Characterization of Bunyamwera, Batai, and Ngari Viruses: Unrecognized Arboviruses of One Health Importance in Rwanda Marie Fausta Dutuze Mdutuz1@Lsu.Edu

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July 2019 Characterization of Bunyamwera, Batai, and Ngari Viruses: Unrecognized Arboviruses of One Health Importance in Rwanda Marie Fausta Dutuze [email protected]

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Recommended Citation Dutuze, Marie Fausta, "Characterization of Bunyamwera, Batai, and Ngari Viruses: Unrecognized Arboviruses of One Health Importance in Rwanda" (2019). LSU Doctoral Dissertations. 4995. https://digitalcommons.lsu.edu/gradschool_dissertations/4995

This Dissertation is brought to you for free and open access by the Graduate School at LSU Digital Commons. It has been accepted for inclusion in LSU Doctoral Dissertations by an authorized graduate school editor of LSU Digital Commons. For more information, please [email protected]. CHARACTERIZATION OF BUNYAMWERA, BATAI, AND NGARI VIRUSES: UNRECOGNIZED ARBOVIRUSES OF ONE HEALTH IMPORTANCE IN RWANDA

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and Agricultural and Mechanical College in partial fulfillment of the requirements for the degree of Doctor of philosophy

The Department of Pathobiological Sciences

by Marie Fausta Dutuze D.V.M., Ecole Inter-Etats des Sciences et Médecine Vétérinaires de Dakar, 2011 M.P.H., Ecole Inter-Etats des Sciences et Médecine Vétérinaires de Dakar, 2013 August 2019

ACKNOWLEDGMENTS

I dedicate this dissertation to my husband Jean de Dieu Ayabagabo and our son J.

Hugo Songa for allowing to sacrifice these years of being apart. Without their continuous understanding and support, I wouldn’t have been able to do this. J.D, thank you for your long-term friendship, your kindness, your sense of humor but most importantly your love and the life we have built together. Also, thank you for being both a father and a mother for our son when I am not there. Songa, your consistent question: “Uzaza ryari ariko mama?” (meaning: “When are you coming back mom?”)-although made me cry several times- has greatly pushed me to work harder. Having you both in my life is the best thing that ever happened to me.

Special thanks to my mom Stéphanie Nikuze for always being a good model for hard work and determination. Thank you for your efforts to keep our family going forward despite the challenges of being single parent. To my dad Faustin Kanamugire in memorium. Although the course of life separated us physically, your advices and guidance remain alive in my everyday life. Continue resting in eternal peace dad, I miss you every day.

To my brothers Fabrice, Festus, Hippolyte, and Cyrille, and my sister Teddy. Being your big sister has always motivated me to work hard and serve as a good example for you all. I hope I try. Thank you for your daily motivation.

To my father, mother, sisters, and brothers-in-law, I am glad you came into my life.

To my mentor, Dr. Rebecca C. Christofferson. Thank you for your guidance, perpetual encouragement, understanding, and flexibility. You have supported me personally and allowed me to grow professionally. I am very grateful for that. Also, thank you for your

ii spontaneity for finding solutions to challenges (even unusual ones) and for teaching me to think out of the box.

To Dr. Christopher N. Mores. Thank you for giving me the opportunity to join PBS graduate program, guiding my first steps as I entered LSU, and supporting me professionally.

To other members of my graduate committee: Dr. Rhonda Cardin and Dr. Mark

Mitchell. Thank you for your valuable comments which have increased the quality of this work.

To Dr. Manassé Nzayirambaho. Thank you for accepting to be my Rwandan advisor.

Special thanks to Dr. Joyoni Dey, for accepting to serve as dean representative on my committee in non-flexible circumstances.

To Dr. Fabio Del Piero. Thank you for conducting histopathology part of this work.

To past and present members of the Christofferson lab: Ania Kawiecki, Handly

Mayton, Chrissy Walsh, Ryan Tramonte, and Austin. I am grateful for each of you for your time, your help, and entertainment very much needed to survive stressful times.

To other PBS graduate students: Ryan Avery, thank you for your technical assistance and for taking me to my first American football game. Although I found it endless, I enjoyed it. Hanna Laukaitis and Paige Allen, thank you for your technical assistance. Krit

Jirakanwisal, Ifeanyi Kingsley U., Natthida Tongluan, thank you for consistent encouragement.

To Dr. Sean Riley and Daniel Garza, thank you for always fixing BSL3 issues.

iii

Special thanks to my fieldwork teammate Elysée Ndizeye for fun times on the field.

Jean Bosco Noheri, thank you for teaching me how to make dry ice. Grace Mukasine, thank you for your assistance in customer service clearing procedures.

To my friends: ActiveLadies (Joyce and Millicent), Arlie, Virgo, Shimayire, Rugori, Eva,

Surani, and Malaika. Thank you for your consistent encouragement.

To my UR colleagues and friends: Maurice, Rosine, Kizito, Richard H., Richard G., and Claude. Thank you for fun times.

To other BHEARD students at LSU: Sarah, Chunala, Bennett, and Fydess. Thank you for making my stay in Baton Rouge enjoyable.

I am very grateful for these organizations and institutions:

- University of Rwanda (UR) for allowing me to pursue this PhD program.

- United States Agency for International Development (USAID) for financing this

PhD program through Borlaug Higher Education for Agricultural Research and

Development (BHEARD) program.

- Louisiana State University (LSU) for giving me opportunity to pursue this PhD

program

- Rwanda Agriculture Board (RAB) for collaboration. Special thanks to Dr. Isidore

Gafarasi, Angelique Ingabire, Jean Claude Tumushime, Evodie Uwimbabazi, and

Rosa for their technical assistance.

- Rwanda Biomedical Center (RBC) for collaboration. Special thanks to Mr.

Emmanuel Munyemana.

TABLE OF CONTENTS

ACKNOWLEDGMENTS……………………………………………………………...... ii

LIST OF TABLES ………………………………………………………………………………vii

LIST OF FIGURES……………………………………………………………………………...ix

ABSTRACT…………………………………………………………………………...... xi

CHAPTER 1. LITERATURE REVIEW………………………………………………………...1 1.1. Introduction……………………………………………………………………………….1 1.2. Bunyavirales order……………………………………………………………..……...... 7 1.3. Bunyamwera, Batai, Ngari viruses…………………………….………………………13 1.4. Bunyavirus environmental suitability in Rwanda……………………………………..31 1.5. Rationale, hypothesis, and objectives……………………………………………...…35 1.6. References…………………………………………...…………..…………...…….…..36

CHAPTER 2. COMPARISON OF IN VITRO INFECTION KINETICS AND EX VIVO STABILITY OF BUNYAMWERA, BATAI, AND NGARI VIRUSES……………………….48 2.1. Introduction…………………………………………………………...…………...... 48 2.2. Material and methods…………………………………………….…………...……….52 2.3. Results………………………………………..………………………………………....64 2.4. Discussion……………………………………………..………………………………..90 2.5. References…………………………………………………………………..………….95

CHAPTER 3. PRELIMINARY IN VIVO INVESTIGATION OF BUNYAMWERS, BATAI, AND NGARI VIRUSES………………………………………………………………………101 3.1. Introduction………………………………………………...…………………………..101 3.2. Material and methods…………………………..…………………………………….104 3.3. Results……………………………………………………..…………………………..108 3.4. Discussion…………………………………………………………………...………...118 3.5. References…………………………………………………...………………………..121

CHAPTER 4. IDENTIFICATION OF ORTHOBUNYAVIRUS INFECTIONS IN CATTLE DURING A RIFT VALLEY FEVER OUTBREAK IN RWANDA IN 2018…………..……127 4.1. Introduction……………………………………………………...……………………..127 4.2. Material and methods……………………………………………...………………….130 4.3. Results………………………………………………..………………………………..134 4.4. Discussion…………………………………………...………………………...... 140 4.5. References………………………………………………...…………………...... 143

CHAPTER 5. GENERAL DISCUSSION AND CONCLUSIONS…………………………149 5.1. Introduction………………………………………………………………..…...... 149 5.2. Summary of results………………………………..………………………………….150

5.3. Conclusions and future perspectives………………….…………………………...155 5.4. References……………………………………………..……………………………..157

APPENDIX A. FRONTIERS COPYRIGHT STATEMENT ……………………………….161

APPENDIX B. AREAS UNDER THE CURVES (AUC) FITTED BY GROWTHCURVER FUNCTION IN R……………………………………………..………162

VITA……………………………………………………………………………………………174

LIST OF TABLES

Table 1.1. The 32 viruses of the Bunyamwera group, geographic areas of origin, main hosts, and principal vectors, including relevant references.....……………………..19

Table 1.2. Number of mosquito species collected by ecological zones during YFV risk assessment in Rwanda in November-December 2012…………………………...... 34

Table 2.1. Volume calculations for BUNV, BATV, and NRIV for 1.02 x 106 PFU considering their titers…………………………..………………...…………………………..54

Table 2.2. Scores attributed to Cytopathic Effect (CPE)……………………………………56

Table 2.3. Sequences of primers and probes used for qRT-PCR……….……………… 60

Table 2.4. Primers for conventional PCR…………………………………………………….62

Table 2.5. Specificity of qRT-PCR primers…………………………………………………..66

Table 2.6. GenBank similarity percentages of BUNV, BATV, and NRIV segment alignments using the designed traditional PCR primers (BUNV: strain 6547-8, BATV:M2222 strain, and NRIV: DAK-AR-D2852) …………………………………..…….70

Table 2.7. Peak days and Doubling Times (DT) for growth curves of BUNV, BATV, and NRIV with standard cell culture conditions (10%FBS) ………………………………74

Table 2.8. Gradual CPE during persistent infections of BUNV, BATV, and NRIV in Vero cells at 10%FBS……………………………………………………………………..…..80

Table 2.9. Peak days and Doubling Times for growth curves of BUNV, BATV, and NRIV in sub-standard cell culture (2%FBS) ……………………………………………..….81

Table 2.10. Comparison of DT for standard and sub-standard cell culture conditions within viruses………………………………………………………………….………………..82

Table 2.11. Viral titers of supernatants collected at 30 dpi………………………………....83

Table 2.12. Analysis of infectivity of supernatant collected at 30 dpi……………………..89

Table 3.1. Homologous and cross-neutralization study plan……….…………………….107

Table 3.2. PRNT50 and PRNT80 titers for homologous and cross-neutralization BUNV, BATV, and NRIV ………………………………………………………….……...... 110

Table 4.1. Diagnostic scheme of BUNV, BATV, and NRIV using M and L primers…… 133

vii

Table 4.2. Sample categories and proportions of positive samples……………………...137

Table 4.3. Potential co-infection of RVF and BUNV, BATV, and NRIV………………….139

viii

LIST OF FIGURES

Figure 1.1. Rift Valley Fever epidemic cycle indicating One Health implications……..….6

Figure 1.2. Bunyavirales genome……………………………………..……………………..10

Figure 1.3. Geographic distribution of BUNV, BATV, and NRIV (1943-2015) …………..16

Figure 1.4. Genomic organization of BUNV, BATV, and NRIV…………………………….23

Figure 1.5. The four ecological zones of Rwanda ………………………………………….33

Figure 2.1. General viral growth curve in mammalian cell …………………………….…..50

Figure 2.2. Comparative plaque morphology for BUNV, BATV, and NRIV……………….65

Figure 2.3. Specificity of designed primers for conventional PCR…………………………68

Figure 2.4. Growth curves of BUNV, BATV, and NRIV in standard cell culture condition (10%FBS), at a MOI of 1 ………………………………………………...…..…...71

Figure 2.5. BUNV, BATV, and NRIV growth kinetics in Vero cells grown in standard conditions (10% FBS) with different inoculation doses…………………………………....73

Figure 2.6. Logic framework of in vitro and ex vivo growth kinetics study of BUNV, BATV, and NRIV……………………………………………………………………………….76

Figure 2.7. BUNV, BATV, and NRIV growth kinetics in Vero cells grown sub-standard conditions (2%FBS) with different inoculation doses……………………...78

Figure 2.8. Comparative infectivity of BUNV after in vitro persistence at 30 dpi from cells both grown in the standard (10%FBS) and sub-standard conditions (2%FBS) ….84

Figure 2.9. Comparative infectivity of BATV after in vitro persistence at 30 dpi from cells both grown in the standard (10%FBS) and sub-standard conditions (2%FBS) ….85

Figure 2.10. Comparative infectivity of NRIV after in vitro persistence at 30 dpi from cells both grown in the standard (10%FBS) and sub-standard conditions (2%FBS) ….86

Figure 2.11. Ex vivo stability of BUNV BATV, and NRIV……….………………………….87

Figure 2.12. Infectivity of BUNV, BATV, NRIV after stability in ex vivo conditions……….88

Figure 2.13. Viral RNA detection of BUNV, BATV, and NRIV after 1% Triton-X-100 inactivation ……………………………………………………………………………………..90

Figure 3.1. Comparative viremia curves of BUNV, BATV, in NRIV in C57bl/6 mice……109

Figure 3.2. Comparative homologous and cross-neutralization profiles for BUNV, BATV, and NRIV for different serum dilutions ………….………………………………...112

Figure 3.3. Dead BUNV infected IRF 3/7 -/--/- mouse with facial swelling…………………113

Figure 3.4. Viremia, weight loss, and survival of BUNV infected IRF 3/7 -/--/- mice……114

Figure 3.5. Necrotic lesions of the liver of BUNV infected IRF3/7 -/--/- mice ……….……115

Figure 3.6. Necrotic and inflammatory lesions of the lung of BUNV infected IRF3/7 -/--/- mice ……………………………………………………………….………….…..116

Figure 3.7. Necrotic lesions of lymphoid organs in BUNV infected IRF3/7 -/--/- mice ….116

Figure 3.8. Necrotic lesions of reproductive system of BUNV infected IRF3/7 -/--/- female mice ………………………………………………………………...…………..……117

Figure 3.9. Lesions associated with facial swelling …………………………………….....117

Figure 4.1. Clinical manifestations of cattle during RVF outbreak……………………….131

Figure 4.2. Administrative map of Rwanda displaying provinces and districts…….…..134

Figure 4.3. Epidemic chronology of RVF outbreak ………………………………………..135

Figure 4.4. Positive samples to BUNV and/or BATV and negative RVFV……………….138

Figure 4.5. Geographic distribution of RVF and Orthobunyaviruses cases …………….142

ABSTRACT

Bunyamwera (BUNV), Batai (BATV), and Ngari (NRIV) are mosquito-borne viruses (Genus: Orthobunyavirus, Family: Peribunyaviridae, Order: Bunyavirales). They have an RNA tripartite genome consisting of small (S), medium (M), and large (L) segments. NRIV is a natural reassortant of BUNV and BATV, with the genome: SBUNV,

MBATV, LBUNV. All three viruses have been associated with disease of varying severity in domestic ruminants and humans. In livestock, infection with these viruses is associated with abortions, while humans may present a mild febrile illness or severe disease such as hemorrhagic fever or meningoencephalitis. In East Africa, BUNV, BATV, and NRIV co- circulate and present similar clinical manifestation with Rift Valley Fever virus (RVFV), another Bunyavirus (Genus: Phlebovirus, Family: Phenuibunyaviridae) of critical importance in this region as it is associated with massive economic losses in livestock production and cases of human death. In Rwanda, although RVFV is known to circulate in livestock, whether these other Bunyaviruses co-circulate remains to be determined.

The overall goal of this research is to describe and compare the in vitro and in vivo characteristics of BUNV, BATV, and NRIV. To achieve this objective, we a) comparatively characterized in vitro infection kinetics in Vero cells, b) comparatively characterized in vivo infection kinetics in C57BL/6 mouse model and quantified potential cross-neutralization of resulting antibody, and c) developed a disease progression model for BUNV infection in IRF3/7-/--/- mice. Further, we hypothesized that these

Orthobunyaviruses circulate in Rwanda and cause infections that are misclassified as

RVFV cases. We found: 1) all viruses are stable in extracellular conditions up to 30 days but inactivation by a commonly used detergent is successful; 2) C57BL/6 mouse infection

xi is not a robust infection model; 3) there are varying degrees of cross-neutralization among the viruses; 4) BUNV infection in IRF3/7-/--/- mice may provide hemorrhagic fever infection model; and 5) these viruses co-circulate with RVFV in Rwanda. This study is the second report of BATV in Africa, the first to detect these viruses in Rwanda, and the first report of co-infection with BUNV and BATV, providing insight into the provenance of the reassortant NRIV.

xii

CHAPTER 1. LITERATURE REVIEW

1.1. Introduction

The Bunyavirales order, includes more than 350 viruses1 across 10 taxonomic families namely Hantaviridae, Nairoviridae, Phenuiviridae, Peribunyaviridae, Cruliviridae,

Fimoviridae, Mypoviridae, Wupedeviridae, Phasmaviridae, and Arenaviridae (Maes et al.

2019). Members of this order are often transmitted by mosquitoes and have been attributed to sometimes severe diseases in humans, livestock, wildlife, and plants worldwide for a long time (Bishop et al. 1980; Calisher 1996; Elliott 2013). Although bunyaviral disease manifestation differs among virus species and affected hosts, the most commonly detected disease is spontaneous abortion in animals (mostly domestic ruminants) while in humans, infections are generally characterized by non-specific febrile illnesses though severe forms can manifest with hemorrhagic fever or neurological complications (Soldan and González-Scarano 2014; Wilson et al. 2015). As these viruses affect both humans and animals, they are of One Health importance and should be studied in the context of their multi-factorial impacts in the affected region (s).

Bunyaviruses have single-stranded RNA genome, which is bi-segmented in the

Arenaviridae family (Small and Large segments) and tri-segmented in other families

(Small, Medium, and Large segments). In the Arenaviridae family, the Small segment (S) encodes major structural proteins (nucleoprotein and glycoproteins) while the Large segment encodes RNA-dependent RNA polymerase and a Zinc-binding protein (Z)

This chapter has been published as: Dutuze, M. F., M. Nzayirambaho, C. N. Mores, and R. C. Christofferson. 2018. "A Review of Bunyamwera, Batai, and Ngari Viruses: Understudied Orthobunyaviruses With Potential One Health Implications." Front Vet Sci 5:69. doi: 10.3389/fvets.2018.00069.

(Lukashevich 2006; Loureiro, D'Antuono, and López 2019; Vogel et al. 2019). For all other families, Small (S), Medium (M), and Large (L) segments encode respectively the nucleoprotein, glycoproteins, and RNA-dependent RNA polymerase. Additionally, S and

M segments of the Peribunyaviridae and Phenuiviridae families encode additional nonstructural proteins (Calisher 1996; Elliott 2013; Soldan and González-Scarano 2014). The genome of Bunyaviruses is prone to various types of mutations and reassortments, which can result in altered phenotypic characteristics compared to parental viruses, such as differences in pathogenesis and even host range (Kolakofsky and Hacker 1991; Elliott

2013; Soldan and González-Scarano 2014; Otieno 2015). This further enhances the one health implications of these viruses, which are already economically and medically important due to their significant number, the wide host range, the broad geographic distribution, and the potentially serious clinical manifestations.

While reassortment among viruses of the Bunyavirales order has been well documented in experimental settings (Borucki et al. 1999; Briese, Calisher, and Higgs

2013) and predicted to occur at relatively high rates in nature (Briese, Calisher, and Higgs

2013), few recognized natural reassortment events are reported (Bowen et al. 2001;

Briese et al. 2006; Aguilar et al. 2011; Veldhuis et al. 2013; Hontz et al. 2015).

In the Peribunyaviridae family, the largest family of the order, several sudden discoveries of new viruses have highlighted the importance of reassortment in this group of viruses. Examples include the identification of the now confirmed reassortant NRIV and yet to be confirmed but suspected reassortants such as Itaya, Iquitos, and

Schmallenberg viruses (Bowen et al. 2001; Briese et al. 2006; Aguilar et al. 2011;

Veldhuis et al. 2013; Hontz et al. 2015). NRIV has been associated with human

2 hemorrhagic fever cases in East Africa in 1998-1999 and with abortive disease in goats in Mauritania in 2010, cementing its place as a One Health threat (Bowen et al. 2001;

Briese et al. 2006; Beer, Conraths, and van der Poel 2013; Eiden et al. 2014). Itaya virus is suspected to be a reassortant between Caraparu virus (S and L segments) and another unknown Orthobunyavirus (M segment) and was associated with human febrile illnesses between 1999 and 2006 in Peru (Hontz et al. 2015). Iquitos virus was identified as a reassortant between Oropouche virus (OROV) (S and L segments) and another unknown

Orthobunyavirus (M segment) and associated with febrile illnesses between 2005 and

2006 in Peru (Aguilar et al. 2011). Schmallenberg virus (SBV) is identified as a reassortant between Shamonda virus (S and L segments) and possibly Sathuperi or Douglas viruses

(for M segment) and has caused severe outbreaks of malformations in ruminants in several countries of Northern Europe between 2011 and 2012 (Garigliany et al. 2012;

Beer, Conraths, and van der Poel 2013, Yanase et al. 2012; Veldhuis et al. 2013; Wernike et al. 2014).

Among those viruses produced by natural reassortment events, confidence into the genetic origins of NRIV is relatively high. Additionally, NRIV could be the reassortant with highest potential to become a major One Health issue, as it has been found to cause disease in both humans and animals like its genetic “parents”, sometimes severe (Bowen et al. 2001; Gerrard et al. 2004; Eiden et al. 2014). Its importance also lies in its broad range as it has been identified several times in different ecological zones (discussed further in this chapter), potentially indicative of a broader host or vector range (Zeller et al. 1996; Gerrard et al. 2004; Aguilar et al. 2011).

NRIV has been associated with highest pathogenicity among the three viruses: NRIV,

BUNV, and BATV (Zeller et al. 1996; Bowen et al. 2001; Briese et al. 2006; Eiden et al.

2014). The disease associated with BUNV is characterized by mild symptoms such as fever, joint pain, and rash in many mammals, including humans (Kokernot et al. 1958;

Ashford 2001; Rodrigues Hoffmann et al. 2013; Odhiambo, Venter, Limbaso, et al. 2014;

Tauro et al. 2015). BATV causes a mild flu-like illness in humans but it is associated with a more severe disease in ruminants where it is manifested by abortions, premature births, and genetic defects (Ashford 2001; Yanase et al. 2006; Medlock, Snow, and Leach 2007).

NRIV has been associated with fatal hemorrhagic fevers in both humans and ruminants

(Bowen et al. 2001; Briese et al. 2006; Beer, Conraths, and van der Poel 2013; Jackel et al. 2013; Eiden et al. 2014). However, these reports represent only a handful of actual isolations and definitive diagnoses and thus, there remains a need to further characterize these viruses, the diseases they cause, and their potential to become public health issues, with emphasis on their One Health importance.

BUNV, because it is the most characterized member of the Bunyavirales order, is known as the prototype of that order (Szemiel, Failloux, and Elliott 2012; Riblett and Doms

2016). BATV has also been studied given its wide distribution across Europe and Asia, while NRIV is the least studied of the three (Briese et al. 2006; Medlock, Snow, and Leach

2007). Studies of NRIV have been focused on genomic comparisons with other viruses of the Peribunyaviridae family (BUNV, BATV, and Ilesha viruses) and the similarity of its clinical manifestations with RVFV (Gerrard et al. 2004; Briese et al. 2006). RVFV is often associated with abortions in ruminants (cattle, goats and sheep) and mild (flu-like) to severe (hemorrhagic fever, encephalitis, and ocular troubles) symptoms in humans and

4 ruminants. RVFV causes large outbreaks on the African continent and Arabian Peninsula and is a major public health and food security concern and is a good example of the One

Health paradigm (Balkhy and Memish 2003; Kifaro et al. 2014; Nanyingi et al. 2015; Bird and McElroy 2016).

The concept of One Health centers on the idea that the health of humans is intimately tied to the health of animals and the environment (Oura 2014; Asokan 2015; Stadtlander

2015; Mwangi, de Figueiredo, and Criscitiello 2016; Sukura and Hanninen 2016;

Havigerova, Dosedlova, and Buresova 2019). In the case of mosquito-borne viruses, such as RVFV and the Orthobunyaviruses of concern here, this concept is easily illustrated by Figure 1.1. Briefly, in developing nations, cattle are often important for food security, personal and familial economy (Schneider 1957; Hall 1986; Barrett 1992). Thus, their health is intimately tied to the well-being of the humans associated with those populations. Further, in zoonotic systems, where viruses may infect both humans and animals, these cattle may serve as reservoirs or amplification hosts, directly contributing to the risk of human disease. Finally, the vectors that transmit these viruses are closely tied to the ecology and environment of the transmission system. Thus, an environmental component can drive and alter transmission and disease risk in both the human and animal populations. This system is a One Health system.

Figure 1.1. Rift Valley Fever epidemic cycle indicating One Health implications. This cycle is applicable to other mosquito-borne zoonotic diseases (modified from (Bird and McElroy 2016)).

NRIV was recognized among etiologies of febrile illnesses in humans in various countries in East Africa. It was identified in Sudan, Somalia, and Kenya outbreaks in 1988,

1997, and 1998, respectively. In these instances, molecular-based techniques such as nucleic acid amplification and sequencing were rarely used, and diagnosis was based on clinical manifestations, thus the disease was diagnosed as the most common febrile illnesses (Nashed, Olson, and el-Tigani 1993; Bowen et al. 2001; Briese et al. 2006;

Groseth et al. 2012). Consequently, in the Sudanese outbreak, the disease was improperly diagnosed as malaria, as in malaria-endemic countries 70% of febrile illnesses are treated as presumptive malaria, often without proper medical examination and a laboratory analysis (Amexo et al. 2004). In Somalia and Kenya, as it was during an ongoing RVFV outbreak, the disease was mistakenly taken for RVFV. In addition to East

Africa, NRIV has also been identified in goats during an ongoing RVF outbreak in

Mauritania as well (Eiden et al. 2014). This shows that the burden of disease of NRIV and

6 likely BUNV and BATV as well is underreported and their public health impact under- appreciated.

In East Africa, despite various isolations of different Bunyaviruses, the status of many countries in that region remains unknown and requires more investigation. In

Rwanda for example, no extensive research has been conducted on potential presence of different members of the Bunyavirales order. Although the environment seems suitable for the transmission of these viruses, the only recognized and surveilled Bunyavirus is

RVFV.

In this literature review, we compile the available knowledge on BUNV, BATV, and NRIV. Emphasis is put on the relationships among molecular characteristics, transmission mechanisms, geographic distributions, and clinical manifestations among the BUNV, BATV, and NRIV, as well as with other related viruses. In addition, we discuss the documented and potential co-circulation of BUNV, BATV, and NRIV with RVFV, in

East Africa and in Rwanda particularly.

1.2. Bunyavirales order

The new Bunyavirales order (according to latest taxonomy as of October 2018) is one of the largest groups of RNA viruses. It counts more than 350 viruses, usually referred to as ‘Bunyaviruses’ and classified within 10 families. Among the ten (10) families: four consist old genera of the former Bunyarividae family considered now as families, five consist of the newly characterized Bunyaviruses, and viruses belonging to Arenaviridae family joined this order. The members of the former Bunyaviridae family are: former

Hantavirus genus, currently named Hantaviridae family; former Nairovirus genus, currently named Nairoviridae family; former Phlebovirus genus, currently named

Phenuiviridae family for the former, and former Orthobunyavirus genus, currently named

Peribunyaviridae family. The fifth genus of the former Bunyaviridae family-Tospovirus genus-was moved to Peribunyaviridae family. The five families consisting the newly characterized Bunyaviruses are Cruliviridae, Fimoviridae, Mypoviridae, Wupedeviridae, and Phasmaviridae families (Maes et al. 2019).

The taxonomy of these viruses is a dynamic process due to their significant high number, reassortment events leading to new viruses, widespread distribution, wide host range, and mostly the non-specific aspect of their diseases.

Except Arenaviridae family which has a bipartite RNA genome, other families of the Bunyavirales order have tripartite RNA genome consisting of small(S), medium (M), and large (L) segments due to their relative sizes which are respectively of 961bp-2,916 bp; 3,616-4,888 bp; and 6,404-12,255 bp (Calisher 1996; Soldan and González-Scarano

2014) (Figure 1.1). These RNA segments are single-stranded, exist in helical formation within the virion, and exhibit a pseudo-circular structure due to each segment’s complementary end. The L and M segments are negative sense. L encodes the RNA dependent RNA-polymerase, necessary for viral RNA replication and mRNA synthesis.

The M segment encodes the viral glycoproteins, which projects from viral surface and help the virus in attaching to and entering the host cell. Additionally, M segment of members of Peribunyaviridae and Phenuiviridae families encodes NSm is thought to play a role in virus assembly and suppression of virus-induced apoptosis (Calisher 1996;

Schmaljohn 2001; Soldan and González-Scarano 2014; Otieno 2015). The S segment encodes nucleocapsid (N) in negative sense. For Peribunyaviridae and Phenuiviridae families, this segment encodes an additional protein; nonstructural protein (NSs) in

8 ambisense (Phenuiviridae) and through overlapping reading frame (Peribunyaviridae)

(Soldan and González-Scarano 2014). NSs has been shown to play an important role in countering the innate immune response to viral infection, serving as a suppressor of RNA silencing and inhibiting host cell protein synthesis (Bridgen et al. 2001; Weber et al. 2002;

Soldan et al. 2005). For the Arenaviridae family, the two RNA genome segments S (3.4kb) and L (7 kb) are all ambisense. The S segment encodes the viral nucleocapsid protein

(NP) and glycoprotein precursor (GPC) which is post-translationally cleaved into Gc and

Gn. L segment encodes Z protein, which binds zinc and acts as a matrix protein, and the

L protein, which is likely to function as the viral polymerase (Figure 1.2) (Hass et al. 2004).

Figure 1.2. Bunyavirales genome. A. Genome of Bunyamwera virus (prototype of the Bunyavirales order), representative of Hantaviridae, Nairoviridae, Peribunyaviridae, Phenuiviridae, Cruliviridae, Fimoviridae, Mypoviridae, Wuperiviridae, and Phasmaviridae families (Bioinformatics Swiss Institute, 2010a), B. Lassa virus genome, representative of the Arenaviridae family (Bioinformatics Swiss Institute, 2010b).

Bunyaviruses are distributed worldwide and cause diseases in a large number of hosts including humans, livestock, and wildlife. The clinical manifestations of diseases associated with these viruses vary but are mainly characterized by abortions in livestock, and mild febrile illness to severe hemorrhagic fever in humans. However, a big proportion of infections are asymptomatic (Calisher 1996; Elliott 2013; Soldan and González-

Scarano 2014; Wilson et al. 2015).

The Bunyavirales order includes highly pathogenic viruses of significant public health concern such as Crimean-Congo hemorrhagic fever virus (CCHFV) in the

Nairoviridae family; Rift Valley fever virus (RVFV), Severe fever with thrombocytopenia syndrome virus (SFTSV), and Toscana virus (TOSV) in the Phenuiviridae family; La

Crosse virus (LACV), Oropouche virus (OROV), Cache Valley virus (CVV), and Akabane virus (AKAV) in the Peribunyaviridae family; as well as Lassa virus (LASV) in the

Arenaviridae family. These viruses are causative of severe and fatal illnesses characterized by hemorrhagic fever, pulmonary disease, congenital abnormalities, and encephalitis (Zeller et al. 1996; Zentis et al. 2012; Soldan and González-Scarano 2014;

McDonald et al. 2016; Dutuze et al. 2018).

With exception of the members of the Hantaviridae family which are rodent- transmitted, Bunyaviruses are principally transmitted by arthropods. Unlike most arboviruses which tend to be adapted to a narrow range of vectors, arthropod-borne

Bunyaviruses are transmitted by a wide range of vectors such as mosquitoes, ticks, flies, sandflies, and biting midges (Calisher 1996; Wertheim 2012; Elliott 2013).

In addition, direct transmission is an important way of transmission in cases of hemorrhagic fever and abortions when the virus is largely being spread in the

11 environment. This has been found to be significantly important in transmission of RVFV, and CCHFV (Shepherd et al. 1989; Balkhy and Memish 2003; Conger et al. 2015;

Pshenichnaya and Nenadskaya 2015; Bird and McElroy 2016).

Like for all RNA viruses, Bunyaviruses are subjects to various mutations (Elliott 1989a;

Weber and Elliott 2002). Additionally, the segmented structure of their genome exposes them to reassortment, a process of genetic recombination exclusive to segmented RNA viruses in which co-infection of a host cell with multiple viruses may result in the shuffling of gene segments to generate progeny viruses with novel genome combinations (Weber and Elliott 2002). Theoretically, co-infection of two tri-segmented Bunyaviruses would lead to the creation of 6 (23-2) reassortants while co-infection of bi-segmented

Bunyaviruses would lead to 2 (22-2) reassortants. However, this process requires genetic compatibility between parental viruses, which occurs in the form of conserved packaging signals, maintenance of RNA, and protein interactions (McDonald et al. 2016). This may explain why even though many Bunyaviruses co-circulate in the same regions and share hosts and vectors, reassortment events are not as many as the theory seems to indicate.

Reassortment may result in novel aspects of pathogenesis, pathogenicity, and virulence within same hosts. This has been seen with the unusually high pathogenicity of NRIV in humans and SBV in ruminants (Bowen et al. 2001; Briese et al. 2006; Garigliany et al.

2012; Yanase et al. 2012; Beer, Conraths, and van der Poel 2013; Wernike et al. 2014).

Due to the significant number of members in this order and to the non-specific disease manifestations, these reassortments are thought to go unnoticed except when they are clinically manifested with severe alarming signs (Briese, Calisher, and Higgs 2013).

One of the most documented naturally occurring reassortement is the creation of

Ngari virus (NRIV) from Bunyamwera virus (BUNV) and Batai virus (BATV), two members of Orthobunyavirus in the Peribunyaviridae family. The three viruses cause diseases in animals and humans and are characterized by different virulence. BUNV is characterized by moderate virulence, BATV by low virulence while NRIV is more virulent than its two

“parents” (Bowen et al. 2001; Briese et al. 2006; Medlock, Snow, and Leach 2007;

Rodrigues Hoffmann et al. 2013; Eiden et al. 2014; Tauro et al. 2015).

Despite their relatively significant medical importance, their presumably related geographic distribution, and the less-characterized conditions of this natural reassortment, these viruses are understudied. This dissertation covers the comparative characterization of BUNV, BATV, and NRIV on different aspects such as in vitro and in vivo infection kinetics, geographic distribution, and (dis) similarities with RVFV especially on clinical manifestation.

1.3. Bunyamwera, Batai, and Ngari viruses

1.3.1. Origins and geographic distributions

BUNV was first isolated in 1943 from Aedes mosquitoes collected in the Semliki Forest of Uganda as part of a yellow fever surveillance effort (Smithburn, Haddow, and Mahaffy

1946; Kokernot et al. 1958; Szemiel, Failloux, and Elliott 2012). BUNV is now considered endemic in several African countries, such as Uganda, Tanzania, Mozambique, Nigeria,

Guinea, South Africa, and Democratic Republic of Congo (Figure 1.2). Recently, Cache

Valley Fever virus was classified as a strain of BUNV, which expands its geographic range to the North and strains of BUNV were isolated in Argentina (Tauro et al. 2015). A recent compilation of studies reported the presence of BUNV in Senegal, Guinea, Ivory Coast,

Nigeria, Cameroon, Central African Republic, Kenya, Uganda, South Africa, and

Madagascar (Wertheim 2012).

Ngari virus was first isolated from Aedes simpsoni mosquitoes in 1979 in Southeastern

Senegal. It was then recovered from several mosquito species in Senegal, Burkina Faso,

Central African Republic and Madagascar (1988–1993) (Zeller et al. 1996). The first suspicion of its potential pathogenicity in humans was when the virus was isolated from two patients in Dakar in October and November 1993 (Wertheim 2012). In Kenya and

Somalia (1998–1999), the Garissa strain of NRIV was identified in what was thought to be a RVFV outbreak, owing to the hemorrhagic manifestations of infection (Bowen et al.

2001). In 2010, during an ongoing RVFV outbreak in Mauritania, NRIV was again isolated in goats (Eiden et al. 2014). NRIV has additionally been isolated in Burkina Faso, Central

African Republic, and Madagascar (Wertheim 2012). In all these cases, microscopy, RT-

PCR, and complete nucleotide sequences were used in the differential diagnostic panels and confirmed the differentiation between other Bunyaviruses (BUNV, BATV, ILEV, and

RVFV) as well as other suspected agents (malaria) (Bowen et al. 2001; Gerrard et al.

2004; Eiden et al. 2014).

Batai virus is one of the most geographically widespread members of the

Orthobunyavirus genus (Figure 1.3) (Briese et al. 2006; Groseth et al. 2012). It also has the broadest vector range as it was isolated from Anopheles maculipennis in

Czechoslovakia in 1950, from Culex gelidus Theo mosquitoes in Malaysia, from

Anopheles barbirostris mosquitoes in the Chittoor district in India, and from A. maculipennis and Aedes punctor mosquitoes collected in what is modern Ukraine (Singh

1966; Gaidamovich et al. 1973; Karabatsos 1985; Briese et al. 2006; Szemiel, Failloux,

14 and Elliott 2012; Liu et al. 2014). In Europe, evidence for circulation exists in Norway,

Sweden, Finland, Slovakia, the Czech Republic, Croatia, Serbia, Bosnia, Montenegro,

Italy, Hungary, Romania, Austria, Portugal, Germany, and Belarus (Hubálek 1996). The seroprevalence reported in humans in Europe is low: less than 1% in Sweden, Finland,

Germany, Austria, and parts of the former Yugoslavia. However, there was a 32% seroprevalence reported in southern Slovakia (Medlock, Snow, and Leach 2007).

Similarly, seroprevalence in animals varied from 1 to 46% (Medlock, Snow, and Leach

2007). Interestingly, BATV is not widely reported on the African continent, having only been reported in Uganda in 1967 based on molecular techniques (PCR) (Briese et al.

2006) and highly suspected in Sudan in 1988 based on serological technique (PRNT)

(Nashed, Olson, and el-Tigani 1993). However, the distribution of NRIV as well as the presence of suitable hosts and vectors suggests that BATV might be present in other

African countries and be underreported, or at least has the potential to be so.

Although no research has been conducted on the asymptomatic presentation of

BUNV, BATV, and NRIV infections in vertebrate hosts, other mosquito-borne viruses across several genera are known to present as mostly asymptomatic or as a mild influenza- like illness (Hollidge, González-Scarano, and Soldan 2010). Therefore, it is possible that NRIV, BUNV, and even BATV were circulating before successful isolation and that they may silently circulate in many areas, where no serological studies have been conducted that may detect asymptomatic or mild, generalized disease.

NRIV was isolated 35 years ago, nearly 30 years after the parental viruses, BUNV and BATV. Given the geographical history of BUNV and BATV, the speculation is that the first reassortment leading to new viral progeny likely resulted from the introduction of

BATV into the population of already-circulating BUNV in Africa. Many factors may have contributed to the introduction of BATV in East Africa, likely long-distance bird migration

(Altizer, Bartel, and Han 2011). But these distinct geographies of BATV and BUNV also suggest that NRIV likely has developed its own niche and propagates as a distinct virus rather than relying on repeated introductions and chance co-infections of BATV and

BUNV in the wild.

Figure 1.3. Geographic distribution of BUNV, BATV, and NRIV (1943-2015). The known distributions of Bunyamwera, Ngari, and Batai viruses in Europe and Africa, where most isolations have occurred. Not shown are periodic isolations in Russia, India, Malaysia, and China, as well as Cache Valley Fever Virus in North America. 16

1.3.2. Classification, genomic characteristics and replication in host cells a. Classification

BUNV, BATV, and NRIV belong to the Bunyamwera group, one of 18 serologically defined arbovirus serogroups belonging to the Orthobunyavirus genus of the family

Peribunyaviridae (Yanase et al. 2006). The 18 serogroups are Anopheles A, Anopheles

B, Bakau, Bunyamwera, Bwamba, group C, Capim, California, Gamboa, Guama,

Koongol, Minatitlan, Nyando, Olifanstlei, Patois, Simbu, Tete, and Turlock (Mohamed,

MacLees, and Elliott 2009). The viruses of this genus have been classified into these different serogroups based on serological relationships of complement fixing antibodies, as well as hemagglutinating assay results and neutralizing antibodies (Calisher 1996).

The classification of the Bunyaviridae family has evolved, since its first organization in

1971 when the Subcommittee on Interrelationships Among Catalogued Arboviruses

(SIRACA) conducted a serological study of 23 viruses. They subsequently classified 14 viruses as belonging to the Bunyamwera group viruses and further identified five subgroups or complexes: Bunyamwera, Cache Valley, Wyeomyia, Kairi, and Guaroa

(Scherer 1976). In 1978, this classification was updated based on the antigenic relationships of the Bunyamwera group viruses established by the cross-neutralization pattern of virus-specific hyperimmune antibodies (Tikasingh, Spence, and Downs 1966).

From this effort emerged three subgroups that diverge largely over geographic origin:

South America, Africa, and Europe/Asia (of which BATV is the only member) (Hunt and

Calisher 1979). While there was very little cross-neutralization among members of the

South-American and African subgroups, BATV is cross-neutralized by other virus-specific antibodies from the other two subgroups (Hunt and Calisher 1979). As of 1996, there are over 30 Bunyamwera serogroup viruses that have been identified and classified (Calisher 17

1996). As mentioned, CVV was also recently reclassified as a strain of BUNV (Tauro et al. 2015). Table 1.1 shows the 32 Bunyamwera group viruses and the geographic areas, where they were first isolated, as well as their main vertebrate hosts and principal vectors.

Most of the Bunyamwera serogroup viruses are etiological agents of human and animal diseases (Southam and Moore 1951; Kokernot et al. 1958; Causey et al. 1961; Casals

1965; Groseth et al. 2012).

The taxonomy of Bunyaviruses has been challenging, given the propensity for reassortment, their general lack of characterization, and the broad geographic ranges of some genera. Often, taxonomy is first developed based on antigenic relationships, but is becoming more molecular-based as genomic methods become cheaper and data becomes available (Kuhn et al. 2016). Recently, the International Committee on

Taxonomy of Viruses proposed the latest classification whereby from the family of

Bunyaviridae, a new virus order of Bunyavirales was created. As previously mentioned, the new Bunyavirales order consists of ten families combining all viruses of the former

Bunyaviridae family, the newly characterized Bunyaviruses, and viruses of the

Arenaviridae family. In this classification, BUNV, BATV, and NRIV belong to

Orthobunyavirus genus of the Peribunyaviridae family (Maes et al. 2019).

Table 1.1. The 32 viruses of the Bunyamwera group, geographic areas of origin, main hosts, and principal vectors, including relevant references

Virus Abbreviation Source, location, year of Main hosts Principal arthropod References first isolation vectors AG83 1746 South America Rodents Mosquitoes (Calisher 1996) Anhembi AM Brasil Not known Mosquitoes (culicine) (Hunt and Calisher 1979; Calisher 1996) Batai BAT Anopheles maculipennis, Humans, Mosquitoes (Hunt and Calisher 1979; Calisher Czechoslovakia, 1950 ruminants, sentinel (anopheline, culicine), 1996; Liu et al. 2014) animals (birds) Biting midges BeAr South America Not known Mosquitoes (Calisher 1996) 3282208 Birao BIR Anopheles pharoensis, Not known Mosquitoes (Hunt and Calisher 1979; Calisher Central Afr. Rep., 1969 (anopheline) 1996; Liu et al. 2014) Bozo Africa Not known Mosquitoes (Calisher 1996) (anopheline, culicine) Bunyamwera BUN Aedes spp., Uganda, Humans Mosquitoes (Hunt and Calisher 1979; Calisher 1943 (anopheline, culicine) 1996; Liu et al. 2014) Cache valley CV Culiseta inornata, Utah, Sheep, cattle, Mosquitoes (Hunt and Calisher 1979; Calisher 1956 marsupials (anopheline, culicine) 1996; Liu et al. 2014) CbaAr426 South America Not known Mosquitoes (Calisher 1996) Fort Sherman Human, Panama, 1985 Humans Mosquitoes (Calisher 1996; Liu et al. 2014) Germiston GER South Africa Humans, rodents, Mosquitoes (culicine) (Hunt and Calisher 1979; Calisher sentinel animals 1996)

(Table 1.1 cont’d)

Virus Abbreviation Source, location, year of Main hosts Principal arthropod References first isolation vectors Guaroa GRO Colombia Humans Mosquitoes (Hunt and Calisher 1979) Iaco South America Not known Mosquitoes (culicine) (Calisher 1996) Ilesha ILE Human, Nigeria, 1957 Humans Mosquitoes (anopheline) (Hunt and Calisher 1979; Liu et al. 2014) Kairi KRI Trinidad Equine, primates, Mosquitoes (culicine) (Calisher 1996) rodents, sentinel animals Lokern LOK USA Cattle, goats, Mosquitoes (anopheline, (Hunt and Calisher 1979; Calisher sheep culicine), Culicoid flies 1996,) Macaua South America Rodents, bats Mosquitoes (culicine) (Calisher 1996) Maguari MAG Aedes scapularis, Sentinel animal, Mosquitoes (anopheline, (Hunt and Calisher 1979; Calisher Ecuador, 1974 cattle culicine) 1996; Liu et al. 2014) Main Drain MD USA Equine Mosquitoes (culicine), (Hunt and Calisher 1979; Calisher Culicoid flies 1996) Mboke Africa Not known Mosquitoes (culicine) (Calisher 1996) Ngari Aedes simpsoni, Humans Mosquitoes (anopheline, (Calisher 1996; Odhiambo, Venter, Senegal, 1979 culicine), ticks Chepkorir, et al. 2014; Odhiambo et al. 2015) Northway NOR Aedes spp., Alaska, Rodents, sentinel Mosquitoes (culicine) (Hunt and Calisher 1979; Liu et al. 1971 animals 2014) Playas South America Not known Mosquitoes (culicine) (Elliott 1997) Potosi Aedes albopictus, Not known Mosquitoes (Elliott 1997; Liu et al. 2014) Missouri,1989

(Table 1.1 cont’d)

Virus Abbreviation Source, location, year of Main hosts Principal arthropod References first isolation vectors Santa Rosa North America Not known Mosquitoes (culicine) (Hunt and Calisher 1979) Shokwe Aedes cumminsii, South Humans, rodents Mosquitoes (anopheline, (Hunt and Calisher 1979; Calisher Africa, 1962 culicine) 1996; Liu et al. 2014) Sororoka SOR Brazil Not known Mosquitoes (culicine) (Hunt and Calisher 1979; Calisher 1996) Tensaw TEN USA Rodents, sentinel Mosquitoes (anopheline, (Calisher 1996) animals, cattle culicine) Tlacotalpan TLA Mexico Not known Mosquitoes (Hunt and Calisher 1979; Calisher 1996) Tucunduba Brazil Not known Mosquitoes (Hunt and Calisher 1979; Calisher 1996) Wyeomyia WYO Wyeomyia Not known Mosquitoes (anopheline, (Hunt and Calisher 1979; Calisher melanocephala, culicine) 1996: Liu et al. 2014) Colombia, 1940 Xingu Human, Brazil, unknown Humans Mosquitoes (Elliott 1997; Liu et al. 2014) date

b. Genomic characteristics

As with all viruses in the Peribunyaviridae family, BUNV which is the prototype of the family has a segmented negative sense RNA genome. As stated, the genome is composed of three segments S, M, and L. Each segment is transcribed individually to give a single mRNA (Figure 1.4). Although there are differences in the sizes of the segments among different species and among strains within species, the averages for

Orthobunyaviruses are 6.9 kb for the L segment, 4.5 kb for the M segment and 1.0 kb for the S segment (Elliott 2014). The S segment encodes the N (nucleocapsid) protein and a NSs, which are translated from overlapping open reading frame in the same mRNA by leaky ribosomal scanning (Fuller and Bishop 1982; Beaty, Fuller, and Bishop 1983; Elliott

1989b; Elliott 2014). The M segment mRNA encodes a polyprotein that is post- translationally cleaved by host proteases into NS, and glycoproteins (Gn and Gc)

(Gentsch and Bishop 1979; Fuller and Bishop 1982; Elliott 1985) and the L segment mRNA encodes the RNA-dependent RNA polymerase (Elliott 1989b; Walter and Barr

2010; Elliott 2014) (Figure 1.4). As in other Bunyavirus families (Nairoviridae and

Phenuiviridae), NSs and NSm have been shown to be closely related to the pathogenicity and propagation of the viruses in vertebrate cells (Kuhn et al. 2016; Phoenix et al. 2016).

Although NSs is encoded by all three viruses, the length of this gene can vary among the genus (Elliott 2014).

The coding sequences are generally less conserved than the untranslated regions

(UTRs). The functional promoter is formed by the 5′ and 3′ UTRs of each segment

(Barr et al. 2003; Kohl et al. 2004; Barr and Wertz 2005; Elliott 2014). 3′ and 5′ UTRs are complementary. The deletion of internal sequences in the UTRs of BUNV resulted in

attenuation of virus replication and loss of cytopathogenicity in mammalian cell culture, but the mechanistic basis of this remains unclear [91, 102].

Figure 1.4. Genomic organization of BUNV, BATV, and NRIV. NRIV takes the L and S segments from BUNV and the M segment from BATV. The M segment transcripts are translated into a polyprotein which is split into Gn, NSm and Gc after post-translational modifications. Nucleotide lengths were taken from specific strains (GenBank accession numbers: NC_001925, NC_001926, NC_001927, JX846597, JX846596, JX846595, KC608152, KC608153 and KC608154 for BUNV-L segment, BUNV- M segment, BUNV- S segment, BATV-L segment, BATV-M segment, BATV-S segment, NRIV-L segment, NRIV M segment and NRIV-S segment respectively). ORF: open reading frame, UTR: untranslated region, NSs: non-structural protein encoded by the S segment, NSm: nonstructural protein encoded by the M segment, Gc and Gn: glycoproteins, nt; nucleotides.

Like other Orthobunyaviruses, Bunyamwera serogroup viruses are able to increase genetic and phenotypic diversity through reassortment of genome segments during mixed infections with viruses of the same group (Borucki et al. 1999; van Knippenberg and Elliott

2015). For example, reassortment between Bunyamwera, Maguari, Batai, and Northway viruses were obtained in cell cultures (Weber et al. 2002). NRIV is a naturally occurring reassortment resulting from a BUNV and BATV co-infection, though it is unknown whether 23

this co-infection was in the vector or a vertebrate host (Zeller et al. 1996; Gerrard et al.

2004). The L and S segments of NRIV are from BUNV and the M segment from BATV

(SBUNV, MBATV, LBUNV) (Zeller et al. 1996). There are 93%, 97–98%, and 89–95% nucleotide sequence homologies between BUNV and NRIV S segments; BUNV and

NRIV L segments; and between BATV and NRIV M segments respectively (Gerrard et al.

2004; Briese et al. 2006). Little is known about the roles of different factors related to the occurrence of the reassortment events, such as host species, viral titers, and viral strains.

Indeed, the lack of understanding of the genomic variability of these viruses, especially

NRIV, means that we are unable to entirely understand the phylogenetic relationships among species and strains of species, and thus we cannot fully appreciate the processes that promote reassortment and the extent of propagation of such strains. Similar challenges exist for related Bunyaviruses, such as Nairovirus, and recent efforts have established new species and that there is some plasticity to classification of Bunyaviruses related in large part to the lack of comprehensive sequence data (Elliott 2014).

c. Replication in Host Cells

Bunyaviridae are enveloped viruses that replicate in the cytoplasm of mammalian cells and bud into the Golgi apparatus (Elliott 1985; Weber et al. 2002). Host cell entrance is by clathrin-dependent endocytosis and vacuolar acidification, though the receptors, cellular factors, and specific pathways are not well understood (Jin et al. 2002; Simon,

Johansson, and Mirazimi 2009; Odhiambo, Venter, Limbaso, et al. 2014). The two envelope glycoproteins Gc and Gn are responsible for viral attachment and acid-activated penetration (Albornoz et al. 2016). It has been reported that the entry of Germiston and

LaCrosse viruses, two other Orthobunyaviruses, is promoted by DC-SIGN (Dendritic cell-

specific intracellular adhesion molecule-3-grabbing non-integrin) receptors, which is seen in other Bunyaviruses, like the Phleboviruses (Phoenix et al. 2016). These receptors are found in dermal dendritic cells and may play a role in viral entry into mammalian cells following transmission by an arthropod bite (Albornoz et al. 2016; Phoenix et al. 2016).

As with many arboviruses, in vitro infection is generally lytic in mammalian cell lines, while these viruses produce a persistent infection in mosquito cells with a lack of cytopathic effect (Newton, Short, and Dalgarno 1981; Bird and McElroy 2016). In vitro infection of

Aedes aegypti cells with BUNV revealed that NSs are essential for viral replication, as these cells were refractory to infection with a recombinant NSs gene deletion strain

(rBUNdelNSs2) (Newton, Short, and Dalgarno 1981; Bird and McElroy 2016). Further infections of Aedes albopictus cell lines with both a wild-type BUNV and another recombinant NSs gene deletion strain showed that the wild-type strain demonstrated greater fitness with 100-fold higher titers (Szemiel, Failloux, and Elliott 2012). In mammalian cell lines, NSs are also implicated in pathogenesis, as NSs induced shut-off of host protein synthesis and resulted in cell death (Nashed, Olson, and el-Tigani 1993;

Szemiel, Failloux, and Elliott 2012; Tauro et al. 2015).

Infection of BHK cells first showed cytopathic effect at 3 h post- infection. Using Vero cells, most BUNV group members produce clear plaques within 4 days (Groseth et al.

2012). BUNV isolates were shown to grow to higher titers than NRIV isolates (Odhiambo,

Venter, Limbaso, et al. 2014). Additionally, after serial passage on Vero cells, NRIV was found to have total conservation of N and NSs proteins, while nucleotide substitutions were observed in both the S and M segments of subsequent passages of BUNV, which

may suggest that the NRIV genome is more stable in Vero cells than its parent BUNV

(Odhiambo, Venter, Limbaso, et al. 2014).

Persistent infection in C6/36 cells is not associated with cytopathic effects and can last indefinitely. Persistent infection in mammalian cells or long-term infection in mammalian cells or culture media without cells has been noted in other Bunyaviridae viruses but has not been reported for Orthobunyaviruses (Elliott 1985, Hardestam et al.

2007).

1.3.3. Transmission and Epidemiology

Bunyamwera virus is likely maintained in nature by blood- feeding mosquitoes and susceptible vertebrate hosts. Evidence suggests the Ae. aegypti might be the primary mosquito vector (Odhiambo, Venter, Chepkorir, et al. 2014; Tauro et al. 2015).

Experimental studies showed that Ae. aegypti was competent to transmit BUNV, but not a competent vector for NRIV. However, Anopheles gambiae Giles was competent for both viruses, while Culex quinquefasciatus did not demonstrate competence to transmit either of the two viruses (Odhiambo, Venter, Chepkorir, et al. 2014). The two identified NRIV outbreaks of human febrile illnesses (Sudan in 1988 and Somalia- Kenya in 1997–1998) coincided with episodes of unusually heavy rains and extensive flooding in areas normally arid (Briese et al. 2006). This seasonal emergence pattern resembles that of RVFV, the primary vector of which is Ae. aegypti (Pepin et al. 2010; Nanyingi et al. 2015).

Ngari virus has been isolated in many mosquito vectors, such as Aedes argenteopunctatus, Aedes minutus, Aedes vexans, Aedes mcintoshi, Anopheles coustani, Aedes neoafricanus, Aedes simpsoni, Aedes vittatus, Anopheles pretoriensis,

Anopheles phar-oensis, Anopheles mascarensis, Culex bitaeniorhynchus, Culex tri-

taeniorhynchus, Culex antennatus, and Culex poicilipes in Senegal during a study period from 1991–1994 (Zeller et al. 1996). These successful isolations suggest a large vector range, which could indicate the potential for widespread geographic distributions as well as potential vertebrate host ranges, given the diversity of feeding preferences of these mosquitoes.

Batai virus has been isolated from cattle and human patients (Nashed, Olson, and el-

Tigani 1993; Yanase et al. 2006; Liu et al. 2014). BATV has been isolated from several mosquito species as described above, and additionally in Anopheles claviger,

Coquillettidia richiardii, Culex pipiens s.i., Ochlerotatus punctor, Ochlerotatus Communis, and Ae. vexans (Francy et al. 1989; Hubálek 2008; Lozach et al. 2010). BATV has also been shown to be transmitted by biting midges (Culicoides spp.) and ticks (Dilcher et al.

2013).

Because the risk for BATV transmission has been correlated with migratory and resident bird population distributions, BATV is thought to be associated with bird-mosquito enzootic cycle similar to WNV, Usutu virus, and Sindbis virus (Hubálek 1996; Lozach et al. 2010).

Indeed, the migratory patterns of competent bird species likely accounts for its wide geographic distribution across Europe and Asia (Hubálek 2008, Hofmann et al. 2015).

BATV infection is less severe in humans, though it occurred epidemically in Scandinavia in the 1960s. In addition, neutralizing antibodies have been detected in cows and one farmer on coastal farms in Finland (Brummer-Korvenkontio 1973; Brummer-Korvenkontio and Saikku 1975; Medlock, Snow, and Leach 2007).

1.3.4. Associated diseases

All three viruses (BUNV, NRIV, and BATV) have been reported to infect humans and other mammals. BUNV causes mild symptoms in humans, such as fever, headache, joint pain, and rash. Symptomatic infections are more often reported in children, and immunocompromised patients may progress to severe encephalitis (Ashford 2001). In domestic animals, especially ruminants, infection leads to severe symptoms, such as spontaneous abortion and teratogenic effects (Rodrigues Hoffmann et al. 2013). In 2013,

BUNV was isolated for the first time in horses in Argentina after two horses developed neurological signs and died (Tauro et al. 2015). CVV is endemic to North America and causes very severe diseases in humans and ruminants characterized by embryonic and fetal death, stillbirths, and multiple congenital malformations especially in ruminants.

Humans are rarely infected by CVV, however, severe headache, nausea, vomiting, fatigue, encephalitis, and multiorgan failure have been reported in patients (Tauro,

Campbell et al. 2006; Almeida, and Contigiani 2009; Armstrong, Andreadis, and

Anderson 2015).

BATV infections in livestock results in abortions, premature births, and congenital defects (Yanase et al. 2006; Dilcher et al. 2013; Rodrigues Hoffmann et al. 2013). In humans, BATV infection has been characterized as a non-descript febrile illness in Africa and Asia and associated with an influenza-like illness in Europe (Lozach et al. 2010).

Febrile disease, bronchopneumonia, exudative pleurisy, catarrhal and follicular tonsillitis, and acute gastritis have all been reported clinical signs associated with BATV infection in humans (Sluka 1969; Medlock, Snow, and Leach 2007).

NRIV is reported to cause severe and fatal hemorrhagic fever (Gerrard et al. 2004).

NRIV clinical manifestations are similar to those of RVFV, as seen in humans in the Kenya and Somalia outbreaks (1998–1999) and in animals in the Mauritania outbreak (2010)

(Bowen et al. 2001; Jackel et al. 2013). NRIV has also been isolated from small ruminants with hemorrhagic manifestations (Jackel et al. 2013).

1.3.5. One Health implications

While BATV is less associated with severe human disease, it can lead to significant economic losses as it infects agriculturally important mammals and bird species (Sluka

1969; Yanase et al. 2006; Medlock, Snow, and Leach 2007; Rodrigues Hoffmann et al.

2013). However, given that RVFV, BUNV, and NRIV present many similarities in their clinical manifestations, co-circulate within the same vector and/or vertebrate host ranges, and share the same ecological distribution, NRIV and BUNV might contribute to outbreaks of hemorrhagic fever in these regions of both cattle and humans. Indeed, there is evidence for co-circulation in Kenya (1997–1998), where RVFV and NRIV were found in

23 and 27%, respectively of human hemorrhagic fever cases tested, which was the first report of BUNV causing human disease (Bowen et al. 2001). In Mauritania (2010), during an ongoing RVFV outbreak in livestock, 163 serum samples (62 from camels, 8 from cattle, and 93 from small ruminants) were tested for NRIV, where two goat samples were positive for NRIV (Eiden et al. 2014). In addition, one of these two samples was also IgM positive for RVFV, suggesting a recent (possibly co-) infection of RVFV (Eiden et al.

2014). These data suggest that in areas, where RVFV is reported but not confirmed or where the etiologic agent has not been molecularly confirmed, BUNV and NRIV should be included in a differential diagnostic panel for hemorrhagic fevers of humans and

animals (Gerrard et al. 2004; Briese et al. 2006). That being said, the diagnostic capabilities available for Orthobunyaviruses in general are limited. Detection of nucleic acid has been utilized and is capable of differentiating between BATV, NRIV, and BUNV, but requires multiple genes to be amplified. Inherent in this method is the assumption that there are no co-infections of NRIV and one of its parental viruses, as theoretically, a co- infection of BATV and BUNV would give the same PCR read-out as NRIV. Thus, serological assays are needed to detect circulation of these viruses. Cross-neutralization among these viruses is not limited to these three, and traditional PRNT methods are likely to be unable to distinguish to the species level (Hunt and Calisher 1979). In addition, there is a commercially available nucleotide and antibody bundle that was developed for Cache

Valley Fever virus (Discovery Antibodies, UK), a strain of BUNV, which purports to cross- react with BATV. Thus, while diagnostic capability for these three Orthobunyaviruses exists, whether or not there is an ability to definitively identify which virus is the agent remains unclear.

In addition to diagnostic capabilities, there is a better need to understand the control and prevention of transmission. A first step in this process is identifying the mosquito vectors of medical and veterinary importance. To that point, the geographic distribution(s) of these three viruses should be better characterized in effort to accurately assess which mosquito vectors could transmit. Additionally, comparative vector competence studies should be conducted in efforts to elucidate the relative transmission potential of these three viruses. Although BATV is mainly defined as European/Asian virus and has only once been isolated in Africa, it may be present in several other countries in Africa, but underreported, since NRIV has only been isolated in Africa and BATV is its presumptive

parent. After identification of likely vectors, appropriate control methods can be implemented, such as additional spraying efforts, behavior modification, and education programs. Since the life cycles of vectors are closely related to climatic conditions, the role of climate change in continued circulation, the potential for altered and expanded geographic distributions, and seasonality of transmission should be considered in all prevention campaigns.

1.4. Bunyavirus environmental suitability in Rwanda

Although important efforts have been done in inventorying arboviruses circulating in

Rwanda, its status remains less characterized. Different arboviruses such as Flaviviruses

(Yellow Fever virus, Zika virus, West Nile virus, and non-identified Flaviviruses), non- identified Alphaviruses, and Bunyaviruses (Rift Valley Fever virus) have been found to circulate in Rwanda (Demanou 2011; Kading et al. 2013; Rwanda Agriculture Board 2013;

Rwanda Ministry of Health 2017; Umuhoza et al. 2017; Seruyange et al. 2018; USAID

2018). In human populations, YFV seroprevalence of 0.16% and ZIKV seroprevalence of

1.4% were reported respectively by PRNT and IgG ELISA (Demanou 2011; Seruyange et al. 2018). In wildlife, a serosurveillance on Alphaviruses and Flaviviruses conducted in the Congo basin region (including DRC, Gabon, Chad, Rwanda, and Zambia), using the

PRNT technique, reported that over 17 gorillas from Rwanda, 2 were positive to non- identified Alphavirus(es), 2 were positive to WNV, and 1 positive to non-identified

Flavivirus (es) (Kading et al. 2013). In livestock, Rift Valley Fever virus is regularly reported, surveilled, and causes significant economic losses in livestock production

(Umuhoza et al. 2017). Although RVFV is the only Bunyavirus reported in Rwanda, different factors such as environmental parameters, proximity and important human and

animal movements between countries such as Kenya and Uganda, where more

Bunyaviruses are regularly reported (Kifaro et al. 2014; Nanyingi et al. 2015; Bird and

McElroy 2016; Dutuze et al. 2018) suggest that Rwanda is exposed to Bunyavirus transmission.

Rwanda is characterized by temperate tropical climate which is known to be favorable for mosquito emergence thus making it suitable for bunyaviral and arboviral disease transmission in general (Loevinsohn 1994). There are four weather seasons in Rwanda: moderate dry season (December-February), intense rainy season (March-May), intense dry season (June-September), and moderate rainy season (October-November).

Geographically, based on rainfall, vegetation, and altitude, Rwanda is divided into 4 ecological zones with the following characteristics (Demanou 2011) (Figure 1.5).

Zone 1. Forested, high level of precipitation, altitude <2,300 m;

Zone 2. Forested, high level of precipitation, altitude >2,300 m in few areas;

Zone 3. Dry, cropland, natural vegetation, altitude <2,300 m;

Zone 4. Dry, altitude <2,300 m, savannah, grasslands, possibly some croplands.

Figure 1.5. The four ecological zones of Rwanda. ArcGIS (ESRI, Redland, CA) mapping of the four ecological zones outlined as polygons (Demanou 2011).

Mosquito species distribution suggests that the most common mosquito species in the descending order of frequency are Culex sp, Aedes sp, Coquillitedia sp, Mansonia sp, and Anopheles sp. All of these species are found in high proportions in zone 4 compared to the other zones (Table 1.2) (Demanou 2011).

Table 1.2. Number of mosquito species collected by ecological zones during YFV risk assessment in Rwanda in November-December 2012 (Demanou 2011)

Thus, zone 4 is more suitable for mosquito-borne diseases. By transposition, this zone corresponds to the Eastern province which is characterized by the highest malaria prevalence in human population (24% in children under 5 and 12% in individuals older than age 15) (Rwanda Ministry of Health 2017; USAID 2018). The transmission is increased during the end of the most intense rainy season and the beginning of the intense dry season (May-July) as this period is characterized by wet and hot conditions associated with high mosquito emergence (Rwanda Ministry of Health 2017; Umuhoza et al. 2017; USAID 2018).

In livestock, for a long time, periods after heavy rain and flooding have been characterized by tick-borne diseases such as Theileriosis, Babesiosis, and Anaplasmosis

(Rwanda Agriculture Board 2013). Although in neighboring countries like Uganda,

Tanzania, and Kenya several Bunyaviruses such as RVFV, BUNV, Ilesha, Germiston

(Kifaro et al. 2014; Nanyingi et al. 2015; Bird and McElroy 2016; Dutuze et al. 2018) have been reported and regularly emerge during this period, none of these were reported in

Rwanda before 2011. The first Bunyavirus to be reported in Rwanda was RVFV in May

2011. This occurred in a cattle farm in Bugesera (zone 4 on Figure 1.5) during unusually massive abortions in livestock (Rwanda Agriculture Board 2013).

It is speculated that RVFV could have been circulating before under less severe and unrecognized clinical manifestation. This assumption is also built on the fact that the main clinical sign of RVF in livestock is abortion which is shared between other common animal diseases found in Rwanda such as Brucellosis and Theileriosis (Rwanda Agriculture

Board 2013). Since RVFV was first confirmed, sporadic cases are reported countrywide with more cases from ecological zone 4 and vaccination campaigns are regularly conducted with emphasis on farms located in this zone. Unfortunately, due to the insufficiency of laboratory facilities, the diagnosis is mainly clinically or serologically based after few molecular confirmations. The identification of RVFV in 2011 showed that

Rwanda constitutes a suitable environment for Bunyavirus transmission, thus other

Bunyaviruses such as BUNV and NRIV found in neighboring countries might be circulating under silent forms.

1.5. Rationale, hypothesis, and objectives

The genetic relationship between BUNV, BATV, and NRIV viruses raises different research questions on the potential parallelism in infection kinetics and transmission dynamics of these viruses. In Rwanda, although RVFV is the only Bunyavirus commonly reported, given that its diagnosis is mainly based on clinical signs which are common to a large number of bunyaviral diseases, the suitable environmental conditions, and the presence of many Bunyaviruses in the neighboring countries, it seems legitimate to

speculate that other Bunyaviruses such as BUNV and NRIV (and probably BATV) circulate in the country.

For the present research project, we hypothesize that given their genetic relationship, infection kinetics of BUNV, BATV, and NRIV might be related. Further, we hypothesized that these Orthobunyaviruses circulate in Rwanda and cause infections that are misclassified as RVFV cases. We decided to test this through the following objectives:

a) Comparatively characterize in vitro infections of BUNV, BATV, and NRIV. This is

addressed in chapter 2 where we offer the comparison of dose-dependent effects

on infection of the three viruses in Vero cells as well as their capabilities to stay

stable after cell death.

b) Comparatively study in vivo infections of BUNV, BATV, and NRIV, and investigate

their potential cross-neutralization. This is covered in Chapter 3 where we

compared the viremia caused by the three viruses in C57Bl/6 mice and quantified

their capabilities to cross-neutralize each other. In addition, we studied the role IRF

3 and IRF 7 in BUNV infection by investigating its pathogenesis IRF 3/7-/--/- mice.

This is was done in order to create a disease progression model for many

Bunyaviruses presenting similar disease manifestations as BUNV.

c) Investigate the presence of BUNV, BATV, and NRIV in Rwanda. For the first time,

we offer the diagnosis for Bunyaviruses other than RVFV in Rwanda.

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CHAPTER 2. COMPARISON OF IN VITRO INFECTION KINETICS AND EX VIVO STABILITY OF BUNYAMWERA, BATAI, AND NGARI VIRUSES

2.1. Introduction

Bunyamwera virus (BUNV), Batai virus (BATV), and Ngari virus (NRIV) are mosquito- borne viruses of the Orthobunyavirus genus in the Peribunyaviridae family, one of the ten virus families comprising the Bunyavirales order (Bowen et al. 2001; Gerrard et al. 2004;

Briese et al. 2006; Groseth et al. 2012; Tauro et al. 2015; King et al. 2018). As with all members of the Peribunyaviridae family, BUNV, BATV, and NRIV are enveloped viruses with single-stranded RNA genome made up with three segments, denoted as Small (S),

Medium (M) and Large (L). The S segment encodes the nucleocapsid, the M segment encodes envelope glycoproteins, and the L segment encodes the polymerase protein (,

Fuller and Bishop 1982; Bowen et al. 2001; Gerrard et al. 2004; Groseth, Weisend, and

Ebihara 2012). NRIV is a natural reassortant of BUNV and BATV and results from a combination of the BUNV S and L segments and the BATV M segment (Gerrard et al.

2004; Briese et al. 2006; Odhiambo et al. 2014). The relationship of these “parental” viruses and the “progeny” poses an interesting question, especially given that there is overlap in their respective transmission and infection ecologies. The existing knowledge regarding the comparison and contrast of these viruses and what is currently known and unknown is reviewed in Chapter 1 (Dutuze et al. 2018), and briefly summarized below.

BUNV, the prototype of the Peribunyaviridae family and Bunyavirales order, is the most characterized of the three, followed by BATV, and lastly NRIV (Briese et al. 2006;

Medlock, Snow, and Leach 2007; Szemiel, Failloux, and Elliott 2012; Riblett and Doms

2016; Dutuze et al. 2018). Investigations of NRIV have mostly focused on the genomic

(dis)similarities with other members of the Orthobunyavirus genus and the similarity of its clinical manifestations with Rift Valley Fever virus (RVFV), an important bunyavirus of the genus Phlebovirus, and family Phenuiviridae (Bowen et al. 2001; Gerrard et al. 2004;

Briese et al. 2006; Jackel et al. 2013, Burleson 2014).

These three Orthobunyaviruses cause disease in domestic ruminants and humans, with NRIV being epidemiologically associated with hemorrhagic fever in humans in East

Africa (Kenya, Somalia, and Sudan) (Bowen et al. 2001; Gerrard et al. 2004; Briese et al.

2006; Odhiambo et al. 2014). BATV has been isolated across Europe, Asia and Africa, while BUNV and NRIV remain sporadically isolated in Africa (Smithburn, Haddow, and

Mahaffy 1946; Singh 1966, Gaidamovich et al. 1973, Nashed, Olson, and el-Tigani 1993;

Zeller et al. 1996; Lambert and Lanciotti 2009; Eiden et al. 2014; Liu et al. 2014; Dutuze et al. 2018). Despite the potential for one or all these viruses to present an emergent public health threat, they remain relatively under characterized. There have been no detections of BUNV and BATV concomitantly, though NRIV itself is evidence that this event must have occurred. BATV was molecularly identified in Uganda in 1967 (East

African Research Institute 1967), highly suspected in Sudan based on serological test in

1988 (Nashed, Olson, and el-Tigani 1993), but otherwise has not been detected in Africa since. BUNV and NRIV have been identified in many African countries between 1943 and

2015 (shown by Figure 1.2. and extensively reviewed in Chapter 1 (Dutuze et al. 2018).

Importantly, though these viruses are linked genetically and ecologically, there have been no studies that comparatively characterize the three viruses, especially given the differences in associated pathogenicity of NRIV compared to BUNV and BATV. Thus, we compared these three viruses in vitro, reporting growth curves in a commonly used

cell culture (Vero cells), designed traditional and qRT-PCR assays and report specificity and sensitivity of such, and describe differences of the in vitro phenotype (plaque morphologies).

Generally, growth curves of viruses in vertebrate cells consist of four phases, which are a) adsorption and penetration, b) eclipse, c) maturation, and d) release (Burleson

2014) (Figure 2.1). Adsorption, penetration and eclipse correspond exclusively with the intracellular phase before the virus starts being detected in both intracellular and extracellular medium. On the curve, these stages are shown by no to low detection before the ascendant phase, which corresponds to maturation. Release is translated by descendant phase on the curve and is followed by no detection of viable virus. This is due to the fact that cells become metabolically and structurally incapable of supporting additional replication after the virus production has reached the plateau (Flint 2000).

Figure 2.1. General viral growth curve in mammalian cell

Preliminary characterization and comparison of these viruses in cell culture is a necessary first step. Specifically, these phases of the growth curve can provide a means of laboratory standardization for growth of virus to match titers, and for general use in experimentation. Additionally, comparative kinetics can give preliminary insights of important epidemiological concepts such as relative incubation and infectious periods, especially when comparing across species of viruses where in vivo kinetics are unknown

(Yin and Redovich 2018). Subsequently, these parameters are useful in estimations of factors important for adequate control measures such as determination of adequate quarantine periods (Farewell et al. 2005; Nishiura 2007; Chan and Johansson 2012), the window of efficacious administration of antiviral medication (Lessler et al. 2009), the identification of potential sources of infection in a population (Lessler et al. 2009), and establishment of accurate transmission models (Chan and Johansson 2012).

Long-term stability has been studied in members of several families of the

Bunyavirales order such as Phenuiviridae, Nairoviridae and Hantaaviridae (Southam

1954; Hardestam et al. 2007). In general, these viruses can persist for long periods in less than ideal conditions (such as cell-free conditions). Only one study in 1954 investigated the ex vivo stability of a Peribunyavirus, where BUNV was reported to quickly disappear from detection (using mouse infectivity titration), though it had been persistently infecting human lymph nodes in cell culture (Southam 1954). There is no information on any of the three viruses of interest here on long-term stability in Vero cells – a commonly used laboratory cell line that is easy to culture and widely used in arbovirology. The knowledge on the duration of infectiousness of virus in extracellular medium provides a foundation for establishment of biosafety measures regarding the handling and safe-

storage of these viruses, as well as inactivation methods. This work provides the first comparative study of in vitro properties of BUNV, BATV, and NRIV.

2.2. Material and methods

2.2.1. Viruses, plaque assay, and comparative plaque morphology

6547-8, MM2222, and DAK-AR D28542 are the strain designations of BUNV, BATV, and NRIV respectively used throughout this study. 6547-8 is the prototype strain of BUNV isolated from Aedes spp. mosquitoes in Uganda in 1943 (GenBank accession numbers:

X14383, M11852, and X73465). MM2222 is the prototype of BATV isolated from Culex spp. mosquitoes in Malaysia in 1955 (GenBank accession numbers: AB257766,

JX8446595-97, and X73464). DAK-AR D28542 strain of NRIV was isolated from Aedes spp. male mosquitoes in Senegal in 1985 (GenBank accession number: AY593728-29).

The viruses were obtained from the World Reference Center for Emerging Viruses and

Arboviruses at the University of Texas Medical Branch (UTMB). All viruses were provided in lyophilized form and were reconstituted by adding 1 mL of cell culture media to the vials. The passage histories at the time they were received were Suckling mice (SM) 47/

Vero 2, SM3/Vero 2, and SM 4/Baby Hamster Kidney cells (BHK) 1/Vero 2 for BUNV,

BATV, and NRIV, respectively. After reconstitution, the viruses were cultured on Vero

African green monkey kidney cells (ATCC) and stocks were collected at 2 dpi, 5 dpi, and

3 dpi for BUNV, BATV, and NRIV respectively, after observation of cytopathic effects

(CPE). Viral titers were then determined by standard plaque assay techniques. The calculated titers were 6.65 x 106PFU/mL at 3 dpi, 2.27 x 107 PFU/mL at 5 dpi, and 5.7 x

107 PFU/mL at 4 dpi for BUNV, BATV, and NRIV, respectively. Pictures of the plaques of each virus were taken in a 6-well plate with a ruler on the side. Plaques were

phenotypically compared and a random sample of 20 plaques per virus were measured for diameter. BUNV and BATV are biosafety level 2 (BSL-2) viruses while NRIV is a BSL-

3 virus (CDC, 2019). To control for variation due to environmental conditions, all experiments were conducted with the LSU School of Veterinary Medicine, Department of

Pathobiological Sciences BSL-3 laboratory.

2.2.2. In vitro growth kinetics and ex vivo stability

a. Comparative growth curves at a Multiplicity of Infection (MOI) of 1

We first studied comparative growth curves of BUNV, BATV, and NRIV grown in Vero cells under standard conditions at a MOI of 1. For that, Vero African green monkey kidney cells (ATCC) were grown in optimal medium (88% Medium 199, Earle's Salts, 10% Fetal

Bovine Serum, and 2% Antibiotic-Antimycotic) in T75 flasks at 37˚C, 5% CO2. At 85-90% confluence, cells were trypsinized and counted using a hemocytometer after being stained by Trypan Blue (Louis 2011). Cells were then seeded in 6-well plates with a count of 1.02 x 106 cells in 2 mL of medium per well. After being observed every hour, they were found to be fully adhered to the flask bottom after 6 hours. Growth media was removed, and each well was inoculated with an appropriate volume of each virus calculated to have

1.02 x 106 PFU equivalent to a MOI = 1 (Table 2.1). The infections were done in triplicates

o per virus. Plates were rocked for 30 mins and incubated at 37 C, 5% CO2 for 30 minutes before 2 mL of new media was added to each well. Supernatants were initially collected from 1 to 7 dpi and stored at -80oC until RNA extraction and amplification. Additional collection at 10 dpi was done after high concentrations of viral RNA were detected at 7dpi.

Table 2.1. Volume calculations for BUNV, BATV, and NRIV for 1.02 x 106 PFU considering their titers

Virus Volume for 1.02 x 106 PFU

BUNV 152µL (titer = 6.65 x 106PFU/mL)

BATV 44.7µL (titer = 2.27 x 107PFU/mL)

NRIV 17.7µL (titer = 5.7 x 107PFU/mL)

b. Dose dependence in vitro growth kinetics under standard growth conditions

Given that mosquitoes inoculate various titers of arboviruses depending on a multitude of factors, we wanted to characterize and compare the dose-dependence of growth kinetics of BUNV, BATV, and NRIV in Vero cells (Styer et al. 2007; Secundino et al. 2017;

Talavera et al. 2018). For that, 85-90% confluent 6 well plates of Vero cells grown in optimal media (88% Medium 199, Earle's Salts,10% Fetal Bovine Serum and 2%

Antibiotic-Antimycotic), and subsequently infected with serially diluted BUNV, BATV, and

NRIV from 106 PFU/mL to 101 PFU/mL. Prior to infection, media was removed from each well (of 6-well tissue culture plates) and 100 µL of each dilution for each of three viruses was inoculated onto individual wells. The wells were then rocked for 30 min and incubated

o for 30 min at 37 C, 5% CO2. After incubation, 2 mL of new media was added to each well

o before incubation again at 37 C, 5% CO2. Negative control wells with M199E medium were prepared following the same protocol. 100 µL of supernatant was collected from each well at 1, 3, 5, 7, 14, and 30 dpi and stored in -80oC until RNA extraction and amplification.

c. Stability in extracellular and cell-free media

Given the previous investigation of BUNV extracellular stability by mouse infectivity titration we wanted to investigate whether BATV or NRIV showed differential stability and further wanted to determine if stability could be assessed in a more accessible method since these viruses often affect resource-poor regions (Southam 1954). We investigated stability by looking at 1) the dose-dependent effect on growth kinetics in Vero cell culture at less than optimal conditions (2% rather than 10% FBS), 2) the association between

CPE occurrence and viral RNA detection, and 3) long-term stability in cell-free media. For that, Vero cells were grown in nutrient reduced medium (96%Medium 199, Earle's Salts,

2% Fetal Bovine Serum, and 2% Antibiotic-Antimycotic) at 37˚C in 5% CO2. This is because with optimal nutrition (10%FBS) cell growth can out-pace cell death, leading to persistence in cell culture as there are continually more cells to infect. At lower cell nutrition availability (2%FBS), cell growth is slowed and thus, when compared to the growth in 10%FBS (see 2.2.2.b), we wanted to determine whether this is persistent infection or simply viral stability.

After serial dilutions (106 PFU/mL to 101 PFU/mL) of each virus, 85-90% confluent 6 well plates were infected with 100 µL of each dilution for each virus using the protocol previously described. 100 µL of supernatant was collected from each well at 1, 3, 5, 7,

14, and 30 dpi and stored in -80oC until RNA extraction and amplification. In addition, cytopathic effect (CPE) was observed under inverted microscope at 20X at various dpi and pictured with the DINOEYE capture function. Different qualitative scores were given to each well at 1-14 and 30 dpi according to the relative proportion of dead cells compared to healthy cells. The scores and description of corresponding CPE are given in Table 2.2.

Table 2.2. Scores attributed to Cytopathic Effect (CPE)

CPE Score Corresponding cell culture description

- No observed CPE

+ Few dead cells

++ Approximately 1/3 cells are dead

+++ Approximately 1/2 cells are dead

++++ Approximately 2/3 cells are dead

+++++ Very few cells still attached as monolayer

++++++ All cells dead (none attached)

To investigate the stability in cell-free media, 100 µL of the each of the three viruses was inoculated into 2 mL of cell-free M199 1X media in 2 mL Eppendorf tubes. As previously mentioned, the titers were 6.65 x 106PFU/mL, 2.26 x 107 PFU/mL, and 5.7 x

107 PFU/mL for BUNV, BATV, and NRIV, respectively. The tubes were incubated at 37oC,

5% CO2 and 100 µL of supernatant was collected from each tube at 1, 3, 5, 7, 14, and 30 dpi and kept in -80oC until viral RNA extraction and amplification. This experiment was also run in triplicates.

d. Comparative inactivation by Triton-X-100

Given the potential for long-term stability of these viruses, we tested the ability of

Triton-X-100 – a commonly used detergent used in the development of ELISAs – to inactivate BUNV, BATV, and NRIV. This was done for potential use of these viruses in the development of serological assays using less biosafety containment facilities especially in developing countries where high containment facilities are unavailable. For that we investigated comparative inactivation of these viruses by Triton-X-100 as this

detergent has been used to inactivate other enveloped viruses such as Ebola virus

(Colavita et al. 2017) and influenza (Jonges et al. 2010). 1% Triton-X-100 was added to the stock of each of the three viruses (5 µL of Triton-X 100 in 495µL viral stock) and the mixture was incubated at room temperature for 1 hour and kept in -80oC until further use.

To test the efficacy of Triton-X-100 inactivation capacity on these viruses, infectivity of treated viruses was investigated via CPE appreciation and RNA detection after the supposedly inactivated viruses were inoculated on Vero cells. Prior to infections, Triton-

X-100 was removed using the column-based absorption Detergent OUT kit according to manufacturer instructions (Millipore sigma, Cat No. 2114). 85-90% confluent 6 well plates of Vero cells were grown in optimal conditions (10%FBS) were used. After removal of the

2 mL growth medium, 100 µL of each virus was inoculated in a well. Plates were then

o rocked for 30 mins and incubated at 37 C, 5%CO2 for 30 minutes before addition of 2 mL of new media. Presence or absence of CPE was observed and 100 µL of the supernatant was collected every day from 1-7 dpi, and RNA extraction and amplification were performed as described below. This experiment was also run in triplicates.

e. Determination of viral infectivity after in vitro and ex vivo persistence

Stability (long-term infectivity) of these viruses was investigated by inoculating supernatants collected at 30 dpi on newly grown Vero cells with optimal medium concentration. Samples from both the 10% and 2% dose-dependence growth studies that were collected at 30 dpi were inoculated onto Vero cells. We tested only the viral doses of 106PFU/mL, 104PFU/mL, and 101PFU/mL of each of the three viruses. 100 µL of 10- fold dilution of the supernatants from each %FBS-dose-replicate were inoculated on 85-

90% confluent 6 well plates of Vero cells. Equivalent experiments were performed for

supernatants collected at 30 dpi from tubes with cell-free media initially infected with viral stocks. Collection of these samples was done at 1, 3, 5, and 7 dpi followed by RNA extraction and amplification. This experiment was also run in triplicates.

2.2.3. Viral RNA detection, qRT-PCR and PCR assay design, and sequencing

RNA was extracted using the KingfisherTM (Thermo-Fisher) automated extraction platform according to the instructions of manufacturer. A total of 50µL of RNA was collected for each sample and stored in -80oC until further use.

Both qRT-PCR and traditional PCR primer sets were designed for these viruses for the M and L segments. This was done because among these Orthobunyaviruses, these two segments can be used to identify BUNV, BATV, and NRIV. Additionally, for complete confirmation and exclusion any potential reassortment event in diagnostic process, S segment primers were designed for traditional PCR. We could not design effective S segment primers for qRT-PCR.

qRT-PCR primers and probes targeting conserved regions of the M and L segments of BUNV, BATV, and NRIV were designed using www.idt.dna PrimerQuest tool (Table

2.3). Prior to being used for the experiments, they were tested on viral stocks for sensitivity and specificity with standard curves generated while titer was determined by plaque assay as described above. Nucleic acid amplification was performed by qRT-PCR using the SuperScript™ III One-Step RT-PCR System with Platinum® Taq DNA

Polymerase (Invitrogen, Cat No. 1.1732-088). A PCR mixture of 10µL 2X RT buffer, 1µL

Forward primer (10 µM), 1µL Reverse primer (10 µM), 0.4 µL Probe (20 µM), 0.4 µL SIII

Taq, 1.4 µL ddH2O, 0.8 µL MgSO4 and 5µL RNA was used on 96-well PCR plates. After being loaded, plates were centrifuged for 2 mins at 2000 rpm prior to actual amplification

on a Roche Lightcycler® 480 II. The following thermal profile as a cycle of reverse transcription for 10 min at 50°C, 15 min at 95°C for reverse transcriptase inactivation and

DNA polymerase activation followed by 40 amplification cycles of 15 sec at 95°C

(annealing-extension step) and 30 sec min 40°C (cooling). These cycling conditions were already used in our laboratory to amplify Zika virus (Faye et al. 2013; Kawiecki et al.

2017). The data were analyzed using Lightcycler® 480 Software.

Table 2.3. Sequences of primers and probes used for qRT-PCR

Virus and segment Primer/probe Sequence GenBank targeted accession No BUNV- M Forward primer 5’-GCT TAT GGA TGG GCG TAC AA-3’ M11852 Reverse primer 5’-GGA GCC ACA GAC ACA ATA TGA-3’ Probe 5’-/5Cy5/ATG CAC TTG CGG ATT GGC AT/3lAbRQSp/-3’ BUNV-L Forward primer 5’-GCC ACT TTG CTG ATT CCT TTG -3’ X14383 Reverse primer 5’-CTA ACC TTG TAG TGC TGG CTA ATA -3’ Probe 5’-/56-FAM/TGG AAG AGG/ ZEN/CAA GCA GAT TGA GCT/3lABkFQ/-3’ BATV-M Forward primer 5’-GCA TGT GGA AAC TCA CCA AAT TA-3’ JX846596.1 Reverse primer 5’-ATT CTT GTG AGG CAG GGA TTA G -3’ Probe 5’-/5Cy5/AAG GGA GAA GTG TGG TGT TCA GGT/3BHQ_2/-3’ BATV-L Forward primer 5’-CAC TCT ACC AGC TGC ATT CTA C-3’ JX846597.1 Reverse primer 5’-GTT GAC CAC GGT TCA CTA CTT-3’ Probe 5’-/56-FAM/ACAGCTGCA /ZEN/GGG ATAATTAACTGG ACC/3lABkFQ/-3’ NRIV-M Forward primer 5’-TAT AGG CCC TTT ACA GCA AGT G-3’ KC608153 Reverse primer 5’-GCT GCA TCC AGG TCT GAT ATT-3’ Probe 5’-/5Cy5/ACA TGC GAC GAT AAA GCA AGC AGA/3lAbRQSp/-3’ NRIV-L Forward primer 5’-GCG AAA CCG TGT AGA AAG TAG A -3’ KC608152 Reverse primer 5’- CCC TGA AAT CAC CGA CCT TTA T -3’ Probe 5’-/56-FAM/AGCTTGTGA/ZEN/AAG TGC TTA TTG TTG TGA TGC/3lABkRQ/-3’

Conventional PCR primers were designed using www.idt.dna PrimerQuest tool as well (Table 2.4). We endeavored to amplify all three gene segments for future potential use in sequencing. From RNA extracted as previously described, cDNA was synthesized using Superscript III First-Strand kit (Invitrogen, Cat No. 18080-051). A mix of 8µL RNA,

1µL of Reverse primer (2µM) and 1 µL dNTPs was done in 200 µL tubes and run on

o thermocycler 65 C for 5 min after what 2µL 10X RT Buffer, 4µL 25m MgCl2, 2µL 0.1mm

DTT, 1µL RNase OUT and 1µL Superscript III RT was added before running it back to the thermocycler for 50oC for 50 min and 85oC for 5 min. 1µL of RNase H was then added to the tube and put back to the thermocycler again at 37oC for 20min. A total of amount

21µL cDNA collected and kept at -20oC until further use. cDNA was amplified in 25 µL

PCR mix consisting of 2.5 µL 10X PCR Buffer, 1.5 µL 50mM MgCl2, 0.55 µL 10mM dNTP,

2 µL Forward primer, 2 µL Reverse primer, 0.1 µL Taq, 15.35 µL ddH2O, and 1µL cDNA.

The amplification on thermocycler was performed with the following cycling parameters:

95oC for 15 min; 35 cycles of 94oC for 30 sec, 54oC for 30 sec, 72oC for 1 min and 72oC for 10 min and 10oC forever. The amplicons were stained by GelRed (Biotum, Cat No.

41003), run on 2% agarose gel with 1X TAE buffer at 100V for 1 hour, and visualized with

Bio View UV Light transilluminator.

Table 2.4. Primers for conventional PCR

Virus and segment Primers Primer sequence Amplicon GenBank targeted size accession No

BUNV-S Forward primer 5’-CTT CCC AGG ATC AGA GAT GTT-3’ 395bp D00353.1 Reverse primer 5’-ATT TAG CCC GCT GTC TTT CT-3’ BUNV- M Forward primer 5’-GCT TAT GGA TGG GCG TAC AA-3’ 651bp NC001926 Reverse primer 5’-GAT GCC TCT GAC CCA GTT AAT-3’ BUNV- L Forward primer 5’-AAT CCA GAG GCC CTA GGT ATA A-3’ 767bp NC001925 Reverse primer 5’-GAC CAA GGC TCT TGC TCT ATC- 3’ BATV-S Forward primer 5’-GAC CCA GAG GTT GCA TAC ATT A-3’ 394bp JX846595.1 Reverse primer 5’-CAG ACC CTG GAA AGA ATG AGA G-3’ BATV- M Forward primer 5’-GAA CTA ATC CCT GCC TCA CAA G-3’ 847bp JX846596.1 Reverse primer 5’-GAC ACT CTC CTC AAC TGC TTT-3’ BATV-L Forward primer 5’- TGT ATC ATC ACC CGG CTT ATT C-3’ 798bp JX846597 Reverse primer 5’-CAG AGG GTT TGA CAG CCT ATA TT-3’ NRIV-S Forward primer 5’-TGC TAA CAC CAG TAC TTT-3’ 504bp JX857325.1 Reverse primer 5’-ACC TCT GTC GCA TTG TCT TT- 3’ NRIV-M Forward primer 5’-TGG GTG CCT TGC TGT AAA TA - 3’ 914bp KC608153 Reverse primer 5’-CAC TGA TCC CTT CAT CCC TAA C- 3’ NRIV-L Forward primer 5’-AGC CAG CAC TAC AAG ATT AGA- 3’ 785bp KC608152 Reverse primer 5’-TCT CCT TGC TCA TCA CCA TTA- 3’

To test the efficiency of our designed traditional PCR primers and confirm the identity of our viral stocks, we performed sequencing. This was done using Big Dye Terminor v.3.1 cycle (Applied Biosystems, Cat No.1701127) according to manufacturer’s instructions and run on the 3130 genetic analyser. Forward and reverse primers for S, M and L segments for each of the three viruses (BUNV, BATV, and NRIV) were used.

Alignments of obtained sequences were viewed using Bioedit software, USA and subjected to BLAST search in NCBI GenBank blast tool to identify similar sequences.

2.2.4. Statistics

To determine differences in plaque morphology/size, the diameters of plaques were compared between viruses by Kruskal-Wallis non-parametric t-test after determining the data were not normally distributed (Shapiro Wilk, p<0.05).

R package Growthcurver was used to analyze the growth kinetics of BUNV, BATV, and NRIV across all growth-curve experiments. Growthcurver determines area under the curve (AUC) and doubling time (DT) used to characterize growth curves. AUC is determined under the logistic function curve in forms of two metrics: 1) the logistic AUC

(AUC-L) that finds the area under the logistic function that is the best-fit to the data and

2) DT, which is the time that a population takes to double the number of individuals.

Growthcurver determines the fastest DT which occurs when the population is maximizing its growth potential (Rockwood 2015; Sprouffske and Wagner 2016). For all growth conditions and for each inoculation dose of each virus, Growthcurver analysis was performed for all replicates individually and analysis of variance (ANOVA) on AUC-L

(Appendix B) and DT values was performed to compare the growth kinetics among the three viruses and inoculation doses and different growth cell conditions. When

significance was determined, Tukey post-hoc test was performed for pairwise comparisons. Additionally, ANOVA was performed to compare DT of the curves between cell growth conditions and inoculation doses among viruses.

A paired, one-tailed t-test was performed to test the infectivity of BUNV, BATV, and

NRIV after Triton-X-100 inactivation and after stability at 30 dpi by comparing the RNA equivalent PFU/mL at day 1 dpi and days 4 or 7 dpi. We determined if the difference of

RNA detected was greater than 0 between 1 and 4 dpi for Triton-X-100 inactivated viruses and between 1 and 7 dpi for supernatant collected at 30 dpi for the in vitro and ex vivo stability experiments, then there is evidence of viral replication, indicating the presence of infectious virus.

2.3. Results 2.3.1. Plaque assay and plaque morphology

BUNV and NRIV produce a mixture of small, medium, and large sized plaques (Figure

2.2 A-1 & A-3). These plaques are harder to see, and we found that increasing the concentration of neutral red stain to 0.8mL/25mL of overlay media optimized visualization.

BATV makes well distinguishable plaques that are easily visible at lower neutral red stain concentration (0.7mL/25mL) (Figure 2.2. A-2). Using Kruskal-Wallis rank sum test, we determined that BATV produced plaques that had a tendency to be larger than either

BUNV or NRIV. The difference between BATV and BUNV-NRIV was significant by Dunn’s post hoc test, while plaque diameter was found not to be significant between NRIV and

BUNV (Figure 2.2.B).

Figure 2.2. Comparative plaque morphology for BUNV, BATV, and NRIV. A-1. BUNV at 4 dpi, A-2. BATV at 5dpi, A-3. NRIV at 4dpi. B. Comparative plaque diameters in mm. There was a significant difference between BATV and BUNV-NRIV while no significant difference was found between NRIV and BUNV.

2.3.2. Specificity and sensitivity of qRT-PCR and PCR

The designed qRT-PCR primers for each virus successfully amplified homologous self-segments. In addition, there was a successful cross-amplification of BUNV-derived

L-segment primers and the NRIV L gene, and vice versa. The cross-amplification of

BATV-derived M primers and the NRIV M segment was also successful; however, NRIV- derived M primers did not amplify the BATV M segment. For our experimental studies, since no co-infections were performed in vitro, we proceeded with using BUNV M and L primers and NRIV M and L primers for the detection of all three viruses. Standard curves generated based on titers calculated from the plaque assays for both M and L primers had a sensitivity of detection of 101 PFU/mL for all three viruses. Thus, we had a qRT-

PCR assay that was sensitive and specific given we knew the identity of the infecting virus of either NRIV or BATV.

Table 2.5. Specificity of qRT-PCR primers

Viruses

Primers BUNV BATV NRIV

BUNV M + - -

L + - +

BATV M - + +

L - + -

NRIV M - - +

L + - +

We also designed traditional primers for each of the three segments (S, M, and L) of each of the three viruses and tested for specificity. There was a successful amplification of virus segments by homologous primers, or primers derived from the matching virus

(Figure 2.3.A). Across the shared segments between BUNV and NRIV (S and L), the cross-amplification was successful as well, while the cross-amplification of the shared segment between BATV and NRIV (M) was not very successful. NRIV M primers weakly amplified the BATV M segment (Figure 2.3.C) while BATV M primers did not amplify the

NRIV M at all (Figure 2.3.B).

Figure 2.3. Specificity of designed primers for conventional PCR. A. Amplification by homologous primers. B. Comparison between amplification of NRIV genome segments by homologous primers and its parental genome segment primers, C. Comparison between amplification of BUNV (S, L) and BATV (M) genome segments by homologous primers and by NRIV primers.

An alignment of the GenBank sequences associated with these viruses after we sequenced using designed primers for traditional PCR (Table 2.4.) showed the similarity percentages displayed in Table 2.6. The low similarity percentages between BATV M and

NRIV M segments (87.23% and 89.55%) might be associated with failure of BATV M and

NRIV M primers to amplify each other.

Table 2.6. GenBank similarity percentages of BUNV, BATV, and NRIV segment alignments using the designed traditional PCR primers (BUNV: strain 6547-8, BATV:M2222 strain, and NRIV: DAK-AR-D2852)

Similarity percentages

BUNV-S BUNV-M BUNV-L BATV-S BATV-M BATV-L NRIV-S NRIV-M NRIV-L

Primer targets

BUNV-S 95.95% 91.67%

BUNV-M 99.66%

BUNV-L 99.59% 97.25%

BATV-S 99.7%

BATV-M 98.02% 87.23%

BATV-L 99.86%

NRIV-S 100% 100%

NRIV-M 89.55% 99.29%

NRIV-L 97.39% 99.79%

2.3.3. In vitro growth kinetics of BUNV, BATV, and NRIV

2.3.3.1. Growth curves under standard conditions

a. Comparative growth curves at MOI of 1

Analysis of variance of AUC-L shows that there was no difference of growth between viruses and target for RNA amplification (M and L segments). Since there was no difference between M and L primers for specificity or for apparent detection in growth curves, we proceeded with the use of M primers for the rest of our study. Figure 2.4. shows the average growth curves for each virus and log (PFU/mL) determined by qRT-

PCR for the M segment (Figure 2.4 A) and the L segment (Figure 2.4 B.). No difference was determined via ANOVA (p>0.05).

Figure 2.4. Growth curves of BUNV, BATV, and NRIV in standard cell culture condition (10%FBS), at a MOI of 1 for quantification by A. M primers B. L primers.

b. Inoculation dose dependence for growth kinetics

A MOI of 1 was equivalent to 106 PFU and may have led to early saturation of cells by viruses, thus masking a critical difference in early eclipse and maturation stages that might be dose-dependent. Thus, we decided to investigate differences due to inoculation dose. Using ANOVA, we found no significant defect of virus identity on the growth curves.

However, a significant effect of inoculation dose was found, though this was not surprising

(p<0.05). Using the Tukey post-hoc comparison, we determined the significant differences to be between 106 and 101PFU/mL (p < 0.0001), 106 and 104PFU/mL

(p=0.041), and 104 and 101PFU/mL (p=0.025). Growth curves for high initial doses peaked earlier than those for low initial doses on average by 1-2 days (Figure 2.5).

Figure 2.5. BUNV, BATV, and NRIV growth kinetics in Vero cells grown in standard conditions (10%FBS) with different inoculation doses. (106-1 PFU/mL). Error bars represent standard errors.

For doubling times, ANOVA using DT values for all replicates, each taken individually did not show any significant difference between viruses. However, there was a significant difference between initial doses. Using the Tukey post-hoc comparison, we determined the significant differences to be between 106 and 101PFU/mL (p < 0.0001), 106 and 104PFU/mL

(p=0.0408), and 104 and 101PFU/mL (p=0.025). The means of DT values and peak days per inoculation dose for each virus are shown in Table 2.7.

Table 2.7. Peak days and Doubling Times (DT) for growth curves of BUNV, BATV, and NRIV with standard cell culture conditions (10%FBS)

Virus Inoculation titer (log PFU/mL) Peak day Doubling time

BUNV 6 4 0.32 5 5 0.24 4 5 0.25 3 5 0.36 2 7 0.28 1 7 0.48 BATV 6 3 0.32 5 4 0.44 4 5 0.25 3 5 0.36 2 7 0.28 1 7 0.63 NRIV 6 3 0.21 5 3 0.27 4 6 0.33 3 6 0.18 2 6 0.29 1 7 0.29

In the course of this investigation, we determined that viral RNA was still detected at 30 dpi when cells were presumably dead. We then hypothesized that there was viral stability of these viruses in the absence of viable cells in which to replicate. This led to a series of experiments to characterize the stability of these viruses. The course and logical framework of the remaining results are depicted in Figure 2.6.

Figure 2.6. Logic framework of in vitro and ex vivo growth kinetics study of BUNV, BATV, and NRIV.

2.3.3.2. Investigation of viral stability

a. Viral kinetics in Vero cells grown in less nutritious medium

Analysis of variance of AUC-L values showed that there were significant effects of both viral identity and inoculation dose. Pair-wise comparison by Tukey post-hoc test on viral identity showed that only BUNV and BATV were different (p=0.009). For post-hoc comparisons of inoculation doses, differences were found between 106 and 101PFU/mL

(p<0.0001), 106 and 104PFU/mL (p=0.0253), and 104 and 101PFU/mL (p< 0.0001).

Average growth curves for each virus at 2% FBS and 106-101 PFU/mL inoculation doses are shown in (Figure 2.7.).

Figure 2.7. BUNV, BATV, and NRIV growth kinetics in Vero cells grown in sub-standard conditions (2%FBS) with different inoculation doses (106-1PFU/mL). Error bars indicate represent standard errors.

Generally, BUNV and NRIV infected cell culture showed faster CPE than BATV infected cell culture. At 10 dpi all cells were visibly not viable after infection with all viruses at all inoculation doses. BATV infected cell culture did not show any signs of CPE at 1 dpi at any inoculation dose, while BUNV and NRIV showed early CPE at higher inoculation doses

(Table 2.8). As shown by CPE visualization, viral RNA was still detected long after all cells of the monolayer were dead.

Table 2.8. Gradual CPE during persistent infections of BUNV, BATV, and NRIV in Vero cells at 10%FBS

Virus Titer at 1dpi 2dpi 3dpi 4dpi 5dpi 6dpi 7dpi 8dpi 9dpi 10dpi 0 dpi (PFU/mL) BUNV 106 + ++ ++++ +++++ +++++ +++++ ++++++ ++++++ ++++++ ++++++ 105 + ++ ++++ +++++ +++++ +++++ ++++++ ++++++ ++++++ ++++++ 104 + + ++++ +++++ +++++ +++++ +++++ ++++++ ++++++ ++++++ 103 - + +++ ++++ ++++ ++++ ++++ ++++++ ++++++ ++++++ 102 - - ++ +++ +++ ++++ ++++ +++++ ++++++ ++++++ 101 - - + ++ ++ +++ ++++ +++++ ++++++ ++++++ BATV 106 - ++ +++ ++++ ++++ +++++ +++++ ++++++ ++++++ ++++++ 105 - + ++ ++++ ++++ +++++ +++++ ++++++ ++++++ ++++++ 104 - + ++ +++ +++ ++++ +++++ +++++ ++++++ ++++++ 103 - + + ++ ++ +++ ++++ ++++ ++++++ ++++++ 102 - + + + + +++ ++++ ++++ ++++++ ++++++ 101 - - + + + ++ +++ ++++ ++++++ ++++++ NRIV 106 + ++ ++++ +++++ +++++ ++++++ ++++++ ++++++ ++++++ ++++++ 105 + ++ +++ ++++ +++++ +++++ +++++ ++++++ ++++++ ++++++ 104 - ++ ++ ++++ ++++ ++++ ++++ ++++++ ++++++ ++++++ 103 - ++ ++ ++ ++ ++++ ++++ +++++ ++++++ ++++++ 102 - + + ++ ++ +++ +++ ++++ +++++ ++++++ 101 - - + ++ ++ ++ +++ ++++ +++++ ++++++

-: No CPE observed +: few dead cell ++: approximately 1/3 of cells are dead +++: approximately ½ of cells are dead ++++: approximately 2/3 of cells are dead +++++: very few cells still attached ++++++: all cells are dead (no cell attached

For doubling times, ANOVA using DT values for all replicates, each taken individually significant differences between viruses and inoculation doses. By Tukey post-hoc test, a difference was found to be between BUNV and BATV (p=0.009), however no difference was found between BUNV and NRIV (p>0.05), and between BATV and NRIV (p>0.05).

Regarding inoculation doses, differences were found between 106 and 101PFU/mL (p <

0.0001), 106 and 104PFU/mL (p=0.025), and 104 and 101PFU/mL (p< 0.0001). The means of the DT values and peak days per inoculation dose for each virus are shown in Table

2.9.

Table 2.9. Peak days and Doubling Times for growth curves of BUNV, BATV, and NRIV in sub-standard cell culture (2%FBS)

Virus Inoculation titer (log Peak day Doubling Time PFU/mL) BUNV 6 4 0.08 5 4 0.09 4 4 0.13 3 4 0.13 2 7 0.23 1 7 0.33 BATV 6 3 0.1 5 7 0.51 4 7 0.43 3 7 0.6 2 7 0.76 1 7 0.78 NRIV 6 3 0.14 5 3 0.09 4 4 0.21 3 4 0.21 2 7 0.18 1 7 0.59

When we compared growth kinetics between optimal conditions (10% FBS) from section 2.2.2 and 2% FBS cell culture conditions within viruses, the analysis of doubling time within viruses showed that there was a significant difference for BUNV.

The DT was doubled when BUNV was inoculated on cells grown in standard conditions compared to sub-standard conditions. No difference was found for BATV and

NRIV between both cell culture conditions (Table 2.10).

Table 2.10. Comparison of DT for standard and sub-standard cell culture conditions within viruses

DT Mean se p-value BUNV 10%FBS 0.32 0.082 0.0156 2%FBS 0.16 0.088 BATV 10%FBS 0.38 0.128 0.2376 2%FBS 0.53 0.229 NRIV 10%FBS 0.26 0.051 0.7553 2%FBS 0.23 0.16

b. Stability of BUNV, BATV, and NRIV

- Infectivity after stability for 30 dpi

BUNV, BATV, and NRIV inoculated in vitro with both optimal (10% FBS) and sub- optimal (2% FBS) conditions were shown to be stable in the absence of replication up to

30 dpi. The titers of supernatant collected from high (106PFU/mL), medium (104PFU/mL), and low (101PFU/mL) inoculation doses at 30 dpi are given in Table 2.11.

Table 2.11. Viral titers of supernatants collected at 30 days post-inoculation

Virus % FBS Inoculation dose (log Viral titer of supernatant collected PFU/mL) at 30 dpi (log PFU/mL) BUNV 10 1 6 10 4 6 10 6 6 2 1 6 2 4 7 2 6 7 BATV 10 1 6 10 4 6 10 6 6 2 1 7 2 4 7 2 6 7 NRIV 10 1 7 10 4 7 10 6 7 2 1 7 2 4 8 2 6 8

Supernatant from the 30 dpi collections were inoculated on Vero cells and collected at several time-points post-inoculation. We used a paired t-test to determine if there was a significant increase in viral titer from 1 dpi to 7 dpi, indicating replication and thus infectious virus. For each virus and %FBS condition, at least 2 out of 3 inoculations showed definite signs of replication (Figure 2.8. and Table 2.12.). This indicates that these three Orthobunyaviruses are stable in extracellular conditions.

Figure 2.8. Comparative infectivity of BUNV after in vitro persistence at 30 dpi from cells both grown in standard (10%FBS) and sub-standard conditions (2%FBS).

Figure 2.9. Comparative infectivity of BATV after in vitro persistence at 30 dpi from cells both grown in standard (10%FBS) and sub-standard conditions (2%FBS).

Figure 2.10. Comparative infectivity of NRIV after in vitro persistence at 30 dpi from cells both grown in standard (10%FBS) and sub-standard conditions (2%FBS).

- Ex vivo viral stability

For the viruses kept in cell-free M199 medium at 37oC, viral RNA was still detected at

30dpi. A decline of the curves was observed, correctly indicating no viral replication without cells; however, the final detected titers at 30 dpi were still relatively high 104-105

PFU/mL for BUNV and BATV, and higher than 105.75PFU/mL for NRIV.

Figure 2.11. Ex vivo stability of BUNV, BATV, and NRIV. The inoculation doses were viral stock titers (106PFU/mL for BUNV, 107PFU/mL for both BATV and NRIV)

Similarly, tests to determine infectiousness of virus collected from the media-only experiment at 30 dpi showed that only BUNV was still infectious at 30 dpi. A paired t-test comparing the replication titers at 1 and 7 dpi of supernatant collected at 30 dpi from cell-

free medium infected by BUNV, BATV, NRIV showed only a significant difference for

BUNV (p=0.03333).

No differences were found for supernatants of BATV and NRIV (Table 2.12). This indicates that BATV and NRIV are probably not replicating after staying in ex vivo conditions for 30 days.

Figure 2.12. Infectivity of BUNV, BATV, NRIV after stability in ex vivo conditions.

Table 2.12. Analysis of infectivity of supernatant collected at 30 dpi

Virus Cell growth Initial inoculation Analysis of infectivity of 30 dpi condition/ ex dose (log supernatant, p-value (paired t test vivo PFU/mL) analysis between 1 and 7 dpi) BUNV 10% 6 0.1972 (NS) 4 0.002565 1 0.03954 2% 6 0.04028 4 0.0005203 1 0.1407 (NS) Ex vivo 6 0.03333 BATV 10% 6 0.01851 4 0.03436 1 0.04964 2% 6 0.0482 4 0.008229 1 0.05556 (NS) Ex vivo 7 0.1387 (NS) NRIV 10% 6 0.03759 4 0.01472 1 0.002535 2% 6 0.007274 4 0.01645 1 0.005696 Ex vivo 7 0.416 (NS)

NS: No significant difference

c. Comparative inactivation by Triton-X-100

No CPE was observed on Vero cell monolayers infected with 1%Triton-X inactivated

BUNV, BATV, and NRIV before 5 dpi. However, for all three viruses, viral RNA was detected up to 7 dpi (Figure 2.13). A paired t test showed that there was no difference between RNA concentration at 1 and 4 dpi indicating that the viruses were not infectious.

Figure 2.13. Viral RNA detection of BUNV, BATV, and NRIV after 1% Triton-X-100 inactivation.

2.4. Discussion

These viruses have the potential to become emergent public health threats to animals and people and gaining a basic understanding of their behaviors in common laboratory systems is a necessary first step in getting “ahead of the curve”. Further, experimental studies in mammalian cell lines give insight into how the viruses may comparatively behave in mammalian host cells. Thus, this study provides a first glimpse into the infection kinetics of BUNV, BATV, and NRIV at different infective doses, and this study is the first to directly compare in vitro growth kinetics of BUNV, BATV, and NRIV.

First, we describe diagnostic assays to be used for field detection and other laboratory studies of these viruses. We showed that our designed BATV primers did not amplify the

NRIV M segment as well as expected, while the BUNV and NRIV cross-amplification consistently performed well. However, the BATV and NRIV M segments have a range of similarity of 89.3-96.0% (Briese et al. 2006) while the BUNV and NRIV S and L segments have a range of similarity of 99.6-99.8% and 98.5-99.8% respectively (Groseth, Weisend, and Ebihara 2012). This potential for lower genetic similarity of BATV and NRIV could be the cause of decreased efficacy of the BATV M primers for NRIV and/or vice versa.

We demonstrated that BUNV and NRIV produce more heterogeneous sized plaques while BATV plaques were more homogeneous, despite NRIV and BATV sharing the gene segments that regulate Bunyavirus entry into the cell (M segment). The M segment encodes the glycoproteins Gn and GC, which are responsible for viral attachment to host cells and may be involved in entry into the cell, though the mechanism of cell entry for

Bunyaviruses remains largely unknown (Albornoz et al. 2016). However, plaque size between NRIV and BATV was not different, while BUNV plaque sized tended to be smaller than the other two viruses. Previous studies identified small and large plaques of

BUNV and NRIV as different isolates and were associated with different pathogenesis profiles in mice (Odhiambo et al. 2014). Large plaque isolates grew to higher titers but were less virulent than small plaque isolates (Odhiambo et al. 2014). Given our results and this study, it is possible that the BUNV and NRIV stocks are a mixture of different isoforms or that these viruses have an inherent propensity to produce different isoforms in mammalian cell systems. In other viruses such as Zika virus, different plaque phenotypes have been associated with passage history or host species ((Moser et al.

2018). The lack of variability in BATV virus plaque morphology might point to a more consistent viremia profile in mammalian systems, especially given its lack of association with severe disease in mammals. It has been proposed that viremia profiles of arboviruses in some hosts are “slow-and-steady” compared to the fast-peak but fast- cessation profile in humans or other vertebrate hosts. This “tortoise-hare” model was proposed as an alternative strategy by the virus to infect as many mosquitoes as possible through longer viremic periods, even with lower systemic titers (Althouse and Hanley

2015). It may be that BATV utilizes the “tortoise” model while BUNV and NRIV are more

“hare”. Ecologically, this could make sense as BATV has been proposed to infect mostly birds, where long and persistent infections have been seen with other arboviruses like

West Nile (Komar et al. 2003; Nemeth et al. 2009; Wheeler et al. 2012; Reisen et al.

2013).

We also found that there were no significant differences among these viruses in their growth kinetics, regardless of cell culture conditions (% FBS) or inoculation doses. Of course, the correlation between the growth curves and clinical manifestation in vivo should also be investigated for direct comparisons and estimations of infectivity in humans and other natural hosts. However, we did determine that to fully assess infectivity, it is better to dilute the virus to a titer below a MOI of 1 (at a titer of 104 PFU/mL per 106 cells, e.g.) as the higher inoculation doses apparently saturated the system.

Viruses are obligate intracellular organisms. Thus, for viral replication, viable host cells are necessary. However, we demonstrated that these viruses were stable outside of viable cell culture and maintained infectiousness for at least 30 dpi under laboratory conditions. The phenomenon of viral stability has been found in other

Bunyaviruses: Rift Valley Fever Virus (Phenuiviridae family) (Billecocq, Vialat, and Bouloy

1996), Toscana virus (Phenuiviridae family) (Verani, Nicoletti, and Marchi 1984) and

Dugde virus (Nairoviridae family) (David-West and Porterfield 1974). In all these cases, viral inocula were enriched in deferring interfering (DI) particles which seem to be associated with persistent infections (David-West and Porterfield 1974; Verani, Nicoletti, and Marchi 1984; Billecocq, Vialat, and Bouloy 1996). Defective interference with large deletions in the L RNA segment has been reported in BUNV infected baby hamster kidney

(BHK) cells (Scallan and Elliott 1992). Further investigations need to be conducted in

Vero cells especially in relation to persistent infections. Toscana infected Vero cells were morphologically similar to parental cells and didn’t show any CPE (Verani, Nicoletti, and

Marchi 1984) contrary to BUNV, BATV, and NRIV infected Vero cells in our study.

Previous studies have reported long-term viral stability in other Bunyaviruses: Sandfly

Fever Sicilian Virus (SFSV), a sandfly-borne virus of the Phenuiviridae family; Hantaan virus, a rodent-borne virus of the Hantaviridae family; and Crimean-Congo Hemorrhagic

Fever Virus (CCHV) of Nairoviridae family (Hardestam et al. 2007). The epidemiological relevance of stability harkens back to RVFV, which has also been found to be stable for long time in the environment (Craig, Thomas, and DeSanctis 1967, Miller 1962). In the case of RVFV, environmental contamination from abortions or slaughter (and thus association with ex vivo stability) is a major source of human-acquired cases (Yuill 1991;

Archer et al. 2011; OIE 2016; Clark et al. 2018).

Given that these Orthobunyaviruses are also associated with abortions and hemorrhagic manifestations similar to RVFV, it could be postulated that dead hosts and

infective tissues might be sources of infectious virus from the environment for relatively long periods, provided permissive conditions (humidity, temperature, etc.).

For the purposes of diagnostic development, we showed that 1%Triton-X-100 can be used to inactivate BUNV, BATV, and NRIV. This could have useful applications such as

ELISA design for detecting exposure to these viruses since there are no such commercially available kits. The use of inactivated viruses is especially important in developing parts of the world which lack biosafety facilities required to handle infectious viruses. At the laboratory level, the combination of viral stability and Triton-X-100 inactiviation should be taken into account in the design and implementation of the laboratory biosafety measures.

Summary

We provide a comparative analysis of the in vitro kinetics of BUNV, BATV, and NRIV and conclude that growth kinetics are not different among them. However, we also provide preliminary characterization of and diagnostic assays for these viruses for laboratories or field studies looking to work with these viruses, so they do not have to

“start from scratch.” We also provide an important biosafety and epidemiological finding of the stability of these viruses and determine the utility of a common detergent in inactivating these viruses for the purpose of further diagnostic development.

Acknowledgements

We would like to thank Dr. Bob Tesh, Adeola Lawal, and Ms. Julie Cherry for their work in obtaining the viruses from the World Reference Center for Emerging Viruses and

Arboviruses.

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CHAPTER 3. PRELIMINARY IN VIVO INVESTIGATION OF BUNYAMWERA, BATAI, AND NGARI VIRUSES

3.1. Introduction

Bunyamwera (BUNV), Batai (BATV), and Ngari (NRIV) viruses are members of

Orthobunyavirus genus in the Peribunyaviridae family of the Bunyavirales order involved in a reassortment process as NRIV is a natural reassortant of the other two (Bowen et al.

2001; Weisend, and Ebihara 2012; King et al. 2018, Groseth; Maes et al. 2019). As mentioned in previous chapters, members of this family are characterized by single- stranded RNA genome partitioned into three segments Small (S), Medium (M), and Large

(L). In the Peribunyaviridae family, S segment encodes nucleoprotein (N) and nonstructural protein (NSs), M segment encodes glycoproteins (Gn and Gc) and nonstructural protein (NSm) while L segment encodes polymerase protein (Elliott 1996;

Schmaljohn 2001; Bioinformatics Swiss Institute 2010; Elliott 2013; Elliott 2014). The

NRIV genome consists of the S and L genome segments from BUNV and the M segment from BATV (Gerrard et al. 2004; Briese et al. 2006). The M segment gene products (Gc and Gn) are known to be major determinants of host range, tissue tropism, transmissibility, neutralization, and hemagglutination (Beaty et al. 1981; Shope, Rozhon, and Bishop 1981; Castrucci 1993). In addition, NSm, another M segment gene product, partially contributes to the virulence (Elliott 1996; Bridgen et al. 2001; Kohl et al. 2003;

Elliott 2013; Soldan and González-Scarano 2014; Otieno 2015).

Subsequently, reassortants have been speculated to share similar aspects with their genetic parents from which they got M segment (Otieno 2015). However, apart from reassortants which share the entirety of M segment with the donor viruses, it has been

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shown that Gn and Gc have conserved regions shared among a large number of viruses within genus. These conserved regions are at the basis of antigenic relationships within the genus, leading to antibody cross-neutralization. For many years, as genome sequences were scarce, Bunyaviruses were classified into serogroups based on these antigenic relationships detected by hemagglutination inhibition and neutralization tests

(Elliott 1996; Cheng et al. 200; Elliott 2013; Otieno 2015).

BUNV and BATV were found to cross-react between themselves and with more than

30 other viruses belonging to the same Bunyamwera serogroup, one of seventeen (17) in Orthobunyavirus genus (Table 1.1) (Eley et al. 1989; Mohamed, McLees, and Elliott

2009; Elliot 2013; Dutuze et al. 2018). However, after NRIV was confirmed as reassortant between BUNV and BATV, no study has been conducted to test the abilities of NRIV to cross-neutralize BUNV and BATV and vice-versa.

BUNV, BATV, and NRIV have all been shown to cause diseases in both humans and animals. However, although they are connected this genetic phenomenon, the diseases associated with these viruses in vertebrate hosts are dissimilar in pathogenesis and virulence. BATV is characterized by low virulence, BUNV by moderate virulence, and

NRIV has been shown to be the most virulent of the three (Southam and Moore 1951;

Kokernot et al. 1958; Nashed, Olson, and el-Tigani 1993; Edwards 1994; Gerrard et al.

2004; Briese et al. 2006; Eiden et al. 2014; Tauro et al. 2015). Unfortunately, these differences in virulence and pathogenesis are compared mainly by observational studies from clinical cases isolated during outbreaks or other field occurrences. Very few experimental studies have been conducted to characterize these aspects adequately.

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Like for other viral infections, immune responses against these Bunyaviruses inckude type I interferons as a come in front line in innate immunity (Garcia-Sastre and Biron 2006;

Stetson and Medzhitov 2006). Type I interferons (IFN-α/β) are produced and secreted by cells in response to viral infection and activate the expression of so-called IFN-stimulated genes (ISGs) in neighboring cells.

Many ISGs encode proteins with the ability to directly or indirectly inhibit virus multiplication. The expression of IFN genes is enhanced by different transcription factors such as IRF3, IRF7, NF-kB and AP-1 (Garcia-Sastre and Biron 2006; Kawai and Akira

2006; Stetson and Medzhitov 2006; Elliott and Weber 2009). The importance of IFNs in bunyaviral infections has been confirmed by studies showing that several Bunyaviruses can be inhibited by IFNs (Luby 1975; Tamura et al. 1987; Morrill et al. 1989; Temonen et al. 1995; Livonesi et al. 2007; Habjan et al. 2009). In addition to mouse models indicating that mice lacking functional type I IFN receptors are highly susceptible to infection with

BUNV, La Crosse virus (LACV), Dugbe virus (DUGV), Hantaan virus (HTNV), and Rift

Valley Fever virus (RVFV) (Hefti et al. 1999; Bouloy et al. 2001; Weber et al. 2001;

Wichmann et al. 2002; Boyd, Fazakerley, and Bridgen 2006; Blakqori et al. 2007).

Although BUNV is the prototype of Bunyavirales order, and therefore should be extensively studied for better understanding the infections by viruses of this order, there are still few available mouse models for infection by this virus. These include the comparison of pathogenesis of wild-type BUNV and BUNdelNSs (NSs deleted mutant) in

5-week-old BALB/c (Bridgen et al. 2001), description of brain lesions in two-day-old ICR

Swiss mice (Murphy, Harrison, and Tzianabos 1968), and comparison of pathogenicity of small and large BUNV plaques in (1–4 day-old) and 6-week-old Swiss Albino suckling

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mice (Odhiambo et al. 2014). All of these models focus on the description of occurrences of clinical manifestations and mortality. Histological lesions for this infection are less investigated as only brain lesions are described only after intracerebral inoculation

(Murphy, Harrison, and Tzianabos 1968; Bridgen et al. 2001).

In this study, we investigated the comparative infection kinetics of BUNV, BATV, and

NRIV in C57BL/6 (wild type) mice and quantified the potential of BUNV, BATV, and NRIV to cross-neutralize each other. In addition, we investigated the role of IRF3 and IRF7 in

BUNV infection by characterizing phenotypically and histopathologically the pathogenesis in IRF3/7 double knock out (-/--/-) mice.

3.2 . Material and methods 3.2.1. Viruses

As previously mentioned in chapter 2, viral stocks used in this research project were

BUNV strain 6547-8 with a titer of 6.65x106PFU/mL, BATV strain MM2222 with the titer of 2.27 x 107PFU/mL, and NRIV strain DAK AR D 2854 with the titer of 5.7x107PFU/mL.

For infection of C57BL/6 mice aimed at comparatively characterizing BUNV, BATV, and

NRIV infection kinetics, the three viruses were used after matching titers at 106 plaque forming units (PFU)/mL. For the characterization of pathogenesis of BUNV in IRF 3/7 -/--/- mice, only BUNV was used at 103PFU/mL.

3.2.1 Mice and infections a. Viremia and neutralization study of BUNV, BATV, and NRIV in wild type mice

Fifteen 10-week-old C57BL/6 mice were put into three groups of five each. Each group was inoculated one of the three viruses (BUNV, BATV, and NRIV). Mice were inoculated by subcutaneous injection with 100µL of 6.65x106PFU/mL, 2.27x106PFU/mL, and

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5.7x106PFU/mL respectively for BUNV, BATV, and NRIV. Inoculated mice bled at 1, 3,

5, 7, and 30 days post-infection (dpi). Blood samples collected at 1, 3, 5, and 7 dpi was used to study comparative viremia of these viruses while blood samples collected at 30 dpi served at serum collection for homologous and cross-neutralization study. Samples were incubated for 30 mins at room temperature and centrifugated for 4 minutes at 6,000 rcf and 4°C. Serum was then separated from clot and kept in -80oC until ready to use. For the viremia study, viral RNA was extracted and amplified while plaque reduction neutralization tests (PRNTs) were performed for serological investigation. b. BUNV pathogenesis in IRF 3/7-/--/- mice

Eight (8) female IRF3/7 double knock out mice were used for these experiments. The mice were 8 to 9 weeks old at the date of inoculation. Prior to the actual infection experiment, a male mouse was subcutaneously inoculated with 100 µL of 6.65 x

105PFU/mL for an infection test. The mouse died at 4 dpi. From that infection test, we decided to infect the 8 females with a lower titer hoping that they would survive longer for more appreciation of clinical manifestations and lesions. Mice were then subcutaneously inoculated with 100 µL of 6.65 x 103 PFU/mL of BUNV. They were weighed every day until 6 dpi and bled every day until 4 dpi. They could not be bled after 4 dpi because of severe vascular lesions. As for C57BL/6 mice infection, blood samples were incubated for 30 minutes at room temperature and serum was collected after centrifugation (6,000 rcf, 4 mins, 40C). Serum was then separated from the clot and stored at -80oC until viral

RNA extraction and amplification.

After death, all mice were opened on the abdominal line for complete multisystemic gross examination. Three mice were placed in 10% neutral buffered formalin and sent to

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a histopathology laboratory for analysis. Tissues were then dehydrated in alcohol and xylene and embedded in paraffin. 5 μm thick tissue sections were then obtained for slide preparation before being stained with hematoxylin and eosin and coverslipped. After slides were examined via light microscopy, pathological changes were recorded by certified pathologists of the American College of Veterinary Pathologists (ACVP) board at the LSU School of Veterinary Medicine.

3.2.2. Viral RNA extraction and qRT-PCR

RNA extraction was performed using the KingfisherTM (Thermo-Fisher) automated extraction platform (according to the instructions of manufacturer) as previously described. Viral RNA was detected by qRT-PCR using the SuperScript™ III One-Step

RT-PCR System with Platinum® Taq DNA Polymerase (Life Technologies) on a Roche

Lightcycler® 480 (Roche). We used primers and probes targeting the M segments (Table

2.3).

3.2.3. Plaque Reduction Neutralization Test (PRNT)

Vero African green monkey kidney cells (ATCC) were grown under optimal growth conditions (88% Medium 199, Earle's Salts, 10% Fetal Bovine Serum, and 2% Antibiotic-

Antimycotic) at 37˚C, 5% CO2 were used to perform PRNT. Serum samples collected at

30 dpi for each mouse were serially diluted in plain M199E as follows: 1:20, 1:40,

1:80, 1:160, 1:320, 1:640, 1:1280, 1:2560, 1:5120, and 1:10240. Before performing cross-neutralization assays, homologous neutralization assays were performed to check the antibody production level for each group. For cross-neutralization study assay, neutralization of each virus by the antibodies produced against the other two was studied in a pairwise way as presented in Table 3.1. PRNTs were performed following WHO

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protocol for Dengue virus (OIE 2017), previously used in our lab for both Dengue and

Zika viruses (Kawiecki and Christofferson 2016; Kawiecki et al. 2017), and briefly described below. Diluted sera were heat inactivated at 56oC for 30 min before, 100µL of viral mix (viral stock diluted in M199E medium) containing 50 PFU was added to 100µL of each serum dilution. The mixes virus/diluted sera samples were then incubated at 37oC for 1 hour. After removal of the maintenance media on confluent 6 well plates, the virus- serum mix was added, with each well corresponding to a specific serum dilution. Plates were then rocked for 30 mins at room temperature and incubated for 30 mins at 37oC. An additional positive control well with only 100 µL of 50 PFU of virus was prepared to serve in the calculation of plaque number reduction as well as a negative control well with only

100 µL of M199E media. Positive and negative controls all received the same treatment as other wells with serum dilutions. 3 mL of overlay media was added to each well and 2 mL of second overlay media was added at 3 dpi for both BUNV and NRIV, and at 4 dpi for BATV.

Table 3.1. Homologous and cross-neutralization study plan

Assay Virus to neutralize Serum used

Homologous neutralization BUNV BUNV BATV BATV NRIV NRIV Cross-neutralization BUNV BATV NRIV BATV BUNV NRIV NRIV BUNV BATV

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3.2.4. Statistics

Weight loss was reported as the percent reduction compared to initial weight. For

BUNV, BATV, and NRIV infected C57BL/6 mice and BUNV infected IRF3/7 mice, viremia titers reported are log10 transformed titers by qRT-PCR.

For homologous and cross-neutralization experiments, PRNT50 and PRNT80 titers were expressed as the reciprocal of the highest dilution at which the 50% and 80% of plaque reduction was achieved.

3.3. Results 3.3.1. In vivo infections kinetics of BUNV, BATV, and NRIV in C57BL/6 mice

a. Viremia curves

Neither BUNV nor BATV were detected after 3 dpi, and only one NRIV infected mouse presented with what looked like viremia peaking at 3 dpi. Although there was no increase in viral titers, for the other four NRIV infected mice, viral RNA was detected up to 5 dpi

(Figure 3.1).

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Figure 3.1. Comparative viremia curves of BUNV, BATV, in NRIV in C57bl/6 mice.

b. Homologous and cross-neutralization experiments

As expected, it was found that BUNV, BATV, and NRIV all cross-neutralize each other.

However, the levels of neutralization differ depending on antibodies and the virus being neutralized. The averages of neutralization titers for all pairs of neutralization are summarized in Table 3.2. NRIV was the most efficient at both self and cross- neutralization, BUNV self and cross-neutralized at a moderate level while BATV was the least efficient at both.

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Table 3.2. PRNT50 and PRNT80 titers for homologous and cross-neutralization BUNV, BATV, and NRIV.

Virus neutralized BUNV BATV NRIV Antibodies PRNT50 PRNT80 PRNT50 PRNT80 PRNT50 PRNT80 BUNV 4736 272 384 128 320 20 BATV 656 <20 <20 <20 <20 <20 NRIV 944 40 2816 1280 >10240 3456

The detailed neutralization profiles for all dilutions of BUNV antibodies for homologous

(neutralization of BUNV) and cross-neutralization (neutralization of BATV and NRIV) are represented by Figure 3.2.

Figure 3.2 also provides detailed neutralization profiles for all dilutions of BATV antibodies for homologous (neutralization of BATV) and cross-neutralization

(neutralization of BUNV and NRIV). It was found that these antibodies did not neutralize any of these viruses to 80% reduction. No reduction of 50% was obtained for cross- neutralization (neutralization of BUNV and NRIV) while for homologous neutralization was achieved at high serum dilutions (PRNT50 of 656 as shown in Table 3.2). This could be an indication that BATV infected C57BL/6 mice did not produce a large amount of antibodies.

NRIV was the most efficient at both self-neutralization and neutralization of its genetic parents as shown by titers in Table 3.2 and in more details provided in Figure 3.4. At dilution of 1:10240 (the last for our experiment), the homologous neutralization was still higher than 50% reduction, showing a strong self-neutralization. NRIV antibodies neutralize BATV more than BUNV. The observed PRNT50 were 2816 and 944 for BATV and BUNV respectively. And the observed PRNT80 titers were 1280 and 40 for BATV

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and BUNV respectively. This is not surprising as BATV and NRIV have similar glycoproteins, which serve as triggers of neutralizing antibody production.

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Figure 3.2. Comparative homologous and cross-neutralization profiles for BUNV, BATV, and NRIV for different serum dilutions.

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3.3.2. Pathogenesis of BUNV infection in IRF 3/7 -/--/- mice

a. Clinical manifestations

By 3 dpi, all mice (100%) had shown signs of illness such as lethargy and hunched posture. Facial swelling was observed in 3 mice (37.5%) (as shown by Figure 3.3).

Mortality was observed starting at 4 dpi and was at 100% at 6dpi.

Figure 3.3. Dead BUNV infected IRF 3/7 -/--/- mouse with facial swelling.

b. Weight loss, viremia, and survival

The highest weight loss was observed from 1 to 3 dpi (Fig 3.4 A). All mice were viremic but peak days were not well identified due to the rapid mortality (Figure 3.4.B).

The survival percentage was 100% up to 3dpi, 37.5% at 4dpi, 12.5% at 5dpi and 0% at

6dpi (Figure 3.4.C).

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Figure 3.4. Weight loss, viremia and survival curves of BUNV infected IRF 3/7 -/--/- mice. A. Weight loss, B. Viremia, C. Survival curves.

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c. Histopathological lesions

The disease was multisystemic causing mainly necrotic lesions in the liver, lungs, spleen, bone marrow, reproductive system (uterus and ovaries), and skin with vascular necrosis.

The liver was affected by severe random acute coagulative necrosis (framed area on

Figure 3.5.A) with multifocal single cell necrosis, (red arrows on Figure 3.7.B) and portal hepatitis (black arrow on Figure 3.5.B).

Figure 3.5. Necrotic lesions of the liver of BUNV infected IRF3/7 -/--/- mice. A. Liver severe ample necrosis (10X), B. Liver single cell necrosis and portal hepatitis (40X).

In the lungs, there was intravascular leukocytosis (black arrow on Figure 3.6.A), cell death (black arrows on Figure 3.6.B), and mild incipient accumulation of inflammatory cells in lung parenchyma (framed on Figure 3.6.B). Additionally, there was a mediastinal steatitis (framed on Figure 3.6.C), and vascular necrosis characterized by loss of structure of endothelial cells (black arrow on Figure 3.6.C).

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Figure 3.6. Necrotic and inflammatory lesions of the lung of BUNV infected IRF3/7 -/--/- mice A. Lung intravascular leukocytosis (40X), B. Lung intravascular leukocytosis and leukocyte death (40X), C. Lung mediastinal steatitis and vascular necrosis (40X).

The lymphoid organs associated with immune response were necrotic (Figure 3.7.A).

The bone marrow was affected by multifocal necrosis and the spleen by a severe perifollicular necrosis (framed on Figure 3.7.B).

Figure 3.7. Necrotic lesions of lymphoid organs in BUNV infected IRF3/7 -/--/- mice. A. Bone marrow necrosis (40X), B. Spleen perifollicular pulp severe necrosis (40X).

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In the reproductive system, there was a multifocal necrotizing oophoritis in the ovary follicle (framed on Figure 3.8.A) and a mild multifocal necrosis in endometrium (black arrows on Figure 3.8.B).

Figure 3.8. Necrotic lesions of reproductive system of BUNV infected IRF3/7 -/--/- female mice, A. Ovary necrotizing oophoritis (20X), B. Mild endometrium necrosis(40X).

The disease was also characterized by necrosis of the skin (Figure 3.9. A). In the eyelids, there was severe diffuse acute edema expanding the loose connective tissue, and dilatation of lymphatics (Figure 3.9.B). These changes were responsible for the prominent facial swelling.

Figure 3.9. Lesions associated with facial swelling. A. Skin necrosis and mast cell degranulation (40X), B. Eyelid severe diffuse edema (20X). 117

No significant lesions were observed in the brain, kidneys, adrenal glands, esophagus, stomach and intestine, skeletal muscle, bone, pancreas, thyroid, or parathyroid.

3.4. Discussion

In this study, we compare in vivo infections of BUNV, BATV, and NRIV, provide a cross-neutralization study including NRIV, and provide multisystemic histological characterization of BUNV associated lesions in IRF3/7 -/--/- mice.

This study shows that immunocompetent 6-week old C57BL/6 mice are not susceptible to either BUNV, BATV, or NRIV infection. Although no clinical sign was noticed, NRIV was the only virus to be detected up to 3 dpi, while BUNV and BATV were only detected at 1 dpi. This extended detection of NRIV could be one indication of its increased virulence compared to BUNV and BATV.

Previous experimental studies have shown that suckling (2 day-old) ICR Swiss mice inoculated intracerebrally by BUNV (infectivity dose of 1010 LD50/mL) started dying after

53 hours after developing severe encephalitis. Viremic titers increased progressively and plateaued at 16h (Murphy, Harrison, and Tzianabos 1968). Another study showed that when BUNV and NRIV (infectivity doses of 109PFU/mL) were used to intraperitoneally infect both 6-week-old and suckling (1-4 days old) Swiss Albino mice, 6-week-old mice were not susceptible to any of the two infections, while suckling mice died between 2 and

6 days after showing neurological symptoms such as limb paralysis, tremors, and disorientation (Odhiambo et al. 2014). All these studies including ours, indicate that suckling wild type inoculated both intracerebrally and intraperitoneally as well as adult wild type mice inoculated intracerebrally are severely susceptible to BUNV and NRIV infections and die quickly starting at 2 dpi.

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However, adult wild type mice inoculated intraperitoneally and subcutaneously are not susceptible to infection BUNV, BATV, and NRIV.

Neutralization assays showed that BUNV, BATV, and NRIV all self and cross- neutralize each other though levels of neutralization differ. BUNV and BATV cross- neutralization is not surprising as they belong to the same Bunyamwera serogroup.

Viruses within this serogroup have previously shown cross-neutralization (Hunt and

Calisher 1979). However, this is the first study including their reassortant in cross- neutralization investigation. NRIV was the most efficient at both self and cross- neutralization, BUNV self and cross-neutralized at a moderate level, and BATV was the least efficient at both. The low homologous neutralization might indicate low antibody production by C57BL/6 mice.

It was noticed that NRIV strongly neutralized BATV, which is not surprising because the two viruses have similar glycoproteins, which serve as triggers for neutralizing antibodies (Elliott 1996; Yandoko et al. 2007; Elliott 2013; Elliott 2014; Soldan and

González-Scarano 2014). However, BATV did not cross-neutralize BUNV and NRIV nor itself very efficiently. As previously mentioned, this could be related to a low level of BATV antibody production by C57BL/6. The epidemiological relevance of this cross- neutralization is that an infection to any of these three viruses will lead to protection of the next infection of any of these three viruses. This is important especially for BUNV and

NRIV which have overlapping distributions in Africa (Zeller et al. 1996; Bowen et al. 2001;

Wertheim 2012; Dutuze et al. 2018). The high neutralization of BATV by both BUNV and

NRIV combined with its low virulence leading to non-specific alarming symptoms, could

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be the reasons why BATV isn’t well investigated in Africa, where BUNV and NRIV are known to circulate.

In previous a study on homologous and heterologous neutralization of viruses of the

Bunyamwera serogroup, it was found that the PRNT90 of homologous neutralizations were 1800 and 5100 for BUNV (RI-1 strain) and BATV (MM2222 strain), respectively. The ratios of homologous: heterologous titers to the nearest whole number were all 91 for

BATV neutralization by BUNV and vice-versa (Hunt and Calisher 1979).

The difference between these findings and ours may come from the difference in mice strains used, though in the study discussed, mice strain used was not specified.

In this study, we also showed that IRF3/7 -/--/- mice are susceptible and severely affected by BUNV infection, contrary to wild type mice, which did not show any sign of sickness, this highlights the importance of type I IFN in fighting BUNV infection. The histological characteristics show that the disease was multisystemic affecting different organs such as the lungs, liver, and organs of the reproductive system (uterus, ovaries).

The main lesion found in necropsy was hemprrhage in abdominal cavity while the main histopathological lesions were necrosis and inflammation. Although BUNV infection has not been associated with hemorrhagic fever in either humans nor animals, hemorrhage in abdominal cavity found in IR3/7 -/--/- shows that this virus has the potential to do so, especially in immunocomprised individuals.

Similar lesions have been observed in an experimental study of another member of

Orthobunyavirus genus, Schmallenberg virus (SBV). When 0.86x103 50% tissue culture infectious doses of SBV was used to subcutaneously inoculate 8-week-old type I IFN receptor knock out (IFNAR-/-), mice presented with diffusely discolored livers, large

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amounts of blood in abdominal cavities, and internal bleeding into the small intestine. In histopathology, the liver presented severe hepatocellular degeneration and necrosis

(Wernike et al. 2012).

These studies confirm the role of type I IFN in immune response to Orthobunyaviruses and indicate that these are highly necrotic viruses that have a high potential to produce hemorrhagic fevers especially in immunocomprimised individuals. As BUNV is the prototype of the Peribunyaviridae family and of Bunyavirales order in general, this study could be a model of Bunyaviral infection in IRF 3/7-/--/- mice.

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CHAPTER 4. IDENTIFICATION OF ORTHOBUNYAVIRUS INFECTIONS IN CATTLE DURING A RIFT VALLEY FEVER OUTBREAK IN RWANDA IN 2018 4.1. Introduction

Members of the Bunyavirales order have caused diseases of zoonotic and economic importance globally since the beginning of the 20th century (Briese, Calisher, and Higgs 2013; Elliott 2013; Clark et al. 2018; Dutuze et al. 2018).

In Africa, the most ancient and most devastating Bunyavirus is Rift Valley Fever Virus

(RVFV) (Genus: Phlebovirus, Family: Phenuiviridae,) was first identified and characterized in 1912 and further detected as etiological agent of fatal epizootic outbreak in Kenya in 1930 (Stordy 1913; Daubney, Hudson, and Garnham 1931; Baba et al. 2016).

RVFV is a vector-borne virus that causes disease in humans, livestock, and wildlife ruminant species (Daubney, Hudson, and Garnham 1931; Ikegami 2011; Bird and

McElroy 2016; Clark et al. 2018). In animals, it is clinically characterized by abortions and stillbirths, hepatitis, and hemorrhagic fever in severe cases (Daubney 1931; Findlay 1931;

Easterday 1965; Ikegami 2011). In humans, the mild form is characterized by self-limiting febrile illnesses, but individuals may progress to severe disease manifesting as hemorrhagic fever, encephalitis, vision loss and conjunctivitis (Freed 1951; van Velden et al. 1977; Laughlin et al. 1978; Salib and Sobhy 1978; Strausbaugh et al. 1978; Siam,

Meegan, and Gharbawi 1980; Deutman and Klomp 1981; Ikegami 2011). These clinical manifestations are common in many diseases caused by Bunyaviruses (Elliott 1996;

Elliott 2013; Soldan and González-Scarano 2014). Although RVFV has a broad range of identified vectors consisting of mosquitoes, ticks, and flies, the primary vectors are Aedes spp and Culex spp mosquitoes (Daubney R. 1932; Lee 1979; Davies and Highton 1980;

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Linthicum, Davies, et al. 1985; Linthicum, Kaburia, et al. 1985; Pepin et al. 2010;

Fontenille et al. 1998; Labuda and Nuttall 2004). RVFV consistently circulates in domestic ruminants and human populations throughout different countries of Africa and the Arabian

Peninsula where it causes significant outbreaks associated with livestock mortality, associated economic losses, and human mortalities (Woods 2002; Balkhy and Memish

2003; Archer et al. 2011; Baba et al. 2016; Samy, Peterson, and Hall 2017; Budasha et al. 2018; Clark et al. 2018; Georges et al. 2018). The virus is maintained during interepidemic periods by transovarial transmission in Aedes mosquitoes and the resurgence of outbreaks is facilitated by periodic rainfall and flooding due to the intertropical convergence of air currents from southern and northern hemispheres which leads to a seasonal emergence of infected Aedes spp. (Linthicum et al. 1999). After the emergence of infected Aedes mosquitoes, an outbreak may be sparked but the epidemic is likely maintained by Culex spp. (Linthicum et al. 1999). In East Africa, this cycling pattern has been recognized since the 1970s with large outbreaks occurring in 1977

(Egypt) (Siam, Meegan, and Gharbawi 1980), in 1997-1998 (Kenya) (Woods 2002), in

2006-2008 (Somalia, Tanzania, and Kenya) (WHO 2007; Anyangu et al. 2010), and 2018

(Kenya, Uganda and Rwanda) (OIE 2018; WHO 2018).

In Rwanda, the only Bunyavirus regularly surveilled is RVFV. The first molecularly confirmed case of RVFV was reported in Bugesera in 2011 after unusually high rate of abortions in domestic ruminants (mainly cattle) during the period of May to June (Rwanda

Agriculture Board 2013; Umuhoza et al. 2017). Since that time, RVFV has become endemic with sporadic cases reported year-round in the Eastern part of the country, and higher intensity outbreaks in May-June and December-January following the rainy

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seasons of March-April and October-November (Rwanda Agriculture Board 2013;

Umuhoza et al. 2017). For prevention, livestock are recommended to be regularly vaccinated with live attenuated Rift Valley Fever vaccine prepared from Smithburn’s attenuated strain of RVF virus (RIFTVAX) (Rwanda Agriculture Board 2013). However, vaccine uptake and compliance rates are unknown.

Although RVF is the most recognized and significant arbovirus in the region, many other zoonotic Bunyaviruses such as Bunyamwera virus (BUNV), Batai virus (BATV), and

Ngari virus (NRIV), circulate in East Africa. BUNV was first identified in Uganda (1943) and since has been regularly found in Tanzania, Kenya, and Uganda between 1945 and

2012 (Kokernot et al. 1958; Wertheim 2012; Otieno 2015; Dutuze et al. 2018). BATV was first identified in Uganda (1966) and highly suspected in Sudan by serological assay

(1988) (Singh 1966; Nashed, Olson, and el-Tigani 1993). NRIV was first identified as an etiological agent of fatal hemorrhagic fever outbreak in Kenya (1997-1998) (Gerrard et al.

2004; Briese et al. 2006; Otieno 2015). Despite their obvious potential to be of public and one health importance, BUNV, BATV, and NRIV are not ordinarily included in diagnostic panels, and could be cryptically circulating throughout Sub-Saharan Africa.

BUNV, BATV, and NRIV belong to the taxonomic genus Orthobunyavirus, and NRIV is the natural reassortant of BUNV and BATV (SBUNV MBATV LBUNV) (Zeller et al. 1993).

BUNV, BATV, and NRIV are transmitted by similar vectors as RVFV, infect similar vertebrate hosts, and infection with these viruses have been characterized with similar clinical manifestations (Gerrard et al. 2004; Briese et al. 2006).

In 2018, there was unusually heavy rainfalls that led to explosion of RVFV in East

African countries (Rwanda, Kenya, Uganda, and Tanzania). This was the most intense

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occurrence of RVF in Rwanda as it caused many cases of death and/or abortion cases in the ruminant population (OIE 2018). In addition, there was likely an underreporting of

RVFV cases of small ruminants due to their low economic and cultural importance in

Rwanda. Diagnosis of cattle was primarily by clinical manifestations, not by molecular techniques.

Thus, we tested samples from acutely ill ruminants from Rwanda during the May-June

2018 outbreak using conventional PCR to molecularly confirm RVFV and to test the hypothesis that BUNV, BATV, or NRIV might be co-circulating (Dutuze et al 2018).

4.2. Material and methods 4.2.1. Sample collection and handling

One hundred and fifty-seven (157) blood samples were collected from cattle and twenty-eight (28) from goats suspected of RVFV infection in different parts of the country between May 29 and July 25, 2018. The inclusion criteria for blood collection were (at least one of them): having aborted in less than a week, presenting signs of hemorrhagic fever, history of death suspected for RVF in same farm in less than 3 days, and history of abortion suspected to be associated with RVF in less than 3 days. Calves whose mothers presented signs of hemorrhagic fever or have died suspected of RVF with 3 days were sampled as well.

Figure 4.1 shows cases of hemorrhagic fever and abortion in cows. Blood was collected in 5mL purple-top Vacutainer tubes (without anti-coagulant). Demographic information of farmers such as address and complete identification of sampled animals

(age, sex, breed, clinical signs, status of RVFV vaccination, and breed) were also collected. From the field, samples were transported to the Virology laboratory of Rwanda

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Agriculture Board immediately at Rubirizi station. Samples were then put into 2 mL

Eppendorf tubes and kept in -80oC and subsequently, samples were shipped to Louisiana

State University for additional analyses.

Figure 4.1. Clinical manifestations of cattle during RVF outbreak. A and B. Case of hemorrhagic fever, C and D. Abortion cases. This picture also shows the risk of exposure from animals to human population as the environment was contaminated by infected tissues.

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4.2.2. Viral RNA extraction and cDNA synthesis

TRI Reagent® was added to each sample homogenate in DNA/RNA Shield™ in a ratio of 3:1 after what RNA extracted was performed using Direct-zol TM MiniPrep Zymo

Research kit (Cat No. R2052) according to manufacture instructions. First-step total cDNA synthesis was performed on RNA extracted from whole blood. 8μL RNA was added to 1μL random hexamers and 1μL dNTPs and run on thermocycler at 65oC for 5min. After that, to this 10μL was added another 10μL consisting of 2μL 10X RT Buffer, 4μL 25mM

MgCl2, 2µL 0.1m DTT, 1µL RNase OUT, 1µL Superscript III RT (Invitrogen kit, Cat No.

18080-051). The new 20μL was run on thermocycler at 25oC for 10min, 50oC for 50min, and 80oC for 5min. 1µL of RNase H were added to collected reactions and put back on thermocycler at 37oC for 20 min. A total of 21µL cDNA was collected and kept in -20oC until ready to use.

4.2.3. DNA amplification

Samples were first tested for RVFV infection. RVFV was detected by conventional PCR using primers targeting conserved 90bp region of L segment (Fwd 5’-

TGAAAATTCCTGAGACACATGG-3’ and Rev 5’- ACTTCCTTGCATCATCTGATG-3’)

(LaBeaud et al. 2011). 1 μL of total cDNA was added to the PCR mixture of Qiagen

HotStar Taq DNA polymerase kit (Cat. No. 203205) containing 2.5 µL 10X PCR Buffer,

0.75 µL 50mM MgCl2, 0.55 µL 10mM dNTP, and 0.1 µL Taq as well as primers (1µL 10μM forward primer and 1 µL 10μM reverse primer) and 18.15 µL ddH2O. A total of this PCR mixture was put on Applied Biosystems 2720 thermal cycler under these cycling

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parameters used were 95oC for 15 min; 30 cycles of 94oC for 30 sec, 50oC for 30 sec,

72oC for 1 min and 72oC for 10 min (LaBeaud et al. 2011).

The amplicons were stained by GelRed (Biotum, Cat No. 41003), run on 2% agarose gel with 1X TAE buffer at 100V for 1 hour, and visualized with Bio View UV Light transilluminator. After samples were tested for RVFV, cDNA of RVF negative samples were pooled by 5-7 according to districts of origin and were tested for BUNV, BATV, and

NRIV using cycling parameters mentioned in Chap 2. Potential co-infection cases of

RVFV and these three Orthobunyaviruses were detected by testing RVF positive samples for BUNV, BATV, and NRIV as well. Detection of the three Orthobunyaviruses was done by targeting all the three segments S, M, and L for each virus by PCR. Primers and cycling parameters used are shown in Chapter 2 (Table 2.3). As mentioned previously in Chapter

2, there was a good cross-reaction between our L primers of BUNV and NRIV but not between M primers of BATV and NRIV. Thus, positivity of M primers was only required for homologous amplification and diagnosis was confirmed as shown in Table 4.1.

Table 4.1. Diagnostic scheme of BUNV, BATV, and NRIV using M and L primers.

BUNV BATV NRIV Conclusion

S M L S M L S M L + + + - - - + - + BUNV + + + + - - - + + + BUNV and NRIV co-infection + + + + + + + + + BUNV, BATV, and NRIV co-infection - - - + + + - - - BATV+ + - + + + + + + + BATV and NRIV co-infection + + + + + + + - + BUNV and BATV co-infection + - + - - - + + + NRIV+

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4.3. Results 4.3.1. Epidemic timing

The outbreak started in Ngoma and Kirehe districts (Eastern province) in the end of

May and many cases were still reported in these two districts at the end of the outbreak in the end of July. The spread continued in other parts of the country starting by other districts of Eastern province (Kayonza, Gatsibo, Rwamagana). The Southern province was also heavily affected followed by Northern province. Western province and Kigali city were affected late and at low level (Figure 4.2 and Figure 4.3). Animal movements were banned from mid-June to the end of July. Human cases were not documented except death of two veterinary officers; Ngoma on June 11 and Kamonyi on July 3. The outbreak transmission was stopped by massive vaccination campaigns of healthy and non- pregnant animals.

Figure 4.2. Administrative map of Rwanda depicting provinces and districts (NSRI 2019).

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Figure 4.3. Epidemic chronology of RVF outbreak.

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4.3.2. Rift Valley Fever prevalence

There were more cows sampled than goats, due to the economic and cultural importance of cows. Over 97% of samples were female, with the exception of 4 male calves included because their mothers had hemorrhagic fever symptoms and mothers have died within the last 3 days (Table 4.2). Approximately 79.5% of samples came from family farms compared to commercial farms. Cattle were identified as either exotic, local, or crossed breed. Almost 65% of cows sample were classified as crossed breed.

However, the study was not designed to test for differences in positivity among breeds.

According to owners, none of the sampled animals were vaccinated for RVF. 30.3%

(56/185) samples were positive to Rift Valley Fever. Of those that were positive for RVFV,

26.7% manifested in hemorrhagic disease while 82% presented with abortion. All cases of hemorrhagic disease were reported deceased 1 to 4 days after sample collection.

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Table 4.2. Sample categories and proportions of positive samples.

Samples categories Total number of RVF positive samples collected samples

Overall prevalence 185 56 (30.27%) Species Cattle 157 44 (28.02%) Goats 28 12 (48.85%) Sex Female 181 55 (30.38%) Male 4 1 (1/4) Type of farm Family 147 41 (27.89%) Commercial 38 15 (39.47%) Breeds (cattle) Local 16 6 (37.5%) Crossed 102 30 (29.41%) Exotic 39 8 (20.51%) Clinical symptoms Abortion 157 46 (29.29%) Hemorrhagic 15 15 (100%) fever Geographic Eastern Ngoma 41 10 (24.39%) distribution province Kirehe 32 7 (21.87%) Kayonza 18 6 (33.33%) Gatsibo 28 12 (42.85%) Rwamagana 5 4 (4/5) Southern Ruhango 17 9 (52.94%) province Kamonyi 15 4 (26.66%) Muhanga 3 - Nyanza 5 - Gisagara 1 1 (1/1) Northern Gakenke 5 - province Gicumbi 10 - Western Ngororero 1 1 (1/1) province Rusizi 1 1 (1/1) Kigali Gasabo 3 1 (1/3) city

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4.3.3. Identification of Orthobunyaviruses in Rift Valley Fever negative samples

Among the 129 RVF negative samples, 7 were positive to BUNV and/or BATV. Two samples were exclusively positive to BUNV, two were exclusively positive to BATV, while three were positive to all segments of both viruses. These 7 samples were either from

Southern (Nyanza, Ruhango, and Kamonyi districts) and Northern province (Gakenke and Gicumbi districts). Figure 4.4 shows the visualization on 2% agarose gel of DNA bands for amplified regions of BUNV and BATV S, M, L segments of these 7 samples.

Figure 4.4. Positive samples to BUNV and/or BATV and negative RVFV. Nyan-61 and Kam-143: positive to BATV only; Gak-84 and Gic-120: positive to BUNV only; Gic-127, Ruh-128, and Ruh-129: positive to both BUNV and BATV Nyan= Nyanza; Gak=Gakenke; Gic=Gicumbi; Ruh=Ruhango; Kam=Kamonyi

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4.3.4. Potential co-infection of Rift Valley Fever virus and Orthobunyaviruses

Ten (10) Rift Valley Fever positive samples had evidence of Orthobunyavirus co- infection. However, based on the diagnostic scheme for our three viruses of interest, none of them were decisively positive to either BUNV, BATV or NRIV. One sample was positive for two segments of BUNV.

Although these results are not conclusive, they are indicative of potential co-infection of multiple Bunyaviruses in Rwanda and potentially previously undetected reassortants.

Table 4.3. Potential co-infection of RVF and Orthobunyaviruses

Sample No RVFV BUNV BATV NRIV Conclusion: potential & District co-infection of RVFV L S M L S M L S M L and either BUNV or/and BATV or/and NRIV Gatsibo- + + + RVF+, BATV+/NRIV+ 102 Rwamagana- + + + RVF+, BUNV+/NRIV+ 106 + Rwamagana- + + + + + + RVF+, 108 BUNV+/BATV+/NRIV+ Rwamagana- + + + + + RVF+, 109 + BUNV+/BATV+/NRIV+ Ruhango- + + + RVF+, BUNV+/NRIV+ 132 + Kamonyi- + + + + RVF+, 144 BUNV+/BATV+/NRIV+ Ngoma- + + RVF+, BUNV+ 160 Ngoma- + + + RVF+, BUNV+/BATV+ 167 Kayonza- + + + RVF+, BATV+/NRIV+ 195 Kayonza- + + + RVF+, BATV+ 198

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4.4. Discussion

In this study, we identified RVFV as the causative agent of hemorrhagic fever and/or abortion in over 30% of cases during the largest outbreak documented in Rwanda. The outbreak highly affected the Eastern province of Rwanda, which was not surprising as it has been shown to be the most suitable region for mosquito life cycles. This region is characterized by multiple lakes, rice plantations, and the lowest altitude of the country

(<2,300m), low precipitation, and banana plantations in close proximity to households

(Demanou 2012). These ecological determinants, coupled with heat and humidity, make this region very suitable for mosquito breeding. For example, a mosquito survey conducted in 2011 showed that this region had the highest density of four of the most

One Health relevant mosquito species (in order of highest density to lowest): Culex spp,

Aedes spp, Coquilletidia spp, and Anopheles spp. Culex and Aedes are presumed primary and secondary vectors of RVFV and have been shown to be competent for these

Orthobunyaviruses (Odhiambo et al. 2014). All districts of Eastern provinces were affected except Bugesera and Nyagatare districts, which have undergone very systematic vaccination campaigns in less than one year before the outbreak occurs. This underscores the need for high coverage vaccination programs to preserve the herd health of ruminants to RVFV.

In addition, we molecularly confirmed the presence of other Orthobunyaviruses, BUNV and BATV. We show that these viruses, in a small number of cases, caused abortions in ruminants, similar to RVFV. This suggests that these viruses are contributing to RVFV- like illnesses in Rwanda and may be present in neighboring countries. This is the first time another Bunyavirus other than RVFV is reported in Rwanda and the second time

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BATV is identified in Africa by molecular technique. BATV mainly circulates in Europe and Asia was isolated in Uganda in 1967 and only suspected in Sudan in 1988 based on serological evidence (East African Research Institute 1967, Nashed, Olson, and el-Tigani

1993).

This study represented the second molecular confirmation of BATV in an acutely ill vertebrate in Africa.

Importantly, this study demonstrated that cows could be co-infected with BATV and

BUNV. The provenance of NRIV is unknown but assumed to have occurred due to a co- infection with BUNV and BATV. This is largely assumed to have happened in the vector

(REFS). However, our study shows that this might have occurred in a cow.

Further, not only did we identify these viruses in Rwanda, but we demonstrate for the first-time co-infection with RVFV and unidentified Orthobunyaviruses. Our study was not sufficient to definitively identify these other Orthobunyaviruses, but this represents an important future direction, as many known and unknown arboviruses continue to cryptically circulate in Africa. In neighboring countries, other Orthobunyaviruses related to BUNV circulate including Ilesha and Germiston viruses (Yandoko et al 2007). This raises the question of whether novel reassortants could be occurring in the region.

This study reports the identification of BUNV and BATV in the following districts:

Gicumbi (BUNV infection and BUNV/BATV co-infection), Gakenke (BUNV infection),

Nyanza (BATV infection), Ruhango (BUNV/BATV co-infection), and Kamonyi (BATV infection) (Figure 4.5). RVFV was spread throughout the entire country, with higher density in the southeast of the country. The Orthobunyaviruses shared a similar distribution with RVFV, though obviously with less density.

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Figure 4.5. Geographic distribution of RVF and Orthobunyaviruses cases. Ortho: BUNV or BATV positive case, RVFV: RVFV positive case, RVFV+uOrtho: potential co-infection between RVFV and BUNV and/or BATV and/or NRIV, mOrtho: BUNV and BATV co-infection cases.

There is a general dearth of knowledge regarding the ecology of arbovirus transmission in Rwanda. Environmental suitability studies on potential vectors and role of environmental parameters should be conducted in order to understand transmission dynamics and offer control solutions. The scope of this study was a detection of viral RNA among acutely ill animals during an outbreak of RVFV-like illness. However, we speculate that seroprevalence will be higher in the general cattle population, and perhaps in the human population. Importantly, though RVFV was assumed ot be the causative agent of disease in this unprecedented outbreak, there was a high proportion of samples negative for all viruses tested here. The etiology of these cases could be due to any number of

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pathogens that share clinical manifestation, namely Brucellosis and anaplasmosis. These are important issues warranting further research in the region.

Overall, we demonstrate that RVFV was responsible for only 30% of tested samples with RVFV-like illness. We also show that BUNV and BATV co-circulate with RVFV and are capable of producing acute illness in ruminants. Further, our data suggests that co- infections among Orthobunyaviruses and between Orthobunyaviruses and RVFV occur, and that there may be other Orthobunyaviruses circulating in the region. Further work is needed to describe the biodiversity of arboviruses in Rwanda, the ecology of such

(including vectors, thermal limits of transmission, reservoirs, etc.), and the possibility of causing human disease.

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CHAPTER 5. GENERAL DISCUSSION AND CONCLUSIONS

5.1. Introduction

Although many laboratory and some natural reassortment events have been observed among members of the Bunyavirales order (Gentsch and Bishop 1976; Borucki et al.

1999; Bowen et al. 2001; Briese et al. 2006; Bodewes 2010; Aguilar et al. 2011, Briese,

Calisher, and Higgs 2013; Eiden, Vina-Rodriguez, El Mamy, Isselmou, Ziegler, Höper, et al. 2014; Hontz et al. 2015), few comparative have been studies are conducted on parental and progeny viruses resulting from the genome reassortments. In this study, we compared in vitro and in vivo kinetics of Bunyamwera (BUNV), Batai (BATV), and Ngari

(NRIV) viruses, three viruses of One Health concern belonging to Orthobunyavirus genus,

Peribunyaviridae family in the Bunyavirales order (Bowen et al. 2001; , Gerrard et al.

2004; Briese et al. 2006; Maes et al. 2019). NRIV is a progeny virus of BUNV and BATV

(Bowen et al. 2001; Briese et al. 2006; Yandoko et al. 2007; Groseth, Weisend, and

Ebihara 2012). In vitro studies were conducted in Vero cells, one of the most commonly used laboratory cell lines, and in vivo studies were conducted by infections in wildtype

C57BL/6 mice. In addition, this research project presents data collected from studies that investigated the hypothesis that these three Orthobunyaviruses co-circulate and contribute to the burden of disease in a Rift Valley Fever (RVF) endemic area, during the

RVFV outbreak of May-July 2018 in Rwanda. This hypothesis was based on evidence of circulation of these viruses in neighboring countries (Kokernot et al. 1958; Nashed, Olson, and el-Tigani 1993; Dutuze et al. 2018; Gerrard et al. 2004; Briese et al. 2006; Wertheim

2012; Otieno 2015), lack of consistent molecular confirmation of RVFV in acutely ill animals, and the documented similarity of clinical manifestations among diseases caused

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by RVFV, BUNV, BATV, and NRIV (Nashed, Olson, and el-Tigani 1993; Elliott 1996;

Briese et al. 2006; Hollidge, Gonzalez-Scarano, and Soldan 2010; Ikegami 2011; Elliott

2013; Eiden, Vina-Rodriguez, El Mamy, Isselmou, Ziegler, Hoper, et al. 2014; Soldan and

González-Scarano 2014).

In this chapter we summarize the findings and provide general conclusions and future perspectives as results from this study constitute a valuable baseline for deeper characterization of these understudied viruses.

5.2. Summary of results 5.2.1. In vitro infection kinetics, in vivo infection kinetics, and ex vivo stability of BUNV, BATV, and NRIV

We conducted in vitro studies which constitute a valuable tool for giving insight into infection kinetics in natural hosts (Chapter 2). Through in vitro studies of BUNV, BATV, and NRIV in Vero cells, we demonstrated that there was 1) similarity of plaque morphology between BUNV and NRIV more than the other pairwise comparisons, 2) stability of these three viruses in extracellular conditions at different inoculation doses up to 30 dpi, and 3) a general lack in differences among the three viruses of growth kinetics in Vero cells.

BUNV and NRIV plaques were found to be similarly heterogenous with a mixture of small, medium, and large plaques, while BATV generally produced homogenously large plaques. Studies have shown that isolates from different sized plaques of BUNV and

NRIV have different pathogenicity, as small BUNV plaques were more lethal in suckling mice while larger NRIV plaques were more lethal (Odhiambo et al. 2014). We speculate that the ability of these viruses to produce these small plaques with differential infectivity might account for the observed higher pathogenesis of BUNV and NRIV compared to

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BATV. This avenue of research warrants further investigation to determine if within host variants (i.e. plaque-size associated phenotypes) do actually cause higher disease.

BUNV, BATV, and NRIV were all found to be stable and infectious in extracellular conditions for up to 30 days. This suggests a potential of these viruses to be directly transmitted via environmental fomites, should they come into contact with mucous membranes or open wounds. Other members of the Bunyavirales order have been found to be directly transmitted in farms where abortive tissues, blood secretions (in cases of hemorrhagic fever), and various equipment play a significant role in transmission especially during outbreaks (Valero 2017, Hartman 2017, Stavropoulou and Troillet 2018,

Pshenichnaya and Nenadskaya 2015). The environmental stability of these viruses in field conditions relevant to the region (such as desiccation, humidity differences, and sunlight exposure) should be tested to provide more information regarding the transmissibility of environmentally stable viruses. The prolonged stability of these viruses led us to investigate the efficacy of inactivation of laboratory detergents. We report that

1%Triton-X-100 after one-hour incubation could inactivate the three viruses.

Comparative in vivo studies (Chapter 4) were conducted in C57Bl/6 mice, to determine if these mice were a good model for infection and/or disease. We compared the viremia status, antibody production, and cross-neutralization of antibodies following infection.

Viremia results showed that none of the BUNV or BATV infected mice were viremic, and only 1/5 NRIV infected mice showed what looked like viremia (higher viral output than initial input), though virus was detected for a much longer period of time in the NRIV infected mice than the other two groups. While these mice do not appear to be good models for infection, these results are suggestive of a potential lower virulence of BUNV

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and BATV compared to NRIV, which has been observed in human populations (Nashed,

Olson, and el-Tigani 1993; Gerrard et al. 2004; Briese et al. 2006).

There was at least some level of cross-neutralization among all pairs of the three viruses. BUNV and BATV had already been shown to cross-neutralize each other as they belong to the same serogroup (Bunyamwera serogroup) (Hunt and Calisher 1979).

However, ours is the first study of cross-neutralization of NRIV, and the first study to compare across both parental viruses and the NRIV progeny. We report a strong neutralization of BATV by NRIV, which was expected as these two viruses share M segment which encodes glycoproteins, which serve as triggers for neutralizing antibody production (Bowen et al. 2001; Gerrard et al. 2004; Groseth, Weisend, and Ebihara 2012).

However, serum from BATV exposed mice did not neutralize NRIV and BUNV well, nor itself. The low level of BATV homologous neutralization is likely indicative of low antibody production by the strain of the mouse strain used in our experiment (C57BL/6) as they quickly cleared the virus (Figure 3.6.A). The cross-neutralization of these viruses is beneficial for exposed individuals living in areas where these viruses co-circulate as one infection could provide protection for future infections of all the three viruses. Additionally, given that neutralizing antibodies are crucial for vaccine-mediated protection against viral diseases (Burton 2002), existence of cross-neutralization may provide the give the possibility of developing common vaccine, though there may be different efficacy between viruses. We speculate that cross-neutralization will likely confound serological diagnostic development and could be a reason why BATV is less reported in Africa.

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5.2.2. Identification of Orthobunyaviruses in Rwanda

The general status of Bunyaviruses and arboviruses in general circulating in

Rwanda is not well characterized. The only Bunyavirus which was known to circulate in

Rwanda before our study was Rift Valley Fever Virus and only with evidence of circulation since 2011 (Rwanda Agriculture Board 2013). In Chapter 4 of our study, we provide evidence that BUNV and BATV both circulate in Rwanda contributing to RVF-like disease.

In addition, we could not rule out the presence of NRIV as co-infection with BUNV and

BATV may mask the additional presence of NRIV. In addition to these confirmed cases, we also report cases of non-conclusive combinations of genome segments of

Orthobunyaviruses, which indicates the presence of these viruses and potentially new reassortment events. Further characterization of these genomes should be made in an effort to determine if they are known virus sequences or unknown serogroup members.

The scarcity of molecular confirmation compared to clinical diagnosis in outbreak settings can lead to misdiagnosis. In the case of these bunyavirus infections (and more generally, arbovirus infections), disease is often characterized by non-specific clinical signs or by signs that are attributed to a known agent, such as RVFV.

RVFV, for example, was confirmed for the first time in 2011 in Rwanda after being suspected due to uncommonly high incidence of abortions concentrated in one part of the country, following the known seasonality of RVFV from other regions (Rwanda Agriculture

Board 2013; Umuhoza et al. 2017). Given that many cases of RVFV infections are characterized by non-specific mild illnesses (Easterday 1965; Ikegami 2011), it is possible that it was already circulating long before 2011 in less intense or even asymptomatic forms. This is very important as education and affordability to veterinary services play

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important role in requesting veterinarian services in rural areas of Rwanda. Most farmers don’t usually notify veterinarians for mild forms of diseases, which means that transmission may go unabated.

In May-July 2018, at the time of our study, an unprecedented outbreak of RVFV-like illness occurred in Rwanda (OIE 2018). The high transmission of RVFV was the consequence of exceptionally heavy rain in March and April causing flooding events. This was commonly seen in other countries in East Africa (Kenya, Uganda, and Tanzania)

(OIE 2018, WHO 2018).

This outbreak allowed us to test our hypothesis that these Orthobunyaviruses may contribute to RVFV-like illness in real-time. We were able to conclusively detect conclusively cases of BUNV (2/127), BATV (2/127), and BUNV/BATV co-infections

(3/127) in RVFV negative samples. Further, there were non-conclusive combinations of

RNA genome segments of BUNV, BATV, and NRIV by PCR in RVFV positive samples

(10/56). While the RVFV cases were predominantly reported in Eastern provinces, all

Orthobunyaviruses conclusive cases were were found in Northern and Southern provinces (map, Figure 4.2) and non-conclusive cases found in Southern and Eastern provinces. Although the overall reported prevalence of Orthobunyaviruses was low, their presence itself is indicative of emergence of subsequent surveillance considerations; and the geographic differences in cases may indicate an ecological niche divergence which could be used for targeted surveillance for Orthobunyaviruses in future.In addition, given these ecological differences, further studies are needed to determine the vector ranges of these viruses, as mosquito biodiversity, populations, and abundance differ across

Rwanda. Further, the actual prevalence of these viruses in Rwanda could be greater than

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what we observed due to the asymptomatic aspect of bunyaviral diseases (Elliott 1996;

Elliott 2013; Soldan and González-Scarano 2014).

In the human population, no evidence of RVFV has been shown in Rwanda. But this is highly suspected due the proximity between humans and animals, especially in rural areas of Rwanda. Fever which is the most common manifestation of Bunyaviral disease in its mild form, is usually diagnosed as malaria without confirmatory laboratory testing

(Amexo et al. 2004). The only systematically studied arbovirus in the human population is yellow fever and was reported at very low seroprevalence (0.2%, 2/1284 participants)

(Demanou 2011). The same assessment study reported a very small population of Aedes spp mosquitoes (190 out of 1971 mosquito collected all over the country (Demanou

2011). Without molecular confirmation, it may be that a proportion of human febrile illnesses in Rwanda is attributable to one of these or other Orthobunyaviruses, and serological surveys could ascertain the exposure rate in human populations as well.

The non-conclusive combinations of genome segments could be indicative of possible reassortment events between these viruses or the presence of other Orthobunyaviruses known to circulate in the region such Ilesha, Germiston, and Shokwe (Yandoko et al.

2007). In addition, the non-conclusive results could also indicate the presence of uncharacterized Orthobunyavirus species or reassortment progeny. Further studies in the region would allow for the identification and study of the relatively neglected group of

Orthobunyaviruses currently circulating in the region.

5.3. Conclusions and future perspectives

Human-animal disease transmission in Rwanda is facilitated by the fact that livestock animals are found in close proximity to many households, especially in rural areas of

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Rwanda. This interaction is also enhanced by some Rwandan Government programs for alleviation of poverty and malnutrition such as “Girinka” (One cow per family) which promotes the owning of cows in poor families to increase access to animal proteins at low cost (Rwanda Government Board 2018). This proximity increases the risk of zoonotic disease transmission while simultaneously providing a benefit to the family.

This work significantly contributes to the increase in knowledge available on

BUNV, BATV, and NRIV as it characterizes the comparative in vitro and in vivo infections of these viruses and contributes to identification of these viruses in new geographic areas.

Based on the findings of our study, we recommend enhanced laboratory inactivation practices when handling these viruses due to their stability in extracellular medium, and we further confirm that the use of 1% Triton-X-100 successfully inactivates these viruses.

We further suggest that, surveillance efforts in Rwanda and other East African countries should allow for pathogen discovery whenever possible, though we recognize the considerable challenges that may be faced.

Finally, we highlight the need for more investigation of these and other under- characterized arboviruses. In infection kinetics, more research is needed to determine the mechanisms for observed higher virulence of NRIV compared to BATV and BUNV. For the transmission dynamics of these viruses in Rwanda, more investigation is needed on the comparative ecologies of these viruses, including the host ranges (humans, livestock species, and wildlife), the role of hosts in epidemiology (reservoirs, dead-end hosts, and amplification hosts), identification of competent vectors, and the identification of other

Bunyaviruses circulating in the region.

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This problem exemplifies the concept of One Health where animal health – here, because of the direct benefit of the animal to the issue of child-growth stunting, and the increase risk of zoonotic transmission – is a direct influencer of human health. Further, as we have shown the geographical distribution of these viruses is different, the environmental factors associated with these geographies likely contributes to the risk of transmission (as with flooding events and increased RVFV transmission). Our study points to the necessity for a One Health approach to disease surveillance and mitigation, especially in developing nations.

5.4. References

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Bodewes, R., Ruiz-Gonzalez, A., Schürch, A. C., Osterhaus, A. D., & Smits, S. L. 2010. "Ngari Virus in Goats during Rift Valley Fever Outbreak, Mauritania." Emerging infectious diseases 20 (12):2172.

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Elliott, R. M. . 1996. The Bunyaviridae. New York, N.Y: Plenum Press.

Elliott, Richard M. 2013. The bunyaviridae: Springer Science & Business Media.

Gentsch, J., and D. H. Bishop. 1976. "Recombination and complementation between temperature-sensitive mutants of a Bunyavirus, snowshoe hare virus." J Virol 20 (1):351-4.

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OIE. 2018 "WAHIS Interface, Weekly disease report ". www.oie.int

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Yandoko, E. N., S. Gribaldo, C. Finance, A. Le Faou, and B. H. Rihn. 2007. "Molecular characterization of African orthobunyaviruses." J Gen Virol 88 (Pt 6):1761-6. doi: 10.1099/vir.0.82643-0.

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APPENDIX A. FRONTIERS COPYRIGHT STATEMENT

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APPENDIX B. AREAS UNDER THE CURVES (AUC) FITTED BY GROWTHCURVER FUNCTION IN R

B.1 AUC of BUNV in standard cell culture conditions (10%FBS)

(Appendix B.1 cont’d.)

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B.2 AUC of BATV in standard cell culture conditions (10%FBS)

(Appendix B.2 cont’d.)

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B.3 AUC of NRIV in standard cell culture conditions (10%FBS)

(Appendix B.3 cont’d.)

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B.4 AUC of BUNV in sub-standard cell culture conditions (2%FBS)

(Appendix B.4 cont’d.)

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B.5 AUC of BATV in sub-standard cell culture conditions (2%FBS)

(Appendix B.5 cont’d.)

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B.6 AUC of NRIV in sub-standard cell culture conditions (2%FBS)

(Appendix B.6 cont’d.)

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VITA

Marie Fausta Dutuze was born and grew up in the beautiful mountains of Shyorongi in the Northern part of Rwanda. She lived in Rwanda until 2006 when she finished high school at Lycée Notre Dame de Cîteaux in Kigali. After being ranked among the top performing students at country level in the National Examination, she was awarded a scholarship of excellence by the Government of Rwanda to pursue veterinary medicine at Ecole Inter-Etats des Sciences et Médecine Vétérinaires de Dakar in Senegal. She then moved to Senegal where she stayed for 7 years and completed DVM and master’s in public health degrees. She graduated of DVM degree in 2011 and master’s degree in

2013. Immediately after accomplishing master’s, she went back to Rwanda and worked as assistant lecturer at the University of Rwanda in the College of Agriculture, Animal

Sciences and Veterinary Medicine (UR/CAVM) where she is still working. In 2015, she was awarded USAID scholarship under a Borlaug Higher Education in Agricultural and

Research Development (BHEARD) program to pursue PhD program at Louisiana State

University. She was fortunate to join Dr. Rebecca C. Christofferson laboratory where she gained significant knowledge and skills in epidemiology of arboviruses. Fausta anticipates graduating in August 2019. After graduation, she will continue her career in academia and research with special interest in epidemiology of infectious diseases in One Health perspective

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