Trna Subcellular Dynamics Dictates Modification and Nutrient Sensing

tRNA subcellular dynamics dictates modification and nutrient sensing

DISSERTATION

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

By Alan Christopher Kessler Graduate Program in Microbiology

The Ohio State University 2018

Dissertation Committee: Dr. Juan D. Alfonzo, Advisor Dr. Jane E. Jackman Dr. Charles Daniels Dr. Patrice Hamel

Alan C. Kessler

2018

Abstract

In all eukaryotes, tRNAs are transcribed in the nucleus and then exported to the cytoplasm to engage in protein synthesis. However, previous work in Saccharomyces cerevisiae showed that tRNAs can also be sent back to the nucleus, and intracellular transport can be altered in response to starvation, leading to nuclear accumulation of tRNA. At least in one case, retrograde nuclear transport from the cytoplasm is necessary for wybutosine formation in tRNAPhe.

Despite the fact that retrograde transport has been firmly established in yeast, it has been difficult to assess whether such a mechanism has a broader evolutionary distribution in eukaryotes.

In the first part of this dissertation, I examined the post-transcriptional modification Queuosine (Q), and its relationship to retrograde transport in

Trypanosoma brucei. Q is found at the first position of the anticodon in several tRNAs (tRNATyr, tRNAAsp, tRNAAsn and tRNAHis) and is presumably important for protein synthesis, although its function is not yet fully understood. Eukaryotes cannot synthesize Q and must rely on uptake and salvage of the free base from either nutrients or from gut microbiota. Following uptake, the enzyme tRNA guanine-transglycosylase (TGT) is responsible for the incorporation of queuine into tRNA by replacing guanine. In this work I show that T. brucei’s TGT (TbTGT) is a nuclear enzyme essential for Q formation in tRNA. One of its natural substrates, tRNATyr, also contains an intron, which must be removed prior to Q formation. In the present work, I show that because essential components of the

ii splicing machinery are cytoplasmic, there is a dynamic interplay between tRNA splicing and modifications, all driven by the intracellular distribution of the different maturation components. Taken together, I demonstrate the existence of a tRNA nuclear retrograde transport pathway in T. brucei akin to what has been described in yeast, but with implications for other eukaryotic systems.

The latter half of the dissertation examines Q and its potential involvement in nutrient sensing in T. brucei. As T. brucei transitions from the procyclic insect stage, to the mammalian bloodstream stage, metabolic reprograming occurs, with concomitant changes in the expression of stage specific genes. Additionally, in T. brucei gene regulation occurs post-transcriptionally. Because of this, post- transcriptional modifications may play critical roles in regulating gene expression.

Here I show that Q modification levels change between the procyclic and bloodstream developmental stages of the parasite. Q levels also fluctuate in response to changes in the availability of a subset of amino acids. These findings have implications for how organisms may use modifications to sense nutrient availability and adjust translational rates accordingly.

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Dedication

I would like to dedicate this work to my loving mother Judy Kessler. Without her

guidance and unwavering support, none of this would have been possible.

Acknowledgments

It would be impossible to acknowledge everyone who had a positive impact on my professional development during graduate school. First off, I would like to thank Dr. Juan Alfonzo and Dr. Mary Anne Rubio. They have both created an environment that has been challenging yet rewarding in all its aspects. From the interesting RNA topics, to the numerous political conversations we have had, it has all been intellectual and enjoyable. I appreciate all the jokes over the years which made failed experiments and mistakes seem light hearted and easier to deal with.

The lab could not be described without the different friends that I have grown accustomed to over the years. One of my best friends in the lab, Katie

McKenney, has been there for me throughout all the ups and downs experienced throughout graduate school. I appreciate all the moments she helped me with experiments, as well as her unwavering positive outlook. I would next like to thank

Dr. Ian Fleming for his guidance and friendship. I will never be able to “pay the bills” as he once put it, but I am thankful for all his assistance in the lab and for

“voting to allow me to say on that side of the lab.” There were also many others in the lab over the years which I have fond memories with including Dr. Zdeněk Paris,

Dr. Paul Sample, Dr. Raphael Sores, Scott Hinger, Caitlin Moore, Gabriel d’Almeida.

None of this would have been possible without the love and support that my family and friends have given me over the years. My mother Judy Kessler, has always encouraged me to find the right path in my life. She has been full of advice v and above all, helped me succeed in nearly every way imaginable. I was fortunate to have Michelle Gibbs always around through most of my graduate schooling.

She has been caring, thoughtful, and made the experience more enjoyable. I would also like to mention my late father Christopher Kessler who has been a major source of inspiration throughout my life and late grandmother Adeline Kessler, who always encouraged me to never stop learning.

Vita

2008 ...... Kane Area High School 2012 ...... B.S. Biology, Indiana University of PA 2012 to present ...... Graduate Teaching/Research Associate, Department of Microbiology, The Ohio State University

Publications

Kessler AC, Kulkarni SS, Paulines MJ, Rubio MAT, Limbach PA, Paris Z, Alfonzo JD. Retrograde nuclear transport from the cytoplasm is required for tRNATyr maturation in T. brucei. RNA Biology. 2017. Sep 13:1-9. ahead of print.

Kessler AC, d’Almeida GS, Alfonzo JD. The role of intracellular compartmentalization on tRNA processing and modification. RNA Biology. 2017. Aug 29:1-13. ahead of print.

Lopes RR, Silveria Gde O, Eitler R, Vidal RS, Kessler A, Hinger S, Paris Z, Alfonzo JD, Polycarpo C. The essential function of the trypanosoma brucei Trl1 homolog in procyclic cells is maturation of the intron-containing tRNATyr. RNA. 2016. 22(8):1190-9.

Lopes RR, Kessler AC, Polycarpo C, Alfonzo JD. Cutting, dicing, healing and sealing: The molecular surgery of tRNA. Wiley Interdiscip. Rev. RNA. 2015. 6(3):337-49.

Fields of Study

Major Field: Microbiology

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Table of Contents

Abstract ...... ii

Acknowledgments ...... v

Vita ...... vii

Publications ...... vii

Fields of Study ...... vii

Table of Contents ...... viii

List of Tables ...... xii

List of Figures ...... xii

Chapter 1 : Introduction ...... 1

1.1 A brief Introduction to Trypanosoma brucei ...... 1

1.2 Gene regulation in T. brucei ...... 4

1.3 Transfer RNA ...... 6

1.4 Role of compartmentalization on tRNA processing and modification ...... 10

1.4.1 Functionality checkpoints: Early steps in tRNA maturation in eukaryotes

...... 16

1.4.2 The cytosolic fate of tRNA modifications ...... 22

1.4.3 The connection between tRNA splicing and modifications ...... 25

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1.4.4 Intracellular transport dynamics that set the order of modifications .... 31

1.4.5 Modifications and the Mitochondria ...... 39

1.5 The Queuosine Modification ...... 46

1.5.1 The Queuosine Pathways of Bacteria and Eukarya ...... 47

1.5.2 TGT in Bacteria and Eukarya ...... 50

1.5.3 The Biological Role of Q ...... 52

Chapter 2 : Retrograde transport is required for tRNA maturation in T. brucei ...... 56

2.1 Introduction ...... 57

2.2 Results ...... 60

2.2.1 T. brucei encodes a TGT homolog ...... 60

2.2.2 TGT localizes to the nucleus ...... 66

2.2.3 Intron containing tRNATyr does not contain Queuosine ...... 68

2.2.4 Retrograde nuclear import is necessary for queuosine formation in

tRNATyr ...... 71

2.3 Discussion ...... 76

Chapter 3 : The sensing of amino acids impacts the tRNA modification ...... 80

3.1 Introduction ...... 80

3.2 Results ...... 85

3.2.1 The Steady-state levels of Queuosine differs between developmental

stages ...... 85

3.2.2 Both Q-containing and non-Q-containing tRNAs partake in translation

...... 89

3.2.3 Changes in glucose levels and other general nutrients do not affect Q

levels ...... 91

3.2.4 The levels of Queuosine change in response to specific amino acids 94

3.3 Discussion ...... 106

Chapter 4 : Concluding remarks and future directions ...... 111

References ...... 119

Appendix A: Materials and Methods ...... 140

A1: RNA purification by guanidine isothiocyanate ...... 141

A2: RNA purification by acid phenol ...... 142

A3: Standard polyacrylamide gel electrophoresis for RNA separation ...... 143

A4: Acid gel electrophoresis for RNA separation ...... 144

A5: APB gel electrophoresis for RNA separation ...... 145

A6: SDS Polyacrylamide gel electrophoresis for protein analysis ...... 147

A7: End labeling of northern blot probes ...... 149

A8: Northern blot utilizing electrotransfer ...... 150

A9: Western blot, utilizing electrotransfer ...... 153

A10: Nuclear and cytoplasmic fractionation ...... 154

A11: Immunofluorescent microscopy ...... 156

A12: Polysome analysis ...... 158

A13: Reverse Transcription-PCR ...... 160

A14: Preparing tRNA for nucleoside analysis: HPLC and/or LC-MS/MS ...... 162

A15: DNA purification from T. brucei ...... 164

A16: Plasmid DNA purification from E. coli ...... 165

A17: Electroporation of T. brucei ...... 166

A18: Preparing competent DH5α and transformation ...... 168

Appendix B: SDM-79 growth and depletion media used ...... 170

B1: Standard SDM-79: ...... 171

B2: Nutrient depleted SDM-79 ...... 176

Appendix C: Supporting figures ...... 179

List of Figures

Figure 1.1: The lifecycle of T. brucei ...... 3

Figure 1.2: Common tRNA depiction ...... 7

Figure 1.3: Universal genetic code ...... 9

Figure 1.4:Tertiary contacts necessary for tRNA structure ...... 11

Figure 1.5:Two major surveillance pathways for defective tRNAs ...... 19

Figure 1.6: Dependence of modification on introns ...... 29

Figure 1.7: Yeast yW biosynthesis pathway ...... 34

Figure 1.8: The biosynthesis of Q in T. brucei ...... 38

Figure 1.9: Mitochondrial import in L. tarentolae and T. brucei use thiolation ..... 45

Figure 1.10: Pathways of Queuosine synthesis ...... 49

Figure 2.1: Queuosine containing tRNA in T. brucei ...... 62

Figure 2.2: TbTGT is necessary for Queuosine ...... 64

Figure 2.3: TbTGT is a nuclear enzyme ...... 67

Figure 2.4: Intron-containing tRNA lack Queuosine ...... 70

Figure 2.5: The nucleus contains significant amounts of spliced tRNA Tyr ...... 73

Figure 2.6: Neither lack of queuosine nor aminoacylation are signals for retrograde transport...... 75

Figure 2.7: Nuclear retrograde transport is necessary for queuosine formation in tRNATyr ...... 79

Figure 3.1:Queuosine modification levels change depending on stage ...... 88

Figure 3.2: Q containing tRNA are used during translation ...... 90 xii

Figure 3.3: Queuosine does not change during general nutrient deprivation ..... 93

Figure 3.4:Cell growth is uninhibited in media reduced for a single amino acid . 96

Figure 3.5: Reduction of specific amino acids causes Q-tRNA increase ...... 98

Figure 3.6: The percent of Q-tRNA decreases with additional amino acids ...... 99

Figure 3.7: The Queuosine level is dependent on amino acids and time ...... 101

Figure 3.8: Polysomes are not affected by the loss of Q-tRNA ...... 104

Figure 3.9: Cells are actively translating when grown in low tyrosine media .... 105

Figure C.1: TbTrl1 knockdown causes intron-containing tRNATyr accumulation180

Figure C.2: Intron-containing tRNA are in the cytoplasm ...... 181

Figure C.3: Additional controls for Figure 2.6 ...... 182

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List of Tables

Table 1: The intracellular localization of a representative set of modification ..... 14

Table 2: Single amino acid reductions cause minor differences in doubling rate 97

Table 3: The antibiotics utilized in T. brucei media ...... 172

Table 4: SDM-79 media composition ...... 173

Table 5: Additional components of SDM-79 ...... 174

Table 6: The common components to all depleted media conditions ...... 177

Table 7: Amino acid concentration of SDM-79 and depleted SDM-79 ...... 178

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Chapter 1 : Introduction

The focus of my research is the study of tRNA nucleotide modification using the model organism Trypanosoma brucei. We seek to understand intricacies of the unique molecular biology of T. brucei. Our focus on modification leads us down many paths, striving to build a detailed story about tRNA, their specific modifications, and their function. This dissertation will attempt to shed light on the modified nucleotide Queuosine, and how it relates to tRNA intracellular dynamics and metabolic sensing in T. brucei.

1.1 A brief Introduction to Trypanosoma brucei

Trypanosoma brucei is a single celled parasitic protozoan, endemic to regions of Africa, and is the causative agent for the disease Human African

Trypanosomiasis (HAT), commonly referred to as African sleeping sickness. It resides in its Tsetse fly vector, through which it is transferred to a human host while the insect takes a bloodmeal, this stage of the parasite is known as the bloodstream form (BF). The life cycle is completed when a second Tsetse fly takes a bloodmeal from the infected host, at which point T. brucei travels to the midgut of the fly where it develops into the procyclic form (PF), the main focus of my work.

This is followed by the eventual migration to the salivary gland of the fly where it awaits transfer back to the mammalian host (Figure 1.1) (Smith et al. 2017).

As T. brucei moves from the midgut to the salivary glands of the Tsetse fly, it expresses a major surface membrane protein called procyclin (Rudenko 2011).

Once it enters the blood stream however, T. brucei stops expression of procyclin and instead switches to express the variant surface glycoproteins or VSGs, creating a dense outer layer (Horn 2014; Vanhamme et al. 2001). VSGs provide an expansive antigen repertoire that allows T. brucei to evade the immune system, but this will not be a major focus of this dissertation.

Figure 1.1: The lifecycle of T. brucei T. brucei is transmitted to the bloodstream by a Tsetse fly harboring the parasite during a bloodmeal. The bloodstream form’s (BF) hallmark is VSG switching, allowing for immune evasion. Traveling through the bloodstream, T. brucei is eventually ingested by a Tsetse fly, marking the beginning of the procyclic form

(PF). As T. brucei travels through the midgut to the salivary gland, it undergoes various morphological changes. Differing from the BF, the PF form mainly expresses the surface coat protein procyclin. From the salivary gland, T. brucei will be transmitted to the next host when the Tsetse fly takes a bloodmeal.

1.2 Gene regulation in T. brucei

T. brucei has also been at the forefront of many discoveries in molecular biology, such as the massive RNA editing events that take place in their mitochondria (Benne et al. 1986; Feagin et al. 1988; Feagin et al. 1987; Shaw et al. 1988). Belonging to the class Kinetoplastida, trypanosomatids are defined by a single mitochondrion, which stretches from the anterior to posterior end of the cell and contains mitochondrial DNA termed kinetoplast DNA or kDNA (Verner et al.

2015). The kDNA is a complex catenated network of DNA composed of maxicircles and minicircles. Maxicircle DNA is found at roughly 40 copies per cell with an approximate size of 30 kb; maxicircles are analogous to the conventional DNA of eukaryotic mitochondria encoding various mitochondrial proteins and ribosomal

RNA (rRNA). Minicircles are found at several thousand copies per mitochondrion and encoding short RNAs of 50 nucleotides average length; these serve as a template which “guide” the extensive nucleotide insertion/deletion editing that takes place in the mitochondrion (Corell et al. 1993; Simpson 1997; Hong and

Simpson 2003).

Throughout its lifecycle, transitioning from insect vector to human host, T. brucei undergoes vast metabolic changes. For example, the mitochondria of T. brucei is used for oxidative phosphorylation in the procyclic insect stage of its lifecycle but enters a reduced functional state in the bloodstream form where the

4 parasite switches to substrate level phosphorylation (Clayton and Michels 1996;

Bochud-Allemann and Schneider 2002).

Traditionally, the bulk of the regulation of gene expression in cells is thought to occur at the transcriptional level. Typically, in eukaryotes mRNAs are transcribed by RNA polymerase II, which requires specific promoters found upstream of genes and also requires a myriad of transcription factors which enhance or repress transcription of individual genes. In T. brucei however, there are no identifiable Pol II promoters and protein-coding genes are transcribed as polycistronic transcripts from which monocistronic RNAs are excised by the process of trans-splicing and poly-adenylation. Trans-splicing involves the attachment of a separately transcribed RNA, known as the spliced leader, to the

5’ end of the nascent mRNA, also providing the cap structure typically found at the

5’ end of mRNAs, while a poly A tail is also simultaneously added to the 3’ end

(Perry et al. 1987).

Due to their prominent lack of transcriptional regulation, T. brucei relies on post-transcriptional mechanisms to regulate gene expression. For example, the stage dependent differential expression of phosphoglycerate kinase is controlled at the level of transcript degradation (Haanstra et al. 2008). We believe that post- transcriptional modifications of tRNA may serve an important role as part of additional regulatory mechanisms that regulate gene expression.

1.3 Transfer RNA

tRNAs are adaptor molecules which allow the transfer of information from mRNA into that ultimately found in proteins; tRNA deliver specific amino acids to the growing polypeptide chain in the ribosome during translation. All tRNAs maintain a three-dimensional L shape fold (Figure 1.2), a structure important for ribosome interaction. Each mature tRNA is approximately 76 nucleotides in length with the canonical CCA sequence either encoded in the tRNA gene or added post- transcriptionally to the 3’ terminus. Finally, one of the twenty standard amino acids is added to its respective tRNA by a specific aminoacyl tRNA synthetases (aaRS), which recognize their tRNAs via identity elements or recognition points.

tRNA are often depicted in their two-dimensional cloverleaf structure highlighting the four paired stems of a tRNA. The D stem and TΨC stem are named after their characteristic modifications Dihydrouridine in the former and 5- methyluridine (T) and pseudouridine (Ψ) in the latter. The acceptor stem brings together the 5’ and 3’ termini of the tRNA and creating the stem that following CCA addition ultimately serves as the amino acid acceptor end via the terminal 3’ or 2’ hydroxyls. The anticodon stem contains the anticodon sequence (positions 34, 35,

36) which directly interacts with codons in mRNA during translation.

Figure 1.2: Common tRNA depiction The 2D and 3D structures of the yeast tRNAPhe is shown. A. In the 2D cloverleaf scheme, the four arms Acceptor, D, TΨC, and Anticodon are outlined. In the

Anticodon Arm positions 34, 35, and 36 are indicated by X, consisting of the anticodon which directly interacts with the mRNA codon. In the acceptor stem, the

CCA denotes the location of amino acid attachment. B. The 3D structure of tRNAPhe is shown, outlining the L shape of the molecule. 3D structure PDB ID

4W2E (Gagnon et. al. 2014)

There are 64 possible codons to code for the 20-22 amino acids; three of these are typically used as stop codons (UGA, UAA, UAG) while the remaining 61 codons are used for protein synthesis (Figure 1.3). Because of redundancy in the genetic code, multiple codons can be used to encode a single amino acid. Along these lines, specific tRNAs are also capable of decoding multiple codons by

“wobbling,” whereby a near cognate codon-anticodon interaction can still be accurately read by the ribosomes. Because of tRNAs’ role in translation, post- transcriptional modifications, which heavily decorate tRNA, can directly influence decoding. This allows cells to use modification as another means of post- transcriptional regulation. To further enhance the ability of cells to regulate via modifications, many modification enzymes are localized to specific cellular compartments whereby successful modification is often dictated by tRNA intracellular transport dynamics (Chapter 1.4).

Figure 1.3: Universal genetic code A chart displaying each codon with its specific amino acid. Codons UAA, UAG, and UGA are typically reserved for stop codons.

1.4 Role of compartmentalization on tRNA processing and modification

Section 1.4 is published as: Kessler AC, Silveria d’Almeida G, Alfonzo JD. (2017), The role of intracellular compartmentalization on tRNA processing and modification. RNA Biol, Online ahead of print. Aug 29, 2017. Doi: 10.1080/15476286.2017.1371402

In eukaryotes, tRNAs are encoded in either the nucleus or one of the other genome-containing organelles; mitochondria, chloroplast or plastids, depending on the organism. Regardless of their site of synthesis, in all cases tRNAs are transcribed as premature molecules that cannot immediately partake in translation.

As tRNAs are transcribed, they begin to form local structures that may include initial folding of the arms; the D-arm, followed by the anticodon arm, and finally the

TΨC arm and the acceptor stem (Figure 1.4). Either during or soon after transcription, tRNAs also undergo various forms of “molecular pedicure” that trims their 5’ and 3’ ends, removes introns (if present) and adds their characteristic CCA tails, where eventually amino acids are attached. Concomitantly, the tRNA is also surveying various folding pathways and eventually assumes its canonical L-shape structure required for translation.

Figure 1.4:Tertiary contacts necessary for tRNA structure A. General tRNA cloverleaf structure highlighting the numbering scheme and important tertiary contacts between tRNA arms shown as dashed lines. Darker circles represent nucleotides, which when altered, disrupts the stability and export of the tRNA. Many nucleotide contacts that disrupt nuclear export are also those involved in inter-arm base pairing between the D and TΨC arms. B. As tRNA tertiary structure is disrupted, certain modifications will be negatively affected while others may not. In blue are positions in which corresponding modification enzymes can tolerate global tertiary structural changes while those shown in red do not tolerate tertiary alteration of the tRNA

At any point in this maturation pathway, post-transcriptional modifications may be added, and those that appear early tend to influence and enforce proper folding. Studies using microinjection of tRNAs into Xenopus oocytes indicated that certain modifications appear before end maturation, in a somewhat specific order.

For example, intron-containing pre-tRNATyr with immature 5’ leader and 3’ trailer sequences acquire pseudouridines (Ψ) in the anticodon and TΨC loop, as well as

5-methylcytidine (m5C) in the variable loop and 1-methyladenosine (m1A) in the

TΨC loop (Nishikura and De Robertis 1981; Johnson and Abelson 1983). As the

5’ and 3’ ends become matured, new modifications are added such as dihydrouridine (D) to the D-loop and additional pseudouridylations. Finally, after intron removal the remaining modifications are added to complete the required set for a particular tRNA (Nishikura and De Robertis 1981), but in eukaryotes intracellular compartmentalization further imposes order to tRNA maturation and to the occurrence of modifications (Table 1).

Beyond the nucleus, tRNAs also undergo further modification in the cytoplasm. Yet the story does not end there; many cellular RNAs, and indeed tRNAs, may travel back and forth across cellular membranes and each movement to a new locale offers the potential to be further modified. If one accepts that many modifications may alter the structure of tRNAs locally and sometimes globally, then what modification enzymes encounter is an ever-changing substrate structural landscape, which in turn may be recognized differently by different enzymes. For example, it is possible that intracellular transport dynamics may lead to the

12 appearance a hypomodified in a cellular compartment (i.e. nucleus or cytoplasm) where such tRNA may now be recognized by a different set of enzymes, which normally do not encounter such a substrate. This may lead to the given tRNA acquiring new modifications that it normally does not get. Taken together, this raises the exciting possibility that the diversity of modifications in tRNA sets varies greatly in the life span of a cell and may be influenced not just by environmental cues but also by transport dynamics. In the following pages, we highlight a number of recent examples of tRNA movements within cells and how intracellular distribution of tRNAs, because of transport, is intricately linked to many processing events and most certainly modifications. In passing, we will also discuss the fates of tRNAs that are recognized by cells as not fully modified and highlight the fact that such surveillance pathways are not so exacting; an observation that should not be surprising given the often subtle effects that modifications have in their substrate targets.

Table 1: The intracellular localization of representative modifications

Modification Enzyme(s) Localization Organism Found in unspliced tRNA Queuosine (Q) TGT Outer Mitochondrial COS-7 No Membrane (Boland et Monkey al. 2009) kidney Queuosine (Q) TGT Nucleus (Kessler et al. T. brucei No 2017) 1-methylguanosine Trm5 Nucleus(Ohira and S. cerevisiae No (m1G37) Suzuki 2011) Wybutosine (yW) TYW1, Cytoplasm/extra- S. cerevisiae No TYW2, nuclear (Noma et al. TYW3, 2006) TYW4 2-thiouridine 34 (S2U) Tuc1 Cytoplasm (Nakai et al. S. cerevisiae No (Ncs6), 2008) Tuc2 (Ncs2) 2-thiouridine 34 (S2U) Mtu1 Mitochondria (Umeda S. cerevisiae, No et al. 2005) Homo sapiens N2, N2-dimethylguanosine Trm1 Nucleus, Mitochondria S. cerevisiae Yes (Nishikura (m2,2G) (Li et al. 1989) and De Robertis 1981) N2-methylguanosine Trm11/Trm Cytoplasm S. cerevisiae No 2 (m G10) 112 (Purushothaman et al. 2005) 1-methyladenosine Gcd10/Gcd Nucleus(Anderson et S. cerevisiae Yes (Nishikura 1 (m A58) 14 al. 1998) and De Robertis 1981) Threonylcarbamoyladenos Sua5/KEO Cytoplasm(Thiaville et S. cerevisiae No ine (t6A) PS al. 2014) Threonylcarbamoyladenos Sua5/Qri7 Mitochondria(Thiaville S. cerevisiae No ine (t6A) et al. 2014) 3 3-methylcitidine (m C32) Trm140 Cytoplasm(Asakura et S. cerevisiae No al. 1998) 5 5-methylcytidine (m C40) Trm4 Nucleus(Wu et al. S. cerevisiae Yes (Masson et (NCL1) 1998) al. 1987) 5 5-methylcytidine (m C34) Trm4 Nucleus(Wu et al. S. cerevisiae Yes (Strobel (NCL1) 1998) and Abelson 1986) Pseudouridine (Ψ32) Pus8 Cytoplasm(Behm- S. cerevisiae No Ansmant et al. 2004) Pseudouridine (Ψ38 and Pus3 Nucleus, S. cerevisiae Yes (Nishikura Ψ39) (Deg1) Cytoplasm(Lecointe et and De al. 1998) Robertis 1981) Pseudouridine (Ψ34 and Pus1 Nucleus(Simos et al. S. cerevisiae Yes (Simos et Ψ36) 1996) al. 1996)

N6-isopentenyladenosine Mod5 Nucleus, Cytoplasm, S. cerevisiae No 6 (i A37) Mitochondria(Tolerico et al. 1999) Wyosine (imG) TYW1S,2,3 Mitochondria(Sample et T. brucei No B al. 2015) Inosine (I34) ADAT2/3 Cytoplasm(Gaston et T. brucei No al. 2007) 3 3-methylcytidine (m C32) ADAT2/3,T Nucleus(Rubio et al. T. brucei No RM140 2017) 3-methyluridine (m3U32) ADAT2/3, Nucleus(Rubio et al. T. brucei No Trm140 2017) 1-methylinosine (m1I37) Trm5 Cytoplasm, S. cerevisiae No Mitochondria(Lee et al. 2007) 5 5-formylcytidine (f C34) Nsun3 Mitochondria (Nakano Homo No et al. 2016) sapiens N2-methylguanosine Trm11/Trm Cytoplasm S. cerevisiae Yes (Nishikura 2 (m G10) 112 (Purushothaman et al. and De 2005) Robertis 1981) 5-carbamoylmethyl-2′-O- Trm7 Cytoplasm(Pintard et S. cerevisiae No methyluridine (ncm5Um) al. 2002) 5- Mss1 Mitochondria(Decoster S. cerevisiae No carboxymethylaminometh et al. 1993) 5 yluridine (cmnm U34)

Modification enzymes localize to various subcellular compartments. These

localizations often are not the same in distinct species. Whether a given

modification occurs in intron-containing tRNAs (unspliced tRNA) is as indicated.

1.4.1 Functionality checkpoints: Early steps in tRNA maturation in eukaryotes

With a starting point set by transcription, modification enzymes have been divided into two major groups based on the way they recognize their substrates: architecture-dependent enzymes requiring a fully folded tRNA for activity, and architecture-independent enzymes, which do not require a full-length tRNA for activity (Grosjean et al. 1996). The influence of tRNA structure on modifications has been examined by introducing a series of structural alterations to

Saccharomyces cerevisiae tRNAAsp followed by microinjection into Xenopus oocytes (Grosjean et al. 1996; Becker, Motorin, Sissler, et al. 1997). When mutations were introduced that changed the 3D structure of tRNAAsp, several

1 modifications such as 1-methylguanosine at position 37 (m G37), Ψ40 and Ψ13 were not formed, leading to the suggestion that the enzymes catalyzing such reactions recognize a fully folded tRNA and are therefore architecture-dependent. On the

5 other hand, global changes in tRNA structure had no major impact on m C49, 5-

5 methyluridine at position 54 (m U54) and Ψ55 (Figure 1.4 B). Unaffected modifications are thus catalyzed by enzymes that recognize local structure such as stem loops or specific sequences; these motifs become available shortly after transcription and before tRNA assumes its L-shape. These enzymes are therefore architecture independent. These, however, still recognize limited structural features; for example, for Ψ55 the presence of 4 G-C pairs in the TΨC loop, while a slightly longer stem was necessary for m5U formation (Becker et al. 1997;

McClain and Seidman 1975). Thus synthesis of modifications may be differently affected by changes to global vs. local structure of tRNA (Grosjean et al. 1996).

Defects in the addition of modifications can impact tRNA stability. An

1 interesting case occurs with m A58; a modification conserved in Bacteria, Archaea,

1 and Eukarya (Björk et al. 1987). In S. cerevisiae m A58 is found in many tRNAs,

Met but it is only essential for tRNAi . Mutation of the Trm6/Trm61 (GCD10/GCD14)

1 Met complex, the methylase responsible for m A58, results in increased tRNAi

Met instability and a lethal reduction in tRNAi pools; a phenotype easily overcome

Met by overexpression of tRNAi (Anderson et al. 1998; Anderson et al. 2000; Kadaba et al. 2004). Further analysis, based on genetic screening, indicated that RRP44

(DIS3), a 3’-5’ exoribonuclease exosome subunit, and TRF4, DNA polymerase

Met with poly(A) polymerase activity, had roles in the observed reduction of tRNAi .

Upon closer investigation, it was apparent that mutation of either TRF4 or RRP44

Met resulted in restoration of tRNAi to levels close to normal, supporting their

Met involvement in the reduction of tRNAi . Furthermore, deletion of the gene for the

Met nucleus-exclusive exosomal subunit RRP6 restored tRNAi levels while overexpression of Trf4 caused an exacerbated reduction (Kadaba et al. 2004).

Met Because of this, Trf4 was suggested to polyadenylate tRNAi , which would then serve as a signal for its targeted degradation by the nuclear exosome. Indeed, exosomal degradation of polyadenylated RNAs requires Trf4 polyadenylation activity (Egecioglu et al. 2006; Kadaba et al. 2006). The polyadenylation of tRNA and its subsequent exosomal degradation became known as the tRNA nuclear

17 surveillance pathway. However, this discovery did not initially address how the

Met hypomodified tRNAi was targeted, as Trf4 does not possess a recognizable

RNA binding domain. To address this, affinity purification of Trf4, and a two-hybrid analysis of the nuclear exosome cofactor Mtr4, led to the identification of the

TRAMP protein complex (LaCava et al. 2005; Vaňáčová et al. 2005). The TRAMP complex consists of three subunits: Trf4 or Trf5, Air1 or Air2, and Mtr4; When in a complex, Air1 or Air2 bind to their RNA target via their RNA binding domain. This is followed by polyadenylation by Trf4 or Trf5 and lastly, Mtr4 binds and facilitates unwinding of the RNA target via its helicase activity (Jia et al. 2012). Taken

1 Met together, the lack of m A58 destabilizes of tRNAi , which is then flagged for degradation by the activity of Trf4 and the TRAMP complex and later degraded by

1 the exosome (Figure 1.5 A). Although m A58 is the best known example of nuclear surveillance, other tRNAs are potentially monitored, since TRF4 deletion strains of yeast begin to accumulate precursor tRNAs in the nucleus (Copela et al. 2008).

Figure 1.5:Two major surveillance pathways for defective tRNAs

Met 1 A. tRNAi lacking m A58 is polyadenylated by the TRAMP complex (Trf4/5,

Air1/2). The helicase Mtr4 assists the TRAMP complex and targets TRAMP to the nuclear endonuclease complex consisting of several structural proteins and a 3’ to

5’ exonuclease. B. The rapid tRNA decay pathway (RTD) degrades various hypomodified or destabilized tRNA such as tRNAVal and tRNASer.

Export is the last major step that takes place in the nucleus and therefore serves as the final checkpoint tRNAs must clear before entering the cytoplasm.

One of the major tRNA exporters, exportin-t, was first discovered when searching for nuclear transporters which interact with RanGTP, the GTPase involved in many transports functions (Kutay et al. 1998). Exportin-t preferentially binds end- processed tRNAs in vitro but does not discriminate between the presence or absence of introns, instead relying on the recognition of a properly folded tRNA backbone (Arts et al. 1998). Lack of discrimination between intron-containing and intronless tRNAs is especially important in cases where the splicing machinery localizes to the cytoplasm, for example, in S. cerevisiae and Trypanosoma brucei

(Lopes et al. 2016; Yoshihisa et al. 2003). In turn, deletion of the S. cerevisiae exportin-t homolog (Los1) causes end-matured, intron-containing tRNA to accumulate in the nucleus but paradoxically is non-essential for viability (Hellmuth et al. 1998; Hopper et al. 1980; Sarkar and Hopper 1998). The plant ortholog is also non-essential indicating the existence of redundant pathways for export from the nucleus to the cytoplasm (Hunter et al. 2003; Li and Chen 2003). The existence of alternative tRNA exporters became evident with the discovery of exportin-5 in vertebrates and its homolog Msn5 in S. cerevisiae. Exportin-5 exports microRNAs and to a varying degree, aminoacylated tRNAs bound to eukaryotic elongation factor 1A (Calado et al. 2002; Yi et al. 2003; Shibata et al. 2006). Although tRNAs interact with exportin-5, they do so with different affinities; consequently, the role of exportin-5 in tRNA export varies among different species. Knockdown of

20 exportin-5 in humans and plants does not affect tRNA levels in the nucleus, while a knockdown in Drosophila, which naturally lacks exportin-t, leads to nuclear tRNA accumulation (Park et al. 2005; Shibata et al. 2006). In S. cerevisiae, Msn5 is the primary re-exporter for tRNAs, which have been retrograde-imported into the nucleus after splicing, and as, it such binds to matured aminoacylated tRNA

(Murthi et al. 2010; Huang and Hopper 2015). It should be noted that the ability for

Msn5 to export tRNAs that do not require splicing has not been ruled out. Although it is clear both Los1 and Msn5 share a role in tRNA export, deletion of Los1 and

Msn5 simultaneously is still not lethal (Takano et al. 2005), indicating further built- in redundancy. However, in some systems nuclear export is not an all or nothing mechanism, but more of a kinetically controlled pathway, where tRNAs will be exported regardless, but exporters facilitate the export of “healthy” tRNAs (Lund and Dahlberg 1998).

When comparing the presence and absence of modifications, modified tRNAs bound with greater affinity to exportin-t than tRNA lacking modifications

(Kutay et al. 1998). Seeing how various modifications give rise to changes in both the structure and stability of tRNAs, it is conceivable that hypomodification could affect the export potential for tRNA from the nucleus. In fact, various point mutations interrupting critical tertiary contacts between the D and TΨC loops caused a decrease in affinity to exportin-t and subsequent nuclear export (Tobian et al. 1985). Disruption of the TΨC stem caused a similar reduction in the binding affinity of exportin-t for tRNA (Lipowsky et al. 1999). Although specific

21 modifications themselves may individually be too small to influence export directly, hypomodification could influence the structure and rigidity of tRNA, which subsequently influences export. Along these lines, organisms have also adopted aminoacylation as a further checkpoint for export. For example, it was discovered that in Xenopus oocytes tRNAs are aminoacylated prior to nucleus-export to the cytoplasm (Lund and Dahlberg 1998). The lack of aminoacylation leads to a decrease in tRNA nuclear export and nuclear accumulation in yeast (Grosshans et al. 2000; Sarkar et al. 1999). The adoption of tRNA aminoacylation as a prerequisite for nuclear export allows further screening of tRNA “healthiness” by aminoacyl tRNA synthetases, as a type of proofreading mechanism to ensure that only “good” tRNAs make it to the cytoplasm.

1.4.2 The cytosolic fate of tRNA modifications

After export from the nucleus, most tRNAs lack a complete set of modifications and still rely on numerous cytoplasmic modification enzymes for the final steps of maturation. In most cases, the intracellular distribution of modification enzymes is not clear, but certainly, many localize to the cytoplasm. The reasons why modification enzymes localize to specific compartments is not well understood, but in some cases localization is in line with the localization of other processing enzymes that are critical for tRNA maturation. For example, modification enzymes that are localized to the cytoplasm may recognize tRNA substrates that have previously undergone splicing, such as tRNA containing

22 ribose methylations catalyzed by Trm44 (Um44), Trm3 (Gm18), or Trm7 (Cm32, Nm34)

(Hopper 2013). Here, it stands to reason that if splicing is cytoplasmic and the given enzyme requires the intron for substrate recognition, then it follows that the modifications will likely be cytoplasmic. However, this is not always the case, for there are examples of modifications that only occur after splicing but reside in the nucleus, as discussed in the following pages. In other cases, there are modification enzymes that do not discriminate based on the presence or absence of an intron, and thus their localization cannot be predicted by looking at the distribution of different processing events.

The differential distribution of modification enzymes within cells could potentially lead to the accumulation of immature tRNAs in the cytoplasm, which could cause a problem during protein synthesis. However, analogous to the nucleus, the cytoplasm also has mechanisms to deal with hypomodified tRNAs.

Studies examining the importance of modifications were at first puzzling, as many genes encoding evolutionarily conserved modifications were found non-essential.

For example, deletion of genes encoding the enzymes responsible for the synthesis of m5C (at positions 34, 40, 48 and 49), D (at positions 16, 17, 20 and

47), Ψ (at positions 13, 31, 35 and 55), 7-methylguanosine (m7G) at position 46,

1 and m G9 had little to no effect on growth (Alexandrov et al. 2005; Ansmant et al.

2001; Becker et al. 1997; Jackman et al. 2003; Ma et al. 2003; Xing et al. 2002).

However, a number of nonessential genes encoding for modification enzymes when deleted in tandem lead to synthetic lethality. For example, deletion of several 23 genes encoding “nonessential” modification enzymes in yeast, led to a significant

Val reduction in the steady-state levels of tRNA AAC (Alexandrov et al. 2006). This observation led to the discovery of the rapid tRNA decay pathway (RTD), which works independently of the nuclear surveillance pathway described previously

(Figure 1.5 A). Work to identify the components of RTD uncovered three genes which affected tRNA degradation: XRN1, RAT1, and MET22. Xrn1 and Rat1 are both 5’ to 3’ exonucleases, which can mediate degradation of destabilized tRNA, while Met22 indirectly modulates Rat1 and Xrn1 activity via accumulation of the inhibitor adenosine 5’,3’ bisphosphate (Figure 1.5 B) (Chernyakov et al. 2008;

Dichtl et al. 1997). Not all tRNAs are affected equally by RTD after multiple “non-

7 5 essential” modification genes are deleted. For example, loss of m G46 and m C49

Val Val in both tRNA AAC and tRNA CAC, results in the targeted degradation of only

Val 7 5 tRNA AAC. This specificity is also observed for other m G46 and m C49 containing tRNAs such as tRNAMet and tRNAPhe neither of which is subsequently degraded following the loss of these specific methylations. Similarly, loss of N4-acetylcytidine

4 Ser at position 12 (ac C12) and Um44 leads to the specific degradation of tRNA

Ser UGA/CGA while tRNA IGA/GCU is unaffected (Chernyakov et al. 2008; Kotelawala et al. 2008). These results point out that simply lacking a set of modifications is not enough to cause RTD but likely leads to a destabilization or disruption of the tRNA structure in certain tRNAs. In favor of this, the nucleotide sequence of the RTD

Ser Ser susceptible tRNA CGA was altered to more closely resemble that of tRNA IGA in

Ser the TΨC and acceptor stem. Unlike before, the resulting mutant tRNA CGA was

24 not degraded by RTD as the altered nucleotide sequences allowed for the stability

4 required to compensate for the loss of ac C12 and Um44 (Whipple et al. 2011). The data suggests that RTD can be triggered by the absence of modifications that alter the structural stability in the TΨC and acceptor stems. The presence of certain pairs of modifications becomes more crucial for tRNAs that may be inherently more structurally unstable and rely on those modifications more heavily for proper folding. The existence of a cytoplasmic tRNA monitoring pathway helps ensure that only properly modified tRNAs participate in translation, while limiting the availability of hypomodified tRNAs, which may cause problems of translational efficiency, accuracy or both.

In general, it is difficult to predict or even tease out the significance of the intracellular compartmentalization for a given modification enzyme. Although, examples above highlight aspects of specific systems where the localization of a given processing event establishes modification enzyme distribution, in reality, there are not many ‘hard and fast’ rules for localization prediction. In the end, it may well be that intracellular localization is dynamic, if not transitory, and cells may exploit intracellular partitioning as a way to control enzyme function and certainly impact substrate availability and recognition.

1.4.3 The connection between tRNA splicing and modifications

In all domains of life, subsets of tRNAs are interrupted by introns; these must be removed before the tRNA can be used for translation (Lopes et al. 2015;

Yoshihisa 2014) There is no conservation regarding the number and distribution of tRNA introns across different species, with the range of intron-containing tRNA genes varying from a single tRNA in Azoarcus sp. and T. brucei, to 5% of all tRNAs genes in humans, mouse and drosophila, 20% in S. cerevisiae, 48% in

Crenarchaeota and 92% in the yeast Cryptococcus neoformans ( Berriman et al.

2005; Goffeau et al. 1996; Krause et al. 2006). The mechanism of tRNA splicing also varies between domains of life: bacterial introns are removed through a self- splicing mechanism, while archaeal and eukaryotic introns are removed strictly through protein-catalyzed reactions involving specialized enzymes that work independently from the mRNA spliceosomes (Krause et al. 2006; Lopes et al.

2015; Barbara and Shub 1992). Despite the fact that tRNA introns are ubiquitous, ascertaining the reasons why they are maintained has proven challenging; some studies indicate certain introns can be removed from the genome with little effect on the organism, while in other cases introns serve as important substrate recognition elements for splicing and modification enzymes (Mori et al. 2011;

Hopper 2013). Early studies in Xeonopus oocytes were used to classify tRNA modifications under three categories, according to their dependence on the presence of an intron (Nishikura and De Robertis 1981; Grosjean et al. 1997). The first category includes modifications that are added only to intron-containing

5 tRNAs, including Ψ (at positions 34-36) and m C (at positions 34 and 40) (Figure

1.6 A). The second involves modifications that are added only to spliced tRNAs,

3 including Ψ32, m C32, Um44 and Gm18 (Figure 1.6 B). The third, and final category,

26 contains modifications that are added to tRNA regardless of the presence of an intron, but do not depend on the intron as a recognition element; these include

1 2 m G9, m G6, 10, D16, 17, 20, and 5-methoxycarbonylmethyl-2-thiouridine at position

5 2 34 (mcm s U34) (Grosjean et al. 1997). Interestingly, studies in which the intron sequence was altered, in an effort to elucidate its role in substrate recognition by the tRNA splicing endonuclease and/or modification enzymes, showed that most mutations do not prevent splicing or modification, as long as they do not interfere with the overall cloverleaf structure of the pre-tRNA (Johnson et al. 1980; Strobel and Abelson 1986). Two noteworthy studies on this regard were performed in yeast, taking advantage of naturally occurring suppressor tRNAs. In the first study, the genomic copy of the tRNATyr gene SUP6 was replaced with a version that lacked the 14 nucleotide-long intron (Johnson and Abelson 1983). SUP6 is a tRNATyr ochre suppressor, responsible for tRNA-mediated nonsense suppression during translation. Analysis of the mutant tRNA showed that it lacked the modification Ψ at the second position of the anticodon (Ψ35), indicating that either the intron was necessary for substrate recognition by the appropriate modification enzyme, or that the splicing pathway was a pre-requisite for the modification to occur (Johnson and Abelson 1983). This mutant tRNA was defective at nonsense suppression, and present in lower concentrations in the cell (Johnson and Abelson

1983). In a following study, the 32 nucleotide-long intron sequence of the tRNALeu gene SUP53, another nonsense suppressor, was either mutated or removed, and the mutant tRNAs analyzed (Strobel and Abelson 1986). The mutant tRNA,

5 transcribed from the intronless gene, lacked the modification m C34, and was defective at tRNA-mediated nonsense suppression (Strobel and Abelson 1986).

Figure 1.6: Dependence of modification on introns A. Certain modifications (as indicated) can only be added to intron-containing tRNA; the intron is an essential recognition element for their respective enzyme.

Refer to Table 1. B. Some modifications are only added after splicing (as indicated).

Unusual examples are the cases of T. brucei and Haloferax volcanii, in the former, the intron sequence must be first edited in two or three positions in order for splicing to take place (Rubio et al., 2013). In the latter, the tRNATrp intron once cleaved becomes a small guide RNA (sRNA) required for ribose methylations to

Trp form Cm34 and Um39 on the cognate spliced tRNA (Clouet d’Orval et al. 2001).

These two examples provide a rationale for intron maintenance and in both cases the splicing of the intron is intricately connected to an additional activity. Notably, in the T. brucei system, and other kinetoplastids, tRNATyr is the only intron- containing tRNA. Therefore, either at some point during their evolution other tRNAs contained introns in these organisms and the intron-containing genes where systematically replaced by their spliced equivalent, or most tRNAs never contained introns. Importantly, despite the variability in the numbers and types of intron- containing tRNAs in many eukaryotes, in all cases tRNATyr always contains an intron, arguing for the importance of that particular intron.

As mentioned above, since in most eukaryotes tRNA splicing takes place in the nucleus, modifying enzymes that act on intron-containing tRNAs also localize to the nucleus, while those that act on spliced tRNAs may also localize to the cytoplasm (Grosjean et al. 1997). There is at least one significant exception to this rule, however, in S. cerevisiae the tRNA splicing machinery localizes to the surface of the mitochondria facing the cytoplasm. In this organism, tRNA splicing takes place in the cytoplasm, and many modifying enzymes that act on both pre-tRNA and spliced tRNA localize to the cytoplasm as well (Grosjean et al. 1997; Hopper

2013; Yoshihisa et al. 2003). These include the cases of SUP6 and SUP53

5 described above, in which the modifying enzymes (Pus7 for Ψ35 and Trm4 for m C) localize to the cytoplasm and require the intron for substrate recognition

(Machnicka et al. 2013).

1.4.4 Intracellular transport dynamics that set the order of modifications

It was always assumed that tRNAs moved unidirectionally from their site of transcription in the nucleus to their site of action in the cytoplasm, and the cytoplasm was the final destination. Work in recent years, however, demonstrated that the cytoplasm is not the last stopping point for tRNAs. Not only can tRNAs be reimported to the nucleus (by retrograde transport) but, in a growing number of organisms, tRNAs are also imported into the organelles, for example mitochondria

(Rubio and Hopper 2012). The latter, as discussed below, serve to complement or complete the tRNA set needed for organellar translation.

The concept of tRNA retrograde transport was first described in S. cerevisiae, based on the observation of significant levels of spliced tRNAs in the nucleus, despite splicing being a cytoplasmic event (Shaheen and Hopper 2005;

Takano et al. 2005). It was difficult to explain why mutations of MES-1 or CCA-1, which led to lack of aminoacylation, caused the accumulation of spliced tRNA within the nucleus (Sarkar et al. 1999; Feng and Hopper 2002). The retrograde nuclear transport pathway was then discovered by means of an elegant heterokaryon assay, where a karyogamy deficient kar1-1 mutant, which prevents

31 nuclear fusion after mating was used. This leads to heterokaryon cells, which harbor two nuclei in a shared cytoplasm. Using heterokaryon assays in combination with fluorescence in situ hybridization (FISH) to label tRNA, the movement of tRNA from a donor to a target nuclei was observed (Shaheen and

Hopper 2005; Takano et al. 2005). Furthermore, spliced endogenous tRNAs accumulated in the nucleus after treatment with the RNA polymerase inhibitor thiolutin, again supporting the movement of tRNA from the cytoplasm to the nucleus. Since then, multiple lines of evidence have supported these initial studies while several nutritional conditions have been outlined that cause nuclear accumulation of tRNA (Grosshans et al. 2000; Hurto et al. 2007; Whitney et al.

2007). The use of heterokaryon assays proved to be instrumental in demonstrating the existence of bidirectional movement of tRNA, further highlighting tRNA intracellular transport dynamics.

It is now clear that retrograde transport occurs constitutively. Thus retrograde transport constantly cycles tRNA in and out of the nucleus, (Murthi, et al. 2010) raising the possibility that it may also play other roles in tRNA maturation.

An interesting example is offered by wybutosine (yW) a bulky hypermodified guanosine found at position 37 in tRNAPhe of most eukaryotic organisms; it provides an important frameshifting-prevention mechanism (Waas et al. 2007). yW biosynthesis involves the formation of a new heterocycle in an otherwise standard guanosine base, this is followed by several enzymatic steps that add various methyl groups and an ACP (aminocarboxypropyl) side chain. It all starts with

1 methylation of G37 to form m G, a reaction catalyzed by Trm5 in eukaryotes. This is followed by serial enzymatic reactions catalyzed by Tyw1, Tyw2, Tyw3, and

Tyw4 (and Tyw5 in some organisms). Although Trm5 methylates several tRNAs, only tRNAPhe gets yW. In yeast, Trm5 localizes to the nucleus, while the remaining

Tyw enzymes are found in the cytoplasm, thus formation of yW involves two different cellular compartments. The story is, however, even more complicated as

Trm5 is not able to methylate intron-containing tRNAPhe and since the splicing endonuclease is tethered to the outer membrane of the mitochondria facing the cytoplasm, the tRNA has to forcibly go back and forth from the nucleus to get yW.

Thus, the separation of these maturation steps by intracellular compartmentalization causes a situation whereby yW synthesis is dependent on retrograde transport. First, intron-containing tRNAPhe is transcribed in the nucleus and receives some modifications and undergoes end maturation. Then intron- containing tRNAPhe is exported to the cytoplasm for splicing, followed by retrograde

1 transport to the nucleus to get m G37, and finally it is re-exported to the cytoplasm to complete the synthesis of yW (Figure 1.7)(Ohira and Suzuki 2011).

Figure 1.7: Yeast yW biosynthesis pathway The biosynthesis of wybutosine (yW) in S. cereviase requires retrograde transport of tRNAPhe to the nucleus. Intron-containing tRNAPhe is exported to the cytoplasm from the nucleus, where the intron is cleaved by the heterotetrameric splicing endonuclease, which localizes to the outer surface of the mitochondria facing the cytoplasm endonuclease. The exons are then processed by the tRNA-splicing ligase (Trl1) and the tRNA phosphotransferase (Tpt1) to complete the splicing reaction. The newly spliced tRNAPhe is imported back into the nucleus (retrograde

1 transport) where a nucleus-localized 1-methyltransferase Trm5 forms m G37. The methylated tRNA is then re-exported to the cytoplasm, where a series of reactions catalyzed by Tyw1, Tyw2, Tyw3, Tyw4 ultimately creates yW37.

In the trypanosome system, we have discovered a second case of retrograde transport impacting tRNATyr and the modified nucleotide queuosine (Q)

(Chapter 2 of this dissertation). This occurs at position 34 of the anticodon of tRNATyr, -Asp, -Asn and -His in all Eukarya, except for S. cerevisae, which naturally lacks this modification pathway. Q formation is catalyzed by tRNA guanine- transglycosylase (TGT), which is strictly a sequence-specific enzyme and only modifies anticodon loops containing the trinucleotide UGU (positions 33-35)

(Carbon et al. 1983; Kung et al. 2000). TGT has a known substrate preference for spliced tRNA as the transglycosylation reaction performed when replacing G with

Q is inhibited by the presence of an intron (Nishikura and De Robertis 1981). In mammals, tRNA splicing is confined to the nucleus and TGT has been localized to the outer membrane of the mitochondria thus clearly Q formation occurs in a spliced tRNA. In T. brucei, however, tRNA splicing, like in yeast, localizes to the cytoplasm, (Lopes et al. 2016) while TGT localizes to the nucleus. In T. brucei as mentioned before, tRNATyr contains an intron, which must be removed before TGT can use it as a substrate. This again creates a similar situation as tRNAPhe in yeast; intron-containing tRNATyr must leave the nucleus for splicing and then re-enter the nucleus for Q addition (Figure 1.8).

Although retrograde transport is necessary for Q formation, its current role in T. brucei is not fully understood. In other organisms, however, studies conducted on the role Q may have on translation suggest the presence of Q can alter the codon bias of Q-containing tRNA. For example, Xenopus microinjections of either

His 3 G34-containing or Q34-containing tRNA aminoacylated with [ H]-labeled histidine indicated a clear preference toward CAC over CAU codons when Q was absent

(Meier et al. 1985). Subsequent work has argued that Q can alter codon preference by altering the speed and accuracy of a given Q-containing tRNA codon, which ultimately shapes the proteome (Zaborske et al. 2014).

Although thus far only yW and Q require retrograde transport, this limited set is more representative of the difficulty in evaluating the intracellular distribution of modifications, rather than an exhaustive list. For example, an interesting possibility exists with the dually localized (nucleus and cytoplasm) modification enzyme Mod5, which generates isopentenyladenosine (i6A), a common modification important for translational accuracy in all domains of life. Addition of

6 i A37 only happens after intron-containing tRNAs are spliced, which as discussed previously occurs in the S. cerevisiae cytoplasm (Chimnaronk et al. 2009). The possibility then exists that after splicing, a portion of the spliced tRNA travels to the nucleus to receive i6A. However, given that significant amounts of Mod5 are detectable in the cytoplasm, it may well be that retrograde transport is not really necessary for i6A formation.

Aside from a direct requirement for retrograde transport to facilitate modifications, such as the case of yW and Q, it is possible that dual localization of modification enzymes serves a purpose as a repair mechanism. One could envision a scenario where the tRNA export rate to the cytoplasm may be faster than the rate of nuclear modification, depending on environmental conditions. It

36 then may be possible that hypomodified tRNAs may make it to the cytoplasm; these either get degraded by the RTD pathway or alternatively may be remodified by dually localized enzymes to set them once more into a fully modified, fully functional state. In this realm, constitutive retrograde transport may serve a similar

“rescue” function. Perhaps the dual localization of Mod5 reflects such a role. Likely, many tRNAs may take advantage of this “second chance” at modification, which may be advantageous if repairing modifications becomes preferable to degradation and re-synthesis.

Figure 1.8: The biosynthesis of Q in T. brucei T. brucei requires retrograde transport of tRNATyr to the nucleus for Q formation.

The only intron-containing tRNA in T. brucei, tRNATyr, is exported to the cytoplasm for splicing by the tRNA splicing-specific endonuclease, ligase (Trl1) and phosphotransferase (Tpt1). After splicing tRNATyr travels back to the nucleus where the nucleus-localized tRNA-guanine transglycosylase (TGT) forms Q34.

1.4.5 Modifications and the Mitochondria

Over the course of evolution, mitochondria have undergone loss of genetic material, becoming dependent on nucleus-encoded genes and protein transport from the cytoplasm for function (Salinas et al. 2008). The number of tRNAs encoded by this organelle varies considerably between species and includes a presumably minimal but complete set in humans, all except those for six amino acids in Arabidopsis thaliana and only three tRNAs in Chlamydomonas reinhardtii

(Salinas et al. 2008). In the most extreme cases, such as the kinetoplastids (e.g.

Leishmania tarentolae and T. brucei) and other protists, no tRNAs are encoded in the mitochondrial genome and the complete set must be imported from the cytosol

(Suyama 1967; Schneider and Maréchal-Drouard 2000). This led to the question of the evolution of mitochondrial tRNA import and how it relates to tRNA demands set by mitochondrial translation. To answer this question, work performed in C. reinhardtii replaced the often-used GGC and GGT codons with the seldom-used

GGG in the mitochondrial genome, increasing the percentage of GGG codons from

Gly 0.1 to 0.45%. Strikingly, no increase in import of the tRNA CCC, now presumably needed in higher amounts, was observed; the mutants showed decreased respiratory rates caused by low activity of complexes I and IV, increased doubling time and reduced mitochondrial protein synthesis. These results suggest that the correlation between mitochondrial tRNA import and codon usage is most probably a result of co-evolution of both import and translation, and that tRNA import cannot

39 be regulated to quickly cope with changes in mitochondrial genome content

(Salinas et al. 2012).

Overall, tRNA import seems to functionally complement what is encoded in the mitochondrial genome by providing the missing tRNA species necessary for mitochondrial translation. There are cases, however, in which the imported tRNAs are already encoded in the mitochondrial genome: in the plant Marchantia

Val polymorpha, tRNA AAC is imported from the cytoplasm, where it coexists with the

Val mitochondrial-encoded tRNA UAC, which alone was considered sufficient for translating all Val codons through wobble pairing (Akashi et al. 1998). In yeast, rat

Gln and human mitochondria, the nucleus-encoded tRNA UUG was shown to co-exist

Gln with the mitochondria-encoded tRNA UUG after being imported through an ATP- dependent mechanism (Rinehart et al. 2005; Rubio et al. 2008). Finally, in S.

Lys cerevisiae, the imported tRNA CUU co-exists with the mitochondria-encoded

Lys tRNA UUU, which was considered sufficient for translating all Lys codons through wobble pairing (Martin et al. 1979). Interestingly, recent work shed some light on to the yeast tRNALys import; under high temperatures (37 °C, as opposed to the

Lys normal 30 °C for yeast), the mitochondria-encoded tRNA UUU becomes hypomodified and incapable of decoding the AAG codon, with the imported

Lys tRNA CUU becoming necessary to correct the translational deficiency, indicating that, perhaps the seemingly redundant imported tRNAs become particularly important under certain growth conditions (Kamenski et al. 2007).

The determinants for mitochondrial import are still poorly understood and do not appear to be conserved even within similar organisms, nevertheless, sequence specificity, editing and modification have been described as determinants in some systems (Alfonzo and Söll 2009). One noteworthy study compared the cytosolic and mitochondrial populations of tRNAGlu, tRNAGln and tRNALys in L. tarentolae. In this organism, populations of tRNAs are not always evenly distributed among the cell compartments, and can be classified accordingly: group I tRNAs are mostly cytosolic, group II, which includes tRNALys, are mostly mitochondrial and group III, which includes tRNAGlu and tRNAGln, are equally distributed between compartments (Kaneko et al. 2003). Their analysis revealed

Glu Gln 5 2 that the cytosolic tRNA and tRNA populations contained mcm s U34, while the mitochondrial populations of the same tRNAs contained only 5- methoxycarbonylmethyl-2′-O-methyluridine (mcm5Um). Furthermore, in vitro assays with isolated mitochondria showed that the thiolated tRNAs were not as efficiently imported into the organelle as the non-thiolated ones, suggesting that L.

2 tarentolae uses s U34 as a negative determinant for mitochondrial import (Figure

1.9 A) (Kaneko et al. 2003). Interestingly, similar work performed in the closely related organism T. brucei showed a different result. Knocked down by RNAi of the cysteine desulfurase (TbNfs), essential for tRNA thiolation, led to

2 disappearance of s U34, as expected, but had no effect on mitochondrial tRNA import in vivo or in vitro (Paris et al. 2009).

Mitochondrial reliance on nucleus-encoded tRNAs has also led to an interesting situation beyond simply supplying tRNAs: Reflecting its bacterial ancestry, the mitochondrial genome is not universal and differs from the nuclear genetic code. For example, in most eukaryotes, with the exception of plants, UGA has been reassigned from a stop codon to now mean tryptophan. In turn, most mitochondrial genomes encode a tRNATrp with anticodon UCA dedicated to reading the reassigned codon. But this raised an interesting question in kinetoplastid protists, which do not encode any tRNAs in their mitochondria, and thus import all tRNAs from the cytoplasm, and encode no gene in the nuclear genome that could act as a potential suppressor. It was first found in L. tarentolae

Trp that these organisms import the standard tRNA CCA into mitochondria and following import approximately 50% of the tRNA is edited from C34 to U34

Trp generating tRNA UCA that can now decode UGA as tryptophan, while maintaining

Trp a significant level of tRNA CCA presumably to decode the UGG codons, which also exist in mitochondria (Wohlgamuth-Benedum et al. 2009; Phizicky and

Alfonzo 2010). The question then is how is the 50/50 ratio between edited and unedited tRNA maintained. It was found that in L. tarentolae, tRNATrp was also thiolated at U33, a position that was supposed to be unmodified in all tRNAs in all organisms. However, only the edited tRNA was thiolated, suggesting a connection between editing and thiolation (Crain et al. 2002). A similar situation was later found in T. brucei, but in this case both the edited and unedited tRNAs were thiolated and in addition there was significant amounts of edited but not thiolated

42 tRNATrp, suggesting that this modification may not be a requirement for editing

(Charrière et al. 2006). Separate studies in T. brucei then showed that thiolation acts as a negative determinant for editing, with the down-regulation of tRNA thiolation in this organism, achieved by knockdown of the conserved cysteine desulfurase Nsf1, leading to an increase in C to U editing levels that resulted in almost 90% of tRNATrp being edited (Wohlgamuth-Benedum et al. 2009).

Curiously, mitochondrial thiolation relies on Nsf1, which is also required for cytoplasmic thiolation, but in addition, mitochondrial thiolation requires the mitochondrial enzymes tRNA-specific 2-thiouridylase, Mtu1, and iron-sulfur biogenesis desulfurase interacting protein, Isd11 (Figure 1.9) (Phizicky and

Alfonzo 2010; Čavužić and Liu 2017). Overall, these findings indicate considerable differences between the cytoplasmic and mitochondrial thiolation pathways, and how thiolation and editing are specifically handled between these two closely related organisms.

Finally, there are cases in which tRNA modifications help with naturally occurring extreme structural changes in tRNAs, such as the truncated mitochondrial tRNAs found in nematodes. These tRNAs are unlike cytoplasmic tRNAs, with many lacking the entire D or T-arms. Recognition by tRNA binding proteins is thus dependent on unique proteins including mitochondria-specific elongation factors (EF-Tu1 and 2) that recognize armless tRNAs only (Ohtsuki et al. 2002; Lorenz et al. 2017). In the nematode Ascaris suum, EF-Tu1 recognition of tRNAMet, which lacks the T-arm, is achieved through unique interactions

43 between the protein C-terminus and the D-arm (Sakurai et al. 2006). This interaction is dependent both on conserved residues in the D-arm, and in the overall structure of the tRNA, which is altered by the presence of the modification

1 m A9. This methylation leads to a different folding pattern in the D-arm and the

1 small loop region that replaces the missing T-arm. Moreover, m A9 was also shown to be important for efficient aminoacylation, as the structural changes it generates also change the distance between the CCA and the anticodon, affecting the binding efficiency of the corresponding aminoacyl tRNA synthetase (Lorenz et al.

2017).

We have previously proposed that post-transcriptional modifications do not occur in isolation but in fact may be part of largely organized and well-orchestrated chemical cascades (Rubio et al. 2017). One modification may in a subtle manner change the local structure of a target substrate creating a transitory substrate structure that will then be slightly changed by a subsequent modification and so on. Such ideas become even more provocative when one considers intracellular transport dynamics and the permeability barriers set forth by cellular membrane systems. It is our view that cellular transport of tRNAs across membranes may be influenced by many factors and transport rates may change in response to environmental cues. We also suggest that such changes will no doubt impact tRNA processing and tRNA modifications. Environmental sensing will no doubt be at the front and center of how cells couple translational responses with metabolite availability via tRNA modifications.

Figure 1.9: Mitochondrial import in L. tarentolae and T. brucei use thiolation A. Thiolation of tRNAGlu and tRNAGln acts as a negative determinant of mitochondrial import in L. tarentolae, but not in T. brucei. B. A portion of tRNATrp is kept for cytoplasmic translation, while another is imported into the mitochondria.

In the mitochondrial lumen, tRNATrp is subjected to thiolation at the unusual position U33 and edited from C to U at position 34. In L. tarentolae, only the edited tRNA gets thiolated whilst in T. brucei, thiolation acts as a negative determinant for editing. The same enzyme Nisf1, is responsible for thiolation of cytoplasmic

Glu Gln tRNA and tRNA at position U34.

1.5 The Queuosine Modification

One of the more structurally complicated nucleotide modifications found in cells is Queuosine (Q). Q is present in nearly all known eukaryotic and bacterial organisms, with a notable exception in S. cerevisiae. Although highly conserved, the exact function of Q has been elusive but various connections have been made with other cellular process, some of which will be further examined later in this section. Given the wide distribution of Q throughout Eukarya, it is intriguing that only bacteria can synthesize the modification. Eukaryotic organisms must rely on bacteria as the initial source of Q and instead of de novo synthesis, salvage Q from their surroundings (Farkas 1980; Reyniers et al. 1981; Marks and Farkas 1997).

In both bacterial and eukaryotic organisms, the hallmark enzyme of the Q pathway is tRNA-guanine transglycosylase (TGT). TGT performs a transglycosylation base exchange reaction at G34 with the nucleotide base queuine in eukaryotes, or a Q precursor, preQ1 in bacteria. In a similar reaction archaeal

TGT utilizes preQ0 in the penultimate step for the synthesis of archaeosine

(Watanabe et al. 1997; Kung and Garcia 1998; Boland et al. 2009). TGT recognizes its tRNA substrate by a U33G34U35 sequence found in the anticodon stem loop of several tRNAs leading to the modification of tRNAsTyr, His, Asp, Asn

(Harada and Nishimura 1972; Kung and Garcia 1998; Okada et al. 1976).

1.5.1 The Queuosine Pathways of Bacteria and Eukarya

The de novo synthesis of Q (2-amino-5-[(3S, 4R, 5S)-4,5- dihydroxycyclopent-1-en-3-ylaminomethyl]-7-( β-D-ribofuranosyl)-pyrrolo-[2,3-D]- pyrimidin-4-one) is a complicated, several step pathway occurring in bacteria.

Unlike most modifications, synthesis takes place mostly outside of tRNA. The first major step of Q biosynthesis is the multi-step conversion of GTP to 7-aminomethyl-

7-deazaguanine (preQ1). Here, N7 contained within the guanine base is lost, creating the deazaguanine base which Q is built upon. In this step, GTP cyclohydrolase I converts GTP to 7, 8-dihydroneopterin triphosphate (H2NTP)

(Phillips et al. 2008) and from there QueD converts H2NTP to 6-carboxy-5,6,7,8- tetrahydropterin (CPH4) (Chalovich and Eisenberg 2005). At this point QueE converts CPH4 into 7-carboxy-7-deazaguanine (CDG) (McCarty et al. 2009) followed by CDG conversion by QueC to 7-cyano-7-deazguanine (preQ0) (Gaur and Varshney 2005). In the final step, QueF converts the nitrile group to an amino group creating the Q precursor preQ1 (Van Lanen et al. 2005). PreQ1 is the substrate for the unique post-transcriptional base exchange of guanine 34 catalyzed by TGT (discussed in the following subsection). PreQ1-containing tRNA is the substrate for QueA to facilitate the isomerization and transfer of the ribose moiety from S-adenosylmethionine to preQ1 ultimately generating epoxy queuosine (oQ) (Quea et al. 2003; Slany et al. 1994) which is then reduced by epoxyqueuosine reductase generating Q (Figure 1.10) (Miles et al. 2011).

In bacteria, Q-containing tRNAAsp can be further modified with the addition of glutamate to the Q modification at C4” or C5” of the cyclopentene ring (Salazar et al. 2004). This is facilitated by the unique Glu-Q-tRNAAsp synthetase (Glu-Q-RS) which is encoded for by the yadB gene. Glu-Q-RS activates glutamate without the requirement of a cognate tRNA and has affinity for the anticodon stem loop of tRNAAsp (Dubois et al. 2004). Interestingly, the nucleotide sequence for the anticodon stem of tRNAAsp shares a six nucleotide stretch identical to the acceptor stem sequence for tRNAGlu potentially sharing some identity elements utilized by

Glu-RS (Blaise et al. 2005).

Figure 1.10: Pathways of Queuosine synthesis A. Bacteria synthesize Q de novo utilizing several steps to produce Q modified tRNAs. B. In eukaryotes, Q (or a form of Q) is taken from the environment before being converted to queuine and inserted into tRNA as eukaryotes do not possess the means for de novo synthesis. The modification Q, shown in C, can be further modified with the addition of either mannose, or galactose in eukaryotes, or glutamine in bacteria, to one of the hydroxyls of the cyclopentane ring. This reaction specifically generates Gal-Q-tRNATyr, Man-Q-tRNAAsp in eukaryotes, and

Glu-Q-tRNAAsp in bacteria. In a similar modification outlined in D, archaea undergoes several initial steps of Q synthesis but diverge to generate archaeosine.

The Q pathway in eukaryotes is not as well defined. A form of Q must be imported into the cell for TGT to utilize queuine in its exchange reaction (Figure

1.10). Specifically in mammals, the source of Q includes both the gut microbiome and the diet (Farkas 1980; Reyniers et al. 1981). Various forms of Q such as Q- containing tRNA itself, Q-5’-phosphate or Q-3’-phosphate have been proposed to serve as sources of Q which must be processed to the free base queuine before being used by TGT (Reyniers et al. 1981; Gündüz and Katze 1982). To identify components of the eukaryotic Q salvage pathway, a combination of computational and phylogenetic approaches were taken. Among the potential gene candidates uncovered, a DNA glycosylase-like homolog family DUF2419 was co-conserved with eukaryotic TGT, and absent from organisms which did not maintain Q.

Deletion and rescue experiments confirmed DUF2419 was essential to Q, albeit for an unknown reason (Zallot et al. 2014). Eukaryotes can also further modify Q- containing tRNAAsp and tRNATyr with the addition of mannose and galactose, respectively (Q*) ( Okada 1977; Okada et al. 1977). The pathway to generate Q* has not been characterized nor has the mannosyl or galactosyl transferase been identified.

1.5.2 TGT in Bacteria and Eukarya

In bacteria, TGT is capable of recognizing and catalyzing the base exchange reaction in a U33G34U35 containing mini-helix substrate demonstrating a fairly minimal recognition motif (Curnow and Garcia 1995). TGT coordinates one

Zn2+ in a C terminal domain with conserved cystine residues 302, 304, 307 and histidine 317 (E. coli numbering), necessary for enzyme structure and tRNA binding. (Chong et al. 1995; Garcia et al. 1996).

Initial crystal structures of Zymomonas mobilis TGT in complex with tRNA and preQ1 pointed to several amino acids that could be involved in catalysis.

Specifically, aspartate 102, 156, 280 (Z. mobilis numbering) were found in close proximity to the active site and conserved throughout all TGTs (Romier et al.

1996a; Romier et al. 1996b). The initial crystal structures allowed for the proposal of a three-step mechanism which was subsequently supported in kinetic studies

(Goodenough-Lashua and Garcia 2003). After TGT binds to its tRNA substrate, the nucleophilic attack of Asp (Originally suspected Asp 102 but later disproven) to the C1’ carbon of G34 ribose was said to release guanine. This mechanism was later supported by chemical trapping and X-ray cystography importantly highlighting Asp 280 as the critical nucleophile necessary for the covalent intermediate established between the RNA and TGT (Xie et al. 2003). After this, the leaving guanine is replaced with preQ1 in the binding pocket. The final step outlined was the nucleophilic attack of preQ1 N9 to the C1’ of the ribose at position

34. This would therefore join preQ1 to the ribose and afterward release the enzyme. Although much of the enzymology of TGT has been conducted on bacterial TGT, the data is still informative for eukaryotes as their active site sequence similarity suggests a similar mechanism.

As previously stated, eukaryotic TGT can directly incorporate queuine into the same tRNA set based on the UGU sequence requirement. Eukaryotic TGT has been classified as a homodimer in wheat germ and a heterodimer in mammals

(Boland et al. 2009; Shindo-Okada et al. 1980; Slany and Müller 1995; Walden et al. 1982). It has been studied more in depth in humans with the subunits termed queuine tRNA-ribosyltransferase 1 (QTRT1) and queuine tRNA-ribosyltransferase domain containing 1 (QTRTD1). QTRT1 has been identified as the catalytic subunit with roughly 42% sequence identity to bacterial TGT while QTRTD1 only retains 23% identity (Boland et al. 2009). QTRT1 retains the catalytic Asp residues and C-terminal zinc coordination site while QTRTD1 only retains the zinc coordination site. Initially, a deubiquitinating enzyme USP14 was identified as an essential non-catalytic subunit of TGT but later work ruled it out (Chen et al. 2010).

1.5.3 The Biological Role of Q

Initial attempts to understand the role of Q in mammals showed that germfree mice fed a special diet lacking Q for one year could survive asymptomatically with all Q-containing tRNAs devoid of Q. Upon feeding a Q diet back to the mice, the Q-tRNAs could regain their modification state (Reyniers et al. 1981). Interestingly, when germfree mice were kept on a Q free diet for only four weeks, tRNAAsn and tRNAHis were devoid of Q while tRNATyr and tRNAAsp remained fully modified (Farkas 1980) possibly suggesting a preference or hierarchy of modification. Further experimentation with Q starvation remarkably

52 demonstrated the non-essential amino acid tyrosine becomes essential when germfree mice are starved of Q (Marks and Farkas 1997). Q-starved mice devoid of tyrosine display symptoms of phenylketonuria disease accompanied by lost activity of phenylalanine hydroxylase (PAH), responsible for conversion of phenylalanine to tyrosine using the cofactor tetrahydrobiopterin (BH4). PAH function depends on an oxidation-reduction cycle of BH4 which needs to be recycled to generate a functional PAH. Mice deficient in Q and tyrosine were found to have inefficient pools of BH4 while displaying an increase in dihydrobiopterin

(BH2) thus rendering PAH ineffective. Due to the loss of PAH activity, the Q free tyrosine free mice lacked the ability to obtain sufficient amounts of tyrosine eventually leading to death (Rakovich et al. 2011).

In cancers, Q often deviates from its normal levels. Hypomodification of Q has been observed in various cancers including lung (Huang et al. 1992), ovarian

(Baranowski et al. 1994), and leukemia (Emmerich et al. 1985). However Q, or lack thereof, is not a prerequisite to cancer as Q-free mice do not promote tumorigenesis (Farkas 1980). Others have demonstrated certain tumors contain completely modified Q-tRNA (Singhal and Vakharia 1983) pointing to the complexity of cancer and suggesting the hypomodification of Q could be a downstream consequence of other factors. Antioxidant defense enzymes were found to have less activity in mice lymphoma liver but activity of the enzymes superoxide dismutase, catalase, glutathione peroxidase and reductase were stimulated by the addition of queuine (Pathak et al. 2008).

As Q occupies position 34 in the anticodon, links between Q and translation have arisen. X-ray crystallography comparing Q34-containing to G34-containing tRNA yielded no major differences in the recognition and binding of mRNA and showed the bulky cyclopentene ring does not remain inside the anticodon loop and instead protrudes outside the loop (Yokoyama et al. 1979). The presence of Q did, however, impact the overall affinity for tRNA to the ribosome. In vitro, the efficiency of tRNA binding to the ribosome was increased when Q-containing tRNA was present compared when non-Q-containing tRNA was present (Noguchi et al.

1982). It was also observed that codon preference itself could be altered by the

His His presence of Q-containing tRNA in Drosophila melanogaster. tRNA GUG prefers

His the codon CAC but can still utilize CAU whereas tRNA QUG does not show much preference (Meier et al. 1985). In another instance of preference, a stop codon in

Tobacco mosaic virus is only recognized when Q-tRNATyr is present but when

Tyr tRNA lack Q, the stop codon is suppressed by G34-tRNA generating a larger protein (Beier et al. 1984).

Another unique aspect of Q is its requirement for virulence in Shigella flexneri. In studying the main pathogenicity island in Shigella, TGT was found essential for virulence but the reason was unclear. When either Q or TGT was lacking, the translation efficiency of the mRNA encoding a virulence transcription regulator, VirF significantly decreased while still maintaining wild type copy level

(Durand et al. 1994, 2000). It was proposed that VirF may require the addition of

Q into the mRNA itself to maintain some structure suitable for initiation. Because

54 the basic requirement for bacterial TGT is a stem loop containing a UGU sequence, a computation approach was used to identify possible Q addition sites within VirF. Surprisingly, E. coli TGT was capable of exchanging queuine for G421 in vitro suggesting the possibility that TGT may have substrates outside of tRNA

(Hurt et al. 2007).

Chapter 2 : Retrograde transport is required for tRNA

maturation in T. brucei

Chapter 2 is published as: Kessler AC, Kulkarni SS, Paulines MJ, Limbach PA, Paris Z, Alfonzo JD. (2017), Retrograde nuclear transport from the cytoplasm is required for tRNATyr maturation in T. brucei. RNA Biol, Online ahead of print. Sept 13, 2017. Doi: 10.1080/15476286.2017.1377878.

Retrograde transport of tRNAs from the cytoplasm to the nucleus was first described in Saccharomyces cerevisiae and most recently in mammalian systems

(Shaheen and Hopper 2005; Takano et al. 2005; Hopper 2013). Although the function of retrograde transport is not completely clear, it plays a role in the cellular response to changes in nutrient availability. Under low nutrient conditions tRNAs are sent from the cytoplasm to nucleus and presumably remain in storage there until nutrient levels improve. However, in S. cerevisiae tRNA retrograde transport is constitutive and occurs even when nutrient levels are adequate. Constitutive transport is important, at least, for the proper maturation of tRNAPhe, which undergoes cytoplasmic splicing, but requires the action of a nuclear modification enzyme that only acts on a spliced tRNA. A lingering question in retrograde tRNA transport is whether it is relegated to S. cerevisiae and multicellular eukaryotes or alternatively, is a pathway with deeper evolutionary roots. In the early branching eukaryote T. brucei, tRNA splicing, like in yeast, occurs in the cytoplasm. In the present report, we have used a combination of cell fractionation and molecular approaches that show the presence of significant amounts of spliced tRNATyr in the nucleus of T. brucei. Notably, the modification enzyme tRNA-guanine transglycosylase (TGT) localizes to the nucleus and, as shown here, is not able to 56 add queuosine (Q) to an intron-containing tRNA. We suggest that retrograde transport is partly the result of the differential intracellular localization of the splicing machinery (cytoplasmic) and a modification enzyme, TGT (nuclear). These findings expand the evolutionary distribution of retrograde transport mechanisms to include early diverging eukaryotes, while highlighting its importance for queuosine biosynthesis.

2.1 Introduction

Following transcription, tRNAs undergo several maturation steps before partaking in protein synthesis. These include trimming of the 5’ and 3’ leader and trailer sequences, (Maraia and Lamichhane 2012; Klemm et al. 2016) 3’-end CCA addition,(Betat and Mörl 2015) and the acquisition of a number of post- transcriptional nucleotide modifications (McKenney and Alfonzo 2016). Some tRNAs also contain introns, which are removed by a specialized tRNA splicing machinery (Lopes et al. 2015). In Eukarya, many tRNA processing events are localized to the nucleus, for example end trimming and some, but not all, modifications; nuclear export may thus serve as an important checkpoint that ensures that only properly processed tRNAs make it to the cytoplasm and engage in translation.

Paradoxically, the intracellular localization of tRNA splicing varies among different organisms, and although in many cases it takes place in the nucleus, in

S. cerevisiae all the factors involved in tRNA splicing reside in the cytoplasm (Huh

57 et al. 2003; Mori et al. 2010; Yoshihisa et al. 2003). Furthermore, the localization of maturation components is intricately connected to dedicated mechanisms that export tRNAs from the nucleus (Calado et al. 2002; Kutay et al. 1998; Lipowsky et al. 1999). In S. cerevisiae tRNAs are also imported back to the nucleus from the cytoplasm (Eswara et al. 2009; Whitney et al. 2007; Shaheen and Hopper 2005;

Takano et al. 2005) by retrograde nuclear transport; a pathway that is generally constitutive. Retrograde nuclear transport is necessary for 1-methylguanosine

(m1G) formation at position 37 of tRNAPhe; a required first step in the synthesis of the hypermodified nucleotide wybutosine (yW) in S. cerevisiae (Ohira and Suzuki

2011). Beyond its role in tRNA modification, retrograde transport has also been implicated in cellular responses to nutrient deprivation. Low levels of certain nutrients (e.g. glucose or amino acids) lead to nuclear tRNA accumulation, potentially serving as a protective mechanism against tRNA degradation (Murthi et al. 2010; Hurto et al. 2007; Whitney et al. 2007; Shaheen and Hopper 2005).

Upon removal of stress, cells can then reverse tRNA nuclear retention, releasing the withheld tRNA to the cytoplasm presumably contributing to a rapid translational response. Although this process has been observed in S. cerevisiae, rat hepatoma, Chinese hamster ovary cells, and humans, little is known about its occurrence in early divergent eukaryotes and/or its evolutionary conservation

(Ghavidel et al. 2007; Barhoom et al. 2011; Miyagawa et al. 2012).

Reminiscent of S. cerevisiae, cytoplasmic localization of tRNA splicing has been recently described in the single-cell protist Trypanosoma brucei, (Lopes et

58 al. 2016)(Appendix Figure C.1, C.2) an organism that encodes a single intron-

Tyr containing tRNA (tRNA tyrosine, tRNA GUA) (Schneider et al. 1993). While studying modifications that target this tRNA, we observed that the native intron- containing tRNA is devoid of modifications, yet the mature tRNA contains the modified nucleotide queuosine (Q) at the first position of the anticodon (Q34) (Rubio et al. 2013). This suggested that this modification only occurs after tRNA splicing.

Q modification in eukarya is catalyzed by the enzyme tRNA guanine- transglycosylase (Figure 2.1 eTGT),(Howes and Farkas 1978; Boland et al. 2009) which in Xenopus has a strict preference for spliced over intron-containing tRNA

(Nishikura and De Robertis 1981). In this manuscript, we focus on TbTGT, a paralog of the canonical eukaryotic TGT and show that, unlike other eukaryotic counterparts, it localizes to the nucleus of T. brucei. In line with this observation, we also show substantial amounts of spliced Q-containing tRNA in nuclear fractions, but importantly, like in other eukaryotes, intron-containing tRNATyr is not a substrate for TGT. Taken together these findings lead to a model for a dynamic interplay between localization, splicing and tRNA modification and provides the first example of tRNA nuclear retrograde transport in an early branching eukaryote; a process that may be more evolutionarily widespread than currently thought.

2.2 Results

2.2.1 T. brucei encodes a TGT homolog

Queuosine formation in Eukarya is catalyzed by tRNA-guanine transglycosylase (TGT), which exchanges guanine for queuine at position 34 of a sub-set of tRNAs encoded with a G at position 34 (tRNAHis, tRNAAsp, tRNAAsn and tRNATyr) (Figure 2.1),(Haumont et al. 1987; Noguchi et al. 1982) while related modifications (pre-Q0 and pre-Q1) are catalyze by similar enzymes in Archaea and

Bacteria respectively (Okada et al. 1979; Watanabe et al. 1997). In general, the sequence U33G34U35 is essential for TGT recognition,(Nakanishi et al. 1994) where the almost universal U33, which forms the U-turn in the anticodon loop in most tRNAs, is part of the recognition motif. It is thought that for activity, TGT requires an unpaired G34 target (Nishikura and De Robertis 1981; Boland et al. 2009).

However, eukaryal tRNATyr universally contains an intron, which forms a base pair critical for splicing with G34 in the exon, suggesting that Q formation occurs after splicing. Prior to this work we showed that in T. brucei the tRNA splicing endonuclease and ligase localize to the cytoplasm; (Lopes et al. 2016) expectedly if Q formation follows splicing, TGT may also be a cytoplasmic enzyme.

To assess the intracellular distribution of Q, we first determined its presence in total T. brucei RNA. In Eukarya and bacteria Q is found at the first position of the anticodon (Noguchi et al. 1982; Haumont et al. 1987). We took advantage of an affinity chromatography system, which exploits the ability of amino-phenyl boronic acid (APB) to bind cis-diols, such as those present in queuosine (Kӧssel

1985). In these gels, an electrophoretic-mobility shift is observed when RNAs contain Q. This shift is in addition to that caused by the cis-diols naturally occurring at the ends of RNAs (i.e. due to the 3’ and 2’ terminal hydroxyls). We separated total RNA from T. brucei by APB-gel electrophoresis, followed by northern analysis with radioactive oligonucleotide probes specific for tRNATyr, tRNAAsp, tRNAAsn and tRNAHis. As expected, tRNATyr, -Asp, -Asn and –His displayed a slower migrating band during electrophoresis, indicative of Q (Figure 2.1 B).

As a negative control, separate samples were treated with the oxidizing agent sodium m-periodate prior to electrophoresis. This treatment converts cis- diols into cis-dialdehydes; the latter have no affinity for boronate (Kӧssel 1985). As

Glu an additional control, we also probed for tRNA , which lacks G34 and is not a known substrate for TGT (Figure 2.1 B). In this case a small shift was observed that is ascribed to that expected from the terminal hydroxyls naturally occurring at the 3’ end of most RNAs, also noticeable when comparing the oxidized control to queuosine-containing tRNAs.

We then used the sequences of the various eukaryotic TGT subunits to search the TriTryp database, revealing the presence of a potential TGT homolog in the T. brucei genome, TbTGT (Tb927.5.3520). To establish the role of TbTGT on Q biosynthesis, a portion of the coding sequence of this protein was cloned into the RNAi vector p2T7-177, transfected into T. brucei 29-13 cells and clonal lines established as described (Wickstead et al. 2002). In these cells, RNAi can be induced by addition of tetracycline to the growth media

Figure 2.1: Queuosine containing tRNA in T. brucei A. Model for the proposed queuosine (Q) insertion pathway in eukaryotes.

Queuine (Q) is taken from an extracellular source (i.e. food or gut microbiome) and inserted into tRNA by the eukaryotic tRNA-guanine transglycosylase (eTGT). B.

Total RNA extracted from T. brucei was analyzed for the presence of Q by APB- gel electrophoresis followed by northern hybridization. Probes corresponding to the four known Q-containing tRNAs in other eukaryotes (tRNAAsp, -Asn, -His and –Tyr) were used to determine the presence (+Q) or absence (-Q) of queuosine in T. brucei. A portion of RNA was oxidized by sodium periodate treatment serving as a negative control (OX); WT refers to the wild type non-oxidized sample. The non-

Q-containing tRNAGlu was used as a loading control.

A growth curve comparing uninduced and RNAi-induced cells did not show a major alteration to growth when TbTGT was knocked down (Figure 2.2 A). Five days post induction, total RNA was collected from both RNAi-induced and uninduced cells and analyzed by APB-gel electrophoresis followed by northern hybridization utilizing the same oligonucleotide probes as in figure 1B (Figure 2.2

B). A substantial reduction in Q levels was observed when comparing the RNAi- induced samples to either the uninduced or the wild type (Figure 2.2 B).

To validate the APB gel and northern results, total RNA isolated from wild type, uninduced and TbTGT RNAi-induced cells was also analyzed by LC-MS/MS

(Ross et al. 2016). A peak eluting at 25 min with a 410 m/z was observed, consistent with the presence of Q in tRNA from wild type or uninduced cells

(Phillipson et al. 1987). This peak was further analyzed by collision-induced dissociation yielding a fragmentation pattern characteristic of Q, including the neutral loss of the C-N bond yielding a m/z of 295 and the glycosidic bond breakage yields the ion 163, confirming the presence of queuosine in total tRNA from T. brucei (Figure 2.2 C-E). A similar peak was absent in RNA isolated from the TbTGT RNAi cells indicating that this enzyme is essential for Q formation in T. brucei (Figure 2.2 C).

Figure 2.2: TbTGT is necessary for Queuosine A. A growth curve of T. brucei cells where TbTGT expression has been knockdown by RNAi (TET+) compared to wild-type cells or an uninduced control (TET-). The inset shows the reduction in TbTGT levels as determined by reverse transcription

PCR (RT-PCR), where RT- refers to a control for DNA contamination where reverse transcriptase was left out of the reaction and RT+ refers to reaction to which reverse transcriptase was added prior to PCR. “Genomic” refers to a DNA positive control for PCR. B. APB-gel northern hybridization showing the effect of

RNAi knockdown of TbTGT on the queuosine content of tRNATyr. Oxidixed RNA was used as a negative control (OX).

(Figure 2.2 Continued)

C. Extracted ion chromatogram for Q 410 m/z comparing samples from TbTGT

RNAi induced (TET+) and uninduced (TET-) conditions to total wild type RNA

(WT). A peak corresponding to m/z of 410 is not (Figure 2.2 Continued)detectable in the RNAi-induced sample. D. Collision induced dissociation of the ion in C (410 m/z) reveals a characteristic fragmentation pattern of Queuosine. E. The neutral loss of C-N bond gives an m/z of 295; glycosidic bond breakage then yields the ion 163.

2.2.2 TGT localizes to the nucleus

Given our previous report that tRNA splicing occurs in the cytoplasm (Lopes et al. 2016) (Appendix Figure C.2) and that intron-containing tRNA, prior to splicing, is devoid of modifications (Rubio et al. 2013), we determined the intracellular localization of TbTGT. This was done with the goal of establishing the order of events that lead to tRNATyr maturation. In eukaryotes, some TGTs localize to the cytoplasm while others are found in the outer mitochondrial membrane

(Boland et al. 2009). TbTGT was tagged with a V5 epitope, expressed in T. brucei and analyzed by immunofluorescence with anti-V5 antibodies (Sample et al. 2015).

These cells were also stained with MitoTracker Red and DAPI, to mark the position of the mitochondrial and nuclear compartments. We found that TbTGT was strictly found in the nucleus (Figure 2.3) as judged by its co-localization with the nuclear

DAPI signal. Importantly, no significant levels of TbTGT were observed in either the cytoplasm or the mitochondrial membrane leading to the conclusion that

TbTGT is a nuclear enzyme (Figure 2.3).

Figure 2.3: TbTGT is a nuclear enzyme Immunofluorescence localization performed with cells expressing a V5-epitope tagged TbTGT. Anti-V5 antibodies were used to detect TbTGT (in green).

Mitotracker was used to stain the mitochondria (red) while DAPI stained the nuclear (N) and mitochondrial DNA (K) (blue). DIC refers to a phase-contrast image. The figure is representative of at least 5 different experiments.

2.2.3 Intron containing tRNATyr does not contain Queuosine

The intracellular distribution of TbTGT and the tRNA splicing machinery to two separate compartments then raises the possibility of a dynamic intracellular distribution of tRNATyr in the process of maturation. Either Q formation takes places in the nucleus prior to cytoplasmic export or it occurs after cytoplasmic splicing.

The former implies that intron-containing tRNA is a substrate for Q formation, contrary to the known specificity of TGT for an unpaired G34; the latter suggests that once splicing takes place in the cytoplasm, the tRNA undergoes retrograde transport to the nucleus to be modified.

To test these possibilities, we exploited the earlier observation that RNAi of

TbTrl1 leads to accumulation of intron-containing tRNA in T. brucei. Total RNA from the TbTrl1 RNAi-induced cells was separated by APB-gel electrophoresis and analyzed by northern hybridization with a probe specific for the 3’ exon of tRNATyr and compared to RNA collected from an RNAi uninduced control. This probe does not discriminate between intron-containing and spliced tRNATyr, but can differentiate between the two based on their size; as such it can detect both species simultaneously. These experiments revealed the presence of two bands in the uninduced sample; a slow migrating band corresponding to Q34-containing

Tyr spliced tRNA and a faster migrating band corresponding to G34-containing spliced tRNATyr (Figure 2.4 A). In the TbTrl1 RNAi-induced sample, a faint band corresponding to spliced Q-containing tRNA is present but in addition a strong band corresponding to intron-containing tRNA is also observed (Figure 2.4 A).

Importantly, the migration of the intron-containing tRNA band does not shift to a position indicative of the presence of Q when compared to the oxidized control when the same membrane is probed with an intron-specific probe (Figure 2.4 B).

This observation supports the view that intron-containing tRNA is not a substrate for TGT and Q formation occurs following splicing. This conclusion is also in line with our previous observation that native intron-containing tRNATyr was devoid of detectable modifications (Rubio et al. 2013).

To rule out the possibility that the lack of Q in intron-containing tRNATyr is due to some general or secondary effect from the TbTrl1 RNAi knockdown, we tested the levels of Q in the non-intron containing tRNAs that are also targets of

TGT. Interestingly, the Q content of these tRNAs appeared to increase after down regulation of TbTrl1 following RNAi (Figure 2.4 C), while the levels of the non-Q containing tRNAGlu remained unaffected (Figure 2.4 D). Although we do not fully understand these results, the observed Q increase in other TbTGT tRNA substrates after TbTrl1 knockdown may be due to one less tRNA competing for

TbTGT or alternatively it may be due to the fact that Q is in limiting concentrations in the cells and the absence of one substrate inherently leads to increase Q availability for others. However, at the moment we cannot discriminate between either possibility. Taken together, these observations support the view that intron- containing tRNATyr is not a substrate for TbTGT and the lack of Q is not due to a more general defect in Q formation related to the down regulation of an important protein.

Figure 2.4: Intron-containing tRNA lack Queuosine A. APB-gel/northern hybridization was performed on total RNA collected from

TbTrl1 RNAi induced (+) and uninduced (-) cells. The arrows indicate spliced tRNA and intron-containing tRNA generated by the RNAi knockdown of TbTrl1. Shifted

Q-containing (+Q) bands and non-Q-containing-bands (-Q) are as indicated.

Samples were treated with sodium periodate to serve as a negative Q control (OX).

The experiment was performed with a probe specific for the 3’ exon of tRNATyr. B.

The same membrane as in (A) was probed with an intron-specific probe to assess whether intron-containing tRNA contained Q. The higher band located above the intron-containing tRNATyr band is not likely related to Q, as it is also found in the oxidized control lanes. C. Detection of Q in other potential Q-containing tRNAs

(tRNAAsp, -Asn and –His) using the samples as in (A). As before, the Q-containing band (+Q) appears as a shifted band as indicated. D. The membrane was further hybridized with a probe specific for tRNAGlu serving as a loading and non-Q containing tRNA control.

2.2.4 Retrograde nuclear import is necessary for queuosine formation in tRNATyr

We then explored the second possibility that because tRNA splicing occurs in the cytoplasm and TbTGT is nuclear, then tRNATyr may require retrograde transport to the nucleus to get modified. We purified total RNA from wild type T. brucei subcellular fractions as previously described (Lopes et al. 2016). The resulting RNA was again analyzed by APB-gel electrophoresis followed by northern hybridization using the same 3’ exon-specific probe, as before.

Surprisingly, we found significant amounts of mature tRNATyr in the nuclear fraction

(Figure 2.5 A).

Given the reported cytoplasmic localization of splicing, this observation is consistent with export of the intron-containing tRNA to the cytoplasm after transcription, followed by cytoplasmic splicing and the subsequent re-import of the spliced tRNA to the nucleus to get modified. Thus, we also analyzed the different fractions for the presence of Q. We found that both non-Q and Q-containing tRNA were present in the nuclear fraction (Figure 2.5 A). Similar results were obtained when the same membrane was probed for tRNAGlu, which is not a TGT substrate, indicating that differences in tRNA levels cannot be ascribed to a general instability of tRNAs during fractionation (Figure 2.5 B). To ensure the purity of the fractions and rule out the possibility of compartment cross-contamination, the fractions were also probed for compartment-specific markers. Northern hybridization with a probe specific for SnoRNA, a nucleolar marker, revealed negligible levels of this RNA in

71 the cytoplasmic fraction with the majority found in the nuclear fraction (Figure 2.5

C). Similarly, Western blot experiments with protein samples from the same fractionation, using antibodies for the compartment-specific protein markers: anti-

Nog1 (nuclear/nucleolar marker) and enolase (cytoplasmic marker), confirmed the purity of the fractions and ruled out the possibility that the observed presence of spliced Q-containing tRNATyr in the nuclear fractions was due to fraction cross- contamination during cell breakage and purification (Figure 2.5 D).

Figure 2.5: The nucleus contains significant amounts of spliced tRNA Tyr A. Total RNA from unfractionated “T,” nuclear “N,” and cytoplasmic “C” subcellular fractions analyzed by APB-gel followed by northern hybridization.

Arrows indicate the spliced Q-containing (+Q) and non-Q-containing (-Q) tRNA bands. A portion of the RNA was treated with sodium periodate and used as a negative control for Q (OX). TbTGT RNAi+ refers to total RNA collected from

TbTGT RNAi induced cells as shown. B. The same membrane was probed for tRNAGlu, a non-Q-containing tRNA, which is used as a loading control. C. The membrane was probed for SnoRNA (nuclear/nucleolus marker) to assess fraction purity. D. Western blot with antibodies against enolase (cytoplasmic marker) and

Nog1 (nuclear/nucleolar marker) and the same subcellular fractions as above used as an additional control for fraction purity.

We also examined the role Q might play in nuclear retrograde transport.

Specifically, we determined if lack of Q was a signal for nuclear import of spliced tRNATyr to obtain Q. We purified RNA from subcellular fractions of the TbTGT RNAi knockdown; if lack of Q were a signal for retrograde transport, we would expect spliced tRNATyr to accumulate in the nucleus with reduced levels in the cytoplasm.

Northern hybridization analysis probing for tRNATyr exon revealed no major differences in the band intensities between TbTGT RNAi induced and uninduced cells (Figure 2.6 A) suggesting that lack of Q was not a signal to send tRNA to the nucleus. If Q was not directly affecting retrograde transport, it may indirectly contribute to it by altering the aminoacylation state of the tRNA. To test this, we purified total RNA from wild type, TbTGT RNAi induced, and uninduced cells under acidic conditions and performed acid-gel electrophoresis followed by northern hybridization to determine if the lack of Q alters the aminoacylation state of the tRNA. By this method, tRNA that retain their amino acid appear as a shifted band compared to deacylated tRNA allowing for visualization of aminoacylation levels.

The acid gel northern hybridization probed for tRNATyr did not reveal major differences in aminoacylation among these samples (Figure 2.6 B). These results, taken together, suggest that the absence of Q is not a signal for retrograde import itself, nor is Q a requirement for aminoacylation.

Figure 2.6: Neither lack of queuosine nor aminoacylation are signals for retrograde transport. A. Nuclear (N) and cytoplasmic (C) subcellular fractions of total RNA isolated from

TbTGT RNAi induced cells (RNAi+) were compared to similar samples isolated from wild type cells and analyzed by northern hybridization with probes specific for either the tRNATyr 3’ exon, SnoRNA or tRNAGlu as indicated. B. The aminoacylation levels of tRNATyr after TbTGT RNAi (RNAi+), an uninduced control (RNAi-) or wild- type cells were determined by acid-gel electrophoresis and northern hybridization using a 3’ exon-specific tRNATyr probe. A portion of RNA was deacylated by in a basic pH buffer and served as a negative control. Shifted bands correspond to the aminoacylated tRNA. (See Appendix C for supplementary Figure C.3)

2.3 Discussion

One of the defining features of eukaryotic cells is their extensive intracellular compartmentalization, whereby membrane boundaries provide a higher order of organization, but make it a requisite to also have specialized transport systems.

The latter ensure the purposeful movement of all sorts of molecules across selectively permeable membranes, driven by the need to provide a given function in a specific compartment. Intracellular transport dynamics may also dictate where and when many macromolecules become fully mature (Rubio and Hopper 2012).

In the case of tRNA, maturation involves the addition of, sometimes, numerous posttranscriptional modifications. These may take place at any step of the maturation pathway: In the nucleus during or immediately following transcription, in the cytoplasm following nuclear export, or even in the genome-containing organelles (plastids or mitochondria). Although for many years tRNA modification was thought to temporally occur in a linear fashion pinpointed by transcription and subsequent nuclear export to the cytoplasm, the discovery that tRNAs could be retrograde transported back to the nucleus raised the possibility that addition of modifications could be more dynamic. For example, in S. cerevisiae methylation at position 37 of tRNAPhe to form 1-methylguanosine (m1G) occurs in the nucleus, the place where the Trm5 enzyme resides. However, since Trm5 cannot act on an intron-containing tRNA and splicing is cytoplasmic, once the intron is removed the tRNA travels back to the nucleus to be modified (Ohira and Suzuki 2011).

Our findings on Q formation in T. brucei are in line with published work showing that TGT is not able to modify an intron-containing tRNA (Nishikura and

De Robertis 1981). Confocal localization studies conducted in monkey kidney cells

(Cos7) showed that endogenous TGT is associated with the outer mitochondrial membrane (Boland et al. 2009). These studies were supported by the observed efficiency in the rates of Q incorporation when tRNAs were microinjected into the cytoplasm of Xenopus laevis (Carbon et al. 1982; Haumont et al. 1987). We now show that in T. brucei a nucleus-localized TGT paralog is involved in Q formation in the nucleus and at least in the case of tRNATyr, Q formation requires the tRNA to travel from and to the nucleus to get spliced and modified (Figure 2.7).

Interestingly, three additional tRNAs are also TGT substrates but do not contain an intron; these may not require retrograde nuclear transport to be modified. It is then feasible that the differential localization of TGT and the splicing machinery, coupled to the necessity to form Q in tRNATyr, has partly provided the selective pressure to maintain a retrograde transport mechanism in this organism. The pathway described here is analogous to that seen in yeast for the biosynthesis of wybutosine providing another example of a modification requiring retrograde transport (Ohira and Suzuki 2011).

Retrograde transport is thought to play a particularly important function during times of nutritional stress. In S. cerevisiae, a decrease in the levels of amino acids, inorganic phosphate, and/or glucose causes tRNA accumulation in the nucleus (Murthi et al. 2010; Whitney et al. 2007; Shaheen and Hopper 2005).

Because retrograde transport is constitutive, such nuclear accumulation has been attributed to a reduction in the rate of re-export when nutrients are low (Takano et al. 2005). Presumably, by withholding tRNAs in the nucleus, cells can avoid tRNA degradation and re-synthesis (Chu and Hopper 2013). Upon removal of stress, however, these tRNAs are rapidly released back to the cytoplasm without the need for re-synthesis ( Murthi et al. 2010; Hurto et al. 2007; Whitney et al. 2007;

Shaheen and Hopper 2005). Our findings presented here only show the connection between retrograde transport and formation of queuosine in one tRNA.

We cannot rule out the possibility that in T. brucei retrograde transport also serves other purposes, for example to better deal with nutritional imbalances. These organisms have two different well-marked developmental stages: one in the insect vector, the other in the mammal host. Each stage demands specific metabolic adaptations from the parasites (Smith et al. 2017). For instance, in the insect T. brucei respires and primary gets all its ATP by oxidative phosphorylation, while in the host, glycolysis and substrate-level phosphorylation take over and mitochondrial functions are down regulated (Clayton and Michels 1996; Bochud-

Allemann and Schneider 2002). It is provocative to think that retrograde transport of tRNAs may play a role in such environmental adaptations.

Figure 2.7: Nuclear retrograde transport is necessary for queuosine formation in tRNATyr The figure shows a model for the maturation of tRNATyr in T. brucei. The tRNA is transcribed in the nucleus containing an 11-nucleotide long intron. In the nucleus, two or three nucleotides within the intron undergo non-canonical editing before export to the cytoplasm; editing is required for intron cleavage. Once in the cytoplasm, the edited intron-containing tRNATyr is spliced and then re-imported back into the nucleus where the nucleus-localized TGT replaces G34 for Q34.

Finally, Q-containing tRNATyr is re-exported to the cytoplasm.

Chapter 3 : The sensing of amino acids impacts the tRNA modification

3.1 Introduction

During growth, cells must withstand and cope with an ever-changing environment; they must also adapt to constant fluctuations in nutrient levels. For the parasite Trypanosoma brucei, progression through its lifecycle leads to vast differences in nutrient availability as it transitions between the mammalian host and the insect vector. To adapt to the mammalian bloodstream or the Tsetse fly, T. brucei must carefully manage differential gene expression. But while most eukaryotes have multiple means for genetic regulation, at the level of transcription, mRNA stability, or translation, kinetoplastids, including T. brucei, generally lack promoters. Transcription initiation in these organisms is thus a stochastic event and occurs in any region of the genome, generating long polycistronic transcripts that are subsequently process into discrete gene units. In turn, the bulk of the regulation of gene expression in trypanosomes occurs at the post-transcriptional level.

During its lifecycle, procyclic form (PF) T. brucei resides in the Tsetse fly vector, where it awaits transfer to a mammalian host during a bloodmeal. The bloodstream form (BF) parasite, following transfer by the insect into the mammalian host, grow extracellularly, until it is taken back up by another Tsetse fly where it can complete its lifecycle. In the bloodstream, there are two major cellular morphologies, the rapidly dividing long slender form, and the non-

80 proliferative short stumpy form which prepares for a transition back into the insect by expressing developmentally regulated genes which pre-adapt it for survival in the Tsetse fly (Vassella et al. 1997). In the transition between different lifecycle stages, T. brucei undergoes major metabolic reprogramming events to facilitate adaptation to its local environment. For example, because the mammalian bloodstream is rich in nutrients such as glucose, ATP generation in T. brucei occurs via substrate-level phosphorylation and mitochondrial function is downregulated. To produce enough ATP via substrate-level phosphorylation, T. brucei compartmentalizes the first seven steps of glycolysis into dedicated organelles called glycosomes. The number of glycosomes varies depending on the lifecycle, with BF cells harboring around 65 glycosomes per cell. This is in contrast to the PF parasites, which encounter a low-glucose environment in the

Tsetse fly, where mitochondrial function is upregulated and the number of glycosomes is reduced to ~5/per cell (Hart et al. 1984; Tetley and Vickerman

1991). The insect also contains high levels of amino acids, such as proline, which is used by T. brucei as a carbon source. The PF cells have a fully active mitochondrion, thus energy generation switches from substrate-level in the mammalian host, to oxidative phosphorylation in the insect. These metabolic changes are also accompanied by changes in mitochondria morphology, in PF cells the mitochondrion occupies approximately 25% of the cell volume, while in the BF cells only 5% (Vertommen et al. 2008). Overall, these changes are

81 reflective of the coping mechanisms to adjust energy generation and metabolic needs.

Clearly, such metabolic changes must involve careful coordination of differential gene expression, but how metabolic reprograming is achieved at a gene level is not fully understood. During its lifecycle, T. brucei must be able to sense certain environmental cues and respond accordingly. For instance, an important signal for differentiation is cold shock, brought on by transitioning from the 37oC mammalian bloodstream, to the 27oC temperature of the Tsetse fly. This temperature shift is necessary for the parasites to detect micromolar concentrations of the metabolites cis-aconitate and citrate, which in vitro can cause the parasite to differentiate into PF form cells (Brun and Schönenberger 1981;

Overath et al. 1986). Cis-aconitate serves as a signal molecule, sensed by the

PAD family of proteins (proteins associated with differentiation), which activates differentiation by inducing a signaling cascade mechanism (Szöor et al. 2006;

Dean et al. 2009). After a bloodmeal, T. brucei moves to the midgut of the fly and eventually to the salivary gland, where it attaches to the gland wall via flagellar outgrowths and prepares for transmission into the mammalian bloodstream, again by differentiating into an infective morphology. Importantly, this differentiation is marked by acquisition of a new variant surface glycoprotein (VSG) coat, so the parasite can survive transfer into the bloodstream. These adaptations to different lifestyles once again demand differences in gene expression.

In trypanosomes, some of the regulation of differential gene expression occurs via the untranslated regions (UTRs) of mRNAs. Specific motifs are found within UTRs which affect mRNA stability, while other motifs may alter translation of some mRNAs (Hehl et al. 1994; Furger et al. 1997; Hotz et al. 1997).

Alternatively, trans-splicing also generates 5’ UTRs with variable length, which often contain upstream open reading frames (uORF). More than 20% of T. brucei open-reading frames contain an upstream AUG codon in the UTR (Vasquez et al.

2014); the presence of uORFs in mammals has been correlated with lower protein expression level (Calvo et al. 2009). Along these lines, removal of such an uORF in a reporter construct generated a seven fold increase in reporter expression

(Siegel et al. 2005) suggesting that the uORFs of T. brucei may also modulate translation. However, for many genes, the UTRs are not well defined and therefore how they may or may not regulate gene expression is not exactly clear (Siegel et al. 2011).

An additional level of regulation may occur during translation beyond the

UTRs. Organisms typically favor a specific set of codons used for translation; this bias is thought to favor the use of highly expressed tRNAs and those which undergo Watson-Crick pairing for all three nucleotides during anticodon-codon interaction (Akashi and Eyre-Walker 1998). Bias for certain codons is also apparent in T. brucei which displays preference for certain codons over other synonymous ones (Horn 2008). For example, two codons, UAU and UAC, both code for tyrosine but in nuclear genes UAC is used in 69% of all tyrosine codons,

UAU in only 31%. Recent reports have suggested that organisms can exploit codon bias to regulate the expression of certain proteins, particularly in response to environmental changes and in this realm tRNA modifications play a role. For example, m5C levels, catalyzed by the enzyme Trm4, increased upon exposure of cells to H2O2 resulting in an increase in translation of UUG containing transcripts such as RRL22A, a key protein used to mediate ROS (Endres et al. 2015). Beyond cellular stress, it was observed that 16 different modifications change levels during cell division, with each division phase displaying changes to tRNA modification levels. One of the modifications altered, mcm5, enhances the translation of RNR1, a necessary component of the RNA complex which regulates cell cycle by its role in dNTP synthesis (Patil et al. 2012). Together, this shows the impact modifications have in cellular processes by taking advantage of codon bias to influence gene expression.

We hypothesize that T. brucei, given its heavy reliance on post- transcriptional gene regulation, may exploit codon-biased translation and differential nucleotide modifications for regulating protein synthesis. To test this, we have focused on the modified nucleotide Queuosine (Q) which, when present in a particular tRNA, enhances translation of near-cognate codons (Okada et al.

1976; Zaborske et al. 2014). Similar to the changing modification levels seen in yeast during cell cycle progression, In Drosophila melanogaster, the levels of tRNA modified with Q change depending on its developmental stage. For example, in the adult fly, nearly 50% of tRNATyr has Q, while in the third instar stage, only 7%

84 of tRNATyr has Q (White et al. 1973; Zaborske et al. 2014). The specific regulation of Q is thought to perhaps influence stage specific expression of genes (White et al. 1973; Zaborske et al. 2014). Furthermore, unlike other eukaryotic nucleotide modifications, eukaryotes do not synthesize Q, and instead rely on extracellular sources for Q, as demonstrated by experiments showing tRNA of a germfree mouse fed a Q free diet eventually become devoid of Q (Farkas 1980). Because

Q is therefore a micronutrient in mammals, and may be important for stage specific differentiation in D. melanogaster, we wanted to determine whether Q may have a similar role in T. brucei, possibly a stage transition and allowing T. brucei to adjust stage specific gene regulation through codon bias. Here we show the percent of

Q-modified tRNA relative to unmodified tRNA, here after referred to as “Q levels” indeed change depending on life stage of T. brucei and are dependent on the level of certain nutrients. Most significantly, Q levels can increase or decrease as a response to changes in specific amino acids, especially noticeable with tyrosine, methionine, cysteine, valine, glutamine, and glutamate. We suggest that the observed Q fluctuations may be part of a mechanism connecting translational rates to environmental cues as a form of rapid adaptation.

3.2 Results

3.2.1 The Steady-state level of Queuosine differs between developmental stages

In most eukaryotes, Saccharomyces cerevisiae being the only known exception, tRNATyr , -Asp, -Asn and -His are encoded with a guanosine at position 34 of

85 the anticodon (G34), which is post-transcriptionally replaced by Queuosine (Q) by the enzyme tRNA-guanine transglycosylase (TGT) (Katze 1978; Singhal and

Vakharia 1983; Walden et al. 1982). While studying the maturation pathway of tRNATyr, a substrate for TGT, in T. brucei, we observed that the steady-state level of the Q34 modification was only 45% (Kessler et al. 2017). This was similarly true for all the Q substrate tRNAs mentioned above. A similar observation was made in D. melanogaster where, like in T. brucei, Q-containing tRNA levels never exceed 50% under normal growth conditions of wild type organisms (White et al.

1973; Zaborske et al. 2014). However, Q levels in D. melanogaster were found to change depending on developmental stage. We decided to test if the same was true in T. brucei. We compared the Q modification levels between PF and BF cells, taking advantage of aminophenylboronic acid (APB) polyacrylamide gel electrophoresis followed by northern blot analysis, as previously described (Kӧssel

1985; Kessler et al. 2017). APB, when polymerized in gels, interacts with cis-diol groups, such as those found on Q, but also found at the 3’ end of most RNAs (i.e. the 2’ and 3’ hydroxyls), and causes a retardation in RNA migration during electrophoresis and therefore a band shift when compared to non-Q containing tRNAs. The shift in migration pattern can then be used to easily quantify relative levels of modification in a particular tRNA by northern blots. In these experiments an important negative control is a similar sample as that tested, but which has been previously treated with sodium periodate. This treatment converts the cis-diols into cis-dialdehydes; the latter have no affinity for APB (Kӧssel 1985). Importantly, in

86 some APB gels a band migrating slightly above non-Q tRNA can be seen. In all calculations, this was considered a Q-containing tRNA as it is not present in the oxidized control lane. We postulate this band could represent the hypermodified galactose-Q-containing tRNATyr and thus have a different migration compared to

Q-containing tRNATyr lacking galactose. We isolated total RNA from PF and BF stage T. brucei cells and analyzed by APB electrophoresis followed by northern hybridization. A portion of each RNA sample was also oxidized with sodium periodate prior to electrophoresis, serving as the negative control. The resulting membranes were hybridized with a probe specific for tRNATyr. In all samples, a shifted band corresponding to Q-containing tRNATyr was observed, but Q formation was not 100% in either sample, indicating that in cells regardless of developmental stage there are two populations of tRNA; one modified and the other unmodified.

Comparing the Q levels of PF and BF, a clear difference in the level of Q can be also observed with PF tRNATyr being only 50% modified as compared to 86% in

BF (Figure 3.1).

Figure 3.1:Queuosine modification levels change depending on stage RNA collected from both procyclic form (PF) and bloodstream form (BF) T. brucei was analyzed for the presence of Q by APB northern blotting (WT lane). A portion of each sample was treated with sodium periodate generating the negative control

(Ox). The membrane was hybridized with a probe corresponding to tRNATyr. The shifted bands represent tRNA with Q. For BF sample, N = 1. (Figure kindly provided by Zdeněk Paris, Institute of Parasitology, Biology Centre, Czech

Academy of Sciences and Faculty of Science, University of South Bohemia, České

Budějovice, Czech Republic)

3.2.2 Both Q-containing and non-Q-containing tRNAs partake in translation

Given that only 50% of PF tRNATyr contains Q, and our previous results showing the absence of Q does not impact aminoacylation (Kessler et al. 2017), we assessed whether both Q-containing and non-Q-containing tRNATyr engage in translation. We performed polysome analysis on PF cells utilizing sucrose gradient sedimentation as previously described (Fleming et al. 2016). APB northern blotting was then used to analyze the resulting fractions corresponding to monosomes and polysomes (Figure 3.2 A). As a control, total unfractionated RNA was used, while a portion of the unfractionated RNA was oxidized and again used as a negative control. Northern blot membranes were hybridized to the same tRNATyr-specific probe as before, revealing that in all polysomes fractions both Q-containing and non-Q-containing tRNAs are present in similar ratios, indicating that both species partake in translation (Figure 3.2 B).

Figure 3.2: Q containing tRNA are used during translation A. Polysome analysis was performed on cells grown in normal media. Polysomes were separated by sucrose density sedimentation. The resulting peaks corresponding to the individual subunits (40S, 60S), monosome (80S), and polysomes are indicated. B. Sedimentation fractions were analyzed for Q- containing tRNA by APB northern blotting. Before polysome purification, a portion of cells was used to generate a total fraction (T). This sample was further oxidized, serving as a negative control. The membrane was hybridized with a probe corresponding to tRNATyr.

3.2.3 Changes in glucose levels and other general nutrients do not affect Q levels

The utilization of both Q-containing tRNATyr, and non-Q-containing tRNATyr in translation, coupled with the fact that PF and BF stages have differing levels of

Q modification, led us to wonder whether Q may have a role in the metabolic reprogramming that occurs as the parasite transitions from one lifecycle stage to another. To support this claim, we tested whether changes in nutrient conditions might alter Q levels. We first tested whether a decrease in availability of glucose or hemin, changing conditions that T. brucei likely encounters in its lifecycle when the parasite leaves the mammalian bloodstream and transitions into the Tsetse fly would affect Q level. To test this, PF cells were first grown in media lacking one of the above-mentioned nutrients and growth was monitored for several days. We also tested growth in media lacking fetal bovine serum (FBS), a major media component. As expected, T. brucei was unable to sustain continuous growth in media lacking either hemin or FBS; both of which are essential to T. brucei growth in culture, but it was able to grow in media in which glucose was reduced (Figure

3.3 A). As described above the amino acid proline can be metabolized by PF T. brucei in the absence of glucose (Mantilla et al. 2017) and PF media is generally rich in proline. Total RNA from cells grown in each condition was then collected and analyzed for the presence of Q-containing tRNA by APB northern blot. No change in the level of Q-containing tRNATyr was observed when cells were in the absence of either hemin or glucose, but a significant decrease in Q is apparent in

91 cells grown in media lacking FBS (Figure 3.3 B, C). Lack of Q in the absence of

FBS is expected for, as previously mentioned, eukaryotes do not synthesize Q and therefore must rely on uptake from their environment, indicating that FBS is the only source of Q in the culture media. This notion is supported by studies that showing animal serum acts as the source of Q for mammalian cells grown in vitro

(Katze 1978; Katze and Farkas 1979).

Figure 3.3: Queuosine does not change during general nutrient deprivation A. Cells were monitored for growth in media depleted for either hemin, fetal bovine serum (FBS), or glucose. Because media without hemin or FBS caused cell death, growth was not monitored after day 7. B. RNA collected during the growth curve was analyzed for Q-tRNA by APB northern blotting. A representative membrane was hybridized with a probe corresponding to tRNATyr. The shifted bands represent tRNA with Q. RNA collected from Normal condition was oxidized (ox) for a negative control. Because FBS and hemin depletion caused cell death, RNA was collected at day 3. C. Average Q levels were calculated from replicate experiments. In the graph, N = 9 for Normal, while N = 4 for the remaining conditions.

3.2.4 The levels of Queuosine change in response to specific amino acids

To further examine the possibility that changes in nutrient availability may drive Q modification, we explored the role amino acid availability might have on Q levels, since T. brucei also encounters changes in amino acid availability throughout its lifecycle. To this end, T. brucei was grown in media in which the levels of one of the twenty amino acids was reduced at a time. Admittedly, it was not feasible to eliminate a particular amino acid completely, given the requirement for FBS (Figure 3.3 A). The specific composition of experimental media can be found in Appendix B. In all instances, cells were grown to mid log (6x106 – 1x107 cells/mL), washed in PBS before transfer to experimental media conditions.

Growth was compared to PF cells grown in amino acid replete media (Normal), as well as PF cells in media reduced for all 20 amino acids simultaneously (AA-).

Growth of T. brucei was monitored over the course of six days in triplicate experiments (Figure 3.4 A, Table 2).

To determine the Q-containing tRNA levels in each condition, RNA was harvested from all conditions at day six and analyzed by APB northern blotting.

Membranes were then hybridized with the same tRNATyr probe. In the majority of cases, reduction in the levels of a particular amino acid led to no significant change in the percent of Q in tRNATyr (Figure 3.5 A-B). However, individual reductions in the concentration of tyrosine, cysteine, methionine, valine, glutamine, and glutamate all resulted in an increase in the percent of Q-modified tRNATyr to 74%

(Tyr), 77% (Cys), 76% (Met), 75% (Val), 77% (Glu), 75% (Gln), respectively when

94 compared to cells grown in normal media (Figure 3.5 A). After replicate experiments, the Q-containing tRNA level was also seen elevated in several amino acid conditions such as proline, serine, and glycine (Figure 3.5 B)

Since reduction of these amino acids increased Q levels, we performed the reciprocal experiment, increasing tyrosine, cysteine, methionine, valine, glutamine, and glutamate amino acids individually. To test this, we grew T. brucei in media in which the six specific amino acids were individually doubled in concentration from that prescribed in the media formulation (See Appendix B for composition). As before, cells were grown to mid log then washed in PBS before being transferred to experimental media conditions. Cells were grown for 72 hours before total RNA was collected and analyzed by APB northern blotting to determine the percent of

Q modified tRNA. For controls, cells were also grown in normal growth media and a portion of this RNA was oxidized, serving as a negative control. Interestingly, when cells were grown in media containing higher concentrations of amino acids, the level of tRNATyr modified with Q decreased. In particular, when amino acids tyrosine, cysteine, and glutamine were increased, the Q modification percentage decreased from 45% in normal media to 23% (Tyr), 24% (Cys), and 25% (Glu)

(Figure 3.6 A,B).

Figure 3.4: Cell growth is uninhibited in media reduced for a single amino acid Cells were monitored for 6 days after transfer to media reduced for one amino acid.

In all cases cells grew as those grown in normal media and are not distinguishable above. Importantly, as a control cells were grown in media depleted for all 20 amino acids simultaneously (-aa). Growth was monitored for 6 days before collection of

RNA. N = 3

Table 2: Single amino acid reductions cause minor differences in doubling rate

Amino acid Doubling Hours Amino Acid Doubling Hours

Ala 10.2 ± 0.7 Met 9.6 ± 0.3

Arg 9.7 ± 0.5 Phe 10.3 ± 0.5

Asn 9.6 ± 0.1 Pro 9.7 ± 0.5

Asp 9.7 ± 0.2 Ser 10.4 ± 0.4

Cys 9.8 ± 0.1 Thr 9.3 ± 0.3

Gln 9.8 ± 0.3 Trp 9.1 ± 0.1

Glu 10.1 ± 0.5 Tyr 9.4 ± 0.2

Gly 9.5 ± 0.1 Val 9.5 ± 0.2

His 9.5 ± 0.2 Normal 9.3 ± 0.1

Ile 9.4 ± 0.2 Glucose 14.3 ± 0.7

Lys 9.3 ± 0.2 All AA 25.4 ± 3.1

Mean doubling rates for tested growth conditions (Figure 3.4) were calculated and are represented in hours (N = 3). The amino acids signify which one was reduced.

For reference, Normal media conditions, reduction of all amino acids simultaneously (All AA), and glucose reduction are included for comparison.

Figure 3.5: Reduction of specific amino acids causes Q-tRNA increase A. RNA collected from cells grown in media with reduced concentrations of amino acids was analyzed by APB northern blotting using a probe corresponding to tRNATyr. Individual amino acid reductions are denoted by their single letter abbreviation. The Q% represents the intensity of the shifted band over the total of each lane. OX represents oxidized total RNA collected from cells grown in Normal media conditions B. The Q level gained from replicate experiments of (A) were graphed to compare the increases seen. N ≥ 3 in all samples tested except for Glu and Gln where N = 2.

Figure 3.6: The percent of Q-tRNA decreases with additional amino acids A. Cells were grown in the presence of higher amounts of amino acids and then the Q% was determined by APB northern blotting with a probe corresponding to tRNATyr. As before, the shifted band represents tRNA with Q. Each lane represents a media containing exactly two times the concentration of the denoted amino acid found in Normal media. Cells were grown in the experimental media for 72 hours before APB analysis. Again, the Ox lane represents oxidized total RNA collected from cells grown in the Normal media condition. B. Replicate experiments of (A) were conducted and are represented. In each tested condition, N ≥ 4.

To determine how rapid the observed changes to the level of Q-tRNA occurred, RNA was collected from cells over a time course either grown in low tyrosine conditions, or in media with high tyrosine. As previously described, cells were first washed in PBS before being transferred to experimental media. Cells were collected 4, 12, 24, and 48 hours after transfer and total RNA analyzed by

APB northern blotting using a probe corresponding to tRNATyr. When cells were grown in low tyrosine media, the percent of Q-containing tRNATyr increased from

50% to 77% after a few hours and did not change in the following time points

(Figure 3.7 A). Interestingly, for cells grown in high tyrosine media, no major change in the percent of Q-tRNA was observed in 48 hours (Figure 3.7 B). As described above, after growth for 72 hours in high tyrosine media, the percent of tRNATyr modified with Q was 23%. To determine if this would change with further incubation, cells were grown in high tyrosine media for 96 hours and RNA was analyzed by APB northern blotting again using the tRNATyr probe. After 96 hours, the percent of tRNATyr modified with Q further decreased to 9% (Figure 3.7 C).

100

Figure 3.7: The Queuosine level is dependent on amino acids and time A. Cells were grown in low tyrosine media and collected over a time course. The numbers above the figure represent the hours of collection while the Q%, found below, represents the percent of Q-modified tRNATyr at each time. B. In a similar experiment, cells were grown in high tyrosine media and the percent of Q modified tRNATyr was again determined. The numbers above the figure represent the hour collected. C. Cells were grown in high tyrosine media for 96 hours and the percent of Q was determined. Membranes were all probed for tRNATyr. (Note: for data represented in this figure N = 1).

101

Since both Q-modified and unmodified tRNAs are used in translation, and the levels of modified tRNATyr are affected by amino acid availability, we examined the impact that the availability of Q-containing tRNA had on translating ribosomes.

To determine this, we performed polysome analysis on PF cells grown in conditions which caused Q-containing tRNA to either decrease or increase. As was demonstrated in a previous report, the RNAi mediated knockdown of TbTGT was sufficient to reduce the level of Q-containing tRNA to negligible levels (Kessler et al. 2017). Therefore, to generate a population of cells lacking Q, TbTGT RNAi knockdown was performed. After knockdown, cells were then used for polysome analysis as described above. Both RNAi induced and uninduced growth conditions were analyzed to determine whether the conditions altered translating ribosomes.

In the samples analyzed, both polysomes and monosomes are detectable and no major difference was seen between induced and uninduced RNAi conditions

(Figure 3.8). Next, cells grown in low tyrosine conditions for six days was then used for polysome analysis to determine how an increase in Q-containing tRNA would affect translation. As a control, cells grown in normal media conditions was used for comparison. For cells grown in low tyrosine conditions, no major difference was detected when comparing to cells grown in normal media (Figure

3.9). Together, the polysome analysis revealed neither the increase nor decrease of Q-containing tRNA levels caused any alterations to the polysome or monosome peaks, suggesting translation is not being inhibited, in agreement with earlier

102 observations of unaltered growth rates during these conditions (Figure 3.4,

Kessler et al., 2017).

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Figure 3.8: Polysomes are not affected by the loss of Q-tRNA A. Polysomes were purified by sucrose density sedimentation and compared in the presence and absence (B) of Q-tRNA by using the RNAi knockdown of TbTGT.

Uninduced conditions (TET-) and induced conditions (TET+) are indicated. The X axis represents fractions that was collected during the purification.

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Figure 3.9: Cells are actively translating when grown in low tyrosine media A. Polysomes were compared by sucrose density sedimentation from cells grown in normal media versus (B) tyrosine reduced conditions. In each case, polysomes were separated by sucrose density sedimentation. The X axis represents fractions that was collected during the purification.

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3.3 Discussion

Recently, it has become clear that some post-transcriptional nucleotide modifications favor the translation of certain mRNAs depending on their codon usage. This form of gene expression regulation effectively produces increased expression of specific genes. Translation of these modification tunable transcripts

(“MoTTs”) is heavily influenced by an organism’s natural codon bias, altering translation of an MoTT depending on the presence or absence of a specific tRNA modification. This can either enhance or repress expression of specific genes. For example, the Trm9 methyltransferase, required for the synthesis of mcm5U in

-Arg, -Gly, -Lys, -Gln, -Glu 5 2 tRNAs at U34, which can be further modified to mcm s in tRNAs

-Lys, -Gln, -Glu, (Kalhor and Clarke 2003; Johansson et al. 2008) has a role in codon bias translation of transcripts enriched with AGA and GAA codons (Deng et al.

2015). In yeast, trm9 deletion resulted in the down regulation of 54 AGA and 45

GAA enriched transcripts while up regulating 10 AGA and 10 GAA enriched transcripts when compared to wild type (Deng et al. 2015).

The levels of several other modifications change depending on the yeast cell cycle phase, and this is thought to drive cellular division forward by selectively enhancing the expression of cell-cycle genes (Patil et al. 2012). Along these lines, in D. melanogaster the levels of Q in tRNAs change depending on the stage of insect maturation (White et al. 1973). It was proposed that D. melanogaster, as in yeast, could be using such modification changes to regulate genes in developmental-state specific manner (Zaborske et al. 2014). Interestingly,

106 however, because eukaryotes do not have a biosynthetic pathway for Q, and they are instead dependent on uptake from the media, this would mean the availability of Q itself then has a role in gene regulation. Indeed, this proposal is supported by the fact that addition of extra Q to the typically low Q level, third larval instar stage of D. melanogaster, caused an increase in Q levels in tRNA (Siard et al. 1991) .

Thus, for organisms which pass through different developmental stages, the availability of Q may direct the regulation of translation of stage specific transcripts suggesting that Q levels, by themselves, may serve as a signal for development.

In the present work we have studied the possible connection between Q levels and nutrient availability in T. brucei and how it may relate to development.

Indeed, our observations show varying levels of Q modification depending on T. brucei lifecycle stage, such that, Q levels increase from 50% in PF (insect form) to nearly 100% in BF (mammal form). For an organism such as T. brucei, the transition between mammalian bloodstream and insect vector may provide differing availability of Q, thus influencing modification levels. We showed the removal of FBS caused a decrease in Q, indicating that T. brucei, like other eukaryotes, cannot synthesize Q de novo and instead relies on uptake from growth media.

Our evidence however, suggests an additional factor influencing levels of

Q-containing-tRNA besides the availability of Q as changing the concentration of several amino acids, including tyrosine, glutamine, glutamate, valine, methionine, and cysteine all resulted in observable differences in the level of Q-containing

107 tRNA. Interestingly, there is a noticeable trend between specific amino acid concentration and Q level. As amino acid concentrations are increased, Q levels decrease, while the opposite effect is seen when amino acid concentrations increase. The fact that the concentration of FBS added to growth media was kept constant argues that the observed results are not due to differences in Q availability. When cells were grown in low tyrosine conditions, the levels of Q in tRNA increased within a 4 hour window, but in high tyrosine conditions Q levels did not change much until 72 hours. This suggests that in cells grown in low tyrosine media, TbTGT itself may be either upregulated, or an inhibitory mechanism is removed, causing Q levels to rise rapidly. The fact that it took 72 hours before a significant decline in Q-containing tRNA was observed may be due to the general stability of tRNAs, with half-lives ranging between 2 to 3 days

(Phizicky and Hopper 2010). This would favor a model where stability rather than direct removal of Q is the determining factor for modification. Alternatively, TGTs can catalyze the insertion of other nucleosides besides queuine into tRNA, such as guanine, pre-Q1, and archaeosine (Okada and Nishimura 1979; Phillips et al.

2012; Stengl et al. 2005). If growth in high tyrosine conditions caused TbTGT to switch preference for another nucleoside, or increased the availability of the other nucleosides, this could likely cause a decrease in observable Q, this however is beyond what the APB-gel/ northern blot can discern. Importantly, once tRNAs are modified with Q, no exchange of Q by another nucleobase has been demonstrated for TGT (Boland et al. 2009). If specific amino acids, such as tyrosine, act as

108 allosteric effector molecules, their cellular concentrations could, however, regulate

TGT specificity allowing it to catalyze the replacement of Q for G. Additionally, as

T. brucei requires uptake of Q from the growth media, any change to the import of

Q can affect modification level. If Q is imported through pathways, that also import certain amino acids, then it is possible that such amino acids may compete with Q for the transporter. Here, changes in Q levels could be the indirect result of amino acid transporters being differentially engaged.

The link between environmental cues and gene expression has been established in T. brucei. The proliferating, long slender form of BF cells eventually transitions to the short stumpy non-proliferating BF form. In this form, cells are sensitive to temperature and the metabolites citrate or cis-aconitate; these act as signals beginning the transition to PF by allowing the expression of stage specific genes. Considering the metabolic differences between insect vector and mammalian host, there likely exist multiple signals that may influence cellular metabolism and gene expression. Metabolically, the nutrients available to T. brucei throughout its lifecycle are drastically different depending on the developmental stage. In the bloodstream, the parasite has access to large amounts of glucose while this is not the case in the midgut of the fly which is an environment low in glucose but abundant in amino acids (Mantilla et al. 2017; Balogun 1974).

Importantly, in our media with reduced glucose concentrations, the percent of Q- containing tRNA remained unchanged, however changes in amino acids did result in significant alterations in the modification of Q. Our data shows that the percent

109 of tRNA modified with Q increases rapidly after exposure to media with reduced amino acids. Therefore, T. brucei may sense amino acid concentration as another means to assess when it is transitioning between lifecycle stages and rapidly assist the expression of stage specific genes by exploiting codon bias. This work raises interesting questions as to whether or not the observed changes in Q can immediately reflect changes in gene expression. It is clear that the parasite can sense changes in certain nutrients in the growth media and in turn change the levels of Q in tRNAs. Currently, we do not know the sensing mechanism but suggest, that like in Drosophila, T. brucei may exploit codon-biased translation to regulate gene expression and cope with environmental changes in the availability of certain nutrients.

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Chapter 4 : Concluding remarks and future directions

4.1 Summary

As T. brucei makes the transition from life in the Tstetse fly to the human bloodstream, it undergoes metabolic reprograming, adapting to the bloodstream environment while only utilizing post-transcriptional mechanisms of gene regulation. How this occurs, and by what means, is not well understood. Here, we propose tRNA maturation and modification may have roles in impacting gene expression in T. brucei. After transcription, a series of processing events, such as splicing and modification, are necessary to render tRNAs functional in translation.

As previously mentioned, tRNATyr contains an intron in T. brucei, therefore splicing is essential. Since essential components of the splicing machinery localize to the cytoplasm, tRNA splicing is a cytoplasmic event (Appendix C). In Chapter 2, we revealed the modification enzyme TbTGT is localized to the nucleus, and because of this, tRNATyr must re-enter the nucleus after splicing to receive the Q modification. This outlined the first instance of tRNA retrograde transport in T. brucei, and provided another example of a modification that is dependent on transport dynamics, suggesting a connection between splicing, intracellular tRNA transport, and the Q modification. In yeast, intracellular tRNA transport can be affected by nutrient availability. Because of this, we addressed whether certain nutrient conditions might influence the Q modification. Unlike the situation in yeast, however, we observed that the availability of specific amino acids influence the level of Q modification seen in tRNAs that are TGT substrates, but how amino acid 111 levels are sensed and such signal transduced into tRNA modification levels remains unclear. Because Q may have a role in codon-biased translation, in

Chapter 3 we therefore hypothesized that changing Q levels may be important for a gene expression regulation mechanism used by T. brucei. More specifically, the splicing, modification, and intracellular transport of tRNATyr are also potential regulation points for tRNATyr function. We believe the work outlined here can provide new insight into how intracellular transport is related to modification, and provide another mechanism T. brucei may use in the transition between insect and mammalian bloodstream. Beyond our findings outlined here however, there are still unanswered questions and interesting connections further discussed below.

4.2 Exploring connections between amino acids that influence Q levels

Typically, in normal growth conditions, roughly 45% of the population of tRNATyr has Q, while the other 55% does not. Similar ratios are observed for the three other TGT substrate tRNAs (Asp, Asn, His). Interestingly, changes in the concentrations of tyrosine, cysteine, glutamine, glutamate, methionine, and valine affect the level of Q modification in these tRNAs while other nutrients only had negligible effects on Q levels.

In humans, phenylalanine, tryptophan, threonine, valine, methionine, leucine, isoleucine, lysine, and histidine are all essential amino acids and must be obtained from the diet. Bioinformatic approaches suggest the same is true in T. brucei, which lacks biosynthetic genes for these 9 amino acids. In addition, different from many eukaryotes, T. brucei also lacks the enzyme phenylalanine 112 hydroxylase (PAH), which converts phenylalanine to tyrosine, therefore tyrosine is also an essential amino acid (Berriman et al. 2005). RNAi mediated knockdown of transporters for arginine and lysine resulted in cell death (Mathieu et al. 2017) while in the related organism, Leishmania major, biosynthetic pathways for the synthesis of arginine and serine or any of the aforementioned 9 essential amino acids could be identified (Payne and Loomis 2006). Furthermore, T. brucei requires cysteine in the media for growth in vitro (Duszenko et al. 1992). Comparative metabolomics of media before and after cell growth supported the requirement for cysteine and further demonstrated a need for glutamine, phenylalanine, and tryptophan (Creek et al. 2013). Taken together, almost all of the amino acids, which affect Q levels

(Tyr, Val, Glu, Gln, Cys, Met), may in fact be essential.

Interestingly, synthesis of trypanothione, necessary for redox homeostasis in T. brucei (Comini and Flohé 2013), involves several of the amino acids which affect Q levels. During synthesis, methionine acts as a sulfur donor for cysteine biosynthesis via the reverse trans-sulfuration pathway (Manta et al. 2013). Once cysteine is obtained it is used along with glycine and glutamate for the generation of glutathione. Trypanosomes however, do not utilize glutathione as a redox- cofactor and they lack traditional enzymes of the pathway. Instead, glutathione and spermidine are precursors for the eventual synthesis of trypanothione (Comini and

Flohé 2013). Considering that previous work has correlated Q levels with redox homeostasis, by promoting the activities of antioxidant enzymes (Pathak et al.

2008), perhaps the same connection exists in trypanosomes. If this were the case,

113 altering the availability of amino acids involved in the synthesis of trypanothione might additionally lead to changes in Q level.

4.3 Potential mechanisms influencing Q levels

As mentioned previously, eukaryotic organisms do not synthesize Q and instead rely on extracellular sources of Q, for example that synthesized de novo by gut bacteria, or from food. Therefore, extracellular Q must be transported across the cell membrane in order for TGT to catalyze the generation of Q-containing tRNA, but the Q transporter has not been identified. If Q comes into cells via the same transporter also used for amino acids, both Q and the amino acids could compete for transport. In a similar situation, growth of Chlamydiales trachomatis in the presence of high amounts isoleucine, leucine, methionine, or phenylalanine is inhibited, as these amino acids act as competitive inhibitors against the transport of valine, an essential amino acid (Braun et al. 2008). Therefore, if specific amino acids, such as glutamine, glutamate, tyrosine, methionine, valine, and cysteine compete with Q, any alteration in their concentrations could then be reflected in the level of Q modification.

Alternatively, it is possible the observed fluctuations in Q levels may have to do with specific membrane transporters being differentially expressed in varying amino acid conditions. The parasitic protozoa, including T. brucei, have lost the ability to synthesize purines as they lack many of the necessary genes, and instead must rely on the salvage of purines from their host (Landfear et al. 2004).

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Therefore, many genes in T. brucei encode potential nucleoside or nucleotide transporters. Transporters can be grouped into two major groups, those expressed in both PF and BF, and those expressed solely in BF (Sanchez et al. 2002). If Q transporter expression is lifecycle dependent, this may explain why the percent of

Q modification is different in PF and BF and may help explain the change in modification levels. Because the availability of amino acids changes during T. brucei lifecycle, this may be one of the key signals T. brucei utilizes to sense its environment, subsequently influencing the expression of a stage specific Q transporter. Further work in the identification of transporters may reveal interesting connections between Q and the currently orphan transporters such as TbNT3 and

TbNT4 (Sanchez et al. 2002). It would be interesting to see how the expression of such transporters changes depending on growth conditions.

We must also consider the possibility that the change in the level of Q- containing tRNA is due to a change in tRNA intracellular transport dynamics. In

Chapter 2, we attempted to answer this by asking whether the lack of Q was a signal for re-import. As described, we concluded that the lack of Q is not a signal for sending the tRNA to the nucleus, but this did not rule out the possibility that other factors regulate tRNA dynamics. In yeast, starvation for nutrients such as amino acids changes the intracellular transport dynamics of tRNA by slowing the re-export of tRNAs to the cytoplasm, thus leading to nuclear retention. If a similar mechanism were to exist in T. brucei, nuclear retention of tRNAs might cause an increase in the percent of tRNA modified with Q. This could potentially explain why

115 the growth of T. brucei in amino acid reduced media conditions leads to an increase in the percent of tRNA modified with Q but does not as easily explain why the reciprocal experiment led to a decrease in Q.

Lastly, it is also possible that difference seen in Q levels is due to the TbTGT enzyme itself. The six amino acids could be interacting with TbTGT modulating its function. This could be tested in vitro by studying the activity of the recombinant enzyme in various concentrations of amino acids.

4.4 The connection between Q and folate metabolism

One of the known interesting connections between metabolism and Q occurs in mice where Q is an nonessential micronutrient, but becomes conditionally essential when mice are simultaneously starved for Tyrosine and Q.

This is due to loss of PAH activity, the enzyme responsible for the conversion of

Phenylalanine to Tyrosine (Farkas 1980; Marks and Farkas 1997; Rakovich et al.

2011). The mechanism behind this is unclear, but studies revealed that loss of

PAH activity was due to accumulation of dihydrobiopterin (BH2), the oxidized form of the essential PAH cofactor tetrahydrobiopterin (BH4) (Rakovich et al. 2011). The enzyme dihydrofolate reductase (DHFR), is capable of salvaging BH2 and catalyzing its reduction to BH4, however its main function is to convert dihydrofolate into tetrahydrofolate. It was therefore postulated that Q may somehow influence folate metabolism, thereby slowing the conversion of BH2 to

BH4, leading to a decrease in activity of PAH and a reduction in tyrosine production

116 from phenylalanine. Because of the complexity and multitude of potential factors influencing PAH, it remains difficult to determine the reason Q is necessary for this pathway. T. brucei perhaps presents a unique situation; there is no identifiable

PAH gene, and therefore no pathway for phenylalanine to tyrosine conversion although they maintain DHFR (Sienkiewicz et al. 2008). In addition, our preliminary data suggests growth of cells in low tyrosine media conditions while simultaneously knocking down TbTGT by RNAi does not produce a growth phenotype. Because of the undefined concentration of tyrosine present in FBS, it would useful to determine the necessity of Q for cells grown in media containing a concentration range of tyrosine. To determine whether there is a connection between folate metabolism and Q, it would be interesting to test the requirement for DHFR for function in T. brucei in the context of a tyrosine depletion experiment and in the presence and absence of TbTGT.

4.5. Our Model

Our overall model involves the maturation of tRNATyr, the only tRNA with an intron, and a Q-containing tRNA. Like other tRNAs, it is transcribed in the nucleus where end maturation occurs. At this point, the intronic sequence of tRNATyr must undergo non-canonical editing in the nucleus at 2-3 positions for the splicing reaction to take place (Rubio et al. 2013). This edited tRNA is exported to the cytoplasm where it encounters the splicing machinery and undergoes splicing

(Lopes et al. 2016). After splicing, tRNATyr is re-imported into the nucleus where

TbTGT catalyzes the modification of Q (Kessler et al. 2017). Finally, tRNATyr is 117 exported back to the cytoplasm for use in translation. As cells encounter changing concentrations of specific amino acids (such as Tyr, Glu, Gln, Cys, Met, Val), they change the modification percent of Q-containing tRNA, which we believe, may influence gene regulation (Chapter 3). We hypothesize that, together, this may assist T. brucei in the transition between host environments.

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Appendix A: Materials and Methods

140

A1: RNA purification by guanidine isothiocyanate

1. Spin cells 5000 RPM for 10 min

2. Rinse pellet twice with 1x PBS (can move pellet to 2 mL tube with 200 µL

PBS)

3. Resuspend pellet in 200 µL PBS first then:

a. 600 µL Solution D (Add this last after pellet is completely resuspended) b. 60 µL 2M NaOAc c. 600 µL H2O-phenol d. 180 µL chloroform/IAA 4. Vortex 1 min+

5. Ice 10 min.

6. Spin 20 min max speed

7. Transfer supernatant to new tube and add:

a. 1 µL glycogen b. Equal volume isopropanol 8. Spin at max speed 30 min

9. Dissolve pellet in 300 µL TE

10. Extract with 300 µL Tris-phenol

11. EtOH precipitate + 4 µL glycogen

12. Resuspend in TE. (generally 20 -100 µL depending on pellet size)

13. Check concentration

Solution D 4M guanidine isothiocyanate 25mM sodium citrate pH 7.0

141

A2: RNA purification by acid phenol

1. Spin cells 5000 RPM for 10 min

2. Rinse pellet twice with 1x PBS in 2 mL tube

3. Resuspend pellet in 100 mM NaOAc pH 4.5

4. Add equal volume of acid phenol equilibrated with 100 mM NaOAc pH 4.5

5. Extract with acid phenol again.

6. Ethanol precipitation

7. Resuspend RNA pellet in 100 mM NaOAc pH 4.5

142

A3: Standard polyacrylamide gel electrophoresis for RNA separation

1. Pour 8% 8M urea polyacrylamide gel (60 µL APS + 10 µL TEMED for

10mL)

2. Typical RNA loading 8 – 12 µg RNA per well. EtOH + glycogen precipitate

to obtain small volume for loading. Resuspend pellet in 7 µL urea load dye

3. Heat samples 90°C for 10 min in urea loading dye. Brief centrifuge before

loading gel.

4. Electrophoresis in 1x TBE or 1x TAE to correlate with polyacrylamide gel.

5. Carefully wash wells with buffer filled syringe multiple times.

6. Dispense sample slowly so dye forms thin layer on bottom of well and

does not spread out.

7. Run gel at 85-100V until bromophenol blue dye is at the bottom of the gel

(1-2 hour). For better separation, run bromophenol blue off the gel and

proceed until xylene cyanol is near the bottom.

8. To visualize RNA, add gel to small dish containing buffer + EtBr for 10

min.

9. Gel can be washed in buffer to remove traces of EtBr before visualization.

143

A4: Acid gel electrophoresis for RNA separation

1. Make 12% 8M acid urea polyacrylamide (30 mL working stock)

a. 7.5 mL 40% acrylamide stock b. 12.6 g Urea c. 3 mL 3M NaOAc d. Bring Volume up to 30 mL 2. Add 150 µl APS and 46 µl TEMED for 10 mL

3. Deacylate negative control RNA in 100mM Tris-HCl pH 9.0, 30min, 37°C.

4. Load 10 µg RNA per well. EtOH + glycogen precipitate to obtain small

volume for loading. Resuspend pellet in 7 µL urea load dye in 100 mM

NaOAc pH 4.5

5. Electrophoresis in 100mM NaOAc pH 4.5 buffer

6. Carefully wash wells with buffer filled syringe multiple times.

7. Pre-run gel for 30min with loading dye.

8. Dispense sample slowly so dye forms thin layer on bottom of well and

does not spread out.

9. Run gel at 50V at 4°C for 48hr

10. To visualize RNA, add gel to buffer + EtBr for 10 min.

11. Gel can be washed in buffer to remove traces of EtBr before visualization.

12. When doing northern blotting, find the target RNA by UV visualization then

place blotting membrane overtop.

13. Northern blotting can be carried out as described in Northern blotting: A9

14. *Important - RNA must remain in acid buffer at all times 144

A5: APB gel electrophoresis for RNA separation

1. Pour APB gel by adding 60 µL APS + 10 µL TEMED to 10 mL 1x APB

mix.

2. Deacetylate all samples (10-15 µg) in 100mM Tris-HCl pH 9 for 30

minutes at 37°C.

3. Ethanol precipitate samples and wash pellets with 70% ethanol.

4. Resuspend pellets in 7µL of urea load.

5. Add oxidized control to gel.

6. Heat RNA for 10 minutes at 90°C.

7. Run gel in 1x TAE at 80V at 4°C until xylene cyanol dye is near bottom

approximately 5 hr.

8. Re-cycle buffer during run time as necessary.

9. APB gel is easily stuck to electrophoresis plates. Separate gel from plates

submerged under 1x TAE buffer.

10. To visualize RNA, add gel to small dish containing buffer + EtBr for 10

min.

11. Gel can be washed in buffer to remove traces of EtBr before visualization.

145

Sodium periodate oxidation of RNA

1. Resuspend deacylated tRNA pellet in H2O and incubate RNA in 50mM

NaOAc pH 4.5-5.2 and 2.5 mM NaIO4 for 2 hr at 37°C in the dark. (we

generally use large amount of RNA so as to not have to make this often;

50uL volume)

2. Quench reaction with 2mM glucose and incubate 30 min 37°C in the dark.

3. Purify by Sephadex G-25 column

4. Ethanol precipitate, wash 70% ethanol, take concentration

5. Take the amount that is needed for the gel when ready.

1x APB mix

42 g UREA 2 mL 50x TAE 31.7 mL 30% acrylamide stock (acrylamide 28.38g/100mL and bis-acrylamide 1.62g/100mL) H2O up to 100 mL

*Dissolve APB for 60mg/10mL of mix

146

A6: SDS Polyacrylamide gel electrophoresis for protein analysis

1. Pour bottom of stacking gel first:

4 mL H2O 3.3 mL 30% acrylamide mix 2.5 mL 1.5 M Tris pH 8.8 0.1 mL 10% SDS 0.1 mL APS 0.01 mL TEMED

2. After pouring, layer 0.2 mL of 0.1% SDS on top of gel to level the bottom

layer

3. Once polymerized, pour off the 0.1% SDS, dry remaining liquid with paper

towel.

4. Pour, then layer the top of the stacking gel:

2.1 mL H2O 0.5 mL 30% acrylamide mix 0.38 mL 1.5 M Tris pH 6.8 0.03 mL 10% SDS 0.03 mL APS 0.003 mL TEMED

5. After polymerized, gel can be run in 1x SDS-PAGE buffer

6. Gels are run for 1-2 hr at 100V, depending on the separation required

7. If doing a western blot, see “Western blot” A10 protocol, if staining,

continue below:

a. Add gel to Fixing solution overnight (50% MeOH, 10% acetic acid v/v) b. Stain gel in Staining solution 30 min to 1 hr. (0.025% Coomassie brilliant blue, 10% acetic acid w/v) 147

c. Transfer gel to De-staining solution and leave with agitation until desired level of destaining (10% acetic acid v/v).

10x SDS PAGE buffer

30.0 g Tris base 144.0 g glycine 10.0 g SDS Bring final volume to 1 L with water

148

A7: End labeling of northern blot probes

1. Mix 1µl of a 40 µM probe with 1µl of gamma ATP, 1 µl PNK buffer, 1 µl

PNK in a 10 µl total volume reaction brought up with water.

2. Leave reaction for 1-2 hr at 37°C.

3. Purify labeled probe by G25 column purification.

4. To make column, add 30 µl of glass beads and 300 µl Sephadex G25 to

500 µl tube placed inside 1.5mL tube. Make sure to poke a hole in the

bottom of the smaller tube with a needle.

5. Centrifuge 4.4RPM for 1min to pack column

6. Discard bottom tube with liquid.

7. Add column to new 1.5 mL tube

8. Add sample to column.

9. Spin again 4.4RPM for 30 seconds.

10. Check the liquid with a Giger counter to make sure probe is there.

11. Before use, heat probe to 100°C for 10 min and chill on ice for 1 min.

149

A8: Northern blot utilizing electrotransfer

1. Electrotransfer in 0.5x TBE (or TAE if used previously). Wet sponges, filter

paper, and Zeta Probe membrane (BioRad) before assembling. Cut the

bottom right corner of the membrane before placing on gel for orientation.

2. Transfer at 80 V for 2 hours.

3. UV crosslink membrane for 1 min.

4. Bake membrane at 80°C for 30 min.

5. Denature SS-DNA for 10 min at 90°C then add to pre-hybridize (1 µL/

1mL) solution on membrane.

6. Pre-hybridize in rotating oven for 30 minutes.

7. Denature probe for 10 min at 90°C. Quickly centrifuge and place on ice for

1 min.

8. Add probe to pre-hybridization solution and leave overnight.

9. Pour out labeled probing solution to small 15 mL conical tube for decay.

10. Add 5mL of Oligo Rinse and briefly wash membrane. Discard oligo rinse

into hot sink.

11. Add 5mL of Wash 1 solution and wash for 20 min in hybridization oven.

12. Dump wash down hot sink.

13. Add 5mL of Wash 2 solution and wash for 20 min in hybridization oven.

14. Dump wash down hot sink.

150

15. Cut filter paper slightly larger than membrane. Wet with water. Place

membrane on top of wet filter paper (Do not let membrane with probe dry

out)

16. Place this in a plastic bag or surround in saran wrap.

17. Place in Phoshpoimager cassette with screen overnight. Screen should be

blanked for 10 min just before use.

18. Visualize using phosphoimager scanner

19. Place membrane in 50 mL conical tube with stripping solution at 90°C for

20 min to remove probe.

20. Membrane can be probed again after stripping.

Northern blot solutions are adapted from Bio-Rad Zeta-Probe

20x SSC 175.3g NaCl 88.2 g Sodium Citrate Adjust volume to 1 L and pH to 7.0

100x Denhardt’s solution 2% Bovine Serum Albumin 2% Polyvinylporrolidone 2% Ficoll

Prehybridization 5x SSC 20 mM Na2HP04, pH 7.2 7% SDS 1x Denhardt’s *SS DNA added separately before hybridization.

151

Wash 1 3x SSC 10x Denhardt’s 5% SDS 25 mM NaH2P04, pH 7.5

Wash 2 1x SSC 1% SDS

Stripping Solution 0.1x SSC 0.5% SDS

152

A9: Western blot, utilizing electrotransfer

1. Protein gel is transferred to a membrane in 1x PAGE, 2hr at 75V.

2. Block the membrane in 5% dry non-fat milk resuspended in 1xPBS-

TWEEN for 30 min with shaking.

3. Add the primary antibody diluted in 1xPBS-TWEEN milk solution for 1 hr

at room temperature with shaking.

4. Remove the primary solution and wash the membrane 3x for 5 min each

with PBS-TWEEN.

5. Add the secondary antibody diluted in 1xPBS-Tween milk solution for 1 hr

at room temperature with shaking.

6. Remove the secondary solution and wash the membrane 3x for 5 min

each with PBS-TWEEN.

7. Remove liquid from membrane and add Clarity Western Solutions in 1:1

ratio.

8. After 5 minutes, remove solution and image.

PTM (PBS- TWEEN MILK) 50 mL PBS Twee 2 g dry milk

PBS TWEEN 1x PBS 0.5% TWEEN

153

A10: Nuclear and cytoplasmic fractionation

1. Harvest 1L of cells at 5 x 106 to 3 x 107 cells/mL. If concentration is too

great, the nuclei are difficult to break.

2. Set aside 50 mL of cells and use for total RNA and protein purification.

3. Pellet 1L of cells at 5,000 RPM for 10 min at 4°C.

4. Wash pellet in 1x PBS and spin again 5,000 RPM for 10 min at 4°C.

5. Resuspend cells in appropriate amount of Lysis Buffer to reach a cell

density of 8x108 cells/mL. Start resuspension with ¾ of the calculated

volume of Lysis Buffer and observe cellular swelling with a microscope.

There should be a mixed population of swollen/ unaffected cells.

6. Slowly add the remaining volume of Lysis Buffer while checking cellular

swelling with microscope. Try to achieve uniform population of cellular

swelling without breaking.

7. Break cells using Stanstedt Machine at 20 psi.

8. Check cellular breakage with microscope. Nuclei are released and can be

seen with cellular debris floating. Do not pass through machine twice

unless absolutely necessary as this will begin to rupture nuclei.

9. Centrifuge lysate at 2,500 x g for 20 min at 4°C.

10. The resulting supernatant is the cytoplasm while the pellet is the nuclei.

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Cytoplasmic purification continued

1. Spin cytoplasm at 25, 000 RPM at 4°C

2. Set aside a portion for protein fraction. Add SDS load and store at -20°C.

3. For RNA, add equal volume of water saturated phenol. This can be carried

out in 2 mL tubes and centrifuged at max speed in a microcentrifuge. Any

remaining unused cytoplasmic fraction can be stored at -20°C.

4. Ethanol precipitate the top layer. The resulting pellet can be resuspended

in TE and used as cytoplasmic RNA.

Nuclei purification continued

1. Resuspend the pellet in 20 mL of Wash Buffer and centrifuge at 2,500 x g.

2. Repeat this wash twice

3. Take a small portion of the pellet into a new 2mL tube. Resuspend in

water and add SDS load. This will be the nuclei protein fraction.

4. The remaining pellet can be resuspended in a small volume of PBS and

used for a guanidine isothiocyanate RNA prep as described previously.

Lysis Buffer 0.2 mL 0.5 M PIPES pH 7.4 0.2 mL 1.0 M CaCl2 6.38 mL 7.825 M Hexylene Glycol Bring volume to 100 mL with water

Nuclei Wash Buffer 0.2 mL 0.5 M PIPES pH 7.4 0.5 mL 1.0 M CaCl2 6.38 mL 7.825 M Hexylene Glycol 42.78 mL 60% sucrose Bring volume to 100 mL with water 155

A11: Immunofluorescent microscopy

1. Centrifuge mid log cells at 1,300 x g for a target density of 2x107 – 1x108

in 1 mL of fresh media.

2. Transfer 1 mL of suspended cells to 9 mL of media.

3. Add 4µl of 500 µM MitoTracker Red to the cells

4. Incubate cells with MitoTracker Red for 30 min at 27°C

5. Centrifuge cells at 1,300 x g for 10 min

6. Wash with 2 mL PBS and centrifuge 1,300 x g for 10 min

7. Remove supernatant and carefully resuspend cells in 200 µl of PBS

8. Add 200 µl of 7.4% formaldehyde. Use PBS to dilute formaldehyde to

working percentage.

9. After adding formaldehyde to cells, gently invert to mix.

10. Add 50 µl of cell mix to a clean microscope glass slide. Spread cells to

cover an area approximately 2x2 cm.

11. Incubate at RT for 15 min inside dark humid chamber. Can use small box

with moist towel. Careful not to let the moist towel come in contact with the

cells.

12. Carefully pipette liquid from slide and begin 3x wash with 100 µl PBS

a. Tilt slide at an angle and let PBS wash cascade over cell spot. b. Pipette off droplets of PBS that remain c. Repeat the washes 3x 13. To permeabilize cells, add 100 µl of 0.1% Triton X-100. Dilute with PBS to

generate the correct dilution. 156

14. Incubate for 10 min in the humid chamber.

15. Pipette off the liquid and begin another series of 3x washes as above.

16. Block cells with 100 µl 5.5% FBS in PBS and 0.05% Tween for 1hr.

17. Pipette the liquid and carry out 2 washes as described above.

18. Add 50 µl of 1° antibody diluted in 3% BSA, 1xPBS, 0.05% Tween

19. Incubate at RT for 1 hr inside humid chamber

20. Pipette off liquid, wash 3x with 100 µl of 1x PBS-Tween

21. Wash 2x with 100 µl 1x PBS

22. Add 50 µl of 2° antibody diluted in 3% BSA, 1xPBS, 0.05% Tween (If DAPI

and MitoTracker Red are used, 2° antibody must be FITC)

23. Incubate at RT for 1hr inside humid chamber

24. Pipette off liquid, wash 3x with 100 µl of 1x PBS-Tween

25. Wash 2x with 100 µl 1x PBS

26. Add 100 µl of diluted DAPI (1:1000 in PBS) for 1min

27. Wash 2x with 100 µl 1x PBS

28. Pipette off liquid and air dry slide in dark.

29. Mount cells with 15 µl of VectaShield onto cells, then add coverslip

30. Make sure to remove any trapped air by gently pressing on coverslip

31. Edge can be sealed with nail polish

32. Completely air dry slide in dark for 30 min

157

A12: Polysome Analysis

1. This procedure can effectively be carried out with 50 mL of starting

culture. Begin with cells at 6 x 106 to 2 x 107 cells/mL.

2. Centrifuge cells for 10 min at 5,000 RPM.

3. Wash pellet in 1x PBS for 10 min at 5,000 RPM.

4. Resuspend pellet in 780 µl 1x polysome buffer with 100 µg/mL

cycloheximide, 2mM DTT, 1x protease inhibitor.

5. Lyse cells with the addition 20 µl 10% NP-40. Incubate 5 min on ice.

6. Centrifuge lysate at 15,000 x g for 10 min at 4°C.

7. Take OD readings of the clarified lysate and calculate loading

concentration if necessary. 500 µg to 1 mg total RNA are typical loading

concentrations.

8. Carefully load on a linear 10-50% sucrose gradient. (To prepare gradients,

layer 4.4 mL 10% onto 4.4 mL 50% sucrose and lay flat at 4°C. Carefully

tilt gradients upright after 4 hr)

9. Centrifuge at 36,000 RPM in SW41 Ti Beckman rotor for 2 hr at 4°C.

10. Gradients can be harvested using syringe pump with UV absorbance

detector (UA-6, ISCO) measuring A254 through the gradient. Fractions can

be collected in 1mL aliquots.

11. If RNA is desired from fractions, Add equal volume Tris-phenol, centrifuge

15 min at 13,200 RPM.

158

12. Ethanol precipitate supernatant with 1/10 volume 3M NaOAc, 2x volume

EtOH, and 3µl glycogen.

10x Polysome Buffer 10 mL 1M Tris-HCl pH 7.5 20 mL 3M KCl 10 mL 1M MgCl2 Bring to 100 mL with water

60% Sucrose (100 mL) 85.71 mL 70% sucrose stock 10 mL 10x polysome buffer Bring to 100 mL with water

50% Sucrose (50 mL) 35.71 mL 70% sucrose stock 5 mL 10x polysome buffer Bring to 50 mL with water

10% Sucrose (50 mL) 7.14 mL 70% sucrose stock 5 mL 10x polysome buffer Bring to 50 mL with water

159

A13: Reverse Transcription-PCR

1. Isolate RNA by guanidine isothiocyanate protocol

2. RQ1 DNAse treatment

a. 5 µl of 10x RQ1 DNAse buffer b. 1 µl of RQ1 DNAse c. 5 µg of RNA d. Bring volume to 50 µl with water 3. Incubate reaction at 37°C for 1hr to overnight

4. Add 50 µl of water, then phenol extract with tris saturated phenol

5. Ethanol precipitate with 2x volume ethanol, 1/10th volume 3M NaOAc, 1 µl

glycogen

6. Resuspend pellet in 20 µl of water

7. Recommended step: attempt a PCR to check the DNAse treatment. If

successful there should be no product. Be sure to include a positive

control.

8. Anneal RT primer. (make sure to calculate how many reactions you need)

a. 1 µl of 2 pmol RT oligo b. 5 µl of RNA c. 5 µl of water 9. 70°C for 10 min followed by chill on ice 15 sec.

10. Add the following master mix to the reaction tube:

a. 4 µl of 1st strand buffer b. 2 µl of 0.1 M DTT c. 1 µl of dNTP’s (10 mM) 11. Set reaction for 2 min at 42°C

160

12. Add 1 µl of Reverse Transcriptase to the RT+ tubes. (You will need an

RT+ and RT- tube for each sample)

13. Leave reaction 42°C for minimum of 1hr

14. 2 µl of reaction can be used as template for a 100 µl PCR reaction

161

A14: Preparing tRNA for nucleoside analysis: HPLC and/or LC-MS/MS

1. Purify total RNA by the guanidine isothiocyanate method

2. Mix RNA (roughly 50-200 µg) with clear urea load buffer

3. Pour an 8% urea acrylamide gel as described in “standard polyacrylamide

gel electrophoresis” but use a comb with large wells, or one very large

well.

4. Load 50-200 µg of RNA in the well depending how large the wells are

5. In a small well next to the wells used for samples, load a small amount of

urea load + dye. This will be used as a marker to judge separation

6. Once adequate separation is achieved (roughly 2 hr at 100V), visualize

RNA bands by UV shadow

7. Cut out bands of interest and place in 0.3 M NaOAc overnight

8. The following day, collect the supernatant and ethanol precipitate. Add 0.3

M NaOAc to the tube once more and repeat this step.

9. Digest pooled RNA samples (in 20 µg reactions) from the 1st and 2nd gel

elution as follows:

a. 10 µl 10x P1 Buffer b. 1 µl Nuclease P1 c. X µl of 20 µg RNA d. Bring final volume to 100 µl 10. Incubate at 37°C over night

11. To remove the phosphate, add the following to the reaction:

a. 100 µl of P1 digest reaction b. 15 µl 10x CIAP buffer c. 2 µl CIAP d. 33 µl water 162

12. Incubate at 37°C for 2 hr

13. Add 50 µl water to the sample and filter with 0.2 micron filter with 1 mL

syringe before HPLC use.

163

A15: DNA purification from T. brucei

1. Pellet 10 ml cells mid log and wash in PBS in 1.5 mL tube

2. Spin at max speed for 1 min and remove supernatant

3. Resuspend pellet in 300 µl of 2x lysis buffer

4. Add 2 µl of 10 mg/ml RNAseA, incubate 1hr at 37°C. Cool down to room

temperature.

5. Add 133 µl of 7.5M ammonium acetate, mix by inversion

6. Spin down 10 min max speed

7. Transfer supernatant to fresh tube and precipitate DNA by adding equal

volume of isopropanol

8. Spin down max speed for 10 min

9. Wash twice with 70% EtOH and let dry

10. Resuspend pellet in TE

2x Lysis Buffer 20 mM Tris pH 8.0 20 mM NaCl 20 mM EDTA 1% SDS

164

A16: Plasmid DNA purification from E. coli

1. Centrifuge 2 mL of mid to late log E. coli at 13.2 RPM for 1 min

2. Dump supernatant and resuspend pellet in 100 µl of Solution 1

3. Rake tubes across tube rack to resuspend

4. Add 200 µl of Solution 2 and invert to mix tubes 3-5 times

5. Add 150 µl of Solution 3 and invert to mix tubes 3-5 times

6. Centrifuge tubes at 13.2 RPM for 15 min

7. Collect supernatant and move to new tube

8. Phenol extract with equal volume Tris saturated phenol

9. Add 2x volume of ethanol and centrifuge at 13.2 RPM for 30 min

10. Remove ethanol and briefly allow pellet to air dry

11. Resuspend in 30-50 µl of water

Solution 1 1.25 mL of 1 M Tris-HCl 1 mL of 0.5 M EDTA Bring volume up to 50 mL with water

Solution 2 1 mL of 10 M NaOH 2.5 mL of 20% SDS Bring volume up to 50 mL with water

Solution 3 14.76 g potassium acetate 5.75 mL glacial acetic acid Bring volume up to 50 mL with water 2.5 mL of 20% SDS Bring volume up to 50 mL with water 165

A17: Electroporation of T. brucei

1. Collect 10 mL mid log cells and centrifuge 1,300 x g . Save supernatant

2. Wash cell pellet with 10 mL ice cold CytoMix buffer

3. Centrifuge again to pellet cells

4. Resuspend pellet in 1 mL of CytoMix buffer

5. Add 10-20 µg of linearized DNA. DNA must be sterile. Ethanol precipitate

DNA and remove ethanol in the cell culturing hood.

6. Add 0.5 mL of cells suspended in CytoMix buffer

7. Electroporation settings: 1,600V, 25Ω, 50 µF.

8. Add cells to 5 mL of conditioned media (supernatant cells were growing in

before first pelleting) and incubate 18hr.

9. Add 5mL of SDM-79 media with correct drug at 2x concentration.

10. Perform limiting dilution in SDM-79 in 24-well plate

a. Add 1.5 mL of culture to first row b. Add 1.0 mL of SDM-79 with drug 1x to second and third rows. c. Add 0.5 mL of SDM-79 with drug 1x to fourth row 11. Transfer 0.5 mL of culture in first row to second, mix by pipette. Transfer

0.5 mL to third row, mix by pipette. Transfer 0.5 mL to fourth row, mix by

pipette. This will generate each row with 1 mL of media and cells.

12. Wrap plates in saranwrap or parafilm to limit evaporation.

13. Dilute wells 1:2 with fresh media once many cells are seen.

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CytoMix buffer, Filter Sterilize before use: 25 mM HEPES pH 7.6 120 mM KCl 0.15 mMCaCl2 10 mM K2HPO4 10 mM KH2PO4 pH 7.6 2 mM EDTA 6 mM Glucose 5 mM MgCl2

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A18: Preparing competent DH5α and transformation

1. Inoculate 100 mL media with 1 mL of overnight cells

2. Grow the culture at 37°C with shaking until OD of 0.25 to 0.3. Do not

overgrow. Roughly 2 hr

3. Make Transformation Buffer, filter sterilize, keep on ice.

4. Once cells are at proper OD, chill on ice

5. Spin 5000 RPM (6,000g) for 7 min

6. Resuspend pellet in 20 mL of transformation buffer

7. Spin at 5000 RPM (6,000g) for 7 min

8. Resuspend pellet in 2.5 mL of transformation buffer, keep on ice.

9. Aliquot cells into microcentrifuge tubes and add 3.6 µL of DMSO/100 µL of

cells

10. Freeze -80 until use.

Transformation buffer (25 mL for 100 mL culture) MnCl2 0.22g CaCl2 anhydrous 0.17g KOAc 0.1g Sucrose 3.75g RbCl .325g Water up to 25 mL

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Transformation:

1. Thaw on ice 20 min

2. Add DNA

3. Let sit on ice 20 min

4. Heat shock 42°C 30 Sec or 37°C 2 min

5. Ice 1 min

6. Recover 20-30 min 37°C with LB

7. Spin 3,000 RPM 1 min

8. Gently pipette to resuspend for spreading on plate

9. 60 µL of xGal IPTG to plate if needed before cell addition.

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Appendix B: SDM-79 growth and depletion media used

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B1: Standard SDM-79:

1. To make 1 L, mix the components in 900 mL and stir for several hours

2. Once completely dissolved, bring final volume to 1 L.

3. pH the media to 7.3 using NaOH. Should turn light pink/red color

4. In the biosafety cabinet, filter sterilize the media with a 0.2 micron filter

5. Place media at 37°C over-night to ensure there is no bacterial

contamination.

6. The following day, move media to 4°C

7. When media is needed, aliquot the amount needed to a separate flask and

add the remaining components:

a. 10 mL fetal bovine serum/L b. 3.8 mL of hemin/L c. 10 mL of Pen-Strep is optional* 8. For RNAi we do not add tetracycline to the media flask, instead we add 1 µl

of Tet (Working Stock 1mg/mL)/1 mL of culture.

9. Antibiotics needed for strain maintenance are added to the aliquoted flask

of complete media

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Table 3: The antibiotics utilized in T. brucei media Antibiotic Stock Final /L G418 100 mg/mL 12 µg/mL 120 µl Hygromycin 100 mg/mL 50 µg/mL 500 µl Phleomycin 20 mg/mL 2.5 µg/mL 125 µl Puromycin 10 mg/mL 1 µg/mL 100 µl

Blasticidin 10 mg/mL 10 µg/mL 1000 µl

Maintaining strains of T. brucei may require the use of various antibiotics added to the media. For example, the Wild type 29-13 cell line uses the antibiotics G418 and hygromycin. Listed are the final concentrations we utilize in the media.

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Table 4: SDM-79 media composition

SDM-79 Media Composition Amount/L 1 MEM Powder 7.0 g 2 Grace's Insect Media 2.0 g 3 50x MEM essential a.a 8.0 mL 4 100x MEM non-essential a.a 6.0 mL 5 Glucose 1.0 g 6 HEPES 7.33 g 7 MOPS 5.0 g 8 Sodium Bicarbonate 2.0 g 9 Sodium Pyruvate 100 mg 10 DL-Alanine 200 mg 11 L-Arginine 100 mg 12 L-Glutamine 300 mg 13 DL-Methionine 70 mg 14 L-Phenylalanine 80 mg 15 L-Proline 600 mg 16 DL-Serine 60 mg 17 Taurine 160 mg 18 DL-Threonine 350 mg 19 L-Tyrosine 100 mg 20 Guanosine 10 mg 21 Folic Acid 4 mg 22 D(+)Glucosamine Hydrochloride 50 mg 23 p-Aminobenzoic acid 2 mg 24 Biotin 0.2 mg 25 Adenosine 20 mg

Listed are the 25 components we add to water to generate SDM-79 media (lacking hemin and 10% FBS) used in the Alfonzo lab. Specific details concerning numbers

1-4 are found in the following table.

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Table 5: Additional components of SDM-79

Grace's 50x 100x non- MEM Insect essential essential Powder Media a.a a.a (mg/L) (mg/L) (mg/L) (mg/L) 1 Glucose 734 31.5 2 DL-Alanine 18 5.34 3 L-Arginine 92.953 31.5 50.56 4 L-Glutamine 214.328 27 5 DL-Methionine 11.01 2.25 6 L- 6 Phenylalanine 23.488 6.75 13 7 L-Proline 15.75 6.9 8 DL-Serine 49.5 6.3 9 DL-Threonine 35.232 7.875 19.04 10 L-Tyrosine 38.168 31.599 17.28 11 Folic Acid 0.734 0.0009 Aminobenzoic 12 acid 0.0009 13 Biotin 0.00045 L-Cysteine 2 14 HCl 22.754 1.125 12.48 L-Histidine 15 hydrochloride 30.828 112.5 16.76 16 L-Isoleucine 38.168 2.25 21 17 L-Leucine 38.168 3.375 21 L-Lysine 18 Hydrochloride 53.215 28.125 29 19 L-Tryptophan 7.34 4.5 4 20 L-Valine 33.764 4.5 18.72 Choline 21 chloride 0.734 0.009 Calcium 22 pantothenate 0.734 23 Niacinamide 0.734 0.0009 Pyridoxal 24 hydrochloride 0.734 25 Riboflavin 0.0734 0.0009 Thiamine 26 hydrochloride 0.734 0.0009 27 i-Inositol 1.468 0.0009 Calcium 28 Chloride 146.8 174

Magnesium 29 Sulfate 71.690 Potassium 30 Chloride 293.6 Sodium 31 Chloride 4991.2 Sodium Phosphate 32 monobasic 102.76 33 Phenol Red 7.34 34 L-Asparagine 15.75 9 35 L-Aspartic Acid 15.75 7.98 L-Glutamic 36 Acid 27 8.82 37 Glycine 29.25 4.5 38 Sucrose 1200.6 39 D-(-)-Fructose 18 D-pantothenic 40 acid 0.0009 41 pyridoxine HCl 0.0009 42 Fumaric acid 2.475 a-ketoglutaric 43 acid 16.65 44 L-Malic Acid 30.15 Succinic Acid 45 free acid 2.7

A breakdown of the contribution Grace’s Insect Media, MEM powder, 50x essential, and 100x non-essential amino acids adds to SDM-79.

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B2: Nutrient depleted SDM-79

When making nutrient depleted SDM-79, various approaches were taken.

1. For depletion of additional media components such as FBS or hemin, the

standard SDM-79 protocol is followed, and they are simply left out.

2. For amino acid depletions, the above SMD-79 was modified slightly to

deplete a specific amino acid.

a. The non-amino acid components were added together first and dissolved in water for several hours. Due to the complexity of MEM powder and Grace’s Insect Media, neither of these were left out of the depleted media. This generated depletion media that still contained trace amounts of amino acids. b. This was then distributed evenly to 20 flasks c. Cumulative concentrations of each amino acid found in 50x essential, 100x non-essential, and exogenously added amino acids was found. d. This value was the specific amount added to maintain the 19 non- depleted amino acids at the concentration used in standard SDM- 79 (as each depletion condition had 19 non-depleted and 1 depleted amino acid). The amino acid being depleted was simply not added to the appropriate flask. e. The 50x essential and 100x non-essential amino acid liquids were not used in the media. f. Bring the media to the volume being made, in our case 100 mL. g. Akin to standard SDM-79, remaining components hemin and FBS were added after the required pH adjustment and filter sterilization.

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Table 6: The common components to all depleted media conditions

Non-amino acid SDM-79 Media Composition Amount/L 1 MEM Powder 7.0 g 2 Grace's Insect Media 2.0 g 3 Glucose 1.0 g 4 HEPES 7.33 g 5 MOPS 5.0 g 6 Sodium Bicarbonate 2.0 g 7 Sodium Pyruvate 100 mg 8 Taurine 160 mg 9 Guanosine 10 mg 10 Folic Acid 4 mg 11 D(+)Glucosamine Hydrochloride 50 mg 12 p-Aminobenzoic acid 2 mg 13 Biotin 0.2 mg 14 Adenosine 20 mg

When nutrient depleted SDM-79 was generated, these 14 common media components were added to all tested conditions. The concentrations used were the same as standard SDM-79 previously described.

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Table 7: Amino acid concentration of SDM-79 and depleted SDM-79

SDM-79 (mg/L) Depleted SDM-79 (mg/L) Alanine 223.34 18 Arginine 275.01376 124.45376 Asparagine 24.75 15.75 Aspartic Acid 23.73 15.75 Cysteine 36.359 23.879 Glutamic Acid 35.82 27 Glutamine 541.328 241.328 Glycine 33.75 29.25 Histidine 160.088 143.328 Isoleucine 61.418 40.418 Leucine 62.543 41.543 Lysine 110.34 81.34 Methionine 89.26 13.26 Phenylalanine 123.238 30.238 Proline 622.65 15.75 Serine 115.8 49.5 Threonine 412.147 43.107 Tryptophan 15.84 11.84 Tyrosine 187.047 69.767 Valine 56.984 38.264

Final known amino acid concentrations are detailed above comparing amino acid depleted media (Depleted SDM-79 column) to standard media (SDM-79 column).

In each depletion tested, only one amino acid was depleted while the remaining

19 amino acids would be at the concentration in the SDM-79 column. For example, the alanine depletion would have a concentration of 18 mg/L of alanine while the remaining 19 amino acids would be at the traditional SDM-79 concentration found in the SDM-79 column. Additionally, unknown trace amounts of amino acids may be found in FBS added to the media as well.

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Appendix C: Supporting figures

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Figure C.1: TbtTrl1 knockdown causes intron-containing tRNATyr accumulation A. Northern blot analysis of TbTrl1 RNAi knockdown, induced by the addition of

TET, reveals processing of tRNATyr. B. To determine positions of intron-containing tRNA, the membrane was hybridized with an intron specific probe. Figure source:

(Lopes et al. 2016).

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Figure C.2: Intron-containing tRNA are in the cytoplasm A. RNA from total (T), nuclear (N) and cytoplasmic (C) fraction of 4 days induced

(+) and uninduced (-) TbTrl1 RNAi cells was extracted and analyzed by northern blotting using a 3’ exon or intron probe. B. The position of intron- containing tRNA is indicated and confirmed by re-hybridizing the membrane with an intron specific probe. C. As a control, the membrane was hybridized with the nuclear specific

SnoRNA. D. Fractions were also analyzed by Western blotting by nuclear specific

NOG1 and cytoplasmic specific Enolase proteins. Figure source: (Lopes et al.

2016).

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Figure C.3: Additional controls for Figure 2.6 A. Fractions from (Figure 2.6 A, lanes “C” and “N”) and (Figure 2.6 B) were analyzed for the presence of Q by APB northern blotting to ensure Q was absent.

B. As a control for Figure 2.6 A, fractions purity was determined by Western blot with NOG1 being a nuclear marker and Enolase, cytoplasmic.

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