Point Mutational Examination of Eukaryotic Initiation Factor 4E (Eif4e)

Iowa State University Capstones, Theses and Graduate Theses and Dissertations Dissertations

2018 Point mutational examination of eukaryotic initiation factor 4E (eIF4E), inclusion bodies analysis, and Barley Yellow Dwarf Virus (BYDV) exonuclease resistant RNA structure mapping and prediction Melissa Kathryn Sheber Iowa State University

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Recommended Citation Sheber, Melissa Kathryn, "Point mutational examination of eukaryotic initiation factor 4E (eIF4E), inclusion bodies analysis, and Barley Yellow Dwarf Virus (BYDV) exonuclease resistant RNA structure mapping and prediction" (2018). Graduate Theses and Dissertations. 16464. https://lib.dr.iastate.edu/etd/16464

This Thesis is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Graduate Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. Point mutational examination of eukaryotic initiation factor 4E (eIF4E), inclusion bodies analysis, and Barley Yellow Dwarf Virus (BYDV) exonuclease resistant RNA structure mapping and prediction

Melissa Sheber

A thesis submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

MASTER OF SCIENCE

Major: Biochemistry

Program of Study Committee: W. Allen Miller, Major Professor Dipali Sashital Cathy Miller

The student author, whose presentation of the scholarship herein was approved by the program of study committee, is solely responsible for the content of this thesis. The Graduate College will ensure this thesis is globally accessible and will not permit alterations after a degree is conferred.

Iowa State University

Ames, Iowa

2018

TABLE OF CONTENTS

Page

LIST OF FIGURES ...... iii

LIST OF TABLES ...... v

ACKNOWLEDGMENTS ...... vi

ABSTRACT ...... vii

CHAPTER 1. A REVIEW OF THE LITERATURE ...... 1 Canonical Eukaryotic Translation Initation ...... 1 Plant Viral Translation Initation: 3’ Cap-Independent Translation Elements (3’CITEs) ...... 5 Natural Resistance Conferred by Eukaryotic Initation Factor 4E (eIF4E) ...... 10

CHAPTER 2. POINT MUTATIONAL ANALYSIS OF EUKARYOTIC INITIATION FACTOR 4E (eIF4E) ...... 15 Introduction ...... 15 Materials and Methods ...... 18 Results ...... 26 Discussion ...... 47

REFERENCES ...... 53

APPENDIX. BARLEY YELLOW DWARF VIRUS (BYDV) SUBGENOMIC RNA3 CONTAINS AN XRN1 RESISTANT RNA (xrRNA) STRUCTURE ...... 67 Abstract ...... 67 Introduction ...... 67 Materials and Methods ...... 75 Results ...... 77 Discussion ...... 86 Acknowledgments ...... 87 References ...... 87

iii

LIST OF FIGURES

Page

Figure 1.1 Eukaryotic translation initiation ...... 6

Figure 1.2 Secondary strucutre of the 7 classes of 3’CITEs ...... 10

Figure 1.3 eIF4E surface map of regions associated with viral resistance...... 14

Figure 2.1 Host mRNA and viral mechanims for circularization and activation for ribosome loading...... 17

Figure 2.2 Secondary strucutre of PTE from Panicum mosiac virus (PMV) and 3D modeling of PTE-eIF4E binding...... 18

Figure 2.3 Wheat eIF4E mutants with a focus on the bottom distal side of the cap binding pocket...... 32

Figure 2.4 Wheat eIF4E mutants with a focus on the top proximal side of the cap binding pocket...... 33

Figure 2.5 Cap binding pocket amino acid coordination to m7GDP with a focus on the top proximal side of the cap binding pocket...... 34

Figure 2.6 Cap binding pocket amino acid coordination to m7GDP with a focus on the bottom distal side of the cap binding pocket...... 35

Figure 2.7 Electrophoretic mobility shift assay (EMSA) of wheat eIF4E with the PTE from Thin paspalum asymptomatic virus (TPAV)...... 39

Figure 2.8 Plasmid map of pGEX-5X-2 with eIF4E insert...... 42

Figure 2.9 Expression of GST-eIF4E soluble and insoluble fractions at 37 °C...... 44

Figure 2.10 Expression of GST-eIF4E soluble and insoluble fractions at 37 °C versus 30 °C...... 45

Figure 2.11 Expression of GST-eIF4E soluble and insoluble fractions at 25 °C...... 46

Figure 3.1 Genome map of Barley yellow dwarf virus (BYDV)...... 71

Figure 3.2 Schematic Xrn1 degradation to generate Barley yellow dwarf virus (BYDV) subgenomic RNA3 (sgRNA3)...... 71

Figure 3.3 Plant mRNA decay...... 72 iv

Figure 3.4 Structure of xrRNA from flaviviruses...... 73

Figure 3.5 Structure of xrRNA from diantoviruses...... 74

Figure 3.6 Map of Barley yellow dwarf virus (BYDV).subgenomic RNA3 (sgRNA3) deletion constructs ...... 82

Figure 3.7 5’ triphosphate transcripts of deletion constructs...... 83

Figure 3.8 Xrn1 assay of 5’ triphosphate deletion constructs...... 83

Figure 3.9 5’ monophosphate transcripts of deletion constructs...... 84

Figure 3.10 Xrn1 assay of 5’ monophosphate deletion constructs...... 84

Figure 3.11 Secondary strucutre prediction of BYDV sgRNA3 xrRNA...... 85

Figure 3.12 RNAalifold results sequence alignment...... 85

LIST OF TABLES

Page

Table 2.1 Forward and reverse primers used in the generation of eIF4E point mutations...... 19

Table 2.2 PCR conditions for site directed mutagensis...... 20

Table 2.3 Insoluble resuspension...... 21

Table 2.4 PCR conditions to amplify eIF4E open reading frame (ORF) and to confirm eIF4E ORF insert presence by colony PCR...... 23

Table 2.5 Wheat and lettuce eIF4E mutations with results of lettuce eIF4E study by German Retana et al., 2008...... 31

ACKNOWLEDGMENTS

I would like to thank my advisor Dr. Allen Miller for giving me the opportunity to work on these projects and grow as a researcher and for his mentorship, support and encouragement throughout my work but especially during troubleshooting efforts. I would also like to thank my committee members Dr. Cathy Miller and Dr. Dipali Sashital for taking the time to provide constructive criticism and guidance and for meeting with me on such short notice. In addition, I would like to thank Dr. Walter Moss for his excellent help and advice with bioinformatics analysis.

I would like to express great appreciation to my mentors Dr. Juliette Doumayrou and Dr. Keisuke Komoda for the time and care they took training me and for their continued support and advice. I am deeply grateful to have had the chance to learn from both of them. I would also like to thank my lab mates Elizabeth Carino, Pulkit Kanodia and Dr. Ryan Lappe for their discussion, support, jokes and camaraderie. I owe special thanks to Elizabeth Carino for her mentorship, guidance and emotional support during difficult times.

I would like to thank my professors, especially Dr. Richard Honzatko, for their stimulating course work and enlightening discussions. Lastly, I would like to thank my family, friends and my partner for their understanding and support and for having confidence in me when I sometimes doubted myself. vii

ABSTRACT

Eukaryotic initiation factor 4E (eIF4E) binds the 5’m7GTP cap structure of mRNA in order to facilitate effective translation. Recessive eIF4E alleles harboring naturally occurring point mutations have been associated with resistance to viral infection, particularly in members of the genus Potyvirus. Resistance is thought to be conferred by disrupting the binding of the 5’ viral-encoded protein (VPg), covalently attached to the 5’ end of the Potyvirus genomic RNA, to eIF4E. RNAs of members of the genus Panicovirus, some Carmoviruses and one Umbravirus however, bind eIF4E through a 3’ cap independent translation element (3’CITE) termed the PTE (Panicum mosaic virus-like translation enhancer). The PTE consists of a T shaped secondary structure with a C-rich region at the branch point between stem loops 1 and 2 and G-rich region in a bulge in the basal stem loop. The C-rich and G-rich regions form a pseudoknot where the G-rich region contains a flexible G that is thought to flip outward and potentially act as a 5’ m7GTP cap analog to bind eIF4E. Point mutations were introduced into wheat eIF4E in order to identify amino acids needed for PTE binding and were designed based on previous studies of recessive resistance-conferring eIF4E alleles, along with the crystal structure of wheat eIF4E bound to m7GDP. However, the chosen method of eIF4E purification involving a glutathione S-transferase (GST) tag repeatedly resulted GST-eIF4E inclusion bodies, which limited the ability to purify mutant eIF4E for binding studies. Therefore a large portion of this work is concerned with attempts to solubilize the GST-eIF4E fusion protein. viii

Additionally, the second portion of this work involves mapping and predicting the structure in Barley yellow dwarf virus RNA (BYDV) that blocks exonuclease Xrn1 to generate BYDV subgenomic RNA3. Deletion analysis places the Xrn1 resistant

(xrRNA) structure within the first 5’ 67 nucleotides of BYDV sgRNA3. Dianthoviruses, related to luteoviruses, have recently been shown to contain an xrRNA structure of which a crystal structure has been obtained. Bioinformatics analysis using the programs

INFERNAL and Dynalign suggest the BYDV sgRNA3 xrRNA structure may be different than that of dianthoviruses. A proposed secondary structure of BYDV sgRNA3 xrRNA was obtained by analysis with the programs DotKnot and RNAalifold. 1

CHAPTER 1. A REVIEW OF THE LITERATURE

Translation, the process of decoding messenger RNA (mRNA) to synthesize protein, is divided into three stages: initiation, elongation and termination. Translation is a highly regulated and intricate process, and a significant body of work has contributed to our understanding of the factors and complex mechanisms involved. The field of translation continues to advance as areas of emerging interest like the diverse roles of the untranslated regions (UTRs) of the message are being investigated (Mayr, 2017; Mignone et al., 2002).

The review will focus on translation initiation and elements of UTRs and their respective roles in plant viral resistance. An understanding of these topics is both necessary and useful for understanding the work in this thesis and the relevance of the work in terms of conferring viral resistance in plants.

Canonical Eukaryotic Translation Initiation

Translation initiation involves several initiation factors and multiple points of regulation (Aitken and Lorsch, 2012; Sonenberg and Hinnebusch, 2009). Canonical translation initiation begins with the formation of the 43S preinitiation complex (43S PIC) and concomitant activation of mRNA. Activated mRNA loads onto the 43S PIC at which time the 43S PIC scans in the 5’ to 3’ direction in search of the AUG start codon.

Identification of the start codon coupled with GTP hydrolysis and release of inorganic phosphate yields the 48S preinitiation complex (48S PIC). Hydrolysis of a second GTP and displacement of initiation factors allows binding of the 60S ribosomal subunit resulting in the elongation competent 80S initiation complex. A schematic of translation initiation described above is shown in figure 1.1. The steps of translation initiation will be discussed in greater detail below. 2

The 43S PIC is composed of the ternary complex (TC), 40S ribosomal subunit and eukaryotic initiation factors (eIFs) eIF1, eIF1A, eIF3, eIF5, where eIF2 bound to GTP and the initiator tRNA, Met-tRNAi comprise the TC (Asano et al., 2000). Before initiation factor functions are later described, an understanding of their location within what is known of the

43S PIC is useful. The structure of the 40S ribosomal subunit can be envisioned by a globular head and body with a platform projection connected via a neck region forming the mRNA channel (Jackson et al., 2010). The A site, P site and E site line the mRNA channel. eIF1 and eIF1A are located near the P site and in the A site respectively (Hinnebusch, 2017;

Lomakin et al., 2003). Met-tRNA is superficially situated in the P site relative to the deeper position that occurs upon anti-codon codon binding (Jackson et al., 2010). The multisubunit initiation factor eIF3 wraps around the solvent side of the 40S with subunits located near the entry and exit of the mRNA channel as well as subunits positioned to enable interactions with multiple initiation factors including eIF1, eIF1A, eIF2, and eIF5 (Hinnebusch, 2006).

Solvent facing subunits of eIF3 are also positioned to take part in facilitating activated mRNA attachment to the 43S PIC (Hinnebusch, 2006).

Cellular mRNAs are co-transcriptionally modified with the addition of a 5’ m7GpppN cap structure and post transcriptionally modified by 3’ polyadenylation forming a

3’ poly(A) tail (Lewis et al., 1995). Activation of mRNA involves binding of the 5’ m7GppppN cap to the eIF4F complex. In mammals, eIF4F is comprised of the cap-binding protein eIF4E, a large scaffold protein eIF4G and an RNA helicase eIF4A (Haghighat and

Sonenberg, 1997; Jackson et al., 2010). The 5’UTRs of mRNA are often highly structured, which necessitates unwinding by the ATP dependent helicase activity of eIF4A aided by another factor, eIF4B (Aitken and Lorsch, 2012; Svitkin et al., 2001). Interestingly, plant and 3 yeast eIF4F are composed of only eIF4E and eIF4G and furthermore, plants have an isoform of eIF4F called eIFiso4F constituted by eIFiso4E and eIFiso4G (Browning, 2004). Though not a part of the eIF4F complex, the poly(A)-binding protein (PABP), which binds the 3’ poly(A) tail, also binds eIF4G resulting in circularization of the mRNA and enhanced translation (Gallie, 1998; Wells et al., 1998). Delivery of activated mRNA to the 43S PIC for attachment and subsequent scanning is mediated by the binding of the scaffold eIF4G of eIF4F to the multisubunit eIF3 of the 43S PIC (Hinnebusch, 2006; Kumar et al., 2016).

Scanning for and accurate selection of the AUG start codon by the 43S PIC requires the coordination of initiation factors as well as favorable start codon sequence context. In eukaryotes, the Kozak sequence GCC(A/G)CCAUGG serves as optimal AUG (shown in bold) context (Kozak, 1986, 2002). Initiation factor eIF1, bound near the P site, promotes scanning by sterically blocking full depth inclusion of Met-tRNA into the P site in the absence of AUG binding (Lomakin et al., 2003). As mentioned previously, eIF1A is located near the A site, however the C terminal tail (CTT) and N terminal tail (NTT) of eIF1A extend into the P site and serve opposing functions (Fekete et al., 2007; Hinnebusch, 2017). The

CTT of eIF1A functions to partially obstruct the P site similar to eIF1 promoting the open scanning conformation (Hinnebusch, 2017). In contrast, the NTT of eIF1A forms interactions to stabilize the closed conformation that occurs upon AUG binding (Fekete et al., 2007;

Hinnebusch, 2017).

Start codon binding results in the formation of the 48S PIC and concomitant stimulation of eIF5, a GTPase activating protein (GAP), which induces GTP hydrolysis by eIF2 (Asano et al., 2001). However, GTP hydrolysis alone is not sufficient to facilitate AUG selection. Rather, anti-codon codon binding in concert with the stabilizing interactions 4 mentioned prior, provide enough energy to displace eIF1, which simultaneously permits the irreversible release of inorganic phosphate resulting in commitment of AUG selection

(Algire et al., 2005). Subsequent dissociation of eIF2-GDP, eIF3, and eIF5 occurs while eIF1A remains bound. eIF5B-GTP binds the 48S PIC and mediates attachment of the 60S subunit (Pestova et al., 2000). Hydrolysis of the second GTP by eIF5B and dissociation of eIF1A and eIF5B-GDP ensues, marking the formation of the elongation competent 80S initiation complex (Jackson et al., 2010).

Initiation factors also serve regulatory roles via phosphorylation. In mammals, phosphorylation of eIF2 results in tight binding and sequestering of eIF2B, where eIF2B normally serves as a guanine exchange factor (Sonenberg and Hinnebusch, 2009).

Sequestering of eIF2B by phosphorylated eIF2 inhibits eIF2B’s guanine exchange factor activity therefore decreasing eIF2-GTP concentrations available for TC formation and consequently decreasing rates of translation initiation (Sonenberg and Hinnebusch, 2009).

However, the homologous kinases responsible for eIF2 phosphorylation in mammals have not yet been expressly identified in plants, except for the nutrient deprivation stimulated kinase GCN2, suggesting that regulation mediated by eIF2 phosphorylation may differ in pathway or may not be as important in plants (Browning and Bailey-Serres, 2015).

Furthermore, homologs to mammalian 4E-BPs (eIF4E binding proteins) have not been specifically identified in plants (Browning and Bailey-Serres, 2015). In mammals, hypophosphorylated 4E-BPs function to compete with eIF4G for eIF4E binding, inhibiting formation of eIF4F complexes, which reduces translation initiation rates (Rong et al., 2008). 5

Though a considerable amount of the mechanism of translation initiation has been elucidated, much more remains to be discovered in terms of the intricacy of regulation and structural interactions in initiation complex formation.

Plant Viral Translation Initiation: 3’ Cap-Independent Translation Elements (3’CITEs)

Most positive sense RNA plant viruses lack a 5’ cap and 3’ poly(A) tail yet must compete with host mRNA for translation machinery, whereas host mRNAs employ both a 5’ cap structure and poly(A) tail to translate efficiently (Dreher and Miller, 2006). Thus genomes of these viral RNAs have evolved alternative means of recruiting initiation factors, or in some cases ribosome subunits (Simon and Miller, 2013). Cap-independent translation enhancers (CITEs) located in their 3’ UTRs are the most common alternative strategy employed for efficient translation (Miller et al., 2007). 3’ CITEs vary in sequence, structure and specific mechanism (Miller et al., 2007). However, overall function similarities emerge in that 3’ CITES bind components of the translation machinery and circularize the message by engaging in interaction with the 5’ UTR, which serves to facilitate recruitment of translation machinery to the 5’ end for scanning (Nicholson and White, 2011). Seven classes of 3’ CITEs have been identified thus far based on secondary structure shown in figure 1.2

(Truniger et al., 2017).

Figure 1.1 Translation initiation beginning with activation of mRNA and formation of the 43S PIC. Scanning in the 5’ to 3’ direction leads to start codon recognition and formation of 48S PIC. GTP hydrolysis and release of factors allows 60S binding. A second GTP hydrolysis and release of eIF5B-GDP results in the elongation competent 80S initiation complex. 7

The first 3’ CITE identified, the translation enhancer domain (TED), was discovered in Satellite tobacco necrosis virus (STNV) (Danthinne et al., 1993; Meulewaeter et al.,

1998). The structure of TED has yet to be experimentally verified but is proposed to form a long stem loop with several bulges, one of which binds a complementary sequence presented in a 5’ UTR hairpin loop (Blanco-Perez et al., 2016; Simon and Miller, 2013). TED binds eIF4F or eIFiso4F facilitating translation initiation presumably in concert with the 5’-3’

RNA-RNA interaction (Gazo et al., 2004).

The BTE (Barley yellow dwarf virus-like translation element) is one of the most characterized 3’ CITEs and was first discovered in Barley yellow dwarf virus (BYDV)

(Treder et al., 2008; Wang et al., 2010). The BTE is found in the genera Luteovirus,

Dianthovirus and Necrovirus (Simon and Miller, 2013; Wang and Miller, 1995). BTE structure consists of a central region radiating either three, four, or six stem loops depending on the genus from which the BTE originates (Simon and Miller, 2013). However, a 17 nucleotide consensus sequence involving stem loop I (SL-I) is observed in all BTEs so far

(Simon and Miller, 2013). BTE mediated translation initiation involves both binding eIF4G of the eIF4F complex and participating in long distance RNA-RNA interaction with the 5’

UTR (Guo et al., 2001; Treder et al., 2008).

Originally identified in Panicum mosaic virus (PMV), the PTE (Panicum mosaic virus-like translation enhancer) consists of three branching helices which form a T shaped secondary structure, where a C rich region at the branch point forms a pseudoknot with a G rich region in a bulge in the stem (Batten et al., 2006; Wang et al., 2009). SHAPE probing revealed a hypermodifiable G in the G rich region putatively involved in the PTE’s unique ability to bind eIF4E with very high affinity (Wang et al., 2011; Wang et al., 2009). Along 8 with binding eIF4E, PTE-mediated translation initiation also likely involves 3’ to 5’ RNA-

RNA interaction (Gao et al., 2012; Nicholson and White, 2011). Though mostly observed in the genus Panicovirus, the Umbravirus Pea enation mosaic virus RNA 2 (PEMV2) and some

Carmoviruses also contain a PTE (Simon and Miller, 2013).

Similar to the PTE, the Y-shaped structure (YSS) 3’ CITE observed in the genus

Tombusvirus consists of three branching helices, however the helices in the YSS are significantly longer than those of the PTE (Simon and Miller, 2013). Also in contrast to the

PTE, the YSS does not have pseudoknot forming C and G rich regions and binds eIF4F or eIFiso4F (Nicholson and White, 2011). A complementary sequence in the 5’ UTR binds the

YSS and 5’ to 3’ long distance interaction between the YSS and 5’ UTR is required for YSS mediated translation initiation (Fabian and White, 2004).

The Tombusvirus Maize necrotic spot virus (MNeSV) and the Carmovirus Melon necrotic spot virus (MNSV) both utilize a class of 3’ CITE termed the I-shaped structure

(ISS) (Truniger et al., 2017). The ISS are among the shortest 3’CITEs identified so far using an average of 60 bases to form a stem loop structure with two bulges, where the sequence of the stem loop structure seems to be less important than the extensive base pairing that conserves the structure (Truniger et al., 2017). As with other 3’ CITEs, ISS-mediated translation initiation involves long distance 3’ to 5’ RNA-RNA interaction and binding of an initiation factor (Truniger et al., 2017). eIF4F serves as the binding partner for the ISS, however two hypermodifiable G bases, G13 and G47, suggests that binding may occur through the eIF4E subunit of eIF4F (Nicholson et al., 2010).

Another class of 3’CITE, first discovered in Turnip crinkle virus (TCV), was termed

TSS (T-shaped structure) (McCormack et al., 2008). As implied by its name, the TSS forms 9 a tertiary T-shaped structure similar to tRNA (Nicholson and White, 2011). In contrast to other 3’CITEs, TSS has not been tested for binding to initiation factors but rather has been shown to bind the P-site of the 80S ribosome or 60S ribosomal subunit directly (Truniger et al., 2017). Furthermore, communication with the 5’ end does not appear to be mediated through 5’ to 3’ RNA-RNA interaction but rather is hypothesized to occur through a protein bridge formed by joining of the TSS bound 60S ribosomal subunit to the 5’ UTR bound 40S ribosomal subunit (Stupina et al., 2011).

The newest class of 3’CITE, CXTE (CABYV-Xinjiang-like translation element), was discovered in a resistance-breaking isolate of MNSV, whereby MNSV obtained the new 3’

CITE from an Asian isolate of Cucurbit aphid-borne yellows virus (CABYV) through recombination (Miras et al., 2014; Truniger et al., 2017). SHAPE probing revealed the structure of the CXTE to consist of two helices radiating from a central region (Miras et al.,

2014). Unlike the PTE, YSS, and other 3’CITEs containing branching helices, the helices of

CXTE do not appear to branch from a basal helix. So far, CXTE binding partners and mode of communication with the 5’ UTR have not been identified (Truniger et al., 2017).

While researchers have made significant advances in the important foundational identification and classification of 3’CITEs presented above, the study of viral 3’CITEs is still in its infancy. Many more classes of 3’CITEs are likely to be discovered. Furthermore, although binding to a component of the translation machinery and communicating with the 5’

UTR appear to be common functions among 3’CITEs discovered thus far, the unique and intricate mechanisms by which different classes achieve translation initiation have yet to be elucidated.

Figure 1.2 Secondary structural representation of the seven classes of 3’ CITEs adapted from Simon and Miller, 2013; Miras et al., 2014.

Natural Resistance Conferred by Eukaryotic Initiation Factor 4E (eIF4E)

Natural viral resistance in plants, particularly to viruses of the genus Potyvirus, has been linked to recessive genes coding for eukaryotic initiation factor 4E (eIF4E) (Robaglia and Caranta, 2006; Truniger and Aranda, 2009). Plants have two forms of eIF4E, eIF4E and eIFiso4E, both of which are members of class I (Browning and Bailey-Serres, 2015). Class I eIF4Es are characterized by conservation of the two cap binding tryptophan residues, which sandwich the cap and stabilize binding via π electron stacking interactions (Dinkova et al.,

2016; Marcotrigiano et al., 1997). Importantly, plants also exhibit varying degrees of gene redundancy for eIF4E and eIFiso4E (Dinkova et al., 2016; Robaglia and Caranta, 2006).

Arabidopsis thaliana for example, has one gene coding for eIFiso4E and three genes coding eIF4E, eIF4E1, eIF4E2 and eIF4E3 (Robaglia and Caranta, 2006). Gene redundancy in both forms of eIF4E may serve to safeguard plants against adverse effects due to loss of function 11 alleles or engineered eIF4E knockout (KO) strains but simultaneously provide the potential for multiple susceptibility factors for viral infection (Bastet et al., 2017).

Viruses exhibit differential eIF4E requirements depending both on the virus and plant host (Robaglia and Caranta, 2006; Truniger and Aranda, 2009; Wang and Krishnaswamy,

2012). For example, Arabidopsis thaliana lacking eIFiso4E are resistant to Turnip mosaic virus (TuMV) but are susceptible to Clover yellow vein virus (ClYVV) (Sato et al., 2005).

Conversely, Arabidopsis thaliana lacking eIF4E are resistant to ClYVV but are susceptible to TuMV (Sato et al., 2005), suggesting ClYVV requires eIF4E while TuMV requires eIFiso4E to infect the same host in the case of Arabidopsis (Sato et al., 2005; Truniger and

Aranda, 2009; Wang and Krishnaswamy, 2012). However, differential eIF4E requirements are also exhibited for the same virus among infection of different hosts (Robaglia and

Caranta, 2006). Lettuce mosaic virus (LMV) for example, requires eIFiso4E for Arabidopsis infection but requires eIF4E for infection of lettuce, pepper and tomato (Robaglia and

Caranta, 2006). Furthermore some viruses are able to use multiple forms of eIF4E during infection (Bastet et al., 2017; Gauffier et al., 2016). For example, Potato virus Y (PVY) and

Tobacco etch virus (TEV) appear to recruit eIF4E2 upon knockout of eIF4E1 (Gauffier et al.,

2016). Differential eIF4E requirements depending on virus-host combinations and the ability of some viruses to use multiple eIF4Es suggest the necessity of multiple eIF4E knockouts to obtain effective resistance. However, pyramiding eIF4E knockout mutants can prove lethal for some host (Bastet et al., 2017). In the case of Arabidopsis, knocking out eIF4E1 and eIFiso4E is lethal, (Bastet et al., 2017; Callot and Gallois, 2014). Consequently, limitations in the utility of knockout strains necessitate the investigation of alternative strategies for engineering viral resistance. 12

Many recessive resistance-conferring alleles of eIF4E have been identified (Bastet et al., 2017; Robaglia and Caranta, 2006; Truniger and Aranda, 2009; Wang and

Krishnaswamy, 2012). Recessive resistant eIF4E alleles identified thus far include: mo1¹ and mo1² in lettuce (German-Retana et al., 2008; Nieto et al., 2006), pvr2² and pvr1 pepper (Kang et al., 2005; Ruffel et al., 2002), sbm1 pea (Gao et al., 2004), nsv in melon (Nieto et al.,

2006), pot1 in tomato (Ruffel 2005), and rym4 and rym5 in Barley (Stein et al., 2005). Many of the recessive resistance alleles appear to mediate resistance by functional eIF4E point mutations (Truniger and Aranda, 2009). Taken together, these point mutations clustered in two regions, inside the cap binding pocket and on the solvent accessible surface near the cap binding pocket shown in figure 1.3 (Robaglia and Caranta, 2006; Truniger and Aranda,

2009). Mutations in these regions are proposed to confer resistance by disrupting the binding of the viral genome-linked protein (VPg) in members of the genus Potyvirus. The 5’ covalently linked VPg is known to bind eIF4E and be important for Potyvirus infection

(Leonard et al., 2000; Shukla et al., 1991). Point mutational analysis of eIF4E guided by resistance alleles discovered thus far may reveal resistance-conferring mutations in crops where natural resistance is lacking or where knockout-mediated resistance is not possible.

Upon identification of such resistance conferring point mutations, the gene editing method

CRISPR/Cas9 could then be used to engineer resistant crops (Bastet et al., 2017; Pyott et al.,

2016). CRISPER/Cas9 has already been effectively used in a transgene-free manner in the genome editing of wheat (Zhang et al., 2016). Resistant crops engineered without the use of transgenes and based on naturally occurring mutations may prove to be very valuable in the current climate of controversy surrounding GMOs (Bastet et al., 2017). 13

Point mutational analysis studies based on natural point mutations have already been conducted for lettuce (German-Retana et al., 2008) and pea (Ashby et al., 2011). Both of these studies investigated point mutations in lettuce eIF4E and pea eIF4E on binding of the

VPg for Lettuce mosaic virus (LMV) and Pea seed-borne mosaic virus (PSbMV) respectively. The work of this thesis seeks to extend the point mutational analysis performed with lettuce eIF4E on VPg binding to wheat eIF4E binding the 3’ cap-independent translation enhancer (3’CITE) Panicum mosaic virus-like translation enhancer (PTE).

Figure 1.3 Surface representation of eIF4E (PDB file 2IDV) bound to m7GDP (shown as a stick representation in grey). Yellow indicates regions directly involved in cap binding. Blue indicates regions inside the cap binding pocket associated with mutations conferring viral resistance. Orange indicates regions on the outer surface associated with mutations conferring viral resistance. A) Focus on the bottom distal side of the cap binding pocket. B) Focus on the top proximal side of the cap binding pocket. 15

CHAPTER 2. POINT MUTATION ANALYSIS OF EUKARYOTIC INITATION FACTOR 4E (eIF4E)

Introduction

Point mutations were introduced into eukaryotic initiation factor 4E (eIF4E) with the goal of investigating the mechanism of viral translation and exploiting possible defects, which may be able to confer viral resistance. Important in the facilitation of translation initiation, the 25 kDa protein eukaryotic initiation factor 4E (eIF4E) binds the 5’ m7GTP cap structure of mRNA (Marcotrigiano et al., 1997; Rhoads, 2009). Two tryptophan residues in the cap binding pocket (W43 and W56 in human eIF4E and W62 and W108 in wheat eIF4E) sandwich the guanine of m7GTP and stabilize cap binding through stacking π bond interactions (Dinkova et al., 2016; Marcotrigiano et al., 1997). During mRNA activation for translation initiation, eIF4E binds the large scaffold protein, eIF4G, to form the eIF4F complex in plants whereas eIF4E, eIF4G and the RNA helicase eIF4A form the eIF4F complex in mammals (Browning and Bailey-Serres, 2015; Jackson et al., 2010). The poly(A) tail binding protein (PABP) binds the poly(A) tail of mRNA and also binds eIF4G resulting in circularization of the mRNA (Gallie, 1998). The ribosome is then able to load onto the activated mRNA and translation ensues upon identification of the start codon AUG (Jackson et al., 2010). However upon viral infection, viruses must compete with host mRNA for translation machinery, specifically for binding to eIF4E in many cases (Leonard et al., 2000;

Nicholson and White, 2011; Simon and Miller, 2013; Wang et al., 2009). The canonical mRNA and viral mechanisms for binding eIF4E important to the work in this thesis are shown in figure 2.1.

Recessive alleles coding for eIF4E have been linked to viral resistance, particularly with respect to members of the genus Potyvirus (Dinkova et al., 2016; Robaglia and Caranta, 16

2006). Potyvirus genome organization consists of a positive sense single stranded RNA with a 5’ covalently linked viral-encoded protein (VPg) and 3’ poly(A) tail (Shukla et al., 1991).

Binding between the viral 5’ VPg and host eIF4E has been shown to occur and to be important for infectivity of the virus (Leonard et al., 2000). Naturally occurring point mutations in eIF4E encoded by recessive resistance alleles have been identified and have been suggested to confer resistance by disrupting the binding of eIF4E to the viral VPg

(Robaglia and Caranta, 2006).

Designing eIF4E point mutations guided by recessive resistance alleles may reveal amino acids involved in VPg binding and thus in conferring viral resistance. A study of this nature has been conducted in lettuce (German-Retana et al., 2008). eIF4E point mutations were designed based in part on the recessive resistance allele of lettuce, pepper and pea in order to identify amino acids important for conferring resistance to the Potyvirus Lettuce mosaic virus (LMV) (German-Retana et al., 2008).

Unlike Potyviruses, which employ the VPg to bind eIF4E, members of the genus

Panicovirus and some Carmoviruses bind eIF4E through a 3’ cap-independent translation element (3’CITE) termed the PTE (Panicum mosaic virus-like translation enhancer) (Batten et al., 2006; Simon and Miller, 2013; Wang et al., 2009). The PTE, located in the 3’UTR, forms a T shaped secondary structure where the branch point of stem loop 1 (SL1) and stem loop 2 (SL2) contain a C rich region that forms a pseudoknot with a G rich region in a basal stem loop bulge shown in figure 2.2 (Wang et al., 2009). eIF4E binds the PTE where a hypermodifiable G in the G rich region is thought to act as a potential cap analog in eIF4E binding shown in 3D modeling in figure 2.2 (Wang et al., 2011). This study sought to extend 17 the point mutational analysis performed with lettuce in order to identify amino acids in wheat eIF4E important for binding of the PTE.

Figure 2.1 Host cellular mRNA and viral mechanisms for circularization and activation for ribosome loading. Host mRNA circularization occurs by the 5’ m7GTP cap structure (labeled m7G) binding eIF4E while the 3’ poly(A) tail binds PABP. Panicoviruses and some carmoviruses bind eIF4E through the 3’ CITE PTE where circularization is facilitated by an RNA-RNA interaction with the 5’ end. Potyvirus achieves circularization by the 5’ linked protein VPG binding to eIF4E while the 3’ poly(A) tail binds the PABP.

Figure 2.2 Secondary structure of the PTE 3’CITE from Panicum mosaic virus (PMV) is shown on the left (Wang et al., 2011). Arrows indicate the C-rich region between stem loops 1 (SL1) and 2 (SL2), and the G-rich region in a bulge of the basal stem loop (Wang et al., 2011). A 3D modeling representation of the PTE binding eIF4E is shown on the right where the label PK indicates the pseudoknot between the C-rich and G-rich regions (Wang et al., 2011).

Materials and Methods

Site Directed Mutagenesis

Plasmids pwheat_eIF4E_PET22b expressing full length wheat eIF4E and pwheat_eIF4E_ΔN-PET15 expressing truncated wheat eIF4E lacking the N-terminal 38 amino acids were obtained from Karen Browning. New England Biolabs Q5® Site-Directed

Mutagenesis Kit (EO554) was used to introduce mutations into the eIF4E ORF of both the full length and truncated plasmids. PCR primers used to construct the mutants are indicated 19 in table 2.1 while PCR conditions are specified in table 2.2. The following modifications were made to the manufacturer’s protocol. 2.5 µL of KLD mixture was added to 25 µL of

TOP 10 competent cells. 200 µL instead of 950 µL of SOC was added after heat shocking at

42 °C for 30 seconds and incubating on ice for 5 minutes. After incubating the transformation mixture at 37 °C for 20 minutes while shaking, 100 µL of the transformation mixture was plated onto 100 µg/ml ampicillin agar plates for overnight growth.

Table 2.1 Mutation location column indicates the region of eIF4E in which the mutation was made. Wheat eIF4E mutation column specifies the mutation where the first letter corresponds to wild type eIF4E amino acid one letter code, the number specifies the location of in the primary sequence of wheat eIF4E, and the last letter corresponds to the amino acid one letter code of the mutation. The third and fourth columns contain the forward and reverse primers, respectively designed to in order to produce every mutation.

Wheat Mutation eIF4E Forward Primer Reverse Primer location mutation eIF4G binding W79A 5' GCG GGC CTT TAC AAC AAT ATC CAT AAC CC 3' 5' GAA GTC CTC GAC GGT GGA GAA GG 3' site W49A 5' GCG TTC GAC AAC CCG CAG G 3' 5' GAA GGT CCA GGC GTT CTC GAG 3'

W62A 5' GCG GGG AGC ACC ATC CAC 3' 5' GGC CAC CTG CCT GGA CTT 3'

W108A 5' GCG GAA GAC CCC ATT TGT GCC 3' 5' TTT TGG CTC AAT CTT GTT CTT GAA GCA ATG G 3' Cap binding E109A 5' GCG GAC CCC ATT TGT GCC 3 5' CCA TTT TGG CTC AAT CTT GTT CTT GAA GCA 3' site R158A 5' GCG CAG AAA CAG GAA AGA GTA GCT ATC T 3' 5' CAC GCT AAC GAC TGC TCC ACA 3'

R163A 5' GCG GTA GCT ATC TGG ACC AAA AAT GCT 3' 5' TTC CTG TTT CTG ACG CAC GCT AAC 3'

W167A 5' GCG ACC AAA AAT GCT GCC AAT GAA GC 3' 5' GAT AGC TAC TCT TTC CTG TTT CTG ACG CAC 3' Outer surface F50A 5' GCG GAC AAC CCG CAG GGC 3' 5' CCA GAA GGT CCA GGC GTT CTC 3' near cap binding H67A 5' GCG CCC ATC CAC ACC TTC TCC ACC 3' 5' GAT GGT GCT CCC CCA GGC CAC CTG 3' site

Table 2.2 PCR conditions used in site directed mutagenesis to generate mutations in wheat eIF4E.

PCR Step Temperature °C Time

Initial denaturation 98 30 seconds

98 10 seconds

25 cycles 54 10 seconds

72 3 minutes

Final extension 72 2 minutes

Hold 4 ∞

Plasmid Purification

Two colonies of every mutant for each plasmid type (full length and truncated) were selected for plasmid purification. Qiagen QIAprep spin Miniprep kit was used to purify plasmids following manufacturer’s protocol. Purified plasmids were checked for quality via

1% agarose gel electrophoresis and quantified by Nanodrop. To verify the correct mutations, plasmids were sent to the Iowa State DNA facility for sequencing using the T7 promoter primer supplied by the facility.

Small Scale Induction

Plasmids coding for eIF4E mutants to be expressed were transformed into E. coli

BL21 competent cells. Five colonies per mutant were selected and transferred to 5 mL of 100

µg/ml ampicillin LB media for overnight shaking at 250 rpm at 37°C. 5 ml of 100 µg/ml ampicillin LB media was inoculated with 50 µL of saturated overnight culture and shaken at 21

250 rpm at 37°C until OD reached 0.8 using a wavelength of 600 nm. Cultures were then induced by addition of 2 µL of 1 M IPTG for a final concentration of 0.5 mM IPTG and shaken at 37°C for an additional 1.5 hours. Samples (1 ml) of induced (+IPTG) and pre- induced (-IPTG) were pelleted by centrifugation at 5000 rpm for 5 minutes. Samples were either subsequently analyzed by SDS-PAGE or stored at -20 °C for analysis the following day.

Soluble-Insoluble Fraction Separation

Induced cell pellets (+IPTG) and pre-induced pellets (-IPTG) were suspended in 100

µL of B-PER Bacterial Protein Extraction Reagent and incubated at room temperature for 10 minutes. Repelleting was accomplished by centrifuging at 5000 rpm for 5 minutes. The soluble supernatant was transferred into clean microfuge tubes. The pellet was resuspended in the insoluble solution indicated by table 2.3. Soluble +IPTG and –IPTG fractions and insoluble +IPTG and –IPTG fractions were analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS PAGE).

Table 2.3 The left hand column indicates the reagents used to make the insoluble fraction resuspension solution. The right hand column indicates the volume of reagent required per cell pellet to be resuspended.

Insoluble Resuspension

Component µl per sample

Water 57

Dnase 10X buffer 6.5

Dnase 1 1

Rnase H 0.5

Total 65

SDS PAGE Analysis

Resuspended cell pellets (6.5 µl per sample) to be analyzed were mixed with 2.5 µl

NuPAGE LDS buffer 4X and 1 µl NuPAGE reducing agent 10X. Samples were incubated at

70 °C for 10 minutes. Samples were loaded onto a NuPAGE 4-12% gel and run for 25 minutes at 200 V. Gels were washed by shaking in water for 10 minutes 3 times. Staining occurred by shaking the gel in GelCode™ Blue Stain Reagent for 1 hour. BioRad

ChemiDoc™ was used to obtain gel images.

Large Scale Induction

Plasmids coding for eIF4E mutants to be expressed were transformed into E. coli

BL21 competent cells. Five colonies per mutant were selected and transferred to 5 mL of 100

µg/ml ampicillin LB media for overnight shaking at 250 rpm at 37 °C. 5 ml of saturated overnight culture was added to 0.5 L of 100 µg/ml ampicillin LB media and shaken at 250 rpm at 37 °C until A600 reached 0.8, at which time they were induced by addition of 250 µL of 1 M IPTG for a final concentration of 0.5 mM IPTG. Induced cultures were shaken at 250 rpm at 37 °C for an additional 1.5 hours. Soluble and insoluble fractions were analyzed by

SDS PAGE as previously described.

Constructing GST-eIF4E Fusion Using pGEX-5X-2

Mutant full length eIF4E PCR

The ORF of all full length eIF4E mutants flanked by a 5’ BamH1 restriction site and

3’ Sal1 restriction site were amplified using Promega GoTaq® Green Master Mix (M712) protocol and the PCR conditions in table 2.4. PCR products were purified using the Qiagen

QIAquick PCR purification kit following manufacturer’s protocol. Quality of PCR products was checked via 1% agarose gel electrophoresis and quantified by Nanodrop. 23

Table 2.4 PCR conditions used to amplify full length eIF4E ORFs of wild type eIF4E and all mutant eIF4E for ligation into pGEX-5X-2. These PCR conditions were also used to check for eIF4E ORF insert presence after ligation in pGEX-5X-2.

PCR Step Temperature (°C) Time (minutes)

Initial denaturation 95 2

95 0.5

25 cycles 55 0.5

72 1

Final extension 72 5

Hold 4 ∞

Double restriction digest

Full length mutant eIF4E PCR products and GST-fusion plasmid PGEX-5X-2-2 were diluted to 5 ng/µL. The restriction digest buffer 3:1 NEBuffer and restriction enzymes

BamH1 and Sal1 HF were purchased from New England Biolabs. The double restriction reaction was incubated for 3 hours at 37 °C. Double restriction digest products were purified by the Qiagen QIAquick PCR purification kit following manufacturer’s protocol and assessed for complete digestion by 1% agarose gel electrophoresis.

Ligation and colony PCR

All full length mutant eIF4E double-digested PCR fragments were inserted into

BamH1 Sal1 HF double-digested PGEX-5X-2-2 plasmid using New England Biolabs T4

DNA ligase (M0202) protocol. A 3:1 vector DNA to insert DNA ratio was used and calculated via the NEBio Calculator. Ligation reactions were transformed using TOP 10 24 competent cells. Four colonies per ligated mutant construct were selected to check for eIF4E insert presence by colony PCR. Colony PCR was performed using Promega GoTaq® Green

Master Mix (M712) and PCR conditions in table 2.4. Colony PCR products were purified by

Qiagen QIAquick PCR purification kit following manufacturer’s protocol. Purified colony

PCR products were checked for quality via 1% agarose gel electrophoresis and quantified by

Nanodrop. Correct insert orientation and presence of desired eIF4E mutants were validated by sequencing performed by the Iowa State DNA sequence facility using the PGEX-5 primer supplied by the facility. eIF4E Purification

A pellet (~1 g) obtained from wild type wheat eIF4E large scale induction was resuspended in B-100 buffer (0.1 M KCl, 0.02 M HEPES-KOH, 0.01 mM EDTA, 1 mM

DTT, 0.1% (v/v) Glycerol, 1 protease inhibitor tablet). Cells were lysed by sonication using a

¼” sonication tip pulsing 30 seconds on and 30 seconds off for 3 minutes at 50% power followed by a 3 minute incubation on ice and then 30 seconds on and 30 seconds off for an additional 3 minutes at 75% power. Cell lysate was centrifuged at 11000 rpm for 30 minutes at 4 °C. Supernatant was subsequently filtered by an Acrodisc syringe filter for purification by Jena Bioscience m7GTP column. The column was prepared by transferring 300 µL of

Jena Bioscience m7GTP agarose beads to a 10 mL column and washing with 5 mL of water followed by 5 mL of B-100 buffer. Filtered sample (5 mL) was loaded on the column and washed twice, each time with 10 mL of B-100 buffer collecting wash flow through in 2 mL increments. Wild type eIF4E was eluted by four aliquots (500 µL) of elution buffer 1 (100

µM GTP in B-100) followed by four aliquots (500 µL) of elution buffer 2 (100 mM GTP in

B-100). Purification results were analyzed using SDS PAGE. 25

Electrophoretic Mobility Shift Assay (EMSA)

TPAV DNA template preparation

Thin paspalum asymptomatic virus (TPAV) template DNA used for in vitro transcription of PTE 3’CITE of TPAV was prepared using New England Biolabs Phusion®

High-Fidelity PCR Kit following manufacturer’s protocol. Template PCR products were purified by Qiagen QIAquick PCR purification kit following manufacturer’s protocol and analyzed for quality by 1% agarose gel electrophoresis. PTE template DNA was quantified by Nanodrop.

TPAV probe preparation

5’ m7GTP capped TPAV probe was radioactively labeled with [α-32P] CTP and prepared using T7 mMessage mMachine [Ambion AM1344] with a ratio of 4:1 for m7GTP to GTP following the manufacturer’s protocol. Uncapped TPAV probe was prepared using

T7 megashortscript [Ambion AM1354]. Transcripts were purified using Bio-Rad Micro Bio-

Spin® columns. Transcripts were quantified using a Liquid Scintillation Analyzer.

Transcripts quality check was achieved by running transcripts on a 10% acrylamide urea gel and visualizing by phosphoimager exposure.

Running EMSA

Concentrated wild type wheat eIF4E was obtained from Elizabeth Carino, Iowa State

University. 5% TBE polyacrylamide gels were poured into Thermofisher Novex Cassettes

(NC2010) and stored at 4 °C. Titration of eIF4E with capped or uncapped TPAV PTE was performed by incubating PTE (~2000 cpm), eIF4E (0 µM, 0.25 µM, 0.5 µM, 0.75 µM, or 1

µM), and EMSA mix [1X binding buffer (10 mM HEPES pH 7.5, 20 mM KCl, 1 mM DTT,

1 mM MgCl₂), 3 mM DTT, 2 µg/µL tRNA, 2 µg/µL BSA, 1 U RNaseout, Tris 10% glycerol] on ice for 25 minutes. Wells of a 5% TBE polyacrylamide gel were loaded with 6.5 µL of 26

EMSA reaction and 2 µL of 2X loading dye. The gel was run for 45 minutes at 110V and was dried for 1 hour at 75°C. After overnight exposure, EMSA gel results were visualized by

Bio-Rad phosphoimager.

Results

German-Retana et al., 2008 performed an innovative experiment to test the ability of lettuce eIF4E harboring point mutations to support VPg mediated Lettuce mosaic virus LMV infection and thus eIF4E-VPg binding. In this system, infectious recombinant LMV engineered to express host mutant eIF4Es or wild type eIF4E were used to infect resistant lettuce varieties. During recombinant LMV infection, eIF4E was expressed as a fusion protein between viral protein P1 and Hc-Pro domains, which is then converted to free eIF4E upon proteolytic cleavage in vivo. Therefore if mutant eIF4E expressed from recombinant

LMV was able to restore LMV susceptibility in resistant lettuce, the point mutation contained in such a mutant eIF4E is not important for VPg-eIF4E binding.

The results obtained by German-Retana et al., 2008 along with the crystal structure of wheat eIF4E provided the basis for designing point mutations in wheat eIF4E for the purpose of investigating amino acids important for PTE binding. The experimental design of this work was such that recombinant wheat eIF4E containing point mutations was to be expressed in E. coli and purified for investigation of PTE binding. Electrophoretic mobility shift assay

(EMSA) was to be used to evaluate relative mutant eIF4E-PTE binding affinity. However, solubility problems arising from the GST fusion method of purification limited the ability to purify mutant eIF4E. Therefore a significant portion of this work is concerned with attempts to solve the GST-eIF4E fusion solubility problem. 27

Designing Mutations of Wheat eIF4E

eIF4E point mutations associated with viral resistance are localized in two regions: inside the cap-binding pocket and on the solvent accessible side bearing the loop between beta sheets S1 and S2 near the cap binding pocket (Truniger and Aranda, 2009). In accordance with the study performed on lettuce eIF4E to test VPg binding, three classes of mutations were also introduced into wheat eIF4E to test PTE binding as shown in table 2.5.

In lettuce, mutant W94A was designed to disrupt the ability of eIF4E to bind eIF4G in order to assess the importance of eIF4F complex formation on eIF4E-VPg mediated LMV susceptibility (German-Retana et al., 2008). However, mutant W94A appeared to only partially disrupt eIF4G binding and was able to restore LMV susceptibility to a similar level as susceptible lettuce eIF4E (German-Retana et al., 2008). Ability to restore LMV susceptibility indicates that W94 is not likely important for LMV resistance (German-Retana et al., 2008). Although eIF4F complex formation was not tested in wheat eIF4E-PTE binding and W94 does not appear to be important for VPg binding, the analogous mutant, W79A, was introduced into wheat eIF4E in order to evaluate the potential involvement of the eIF4G binding site in PTE binding.

The class of mutants in the wheat eIF4E cap binding pocket include W49A, W62A,

W108A, E109A, R158A, R163A, and W167A where mutants W49A, W62A, W108A,

R158A, and W167A are based on analogous mutations made in lettuce eIF4E shown in table

2.5 and mutants E109A and R163A are based on the m7GDP crystal structure of wheat eIF4E (German-Retana et al., 2008; Monzingo et al., 2007a). All mutated wheat eIF4E residues are shown in figures 2.3 and 2.4 where cap binding eIF4E residues coordinated to m7GDP are shown in figures 2.5 and 2.6. 28

Two tryptophan residues in the cap binding pocket, W77 and W123 in lettuce and

W62 and W108 in wheat eIF4E, sandwich and bind the cap through stacking π bond interactions (Marcotrigiano et al., 1997). Mutant W123A was able to restore LMV susceptibility in lettuce indicating it is not important for VPg-eIF4E binding whereas the lettuce mutant W77L was shown to partially disrupt restoration of LMV susceptibility

(German-Retana et al., 2008). Differential LMV susceptibility restoration between these two cap binding tryptophans is not entirely surprising due to the positions of W123 and W77 and the observation that the VPg likely does not bind in a manner consistent with a cap analog.

Specifically, W123 is distal to the surface region associated with viral resistance whereas

W77 is proximal to this region as observed in figures 2.3 and 2.4 by the distance of wheat

W108 (homologous to lettuce W123) to surface mutants F50 and H67. In contrast to the

VPg, the PTE contains a hypermodifiable G that may act as a putative cap mimic in eIF4E binding (Wang et al., 2011; Wang et al., 2009). Therefore both mutants W62A and W108A, analogous to W77L and W123A in lettuce respectively, were made in wheat eIF4E with the expectation that both may impact PTE binding in a case such that the hypermodifiable G in the PTE acts as a true cap analog.

Mutants E109A, W167A, R158A, and R163A were made for their involvement in stabilizing cap binding and for the disrupted ability of their lettuce eIF4E analogous mutations (shown in table 2.7) to restore LMV infection in lettuce (German-Retana et al.,

2008; Marcotrigiano et al., 1997; Monzingo et al., 2007a). In murine eIF4E, E109 epsilon oxygen atoms OE1 and OE2 are proposed to hydrogen bond to the guanine nitrogen atoms

N1 and N2 respectively (Marcotrigiano et al., 1997) whereas wheat eIF4E E109 epsilon oxygen atom OE2 is located such that hydrogen bonding with N1 or N2 of guanine is 29 probable as shown in figure 2.5 (Monzingo et al., 2007a). A homologous mutation of wheat eIF4E E109A made in human eIF4E resulted in a loss of cap binding (Morino et al., 1996) supporting the hypothesis that PTE binding may be disrupted or negatively impacted by mutant E109A if the PTE acts as a cap analog. Situated in the bottom of the cap binding pocket, residue W167 is proposed to stabilize cap binding by forming van der Waals interactions with the methyl group bound to guanine nitrogen N7 shown in figure 2.5

(Marcotrigiano et al., 1997). The mutation W182A in lettuce (corresponding to W167A in wheat) disrupted cap binding (German-Retana et al., 2008) suggesting that the corresponding mutation in wheat may disrupt PTE-eIF4E binding. Mutant W49A was made based in part on the disrupted ability of the analogous lettuce mutation, W64A, to restore LMV infection

(German-Retana et al., 2008). Interestingly, W49 is partially surface accessible while also being in the cap binding pocket. Both cap binding tryptophans W62 and W108 are on opposing flexible loops, where W62 is on the flexible loop proximal to W49 shown in figure

2.3 and 2.4. It is therefore conceivable that the flexibility of the loop containing W62 and the accessibility of W49 may allow π electron stacking interactions, analogous to those that occur in W62 and W108 mediated cap binding, to occur between W62 and W49 and the hypermodifiable G of the PTE in PTE binding. Alternatively, it is possible W62 and W49 otherwise coordinate to form stabilizing hydrophobic interactions with the hypermodifiable

G of the PTE to facilitate PTE binding.

Proximal residues in eIF4E tertiary structure, residues R158 (R173 in lettuce) and

R163 are proposed to stabilize cap binding by interacting with the alpha and beta phosphate oxygen atoms of m7GDP shown in figure 2.5 and 2.6 (Marcotrigiano et al., 1997; Monzingo et al., 2007a). In contrast to cap binding mutants discussed prior, lettuce eIF4E mutant 30

R173A (R158A in wheat) allowed cap binding (German-Retana et al., 2008). Interestingly however, the ability of lettuce R173A to restore LMV susceptibility was disrupted (German-

Retana et al., 2008). Consequently, wheat eIF4E mutants R158A and R163A may not disrupt cap binding and therefore may not disrupt PTE binding if the PTE acts as a true cap analog.

However, wheat mutants R158A and R163A may disrupt PTE-eIF4E binding if the PTE binds eIF4E in a way such that positive charges on R163 and R158 are positioned to stabilize phosphate oxygen atoms on a neighboring portion of the PTE phosphate backbone.

The last class of wheat eIF4E mutants, consisting of H67A and F50A, were made on the surface region of eIF4E near the cap binding pocket associated with resistance (German-

Retana et al., 2008; Robaglia and Caranta, 2006; Truniger and Aranda, 2009). Wheat eIF4E mutation H67A corresponds to lettuce eIF4E mutation R82L (German-Retana et al., 2008), both of which are positively charged. R82L allowed cap binding and was able to restore

LMV susceptibility suggesting that R82 is neither important for cap binding nor for VPg binding (German-Retana et al., 2008). However, the homologous wheat eIF4E mutant H67A was still made because the positive charge may function in stabilizing PTE binding by coordinating with the negatively charged RNA phosphate backbone. A crystal structure of

Xenopus laevis RNA-binding protein A (Xlrbpa) bound to RNA in a non-sequence specific protein-dsRNA binding study revealed that positively charged Xlrbpa residue H141 does in fact coordinate with a negatively charged oxygen atom of the RNA phosphate backbone

(Ryter and Schultz, 1998). Furthermore wheat eIF4E mutant H67A is located not only in a region previously associated with resistance but also in a location such that coordination with the phosphate backbone is likely based on computer modeling studies of PTE-eIF4E binding shown in figure 2.2 (Wang et al., 2011). Lettuce eIF4E mutant F65A (F50A in wheat) 31 disrupted LMV susceptibility restoration and partially disrupted cap binding (German-Retana et al., 2008). It is noteworthy and very interesting that F65A located on the outer surface of eIF4E negatively impacted cap binding, which occurs inside the cap binding pocket. Analysis of the wheat eIF4E crystal structure by German-Retana et al., 2008 led to the proposal of a mechanism by which mutation of lettuce eIF4E F65A impacts cap binding where F65A might alter the orientation of other residues such that the bottom of the cap binding pocket is altered. Therefore the homologous wheat eIF4E mutant F50A may also negatively impact

PTE binding if the PTE acts as a true cap analog.

Table 2.5 Lettuce eIF4E mutations and results adapted from German-Retana et al., 2008 with the addition of parallel wheat eIF4E mutations. The cap binding tryptophans are indicated in bold. Wheat mutants E109A and R163A were made based on the wheat eIF4E crystal structure and do not have homologous lettuce eIF4E mutations.

Restore LMV Wheat Lettuce Cap binding susceptibility eIF4G binding W79A W94A + +

W49A W64A - - W62A W77L ± - W108A W123A - + Cap binding E109A

R158A R173A + - R163A

W167A W182A - -

F50A F65A ± - Outer Surface H67A R82L + ±

Figure 2.3 Identification of mutants made in wheat eIF4E (PDB file 2IDV) bound to m7GDP (shown in white) focusing on the bottom distal side of the cap binding pocket. The cap binding tryptophans W62 and W108 are shown in yellow. Blue indicates positively charged amino acids while red indicates negatively charged amino acids. Purple indicates neutral amino acids with the exception of W79, which is shown in orange to indicate its location in the eIF4G binding region. 33

Figure 2.4 Identification of mutants made in wheat eIF4E (PDB file 2IDV) bound to m7GDP (shown in white) focusing on the top proximal side of the cap binding pocket. The cap binding tryptophans W62 and W108 are shown in yellow. Blue indicates positively charged amino acids while red indicates negatively charged amino acids. Purple indicates neutral amino acids with the exception of W79, which is shown in orange to indicate its location in the eIF4G binding region.

Figure 2.5 m7GDP coordinated to eIF4E cap binding residues (PDB file 2IDV) with top proximal cap binding focus. m7GDP coloring is as follows: carbon is green, nitrogen is blue, oxygen is red, and phosphorous is orange. The epsilon oxygen of Glu109 (shown in red) is within hydrogen bonding distance of the hydrogen atoms bound to guanine nitrogen atoms N1 and N2. Try167 (shown in purple) is positioned such to form Van der Waals interactions the methyl group (CM7) on guanine N7. Amines of Arg163 and Arg158 (shown in blue) are within hydrogen bonding distance to oxygen atoms of the pyrophosphate. 35

Figure 2.6. m7GDP coordinated to eIF4E cap binding residues (PDB file 2IDV) with bottom distal cap biding focus. m7GDP coloring is as follows: carbon is green, nitrogen is blue, oxygen is red, and phosphorous is orange. The epsilon oxygen of Glu109 (shown in red) is within hydrogen bonding distance of the hydrogen atoms bound to guanine nitrogen atoms N1 and N2. Try167 (shown in purple) is positioned to form Van der Waals interactions with the methyl group (CM7) on guanine N7. The amine of Arg163 and the hydrogen on the epsilon nitrogen (NE) are both within hydrogen bonding distance to an oxygen (02B) on the beta phosphorous atom. The amine of Arg158 is within hydrogen bonding distance to an oxygen atom (O1A) on the alpha phosphorous atom.

Electrophoretic Mobility Shift Assay (EMSA) For Investigation Of eIF4E-PTE Binding

Electrophoretic mobility shift assay is a method used to detect protein-nucleic acid complexes based on differences in electrophoretic mobility of unbound nucleic acid molecules to nucleic acid bound to protein (Carey et al., 2013; Hellman and Fried, 2007).

Unbound RNA or DNA migrate to a greater extent compared to the migration of protein-

RNA or protein-DNA complexes (Carey et al., 2013; Hellman and Fried, 2007). Probe RNA or DNA are radioactively labeled and visualized via exposure to a phosphoimager. Thus relative fractions of bound and unbound probe can be quantified using programs such as Bio-

Rad Quantity One® 1-D analysis software. Consequently RNA or DNA probes can be titrated with increasing concentrations of protein. Fraction of probe bound vs protein concentration and be plotted to allow approximation of the dissociation constant Kd (Carey et al., 2013; Fried and Crothers, 1981). However, EMSA is most commonly used for qualitative binding studies (Hellman and Fried, 2007).

The EMSA method was chosen for this work with the intention of qualitatively identifying binding of the PTE from Thin paspalum asymptomatic virus (TPAV) to eIF4E harboring point mutations designed to affect PTE-eIF4E binding. Previous work performed to optimize TPAV EMSA conditions yielded conditions sufficient for this work (Sung Ki

Cho, Elizabeth Carino unpublished data). To confirm sufficiency of EMSA conditions for qualitative assessment of eIF4E-PTE binding, EMSA was first performed with wild type eIF4E titrating both a functional PTE and mutated PTE unable to bind eIF4E. The mutant

TPAV PTE (TPAV-m2) contains mutations in the C-rich region at the branch point of the

PTE stem loops similar to the mutant PEMV-m2 of the PTE in Pea enation mosaic virus

(PEMV) (Wang et al., 2009, Wang unpublished data). It is thought that loss of the mutant

PTEs’ ability to bind eIF4E is due to the disruption of the pseudoknot formed by the C rich 37 region at the stem loop branch point and the G-rich region in a bulge of the basal stem loop shown in figure 2.2, the G rich region of which contains the hypermodifiable G putatively involved in eIF4E binding (Wang et al., 2009).

The results of EMSA performed with functional TPAV PTE and nonfunctional

TPAV-m2 PTE are shown in figure 2.7B. A shift is observed in the titration of TPAV PTE with concentrations of wild type wheat eIF4E ranging from 0.25 µM to 1 µM increasing at increments of 0.25 µM relative to the migration of the TPAV PTE with 0 µM of eIF4E. As expected, no shift is observed in the titration of the TPAV-m2 PTE with concentrations of wild type eIF4E of the same range. Consequently EMSA conditions appear sufficient to differentiate between a positive and negative result of PTE-eIF4E binding. Although sufficient, further optimization could be done to better resolve the band smearing of TPAV-

PTE bound to eIF4E.

The ability of EMSA conditions to sufficiently differentiate between relatively strong and weak binding was also tested. TPAV PTE with a 5’ m7GTP cap translates with greater efficiency than uncapped TPAV PTE (Elizabeth Carino unpublished data). Addition of a 5’ m7GTP cap restores the ability of TPAV-m2 PTE to translate although with less efficiency than both capped TPAV PTE and uncapped TPAV PTE suggesting that capped TPAV-m2

PTE binds eIF4E with less affinity than capped TPAV PTE (Elizabeth Carino unpublished data). Therefore capped TPAV PTE and capped TPAV-m2 PTE were titrated with eIF4E to establish the ability of the EMSA conditions to differentiate between relatively strong and weak binding shown in figure 2.7A. A shift is observed upon titration of capped TPAV PTE with eIF4E ranging from 0.25 µM to 1 µM at increasing increments of 0.25 µM relative to free capped TPAV PTE. The same shift is observed for capped TPAV-m2 PTE for eIF4E 38 concentrations of 0.25 µM to 1 µM increasing at increments of 0.25 µM eIF4E. Qualitatively comparing the band intensities of bound to unbound probe show the ratio of bound capped

TPAV-m2 PTE to unbound capped TPAV-m2 PTE appears lower than the ratio of bound capped TPAV PTE to unbound capped TPAV PTE. The lower ratio of bound to unbound capped TPAV-m2 PTE relative to the ratio of bound to unbound capped TPAV PTE would normally be consistent with a lower binding affinity of TPAV-m2 PTE relative to TPAV

PTE. However, the band intensities of increasing concentrations of eIF4E appear the same indicating saturation at the lowest concentration of eIF4E. Consequently, the unbound bands likely correspond to a subpopulation of RNA in alternate conformations unable to bind eIF4E. Therefore relative binding strengths cannot be determined using these EMSA conditions. Further optimization is needed to both reduce band smearing and to allow for qualitative determination of relative binding strengths.

GST-eIF4E Fusion: Cloning eIF4E Mutants Into pGEX-5X-2 Vector

m7GTP-sepharose affinity chromatography is a common method used for purification of untagged eIF4E where eIF4E binds the column via m7GTP moieties and is eluted by 100

µM-100 mM GTP after adequate washing (Grzela et al., 2006; Mayberry et al., 2007; Rau et al., 1996; Salaün et al., 2003). Wild type wheat eIF4E was purified using an m7GTP- sepharose affinity column. However, eIF4E cap binding mutants were made such that the ability of some mutants to bind the cap analog would be putatively disrupted making purification by m7GDP column not feasible. An alternative method of purification was sought by cloning mutant eIF4E ORFs from the previous vector into the pGEX-5X-2 vector.

The pGEX vectors express the protein of interest N-terminally fused to glutathione S- transferase (GST) (Frangioni and Neel, 1993b). 39

4E µM Cap

Figure 2.7 Electrophoretic mobility shift assay (EMSA) of wild type wheat eIF4E and the PTE 3’CITE from Thin paspalum asymptomatic virus (TPAV). A) eIF4E concentrations are indicated by the row 4E mM and increase from 0 mM to 1 mM. The PTE from TPAV is the gel shift on the right labeled TPAV whereas the gel shift on the left is the mutant TPAV PTE m2 labeled TPAV-m2. The + indicates both the TPAV PTE and TPAV-m2 PTE have a 5’ m7GTP cap B) eIF4E concentrations are indicated by the row 4E mM and increase from 0 mM to 1 mM. The PTE from TPAV is the gel shift on the right labeled TPAV whereas the gel shift on the left is the mutant TPAV PTE m2 labeled TPAV-m2. The - indicates both the TPAV PTE and TPAV-m2 PTE do not have a 5’ m7GTP cap.

GST is a 26 kDa protein that binds glutathione with high affinity (Frangioni and Neel,

1993b). Thus affinity columns containing glutathione can isolate GST fusion proteins

(Frangioni and Neel, 1993b). A Factor Xa proteolytic cleavage site between the C-terminus of GST and the N-terminus of the protein of interest (eIF4E in this case) allows elution of the purified protein of interest by Factor Xa cleavage while GST remains bound to the column

(Frangioni and Neel, 1993b).

Sequencing upstream and downstream of the mutant eIF4E ORFs in their original vector showed that the ORFs are flanked by a 5’ BamH1 restriction site and a 3’ Sal1 restriction site. pGEX-5X-2 has a GST C-terminally situated multiple cloning site importantly containing the restriction site BamH1 upstream of the restriction site Sal1.

Therefore a double restriction digest using the restriction enzymes BamH1 and Sal1 could be performed to ligate eIF4E ORFs into pGEX-5X-2. However, analysis of the resultant reading frame revealed eIF4E ORFs would be out of frame compromising correct translation of eIF4E. Consequently, PCR was used to amplify eIF4E ORFs with a forward primer 5’ AA

TAA GGA TCC CCA TGG CCG AGG ACA CG 3’ containing an extra C base to adjust the reading frame where the added C is underlined and the BamH1 site is indicated in bold.

Additionally, 3 to 4 nucleotides were added upstream of the restriction site in both the forward and reverse primer to ensure proper substrate length for the respective restriction enzymes BamH1 and Sal1 upon double digestion of eIF4E ORF PCR products.

A vector map of pGEX-5X-2 with eIF4E ORF insert is shown in figure 2.8. eIF4E

ORF insert ligation for all mutants and wild type eIF4E into pGEX-5X-2 was confirmed by colony PCR. Four colonies per construct were chosen to check eIF4E ORF insert presence.

Two colonies per construct were chosen for sequencing to confirm the correct orientation of 41 the eIF4E ORF insert and ensure that the desired mutations were maintained for all constructs. All sequenced colonies showed the correct orientation of eIF4E insert and maintenance of desired mutation with the exception of two colonies. Colony 2 of mutant

W167A contained extra mutations K185E and K169E where lysine residues at positions 185 and 169 were both mutated to glutamate. Colony 1 of mutant W62A contained the extra mutation H40L where the histidine residue at position 40 was mutated to leucine. Therefore colonies 3 and 4 of mutant constructs W167A and W62A were also sequenced and were found to contain the desired mutation with no extraneous mutations.

GST-eIF4E Fusion Inclusion Bodies

pGEX-5X-2 is under the control of the tac promotor shown in figure 2.8 and is inducible by IPTG (Harper and Speicher, 2008). Previous studies revealed optimal induction of untagged eIF4E in the IPTG inducible pET plasmid under the control of the lac promotor occurs at 37 °C at an A600 of 0.9 using an IPTG concentration of 0.5 mM (Mayberry et al.,

2007). However, induction of GST-eIF4E fusion of wild type, mutant W49A and mutant

F50A under these conditions resulted in GST-eIF4E fusion in the insoluble fraction in the form of inclusion bodies. As shown in figure 2.9B expression bands of expected GST-eIF4E fusion size (~50 kDa) were observed in the insoluble fraction for lanes induced with IPTG.

Expression bands of ~0.25 kDa were also observed in the soluble fraction for lanes induced with IPTG shown in figure 2.9A suggesting premature proteolytic cleavage of the GST (26 kDa) and eIF4E (25 kDa) fusion protein consistent with cellular stress response (Hannig and

Makrides, 1998).

Figure 2.8 Plasmid map of pGEX-5X-2 with eIF4E ORF insertion. The 5609 base pair pGEX-5X-2_eIF4E plasmid has ampicillin resistance and 5 open reading frames (ORFs). The ORF of GST is indicated in light pink. The inserted eIF4E ORF is indicated in green. The factor Xa cleavage site is shown in light pink between the GST and eIF4E ORFs. The GST-eIF4E fusion ORF is under the control of the tac promotor. Restriction sites BamH1 and Sal1 are indicated at positions 934 and 1589 respectively. The lacI and lacZ ORFs, shown in dark pink, are under control of the lacIq and lac promotor respectively. The lac operator is shown in blue and the origin of replication is shown in yellow.

Stress response can be stimulated by protein aggregates induced by high temperatures through favoring hydrophobic interactions (Rosano and Ceccarelli, 2014; Schein and

Noteborn, 1988). Therefore the induction temperature was lowered to 30 °C in attempt to resolve the GST-eIF4E insolubility problem.

As shown in figure 2.10 the GST-eIF4E fusion protein was still observed entirely in the insoluble fraction upon induction at 30 °C. The bands putatively correlated to premature proteolytic cleavage of GST-eIF4E are also still observed upon induction at 30 °C. Therefore the induction temperature was further reduced to 25 °C according to previous success with aggregate prone recombinant protein expressed at 25 °C (Rosano and Ceccarelli, 2014 and references therein). Induction at 25 °C did not improve solubility as the GST-eIF4E fusion protein was still observed in the insoluble fraction shown in figure 2.11B. However, less premature proteolytic cleavage appears to have occurred evidenced by the partial disappearance of the second ~25 kDa band in the soluble fraction IPTG induced lanes shown in figure 2.11A. Less proteolytic cleavage indicates a reduced stress response triggered by misfolded protein (Hannig and Makrides, 1998). However, the reduction of stress response is likely due to a lower production of GST-eIF4E fusion protein at 25 °C rather than correct folding as the GST-eIF4E fusion protein still appears entirely in the insoluble fraction.

Figure 2.9 Expression of GST-eIF4E fusion at 37 °C. WT refers to wild type eIF4E whereas W49A and F50A refer to the respective eIF4E mutants. The – and + indicate pre-induced or induced expression by IPTG respectively. A) Soluble fraction in which the black arrow indicates the expected size of the GST-eIF4E fusion protein where no expression band is observed. The grey arrow indicates the expression bands likely due to premature proteolytic cleavage of the GST-eIF4E fusion protein due to E. coli stress response. B) Insoluble fraction in which the black arrow indicates the GST-eIF4E fusion protein. The * indicates W49A IPTG induced sample was compromised in preparation.

Figure 2.10 Expression of wild type eIF4E GST-eIF4E fusion protein at 37 °C and 30 °C. The – and + indicate pre-induced or induced expression by IPTG respectively. The soluble fraction is shown on the left side of the gel while the insoluble fraction is shown on the right side. The black arrow indicates the GST-eIF4E fusion protein in the insoluble fraction. The grey arrow indicates the expression bands in the soluble fraction likely due to premature proteolytic cleavage of the GST-eIF4E fusion protein due to E. coli stress response. It appears the E. coli stress response was reduced at 30 °C evidenced by the reduction in the probable premature proteolytic cleavage bands.

Figure 2.11 Expression of GST-eIF4E fusion at 25 °C. WT refers to wild type eIF4E whereas W49A and F50A refer to the respective eIF4E mutants. The – and + indicate pre-induced or induced expression by IPTG respectively. A) Soluble fraction in which the black arrow indicates the expected size of the GST-eIF4E fusion protein where no expression band is observed. The grey arrow indicates the expression bands likely due to premature proteolytic cleavage of the GST-eIF4E fusion protein due to E. coli stress response. B) Insoluble fraction in which the black arrow indicates the GST-eIF4E fusion protein. 47

Discussion

Further EMSA Optimization

Though EMSA conditions previously established are sufficient to determine binding versus no binding, further EMSA optimization are needed to allow for qualitative determination of binding strengths and to reduce band smearing. EMSA band resolution of eIF4E-PTE complexes could potentially be improved by lower salt concentration and running the gel for a shorter time in reduced temperatures (Hellman and Fried, 2007). The band smearing is likely due to eIF4E-PTE complex dissociation while the gel is running.

Running the gel for a shorter duration and in reduced temperatures may lower rates of dissociation while lower salt concentrations may increase electrostatic stabilization of the eIF4E-PTE complex (Hellman and Fried, 2007). To allow for assessment of relative binding strength, eIF4E concentration range should be greatly reduced such that a clear difference between fractions of PTE bound to fractions of PTE unbound is observable between increasing increments of lowered eIF4E concentrations. Such adjustments should allow better qualitative identification of mutant eIF4E constructs that partially impair binding of the

PTE and thus lower the PTE binding affinity. Obtaining relative binding strength information particularly for mutant eIF4E constructs that lower PTE binding affinity may be useful in deciding which constructs to continue for future experimentation as weak PTE binding may have be sufficient for conferring viral resistance while maintaining functional cap binding ability.

Solubility Problem

The solubility problem of GST-eIF4E fusion needs to be resolved for purification of mutant eIF4E. Although GST is highly soluble when expressed alone, GST fusion proteins can often be partially or completely insoluble upon lysis (Frangioni and Neel, 1993a). Such is 48 the case for the GST-eIF4E fusion protein, where reducing the temperature from 37°C to

30°C to 25°C did not result in soluble fusion protein. However, reducing the temperature further in concert with multiple alternative solubilization approaches may be taken in attempts to solve the solubility problem. A further reduction of the temperature to 20°C or

15°C while decreasing the IPTG concentration may improve solubility (Donovan et al.,

1996; Harper and Speicher, 2008). Additional decrease of the induction temperature may disfavor the hydrophobic interactions important for inclusion body formation sufficiently enough to allow solubilization of the fusion protein (Donovan et al., 1996; Harper and

Speicher, 2008; Rosano and Ceccarelli, 2014).

Decreasing the IPTG concentration may improve solubility by reducing transcription rates which accommodate the facilitation of proper folding (Donovan et al., 1996). The pGEX-5X-2 is under the control of the tac promoter, which was derived from the lacUV5 promoter and the trp promoter (de Boer et al., 1983; Harper and Speicher, 2008).

Importantly, the tac promoter is 7 to 10 times more efficient than the parental lacUV5 promoter (which is stronger than the lac promoter) and 2 to 3 times more efficient than the parental trp promoter (de Boer et al., 1983). Therefore reduced concentrations of IPTG may be more than sufficient relative to the previously optimized eIF4E induction conditions under the control of the lac promoter using 0.5mM IPTG as described by Mayberry (Mayberry et al., 2007). Additionally, to reduce promoter leakiness, pGEX-5X-2 contains the repressor molecule coding gene, lacI (Harper and Speicher, 2008), meaning the concentration of IPTG required for a desired level of induction is dependent on the cellular plasmid copy number

(Donovan et al., 1996). The concentration of IPTG required in such cases can range as low as

0.005-0.1mM (Donovan et al., 1996). In fact, a concentration as low as 0.062mM was 49 determined to be optimal for expression of GST under the control of the tac promoter in

DH5α cells (Donovan et al., 1996). Therefore, although BL21 cells were used in this study, the GST-eIF4E fusion has the potential to greatly benefit from lower IPTG concentrations in terms of solubility.

Furthermore, inducing at a lower OD may improve solubility by decreasing the metabolic burden affecting translation and also decreasing the proteolytic stress response triggered in part by nutrient deficiency (Donovan et al., 1996). Expression of eIF4E fused to maltose-binding protein (MBP) under the control of the tac promoter was induced at an OD as low as 0.2 (Morino et al., 1995).

Co-expression of chaperones may also useful in improving solubility. The E. coli chaperone protein teams DnaK/DnaJ/GrpE and GroEL/GroES function to refold misfolded proteins (Baneyx and Mujacic, 2004; Nishihara et al., 1998). Plasmids co-expressing these chaperone teams have been constructed including pKJE8 co-expressing DnaK/DnaJ/GrpE, pGro12 co-expressing GroES/GroEL, and pG-KJE7 co-expressing DnaK/DnaJ/GrpE and

GroES/GroEL (Nishihara et al., 1998). Therefore BL21 competent cells lines containing one or more of these plasmids may be useful in attempts to increase solubility by co-expression of chaperone proteins during GST-eIF4E expression to refold misfolded GST-eIF4E fusion proteins.

Correct folding of wheat eIF4E involves a novel disulfide bond between cysteine 113 and cysteine 151 (Monzingo et al., 2007b). However, the reducing environment of E. coli cytoplasm inhibits the formation of disulfide bonds (Baneyx and Mujacic, 2004).

Consequently, the lack of disulfide bond formation in eIF4E likely contributes to GST-eIF4E misfolding such that inclusion bodies are formed. Thioredoxin reductase is mutated in the E. 50 coli strain Rosetta gami (DE3) allowing cytoplasmic disulfide bond formation (Schagerlof et al., 2006). Therefore attempting GST-eIF4E expression in Rosetta gami DE3 cells in combination with alternative methods discussed prior may result in improved solubility.

In the case that sufficient GST-eIF4E solubilization does not occur by these methods, a different fusion tag will be necessary. Many different fusion or affinity tags have been developed for commercial use including polyarginine-tag (Arg-tag), polyhistidine-tag (His- tag), Strep-tag, calmodulin-binding peptide, FLAG, SBP, chitin-binding domain, and maltose-binding protein (Terpe, 2003). However, the maltose-binding protein (MBP) fusion tag should be chosen in the case where GST-eIF4E fusion cannot be solubilized as MBP- eIF4E fusions have already been successfully utilized in expression of synthetic gene coding for human eIF4E for purification (Morino et al., 1995). Additionally, MBP has been shown to increase solubility of fusion proteins (Sachdev and Chirgwin, 1999; Terpe, 2003).

Future Directions

If eIF4E point mutations disrupting or reducing PTE binding are able to be identified upon solubility problem resolution and continued testing, the basis of eIF4E-PTE binding disruption should be analyzed and used to design more eIF4E point mutations in efforts to disrupt PTE binding while maintaining cap binding. Among future wheat eIF4E point mutations to design are mutations similar to the mutation in melon eIF4E recessive resistance allele nsv where His228 is mutated to leucine (Nieto et al., 2006). Making similar mutations in wheat eIF4E is of great significance and interest because the nsv allele confers resistances to Melon necrotic spot virus (MNSV), which uses a 3’ CITE termed the I-shaped structure

(ISS) where all other recessive resistance alleles discovered thus far confer VPg-eIF4E mediated potyvirus resistance (Nieto et al., 2006; Truniger et al., 2017). Furthermore, the ISS binds eIF4F likely through the eIF4E subunit and contains a highly flexible G while the PTE 51 is known to bind eIF4E and is thought to bind eIF4E through a hypermodifiable G

(Nicholson and White, 2011). Perhaps the positive charge of melon eIF4E His228 functions to stabilize eIF4E-ISS binding by interacting with the negatively charged phosphate backbone as His228 is located on the rim of the cap binding pocket. In contrast however, wheat eIF4E contains a glycine residue at the position homologous to melon eIF4E His228.

Therefore searching for and mutating histidine residues or other positively charged residues lining the rim of the wheat eIF4E cap binding pocket to neutral residues may be of great interest in designing future mutants in attempts to disrupt wheat eIF4E-PTE binding while maintaining eIF4E cap binding ability.

Isolating eIF4E mutants that maintain cap binding ability while disrupting PTE binding are important for identifying functional eIF4E point mutations that confer viral resistance versus resistance conferring mutations that are nonfunctional and thus also act as eIF4E knockout (KO) mutants. Functional resistance is preferred over loss of function mutations due to the observation that KO alleles may limit plant growth or result in lethality in some cases (Bastet et al., 2017; Callot and Gallois, 2014). Importantly, plants have two forms of eIF4E, eIF4E and eIFiso4E, both of which exhibit gene redundancy (Bastet et al.,

2017; Dinkova et al., 2016). Consequently, plants have multiple viral susceptibility factors in terms of eIF4E and thus may need multiple eIF4E KO alleles to obtain effective resistance.

For example Arabidopsis has three genes coding eIF4E, eIF4E1, eIF4E2, and eIF4E3 and one gene coding for eIFiso4E (Robaglia and Caranta, 2006). Turnip mosaic virus (TuMV) requires eIF4E1 to infect Arabidopsis and Clover yellow vein virus (ClYVV) requires eIFiso4E (Robaglia and Caranta, 2006; Sato et al., 2005) yet double KO of Arabidopsis eIF4E1 and eIFiso4E results in lethality (Callot and Gallois, 2014). These observations 52 demonstrate that identifying functional resistance conferring point mutations in eIF4E are of greater significance over point mutations that disrupt both PTE binding and cap binding.

The ultimate goal of investigating eIF4E for mutations disrupting PTE-eIF4E binding and conferring functional resistance is to use gene editing tools such as CRISPR/Cas9 to engineer resistant crops (Chandrasekaran et al., 2016; Pyott Douglas et al., 2016).

CRISPER/Cas9 has already been used effectively in the transgene-free genomic editing of wheat (Zhang et al., 2016). In this way, using a transgene-free gene editing tool such as

CRISPER/Cas9 to engineer resistant crops, which rely on functional resistance conferring mutations, is important not only to avoid the complications of resistance mediated by KO alleles, but also may be of rising importance in the current climate of controversy surrounding genetically modified organisms (GMO).

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APPENDIX. BARLEY YELLOW DWARF VIRUS (BYDV) SUBGENOMIC RNA3 CONTAINS AN XRN1 RESISTANT (xrRNA) STRUCUTRE

Abstract

The luteovirus Barley yellow dwarf virus (BYDV) subgenomic RNA3 (sgRNA3) is able to be generated by incomplete Xrn1 degradation and thus contains an Xrn1 resistant

RNA (xrRNA) structure. Deletion analysis places the xrRNA structure within the first 5’ 67 nucleotides of BYDV sgRNA3. Dianthoviruses, related to luteoviruses, have been shown to contain an xrRNA structure of which a crystal structure has been obtained recently.

Bioinformatics analysis using the programs INFERNAL and dynalign suggest the BYDV sgRNA3 xrRNA structure may be different from that of dianthoviruses. A proposed secondary structure of BYDV sgRNA3 xrRNA was obtained by analysis with the programs

DotKnot and RNAalifold.

Introduction

Barley yellow dwarf virus (BYDV) is a positive sense single stranded RNA virus with a genome of 5.6 kb belonging to the genus Luteovirus (Miller et al., 1995; Miller et al.,

1988). BYDV does not have a 5’ cap or a poly(A) tail but instead facilitates translation by a

3’ cap-independent translation element (3’CITE) termed the BYDV-like cap-independent translation element (BTE) (Guo et al., 2001). 180 protein subunits of T = 3 icosahedra arrangement form the 25-28nm diameter BYDV virions (Ali et al., 2014). BYDV infects cereals such as wheat, barley, and oats and in some cases rice and maize with BYDV infection causing average yield losses of 11 to 33% (Miller and and Rasochová, 1997).

The genome organization of BYDV consists of 7 open reading frames (ORFs) shown in figure 3.1 (Miller et al., 1988; Smirnova et al., 2015). ORFs 1 and 2, coding for 39 kDa and 60 kDa protein respectively, are sometimes translated together via an infrequent -1 68 ribosomal frameshift event (Barry and Miller, 2002). ORF 2 codes for the RNA-dependent

RNA polymerase (Miller et al., 1988) where the exact function of the protein encoded by

ORF 1 is not known but is proposed to be a helicase (Habili and Symons, 1989; Miller and and Rasochová, 1997). BYDV produces three 3’ co-terminal subgenomic RNAs

(sgRNA). sgRNA1 is mRNA for ORFs 3, 3a, 4, and 5, and sgRNA2 is mRNA for ORF 6, while sgRNA3 does not contain any ORFs (Kelly et al., 1994). ORF 4, coding for a 17 kDa movement protein, overlaps and is contained entirely within ORF 3 (Miller et al., 1988). The

22 kDa coat protein (CP) is encoded by ORF 3 (Miller et al., 1988) and in-frame readthrough of ORF 3 stop codon results in CP with the 60 kDa read through domain (RTD) as the C terminus (Brown et al., 1996). However, proteolytic cleavage of a C-terminal portion of the

RTD occurs resulting in a truncated 51-58 kDa CP-RTD (Miller et al., 1995). The RTD is important for aphid transmission (Chay et al., 1996). ORF 3a is required for long-distance movement and is predicted to produce a protein ranging from 4.8-5.3 kDa (Smirnova et al.,

2015). ORF 6 codes for a 4 to 7 kDa protein of unknown function in vitro, which has not been detected in vivo despite much effort (Shen et al., 2006). Although no ORFs are contained, sgRNA3 is produced abundantly during infection and encapsidated (Kelly et al.,

1994). Interestingly, sgRNA3 has been shown to be produced from genomic RNA, sgRNA1, and sgRNA2 via incomplete degradation by the exoribonuclease Xrn1 in vitro shown schematically in figure 3.2 (Keisuke Komoda, unpublished data).

The main plant mRNA decay pathways are shown in figure 3.3. The normal mRNA decay pathway in both plants and mammals begins with shortening of the poly(A) tail by the deadenylation activity of CCR4-NOT or PARN (Belostotsky and Sieburth, 2009; Garneau et al., 2007). At this point, mRNA degradation can either proceed in the 5’ to 3’ direction or the 69

3’ to 5’ direction (Garneau et al., 2007). The exosome, a 10 to 12 subunit macromolecular complex, degrades mRNA in the 3’ to 5’ direction leaving an oligomer with a 5’ cap, which is degraded by the scavenger decapping enzyme DcpS in mammals (Garneau et al., 2007).

Conversely, degradation in the 5’ to 3’ direction begins with decapping by the dimeric decapping enzyme composed of DCP1 and DCP2 in mammals (Garneau et al., 2007) or the trimeric decapping complex composed of DCP1, DCP2 and the scaffold VCS in plants

(Belostotsky and Sieburth, 2009). Decapping in mammals is aided by the heptameric Lsm complex associated with the 3’ end of mRNA which is to be degraded (Garneau et al., 2007).

Upon decapping, 5’ to 3’ degradation is carried out by the exoribonuclease Xrn1 or Xrn4 in mammals or plants respectively (Belostotsky and Sieburth, 2009; Garneau et al., 2007).

Xrn1-resistant RNA (xrRNA) has previously been identified in flaviviruses and have been termed subgenomic flavivirus RNA (sfRNA) (Chapman et al., 2014; Pijlman et al.,

2008). sfRNAs are generated by incomplete 5’ to 3’ degradation where a sfRNA 5’ highly structured element functions to impede Xrn1 (figure 3.4) (Chapman et al., 2014; Moon et al.,

2012; Pijlman et al., 2008). A crystal structure of the sfRNA xrRNA element from Murray

Valley Encephalitis virus has been solved, elucidating the structural basis for Xrn1 resistance, along with the xrRNA structure from Zika Virus and West Nile Virus (Akiyama et al., 2016; Chapman et al., 2014; Pijlman et al., 2008). The 5’ end of sfRNA is passed through a ring-like structure formed by a three-way helical junction and stabilizing pseudoknot (Chapman et al., 2014). The 5’ end is unable to be brought through the ring-like structure (Chapman et al., 2014). In this way, the ring-like structure acts as a mechanical brace impeding degradation by Xrn1 (Chapman et al., 2014). sfRNA is proposed to play a role in mitigating the host immune response by inhibiting pathways involved in RNAi, 70 interferon response stress granules, and RIG-I signaling (Göertz and Pijlman, 2015;

Manokaran et al., 2015).

Until recently, no xrRNA structure had been identified in plant viruses. Plant viruses of the genus dianthovirus were investigated for and found to contain xrRNA (Jeffrey Kieft, data pending publishing). A crystal structure of the xrRNA of dianthovirus Sweet clover necrotic mosaic virus (SCNMV) revealed a structure different from that of flaviviruses yet similar in the formation of a protective ring (Jeffery Kieft, data pending publication). The ring-like structure in SCNMV xrRNA is facilitated by unwinding of a 5’ helix where the 3’ end of the unwound helix wraps around the 5’ end forming a pseudoknot with an apical loop shown in figure 3.5 (Jeffery Kieft, data pending publication).

The genera of dianthovirus and luteovirus are related through recombination (Miller et al., 1995), so we tested whether sgRNA3 of BYDV is generated by Xrn1. Indeed, BYDV has been shown to contain an xrRNA structure involved in the generation of BYDV sgRNA3

(Keisuke Komoda, unpublished data). Interestingly, sgRNA3 accumulates abundantly during

BYDV infection and is encapsidated by virions (Kelly et al., 1994). The work contained in this appendix is a repetition and extension of the work performed by Keisuke Komoda on

BYDV sgRNA3 xrRNA structure.

Figure 3.1 Genome organization of Barley yellow dwarf virus (BYDV). Boxes indicate open reading frames (ORFs). Numbers on the right indicate 5’ nucleotide position. Numbers on the left indicate 3’ nucleotide position. Subgenomic RNA1 (sgRNA1) contains ORFs 3 (coat protein CP), 4, 5, and 6. Subgenomic RNA2 (sgRNA2) contains ORF 6. Subgenomic RNA3 (sgRNA3) does not contain any ORFs.

Figure 3.2 Generation of BYDV sgRNA3 by incomplete Xrn1 mediated 5’ to 3’ degradation of gRNA, sgRNA1, sgRNA2. Xrn1 is shown in blue while sgRNA3 is shown in red. The black bars on the 5’ end of sgRNA3 indicate the 5’ structure impeding Xrn1 degradation. 72

Figure 3.3 Plant mRNA decay. The 5’ m7G cap structure is shown in yellow. The 5’ untranslated region (5’ UTR) is indicated by brackets while the black box indicates the open reading frame (ORF). The 3’ UTR is indicated by brackets and contains the poly(A) tail. Deadenylation occurs by CCR4-NOT or PARN shown in dark green. 5’ to 3’ mRNA decay pathway starts with decapping by the decapping complex shown in purple, which is composed of DCP2, DCP1 (conserved in mammals) and the scaffold protein (VCS). Upon decapping, Xrn4 shown in blue, degrades in the 5’ to 3’ direction. 3’ to 5’ degradation is performed by the exosome, a 10 to 12 subunit macromolecular complex shown in light green.

Figure 3.4 Structure of xrRNA from flavivirus Murray Valley Encephalitis (MVE) Virus (Chapman et al., 2014). The Xrn1 blocking ring surrounding the 5’ end is formed by helices P1 and P3 (Chapman et al., 2014). The pseudoknot forms between helix P4 and hairpin loop L3 (Chapman et al., 2014).

Figure 3.5 Modified from Jeffery Kieft pending publication. Structure of xrRNA from dianthovirus Sweet clover necrotic mosaic virus (SCNMV) (shown with kind permission of Jeffery Kieft). The 5’ end helix unwinds putatively allowing sequence downstream of the hairpin to form a pseudoknot with the hairpin apical loop. The crystal structure in the open conformation is shown in the bottom left where the 5’ end is in red, the sequence involved in the putative pseudoknot is in orange (Jeffery Kieft data pending publication). The bottom right shows the model of the closed conformation where the 5’ end is enclosed in the putative Xrn1 blocking loop formed by the pseudoknot shown in orange (Jeffery Kieft, manuscript submitted for publication).

Materials and Methods

In Vitro Transcription

DNA templates of sgRNA deletion constructs in the context of sgRNA2 were obtained from Keisuke Komoda. DNA templates for deletion constructs 1 to 6 were made by linearization by Sma1 whereas DNA templates for deletion constructs 7 to 9 were made by

PCR. The MEGAscript® T7 transcription kit was used following manufacturer’s protocol to generate [α-32P] CTP radioactively labeled transcripts for degradation by Xrn1. The following modifications were made to the manufacturer’s protocol: reaction volumes of all reaction components were halved, 0.5 µl of [α-32P] CTP and 0.5 µl of RNase Out were added per reaction, 50 ng of template DNA was used per reaction, and incubated at 37 °C for 2 hours. Transcripts were purified by Bio-Rad spin columns and assessed for quality by 8M urea 6% polyacrylamide gel electrophoresis and exposing to a phosphoimager for visualization.

Xrn1 Assay

The Xrn1 degradation assay master mix was prepared using the following components and volumes per reaction: 0.25 µl 10 mg/ml CPK (10 mg creatine kinase, 20 mM HEPES-KOH pH 7.4, 5 mM DTT, 50% v/v glycerol), 1.25 µl 10X sub mix (7.5 mM

ATP, 1 mM GTP, 250 mM creatine phosphate, 0.25 mM amino acids minus cysteine, 0.25 mM amino acids minus leucine, 0.8 mM spermine), 0.25 µl RNase Out, 9.25 µl TR buffer

(25 mM Tris, 192 mM Glycine), and 0.5 µl Xrn1 obtained from NEB. Radio-labeled deletion construct transcripts (1 µl) were added to 11.5 µl of Xrn1 assay master mix and incubated for

3 hours at room temperature. The reaction was stopped by cooling on ice. The resulting RNA degradation was assessed by 8 M urea 6% polyacrylamide gel electrophoresis and exposure to a phosphoimager for visualization. 76

5’ Monophosphate Preparation

RNA 5’ pyrophosphohydrolase (RppH) was obtained from New England Bio labs to remove the 5’ pyrophosphate from the 5’ ends of the transcripts. Deletion transcripts were incubated with RppH to yield 5’ monophosphate (the usual substrate for Xrn1), following the manufacturer’s protocol. The resulting 5’ monophosphate deletion transcripts were purified using Bio-Rad spin columns.

INFERNAL

The xrRNA sequence of Sweet clover necrotic mosaic virus (SCNMV) shown in figure 3.4 (Jeffery Kieft data pending publication) was blasted using NCBI blast tool to identify homologous sequences. Four sequences with percent sequence homology ranging from 86% to 100% were aligned and annotated with structure and were then used as input for

INFERNAL to create a covariance model (CM). The CM was used by INFERNAL to search an alignment of 75 BYDV sgRNA3 sequences for similar sequence and structure.

Dynalign

The xrRNA sequence of Sweet clover necrotic mosaic virus (SCNMV) shown in figure 3.4 (Jeffery Kieft ,data pending publication) and the BYDV sgRNA3 67 nucleotide fragment were used as input for the program Dynalign in attempts to identify common structures within the given low sequence homology between the two input sequences.

Pseudoknot Prediction

The sequence comprising the sg3 portion of deletion mutant Δ9 [nucleotides 5348 to

5415] was used as input for the program DotKnot to identify structures capable of forming pseudoknots. DotKnot analysis identified one structure containing one pseudoknot.

RNAalifold

The sg3 Δ9 sequence [nucleotides 5348 to 5415] was blasted using NCBI Blast tool to identify similar sequences. Results were comprised entirely of BYDV isolates. Six BYDV isolates with sequence homology ranging from 87% to 97% were chosen as input for

RNAalifold along with the BYDV sg3 Δ9 sequence for determination of a consensus sequence and consensus structure.

Results

BYDV subgenomic RNA3 (sgRNA3) deletion constructs were designed by Keisuke

Komoda based on truncations of sgRNA3 secondary structure prediction. Truncations of putative secondary structure were made to determine if long distance interactions in sgRNA3

3’ end are needed to form the Xrn1 resistant RNA (xrRNA) structure located at the 5’ end of sgRNA3. The deletion constructs, shown in figure 3.6, were made in the context of sgRNA2 to allow substrate length adequate for comparison of degree of degradation in Xrn1 assays.

Xrn1 Assay

Deletion construct transcripts are shown in figure 3.7. An extra band of smaller length is observed in sg3, Δ7, Δ8 and Δ9. The migration pattern of the weak extra band observed in sg3, Δ7, Δ8 and Δ9 is consistent with early termination of T7 polymerase (Macdonald et al.,

1993). The same early termination band is likely present for sg2, Δ1, Δ2, Δ3, Δ4, Δ5, and Δ6, although not observed due to the decreased transcript concentration of these constructs relative to the transcription efficiency of sg3, Δ7, Δ8 and Δ9. The observed difference in transcript concentration is likely attributed to a difference in template concentration.

Template DNA for sg2, Δ1, Δ2, Δ3, Δ4, Δ5, and Δ6 was prepared by plasmid linearization using Sma1 whereas template DNA for sg3, Δ7, Δ8 and Δ9 was prepared by PCR. 50ng of 78 template DNA was used for in vitro transcription resulting in higher DNA template concentrations in terms of molarity for constructs made by PCR.

Xrn1 assay of all deletion constructs shown in figure 3.8 resulted in a band generated by impeded degradation by Xrn1 for all constructs as shown in figure 3.9. However, a sgRNA3 band was consistently absent in sgRNA2 input RNA likely indicating a problem with the template DNA sequence of sgRNA2. Additionally, bands generated by impeded

Xrn1 of deletion constructs could only be observed upon digital overexposure of the gel image indicating a low concentration of incomplete Xrn1 degradation products. Consistent with this observation is the presence of a strong band in wells with Xrn1 of equal size to lanes without Xrn1 for every construct indicating a high concentration of transcripts not degraded by Xrn1. The decreased activity of Xrn1 is likely due to improper substrate composition. The normal substrate of Xrn1 is mRNA with a 5’ monophosphate (Garneau et al., 2007) however, in vitro transcription results in transcripts with a 5’ triphosphate.

Therefore the 5’ triphosphate of deletion construct transcripts needed to be cleaved to a 5’ monophosphate in order to return Xrn1 activity to appropriate levels.

Sg2 and deletion constructs Δ1, Δ2, Δ3, Δ4, Δ5, Δ6 and Δ9 were chosen for continued analysis as the sgRNA3 xrRNA structure likely does not involve long distance interactions with the more distal 3’ end. The requirement of only 5’ proximal nucleotides is evidenced by the observation that deletion construct Δ9 was sufficient to produce a band generated by incomplete Xrn1 degradation shown in figure A.9.

RNA 5’ pyrophosphohydrolase (RppH) is a bacterial enzyme that cleaves the pyrophosphate of 5’ triphosphate RNA resulting in RNA with a 5’ monophosphate (Deana et al., 2008). RppH was used to prepare transcripts of sg2, Δ1, Δ2, Δ3, Δ4, Δ5, Δ6 and Δ9 with 79 a 5’ monophosphate. 5’ monophosphate transcripts, shown in figure A.9, appear too weak due to 5-fold dilution needed for the RppH reaction relative to the preparation volume of 5’ triphosphate transcripts. Xrn1 assay with 5’ monophosphate deletion construct transcripts as input RNA is shown in figure 3.10. Nearly all input RNA was degraded to a large degree by

Xrn1 indicated by the very faint strength or absence of the band in lanes with Xrn1 of equal migration distance to wells without Xrn1. Therefore Xrn1 activity appears to have been greatly improved by use of 5’ monophosphate input RNA compared to 5’ triphosphate input

RNA. However, the low concentration of input RNA in concert with an excessive Xrn1 concentration optimized for 5’ triphosphate transcripts resulted in complete degradation of transcripts with the exception of Δ9, which had a higher input concentration indicated by the slightly greater strength of the input RNA Δ9 band. The grey arrow in figure 3.10 points to the faint Δ9 band generated by incomplete Xrn1 degradation. The faint broad bands at the bottom of figure are likely free CTPs.

Bioinformatics Analysis

Deletion construct Δ9 was sufficient to produce a band generated by incomplete Xrn1 degradation in every Xrn1 assay performed. Therefore it appears the xrRNA structure of sgRNA3 is located within the 67 nucleotide segment from nucleotide position 5348 to 5415.

As dianthoviruses and luteoviruses are related (via proposed recombination (Miller et al.,

1995)), it is conceivable that the xrRNA structure of the luteovirus BYDV sgRNA3 might be similar to that of dianthoviruses. INFERNAL is a program that creates a covariance model

(CM) from a structurally annotated alignment of sequences and uses the CM to search other alignments for sequences that fit the CM in terms of sequence and structure (Nawrocki and

Eddy, 2013). An alignment of homologous sequences to the xrRNA sequence of the dianthovirus Sweet clover necrotic mosaic virus (SCNMV) was annotated with the xrRNA 80 structure and used as input for INFERNAL to create a CM. INFERNAL then used the CM to search an alignment of 75 BYDV sgRNA3 sequences. No hits were found, meaning either the xrRNA of BYDV sgRNA3 does not form a similar structure to that of dianthovirus or dianthovirus xrRNA and BYDV sgRNA3 xrRNA do form a similar structure but the sequence divergence between them is too great for INFERNAL to identify the common xrRNA structure in BYDV sgRNA3.

Dynalign is a program that searches for common structures and aligns sequences simultaneously therefore identifies common structures without the need for high sequence homology (Mathews and Turner, 2002). The sequence of the SCNMV xrRNA and the sequence of the BYDV sgRNA3 67 nucleotide fragment were used as input to Dynalign in attempts to find a similar structure in BYDV sgRNA 3 despite high sequence divergence. No similar structure was identified indicating that the xrRNA structure of BYDV of sgRNA3 is likely different than that of dianthoviruses.

The ring structure that forms around the 5’ end of flavivirus sfRNA to block Xrn1 degradation is stabilized by a pseudoknot between helix P4 and hairpin loop L3 shown in figure 3.3 (Chapman et al., 2014), while the ring structure in xrRNA structure of dianthovirus

SCNMV putatively requires a pseudoknot between the stem loop and sequence downstream of the stem loop shown in figure 3.4 (Jeffery Kieft, data pending publication). Thus it is a reasonable assumption that the xrRNA structure of BYDV sgRNA3 might require a pseudoknot. To identify structures in the sgRNA3 fragment of interest capable of forming pseudoknots, the sequence of the 67 nucleotide fragment [5348 to 5415] was used as input for the program DotKnot. DotKnot is a pseudoknot prediction program that selects regions of the secondary structures obtained from probability dot plots to construct pseudoknots using 81 pseudoknot free energy parameters (Sperschneider and Datta, 2010). Shown in figure 3.12, one structure containing one putative pseudoknot was obtained from the sgRNA3 67 nucleotide fragment using DotKnot. The resultant structure contains three stem loops (SL) where SL1 and SL2 form the pseudoknot. The sequence UGGU in the apical loop of SL1 is proposed to form a kissing stem loop pseudoknot with the sequence ACCA in the apical loop of SL2.

To evaluate the plausibility of the structure and potential pseudoknot identified by

DotKnot, sequence and structure conservation analysis was performed. The sgRNA3 67 nucleotide fragment was blasted using NCBI blast tool to obtain homologous sequences.

Blast results were comprised entirely of BYDV isolates. Five BYDV isolate sequences with percent sequence homology to the input sequence (BYDV sgRNA3 67 nucleotide fragment) of 87, 88, 90, 91, and 97% respectively, were chosen for analysis with the program

RNAalifold. The sequences chosen comprise the maximum range of percent sequence homology within the NCBI blast results. Selecting maximum range of percent sequence homology is important for the function of RNAalifold, which uses sequence covariation and thermodynamic parameters to predict a consensus sequence and consensus structure for a given set of aligned sequences (Hofacker, 2008).

RNAalifold analysis of the BYDV sequences resulted in a consensus structure identical to the structure predicted by DotKnot from nucleotides 1 to 52 in figure 3.11, importantly preserving SL1 and SL2. However, the consensus sequence determined by

RNAalifold, shown in figure 3.12, revealed a sequence variation weakening the potential pseudoknot between SL1 and SL2. The consensus sequence contains a UAGU in the apical loop of SL1 instead of UGGU requiring a noncanonical AC base pair to form the full four 82 base pair predicted pseudoknot with the ACCA sequence in the apical loop of SL2. The noncanonical AC base pair weakens but does not abolish the probability of the putative pseudoknot. At least two different geometric classifications of the noncanonical AC base pair exist and both have been observed in the crystal structure of the 4.5S RNA in bacterial SRP

(signal recognition particle) (Jovine et al., 2000; Leontis and Westhof, 2001). Therefore the complete conservation of SL1 and SL2 in terms of structure and the conservation of the putative pseudoknot in terms of sequence, albeit weaker than structure conservation, provide promise for the possibility of the proposed BYDV sgRNA3 putative xrRNA structure.

However, biochemical analysis should be performed to further support or to refute the proposed structure.

Figure 3.6 Deletion constructs of sgRNA3 in the context of sgRNA2. The 5’ end of sgRNA2 is indicated by nucleotide position 4809 on the left and extends to the 3’ end nucleotide position 5677 on the right. The 5’ end of sgRNA3 starts at nucleotide position 5348 and extends to the 3’ end at nucleotide position 5677. Deletion constructs are indicated on the far left labeled Δ1 through Δ9. The deleted fragment from sgRNA3 of every construct is indicated by the nucleotide position on the left side of the gap to the nucleotide position on the right side of the gap. 83

Figure 3.7 5’ triphosphate sgRNA3, sgRNA2 and deletion construct transcripts produced by in vitro transcription. Transcripts were run on 8M urea 6% polyacrylamide gel at 65 Watts for 45 minutes.

Figure 3.8. Xrn1 assay of 5’ triphosphate sgRNA3, sgRNA2 and deletion constructs Δ1 through Δ9 digitally overexposed. In the row labeled Xrn1, the – indicates Xrn1 was not added to the assay while the + indicates Xrn1 was present in the assay. The row labeled input RNA indicates the construct assayed with and without Xrn1.

Figure 3.9 5’ monophosphate sgRNA3, sgRNA2 and deletion construct transcripts produced by in vitro transcription and assayed with RppH to remove the 5’ pyrophosphate. Transcripts were run on 8M urea 6% polyacrylamide gel at 65 Watts for 45 minutes

Figure 3.10 Xrn1 assay of 5’ monophosphate sgRNA3, sgRNA2 and deletion constructs Δ1 through Δ6 and Δ9 digitally overexposed. In the row labeled Xrn1, the – indicates Xrn1 was not added to the assay while the + indicates Xrn1 was present in the assay. The row labeled input RNA indicates the construct assayed with and without Xrn1. The arrow indicates the faint Δ9 band generated by incomplete Xrn1 degradation.

Figure 3.11 Secondary structure of putative xrRNA BYDV sgRNA3 67 nucleotide fragment. Stem loop 1 (SL1) is comprised of nucleotides 1 to 21. Stem loop 2 (SL2) is comprised of nucleotides 30 to 52. Stem loop 3 (SL3) is comprised of nucleotides 54 to 65 and was not conserved in the RNAalifold consensus structure. A putative pseudoknot forms between the circled UGGU sequence in apical loop SL1 and the circled ACCA sequence in the apical loop of SL2.

Figure 3.12 RNAalifold results of putative xrRNA BYDV sgRNA3 67 nucleotide fragment. The BYDV sgRNA3 67 nucleotide fragment sequence is shown in blue. The RNAalifold consensus sequence is shown in green. RNAalifold consensus sequence shows UAGU instead of UGGU at nucleotide position 7 to 11 (boxed) in the apical loop of SL1, which weakens the pseudoknot formed with the conserved ACCA at nucleotide position 42 to 45 (boxed) in the apical loop of SL2.

Discussion

Visualization of Xrn1degradation assay by gel electrophoresis results should be improved to confirm the generation of sgRNA xrRNA fragments in deletion constructs. To improve transcription efficiency to a level similar to sg3 and Δ7 through Δ9, the template

DNA for constructs Δ1 through Δ6 and sg2 should be prepared by PCR. The transcripts should be purified by gel electrophoresis rather than by Bio-Rad spin columns in order to remove the extra band presumably generated by early termination of T7 polymerase. Upon removing the 5’ pyrophosphate of purified transcripts, the 5’ monophosphate transcripts should be concentrated by ethanol precipitation to avoid poor band visibility by over dilution.

The putative BYDV sgRNA3 xrRNA secondary structure and pseudoknot identified by DotKnot and supported by RNAalifold analysis should be further supported or refuted by biochemical analysis. Selective 2’-hydroxyl acylation analyzed by primer extension sequencing (SHAPE-seq) is a method whereby the 2’-hydroxyl of flexible, unpaired or unblocked nucleotides is modified by a shape reagent such as 1-methyl-7-nitroisatoic anhydride (1M7) in a non-denaturing buffer solution (Watters 2016). The modified RNA can then be reverse transcribed where the reverse transcription machinery falls off more often at modified nucleotide positions creating a DNA library of different lengths reflecting modified nucleotide positions (Watters 2016). After Illumina sequencing of the DNA primer extension products, reactivity data per nucleotide position can be obtained by analyzing the DNA library sequences with the program Spats (Watters 2016). Therefore SHAPE-seq can be used to support or refute the candidate secondary structure and potential pseudoknot of xrRNA

BYDV sgRNA3. Nucleotides involved in the helix of SL1 and SL2 and nucleotides in the loops of SL1 and SL2 involved in the pseudoknot shown in figure 3.12 should have poor

SHAPE-seq reactivity to support the xrRNA structure of BYDV sgRNA3 proposed. 87

However, the xrRNA secondary structure of BYDV sgRNA3 proposed does not explicitly explain how Xrn1 may be impeded. X-ray crystallography to obtain a tertiary structure is one method by which this question could be addressed. If the BYDV sgRNA3 xrRNA secondary structure proposed by DotKnot is supported by SHAPE-seq, it would be interesting to see if the helix of SL1 containing the 5’ end partially unwinds like that of dianthovirus (Jeffery Kieft data pending publishing). Perhaps then, the 5’ end may be passed through a loop formed by the pseudoknot of SL2 and the apical loop of partially unwound

SL1, thereby creating a ring-like structure that would block Xrn1.

Acknowledgments

I thank Jeffrey Kieft for allowing me to show unpublished structure, Dr. Keisuke

Komoda for advice on Xrn1 assays and Dr. Walter Moss for advice on RNA structure prediction.

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