATPγS (hUpf1hd-ATPγS) has also been

solved. Structural comparison showed that the overall structures of hUpf1hd-AMPPNP

and hUpf1hd-ATPγS, and the conformational changes induced by bound AMPPNP and

ATPγS are very similar (Figure 5-9). These observations suggest that AMPPNP and

ATPγS bind to hUpf1hd with a similar affinity, and have similar effects on ssRNA

binding. 141

Figure 5-9 Structure comparison of hUpf1hd-AMPPNP with hUpf1hd-ATPγS. (A) Stereo view of superposition of domain 1A of hUpf1hd-AMPPNP with that of hUpf1hd-ATPγS. (B) Stereo view of superposition of ligand binding sites of hUpf1hd-AMPPNP and hUpf1hd-ATPγS. The AMPPNP- and ATPγS-bound forms of hUpf1hd are shown in pink and lime, respectively. AMPPNP, ATPγS, phosphate ion and the residues involved in interaction with ligands or Mg2+coordination are shown in stick models. Mg2+ ion from AMPPNP-bound and ATPγS-bound are shown as spheres with pink and lime, respectively.

5.1.5 Differential effects of Upf1 mutants on P-body formation

The above results begin to define a cycle of conformational changes in Upf1 that

occur during the process of NMD and are likely to be coupled to, and influenced, by other protein-protein and protein-RNA interactions. Recent results have indicated that the process of NMD can occur within cytoplasmic processing bodies (P-bodies), with

Upf1 being required for targeting of aberrant mRNAs to P-bodies, while the ATPase activity of Upf1 was required at a later stage of NMD to allow mRNA decapping and degradation (Sheth and Parker, 2006). Thus, the accumulation of P-bodies identifies an 142

intermediate stage in the NMD pathway and thereby can be used to map specific aspects

of Upf1p function to upstream or downstream of the formation of the mRNP that can accumulate within P-bodies. Given these results, the effects of different mutations of

Upf1 on the formation of yeast P-bodies were assessed by showing the cellular localization of a Dcp2-GFP reporter protein.

These experiments revealed that mutations (specifically, K436A, DE572AA,

Q601A, and R639A, corresponding to K498A, DE636AA, Q665A, and R703A in hUpf1, respectively) in Upf1 that inhibit the ATPase activity of Upf1, but still allow RNA binding (Figure 5-6D), all led to the accumulation of large P-bodies (Figure 5-10). This is consistent with earlier results with the DE572AA allele and suggests that the ATPase activity of Upf1 functions after the formation of a translationally repressed mRNP, which can accumulate in P-bodies (Sheth & Parker, 2006). In contrast, mutations that inhibit

RNA binding in Upf1, such as deletion of domains 1B or 1C, or mutation of Arg801

(Arg865 in hUpf1) to Ala (Wang et al., 1996; Figure 5-8), do not cause enhanced P-body

accumulation, even though they inhibited the process of NMD as assessed by mRNA

levels (Figure 5-3E). These results reveal allele specific effects on the Upf1p dependent

accumulation of P-bodies. Most notably, the absence of P-body accumulation in the 1C∆

suggests that RNA binding is either required early in the process of NMD before

formation of the mRNP that can accumulate and form P-bodies, or RNA binding is

required following ATP hydrolysis by Upf1p for stable mRNA accumulation in P-bodies.

Interestingly, when the 1C∆ was combined with the DE572AA allele, which blocks the

ATPase and leads to the accumulation of P-bodies, we observed an accumulation of P-

bodies similar to what was seen in the DE572AA strain. This indicates that the function 143

of domain 1C is not required to form P-bodies and implies that RNA binding to Upf1p

occurs after targeting of the RNA to P-bodies. This is consistent with a model of Upf1p

function wherein RNA binding is important at a late stage of the process following ATP

hydrolysis (see discussion).

Figure 5-10 Visualization of Dcp2-GFP in live yeast strains expressing Upf1 mutants.The panels show microscopic accumulation of a Dcp2-GFP fusion protein as a marker for P-bodies. Cells are an upf1∆ strain expressing either the wild-type (WT) Upf1 protein or specific mutant proteins of Upf1 as designated above each panel.

5.2 Discussion

The structure of hUpf1hd described here revealed that hUpf1hd comprised two

“RecA-like” domains with the relative orientations of these two domains similar to those

observed in PcrA, Vasa and Rep helicases. One striking structural feature of hUpf1hd is

the existence of two subdomains 1B and 1C in domain 1. Two lines of evidence suggest

that these unique domains are important in the function of Upf1p. First, in vitro domain

1C is required for RNA binding and ATP hydrolysis, and domain 1B affects the strength of RNA binding (Figures 5-6D). In addition, deletion analysis indicates that domains 1B and 1C are required for Upf1p function in vivo (Figure 5-3E). Moreover, domain 1B 144 appears to play a role in modulating the effects of ATP binding to Upf1p on RNA affinity because deletion of domain 1B or smaller deletions within the domain 1B 349-355 loop allow RNA binding to be relatively insensitive to ATP (Figure 5-8). Interestingly, structural comparisons showed that domains 1B and 1C of hUpf1hd undergo conformational changes due to the closure and opening of the cleft between domains 1A and 2A induced by nucleotide binding and/or hydrolysis.

A common theme in Superfamily 1 helicases is that nucleotide cofactors allosterically regulate nucleic acid binding (Korolev et al, 1997; Velankar et al, 1999).

However, the details of the underlying mechanisms for hUpf1, PcrA and Rep appear to be somewhat different. In Rep helicase, ADP favors the binding of ssDNA whereas

AMPPNP favors simultaneous binding of duplex DNA and ssDNA (Wong et al, 1992).

Moreover, the Rep helicase crystal structure showed that the allosteric effect of ADP binding is transduced to the ssDNA binding site through motif II (Korolev et al, 1997).

The conformational change induced by ADPNP binding in PcrA also favors duplex DNA binding, although motif VI rather than II appears to mediate the transduction of nucleotide-induced conformational change to the DNA binding site (Velankar et al,

1999). For Upf1, biochemical data showed that ATP binding rather than its hydrolysis reduced the affinity for ssRNA (Weng et al, 1998). These differences illustrate how the basic and flexible module of two RecA domains can be coupled to ATP dependent changes in affinity for different nucleic acids.

The structures of hUpf1hd presented here suggest a mechanistic model for how

ATP binding and hydrolysis alter the affinity of Upf1p for RNA. In brief, the binding of

ATP to Upf1p would stabilize a closed conformation of domain 1A and 2A. This closed 145 conformation would reduce the width of the channel between domains 1B and 1C, and stabilize the loop 349-355, which would also sterically hinder the Upf1p-RNA interaction. These ATP induced conformational changes would reduce but not abolish the binding of ssRNA to Upf1, consistent with our SPR data (Figure 5-8) and previous biochemical data by Weng et al., (1998). Following ATP hydrolysis, the movement of the released phosphate to the position seen in the ADP bound structure would disrupt interactions with the γ-phosphate and thereby destabilize the network of interactions holding domains 1A and 2A in close proximity. The resulting opening of the cleft between domains 1A and 2A and the disordering of loop 349-355 would thereby allow enhanced RNA binding.

An unresolved issue is how RNA enhances the ATPase activity of Upf1p. One possibility is that transient binding of RNA to the ATP bound Upf1p triggers the ATPase, which could then lead to tighter binding of that RNA to Upf1p complexed with ADP.

Alternatively, RNA might stimulate the ATPase activity by keeping the cleft between domains 1A and 2A open, thereby stimulating the ATPase activity by facilitating the release of ADP and free phosphate.

An interesting issue is how ATP hydrolysis by Upf1p and changes in RNA binding is coupled with other events in the pathway of NMD. We have used the accumulation of P-bodies as a marker for an intermediate step in the pathway of NMD to reveal the order of events in NMD. Our results indicate that any mutation that blocks

Upf1p ATPase activity leads to increased accumulation of P-bodies, which suggests that

ATP hydrolysis by Upf1p is required after targeting of mRNAs to P-bodies. Our results also argue that RNA binding is only required after targeting of RNAs to P-bodies, and are 146 consistent with a model of NMD wherein Upf1p promotes the targeting of mRNA to P- bodies, followed by ATP hydrolysis, leading to enhanced RNA binding and ultimately mRNA decapping. An important goal of future work will be to understand how other protein-protein interactions occurring during NMD either modulate, or are affected by, this cycle of ATP hydrolysis and conformational changes.

147

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Appendix I

MOPS Minimal Medium (Neidhardt et al., 1974)

(1) 10x MOPS buffer (1 liter): Adjust pH to 7.4 with KOH, sterilize with 0.22μm filter, and store in 50ml aliquots at -20°C.

Reagents Concentration (mM) Quantity (gram) MOPS 400 83.72 Tricine 40 7.17 FeSO4•7H2O 0.1 0.028 NH4Cl 95 5.08 K2SO4 2.76 0.48 CaCl2 0.005 0.000555 MgCl2 5.3 0.0011 NaCl 500 29.22 (2) 5x Amino acids stock (1 liter): Sterilize with 0.2μm filter.

Reagents Concentration (mM) Quantity (mg) Ala 4 356.4 Arg 2.5 526.75 Asn 2 264.2 Asp 2 266.2 Cys 0.5 60.6 Glu 3 441.3 Gln 3 438.3 Gly 4 300.4 His 1 155.2 Ile 4 524.8 Leu 8 1049.6 Lys 4 730.3 Phe 2 330.4 Pro 2 230.2 Ser 18.3 1922.6 Thr 2 238.2 Trp 0.5 102.1 Tyr 2 362.4 Val 6 703.2 Adenine 1 171.6 Guanine 1 151.1 Cytosine 1 111.1 Uracil 1 112.1 Thymine 1 126.1 169

(3) 5000x Micronutrients (500ml)

Reagents Quantity (gram) Na2MoO4•2H2O 0.463 H3BO3 0.618 CoCl2 0.178 CuSO4 0.043 MnCl2 0.396 Zn(OAc)2 0.072 (4) 2000x Vitamin mix (10ml): Add a little NaOH to improve the solubility, sterilize with 0.2μm filter.

Reagents Quantity (gram) Thiamine 67.4 Pantothenic acid 47.6 p-hydroxybenzoic acid 32 p-aminobenzoic acid 27.4 2,3-dihydroxybenzoic acid 30.8

The recipe of MOPS minimal medium (10 liter)

Reagents or buffers Volumn (ml) 10x MOPS buffer 1000 20% Glucose 300 5x Amino acid stock 2000 Water 6600 0.132 M K2HPO4 (be added after water) 100 2000x Vitamin stock 5 5000x Micronutrients 0.2

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Appendix II

Publication list

1) Cheng Z, Liu Y, Wang C, Parker R, Song H. Crystal structure of Ski8p, a WD- repeat protein with dual roles in mRNA metabolism and meiotic recombination. Protein Sci. 2004 Oct;13(10):2673-84. (Cover story)

2) Cheng Z, Coller J, Parker R, Song H. Crystal structure and functional analysis of DEAD-box protein Dhh1p. RNA. 2005 Aug;11(8):1258-70.

3) Cheng Z, Muhlrad D, Lim M, Parker R, Song H. Structural and functional insights into the human Upf1 helicase core. EMBO. J 2007 Jan 10;26(1):253-64.

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