Reverse Genetic Strategies to Interrogate Reassortment Restriction Among Group a Rotaviruses

Home , NSP6 (rotavirus), Reassortment

REVERSE GENETIC STRATEGIES TO INTERROGATE REASSORTMENT RESTRICTION AMONG GROUP A ROTAVIRUSES

JESSICA R. HALL

A Thesis Submitted to the Graduate Faculty of

WAKE FOREST UNIVERSITY GRADUATE SCHOOL OF ARTS AND SCIENCES

in Partial Fulfillment of the Requirements

for the Degree of

MASTER OF SCIENCE

Biology

August 2019

Winston-Salem, North Carolina

Approved By:

Sarah M. McDonald, Ph.D., Advisor

Gloria K. Muday, Ph.D., Chair

Douglas S. Lyles, Ph.D.

James B. Pease, Ph.D.

ACKNOWLEDGEMENTS

This body of work was made possible by the influences of countless people including, but not limited to, those named here. First, I am thankful for my advisor, Dr.

Sarah McDonald, who paved way for this opportunity. Her infectious enthusiasm for science served as a consistent reminder to “keep getting back on the horse”. I am grateful for my committee members, Drs. Douglas Lyles, Gloria Muday, and James Pease whose input and advice aided in my development into a careful scientist who thinks critically about her work. I am also thankful for continued mentorship from my undergraduate research advisor, Dr. Nathan Coussens, whose enthusiasm for science inspired mine.

I am appreciative of all of my friends for their unyielding support. I am blessed to have a solid foundation in my family, both blood and chosen. I am thankful for my mother,

Liana Hall, who instilled in me my work ethic and whose pride in me has inspired me to keep pursuing my dreams. I am forever indebted to Dr. Diana Arnett, who has served as my personal and scientific role model; thank you for believing in me, investing in me, and shaping me into the person and scientist I am today.

This thesis is dedicated to all of you.

TABLE OF CONTENTS

ACKNOWLEDGEMENTS...... ii

TABLE OF CONTENTS...... iii

LIST OF FIGURES AND TABLES...... vi

LIST OF ABBREVIATIONS...... viii

ABSTRACT...... x

INTRODUCTION...... 1

Rotavirus as a Pathogen...... 1

RV Genome and Capsid Structure...... 2

RV Classification...... 6

RVA Lifecycle and Assortment of Viral RNAs...... 10

RVA Reassortment...... 15

RVA Reverse Genetics...... 16

RVA NSP5...... 22

Research Aims...... 25

METHODS...... 26

Phylogenetic Analysis of Modern Human, Wa-like RVA Strains...... 26

Cell Culture...... 26

Plaque Assays...... 27

Preparation of Viral RNAs...... 28

Plasmid Construction...... 29

Generation of Recombinant RVAs using an All Plasmid Reverse Genetics

System...... 33

iii

Imaging of BHK-T7 Cells Transfected with Plasmid Encoding Green

Fluorescent Protein (GFP)...... 34

Growth of High-Titer Viral Stocks...... 34

Quantification of Viral Growth...... 34

Immunoblotting...... 35

Sequencing of NSP2 and NSP5 cDNAs...... 36

Nucleotide Alignments...... 38

RNA Folding Analysis...... 38

Protein Alignments...... 38

RESULTS...... 39

The laboratory strain P is genetically similar to modern, Wa-like RVAs...... 39

Wa grows to a higher titer than P in cell culture...... 42

BHK-T7 cells are transfectable and express T7 RNA polymerase...... 42

MA104 cells are permissive to RV infection...... 45

Modifications to the existing all plasmid-based reverse genetics system yielded

successful rescue of wildtype rRV SA11...... 45

Six Wa genome segments were successfully cloned into reverse genetics

vectors...... 50

rSA11-g11-Wa exhibits a modest replicative defect relative to wildtype...... 50

Wa and SA11 g11 share 88% nucleotide identity...... 53

Wa and SA11 g11 +RNAs are predicted to form different secondary structures

...... 55

Wa and SA11 NSP5 share 87% amino acid identity...... 57

Intergenotypic variation among NSP5 amino acid sequences lies in important

functional regions...... 59

Western blot of Wa and SA11 NSP5 reveals differential banding patterns at 8 h.p.i.

...... 61

Sequencing of NSP5 and NSP2 reveal no compensatory mutations...... 63

DISCUSSION...... 64

REFERENCES...... 73

APPENDIX...... 82

CIRICULUM VITAE...... 83

LIST OF FIGURES AND TABLES

Figure 1. Mature RV virion and genome composition...... 5

Figure 2. Human RVA genotype constellations...... 8

Figure 3. Overview of RVA lifecycle...... 14

Figure 4. SA11 H-96 reverse genetics system...... 18

Figure 5. Human vs. simian RVA genotype constellations...... 20

Figure 6. Important functional regions of NSP5...... 24

Figure 7. Cloning scheme for reverse genetics constructs containing Wa genes...... 32

Figure 8. Figure 8. Phylogeny of archival and modern RVA VP4 genotype sequences...... 41

Figure 9. BHK-T7 cells can uptake and express a plasmid encoding GFP...... 44

Figure 10. RT-PCR validation of successful wildtype rSA11 RVA rescue using all plasmid- based reverse genetics system...... 47

Figure 11. Growth comparison of rSA11-wt and rSA11-g11-Wa during one cycle of infection...... 52

Figure 12. Pairwise nucleotide alignments of Wa and SA11 genome segment 11...... 54

Figure 13. Predicted secondary structures of Wa and SA11 g11 +RNAs...... 56

Figure 14. Pairwise amino acid alignments of Wa and SA11 NSP5 and NSP6...... 58

Figure 15. Intergenotypic variation among NSP5 amino acid sequences...... 60

Figure 16. Western blot of Wa and SA11 NSP5 reveals differential banding patterns.....62

Figure 17. NSP5 amino acid changes and important functional regions...... 70

Table 1. RV proteins, functions, genotypes, and percent identity cut-off values...... 7

Table 2. Primer sequences for Wa reverse genetics constructs...... 30

Table 3. Primer sequences for NSP5 and NSP2 compensatory mutation screen...... 37

Table 4. Wa-like RVA strains that cluster with modern strains...... 40

Table 5. MA104 cell overseed experiment results...... 49

vii

LIST OF ABBREVIATIONS

AGE...... Acute gastroenteritis

AHSV...... African horse sickness virus

BHK...... Baby hamster kidney

DLP...... Double-layered particle c...... Complementary

CPE...... Cytopathic effect

DMEM...... Dulbecco’s Modified Eagle Medium

DNA...... Deoxyribonucleic acid ds...... Double-stranded

EE...... Early endosome

ER...... Endoplasmic reticulum

GFP...... Green fluorescent protein

HI-FBS...... Heat-inactivated fetal bovine serum h.p.i...... Hours post-infection kb...... Kilobase kDa...... Kilodalton

MOI...... Multiplicity of infection

NCBI...... National Center for Biotechnology Information

NSP...... Non-structural protein

ORF...... Open reading frame

PAGE...... Polyacrylamide gel electrophoresis

PCR...... Polymerase chain reaction

PFU...... Plaque-forming unit

viii r...... Recombinant

RCWG...... Rotavirus Classification Working Group

RdRp...... RNA-dependent RNA polymerase

RI...... Replicase intermediate

RNA...... Ribonucleic acid

RT...... Reverse transcriptase

RV...... Rotavirus

RVA...... Group A Rotavirus

SDS...... Sodium dodecyl sulfate

TLP...... Triple-layered particle

UTR...... Untranslated region

VP...... Viral protein

WHO...... World Health Organization wt...... Wildtype

ABSTRACT

Rotavirus (RV) is an 11-segmented, double-stranded RNA virus and gastrointestinal pathogen that infects a wide variety of mammalian and avian hosts.

Because RV’s genome is segmented, when a host cell is co-infected by more than one strain of RV, homologous genome segments belonging to different strains could be exchanged, or reassort. In theory, this process could generate up to 211 possible genetically distinct progeny with genome segments derived from both co-infecting parental viruses. In reality, though, genomic analyses of clinically isolated RVs indicate that reassortment occurs less frequently than what would be predicted due to chance alone. It is hypothesized that reassortment is restricted by biochemical incompatibilities between two RV strains’

RNAs or proteins. This hypothesis could be tested using a reverse genetics approach. While a reverse genetics system for simian RV strain SA11 has been developed, is poses significant technical challenges to successful rescue of recombinant RVs. After achieving functionality of the existing reverse genetics system, it was used to generate a reassortant virus containing genome segment 11 (g11) from the genetically divergent human RV strain, Wa. The mono-reassortant RV exhibited a moderate replicative defect relative to wildtype SA11, demonstrating that not only can the existing reverse genetics system be used to rescue mono-reassortant RVs, but differences between SA11 and Wa g11 RNAs and proteins are important for viral replication in cell culture. Because human RVs are so genetically divergent from strain SA11, the development of a reverse genetics system for a Wa-like human RV would provide a critical platform upon which human RV biology could be studied. This work is significant because it furthers understanding of how genotypic differences between RV strains contribute to their biology.

INTRODUCTION

Rotavirus as a Pathogen

As estimated by the World Health Organization’s (WHO) latest surveillance data released in 2017, 1.7 billion cases of diarrheal disease were reported in children worldwide.

Diarrheal disease is a leading cause of death among children globally, leading to greater than 525,000 childhood mortalities per year (Worth Health Organization, 2017; Troegen et al, 2017). Diarrheal disease can result from various viral, bacterial, and parasitic origins including—but not limited to—rotavirus, Clostridium difficile, and Cryptosporidium, all of which can lead to dehydration and ultimately death if left untreated (World Health

Organization, 2017). While most diarrheal disease can be treated effectively using supportive oral and intravenous rehydration therapies and zinc supplementation, it remains a significant cause of death among children in developing countries where access to appropriate and timely intervention is limited (Nunan et al., 2017).

Rotavirus (RV) is a leading cause of acute gastroenteritis (AGE) in young children.

RV also infects a wide variety of other mammalian hosts, as well as avian hosts. RV replicates within host enterocytes and can be shed via stool and transmitted by fecal-oral route (Ramig, 2004). RV particles are highly environmentally stable, and spread of the virus is poorly controlled despite sanitation efforts (De Zoysa and Freacham, 1985). Posing considerable morbidity as a common viral infection of childhood, RV infects virtually all unvaccinated children aged 5 and younger globally (Burnett et al., 2018). Four oral, live- attenuated vaccines have been recommended for RV prophylaxis by WHO, and national immunization programs in more than 80 countries have adopted RV vaccine implementation (RotarixTM, GSK Biologics; RotaTeqTM, Merck and Co.; RotavacTM,

Bharat Biotech; and RotaSiilTM, Serum Institute of India). These vaccines have been

1 deemed largely efficacious by the scientific community; when comparing RV-induced,

AGE-related hospitalizations and direct medical costs between pre- and post-vaccine eras, it was estimated that 382,000 RV-induced AGE-related hospitalizations and $1.228 billion in medical costs were prevented between 2008 and 2013 (Leshem et al., 2018). These estimations, however, principally represent the effects of vaccine administration in medically privileged countries, such as the United States, as well as some developing countries. As of 2013, RVs were still responsible for over 215,000 deaths per year worldwide (Tate et al., 2016). Several studies have shown differential efficacy of the RV vaccines in some low-income countries, where children are most vulnerable to RV-related morbidity and mortality due to limited access to medical intervention (Marcy and Partridge,

2017). Augmenting this issue, RVs shed in fecal specimens of babies vaccinated with the pentavalent RV vaccine, RotaTeqTM, were detected up to 9 days post-vaccination and could be cultivated, suggesting potential for transmission of vaccine strains (Yen et al., 2011).

From a translational science standpoint, studying RV biology is significant because of its impact on the health of children worldwide.

RV Genome and Capsid Structure

Early electron microscopic images informed RV particle morphology. A “wheel”- like smooth outer edge adorned with spike-like projections inspired the virus’ name, which stems from Latin root “rota” (Flewett et al., 1974). Further analysis revealed that RVs are nonenveloped icosahedrons that encapsidate 11 individual genome segments (Davidson et al., 1975; Schnagl and Holmes, 1976; Prasad et al., 1988).

The 11-segmented RV genome totals ~23 kilobases (kb) and consists of double- stranded (ds) ribonucleic acid (RNA). Ranging in length from about 0.7 to 3.3 kb, each

2 linear segment of dsRNA is numbered (g1-g11) based on its relative migration during polyacrylamide gel electrophoresis. All 11 genome segments’ positive-sense (+)RNAs contain a central open reading frame (ORF) flanked by 5 and 3’ untranslated regions

(UTRs). While +RNAs’ UTRs vary in length, their extreme ends are highly conserved and are thought to contain signals important for initiation of viral genome replication (reviewed in Estes and Cohen, 1989; Tortorici et al., 2003). Each +RNA is monocistronic, with the exception of g11, which contains an alternative ORF that encodes for an additional protein,

NSP6, only expressed in some RV strains (Rainsford and McCrae, 2007).

RV’s protein capsid consists of viral proteins (VPs) 1-4, 6, and 7. The mature, infectious RV virion is a triple-layered particle, or TLP (Figure 1). The outermost layer of the TLP consists of a smooth VP7 decorated with VP4 spike proteins. Underneath VP7 and

VP4 lies intermediate-layer protein, VP6, which forms the outer layer of the double-layered particle (DLP) (reviewed in Desselberger, 2014). Finally, the core shell, or innermost layer of the mature virion is comprised of conformational heterodimers of VP2 arranged into flower-like decameric units. Positioned just off-center from most, if not all, of the 12, five- fold icosahedral vertices of the VP2 lattice, are single copies of VP1, the viral RNA- dependent RNA polymerase (RdRp) (Estrozi et al., 2013). Also thought to be associated with VP2-VP1 complexes is VP3, viral guanylyl/methyl transferase. The remainder of the virus’ proteome consists of nonstructural proteins (NSPs) 1-6 (Figure 1). While VPs contribute to the structure of the protein capsid, NSPs are not packaged into the mature virion; instead, NSPs are only expressed in host cells during the viral replication cycle

(reviewed in Desselberger, 2014). For the remainder of this thesis, RV +RNAs will be referred to as their respective gene numbers (g1-g11), and proteins will be referred to as

3 their respective protein names (VP1-4, VP6-7, and NSP1-5); for example, genome segment

11, or g11, encodes two proteins, NSP5 and sometimes NSP6.

Figure 1. Mature RV virion and genome composition

(A) The mature RV virion and its structural and genetic components, including VP4

(red), VP7 (yellow), VP6 (green), VP2 (blue), VP3 (purple), VP1 (pink), and genomic dsRNA (gray). (B) Proteins encoded by respective genome segments, g1-g11, and their lengths (in amino acid residues), with structural proteins colored as in panel A and nonstructural proteins in white. Figure made by Courtney Steger, Ph.D.

RV Classification

RV is prototypic member of the Reoviridae family of viruses, which describes a group of viruses characterized by icosahedral capsids that enclose 9-12 dsRNA genome segments. RVs are classified at several levels. The first tier of RV classification describes

RV “species”—namely, groups—that were initially formed based on differential antigenic properties conferred by VP6 and later assigned based upon sequence analysis of the VP6- coding gene (Hoshino and Kapikian, 1996; Matthijnssens, 2012). Although more than eight genetically-distinct RV groups have been identified in nature (groups A-H and tentative groups I and J), humans are infected almost exclusively by Group A RVs (RVAs), with only rare infections caused by Groups B and C RVs (Estes and Kapikian, 2007; Mihalov-

Kovács et al., 2015; Bányai et al., 2017). RVAs are most well-studied and are the focus of this thesis.

Because its genome is segmented and individual segments can independently reassort (see RV Reassortment), RVA is further classified at the genic level. Historically, classification of RVAs involved serotyping, limited to the detection of RVA outer-capsid proteins VP4 and VP7 in patient serum by neutralizing antibodies (Greenburg, 1983).

Currently, individual genome segments are differentiated by various genotypes that are determined by specific nucleotide percent identity cutoff values for each segment. For each each genome segment, 20-51 genotypes have been established by the RV Classification

Working Group (Matthijnssens et al., 2011). In this system, each genome segment is designated by a letter, which corresponds to the genome segment’s encoded protein’s function. For example, H denotes the genome segment that encodes NSP5, which is hyperphosphorylated. 22 different genotypes for the NSP5-encoding gene (H1-H22), which are each less than 91% identical to each other in sequence similarity (Table 1).

Strung together, these genotypes inform genotype constellations that take the form of: Gx-

P[x]-Ix-Rx-Cx-Mx-Ax-Nx-Tx-Ex-Hx, wherin “x” represents the genotype as an integer.

Table 1. RV proteins, functions, genotypes, and percent identity cut-off values

Protein Function(s) Genotypes Cut-Off Value (%) VP1 RNA-dependent RNA polymerase R1-22 83 VP2 Core shell protein C1-20 84 VP3 mRNA capping enzyme M1-20 81 VP4 Protease-sensitive spike P1-51 80 NSP1 Antagonist of innate immunity A1-26 79 VP6 Intermediate layer I1-36 85 VP7 Glycoprotein G1-31 80 NSP2 NTPase, viroplasm N1-22 85 NSP3 Translation enhancer T1-22 85 NSP4 Enterotoxin E1-27 85 NSP5 Hyperphosphorylation, viroplasm H1-H22 91

RVA strains are defined by their genotype constellations. For human RVAs, genotype constellations inform the designation of genogroups, which are groups of individual strains that are genetically similar in their genotype constellations (Nakagomi and Nakagomi, 2002). Whole genome classification of RVAs supports the existence of two major genogroups that dominate human infections, Wa-like and DS-1-like (Figure 2). Wa- like RVs, of which the laboratory-adapted strain Wa is the reference, exhibit constellations of genotype 1 internal genes (or those encoding for non-outer-capsid proteins) (I1-R1-C1-

M1-A1-N1-T1-E1-H1) and outer-capsid protein-encoding genotype combinations G1P[8],

G3P[8], G4P[8], and G9P[8] (Wyatt et al., 1974; Heiman et al., 2008). The Wa-like RVA genogroup also contains laboratory reference strains P and D. In contrast, DS-1-like RVAs, of which the laboratory strain DS-1 is representative, exhibit constellations of genotype 2 internal genes (I2-R2-C2-M2-A2-N2-T2-E2-H2) and an outer-capsid protein-encoding genotype combination G2P[4] (Figure 2) (Heiman et al., 2008).

Figure 2. Human RVA genotype constellations

Representation of human RVA genotype constellations. Each column represents a genome segment and is labeled according to respective encoded protein(s). Rows depict genotype constellations as strings of individual genotypes differentiated by color. A description of each strain can be found to the left, and each lists the strain name using the convention: RV group/host and source of isolation/country of isolation/common strain name/year of isolation. Wa-like genotype constellations (green) are shown atop DS-1-like constellations

(red).

RVA Lifecycle and Assortment of Viral RNAs

Initial attachment of RVA particles to intestinal epithelial cells is mediated by the viral spike protein, VP4. First, trypsin-like proteases within the host digestive tract cleave

VP4 into fragments known as VP5 and VP8. The VP8 fragment of VP4 mediates cell surface attachment (Dormitzer et al., 2002). Strains with different P-types have been shown to bind different cell-surface glycans—including neuraminidase, sialic acid, and histo- blood group antigens—as a first step in cell entry (Ciarlet et al., 2001; Huang et al., 2012).

It is likely that initial binding of an RVA to cell types of different hosts is limited by its P- type, and this idea presents considerable implications for RVA transmission and vaccine response (Ramani et al., 2016).

Upon contact with the host cell, RV’s outer capsid protein, VP7, interacts with various co-receptors, initiating one of several possible endocytic pathways—a process also thought to be governed by P-type (Diaz-Salinas et al., 2014). Regardless of the pathway used, vesicles containing virus particles fuse with the early endosome (EE). The calcium- depleted, low pH environment of the EE induces shedding and solubilization of VP7, exposing VP6 and forming the DLP transcriptase complex (Gerasimenko et al., 1998;

Ludert et al., 1987). Solubilized VP7 may lyse the endosomal membrane, but more investigation into the precise mechanism of DLP release into the cytosol is still needed

(Ruiz et al., 1997; Abdelhakim et al., 2014).

Within the DLP, viral RdRp VP1 unwinds the viral dsRNA genome segments and uses negative-sense (-)RNAs as templates for the transcription of +RNAs. Structural analysis of VP1 reveals two RNA exit tunnels, the first of which directs nascent transcripts towards a pentameric gauntlet opening into the cytosol following the addition of a 5’ m7GpppG cap structure (Lu et al., 2008; Lawton et al., 1997). Once viral +RNAs enter the

10 cytosol, some are translated by host ribosomes with the aid of NSP3. While bearing 5’ caps, RVA +RNAs lack polyadenylation signals necessary for translation in eukaryotic cells. NSP3 facilitates translation of viral +RNAs by binding a 3’ consensus sequence of viral +RNAs and interacts with eukaryotic initiation factor 4G, evicting its poly(A)-binding protein and enhancing RVA +RNAs’ competition for host ribosomal machinery (Piron et al., 1998). Untranslated +RNAs are sequestered into nascent viroplasms—or viral replication factories—where new virions begin assembling and replicating their genomes using the +RNAs as templates (Silvestri et al., 2004).

The segmented nature of the RVA genome poses a significant challenge to nascent virions: How are all 11 segments of the RVA genome packaged into a single viral particle?

Some segmented RNA viruses are promiscuous in their selection of RNAs, resulting in random packaging into defective interfering particles incapable of propagation (Luque et al., 2009). Reoviridae family members are thought to have evolved a more sophisticated packaging mechanism, in which RNAs are recognized by proteins involved in replication and assembly and packaged selectively, or assorted (reviewed in McDonald and Patton,

2010). Coupled with the fact that there have been no reports of infectious RVAs encapsidating greater or fewer than 11 full-length segments, evidence in support of selective packaging and assortment of +RNAs stems from analysis of RVA +RNA sequences and predicted secondary structures that are thought to contain important cis- acting functional elements. While specific assortment signals remain unclear due to limited in vitro assays, available data supports a model in which sequence specificity between

RVA +RNAs promotes inter-segment interactions to form a supramolecular complex

(Borodavka et al., 2017). Stable stems have been predicted to form from sequence complementarity between UTR sequences immediately flanking ORFs of viral +RNAs,

11 but introduction of oligos complementary to the extreme 3’ ends of the +RNAs were shown to ablate -RNA synthesis in vitro, suggesting the importance of an accessible 3’ tail in the selection of +RNAs for genome replication (Chen and Patton, 1998). In addition, RVAs whose genome segments lack ORFs have been identified, further suggesting that UTRs of

RVA +RNAs may function as assortment signals that may be necessary for replication of the RV genome (reviewed in Desselberger, 1996). RVA proteins are thought to play roles in the recognition and sequestration of RVA +RNAs into the core replicase intermediate

(RI) in preparation for genome replication (reviewed in McDonald and Patton, 2011). Both

NSP2 and NSP5 have been shown to possess single- and double-stranded RNA binding capabilities (Taraporewala et al., 1999; Vende et al., 2002). The N-terminal domain of VP2 alone (residues 1-132) has been shown to bind RNA, and N-terminal truncation mutants exhibit diminished dsRNA synthesis when paired with VP1 in vitro (Labbé et al., 1994;

McDonald and Patton, 2011). These results, coupled with the observation that the N- terminal domain of VP2 contains highly conserved positively-charged residues, suggest that the core shell protein is involved in RNA binding and might be important for assortment of viral +RNAs. Evidence also supports the important role of NSP2 in the assortment process. More specifically, NSP2 is thought to induce +RNA remodeling such that specific RNA-RNA contact sites are exposed for interaction with other viral +RNAs, leading to the formation of a supramolecular complex of viral +RNAs (Borodavka et al.,

2017).

RV genome replication (dsRNA synthesis) by VP1 is dependent on putative interaction with core shell protein, VP2, which is thought to induce a conformational switch that activates the RdRp to initiate dsRNA synthesis within the confines of the core RI

(Patton et al., 1997; Lu et al., 2008). It has been demonstrated that NSP5 binds VP2, but

12 not in the presence of VP6; this result, paired with outcomes of studies using reovirus, suggests that NSP5 may engage VP2 as a control mechanism to prevent premature DLP and transcriptase complex formation during dsRNA synthesis (Berois et al., 2003). In a similar manner, the TLP cannot be formed until the DLP assembles and encounters the outer-capsid glycoprotein, VP7, at the endoplasmic reticulum (ER) lumen, the site of mature RV virion formation (reviewed in Trask et al., 2012). This process relies on NSP4, which serves as a viral ER membrane-bound receptor (O’Brien et al., 2000; Taylor et al.,

1993). Thought to act as a chaperone, NSP4 traffics not only nascent DLPs, but also cytosolic VP4, into the ER lumen (Trask and Dormitzer, 2006). Although the precise mechanism through which NSP4 elevates cytosolic calcium concentrations is unknown,

VP7 outer-capsid formation and stabilization is dependent on this phenomenon (Tian et al.,

1994). In addition to usurping of ER functions, NSP4 has been shown to trigger autophagic cascades (Crawford et al., 2012). Like entry, mechanisms of RV egress appear non- uniform. Early studies demonstrated that RVs are released via cell lysis, while more recent studies using polarized intestinal epithelial cells suggest the use of secretory exocytosis

(Figure 3) (Musalem and Espeio, 1985; Jourdan et al., 1997).

Figure 3. Overview of RVA lifecycle

RVA adheres to host cells via binding myriad potential cell-surface glycans. Endocytosis and subsequent fusion to the EE promotes VP7 dissociation from VP6, leading to the formation of a transcriptionally active DLP. Some viral transcripts are translated on host ribosomes, while others are sequestered into viroplasms, sites of viral +RNA assortment and genome replication, along with some nascent viral proteins. RVA matures as its trafficked through the ER and leaves the cell either via lysis or exocytosis. Figure made by

Courtney Steger, Ph.D.

RVA Reassortment

Capacity for assortment is a shared feature of viruses with segmented genomes.

While viral assortment refers to the selective packaging of a full complement of genome segments into nascent virions, reassortment describes the capacity of two (or more) parental strains to exchange genome segments, resulting in hybrid progeny that are genetically distinct from either individual parental strain (reviewed in McDonald et al.,

2016). While specific packaging and assortment signals have yet to be identified, theoretically, two strains of RV whose +RNAs share 5’ and 3’ UTR sequence similarity and predicted +RNA stem-loop structures could be compatible with RNAs and proteins of either strain. Considering the breadth of RV genotype constellations found in nature and culture, reassortment occurring in nature is conceivable. Such events could confer selective advantages to reassortant progeny, such as the ability to infect a new host and evade its immune response. Although inter-group reassortment has never been detected, it was widely assumed that intra-group reassortment of RVAs occurs readily. This idea would support random association of RVA genome segments due to chance encounters and the existence of countless genetically divergent strains. When RVAs are isolated from stool samples of infected children and sequenced, however, stable genotype constellations that adhere to strict patterns predominate, with reassortants documented infrequently (see

Figure 2) (Heimann et al., 2008; McDonald et al., 2009; reviewed in McDonald et al.,

2016). Further, the majority of reassortants identified in nature have been limited to genome segments that encode outer-capsid proteins VP4 and VP7, but even these genome segments tend to pair preferentially (reviewed in Cunliffe et al., 2002). Reassortment is also less common than predicted in culture; cells co-infected with RVAs from human and animal hosts produced few reassortants relative to what is expected due to chance alone

(Graham et al., 1987). These outcomes suggest that the capacity for RVAs to reassort depends on genetic and/or biochemical compatibility between viral RNAs and proteins.

Further, nucleotide and amino acids differences between strains may inform interactions between viral RNAs and proteins critical for the formation of nascent virions.

RVA Reverse Genetics

Historically, reassortment was studied using partial plasmid-based reverse genetics systems that were dependent on helper viruses and limited to the rescue of RVAs containing only one recombinant gene. With this approach, rescue of recombinant (r)RVAs was limited to those fit enough to be selected for in the presence of helper viruses, which posed significant challenges to the study of reassortant viruses that bore even moderate replicative defects (Komoto et al., 2006; Trask et al., 2010; Troupin et al., 2010; Johne et al., 2015; Mingo et al., 2017). In 2017, an all plasmid-based reverse genetics system for

RVA was developed for simian strain, SA11 H-96 (Kanai et al., 2017). 11 plasmids encoding for each SA11 H-96 genome segment (i.e. +RNA), as well as 3 accessory plasmids, encoding an orthoreovirus fusion-associated small transmembrane (FAST) protein and two subunits of a vaccinia virus capping enzyme, D1R and D12L, are transfected into baby hamster kidney (BHK) cells stably expressing T7 polymerase (BHK-

T7 cells). Genes encoded by each plasmid are flanked by a 5’ T7 promoter and 3’ antigenomic hepatitis delta virus (HDV) ribozyme sequence. BHK-T7 cells stably express

T7 polymerase that recognizes the T7 promoters of transfected plasmids, and the HDV ribozyme serves to cleave transcripts at their 3’ ends, yielding full-length RV messages.

The FAST protein functions to fuse BHK-T7 cells and increase the likelihood that all 11

RV transcripts are contained within a single plasma membrane (Kanai et al., 2017). In the

16 original system, D1R and D12L act in concert to cap the 5’ ends of +RNAs derived from

T7 transcription. The Patton lab (Indiana University) has replaced these two constructs with just one African horse sickness virus (AHSV) construct, NP868R, which enhanced rescue of wildtype recombinant SA11 (rSA11-wt) when tested against D1R and D12L

(Figure 4) (personal communication).

Figure 4. SA11 H-96 reverse genetics system

Brief overview of reverse genetics system for SA11 H-96. 11 plasmids each code for one genome segment, and two accessory plasmids code for a FAST fusion protein and an

NP868R capping enzyme. Each gene is driven by a T7 promoter sequence, and each RVA gene is followed by an antigenomic HDV ribozyme sequence. The aforementioned plasmids are transfected into BHK-T7 cells that stably express T7 RNA polymerase. Gene expression yields +RNAs capable of launching infection and production of recombinant, wildtype SA11 H-96 or engineered mutants. Recombinant RVAs are then passaged in

RVA-permissive MA104 cells. Adapted from Kanai, et al. (2017).

Since its creation, improvements to the system’s first iteration have been published.

One study demonstrated that the plasmid burden could be reduced to just 11 cDNA plasmids encoding each of the 11 RV genome segments by increasing the concentrations of plasmids encoding NSP2 and NSP5 3-fold relative to the concentrations used in the original system, suggesting the importance of viroplasm formation to the system’s ability to produce recombinant RVs (Komoto et al., 2018). Stable reporter RVs expressing luciferase or green or red fluorescent proteins from within a truncated NSP1 ORF were generated using this optimized system (Kanai et al., 2019). While changes have improved the system’s ability to produce rRVAs capable of initiating productive infection, its rescue efficiency is relatively low. Thus, the first aim of this work was to attain system functionality in the McDonald lab and optimize its rescue efficiency.

While the development of this system constitutes a significant advancement for the study of RV biology in general, knowledge gained by the utilization of the current system is limited to understanding of simian strain SA11. Exhibiting robust growth in culture,

SA11 was a clear candidate for the creation of an RV reverse genetics system (Estes et al.,

1979). Infection of cultured cells with cell culture-adapted human RV strains is not as productive as SA11 infection. However, to understand human RV replication and pathogenesis, the development of a reverse genetics system utilizing human RV genotype constellations would be key (Figure 5). As a result, the second aim of this work was to attempt to develop a reverse genetics system using a Wa-like RVA strain with the rationale that Wa-like RVAs dominate human infections. While working towards this project goal, a reverse genetics system that utilized an additional cell line (CV-1 cells) and roller bottles for recombinant RV infection and propagation resulted in the successful rescue of infectious recombinant human RV strain KU (Komoto et al., 2019).

Figure 5. Human vs. simian RVA genotype constellations

Representation of RV genotype constellations. Each column represents a genome segment and is labeled according to respective encoded proteins. Rows depict genotype constellations as strings of individual genotypes differentiated by color. A description of each strain can be found to the left, and each lists the strain name using the convention: RV group/host and source of isolation/country of isolation/common strain name/year of isolation. Wa-like and DS-1-like genotype constellations represent the vast majority of human RV infections, and both are shown above genotype constellation examples of simian strain, SA11. G3-P[2]-I2-R2-C5-M5-A5-N2-T5-E2-H5 is the constellation used for the creation of a reverse genetics system for RV, and diverges vastly from constellations of strains that infect humans.

The current SA11 H-96-based system is useful for probing RVA reassortment.

Mono-reassortant viruses could be created, or rescued, simply by exchanging an SA11 plasmid with another encoding for the same genome segment of a different strain. Kanai et al. developed a mono-reassortant virus in which the SA11 VP6 genome was exchanged with that of human RV, KU. In culture, the KU VP6 reassortant virus exhibited only a mild replicative defect detected by a multicycle growth analysis (Kanai et al., 2017). While this result demonstrated that the reverse genetics system can be used to rescue a mon- reassortant RV bearing a mild replicative defect, it was hypothesized that it could be used to rescue other mono-reassortant RVs that may exhibit more substantial replicative defects.

The third aim of this work was to create a different mono-reassortant RVA and analyze its growth in culture. The rescue of a mono-reassortant RVA would indicate that the nucleotide and amino acid differences between the genome segment exchanged would not be sufficient to ablate RNA-RNA, RNA-protein, and protein-protein interactions critical for the formation of an infectious reassortant virus. Specifically, it was hypothesized that an H1 genotype NSP5-encoding genome segment from human rotavirus strain, Wa, could functionally replace that an H5 genotype in the genetic context of SA11 and that this mono- reassortant virus, rSA11-g11-Wa, could be rescued. A g11 reassortant provided a candidate for likely successful rescue and functional compatibility between Wa and SA11 because among all RV genome segments, g11 exhibited the highest sequence identity between the two genotypes. Rescue of rSA11-g11-Wa would validate the system’s ability to successfully produce mono-reassortant viruses, which would be necessary for the creation of future mono-reassortant RVAs to study their biology. The reassortant virus’ growth in culture relative to wildtype SA11 would be informative of potential replicative defects conferred by the nucleotide and amino acid differences between H1 and H5.

RVA NSP5

The primary role that NSP5 serves in the infectious RV lifecycle is formation of the viroplasm. Early experiments demonstrated that it binds NSP2 following chemical crosslinking in vivo (Afrikanova et al., 1998). Also, when co-transfected, plasmids expressing these NSP5 and NSP2 formed viroplasm-like structures in vivo (Fabretti et al.,

1999). While no high-resolution structure exists, cryo-electron microscopy has captured a truncated NSP5 (residues 68-188) in complex with an NSP2 octamer (Jiang et al., 2006).

Martin et al. mapped the site of specific interaction with NSP2 to amino acid residues 132-

146; this NSP2 binding region is also contained within a larger region important for dimerization of the NSP5 protein (Figure 6) (Matrin et al., 2011). As described previously,

NSP5 is known to interact with VP2 and is thought to stall the formation of the core RI during +RNA assortment in preparation for genome replication (Berois et al., 2003).

Analysis of NSP5 C-terminal amino acid sequence predicts an amphipathic alpha helix, which could serve various functional roles within the viral lifecycle. Specifically, C- terminal amino acid residues 187-198 were found to be necessary for multimerization of the NSP5 protein (Figure 6) (Torres-Vega et al., 2000). One study found that the C-terminal domain of NSP5 alone was sufficient to bind VP1 (Arnoldi et al., 2006). Another study, noting the morphological similarities between viroplasms and lipid droplets, hypothesized that the putative amphipathic NSP5 C-terminal helix serves to anchor nascent virions to lipid droplets (Cheung et al., 2010). Finally, NSP5 is known to interact with NSP6. As mentioned previously, NSP6 is only expressed by some strains of RV and not others, and results of competitive growth assays of viruses engineered to lack the NSP6 ORF demonstrate that NSP6 does not play an essential role in the RV lifecycle (Komoto et al.,

2017).

Each of the aforementioned protein interactions facilitate NSP5’s chief functional role as driver of viroplasm formation, in tandem with NSP2. NSP5’s function is thought to be dependent on hyperphosphorylation by NSP2, cellular kinases, and its own autokinase activity (Afrikanova et al., 1996; Blackhall et al., 1998). NSP5 phosphorylation is thought to function as a conformation switch, prerequisite for the formation of viroplasms. The current model for NSP5 phosphorylation suggests that NSP2 binds the 26-kDa form of the protein and phosphorylates it, producing a 28-kDa form. At this point, the 28-kDa NSP5 in complex with NSP2 is recruited to the viroplasm, and cellular casein kinase 1 a (CK1a) is thought to play a role in NSP5 hyperphosphorylation, producing 32- and 35-kDa forms

(Criglar et al., 2018). Several conserved candidate phosphorylation sites have been determined, and all map to serine residues, including Ser67, Ser153, Ser155, Ser163, Ser165

(Eichwald et al., 2002). Within the viroplasm, NSP5 is thought to play an important role in

+RNA assortment and sequestration to and within the viroplasm due to its single-stranded and dsRNA binding capabilities (Vende et al., 2002). If rSA11-g11-Wa could be rescued, it would indicate that nucleotide and amino acid residue differences between SA11 and Wa

NSP5 would be insufficient to ablate RNA-RNA, RNA-protein, and protein-protein interactions necessary for assortment and replication to the extent that these processes depend on NSP5.

Figure 6. Important functional regions of NSP5

Linearized protein schematic of SA11 and Wa NSP5. In both proteins, amino acid residues

103-146 are important for initial dimerization of the NSP5 protein, residues 132-146 are important for binding NSP2, and C-terminal residues 187-198 are important for multimerization.

Research Aims

The goals of this body of work were to: (1) attain functionality of and optimize the current reverse genetics system for simian RV strain SA11 H-96, (2) develop a reverse genetics system for human RV Wa, and (3) using the existing reverse genetics system, create a reassortant rRV that contains g11/NSP5 from human strain Wa within the SA11 H-96 genetic backbone (rSA11-g11-Wa) and test its growth in culture relative to wildtype rSA11 H-96 (rSA11-wt). Both human and simian-based reverse genetics systems for RVAs could be used to address the hypothesis that reassortment among

RVAs is restricted by biochemical incompatibilities between heterologous strains’ RNAs and proteins. Nucleotide and amino acid differences sufficient to ablate viral replication in cell culture could yield insight into which RNA-RNA, RNA-protein, or protein-protein interactions are critical for the formation of nascent virions. Replicative defects displayed by reassortant RVAs could be indicative of indirect restrictions on reassortment.

METHODS

Phylogenetic Analysis of Modern Human, Wa-like RVA Strains

Complete sequences for archival, culture-adapted RVA strains Wa, P, D, and KU were obtained using the National Center for Biotechnology Information (NCBI)’s nucleotide database for each of the Wa-like genotypes (G1 or G3, P[8], I1, R1, C1, M1,

A1, N1, T1, E1, and H1). Complete sequences for archival DS-1 genotypes (G2, P[4], I2,

R2, C2, M2, A2, N2, T2, E2, and H2) were obtained in a similar manner to serve as outgroup representatives. Accession numbers for archival strain sequences used can be found in the appendix. Other archival sequences of RVs isolated from patients at DC

Children’s Hospital were included in this analysis for comparison to both culture-adapted archival strains and modern strains (Heiman et al., 2008). The NCBI’s Virus Variation RV database was used to obtain an average of 160 complete gene sequences for each of the aforementioned Wa-like genotypes that were isolated from RVA-infected humans in various geographic regions from 2010 to present. For each genotype, Geneious Pro v10.2.3

(BioMatters) software was used align nucleotide sequences using the CLUSTALW cost matrix. Alignments were trimmed to boundaries of shared complete sequence. Final consensus trees were constructed using Geneious Tree Builder’s neighbor-joining method with a Jukes-Cantor genetic distance model, including 1,000 bootstrap replicates and a random seed of 98,101. FigTree v1.4.3 was used to collapse monophyletic branches and create figures.

Cell Culture

African green monkey kidney epithelial (MA104) cells were maintained as described by Arnold et al. in medium M199 (Gibco) that was supplemented to contain

5% heat-inactivated fetal bovine serum (HI-FBS) (Atlanta Biologicals), 100 U/mL penicillin, and 100 µg/mL streptomycin (Gibco) and split once they reached approximately 90% confluence at dilutions ranging from 1:2 to 1:10 every 3-7 days

(2009).

Baby hamster kidney epithelial cells stably expressing T7 RNA polymerase

(BHK-T7) were kindly gifted by Dr. John Patton (Indiana University) who received cell stocks from Dr. Peter Collins (National Institutes of Health). BHK-T7 cells were maintained in Dulbecco’s Modified Eagle Medium (DMEM) that was supplemented with 5% HI-FBS, 10% tryptose broth (Gibco), 1% 200 mM L-glutamine (Invitrogen),

100 U/mL penicillin, and 100 µg/mL streptomycin. Cells stably expressing T7 RNA polymerase were selected for by supplementing media with 2% 50 mg/mL G418 (Gibco) every other passage. Cells were maintained at 37°C and 5% CO2 and split once they reached approximately 90% confluency at a 1:20 dilution every 2-3 days.

Plaque Assays

RV-infected MA104 cell lysates were clarified via centrifugation at 1,808 x g for 5 minutes. Trypsin, porcine pancreatic type IX (Sigma-Aldrich), was added to the clarified supernatant at a final concentration of 10 µg/mL. Following activation at 37°C for 1 hour, the supernatant containing RV was serially diluted in 10-fold in serum-free M199 to yield virus concentrations ranging from 10-1 to 10-8. Once MA104 cells seeded in 6- well plates form tight monolayers, each dilution was used to infect duplicate wells of cells at 37°C in 5% CO2 for 1 hour. Following infection, inoculum was removed and cells were washed once with serum-free M199. 2 mL of a 1:1 mixture of 2% SeaPlaque agarose (Lonza) and serum-free 2X E-MEM (VitaScientific) containing 0.5 µg/mL of

27 trypsin was used to overlay infected MA104 cells in each well. Once agarose mixtures solidified, cells were placed at 37°C in 5% CO2. Approximately 3 days post-infection, plaques, or concentrated areas of cell death, could be visualized, and 3 mL of a 1:1 mixture of 1.2% agarose and serum-free 2X E-MEM containing 50 µg/mL of viability stain neutral red (Sigma-Aldrich) was used to overlay infected MA104 cells in each well. Once agarose mixtures solidified, cells were placed at 37°C in 5% CO2 until plaque forming units

(PFUs) were counted 4-24 hours later.

Preparation of Viral RNAs

Viral RNAs were extracted from Wa-infected MA104 cell lysate to serve as a template for the generation of authentic, full-length cDNAs via RT-PCR. Specifically, 250

µL of lysate was added to 750 µL of TRIzol LS Reagent (Invitrogen). To the lysate/TRIzol mixture, 200 µL of chloroform was added, mixed vigorously, and incubated at room temperature for 15 minutes prior to centrifugation at 16,873 x g for

10 minutes. The aqueous layer was transferred, and one-tenth volume of 3M sodium acetate, pH 5.2 and one volume of isopropanol was added, and the mixture was incubated overnight at -20°C to precipitate the RNA. The mixture underwent centrifugation at 16,873 x g for 30 minutes. Supernatant was discarded, and the pellet was washed with 1 mL of 70% ethanol prior to centrifugation at 16,873 x g for 10 minutes. Supernatant was discarded, and residual ethanol was discarded after centrifugation at 16,873 x g for 1 minute. The pellet was air-dried and resuspended in

20 µL of DEPC-H20.

Plasmid Construction

An existing reverse genetic vector containing a T7 promoter and an antigenomic hepatitis delta virus (HDV) ribozyme sequence was purchased from AddGene (Kanai et al., 2017) and amplified via outward PCR using AccuPrime Pfx SuperMix according to the manufacturer’s protocol (Invitrogen). Following denaturation of viral dsRNAs by 1:1 dilution with DMSO at 94°C for 10 minutes, RT-PCR amplification of RVA genome segments was carried out using SuperScript OneStep RT-PCR System with Platinum Taq

DNA polymerase according to the manufacturer’s protocol (Invitrogen). Primers used for

(RT-)PCR amplification are listed in Table 2.

Table 2. Primer sequences for Wa reverse genetics constructs

Primer Sequence

pT7-Forward GCATGGTGCAGAGTCGACTATCGTA—GG__TCGGCATGGCATCTCCACCTCCTCGC

pT7-Reverse 1 GCATGGTGCAGAGTCGACTATCGTA__AA—GCCTATAGTGAGTCGTATTAACCGGG

NSP5-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TAAAAGCGCTACAGTGATGTCTCTCA

NSP5-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACAAAACGGGAGTGGGGAGCT

NSP4-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TTAAAAGTTCTGTTCCGAGAGAGCGC

NSP4-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA— CC— CGGTCACACTAAGACCATTCCTTCC

NSP3-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TTAATGCTTTTCAGTGGTTGTTGCTCAAGATGG

NSP3-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATAACGCCCCTATAGCCATTTAGG

NSP2-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TTAAAGCGTCTCAGTCGCCGTTTGAGCC

NSP2-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATAAGCGCTTTCTATTCTTGC

VP7-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TAAAAGAGAGAATTTCCGTCTGGCTAACG

VP7-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATCGAACAATTCTAATCTAAG

VP6-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TAAAACGAAGTCTTCGACATGGAGGTTC

VP6-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATCCTCTCACTACATCATTC

NSP1-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TTTTTATGAAAAGTCTTGTGGAAGCCATGG

NSP1-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATTTTATGCTGCCTAGGCGC

VP3-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TT__TTAAAGCAGTACTAGTAGTGCGTTTTACC

VP3-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATCATGACTAGTGTGTTAAGTTTC

pT7 Reverse 2 GCATGGTGCAGAGTCGACTATCGTA__TA—TAGTGAGTCGTATTAACCCGGGATCG

VP4-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TA__GGCTATAAAATGGCTTCGCTCATTTATAG

VP4-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATCCTCAATAGCGTTCTCAC

VP2-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TA__GGCTATTAAAGGCTCAATGGCGTACAGG

VP2-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCATATCTCCACAGTGGGGTTGGCG

VP1-Wa Forward GCATGGTGCAGAGTCGACTATCGTA—TA__GGCTATTAAAGCTGTACAATGGGGAAGTAC

VP1-Wa Reverse GCATGGTGCAGAGTCGACTATCGTA__CC—CGGTCACATCTAAGCACTCTAATCTTGAAAG

Primers used for RT-PCR or PCR were designed such that a BsgI recognition site

(emboldened and underlined) was flanked by 5 randomly chosen nucleotides on the 5’ end and 14 randomly chosen nucleotides on the 3’ end, leading to the site of cleavage by the restriction endonuclease. Two-base overhangs left by the restriction endonuclease are noted by dashes. The remainder of each sequence was complementary to plasmid or Wa sequence. “pT7-Reverse 1” was used to clone NSP5, NSP4, NSP3, NSP2, VP7, VP6, and

VP3, while “pT7-Reverse 2” was used to clone VP4, VP2, and VP1 reverse genetics constructs due to differential sequence complementary in overhang regions of these genes.

RT-PCR and PCR products were spin-column purified (QIAGEN) and digested with restriction endonuclease BsgI according to the manufacturer’s instructions (NEB).

Vector PCR products were also digested with DpnI to eliminate methylated template as described by the manufacturer (NEB). Digested insert and vector were gel purified

(Invitrogen) and ligated using T4 DNA ligase (NEB) at a 10:1 molar ratio. Ligation products were transformed into One Shot TOP10 Chemically Competent E. coli cells according to the manufacturer’s protocol (Invitrogen), plated on medium containing Luria broth (LB) base, Miller (Sigma-Aldrich), bacteriological agar (Sigma-Aldrich), and 100

µg/mL of ampicillin, and grown at 37°C for 16 hours. Bacterial cultures of individual colonies were grown in LB base, Miller and 100 µg/mL ampicillin at 37°C for 16 hours and miniprepped (QIAGEN) to extract plasmid DNA. Plasmids were then analyzed via

Sanger sequencing. Figure 5 shows an overview of the cloning scheme used to generate reverse genetics constructs containing Wa genes.

Figure 7. Cloning scheme for reverse genetics constructs containing RVA strain Wa genes

Using NSP5 as an example, following (RT-)PCR amplification Wa RNA and pT7 vector

DNA, cDNAs were digested with restriction endonuclease, BsgI. The enzyme cleaves dsDNA 14 base pairs downstream of its recognition site, leaving a two-base overhang.

Two-base overhangs at 5’ and 3’ ends of inserts were complementary to 3’ and 5’ ends of the vector, respectively.

Generation of Recombinant RVAs using an All Plasmid Reverse Genetics System

BHK-T7 cells were seeded at a densities of 5 x 104 or 105 cells in 2 mL of complete

DMEM per well of a 6-well plate. 48 hours later, cells were washed twice with complete

DMEM, and media was replaced with 2 mL of complete DMEM. Cells were then transfected with mixtures containing 1 µL of each plasmid at concentrations of 0.15 µg of pT7-FAST and 0.8 µg of each of the remaining 12 plasmids, 250 µL of OptiMEM

(Gibco), and 19.2 µL of TransIT-LT1 Transfection Reagent (Mirus) after a 20-minute incubation at room temperature; the negative control lacked the pT7-NSP2 plasmid, of which the encoded protein is necessary for viral replication. 48 hours post-transfection,

MA104 cells were split and pelleted at 200 x g for 5 minutes, washed with 10 mL of serum-free DMEM, and re-pelleted at 200 x g for 5 minutes. Following resuspension in serum-free DMEM, MA104 cells were counted using an automatic cell counter (Life

Technologies) and seeded at a density of 105 cells in 2 mL of serum-free DMEM and

0.5 µg/mL trypsin per well. 72 hours post-MA104 overseed, BHK-T7 and MA104 cells were lysed by 3 rounds of freeze-thaw, and virus was clarified by centrifugation at 1,808 x g for 5 minutes. Following activation with 10 µg/mL of trypsin at 37°C for 1 hour, 1 mL of virus was used to infect a new 6-well plate of MA104 cells. Following infection at 37°C and 5% CO2 for 1 hour, cells were washed once with serum-free M199, and media was replaced with serum-free M199 with 0.5 µg/mL trypsin. Cytopathic effect

(CPE) was observed 1-4 days following infection.

Imaging of BHK-T7 Cells Transfected with Plasmid Encoding Green Fluorescent

Protein (GFP)

BHK-T7 cells were seeded at densities of 0.5, 1, and 2 x 105 cells per well of a 6- well plate. Cells were transfected with plasmids encoding for each of the 11 RV genome segments, the FAST protein, and the AHSV capping enzyme as described above, in addition to 1 µL (0.8 µg) of a GFP-encoding plasmid driven by a T7 promoter. 48 hours post-transfection, cells were imaged using the Echo Revolve4 Microscope (VWR) with assistance from Dr. Heather Brown-Harding.

Growth of High-Titer Viral Stocks

To harvest enough virus for use in single-cycle growth assays, high-titer stocks of each recombinant RV (rSA11 or rSA11-g11-Wa) were grown by serial passage in MA104 cells as described previously by Arnold et al., 2009. Briefly, for each virus, a confluent T-

150 flask of MA104 cells was infected with 100 µL of the passage 1 (P1) stock for a P2 stock. P2 virus was harvested by lysing cells with 3 rounds of freeze-thaw. 1 mL of clarified supernatant from each P2 stock was then used to infect each of 4 confluent T-

150 flasks of MA104 cells for a P3 stock, yielding 100 mL of each virus. Stocks were titered by plaque assays as described above.

Quantification of Viral Growth

When MA104 cells in 6-well plates reached confluence, 6 wells of an additional plate were counted using an automatic cell counter; these counts were averaged and used to determine PFUs of each virus (wildtype or rSA11-g11-Wa) needed for infection of cells at a multiplicity of infection (MOI) of 5 (Arnold et al., 2009). Following viral activation

34 and infection with P3 virus as described above, cells were washed with complete M199, and media was replaced with complete M199 to neutralize trypsin left over from the infection and prevent further viral entry into new cells. Viruses and cells were harvested at

0, 8, and 16 hours post-infection (h.p.i.) by freezing. Virus was harvested by lysing cells with 3 rounds of freeze-thaw, and viral titers in each clarified lysate were determined via plaque assay as described above. This experiment was carried out in technical duplicate and was repeated 4 times.

All comparisons were analyzed using a one-tailed Student’s t-test in Microsoft

Excel. Outliers were determined and excluded from the analysis using a Dixon’s Q test in

Microsoft Excel.

Immunobloting

Infections of MA104 cells with rSA11-wt or reassortant (rSA11-g11-Wa) were carried out at an MOI of 5 as described above. Mock- and virus-infected cells to be harvested at 0 h.p.i. were washed once with 1X PBS and lysed with 500 µL of cold RIPA buffer [10 mM Tris, pH 8; 1% Triton X-100; 0.1% sodium deoxycholate; 150 mM sodium chloride (NaCl); and 1 Pierce Protease Inhibitor Tablet Mini (Roche)], and the lysate was placed a -20°C. Cells to be harvested at 8 h.p.i. were incubated in complete M199 at 37°C and 5% CO2 for 8 hours prior to being lysed via freezing.

Lysate was thawed on ice and centrifuged at 16,873 x g for 1 minute to pellet cell debris. Proteins contained in 15 µL of lysate were denatured 1:1 in 2X Laemmli Sample

Buffer (Bio-Rad) with 5% 2-mercaptoethnaol at 95°C for 5 minutes. Proteins were separated on a 4-15% sodium dodecyl sulfate (SDS)-polyacrylamide gel (Bio-Rad) at 100

V for 5 minutes, then 135 V for 70 minutes. Proteins were transferred to nitrocellulose

35 membranes at 100 V for 2 hours in cold transfer buffer (25 mM Tris, 190 mM glycine,

20% methanol, and 0.1% SDS). Membranes were blocked in TBST (20 mM Tris, pH

7.5; 150 mM NaCl; and 0.1% Tween-20) with 5% dry, non-fat Carnation milk at room temperature for 1 hour on a platform rocker. Primary guinea pig antibodies against VP6 and VP1/2 were used at 1:1000 in TBST with 5% milk, and a primary antibody against

NSP5 was used at 1:1000 in TBST with 3% bovine serum albumin and incubated with respective membranes at 4°C overnight on a platform rocker. Membranes were washed

5 x 7 minutes with TBST at room temperature on a platform rocker. An anti-guinea pig horseradish peroxidase-conjugated secondary antibody was used at 1:10,000 in TBST with 5% milk and incubated with membranes at room temperature for 1 hour on a platform rocker. Membranes were again washed 5 x 7 minutes with TBST at room temperature on a platform rocker prior to a 5-minute incubation with 1:1 mixtures of

Pierce ECL Detection Reagents 1 Peroxide Solution and 2 Luminol Enhancer Solution

(Invitrogen) and subsequent imaging using an Amersham Imager 600 (GE Healthcare).

Sequencing of NSP2 and NSP5 cDNAs

Following RNA extraction from wildtype and rSA11-g11-Wa-infected MA104 cell lysates using TRIzol as described above, NSP5 and NSP2 cDNAs were generated via RT-

PCR using SuperScript OneStep RT-PCR System with Platinum Taq DNA polymerase according to the manufacturer’s protocol (Invitrogen) using primer sequences conserved between SA11 and Wa (Table 3). cDNAs were gel purified (Invitrogen) and ligated into pGEM® T Vectors (Promega) via T4 DNA ligase (NEB) at 3:1 molar ratios. Following transformation in E. coli as described above, cells were plated on medium containing LB base, Miller (Sigma-Aldrich), bacteriological agar (Sigma-Aldrich), 100 µg/mL of

36 ampicillin, and 100 µL ChromoMax™ IPTG/X-Gal Solution (Invitrogen) and grown for

16 hours at 37°C. DNA was miniprepped (Invitrogen) from white colonies and sequences were determined via Sanger sequencing as described above.

Table 3. Primer sequences for NSP5 and NSP2 compensatory mutation screen

Primer Sequence

NSP5 Forward GCGCTACAGTGATGTCTCTC

NSP5 Reverse GTGGGGAGCTCCCTAGTG

NSP2 Forward GGCTTTTAAAGCGTCTCAGTC

NSP2 Reverse CATAAGCGCTTTCTATTCTTGC

Nucleotide Alignments

Using Geneious Pro v10.2.3 software, H1 and H5 nucleotide sequences were pairwise aligned using an IUB cost matrix.

RNA Folding Analysis

Predicted RNA secondary folding structures were determined using the Geneious

Pro v10.2.3 RNA fold tool by applying the Turner free-energy model at a simulated temperature of 37°C (Matthews et al., 2004).

Amino Acid Alignments

Wa and SA11 NSP5 (H1 and H5, respectively) amino acid sequences were compared both to each other and to other mammalian H genotypes using Geneious Pro v10.2.3 software. For the H1 and H5 alignment, respective RNA sequences were trimmed to NSP5 and NSP6 ORF sequences and translated. Translated sequences were pairwise aligned using the BLOSUM62 cost matrix. For the intergenotypic analysis, representative amino acid sequences were chosen for each available NSP5 genotype from the NCBI’s

Virus Variation RV database. For human RVA H-types, strains Wa, DS-1, and AU-1 were chosen to represent H1-3, respectively. SA11 H-96 was chosen to represent H5. Amino acid sequences were aligned using the BLOSUM62 cost matrix.

RESULTS

The laboratory strain P is genetically similar to modern, Wa-like RVAs

Human RVAs are difficult to cultivate from fecal specimens, but several Wa-like strains have been adapted to grow in culture (strains Wa, P, D, and KU). To choose a culture-adapted strain that would form the genetic basis of a reverse genetics system for a

Wa-like human RVA, a phylogenetic approach was employed to determine which of the four aforementioned culture-adapted strains is most genetically related to current RVAs that circulate within the human population. Neighbor-joining phylogenies limited to Wa- like genotype constellation G1- or G3-P[8]-I1-R1-C1-M1-A1-N1-T1-E1-H1 were generated for each genome segment. Included in each phylogeny were archival sequences of culture-adapted Wa-like strains Wa, P, D, and KU, in addition to an archival DS-1 sequence to serve as an outgroup. RVA sequences isolated from patients in 2010 or later were considered modern.

Phylogenies generated using VP2, VP3, VP4, VP6, NSP2, and NSP5 sequences showed that the sequence for archival strain P tended to cluster in branches containing more modern RVA sequences than the sequence for archival strain Wa. While the majority of the phylogenies demonstrated that P more closely resembles sequences of modern

RVAs, in some of these phylogenies, P clusters with archival strains but still lies within a larger monophyletic grouping with modern strains. The phylogeny generated for VP4 serves as an example of this phenomenon (Figure 8). A comparison of the phylogenies generated using VP7-G1 and VP7-G3 sequences also showed that P clustered with more modern strains than did Wa. In phylogenies generated using NSP3 and NSP4 sequences, archival strains clustered with each other and not with modern strains. In one phylogeny generated using VP1 sequences, the archival Wa strain sequence clustered with more

39 modern sequences than did P, and in another phylogeny generated using NSP1 sequences, the archival D strain sequence clustered with more modern sequences than did P or Wa.

Interestingly, in none of the phylogenies did KU cluster with more modern strains than P,

Wa, or D. These results are summarized in Table 4.

Table 4. Wa-like RVA strains that cluster with modern strains

VP1 VP2 VP3 VP4 VP6 VP7 NSP1 NSP2 NSP3 NSP4 NSP5 Wa P P P P P D P None None P

Figure 8. Phylogeny of archival and modern RVA VP4 genotype sequences

Complete archival sequences for culture-adapted RVA strains Wa, P, D, KU (Wa-like strains), and DS-1 (outgroup) were obtained using the NCBI’s nucleotide database for each of the Wa-like and DS-1 genotypes, with P1 shown as an example in this figure. The

NCBI’s Virus Variation RV database was used to obtain 147 complete P1 sequences that were isolated from RVA-infected humans in various geographic regions from 2010 to present. Geneious software was used generate sequence alignments that were trimmed to boundaries of shared sequence. A final consensus tree was created using Geneious, and

FigTree was used to collapse monophyletic branches. Archival sequences of culture adapted strains are colored: red (DS-1), yellow (KU), green (Wa), blue (P), and purple (D).

Black denotes clusters containing archival sequences, and white denotes clusters containing modern sequences.

Strain Wa grows to a higher titer than P in MA104 cell culture

In terms of public health relevance, the Wa-like culture-adapted strain P provided a likely candidate for use in a reverse genetics system for human RV because it more closely resembled modern human RVA strains. However, to produce a robust reverse genetics system capable of reliable recombinant virus rescue, the ability of a culture- adapted virus to propagate in culture must be considered. Following passage of strain P in a T-150 flask of MA104 cells, its titer was calculated to be 2 x 103 PFU/mL, almost two logarithmic units lower than Wa, titered at 1.8 x 105 PFU/mL. Moreover, the plaques produced by strain P developed 4 days slower than did those produced by strain Wa.

Therefore, even though it was not as close a genetic match as was strain P, Wa was chosen for the generation of clones and the development of reverse genetics because of its improved growth in culture relative to strain P.

BHK-T7 cells are transfectable and express T7 RNA polymerase

The reverse genetics system developed by Kanai, et al. requires transfection of 14 plasmids into a single cell (2017). While the plasmid burden was reduced to 13 by replacing two vaccinia virus capping enzyme constructs (D1R and D12L) with just one African horse sickness virus construct (NP868R), successfully introducing 13 plasmids into one cell poses considerable technical challenges. When multiple attempts using the original protocol described in Kanai, et al. failed to produce a wildtype recombinant SA11 H-96 virus in the McDonald lab, the transfectability of the BHK-T7 cells was assessed first

(2017). Transfecting BHK-T7 cells with a plasmid encoding GFP under the control of a T7 promoter served the dual purpose of assessing the cells’ ability to uptake plasmid and

42 express messages and proteins encoded by the plasmids using a functional T7 RNA polymerase.

BHK-T7 cells were seeded at densities of 0.5, 1, and 2 x 105 cells per well of a 6- well plate. 48 hours later, cells were transfected under the same conditions as used for reverse genetics experiments, including 1 µL (0.8 µg) of a plasmid encoding green fluorescent protein (GFP). Images were taken 48 hours post-transfection and visualized for the expression of GFP. Green cells could be visualized under each condition (Figure

9). This experiment revealed that cells were able to successfully uptake plasmid and still expressed T7 polymerase.

Figure 9. BHK-T7 cells can uptake and express a plasmid encoding GFP

Various densities of BHK-T7 cells were transfected with a plasmid expressing GFP under the control of a T7 promoter and imaged 48 hours post-transfection for GFP expression.

Images were generated by Taylor Shue, with assistance from Dr. Heather Brown-Harding.

MA104 cells are permissive to RV infection

In addition to assessing the functional role of BHK-T7 cells in the all plasmid-based reverse genetics system, the ability of RV to propagate in MA104 cells was tested. Upon reaching confluence, MA104 cells were infected with human RV strain Wa at an MOI of

0.5 or 2. Wa was used instead of SA11 for this experiment with the rationale that use of wildtype SA11 in the cell culture hood could potentially lead to later contamination of

MA104 cells with this robust virus. Both MOIs produced CPE 3 days post-infection, indicating that the MA104 cells were still permissive to RV infection.

Modifications to the existing all plasmid-based reverse genetics system yielded successful rescue of wildtype rRV SA11

Previous iterations of the all plasmid-based reverse genetics protocol seeded higher densities of BHK-T7 cells (2 and 3 x 105 cell per well), but it was observed that CPE of

MA104 cells infected with virus harvested from lower densities of BHK-T7 cells could be visualized up to 48 hours before CPE of MA104 cells infected with virus harvested from higher densities of BHK-T7 cells. Additionally, work in the Patton lab demonstrated that the addition of tryptose broth to the BHK-T7 cells’ media enhanced rescue of recombinant wildtype SA11 H-96 from the system (personal communication). Personal communication with Dr. Patton revealed that the BHK-T7 cells stocks given to the McDonald lab were contaminated with mycoplasma. Together, the reduction of the seeding density of BHK-

T7 cells to 0.5 and 1 x 105, addition of tryptose broth to complete BHK-T7 cell media, and use of fresh stocks of BHK-T7 cells resulted in visualization of CPE in MA104 cells.

To determine whether observed CPE was caused by rRV SA11 generated by the reverse genetics system, viral RNAs were TRIzol-extracted from freeze-thawed MA104-

45 infected cell lysate generated by the reverse genetics experiment. Primers complementary to a region of VP4 from SA11-H-96 (strain used in reverse genetics system) and putative

VP4 reassortant SA11 variant, 4F, were used for RT-PCR amplification of viral RNAs

(Small et al., 2007). Presence of SA11-4F was tested with the rationale that observed CPE could potentially have been a result of contamination from SA11-4F, the “wildtype” strain used by the McDonald lab. Reactions containing SA11 H-96 primers and RNA extracted from the reverse genetics experiment produced a 322 base pair (b.p.) product, corresponding to amplification of SA11-H-96 that was detectable in all three experimental conditions, as well as one control condition. SA11 H-96 primers tested with SA11 4F RNA produced no detectable band, indicating that SA11 H-96 primers did not amplify SA11 4F

RNA. An RT-PCR control consisting of primers and RNA used and amplified previously produced a band of expected size (1,079 b.p.) (Figure 10A). Reactions containing SA11

4F-specific primers and RNA extracted from the reverse genetics experiment produced no detectable 241 b.p. fragments, which would correspond to SA11-4F amplification, but the reaction containing SA11-4F RNA and SA11-4F specific primers produced a 241 b.p. product, indicating that 4F primers successfully amplify SA11-4F RNA (Figure 10B).

Altogether, these results suggest that rRV SA11-H-96 was successfully rescued using the modified reverse genetics system.

Figure 10. RT-PCR validation of successful wildtype rSA11 RVA rescue using all plasmid-based reverse genetics system

Viral RNAs were extracted from MA104 control- and virus-infected cell lysate, and SA11-

H-96 (A) and -4F (B) specific primers were used to amplify respective 322 b.p. or 241 b.p. regions of the VP4 genome segment via RT-PCR. Control wells represent RNA extracted from MA104 cells infected with clarified supernatant from BHK-T7 cells that were transfected with solutions excluding the NSP2 construct, which is vital for RV replication and viroplasm formation.

Following two successful recombinant wildtype SA11 H-96 rescues using the three aforementioned modifications, two subsequent attempts to rescue recombinant wildtype

SA11 H-96 in parallel with mono-reassortant and mutant rRVs failed. In an effort to further troubleshoot the reverse genetics system, Minimum Essential Medium-Non-Essential

Amino Acids Solution (MEM-NEAA) (Gibco) was added to supplement BHK-T7 cell media after personal communication with Dr. Patton revealed that this modification enhanced rescue of rRVs from reverse genetics experiments in his lab. BHK-T7 cells were thawed from liquid nitrogen stocks and grown for two passages in complete DMEM supplemented to contain MEM-NEAA; however, a reverse genetics experiment using these cells failed to produce CPE in MA104 cells.

In the existing protocol, just prior to overseeding MA104 cells, serum-free medium is removed from the transfected BHK-T7 cells. It was postulated that this medium may already contain viral particles that would be removed with this step, reducing the number of virus particles available to infect the MA104 cells. To test this idea, BHK-T7 cells were seeded at densities of 0.5 and 1 x 105 cells per well, and MA104 cells were either overseeded at 105 cells per well by replacing the old media (adhering to the original protocol) or by overseeding either 105 or 2 x 105 cells in a total volume of 250 µL directly to the serum-free media. Four days post infection of a new monolayer of MA104 cells, no

CPE was observed with the lower BHK-T7 cell seeding density under any condition tested.

However, with the higher BHK-T7 cell seeding density, CPE was observed when the old media was maintained and a higher number of MA104 was overseeded (Table 5).

Table 5. MA104 cell overseed experiment results

Old media Old media Old media removed, maintained, maintained, 105 MA104 cells 105 MA104 cells 2 x 105 MA104 overseeded overseeded cells overseeded 0.5 x 105 BHK-T7 No CPE No CPE No CPE cell seeding density 105 BHK-T7 cell No CPE No CPE CPE seeding density

Six Wa genome segments were successfully cloned into reverse genetics vectors

The second aim of this work was to develop a reverse genetics system for human

RV strain, Wa. To reconstitute the existing reverse genetics system for human RV Wa,

SA11 genome segments within reverse genetics vectors would need to be replaced with those of Wa.

First, Wa NSP5 was successfully amplified via RT-PCR, and the pT7 reverse genetics vector was successfully amplified via PCR. Ultimately, Sanger sequencing revealed the insertion of the Wa NSP5 gene was identical to that of the authentic human

RV Wa NSP5. In addition to cloning Wa NSP5 into its reverse genetics vector, Wa NSP4,

NSP3, NSP2, NSP1, and VP6 were successfully cloned into reverse genetics vectors. The

VP6 genome segment had two mutations at b.p. positions 180 and 335, and the NSP2 genome segment had one mutation at b.p. position 164 relative to known Wa sequence.

While Wa VP7 and VP3 were successfully amplified via RT-PCR, multiple attempts at inserting these genome segments into reverse genetics vectors failed, producing what appeared to be empty vectors when analyzing Sanger sequencing results. Wa VP2 and VP1 were also successfully amplified via RT-PCR, but the primers used to amplify the pT7 reverse genetics vector with sequence complementarity to these genome segments were unsuccessful.

rSA11-g11-Wa exhibits a modest replicative defect relative to rSA11-wt

While troubleshooting the reverse genetics protocol in the McDonald lab, the reverse genetics vector containing Wa NSP5 was sent to the Patton lab. They were able to successfully rescue an NSP5-Wa mono-reassortant virus, rSA11-g11-Wa, along with a wildtype control, rSA11-wt.

To assess the replicative capacity of rSA11-g11-Wa relative to rSA11-wt, high-titer stocks of each virus were grown by serial passage in T-150 flasks and titered. MA104 cells were infected with each virus at an MOI of 5 for 1 hour. Following infection, inoculant was replaced with media containing FBS to neutralize trypsin, preventing further viral entry into cells. In this way, this experiment measures how many progeny one virion can make just by replicating inside of a cell. Virus was harvested at 0, 8, and 16 h.p.i. at -20°C, and viral titers were determined by plaque assay. The growth experiment demonstrated a modest replicative defect exhibited by rSA11-g11-Wa relative to rSA11-wt (Figure 11A).

This defect is further reflected by a statistically significant difference in peak viral titers reached by rSA11-g11-Wa relative to rSA11-wt (Figure 11B).

Figure 11. Growth comparison of rSA11-wt and rSA11-g11-Wa during one cycle of infection

(A) Virus was harvested at 0, 8, and 16 h.p.i., and viral titers were determined via plaque assay. Average titers were plotted on a logarithmic scale, and error bars represent standard deviation. (B) Peak viral titers were compared and determined to be statistically significant

(p < 0.01) by a one-tailed Student’s t-test.

Wa and SA11 g11 share 88% nucleotide identity

The observed difference in growth of rSA11-g11-Wa relative to rSA11-wt could reflect differences in nucleotide or amino acid residues important for interaction with other viral RNAs or proteins. To examine nucleotide differences between Wa and SA11 g11, the nucleotide sequences of Wa and SA11 genome segment 11 were compared in a pairwise alignment. In total, there are 79 nucleotide differences between Wa and SA11 genome segment 11, as well as 3 nucleotide deletions in the Wa sequence relative to SA11 (Figure

12).

Figure 12. Pairwise nucleotide alignments of Wa and SA11 genome segment 11

RNA sequences of Wa and SA11 genome segment 11 were compared in a pairwise alignment. Nucleotide differences are shown in white, and shared sequence in gray. 5 and 3’ UTRs are labeled.

Wa and SA11 NSP5 +RNAs are predicted to form different secondary structures

Because Wa and SA11 g11 +RNA sequences differ by 82 nucleotides total, it was postulated that differences in nucleotide identity between Wa and SA11 genome segment

11 RNAs might have contributed to the phenotypic difference observed in rSA11-g11-Wa relative to rSA11-wt. Differences at the +RNA level that could be important to viral replication could be observed in terms of differential secondary folding structures. Using

Geneious software, RNA folding structures of independent Wa and SA11 +RNAs can be predicted. While the 5’ and 3’ ends of the RNAs appear to adopt similar stem structures, the stem and loop structures formed by Wa and SA11 NSP5’s ORFs appear to differ in structure (Figure 13). Interestingly, the free energy changes between optimal Wa and SA11 g11 +RNA structures are comparable with values of -158.50 and -159.50 kcal/mol, respectively, but the ensemble diversities differ widely with values of 143.43 and 106.09, respectively; this result indicates that Wa’s g11 +RNA sequence has the capacity to assume a more diverse set of secondary structures than SA11 g11. Together, these results raise questions about the stem-loop structures formed by different RVA RNAs and whether they serve functionally significant roles in RVA RNA assortment.

3’ UTR

Wa g11

5’ UTR

3’ UTR

SA11 g11

5’ UTR

Figure 13. Predicted secondary structures of Wa and SA11 g11 +RNAs

The predicted secondary structures of Wa (top) and SA11 (bottom) +RNAs that encode

NSP5 were determined using Geneious software; bases colored according to base-pair probabilities, wherein warmer colors represent greater favorability of interaction, and cooler colors represent lesser favorability of interaction.

Wa and SA11 NSP5 share 87% amino acid identity

While the observed phenotypic difference between rSA11-g11-Wa and rSA11-wt could reflect differences in nucleotide identity and +RNA secondary structure formation, it could also reflect differences in amino acid identity and protein structure. To investigate amino acid differences between Wa and SA11 NSP5 and NSP6, respective genome segment 11 RNA sequences were trimmed to ORF length and translated. Translated sequences were pairwise aligned. Comparing the NSP5 amino acid sequences, there are 25 amino acid differences between Wa and SA11, as well as 1 amino acid deletion in the Wa sequence relative to SA11 (Figure 14A). In contrast, there are 11 amino acid differences between Wa and SA11 NSP6 proteins (Figure 14B).

SA11-H5 Wa-H1

Figure 14. Pairwise amino acid alignments of Wa and SA11 NSP5 and NSP6

Amino acid sequences of Wa and SA11 NSP5 (A) and NSP6 (B) were compared in a pairwise alignment. Amino acid differences are shown in white, and shared sequence in gray.

Intergenotypic variation among NSP5 amino acid sequences lies in important functional regions

While the pairwise alignment comparing Wa and SA11 (H1 and H5, respectively) NSP5 amino acid sequences directly was useful to determine which residues differ between the two proteins, it was thought that a multiple alignment comparing amino acid sequences of different H genotypes could yield insight into the relative conservation of amino acid residues at the sites of divergence between H5 and

H1. To evaluate the differences between SA11 and Wa NSP5 on an amino acid level in the context of other NSP5-coding genotypes, a mammalian amino acid sequence representative of each genotype was downloaded from the NCBI’s Virus Variation RV database, and these sequences were aligned using Geneious software and a BLOSUM62 cost matrix. There are 2 amino acid differences unique to SA11 NSP5: Pro14 and Thr16.

At both of these positions in the remainder of the sequences are highly conserved S residues. In contrast, there are 3 amino acid differences unique to Wa NSP5: Asn37,

Cys125, and Glu163. At positions 37 and 125 in the remainder of the amino acid sequences is a highly conserved S residue. In the Wa amino acid sequence, Tyr162 substitutes a relatively conserved S residue that serves as a phosphorylation site in other sequences

(Figure 15). The only difference unique to both SA11 and Wa NSP5 is Arg155, which contrasts to a relatively conserved K in the remainder of the amino acid sequences. 13 of the 25 of the amino acid changes between SA11 and Wa NSP5 map to an NSP5 dimerization region—residues 103-146. Interestingly, this functional region also appears to contain the majority of the sequence variation (Figure 15). There are no differences in

SA11 or Wa amino acid sequences relative to the consensus sequence in the region important for NSP5 decamerization (Figure 15).

Figure 15. Intergenotypic variation among NSP5 amino acid sequences

A representative amino acid sequence from each genotype was selected from the NCBI

Virus Variation rotavirus database and aligned using the BLOSSUM62 cost matrix in

Geneious software. SA11 and Wa are represented as H5 and H1, respectively. Percent identity (top) is represented by a colorized graph in which green bars represent residues with high intergenotypic conservation, yellow bars represent residues with intermediate conservation, and red bars represent high intergenotypic variation. Non-consensus sequence is shown in white, and consensus sequence is shown in gray. Dashes indicate gaps in protein sequence. Arrows represent sites that differ between SA11 and Wa NSP5 amino acid sequences. Asterisks represent conserved phosphorylation sites.

Western blot of Wa and SA11 NSP5 reveals differential banding patterns at 8 h.p.i.

In effort to investigate a potential cause for the observed replicative defect of rSA11-g11-Wa relative to wildtype rSA11-wt, a Western blot was performed using mock-

, rSA11-g11-Wa-, or rSA11-wt-virus-infected MA104 cell lysate harvested at 0 and 8 h.p.i.

Proteins were separated by SDS-PAGE and transferred to a nitrocellulose membrane. The blot was probed with a-VP1/2 or a-VP6 antibodies to verify equivalent expression of viral proteins. An a-NSP5 antibody was used to blot for expression SA11 and Wa NSP5. Wa

NSP5 demonstrated a differential banding pattern when compared to wildtype rSA11

(Figure 11). This result is consistent with previous experiments demonstrating phosphorylation patterns of NSP5, which suggests that Wa NSP5 could be differentially phosphorylated relative to SA11 (Afrikanova et al., 1996; Blackhall, et al., 1998).

Figure 16. Western blot of Wa and SA11 NSP5 reveals differential banding patterns

Western blotting was performed on mock-, rSA11-wt, and rSA11-g11-Wa-infected cell lysate extracts at 0 and 8 h.p.i. a-VP1/2 and -VP6 antibodies were used as controls, and an a-NSP5 antibody was used to detect NSP5 from rSA11-wt and rSA11-g11-Wa.

Sequencing of NSP5 and NSP2 reveal no compensatory mutations

Because both the wildtype and reassortant virus underwent serial passage in

MA104 cells to generate high-titer stocks for use in single-cycle growth assays, it was thought that compensatory mutations could have arisen in the reassortant virus. Such mutations could be informative of nucleotide or amino acid residues important for RNA or protein contacts made during the viral replication cycle. Because NSP5’s function depends on its interaction with NSP2, viral RNAs were extracted from rSA11-g11-Wa- or wildtype rSA11-infected MA104 cell lysate and amplified via RT-PCR using primers complementary to conserved regions of both NSP5 and NSP2. Genome segments were

TA-cloned and sequenced for compensatory mutations. No mutations were detected in

NSP5 or NSP2 genome segments amplified from either virus, meaning that NSP5- and

NSP2-coding RNAs extracted from the reassortant virus were identical to the original Wa reverse genetics plasmids (data not shown).

DISCUSSION

The main objectives of this research were to: (1) successfully rescue wildtype rRVA SA11 using the existing reverse genetics system, (2) develop a reverse genetics system for a Wa-like human RVA, and (3) utilize the existing reverse genetics system to rescue and test the replicative capacity of a mono-reassortant rSA11 RVA containing the

NSP5-coding genome segment human strain, Wa. The second and third goals were in part dependent on the first, as developing a functional protocol for the reliable rescue of wildtype rSA11 RVAs proved challenging. Multiple attempts adhering to the protocol described in Kanai, et al. failed to produce wildtype rRVAs (2017). Initial troubleshooting experiments involved testing functionality of the cell lines utilized by the system. First, the ability of BHK-T7 cells to successfully transfect and express protein from plasmid DNA was tested by introducing a GFP-encoding plasmid under the control of a T7 promoter.

Results showed that the cells were not only capable of successful transfection, but they were also still expressing T7 RNA polymerase. BHK-T7 cell densities were too high to count at the time of imaging, so transfection efficiency could not be quantified; however, transfection efficiency of 13 plasmids could not be inferred from measured transfection efficiency of one plasmid. Instead, time required for observation of CPE in rRVA-infected

MA104 cells was used as a proxy for assessing transfection efficiency of BHK-T7 cells.

Later experiments showed that MA104 cells infected with lysate harvested from the lowest

BHK-T7 cell seeding density displayed total CPE only 24 h.p.i., in contrast to MA104 cells infected with lysate harvested from the intermediate and high seeding densities that displayed CPE 72 h.p.i. From this observation, one could infer that a larger number of infectious virus particles were initially generated from the BHK-T7 cells seeded at lower densities, thus resulting in more efficient propagation in MA104 cells.

In addition to lowering the seeding density of BHK-T7 cells, two additional alterations were made to the existing protocol; together, both modifications resulted in the successful rescue of a recombinant wildtype rRVA SA11. BHK-T7 cell stocks gifted for use in reverse genetics experiments were discovered to be contaminated with mycoplasma.

Mycoplasma has been shown to affect cultured cells’ metabolism, including DNA, RNA, and protein production (Barile, 2014). Use of fresh, mycoplasma-free BHK-T7 cells likely enhanced the cells’ ability to transcribe multiple plasmids and translate viral proteins, which could have resulted in the production of more infectious virus particles by the cells.

Additionally, tryptose broth was added to the BHK-T7 cells’ growth medium as a supplementary source of nutrients and growth factors, which may have also served to enhance transcription and translation in these cells. When the BHK-T7 cell media was also supplemented to contain MEM-NEAA, the system, again, failed to produce detectable wildtype rRVA SA11. While consistently diluting the cells 1:20 with each passage, cells propagated in medium containing MEM-NEAA reached confluence a day later than expected. In addition, when these cells were used in reverse genetics experiments, they appeared sparse just prior to transfection. It is possible that the MEM-NEAA slowed their growth, and there appears to be a density threshold necessary for successful expression of rRVAs by the BHK-T7 cells. Future experiments should seek to define not only an optimal

BHK-T7 cell seeding density, but also an optimal density for transfection and generation of rRVAs.

Steps of the reverse genetics protocol following BHK-T7 cell seeding and transfection also require further optimization. 48 hours post-transfection, visually, BHK-

T7 cells exhibit a shrunken monolayer, with several dead cells floating in the media.

While this observation could be an effect of lipofectamine treatment, it could also be CPE

65 from the generation of rRVAs. RV SA11-H-96 entry into cultured cells is dependent on trypsin, and trypsin is thought to be necessary for viral egress in some cell lines. By removing media containing dead BHK-T7 cells prior to MA104 overseed and the addition of trypsin, it is possible that virus particles trapped within cells or adhered to cellular membranes were also discarded. This idea was supported by the results of the experiment testing different MA104 overseed conditions, which showed that CPE observed was limited to the condition in which BHK-T7 cell media was maintained and

MA104 cells were overseeded at the highest density tested. To test whether the number of MA104 cells overseeded has an effect on successful rescue of rRVAs, this experiment should be repeated to include various densities in both old and new media. Still, in this experiment, conditions that yielded successful rescue of rRVAs previously failed to produce the same outcomes. Further optimization of the system would benefit from determining sources of stochasticity within the method.

Because the all plasmid reverse genetics system was failing to produce recombinant viruses for several months, the Patton lab kindly rescued the rSA11-g11-Wa reassortant virus. Successful rescue of the virus suggests that nucleotide and amino acid differences between SA11 H5 and Wa H1 were not sufficient to ablate RNA-RNA, RNA-protein, and protein-protein interactions critical for reassortment or nascent virion assembly. More specifically, these results indicate that SA11 VP1 is capable of recognizing the H5 dsRNA segment for transcription and its +RNA for dsRNA synthesis. While the precise order of events that de novo virion assembly follows is unclear, recombinant virions produced by

BHK-T7 cells must replicate within the MA104 cells successfully to produce CPE. This would suggest that SA11 VP3 is also capable of recognizing H5 +RNA for capping. To form a functional viroplasm capable of producing RV progeny, NSP5 must be able to

66 interact with NSP2. Again, the rescue of rSA11-g11-Wa suggests that the putative binding interface between SA11 NPS2 and Wa NSP5, in any conformation, was not disrupted by differences in the amino acid residues between SA11 and Wa NSP5. NSP5 is also responsible for RNA binding and postulated to play an important role in assortment of viral

RNAs. The ability of rSA11-g11-Wa to replicate indicates that Wa NSP5 is capable of recognizing SA11 +RNAs for selective packaging into nascent virions. Overall, this result suggests that nucleotide and amino acid changes between H5 and H1 do not contain biochemical determinants for reassortment restriction.

Nevertheless, it was still possible that rSA11-g11-Wa harbored a replicative defect relative to wildtype SA11. To test this possibility, high-titer stocks of rSA11 and rSA11- g11-Wa were competed in a single-cycle growth assay that revealed a modest replicative defect in the reassortant virus relative to wildtype. While the 82 total nucleotide differences between H5 and H1 do not preclude selective packaging of H1 by the SA11 virus, it is possible that these differences affected the overall ability of the rSA11-g11-Wa to replicate.

While the RNA folding analysis revealed comparable stem structures between H5 and H1

+RNA UTRs, loop structures found in the ORFs differ. The role of stem-loop structure motifs in +RNA assortment has not been widely examined, but in silico analysis has revealed a potential functional purpose (Borodavka et al., 2017). It is possible that differences in H5 and H1 +RNA stem-loop structures stalled the assortment process.

Testing this hypothesis, however, would require molecular dynamics simulations that can predict these secondary structures in the context of other viral +RNAs. More specifically, it is possible that SA11 NSP2 does not readily engage g11-Wa, delaying the formation of a supramolecular complex. RNA-RNA interactions can influence secondary structure, and secondary structure predictions of H5 and H1 in this investigation are limited to

67 independent folding analyses; these predicted structures also did not contain free extreme

3’ termini, which have been shown to be critical for recognition by VP1 (Chen and Patton,

1998). Future experiments could assess the compatibility of H1 +RNA with SA11 VP1 and

VP2 using an in vitro replicase assay. Since the N-terminal domain of VP2 is hypothesized to bind +RNAs and bring them in proximity to VP1 for dsRNA synthesis, this experiment could reveal whether sequence or stem-loop structural differences between H5 and H1 stall recognition by VP2.

On a protein-level, 26 amino acid residues differed between SA11 and Wa NSP5, including 1 deletion in the Wa sequence relative to that of SA11. While no high-resolution structure of the NSP5 protein exists, various studies have linked important regions to function in the RV lifecycle and circular dichroism has revealed that the protein is highly disordered (Martin et al., 2011). 9 of the amino acid changes between SA11 and Wa NSP5 are located in the N-terminal region of the protein—comprised of residues 1-98—which is the protein’s most disordered region (Martin et al., 2011). Two of these changes include serine to proline and vice versa—two residues known to disrupt helices. The majority of the amino acid changes between SA11 and Wa NSP5 map to an NSP5 dimerization region—residues 103-146—which harbors 13 amino acid changes. To test whether the differences in this region between SA11 and Wa NSP5 affected the growth phenotype of rSA11-g11-Wa, a gain-of-function approach could be employed. Specifically, a recombinant RV containing a chimeric NSP5 protein with SA11 amino acids 103-146 flanked by NSP5 sequence could be rescued using reverse genetics and tested for its replicative capacity in a single cycle growth assay. If the replicative capacity of this virus is restored relative to wildtype, it would indicate that the changes in this region contributed to the replicative defect demonstrated by rSA11-g11-Wa. Interestingly, the intergenotypic

68 amino acid alignment also revealed that the majority of the intergenotypic variation in

NSP5 amino acid sequences lies within this region (Figure 16). Martin et al. also mapped the site of specific interaction with NSP2 to amino acid residues 132-146 (2011). In this region, there are 5 amino acid differences between SA11 and Wa NSP5. It would be intriguing to test the replicative phenotype of a double-reassortant RV containing Wa NSP5 and NSP2 in an SA11 genetic background in a single-cycle growth assay. If the replicative capacity of this double-reassortant virus is restored relative to wildtype, it would indicate that the changes in this region contributed to the replicative defect demonstrated by rSA11- g11-Wa. The C-terminal region—residues 180-198—contains no differences between

SA11 and Wa NSP5 and is relatively conserved among other mammalian NSP5 amino acid sequences (Figure 16). Phenotypic differences between Wa and SA11 NSP5 could be the result of amino acid changes located in these regions and perhaps reduce Wa NSP5’s ability to function within the SA11 background.

Figure 17. NSP5 amino acid changes and important functional regions

Linearized protein schematic comparing SA11 and Wa NSP5 and the amino acid changes between them. Amino acids are colored according to polarity. A region known to be important for NSP5 dimerization contains 13 amino acid changes. The NSP2 binding region contains 5 amino acid changes. The dimerization region contains amino acids conserved between both H5 and H1 genotypes. “X” represents a deletion.

The replicative defect displayed by rSA11-g11-Wa could also stem from the differential banding pattern of the Wa NSP5 protein relative to SA11 NSP5 demonstrated by Western blot. Based on molecular weight calculations, Wa and SA11 NSP5 proteins differ by 0.1 kDa—a difference that would not be reflected by SDS-PAGE separation. The smear-like pattern exhibited by the blot of both proteins is consistent with previous experiments demonstrating phosphorylation patterns of NSP5 (Afrikanova et al., 1996;

Blackhall, et al., 1998). Thus, it is possible that the differential banding pattern exhibited by these proteins could be due to differential phosphorylation of Wa NSP5 in the context of SA11 NSP2. NSP5 Ser155 is a known phosphorylation site that is relatively conserved.

In place of a serine at this position is a tyrosine in Wa NSP5. Serine kinases constitute a different class of kinases than those capable of phosphorylating tyrosine residues. While one study demonstrated that SA11 NSP5 is still hyperphosphorylated when this position is mutated to an alanine, the loss of this site could explain the delay in phosphorylation of this reassortant virus (Eichwald et al., 2002). Since viroplasm formation is dependent on hyperphosphorylation of NSP5, this could account for the replicative defect exhibited by rSA11-g11-Wa. Phosphorylation of NSP5 has previously been studied using simian RV strains only; it is conceivable that Wa infections in human cells could usurp a different class of kinases, like tyrosine kinases.

Despite a clear replicative defect observed in the reassortant virus, both rSA11 and rSA11-g11-Wa underwent serial passage in MA104 cells prior to single cycle growth analysis. With each passage, mutations in the viral genome can not only occur but be selected for, and any mutations present following serial passage could be compensatory in nature and inform nucleotide or amino acid residues that interact with other viral RNAs proteins. Sanger sequencing of NSP5 and NSP2 cDNAs isolated from both rSA11 and

71 rSA11-g11-Wa revealed no mutations, suggesting that the replicative defect displayed by rSA11-g11-Wa could be representative of other potential RNA and protein interactions. It is also possible that compensatory mutations occurred in other genes.

Determining biochemical determinants restricting reassortment among RVAs would require (1) a further optimized reverse genetics protocol, (2) further experiments attempting to rescue mono-reassortant RVs using the existing system, and (3) the generation of a reverse genetics system for human RV. While a reverse genetics system for human RV strain KU was published during the preparation of this thesis, a Wa-based system may still be beneficial for the study of human RVs. Following the publication of the KU system, the phylogenetic analysis comparing culture-adapted and modern strains was redone including KU sequences. The analysis revealed that KU tended to cluster with other archival strains, meaning that it may not be as biologically relevant as other Wa-like

RV strains. Nevertheless, this recent development constitutes a significant advancement in the field of rotavirus biology and could provide a proof-of-concept platform for next generation vaccine design.

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APPENDIX

Table 7. Accession numbers of archival sequences for phylogenetic analysis

Wa P D KU DS-1 VP1 DQ490539 EF583037 EF583021 AB022765 EF990691 VP2 X14942 EF583038 EF583022 AB022766 EF990692 VP3 AY267335 EF583039 EF583023 AB022767 EF990693 VP4 L34161 EF672598 EF672570 AB222784 EF672577 VP6 KO2086 EF583040 EF583024 AB022768 EF990694 VP7-G1 M21843 X EF672574 D16343 X VP7-G2 X X X X EF672581 VP7-G3 X EF672602 X X X NSP1 L18943 EF672599 EF672571 AB022769 EF672578 NSP2 L04534 EF672601 EF672573 AB022770 EF672580 NSP3 X81434 EF672600 EF672572 AB022771 EF672579 NSP4 AF093199 EF672603 EF672575 AB022772 EF672582 NSP5/6 AF306494 EF672604 EF672576 AB022773 EF672583

CIRICULUM VITAE Jessica R. Hall

Department of Biology [email protected] Wake Forest University Wake Downtown 455 Vine Street Winston-Salem, NC 27101-4315

Education 2017-Present M.S., Biology Wake Forest University Graduate School of Arts and Sciences, Winston-Salem, NC “Reverse genetic strategies to interrogate reassortment restriction among group A rotaviruses” Anticipated August 2019

2013-2017 B.S., Biology Wake Forest University, Winston-Salem, NC Magna Cum Laude

Relevant Research Experience 2017-Present Graduate Research Student Lab of Dr. Sarah M. McDonald Wake Forest Graduate School of Arts and Sciences, Winston-Salem, NC - Optimized reverse genetics system for recovery of infectious simian rotavirus strain SA11 from transfection of 13 cDNA plasmids - Characterized mutant and reassortant rotaviruses recovered using reverse genetics system - Maintained MA104, BHK-T7, and COS-7 mammalian cell lines - Cloned human rotavirus genome segments into vectors for expression in reverse genetics system via RT-PCR amplification of viral genes - Mentored 1 postdoctoral fellow and 5 undergraduate students

2016 Intern Mentee of Drs. Nathan P. Coussens and Matthew D. Hall National Center for Advancing Translational Sciences, National Institutes of Health, Rockville, MD - Optimized a biophysical assay for high-throughput screening to identify inhibitors of a protein-protein interaction important for cancer metastasis

Publications Roy, NS; Jian, X; Soubias, O; Zhai, P; Hall, JR; Dagher, JN; Coussens, NP; Jenkins, LL; Hall, MD; Byrd, RA; Yohe, ME; and Randazzo, PA. The N-terminus of Arf1 interacts with the PH domain of the Arf GAP ASAP1 under control of phosphatidylinositol 4,5- bisphophate. Journal of Biological Chemistry. In review.

Technical Skills - Mammalian and bacterial cell culture - RNA and DNA isolation and analysis - Recombinant DNA techniques, such as RT-PCR, PCR, cloning, and mutagenesis - Protein isolation and analysis, including SDS-PAGE and Western blotting - Preparation of viral stocks - Plaque assays - High-throughput data collection using AlphaLISA to detect protein-protein interactions