Fluor-Labeling of RNA and Fluorescence-Based Studies of Precursor-Trna Cleavage by Escherichia Coli Ribonuclease P

Fluor-labeling of RNA and Fluorescence-based Studies of Precursor-tRNA Cleavage by Escherichia coli Ribonuclease P

THESIS

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in the Graduate School of The Ohio State University

Andrew Wallace

Graduate Program in Biochemistry

The Ohio State University

2013

Master's Examination Committee:

Venkat Gopalan, Advisor

Edward J. Behrman

Andrew Wallace

2013

Abstract

RNase P participates in tRNA biogenesis by removing the 5′-leader of precursor- tRNAs (pre-tRNAs). A possible relic of the RNA world, the ribonucleoprotein form of

RNase P performs pre-tRNA processing using an RNA catalyst, associated with a different number of essential protein cofactors in the three domains of life. In addition to being a paradigm for studies of protein-aided RNA catalysis, there is some interest in exploiting structural differences between the bacterial and eukaryotic variants to develop new antibacterial drugs. Facile high-throughput assays would aid in the identification of such agents. Since its discovery four decades ago, RNase P activity has typically been assessed by monitoring the cleavage of internally or terminally 32P- labeled pre-tRNAs. Following denaturing polyacrylamide gel electrophoresis to resolve the substrate and products, the extent of cleavage is quantitated using a phosphorimager. Fluor-labeled pre-tRNAs provide an attractive non-radioisotopic alternative while retaining high sensitivity and offering the potential for high-throughput assays. With some modifications to the copper(I)-mediated alkyne-azide cycloaddition, we have generated at low cost 5′-fluor-labeled pre-tRNAs. An in vitro transcription primed with 5′-azido-5′-deoxyguanosine (AzG) yielded a 5′-azide-bearing pre-tRNA, which was then coupled to either a homemade alkyne-bearing carboxyfluorescein derivative or commercially available alkyne derivatives of Cy3 (Cyanine 3, where 3 refers ii to the number of carbon atoms in the polyethylene region) and Alexa Fluor® 647. I have assessed the extent of AzG incorporation at the 5′-terminus and found it to be 87 ± 2% in a model RNA tetramer, with the remainder accounted for by GTP, and I have found the extent of fluor labeling to be 65% in a pre-tRNA substrate. My experiments revealed that several such economically prepared fluor-labeled pre-tRNAs are cleaved by in vitro reconstituted Escherichia coli RNase under multiple-turnover conditions accurately and with efficiency similar to that of radio-labeled pre-tRNAs. Moreover, the fluor-labeled pre-tRNA and products could also be separated and quantitated by laser-induced fluorescence capillary electrophoresis (LIF-CE) in an automated DNA sequencer (ADS).

LIF-CE with an ADS not only allows the assay to be high-throughput, but also enables facile detection of small amounts of mis-cleavage products when non-consensus pre- tRNAs are processed by RNase P. An advantage with fluor-labeled pre-tRNAs, not afforded by radiolabeled counterparts, is the ability to assay in the same reaction two or more substrates of the same length but different composition, an important consideration for substrate-recognition studies. This new approach will provide an experimental handle to better understand the rates at which RNase P processes various pre-tRNAs in vivo, an aspect that remains to be fully investigated.

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Dedication

This document is dedicated to my family and to Elizabeth. I am ever grateful for their

love and support.

Acknowledgments

I thank both my advisor Dr. Venkat Gopalan and Dr. Edward J. Behrman for their support and mentorship, and for their herculean patience with the tribulations of my passage. I also thank the members of the Gopalan laboratory, in particular Dr. Lien B. Lai, Dr.

Gireesha Mohannath, Stella Lai, and Tien-Hao Chen for their cameradie as well as their constant willingness to discuss experiments and teach me new things. For innumerable discussions on all things science and politics, I also thank Sathiya Manivannan.

Further, I thank Michael Zianni (OSU PMGF) for his tutelage and engagement – without his assistance, the LIF-CE studies would not have happened. I also thank Dr.

Carolina Barillas-Mury (Chief, Mosquito Immunity and Vector Competence Section,

Laboratory of Malaria and Vector Research, National Institutes of Health, Rockville, MD) for generously providing the plasmid for our catalase purification. I am also most grateful for the suggestion by Professor Craig Martin (Department of Chemistry,

University of Massachusetts, Amherst) that led to our successful measurement of AzG incorporation in a model RNA.

Vita

September 24, 1988 ...... Born – Norwalk, Connecticut

2012 ...... B.S. Chemistry, The Ohio State University

2012 to present ...... Graduate Teaching/Research Associate,

Department of Chemistry and Biochemistry,

The Ohio State University

Fields of Study

Major Field: Biochemistry

Table of Contents Abstract ...... ii

Dedication ...... iv

Acknowledgments...... v

Vita ...... vi

Fields of Study ...... vi

Table of Contents ...... vii

List of Tables ...... x

List of Figures ...... xi

Abbreviations ...... xiii

Chapter 1: Introduction ...... 1

1.1 RNase P ...... 1

1.2 Using Fluorescence to Study pre-tRNA Binding and Processing by RNase P...... 5

1.3 Bioconjugation and Bioorthogonality ...... 8

1.4 Click chemistry offers versatile tools for bioconjugation ...... 10

1.5 Application of CuAAC to RNA derivatization ...... 13

Chapter 2: Results ...... 21 vii

2.1 Synthesis of 5′-azido-5′-deoxyguanosine via 5′-iodo-5′-deoxyguanosine ...... 21

2.2 Incorporation of AzG in RNA transcripts ...... 21

2.3 Synthesis of 5(6)-propargylamidofluorescein ...... 23

2.4 CuAAC-mediated 5′-terminal labeling of RNA ...... 24

2.5 Periodate oxidation and the CuAAC are orthogonal, enabling one-pot dual-5′- and-

3′-terminal labeling of RNA ...... 25

2.6 5′-labeled precursor tRNA substrates can be used to measure RNase P activity ... 27

2.7 Labeling different substrates with different fluorophores allows simultaneous

quantitation of RNase P processing of multiple substrates ...... 29

2.8 A capillary electrophoresis-based automated DNA sequencer is a convenient

format for assessing RNase P cleavage assays ...... 31

Chapter 3: Discussion ...... 53

3.1 Incorporation of AzG at the 5-terminus of RNAs transcribed in vitro ...... 53

3.2 CuAAC-mediated 5-fluor labeling of RNA ...... 56

3.3 Periodate oxidation and the CuAAC are orthogonal, enabling one-pot dual-5′- and-

3′-terminal labeling of RNA ...... 59

3.4 A capillary electrophoresis-based automated DNA sequencer is a convenient

format for assessing RNase P cleavage assays ...... 60

3.5 Concluding remarks ...... 62 viii

Chapter 4: Summary ...... 66

Chapter 5: Materials and Methods ...... 68

5.1 Synthesis of 5′-azido-5′-deoxyguanosine via 5′-iodo-5′deoxyguanosine ...... 68

5.2 Preparation of fluor-labeled and biotinylated pre-tRNAs ...... 70

5.3 Synthesis of 5(6)-propargylamidofluorescein ...... 72

5.4 Preparation of M1 RNA ...... 73

5.5 Preparation of C5 protein (procedure not performed by this author) ...... 73

5.6 Eco RNase P activity assays ...... 73

5.7 Assay separation and quantitation by laser-induced fluorescence-based automated

DNA sequencer ...... 74

5.8 – Assessment of AzG incorporation ...... 75

References ...... 82

List of Tables

Table 1: Eco RNase P processing of pre-tRNATyr substrates either bearing the 3′-terminal-

ACCA sequence (+CCA) or lacking it (-CCA), and labeled with either Cy3 or FAMA...... 48

List of Figures

Figure 1: Processing of a precursor-tRNA by RNase P...... 16

Figure 2: Crystal structure of the bacterial RNase P holoenzyme from Thermotoga maritima in complex with a mature tRNA...... 17

Figure 3: Copper(I)-catalyzed azide-alkyne cycloaddition...... 18

Figure 4: The mode of hydroxyl-radical generation by copper(I)...... 19

Figure 5: Hydroxyl radical-initiated strand-scission...... 20

Figure 6: Synthesis of 5′-azido-5′-deoxyguanosine via 5′-iodo-5′-deoxyguanosine ...... 36

Figure 7: In vitro transcription initiated with AzG...... 37

Figure 8: Agarose gel depicting in vitro transcriptions of 5′-N3-RNAs...... 38

Figure 9: Assessment of AzG incorporation relative to GTP at the first position of tetramer 5′-GAAC-3′...... 39

Figure 10: Synthesis of 5(6)-propargylamidofluorescein from 5(6)-carboxyfluorescein. 40

Figure 11: 5′-modification of RNA using the CuAAC...... 41

Figure 12: pre-tRNAPheTyr 5′-labeled with FAMA...... 42

Figure 13: Ultraviolet and visible spectrum of pre-tRNAPheTyr-CCA 5′-labeled with Alexa

Fluor® 647 (AF 647)...... 43

Figure 14: Streptavidin-mediated gel-shift demonstrating fluor-labeling and biotinylation of the pre-tRNA in a one-pot reaction...... 44

Figure 15: Cleavage of 5′-32P- (A) and 5′-FAMA- (B) pre-tRNATyr processing by Eco RNase

P in vitro...... 45

Figure 16: Cleavage of 5′-P32- (left) and 5′-FAMA- (right) pre-tRNATyr processing by Eco

RNase P in vitro...... 46

Figure 17: Eco RNase P processing of 5′-FAMA-pre-tRNATyr+CCA and 5′-Cy3-pre-tRNATyr-CCA

(which lacks the 3′-terminal-CCA) assessed in the same reaction...... 47

Figure 18: Eco RNase P processing of both 5′-AF647 and 5′-FAMA-pre-tRNAPheTyr+CCA in the same reaction...... 49

Figure 19: LIF-CE assessment of pre-tRNATyr cleavage by Eco RNase P...... 50

Figure 20: Mis-cleavage of non-canonical pre-tRNAs by Eco RNase P...... 51

Figure 21: PAGE-band assessment of pre-tRNAGln cleavage by Eco RNase P compared to the LIF-CE experiment...... 52

Figure 22: Nujol mull FT-IR spectrum of 5′-iodo-5′-deoxyguanosine...... 76

Figure 23: 1H NMR spectrum of 5′-iodo-5′-deoxyguanosine...... 77

Figure 24: Nujol mull FT-IR spectrum of 5′-azido-5′deoxyguanosine...... 78

Figure 25: 1H NMR spectrum of 5′-azido-5′-deoxyguanosine...... 79

Figure 26: Nujol mull FT-IR spectrum of 5(6)-propargylamidofluorescein...... 80

Figure 27: 1H NMR spectrum of 5(6)-propargylamidofluorescein...... 81

xii

Abbreviations

ADS – automated DNA sequencer

AF647 – Alexa Fluor® 647 alkyne

ATP – adenosine-5′-triphosphate

AzG – 5′-azido-5′-deoxyguanosine

CE – capillary electrophoresis

CTP – cytidine-5′-triphosphate

CuAAC – copper(I)-catalyzed alkyne-azide cycloaddition

Cy3 - Cyanine 3

Cys – cysteine

DMSO – dimethyl sulfoxide

DNA – deoxyribonucleic acid

Eco – Escherichia coli

EDTA – ethylene diamine tetra acetate

FAMA – 5(6)-propargylamidofluorescein

FRET - Förster resonance energy transfer

FT-IR – Fourier transform-infrared

GTP – guanosine-5′-triphosphate k-turn – kink-turn

LIF-CE - laser-induced fluorescence capillary electrophoresis

xiii mRNA – messenger RNA

NMR – nuclear magnetic resonance

NTP – nucleoside-5′-triphosphate

PA – polyacrylamide

PAGE – polyacrylamide gel electrophoresis pk-turn – “p” refers to “RNase P” pre-tRNA – precursor transfer RNA

PRORP – proteinaceous RNase P

RNA – ribonucleic acid

RNase P – ribonuclease P

RNP – ribonucleoprotein

ROS – reactive oxygen species

SAM – S-adenosyl-methionine

Tma – Thermotoga maritima tRNA – transfer RNA

UTP – uridine-5′-triphosphate

xiv

Chapter 1: Introduction

1.1 RNase P

Ribonuclease P (RNase P) is an essential and nearly ubiquitous endoribonuclease activity which cleaves the 5′-leader from a nascent precursor-transfer RNA (pre-tRNA), an essential step in tRNA biogenesis (Lai et al. 2010 and references therein) (Figure 1).

RNase P function is performed by a ribonucleoprotein (RNP) complex in bacteria and archaea. In these domains, the holoenzyme consists of a single catalytic RNA subunit along with varying numbers of protein cofactors (Lai et al. 2010, McClain et al. 2010 and references therein) (Figure 2). The finding that the catalytic component of the bacterial

RNase P holoenzyme is the RNA subunit resulted in the first report of a ribozyme, and this “ribozyme” turned out to be one of a small number of true multiple-turnover RNA enzymes that process substrates in trans (Guerrier-Takada et al. 1983, Willkomm et al.

2007 and references therein). Bacterial RNase P is associated with a single protein cofactor, while archaeal RNase P has up to five (Lai et al. 2010 and references therein,

Cho et al. 2010, Fukuhara et al. 2006). Despite the catalytic competence of the RNA subunit in vitro, all protein subunits in the RNP RNase P are required for organismal viability in bacteria and eukaryotes (Reich et al. 1988, Chamberlain et al. 1998).

In eukaryotes, RNase P exists in at least two forms: while many organisms have been shown to possess a canonical RNA-powered RNP performing RNase P function in the nucleus, a number of organisms have been shown to possess an alternate form of

1 the enzyme in their organelles. In the nucleus of a number of well-studied eukaryotes such as Homo sapiens and Saccharomyces cerevisiae, RNase P is a RNP consisting of a single catalytic RNA and up to 10 protein subunits (Lai et al. 2010, McClain et al. 2010 and references therein, Chamberlain et al. 1998, Xiao et al. 2002). However, RNase P activity is attributed to a protein-only variant of the enzyme in the mitochondria and chloroplasts of many eukaryotes, and this variant appears to have arisen separately from the RNP form (Lai et al. 2010 and references therein, Walker et al. 2008, Gobert et al. 2010, Howard et al. 2012). It has been suggested that in organisms such as

Arabidopsis thaliana and Trypanosoma brucei, proteinaceous RNase P may have entirely replaced the RNP variant (Taschner et al. 2012, Gutmann et al. 2012). The existence of proteinaceous RNase P provides a unique and fascinating opportunity to study the same enzyme activity catalyzed by protein or RNA and gain insights into the possible evolution from RNA to protein-based catalysis.

The discovery of RNase P and its catalytic RNA led to a paradigm shift in our understanding of the cellular roles of RNA. We now appreciate that in addition to being genetic material and a carrier of information, RNAs play catalytic, structural, and regulatory roles. Moreover, the catalytic potential of RNA offers insight into a possible primordial RNA world, wherein RNA performed roles now overtaken by DNA and protein.

Like its role in RNase P, a catalytic RNA is believed to drive the peptidyl-transfer reaction in the ribosome and the phosphoryl-transfer reactions in the spliceosome.

Study of catalysis in the latter enzymes, however, is complicated by their unwieldy nature arising from tens of subunits. RNase P offers a more tractable system to study protein-aided RNA catalysis. By deriving general principles from studies that unravel

RNase P structure-function relationships, one can in principle extrapolate to more complex RNPs such as the ribosome and the spliceosome. Such a claim is exemplified by the finding of a common structural motif present in RNase P, the ribosome, and T- box leaders of messenger RNA (mRNA), which appears to bind the “elbow” region of in a tRNA (Lehmann et al. 2013). This motif was first characterized in RNase P, and was then shown to exist in the other two classes of RNA. In another example, a novel type of kink turn (k-turn) motif – termed a pk-turn (where “p” refers to RNase P) was identified in

RNase P by examining its crystal structure (Meyer et al. 2012, Reiter et al. 2010). This motif has been shown to adopt a geometry identical to that of the canonical k-turn, and its identification is expected facilitate the development of sequence-based structure- prediction tools for RNA (Meyer et al. 2012). It was later shown that the k-turn and the pk-turn are functionally exchangeable in both RNase P and the SAM-I riboswitch

(Daldrop et al. 2013). The simplicity and relevance of the functional tests of RNase P suggests its utility as a platform for studying RNA structure-function correlates. These results illustrate the value of studies on RNase P RNA structure and RNA catalysis for providing a broader view of RNA structure and function.

Perhaps the most valuable characteristic of RNase P as a model system is our ability to reconstitute partial and putatively complete complexes in vitro, and to easily assess their activity (Kole et al. 1981, Chen et al. 2012, Tsai et al. 2006). Reconstitution of partial complexes and assessment of their partial catalytic potential relative to the fully assembled holoenzyme provides insight into the specific role of each of the holoenzyme subunits. Moreover, given the relative ease of reconstituting RNase P and measuring its activity, one can assess the functional effects of mutation on the holoenzyme, and thereby gain insight into the role and relative importance of particular structural modules. Reductionist and reconstitution approaches have enabled biochemists to elucidate the role of individual subunits in multicomponent macromolecular machines. The complexity of machines like the ribosome and the spliceosome renders this strategy difficult. While the ribosome, for example, has been reconstituted, it has not been reconstituted from highly purified recombinant proteins and in vitro transcribed RNA (Traub et al. 1968, Mangiarotti et al. 1997, Bunner et al.

2010). In this regard, RNase P therefore serves as a simpler and more malleable model which is generalizable to much more complicated systems due to the ability to reconstitute it from highly purified recombinant proteins and in vitro transcribed RNA, and the accompanying ability to manipulate those components by mutagenesis or other approaches.

1.2 Using Fluorescence to Study pre-tRNA Binding and Processing by RNase P

For much of the period since its discovery four decades ago, the activity of RNase

P has been measured using assays which monitor cleavage of 32P-labeled pre-tRNA.

Following incubation with a radiolabeled pre-tRNA, the reaction is quenched and the contents separated by denaturing polyacrylamide gel electrophoresis (PAGE). The extent of cleavage is measured using autoradiography or phosphorimaging (Göβringer et al. 2012). While radioisotopic labeling offers exquisite sensitivity, it suffers from drawbacks associated with safety and environmental concerns. Further, the need for exposure to achieve high sensitivity renders it unsuitable for high-throughput or real- time applications. Microplate scintillation counters provide one possible format for a high-throughput radiometric assay, but rapid quantitation may result in poor sensitivity.

Moreover, samples may need to be dried prior to exposure to the scintillation cocktail

(Nassar et al. 2004). Also, the very nature of high-throughput work may not be compatible with the care and caution required to safely work with radioisotopes.

In contrast to radioisotopes, fluorophores offer a number of advantages. For traditional gel-based assays, sensitivity suffers with the use of fluorophores, but increased safety, convenience, affordability (in many, but not all cases), and increased shelf-life all provide a positive counterbalance. Unlike radioisotopes, fluorophores are not subject to the same rapid decay that limits commonly used isotopes such as 32P.

Fluorophores truly shine, however, when used for real-time or high-throughput 5 applications. Fluorescing molecules can be detected directly, without any need for exposure or a secondary step (as in scintillation counting), and so relatively simple detectors can monitor their dynamics in real-time. This allows researchers to assess the time-dependence of phenomena such as folding, binding, or other macromolecular dynamics in solution (Lillo et al. 1997). Understanding dynamic behavior of enzymes and other macromolecules in solution is critical to understanding their function.

The same characteristics of fluorophores that allow real-time measurement allow rapid high-throughput measurement. Fluorescence-based high-throughput assays already see widespread and expanding use in the search for enzyme inhibitors, which can then be used both for detailed biochemical elucidation of the associated pathways, and for the development of novel antibiotics (Gribbon et al. 2003, Lea et al. 2011).

High-throughput screening is increasingly an indispensable tool for the identification of effective and specific inhibitors of enzymes (Gribbon et al. 2003, Lea et al. 2011).

Powerful inhibitors of RNase P might serve as leads to develop novel antibacterial agents, and – if specific for particular variants – would provide valuable biochemical tools for studying the role of the enzyme both in vitro and ultimately in vivo.

Several investigators have explored RNase P as a drug target due to the obvious structural differences between the bacterial and eukaryal forms, its low copy number in bacterial cells, and its essentiality for viability (Eder et al. 2003, Willkomm et al. 2010).

Moreover, a high-throughput assay will expedite fundamental studies by permitting rapid and facile quantitation of rate measurements. Real-time assays, performed using fluorescently-labeled pre-tRNA, have already been shown to be useful in measuring the kinetics of the enzyme-substrate complex formation (Hseih et al. 2010). In single- molecule experiments, fluorescence-based real-time assays will eventually allow the study of RNase P complex assembly and solution dynamics through phenomena such as

Förster Resonance Energy Transfer (FRET). Furthermore, the use of fluorescently- labeled pre-tRNA allows the use of fluorescence anisotropy – a well-established technique to determine the affinity constant (KD) of the enzyme for its substrate.

Indeed, this technique has been widely employed to measure KD values of RNA-protein complexes, including that of Arabidopsis thaliana mitochondrial proteinaceous RNase P

1 (PRORP1) for a pre-tRNACys substrate (Howard et al. 2012).

The Fierke laboratory (University of Michigan, Ann Arbor) has developed a fluorescence anisotropy-based cleavage assay, which uses the post-cleavage change in anisotropy of a 5′-fluorophore attached to a pre-tRNA (Liu 2013). This assay is quite similar to a previously developed assay which uses a fluor-labeled DNA oligomer complementary to a region spanning the cleavage site in the pre-tRNA substrate. This oligomer has higher affinity for the pre-tRNA than the mature tRNA or the 5′-leader, and thus experiences a change in anisotropy which decreases with increasing extent of cleavage (Gopalan et al. 2005). While these assays show promise for high-throughput

7 screening applications, they suffer from a number of drawbacks. In the case of direct anisotropy measurement of a fluor-labeled pre-tRNA, the unstructured nature of many long 5′-leaders may result in a small change in post-cleavage anisotropy because the pre-cleavage leader already provides substantial rotational freedom to the fluorophore

(the so-called “propeller effect”). Therefore, the assay might be restricted to substrates with short leaders. Further, the nature of the product cannot be reliably identified, meaning that miscleavage by RNase P (in the presence of an inhibitor), chemical cleavage by an unknown constituent of a crude cell extract (or due to an unpredicted property of a compound being screened), or cleavage by a contaminating non-target

RNase would all be interpreted as RNase P cleavage. Indirect anisotropy measurement, using a complementary fluor-labeled DNA oligomer, requires an additional step (i.e. oligomer annealing) and also suffers from an inability to identify the nature of the products as gel or capillary electrophoresis can.

1.3 Bioconjugation and Bioorthogonality

Chemical derivatization of biomolecules (bioconjugation) presents a number of unique challenges to chemists. I first to differentiate chemical from enzymatic derivatization – in the latter, biomolecular catalysts (either natural or engineered) often dramatically accelerate the desired chemistry. Chemical derivatization, albeit frequently much slower, lacks the often inconvenient requirement for the reacting molecules to meet the geometric requirements of an enzyme′s active site (Dirksen et al.

2008). This allows any biomolecule with an appropriately reactive moiety to react with any other molecule bearing its counterpart. For this strategy to be successful, however, the reactive moiety and its counterpart must not undergo any significant reaction with other functionalities in target biomolecules. Given the chemical diversity of biomolecules – proteins in particular – identifying reactions which are “orthogonal” to the functional groups has proven difficult. Nonetheless, a number of reactions which proceed selectively and efficiently even in the complex environment of a live cell/organism have been identified. Such reactions are often identified by the term

“bioorthogonal” – meaning they proceed with specificity and efficiency even in the overwhelming chemical labyrinth of life (Sletten et al. 2011, Zeglis et al. 2013, Neves et al. 2013).

In addition to the above mentioned requirements, these bioconjugations must proceed under mild, aqueous conditions. pH and temperature must usually be moderate. Essentially, the conditions must allow the biomolecule to survive the labeling intact and, ideally, retaining the native functional conformation. Certain exceptions to these requirements exist, particularly for in vitro labeling of purified biomolecules, but for in vivo labeling of live organisms they may be absolute.

1.4 Click chemistry offers versatile tools for bioconjugation

“Click” chemistry was coined by Sharpless (Kolb et al. 2001), who defined click chemistry as a set of reactions which must be…“…modular, wide in scope¸ give very high yields, generate only inoffensive byproducts that can be removed by non- chromatographic methods, and be stereospecific (but not necessarily enantioselective).

The required process characteristics include simple reaction conditions (ideally, the process should be insensitive to oxygen and water), readily available starting materials and reagents, the use of no solvent or a solvent that is benign (such as water) or easily removed, and simple product isolation. “

Sharpless suggested in this review that this concept should become a new reaction paradigm in organic synthesis where a novel or generally useful function is the desired outcome. He argued that the exploration of structure and function space essentially takes too long if organic chemists limit themselves to traditional techniques and reproduction/modification of secondary metabolites. Instead, he opined that they should focus on facile reactions which they can use to generate (and modify) large libraries of compounds with ease. These reactions include oxidative modifications of olefins, nucleophilic opening of strained rings, certain dipolar cycloadditions, and a few other heteroatom-carbon electrophile fusion reactions. Even with this relatively small arsenal of near-“perfect” reactions, chemists can relatively easily synthesize compounds

10 of impressive variety, and then use derivatives of hit compounds to assess structure- activity relationships.

The covalent attachment of molecular probes or other modules to biomolecules facilitates delineation of structure-function correlates. Many of the characteristics which make these reactions “easy” to employ in organic synthesis make them desirable in this alternate context. The introduction of a molecular probe into a biomolecule must ideally proceed specifically and efficiently at low reactant concentrations in mild, aqueous conditions. Specificity is frequently crucial – one must be able to target a probe to a specific region of a macromolecule in order to acquire interpretable data.

The need for efficiency at low reactant concentrations arises from the difficulty in obtaining certain macromolecules in large amounts and keeping them soluble in high concentrations, and also from the relative expense of certain probes. Mild and aqueous conditions are crucial to maintain the covalent integrity and supramolecular structure of macromolecules. The high thermodynamic driving force and aqueous compatibility of many Sharpless-type “click” reactions fulfill these requirements.

In 2002, both the Sharpless and the Meldal groups independently published papers identifying a new reaction: the copper(I)-catalyzed azide-alkyne cycloaddition

(CuAAC) (Rostovtsev et al. 2002, Tornøe et al. 2002) (Figure 3). This reaction exemplifies all of the characteristics of a “click” reaction, and all of the more stringent

11 characteristics required of bioconjugation reactions. This reaction, alternately referred to as the Click reaction, the Fokin-Sharpless-Meldal reaction, or the Huisgen-Sharpless cycloaddition, is a stepwise variant of the Huisgen cycloaddition (Huisgen, 1961), which was originally identified by Sharpless as “click” chemistry. Here, I will refer to this reaction as the CuAAC. Despite its high degree of orthogonality – meaning that the functional groups involved do not react with non-target functional groups – the original

Huisgen cycloaddition requires temperatures that are incompatible with most biological macromolecules from mesophiles (Huisgen, 1961; Sharpless et al. 2001). The copper- catalyzed variant, on the other hand, exhibits impressive kinetics (≈105 M-1s-1 per M copper) and can proceed efficiently at room temperature (≈25oC) (Presolski et al. 2010).

The thermodynamic driving force (-84 kJ/mol) for the formation of the highly stable

1,2,3- triazole is such that the reaction is often near-quantitative even at low reactant concentrations (Walter et al. 2012).

Since the discovery of this reaction, its application to bioconjugation has been impressive both in scope and utility. The CuAAC has been used to ligate, decorate, or otherwise derivatize all classes of biomacromolecules (Paredes et al. 2011, Thiele et al.

2012, Slade et al. 2012, Fekner et al. 2009, El-Sagheer et al. 2010). Despite difficulties associated with copper-mediated oxidative damage to the reactants, its presence at the frontlines of bioconjugation is justified by the ability to achieve a high efficiency of modification with low reactant concentrations (Figures 4 and 5). While the click

12 chemistry philosophy may not have substantially changed overarching attitudes towards synthesis of medically relevant molecules, it has indeed significantly affected drug discovery as a whole – in part because of its utility in bioconjugation (Kolb et al. 2003,

Thirumurugan et al. 2013).

1.5 Application of CuAAC to RNA derivatization

Despite the discovery of the CuAAC in 2002, and its application to DNA labeling in 2006, the reaction was not successfully employed for RNA functionalization until

2008, when it was used for fluorescent labeling of 5-ethynyl uridine to study transcription rates in mammalian cells (Gierlich et al. 2006, Jao et al. 2008). Following this initial application, Xu and colleagues at the University of Tokyo creatively demonstrated the existence of a DNA-RNA G-quadruplex by crosslinking a 5′-azido- oligodeoxyribonucleotide with a 5′-alkynyl-oligoribonucleotide (Xu et al. 2009). The reactive moieties are in close proximity due to the formation of the quadruplex, and the crosslinking event is catalyzed by Cu(I). In addition to their identification of a novel hybrid secondary structure, the authors were also the first to demonstrate the utility of the CuAAC for derivatization of intact RNA.

Paredes and Das subsequently speculated that the reason for the delay in application of the CuAAC to RNA was concern for the higher lability of RNA relative to

DNA (Paredes et al. 2011). This may be true, but it is worth pointing out that the

13 difference in chemical rather than enzymatic lability between DNA and RNA is the increased susceptibility of the latter to hydrolysis. It is not clear whether RNA breakdown in the presence of copper(I) would primarily result from metal-assisted hydrolysis or from hydroxyl radical-mediated oxidative damage. The latter mechanism, given the instability of copper(I) in aqueous media and its propensity to participate in

Fenton-like reactions, seems a likely candidate (Figure 4). Interestingly, it has been argued that RNA actually exhibits greater stability to oxidative damage than DNA (Thorp et al. 2000). This, in conjunction with data suggesting that RNA’s conformational properties render it less labile to UV-induced photochemistry, has interesting implications for the possible dual role of RNA as information storage vehicle and catalyst in the ozone-free atmosphere of the early earth (Kundu et al. 2004). Nonetheless, it is clear that RNA is indeed vulnerable to strand scission induced by oxidative modifications either to the sugar or the base, and that this damage is likely to be an issue in applying the CuAAC to RNA modification (Burrows et al. 1998, Pogozelski et al. 1998) (Figure 5).

In spite of this challenge, two groups successfully demonstrate that the Cu(I)-mediated damage can be reduced under certain conditions (Xu et al. 2009, Paredes et al. 2011).

In spite of the availability of numerous chemical strategies for the modification of RNA, none of them combine all of the positive characteristics of the CuAAC – speed, specificity, and stability of the resulting linkage. Moreover, as the CuAAC is orthogonal to all other RNA modification reactions currently in use (though the presence of thiols

14 may interfere with the CuAAC), it can be readily employed in one-pot dual labeling strategies, substantially improving the final yield. While the oxidative damage intrinsic to the CuAAC may render it less than perfect, it is always useful to add another reaction to the RNA modification toolkit.

Here, I show that the CuAAC can be used to incorporate fluorophores at the 5′- end of pre-RNAs and that these labeled pre-tRNAs can be processed by Escherichia coli

(Eco) RNase P. Moreover, I show that laser-induced fluorescence capillary electrophoresis (LIF-CE), performed in an automated DNA sequencer (ADS), may provide a convenient format for assessing endonuclease activity in a medium to high-throughput format, and finally that multiplexing with different fluorescent labels expands the scope of possible studies by permitting the assessment of concomitant RNase P-mediated processing two pre-tRNAs of identical length but different composition.

Figure 1: Processing of a precursor-tRNA by RNase P.

RNase P cleaves the pre-tRNA (left) to generate the mature tRNA, and the 5′-leader (right). Arrow indicates site of cleavage in the pre-tRNA (left).

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Figure 2: Crystal structure of the bacterial RNase P holoenzyme from Thermotoga maritima in complex with a mature tRNA.

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18 Figure 3: Copper(I)-catalyzed azide-alkyne cycloaddition.

Note that the participating alkynes must be terminal.

Figure 4: The mode of hydroxyl-radical generation by copper(I).

Reactions described in Burrows et al. 1998.

Figure 5: Hydroxyl radical-initiated strand-scission.

One possible route to hydroxyl radical-initiated strand-scission. Hydroxyl radical abstraction of the 1′-hydrogen leads to the formation of a labile ribonolactone, which is subject to scission by elimination via 4′-proton abstraction (Burrows et al. 1998).

Chapter 2: Results

2.1 Synthesis of 5′-azido-5′-deoxyguanosine via 5′-iodo-5′-deoxyguanosine

5′-iodo-5′-deoxyguanosine was made by a published procedure (McGee et al.

1986). Molecular iodine and triphenylphosphine were dissolved along with guanosine in

1-methyl-2-pyrrolidinone in the presence of imidazole (Figure 6). Recovery of 5′-iodo-5′- deoxyguanosine after the reaction was accomplished by simple dilution with a mixture of dichloromethane and water, during which the product precipitates at the interface and can be recovered by filtration as a yellow solid. The 1H NMR spectrum of the product agrees with reported literature values, and we additionally contribute an FT-IR spectrum of the non-crystalline solid (Figures 22 and 23; see Methods section).

AzG can then be synthesized by simple SN2 displacement of iodide (from 5′-iodo-

5′-deoxyguanosine) by inorganic azide in water (yield ≈30%). AzG precipitates from the aqueous solution upon cooling. The FT-IR and 1H NMR spectra of the product matched those reported in the literature (Brear et al. 2009, McGee et al. 1986) (Figures 24 and

25; see Methods section).

2.2 Incorporation of AzG in RNA transcripts

There are two reports on the successful incorporation of AzG in in vitro transcripts made using T7 RNA polymerase (Paredes et al. 2011, Williamson 2007). I

21 indirectly confirmed this finding with successful CuAAC-mediated attachment of a fluorophore exclusively to the 5′-end. Given the reported orthogonality of the CuAAC, the successful reaction of RNA with an alkyne-bearing fluorophore is evidence of azide

(via AzG) incorporation. However, I was interested in the extent of AzG incorporation in order to both quantitatively assess the efficiency of the CuAAC reaction and ultimately to determine the viability of the labeling strategy for single-molecule applications where near-quantitative reactions may be essential.

In any in vitro transcription of RNAs containing internal guanosine residues, guanosine-5′-triphosphate (GTP) is needed. However, GTP will compete with AzG for the initiating position in the transcript (Figure 7). Therefore, the extent to which AzG is incorporated instead of GTP will limit the final efficiency of subsequent CuAAC-mediated derivatization. After confirming the success of in vitro transcription reactions in the presence of AzG (Figure 8), I sought to investigate this question. While derivatization of the azide could provide an easy way to assess the extent of AzG incorporation, I attempted to determine the value directly. Initial attempts (performed by undergraduate researcher Eric Reville) to determine AzG incorporation efficiency by thin-layer chromatography of RNase T2-digested 32P-internally labeled transcripts were complicated by difficulties related to resolving AzG-3′-monophosphate from the other nucleoside-3′-monophosphates. Subsequently, we pursued an alternative approach motivated by a suggestion from Professor Craig Martin (Department of Chemistry,

University of Massachusetts, Amherst) who pointed out that in a very short transcript, the 5′-azide and a 5′-triphosphate may have a substantially different electrophoretic profile. He had previously demonstrated the ability of T7 RNA polymerase to generate transcripts as short as four nucleotides from synthetic templates (Martin et al. 1987).

Therefore, I transcribed the 5′-GAAC-3′ tetramer in the presence of GTP and AzG as well as [α-32P]-ATP for visualization, and used PAGE to resolve the two expected products.

The 5′-azide and the 5′-triphosphate products were well separated on a 15% polyacrylamide, 7 M urea gel, allowing facile quantitation of the extent of incorporation.

(Figure 9) I determined that AzG initiates 87 ± 2% of all transcripts under the conditions

I initially tested (1.2 mM GTP, 6 mM adenosine-5′-triphosphate, 6 mM cytidine-5′- triphosphate, 6 mM uridine-5′-triphosphate, 4.8 mM AzG). This result was obtained from four independent trials. However, when AzG and GTP are present at an equal concentration (both 1 mM), the extent of AzG incorporation decreased to 37%

(observed in only one trial).

2.3 Synthesis of 5(6)-propargylamidofluorescein

To fluor-label pre-tRNA with the CuAAC, I synthesized my own alkyne-derivative of fluorescein. Carboxyfluorescein, which may be readily synthesized from resorcinol and trimellitic anhydride, has a convenient reactive handle in the form of a carboxyl group, which may be derivatized using standard peptide chemistry techniques (Angell et al. 1994). In our particular case, we used diisopropyl carbodiimide as an electrophile to

23 activate the free carboxyl group. This activated carboxyl group is then attacked by N- hydroxybenzotriazole, which serves as a nucleophilic catalyst and is ultimately displaced by propargylamine. Recovery of the product is readily accomplished by extraction of the crude solid, dissolved in phosphate pH 7, with ethyl acetate, followed by acidification of the aqueous phase, which results in precipitation of 5(6)- propargylamidofluorescein (FAMA). This is a convenient and inexpensive synthesis of a reactive fluorophore which afforded me convenient access to the means to test CuAAC- mediated RNA labeling (Figure 10).

2.4 CuAAC-mediated 5′-terminal labeling of RNA

We have confirmed the ability of the CuAAC to efficiently modify azide-bearing

RNA in the presence of acetonitrile (Figure 11). With 100 µM 5′-azido-RNA and 300 µM

Alexa Fluor® 647 fluorophore, I achieved 64.6 ± 0.9% (standard deviation is from three technical repeats of the measurement method) labeling of total RNA, as indicated by the ratio of the absorbance at the respective maxima of the fluorophore and the RNA.

Other labeling reactions resulted in a similar extent of incorporation (Figure 13). 5’-

Labeling is confirmed by digestion of the pre-tRNA transcript with RNase P, after which the 5′-leader fluoresces but the mature tRNA does not. Given the 5′-location of the label, only the pre-tRNA and the resulting 5′-leader should be visible, as confirmed by my results (Figure 12).

I am able to achieve near-homogeneity of labeled RNA prior to gel purification – with the exception of the labeled putative early-termination products – by the inclusion 24 of a substantial quantity of dimethyl sulfoxide (DMSO). DMSO is a well-characterized hydroxyl radical scavenger, and its reaction with the hydroxyl radicals prevents their eventual cleavage of the RNA (Repine et al. 1981, Eberhardt et al. 1988). The use of 50%

(v/v) DMSO as a co-solvent allows this reaction to be performed without any prior degassing, and has no apparent effect on the efficiency of the reaction. Short products are clearly visible after PAGE separation, but these might be early termination transcription products.

2.5 Periodate oxidation and the CuAAC are orthogonal, enabling one-pot dual-

5′- and-3′-terminal labeling of RNA

Despite difficulties associated with ROS-mediated modification and breakdown, one significant advantage of adding the CuAAC to the repertoire of reactions used to derivatize RNA is that it is orthogonal to essentially all other modification reactions one might employ. RNAs have been labeled with amine and sulfur-selective electrophiles, as well as with hydrazides and thiosemicarbazides after periodate oxidation (Proudnikov et al. 1996, Rueda et al. 2005, Zhang et al. 2001, Williamson et al. 2007). The introduction of multiple labels in a single macromolecule is increasingly used in studies of macromolecular dynamics by FRET. The ability to introduce two or more labels in a one- pot reaction would substantially improve final product recovery as compared to a sequential process separated by a purification step (e.g., gel purification).

In the interest of exploring possible one-pot combinations of the CuAAC, I investigated the possibility of simultaneously introducing a 5′-fluorophore and a 3′- biotin using the CuAAC and aldehyde-hydrazide condensation. I treated periodate- oxidized 5′-azido-RNA with a 5-fold excess of biotin-hydrazide at pH 4.5 in the presence of aniline, which acts as a nucleophilic catalyst for the condensation. After a 20 min incubation at room temperature (≈22oC), I titrated the reaction to pH 7.5 and added the catalyst and alkyne-bearing fluorophore for the CuAAC. The modified RNA was subsequently gel-purified, and assayed by an electrophoretic mobility-shift assay in the presence and absence of streptavidin (Figure 14). A clear retardation of the fluorescent

RNA band was observed in the presence of streptavidin, confirming the successful modification with both fluorophore and biotin at the 5′-and 3′-termini respectively. The extent of biotin modification appears to be only 8%, suggesting the need for further optimization. However, previous success with the CuAAC and the extensive literature on the aldehyde-hydrazide condensation suggest that quantitative one-pot dual labeling is achievable. Among the problems may have been the addition of the basic aniline without correcting the pH. Further, considerable time passed between the initial reaction and the final assessment of the biotin-conjugation, and it is possible that the labile hydrazone linkage may have undergone substantial hydrolysis (Kalia et al. 2008).

2.6 5′-labeled precursor tRNA substrates can be used to measure RNase P activity

While 5′-labeled fluorescent substrates have been used in several studies on

RNase P, the covalent introduction of a bulky fluorophore may affect substrate recognition and processing. In fact, at least one case has been reported wherein the introduction of a bulky chemical cross-linker substantially increased substrate affinity relative to the unmodified substrate – the KM of a 3′-modified substrate lacking the 3′- terminal-CCA sequence decreased 2.6-fold in the presence of the cross-linker (Oh et al.

1994). Further, it has been demonstrated that the introduction of fluorescein at the 5′- terminus of a pre-tRNA with a short leader does in fact affect its affinity for the RNase P holoenzyme in Bacillus subtilis (i.e. a 5′-fluorescein on a 3 or 4-nt leader decreases the

KD by a factor of 2 to 4) (Hsieh et al. 2009). Given this caveat, it is critical to demonstrate that such a substantial chemical alteration does not generate misleading results. I first showed that in vitro reconstituted Eco RNase P is able to cleave the 5′- fluor-labeled substrate. RNase P is clearly capable of processing a variety of 5′-fluor- labeled substrates with similar efficiency to values reported for 32P-labeled RNA. While it is easy to imagine that a fluorophore might have some effect such as altering the folding equilibrium of the substrate or mediating non-specific interactions with the enzyme, the radio-labeled substrate is expected to behave identically to the unlabeled substrate. Comparison of RNase P processing of fluor-labeled versus radio-labeled substrate is therefore a reasonable litmus test for the former’s utility. I demonstrate 27 here that the same stock of reconstituted Eco RNase P processes the fluor- and radio- labeled substrates with similar efficiency (Figures 15 and 16). This result was expected, as the position of the fluorophore at the end of the leader is many nucleotides distal to the cleavage site (between positions -1 and +1 in the pre-tRNA). Moreover, the fluorophore is unlikely to interfere with recognition of nucleotides -3 through -7 by the protein cofactor (Rueda et al. 2005).

It is interesting to note that I was not able to achieve the above-mentioned results until I avoided UV-irradiation of the fluor-labeled substrate during gel- purification. It is possible that UV-induced crosslinks or base-modifications, which are known to occur during UV-shadowing of RNA, interfered with the ability of the enzyme to efficiently bind and process the entire population of labeled RNA (Kladwang et al.

2012). Nonetheless, switching to a visible stain (methylene blue) for the visualization of the fluor-labeled RNA during the purification process solved the difficulties, and RNase P is able to process 5′-fluor-labeled pre-tRNATyr with similar efficiency to that with which it processes the 5′-terminally radio-labeled counterpart. It is useful to note that purification large amounts of RNA labeled with brightly colored fluorophores, such as

Cy3, does not require any stain as the migrating band is visible to the eye.

2.7 Labeling different substrates with different fluorophores allows simultaneous quantitation of RNase P processing of multiple substrates

RNase P activity is typically studied by assessing the processing of a single substrate in vitro. However, RNase P has the task of processing the entire ensemble of pre-tRNAs in most organisms, in addition to a growing number of non-pre-tRNA substrates (Jarrous et al. 2010). Therefore, it is worth investigating the simultaneous processing of multiple substrates in vitro, as it better represents the task of the enzyme in vivo. In a recent study, the authors grappled with the difficulty of studying the simultaneous processing of two different radio-labeled substrates of the same length and therefore unresolvable by gel electrophoresis (Yandek et al. 2013). The authors successfully addressed this challenge by inserting two terminal G-residues in one substrate and therefore increasing its length. Here, I offer an alternative strategy: the independent quantitation of two substrates unresolvable by PAGE by labeling them two different fluorophores.

As a model for this demonstration, I chose to assess RNase P processing of pre- tRNATyr both with and without the 5′-ACCA-3′ sequence at the 3′-end. This sequence is known to interact with the P15-17 region of the catalytic domain of the in RNase P RNA

(Figure 20), and to contribute 2 kcal/mol to the total binding energy (Oh et al. 1994).

RNase P is capable of cleaving substrates lacking the sequence, but the KM value for these substrates is 3 to 6-fold higher than the wild-type substrates (Altman 1999), and in 29 some cases the enzyme is unable to position the pre-tRNA correctly and mis-cleaves the substrate. It is noteworthy that disruption of the interaction in vivo is fatal to organisms in which it has been attempted (Wegscheid et al 2006, Wegscheid et al. 2007). While a pair of pre-tRNAs differing only in the presence of 3′-terminal-ACCA can be resolved, successful resolution would likely require the use of a sequencing-type polyacrylamide gel, which would not be ideal for routine assays. Thus, it would be convenient to use two different fluorophores for this task.

I attempted to demonstrate the principle of assessing simultaneous processing with orthogonal fluorophores using two different substrates labeled with either FAMA or Cy3. It is unfortunate that while FAMA does not appear to interfere with RNase P processing, Cy3 in fact does so to some extent. In many cases, I found a modest decrease in apparent kcat for Cy3-labeled substrates relative to their FAMA-labeled counterparts, though with certain substrates the opposite was found to be true. To circumvent this difficulty, I attempted to demonstrate the proof of concept by using both combinations of fluorophore and substrate (Figure 17). First, I determined the kcat values by assessing the rate of processing of each substrate at saturation (based on previously reported KM values), and then I calculated an expected kcat value based on the competitive-inhibition-like process predicted to occur when an enzyme processes two substrates (Altman 2002). The experimentally determined values were similar to the predicted values. (Table 1)

I surmised that the differences observed between processing of Cy3-labeled substrates and FAMA-labeled substrates by RNase P resulted from the cationic nature of the Cy3 fluorophore. To circumvent this problem, I labeled pre-tRNAPheTyr+CCA both with

FAMA and with AF647. While AF647 is a cyanine fluorophore it is polyanionic owing to its four sulfonyl groups. I compared Eco RNase P-mediated cleavage of 100 nM 5′-FAMA- and 5′-AF647 using pre-tRNAPheTyr+CCA by incubating both substrates with Eco RNase P (1 nM) (Figure 18). I observed that RNase P cleaves the FAMA-labeled substrate about

15% faster than the AF647-labeled substrate. It is unclear why this is the case, but this is only a modest difference and suggests that the two fluorophores could still be used for the purpose of ratiometric analysis in high-throughput screening applications.

2.8 A capillary electrophoresis-based automated DNA sequencer is a convenient format for assessing RNase P cleavage assays

To identify a convenient format for high-throughput assessment of RNase P processing, I chose to investigate the utility of LIF-CE in an ADS (ABI 3730). I first attempted to examine a time-course reaction with Eco RNase P and pre-tRNATyr. These aliquots were subjected to desalting by drop dialysis, and then electrokinetically injected into the sequencer in the presence of DNA size standards. This initial run was a success as it demonstrated the clear separation of the fluor-labeled pre-tRNA and 5′- leader. However, quantitation of precursor and leader peak intensities did not produce

31 values that matched those obtained from the gel image (trending lower than expected).

Various possible explanations exist, but the two most likely are the following: bias during electrokinetic injection or biased sample loss during desalting.

This trend continued when I injected the contents of an entire RNase P assay, after desalting by dialysis against water. The same assay when assessed by PAGE

-1 -1 exhibited a kcat of 34 ± 2 min , while LIF-CE ADS yielded a kcat of only 28 ± 2 min (Figure

19). While the source of the bias in this particular experiment remains unclear, it seems likely that dialysis using a 0.25 µm membrane was responsible for a selective loss of the

5′-leader. If the bias resulted solely from electrokinetic injection, one would have expected its direction to be opposite, as the smaller electrophoretic profile of the 5′- leader should have resulted in its preferential injection.

This prediction was borne out when I used LIF-CE to assess RNase P cleavage without any prior desalting procedure. Instead, I attempted to mitigate salt-mediated risk to the capillaries by diluting the assays 3-fold, 10-fold, or 100-fold. All three dilutions resulted in the same trend – an apparent rate consistent with kcat value of 23.4

-1 -1 ± 0.3 min , as compared to a kcat value of 21 ± 1 min when the same assays were measured after separation by PAGE – an 11% increase in apparent kcat. This increase is expected due to the smaller electrophoretic profile of the 5′-leader. The magnitude of this modest increase is expected to increase with decreasing leader length.

Nonetheless, for many applications, such a small increase in apparent rate may be acceptable, particularly when one is primarily interested in a comparison of observed rates in the presence and absence of an inhibitor. Given the large quantity of salt used in the cleavage reactions (>400 mM), one might expect that the injection of the untreated sample to be unsuitable for LIF-CE. The samples are still diluted, in order to decrease risk of salt deposition in the capillaries, but the lack of any purification step renders this method more attractive for medium or high-throughput applications.

I also compared rates calculated from PAGE separation of assays before and after ethanol precipitation, and found that, in the case of pre-tRNATyr with a 43- nucleotide leader, the apparent rate actually increased after precipitation. In another case, I used ethanol precipitation to desalt assays performed with cyanobacterial pre- tRNAGln (discussed in more detail on p. 35). In this instance, I did not re-assess the samples using PAGE after precipitation, and only compared the pre-precipitation PAGE and post-precipitation LIF-CE ADS values. I found that the apparent rate, according to the LIF-CE assay, decreased moderately. While I cannot clearly identify the relative contributions of precipitation and electrokinetic injection to the bias in the latter case, the substantially smaller electrophoretic profile of the 10 and 11-nucleotide pre-tRNAGln leaders suggests that if a bias resulted from injection, it should have caused a substantial increase in apparent cleavage rate. It is difficult to rationalize these contradictory results. In one case, ethanol precipitation appears to favor the recovery

33 of the leader relative to the precursor, while in another the opposite is true. Perhaps these results stem from differences the in solid-phase packing efficiencies of the two leaders. While desalting procedures may be attractive for increasing signal intensity

(and perhaps also for increasing the lifespan of the capillaries), inefficient recovery of the actual analytes given their low starting concentrations under our conditions renders this approach less appealing. Moreover, as discussed above, desalting does not appear to be necessary for acceptable signal strength. In future studies, it will be interesting to further explore various possibilities for high-throughput desalting, such as the use of C-

18 pipette tips with a multichannel pipette. Results from such investigations could expand the scope of this work.

LIF-CE-based ADS instruments have advantages beyond their high-throughput nature. They offer the high resolution of capillary electrophoresis and facile detection of multiple fluorescent labels. Moreover, software such as PeakScanner (Applied

Biosystems Instruments) renders quantitation of fluorescence peaks from multiple labels relatively straightforward. Due to these advantages, we have pursued the sequencer as a format for resolution of products generated by RNase P mis-cleavage, and for the assessment of products generated by simultaneous RNase P processing of multiple similarly-sized substrates (distinguished by different fluorophores).

My initial mis-cleavage experiments measured Eco RNase P processing of pre- tRNAGln, whose uridine residue at the +1 position (relative to the canonical cleavage site) results in a mis-cleavage between the +1 and +2 positions (Kikovska et al. 2005). In consensus pre-tRNA substrates, a -1 uridine residue is known to interact with the adenosine-248 in the M1 RNA. A +1 G-C base-pair and prevents mis-cleavage between the +1 and +2 positions (Chen et al. 2012). In pre-tRNAGln, the lack of these attributes and the presence of a +1 uridine cause misalignment of the active site, and this promotes mis-cleavage (Chen et al. 2012) (Figure 20). I measured Eco RNase P processing of fluor-labeled pre-tRNAGln by PAGE and then using LIF-CE ADS. I found that

-1 quantitation of the separated peaks results in a kcat of 55 min from PAGE, and 45 ± 1 min-1 by the sequencer (Figure 21). As discussed on p. 34, this bias is expected to be primarily due to the precipitation step rather than the electrokinetic injection, on account of the direction – electrokinetic injection would be expected to cause an increase in apparent kcat due to the smaller size of the 5′-leader. The sequencer did indeed successfully resolve the mis-cleavage, though the determined extent of mis- cleavage was slightly higher based on the sequencer data. This difference appears likely to be due to the incomplete resolution of the two leader peaks. In future runs, voltage and run-time will have to be optimized to address this concern of incompletely resolved peaks (Figure 28).

Figure 6: Synthesis of 5′-azido-5′-deoxyguanosine via 5′-iodo-5′-deoxyguanosine

Figure 7: In vitro transcription initiated with AzG.

Typical conditions involve 1.2 mM GTP, 4.8 mM AzG, and 6 mM of each of the remaining NTPs.

Figure 8: Agarose gel depicting in vitro transcriptions of 5′-N3-RNAs.

Tyr Gly Gly (Lane 1) 5′-N3-pre-tRNA , (Lane 3) 5′-N3-pre-tRNA , (Lane 4) and 5′-ppp-pre-tRNA . (Lane 2double-stranded DNA size marker (in base pairs).

Figure 9: Assessment of AzG incorporation relative to GTP at the first position of tetramer 5′-GAAC-3′.

(Lane 1) AzG is present but not GTP; (Lane 2) GTP is present but not AzG; (Lanes 3 – 5) 4.8 mM AzG, 1.2 mM GTP (three independent replicates). Reactions were separated on a 15% polyacrylamide, 7 M urea gel. The amounts of radioactivity in each band was determined using a phosphorimager.

Figure 10: Synthesis of 5(6)-propargylamidofluorescein from 5(6)-carboxyfluorescein.

Figure 11: 5′-modification of RNA using the CuAAC.

R = Fluorophore

1 2 1 2

pre-tRNA mature tRNA

5'-leader

Figure 12: pre-tRNAPheTyr 5′-labeled with FAMA.

The substrate was either incubated in RNase P reaction buffer (Lane 1) or digested with RNase P (Lane 2), and then separated on a 10% polyacrylamide 7 M urea gel. Lanes 1 and 2 (left): gel excited with a 488 nm laser and fluorescence detected through a 520 nm emission filter. Lanes 1 and 2 (right): Gel stained with ethidium bromide, then excited with a 532 nm laser and detected through a 610 nm filter. Note that the mature tRNA is visible only after staining with ethidium bromide as the 5′-label has been removed along with the 5′-leader by RNase P. The small (15 nucleotide) 5′-leader cannot be detected in the ethidium bromide scan, possibly due to the low sensitivity and small size of the RNA.

For clarity, please note that the two lanes were not originally adjacent on the gel – this image was created by splicing together the two lanes.

Figure 13: Ultraviolet and visible spectrum of pre-tRNAPheTyr-CCA 5′-labeled with Alexa Fluor® 647 (AF 647).

The ratio of the AF 647 peak to the RNA peak indicates of 64.6 ± 0.9% labeling (from three technical repeats of the measurement of one RNA). The sharp peak at ≈230 nm suggests the presence of some organic contamination.

Figure 14: Streptavidin-mediated gel-shift demonstrating fluor-labeling and biotinylation of the pre-tRNA in a one-pot reaction.

Separation performed on a 12% polyacrylamide gel. (Lane 1) pre-tRNA in water. (Lane 2) pre-tRNA in 2 M urea; (Lane 3) pre-tRNA and streptavidin in water; and (Lane 4) pre- tRNA and streptavidin in 2 M urea.

A B

SC Incubation time SC Incubation time pre-tRNA

5′-leader

Figure 15: Cleavage of 5′-32P- (A) and 5′-FAMA- (B) pre-tRNATyr processing by Eco RNase P in vitro.

SC: substrate control

Figure 16: Cleavage of 5′-P32- (left) and 5′-FAMA- (right) pre-tRNATyr processing by Eco RNase P in vitro.

The kcat values were calculated based on saturating conditions at 100 nM, an assumption justified by the reported KM of 30 nM for pre-tRNATyr (Altman 1999).

Figure 17: Eco RNase P processing of 5′-FAMA-pre-tRNATyr+CCA and 5′-Cy3-pre-tRNATyr-CCA (which lacks the 3′-terminal-CCA) assessed in the same reaction.

Left: Gel scanned for FAMA. Right: Gel scanned for Cy3.

Alone FAMA+CCA Cy3+CCA FAMA-CCA Cy3-CCA -1 kcat (min ) 25 28 132 74 Mixed FAMA+CCA Cy3+CCA FAMA-CCA Cy3-CCA -1 kcat (min ) 16 12 53 20 v observed 0 16 12 53 20 (nM/min) v predicted 0 10 11.2 63.36 35.52 (nM/min)

Table 1: Eco RNase P processing of pre-tRNATyr substrates either bearing the 3′-terminal- ACCA sequence (+CCA) or lacking it (-CCA), and labeled with either Cy3 or FAMA.

( )

Figure 18: Eco RNase P processing of both 5′-AF647 and 5′-FAMA-pre-tRNAPheTyr+CCA in the same reaction.

Each substrate at 100 nM concentration, for a total of 200 nM pre-tRNAPheTyr+CCA. The kcat values on the plot are apparent values calculated by assuming the rate of cleavage for each substrate to be representative of the total rate. I am assuming 200 nM substrate to be saturating. FAMA-labeled substrate is consistently cleaved about 15% faster than the AF647-labeled substrate. This is a plot from one experiment. The same trend was observed in another trial with the +CCA substrate, and also with the –CCA substrate.

Figure 19: LIF-CE assessment of pre-tRNATyr cleavage by Eco RNase P.

-1 This assessment was performed after drop-dialysis to desalt the RNase P assay samples. The kcat of 28 ± 2 min is the mean -1 from three independent experiments, and is comparable to the kcat of 34 ± 2 min measured for the same samples using PAGE (Figure 16).

Figure 20: Mis-cleavage of non-canonical pre-tRNAs by Eco RNase P.

Gln Pre-tRNA from Synechocystis is mis-cleaved due to its U+1 and G+2-C+69 basepair.

Figure 21: PAGE-band assessment of pre-tRNAGln cleavage by Eco RNase P compared to the LIF-CE experiment.

M+1 = Amount of incorrect leader (formed through mis-cleavage). C0 = amount of correct leader. The kcat values reported

represent the means from three independent trials.

Chapter 3: Discussion

3.1 Incorporation of AzG at the 5-terminus of RNAs transcribed in vitro

While various strategies exist for RNA labeling, a number of them are unattractive due to their lack of specificity arising from the use of a large excess of modifying agent. Thus, the quantitative CuAAC-mediated labeling of an azide-bearing small RNA with only a three-fold excess of an alkyne-conjugated fluorophore and recovery of homogeneously labeled RNA is noteworthy (Paredes et al., 2011).

Motivated by these results and the economy of this method, I considered a similar approach to attach a fluorophore to the 5′-terminus of in vitro transcribed pre-tRNAs.

Using existing methods, I first synthesized 5′-azido-5′-deoxyguanosine (AzG) in a two-step process via 5′-iodo-5′-deoxyguanosine (Figure 6). I was able to incorporate

AzG at the 5′-terminus of various pre-tRNAs by including it in T7 RNA polymerase-based runoff in vitro transcription reactions. It was important to determine the extent of AzG incorporation, as it provides a maximum threshold for RNA modification with CuAAC.

Williamson (2007) quantitated the incorporation of AzG during in vitro transcription by first reducing the azide with tris(2-carboxyethyl)phosphine and then derivatizing the resulting 5′-amine with an appropriately reactive biotin. Such an indirect approach for quantitating azide incorporation will yield accurate values only if the reduction and the

53 biotinylation each proceed with 100% efficiency, which is not the case. I used a simpler and more direct approach (see p. 21) to determine 87 ± 2% AzG incorporation at the 5- terminus of an RNA tetramer transcribed in vitro with 4.8 mM AzG, 1.2 mM GTP and 6 mM ATP/CTP/UTP.

A priori, the 4:1 ratio of AzG:GTP might suggest that only 80% of the transcripts could be initiated with AzG. There are at least two reasons that might account for the

87% observed in my experiments. First, the 5-triphosphate is likely to have a slightly unfavorable interaction with the T7 RNA polymerase active site during initiation, as evidenced by its KM being higher than that of guanosine or guanosine-5- monophosphate during initiation (Martin et al. 1989); a similar bias in favor of AzG over

GTP is possible. Second, given GTP’s = 0.6 mM, the use of 1.2 mM GTP might not be sufficient to operate under saturating conditions (Martin et al. 1989). While

there are no literature values for AzG’s , I expect 4.8 mM AzG to permit

saturation (based on guanosine’s = 0.46 mM), or at least render initiation with

4.8 mM AzG more favorable than 1.2 mM GTP. The latter expectation is supported by my finding that AzG is incorporated in only 37% of RNAs when transcription is performed with 1 mM GTP and 1 mM AzG.

Given that in vitro transcriptions of internal-G containing RNAs will always require GTP, it is important to consider approaches to eliminate those RNAs initiated 54 with GTP such as those remaining have a 5-AzG. Indeed, I have gathered preliminary data suggesting the utility of tobacco acid pyrophosphatase and XRN-1 to remove transcripts initiated with GTP (or any other NTP) as opposed to the desired modified nucleoside (data not shown). In principle, tobacco acid pyrophosphatase will cleave pyrophosphate from any 5-triphosphate, generating a 5-monophosphate, which can then be recognized by XRN-1, an exonuclease. This approach, if optimized, would ensure that all the RNAs carried forward to CuAAC have an azide at the 5-terminus.

The model RNA tetramer used for this analysis might not reflect conditions that will be encountered during in vitro transcription of longer RNAs, which will likely contain a number of internal guanosine residues. In such an instance, GTP will be depleted at a much faster rate than AzG thus increasing the efficiency of initiation with AzG; however, this gain is counter-balanced by early termination that might ensue upon rapid depletion of GTP (whose initial concentration is typically four-fold lower than AzG).

Thus, identifying the most favorable conditions for obtaining full-length, long RNAs with

5-AzG will require additional work; in particular, empirically fine-tuning rNTP concentrations to ensure their near-even depletion might nullify concerns regarding synthesis of partial transcripts. In the meantime, our studies with a model RNA provide a useful framework for quantitating AzG-initiated transcription.

3.2 CuAAC-mediated 5-fluor labeling of RNA

A difficult aspect of working with commercial fluorescent probes is the cost. This is particularly true for alkyne-bearing fluorophores, which seem to be less popular – and therefore less commercially available – than their azide counterparts. Despite the availability of a variety of cheap fluorescein derivatives for bioconjugation (including an azide conjugate), a fluorescein-alkyne is not commercially available. I have shown here that 5(6)-propargylamidofluorescein can be readily synthesized from inexpensive components using standard peptide chemistry techniques. Moreover, I have used only a modest excess of this fluorophore to label 5-azide-bearing pre-tRNAs by CuAAC.

Despite many appealing attributes, some challenges remain with CuAAC-mediated labeling of RNAs.

The kinetics of the CuAAC reaction are attractive, particularly given the ability to accelerate the reaction dramatically by increasing the concentration of the copper(I) catalyst. While most work with this reaction employs a variety of specific Cu(I) ligands, which serve to accelerate the reaction and prevent Cu(I)-mediated oxidative side- reactions (by acting as sacrificial reductants in lieu of the reactants and product),

Paredes and Das (2011) reported that the use of acetonitrile as a co-solvent can mimic the protective effects of these ligands. Further, they argue that the rate enhancement mediated by specific ligands is unnecessary under excess copper(I) conditions typically used in bioconjugation. Paredes and Das (2011) report, impressively, that under their

56 conditions they achieve 100% modification of azide-bearing RNA (Paredes et al. 2011).

Later, they reported that 83% modification of alkyne-bearing RNA can be achieved in 30 min with micromolar concentrations of both reactants and catalyst, and that this modification occurs with little oxidative breakdown of RNA (Paredes et al. 2012). I am able to confirm the efficacy of the ligand-free CuAAC, achieving 64.6 ± 0.9% modification

(where the % modification is the mean of three independent measurements of the same labeled RNA) at 100 µM 5-azide-bearing pre-tRNA and 300 µM 5(6)- propargylamidofluorescein. A key reason for the 65% incorporation (compared with the

87% predicted based on AzG incorporation) is the possible reduction of the azide to an amine by dithiothreitol, which is present in the in vitro transcription reaction.

Moreover, we emphasize at least two sources of uncertainty associated with the 65% estimate. First, we have not accounted for hypochromic effects associated with π- stacking, which might result in a suppression of the apparent extent of fluor labeling.

Second, we do not know the exact extent of azide incorporation in our full-length pre- tRNA transcripts (even though we have ascertained the same in model RNAs).

I was able to circumvent the oxidative degradation of RNA with the use of DMSO, a well characterized hydroxyl radical scavenger. However, a number of caveats to the general use of DMSO exist. Foremost, the reaction of DMSO with hydroxyl radicals produces methyl radicals, which themselves can undergo reactions with the RNA – they can induce RNA strand scission and methylate cytidine residues (Kasai et al. 2009, Kawai

57 et al. 2010). Strand-scission products are separated and easily visible (above the limit of detection) after denaturing PAGE. Even those products which are not visible will still be efficiently removed by gel-purification. Base modifications, such as cytosine methylation, however, are not obvious and will not be removed by gel purification. This concern exists also in the absence of DMSO, as reactive oxygen species (ROS) are capable of base modification (such as the formation of 8-oxo-guanosine) as well. In light of these concerns, it seems that thorough degassing of the reactions and the absence of any chemical ROS scavenger might be ideal. However, I have found it difficult to achieve sufficiently thorough degassing, and I imagine that the apparatus necessary to achieve this might render the chemistry inconvenient to many prospective users. For many ensemble assays, a comparison of enzyme activity on labeled versus unlabeled substrate may be sufficient to determine the extent of problematic base modification. For single- molecule assays, on the other hand, variation from molecule to molecule caused by base-modification might complicate results, although grouping based on individual behavior might help resolve such heterogeneity. To entirely avoid this issue of oxidative degradation of RNAs, I propose an alternative below.

Catalase, which has been shown to protect DNA from copper-generated ROS

. damage, could be used to eliminate H2O2, and prevent its reduction to OH (Oikawa et al. 1996). This strategy has been previously attempted in the context of the CuAAC without success, although no data were provided (Hong et al. 2009). Nonetheless, if its

58 use can be optimized catalase promises to be a superior approach to chemical scavengers such as DMSO as it “detoxifies” H2O2 without releasing any reactive intermediates. I conducted preliminary experiments with a crude preparation of

Anopheles gambiae catalase but was unable to demonstrate a protective effect. This experiment needs to be repeated with a highly purified preparation of recombinant catalase. Moreover, I expect that for catalase treatment to be effective, copper concentrations must be quite low to avoid copper-mediated destruction of catalase (Orr

1967). In a related vein, recent advances such as chelation-assisted CuAAC might also improve the utility of catalase.

3.3 Periodate oxidation and the CuAAC are orthogonal, enabling one-pot dual-5′- and-

3′-terminal labeling of RNA

Despite difficulties associated with ROS-mediated modification and breakdown, one significant advantage of adding the CuAAC to the repertoire of reactions used to derivatize RNA is that it is orthogonal to essentially other RNA modification reactions one might employ. RNA has been labeled with amine and sulfur-selective electrophiles, as well as with hydrazides and thiosemicarbazides after periodate oxidation (Proudnikov et al. 1996, Rueda et al. 2005, Zhang et al. 2001, Williamson et al. 2007). The introduction of multiple labels in a single molecule is increasingly important for studying macromolecular dynamics by FRET. The simultaneous introduction of multiple labels at specific sites in an RNA in a one-pot reaction would substantially improve final product recovery as compared to a sequential process. It is for these reasons that I have 59 demonstrated the ability to combine the CuAAC with the aniline-catalyzed hydrazone ligation. While my single experiment in this regard has provided proof of principle for a one-pot, dual-modification reaction, efficient dual labeling will require further optimization.

3.4 A capillary electrophoresis-based automated DNA sequencer is a convenient format for assessing RNase P cleavage assays

The need for high-throughput RNase P activity assays is clearly demonstrated by the interest RNase P as an antibacterial target (Eder et al. 2003, Willkomm et al. 2010), and by its utility as a model for studying RNP assembly and catalysis. Previous work using capillary electrophoresis with an absorbance-based detector to assess RNase P processing (Lazard et al. 1998) raised the possibility of using an ADS instrument for medium to high-throughput assessment of cleavage reactions. An ADS has dense capillary arrays and permits as many as 384 sample injections in a single run. These instruments use laser-induced fluorescence, enabling high sensitivity and multiplexing

(i.e., simultaneously assessing output from multiple fluorophores). My demonstration of the power of multiplexing using PAGE illustrates such potential, especially when used in a high-throughput ADS format.

Moreover, CE has spectacular separation efficiency – to the extent that it has been considered likely to largely replace high-performance liquid chromatography

(Weinberger 2000). Its high resolving potential, stemming (in the case of nucleic acids) 60 largely from the capacity to perform electrophoresis at extremely high voltages, is such that it can achieve single-nucleotide resolution of large nucleic acids (Dovichi et al.

1997). This essential characteristic, which is the underpinning for accurate DNA sequencing, also has utility for DNA/RNA processing assays. Facile single-nucleotide resolution would permit assessment of mis-cleavage by RNase P and other similar endonucleases. In addition to improving our fundamental understanding of enzyme characteristics such as substrate recognition, ready characterization of mis-cleavage could allow identification of small-molecule inhibitors (from large libraries) that function primarily by promoting mis-cleavage rather than inhibiting all processing. A technique such as fluorescence anisotropy, which has been used before to screen for inhibitors of

RNase P, would fail to identify such an inhibitor and contribute to a high number of false negatives.

As pointed out earlier, further work is required to allow LIF-CE to be useful in quantitative fundamental studies. In particular, biases associated with desalting can be avoided by diluting samples; biases associated with electrokinetic injection (and the physical dependence of this process on the different electrophoretic profiles on the precursor and the 5-leader) are small, and can be quantified and corrected. If biases cannot be eliminated, they must be thoroughly characterized such that one can quantitatively correct for them.

3.5 Concluding remarks

(a) CuAAC for RNA modification

Methods for introducing a reactive functionality and ultimately a probe at the 5- end of an in vitro transcribed RNA are diverse and efficient (Zhang et al. 2001, Wu et al.

2001, Williamson et al. 2007). These approaches allow the introduction of a thiol, an amine, a hydrazine, a monothiophosphorate, an azide, an alkyne and many other functional groups. Although notable exceptions do exist, including some elegant kinetic studies on RNase P (Rueda et al. 2005), in several instances where important biological questions are addressed using fluorescent tools, probes are still predominantly introduced into RNAs by enzymatic ligation of a synthetic oligomer containing the probe or a reactive functionality (such as an amine) to ensnare a complementary probe. This observation is echoed in the paper introducing the RNA community to the CuAAC, where Paredes and Das (2011) commented that derivatization of RNA was largely performed with solid-phase synthesis (frequently purchased) followed by ligation to in vitro transcribed RNA. This trend is likely to change as researchers become increasingly aware of the 5-modified nucleoside initiator-mediated labeling strategy for RNA, especially its lower cost relative to the ligation strategy, and of the ability to generate near-quantitatively labeled RNA with little need for a chromatographic separation.

There is also increasing appreciation of new and more efficient chemical strategies, and the superiority (and proper use) of existing chemical strategies. A

62 number of researchers report using vast excesses of reactants in an attempt to achieve appreciable modification (Chan et al. 1999, Rueda et al. 2005). Some of these reports involve the use of chemistry, such as hydrazide-aldehyde coupling, which is indeed sufficiently efficient such that only moderate excess (or no excess) is required to achieve quantitative coupling within an acceptable time period (Dirksen et al. 2008). However, others, such as couplings between iodoacetamide-bearing labels and guanosine-5- monothiophosphorate, may reflect intrinsic kinetic limitations of the reaction (though I am not familiar with any rate measurements of this specific reaction). Thus, the continued introduction of more efficient chemical strategies will enable the RNA field to access cheaper routes to efficiently labeled molecules.

Despite its shortcomings, the CuAAC offers one additional route to efficiently labeled RNA. Incorporation of azides and alkynes in synthetic RNA has long been possible, and it has now been rigorously demonstrated that one can readily incorporate an azide at the 5- or the 3-termini using T7 RNA polymerase and poly(A) polymerase respectively (Paredes et al. 2011, Winz et al. 2012). A variety of chemical and enzymatic methods have been shown to permit the introduction of an azide or an alkyne at internal positions of in vitro transcribed RNA. Mitigating oxidative strand scission with

DMSO enables one to achieve very high reaction rates by increasing the catalyst concentration. Also, the high stability of the resulting 1,2,3-triazole ensures that the reaction will go to completion without the need for substantial excess of either reactant.

The latter attribute makes this reaction economically attractive, especially when one considers the cost of quality probes such as the Alexa Fluor® dyes. Moreover, compared to the un-catalyzed strain-promoted version of the Huisgen cycloaddition or the inverse electron demand Diels-Alder cycloaddition, azides and terminal alkynes are small (and therefore relatively unobtrusive in the context of biomolecules), synthetically available, and stable.

(b) Fluor-labeled pre-tRNAs and their utility in studies of RNase P

A significant caveat to working with fluor-modified biomolecules is the possibility that the fluorescent probe will interfere with the normal functioning of the biomolecule.

Indeed, I have shown that in some cases, this appears to be true, while in others it does not (FAMA at the 5does not affect RNase P processing of pre-tRNAs, while Cy3 and

AF647 do so to a varying extent). For certain experiments, fluor-dependent effects are unavoidable, but one hopes that the qualitative nature of – for example – a specific binding event does not substantially change. It is therefore important to find methods to assess the effects of the modification. It is my hope that automated DNA sequencers

(and perhaps other CE-based formats) may soon serve as platforms for such assays. Of course, it should be noted that capillary electrophoresis has already been used for the assessment of RNase P activity, though in that study the authors were using UV absorbance as their mode of detection, and their apparatus cannot be described as high-throughput (Lazard et al. 1998).

As discussed earlier, and as demonstrated by my results, fluorescence further offers the opportunity for multiplexing, which permits the assessment of simultaneous processing of different, but unresolvable substrates. Additionally, the impressive sensitivity of LIF-CE (in the absence of salt which interferes with electrokinetic injection) suggests the possibility of isolating 5-fluor-bearing pre-tRNA after transfection into live cells, and assessing RNase P processing in vivo. This could address concerns regarding the ability of in vitro studies to faithfully reflect RNase P processing in vivo. The use of fluorescence rather than radioactivity for such a study would allow the pairing of in vivo activity measurements with fluorescence microscopy studies on location and transport of tRNAs within a cell. Moreover, the presence of the 5-fluorophore provides a convenient 5-cap, preventing non-specific 5-exonuclease-mediated degradation. A 3- fluorophore could be additionally introduced to render the substrate completely resistant to 3-exonucleases prior to RNase P processing. The exquisite sensitivity of LIF-

CE (in contrast to PAGE) could be exploited to detect small amounts of pre-tRNA or 5- leader recovered from a cell lysate.

Chapter 4: Summary

In this work, I have demonstrated the ease of efficiently introducing an azide at the 5-end of RNA, confirming earlier work (Paredes et al. 2011, Williamson 2007) and importantly providing additional quantitative data. Under my in vitro transcription conditions, I show 87 ± 2% incorporation of AzG, a counter-intuitive finding given the use of 4:1 AzG:GTP (Figure 2). Incorporation of 5-modified nucleosides has not previously been quantitated by direct methods. The straightforward method used here to assess the extent of 5-AzG incorporation through PAGE analysis of model oligomers should aid in assessment of new RNA modification strategies. Moreover, I determined that the in vitro transcription yields are comparable for the same RNA with a 5-N3 or a

5-ppp (i.e., transcriptions with or without AzG, respectively).

I have shown that bacterial RNase P processes 5-fluor-labeled pre-tRNAs with similar efficiency as the radiolabeled counterparts (Figure 16). Moreover, these fluor- labeled pre-tRNAs provide a means to assess RNase P activity in a high-throughput format, and to this end I have shown that a LIF-CE ADS can serve as such a format

(Figure 19). Further, concomitant processing of different pre-tRNAs of similar or identical size can be assessed by multiplexing – labeling each substrate with a different fluorophore – and allowing RNase P to process both substrates at the same time. This ability, which is potentiated by the sequencer optics and software, can be employed to

66 assess competing cleavage of multiple substrates by RNase P. Such experiments will permit a better understanding of how RNase P processes a large number of different substrates in vivo. Moreover, multiple assays with different fluors can be combined and measured in a single capillary, increasing the overall throughput of this application. I hope that these studies with RNase P will motivate broader usage of fluorescence in investigations of other RNAs.

Chapter 5: Materials and Methods

5.1 Synthesis of 5′-azido-5′-deoxyguanosine via 5′-iodo-5′deoxyguanosine

5′-deoxy-5′-azidoguanosine was synthesized first by conversion of guanosine to the 5′-deoxy-5′-iodo derivative following the procedure of McGee et al. (1986). My 1H

NMR spectrum is in agreement with theirs (Figure 23). I add the following IR data to their excellent characterization: Nujol, 1704, 1628, 1565, 1529, 1349, 1244, 1161, 1120,

1008, 941, 826, 723, 683 cm-1 (Figure 22). I also add an observed rotation of -0.053 ±

0.003o for a solution with estimated concentration 3.9 mM in 50 mM bicarbonate pH 10,

o o which corresponds to a specific rotation of [α]D = -35 ± 2 at 25 C. Note that rotations for guanosine and its derivatives are pH dependent. I further add a UV absorbance

-1 -1 maximum at 253 nm (ε253 = 13,200 M cm ) in water, and a shoulder at 273 nm.

The 5′-iodo-5′-deoxyguanosine was then converted to the 5′-azido derivative by the aqueous method of Brear et al. (2009). My 1H NMR spectra are in agreement with theirs and those of Dean (2002) (Figure 25). I also report IR data in Nujol: 2104, 2035,

1694, 1612, 1533, 1177, 1122, 1042, 802, 778 cm-1 (Figure 24). I also measured a rotation of +0.012 ± 0.001o for a solution with estimated concentration 820 µM, which

o o corresponds to a specific rotation of [α]D = +48 ± 4 at 25 C. I report a UV absorbance

-1 -1 maximum observed at 253 nm (ε253 = 12,800 M cm ) measured in water (not buffered

– pH not determined), and a shoulder at 278 nm.

In the synthesis of 5′-iodo-5′-deoxyguanosine, triphenylphosphine has the unusual role of serving as both a nucleophile, generating iodide by nucleophilic attack on iodine, and an electrophile, turning the 5′-hydroxyl of guanosine into a good leaving group for displacement by iodide. This reaction is similar to both the Mitsunobu reaction, which uses diethyl azodicarboxylate to activate triphenylphosphine for nucleophilic attack by a primary alcohol, turning it into a good leaving group, and the

Appel reaction, which uses triphenylphosphine both to generate a chloride nucleophile from carbon tetrachloride, and to activate the alcohol for nucleophilic displacement

(Roper et al. 2011, Mitsunobu et al. 1981). Remarkably, the 2′ and 3′-hydroxyls remain unmodified, despite the reported ability of this reaction to iodinate secondary alcohols.

It has been demonstrated that triphenylphosphine forms a cyclic protecting group (a

2′,3′-oxyphosphorane) in situ, preventing irreversible displacement of the hydroxyls by iodide by retaining of the leaving group in close proximity to the electrophile (Nakagawa et al. 1983). This synthesis, by McGee et al. (1986) represents a substantial improvement over previously reported methods to synthesize 5′-iodo-5′- deoxynucleosides, which required three synthetic steps and a chromatographic separation step (Dimitrijevich et al. 1979).

I then used an aqueous displacement of iodide by inorganic azide to generate the 5′-azido-5′-deoxyguanosine, as described by by Brear et al. (2009). Although

69 superior yields have been reported using dimethylformamide as a solvent, the high boiling point of this solvent renders it difficult to remove (Dean et al. 2002).

5.2 Preparation of fluor-labeled and biotinylated pre-tRNAs

pre-tRNATyr (with 5′-ACCAUCA-3′ at the 3′-end) from Eco and pre-tRNAGln (with

5′-ACCA-3′ at the 3′-end) from Synechocystis, and pre-tRNAGly (with 5′-ACCA-3′ as the 3′- terminal sequence) from the chloroplast of Nicotiana tabacum (Nta) were transcribed in vitro by run-off transcription of linear templates generated by PCR amplification of pUC19 clones using T7 RNA polymerase (Chen et al. 2012). PCR amplifications were performed with a universal forward primer (5′-CGACGTTGTAAAACGACGGCCAG-3′) and universal reverse primer (5′-GGAAACAGCTATGACCATGAT-3′), and the products were digested with either Fok I (for pre-tRNATyr with 3′-terminal-ACCAUCA) or Bst NI (for pre- tRNAGln and pre-tRNAGly) to generate the desired termini. For transcription of pre- tRNAPheTyr (a chimeric substrate combining the 5′-leader of pre-tRNAPhe and the mature tRNATyr, both from Eco), pBT7-pre-tRNAPheTyr was linearized using either Bst NI (for 3′-

ACCA) or Fok I (for no 3′-ACCA), and T7 RNA polymerase was used for run-off in vitro transcription in 100 mM Tris-HCl pH 7.6, 24 mM MgCl2 or Mg(OAc)2, 20 mM spermidine,

0.01% (v/v) Triton X-100, 10 mM dithiothreitol at 37oC for 16 h (Tsai et al. 2002).

Transcripts were purified by DNase I treatment, phenol chloroform extraction, dialysis,

o and ethanol precipitation. 5′-N3-RNA was subsequently incubated at 37 C for 40 min in

70 the presence of 3 to 6 equivalents of FAMA, Cy3 alkyne (Lumiprobe), or Alexa Fluor®

647 alkyne (Molecular Probes), 2% (v/v) acetonitrile, 4 mM Cu(OAc)2, 40 mM sodium ascorbate, and 100 mM Tris-HCl pH 7.5. The labeled RNA was then separated by denaturing polyacrylamide gel electrophoresis (PAGE) (10% with 7 M Urea), visualized with a methylene blue stain, excised from the gel using a razor blade and eluted into elution buffer (20 mM Tris-HCl, 100 mM NaCl, 1 mM EDTA, and 0.01% SDS) in some cases in the presence of 0.1 volume 1:1 (v:v) phenol:chloroform (or in its absence – no difference was observed between the two cases) by shaking at 37oC for several hours.

Following a final phenol chloroform extraction, the eluent was filtered to remove polyacrylamide fragments (0.2 µm cellulose acetate centrifugation filters) and precipitated using ethanol. The recovered RNA was re-suspended in water and quantitated by measuring solution absorbance at 260 nm. This methodology was adapted from that of Paredes et al. 2011. In subsequent trials, the Cu(OAc)2 concentration was increased to 10 mM, and 50% (v/v) DMSO was used.

For 3′-biotinylation, the substrate RNA was incubated in 20 mM NaIO4 for 1 h in the dark, after which the sample was ethanol precipitated. The reaction was performed in the presence of 100 mM NaOAc pH 4.5, 100 mM aniline, and a 6-fold molar excess of biotin-LC-hydrazide (Thermoscientific) at 37oC for 20 min. Subsequently, the reaction was titrated to pH 7.5 with NaOH, then subjected to the CuAAC conditions described

71 above. The product was gel purified, then stored at -20oC for 4 months (unintended) prior to assessment.

5.3 Synthesis of 5(6)-propargylamidofluorescein

5(6)-carboxyfluorescein was prepared as described by Ueno et al. (2004). A mixture of 5(6)-carboxyfluorescein (0.4 g, 1 mmol), hydroxybenzotriazole (HOBt) (0.3 g,

2 mmol), and diisopropylcarbodiimide (0.3 mL, 2 mmol) were combined in THF (10 mL) to form a dark brown/orange solution, which was degassed with N2. After stirring at room temperature for 35 minutes, propargylamine (0.2 mL, 3 mmol) was added. The reaction was left overnight, after which the solvent was removed under reduced pressure to yield crude product as a brown solid (1 g). A portion of the crude product

(100 mg) was dissolved in ethanol (10 mL) and diluted with phosphate buffer (500 mL,

20 mM, pH 7). A portion of the solution (400 mL) was extracted with ethyl acetate (100 mL) and stored overnight at 5oC resulting in a red precipitate, which was worked up in hydrochloric acid to yield a red solid (40 mg). I report IR data in Nujol: 3283, 2727, 2350,

1 2125 (thin film neat), 1741, 1637, 1593, and H NMR data (600 MHz, DMSO-d5): δ 9.32

(1.4 H, t), δ 9.2 (0.4 H, t), δ 9.16 (1 H, t), δ 9.05 (0.3 H, t), δ 8.47 (1.2 H, s), δ 8.34 (0.36 H, s), δ 8.26 (1.2 H, dd), δ 8.18 (0.9 H, d), δ 8.09 (0.9 H, d), δ 8.04 (0.8 H, triplet), δ 7.93 (0.3

H, t), δ 7.7 (1 H, s, δ 7.52 (0.3 H, s), δ 7.39 (1.2 H, d), δ 7.16 (0.4 H, d), δ 6.7 (4.2 H, dd), δ

6.58 (10.3 H, m), δ 6.46 ( 1.6 H, m), δ 6.4 (1.4 H, dd), δ 4.73 (4.7 H, dq), δ 3.97 (4.2 H, dq), δ 3.81 (2.9 H, dd), δ 3.20 (1.9 H, t), δ 3.17 (1 H, t), δ 3.09 (1.4 H, t), δ 3.07 (0.8 H, t),

δ 2.73 (0.8 H, s), δ 2.61 (0.4 H, s), δ 2.87 (0.4 H, s) (Figures 26 and 27). Resonances with

δ 6.58, 6.7, 7.16, 7.39, 8.09, and 8.26 correspond with the spectrum of 5(6)- carboxyfluorescein in NaOD-D2O reported by Ueno et al. (2004). The singlet with δ 2.73 may correspond to a terminal alkyne proton. The presence of a C-C triple bond stretch is also suggested by the 2125 cm-1 peak observed in the neat FT-IR spectrum.

5.4 Preparation of M1 RNA

M1 RNA (Eco RNase P RNA) was prepared by run-off in vitro transcription of a linear template generated by PCR amplification of the gene from pJA2′ (Vioque 1989).

The amplification was performed with a universal forward primer (5′-CGACGT TGT

AAAACGACGGCCAG-3′) and universal reverse primer (5′-GGAAACAGCTATGACCATGAT-

3′), and the product was digested with FokI to ensure a transcript with the desired 3′- terminus.

5.5 Preparation of C5 protein (procedure not performed by this author)

Performed as described in Gopalan et al. (1997).

5.6 Eco RNase P activity assays

M1 RNA was folded as follows: the RNA was denatured by incubation at 50oC for

50 minutes, allowed to fold for 10 min at 37oC in water, and then allowed to fold further in binding buffer (100 mM Tris-HCl pH 7.5 or 20 mM HEPES-KOH pH 7.5, 400 mM

NH4OAc, 10 mM Mg(OAc)2, 5% (v/v) glycerol, and 0.1% (v/v) IGEPAL (NP-40)) for 30 minutes at 37oC. Folded RNA was diluted to 10 nM then stored as aliquots at -80oC. Eco 73

RNase P was reconstituted by combining M1 RNA (1 nM) and C5 protein (10 nM in all experiments save the comparison between radio-labeled substrate and fluor-labeled substrate, in which 100 nM was used for the reactions with the fluor-labeled substrate) and incubating in binding buffer for 30 to 60 minutes at 37oC. Assays were conducted in binding buffer with an RNase P concentration of 1 nM, and substrate concentration as indicated in the respective figure captions. Aliquots were removed at the desired time intervals and quenched with a solution of 10 M urea and 50 – 100 mM EDTA, or with 5

M urea. Aliquots corresponding to each time-point were then separated by denaturing

PAGE (see figure legends for additional details). After PAGE, assays using radio-labeled

RNA were exposed to a phosphorimaging screen which was then scanned using a

Typhoon Trio (GE Healthcare). Fluorescent gels were directly scanned using the same instrument. All images were quantitated using ImageQuant TL.

5.7 Assay separation and quantitation by laser-induced fluorescence-based automated DNA sequencer

Samples were prepared by drop dialysis against water, ethanol precipitation, or dilution. In drop dialysis, samples were placed atop a 0.25 µm membrane floating on distilled water. The samples were left unattended for 2 h, after which they were loaded on a microplate for injection. The sample volumes increased up to 10-fold in this 2-h period. As we did not directly assess salt concentrations, it is possible that this method primarily served as a dilution. For ethanol precipitation, the samples were precipitated

74 once in 300 mM NaOAc pH 5.2, then rinsed thrice with 1 mL 70% (v/v) ethanol to remove traces of salt. For dilution, the samples were diluted 3-fold, 10-fold, or 100-fold of their original quenched-volume with 5 M urea. The samples were injected at 1600 V for 15 s, and electrophoresed at 15,000 V using an Applied Biosystems Instruments 3730 automated DNA sequencer. The peaks were quantitated using Peak Scanner software

(Applied Biosystems).

5.8 – Assessment of AzG incorporation

A synthetic template coding for sequence 5′-GAAC-3′ was hybridized in 5x in vitro transcription buffer. The template sequence was as follows: noncoding - 5′-

TAATACGCCTCACTATAGAAC-3′, coding - 3′-ATTATGCGGAGTGATATCTTG-5′. The tetramer was transcribed using T7 RNA polymerase in the presence of 6 mM ATP, CTP, and UTP, 1.2 mM GTP, and 4.8 mM AzG, as well as [α-32P]ATP. The products were separated on a 15% polyacrylamide, 7 M urea gel, after which the gel was exposed to a phosphorimaging screen. The screen was scanned and the image quantitated using

ImageQuant TL.

Figure 22: Nujol mull FT-IR spectrum of 5′-iodo-5′-deoxyguanosine.

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Figure 23: 1H NMR spectrum of 5′-iodo-5′-deoxyguanosine.

Figure 24: Nujol mull FT-IR spectrum of 5′-azido-5′deoxyguanosine.

Note the strong 2100 cm-1 peak indicating the presence of the 5′-azide.

Figure 25: 1H NMR spectrum of 5′-azido-5′-deoxyguanosine.

Figure 26: Nujol mull FT-IR spectrum of 5(6)-propargylamidofluorescein.

A faint peak at 2125 cm-1 (not labeled) corresponds to the terminal alkyne C-C stretch. This peak was more clearly identified in a

neat spectrum.

Figure 27: 1H NMR spectrum of 5(6)-propargylamidofluorescein.

References

Altman, S.; Kirsebom, L.A. Ribonuclease P. The RNA World, 2nd edition, Ed. Gesteland,

R.F.; Cech, T.R.; Atkins, J.F. Cold Spring Harbor: Cold Spring Harbor Laboratory Press,

1999. 351 – 380

Angell, Y.M.; Garcia-Echeverria, C.; Rich, D.H. Comparative Studies of the Coupling of N-

Methylated, Sterically Hindered Amino Acids During Solid-Phase Peptide Synthesis.

Tetrahedron Lett. 1994, 35, 5981 – 5984

Brear, P.; Freeman, G.R.; Shankey, M.C.; Trmčić, M.; Hodgson, D.R.W. Aqueous methods for the preparation of 5′- substituted guanosine derivatives. Chem. Commun. 2009, 4980

– 4981

Bunner, A. E.; Nord, S.; Wikström, P.M.; Williamson, J.R. The Effect of Ribosome

Assembly Cofactors on In Vitro 30S Subunit Reconstitution. J. Mol. Biol. 2010, 398, 1 – 7

Burgin, A.B.; Pace, N.R. Mapping the active site of ribonuclease P RNA using a substrate containing a photoaffinity agent. EMBO J. 1990, 9, 4111 – 4118

Burrows, C.J.; Muller, J.G. Oxidative Nucleobase Transformations Leading to Strand

Scission. Chem. Rev. 1998, 98, 1109 – 1151

Chamberlain, J.R.; Lee, Y.; Lane, W.S.; Engelke, D.R. Purification and characterization of the nuclear RNase P holoenzyme complex reveals extensive overlap with RNase MRP.

Genes & Dev. 1998, 12, 1678 – 1690

Chan, B.; Weidemaier, K.; Yip, W.-T.; Barbara, P.F.; Musier-Forsyth, K. Intra-tRNA

Distance Measurements for Nucleocapsid Protein-Dependent tRNA Unwinding During

Priming of HIV Reverse Transcription. Proc. Natl. Acad. Sci. 1999, 96, 459 – 464

Chen, W.Y.; Singh, D.; Lai, L.B.; Stiffler, M.A.; Lai, H.D.; Foster, M.P.; Gopalan, V. Fidelity of tRNA 5′-maturation: a possible basis for the functional dependence of archaeal and eukaryal RNase P on multiple protein cofactors. Nucleic Acids Res. 2012, 40, 4666 – 4680

Cho, I-M.; Lai, L. B.; Susanti, D.; Mukhopadhyay, B.; Gopalan, V. Ribosomal protein L7Ae is a subunit of archaeal RNase P. Proc. Natl. Acad Sci. 2010, 107, 14573 – 14578

Daldrop, P.; Masquida, B.; Lilley, D.M.J. The functional exchangeability of pk- and k-turns in RNA structure. RNA Biol. 2013, 10, 445 – 452

Dean, D.K. An Improved Synthesis of 5′-amino-5′-deoxyguanosine. Synth. Commun.

2002, 32, 1517 – 1521

Dimitrijevich, S.D.; Verheyden, J.P.H.; Moffat, J.G. Halo Sugar Nucleosides. 6. Synthesis of Some 5′-deoxy-5′-iodo and 4′-5′-Unsaturated Purine Nucleosides. J. Org. Chem. 1979,

44, 400 – 406

Dirksen, A.; Dawson, P.E. Rapid Oxime and Hydrazone Ligations with Aromatic

Aldehydes for Biomolecular Labeling. Bioconjug. Chem. 2008 19, 2543 – 2548

Dovichi, N.J. DNA sequencing by capillary electrophoresis. Electrophoresis 1997, 18,

2393 – 2399

Eberhardt, M.K.; Colina, R. The Reaction of OH Radicals with Dimethyl Sulfoxide. A

Comparative Study of Fenton′s Reagent and the Radiolysis of Aqueous Dimethyl

Sulfoxide Solutions. J. Org. Chem. 1988, 53, 1071 – 1074

Eder, P.S.; Hatfield, C.; Vioque, A.; Gopalan, V. Bacterial RNase P as a potential target for novel anti-infectives. Curr. Opin. Investig. Drugs 2003, 4, 937 – 943

El-Sagheer, A.H.; Brown, T. Click chemistry with DNA. Chem. Soc. Rev. 2010, 39, 1388 –

1405

Fekner, T.; Li, X.; Lee, M.M.; Chan, M.K. A pyrrolysine analogue for protein click chemistry. Angew. Chem. Int. Ed. Engl. 2009, 48, 1633 – 1635

Fukuhara, H.; Kifusa, M.; Watanabe, M.; Terada, A.; Honda, T.; Numata, T.; Kakuta, Y.;

Kimura, M. A fifth protein subunit Ph1496p elevates the optimum temperature for the ribonuclease P activity from Pyrococcus horikoshii OT3. Biochem. Biophys. Res.

Commun. 2006, 343, 956 – 964

Gierlich, J.; Burley, G.A.; Gramlich, P.M.E; Hammond, D.M.; Carell, T. Click Chemistry as a

Reliable Method for the High-Density Postsynthetic Functionalization of Alkyne-

Modified DNA. Org. Lett. 2006, 8, 3639 – 3642

Gobert, A.; Gutman, B.; Taschner, A.; Gössringer, M.; Holzmann, J.; Hartmann, R.K.;

Rossmanith, W.; Giegé, P. A single Arabidopsis organellar protein has RNase P activity.

Nat. Struct. Mol. Biol. 2010, 17, 740 – 744

Göβringer, M.; Helmecke, D.; Hartmann, R.K. Characterization of RNase P RNA activity.

Methods Mol. Biol. 2012, 848, 61 – 72

Gopalan, V.; Baxevanis, A.D.; Landsman, D.; Altman, S. Analysis of the Functional Role of

Conserved Residues in the Protein Subunit of Ribonuclease P from Escherichia coli. J.

Mol. Biol. 1997, 267, 818 – 829

Gopalan, V.; Jovanovic, M.; Eder. P.S.; Giordano, T.; Powers, G.D.; Xavier, K.A. Bacterial

RNase P proteins and their use in identifying antibacterial compounds. U.S. Patent

6,936,432 assigned to Message Pharmaceuticals August 30, 2005

Gribbon, P.; Sewing, A. Fluorescence readouts in HTS: no gain without pain? Drug

Discov. Today 2003, 8, 1035 – 1043

Guerrier-Takada, C.; Gardiner, K.; Marsh, T.; Pace, N.; Altman, S. The RNA moiety of ribonuclease P is the catalytic subunit of the enzyme. Cell 1983, 35, 849 – 857

Gutmann, B.; Gobert, A.; Giegé, P. PRORP proteins support RNase P activity in both organelles and the nucleus in Arabidopsis. Genes Dev. 2012, 26, 1022 – 1027

Hong, V.; Presolski, S.; Ma, C. Analysis and optimization of copper-catalyzed azide- alkyne cycloaddition for bioconjugation. Angew. Chem. Int. Ed. 2009, 48, 9879 – 9883

Howard, M.J.; Lim, W.H.; Fierke, C.A.; Koutmos, M. Mitochondrial RNase P structure provides insight into the evolution of catalytic strategies for precursor-tRNA 5′ processing. Proc. Natl. Acad. Sci. 2012, 109, 16149 – 16154

Hsieh, J.; Fierke, C.A. Conformational change in the Bacillus subtilis RNase P holoenzyme-pre-tRNA complex enhances substrate affinity and limits cleavage rate.

RNA 2009, 15, 1565 – 1577

Hsieh, J.; Koutmou, K.S.; Rueda, D.; Koutmous, M.; Walter, N.G.; Fierke, C.A. A divalent cation stabilizes the active conformation of the B. subtilis RNase P-pre-tRNA complex: a role for an inner-sphere metal ion in RNase P. J. Mol. Biol. 2010, 400, 38 – 51

Huisgen, R. 1,3-Dipolar Cycloadditions. Proc. Chem. Soc. 1961, 357 - 396

Jao, C.Y.; Salic, A. Exploring RNA transcription and turnover in vivo by using click chemistry. Proc. Natl. Acad. Sci. 2008, 105, 15779 – 15784

Jarrous, N.; Gopalan, V. Archaeal/eukaryal RNase P: subunits, functions and RNA diversification. 2010 38, 7885 – 7894

Kalia, J.; Raines, R.T. Hydrolytic Stability of Hydrazones and Oximes. Angew. Chem. Int.

Ed. 2008, 47, 7523 – 7526

Kasai, H.; Kawai, K. DNA methylation at the C-5 position of cytosine by methyl radicals: a possible role for epigenetic change during carcinogenesis by environmental agents.

Chem. Res. Toxicol. 2009, 22, 984 – 989

Kawai, K.; Li, Y.S.; Song, M.F.; Kasai, H. DNA methylation by dimethyl sulfoxide and methionine sulfoxide triggered by hydroxyl radical and implications for epigenetic modifications. Bioorg. Med. Chem. Lett. 2010, 20, 260 – 265

Kikovska, E.; Brӓnnnvall, M.; Kufel, J.; Kirsebom, L.A. Substrate discrimination in RNase P

RNA-mediated cleavage: importance of the structural environment of the RNase P cleavage site. Nucleic Acids Res. 2005, 33, 2012 – 2021

Kladwang, W.; Hum, J.; Das, R. Ultraviolet Shadowing of RNA Can Cause Significant

Chemical Damage in Seconds. Sci. Rep. 2012, 2, 1 – 7

Kolb, H.C.; Finn, M.G.; Sharpless, K.B. “Click” Chemistry: Olefins, spring-loaded electrophiles, and heteroatom connections are key elements in a fast, modular, process- driven approach to molecular discovery. Angew. Chem. Int. Ed. 2001, 40, 2004 – 2021

Kolb, H.C.; Sharpless, K.B. The growing impact of click chemistry on drug discovery.

Drug Discov. Today 2003, 8, 1128 – 1137

Kole, R.; Altman, S. Properties of Purified Ribonuclease P from Escherichia coli.

Biochemistry 1981, 20, 1902 – 1906

Kundu, L.M.; Linne, U.; Marahiel, M.; Carell, T. RNA Is More UV Resistant than DNA: The

Formation of UV-Induced DNA Lesions is Strongly Sequence and Conformation

Dependent. Chem. Eur. J. 2004, 10, 5697 – 5705

Lai, L. B.; Vioque, A.; Kirsebom, L. A.; Gopalan, V. Unexpected diversity of RNase P, an ancient tRNA processing enzyme: challenges and prospects. FEBS Lett. 2010 584, 287 –

296

fMet Lazard, M.; Meinnel, T. Role of base G-2 of pre-tRNA in cleavage site selection by

Escherichia coli RNase P in vitro. Biochemistry 1998, 37, 6041 – 6049

Lea, W.A.; Simeonov, A. Fluorescence Polarization Assays in Small Molecule Screening.

Expert Opin. Drug Discov. 2011, 6, 17 – 32

Lehmann, J.; Jossinet, F.; Gautheret, D. A universal RNA structural motif docking the elbow of tRNA in the ribosome, RNAse P and T-box leaders. Nucleic Acids Res. 2013, 41,

5494 – 5502

Liu, X. Molecular Recognition of Inhibitors, Metal Ions and Substrates by Ribonuclease P.

PhD Dissertation 2013, University of Michigan, Ann Arbor.

Lillo, M.P.; Szpikowska, B.K.; Mas, M.T.; Sutin, J.D.; Beechem, J.M. Real-Time

Measurement of Multiple Intramolecular Distances during Protein Folding Reactions: A

Multisite Stopped-Flow Fluorescence Energy-Transfer Study of Yeast Phosphoglycerate

Kinase. Biochemistry 1997, 36, 11273 – 11281

Mangiarotti, G.; Chiaberge, S. Reconstitution of Functional Eukaryotic Ribosomes from

Dictyostelium discoideum Ribosomal Proteins and RNA. J. Biol. Chem. 1997, 272, 19682

– 19687

Martin, C.T.; Coleman, J.E. Kinetic Analysis of T7 RNA Polymerase-Promoter Interactions with Small Synthetic Promoters. Biochemistry 1987, 26, 2690 – 2696

Martin, C.T.; Coleman, J.E. T7 RNA Polymerase Does Not Interact with the 5′-Phosphate of the Initiating Nucleotide. 1989, 28, 2760 – 2762

McClain, W. H.; Lai, L. B.; Gopalan, V. Trials, travails and triumphs: an account of RNA catalysis in RNase P. J. Mol. Biol. 2010, 397, 627 – 646

McGee D.P.C.; Martin, J.C. Acyclic nucleoside analogues: methods for the preparation of 2′,3′-secoguanosine, 5′-deoxy-2′-3′-secoguanosine, and (R,S)-9-[1-(2-hydroxyethoxy)-

2-hydroxyethyl]guanine. Can. J. Chem. 1986, 64, 1885 – 1889

Meyer, M.; Westhof, E.; Masquida, B. A structural module in RNase P expands the variety of RNA kinks. RNA Biol. 2012, 9:3, 254 – 260

Mitsunobu, O. The Use of Diethyl Azodicarboxylate and Triphenylphosphine in the

Synthesis and Transformation of Natural Products. Synthesis 1981, 1 – 28

Nakagawa, I.; Aki, K.; Hata, T. Synthesis of 5′-Alkylthio-5′-deoxyucleosides from

Nucleosides in a One-pot Reaction. J. Chem. Soc. Perkin Trans. 1983, 1, 1315 – 1318

Nassar, A.-E.F.; Parmentier, Y.; Martinet, M.; Lee, D.Y. Liquid Chromatography-Accurate

Radioisotope Counting and Microplate Scintillation Counter Technologies in Drug

Metabolism Studies. J. Chromatogr. Sci. 2004, 42, 348 – 353

Neves, A. A.; Stöckmann, H.; Wainman, Y.A.; Kuo, J.C-H.; Fawcett, S.; Leeper, F.J.;

Brindle, K.M. Imaging Cell Surface Glycosylation in Vivo using “Double Click” Chemistry.

Bioconj. Chem. 2013, 24, 934 – 941

Oh, B.-K.; Pace, N.R. Interaction of the 3′-end of tRNA with ribonuclease P RNA. Nucleic

Acids Res. 1994, 22, 4087 – 4094

Oikawa, S.; Kawanishi, S. Copper-mediated DNA damage by metabolites of p- dichlorobenzene. Carcinogenesis 1996, 17, 2733 – 2739

Orr, C.W.M. Studies on Ascorbic Acid. II. Physical Changes in Catalase Following

Incubation with Ascorbate or Ascorbate and Copper(II). Biochemistry 1967, 6, 3000 –

3006

Paredes, E.; Das, S.R. Click Chemistry for Rapid Labeling and Ligation of RNA.

ChemBioChem 2011, 12, 125 – 131

Paredes, E.; Das, S.R. Optimization of acetonitrile co-solvent and copper stoichiometry for pseudo-ligandless click chemistry with nucleic acids. Bioorg. Med. Chem. Lett. 2012,

22, 5313 – 5316

Pogozelski, W.K.; Tullius, T.D. Oxidative Strand Scission of Nucleic Acids: Routes Initiated by Hydrogen Abstraction from the Sugar Moiety. Chem. Rev. 1998, 98, 1089 – 1107

Presolski, S.I.; Hong, V.; Cho, S-H.; Finn, M.G. Tailored Ligand Acceleration of the Cu-

Catalyzed Azide-Alkyne Cycloaddition Reaction: Practical and Mechanistic Implications.

J. Am. Chem. Soc. 2010 132, 14570 – 14576

Proudnikov, D.; Mirzabekov, A. Chemical Methods of DNA and RNA Fluorescent

Labeling. Nucleic Acids Res. 1996, 24, 4535 – 4542

Reich, C. Olsen, G.J. Pace, B.; Pace, N.R. Role of the protein moiety of ribonuclease P, a ribonucleoprotein enzyme. Science 1998, 239, 178 – 181

Reiter, N. J.; Osterman, A.; Torres-Larios, A.; Swinger, K.K.; Pan, T.; Mondragon, A.

Structure of a bacterial ribonuclease P holoenzyme in complex with tRNA. Nature 2010,

468, 784 – 789

Repine, J.E.; Pfenninger, O.W.; Talmage, D.W.; Berger, E.M.; Pettijohn, D.E. Dimethyl sulfoxide prevents DNA nicking mediated by ionizing radiation or iron/hydrogen perioxide-generated hydroxyl radical. Proc. Natl. Acad. Sci. 1981, 78, 1001 – 1003

Roper, K.A.; Lange, H.; Polyzos, A.; Berry, M.B.; Baxendale, I.R.; Ley, S.V. The application of a monolithic triphenylphosphine reagent for conducting Appel reactions in flow microreactors. Beilstein J. Org. Chem. 2011, 7, 1648 – 1655

Rostovtsev, V.V.; Green, L.G.; Fokin, V.V.; Sharpless, K.B. A Stepwise Huisgen

Cycloaddition Process: Copper(I)-Catalyzed Regioselective “Ligation” of Azides and

Terminal Alkynes. Angew. Chem Int. Ed. 2002, 41, 2596 – 2599

Rueda, D.; Hsieh, J.; Day-Storms, J.J.; Fierke, C.A.; Walter, N.G. The 5′-Leader of

Precursor tRNAAsp Bound to the Bacilus subtilis RNase P Holoenzyme Has an Extended

Conformation. Biochemistry 2005, 44, 16130 – 16139

Slade, P.G.; Hajidandi, M.; Bartel, C.M.; Gorfien, S.F. Identifying the CHO secretome using mucin-type O-linked glycosylation and click-chemistry. J. Proteome Res. 2012, 11,

6175 – 6186

Sletten, E.M.; Bertozzi, C.R. From Mechanism to Mouse: A Tale of Two Bioorthogonal

Reactions. Acc. Chem Res. 2011, 44, 666 – 676

Taschner, A.; Weber, C.; Buzet, A.; Hartmann, R.K.; Hartig, A.; Rossmanith, W. Nuclear

RNase P of Trypanosoma brucei: A Single Protein in Place of the Multicomponent RNA-

Protein Complex. Cell. Rep. 2012, 2, 19 – 25

Thiele, C.; Papan, C.; Hoelper, D.; Kusserow, K.; Gaebler, A.; Schoene, M.; Piotrowitz, K.;

Lohmann, D.; Spandl, J.; Stevanovic, A.; Shevchenko, A.; Kuerschner, L. Tracing fatty acid metabolism by click chemistry. ACS Chem. Biol. 2012, 7, 2004 – 2011

Thirumurugan, P.; Matosiuk, D.; Jozwiak, K. Click Chemistry for Drug Development and

Diverse Chemical-Biology Applications. Chem. Rev. 2013, 113, 4905 – 4979

Thorp, H.H. The importance of being r: greater oxidative stability of RNA compared with

DNA. Chem. Biol. 2000, 7, R33 – R36

Tornøe, C.W.; Christensen, C.; Meldal, M. Peptidotriazoles on Solid Phase: [1,2,3]-

Triazoles by Regiospecific Copper(I)-Catalyzed 1,3-Dipolar Cycloadditions of Terminal

Alkynes to Azides. J. Org. Chem. 2002, 67, 3057 – 3064

Traub, P.; Nomura, M. Structure and function of E. coli ribosomes. V. Reconstitution of fully active 30S ribosomal particles from RNA and proteins. Proc. Natl. Acad. Sci. 1968,

59, 777 – 784

Tsai, H-Y.; Lai, L.B.; Gopalan, V. A Modified pBluescript-Based Vector for Facile Cloning and Transcription of RNAs. Anal. Biochem. 2002, 303, 214 – 217

Tsai H.Y.; Pulukkunat D.K.; Woznick W.K.; Gopalan, V. Functional reconstitution and characterization of Pyrococcus furiosus RNase P. Proc. Natl. Acad. Sci. 2006, 103, 16147-

16152

Ueno, Y.; Jiao, G-S.; Burgess, K. Preparation of 5- and 6-carboxyfluorescein. Synthesis

2004, 15, 2591 – 2593

Vioque, A. Protein synthesis inhibitors and catalytic RNA. Effect of puromycin on tRNA precursor processing by the RNA component of Escherichia coli RNase P. FEBS Lett.

1989, 246, 137 – 139

Walker, S.C.; Engelke, D.R. A Protein-Only RNase P in Human Mitochondria. Cell 2008,

3, 412 – 414

Walter, M.V.; Malkoch, M. Simplifying the synthesis of dendrimers: accelerated approaches. Chem. Soc. Rev. 2012, 41, 4593 – 4609

Wegscheid, B.; Hartmann, R.K. The precursor tRNA 3′-CCA interaction with Escherichia coli RNase P RNA is essential for catalysis by RNase P in vivo. RNA 2006, 12, 2135 – 2148

Wegscheid, B.; Hartmann, R.K. In vivo and in vitro investigation of bacterial type B

RNase P interaction with tRNA 3′-CCA. Nucleic Acids Res. 2007, 35, 2060 – 2073

Weinberger, R. Practical Capillary Electrophoresis, 2nd Edition, San Diego: Academic

Press, 2000.

Williamson, D. The enzymatic synthesis and bioconjugation of 5′-AzaRNAs. Durham theses 2007, Durham University. Available at Durham E-Theses Online: http://etheses.dur.ac.uk/2560/

Williamson, D.; Cann, M.J.; Hodgson, D.R.W. Synthesis of 5′-amino-5′-deoxyguanosine-

5′-N-phosphoramidate and its enzymatic incorporation at the 5′-termini of RNA molecules. Chem. Commun. 2007, 47, 5096 – 5098

Willkomm, D.K.; Hartmann, R.K. An Important piece of the RNase P jigsaw puzzle solved. Trends Biochem. Sci. 2007, 6, 247 – 250

Willkomm, D.K.; Pfeffer, P.; Reuter, K.; Klebe, G.; Hartmann, R.K. RNase P as a Drug

Target. Protein Reviews, 2010, 10, 235 – 256

Winz, M.L.; Samanta, A.; Benzinger, D.; Jӓschke, A. Site-specific terminal and internal labeling of RNA by poly(A) polymerase tailing and copper-catalyzed or copper-free strain-promoted click chemistry. Nucleic Acids Res. 2012 40, e78

Wu, C-W.; Eder, P.S.; Gopalan, V.; Behrman, E.J. Kinetics of coupling reactions that generate monothiophosphate disulfides implications for modification of RNAs.

Bioconjug. Chem. 2001, 12, 842 – 844

Xiao, S.; Scott, F.; Fierke, C.A.; Engelke, D.R. Eukaryotic Ribonuclease P : A Plurality of

Ribonucleoprotein Enzymes. Annu. Rev. Biochem. 2002, 71, 165 – 189

Xu, Y.; Suzuki, Y.; Komiyama, M. Click Chemistry for the Identification of G-Quadruplex

Structures: Discovery of a DNA-RNA G-Quadruplex. Angew. Chem. Int. Ed. 2009, 48

3281 – 3284

Yandek, L.E.; Lin, H.C.; Harris, M.E. Alternative substrate kinetics of Escherichia coli ribonuclease P: determination of relative rate constants by internal competition. J. Biol.

Chem. 2013, 288, 8342 – 8354

Zeglis, B.M.; Sevak, K.K.; Reiner, T.; Mohindra, P.; Carlin, S.D.; Zanzonico, P.; Weissleder,

R.; Lewis, J.S. A Pretargeted PET Imaging Strategy Based on Bioorthogonal Diels-Alder

Click Chemistry. J. Nucl. Med. 2013, 54, 1 – 8

Zhang, L.; Sun, L.; Cui, Z.; Gotlieb, R.L.; Zhang, B. 5′-Sulfhydryl-modified RNA: initiators synthesis, in vitro transcription and enzymatic conjugation. Bioconjug. Chem. 2001, 3,

275 – 278

100