A Dissertation Submitted to the Faculty of the Graduate School of Arts And

CHARACTERIZATION OF GII NOROVIRUS VPG NUCLEOTIDYLYLATION AND NUCLEOSIDE TRIPHOSPHATE BINDING

A Dissertation submitted to the Faculty of the Graduate School of Arts and Sciences of Georgetown University in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Microbiology

Alexei Medvedev, B.S.

Washington, DC December 6, 2016

CHARACTERIZATION OF GII NOROVIRUS VPG NUCLEOTIDYLYLATION AND NUCLEOSIDE TRIPHOSPHATE BINDING

Alexei Medvedev, B.S.

Thesis Advisor: Dr. Brent Korba, Ph.D.

ABSTRACT

Human Norovirus (hNoV) is a single-stranded positive-sense RNA virus of Caliciviridae family. It is the major cause of gastroenteritis outbreaks in the United States. The VPg protein of hNoV is a multi-functional protein essential for virus replication. Sequential and single-point alanine substitutions revealed that several positively charged amino acids in the N-terminal region of hNoV VPg regulate its nucleotidylylation. I provide evidence that hNoV VPg directly binds nucleoside triphosphates (NTPs), that inhibition of binding inhibits nucleotidylylation, and that the NTP binding appears to involve the first 13 amino acids of the protein. Substitution of multiple positively charged amino acids within the first 12 amino acids of the N-terminal region reduces nucleotidylylation without affecting binding. Substitution of only Lys20 abolishes nucleotidylylation, but not NTP binding. These studies indicate that positively charged amino acids in the first 20 amino acids of hNoV VPg regulate its nucleotidylylation through several potential mechanisms. Additionally, I demonstrate that the un-cleaved viral precursor protein,

ProPol (NS5-6) is 100-fold more efficient in catalyzing VPg nucleotidylylation than the mature polymerase (Pol, NS6), suggesting a specific role for ProPol during the hNoV infection.

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The research and writing of this dissertation is a monument to everyone who helped me along the way.

First and foremost, I would like to thank my amazing mentor, Dr. Brent Korba. It is hard to imagine that I could have done my thesis work anywhere else, but your lab. You instilled in me a special kind of appreciation for the scientific process, one that is not always straightforward, having a distinct set of guidelines, but, rather, one that requires creative thinking, ability to adjust on the fly, and just a tad of luck. Thank you for letting me run wild with my ideas and for always finding positives in every outcome, good or bad. Most importantly, thank you for being a great fatherly figure, caring and understanding, especially in the last (and very difficult) year of my thesis work. I can proudly admit that I never had to “work” in the lab under your guidance.

Thank you, my lab-mates and co-mentors, Drs. Jared May and Prasanth Viswanathan. Your friendship and scientific advice were invaluable in my growth as a scientist. My sanity is relatively intact, all thanks to the fun and collaborative environment you helped me create in our lab. I also would like to acknowledge Dr. Brittany Griffin, who was very instrumental in my development as a scientist in the early stages of my PhD education. Thank you for always fighting for my right to do science. I think the scientific community suffered a tremendous loss when you hung up your lab coat, and I hope you can find your way back one day.

Dear members of my Thesis Committee, Drs. John Casey, William Fonzi, Kim Green, and Radhakrishnan Padmanabhan, thank you for guiding me along on this journey. Thank you for pushing me forward when I needed it and for acknowledging my progress when I did something worthy. You helped me create high standards for my scientific work and were instrumental in making sure that I do not fall short of expectations.

Big thanks to my friends Vanya Kamyshko, Zo Noor, and Roma Dogadin. I will never be able to thank you enough for your support in my work, but, even more importantly, in my life outside the lab. You have come through for me on more than one occasion and helped me keep a clear and calm mind in the darkest, most turbulent times. With you, I never feel alone.

Thank you, Katerina Satanovsky and the rest of the Satanovsky clan for helping me start this journey and for supporting me along the way. Most importantly, thank you, Anya Stolpe, for helping me complete that journey. Your unprecedented support and believe in my abilities have given me strength and confidence to finish my work. You make me a better man.

My family. Where would I be without you? Great many thanks to my mother, Tamara Medvedeva, for allowing her little boy to leave for America, an impossible decision for any parent. Thank you, my grandparents, Viktor and Valentina Palanevich, as well as Tamara and Gennady Medvedev for stoking fires of curiosity and instilling great human values in me from an early age. The rest of my Belarussian, Canadian, German, and American relatives, thank you very much for always keeping me in your thoughts and checking up on me from time to time. My mom-away-from-mom, Natasha Medvedev, thank you for taking care of me like your own son. My little siblings, Maxim, Nikita, and Ksenya, thank you for showing me how to be cool and how to have fun. My lovely sister Anastasia, I am in awe of your superhuman abilities to do everything you do. Thank you for inspiring me to work harder, and a special thank you for bringing Milena, a constant beacon of light, into this life.

Finally, thank you, my father, Vladimir Medvedev. None of this would be possible without you. Period. Your relentless scientific and academic pursuits opened up a path to America for me and the rest of our family. You are the most brilliant, driven, and yet humble man I have ever encountered. I have looked up to you my whole life and I would be extremely happy someday to be even half the man and scientist you are. Thank you for your continued support in my work and in my life, and thank you for being the best father a son could ever ask for. I dedicate this work to you.

Many thanks,

ALEXEI MEDVEDEV

TABLE OF CONTENTS

I. Introduction ...... 1

II. Background ...... 5

A. Picornavirus VPg Biology ...... 5

B. Potyvirus VPg Biology ...... 7

C. Calicivirus VPg Biology ...... 11

III. Materials and Methods ...... 14

A. Cloning and Mutagenesis ...... 14

B. Protein Expression ...... 16

C. hNoV Polymerase (Pol) Assay ...... 18

D. hNoV VPg Nucleotidylylation Assay ...... 19

E. hNoV VPg NTP Binding Assay ...... 21

F. Structure Modeling ...... 21

IV. Characterization of Human Norovirus VPg ...... 23

A. hNoV VPg Nucleotidylylation Occurs Through Covalent Linkage to the α-Phosphate

of the Incoming Nucleotide ...... 24

B. Nucleotidylylation of Native hNoV VPg by hNoV ProPol Is Significantly Greater Than

Nucleotidylylation Catalyzed by hNoV Pol ...... 26

C. GMP and UMP Are Preferentially Added to hNoV VPg ...... 28

D. Positively-Charged Amino Acids in the N-Terminus of hNoV VPg Regulate Its

Nucleotidylylation by hNoV ProPol ...... 28

E. The Central Core Region of hNoV VPg Does Not Contain Amino Acids Required

for the Stability of the Nucleotidylylation Reaction ...... 31

F. hNoV VPg Binds NTPs in the Absence of Polymerase ...... 33

V. Future Work ...... 35

A. Plasmon Resonance Required to Study NTP Binding by hNoV VPg ...... 35

B. hNoV VPg Is a Protein Primer for Initiation of hNoV Genomic RNA Synthesis ...... 36

C. Protease Domain of Intact hNoV ProPol Is Required for Interaction with hNoV VPg

and Its Subsequent Nucleotidylylation ...... 39

VI. Conclusions ...... 41

Appendices ...... 46

Appendix A: Assay Optimization ...... 46

hNoV Polymerase Assay Optimization ...... 46

hNoV VPg Nucleotidylylation Assay Optimization ...... 49

Appendix B: Cloning Strategy for Tag-Less hNoV VPg Constructs ...... 51

Bibliography ...... 53

LIST OF TABLES

Table 1: Amino Acid Substitutions in hNoV VPg ...... 29

Table 2: Tag-Less hNoV VPg Cloning Primers ...... 52

LIST OF FIGURES

Figure 1: Overview of hNoV Genome Structure and Transcription/Translation Products ...... 2

Figure 2: Amino Acid Sequence Alignment of VPg Proteins from Various NoV Genogroups ...... 24

Figure 3: hNoV Nucleotidylylation with Either α-32P GTP or γ-32P GTP ...... 25

Figure 4: Nucleotidylylation of His6-Labeled and Native hNoV VPg ...... 26

Figure 5: Nucleotidylylation of Native hNoV VPg by hNoV Pol and hNoV ProPol ...... 27

Figure 6: Nucleotidylylation of Native hNoV VPg by Different NMPs ...... 28

Figure 7: Nucleotidylylation of Alanine-Substituted hNoV VPg ...... 30

Figure 8: Predictive Homology Model of hNoV VPg ...... 32

Figure 9: hNoV VPg Structural Disorder Analysis ...... 33

Figure 10: NTP Binding by hNoV VPg ...... 34

Figure 11: GTP Binding of Alanine-Substituted hNoV VPg ...... 35

Figure 12: hNoV Genomic RNA Replication ...... 36

Figure 13: Effects of His6-VPg on RNA Synthesis by Pol and ProPol ...... 38

Figure 14: Nucleotidylylation of hNoV VPg Derived from C-Terminally Placed Intein Construct ..... 40

Figure 15: The Relationship Between hNoV VPg N-Terminal Charge and Nucleotidylylation ...... 43

Figure 16: hNoV Polymerase (Pol) Assay Optimization ...... 47

Figure 17: hNoV VPg Nucleotidylylation Assay Optimization ...... 49

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I. INTRODUCTION

Norovirus (NoV) is a non-enveloped, single-stranded, non-segmented, positive-sense

RNA virus belonging to the Caliciviridae family [43, 44, 56]. There are 7 NoV genogroups

(GI-GVII) that have been recognized based on the amino acid identity of the major viral capsid protein, with 3 genogroups capable of infecting humans (GI, GII, GIV) [124, 136]. As such, human NoV (hNoV) is the predominant cause of gastroenteritis worldwide, responsible for 21 million cases per year in the United States, and accounts for 10-15% of severe cases of gastroenteritis in children aged less than 5 years old [38, 48, 49]. Of the three hNoV genogroups

(GI, GII, GIV), GII hNoV has been the major cause of gastroenteritis outbreaks in the United

States since 2001 [124]. hNoV infections annually result in 70,000 hospitalizations, 800 deaths, and an estimated $5.7 billion loss [48, 49]. hNoV is the most frequent hospital-acquired infection stateside, accounting for 65% of all hospital unit closures [108]. hNoV is of particular danger to immunocompromised patients, causing significant morbidity and mortality [8]. hNoV has been shown to remain infectious in groundwater for at least 61 days [123], which poses a great threat to public health, especially in developing countries where hNoV is estimated to cause 218,000 deaths among children each year [92]. To date, the inability to culture hNoV has hampered efforts to study details of its lifecycle, as well as to find effective antiviral treatments.

hNoV genome is 7.7 kilobases in length, has a 4 nucleotide 5’ non-translated region

(NTR), a 44 nucleotide 3’ NTR, and a 21 nucleotide-long poly-A tail at the 3’ terminus [43, 44].

Additionally, hNoV genome possesses a 5’ terminal cap composed of virus protein (VPg) linked to the genome. The hNoV genome consists of 3 open reading frames (ORFs), corresponding to 3 peptides produced as a result of viral gene expression [6, 56]. ORF2 and ORF3 encode structural proteins VP1 (major) and VP2 (minor), respectively, responsible for making the viral capsid.

ORF1 encodes a polyprotein that becomes proteolytically cleaved to give rise to several important non-structural proteins, including 3C-like protease (Pro), RNA-dependent RNA polymerase (Pol), and virus protein linked to genome (VPg). Please refer to Figure 1 for a summary of the hNoV genome structure and its encoded products.

Figure 1: Overview of hNoV Genome Structure and Transcription/Translation Products. hNoV enters the cell as a single-stranded, positive-sense RNA. As such, the cell is immediately able to translate its ORF1 into a polyprotein that will be further processed by autocatalytic cleavage to yield several non-structural proteins. Upon transcription, a new copy of the genome will be synthesized (gRNA), as well as a subgenome (sgRNA) that will give rise to capsid proteins when translated.

hNoV VPg (NS5) is a 15.8 kDa multifunction protein covalently linked to the 5’ end of hNoV genome [5]. Lack of VPg capping has been shown to be detrimental to viral infectivity, rendering viruses non-infectious [51]. At its basic role, VPg has been proposed to serve as a protective cap for the hNoV genome against detection by the host immune system [40]. Another role of VPg is its ability to serve as a recruiter of the cellular translation machinery by interacting with eukaryotic translation factors eIF3 and eIF4E [17, 24, 41], a process that was recently linked to the C-terminal end of the VPg protein [19]. A potential role in packaging of the newly synthesized viral genomes has also been hypothesized [58]. Additionally, VPg has been shown to be a primer for the initiation of hNoV RNA synthesis when nucleotidylylated by either one of

2 hNoV’s two equally active versions of polymerase, the mature polymerase (NS6, Pol) and a protease-polymerase precursor protein (NS5-6, ProPol) that contains both protease (Pro) and Pol activities [43, 44, 114].

Previous studies of hNoV VPg have primarily been modeled on work done with

Picornaviruses, of which Poliovirus (PV) is a prototype organism. This is due to similarities between the characteristics and functions of several genes of these two single-stranded positive- sense viruses. However, there are significant differences between VPg of these two virus families. For one, PV VPg is a small protein, only 22 amino acids in size, compared to 133 of hNoV VPg. PV VPg is not required for initiation of protein synthesis [98, 115], whereas hNoV

VPg is absolutely essential for translation [17]. Nucleotidylylation of hNoV VPg occurs in the absence of RNA while uridylylation of PV VPg is RNA template-dependent [71, 84, 87, 96,

111]. For these reasons, it is important to look at additional systems for modeling hNoV VPg research.

The VPg of Potyviruses, such as Potato Virus A (PVA), shares more similarity to the

VPg of hNoV with regard to size (189 amino acids vs. 133, respectively), a requirement for translation initiation, and nucleotidylylation by all four nucleotides in the absence of RNA [100].

The PVA VPg is also capable of directly binding nucleoside triphosphates, a process that was found to serve as a pre-requisite for its nucleotidylylation [100], which has not been studied for hNoV VPg. A nucleotide-binding sequence containing several lysine residues (38AYTKKGK44) is present upstream of the PVA VPg nucleotidylylation site and is required for both binding of

UTP and successful uridylylation [100, 104]. This amino acid sequence was also shown to be necessary for RNA and protein binding properties of PVA VPg [100, 104].

A closer look at hNoV VPg N-terminus upstream of the nucleotidylylation site revealed a region rich in lysine residues within the first 20 amino acids of the VPg protein. Additionally,

3 sequence analysis of the same region showed the presence of a possible nucleotide-binding site similar to that of PVA (1GKKGKNK7). This possibility was further reinforced by a study previously reported in the literature where the first 3, 8, or 20 amino acids of the hNoV VPg

N-terminus containing positively-charged lysine residues were deleted to reveal a progressive reduction in VPg nucleotidylylation levels [5]. This reduction was speculated to have originated from a potential loss of yet undetermined NTP-binding capability of hNoV VPg, but no follow-up studies were performed.

In this body of work, I further characterize hNoV VPg nucleotidylylation, and I investigate the role of specific amino acids in the lysine-rich region comprising the first 20 amino acids of hNoV VPg in the regulation of its nucleotidylylation. I provide the first evidence that hNoV VPg directly binds nucleoside triphosphates and that prevention of binding inhibits nucleotidylylation of the protein. I also demonstrate that the un-cleaved hNoV precursor protein,

ProPol (NS5-6) [43, 44], which possesses both the protease and RNA-dependent RNA polymerase functions of the mature protease (Pro, NS5) and polymerase (Pol, NS6), is vastly superior to Pol with respect to nucleotidylylation of hNoV VPg, suggesting a role for this precursor protein in viral replication.

II. BACKGROUND

The investigations detailed in this document are based on principles firmly established in the literature. As such, it is necessary to review the biology of hNoV and the viruses on which most of the current work is based on.

A. Picornavirus VPg Biology

The majority of initial hNoV research was modeled on the studies performed in

Picornaviruses, with Poliovirus (PV) being the most characterized organism in this family.

In fact, some of the most prominent Calicivirus investigators either concurrently work with

Picornaviruses or began their scientific endeavors studying Picornavirus biology. This is primarily due to the fact that both of these organisms are single-stranded positive-sense RNA viruses and share several similarities with regard to genome organization and gene function.

PV VPg (3B) is a rather small protein – only 22 amino acids and 2.4 kDa in size [101].

3B protein is covalently linked to the 5’ end of the PV genome through its only tyrosine residue in position 3 of the protein [1, 33, 116]. 3B plays an indispensable role in replication of PV RNA genome by serving as a protein primer for PV polymerase (3Dpol) [93]. 3AB protein, a precursor of the 3B primer, is actually the protein responsible for initiation of PV replication by recruiting the replication complex to the cytoplasmic surface of the double-membrane vesicles formed by the destruction of the ER and Golgi apparatus during the PV infection [7, 18, 22, 29, 122].

A strongly hydrophobic C-terminus of the 3A domain of the 3AB protein is responsible for anchoring PV VPg within the membrane vesicles where replication complexes assemble

[26, 129, 134] and mutations within this region result in viruses incapable of genomic replication

[37, 64]. The recombinant 3AB protein has been shown to greatly stimulate in vitro activity of

5 the 3Dpol [94, 99, 109]. Once the replication complex is assembled, PV protease precursor protein (3CDpro) cleaves the membrane-bound 3AB to give rise to 3A and VPg proteins [101]. It is important to note that while 3CDpro is a potent protease, it has no polymerase activity, unlike hNoV ProPol [4], and requires further proteolytic processing to generate active PV 3Dpol.

Before 3B can act as a primer for PV genome replication, it must undergo uridylylation by 3Dpol in a template-dependent manner. Uridylylation is a process by which two uridylic residues are covalently linked to Tyr3 of the PV VPg by 3Dpol [1, 21, 33, 116]. The template for the reaction is a cis-acting replication element known as cre that is located in the coding sequence of PV protein 2C [71, 84, 87, 96, 111, 128, 143]. VPg, 3Dpol, and 3CDpro bind cre RNA and two uridylic residues [21, 132] are added to VPg using the AAACA sequence of cre as a template [97]. The function of 3CDpro in this case is to enhance the binding of the VPg-3Dpol complex to the cre sequence. It is evident from the literature that the recruitment of replication machinery to the double-membrane vesicles and VPg uridylylation processes occur simultaneously and not in a step-wise fashion. Following uridylylation, the replication complex is transferred from cre to the 3’ end of the PV genomic RNA where 3Dpol utilizes uridylylated

VPg as a primer to initiate PV RNA synthesis. Although uridylylated PV VPg primes both positive and negative strand replication, the cre-mediated VPg uridylylation has only been implicated in the synthesis of the positive-sense RNA, and was not required for the synthesis of the negative-sense strand [42, 84, 87]. It was proposed that the poly-A tail of the PV genome can serve as a template in such a case.

In addition to Tyr3 residue of PV VPg being uridylylated by 3Dpol, an arginine at position

17 was shown to be essential for the uridylylation reaction [93, 95]. Substitutions of this residue to either negatively charged glutamic acid or uncharged glutamine abolished VPg uridylylation at Tyr3. However, substitution to a positively charged lysine preserved the majority of the

6 wild-type PV VPg uridylylation levels. Of additional note was a discovery that double lysine substitutions (K9A, K10A) had a deleterious effect on VPg uridylylation as well [95].

Although PV VPg serves as a 5’ cap for the viral genome, it has no role in translation of viral RNA into its functional proteins, and is not required for viral infectivity [33, 89, 140].

In fact, only the virion RNA is covalently linked to VPg, which becomes cleaved off by the cellular unlinking enzyme upon viral entry into the host cell [2]. Instead, PV utilizes internal ribosome entry site (IRES) sequences that form highly ordered RNA structures in its

5’-untranslated region (UTR) to recruit the 40S ribosomal subunit of the host protein synthesis machinery [32, 98, 112, 115, 126]. It is unclear whether the 40S ribosomal subunit binds directly to the IRES or whether it is recruited by the interaction with eIF3 bound to the C-terminal fragment of eIF4G, which is the translation initiation protein that would bind IRES directly in such a scenario [10, 90, 98].

B. Potyvirus VPg biology

Potyviruses make up the largest genus of plant viruses [54]. Like Picornaviruses,

Potyviruses have a single-stranded positive-sense RNA genome with a single open reading frame that generates a large polyprotein upon translation, which must then undergo proteolytic cleavage by one of the three Potyvirus proteases to yield individual viral proteins [13, 14, 36, 50,

65, 78]. More than that, the RNA replication module of Potyvirus polyproteins (CI-6K-NIa-NIb) closely resembles a similar region of Picornavirus polyprotein (2C-3A-3B-3C-3D) in sequence and organization [110], prompting the scientific community to propose placing the two into a superfamily of Picorna-like plant and animal viruses [39, 65]. Tobacco Etch Virus (TEV),

Tobacco Vein Mottling Virus (TVMV), and Potato Virus A (PVA) are some of the best characterized Potyviruses at the molecular level [65].

The 5’ end of all Potyvirus RNA genomes is capped off by VPg [65]. Unlike the small

Poliovirus (PV) VPg, whose sole purpose is to serve as a primer for the replication of PV RNA by 3Dpol [93], Potyvirus VPg is a rather large protein, composed of 189 amino acids, and weighs approximately 22 kDa [65]. The large size of Potyvirus VPg enables it to be a multifunctional protein, participating in numerous viral activities, including cell-to-cell and long-distance movement, phloem loading, gene silencing suppression, RNA replication, and translation [9, 59,

67, 102, 119, 121]. The link to Potyvirus genome occurs through tyrosine amino acid residues located at position 60 for TVMV, 62 for TEV, and 63 for PVA [3, 85, 86, 91]. Mutations in

Tyr60 and Tyr62 residues of TVMV and TEV, respectively, resulted in inability of the viruses to amplify their genomes [85, 86, 119]. Interestingly, substitutions in Tyr63 of PVA had no effect on viral replication [100, 104]. It was proposed that an alternative site for PVA RNA linkage exists at Tyr119, but mutational analysis of this amino acid residue showed minimal differences from the wild-type protein [104].

Potyvirus VPg spends the majority of the viral lifecycle as a part of the N-terminal domain of the NIa protease that resembles PV 3Cpro [65]. The C-terminus of NIa contains the protease function of the protein. The proteolytic processing between the two termini that eventually releases VPg from the precursor protein is slow and has important implications for viral multiplication. The VPg domain of NIa contains a nuclear localization signal (NLS) [106].

Large amounts of capsid protein (CP) are required to create virion particles, resulting in a large amount of viral polyprotein synthesis to achieve required levels of CP [65]. A large amount of

NIa protease, as well as NIb polymerase, is generated at the same time. The NLS located within the VPg domain of the unprocessed NIa targets the protease for accumulation within the cell nucleus, thus downregulating the level of protease activity in the cytoplasm, where viral replication and virion assembly take place. Similarly, NIb has internal NLS sequences that cause

8 it to migrate to the nucleus, decreasing the amount of polymerase in the cytoplasm late in the infection cycle to favor viral genome encapsidation for movement and transmission [69].

Less clear is the role Potyvirus VPg plays in the viral genome replication. Much like in

PV, the replication of Potyvirus genome occurs in association with ER-derived membranes

[65, 110]. A 6K2-NIa precursor protein localizes to the membranes by utilizing yet uncleaved

6K2 transmembrane protein domain as the anchor [107, 120, 139], similar to the function of the

3A protein within the 3AB polyprotein in PV [26, 129, 134]. NIa protein then recruits RNA and

NIb into the replication complex, an interaction that is absolutely essential for viral replication

[23, 31, 69]. The recruitment of NIb to the replication complex positions it in close proximity of template RNA and VPg domain of the NIa, stimulating RNA genome synthesis. Although it was shown that Potyvirus VPg is able to enhance NIb activity [31, 53], the exact mechanism is unclear and it remains to be demonstrated whether Potyvirus VPg truly acts as a protein primer, much like the VPg of PV does [93].

Unlike PV VPg, the VPg of Potyviruses has been found to form complexes with translation initiation factors eIF4E, eIF4F, and eIF4G, and thus enhance the translation of viral proteins [30, 47, 55, 60, 79, 82, 117, 141]. At the same time, the 5’ UTR region of Potyviruses has been shown to contain translation enhancer sequences essential for viral pathogenesis [70].

There is non-definitive evidence to suggest that these sequences might encode IRES structures similar to PV 5’ UTR [15, 88], although these are much less structured than IRES of PV [74].

The relationship between the two factors affecting Potyvirus translation has not been clearly established.

In contrast to strict requirement for an RNA template in PV VPg uridylylation, it was demonstrated that PVA VPg nucleotidylylation is independent of any RNA templates and all four nucleoside triphosphates can participate in the reaction, although uridylic acid residues were

9 added most efficiently [100]. Using Prosite online software (us.expasy.org), the same manuscript proposed a putative nucleotide-binding site within PVA VPg mapping to a 38AYTKKGK44 lysine-rich motif upstream of the Tyr63 nucleotidylylation site. Deletion of this domain led to complete loss of nucleotide binding by PVA VPg and a drastic reduction in VPg nucleotidylylation in vitro. To ensure that the observed effect was not due to structural VPg changes, extensive circular dichroism spectroscopy of the two proteins was conducted and no significant structural differences between wild-type and deletion mutant VPg were discovered.

In addition to cessation of NTP binding and severe reduction in nucleotidylylation, it was later demonstrated that the Δ38-44 mutation of PVA VPg also reduced the ability of VPg to bind

RNA in sequence-independent manner and prevented cell-to-cell spread of PVA [104].

To elucidate further details regarding the putative NTP binding site of PVA VPg, a homology structural analysis of PVA VPg was performed utilizing LOMETS metaserver

(University of Michigan) and PONDR structural disorder analysis software (Molecular Kinetics,

Inc.) [68, 103-105, 142]. The analysis revealed that PVA VPg is a highly disordered globule-like protein with a hydrophobic core, a property that enables it to be structurally flexible, allowing

PVA VPg to adapt to its interacting partners and carry out its numerous functions.

The lysine-rich putative binding site 38AYTKKGK44 was mapped to a positively charged contact surface on the same face of PVA VPg protein that houses the Tyr63 nucleotidylylation site.

It was suggested through modeling that the Δ38-44 mutation causes a drop in the surface charge of the PVA VPg due to the loss of positively charged lysines. The authors proposed that this loss of surface charge causes a reduction in attraction between negative charge of NTPs, or RNA, and the positively charged surface of PVA VPg, leading to reduced association and, thus, abolition of

NTP binding and a decrease in nucleotidylylation activity at the nearby Tyr63 residue of PVA

VPg. It was also pointed out that due to the positioning of the binding motif relative to the Tyr63

10 nucleotidylylation site, a structural rearrangement of PVA VPg was necessary to allow the formation of a covalent bond between incoming NMP and the tyrosine amino acid residue.

C. Calicivirus VPg Biology

Caliciviruses are a diverse family of viruses that have a single-stranded positive-sense

RNA genome similar to Picornaviruses and Potyviruses described above with the exception of encoding three open reading frames (ORFs) instead of one [43, 44]. Much of Calicivirus research has been based on previous studies of Picornavirus and Potyvirus biology. Feline Calicivirus

(FCV) and Murine Norovirus (MNV) are two of the most studied Caliciviridae. The focus of this body of work, however, is on characterization of VPg nucleotidylylation of the human Norovirus

(hNoV), probably one of the least studied Caliciviruses due to its inability to be grown in cell culture [27].

Linked to the 5’ end of the hNoV genome, VPg (NS5) falls somewhere between PV and

PVA VPg proteins in size and function, being 133 amino acids and 15.8 kDa in size [43, 44] and taking part in genome replication and translation into functional protein products [12, 28, 51,

114]. Similar to PV VPg, nucleotidylylated hNoV VPg has been shown to prime RNA synthesis

[114, 131]. In FCV, replication of viral RNA has been linked to association with host cell membranes [45], much like it was for PV [7, 18, 22, 29, 122]. A stable VPg precursor protein,

NS4-5 in hNoV and p43 in FCV, similar to the 3AB VPg precursor in PV, is thought to serve as a hydrophobic membrane anchor and recruiter of the Norovirus replication complex [26, 127,

129, 134]. Additionally, there is evidence that FCV and MNV [40], as well as hNoV [5], are able to interact with RNA in vitro , similar to PVA VPg [77, 104], although it is not yet clear how this might affect viral replication. Either one of the two hNoV polymerases, the mature polymerase

(NS6, Pol) and a stable protease-polymerase precursor protein (NS5-6, ProPol) that contains both

11 protease and Pol activities, is able to synthesize new viral RNA [4]. The requirement for having two forms of viral polymerase is not evident.

Unlike 5’ UTR of PV and PVA genomes, hNoV 5’ UTR is very short, composed of only

4 nucleotides [44], excluding the possibility of having an IRES-mediated translation initiation

[32]. Nevertheless, secondary structures found in the 5’ end of Calicivirus genomes have been suggested to have a possible role in regulation of viral RNA translation [125] and it was shown that the first 110 nucleotides of the hNoV genome are capable of binding cellular factors known to participate in IRES-mediated translation initiation [46]. Still, no experimental evidence for the involvement of an IRES structure in the initiation of hNoV RNA translation has been reported.

Thus, VPg covalently linked to the 5’ end of the hNoV genome plays an essential role in the initiation of viral protein synthesis, as was shown by an experiment involving destruction of

FCV VPg with Proteinase K treatment of VPg-linked RNA, which subsequently abolished translation of the viral RNA [51]. It was later demonstrated that VPg is able to interact with eIF3, eIF4GI, eIF4E, and eIF2a translation initiation factors, as well as S6 ribosomal protein [17, 24,

25, 41]. This interaction was mapped to the C-terminus of norovirus VPg [19]. Based on this data, VPg is likely to play an essential role in initiation of viral RNA translation, although the exact nature of that function remains elusive to this day.

hNoV VPg nucleotidylylation occurs on tyrosine 27 amino acid and can be catalyzed by either Pol or ProPol, although it has been suggested that ProPol might be more efficient, even though no data was presented [5]. Modification of the tyrosine residue covalently linked to NMP has been shown to drastically affect viral replication in FCV [81]. All four nucleotides can be covalently linked to Tyr27, with G being preferred over U, which, in turn, is added more efficiently than either A or C [5, 73]. Experimental evidence shows that two nucleotides are added during the process of nucleotidylylation [5], similar to what was demonstrated for PV [21,

132]. hNoV VPg nucleotidylylation occurs in the absence of a template, although an enhancing element was found in the 3’ end of the viral RNA genome [5]. Deletions of the first 3, 8, or 20 amino acids from the N-terminal end of hNoV VPg have been shown to progressively abrogate nucleotidylylation, suggesting that the first 20 amino acid residues of hNoV protein might have a regulatory role in the process of covalently linking nucleotides to Tyr27 of VPg [5], a phenomenon resembling discovery of an NTP-binding site in PVA [100, 104].

Similar to structure modeling of PVA VPg [104, 105], solution structures of both FCV and MNV VPg were determined with the aid of NMR spectroscopy [66]. A compact hydrophobic core composed of three α-helices for FCV and a double α-helix for MNV was found at the center of the VPg protein. The termini of both proteins had to be removed in order for the solution structures to work, as they were rather disordered, a phenomenon also observed while modeling PVA VPg [104, 105]. Destabilizing mutations of the hydrophobic Calicivirus

VPg core abolished its activity in the majority of cases, with the effect being particularly severe for FCV [66]. Computer simulated docking between proposed VPg structures and a crystal structure of Pol suggested that interactions between Tyr where nucleotidylylation takes place and the active site of Pol would be impossible without VPg undergoing conformational changes.

This observation falls in line with the notion of PVA VPg being a disordered protein able to adjust to its numerous interacting partners, supporting its ability to perform multiple functions

[104, 105].

III. MATERIALS AND METHODS

Most methods utilized in this investigation have previously been described in the literature. Where necessary, I performed optimizations to better suit the needs of the current project. Please refer to Appendix A: Assay Optimization for a detailed report of the steps taken to improve testing conditions.

A. Cloning and Mutagenesis

Human Norovirus (hNoV) genes NS6 (Pol, RdRp), NS5-6 (ProPol), and NS4 (VPg) were cloned into the pET32a vector (Millipore, Inc.). An N-terminal His6 tag was engineered into the constructs using cloning primers. The cloning methodology for hNoV ProPol was previously described [76]. The hNoV Pol was cloned from a hNoV GII.4 construct (GenBank:

AY032605.1) provided by K. Green (NIAID, Bethesda, MD) using 5’-TCGATCCATATGCAT

CACCATCACCATCACGGTGGTGACAGTAAGGGAACA-3’ as the forward primer and

5’-ACGATCGAATTCATTCGACGCCATCTTCATTCAC-3’ as the reverse primer. An NdeI restriction recognition site introduced in the forward primer and an EcoRI site introduced in the reverse primer were utilized to clone hNoV Pol into the pET32a vector. Similarly, His6-labeled hNoV VPg was cloned from the same source into a pET32a vector using

5’-GCATCTAGAAAGGAGATATACATATGCACCATCATCATCATCATGGTAAGAAAGG

GAAGAACAAAACTGG-3’ as a forward primer and 5’-GCACTCGAGCTACTCAAAACTGA

GTTTCTCATTGTAGTCC-3’ as a reverse primer. XbaI and XhoI restriction sites were utilized in the two primers, respectively, for cloning. Constructs were transformed into DH5α chemically-competent E. coli cells (New England Biolabs, Inc.). For protein expression, all plasmid constructs were transformed into Rosetta pLysS competent cells (Millipore, Inc.).

Majority of the studies described required the use of a tag-less (native) hNoV VPg protein. An Intein Mediated Purification with an Affinity Chitin-binding Tag kit (IMPACT, New

England Biolabs, Inc.) was utilized to generate hNoV VPg lacking any tags. Manufacturer’s directions were followed in making the constructs, with some modifications. Initially, hNoV

VPg was cloned into the supplied pTXB1 vector with 5’-GGTGGTCATATGGGTAAGAAAGG

GAAGAACAAAACTGG-3’ as the forward primer and 5’-GGTGGTTGCTCTTCCGCACTCA

AAACTGAGTTTCTCATTGTAGTCC-3’ as the reverse primer, containing NdeI and SapI restriction sites, respectively. Cloning into the pTXB1 plasmid created an Intein tag at the

C-terminal end of the protein upon expression based on the manufacturer’s recommendation that

C-terminal Intein-tagged constructs provide the highest yield of active protein. The Intein tag is later cleaved during the protein purification process, leaving the protein of interest tag-less (see below). However, it was observed that hNoV VPg with an Intein tag at its C-terminus becomes misfolded during expression, which required a subsequent switch of the Intein tag to the

N-terminal end of VPg. To accomplish this, hNoV VPg was re-amplified with 5’-GGTGGTTGC

TCTTCCAACGGTAAGAAAGGGAAGAACAAAACTGG-3’ as a forward and

5’-GGTGGTCTGCAGTCACTCAAAACTGAGTTTCTCATTGTAGTCC-3’ as a reverse primer, containing SapI and PstI restriction sites, respectively. The hNoV VPg was then cloned into the pTYB21 vector (New England Biolabs, Inc.) that placed an Intein tag at the N-terminus of the protein upon expression. Although the yield of the protein expressed off the pTYB21 construct was reduced, protein folding was proper.

Mutations were introduced into hNoV VPg using the Q5 Site-Directed Mutagenesis Kit

(New England Biolabs, Inc.). Multiple mutations (see Table 1) in hNoV VPg were introduced sequentially following manufacturer’s instructions. Mutagenesis primers for constructs of interest were designed using NEBaseChanger online software (New England Biolabs, Inc.).

All cloning and mutagenesis work was confirmed by a commercial DNA sequencing service

(GeneWiz, Inc.). Table 1 contains the positions of various mutations in hNoV VPg and the identifying designations. Please refer to Table 2 in Appendix B: Tag-Less VPg Cloning Primers for further details regarding cloning of the native VPg proteins used in the study.

B. Protein Expression

Bacterial starter cultures (10 mL) from individual colonies were used for each expressed protein and incubated overnight at 30°C in the presence of 150 µg/mL Ampicillin and 50 µg/mL

Chloramphenicol. Overnight Express Instant LB medium (Millipore, Inc.) (300mL each) was prepared according to manufacturer instructions and inoculated with overnight LB cultures. Cells were grown to OD600 = 1.0 in the presence of 75 µg/mL Ampicillin and 25 µg/mL

Chloramphenicol at 37°C. IPTG was then added to 0.02 mM, and the cultures were incubated for

48 hours at 15°C. Cells were harvested by centrifugation at 5000 RPM for 20 minutes at 4°C, and cell pellets were frozen at -70°C.

For His6-tagged proteins [137], cell pellets were resuspended in 40 mL of Lysis Buffer

(50 mM HEPES (pH 7.3), 500 mM NaCl, 20% Glycerol, 1% NP-40 detergent, 10 mM

β-Mercaptoethanol, Lysozyme). The mixture was then sonicated (Branson Ultrasonics Corp., model 150I) for 30 minutes on ice with cycles of 15 seconds on, 45 seconds off, and an amplitude of 70%. The cell lysate was then centrifuged at 18000 RPM for 60 minutes at 4°C.

Co2+-based Talon Metal Affinity Resin (Clontech, Inc.) columns were prepared by washing twice with 20 mL of Lysis Buffer and centrifugation at 5000 RPM for 15 minutes at 4°C.

Per manufacturer instructions, the resin was utilized at 1 mL slurry per 500 mL of cell pellet culture. Cell lysis supernatant was combined with the resin and incubated with shaking for 2 hours at 4°C. The resin slurry-cell lysis mixture was centrifuged at 4500 RPM for 10 minutes at

4°C. The pellet was washed 3 times in 10 mL Wash Buffer (50 mM HEPES (pH 7.3), 500 mM

NaCl, 20% glycerol, 0.1% NP-40 detergent, 10 mM β-Mercaptoethanol, 10 mM Imidazole) with centrifugation for 10 minutes at 4500 RPM at 4°C. Following the wash steps, the resin was resuspended in 10 mL Wash Buffer and packed into a chromatography column (BioRad, Inc.) by gravity flow. The Wash Buffer was then replaced with Elution Buffer (50 mM HEPES (pH 7.3),

500 mM NaCl, 20% Glycerol, 0.1% NP-40 detergent, 10 mM β-Mercaptoethanol, 250 mM

Imidazole). The column was incubated for 15 minutes on ice, and His6-labeled protein was then eluted off the column with Elution Buffer.

For tag-less protein production, manufacturer’s instructions for the IMPACT Kit (New

England Biolabs, Inc.) were followed with modifications. Frozen cell pellets were resuspended in ice-cold Column Buffer lacking Lysozyme (20 mM HEPES (pH 7.3), 1 M NaCl, 1 mM

EDTA, 1 mM TCEP, 0.2% Tween 20 detergent). The cell suspension underwent 3 cycles of freezing-thawing using an ethanol-dry ice bath and a 37°C water bath (10 minutes each), with vortexing between each cycle. Samples were then subjected to two 30 minute cycles of sonication followed by centrifugation as described above for the His6 protein purification.

Chitin-based resin (New England Biolabs, Inc.) was prepared by two consecutive washes in 25 mL of Column Buffer lacking Lysozyme and centrifugation at 5000 RPM for 15 minutes at 4°C.

Per manufacturer instructions, the cell lysates were mixed with 10 mL resin slurry per 1 L of expression cell culture. A chromatography column was packed with washed resin-cell lysate slurry using 25 mL of Column Buffer without Lysozyme by gravity flow. The column was then washed 3 times with 3 column-volumes of Cleavage Buffer (20 mM HEPES (pH 7.3), 1 M

NaCl, 1 mM EDTA, 50 mM DTT). The column was then filled to the top with additional

Cleavage Buffer, capped, and incubated at 4°C for 40 hours, allowing DTT to induce sufficient

Intein tag self-cleavage. The tag-less protein was then eluted off the column in Cleavage Buffer.

The proteins were eluted from the columns in 100 µL fractions. Protein concentration in each aliquot was evaluated by adding 10 µL of each fraction to 200 µL of the Bradford Reagent

(BioRad, 500-0205). Fractions displaying brilliant blue color were pooled and dialyzed three times against one liter of storage buffer (50 mM HEPES (7.3 pH), 50% glycerol, 10 mM DTT) at

4°C as previously described (twice for two hours each, a third overnight) [137]. Float-A-Lyzer

G2 Dialysis Devices of appropriate kD rating (Spectrum Labs, Inc.) were prepared and utilized according to the manufacturer’s instructions. The dialyzed protein was centrifuged for 20 minutes at 13000 RPM at 4°C to pellet out any precipitate. Protein concentrations in the supernatant were determined using a NanoDrop Lite spectrophotometer (Thermo Scientific,

Inc.), and the quality and purity of the protein was assessed by SDS-PAGE gel electrophoresis.

C. hNoV Polymerase (Pol) Assay

An in vitro polymerase (Pol) assay was utilized to compare the RNA synthesis activities of hNoV Pol and hNoV ProPol proteins before being utilized in the hNoV VPg nucleotidylylation assay. The assay used was based on previously reported protocols [4, 34, 35,

72, 80, 113], with minor modifications to optimize the reaction yields (see Appendix A). The Pol reaction buffer was: 1 mM MnCl2, 10 mM DTT, 50 mM HEPES (pH 7.3), 100 µM of each cold

NTP (ATP/CTP/GTP/UTP), 2 µCi of 3000 Ci mmol-1 α-32P GTP, 50 units of RNAse OUT recombinant ribonuclease inhibitor (Invitrogen, Inc.). For the Pol reaction, 1 µM of RNA template (terminal 100nt of the GII.4 hNoV anti-genomic strand) was incubated with 1 µM enzyme (Pol or ProPol), in Pol reaction buffer at 37°C for 2 hours (20 µL reaction volumes).

Reactions were terminated by the addition of a 2x TBE-Urea Buffer (Invitrogen, Inc.) and heating at 70°C for 5 minutes. RNA products were resolved by gel electrophoresis in 1x TBE buffer using precast 6% TBE-Urea gels (Invitrogen, EC6865BOX), dried, and exposed overnight

18 onto a storage phosphor screen (GE Healthcare, Inc.) to be scanned by a phosphorimager

(GE Healthcare, Storm 840). Semi-quantitative analysis of the RNA products was performed using GelQuant.Net software (BiochemLabSolutions.com).

A sequence representing hNoV antigenomic RNA utilized in the assay was synthesized with the HiScribe T7 Quick High Yield RNA Synthesis Kit (New England Biolabs, Inc.). The template for T7 synthesis reaction was a PCR fragment generated from a pET32a plasmid containing the entire hNoV GII.4 genome utilizing 5’-GAAATTAATACGACTCACTATAGGG

CATGCTAGAAAGCACTCCG-3’ as the forward primer and 5’-GTGAATGAAGATGGCGTC

TAAC-3’ as the reverse primer. The forward primer introduced a T7 Pol recognition sequence required for T7-driven RNA synthesis. The RNA fragment synthesized by the T7 Pol in a runoff synthesis reaction corresponded to the terminal 100nt of the hNoV anti-genome. The subsequent use of such template in a hNoV Pol assay resulted in synthesis of a product RNA corresponding to the first 100nt of the hNoV genome.

D. hNoV VPg Nucleotidylylation Assay

The hNoV VPg nucleotidylylation assay was based on previously reported in vitro assays

[5, 35, 72, 73, 114], with modifications to optimize the reaction yields (See Appendix A). For the in vitro assay, 6.3 µM hNoV VPg was incubated with 1.3 µM hNoV ProPol or Pol

(approximately 1:5 enzyme:VPg ratio) in a reaction buffer (1 mM MnCl2, 10 mM DTT, 50 mM

HEPES (pH 7.3), 100 µM cold GTP, 1 µCi 3000 Ci mmol-1 α-32P GTP) (20 µL reaction volumes). For experiments involving other labeled NTPs, corresponding cold and radioactive

NTPs were substituted. Reactions were incubated for 2 hours at 37°C. Reactions were terminated by adding 20 µL of 2x Laemmli buffer (BioRad, Inc.) and heating at 95oC for 5 minutes.

Protein products were resolved on a pre-cast 12% SDS-PAGE gel (BioRad, Inc.) at 100 volts for

19 approximately 90 minutes using 10 µL of the sample-buffer mixtures. It was imperative that the dye front was evacuated from the gel completely as it corresponds well to the migration of the unincorporated radiolabeled nucleotides, which have a potential to obscure nucleotidylylation analysis in the steps that follow. Gels were then stained for 10 minutes in Coomassie Stain

(BioRad, Inc.) and destained for at least 2 hours using Destaining Solution (BioRad, Inc.) to reveal the pattern of hNoV VPg band migration. This was done to verify that the amount of VPg and polymerase in each reaction was equivalent and to facilitate quantitation of the radioactive nucleotidylylated products (see below). Gels were then dried for 30 minutes and exposed onto a phosphor screen to qualitatively evaluate the outcome of the nucleotidylylation reaction.

Once an autoradiography image was obtained, the individual hNoV VPg gel bands were excised from the dried gel and radioactivity was measured using a liquid scintillation counter for quantitative analysis of nucleotide incorporation into hNoV VPg. Quantitation was accomplished by converting the DPM output from the scintillation counter to pmol of α-32P GMP incorporated per µg of VPg. Guidelines from Perkin Elmer website were followed to make the conversion

(http://www.perkinelmer.com/lab-products-and-services/application-support-knowledgebase/ radiometric/radiochemical-calculations.html). Briefly, the DPM was first converted to Curies according to the formula: 1 Curie (Ci) = 2.22 x 1012 DPM. Curies were then converted to moles using the specific activity (SA) of the radioisotope on the day the calculations were made:

Ci x (1 mmol/SA Ci) = mmol. To obtain SA for isotope on the day of the calculations, a Perkin

Elmer Radioactive Decay Calculator was used (http://www.perkinelmer.com/tools/calculatorrad#

/product). This was an essential step in calculations, to directly compare experimental results obtained on different days, as it normalized the effect of the radio decay and resultant loss of signal in gel bands. As a final step, the dilution factor of hot to cold nucleotides was incorporated into the calculations. Millimoles of incorporated α-32P GMP were converted to picomoles and

20 divided by the number of micrograms of VPg. Statistical analysis of hNoV VPg nucleotidylylation levels was performed using student’s two-tailed t-test in Prizm (GraphPad

Software, Inc.). Semi-quantitative analysis of the nucleotidylylation products was performed using GelQuant.Net software (BiochemLabSolutions.com) where scintillation analysis was not available.

E. hNoV VPg NTP Binding Assay

The NTP binding assay (affinity labeling reaction), has been previously described [20,

100]. NTPs were oxidized by incubating 2 µCi of 3000 Ci mmol-1 α-32P GTP (or other labeled

NTPs) in the presence of 0.5 mM HCl, 0.75 mM sodium periodate, and water (5 µL total volume) at room temperature in the dark for 20 minutes. Surplus sodium periodate was reduced with 0.2 volumes of 50% glycerol for 20 minutes to prevent additional oxidation. For the binding reaction, 0.6 µL of the oxidized nucleotide mixture (0.2 µCi of 3000 Ci mmol-1 α-32P GTP or other NTPs) was incubated with 3 µg of hNoV VPg in the presence of 3 mM sodium cyanoborohydrite and 10 mM HEPES (7.3 pH), 1 mM DTT, 5 mM MnCl2, and dH2O (30 µL total reaction volume) for 60 minutes in an ice bath (0°C). These conditions crosslink NTPs that are bound to the protein. The reactions were terminated by the addition of an equal volume of

2x Laemmli buffer and heating to 95oC for 5 minutes, and analyzed by gel electrophoresis as described above for the VPg nucleotidylylation assay.

F. Structure Modeling

To gain additional insight into potential amino acid associations within the structured core of hNoV VPg, homology modeling analogous to the structure analysis reported for PVA

VPg was performed [104]. The hNoV VPg sequence was submitted to the LOMETS metaserver

[142], which produced an output of top 10 predicted models using the wdPPAS algorithm. The top resulting model for my analysis was the solution structure of the core domain of the MNV

VPg protein [66].

IV. CHARACTERIZATION OF HUMAN NOROVIRUS VPG

As can be seen from the literature, human Norovirus (hNoV) shares many features with

Poliovirus (PV) when it comes to genome replication, and with Potato Virus A (PVA) in terms of interactions with viral RNA translation machinery. Similar to PV VPg, hNoV VPg serves as a protein primer for initiation of viral genome replication [42, 93, 114, 133]. Like PVA VPg, hNoV VPg demonstrates interactions with eIF4E and other eukaryotic translation initiation factors, affecting viral protein production [17, 19, 30, 41, 47, 61, 79]. Additionally, hNoV VPg shares certain features of both PV and PVA in terms of VPg nucleotidylylation activities, with possible RNA and ProPol requirements for the enhancement of nucleotidylylation similar to PV

[5, 95, 96, 128, 143], and the ability to accept all four nucleotides in a template-independent manner like PVA [5, 100].

While nucleotidylylation of Tyr27 in hNoV VPg has been well-characterized [5], the concept of nucleoside triphosphate binding by hNoV VPg as a prerequisite to nucleotidylylation, similar to the process described in PVA [100, 104], is a fascinating new direction for Calicivirus research. Evidence suggests that the potential nucleotide-binding site could be located in the lysine-rich high-positive-charge N-terminus of hNoV VPg [5]. Additionally, other charged amino acid residues could participate in nucleotidylylation, as was suggested for PV VPg [95].

Sequence analysis of the GII.4 hNoV VPg N-terminus revealed a lysine-rich sequence,

1GKKGKNK7, similar to nucleotide binding motif of PVA VPg, 38AYTKKGK44. Protein sequence alignment of VPg from representative genogroup strains of Norovirus revealed strong conservation of the KKGK motif in the N-terminal end of the protein (Figure 2). Furthermore, the conservation of positively charged residues extends to amino acid position 20 of the

GII hNoV (Figure 2), a region that was previously shown to be essential for hNoV VPg

23 nucleotidylylation [5]. Together, these findings present a great target for evaluating nucleotide binding capacity of hNoV VPg.

Figure 2: Amino Acid Sequence Alignment of VPg Proteins from Various Norovirus Genogroups. VPg amino acid sequences for representative strains from each Norovirus genogroup were taken directly from GenBank. Please note that GVII VPg amino acid sequence was not available from GenBank at the time. The alignments were constructed utilizing Geneious software (Biomatters, Ltd.). The alignment reveals a high degree of conservation of positively-charged residues, K and R, within the N-terminus of Norovirus VPg.

For the work described here, I substituted the positively charged residues in the first 20 amino acids of the hNoV VPg N-terminus to neutral alanine, sequentially and individually, to reveal residues required for nucleoside triphosphate binding and subsequent nucleotidylylation of VPg at Tyr27. Using homology structure modeling, I identified additional amino acids within the hNoV VPg protein core to potentially be involved in the nucleotidylylation reaction, similar to Arg17 of PV [95]. I mutated these residues to study their effects on VPg nucleotidylylation.

A. hNoV VPg Nucleotidylylation Occurs Through Covalent Linkage to the α-Phosphate of

the Incoming Nucleotide

To ensure that VPg nucleotidylylation reaction occurred under my experimental conditions, I performed a VPg nucleotidylylation assay, previously described in Materials and

Methods, with two different forms of radiolabeled GTP, α-32P GTP and γ-32P GTP.

This experiment, along with the use of sequential nuclease digests, has been previously utilized to demonstrate that nucleotidylylation reaction occurred where the linkage of an incoming

24 nucleotide to the Tyr27 of hNoV His6-VPg was covalent and the attachment took place through an α-phosphate of the radiolabeled GMP via a 5’phosphodiester bond [5].

As can be seen in Figure 3, nucleotidylylation with α-32P GTP produces a strong band,

in a denaturing gel, corresponding to the radiolabeled

32 His6-VPg, whereas reaction with γ- P GTP produced a

very faint band where the nucleotidylylated His6-VPg

would be expected (Figure 3, lanes 2 and 4,

respectively). This suggests that the covalent linkage of

GMP to hNoV His6-VPg occurs through the α-

phosphate, as the γ-labeled phosphate is lost with the

pyruvic acid residue released during the formation of a

5’phosphodiester bond between GMP and Tyr27 of

His6-VPg. Unexpectedly, negative control reactions

lacking ProPol produced faint labeling of His6-VPg

(Figure 3, lanes 1 and 3, respectively). This is likely

caused by tight interactions between radiolabeled GTP Figure 3: hNoV Nucleotidylylation with Either α-32P GTP or γ-32P GTP. VPg nucleotidylylation assay described in and His6-VPg and is similar to results previously Materials and Methods was performed in the presence of either α-32P GTP or demonstrated for PVA VPg, where VPg dimers were γ-32P GTP. Absence of ProPol from the reaction was used as a negative control for present after resolution of the proteins on a highly VPg nucleotidylylation. As can be seen, α-32P GTP is the only nucleotide able to label hNoV His6-VPg, suggesting that the denaturing gel, suggesting strong interactions between 5’phosphodiester bond is formed through the α-phosphate of the GMP. VPg protein and its other interacting partners [100].

B. Nucleotidylylation of Native hNoV VPg by hNoV ProPol Is Significantly Greater Than

Nucleotidylylation Catalyzed by hNoV Pol

The primary purpose of these studies was to examine the influence of amino acids near the amino terminal end of the hNoV VPg on its nucleotidylylation by its polymerases.

All previous studies of hNoV VPg nucleotidylylation utilized a His6-tag on the amino terminal end of the protein. For my studies, I deemed it essential to remove amino terminal tags from hNoV VPg to eliminate any potential effects of additional amino acids on the reaction.

Initial studies quantitatively compared the hNoV VPg nucleotidylylation activities of

both versions of the hNoV polymerase. Figure 4: Nucleotidylylation of His6-Labeled and Native hNoV VPg. Reactions were conducted as Nucleotidylylation of amino terminal described in the methods section using α-32P GTP. Top Panel: autoradiography image: Bottom Panel: SDS PAGE gel in Panel A stained with Coomassie His6-tagged hNoV VPg by hNoV Pol or hNoV dye. Lanes 1-3, His6-hNoV VPg; Lanes 4-9, native hNoV VPg. Reaction negative controls: Lanes 1 and ProPol was essentially equivalent (Figure 4, 4, reactions lacking either of the hNoV polymerases. Lanes 6 and 8, reactions containing inactive hNoV lanes 2 and 3), consistent with previous reports polymerases (active site motifs mutated from 342GDD344 to 342GGG344). Lane 9, native hNoV VPg with Tyr27 substituted by Ala (nucleotidylylation [5]. Nucleotidylylation reactions of His6-tagged site); compare directly to Lane 7, hNoV ProPol catalyzed reaction using native hNoV VPg. hNoV VPg and tag-less (native) hNoV VPg, when catalyzed by hNoV Pol, revealed no significant difference in the levels of nucleotidylylation (Figure 4, lanes 2 and 5). However, nucleotidylylation of native hNoV VPg by

26 hNoV ProPol was 70 times greater than that observed for His6-tagged hNoV VPg (Figure 4, lanes 3 and 7).

In time course studies, hNoV ProPol was approximately 100 times more efficient at nucleotidylylation than hNoV Pol (Figure 5, Panel A), although the de novo RNA synthesis activities of both polymerases were comparable (Figure 5, Panel B). The overall levels of nucleotidylylation of native hNoV VPg catalyzed by hNoV Pol were comparable to those previously reported for His6-tagged hNoV VPg [5]. As previously reported [5], nucleotidylylation of hNoV VPg required an active hNoV polymerase, as substitutions of the

342GDD344 motif in the active site with 342GGG344, which eliminates hNoV Pol activity [135], inhibited hNoV VPg nucleotidylylation by either hNoV Pol or ProPol (Figure 4, lanes 6 and 8).

The tyrosine at position 27 of hNoV VPg has been reported to be the preferred nucleotidylylation

Figure 5: Nucleotidylylation of Native hNoV VPg by hNoV Pol and hNoV ProPol.

Panel A. Nucleotidylylation reactions and quantitation of α-32P GMP addition to hNoV VPg were performed as described in the Methods section. Each data point represents triplicate reactions.

Panel B. RdRp activities of hNoV Pol and hNoV ProPol. RNA synthesis reactions using equimolar amounts of the hNoV polymerases of a 100 nucleotide template were conducted as described in the Methods section to assess the relative RNA polymerase activity of the two enzymes. Reaction negative controls: Lane 1, reaction containing no polymerase. Lanes 3 and 5, reactions containing inactive hNoV polymerases (see Figure 4 legend for details).

27 site [5]. Substitution of an alanine at position 27 in native hNoV VPg essentially abolished nucleotidylylation to a level consistent with previous reports (Figure 4, lane 9). Based on these data, all further studies on hNoV nucleotidylylation were carried out using native hNoV VPg and hNoV ProPol.

C. GMP and UMP Are Preferentially Added to hNoV VPg

It was previously reported that there is a preferential addition of specific nucleotides to hNoV VPg by hNoV ProPol, with GMP being added most efficiently, and AMP being added less efficiently than UMP or CMP to His6-tagged hNoV VPg [5]. In my studies, hNoV ProPol added

GMP to native hNoV VPg at a 2-fold higher level than UMP, and approximately 4-fold higher levels than CMP or AMP (p<0.001) (Figure 6). UMP was added significantly more than either

CMP or AMP (p<0.001), both of which were approximately equivalent to each other with

respect to their ability to become attached to hNoV VPg during the

nucleotidylylation reaction (p=0.750) (Figure 6), consistent with

previously reported patterns using hNoV ProPol [5].

Figure 6: Nucleotidylylation of Native hNoV VPg by Different NMPs. Nucleotidylylation reactions and quantitation of α-32P NMP addition to native hNoV VPg by hNoV ProPol were performed as described in the Methods section. Data were compiled from three separate experiments, each conducted in triplicate for each NMP. GMP incorporation vs. UMP (p=0.001) and UMP incorporation vs. CMP or AMP (p=0.001) (two-tailed student’s t test (Prism, GraphPad Software, Inc.)).

D. Positively-charged Amino Acids in the N-terminus of hNoV VPg Regulate Its

Nucleotidylylation by hNoV ProPol

Previous studies demonstrated that removal of the first 3, 8, or 20 amino acids of hNoV

VPg had a progressively deleterious effect on its nucleotidylylation by hNoV ProPol [5].

This region contains several charged amino acids (Table 1), which were hypothesized to be

28 responsible for the observed inhibition of nucleotidylylation [5]. Amino acid deletion studies of the Potato Virus A VPg have demonstrated that an amino acid sequence, 38AYTKKGK44, in the amino terminal region of the protein was responsible for its nucleotidylylation and for UTP binding [100]. Furthermore, this sequence is well conserved among PVA and the VPg of several other viruses [100], and a portion of this sequence (underlined above), 2KKGK5, is present in hNoV VPg (Table 1). For PV, double lysine substitutions (K9A, K10A) have a deleterious effect on VPg uridylylation [95]. In hNoV VPg, double lysines are present at two positions in the amino terminal region (K2, K3 and K12, K13).

I sought to more extensively examine the role of individual positively charged amino acids in the amino terminal region of hNoV VPg in regulating its nucleotidylylation. A panel of hNoV VPg containing consecutive single point mutations replacing lysines and a single arginine with alanines was created (Table 1). Constructs K2-3A, K2-7A, and K3-20A were designed to remove the lysines and arginine removed in the 3, 8, and 20 amino acid deletion mutants of hNoV VPg previously reported [5]. In addition, VPg constructs containing only single substitutions of each of the individual lysines and arginine were created (Table 1). A construct

Table 1: Amino Acid Substitutions in hNoV VPg. The first 20 amino acids of the N-terminus of hNoV VPg are presented. Wild-type (WT) amino acids of interest are shown in blue. Table A: consecutive alanine substitutions (in red) of positively charged amino acids introduced for these studies. K2-3A, K3-7A, and K2-20A correspond to the 3, 8, and 20 amino acid deletion mutants previously reported. Table B: individual amino acid substitutions. K12-13A mutant was included in the study due to the importance of consecutive lysine residues in VPg nucleotidylylation of poliovirus.

29 containing substitution of the double lysine at positions 12 and 13 was also included (Table 1).

For the complete cloning strategy and primers utilized to generate mutant constructs, please refer to Appendix B: Cloning Primers and Strategy.

The cumulative removal of charged amino acids in the amino terminal region of hNoV

VPg resulted in a progressive loss of nucleotidylylation by hNoV ProPol (Figure 7, panel A), consistent with previous observations using hNoV VPg deletion mutants [5]. No change in nucleotidylylation was observed by the removal of a single lysine at position 2 (K2A).

Figure 7: Nucleotidylylation of Alanine-Substituted hNoV VPg.

Nucleotidylylation reactions and quantitation of α-32P GMP addition to hNoV VPg by hNoV ProPol were performed as described in the Methods section. Data for each VPg were compiled from at least two independent experiments (with triplicate reactions each). Descriptions of mutated VPg are contained in Table 1. Data for reactions using native hNoV VPg (WT) and native hNoV VPg containing a Y27A substitution are displayed in each panel for direct comparison. Note that for these two VPg, the same data is displayed in each panel. Bars represent standard deviations.

Panel A: Nucleotidylylation levels of native hNoV VPg containing consecutive mutations within the first 20 amino acids.

Panel B: Nucleotidylylation levels of native hNoV VPg containing individual mutations within the first 20 amino acids.

Panel C: Nucleotidylylation levels of native hNoV VPg containing individual mutations within the central core region surrounding Tyr27.

Substitution of only the lysines at positions 2 and 3 (K2-3A), produced a significant, 2-fold reduction (p=0.029) in nucleotidylylation. Additional cumulative substitutions up to position 12 induced a trend of gradual, additional loss of nucleotidylylation, but this was not significantly different from the initial loss observed for the K2-3A substitutions. The addition of substitutions at positions 13 (K2-13A) and 20 (K2-20A) reduced nucleotidylylation to essentially undetectable levels, significantly lower than that observed for the K2-5A construct (p=0.001). Substitution of only the two lysines at positions 12 and 13 (K12-13A) also reduced its nucleotidylylation significantly (p=0.029, Figure 7, Panel A) to a level essentially identical to that observed for the

K2-3A mutant.

Substitutions of individual lysine and arginine residues within the first 13 amino acids of hNoV VPg did not reduce its nucleotidylylation, demonstrating that the inhibition observed previously was a result of the cumulative removal of these charged amino acids (Figure 6, Panel

B). In fact, most of these single amino acid substitutions resulted in increased levels of nucleotidylylation over the wild-type protein. Surprisingly, substitution of the single lysine at position 20 (K20A) abolished nearly all nucleotidylylation of hNoV VPg (Figure 7, Panel B).

I was not able to examine the effect of a single substitution of lysine at position 5 since I was unable to express this protein despite numerous attempts and expression conditions.

E. The Central Core Region of hNoV VPg Does Not Contain Amino Acids Required for

the Stability of the Nucleotidylylation Reaction

For the Poliovirus VPg, an arginine at position 17 was shown to be essential for the nucleotidylylation of tyrosine at position 3, and the presence of a charged amino acid at this position was required for nucleotidylylation [93, 95]. Other than an examination of the role of individual tyrosines [5], there have been no studies to determine the involvement of other

Figure 8: Predictive Homology Model of hNoV VPg. LOMETS metaserver was utilized to generate the most likely homology structure of hNoV VPg using the wdPPAS algorithm as described in the Methods section to be used as a guide for selection of candidate amino acids for substitution studies. The modeling template was a solution structure of the Murine Norovirus VPg core domain. Panel A: N-terminus and central core of hNoV VPg (amino acids Gly1-Glu56). Panel B: central core domain of hNoV VPg (amino acids Asp24-Glu56) displaying the proximity to Tyr27 of positively charged amino acids selected for substitution studies. The predicted core is likely composed of a double helix structure with the helices at approximately 90 degrees to each other as shown for the Murine Norovirus VPg. positively charged amino acids within the central core of hNoV VPg surrounding Tyr27 on hNoV VPg nucleotidylylation.

To determine if positively charged amino acids in the central core region of hNoV VPg could potentially be candidates for participation in the nucleotidylylation reaction, I required a reasonable predictive structural model of this region. The PVA VPg is an intrinsically disordered globule-like protein that has a hydrophobic core with disordered amino and carboxy-terminal ends [104]. Analysis by solution NMR of Feline and Murine Norovirus VPg have shown analogous structures [66]. Utilizing homology modeling based on these structures (see Materials and Methods section for details), a model of hNoV VPg closely matched the MNV VPg structure

(Figure 8). For hNoV, a stable core from amino acids Asp24 to Glu56 was predicted, and both termini appeared to be in a high state of disorder and could not be resolved, which was

32 confirmed by PONDR analysis (Molecular Kinetics, Inc.) (Figure 9) [105]. Based on probable proximity to Tyr27, amino acids Lys31, Arg32, Lys34, and Arg50 were selected as having a potential role in conferring stability during nucleotidylylation. Substitution of any of these amino acids with alanine (see Table 1) did not reduce nucleotidylylation of hNoV VPg (Figure 7,

Panel C). In fact, these single amino acid substitutions typically induced enhanced nucleotidylylation.

Figure 9: hNoV VPg Structural Disorder Analysis. PONDR software (Molecular Kinetics, Inc.) was utilized to generate a map of structural disorder for hNoV VPg. The graph and the statistical readout indicate the presence of a stable protein core A16-E64, which roughly coincides with the LOMETS homology predicted structure, showing a double helix core encompassing amino acid residues D24-E56 (Figure 8). The termini at both ends appear to be in the state of high disorder.

F. hNoV VPg Binds NTPs in the Absence of Polymerase

The VPg of PVA has been shown to directly bind NTPs and deletion of an identified

NTP binding sequence (38AYTKKGK44) abolished nucleotidylylation of that protein by its polymerase [100]. To determine whether hNoV VPg was capable of directly binding NTPs,

I performed an affinity labeling assay [20, 100] with radiolabeled NTPs in the absence of any polymerase. Native hNoV VPg was able to bind all 4 individual NTPs at essentially the same level (Figure 10, Panel A), providing the first demonstration of direct NTP binding by a

Calicivirus VPg. GTP binding activity was noticeably reduced only for the K2-13A and K2-20A mutants (Figure 10, Panel B). GTP binding by all of the other consecutive VPg mutants,

33 including K2-12A, as well as the double substitution mutant, K12-13A, was also not reduced

(data not shown). GTP binding was not reduced for the single substitution mutant, K20A, which abolished nucleotidylylation (Figure 10, Panel B). GTP binding for the K2-3 or K2-5 mutants, which encompass a putative NTP binding motif (2KKGK5) discussed above, did not appear to be reduced (Figure 10, Panel C).

Figure 10: NTP Binding by hNoV VPg. Reactions were performed in the absence of polymerase as described in the Materials and Methods section.

Panel A. Native hNoV binding of different α-32P NTP.

Panel B. Binding of α-32P GTP by native hNoV VPg (WT) and native hNoV VPg with alanine substitutions (see Table 1 for descriptions). The migration differences of the VPg molecules are likely due to the reduced charge of the K2-13A and K2-20A proteins compared to WT or K20A containing a single substitution.

Panel C. Binding of α-32P GTP by native hNoV VPg (WT) and native hNoV VPg with alanine substitutions encompassing the 2KKGK5 domain.

V. FUTURE WORK

Additional aspects of hNoV VPg biology and nucleotidylylation were studied over the course of this project. Because of time constraints or technical difficulties, they remain unsolved, but would add greatly to the field of Calicivirus research.

A. Plasmon Resonance Required to Study NTP Binding by hNoV VPg

Whereas incorporation of nucleotides during VPg nucleotidylylation reaction was in the pmol range, the binding interaction between radiolabeled nucleotides and hNoV VPg was at amol (attomolar) level. Even though NTP binding by hNoV VPg was evident from affinity binding studies presented, it suffered greatly from inconsistencies as a result of being an inefficient reaction (Figure 11), except for the very extremes of the progressive amino acid substitution mutant spectrum – WT and K2-20A (Figure 10, Panel B).

The alternative to utilizing affinity labeling method presented here and in PVA literature

[100] is surface plasmon resonance [62, 118]. During a plasmon resonance experiment, a target

Figure 11: GTP Binding of Alanine-Substituted hNoV VPg. Presented is the summary of 5 separate GTP binding attempts with hNoV VPg consecutive alanine substitution mutants. The reactions were setup in identical fashion, utilizing a protocol outlined in Materials and Methods. Please note relative inconsistencies of outcomes, as a result of the reaction’s very low efficiency. The only constant outcome appears to be at the two extremes of the mutation spectrum, WT and K2- 20A mutant.

(VPg) is attached to the dextran molecule conjugated to a metal surface on a chip. The ligand’s interacting partner (NTP) is then flushed through the system and the two partners bind.

The binding interaction changes the resonance angle of the reflected light from a laser that is being directed at the metal surface, which becomes noted by a detector, thus recording interactions between the target and its interacting partner. This method has a high degree of sensitivity and would be better suited for studying interactions of the attomolar magnitude.

B. hNoV VPg Is a Protein Primer for Initiation of hNoV Genomic RNA Synthesis

One of the initial aims of this body of work was to demonstrate the ability of hNoV VPg to serve as a protein primer for initiation of hNoV genomic RNA synthesis.

Important details of hNoV genomic replication remain poorly understood, and not definitively proven by experimental data in the literature. In view of that fact, and based on information detailed within a core set of critical Calicivirus publications [4, 5, 11, 34, 35, 72, 83, 113,

114, 138], I proposed a candidate genomic replication scheme for hNoV Figure 12: hNoV Genomic RNA Replication. Top: Pol uridylylates VPg, which serves as a primer for Pol (Figure 12). by binding to the poly-A tail of the gRNA. Pol then extends the VPg primer and synthesizes the complimentary strand (agRNA). hNoV replicates its positive-sense Bottom: VPg is nucleotidylylated and used as a primer to initiate synthesis of gRNA by primer extension, resulting in genomic RNA (gRNA) by first making a covalent linkage of VPg to the 5’ end of the viral genome.

36 negative-sense antigenomic RNA (agRNA) copy utilizing Pol and uridylylated VPg as a primer

[5, 114] (Figure 12, top panel). Uridylylated VPg anneals to the poly-A tail of hNoV gRNA and serves as the primer for Pol binding and subsequent synthesis of the agRNA strand [114].

While the scheme for agRNA synthesis is well supported by literature, no mechanism for gRNA synthesis has been proven, hindering the creation of a complete hNoV replication scheme

(and for Caliciviridae in general).

It was demonstrated in cell culture that VPg can enhance gRNA synthesis [131] and it is well accepted that it serves as a protein primer for initiation or gRNA synthesis, even though direct evidence for this is lacking. hNoV genome begins with a GU and since hNoV can be nucleotidylylated with any of the four nucleotides, showing preference for G and U [5], hNoV

VPg likely becomes linked to a G or a GU combination of nucleotides, allowing it to serve as a primer for initiation of gRNA synthesis (Figure 12, bottom panel).

In my work, I performed a polymerase assay, as described in Materials and Methods, with an addition of 3 µg of hNoV His6-VPg to the reaction, in order to evaluate the effect VPg has on RNA synthesis activity (Figure 13). Addition of His6-VPg to the RNA synthesis reaction caused an upward shift in the migration of the radiolabeled RNA product, compared to a de novo reaction lacking a potential protein primer (Figure 13, panel A, lanes 4 and 5). It was noted that essentially all of the newly synthesized radiolabeled RNA was interacting with His6-VPg.

Subsequent treatment of His6-VPg-RNA with Proteinase K (PK) degraded His6-VPg, releasing newly synthesized RNA, allowing it to migrate at the level of the de novo product (Figure 13, panel A, lane 6).

A follow up study comparing Pol and ProPol RNA synthesis activities in the presence or absence of hNoV His6-VPg over a period of four hours was conducted (Figure 13, panel B).

The de novo activities of the two polymerases were rather comparable over the course of the

37 experiment (Figure 13, panel B). Addition of His6-VPg to the reaction improved RNA yield of the ProPol-catalyzed reaction two-fold, on average, and had very little effect on the

Pol-catalyzed RNA synthesis for the first two hours, with a reduction of synthesis activities thereafter, as compared to the de novo timecourse (Figure 13, panel B).

Figure 13: Effects of His6-VPg on RNA Synthesis by Pol and ProPol. RNA synthesis assays were conducted as described in Materials and Methods, except 3 µg of His6-VPg was added to the reactions where indicated. Panel A: His6-VPg caused a shift in RNA product migration on a 7M denaturing urea gel (lane 5 vs. 4). Subsequent treatment of the reaction with Proteinase K (PK) degraded His6-VPg protein and released new RNA (lane 6), allowing it to migrate at the level of a de novo reaction product (compare lanes 4 and 6). Panel B: Timecourse data of Pol and ProPol activities in the absence and presence of His6-VPg. Pol and ProPol synthesis activities were comparable in the absence of His6-VPg. Addition of the potential protein primer enhanced ProPol-catalyzed RNA product formation, while diminishing Pol RNA synthesis activity. Timecourse data indicates that hNoV VPg has a stimulatory effect on RNA synthesis by

ProPol. The addition of hNoV His6-VPg to the Pol RNA synthesis reaction appears to be inhibitory to its activity. Considering data from the native hNoV VPg nucleotidylylation timecourse that compared Pol and ProPol activities over time (Figure 5, panel A), where Pol was found to be 100-fold less efficient at nucleotidylylation than ProPol, it appears that the two competing reactions, His6-VPg nucleotidylylation and RNA synthesis, are splitting Pol activity, causing an overall decrease in RNA product formation (Figure 13, panel B). Since hNoV ProPol is so much more efficient at VPg nucleotidylylation than Pol (Figure 5, panel A), the competitive

38 nature of the two reactions, nucleotidylylation and polymerization, becomes greatly diminished and, instead, VPg is able to stimulate ProPol RNA synthesis activity (Figure 13, panel B).

Because hNoV VPg was found to bind RNA [5, 40] and because VPg interactions with its binding partners were shown to withstand denaturing gel conditions in PVA [100, 104], it is impossible to deduce, even in a highly denaturing 7M urea gel, whether hNoV VPg is acting as a primer or is simply binding RNA during the course of the RNA synthesis reaction (Figure 13, panel A, lane 5), although there is no doubt that VPg enhances hNoV genome replication activity

[131]. It would be prudent to replicate VPg-priming studies with the native hNoV VPg, as the removal of the His6 tag increased nucleotidylylation efficiency by ProPol several fold (Figure 4, lanes 3 and 7) and might have an even greater effect on its primed RNA synthesis activity.

C. Protease Domain of Intact hNoV ProPol Is Required for Interaction with hNoV VPg

and Its Subsequent Nucleotidylylation

As was briefly mentioned in Materials and Methods, initial attempts to purify tag-less

(native) hNoV VPg by utilizing a C-terminally placed Intein construct resulted in a production of a misfolded protein. The nature of the misfolding was the presence of two equally prevalent VPg proteins, presumably two isoforms of the same protein with one being folded properly and the other one improperly (Figure 14, lane 1). I believed that the identity of the two proteins was identical because the two-band configuration disappeared upon nucleotidylylation reaction in the presence of ProPol, instead showing up as a single band with the additive intensity of the two separate ones when examined in a coomassie-stained SDS-PAGE gel (Figure 14, lanes 1 and 5).

Same reaction with Pol had no conformational effect on VPg (Figure 14, lane 3). Since the only difference between Pol and ProPol is the presence of the Pro domain on the latter protein, I proceeded to study the effect of Pro on conformational changes of misfolded VPg. In the absence

Figure 14: Nucleotidylylation of hNoV VPg Derived from C-Terminally Placed Intein Construct. The native VPg was isolated and utilized in a nucleotidylylation assay according to protocols presented in Materials and Methods section. 1 µM amount of Pro (lanes 2 and 4) was used to match 1 µM of ProPol (lane 5). In the presence of either hNoV ProPol or the Pro domain of the hNoV ProPol, the misfolded VPg protein assumed its proper conformation (lanes 2, 4, and 5). Interaction with hNoV Pol had no effect on hNoV conformation. The presence of hNoV Pro did not restore nucleotidylylation activity of the hNoV Pol to the hNoV ProPol levels (lanes 4 and 5).

of either Pol or ProPol, Pro was able to alter VPg conformation into

a single protein, much like ProPol did (Figure 14, lane 2). Adding

Pro to the nucleotidylylation reaction catalyzed by Pol caused VPg

to fold properly, but did not stimulate Pol-driven nucleotidylylation of VPg (Figure 14, lane 4) when examined by autoradiography, suggesting that intact ProPol protein is absolutely required for proper hNoV VPg folding and nucleotidylylation (Figure 14, lane 5).

Although the use of the N-terminal Intein construct provided us with a properly folded native hNoV VPg, the questions regarding the nature of the double-banded hNoV VPg generated from the C-terminal Intein setup remain. A mass spectrometry analysis of the two bands would definitively reveal the identity of the hNoV VPg forms seen in the denaturing SDS-PAGE gel

(Figure 14, lane1). If confirmed that the two proteins are identical, the reaction should be studied in a native non-denaturing gel to reveal the details of the hNoV VPg interaction with hNoV Pro and Pro domain of hNoV ProPol. The studies would then greatly benefit from a crystallography analysis of VPg-Pro, VPg-ProPol, and VPg-Pol interactions to elucidate the mechanism of hNoV

VPg conformation alteration and its subsequent nucleotidylylation.

VI. CONCLUSIONS

The hNoV VPg is a multifunctional protein that plays essential roles during viral RNA synthesis [114], translation initiation [17, 24, 41], and potentially packaging of genomes into new virions [58]. In this body of work, I demonstrated that the un-cleaved viral precursor protein, ProPol (NS5-6), is likely the primary polymerase responsible for nucleotidylylation of hNoV VPg, providing a potential, discrete, role for this long-lived intracellular protein.

Nucleotidylylation of hNoV VPg by ProPol was 100-fold greater than that observed for Pol in my studies. Some members of the Caliciviridae family, such as FCV utilize only ProPol due to a lack of a proteolysis cleavage site [63, 127]. For PV, although Pol is the only form of viral polymerase capable of VPg uridylylation, ProPol is absolutely required, as it stabilizes the template-dependent addition of uridylic residues to the Tyr3 of PV VPg [75, 96]. PVA does not produce a form of its polymerase analogous to ProPol [100, 104].

In this manuscript, I provide the first evidence that hNoV VPg directly binds nucleoside triphosphates and is similar to the PVA VPg [100, 104], where NTP binding was correlated with nucleotidylylation. When NTP binding was reduced in the K2-13A hNoV mutant in my studies, nucleotidylylation was essentially eliminated. However, nucleotidylylation was significantly reduced for a number of other hNoV VPg mutants, which did not show reductions in NTP binding, indicating that more than one mechanism is involved with the control of hNoV VPg nucleotidylylation by the N-terminal region of the protein.

Identification of a discrete, small, amino acid sequence involved in NTP binding by hNoV VPg remains elusive. Alanine substitutions up to position twelve, including K2-12A, did not demonstrate reductions in NTP binding, despite showing significant reductions in VPg nucleotidylylation. The hNoV VPg mutants with cumulative substitutions including position

41 thirteen (K2-13A and K2-20A) displayed significant reductions in GTP binding activity, which was accompanied by a further reduction in nucleotidylylation to nearly undetectable levels.

Reductions in nucleotidylylation for the K12-13A double substitution mutant were not correlated with reductions in GTP binding. The K2-3A and K2-5A mutants, which removed lysines in a putative NTP binding site (2KKGK5) suggested by PVA nucleotidylylation and binding studies

[100, 104], did not demonstrate reduced GTP binding despite displaying significantly reduced nucleotidylylation. These data indicate that this five amino acid sequence is likely not essential, by itself, to constitute an NTP binding site for the hNoV VPg, and that NTP binding appears to involve a larger region of at least 13 amino acids. Since no structures for this region of the protein are currently available, it is not known if this group of amino acids potentially comprise an NTP binding pocket or if the alanine substitutions collectively induce conformational changes that prohibit NTP binding.

The preferential addition of GMP or UMP to hNoV VPg (this report and [5]) is not explained by its NTP binding reaction as all four NTPs are bound with equal efficiency. A recent report characterizing hNoV Pol indicated that the binding affinity of hNoV Pol does not add insight to the nucleotidylylation preference since the highest NTP affinity for enzyme is for CTP

[57]. Since ProPol appears to be the primary enzyme involved in hNoV nucleotidylylation, relative affinity of NTPs for ProPol and/or a direct comparison of nucleotidylylation preferences for Pol will need to be established to determine if ProPol is responsible for the observed NMP nucleotidylylation preference.

The reduction of nucleotidylylation by substitutions of the double lysines at positions two and three (K2-3A), and at positions twelve and thirteen (K12-13A), is consistent with observations of the interference of PV VPg uridylylation by elimination of double lysines [95], although no specific mechanism for this interference has been determined.

Sequential substitution of charged amino acids with alanine within the first 20 positions of the protein progressively reduced and eliminated nucleotidylylation, consistent with previous observations that deletion of the first 3, 8, or 20 amino acids of hNoV VPg progressively reduced

its nucleotidylylation [5]. The current data

reduce the probability that potential

conformation changes caused by large

deletions were responsible for interference with

nucleotidylylation, although changes in

conformation due to the alanine substitutions

cannot not be ruled out. My current data is also

Figure 15: The Relationship Between hNoV VPg consistent with studies of the PVA VPg, which N-Terminal Charge and Nucleotidylylation. The charge of the first 20 amino acids within the indicated that the degree of nucleotidylylation positively-charged N-terminus of hNoV VPg is plotted against the radiolabeled GMP incorporation by progressively mutated hNoV VPg proteins. The was correlated to the overall surface charge of identity of proteins is labeled for each data point. A clear relationship between charge and degree of the region of that protein in regulating its hNoV VPg nucleotidylylation is apparent. nucleotidylylation [100, 104]. A graphic plot of charge within the first 20 amino acids of hNoV VPg against the nucleotidylylation intensity of the mutant VPg proteins revealed a distinct correlation between the number of charged amino acid residues within the N-terminus of hNoV VPg and the protein’s ability to incorporate radiolabeled GMP (Figure 15), providing further evidence that surface charge might play an important role in hNoV VPg nucleotidylylation, much like it did for PVA [100, 104].

The presence of six additional histidines in the amino terminal end of hNoV VPg had a profound inhibitory effect on nucleotidylylation by ProPol, but not Pol. This observation may indicate that there is an additional, preferred, interaction between the amino terminal domain of

VPg and the Pro domain of ProPol, or that conformation changes in Pol induced by the presence

43 of a covalently linked Pro sequence are responsible for an enhanced interaction. This idea was additionally reinforced by the data obtained from my study of the misfolding native VPg protein, where interaction with either Pro or Pro domain of ProPol caused a conformational shift within the double-banded hNoV VPg. The reduction of nucleotidylylation by sequential alanine substitutions up to position 12 and the effect of the substitutions of either set of double lysines on nucleotidylylation, in the absence of any effect on NTP binding, may also indicate a direct interaction of the amino terminal end of VPg with ProPol. Protein-protein binding studies will be required to determine if such interactions are present.

An arginine at position 17 of PV VPg was found to provide a role in its uridylylation, and that substitution with uncharged amino acids eliminated the reaction but substitution with a lysine was permissive [95]. Alanine substitution of several positively charged amino acids within the structured core of hNoV VPg that were potential candidates to serve an analogous role to the

PV VPg Arg17 failed to induce a detrimental effect on nucleotidylylation. Of note is that the single substitution of K20 of hNoV VPg with alanine eliminated nucleotidylylation. Since this mutant binds GTP as well as the native VPg, and due to its potential proximity to Tyr27 being located on an apparently flexible region of the protein, K20 may potentially serve an analogous role in nucleotidylylation of hNoV VPg to an arginine or lysine at position 17 of PV VPg.

In this work, I explored the influence of charged amino acids within the lysine-rich

N-terminus of hNoV VPg in its nucleotidylylation. This region of the protein exerts substantial control on the nucleotidylylation process, most likely through several mechanisms. I showed that the un-cleaved viral precursor protein, ProPol is likely the primary polymerase responsible for hNoV nucleotidylylation, and that hNoV VPg can directly bind nucleotide triphosphates. There appears to be no role of charged amino acids within the structured core of hNoV VPg in stabilizing the nucleotidylylation reaction, but the lysine at position 20 of the unstructured

N-terminal portion of the protein is a candidate for such a role. Structural analysis of the protein coupled to nucleoside triphosphates should help address a number of outstanding questions.

It is quite clear from the literature overview and experimental results presented here that hNoV VPg plays an essential role in hNoV replication and viral protein expression, and that VPg nucleotidylylation and NTP binding are essential, albeit complex, processes governing the activity of hNoV VPg. It is worthwhile to note that the biology of the stable hNoV precursor proteins, NS4-VPg and VPg-ProPol [6], has been majorly overlooked. Like 3AB VPg precursor responsible for localization of the replication complex to lipid membranes in PV [26, 29, 94, 99,

109, 129, 130, 134], or the VPg-Pro precursor, NIa, responsible for important proteolytic and protein localization activities during PVA replication [16, 23, 31, 53, 69, 106, 107, 120], hNoV precursor VPg proteins might have activities and effects different than those of the fully processed protein. It would be of great benefit to the field of hNoV research (and Caliciviridae in general) to study the effects NS4-VPg and VPg-ProPol proteins might have on nucleotidylylation and RNA synthesis priming qualities of hNoV VPg.

APPENDIX A: ASSAY OPTIMIZATION

All methods utilized over the course of this study are firmly based on protocols previously described in literature. In some cases, alternate testing conditions were presented by different research groups for the same assay. In order to generate optimal assay conditions to meet the needs of my research, and in order to learn more about the properties of the proteins being studied, I performed several experiments to improve my testing methods. The complete and fully optimized assay conditions could be found in Materials and Methods section.

hNoV Polymerase Assay Optimization

The polymerase (Pol) assay was based on a core set of research publications [4, 11, 34,

35, 52, 72, 80, 113, 114, 138] and optimized for the activity of the hNoV Pol enzyme. The Pol reaction utilized by Dr. Kim Green’s laboratory was used for general assay guidelines and was composed of 3 mM MnCl2, 10 mM DTT, 50 mM HEPES (pH 7.3), 100 µM of each cold NTP

(ATP/CTP/GTP/UTP), 1 µM of 400 Ci mmol-1 α-32P UTP, 10 µM of RNA template, and 1 µM

Pol, incubated at 30°C for 60 minutes in a total 15 µL reaction volume. The use of 50 mM

HEPES and 10 mM DTT was universal to most, if not all, protocols presented in literature and did not need to be further evaluated. As I was working with RNA products, I included 50 units of

RNAse OUT recombinant ribonuclease inhibitor (Invitrogen, Inc.) to avoid degradation.

The reaction volume was adjusted to 20 µL and amounts of the reaction components were altered accordingly to preserve the concentration and to improve the convenience of working with smaller reagent quantities. Because of a desire to establish Pol assay interchangeable with hNoV

VPg nucleotidylylation assay (described later), a switch to 3000 Ci mmol-1 α-32P GTP was implemented. 2 µCi (33.3 nM final amount) of the radiolabeled GTP was utilized due to

46 convenience, being the lowest pipetting limit possible, while generating a strong signal during autoradiography analysis. Similarly, 3 mM MnCl2 was replaced with 1 mM MnCl2 to make Pol assay more comparable with VPg nucleotidylylation assay. Quantification and comparison of the assay products were performed as described in Materials and Methods.

The majority of Calicivirus publications utilized 30ºC as incubation temperature, with some groups preferring physiological temperature of 37ºC. In my assay, there was no significant

Figure 16: hNoV Polymerase (Pol) Assay Optimization. The Pol assay was based on a core set of research publications and optimized for the activity of the hNoV Pol enzyme. The following conditions were set as reference and were adjusted accordingly as assay characterization progressed further: 1 mM MnCl2, 10 mM DTT, 50 mM HEPES, 100 µM of each cold NTP (ATP/CTP/GTP/UTP), 2 µCi of 3000 Ci mmol-1 α-32P GTP, 1 µg of RNA template, and 1 µg of Pol, incubated at 37°C for 60 minutes in a total 20 µL reaction volume. Panel A: Effect of temperature on hNoV RNA synthesis reaction. Panel B: Divalent cation requirement for the hNoV Pol RNA synthesis activity. Panel C: Optimal cold NTP concentration for the RNA synthesis reaction by hNoV Pol. Panel D: Comparison of Pol and ProPol RNA synthesis activities over time. 1 µg of Pol was used to 1.3 µg of ProPol in order to keep the enzyme concentrations equimolar. Panel E: Effect of Pol amount on the amount of labeled RNA product formed. RNA template was held constant at 1 µg. Panel F: Effect of RNA template used on the amount of labeled RNA product produced. Pol quantity was held constant at 1 µg.

47 difference between the two temperatures (p=0.495) (Figure 16, panel A) and 37ºC was picked for all future assays due to its physiological relevance. The literature differed with regard to the cation use in the Pol reactions with 3 mM MgCl2, 3 mM MgOAc2, and 3 mM MnCl2 being most common. My tests revealed 1 mM MnCl2 (utilized instead of 3 mM MnCl2 for compatibility with hNoV VPg nucleotidylylation assay) promoted significantly more activity than the next best option (p=0.008) (Figure 16, panel B) and all future assays were adjusted accordingly. The use of cold NTPs varied greatly from 100 µM to 500 µM, as reported by other labs. In my analysis,

100 µM produced the best results (p<0.050) (Figure 16, Panel C) and was picked as the optimal

NTP concentration to utilize in all future Pol assays. A timecourse experiment comparing Pol and ProPol activities over a 5-hour incubation period revealed that the activities of the two enzymes were comparable and that the relationship between time and amount of radiolabeled product is directly proportional (Figure 16, Panel D). Please note that 1.3 µg of ProPol was required to preserve an equimolar amount of enzyme in the assay, as compared to Pol. In order to produce the best signal possible without exceeding Pol assay times reported in literature, two hours was chosen for all future assays. Increasing the amount of hNoV Pol enzyme used in the Pol assay progressively increased the yield of the labeled RNA product (Figure 16, panel E).

Because working with 1 µg of Pol (0.7 mM) was convenient and because this amount was close to the amount most commonly used in literature (1 mM), it was set as the preferred amount of enzyme to be used in the Pol assay, even though utilizing more would have yielded extra radiolabeled RNA product. 1 µg of RNA template (1.25 mM in my case) was the most common amount reportedly used in aforementioned literature and appeared to be right at the inflection point of the template RNA amount curve in my studies (Figure 16, Panel F), making it ideal for all further work with Pol-catalyzed RNA synthesis.

48 hNoV VPg Nucleotidylylation Assay Optimization

The hNoV nucleotidylylation assay was closely based on the polymerase assay and its optimized conditions described in the previous section, while following general guidelines established by Dr. Kim Green’s laboratory [5]. The reference assay was set as follows:

2 µg hNoV His6-VPg was incubated with 1 µg hNoV Pol in a reaction buffer of 1 mM MnCl2, 60

-1 µM ZnCl2, 10 mM DTT, 50 mM HEPES (pH 7.3), 100 µM cold GTP, and 2 µCi 3000 Ci mmol

α-32P GTP, all in a 20 µL reaction volume, and incubated for 2 hours at 37°C. 2 µg hNoV

His6-VPg was utilized as the lowest amount possible to be easily resolved on an SDS-PAGE gel when a quarter of the reaction volume is loaded into each well, and fell within the range of the

VPg amounts previously used [5].

Although hNoV VPg nucleotidylylation reaction appeared to produce more radiolabeled

VPg at 30ºC, the difference was not statistically significant from 37ºC (p=0.276) (Figure 17, panel A). I chose to keep 37ºC as the default temperature due to physiologic considerations.

The study of cation requirements for the hNoV VPg nucleotidylylation revealed greatest

Figure 17: hNoV VPg Nucleotidylylation Assay Optimization. hNoV nucleotidylylation assay was closely based on the hNoV Pol assay, except with the addition of 2 µg of hNoV His6-VPg. The reference assay was set to include 2 µg hNoV His6-VPg, 1 µg hNoV Pol, 1 mM MnCl2, 60 µM ZnCl2, 10 mM DTT, 50 mM HEPES, 100 µM cold GTP, and 2 µCi 3000 Ci mmol-1 α-32P GTP in a 20 µL reaction volume, all incubated for 2 hours at 37°C. Panel A: Temperature requirement of the hNoV VPg nucleotidylylation reaction. Panel B: The effect of divalent cations on the hNoV VPg ability to incorporate radiolabeled nucleotides. Panel C: A one-hour timecourse comparing Pol and ProPol VPg nucleotidylylation activities. 1 µg of Pol was used to 1.3 µg of ProPol in order to keep the enzyme concentrations equimloar. Panel D: 24-hour extension of the timecourse presented in panel C.

49 preference for 1 mM MnCl2 with 3mM MgCl2 having an inhibitory effect on hNoV VPg nucleotidylylation, as was evident from the reaction where both MnCl2 and MgCl2 were utilized

(Figure 17, panel B). While 60 µM ZnCl2 was previously included concurrently with 1 mM

MnCl2 in hNoV VPg nucleotidylylation reactions [5], its purpose was not evident, as its addition to the reaction did not yield a statistically different amount of radiolabeled hNoV VPg, compared to 1 mM MnCl2 alone (p=0.756). As a result, I decided to exclude ZnCl2 from the reaction altogether. Two timecourse experiments were performed to study the difference between hNoV

Pol and hNoV ProPol VPg nucleotidylylation abilities over time. The short one-hour timecourse

(Figure 17, panel C) and a longer timecourse spanning 24 hours (Figure 17, panel D) revealed that ProPol is likely the preferred enzyme for hNoV VPg nucleotidylylation.

Two-hour incubation roughly appeared to be the inflexion point for both Pol and ProPol nucleotidylylation activities. As a result of these finding, ProPol was chosen as the preferred enzyme for further hNoV nucleotidylylatyion studies and two hours was used as the optimal amount of incubation time, perfectly matching hNoV Pol RNA synthesis assay parameters described in the previous section.

APPENDIX B: CLONING STRATEGY FOR TAG-LESS HNOV VPG CONSTRUCTS

As mentioned earlier in the Materials and Methods section, tag-less (native) hNoV VPg proteins were required to study the effects of positively-charged amino acid residues within the

N-terminus of hNoV VPg on its nucleotidylylation and NTP binding capabilities. Cloning of native hNoV VPg constructs relied heavily on the Intein Mediated Purification with an Affinity

Chitin-binding Tag kit (IMPACT, New England Biolabs, Inc.), as well as the Q5 Site-Directed

Mutagenesis Kit (New England Biolabs, Inc.).

The gene encoding hNoV VPg was first cloned into the pTXB1 vector, as described in

Materials and Methods. This allowed for the placement of an Intein tag at the C-terminus of the expressed hNoV VPg. The tag was later removed during the protein purification steps, leaving native hNoV VPg behind. hNoV VPg constructs detailed in Table 1 were then generated using mutagenesis primers from Table 2 (C-tagged N-terminal mutants and C-tagged core mutants) and the mutagenesis kit.

hNoV VPg Intein-tagged at its C-terminus during protein purification suffers greatly from being misfolded (Figure 13, lane 1). As a result, hNoV VPg constructs had to be re-cloned from their respective pTXB1 vectors into pTYB21 vectors encoding N-terminal Intein tags.

Rather than going through the tedious mutagenesis process, DNA encoding mutant hNoV VPg proteins with a C-terminally placed Intein tag was cloned using additional primers from Table 2

(N-tagged N-terminal mutants, and N-tagged core mutants). The resultant mutants underwent proper folding and were utilized for the majority of the hNoV characterization work presented in this manuscript.

Table 2: Tag-Less hNoV VPg Cloning Primers. The gene encoding hNoV VPg was first cloned into pTXB1 vector as described in Materials and Methods. Using C-tagged N-terminal and Core mutant primers, hNoV VPg progressive mutant proteins were generated (Table1). Unfortunately, the expressed proteins suffered misfolding issues and had to be cloned into the pTYB21 vector for proper expression and folding. This was accomplished with cloning primers for N-tagged N-terminal and core mutants in conjunction with the pTXB1 mutant constructs generated earlier, in order to avoid repeating the mutagenesis procedure.

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