DIVISION OF LABOR AMONG PROTEIN SUBUNITS THAT AID RNA

CATALYSIS IN ARCHAEAL RNASE P

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of

Philosophy in the Graduate School of The Ohio State University

By

Wen-Yi Chen, B.S.

Graduate Program in Molecular, Cellular and Developmental Biology

The Ohio State University

2010

Dissertation committee:

Dr. Venkat Gopalan, Advisor

Dr. Paul Herman

Dr. James Hopper

Dr. John Reeve

Copyright by

Wen-Yi Chen

2010

ABSTRACT

Ribonuclease P (RNase P), a catalytic ribonucleoprotein (RNP) complex, functions in 5′ maturation of tRNAs by removing the 5′ leader from precursor tRNAs

(pre-tRNAs). RNase P contains a catalytic RNase P RNA (RPR) and a varying number of RNase P proteins (RPPs) depending on the source: one RPP in bacteria, at least four in archaea, and at least nine in eukarya. While the four archaeal RPPs

(POP5, RPP30, RPP21, and RPP29) share homology with their eukaryal counterparts, the bacterial RPP shares no sequence homology with archaeal/eukaryal RPPs. Therefore, archaeal RNase P can be used as an experimental surrogate for eukaryal RNase P, which has proven biochemically intractable so far. This dissertation documents a division of labor among archaeal

RPPs that aid the RPR’s catalysis and highlights their roles in chemical cleavage, substrate binding, cleavage-site selection and Mg2+ coordination.

Previous biochemical studies revealed that archaeal RPPs function in pairs;

POP5 dimerizes with RPP30 (POP5RPP30) while RPP21 dimerizes with RPP29

(RPP21RPP29). Individual archaeal RPPs have no influence on the RPR-mediated processing of pre-tRNA under multiple-turnover conditions suggesting that hetero- dimerization is important for the function of RPPs. Furthermore, POP5RPP30 has

-1 -1 been shown to increase RPR’s kcat (from 0.4 min to 10~12 min , pH 7) under

-1 multiple-turnover conditions and to increase RPR’s maximal kobs (from 0.05 min to 5

ii min-1, pH 5.4) when the substrate is provided in cis. The increases observed with

POP5RPP30 parallel those observed with RPR + 4 RPPs indicating that

POP5RPP30 is the sole pair responsible for increasing the rate of reaction.

RPP21RPP29 has been shown to have little influence either on the RPR’s kcat or maximal kobs for cis cleavage while it is able to lower by the KM by 5-fold under multiple-turnover conditions.

Despite these advances made possible by reconstitution of archaeal RNase P, several questions remain unanswered. First, if product release is the rate-limiting step, it is not possible to determine the roles of RPPs in the cleavage step under multiple- turnover conditions. Second, if RPPs play a role in substrate binding and positioning, factors expected to affect the rate of cleavage, such an effect would only manifest in a trans rather than a cis reaction where the substrate is already docked. In this study, I first assessed the effect of each RPP pair on only the cleavage step by using single- turnover kinetic measurements, wherein the enzyme is provided in excess over that of the substrate. Comparison of the kinetic parameters with and without RPPs showed that while POP5RPP30 is solely responsible for enhancing the RPR’s rate of pre-tRNA cleavage by 60-fold, RPP21RPP29 increases substrate affinity by 15-fold.

The protein:RNA molecular mass ratio in bacterial, archaeal, and eukaryal

RNase P is 10%, 50%, and 70%. Whether increased protein content in archaeal/eukaryal RNase P is accompanied by functional gains remains to be proven.

Here, we tested this premise that the increased protein content might influence processing fidelity rather than contribute to drastic alterations in catalytic efficiency.

Indeed, we found that a series of pre-tRNAGln derivatives, which lack consensus

iii structure/sequence elements and are mis-cleaved by bacterial RNase P to generate aberrant tRNA products, are cleaved accurately (~80 to 100% frequency) by the protein-rich archaeal/eukaryal RNase P variants. Exploiting our ability to assemble functional archaeal RNase P in vitro, I sought to dissect which RPPs contribute to this increased fidelity. Towards this goal, I examined trans cleavage of pre-tRNAGln by

Pyrococcus furiosus RPR and cis cleavage of pre-tRNAGln by Methanocaldococcus jannaschii RPR; the latter approach allowed us to focus on the chemical cleavage step. My results show that RPP21RPP29 reduced the RPR’s mis-cleavage (from

80% to ~50%) while leaving the RPR’s rate of cleavage unaffected; in addition to promoting the RPR’s correct cleavage (from 20% to ~50%), POP5RPP30 enhanced cleavage rate by 600-fold regardless of whether it was correct or aberrant cleavage.

4RPPs together increased the correct cleavage to 80%. Finally, I showed that in the presence of POP5RPP30, RPP21RPP29 specifically increases the rate of correct cleavage by ~2.7-fold indicating a synergistic effect between two RPP pairs.

Collectively, these results suggest that protein-rich RNase P confers better fidelity while allowing flexibility in substrate recognition.

Mg2+ plays structural as well as catalytic roles in RNase P catalysis. It remains unclear whether RPPs lower the optimal Mg2+ concentrations by increasing the RPR’s affinity for catalytic and/or structural Mg2+. In this study, I determined that deletion of a universally conserved uridine (ΔU) in a bulge-helix structure in archaeal RPR weakens Mg2+ coordination and lowers activity, a defect completely rescued by their cognate RPPs under multiple-turnover conditions. Cis cleavage of pre-tRNATyr by Mja

ΔU RPR revealed an 80-fold decrease in the cleavage rate compared to the wildtype; both Mja RPP pairs partially ameliorate this defect and narrow the difference to ~12-

iv fold, indicating their ability to coordinate catalytically relevant Mg2+ ions. In addition to archaeal RPPs rescuing an RPR mutation, we (in collaboration with the laboratory of

Dr. Mark Foster, OSU) found that an N-terminal deletion mutant of RPP29, which fails to bind RPP21, is functional upon addition of RPR. Together, these findings illustrate the cooperative subunit interactions critical for generating the functional structure of archaeal RNase P.

Collectively, these results demonstrate the division of labor and the functional coordination among different RPPs, and suggest possible reasons for why these distinct cofactors were recruited by an ancient RNA enzyme during the transition from a primordial RNA to an RNP world.

v

DEDICATION

Dedicated to my wife, Pao-Yin Fan, for her unconditional love, support and

encouragement provided throughout my graduate career.

vi

ACKNOWLEDGEMENTS

Many, many thanks to my advisor, Dr. Venkat Gopalan, for everything he has taught me. I have learned from him not only the pursuit of science but also the integrity of being a good scientist.

I also thank Drs. Paul Herman, James Hopper and John Reeve who have served in my dissertation committee and provided valuable feedback. In addition, I am indebted to Dr. Mark Foster for his valuable insights into the tertiary structure and

Kinetic mechanism of RNase P.

I am also grateful to Dr. Deepali Singh who contributed to the initial stages of the fidelity project described in this dissertation.

I want all the former and current members of the Gopalan laboratory to know how much I enjoyed your company and how helpful you all have been. In particular, I am grateful to Drs. Lien Lai and Dileep Pulukkunat for their valuable insights and expertise in science; I-Ming Cho and Cecilia Go for comradeship; Dr. Anil Challa and

Sathyianarayanan Manivannan for many inspiring chats; Stella Lai for bringing home- baked desserts to every lab meeting; and all the undergraduates, Emily Wong, Derek

Smith, Chigo Ekeke and Andrew Merriman for their support.

vii I would also like to acknowledge the OSU Molecular, Cellular and

Developmental Biology graduate program and the OSU Department of Biochemistry for providing funding support to attend meetings.

Lastly, I would like to thank my parents for allowing me to realize my own potential and for their love and support. Without their help, this degree would not have been possible.

viii

VITA

1994 - 1998...... B.S. (Zoology), National Taiwan University

2000 - 2001...... Research Assistant, National Academic

Sinica, Taipei, Taiwan

2001- present ...... Graduate Teaching and Research

Associate, The Ohio State University,

Columbus, Ohio

Publications

1. Chen, WY*, Pulukkunat, DK*, Cho, IM, Tsai, HY and Gopalan, V (2010).

Dissecting functional cooperation among protein subunits in archaeal RNase P.

Nucleic Acids Res. In press. doi: 10.1093/nar/gkq668.

2. Lai, LB, Cho, IM, Chen, WY and Gopalan, V (2010). Archaeal RNase P:

A mosaic of its bacterial and eukaryal relatives. In Liu, F and Altman, S (Eds.),

Ribonuclease P. Protein Reviews Series (Volume 10, pp. 153-172). Springer-

Verlag, New York, NY.

3. Wenzel, PL, Wu, L, de Bruin, A, Chong, JL, Chen, WY, Dureska, G, Sites, E,

Pan, T, Sharma, A, Huang, K, et al. (2007). Rb is critical in a mammalian tissue

stem cell population. Genes Dev. 21, 85-97.

ix 4. Chen, WY, Yang, YM and Chuang, NN (2002). Selective enhanced

phosphorylation of shrimp beta-tubulin by PKC-delta with PEP (taxol), a

synthetic peptide encoding the taxol binding region. J. Exp. Zool. 292, 376-383.

5. Huang, CF, Chen, WY and Chuang, NN (2000). Differential expression of ras in

organs and embryos of shrimp Penaeus monodon (Crustacea: Decapoda).

Comp. Biochem. Physiol. B Biochem. Mol. Biol. 125, 307-315.

* indicates equal contribution

Field of study

Major Field: Molecular, Cellular and Developmental Biology

x

TABLE OF CONTENTS

ABSTRACT ...... ii DEDICATION ...... vi ACKNOWLEDGEMENTS ...... vii VITA ...... ix LIST OF TABLES ...... xiv LIST OF FIGURES ...... xv LIST OF ABBREVIATIONS ...... xviii

CHAPTERS

1. INTRODUCTION ...... 1

1.1 Prelude ...... 1 1.2 The subunit makeup of RNase P ...... 3 1.2.1 Bacterial RNase P ...... 3 1.2.1.1 The RNA subunit ...... 3 1.2.1.2 The protein subunit ...... 6 1.2.2 Archaeal RNase P ...... 7 1.2.2.1 The RNA subunit ...... 8 1.2.2.2 Protein subunits ...... 9 1.2.2.3 Structures of the protein subunits of archaeal RNase P ...... 10 1.2.2.3.1 POP5 ...... 10 1.2.2.3.2 RPP30 ...... 11 1.2.2.3.3 RPP21 ...... 12 1.2.2.3.4 RPP29 ...... 12 1.2.2.4 Protein-protein interactions in archaeal RNase P ...... 13 1.2.3 Eukaryal RNase P ...... 14 1.2.4 Organellar RNase P ...... 15 1.3 In vitro reconstitution of RNase P ...... 17 1.4 Research objectives ...... 19 1.4.1 Elucidate the roles of archaeal RPPs in aiding RPR-mediated cleavage ...... 19 1.4.1.1 Dissect the roles of archaeal RPPs in substrate binding and chemical cleavage ...... 20 1.4.1.2 Delineate the roles of archaeal RPPs in influencing the RPR’s cleavage-site selection ...... 20 1.4.1.3 Elucidate the roles of archaeal RPPs in coordinating catalytic Mg2+ ...... 21

xi 2. SINGLE-TURNOVER KINETIC STUDIES TO ELUCIDATE THE ROLES OF ARCHAEAL RPPS IN AIDING RNA-MEDIATED CLEAVAGE OF PRECURSOR TRANSFER RNA ...... 38

2.1 Introduction ...... 38 2.2 Materials and methods ...... 39 2.2.1 Generation of Methanothermobacter thermautotrophicus RPR using in vitro transcription ...... 39 2.2.4 Mth RNase P single-turnover assays...... 42 2.2.5 Mth RNase P single-turnover data analysis ...... 43 2.2.6 Pulse-chase experiments with Mth RNase P to demonstrate that the dissociation rate (k-1) is greater than the rate of chemical cleavage (k2) under single-turnover conditions...... 43 2.3 Results ...... 44 2.3.1 Rationale ...... 44 2.3.2 Purification of the subunits of Mth RNase P...... 45 2.3.3 Optimal assay conditions for single-turnover kinetic measurements of RPR and RNPs ...... 45 2.3.4 Single-turnover assays of partially and fully reconstituted Mth RNase P holoenzymes...... 46 2.3.5 Comparison of partially and fully reconstituted Mth RNase P at 30 mM Mg2+ ...... 48 2.4 Discussion ...... 48 2.4.1 Role of protein cofactors in Mth RNase P catalysis ...... 48 2.4.2 The importance of RPP21RPP29 in vivo ...... 50 2.4.3 Comparison of Mth RPR data to the rates of pre-tRNA cleavage by bacterial and archaeal RPRs reported in previous studies ...... 51 2.4.4 Summary ...... 51

3. ELUCIDATING THE ROLE OF ARCHAEAL RPPS IN INFLUENCING THE RPR’S CLEAVAGE-SITE SELECTION ...... 63

3.1 Introduction ...... 63 3.2 Materials and methods ...... 66 3.2.1 Pfu RNase P assay ...... 66 3.2.2 Cloning and in vitro transcription of pre-tRNAGln-Mja RPR ...... 66 3.2.3 Kinetic studies with pre-tRNAGln-Mja RNase P ...... 68 3.2.4 Data analyses ...... 69 3.3 Results ...... 70 3.3.1 The significance of N+1N+72 for pre-tRNA for cleavage-site selection decreases from bacterial > archaeal > eukaryal RNase P ...... 70 3.3.2 Stable tertiary structure of pre-tRNAGln is important for correct cleavage-site selection by bacterial RNase P but not the archaeal or eukaryal variants ...... 71 3.3.3 The length of the T-stem and the acceptor stem of pre-tRNAGln affects cleavage-site selection ...... 72 3.3.4 Archaeal RPPs affect cleavage-site selection ...... 73 3.3.5 Effect of Mja RPPs on cleavage-site selection and the rate of pre- tRNAGln-Mja processing ...... 74 3.4 Discussion ...... 76 xii 3.4.1 Nucleotide identity at position +1 in pre-tRNAGln ...... 76 3.4.2 D-stem, T-stem, and anticodon stem in pre-tRNAs...... 77 3.4.3 Archaeal RPPs influence cleavage-site selection and the rate for processing pre-tRNAGln ...... 78 3.4.4 Protein-rich RNase P confers more flexibility ...... 81

4. COOPERATIVE RNP ASSEMBLY: COMPLEMENTARY RESCUE OF STRUCTURAL DEFECTS BY PROTEIN AND RNA SUBUNITS OF ARCHAEAL RNASE P ...... 92

4.1 Introduction ...... 92 4.2 Materials and methods ...... 94 4.2.1 Cloning and expression of the genes encoding the RNA and protein subunits of Mth and Mja RNase P ...... 94 4.2.2 Construction of mutant derivatives of archaeal RPRs ...... 95 4.2.3 In vitro transcription of RPRs used in this study ...... 96 4.2.4 RNase P assays ...... 96 4.2.5 RNase P activity data analysis ...... 98 4.3 Results ...... 99 4.3.1 Mutations in the P4 helix of archaeal RPRs decrease activity ...... 99 4.3.2 Archaeal RPPs can rescue mutations in the P4 helix of cognate RPRs ...... 99 4.3.3 RPP21•RPP29 and POP5•RPP30 both rescue the decrease in cleavage rate caused by the ΔU mutation ...... 100 4.4 Discussion ...... 102 4.4.1 Evolutionarily and functionally conserved motifs in both bacterial and archaeal RPRs ...... 102 4.4.2 Archaeal RPPs can mitigate catalytic defects in the cognate RPR . 103 4.4.3 Archaeal RPR can rescue the deleterious effects of a large deletion in a cognate RPP ...... 106

5. FUTURE DIRECTIONS ...... 112

5.1 Investigating the roles of L7Ae in archaeal RNase P catalysis ...... 112 5.2 In vivo mapping of the 5′ end of tRNAs ...... 113 5.3 Which RPPs contribute to increased affinity for metal ions in active site? . 114 5.3.1 Introducing the PS mutation in pre-tRNATyr-Mja ...... 115

REFERENCES ...... 119

xiii

LIST OF TABLES

Table 2.1. Characteristics of RPPs from Methanothermobacter

thermautotrophicus ...... 53

Table 2.2. Effect of Mth RPPs on the ionic requirements and rate of cleavage of

pre-tRNATyr by Mth RPR at 55C ...... 54

Table 3.1. Effect of Mja RPPs on the rate of cleavage and cleavage-site selection

of pre-tRNAGln-Mja at 55C at pH6 ...... 83

Table 3.2. Oligonucleotide primers used to construct pre-tRNAGln-Mja RPR...... 84

Table 4.1. Oligonucleotide primers used to construct mutant archaeal RPRs... 107

Table 4.2. Effect of Mja RPPs on the ionic requirements and rate of cleavage of

pre-tRNATyr-Mja and pre-tRNATyr-MjaU at 55C ...... 108

xiv

LIST OF FIGURES

Figure 1.1. Role of RNase P in tRNA maturation ...... 22

Figure 1.2 A cladogram depicting a possible scheme for evolution of RNA-based

RNase P ...... 23

Figure 1.3. Secondary structures of bacterial RPRs ...... 24

Figure 1.4. A minimal bacterial consensus RPR deduced from phylogenetic

analysis ...... 26

Figure 1.5. Structures of the specificity domain of type A and B bacterial RPRs .. 27

Figure 1.6. Inter-domain interactions in bacterial RPR ...... 29

Figure 1.7. Tertiary structure of a bacterial RPP ...... 30

Figure 1.8. Model of the RNase P holoenzyme ...... 31

Figure 1.9. Comparison of secondary structures of RPRs in all three domains of

life ...... 32

Figure 1.10. Tertiary structures of archaeal RPPs ...... 33

Figure 1.11. Simularities and differences in structures of bacterial RPP and

archaeal POP5 ...... 34

Figure 1.12. Tertiary structures of archaeal RPP binary complexes ...... 35

Figure 1.13. Electrostatic potential surface map of POP5RPP30 ...... 36

Figure 1.14. Electrostatic potential surface map suggests two RNA binding regions

in the Pfu RPP21RPP29 complex ...... 37

Figure 2.1. Purification of recombinant Mth RPPs as binary complexes ...... 55

xv Figure 2.2. Effects of Mth RPPs on the single-turnover rate of RPR-catalyzed pre-

tRNA cleavage ...... 56

Figure 2.3. Pulse-chase experiments with Mth RNase P to demonstrate that the

dissociation rate (k-1) is greater than the rate of chemical cleavage (k2)

under single-turnover conditions ...... 58

Figure 2.4. Effects of Mth RPPs on the single-turnover rate of RPR-catalyzed pre-

tRNA cleavage ...... 59

Figure 2.5. Dependence of kobs on assay pH for Mth RPR + 4 RPPs ...... 60

Figure 2.6. Kinetic scheme as a framework to interpret the contribution of the two

binary RPP complexes to archaeal RNase P catalysis ...... 61

Figure 2.7. Schematic drawing of the RPP footprinting data depicted on a

secondary-structure model of Methanocaldococcus jannaschii (Mja)

RPR and possible interactions between RPR and pre-tRNA ...... 62

Figure 3.1. (A) Schematic drawings depicting pre-tRNAGln and its derivatives. (B)

Possible interactions between nucleotides in typical or atypical pre-

tRNAs and the bacterial RPR at the cleavage site ...... 85

Figure 3.2. Mapping the cleavage sites of pre-tRNAGln and its mutant derivatives

by RNase P from (A) Escherichia coli (Eco), (B) Pyrococcus furiosus

(Pfu), (C) Arabidopsis thaliana (Ath)...... 86

Figure 3.3. Comparison of the correct and mis-cleaved products of pre-tRNAGln

processed by RNase P from Escherichia coli (Eco), Pyrococcus

furiosus (Pfu), Arabidopsis thaliana (Ath)...... 87

Figure 3.4. Archaeal RPPs affect the RPR’s cleavage-site selection ...... 88

Figure 3.5. Pfu RPPs affect cleavage-site selection by the cognate RPR ...... 89

xvi Figure 3.6. Effect of Mja RPPs on cleavage-site selection and the rate of self-

cleavage of pre-tRNAGln-Mja ...... 90

Figure 3.7. Kinetic scheme as a framework to interpret the contribution of the two

binary RPP complexes to cleavage-site selection and the rate of pre-

tRNAGln-Mja ...... 91

Figure 4.1. Secondary structures of RPRs from Bacteria and Archaea...... 109

Figure 4.2. Comparison of the Mg2+ dependence of wildtype (WT) and mutant Mth

and Mja RNase P ...... 110

Figure 4.3. Effects of Mja RPPs on the rate of pre-tRNATyr-MjaΔU RPR self-

cleavage ...... 111

Figure 5.1. The role of Mg2+ ions in RNase P-mediated catalysis ...... 117

Figure 5.2. pre-tRNATyr-Mja RPR carrying a phosphorothioate modification at the

RNase P cleavage site ...... 118

xvii

LIST OF ABBREVIATIONS

°C Degree Celsius

Abs600 Absorbance at 600 nm

Abs260 Absorbance at 260 nm

α Alpha

 Beta

Ath Arabidopsis thaliana

Afu Archaeoglobus fulgidus

Bsu Bacillus subtilis

C domain Catalytic domain

C-terminal Carboxyl terminal

DNA Deoxyribonucleic acid

DTT Dithiothreitol

Eco Escherichia coli

EDTA Ethylenediaminetetracetic acid

 Gamma g Gram(s) h Hour(s)

IPTG Isopropyl-β-D-thiogalactopyranoside kDa Kilodalton

L Liter

xviii LB Luria-Bertani

M Molar

MES 2-[N-Morpholino]ethanesulfonic acid mg Milligrams min Minutes

Mja Methanocaldococcus jannaschii ml Milliliter mM Millimolar

Mma Methanococcus maripaludis

Mth Methanothermobacter thermautotrophicus

N-terminal Amino terminal

NAIM Nucleotide analog interference mapping ng Nanograms nm Nanometer nM Nanomolar

NMR Nuclear magnetic resonance

PCR Polymerase chain reaction

Pfu Pyrococcus furiosus

Pho Pyrococcus horikoshii pre-tRNA Precursor transfer ribonucleic acid

RNA Ribonucleic acid

RNase P Ribonuclease P

RNP Ribonucleoprotein rpm Revolutions per minute

RPP RNase P protein

xix RPR RNase P RNA

S domain Specificity domain

SDS-PAGE Sodium dodecyl sulfate-polyacrylamide gel electrophoresis

Tma Thermotoga maritima

Tris Tris-(hydroxymethyl) aminomethane tRNA Transfer ribonucleic acid

UV Ultraviolet

μg Microgram(s)

μl Microliter(s)

μM Micromolar

xx

CHAPTER 1

INTRODUCTION

1.1 Prelude

Nearly five decades ago, Francis Crick hypothesized that there must be transfer RNA (tRNA) adaptors designed to capture amino acids and to transport them to the ribosomes where proteins are synthesized according to the codons in messenger RNAs. The historical studies which validated this premise fueled subsequent studies on the myriad roles of tRNAs in a wide variety of cellular processes. Deciphering the biogenesis and functions of tRNA is an essential part of modern molecular biology that seeks to fully understand the regulation of gene expression. Studies of phage-encoded tRNATyr suppressor-negative mutants revealed accumulation of precursors longer at the 5′ end than the mature counterparts thus implicating the need for 5′ processing during their biogenesis (1). Subsequent experiments revealed that ribonuclease P (RNase P) is responsible for 5′ end maturation and catalyzes the removal of 5′ leaders from pre-tRNAs [Fig. 1.1; (2-5)]. In the process of uncovering the true identity of RNase P, Sidney Altman and co- workers fortuitously discovered the first true RNA enzyme (6) and the following biochemical characterization studies revealed the diverse variants of RNase P in different life forms (7-10) .

1 Although the primary function of RNase P is to cleave pre-tRNAs, the subunit makeup of RNase P from all three domains of life is quite different. It contains one

RNase P RNA (RPR) and a varying number of RNase P proteins (RPPs) depending on the source: one in bacteria, at least four in archaea and up to 10 in eukarya [Fig.

1.2; (2-5)]. The increased protein content in these RNase P variants (10% in bacteria,

50% in archaea, and 70% in eukarya) is inversely correlated with the catalytic activity of RPRs (bacterial > archaeal > eukaryal) (6,11-14). Although the RPR is the catalytic moiety in all life (6,12,13), the RPPs are essential in vivo (9,15,16). These observations raise some interesting questions: (i) Why did nature retain RNA-based catalysis in a cellular milieu where proteins predominantly carry out the biological functions? (ii) What are the roles of protein cofactors in aiding RNA catalysis?

Although protein enzymes exhibit higher catalytic efficiency and versatility, many important cellular machineries still rely on RNA as macromolecular catalysts. For example, the active site of ribosome is made only of RNA (17). Similarly, small nuclear RNAs (snRNA) can catalyze the formation of a lariat structure (a key intermediate during mRNA splicing) in the absence of spliceosomal proteins (18).

Despite the central role of RNA in catalysis of the ribosome, splicesome and RNase P, these macromolecules function as ribonucleoproteins (RNPs) in vivo. If RNAs were the first macromolecules to serve as genetic material and execute catalytic functions, why and how were protein cofactors recruited to participate in RNA-mediated catalysis in RNP complexes? The focus of this dissertation is to understand the possible reassignments of structural and functional attributes of RNAs to protein cofactors and deduce the functional gains conferred by such a transition.

2 1.2 The subunit makeup of RNase P

Comparing RNase P variants with different subunit compositions in the three domains of life is likely to shed light on the structural and functional alterations that accompanied the transformation of RNase P from an ancient RNA catalyst to the extant protein-rich RNP enzyme. The following sections highlight the commonality and differences among RPRs and RPPs from different life forms.

1.2.1 Bacterial RNase P

Temperature-sensitive mutants for bacterial RNase P activity map to two distinct gene loci in the Escherichia coli (Eco) genome implicating that bacterial

RNase P contains two subunits. Subsequent studies revealed that these two loci

(rnpA and rnpB) encode the protein and RNA subunits, respectively (19-21).

Biochemical purification and reconstitution studies have shown that bacterial RNase

P is a hetero-dimer consisting of one essential RNA subunit (~400 nts; 125 kDa) and one protein cofactor (~120 amino acid residues; 15 kDa), and that the RNA is the catalytic moiety (7,22,23). Bacterial RPR was the first true RNA enzyme proven to support multiple-turnover at non-physiological ionic conditions in vitro (6). However, both the RNA and protein subunits are absolutely essential for cell viability in vivo

(15,16,24) highlighting the essential roles of bacterial RPP in supporting RPR- mediated catalysis inside the cell.

1.2.1.1 The RNA subunit

Loops (L) and helices (P, paired regions) in the bacterial RPRs were first determined based on susceptibility of distinct regions in the RPRs to cleavage by nucleases. For example, ribonucleases T1 and V1 specifically cleave single- and

3 double-stranded regions of RNA, respectively. Secondary structure models were built by pairing the complementary bases in the helix region. As the number of RPR sequences increased, these models were subsequently confirmed and refined based on results from phylogenetic covariation analysis of several bacterial RPR sequences

[Fig 1.3; (25-28)]. They categorized bacterial RPRs into types A and B depending on the presence of different structural elements, and identified universally conserved nucleotides (Fig. 1.4) that are otherwise difficult to identify by alignment of the RPR primary sequences. Furthermore, based on covariations of two GNRA tetraloop:helix docking interactions [base triples formed by either G (loop):A/U (helix) or A (loop):G/C

(helix)], Brown et al (29) predicted tertiary contacts between L18:P8 and L14:P8 in type A RPR (Fig 1.3), which were later confirmed by high resolution crystal structures

(30). Although the type B RPR is missing P14 and P18 to form these interactions, phylogenetic studies (27,31) have predicted the L5.1:L15.1 interaction as an alternative in type B RPR (Fig 1.3). By examining the length and sequence covariation between the P9/L9 stem–loop and the P1 helix in both type A and B RPRs,

Westhof and co-workers identified a common tertiary interaction between the L9 tetraloop and two specific base-pairs in P1 [Fig. 1.3; (32)]. Mutagenesis studies have supported the idea that these tertiary interactions contribute to the overall structural stability of the bacterial RPRs (33).

To identify the catalytic core and nucleotides essential for pre-tRNA binding and catalysis, mutagenesis, crosslinking and nucleotide analog interference mapping

(NAIM) studies (34-43) on both type A and B RPRs have been performed and proven fruitful. Biochemical studies on bacterial RPRs identified two independently folding modules (37,44,45): a specificity (S) domain that contains conserved nucleotides recognizing the T stem–loop of the pre-tRNA (46), and a catalytic (C) domain that can 4 cleave the pre-tRNA leader sequence while binding the leader (47), acceptor stem

(48), and the 3′-RCCA sequence (R represents purine) (49-51). Tertiary interactions between these two domains are evident by the fact that these two domains could be assembled into a functional RPR even when they are synthesized and folded separately. These interactions juxtapose these two domains and allow the S domain to interact with T stem-loop of the pre-tRNA while orienting the pre-tRNA cleavage site near the active site in the C domain. Despite tertiary interactions mediated by different peripheral elements in type A and B RPRs, these RPRs share a conserved structural core capable of performing catalysis in vitro [Fig. 1.4; (34)]. The consensus structure consists of P1 to P15 with a majority of conserved nucleotides clustered in the P4 region; the cruciform structure (consisting of P7 to P11) and J11/12 (J for joining, and numbered by the flanking helices) are also common among all bacterial

RPRs with a conserved adenosine in J11/12 (25). The C domain contains the minimal structural elements required to perform catalytic activity (52).

Crystal structures of two different bacterial RPRs have validated these secondary structure models and proposed tertiary interactions. The initial structures were reported for the S domain alone [Fig. 1.5; (53,54)] and then for the full length

RPRs from both types A [Fig. 1.6; (30)] and B (55). Type A RPR consists of two layers of coaxially stacked helices: layer 1 contains both the substrate-binding regions and the catalytic site; layer 2 serves as an assembly organizer with P8/P9 stack in the middle, and P13/P14 and P18 on each side, held by L18:P8, L14:P8, and L9:P1 tetraloop:helix interactions (Fig. 1.6). Type B RPR displays a remarkably flat face formed by a spatial arrangement of coaxially stacked helices. Inter-domain interactions such as L5.1:L15.1 and L8:P4 were evident in the crystal structure (55).

5 Interestingly, although the type B RPR uses different peripheral elements compared to the type A relative to stabilize the overall structure, the two RPRs share some major common features: (i) a cleft, formed by P9, P10 and P11, with nucleotides shown to be involved in direct interactions with the substrate, and the non-helical module L11/12-L12/11 [Fig. 1.5; (53)], and (ii) a similar arrangement of the putative catalytic site consisting of P4 and the adjacent inter-helical joining regions (e.g. J5/15 and J18/2).

1.2.1.2 The protein subunit

The sole bacterial RPP plays multiple roles (56-64): (i) increasing the RPR’s affinity for pre-tRNA over mature tRNA by directly binding the 5′ leader sequence of pre-tRNA, (ii) enhancing RPR’s rate of cleavage presumably through modulating

RPR’s structure at catalytic site, and (iii) promoting RPR’s affinity for Mg2+ to allow the

RPR to function at physiological Mg2+ concentrations.

The tertiary structures of bacterial RPP from Bacillus subtilis (Bsu),

Thermotoga maritima, and Staphylococcus aureus have been determined using X-ray crystallography and NMR spectroscopy (65-67). All bacterial RPPs exhibit an α-β sandwich fold comprising a four-stranded β-sheet surrounded by two α-helices on one face and one α-helix on the other (Fig. 1.7). The face of the β-sheet packing with helix

α1 forms a cleft, which has been shown to bind the pre-tRNA leader (57,60,68-71).

Stacking interactions between the leader of pre-tRNA and the conserved aromatic residues in the cleft likely contribute to pre-tRNA binding, although there is some evidence that hydrogen bonding between the bacterial RPP and the pre-tRNA leader may also play a role in the cleft–5′-leader interactions (70). The other face of the β-

6 sheet packs with helices α2 and α3 to form the hydrophobic core. RPR binding is mediated by a highly basic region, the RNR motif (AHxxRNRxxKRLxR, where x is any amino acid residue) located in helix α2 (68,71-73). This dual RNA binding property of the RPP explains its ability to enhance RPR’s substrate binding and RPR-mediated cleavage.

Footprinting and crosslinking studies (68,71,74) on both type A and B RNase

P holoenzymes have shown that the bacterial RPP interacts exclusively with the

RPR’s C domain and that the central cleft of RPP directly contacts the leader of pre- tRNA. Specifically, Tsai et al [Fig. 1.8; (68)] used EDTA-Fe based footprinting to show that helix α1, the unique left-handed cross-over loop, and the RNR motif in helix α2 of

Eco RPP are proximal to the P3, P4 and J18/2 regions of its cognate RPR, respectively; some residues in helix α2 are also close to J2/4. Niranjanakumari et al

(71), on the other hand, have shown that in Bsu RNase P, the metal-binding loop, which is capable of binding to Zn2+ (67), and the N terminus are near the P3 stem- loop; the RNR motif is proximal to P4 and the pre-tRNA cleavage site; β3 is proximal to J2/3 region. Collectively, these data have helped build computer-aided three dimensional models of the bacterial RNase P holoenzyme with and without the pre- tRNA, and provide valuable insights into the functioning of the bacterial RPP (Fig. 1.8).

However, atomic-level details of the interactions among the RPR, RPP, and pre-tRNA await a crystal structure of the holoenzyme-substrate complex.

1.2.2 Archaeal RNase P

Most of our current understanding about RNase P comes from studies on the bacterial RNase P. However, it is a much simpler variant, consisting of one RNA and

7 only one protein, compared to the archaeal and eukaryal versions, which have at least four and nine RPPs, respectively. In addition, the bacterial RPP does not share homology with archaeal and eukaryal RPPs. Therefore, we use archaeal RNase P as an experimental surrogate for the biochemically intractable eukaryal RNase P. There are several advantages to studying archaeal RNase P: (i) archaeal and eukaryal

RPPs share high sequence homology (8), therefore, findings on archaeal RNase P could be extended to the eukaryal counterpart; (ii) archaeal RNase P, with fewer number and smaller size of RPPs, may represent the minimal catalytic RNP core in archaeal/eukaryal RNase P; and (iii) archaeal RPPs are strongly basic and thermostable, making them better suited for purification, high-resolution structural studies and functional reconstitution.

1.2.2.1 The RNA subunit

The predicted secondary structures of archaeal RPRs are based on bacterial

RPRs (Fig. 1.9). In fact, the minimal consensus secondary structures of bacterial and archaeal RPRs are very similar (75-77). Archaeal RPRs fall into two distinct groups, types A and M [Fig. 1.9; (77-79)]. Type A RPR is apparently the more ancestral structural class, exemplified by Methanothermobacter thermautotrophicus (Mth), which is strikingly similar to ancestral bacterial type A RPRs. The major differences between archaeal and bacterial type A RPRs are the loss of P18 and P13/P14 in the former (Fig. 1.9). Therefore, the tertiary contacts in layer 2 of bacterial type A RPRs cannot be formed in archaeal RPRs (Fig. 1.6), which may explain their decreased catalytic activity and more acute dependence on RPPs.

8 Type M RPRs, exemplified by Methanocaldococcus jannaschii (Mja), in addition to the loss of P13/P14 and P18, differ from archaeal type A RPRs due to (i) the absence of P8 [known in bacteria to be involved in pre-tRNA binding (80,81)], and

(ii) everything distal to P15, including L15 [known in bacteria to be involved in substrate 3-RCCA recognition (49,50,82-85) and Mg2+ ion coordination (86)] (Fig.

1.9). Also the cruciform formed by helices P7–P11 of these RNAs has undergone a significant rearrangement in addition to the loss of P8. The changes include lengthening of P7, P9, and P10/11, and the loss of extrahelical nucleotides from

P10/11 (77). The loss of L15/P16/P17 also eliminates the P6 pseudoknot (77) (Fig.

1.9). Type M RPRs have been found only in Methanococcales and Archaeoglobales.

Archaeal type A RPRs display weaker activity compared to bacterial RPRs

+ 2+ and require 2 M NH4 ion and 0.5 M Mg for optimal enzymatic activity as opposed to

+ 2+ 0.4 M NH4 ion and 0.1 M Mg for bacterial RPRs (12,87) presumably due to lack of structural elements necessary for overall stability (11) and substrate binding (14,88).

Type M RPRs are catalytically inactive without the presence of RPPs (12,88). Mja

RPR can cleave pre-tRNATyr when provided in cis (88) suggesting that its inability to catalyze pre-tRNA processing is largely due to a substrate-binding defect.

1.2.2.2 Protein subunits

In contrast to an apparently common ancestry for RPRs, especially in the putative active site, the currently known four archaeal RPPs share no homology with bacterial RPP (8). However, they share homology with eukaryal RPPs reflecting the common evolutionary origin of archaea and eukarya [Fig. 1.2; (8)]. In fact, the archaeal RPPs were first computationally identified based on their homology to

9 eukaryal counterparts (8). They are termed POP5, RPP30, RPP21, and RPP29

(based on names of the corresponding human RPP homologs). Association of these four RPPs with partially purified archaeal RNase P activity was demonstrated by western blotting and immunoprecipitation (8). Subsequent in vitro reconstitutions using these RPPs further confirmed their ability to enhance RPR’s catalytic activity

(87-90). The archaeal RPPs, particularly RPP29 and RPP30, are smaller than their eukaryal homologs. The additional sequences present in eukaryal RPPs might not be vital for catalysis but rather needed for nuclear localization (91) and/or for interacting with RPPs not present in archaea (92,93).

While the approaches of combined homology searches and partial purification have proven fruitful and have identified four bona fide archaeal RPPs, elucidating the complete subunit makeup will await successful purification and characterization of a native archaeal RNase P holoenzyme.

1.2.2.3 Structures of the protein subunits of archaeal RNase P

Although some RPPs were solved by both NMR spectroscopy and X-ray crystallography (Fig. 1.10) and some differences were reported between the solution and crystal structures, the core structures are identical (as elaborated below).

1.2.2.3.1 POP5

The tertiary structure of Pyrococcus furiosus (Pfu) POP5 alone (94) and

Pyrococcus horikoshii (Pho) POP5 in complex with RPP30 have been solved by X- ray crystallography (95). The secondary structures of archaeal POP5 (βααββαβα) and bacterial RPPs (αβββαβα) are different suggesting that the two proteins may have

10 evolved from different origins (94). POP5 adopts an α-β sandwich fold with a four- stranded antiparallel β-sheet in the middle surrounded by four α helices, similar to an

RNA recognition motif (RRM) (Fig. 1.10 and 1.11). Helices α1–α3 pack against one face of β sheet to form a hydrophobic core; this face has been shown to interact with

RPP30 by NMR chemical-shift perturbations (94) and crystallography (95). Helix α4 packs against the other face of the β sheet; this side is more exposed to solvent and has some loosely packed apolar residues that could engage in stacking interactions with RPR/pre-tRNA (Fig. 1.10 and 1.11). There are major differences between POP5 and bacterial RPPs: (i) the central cleft in bacterial RPP, which is thought to interact directly with the 5′-leader of pre-tRNA, was not seen in POP5 (helix α4 in Pfu POP5, which corresponds to α1 in bacterial RPP, does not directly pack against the β-sheet to form a central cleft [Fig. 1.11; (94)]; (ii) the RNR motif in α2 of bacterial RPPs, which is conserved among all the bacterial RPPs is absent in POP5; and (iii) the unique left-hand crossover structure in bacterial RPP is also not observed in POP5.

Instead, POP5 contains one of the most abundant structural motifs in eukarya, the

RNA recognition motif (RRM), which are found in a variety of RNA-binding proteins involved in post-transcriptional events, including RNA processing, splicing and editing (96).

1.2.2.3.2 RPP30

The tertiary structure of Pho RPP30 was solved with and without POP5 by X- ray crystallography (95,97). The structure of RPP30 is the same in both cases. It contains10 α-helices and 7 β-strands in an α/β barrel structure, similar to the the well- known triose phosphate isomerase (TIM) structure (98). The major difference between RPP30 and typical TIM barrels is that RPP30 has helix α10 acting as a lid to

11 the barrel pore (Fig. 1.10). Among the RNA-binding proteins, RPP30 is similar to tRNA-guanine transglycosylase (TGT), which has a TIM barrel structure with tRNA binding sites located between α7/β7 and α8/β8. It is possible that RPP30 might have similar binding sites for pre-tRNA as does TGT (99).

1.2.2.3.3 RPP21

The structures of Pho and Pfu RPP21 were solved by X-ray crystallography

(100) and NMR (101), respectively. These proteins have a zinc-ribbon motif, where a single Zn2+ is coordinated by the sulfur atom of four invariant Cys residues. RPP21 forms an L-shaped structure. One arm comprises helices α1 and α2 in the N-terminal domain and the other arm contains three anti-parallel β-strands in the C-terminal domain. An unstructured loop connects these two arms (Fig. 1.10). Mutation of any of these invariant Cys residues results in loss of structure and activity suggesting the importance of Zn2+ for the structure and function of RPP21 (100). In fact, the only conserved sequence among Zn2+-binding proteins appears to be the two CXXC motifs to coordinate Zn2+. Zn2+-binding proteins, such as RPP21, typically use metal- ion binding to bring together structural elements and stabilize the overall tertiary structure.

1.2.2.3.4 RPP29

The structures of RPP29 from Mth, Pho and Archaeoglobus fulgidus (Afu) were solved by both NMR and X-ray crystallography (102-105). The central feature of

RPP29 shown in all these high-resolution studies is a sheet of six anti-parallel β- strands wrapped around a conserved hydrophobic core (Fig 1.10). Although crystal structures reveal well-formed helices in the N- and C-termini, which were not seen in

12 NMR structures, the overall structure resembles the Sm/Sm-like proteins and Hfq

(103,104), both are known to bind to RNAs (106,107).

1.2.2.4 Protein-protein interactions in archaeal RNase P

Yeast two-hybrid data on Mth (108) and Pho (109) RPPs and biochemical reconstitution studies with Pfu RNase P (87) revealed that the four RPPs function as pairs: (i) POP5 with RPP30 (POP5RPP30) and (ii) RPP21 with RPP29

(RPP21RPP29). These results are consistent with yeast two-hybrid analysis using human (92) and yeast RPPs (93). NMR spectroscopy studies on the Pfu

POP5RPP30 (94) and RPP21RPP29 (101) complexes revealed significant chemical-shift perturbations in the HSQC spectrum of each member of each pair upon addition of its corresponding partner. Moreover, the structure of each RPP pair has been solved by NMR spectroscopy and X-ray crystallography.

POP5RPP30: The crystal structure of POP5RPP30 was obtained as a heterotetramer consisting two copies of this binary complex in the asymmetric unit

[Fig. 1.12; (95)]. POP5 formed a homodimer in the middle with one copy of RPP30 placed symmetrically on either side. The homodimer is held by hydrogen-bonding while the heterodimer interface (POP5RPP30) consists of salt bridges and hydrophobic interactions. Interestingly, mutations disrupting homodimerization of

POP5 resulted in breakup of the heterotetramer into a heterodimer (judging by size exclusion chromatography) with weakened enzymatic activity (95). However, the basis of this reduced activity is not known. The stoichiometry of RPPs in RNase P holoenzyme awaits further investigation.

13 RPP21RPP29: The crystal structure of Pho RPP21RPP29 complex (Fig.

1.12) revealed that RPP21 uses helices (1 and 2) to interact with the N-terminal unstructured region, 2, and C-terminal helix 3 of RPP29 (110). The protein–protein interface is dominated by hydrogen bonds and salt bridges.

The structures of binary complexes have provided some testable hypotheses regarding RPP regions involved in recognition of RPR and pre-tRNA by RPPs. For example, the electrostatic surface maps of both POP5RPP30 and RPP21RPP29 complexes show an unequal charge distribution and predict the positively charged surfaces that may be involved in RNA binding (Fig 1.13 and 1.14).

1.2.3 Eukaryal RNase P

In eukarya, RNase P exists as multiple forms. In addition to a nuclear isoform,

RNase P variants are found in mitochondria and chloroplasts (discussed in section

1.2.4). Purification and characterization of native yeast and human nuclear RNase P have revealed that they contain one RPR and nine and ten RPPs, respectively

(9,111). Genetic depletion of all nine RPPs individually in yeast has demonstrated their essentiality for pre-tRNA processing and cell viability (9). Human RPPs are named RPP14, RPP20, RPP21, RPP25, RPP29, RPP30, RPP38, RPP40, POP5 and

POP1, where archaeal RPPs share homology with RPP21, RPP29, RPP30, and

POP5. In vitro pull down/cross-linking, yeast two hybrid, and yeast three hybrid experiments identified some protein-protein and protein-RNA interactions in the

RNase P holoenzyme (92,93,112-114). However, the specific roles of RPPs remain elusive due to the lack of a robust in vitro functional reconstitution and tertiary structures.

14 Interestingly, another essential endoribonuclease, RNase MRP, which exists only in eukarya, appears to be a paralog of RNase P. RNase MRP cleaves distinct substrates from RNase P (115), including the precursor of 5.8S rRNA (116), the mitochondria RNAs for generating the primers for mitochondria DNA replication (117), and the B-type cyclin mRNA (118). RNase MRP RNA shares a similar core structure with RNase P RNA, strongly suggesting that the RNA is the catalytic moiety in RNase

MRP (115). In yeast, RNase MRP has been biochemically purified and shown to share eight protein subunits with yeast nuclear RNase P except RPP21 (3), but contains two proteins that are unique to RNase MRP, Snm1 and Rmp1 (119,120).

The differences in substrate specificities between RNase P and RNase MRP could result from these unique proteins or structural changes between the RNAs or a combination of both (3).

1.2.4 Organellar RNase P

As a remnant possibly from their endosymbiotic predecessors, mitochondria and chloroplasts have retained a small and closed circular genome, where some organellar tRNAs are encoded. RNase P activity has been detected in both organelles, consistent with the need for tRNA 5′ maturation, and appears to be distinct from nuclear RNase P activity in some cases (121-124).

In yeast (Saccharomyces cerevisiae), mitochondrial RNase P is made of mitochondrially encoded essential RNA subunit (Rpm1) and a nuclearly encoded protein subunit (Rpm2) (125-128). Although organelles are thought to have evolved from purple bacterial or cyanobacterial endosymbionts, it appears that Rpm2 does not share homology to any known bacterial, archaeal or eukaryal RPPs (128). In addition,

15 Rpm1 homologs have been identified from several genome sequences of

Ascomycetes, a phylum of the kingdom Fungi, and have shown large variations in the

RNA size and secondary structure from bacterial RPRs (129,130).

Surprisingly, it seems an exception rather than the rule for yeast to still retain an RNA-based organellar RNase P, which was possibly inherited from its endosymbiotic ancestors. Mitochondrial RPR is absent from most of other eukaryal major branches and the chloroplast RPR gene is generally lost from the chloroplast genome of land plants (131). In recent studies, a new class of protein metallonucleases, proteinaceous RNase P (PRORP), has been suggested to carry out the function of RNase P in human mitochondria, and plant mitochondria and chloroplasts (132,133). PRORP contains a metallonuclease domain and two or three pentatricopeptides (PPR) motifs, which were implicated to bind to RNA (134). PRORP exhibits RNase P activity as a single protein in plant organelles. Moreover, this protein can replace bacterial RNase P (an RNP) in vivo (133). Since proteinaceous

RNase P is fully capable of processing pre-tRNAs in vivo, it provides a convincing piece of evidence for the notion that the functions of extant protein enzymes were once performed by RNA-based enzymes.

Interestingly, while PRORP is sufficient to process pre-tRNA in plant mitochondria and chloroplasts as a single protein (133), human mitochondrial RNase

P appears to be a patchwork of three unrelated proteins (132): (i) mitochondrial

RNase P protein 1 (MRPP1) is a homolog to a yeast methyltransferase, Trm10p, which methylates guanosine at position 9 of tRNA (135), suggesting it may be involved in substrate recognition; (ii) MRPP2 is a well known enzyme named

16 hydroxysteroid dehydrogenase 10, which is involved in branched-chain fatty acid β- oxidation (136); and (iii) MRPP3/PRORP is thought to be the catalytic center because its C-terminal domain is reminiscent of a metallonuclease active site. While proteinaceous RNase P have been found in organelles, recent studies have shown that the import of nuclear RPR into mitochondria is dependent on polynucleotide phosphorylase (PNPASE) expression and that decreased PNPASE activity in human mitochondria impaired mitochondrial tRNA processing (137). These results suggest that both RNA-based and proteinaceous RNase P might coexist in human mitochondria. The specific roles of these two RNase P variants in mitochondria await further investigation.

1.3 In vitro reconstitution of RNase P

The reaction mechanism of RNase P entails several steps: substrate binding,

Mg2+ coordination, chemical cleavage, and product release. An essential first step to dissect the function of individual subunits contributing to different steps is to reconstitute and study the different RNase P holoenzymes in vitro. Understanding the functions of RPR and RPPs mostly comes from the in vitro studies of the bacterial version. However, bacterial RNase P is much simpler and contains only one RPR and one RPP, which does not share sequence homology with archaeal and eukaryal

RPPs. Therefore, to study the functions of individual subunits in catalysis mediated by

RNase P with multiple RPPs, our laboratory reconstituted in vitro the archaeal variant as an experimental surrogate for the currently intractable eukaryal cousin [For bacterial RNase P, see section 1.2.1; (138)].

17 Functional reconstitution has been achieved for Mth, Pfu, and Pho RNase P by using in vitro transcribed RPR and recombinant RPPs purified from Escherichia coli (87,90,102). The results from multiple-turnover kinetic studies (87) have revealed that (i) while Pfu RPR is capable of multiple-turnover reactions, its activity can be further stimulated by addition of either binary complexes, POP5RPP30 or

RPP21RPP29, to RPR; (ii) addition of RPP pairs lowers the requirement of monovalent and divalent ions for pre-tRNA cleavage activity; (iii) POP5RPP30 increases the kcat of RPR by ~30-fold to the same extent of all four RPPs together, while RPP21RPP29 only increases kcat by 1.6-fold; (iv) addition of either

RPP21RPP29 or POP5RPP30 to RPR decreases KM by 5- or 3-fold, respectively; and (v) all four RPPs together cause a 25-fold increase in kcat and a 170-fold decrease in KM.

Since kcat represents the rate of the slowest step after substrate binding,

POP5RPP30 must play a vital role in chemical cleavage and/or product release under multiple-turnover conditions. A self-cleaving RPR, in which pre-tRNATyr is conjugated with Mja RPR in cis, was used as a model to focus on chemical cleavage step without influence from substrate binding and product release (88). This study showed that POP5RPP30, but not RPP21RPP29, can increase kobs by ~100-fold

(i.e. the rate of chemical cleavage) (88). However, the cis construct could not dissect the contribution of RPPs in substrate binding because the substrate is already conjugated with the RPR.

18 Eukaryal RNase P reconstitution has not been successful. Nevertheless, some progress has been made. Very weak activity of human RPR (H1 RNA) has

-5 -1 been detected (kobs = 10 min , pH 6) (13). The ability of RPP21 and RPP29 to reconstitute with H1 RNA also demonstrated their ability to enhance activity (139).

The observed increase could be attributed to the ability of RPP21 and RPP29 in stabilizing RPR folding and/or substrate binding (139). Although the protein subunit composition of RNase P in eukarya is significantly more complex than that in bacteria and archaea, it appears that not all of them are required for pre-tRNA cleavage. For example, a precursor form of yeast RNase P containing all RPPs but without RPP21 and RPP38 displayed the same steady-state rate for pre-tRNA processing as the mature RNase P containing all nine RPPs (140). We are therefore exploring if the insights from studies of the hierarchy of assembly in archaeal RNase P might be applicable to eukaryal RNase P.

1.4 Research objectives

1.4.1 Elucidate the roles of archaeal RPPs in aiding RPR-mediated cleavage

By studying bacterial RNase P, we have learned that RPP plays multiple roles in substrate binding, chemical cleavage, cleavage-site selection (fidelity) and Mg2+ coordination. However, very little is known about the functions of archaeal and eukaryal RPPs. In this thesis, I sought to extend our understanding of the roles of

RPPs in the protein-rich archaeal RNase P. Since archaeal RPPs share homology with eukaryal counterparts, our findings on RPPs could lay a foundation for investigating the currently intractable eukaryal RNase P.

19 1.4.1.1 Dissect the roles of archaeal RPPs in substrate binding and chemical cleavage

We first sought to dissect the roles of RPPs in catalysis by measuring maximal kobs and KM for pre-tRNA cleavage by RPR  RPPs under single-turnover conditions.

By comparing the kinetic parameters exhibited by the RPR assembled with different

RPPs, we sought to derive insights into how RPPs aid RNA catalysis. Single- turnover measurements allowed us to focus on steps before product release (i.e. substrate binding and chemical cleavage) and helped us to delineate the functions of

RPPs. We were also able to formulate a kinetic framework to better illustrate the functional cooperation among RPPs (see chapter 2).

1.4.1.2 Delineate the roles of archaeal RPPs in influencing the RPR’s cleavage- site selection

RNase P typically hydrolyzes the phosphodiester bond between the first nucleotide (N+1) of mature tRNAs and the preceding nucleotide in the leader sequence (N-1) (Fig. 1.1). We sought to determine the contribution of RPPs to cleavage-site selection by using a non-consensus substrate, pre-tRNAGln, harboring

A-1 and U+1 instead of U-1 and G+1 that are typically found in most pre-tRNAs (see

Fig. 3.1 and chapter 3). It has previously been shown that bacterial RNase P mis- cleaves bacterial pre-tRNAGln (141). To gain mechanistic insights into how archaeal

RPPs affect cleavage-site selection of this non-consensus substrate, we tethered pre- tRNAGln to Mja RPR, and measured both the rate of cleavage as well as the amount of both correct and mis-cleaved products generated. The results of this study, together with prior footprinting data which demonstrated that RPPs bind to distinct

20 sites on the RPR (142), shed light on how archaeal RPPs interact with pre-tRNAs and with RPRs to influence cleavage-site selection (chapter 3).

1.4.1.3 Elucidate the roles of archaeal RPPs in coordinating catalytic Mg2+

Previous studies have shown that P4 represents a universally conserved structure with a helix interrupted by a bulged uridine (Fig. 1.9) in RPRs from all three domains of life (78). Helix P4 has also been shown to be a Mg2+-binding site (143-

145). Alterations in this region caused a severe defect in the ability of bacterial RNase

P to coordinate Mg2+ important for cleavage (143). Since RNase P mediated catalysis is dependent on Mg2+, we decided to investigate the roles of archaeal RPPs in active site Mg2+ coordination by using mutant archaeal RPRs, where the geometry of the P4 region had been changed by deleting the bulged uridine (ΔU) or adding an additional uridine to the bulge (+U).

We first inquired whether archaeal RPPs can rescue the activity of mutant

RPRs (ΔU and +U) under multiple-turnover conditions. Second, to examine which archaeal RPP pair increases the RPR’s affinity for catalytically important Mg2+ ions, we utilized a cis construct, where pre-tRNATyr is conjugated to a mutant Mja RPR

(pre-tRNATyrMjaU), to determine the effect of individual RPP pairs in affecting coordination of Mg2+ ions critical for the chemical cleavage step (chapter 4).

21

Figure 1.1. Role of RNase P in tRNA maturation. RNase P hydrolyzes the phosphodiester bond between the first nucleotide (+1) in the mature tRNA and the last nucleotide in the leader sequence (-1) to remove the 5 leader sequence from precursor tRNAs and thus generate mature tRNAs. This hydrolysis is dependent on Mg2+.

22

Figure 1.2 Cladogram depicting a possible scheme for evolution of RNA-based RNase P. Scales are arbitrary. LUCA is the acronym from last universal common ancestor; RPR and RPP stand for RNase P RNA and RNase P protein, respectively. This figure is reproduced from ref (5).

23 Figure 1.3. Secondary structures of bacterial RPRs. Two different representations of the secondary structures of Escherichia coli (type A) RPR and Bacillus subtilis (type B) RPR. Nucleotides are designated by upper case letters and numbered from the 5′ end of the RPRs. Individual helices are given the designation P, for paired region. P4 and P6 pseudoknots are connected by square brackets. Non-paired regions between helices are given the designation J, for joining segments, and numbered with respect to the helices they connect. Dashed lines indicate the separation of two independently folding domains: specificity (S) domain and catalytic (C) domain. In the lower panels, thick lines connect adjacent nucleotides with arrows indicating 5′ to 3′ polarity. Nucleotides involved in tertiary interactions are boxed and linked by thin, dotted lines. Open and close circles indicate known non-canonical pairings. The same helices in each of these two representations are indicated by the same color. This figure is adapted from ref (31).

24

Figure 1.3.

25

Figure 1.4. A minimal bacterial consensus RPR deduced from phylogenetic analysis. Uppercase letters indicate universally conserved nucleotides; lowercase letters denote those which are at least 80% conserved but are not invariant. Filled circles represent nucleotides which are not conserved in identity but are present in all sequences; open circles depict those which are present in at least 80% sequences, but are absent in at least one. Base pairs in P4 are shown as squares and lines. This figure is reproduced from ref (34).

26 Figure 1.5. Structures of the specificity domain of type A (A) and type B (B) bacterial RPRs. The left panel shows the secondary structures of the RPR from Thermus thermophilus (Tth; A) and Bacillus subtilis (Bsu; B). The nucleotides in S domain are designated by upper case letters. Important tertiary contacts in the S domain are indicated by lines and arrows. The right panel shows the crystal structures of (A) Tth and (B) Bsu specificity domains. The figure is reproduced from ref (53).

27

Figure 1.5.

28

Figure 1.6. Inter-domain interactions in bacterial RPR. Paired regions are indicated by the prefix P, such as P2, P3 etc. The loops which cap a helix are designated by L and numbered according the capped helix. Bacterial RPR consists of two layers. Layer 1 contains the active site in the C domain and P7-P12 of the S domain [known to interact with T stem-loop of precursor tRNA (46)]. Layer 2 includes the P8/P9 coaxial stacked helix, which serves as a central organizer, held by loop:helix interactions between L18:P8 and L14:P8. This figure is reproduced from ref (30).

29

Figure 1.7. Tertiary structure of a bacterial RPP. Bacillus subtilis RPP (67). Red dashed lines indicate the central cleft, which is formed by one face of the central - sheet packing with helix α1 and is thought to interact directly with the pre-tRNA leader. The other face of β-sheet packs with helices α2 and α3 to form the hydrophobic core. The RNR motif in helix α2, which contains many conserved basic residues, is thought to bind to bacterial RPR. This picture is modified from ref (2).

30

Figure 1.8. Model of the RNase P holoenzyme. A stereoview of the tertiary structure model of Escherichia coli (Eco) RNase P (68). The docking of Eco RPP on its cognate RPR was based on a set of distance constraints, which were obtained from affinity cleavage of the RPR by EDTA-Fe covalently linked to the sole thiol in different single cysteine-substituted mutant derivatives of Eco RPP. This figure is reproduced from ref (68).

31

Figure 1.9. Comparison of secondary structures of RPRs in all three domains of life. The conserved nucleotides are shown in upper case letters. The dots refer to nucleotides which are not conserved but whose presence helps to preserve the arrangement of neighboring conserved nucleotides. This figure is reproduced from ref (11).

32

Figure 1.10. Tertiary structures of archaeal RPPs. Ribbon diagrams of Pfu POP5 (94), Pho RPP30 (97), Pho RPP21 (100), and Afu RPP29 (104). -helices and - strands are labeled in red and in blue, respectively.

33

Figure 1.11. Similarities in structures of bacterial RPP and archaeal POP5. Crystal structures of (A) Bacillus subtilis RPP (67) and (B) Pyrococcus furiosus POP5 (94). α-Helices, β-strands, and loops are colored red, blue, and grey, respectively. Although the primary and secondary structures of bacterial RPP and archaeal POP5 are different, they adopt similar tertiary structures.

34

Figure 1.12. Tertiary structures of archaeal RPP binary complexes. Pyrococcus horikoshii (Pho) POP5RPP30 (95) and Pho RPP21RPP29 (110). In each binary complex, one interacting partner is in blue and the other is in red. The POP5 homodimer-interface is shown by a shaded rectangle, and the two different POP5RPP30 interfaces by shaded ovals. Zn2+ and its coordinating cysteine side chains are depicted by a gray sphere and the red sticks in RPP21, respectively. The figure is reproduced from ref (79). 35

Figure 1.13. Electrostatic potential surface map of POP5RPP30. The POP5RPP30 binary complex crystallized as a heterotetramer in the crystallographic asymmetric unit. The surface potential is displayed as a color gradient from red (negative) to blue (positive), showing the relative strong electropositive character of the putative RNA binding sites. This figure is reproduced from ref (95). .

36

Figure 1.14. Electrostatic potential surface map suggests two RNA binding regions in the Pfu RPP21RPP29 complex. The surface potential is displayed as a color gradient from red (negative) to blue (positive). Left panels depict the electrostatic potential maps of the Pyrococcus furiosus (Pfu) RPP21RPP29 complex. There are two electropositive regions indicating two possible RNA-binding sites. Site 1 consists of residues from both proteins; site 2 is located only on RPP21. Right panel is the ribbon diagrams of Pfu RPP29 (red) and Pfu RPP21 (green) with highly conserved basic residues labeled. The orientation is the same for both panels. This figure is reproduced from ref (142).

37

CHAPTER 2

SINGLE-TURNOVER KINETIC STUDIES TO ELUCIDATE THE ROLES OF

ARCHAEAL RPPS IN AIDING RNA-MEDIATED CLEAVAGE OF PRECURSOR

TRANSFER RNA

2.1 Introduction

Several years after the remarkable finding that the bacterial RPR is a true RNA enzyme in the presence of Mg2+ and monovalent ions (6), archaeal and eukaryal

RPRs were also shown to be catalytically active in vitro (12,13). This common attribute of evolutionarily divergent RPRs was anticipated from their shared ancestry, attested by sequence and structural similarity of their putative catalytic core (11,27-

29,34,76,77,146). However, dramatic variations (106-fold) in catalytic potential indicate that not all RPRs are equal: activity of RPRs from bacteria > archaea > eukarya (6,12,13). Although RPRs display activity in vitro without RPPs, they are dependent on their cognate protein cofactors for cellular function. Interestingly, there is an inverse relationship between RPR activity and RNP composition: the protein:RNA mass ratio is 70% in eukaryal and 50% in archaeal RNase P compared to 10% in their bacterial counterpart. Thus, elucidating the intimate cooperation between the RNA and protein subunits of RNase P variants with differing RNP make- up offers a paradigm to understand how structural and functional attributes of RNAs might have been reassigned to protein cofactors during the evolutionary transition

38 from an RNA to RNP world (2-4,11). In this study, we focused our efforts on the simpler and biochemically tractable archaeal version, especially as an experimental surrogate for the eukaryal relative which is yet to be reconstituted in vitro. Rapid advances in functional reconstitution (87,88,90,102) and structural studies

(94,95,97,100-105,110,142) have validated this choice.

It has been demonstrated earlier that robust RNase P activity could be obtained from assembling recombinant subunits of Pyrococcus furiosus (Pfu; type A), and that the four RPPs functioned as two binary complexes (POP5RPP30 and

RPP21RPP29) (87). Although a previous study from our laboratory showed that only the addition of Pfu POP5RPP30 to Pfu RPR increases its kcat (87), if product release is rate limiting [as demonstrated for bacterial and yeast RNase P (59,147)], it is not possible to determine RPP-mediated increases in the rate of cleavage under multiple- turnover conditions. Therefore, we have now employed single-turnover kinetic studies to gain insights into the roles of archaeal RPPs. Results from these new studies permit formulation of a kinetic framework to highlight the functional cooperation among RPPs. These findings, together with the structures of the binary RPPs

(95,110,142), should aid efforts to establish structure-function correlations in archaeal

RNase P.

2.2 Materials and methods

2.2.1 Generation of Methanothermobacter thermautotrophicus RPR using in vitro transcription

The cloning and transcription of Methanothermobacter thermautotrophicus

(Mth) RPR is described elsewhere (102). The Mth RPR was then generated using

39 EcoRI-linearized pUC19-Mth RPR as the template DNA for T7 RNA polymerase- mediated run-off transcription (87). The RNA thus generated was subjected to dialysis to remove unincorporated rNTPs and the concentration determined from Abs260 measurements and its extinction coefficient.

2.2.2 Cloning the genes encoding Mth RPPs

The cloning of Mth RPPs is described elsewhere (Chen at el. 2010).

2.2.3 Overexpression and purification of protein subunits of Mth RNase P

The overexpression and purification of Mth RPPs was initially developed by I-

Ming Cho and Dr. Venkat Gopalan. I optimized and fine-tuned the procedures described here. For the preparation of Mth RPPs: Escherichia coli BL21(DE3) cells freshly transformed with MthRPPTC1 (a plasmid which encodes Mth POP5 and

RPP30 in tandem) or MthRPPTC2 MthRPPTC1 (a plasmid which encodes Mth

RPP21 and RPP29 in tandem) were inoculated into 5 mL Luria Broth (LB) media containing 35 µg/mL kanamycin, and grown overnight at 37C with shaking. These overnight cultures were used to inoculate 500 ml of fresh LB media containing 35

µg/mL kanamycin. The cells were grown at 37°C with shaking until the cells reached an optical density at 600 nm of 0.6-0.8. Protein overexpression was induced by adding isopropyl--D-thiogalactopyranoside (IPTG) to a final concentration of 1 mM.

To the RPP21RPP29 expressing cultures, ZnCl2 was also added to a final concentration of 1 mM at the time of addition of IPTG. The cells were then allowed to grow at 37°C for another 3 h (RPP21RPP29) or room temperature for 15 h

(POP5RPP30) and harvested by centrifugation. The cell pellets were stored at -80C until further use.

40 The whole purification process was performed on ice. Frozen cells were re- suspended in 20 mL of buffer A [25 mM Tris-HCl (pH 7.5), 5 mM DTT, 0.1 mM PMSF].

Five mL of buffer S [25 mM Tris-HCl (pH 7.5), 5 mM DTT, 0.1 mM PMSF and 5 M

NaCl] was added and the cells were lysed by sonication. Cell debris was pelleted by centrifugation (9,000 g, 15 min, 4C). Polyethylenimine (PEI) was added to the cleared supernatant to a final concentration of 0.025% (v/v) and incubated on ice for

30 min. The PEI-precipitated nucleic acid was pelleted by centrifugation (9,000 g, 15 min, 4C). To the resulting soluble fraction, finely powdered (NH4)2SO4 (to a final saturation of 40%) was added slowly. The precipitation was performed over a period of 60 min with constant stirring on ice. The precipitated RPPs were recovered by centrifugation (9,000 g, 15 min, 4C). The POP5RPP30 precipitate was dissolved 10 ml of 25 mM Tris-HCl (pH 7.5) containing 375 mM (NH4)2SO4, 5 mM DTT, and 0.1 mM PMSF. The RPP21RPP29 precipitate was dissolved 10 ml of 25 mM Tris-HCl

(pH 7.5) containing 250 mM (NH4)2SO4, 5 mM DTT, and 0.1 mM PMSF. The solution was filtered through a 0.4 m filter and loaded on a pre-equilibrated SP-Sepharose

(GE Healthcare) column. The bound proteins were eluted with a 0 to 2 M NaCl gradient in buffer A. Fractions containing the archaeal RPPs were identified by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) followed by

Coomassie Blue staining. Binary complexes were typically eluted between 1.2 and

1.5 M NaCl. Using the extinction coefficients of the binary RPP complexes and their absorption at 280 nm, the concentration of each binary complex was calculated. The purified RPP complexes were dialyzed against buffer containing 50 mM Tris-acetate

(pH 5.8), 0.8 M NH4OAc, and 10 mM MgCl2 and stored at room temperature until further use.

41 2.2.4 Mth RNase P single-turnover assays

All reconstitutions and assays were performed in a thermal cycler. The Mth

RPR was folded as follows: incubation at 50°C for 50 min in water followed by 37°C for 30 min in 2-(N-morpholino)ethanesulfonic acid (MES)-HCl, pH 5.8 (unless otherwise indicated) containing 10 mM MgCl2 and 800 mM NH4OAc. The folded RPR was assayed either alone or with a two-fold molar excess of RPPs using the optimal concentrations of MgCl2 and NH4OAc for each RNP (Table 2.2). Optimal RNP assembly entailed successive incubations at 37°C and 55°C for 10 min each. Assays were initiated by adding trace amount Escherichia coli pre-tRNATyr labeled with [α-P32]

GTP, which had been pre-incubated at 55°C for 2 min. In the case of RNPs, the amount of RPR in the assay was used as the concentration of enzyme based on the assumption that all of the RPR is assembled into RNPs under the conditions employed.

To determine the maximal kobs (max. kobs) under single-turnover conditions at

55oC, we incubated ~2 nM pre-tRNATyr with a range of enzyme concentrations: RPR,

0.3 to 20 M; RPR + RPP21RPP29, 0.5 to 3 M; RPR + POP5RPP30, 1 to 10 M; and RPR + 4 RPPs, 0.3 to 3 M. All assays involved incubation for a specified period at 55°C in a thermal cycler. Reactions were terminated by adding an equal volume of stop solution [10 M urea, 5 mM EDTA, 0.05% (w/v) bromophenol blue, 0.05% (w/v) xylene cyanol, 10% (v/v) phenol]. For short incubations (e.g., 5 s), reactions were first terminated by immersing the reaction tubes in liquid nitrogen before adding stop solution. The reaction contents were separated using denaturing PAGE [8% (w/v) polyacrylamide, 7 M urea]. Because the Mth RPR + 4 RPPs reaction is too rapid at pH 5.8 (t1/2 ≤ 8 s) when [E] > 1 M, we could obtain reliable data only by decreasing

42 the assay pH to 5.4. To establish the dependence of the rates of product formation catalyzed by Mth RPR + 4 RPPs on assay pH from 5.4 to 6.15, the assays were performed in 20 mM instead of 30 mM Mg2+ (Fig. 2.5).

2.2.5 Mth RNase P single-turnover data analysis

After denaturing PAGE, the reaction products were visualized by phosphorimaging on the Typhoon (GE Healthcare). The resulting bands were quantitated by ImageQuant (GE Healthcare) to assess the extent of substrate cleaved.

To obtain the rate of product formation (kobs) in single-turnover reactions, the %

-kt product formed at time t (Pt) was fit to Pt = P∞(1 - e ) using Kaleidagraph software

(Synergy). For optimal curve fits, the amplitudes were defined based on experimentally-observed values (Fig. 2.2). The individual curve-fit errors for kobs did not exceed 8%. For all kinetic studies, at least three replicates were performed to obtain the mean and standard deviation values.

The plot of kobs versus [E0] displayed hyperbolic dependence on the Mth RPR in the absence and presence of its RPPs (Fig. 2.4). Kaleidagraph was used to fit these data to:

Max. kobs[E0 kobs M( O) [E0 to derive values for max. kobs and KM(STO).

2.2.6 Pulse-chase experiments with Mth RNase P to demonstrate that the dissociation rate (k-1) is greater than the rate of chemical cleavage (k2) under single-turnover conditions.

43 These pulse-chase experiments were performed essentially as described elsewhere for studies on the hammerhead ribozyme (148). RPR folding, RNP assembly and assay conditions are as described in section 2.2.4. Mth RPR (16.6 µM) was mixed with a trace amount of internally labeled pre-tRNATyr (~2 nM) and incubated at 55°C in a final reaction volume of 20 µl. From this mixture, two 5-µl aliquots were diluted 100-fold in 55°C pre-warmed assay buffer [50 mM MES (pH 5.8),

500 mM MgCl2, 2 M NH4OAc], one at 20 min and the other at 30 min after the initiation of the reaction; the remainder proceeded without any dilution. Similarly, for

Mth RPR + RPP21RPP29 (3 M), two 5-µl aliquots were diluted 100-fold in pre- warmed assay buffer [50 mM MES (pH 5.8), 120 mM MgCl2, 800 mM NH4OAc], one at 5 min and the other at 10 min after the initiation of the reaction. From both the undiluted and diluted reactions, aliquots were removed at defined time intervals to measure the product formed. Reactions were terminated by adding stop solution [10

M urea, 5 mM EDTA, 0.05% (w/v) bromophenol blue, 0.05% (w/v) xylene cyanol, 10%

(v/v) phenol]. The reaction products were separated on denaturing PAGE and quantitated as described.

2.3 Results

2.3.1 Rationale

We purified archaeal RPPs as two binary complexes because functional assays, yeast-two-hybrid and high-resolution structural studies had already established the pair-wise interactions between POP5RPP30 and RPP21RPP29

(See chapter 1). We have now employed single-turnover kinetic measurements, wherein enzyme concentrations far exceed that of the substrate, to specifically

44 measure the rate of cleavage by the RPR either alone or when aided by each RPP pair.

2.3.2 Purification of the subunits of Mth RNase P

Mth RPR was generated using run-off in vitro transcription as described. The four protein subunits (Table 2.1) were overexpressed in Escherichia coli BL21(DE3) cells and purified using cation-exchange chromatography (Fig. 2.1).

2.3.3 Optimal assay conditions for single-turnover kinetic measurements of

RPR and RNPs

To determine the role of each RPP pair in aiding RPR’s catalysis, we compared the first-order rate constant (kobs) of pre-tRNA processing by Mth RPR either alone or in the presence of two- and four-protein combinations. Before we initiated such comparative experiments, we first sought to establish the optimal conditions for each RNP complex.

Like bacterial and eukaryal RNase P, archaeal RNase P requires a

+ monovalent ion, such as NH4 , for RPR folding and a divalent metal ion, preferably

Mg2+, for RPR folding and catalysis (87,88,90). Therefore, we tested different

2+ + concentrations of Mg in combination with different concentrations of NH4 for the

RPR  RPPs to determine the maximal activity under single-turnover conditions. The

2+ + optimal concentrations of Mg and NH4 for Mth RPR alone reaction is 500 mM and 2

M, respectively, which are identical to what have been reported earlier for Pfu RPR

+ alone under multiple turn-over condition (87). While 2 M NH4 is needed for the RPR-

+ alone reaction, 2 M NH4 inhibits the activity of RNPs, presumably by interfering with

45 + RNA-protein interactions. The optimal concentration of NH4 for all RNPs is 800 mM and consistent with previous reports (87,88). RPPs are able to lower the requirement of Mg2+: the optimal Mg2+ concentrations was lowered to 120 mM for RPR +

POP5RPP30 and RPR + RPP21RPP29, and further down to 30 mM for RPR +

4RPPs (data not shown).

Since we failed to determine the KD for binding of the binary RPPs to RPR by electrophoresis mobility-shift assays (EMSA), we determined the optimal stoichiometry of RPPs to RPR by titrating different amount of RPPs to a fixed amount of RPR and measuring the rate. The ratio of RPPs:RPR which resulted in maximal activity was considered the optimal ratio required to ensure the near-complete assembly of RPR to form RNP complexes. We observed that a 2-fold molar excess of

RPPs to RPR is adequate for most RNP assembly (data not shown). Note that this ratio was empirically determined depending on the RPR (enzyme) concentration used.

2.3.4 Single-turnover assays of partially and fully reconstituted Mth RNase P holoenzymes

Once optimal assay conditions were obtained, we then performed comparative single-turnover kinetic measurements to compare the RPR’s rate of pre- tRNA cleavage with and without RPPs. To facilitate manual single-turnover measurements, we slowed down the reaction rate by decreasing the assay pH from the typical 7.5 to 5.8, since the hydroxide nucleophile, which attacks the scissile phosphodiester linkage in the pre-tRNA, is believed to result from deprotonation of a hydrated Mg2+ ion in the RPR’s active site (149-152).

46 An excess of enzyme over substrate ([E] = 0.3-20 M, [S] ~ 2 nM) was used in our single-turnover studies with Mth RNase P. We found that a single-exponential function describes the rate of product formation by the Mth RPR  RPPs (Fig. 2.2).

We use max. kobs to indicate rates determined at saturating concentrations of the RPR

2+ +  RPPs in the presence of optimal levels of Mg and NH4 . The term KM(STO) is used to refer to the KM calculated under single-turnover conditions.

To justify using KM(STO) [(k-1 + k2)/k1] as a measure of KS (k-1/k1), we performed a pulse-chase experiment to investigate if the ES complex dissociates faster than substrate cleavage (i.e., k-1 >> k2; Scheme I) under the single-turnover conditions used. Through large dilution of a pre-formed Mth RNase P–pre-tRNATyr complex, we dissociated the substrate from the enzyme and expected a plateau in product formation post-dilution if k-1 >> k2 (due to Scheme I the pre-tRNA’s inability to rebind for cleavage). Indeed, when the reactions catalyzed by Mth RPR  RPP21RPP29 were diluted few minutes after mixing with pre-tRNATyr, product formation did not increase post dilution (Fig. 2.3).

At pH 5.8 and 55oC, both Mth POP5RPP30 and RPP21RPP29 decreased

2+ + Tyr the concentration of Mg and NH4 required for RPR-mediated pre-tRNA processing; the Mg2+ requirement decreased from 500 to 120 mM with either binary complex and to 30 mM with both (Table 2.2A). The max. kobs for processing of pre- tRNATyr by Mth RPR increased 60-fold upon addition of POP5RPP30 with no further change upon inclusion of RPP21RPP29 (Table 2.2A; Fig. 2.4). Also, RPP21RPP29 alone was unable to increase the RPR’s max. kobs. Despite the inability of

47 RPP21RPP29 to increase the max. kobs, it was able to reduce KM(STO) from 21.3 M to 1.4 M (to the same extent as four RPPs; Table 2.2A). These results demonstrate the importance of POP5RPP30 in cleavage and RPP21RPP29 in substrate binding

(however, see Discussion for additional comments on the role of POP5RPP30).

2.3.5 Comparison of partially and fully reconstituted Mth RNase P at 30 mM

Mg2+

The above studies were performed at the optimal Mg2+ concentration for each of the RNP complexes assembled with Mth RPR. We inquired if the results would be different if the Mth RPR was assayed with and without each binary complex at 30 mM

2+ + Mg and 800 mM NH4 , a condition optimal for the holoenzyme assembled with all four RPPs but not for the partial RNPs (Table 2.2B). There is no detectable activity in the RPR-alone reaction and weak (not reliably quantifiable) activity with RPR +

-1 RPP21RPP29. In contrast, the RPR + POP5RPP30 exhibits a kobs of 0.016 min , which is increased to 5.5 min-1 (344-fold) upon addition of RPP21RPPP29, suggesting that the latter significantly facilitates catalysis at lower Mg2+ concentrations.

2.4 Discussion

2.4.1 Role of protein cofactors in Mth RNase P catalysis

The first hint into the role of POP5RPP30 in promoting pre-tRNA cleavage came from multiple-turnover studies on Pfu RNase P where it was shown to increase

Tyr the RPR’s kcat by 25-fold (87). A self-cleaving RPR, where pre-tRNA was conjugated to Mja RPR in cis as a model system for studying rate-limiting chemistry, was used and revealed that the rate of self-cleavage of this conjugate was accelerated ~100-fold by Mja POP5RPP30, but not at all by RPP21RPP29 (88).

However, if RPP21RPP29 plays a role in substrate positioning that affects the rate of

48 cleavage, such an effect would manifest during pre-tRNA cleavage in a trans rather than in a cis reaction where the substrate is already docked. The single-turnover studies with Mth RNase P now provide new insights on the division of labor between the two binary RPPs. Only RPP21RPP29 decreased the KM(STO) of Mth RPR, while only POP5RPP30 elicited a 60-fold increase in the max. kobs (Table 2.2A, Fig. 2.4).

Our archaeal RNase P data could be interpreted by a framework similar to that used to describe the role of the bacterial RPP (Scheme I; Fig. 2.6). Various kinetic and structural studies on bacterial RNase P have indicated that subsequent to substrate binding, a conformational change converts ES to ES* (defined by the

2+ equilibrium constant Kconf) and optimally positions the pre-tRNA and catalytic Mg ions for cleavage (at rate kc) (62,153-155). Adding to a growing body of evidence supporting such a two-step mechanism, data from recent stopped-flow kinetic studies

(155) confirm an initial bi-molecular collision (E+SES) followed by a uni-molecular conformational change (ESES*). Thus, under single-turnover conditions with saturating concentrations of enzyme, max. kobs = kc(Kconf/1+Kconf). Based on kinetic studies with bacterial RPR ± RPP, Sun et al. (62) concluded that the RPP enhances the RPR’s rate of pre-tRNA cleavage by increasing Kconf and not kc, an inference supported by a recent finding that the bacterial RPP slows the reverse isomerization step (i.e., ES*ES) (155).

Since Mth RPR is capable of pre-tRNA processing in the absence of RPPs, a direct increase in kc upon addition of Mth RPPs is less likely because it would require an alternative mechanism for the RNP compared to the RPR, an unsupported premise. Therefore, we hypothesize that increased conversion of ESES* must underlie the ability of cognate POP5RPP30 to elicit a 60- and 100-fold increase, 49 Tyr respectively, in the Mth RPR’s max. kobs (Fig. 2.4) and the pre-tRNA —Mja RPR self- cleavage rate (88). Because POP5RPP30 significantly increases the rate of the cleavage step, we cannot assess if KM(STO)~KS (as we did for RPP21RPP29; Fig. 2.3).

Thus, without determining the microscopic rate constants that contribute to KS, we cannot rule out the role of POP5RPP30 in substrate binding. Since both bacterial

RPPs and POP5 adopt an RRM fold (67,94), and the central cleft in the -sandwich structure of the bacterial RPP binds the pre-tRNA leader (60), it is likely that POP5 (or

POP5RPP30) performs a similar role.

2.4.2 The importance of RPP21RPP29 in vivo

Although an analysis of maximal kobs values under non-physiological assay conditions is useful in assessing the maximum catalytic potential for the RPR and each RNP assembled with it, such an approach is likely to undervalue the contribution of RPPs under conditions resembling those found in vivo. For example, the failure of

Mth RPP21RPP29 to enhance the RPR’s max. kobs indicates that it does not favorably alter ESES*, however, it enhances the affinity of the RNP for Mg2+ as evidenced by its ability to increase by 344-fold the kobs of Mth RPR + POP5RPP30 at

30 mM Mg2+ but not at 120 mM Mg2+ (Table 2.2B). We chose this lower Mg2+ concentration as it is optimal for the activity of the holoenzyme in our reconstitution system.

In addition, its key role in substrate binding is borne out by its ability to decrease KM(STO) (i.e., KS; Table 2.2A and Fig. 2.4). In fact, human RPP21 has been shown to bind pre-tRNA (156), and the solution structure of Pfu RPP21RPP29 reveals two electropositive surfaces, with the smaller one (exclusively in RPP21)

50 postulated to bind pre-tRNA (142). Collectively, these results highlight its ability to lower the requirement of Mg2+ and the essentiality of RPP21RPP29 in vivo.

2.4.3 Comparison of Mth RPR data to the rates of pre-tRNA cleavage by bacterial and archaeal RPRs reported in previous studies

Gly Although Li et al. (14) did not report a max. kobs, they documented pre-tRNA cleavage by 10 M Mth RPR at pH 6 with a rate of 0.034 min-1, which is similar to the

0.04 min-1 that we observed for cleavage of pre-tRNATyr by 10 M Mth RPR at pH 5.8; this coincidence is reassuring given the different substrates and assay conditions

Tyr used in these two studies. Second, the max. kobs for pre-tRNA cleavage by Mth

RPR is 0.13 min-1 at pH 5.8 and 55oC (Table 1) compared to the bacterial RPR’s max.

-1 o kobs of ~5 min at pH 6 and 37 C. These data, together with earlier studies on chimeric Mth:Eco RPRs (14), reaffirm the idea that the bacterial RPR structural elements which are missing in the type A archaeal RPR contribute to both substrate binding and cleavage rate.

2.4.4 Summary

High-resolution structures reveal that the bacterial RPR is arranged as two one-helix thick layers, with the larger layer 1 juxtaposing the S domain’s substrate- recognition elements and the C domain’s active site (30,55). Intra-molecular braces in layer 2 are vital for the precise orientation of the S and C domains, whose cooperation is essential for efficient catalysis. Although the bacterial RPR is capable of generating a functional tertiary fold in vitro at high ionic strength, association with a single RPP enhances its activity at lower (physiological) Mg2+ concentrations and broadens its substrate-specificity likely due to a gain of new substrate-recognition

51 determinants (62,70,157). One might expect that some of the archaeal/eukaryal

RPP(s) would exhibit equivalent catalytic functions as the bacterial RPP. In addition, since the bacterial RPR’s layer 2 struts are missing in archaeal/eukaryal RPRs, some archaeal/eukaryal RPPs would be predicted to substitute for these missing RNA-RNA tertiary contacts and play structural roles (11).

Footprinting studies provide a structural perspective to better appreciate the functional coordination among the two RPP complexes in aiding the RPR. Those results indicate that POP5RPP30 and RPP21RPP29 interact almost mutually exclusively with the C and S domains, respectively [Fig. 2.7; (87,142)]. Our finding that archaeal POP5RPP30 could increase the max. kobs is consistent with the footprint of POP5RPP30 on RPR’s C domain (68,87,142).

We postulate that one function of archaeal RPP21RPP29 is to substitute for the intra-molecular struts found in bacterial RPR, therefore reduce the need for structural Mg2+. Such a premise is consistent with its inability to alter the rate and its ability to lower the Mg2+ requirement. Results from an independent substrate- recognition study examining the ratios of correct:aberrant cleavages of model substrates indicate that binding of RPP21RPP29 to the archaeal RPR’s domain is critical for optimal recognition of the T stem-loop (TSL) region in the pre-tRNA [Fig.

2.7; (158)]. Akin to the bacterial RPR scenario, this productive TSL-S domain interaction might elicit a conformational change that aids catalysis by positioning the chemical groups and Mg2+ near the cleavage site in the C domain (46,51,153,159).

52 Isoelectric Predicted RPP Gene ID point (pI) mass, Da POP5 MTH687 10.5 14,568 RPP21 MTH1618 11.1 16,936 RPP29 MTH11 11.4 10,722

RPP30 MTH688 8.2 27,662

Table 2.1. Characteristics of RPPs from Methanothermobacter thermautotrophicus Note: Gene IDs and sequences from the Comprehensive Microbial Resource, J. Craig Venter Institute (http://cmr.jcvi.org).

53 A. Assayed under Relative the optimal Max. k *, min-1 K *, M [NH +], M [Mg2+], M max. condition for each obs M(STO) 4 k catalytic entity obs Mth RPR 0.13 ± 0.01 21.3 ± 2.6 2.0 0.50 1 + RPP21RPP29 0.13 ± 0.01 1.4 ± 0.2 0.8 0.12 1 + POP5RPP30 7.9 ± 1 11.8 ± 2.3 0.8 0.12 ~60 + Both binary 0.8 0.03 ~60 7.5 ± 0.14** 1.2 ± 0.1 RPP complexes

B. Assayed under the optimal condition for -1 the holoenzyme with four RPPs kobs, min Relative kobs 2+ + (30 mM Mg , 800 mM NH4 ) Mth RPR (3 M) *** - + RPP21RPP29 *** - + POP5RPP30 0.016 ± 0.0004 1 + Both binary RPP complexes 5.5 ± 0.18** ~344

Table 2.2. Effect of Mth RPPs on the ionic requirements and rate of cleavage of pre-tRNATyr by Mth RPR at 55C* * The standard errors of the curve fits shown in Figure 2.4 are indicated in the

estimates of max. kobs and KM(STO). ** All experiments were performed at pH 5.8, except for Mth RPR reconstituted with both binary RPP complexes, which was assayed at pH 5.4. After establishing a

slope of ~1 in a plot of log kobs versus pH (see Fig. 2.5), we multiplied the maximal rate observed at pH 5.4 by 2.5 to obtain the rate that would have been observed at pH 5.8 should it have been measurable. *** Reliable data could not be obtained due to weak or negligible activity.

54

Figure 2.1. Purification of recombinant Mth RPPs as binary complexes: POP5RPP30 (A), RPP21RPP29 (B). M indicates size marker. UI and I denote the crude extracts from un-induced and induced cultures, respectively. IP and FT represent the ammonium sulfate-fractionated sample that was subjected to the SP- sepharose column chromatography and the flow-though, respectively.

55 Figure 2.2. Effects of Mth RPPs on the single-turnover rate of RPR-catalyzed pre-tRNA cleavage. (A) Increased product formation as a function of incubation time is illustrated in a representative time-course of pre-tRNATyr processing by the Mth RNase P holoenzyme. (B) Examples of the curve fits of the rate of product formation by Mth RPR with and without RPPs under their respective optimal conditions. The concentrations of Mth RPR, RPR + POP5RPP30, RPR + RPP21RPP29 and RPR + 4 RPPs used were 1.5 M, 1.2 M, 1.2 M and 0.5 M, respectively. Most of these ES complexes are productive as indicated by their ≥ 90% amplitudes, except for the reaction with RPR + RPP21RPP29. Since we use a two-fold stoichiometric excess of RPPs:RPR to facilitate assembly of the respective RNP, it is possible that any RPP21RPP29 uncomplexed with the RPR engages in RPR-independent interactions with the pre-tRNA substrate and lowers the amplitude to 40%; however, this problem does not exist when RPP21RPP29 is used in the context of 4 RPPs, presumably due to differences in assay conditions and/or the presence of POP5RPP30.

56

Figure 2.2.

57

Figure 2.3. Pulse-chase experiments with Mth RNase P to demonstrate that the dissociation rate (k-1) is greater than the rate of chemical cleavage (k2) under single-turnover conditions. (A) Mth RPR (16.6 M) and pre-tRNATyr (2 nM) assayed in 50 mM MES (pH 5.8), 2 M NH4OAc and 500 mM MgCl2: the rate of chemical cleavage was measured without dilution () and after dilution at 20 min () or 30 min (). (B) Mth RPR + RPP21RPP29 (3 M) and pre-tRNATyr (2 nM) assayed in 50 mM

MES (pH 5.8), 800 mM NH4OAc and 120 mM MgCl2: the rate of chemical cleavage was measured without dilution () and after dilution at 5 min () or 10 min (). The green and blue lines indicate the plateau theoretically expected post-dilution when k-1

>> k2.

58

Figure 2.4. Effects of Mth RPPs on the single-turnover rate of RPR-catalyzed pre-tRNA cleavage. The rates of product formation (kobs) by Mth RPR with and without RPPs were determined under single-turnover conditions and plotted as a function of the concentration of the respective catalytic entity to obtain the max. kobs and KM(STO) reported in Table 2.2.

59

Figure 2.5. Dependence of kobs on assay pH for Mth RPR + 4 RPPs. These single- turnover assays were performed with a vast excess of enzyme (3 M) over the substrate (2 nM), and in 50 mM MES (indicated pH), 800 mM NH4OAc and 20 mM 2+ 2+ MgCl2. Note that the assays were performed in 20 mM Mg and not 30 mM Mg , which is optimal for the RPR + 4 RPPs; this change lowered the rate by ~6-fold and permitted manual rate measurements over the pH range used in this experiment.

60

Figure 2.6. Kinetic scheme as a framework to interpret the contribution of the two binary RPP complexes to archaeal RNase P catalysis. For simplicity, the scheme does not take into account possible thermodynamic coupling in the binding of the RPPs (or pre-tRNA) and Mg2+ to the RPR. While the role of RPPs to binding of Mg2+ ions is indicated, no distinction is made between metal ions essential for RPR folding and catalysis. Although POP5RPP30 reduced KM(STO) by two-fold in our single-turnover kinetic studies, in the absence of detailed binding studies, we are unable to comment on the role of this binary complex in substrate binding; this uncertainty is indicated by a question mark.

61

Figure 2.7. Schematic drawing of the RPP footprinting data depicted on a secondary-structure model of Methanocaldococcus jannaschii (Mja) RPR and possible interactions between RPR and pre-tRNA. The ”outlined” letters represent universally conserved nucleotides among all RPRs. Circled and boxed nucleotides depict protection to RNase T1 and RNase V1, respectively (142); blue and red colors indicate protection by POP5RPP30 and RPP21RPP29, respectively. Green color refers to an RPR position that showed an increased susceptibility of to RNase T1 in the presence of POP5RPP30. The dashed red line separates the S and C domains of Mja RPR. The solid red lines indicate the proposed interaction between the RPR’s S domain and the pre-tRNA’s T-stem and loop. The solid blue lines indicate the proposed interaction between the RPR’s C domain and the pre-tRNA’s acceptor stem, 3′- and 5′-termini. Scissors indicate the cleavage site in the pre-tRNA. This figure is adapted from (142).

62

CHAPTER 3

ELUCIDATING THE ROLE OF ARCHAEAL RPPS IN INFLUENCING THE RPR’S

CLEAVAGE SITE SELECTION

3.1 Introduction

Transfer RNAs (tRNAs) play a crucial role in translation. Their biogenesis involves many steps including 5′ and 3′ maturation, intron splicing (where applicable), and nucleotide modification before their use in translation (160). Ribonuclease P

(RNase P) catalyzes removal of the 5′-leader from precursor tRNAs (pre-tRNAs) in all three domains of life (2-5). In addition to pre-tRNAs as its primary substrates, RNase

P is also responsible for processing other non-coding RNAs, such as precursors to

4.5S RNA, tmRNA, some viral RNAs, C4 antisense RNA from bacteriophage P1 and

P7, and metastasis-associated lung adenocarcinoma transcript 1 (MALAT1), a long non-coding RNA known to be upregulated in many human cancers (161-164).

RNase P is a ribonucleoprotein (RNP) complex containing one catalytic

RNase P RNA (RPR) and a varying number of RNase P proteins (RPPs): one RPP in bacteria, at least four RPPs in archaea, and up to 10 RPPs in eukarya (2-5). Although pre-tRNA cleavage is associated with the RPR (6,12,13), both RPR and RPP(s) are essential for cell viability in vivo (9,15,16). While the bacterial RPP does not share homology with RPPs in archaea and eukarya (94), all four archaeal RPPs (POP5,

63 RPP30, RPP21, and RPP29) share homology with eukaryal RPPs and function as binary complexes (POP5RPP30 and RPP21RPP29) (8,87,88). Concomitant with the increasing protein content (10% in bacterial, 50% in archaeal and 70% in eukaryal

RNase P) (11), the activity of RPRs also decreases dramatically (106-fold; bacterial > archaeal > eukaryal) (6,12,13). Therefore, it is conceivable that some functional attributes may have been reassigned from RNA to protein subunits in archaeal/eukaryal RNase P, compared to their bacterial cousin (2-4,11).

Bacterial RNase P typically hydrolyzes the phosphodiester bond between the first nucleotide (N+1) in the mature tRNA and its preceding nucleotide in the 5′-leader

(N-1) (Fig. 1.1), and generates a 7-base pair (bp) amino acid acceptor stem; the

His SeCys exceptions are pre-tRNA and pre-tRNA , which are cleaved between N-2 and N-1 to generate an 8-bp acceptor stem (165,166). The number of bp in the acceptor stem of mature tRNAs is an important recognition-determinant for a tRNA to be charged with an amino acid by aminoacyl tRNA synthetases. Therefore, the ability of RNase P to select the correct cleavage site in pre-tRNAs is critical for subsequent utilization of tRNAs.

Chemical modification interference mapping (167,168), cross-linking

(35,36,80), footprinting (169), nucleotides analog interference mapping

(38,39,42,170-174) and mutagenesis (41,51,175) have revealed the functional groups that are important for the interactions between RNase P and pre-tRNAs. Many of the important nucleotides in the pre-tRNAs interacting with RNase P are clustered along the coaxially stacked helix consisting of the acceptor stem and T stem-loop (Acc-T- helical stack) (176). This Acc-T-helical stack engages in the following interactions with

64 bacterial RPRs: (i) the T-stem loop (TSL) region in the pre-tRNA interacts with P7-

P11 region in the RPR, which is referred to the TSL-binding site (TBS) (46,51,159); (ii)

3′-RCC sequence of the pre-tRNA interacts with a conserved GGU sequence in the

L15 loop of RPR through Watson-Crick interactions (49); and (iii) the interaction between the nucleotide at position -1 (N-1) in the pre-tRNA and the adenosine at position 248 (A248) in the RPR (Escherichia coli RPR numbering). Note: The N-1A248 interaction is consistent with canonical Watson-Crick or cis Watson-Crick/Watson-

Crick complementarity although the function is not solely dictated by base-pairing geometry (47,177). The bacterial RPP increases the affinity for pre-tRNA by binding to the 5′-leader of pre-tRNAs (57,60,69) and influences the RPR’s cleavage-site selection (178-180).

It has been shown that the kinetics and cleavage-site selection of pre-tRNAs by bacterial RNase P is mainly dependent on N+1 and the structure of the amino acid acceptor stem (178,180-182). Interestingly, results from several studies indicate that eukaryal RNase P may recognize its substrates differently from bacterial RNase P.

For example, a model substrate comprising a 12-bp stem (similar to the Acc-T-helical stack) capped with a terminal loop and terminating with a 3′-RCCA trailer and be cleaved by bacterial but not by eukaryal RNase P (183-185). However, if this minimal substrate contains even a 1-nt bulge at the linker region between the T and acceptor stems, it can be recognized and cleaved by eukaryal RNase P (183,184).

Furthermore, bacterial RNase P cleaves pre-tRNAHis to generate an 8-bp acceptor stem while eukaryal RNase P cleaves the same substrate to generate a 7-bp acceptor stem (180,181). Therefore, it is reasonable to hypothesize that the higher protein content in archaeal and eukaryal RNase P may alter the mode of specific

65 recognition of RPR for pre-tRNAs and/or provide functional groups as ligands for binding to substrates. In this study, we show that archaeal and eukaryal RNase P exhibit better fidelity than bacterial RNase P in cleavage-site selection especially towards non-consensus substrates, even when pre-tRNAs deviate from the consensus tRNA structure. We also provide a kinetic framework to discuss how archaeal/eukaryal RPPs can alter the cleavage-site selection by their cognate RPRs.

3.2 Materials and methods

3.2.1 Pyrococcus furiosus (Pfu) RNase P assay

Pfu RNase P assays were performed essentially as described elsewhere

(87,186) with the some modifications. All Pfu RNase P assays were carried out under single-turnover conditions with excess amount of enzyme and ~2 nM 5′-[P32]-labeled pre-tRNAGln. For the assays conducted with Pfu RPR alone and Pfu RPR + RPPs, the enzyme concentrations were 15 M and 200 nM, respectively.

3.2.2 Cloning and in vitro transcription of pre-tRNAGln-Mja RPR

pBT7-pre-tRNAGln-UAU-Mja RPR: Synechocystis pre-tRNAGln was conjugated with a linker to L15 of Mja RPR by overlap-extension PCR (5′-UAU-3′ indicates the 3- nt sequence linking pre-tRNAGln to Mja RPR). First, two fragments, one encoding the

Synechocystis pre-tRNAGln and the other encoding Mja RPR, were obtained separately by PCR using primer pairs pGln-S3-M GF + pGln-S3-M GR and pGln-S3-M

MF + pGln-S3-M MR, respectively (Table 3.2); the plasmids pT7Gln (187) and pBT7-

Tyr pt -S3-M RPR (88) served as the respective templates. These two PCR products, which have a 19-nt overlap, were annealed and extended. Finally, the extended product was digested with BamHI and ligated to pBT7 digested with StuI and BamHI.

66 The resulting plasmid was named pBT7-pre-tRNAGln-UAU-Mja RPR.

pBT7-pre-tRNAGln-AAU-Mja RPR and pBT7-pre-tRNAGln-GCCA-Mja RPR: We then generated two other cis constructs that differ in the linker sequence, pBT7-pre- tRNAGln-AAU-Mja RPR and pBT7-pre-tRNAGln-GCCA-Mja RPR (5′-AAU-3′ and 5′-

GCCA-3′ indicate the linker sequences in these self-cleaving constructs). The primer pair, ptGln-AAU-M F and ptGln-linker-M R (Table 3.2), was used to generate pBT7- pre-tRNAGln-AAU-Mja RPR; the primer pair, ptGln-GCCA-M F and ptGln-linker-M R

(Table 3.2), was used to generate pBT7-pre-tRNAGln-GCCA-Mja RPR. These primers flank the linker nucleotides to be changed and orient outward to ensure amplification of the entire pBT7-pre-tRNAGln-UAU-Mja RPR plasmid (except the linker being modified). The resulting PCR product was circularized by ligation with T4 DNA ligase and transformed into E. coli DH5α. ransformants were then screened to identify those harboring desired constructs.

In vitro transcription of cis constructs: All three self-cleaving RPRs were generated using the corresponding template DNA for T7 RNA polymerase-mediated run-off transcription (102). In all cases, the templates for transcription were generated by using a high-fidelity PCR [using Phusion DNA polymerase (New England Biolabs)] performed with 5′-TAATACGACTCACTATAGGTTAATCAATGGGGTGTAG-3′

(forward) and 5′-CTATTTCGGCTTGCACCCC-3′ (reverse) as primers. These primers generate PCR products containing the coding sequence of pre-tRNAGln linked to Mja

RPRs with different linkers under the control of a T7 RNA polymerase promoter

(whose sequence is underlined in the forward primer). Following in vitro transcription, all RNAs were subjected to extensive dialysis to remove unincorporated rNTPs and

67 their concentrations were determined from their extinction coefficients at Abs260.

3.2.3 Kinetic studies with pre-tRNAGln-Mja RNase P

The self-cleavage rates of pre-tRNAGln-Mja RPR were determined in the absence and presence of Mja RPPs. In vitro transcribed pre-tRNAGln-Mja RPR was first folded as follows: incubation at 50°C for 50 min in water followed by 37°C for 30 min in 50 mM Tris-acetate (pH 8), 800 mM NH4OAc and 10 mM Mg(OAc)2. For all cis cleavage reactions, we mixed 25,000 dpm of folded 5′-[P32]-labeled pre-tRNAGln-Mja

RPR with 50 nM of the same unlabeled, folded transcript. Assays were performed in two phases: a pre-incubation followed by a cleavage time-course. To minimize self- cleavage of the reactions involving POP5RPP30 during pre-incubations at 55°C, which was required for temperature equilibration and for reconstitution of the RPR with RPPs (see description below), the pre-incubation step was performed in the presence of 25 mM Ca2+ instead of 10 mM Mg2+. In all cases, once cleavage was initiated (as described below), aliquots were removed at defined time intervals and the reactions were terminated by adding an equal volume of stop dye (10 M urea, 5 mM EDTA, 0.05% (w/v) xylene cyanol, 0.05% (w/v) bromophenol blue, 20% (v/v) phenol). For short incubations, reactions were first terminated by immersing the reaction tubes in liquid nitrogen before adding stop dye.

RPR-alone: The folded RPR [50 nM in 20 μl of 50 mM 2-(N- morpholino)ethanesulfonic acid (MES) pH 6, 2.5 M NH4OAc, 10 mM Mg(OAc)2] was incubated at 55°C for 5 min. Cleavage at 55°C was then initiated by addition of an equal volume of identical buffer containing 1 M Mg(OAc)2, which had been pre- warmed to 55°C.

68 RPR + RPPs: The reconstitutions with different combinations of RPPs involved variations in divalent ions used since care had to be taken to minimize self- cleavage while maximizing RNP assembly and product formation. We empirically determined these optimal conditions, although they largely mirrored those in our earlier study when we examined self-cleavage of pre-tRNATyr-Mja RPR (88). For determining the self-cleavage rates of pre-tRNAGln-Mja RPR in the presence of RPPs, we first mixed the RPR with RPPs (final concentration of 50 and 500 nM, respectively) at 37°C for 10 min followed by 55°C for 10 min. With pre-tRNAGln-Mja RPR +

RPP21RPP29, the RNP was first reconstituted in 20 μl of 50 mM MES (pH 6), 800 mM NH4OAc and 10 mM Mg(OAc)2; at the end of this pre-incubation, cleavage was initiated by addition of 20 μl of 50 mM ME pH 6, 800 mM NH4OAc and 400 mM

Mg(OAc)2. In the case of RPR + POP5RPP30 or 4RPPs, the reconstitution was in 20

μl of 50 mM MES (pH 6), 800 mM NH4OAc and 25 mM Ca(OAc)2 and cleavage initiated by addition of 20 μl of 50 mM ME (pH 6), 800 mM NH4OAc and 200 mM

Mg(OAc)2.

3.2.4 Data analyses

The salient aspects of data analyses are described in Chapter 2.2.5. Some additional details pertinent to this study are provided here. To obtain the rates of correct product formation (kobs,c) under single-turnover conditions, the % correct

-kobs,c˙t product formed at time t (Pt,c) was fit to Pt,c = Ampc˙(1 – e ) using Kaleidagraph

software (Synergy) where Ampc is the amplitude of the correct cleavage defined based on experimentally observed values for optimal curve fits (Table 3.1 and Fig.

3.6). The same analysis was applied to find the rates and the amplitudes of mis- cleaved product formation (kobs,m and Ampm, respectively). The standard errors for the

69 best-fit values of kobs did not exceed 8%. The reported kinetic parameters are the mean and standard deviation values calculated from at least three independent experiments.

3.3 Results

This project was initiated by Dr. Deepali Singh and some of the results presented here are from the joint efforts of Dr. Deepali Singh, Dr. Lien Lai and myself.

In Figures 3.2 and 3.3, data corresponding to cleavage of pre-tRNAGln by the

RNase P holoenzymes from Escherichia coli (Eco) and Pyrococcus furiosus (Pfu) were generated by Dr. Deepali Singh, and the RNase P holoenzyme from

Arabidopsis thaliana (Ath) was purified and provided by Dr. Lien Lai. The results presented in sections 3.3.4 and 3.3.5 did not involve any collaborative efforts and represent solely my contributions.

3.3.1 The significance of N+1N+72 for pre-tRNA for cleavage-site selection decreases from bacterial > archaeal > eukaryal RNase P

The majority of bacterial pre-tRNAs possess a G+1C+72 pair and U-1 (188,189).

Gln One of the exceptions is pre-tRNA as it harbors U+1A+72 and A-1 (Fig. 3.1). Previous studies with bacterial RNase P have shown that changing the nucleotide identity at or near the cleavage site affects the cleavage-site selection (141,178,190,191). To understand the influence of the nucleotide identity of N+1N+72 in a pre-tRNA in cleavage-site selection by RNase P from all three domains of life, we first mapped the cleavage site of pre-tRNAGln by the RNase P holoenzymes from Escherichia coli (Eco),

Pyrococcus furiosus (Pfu), and Arabidopsis thaliana (Ath). Eco, Pfu, and Ath RNase P

Gln cleaved pre-tRNA at the correct cleavage site (between N+1 and N−1; designated C0)

70 70, 85, and 100%, respectively, and mis-cleaved at the incorrect site (between N+1 and N+2; designated M+1) 30, 15, and 0%, respectively (Figs. 3.1A, 3.2, and 3.3).

Most pre-tRNAs contain G+1, where the exocyclic amine of G+1 contributes to cleavage-site selection and catalysis (191). Therefore, we replaced U+1A+72 in pre-

Gln Gln tRNA with G+1C+72 to generate pre-tRNA U-G (Fig. 3.1) and then subjected pre- tRNAGlnU-G to RNase P-mediated cleavage under multiple-turnover conditions.

Gln Interestingly, replacing U+1A+72 with G+1C+72 in pre-tRNA resulted in a higher proportion of correct cleavage and a ~3-fold increase in cleavage rate for Eco and Pfu

RNase P (Figs. 3.2 and 3.3; data not shown). In striking contrast, replacing U+1A+72

Gln with G+1C+72 in pre-tRNA did not have any influence on the cleavage rate for Ath

RNase P-mediated catalysis (data not shown).

3.3.2 Stable tertiary structure of pre-tRNAGln is important for correct cleavage- site selection by bacterial RNase P but not the archaeal or eukaryal variants

We then examined other interactions, which may be used by RNase P for cleavage-site selection. One of the established interactions between bacterial

RNase P and pre-tRNAs is the interaction between the T-stem loop (TSL) region in the pre-tRNA and the cruciform region in the RPR consisting P7-P11, which is referred to as the TSL-binding site (TBS). The TSL-TBS interaction has been shown to affect cleavage-site selection (159). The characteristic structural architecture of the

D-/T-loop interaction in a pre-tRNA plays an important role in stabilizing the overall structure and presenting functional groups that allow a productive interaction with

RNase P (46,159,167). Previous studies have shown that in order to maintain D-/T- loop interaction in archaeal selenocysteine tRNA, which has an extended D-stem of 7

71 bp, the T-stem is reconfigured to contain only contains 4 bp, one bp less than most tRNAs (192). This finding suggests structural compensation between the D- and T- stems to maintain the D-/T-loop interaction. Therefore, we decided to disrupt the D-/T- loop interaction by extending the D stem by one to six bp (pre-tRNAGlnD+ variants) and by deleting the D stem-loop (pre-tRNAGlnD). We also examined the contribution of anticodon stem by removing the anticodon stem-loop altogether (pre-tRNAGlnAC)

(Fig. 3.1).

pre-tRNAGlnD+ variants, pre-tRNAGlnD and pre-tRNAGlnAC were cleaved by

Gln Eco RNase P at C0 only 20-30% compared to pre-tRNA which was cleaved 70% correctly (Figs.3.2 and 3.3). In marked contrast, Pfu and Ath RNase P cleaved pre- tRNAGlnD+ variants, pre-tRNAGlnD, and pre-tRNAGlnAc correctly ~80 and 100%, respectively, suggesting that bacterial RNase P relies on the native tertiary structure for correct cleavage-site selection in pre-tRNAGln while archaeal and eukaryal RNase

P might be more tolerant of structural aberrations.

3.3.3 The length of the T-stem and the acceptor stem of pre-tRNAGln affects cleavage-site selection

One of the proposed recognition mechanisms is the so called “ruler mechanism”, which postulates that RNase P recognizes the highly conserved tertiary- structural arrangement at the bottom of the Acc-T-helical stack, involving the

D−/T−loops. Through specific recognition of functional groups in the T-stem loop,

RNase P is believed to measure 12 bp to guide its positioning of metal ions for site- specific cleavage between the leader and the Acc-T-helical stack (167,168,193). The mis-cleavage observed during processing of pre-tRNAGlnD+ variants, pre-tRNAGlnD

72 and pre-tRNAGlnAC by bacterial RNase P might reflect the disruption of the ruler mechanism resulting from perturbations in the TSL-TBS interaction.

To test whether the ruler mechanism plays a role in cleavage-site selection in pre-tRNAGln, we extended the T- or the acceptor- stem by either one (pre-tRNAGlnT+1 and pre-tRNAGlnAcc+1; Fig. 3.1) or two bp (pre-tRNAGlnT+2 and pre-tRNAGlnAcc+2;

Fig. 3.1). Eco and Pfu RNase P cleaved pre-tRNAGlnT+1 or pre-tRNAGlnAcc+1 mostly between U+1 and G+2 (M+1) and generated a 12-bp Acc-T-helical stack. In striking

Gln Gln contrast, eukaryal RNase P cleaved pre-tRNA T+1 or pre-tRNA Acc+1 at C0 ~40% or 70%, respectively, and generated a 13-bp Acc-T-helical stack (Figs. 3.2 and 3.3).

These results suggest that bacterial and archaeal RNase P might rely on a ruler mechanism to cleave pre-tRNAGln at the correct site more so than eukaryal RNase P.

However, when we tested Eco, Pfu, or Ath RNase P with pre-tRNAGlnT+2 or pre-

Gln tRNA Acc+2, majority of the cleavage occurred predominantly at M+1, generating a

13-bp Acc-/T-helical stack (Figs. 3.2 and 3.3), perhaps reflecting that the ruler mechanism is not implemented rigidly.

3.3.4 Archaeal RPPs affect cleavage-site selection

One possible explanation for the differences in cleavage-site selection between bacterial and archaeal/eukaryal RNase P is that some functional attributes for cleavage-site selection may have been transferred from the RPR to RPPs during evolution. We used in vitro reconstituted RNase P from Pyrococcus furiosus (Pfu), an archaeon, to test whether archaeal RPPs really influence cleavage-site selection.

Interestingly, when we used the Pfu RPR alone to cleave pre-tRNAGln, the majority of cleavage occurred at M+1 (~80%) (Figs. 3.4 and 3.5), whereas Eco RPR alone

73 Gln cleaved pre-tRNA mostly at C0 (~85%) (data not shown). These results suggest that Pfu and Eco RPRs use distinct recognition modes during pre-tRNAGln cleavage.

When either Pfu RPP21RPP29 or POP5RPP30 was added, the cleavage for pre-

Gln tRNA by Pfu RPR at C0 increased from 20 to 60% (Fig.3.4, lanes 3 and 4 and Fig.

3.5). The presence of all four RPPs with Pfu RPR increases cleavage for pre-tRNAGln at C0 to ~90% (Fig. 3.4, lane 5 and 3.5) suggesting that the effect of RPP21RPP29 or POP5RPP30 are additive and possibly work by promoting recognition/docking of different sites on the pre-tRNA (see discussion).

3.3.5 Effect of Mja RPPs on cleavage-site selection and the rate of pre-tRNAGln-

Mja processing

To further understand the contribution of archaeal RPPs to the cognate RPR’s cleavage-site selection and the chemical cleavage of pre-tRNAGln, we constructed three self-cleaving RPRs, which differ only in the linker region. As there was precedence for Mja RPR supporting cis cleavage of an Eco pre-tRNATyr (88), we conjugated a pre-tRNAGln to Mja RPR. The covalent tethering of a pre-tRNA substrate to the RPR allows us to focus solely on the cleavage step (87,88,186).

We tested all three self-cleaving RPRs in cleavage-site selection experiments.

While pre-tRNAGln-AAU-Mja RPR and pre-tRNAGln-GCCA-Mja RPR behaved similarly,

Gln pre-tRNA -UAU-Mja RPR supported an additional mis-cleavage between A-1 and A-2.

We reasoned that the first uridine in UAU linker of pre-tRNAGln-UAU-Mja RPR could

Gln potentially base pair with A-1 in the leader sequence of pre-tRNA thus creating an aberrant mis-cleavage site. Therefore, we focused our subsequent efforts on pre- tRNAGln-AAU-Mja RPR (although the GCCA linker-bearing construct might have been

74 equally appropriate). Also, the pre-tRNAGln-AAU-Mja RPR will be referred to as pre- tRNAGln-Mja RPR for simplicity.

The amplitudes of correct and mis-cleavage are defined based on experimentally-observed values (Ampc and Ampm; Table 3.1). Fc refers to the fraction of total cleavage taking place at the correct site. The rate of correct and mis-cleaved product formation (kobs,c and kobs,m; Table 3.1 and Fig. 3.7) were calculated by analyzing the time course for both products (see method). pre-tRNAGln-Mja supported self-cleavage at C0 only ~20% of the total cleavage (Fc = ~20%; Fig. 3.4, lane 7 and

Fig. 3.6). Both POP5RPP30 and RPP21RPP29 increased Fc from ~20% to ~50%

(Fig. 3.4, lanes 8 and 9 and Fig. 3.6). In addition, POP5RPP30, but not

RPP21RPP29, increased the cleavage rate of pre-tRNAGln-Mja by 600-fold regardless of cleavage taking place at the correct or incorrect site (kobs,c from 0.005 to

-1 -1 ~3.1 min and kobs,m from 0.004 to 2.5 min , at pH 6 and 55C; Table 3.1).

2+ Furthermore, varying the Mg concentrations in the assay did not affect the Fc of pre- tRNAGln-Mja + POP5RPP30 despite the fact that the rates were lower when the Mg2+ concentration was reduced from 100 mM to 50 mM or 25 mM (data not shown).

We next determined the effect of all four RPPs together on the Fc and the kobs for both correct and mis-cleavage. Interestingly, addition of RPP21RPP29 to pre-

Gln tRNA -Mja RPR + POP5RPP30 resulted in a further 2.7-fold increase in kobs,c (from

3.1 to ~ 8 min-1) to a level even greater than that observed with just POP5RPP30 while kobs,m remains unchanged (Table 3.1), suggesting a synergistic effect of

RPP21RPP29 and POP5RPP30 on the rate of correct cleavage. Moreover, adding

Gln RPP21RPP29 to pre-tRNA -Mja RPR + POP5RPP30 further increases the Fc from 75 50% to 80% (Fig. 3.4, lane 10 and Fig 3.6) indicating an additive effect of

RPP21RPP29 and POP5RPP30 on the RPR’s cleavage-site selection.

3.4 Discussion

The plurality of substrates for RNase P illustrates nature’s ability to simplify complex tasks. How RNase P recognizes its many substrates, which differ in sequence and structure, remains of great interest to the field. In this study, we used pre-tRNAGln and its derivatives with mutations, insertions, or deletions in different structural elements to dissect the recognition determinants in a pre-tRNA that might be used by Eco, Pfu, or Ath RNase P (representatives of bacterial, archaeal, or eukaryal RNase P, respectively), and have gained valuable insights into the mechanism of cleavage-site selection.

3.4.1 Nucleotide identity at position +1 in pre-tRNAGln

It has been demonstrated that U-1 in a typical pre-tRNA substrate interacts with A248 of bacterial RPR [Fig. 3.1B; see page 65; (47)]. Therefore, the presence of

Gln U+1 and G+2 in pre-tRNA might cause a local rearrangement at the cleavage site with U+1 interacting with A248 in the bacterial RPR thus shifting the site of cleavage by one nucleotide 3′ to the canonical cleavage site [Fig. 3.1; (141)]. This effect is accentuated for bacterial RNase P when the native tertiary structure of pre-tRNAGln is perturbed (e.g. pre-tRNAGlnD+, pre-tRNAGlnD, and pre-tRNAGlnAC) (Figs. 3.2 and

3.3) suggesting that the tertiary structure of pre-tRNAGln is an important determinant for cleavage at C0 by bacterial RNase P. In contrast, cleavage-site selection by archaeal or eukaryal RNase P was only slightly or not at all affected by the presence

Gln of U+1 and G+2 in pre-tRNA or by alterations to the native tertiary structure.

76 Interestingly, two studies have separately documented the differential effects of deleting the D-stem in a pre-tRNA on the cleavage-site selection by bacterial and human RNase P. Deleting the D-stem in pre-tRNAs caused bacterial RNase P to mis- cleave but had no effect on cleavage-site selection by human RNase P (183,194)

Collectively, these results highlight the higher fidelity and tolerance of structural variations of protein-rich archaeal/eukaryal RNase P compared to their bacterial cousin in processing pre-tRNAs with a non-native structure (Figs. 3.2 and 3.3).

3.4.2 D-stem, T-stem, and anticodon stem in pre-tRNAs

We show that pre-tRNAGlnD and pre-tRNAGlnAC variants can be cleaved by

RNase P across three domains of life suggesting that the D- and anticodon- stems are dispensable for RNase P-pre-tRNA interaction (Figs. 3.2 and 3.3). In contrast, pre-tRNAGlnT, in which the T-stem is deleted, could not be recognized and cleaved by any RNase P highlighting the critical role of T stem-loop in RNase P-pre-tRNA recognition in all three domains of life (Figs. 3.2 and 3.3). RNase P-mediated cleavage at the 5′ end of pre-tRNAs has been suggested to precede 3′ maturation and intron splicing (195). Since RNase P has to recognize all pre-tRNAs, the major recognition motif should be conserved in all pre-tRNAs. Therefore, our results perhaps reflect the fact that the position of an intron, if present, is usually located in the anticodon loop of pre-tRNAs (160), occasionally in the D- and variable loops, and only very rarely in the T-loop (196).

77 3.4.3 Archaeal RPPs influence cleavage-site selection and the rate for processing pre-tRNAGln

The structure of bacterial RPRs can be divided into two independently folding modules (37,44,45): (i) specificity (S) domain which contains conserved nucleotides recognizing the T stem-loop (TSL) in the pre-tRNA (46), and a catalytic (C) domain that cleaves the pre-tRNA leader while binding the 5′-leader (47), the acceptor stem

(48) and the 3′-RCCA (R represents purine) (49-51). Archaeal RPRs can also be demarcated into S and C domains (87,88,197). However, despite the similarity of the secondary structure of archaeal and bacterial RPRs, archaeal RPR cleaves pre-

Gln tRNA primarily at M+1 whereas bacterial RPR mostly cleaves at C0 (Fig. 3.4, lanes 2 and 7; not shown).

Archaeal POP5RPP30 and RPP21RPP29 can independently shift the cleavage-site from M+1 to C0, and their effect on cleavage-site selection is additive

(Table 3.1 and Figs. 3.5 and 3.6). We have shown that archaeal POP5RPP30 footprints on the RPR’s C domain (87,142) and archaeal RPP21RPP29 footprints on the RPR’s S domain (142). Therefore, the additive effect on cleavage-site selection perhaps reflects the interaction of RPP binary complexes with different parts of the pre-tRNA. A previous study examining the ratios of correct:aberrant cleavages of model substrates with an intact T-loop or a GAAA tetraloop by Pfu RPR indicates that the archaeal RPR’s S domain could not recognize the TSL region in these substrates

(158). However, binding of RPP21RPP29 to the archaeal RPR’s domain renders the RPR capable of recognizing the TSL in these model substrates. Therefore, it is conceivable that RPP21RPP29 affects cleavage-site selection by modulating the

78 TSL region of pre-tRNAs while POP5RPP30 interacts with the regions near the cleavage site in the pre-tRNA.

Here we provide a kinetic framework to rationalize how archaeal

POP5RPP30 and RPP21RPP29 might influence the RPR’s cleavage-site selection and cleavage rate (Fig. 3.7). We postulate that there is a rapid substrate-docking step

(E-S ⇄ ESC0 and E-S ⇄ ESM+1, where E-S refers to the cis conjugate) preceding the conformational change from ES to ES*, which helps to position the pre-tRNA and catalytic metal ions optimally for cleavage. These fast substrate-docking steps dictate

the amplitudes of the correct cleaved product (PC0) and mis-cleaved product (PM+1).

Gln Therefore, the Fc value of ~20% observed during self-cleavage of pre-tRNA -Mja

could be explained by a model in which KM+1 (the equilibrium constant of E-S ⇄ ESM+1)

is 4-fold greater than KC0 (the equilibrium constant of E-S ⇄ ESC0). The presence of

either RPP pair could reduce KM+1 and/or increase KC0, thus shifting the equilibrium

favorably toward ESC0 formation resulting in a higher proportion of correct cleavage.

When both RPP pairs are present, their effects on increasing KC0 and decreasing KM+1 are additive.

We previously showed that POP5RPP30 is the sole RPP pair responsible for increasing the RPR’s cleavage rate [Chapter 2; (88,186)]. Consistent with previous findings, POP5RPP30, but not RPP21RPP29, can increase the rate of cleavage presumably by promoting ES* formation in the conformational change step for both

correct cleavage and mis-cleavage (ESC0 ⇄ ESC* 0 and ESM+1 ⇄ ESM* +1). If the conformational change step is faster than the substrate docking step, the amplitudes of correct cleavage and mis-cleavage would be directly proportional to their

79 corresponding rates. However, this was not the case. Even though we observed equal rates of correct cleavage and mis-cleavage for RPR, RPR + RPP21RPP29, and RPR + POP5RPP30, the amplitudes of correctly cleaved and mis-cleaved products were not equal. In contrast, if the substrate-docking step is faster than conformational change step, the amplitude of correct cleavage and mis-cleavage could be uncoupled from the rates, a scenario consistent with our data.

The most intriguing findings come from the addition of RPP21RPP29 to pre- tRNAGln-Mja + POP5RPP30. Although RPP21RPP29 alone did not influence the

ES ⇄ ES* equilibrium (Fig. 3.7), adding RPP21RPP29 to pre-tRNAGln-Mja +

POP5RPP30 resulted in a 2.7-fold increase in the rate of correct cleavage to 8 min-1 while the rate for mis-cleavage still remains the same (Table 3.1 and Fig. 3.7). How does RPP21RPP29 increase the cleavage rate despite binding to the RPR’s S domain? It might do so by helping to position the substrate optimally for catalysis to occur at the correct cleavage site, and this effect manifests only when POP5RPP30 is present. Collectively, we show that the presence of both RPP pairs not only favors

formation of ESC0 but also increases the rate of the ESC* 0 formation.

Finally, our studies revealed that archaeal RPPs can increase the RPR’s cleavage efficiency towards an atypical pre-tRNA substrate to that observed with a typical pre-tRNA substrate. The rate of processing pre-tRNAGln by Mja RPR alone is

40-fold slower than that for pre-tRNATyr, a substrate with consensus elements for optimal recognition by RNase P (Table 3.1). However, when all RPPs are present, the rates of processing pre-tRNAGln and pre-tRNATyr only differ by 2.7-fold (Table 3.1).

These findings mirror the observation that bacterial RPP normalizes the cognate

80 RPR’s rate of cleavage for non-consensus pre-tRNA substrates by altering its energetic contributions to substrate binding and thereby enhancing the rate of RPR- mediated cleavage (62). Our results now highlight the roles of archaeal RPPs in ensuring that processing of different pre-tRNA substrates by the archaeal RNase P holoenzyme occurs at similar rates and with high fidelity.

3.4.4 Protein-rich RNase P confers more flexibility

Why did RNase P evolve into a protein-rich RNP in higher eukaryotes and what are the roles of its protein cofactors in aiding RNA catalysis? In this study, we observed an increase in the fidelity of cleavage with a concomitant increase in the protein content of the RNase P holoenzyme. We also demonstrated that archaeal

RPPs affect the RPR’s cleavage-site selection. Perhaps one of the reasons that archaeal/eukaryal RNase P is associated with more protein subunits is to deal with a wider array of substrates. In striking contrast to bacterial RNase P, eukaryal RNase P exhibits better tolerance of nucleotide identities at position -1 and +1 as well as structural aberrations in substrates and displays higher fidelity in cleavage-site selection. Indeed, by examining nucleotides at N-1 and N+1 in pre-tRNAs using the genomic tRNA database (http://lowelab.ucsc.edu/GtRNAdb) (189), we observed that the nucleotide identity is less biased in eukaryotes than in bacteria: U-1 together with

G+1 is present 43% of all pre-tRNAs in bacteria compared to 28% in eukaryotes; U-1 is

54% in bacteria compared to 40% in eukaryotes. As the transcriptomes became more complex in higher eukaryotes, lessening the constraints on the nucleotide identities and the structural architecture of substrates would have enabled RNase P to participate in the biogenesis of pre-tRNA variants and even other non-coding RNAs.

For example, metastasis-associated lung adenocarcinoma transcript 1 (MALAT1), a

81 long non-coding RNA known to be upregulated in many human cancers, is processed by RNase P (161). Another long non-coding RNA, Men , involved in the formation of paraspeckles, is also processed by RNase P (198). Both these ncRNAs contain a tRNA-like structure at their 3 end. Although these structures have no variable loop and a poorly conserved anticodon stem-loop, RNase P recognizes the tRNA-like motifs and cleaves precisely between A-1 and G+1.

82 A. Assayed under -1 -1 optimal conditions for kobs,c, min * Ampc,% kobs,m, min * Ampm,% each catalytic entity Gln pre-tRNA -Mja RPR 0.005 ± 0.001 15 ± 1 0.004 ± 0.0007 75 ± 6 + RPP21RPP29 0.005 ± 0.0001 38 ± 1 0.004 ± 0.0001 46 ± 3 + POP5RPP30 3.1 ± 0.11 41 ± 1 2.5 ± 0.14 56 ± 1 + Both binary RPP 7.96 ± 0.28 61 ± 4 2.75 ± 0.83 14 ± 1 complexes

B. Assayed under C. Assayed under Relative optimal conditions for k , min-1** optimal conditions for obs activity*** each catalytic entity** each catalytic entity Tyr pre-tRNA -Mja RPR 0.2 ± 0.04 RPR 40 + RPP21RPP29 0.24 ± 0.04 + RPP21RPP29 48 + POP5RPP30 20.5 ± 0.32 + POP5RPP30 6.6 + Both binary RPP 21.7 ± 0.16 + Both binary RPP 2.7 complexes complexes

Table 3.1. Effect of Mja RPPs on the rate of cleavage and cleavage-site selection of pre-tRNAGln-Mja at 55C at pH6 * The standard errors of the curve fits shown in Figure 3.6 are indicated in the

estimates of kobs and amplitudes (Amp). ** The data for pre-tRNATyrMja RPR experiments are reproduced from Table 1 in ref (88). In this earlier publication (88), the rates reported for a self-cleaving pre- tRNATyrMja RPR were at pH 5.4 and not pH 5.1 as was mistakenly reported. To facilitate comparison of the pre-tRNAGlnMja RPR and pre-tRNATyrMja RPR cleavage experiments, the rates observed at pH 5.4 with pre-tRNATyrMja RPR were multiplied by 4 to obtain rates that would have been observed at pH 6 should they have been measurable. We demonstrated previously a linear relationship

between log kobs and pH (88). Tyr *** The relative activity is obtained by dividing the kobs of pre-tRNA -Mja by the rate Gln of correct cleavage (kobs,c) of pre-tRNA -Mja.

83 pGln-S3-M MF 5′-GAATCCTAGCACCCCATATCGGCGCTTAGCC-3′ pGln-S3-M MR 5′-GGGGATCCGTCTCG CTATTTCGGCTTGCACCC-3′ pGln-S3-M GF 5′-TTAATCAATGGGGTGTAGCC-3′ pGln-S3-M GR 5′-GGCTAAGCGCCGATATGGGGTGCTAGGATTC-3′ ptGln-AAU-M F 5′-AATCGGCGCTTAGCC-3′ ptGln-GCCA-M F 5′-GCCACGGCGCTTAGCC-3′ ptGln-linker-M R 5′-TGGGGTGCTAGGATTC-3′

Table 3.2. Oligonucleotide primers used to construct pre-tRNAGln-Mja RPR

84

Figure 3.1. (A) Schematic depicting pre-tRNAGln and its mutant derivatives used in this study. (B) Possible interactions between nucleotides in typical (consensus) or atypical (non-consensus) pre-tRNAs and the bacterial RPR at the cleavage site.

85

Figure 3.2. Mapping the cleavage sites of pre-tRNAGln and its mutant derivatives by RNase P from (A) Escherichia coli (Eco), (B) Pyrococcus furiosus (Pfu), (C) Arabidopsis thaliana (Ath). The top panels depict the cleavage of pre-tRNAGln and its mutant derivatives by different RNase P holoenzymes. The bottom panels magnify the region in the sequencing gels to better depict the 5′-leaders generated from correct and mis-cleavage. Gln indicates wildtype pre-tRNAGln, while the labels on individual mutant derivatives are self-explanatory (see Fig. 3.1 for additional details). For example, U-G represents pre-tRNAGlnU-G. M indicates the ladder obtained from Gln alkaline hydrolysis of pre-tRNA . M+1 refers to the 5′-leader generated from mis- Gln cleavage between positions +1 and +2 in pre-tRNA while C0 refers to the 5′-leader from cleavage at the correct site between positions -1 and +1 in pre-tRNAGln. 86

Figure 3.3. Comparison of the fraction of correct and mis-cleaved products generated by processing of pre-tRNAGln by RNase P from Escherichia coli (Eco), Pyrococcus furiosus (Pfu), Arabidopsis thaliana (Ath). Black and white shading represent the percentage of correct and mis-cleaved products. ND denotes not determined.

87

Figure 3.4. Archaeal RPPs affect the RPR’s cleavage-site selection. T1 represents the G ladder generated by subjecting pre-tRNAGln to digestion with RNase Gln T1. Alk indicates the ladder obtained from alkaline hydrolysis of pre-tRNA . M+1 refers to the 5′-leader generated from mis-cleavage between positions +1 and +2 in Gln pre-tRNA while C0 refers to the 5′-leader from cleavage at the correct site between positions -1 and +1 in pre-tRNAGln. Uncleaved pre-tRNAGln is shown in lane 1. Cleavage of pre-tRNAGln by Pfu RPR, RPR + RPP21RPP29, RPR + POP5RPP30 and RPR + 4 RPPs is shown in lanes 2-5, respectively. Lanes 6 and 7 represent uncleaved and self-cleaved pre-tRNAGln-Mja RPR, respectively. The cleaved 5′-leader products generated by addition of Mja RPP21RPP29, POP5RPP30 and 4 RPPs to pre-tRNAGln-Mja are shown in lanes 8-10.

88

Figure 3.5. Pfu RPPs affect cleavage-site selection by the cognate RPR. Quantitation of the cleavage taking place at the correct and mis-cleavage site in pretRNAGln by different enzyme entities. Black and white shading represent the percentage of correct and mis-cleaved products. Both Pfu RPP pairs increase the fraction cleaved at the correct site. The mean and standard deviation values were calculated from three independent experiments. A representative gel is shown in Figure 3.4, lanes 1-5.

89

Figure 3.6. Effect of Mja RPPs on cleavage-site selection and the rate of self- Gln cleavage of pre-tRNA -Mja. Plots show cleavage at the correct (C0) site (circles), the mis-cleavage site one nucleotide 3′ to the correct site (M+1) site (squares), and the fraction of total cleavage taking place at the correct site (Fc; triangels). By analyzing the time-course for formation of correct and mis-cleaved products, the kobs values for the reactions catalyzed by the RPR with and without RPPs were determined for both correct and mis-cleavage (kobs,c and kobs,m; see Table 3.1). The mean and standard deviation values were calculated from three independent experiments.

90

Figure 3.7. Kinetic framework to interpret the contribution of the two binary RPP complexes to cleavage-site selection and the rate of self-cleavage of pre- tRNAGln-Mja. RPR-S refers to the pre-tRNAGln-Mja cis construct. E denotes different enzyme entities (Mja RPR alone or in the presence of different combinations of RPPs). Although both archaeal RPP pairs increase the percentage of total cleavage taking place at the correct site, we are unable to comment on their specific roles in influencing KC0 and KM+1 (i.e., archaeal RPP pairs could promote correct cleavage by

either increasing KC0 or decreasing KM+1 or both). POP5RPP30 increases the reaction rate for both correct and mis-cleavage presumably by influencing Kconf (88,186).

91

CHAPTER 4

COOPERATIVE RNP ASSEMBLY: COMPLEMENTARY RESCUE OF

STRUCTURAL DEFECTS BY PROTEIN AND RNA SUBUNITS OF

ARCHAEAL RNASE P

4.1 Introduction

Sequence comparison of RNase P RNAs (RPRs) from all three domains of life has led to the identification of their conserved sequences and secondary structures

[Fig. 4.1; (77-79)]. Archaeal RPRs have been categorized into two distinct structural types: A and M (77). Type A, exemplified by Methanothermobacter thermautotrophicus (Mth), is considered to be more ancestral due to the fact archaeal type A RPRs are very similar to bacterial type A RPRs. They are distinct from each other primarily because archaeal type A RNase P RNAs lack P18 and P13/14 [Fig.4.1;

(77)]. In addition, archaeal type A RPRs have been shown to be active on their own like bacterial type A RPRs (12,87). On the other hand, archaeal type M RPRs display activity only in the presence of RPPs [Fig.4.1; (12,77)]. Type M RPRs are exemplified by Methanocaldococcus jannaschii (Mja), in addition to the loss of P18 and P13/14, differ from archaeal type A RPRs due to (i) lack of P8, which is known in bacteria to be involved in pre-tRNA recognition (80,81) , and (ii) everything distal to P15 including

L15, which is known to be base paired with 3-CCA of tRNA (49,83,84). The loss of

92 P15/P16/P17/L17 also eliminates formation of P6 since one strand of this pseudoknot is on L17 in bacterial RPRs [Fig.4.1; (77)].

Despite these structural differences, all RPRs share universally conserved sequences and structures. For instance, a conserved feature in all RPRs is a bulge- helix structure in the P4 paired region (Fig. 4.1). The geometry of this P4 bulge-helix structure in bacterial RPRs is important for Mg2+ coordination, which in turn is critical for pre-tRNA binding and cleavage (199). Disrupting this bulge-helix structure either by eliminating the bulged uridine (ΔU) or inserting even one additional uridine in the bulge (+U) decreases activity. For example, deleting this bulged uridine (U69) in

Escherichia coli (Eco) RPR results in a 100-fold lower single-turnover reaction rate compared to the wild-type (WT) RPR even at saturating (300 mM) concentrations of

Mg2+ (199). Eliminating the bulge in the Eco RPR weakened the apparent affinity for

Mg2+ and reduced the cooperativity for Mg2+ (the Hill coefficient, nH, decreased from

2.2 to 1.5) (199). Replacement of non-bridging oxygen atoms with sulfur at positions proximal to U69 in Eco RPR (e.g., A67) lowered the cleavage rate by three to four orders of magnitude, with the deleterious effect in some instances largely rescued by thiophilic metal ions like Mn2+ (143,200); similar results were reported with Bacillus subtilis (Bsu) RPR (201). Such findings provide evidence for the coordinated binding of catalytically important Mg2+ ions by the bulged uridine and the neighboring phosphate backbone in the P4 helix-bulge structure. Moving the bulge away from the nearby sites of metal ion coordination also decreased activity by 70-fold indicating that the position of the bulge in P4 is important (199).

93 To elucidate similarities in the RNA-mediated catalytic mechanism in bacterial and archaeal RNase P, we investigated if a universally conserved, bulged uridine shown to participate in binding catalytically important Mg2+ ions in bacterial RPRs is also vital in the archaeal relative. Indeed, we found severe catalytic defects in archaeal RPRs in which this bulged uridine was mutated; however, the weakened binding of active-site Mg2+ in these mutant archaeal RPRs is rescued (but not completely) by their RPPs.

In addition to the findings that archaeal RPPs can rescue the mutations in

RPR, we have also obtained the evidence that archaeal RPR can rescue the mutations in RPPs. In separate structural studies of archaeal RPP29 conducted in the laboratory of Dr. Mark Foster (OSU), an N-terminal deletion mutant of RPP29

(RPP29Δ24) which fails to bind its partner RPP21 [as judged by isothermal titration calorimetry (ITC) and NMR spectroscopy] was identified. Despite its inability to form a functional heterodimer with RPP21, in vitro biochemical studies demonstrated that

RPP29Δ24 is functional upon addition of RPR reflecting the RPR’s ability to restore the interactions between RPP29Δ24 and RPP21. These findings collectively illustrate the reciprocal subunit interactions vital for driving archaeal RNase P towards its functional structure.

4.2 Materials and methods

4.2.1 Cloning and expression of the genes encoding the RNA and protein subunits of Mth and Mja RNase P

Details are provided in ref (186).

94 4.2.2 Construction of mutant derivatives of archaeal RPRs

The genes encoding the different single-nucleotide deletion or insertion archaeal RPR mutants were generated using two different PCR-based mutagenesis approaches. The first was used to construct MthΔU, Mth+U, MjaΔU and Mja+U RPRs, while the second was used to make pre-tRNATyr-MjaΔU RPR. he sequences of primers used in these different PCRs are listed in Table 4.1.

The first approach entailed a QuikChange (Stratagene)-like mutagenesis using two primers complementary to each other. The Mth RPR mutant derivatives were made using pUC19-Mth RPR (102) as the template. To make pUC19-MthΔU

RPR, Mth RPRΔU-F and Mth RPRΔU-R were used as the primers; for pUC19-Mth+U

RPR, the primer pair was Mth RPR+U-F and Mth RPR+U-R. The Mja RPR mutant derivatives were made using pBT7-Mja RPR (88) as the template. Either Mja

RPRΔU-F and Mja RPRΔU-R or Mja RPR+U-F and Mja RPR+U-R were used as the primer pair to make pBT7-MjaΔU RPR or pB 7-Mja+U RPR, respectively.

Tyr Tyr To generate pBT7-pre-tRNA -MjaΔU RPR, pB 7-pt -S3-M RPR (88) was utilized as the template. T4 polynucleotide kinase was used to phosphorylate the primers pTyr-S3-Mja RPR ΔU-F and pTyr-S3-Mja RPR-R; these primers flank the nucleotide to be deleted and are oriented outward to ensure amplification of the entire

Tyr pBT7-pt -S3-M RPR plasmid (except the single position to be deleted). The resulting

PCR product was circularized by ligation with T4 DNA ligase and transformed into E. coli DH5α. ransformants were then screened to identify those harboring pBT7-pre- tRNATyr-MjaΔU.

95 4.2.3 In vitro transcription of RPRs used in this study

Mth RPR WT and mutant derivatives were generated using the corresponding

EcoRI-linearized pUC19 plasmids as the template DNA for T7 RNA polymerase- mediated run-off transcription (102). In the case of Mja RPR WT and mutant derivatives, the corresponding pBT7 plasmids were linearized with BsmAI (88). For the cis-cleaving Mja ΔU RPR, we generated the RNA using the appropriate PCR product as the template for transcription. A high-fidelity PCR [using Phusion DNA polymerase (New England Biolabs)] was performed with pBT7-pre-tRNATyr-Mja ΔU

RPR as the template, and 5′-TAATACGACTCACTATAGGGAGCAGGCCAGTAAA-3′

(forward) and 5′-CTATTTCGGCTTGCACCCC-3′ (reverse) as primers. hese primers generate a PCR product containing the coding sequence of pre-tRNATyr-MjaΔU RPR under the control of a T7 RNA polymerase promoter (whose sequence is shown in underline in the forward primer). Following in vitro transcription, all RNAs were subjected to extensive dialysis to remove unincorporated rNTPs and their concentrations determined from their extinction coefficients at Abs260.

4.2.4 RNase P assays

All assays were performed in a thermal cycler. While the various reactions described below were set up differently, they were always terminated using a stop dye [10 M urea, 1 mM EDTA, 0.05% (w/v) xylene cyanol, 0.05% (w/v) bromophenol blue, 20% (v/v) phenol]. The reaction products were then subjected to 8% (w/v) polyacrylamide/7 M urea gel electrophoresis and cleavage rates analyzed as described in the next section.

96 Multiple-turnover reactions

While multiple-turnover trans-cleavage reactions with Pfu RNase P were performed essentially as described elsewhere (87,186), those with Mth and Mja

RNase P were carried out as follows. Mja or Mth RPRs (re-suspended in water) were first folded by incubating for 50 min at 50°C, 10 min at 37 °C, and then for 30 min at

37°C after addition of an equal volume of 100 mM Tris-HCl (pH 8),1.6 M NH4OAc and

20 mM MgCl2. Typically, these folded RPRs were kept at stock concentrations of 4

μM. When holoenzymes were tested, they were first reconstituted by pre-incubating the folded RPR with cognate RPPs (final concentration of 10 and 100 nM, respectively) in assay buffer [50 mM Tris-HCl (pH 7.1), 800 mM NH4OAc, and the concentrations of MgCl2 indicated in Fig. 4.2] for 10 min at 37°C followed by 10 min at

55°C. Subsequently, the pre-tRNA processing assays at 55°C were initiated by adding to the reconstituted holoenzymes 500 nM E. coli pre-tRNATyr, a trace amount of which was internally labeled with [α-P32]-GTP (202). Aliquots were removed at defined time intervals and quenched with an equal volume of stop dye. [Note: For assays with the Mth RPR (i.e., in the absence of RPPs), we used 50 mM Tris-HCl (pH

7.1), 2 M NH4OAc and the concentrations of MgCl2 indicated in Fig. 4.2].

Single-turnover reactions

Details regarding RNA folding and RPR-mediated cis cleavage reactions are provided in section 3.2.3. For the RNP experiments, the reconstitutions with different combinations of RPPs involved variations in pH and divalent ions used since care had to be taken to minimize self-cleavage while maximizing RNP assembly and product formation. Although the procedures are similar to those with pre-tRNAGln-Mja in

Chapter 3, we empirically determined these optimal conditions. For determining the

97 self-cleavage rates of pre-tRNATyr-MjaΔU RPR in the presence of RPPs, we first mixed the RPR with RPPs (final concentration of 50 and 500 nM, respectively) at

37°C for 10 min followed by 55°C for 10 min. With pre-tRNATyr-MjaΔU RPR +

RPP21RPP29, the RNP was first reconstituted in 20 μl of 50 mM MES (pH 6), 800 mM NH4OAc and 10 mM Mg(OAc)2; at the end of this pre-incubation, cleavage was initiated by addition of 20 μl of 50 mM ME pH 6, 800 mM NH4OAc and 200 mM

Mg(OAc)2. In the case of RPR + POP5RPP30, the reconstitution was in 20 μl of 50 mM MES (pH 6), 800 mM NH4OAc and 25 mM Ca(OAc)2, and cleavage initiated by addition of 20 μl of 50 mM ME (pH 6), 800 mM NH4OAc and 200 mM Mg(OAc)2.

When all four RPPs were present, the reconstitution was in 20 μl of 50 mM ME (pH

5.4), 800 mM NH4OAc and 1 mM Mg(OAc)2, and cleavage initiated by addition of 20

μl of 50 mM MES (pH 5.4), 800 mM NH4OAc and 200 mM Mg(OAc)2.

4.2.5 RNase P activity data analysis

Regardless of trans or cis cleavage reactions, the reaction products were separated by denaturing PAGE and visualized by phosphorimager analysis (Typhoon,

GE Healthcare). The resulting bands were quantitated with ImageQuant (GE

Healthcare) to assess the extent of substrate cleaved. For the trans cleavage reactions, we calculated the initial velocities by determining the product formed during time course; typically, the times of incubation were chosen to restrict the amount of substrate cleaved to < 30%. To obtain the rate of product formation (kobs) in cis

-kt reactions, the percent of product formed at time t (Pt) was fit to Pt = P(1 - e ) using

Kaleidagraph software (Synergy). The standard errors for the best-fit values of kobs did not exceed 20%. The reported kinetic parameters are the mean and standard deviation values calculated from at least three independent experiments.

98 4.3 Results

4.3.1 Mutations in the P4 helix of archaeal RPRs decrease activity

We tested the importance of this conserved bulge-helix structure in P4 of archaeal RPRs by either increasing the size of the bulge form one to two uridines (+U) or deleting the bulge (ΔU) in the P4 helix of Mth and Mja RPRs. We chose these two

RPRs as they represent the currently accepted two broad classes of euryarchaeal

RNase P: types A (Mth) and M (Mja) (77), and because their corresponding holoenzymes have been reconstituted (88,186). Although the P4 helix bulge is present in both type A and M RPRs, other notable differences in their structural elements (Fig. 4.1) have important functional consequences: the type A RPR displays pre-tRNA processing activity in the absence of RPPs in contrast to the type M RPR

(12,88). Hence the RPR-alone studies described below pertain only to the Mth RPR.

We determined the initial velocities of WT and mutant Mth RPRs without the cognate RPPs under multiple-turnover conditions (1 μM RPR and 5 μM Eco pre- tRNATyr) and as a function of Mg2+ concentration. Our data show that the reaction rates for MthΔU and Mth+U RPRs were 6- and 8-fold lower, respectively, than Mth

(WT) RPR regardless of the Mg2+ concentration used (Figs. 4.2A and 4.2D).

4.3.2 Archaeal RPPs can rescue mutations in the P4 helix of cognate RPRs

We tested the activity of the P4-mutated archaeal RPRs in the presence of their cognate RPPs since there is considerable precedent for the bacterial RPP influencing substrate recognition, cleavage and metal ion coordination (57-

59,63,68,157,164,201,203). For example, the absence or presence of the RPP caused differences in the ability of thiophilic metal ions to rescue the adverse effects

99 caused by replacing different non-bridging oxygens with sulfur in the P4 helix of Bsu

RPR (201). This observation taken together with subsequent studies (58,63) supported the idea that the bacterial RPP increases the affinity of the RPR for catalytically relevant metal ions. Also, the aberrant cleavage of select pre-tRNAs by the EcoΔC92 RPR mutant was completely rescued by the Eco RPP, suggesting that the protein cofactor can alleviate RPR catalytic defects including altered substrate positioning (203).

After measuring the initial velocities of the Mth and Mja RNase P holoenzymes under multiple-turnover conditions (10 nM enzyme and 500 nM Eco pre-tRNATyr) at different concentrations of Mg2+, we determined that both ΔU and +U RPRs in type A and M holoenzymes resulted in reduced activities at 2.5 and 5 mM Mg2+ (Figs. 4.2B and 4.2C). For example, cleavage rates for the MthΔU, Mth+U, MjaΔU, and Mja+U were 4-, 9-, 2-, and 14-fold lower at 5 mM Mg2+ concentration, respectively, compared to their respective WT counterpart (Figs. 4.2B and 4.2C). However, between 30 to 50 mM Mg2+, the activity of the mutant RNase P holoenzymes is indistinguishable from the WT suggesting that the defect caused by P4 mutations manifests only at low Mg2+ concentrations (Figs. 4.2E and 4.2F).

4.3.3 RPP21•RPP29 and POP5•RPP30 both rescue the decrease in cleavage rate caused by the ΔU mutation

The inability of archaeal RPRs with P4 mutations to coordinate catalytically important Mg2+ could affect substrate binding, chemical cleavage and/or product release. We first sought to determine if the difference in activity between the WT and mutant Mth/Mja RNase P holoenzymes is attributable to a defect in substrate binding.

100 Our previous steady-state kinetic measurements revealed that the KM for processing pre-tRNATyr by a type A (Pfu) RNase P holoenzyme is ~200 nM (87); even if a P4 mutation (e.g., ΔU) increased the M by 20-fold, we reasoned that such a large

Tyr change in KM could be overcome by using [pre-tRNA ] > 4,000 nM. Therefore, we tested the activity of W and ΔU Mth RNase P holoenzymes using 7 μM substrate at

10 mM Mg2+; however, this increased concentration of substrate did not alleviate the effects of the ΔU mutation in the Mth RPR (data not shown), suggesting that altered

Mg2+ coordination (due to the ΔU mutation) likely affected cleavage or product release.

o focus solely on the chemical cleavage step, we introduced the ΔU mutation

Tyr Tyr in a cis construct, pre-tRNA -S3-Mja RPR (hereafter referred to as pre-tRNA -Mja

RPR for simplicity). In this self-cleavage construct, the pre-tRNATyr substrate is tethered via a 3-nt spacer to Mja RPR to help overcome the type M RPR’s substrate- binding defects (12,77,88), which typically prevent pre-tRNA cleavage in trans. We previously showed this cis conjugate to be a good model for studying the chemical cleavage step [based on a slope of ~1 in plots of log(kobs) vs. pH; (88)]. We also demonstrated that addition of Mja POP5RPP30 enhanced kobs for this self-cleavage by ~100-fold while RPP21RPP29 had no effect; however, both binary RPP complexes reduced the monovalent and divalent ionic requirement. Moreover, these cis construct findings were mirrored in single-turnover, trans-cleavage studies with a type A (Mth) RNase P (186).

We constructed pre-tRNATyr-MjaΔU RPR by deleting the bulged uridine in the

RPR and measured the rates of cleavage [Table 4.2 and Fig. 4.3; (88)]. The kobs for self-cleavage of pre-tRNATyr-MjaΔU RPR is 0.0025 min-1 (at pH 6), 80-fold lower than

101 WT pre-tRNATyr-Mja RPR. When RPP21RPP29 was added, the rate increased by

~7-fold. Notably, the rate increased by ~600-fold in the presence of POP5RPP30.

When all four proteins were present, the rate is ~750-fold higher compared to the

RPR-alone reaction. However, unlike the multiple-turnover reaction (Fig. 4.2C), even the presence of all four RPPs was unable to raise the mutant RPR’s activity to that observed with the WT regardless of the Mg2+ concentration used (Table 4.2; not shown). The activity of the mutant holoenzyme is still 12-fold weaker than the WT

(Table 4.2).

4.4 Discussion

4.4.1 Evolutionarily and functionally conserved motifs in both bacterial and archaeal RPRs

Despite the remarkable differences in the subunit composition of bacterial, archaeal and eukaryal RNase P, there is a striking conservation of various RPR features, notably the P4 bulge-helix motif (Fig. 4.1). Altering the geometry of this bulge-helix structure in bacterial RPRs affects substrate binding, catalysis and metal ion interactions (143,199-201). We have now demonstrated that mutagenesis of this universally conserved bulge-helix P4 motif in archaeal RPRs (types A and M) results in loss of function even at 500 mM Mg2+. With archaeal ΔU RPRs, we found an 8-fold decrease in turnover number under steady-state conditions (Fig. 4.2) and a larger 80- fold decrease in cleavage rate under single-turnover conditions (Fig. 4.3; Table 4.2).

These findings mirror the 100-fold decrease in cleavage rate reported for the bacterial

(Eco) ΔU RPR and the complete loss of activity upon deletion of three nucleotides in the P4 helix of human RPR (13,199). Cumulatively, these data indicate a common role for the P4 helix-bulge structure (in all RPRs) in binding metal ions required for

102 catalysis. A recent crystal structure of the bacterial RNase P holoenzyme-tRNA (ES) complex provides additional support for this idea (204). Some attributes of the RNase

P catalytic site environment could be inferred when a pre-tRNA 5′-leader was soaked into a crystal of the Thermotoga maritima RNase P-pretRNAPhe complex, with and without Sm3+ ions which were used as Mg2+ mimics. Notably, a metal ion, likely to generate the attacking hydroxide nucleophile, is held in position by interactions with the O4 oxygen of the universally conserved bulged uridine and its proximal phosphate backbone. Regardless of the interplay between RPRs and RPPs that might be essential to formulate a similar active-site architecture in RNase P variants from all three domains of life, it seems likely that the central role of the conserved bulged uridine in coordinating a catalytically relevant metal ion might be preserved.

4.4.2 Archaeal RPPs can mitigate catalytic defects in the cognate RPR

In contrast to the RPR-alone reactions under multiple-turnover conditions, archaeal ΔU and +U RPRs (types A and M) in the presence of RPPs do not exhibit any catalytic defects at 30 mM Mg2+, although the RPP(s)-mediated rescue is incomplete at 2.5, 5 and 10 mM Mg2+ (Fig. 4.2). This observation reflects the ability of archaeal RPPs to fully restore the catalytic activity of the cognate ΔU/+U RPRs even though they do not promote WT-like metal ion-binding affinities. In fact, another study where the P4 region in a different type A (Pyrococcus horikoshii, Pho) RPR was mutated drew similar conclusions (205); since these Pho RPR mutants were not assayed alone, rescue by RPPs was not documented.

To decipher both the nature of the catalytic defect engendered by the ΔU mutation and the magnitude of the rescue by RPPs, we tested a cis-cleavage

103 construct (i.e., pre-tRNATyr-MjaΔU RPR) with and without RPPs. By comparing these kinetic data with those reported earlier for the WT RPR, we determined that the ΔU mutation caused a 80-fold decrease in the cleavage rate. When this ΔU cis-cleaving construct was assembled with RPP21RPP29 or POP5RPP30 or all four RPPs, the differences in self-cleavage rates were narrowed to an order of magnitude of those observed with the WT RPR with the corresponding RPPs (Table 4.2). These results proved instructive for several reasons. First, since the RPPs did completely alleviate the catalytic defects of the ΔU mutant RPR under multiple-turnover conditions (Fig.

4.2), their failure to do so under conditions where the rate was limited by chemical cleavage (Table 4.2) suggests that the rate-limiting step in the catalytic mechanism of archaeal RNase P must be product release at pH 7, highlighting another important parallel with bacterial and eukaryal RNase P (57,59,147). Second, these results helped uncover a functional redundancy between RPP21RPP29 and the P4 helix, an inference that merits elaboration.

Our previous kinetic studies of trans cleavage of a pre-tRNA by Mth (type A)

RNase P and cis cleavage of a pre-tRNA in the self-processing pre-tRNATyr-Mja (type

M) RPR revealed that POP5RPP30 is solely responsible for increasing the rate of chemical cleavage while RPP21RPP29 plays an essential role in substrate binding

(88,186). Therefore, it is surprising that Mja RPP21RPP29 which does not increase the self-cleavage of pre-tRNATyr-Mja RPR does so by 7-fold for the ΔU counterpart. In the latter instance, RPP21RPP29 must somehow help position the substrate optimally for catalysis to occur, and that this effect (likely mediated by Mg2+ binding) manifests only when the conserved uridine in P4 is absent.

104 Bacterial RPRs can be divided into two independently-folded domains: a specificity (S) domain with conserved nucleotides making up a T-stem-loop binding site (TBS) that recognizes the T stem-loop (TSL) of the pre-tRNA, and a catalytic (C) domain that cleaves the pre-tRNA while binding to the 5'-leader, acceptor stem and the 3′-RCCA sequence (30,45,47,49,51,55,83,159,193). Results from various studies indicate that a similar demarcation is likely in archaeal RPRs (87,88,186,205).

Moreover, footprinting studies indicate that POP5RPP30 binds to the C domain and

RPP21RPP29 binds to the S domain (87,142); such a delineation is consistent with the notion that binding of POP5RPP30 in the vicinity of the active site contributes directly to catalysis, and binding of RPP21RPP29 to the S domain contributes to substrate binding presumably by facilitating the TSL (pre-tRNA)−TBS (RPR) interaction (159,193). Given that P4 is in the C domain (Fig. 4.1) and RPP21RPP29 footprints on the S domain, this binary complex may affect Mg2+ coordination in the active site by enabling long-distance interactions between the S and C domains.

Integrating two earlier observations provides support for this notion. First, in bacterial

RPRs, the TSL-S domain interaction is believed to trigger a conformational change that aids catalysis by positioning the chemical groups and catalytically important Mg2+ near the cleavage site in the C domain (159,193). Second, cleavage-site selection in model substrates revealed that when archaeal RPRs are bound to RPP21RPP29, their substrate-recognition properties coincide with those of bacterial RPRs (158). The inter-domain crosstalk mediated by RPP21RPP29 and its direct relevance to catalytic metal ion interactions remain to be deciphered.

105 4.4.3 Archaeal RPR can rescue the deleterious effects of a large deletion in a cognate RPP

In addition to archaeal RPPs rescuing an RPR mutation, there is also an example of archaeal RPR rescuing an RPP mutation. Joy Xu, a former graduate student in the laboratory of Dr. Mark Foster (OSU), demonstrated that the removal of the first 24 residues in Pfu RPP29 (RPP29Δ24) largely abolishes its ability to bind

RPP21. Interestingly, our laboratory demonstrated that the holoenzyme assembled in vitro using RPP29Δ24 retains ~60% of the activity of wild-type Pfu RNase P (data not shown). This near-normal functional behavior, together with the previous finding that neither RPP21 nor RPP29 can individually activate the RPR, likely reflects the ability of the RPR to restore the interactions between RPP29Δ24 and RPP21.

These observations of mutual rescue between archaeal RPR and RPPs reveal an intimate, inter-dependent relationship among the subunits in RNase P.

These results are consistent with hierarchy and redundancy in induced-fit mechanisms, a recurring theme in large dynamic RNPs (206,207).

106 MthdUF 5′-AGGGGCTGAGGAAACCCACCCATCATACAG-3′ MthdUR 5′-CTGTATGATGGGTGGGTTTCCTCAGCCCCT-3′ MthUUF 5′-AGGGGCTGAGGAAACTTCCACCCATCATACAG-3′ MthUUR 5′-CTGTATGATGGGTGGAAGTTTCCTCAGCCCCT-3′ MjadUF 5′-AAGAGGGGAGGAAGTCCGCCCACCCCATTT-3′ MjadUR 5′-AAATGGGGTGGGCGGACTTCCTCCCCTCTT-3′ MjaUUF 5′-AAGAGGGGAGGAAGTTTCCGCCCACCCCATT-3′ MjaUUR 5′-AATGGGGTGGGCGGAAACTTCCTCCCCTCTT-3′ pTyr-S3-M-dU-F 5′-TCCGCCCACCCCATTTAT-3′ pTyr-S3-M-U-R 5′-CTTCCTCCCCTCTTAAAG-3′

Table 4.1. Oligonucleotide primers used to construct mutant archaeal RPRs

107 + Assayed under [NH4 ]; 2+ kobs(WT), Rel. -1 Rel. kobs(WT)/ optimal conditions for [Mg ], -1 k , min ** min * k obs(MT) k k each catalytic entity M obs(WT) obs(MT) obs(MT) pre-tRNATyr-Mja RPR (WT or U MT) 2.5; 0.5 0.2  0.04 1 0.0025  0.0002 1 80 + RPP21RPP29 0.8; 0.1 0.2  0.04 1 0.017  0.0014 1 14 + POP5RPP30 0.8; 0.1 20  0.32 100 1.46  0.12 584 14 + Both binary RPP 0.8; 0.1 22  0.16 105 1.84  0.18* 736 12 complexes

Table 4.2. Effect of Mja RPPs on the ionic requirements and rate of cleavage of pre-tRNATyr-Mja and pre-tRNATyr-MjaU at 55C * The rates at pH 6 were extrapolated from those determined at pH 5.4 and reported by us previously. We had also earlier established the direct relationship

between log (kobs) versus pH. ** All experiments were performed at pH 6, except for pre-tRNATyr-MjaU RPR reconstituted with both binary RPP complexes, which was assayed at pH 5.4. The rate at pH 6 for the reaction with four RPPs was obtained by extrapolation from 5.4. The standard errors of the curve fits shown in Figure 4.3 are indicated

in the estimates of kobs.

108

Figure 4.1. Secondary structures of RPRs from Bacteria (e.g., Escherichia coli) and Archaea [e.g., Pyrococcus furiosus (Pfu) or Methanothermobacter thermautotrophicus (Mth; type A), and Methanocaldococcus jannaschii (Mja; type M)] (28,31). The universally conserved bulged uridine in the P4 helix of all RPRs is enclosed in a colored circle. Deletion (ΔU) or insertion (+U) mutations made at this position in the Mth and Mja RPRs are also indicated.

109

Figure 4.2. Comparison of the Mg2+ dependence of wildtype (WT) and mutant Mth and Mja RNase P. Multiple-turnover reaction rates for the WT and mutant Mth RPR (A) and the Mth, Mja RNase P holoenzymes (B, C) are plotted as a function of the Mg2+ concentration in the assay. To better illustrate the activity changes as a 2+ function of Mg concentration, we have also plotted krel values, i.e., the reaction rates for the mutant RPRs (D) and holoenzymes (E, F) relative to the WT. The mean and standard deviation values were calculated from three independent experiments.

110

Figure 4.3. Effects of Mja RPPs on the rate of pre-tRNATyr-MjaΔU RPR self- cleavage. By analyzing the time course for product formation, the kobs values for the reactions catalyzed by the RPR with and without RPPs were determined (see Table 4.2). The mean and standard deviation values were calculated from three independent experiments.

111

CHAPTER 5

FUTURE DIRECTIONS

5.1 Investigating the roles of L7Ae in archaeal RNase P catalysis

Archaeal L7Ae, a ribosomal protein, is involved not only in translation (as part of ribosomes) but also in RNA modification (as part of H/ACA and C/D snoRNPs).

Our laboratory only recently demonstrated that L7Ae is also an archaeal RPP (208).

L7Ae was shown to co-elute with Methanococcus maripaludis (Mma) RNase P activity (208). Furthermore, addition of L7Ae to Mma RPR + POP5RPP30 +

RPP21RPP29 increases the optimal temperature for activity by 12C and kcat/KM for pre-tRNA cleavage by ~360-fold, mimicking parameters observed with partially purified native Mma RNase P (208). Together, these data support the idea that L7Ae plays an important role in archaeal RNase P catalysis.

In this dissertation, we have dissected the roles of four known RPPs in chemical cleavage, substrate binding, cleavage-site selection, and Mg2+ coordination.

To gain further insight into the roles of L7Ae in these aspects, it is worth assessing the effect of Mja L7Ae on different self-cleaving constructs (pre-tRNATyr-Mja, pre- tRNATyr-MjaU or pre-tRNAGln-Mja) with or without the two binary Mja RPP complexes

(Chapter 3 and 4).

112 5.2 In vivo mapping of the 5′ end of tRNAs

We have shown that bacterial and archaeal RNase P mis-cleave

Synechocystis pre-tRNAGln in vitro ~30% and 15%, respectively, and generate a mature tRNAGln shorter by one nucleotide at the 5′ end (Chapter 3). These observations raise an important question: does RNase P mis-cleave its substrates in vivo? If so, is there any repair mechanism to correct this mistake?

Since Eco RNase P displayed the most prominent mis-cleavage, and it has been shown that in vitro reconstituted Eco RNase P mis-cleaves Eco pre-tRNAGln

(141), it would be worth mapping the 5′ end of Eco tRNAs by using primer extension analysis of RNA extracted from Eco to examine that whether mis-cleavage indeed happens in vivo. However, it seems unlikely that a cell would tolerate such a high frequency of mis-cleavage (e.g., 30%). If we do not observe the same amount of mis- cleavage in vivo, it is possible that there is a repair enzyme to correct the mis- cleavage. One of the possible candidates for repairing such mis-cleavage is a 3′ to 5′ nucleotidyltransferase.

A recently report suggests that a Thg1p-like-protein (TLP), an ortholog of eukaryal Thg1p (tRNAHis guanylyltransferase), in bacteria and archaea may play such a role to add a single nucleotide at the 5′ end of mis-cleaved tRNAs (209). Eukaryal

His Thg1p is responsible for the G-1 addition to the 5′ end of cytoplasmic tRNA whereas

His in bacteria and archaea, G-1 is genomically encoded and retained in mature tRNA after the RNase P cleavage. Therefore it eliminates the need for the G-1 addition to the 5′ end of tRNAHis. Furthermore, unlike eukaryal Thg1p which only adds guanosine, bacterial and archaeal TLP have been shown that their ability to add a nucleotide is

113 highly template-dependent. This characteristic makes TLP well suited for repairing mis-cleavage by bacterial and archaeal RNase P. In fact, the laboratory of Dr. Jane

Jackman (Department of Biochemistry, The Ohio State University) has already obtained preliminary data showing that bacterial TLP can indeed add U+1 to the 5′ end of tRNAGln mis-cleaved by Eco RNase P.

5.3 Which RPPs contribute to increased affinity for metal ions in active site?

We show that POP5RPP30 and RPP21RPP29 can coordinate catalytically relevant Mg2+ ions (Chapter 4). However, Mg2+ plays two roles in RNase P-mediated catalysis: (i) it can help RPR folding (structural Mg2+), and (ii) it is involved in generating nucleophile in the active site of RNase P (catalytic Mg2+; Fig. 5.1).

Therefore, the experiments in chapter 4, albeit supportive of a role for RPPs in coordinating catalytically relevant Mg2+, do not reveal which class of Mg2+ ions are affected by RPPs.

To focus on only the catalytic Mg2+, we will use a method which utilizes phosphorothioate (PS) substitutions. The pro-Rp oxygen of the scissile phosphate in a pre-tRNA is replaced with sulfur, thus decreasing the affinity for Mg2+ (150,151,210).

The presence of Rp-PS in pre-tRNA reduced the rate of processing by bacterial RPR at least 1000-fold under conditions where chemical cleavage is rate-limiting (150), and this effect could be largely rescued by addition of thiophilic ions such as Cd2+ or

Mn2+. By analyzing the rescue of cleavage for this modified substrate as a function of

[Cd2+] with either partially or complete assembled archaeal RNase P holoenzyme, we should be able to dissect which RPP is responsible for coordinating to catalytic Mg2+.

114 5.3.1 Introducing the PS substitute in pre-tRNATyr-Mja

Unlike bacterial RPR, archaeal RPRs have a severe substrate-binding defect thus rendering difficult the direct analysis of solely the cleavage step (14,88). To overcome this roadblock, we introduced a PS-modification to the Rp or Sp location of the scissile phosphate of pre-tRNATyr-Mja, a self-cleaving construct. In the rescue experiments, a low concentration of Mg2+ will be added to assist RPR folding. To

2+ evaluate the specific effect on cleavage induced by Cd , krel will be calculated

Cd Mg Cd Mg Cd according to this formula krel = (ks /ks )/(ko /ko ) (211), where ks indicates the kobs

2+ Mg obtained by using PS-modified substrate supplemented with Cd ; ko indicates the

2+ kobs obtained by using a non-modified substrate with Mg . We will then plot krel as a function of [Cd2+] to calculate the apparent affinity for catalytic metal ions and the Hill coefficient to determine the cooperative dependence (if any) on metal ions.

We obtained an RNA oligo (RNA 1; Fig. 5.2) containing either Rp- or Sp-PS from Sigma. HPLC will be employed to separate these two diastereomers. RNA 1 (a

13-nt RNA containing the 5’-leader sequence and part of tRNAGln) will then be ligated with RNA 2 (a 10-nt RNA consisting of one strand of the D-stem and flanking regions) and RNA 3 (a RNA molecule including the rest of tRNAGln and Mja RPR). To accomplish this goal, we will employ splinted ligation, where a 40−nt complementary

DNA oligo spanning two ligation sites is designed to hybridize and hold together all three RNAs so that T4 DNA ligase can ligate these three RNAs.

As proof of principle, even without separating the Rp and Sp isomers, we tested the ligation method. Indeed, we have successfully ligated all three RNAs. The preliminary experiments involve subjecting these ligated RNAs to different types and

115 concentrations of thiophilic ions. Once we determine the optimal conditions for thiophilic-ion rescue, we will proceed to determine the effect of archaeal RPPs on coordinating catalytic metal ions. Such a strategy should help identify the RPP pair that contributes to coordination of active site metal ions.

116

Figure 5.1. The role of Mg2+ ions in RNase P-mediated catalysis. General two- metal ion mechanism proposed for RNase P-mediated catalysis. One metal ion is responsible for deprotonating a water molecule to generate the nucleophile, which attacks the scissile bond in the reactive phosphate; the other metal ion is involved in stabilizing the 3 leaving oxyanion group. Figure reproduced from ref (150).

117

Figure 5.2. pre-tRNATyr-Mja RPR carrying a phosphorothioate modification at the RNase P cleavage site. The sites of ligation are marked by gray ovals. The arrow indicates the canonical RNase P cleavage site and the position of phosphorothioate modifications. Individual RNA fragments are colored red, blue and black.

118

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