Modulation of Inflammatory Cytokine, Chemokine, and Toll

University of Kentucky UKnowledge

Theses and Dissertations--Veterinary Science Veterinary Science

2019

MODULATION OF INFLAMMATORY CYTOKINE, CHEMOKINE, AND TOLL-LIKE RECEPTOR GENES AND TRANSCRIPTOME ANALYSIS OF EQUINE ENDOTHELIAL CELLS FOLLOWING INFECTION WITH EQUID HERPESVIRUS-1, AND EQUINE ARTERITIS VIRUS.

Saranajith Wangisa Dunuwille University of Kentucky, [email protected] Digital Object Identifier: https://doi.org/10.13023/etd.2019.388

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Recommended Citation Dunuwille, Saranajith Wangisa, "MODULATION OF INFLAMMATORY CYTOKINE, CHEMOKINE, AND TOLL- LIKE RECEPTOR GENES AND TRANSCRIPTOME ANALYSIS OF EQUINE ENDOTHELIAL CELLS FOLLOWING INFECTION WITH EQUID HERPESVIRUS-1, AND EQUINE ARTERITIS VIRUS." (2019). Theses and Dissertations--Veterinary Science. 44. https://uknowledge.uky.edu/gluck_etds/44

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The document mentioned above has been reviewed and accepted by the student’s advisor, on behalf of the advisory committee, and by the Director of Graduate Studies (DGS), on behalf of the program; we verify that this is the final, approved version of the student’s thesis including all changes required by the advisory committee. The undersigned agree to abide by the statements above.

Saranajith Wangisa Dunuwille, Student

Dr. Udeni Balasuriya, Major Professor

Dr. Daniel K. Howe, Director of Graduate Studies

MODULATION OF INFLAMMATORY CYTOKINE, CHEMOKINE, AND TOLL- LIKE RECEPTOR GENES AND TRANSCRIPTOME ANALYSIS OF EQUINE ENDOTHELIAL CELLS FOLLOWING INFECTION WITH EQUID HERPESVIRUS- 1, AND EQUINE ARTERITIS VIRUS.

______

DISSERTATION ______

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the College of Agriculture, Food and Environment at the University of Kentucky

By Saranajith Wangisa Dunuwille Lexington, Kentucky Co- Directors: Dr. Udeni Balasuriya Professor of Virology and Dr. Peter J. Timoney Professor of Veterinary Science Lexington, Kentucky 2019

ABSTRACT OF DISSERTATION

MODULATION OF INFLAMMATORY CYTOKINE, CHEMOKINE, AND TOLL- LIKE RECEPTOR GENES AND TRANSCRIPTOME ANALYSIS OF EQUINE ENDOTHELIAL CELLS FOLLOWING INFECTION WITH EQUID HERPESVIRUS- 1, AND EQUINE ARTERITIS VIRUS.

EHV-1 is a double-stranded DNA virus whereas EAV is a positive sense, single- stranded RNA virus. Therefore, genetically, they are very different from one another. However, both these viruses are endotheliotropic and thus, infect and replicates in equine endothelial cells resulting in vasculitis. Vasculitis is central to the pathogenesis of these two viruses. Thus, the main objective of this thesis was to investigate the inflammatory and innate immune responses of EECs that contributes towards the development of vasculitis following infection with EHV-1 and EAV in-vitro. Since proinflammatory cytokines and chemokines produced by endothelial cells play a significant role in the development of vasculitis, we investigated their gene expression as well as secretion. Results from this study showed that the proinflammatory response of EECs induced by EAV is relatively less when compared with the corresponding results from EHV-1 infected EECs. Furthermore, EAV elicits a lower type I interferon response in EECs when compared with EHV-1. Further investigations revealed an active role played by TLR 3 in inducing the proinflammatory response in EHV-1 infected EECs during the first 6 hours of infection but not in EAV infected EECs. Analyzing the whole transcriptome of EHV-1 and EAV infected EECs revealed a complex pattern of gene regulation and cellular pathways related to cellular immune, inflammatory and apoptotic responses. Finally, we investigated host genetic factors associated with EHV-1 induced myeloencephalopathy but found no evidence for a recessive allele influencing the development of EHM following EHV-1 infection for any genetic locus was identified. However, more complex host-pathogen interactions are possible. Key words: EHV-1, EAV, TLR3, Type I interferons, RT-qPCR, Luminex, si-RNA, RNAi, GWAS, RNA-seq

Saranajith Wangisa Dunuwille

August 30th, 2019

MODULATION OF INFLAMMATORY CYTOKINE, CHEMOKINE, AND TOLL- LIKE RECEPTOR GENES AND TRANSCRIPTOME ANALYSIS OF EQUINE ENDOTHELIAL CELLS FOLLOWING INFECTION WITH EQUID HERPESVIRUS- 1, AND EQUINE ARTERITIS VIRUS.

By Saranajith Wangisa Dunuwille

Dr. Udeni Balasuriya Co-Director of Dissertation

Dr.Peter J. Timoney Co-Director of Dissertation

Dr. Daniel Howe Director of Graduate Studies

08/30/2019

ACKNOWLEDGEMENTS

It is with great pleasure that I acknowledge the roles of several individuals who were instrumental in providing me with guidance to complete my Ph.D. research work.

Firstly, I would like to express my most sincere gratitude to my advisor, Dr. Udeni B.R.

Balasuriya, for his continuous support throughout my Ph.D. program and research work, for his patience and motivation. I am forever grateful for him for granting me the opportunity to learn and grow as a researcher under his guidance and I could not have imagined having a better advisor and a mentor for my Ph.D. study than Dr. Balasuriya.

I would also like to express my deepest gratitude to Dr. Peter J. Timoney for his advice, motivation and generous support throughout my Ph.D program which helped me immensely in completing my Ph.D. study successfully and in a timely manner.

I would like to express my sincere gratitude to my Ph.D. dissertation committee members:

Dr. Rebecca Dutch, Dr. Thomas Chambers, Dr. Frank Cook and Dr. Kristine Urschel for their time, interest and helpful guidance that paved the way for the successful completion of my Ph.D. research work as well as the Ph.D. examination.

My most sincere thanks also go out to Dr. David W. Horohov and Dr. Amanda Adams for generously providing me access to their lab space and equipment which helped me immensely in completing my Ph.D. research work.

I am indebted to Dr. Ernest Bailey, whom without the GWAS analysis mentioned in this dissertation would not have been possible.

My most sincere gratitude also goes out to Dr. Bettina Wagner at the Cornell University,

Ithaca, NY. in helping me with Luminex assays.

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I would also like to thank Dr. Pouya Dini in helping me with completing the transcriptome data analysis mentioned in this dissertation.

My sincere gratitude also goes out to Dr. Michael Singleton for helping me with the statistical analysis mentioned in chapter 2 of this dissertation.

I am very grateful to Dr. Daniel Howe for all the help and guidance he gave me in the capacity as the director of graduate studies in order to complete my Ph.D. program successfully.

I would also like to thank my fellow lab mates, Dr. Mariano Carossino, Bora Jenny Nam and Annet Kyomuhangi for all the help and support given to me to make my Ph.D. research work a success.

I would also like to express my heartfelt gratitude to Dr. Sanjay Sarkar, Dr. Shankar

Mondal and Dr. Lakshman Chelvarajan for all the guidance and help they gave me when I first joined Dr. Balasuriya’s lab. Their guidance was instrumental for me to learn laboratory as well as molecular biology techniques and methods which helped me in completing my

Ph.D. research work.

Finally, I would like to express my most sincere gratitude to Pamela Henney, Kathleen M.

Shuck, Ashley Skillman, Day Barker and Craig Stewart for helping and guiding me around many a strange laboratory equipment and protocols that was an immense help in completing my Ph.D. research work successfully.

Last but not least, I would like to express my heartfelt gratitude to my parents for all the love and support they have given me my whole life.

TABLE OF CONTENTS Page ACKNOWLEDGEMENTS ...... iii LIST OF TABLES ...... viii LIST OF FIGURES ...... x LIST OF ABBREVIATIONS ...... xiii

CHAPTER

1 Literature Review...... 1 1.1 History of Horses ...... 1 1.2 Equine industry of the United States of America ...... 2 1.3 Impact of infectious diseases on the equine industry ...... 3 1.4 Viruses that cause respiratory and reproductive diseases of the horse ..4 1.4.1 Equine Influenza virus (EIV) ...... 4 1.4.2 Equine rhinitis virus (ERV) ...... 5 1.4.3 Equine arteritis virus (EAV) ...... 5 1.4.4 Equine herpesvirus-1 ...... 6 1.4.5 Equine herpesvirus-2 (EHV-2) ...... 6 1.4.6 Equine herpesvirus-4 (EHV-4) ...... 7 1.4.7 Equine herpesvirus-5 (EHV-5) ...... 8 1.4.8 Equine Adenovirus type-1 (EAdV-1) ...... 8 1.5 Viruses that cause neurological conditions in horses ...... 9 1.5.1 West Nile Virus (WNV) ...... 9 1.5.2 Equine Rabies ...... 10 1.5.3 Equine herpesvirus myeloencephalopathy (EHM) ...... 10 1.6 Persistent viral infections ...... 11 1.6.1 EAV persistent infection ...... 13 1.6.2 EHV-1 latent life cycle ...... 16 1.7 Equine arteritis virus ...... 18 1.7.1 EAV classification ...... 18 1.7.2 EAV virion architecture ...... 18 1.7.3 EAV genome organization ...... 19 v

1.7.4 EAV viral protein ...... 21 1.8 Equine viral arteritis ...... 22 1.8.1 Clinical signs ...... 23 1.8.2 Acute infection ...... 24 1.8.3 Reproductive consequences...... 25 1.8.4 EAV vaccination and management ...... 26 1.9 Host immune response against EAV ...... 28 1.9.1 Innate immune response ...... 28 1.9.2 Humoral immune response ...... 30 1.9.3 Cell mediated immune response ...... 31 1.10 Equine herpesvirus-1 (EHV-1) ...... 32 1.10.1 EHV-1 classification ...... 32 1.10.2 EHV-1 virion architecture ...... 32 1.10.3 EHV-1 genome organization ...... 34 1.10.4 EHV-1 viral proteins ...... 35 1.11 EHV-1 infection in horses and diseases ...... 37 1.11.1 EHV-1 respiratory disease ...... 37 1.11.2 Abortion and neonatal diseases ...... 38 1.11.3 Equine herpesvirus myeloencephalopathy (EHM) ...... 39 1.11.4 Vaccination and management...... 41 1.12 Host immune response against EHV-1 ...... 43 1.12.1 Innate immune response ...... 43 1.12.2 Humoral immune response ...... 45 1.12.3 Cell mediated immune response ...... 46 1.13 Inflammation and its mediation ...... 47 1.13.1 Cytokines ...... 48 1.13.2 Chemokines ...... 49 1.13.3 Toll like receptors ...... 50 1.13.3.1 Equine Toll like receptors ...... 51 1.14 Genomics in infectious diseases ...... 52 1.14.1 Transcriptome in infectious diseases ...... 53

1.14.2 Genome wide association studies (GWAS) in infectious diseases ...... 55 1.15 Scope and outline of dissertation ...... 56

2 Comparison of the pro-inflammatory gene modulation in equine endothelial cells by equid herpesvirus-1, equid herpesvirus-4 and the equine arteritis virus

2.1 Summary ...... 60 2.2 Introduction ...... 61 2.3 Material and methods ...... 63 2.4 Results ...... 70 2.5 Discussion ...... 76 Figures and tables ...... 81

3 RNA-Sequence Analysis of Equine Endothelial Cells Following In Vitro Infection with Equine Herpesvirus-1 and Equine Arteritis Virus Strains

3.1 Summary ...... 88 3.2 Introduction ...... 89 3.3 Materials and methods ...... 91 3.4 Results ...... 95 3.5 Discussion ...... 106 Figures and tables ...... 111

4 Involvement of Toll-like receptors in the innate immune response of equine endothelial cells following EHV-1 and EAV infections

4.1 Summary ...... 124 4.2 Introduction ...... 125 4.3 Materials and methods ...... 127 4.4 Results ...... 131 4.5 Discussion ...... 134 Figures and tables ...... 138

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5 Genome-wide association study (GWAS) to identify host genetic factors associated with equine herpesvirus type-1 (EHV-1) induced myeloencephalopathy (EHM) in horses

5.1 Summary ...... 141 5.2 Introduction ...... 142 5.3 Materials and methods ...... 143 5.4 Results ...... 144 5.5 Discussion ...... 146 Figures and tables ...... 148 6 General Discussion ...... 152 7 Appendix 1 ...... 159 8 References ...... 162 9 Vita ...... 195

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LIST OF TABLES Page Table 2.1: Average virus titers with the standard deviations of the four virus strains used in this study at different timepoints. These values were obtained from 3 independent experiments...... 81

Table 2.2: The p values of statistically significant virus titers (p<0.05) among different virus strains at different timepoints are shown. Statistical analysis was done using the natural log values of the log 10 TCID50 values obtained from cell culture supernatants. NS indicates timepoints with no statistical significance...... 81

Table 2.3: Average fold change in gene expression ± SD in EECs following EHV-1 (T953 and T262), EHV-4 (T445) and EAV. This data is from three independent biological replicates for each of the virus strains. Bold texts show where the increase of fold change from 6 to 12 hpi was statistically significant (p≤0.05) ...... 82 Table 2.4: Average gene expressions that are significantly different between EHV-4 (T445) and EAV at 6 and 12 hpi. Bold texts indicate the higher fold change ...... 83 Table 2.5: Average gene expressions that are significantly different between EHV-1 (T262) and EAV at 6 and 12 hpi. Bold texts indicate the higher fold change ...... 83 Table 2.6: Average gene expressions that are significantly different between EHV-1 and EAV at 6 and 12 hpi. Bold texts indicate the higher fold change ... 83 Table 3.1: List of the top 5 pathways and the differentially expressed genes in EAV 6 hpi vs mock 6 hpi comparison ...... 111

Table 3.2: List of the top 5 pathways and the differentially expressed genes in EAV 12 hpi vs mock 12 hpi ...... 112 Table 3.3: List of the top 5 pathways and the differentially expressed genes in EAV 6 hpi vs EAV 12 hpi (A) and EHV 6 hpi vs mock 6 hpi (B) comparison.113

Table 3.4: List of the top 5 pathways and the differentially expressed genes in EHV 12 hpi vs mock 12 hpi (A) and EHV 6 hpi vs EHV 12 hpi (B) comparison...... 115

Table 3.5: List of the top 5 pathways and the differentially expressed genes in EHV 6 hpi vs EAV 6 hpi (A) and EHV 12 hpi vs EAV 12 hpi (B) comparison...... 117

LIST OF FIGURES Page

Figure 1.1: EHV-1 latent cycle. (A) Following the production of progeny virions, virus particles enter innervating sensory neurons and migrates to the nucleus of the neuron. The viral DNA gets released and circularized once inside the nucleus and latency associated transcripts are expressed. (B) During reactivation, the genes that promote the lytic life cycle are expressed promoting the production of infectious virus particles. Infectious virus particles are released from the axon resulting in recurrent infection and virus shedding. Adapted and reproduced with permission from Springer Nature: Nature Reviews Microbiology [2], copyright 2008 ...... 17 Figure 1.2: Virion architecture of EAV consists of a nucleocapsid and 7 envelope proteins. Major envelope proteins (GP5 and M) forms dimers while 3 of the 5 minor envelope proteins (GP2, GP3 and GP4) forms trimers. E and ORF5a are also minor envelope proteins. Adapted and reproduced with permission from Elsevier: [Vet Clin North Am Equine Pract.][5], copyright 2014 ...... 19

Figure 1.3: Genome organization of EAV depicting the ORFs and the respective proteins coded by them. The 5' proximal three-quarters of the genome is occupied ORF1a and ORF1b which gets translated into two polyproteins which gets autocatalytically processed by three viral proteases (nsp1, 2 and 4) to generate EAV nonstructural proteins. The nested set of mRNAs are also indicated. RNA 1 is identical to the EAV genome and RNAs 2-7 code for the EAV structural proteins. The pink boxes represent the body TRS sequences and the light blue box at the 5' end of each sg mRNA represents the common leader sequence. The sgmRNAs 2 and 5 are bicistronic while the rest are functionally monocistronic. Adapted and reproduced with permission from Elsevier: [Veterinary Microbiology][6], copyright 2013...... 20 Figure 1.4: Virion architecture of EHV-1 consists of a core containing viral genomic DNA, a capsid, tegument and an envelope with 12 embedded glycoproteins. Adapted and reproduced with permission from Bentham Open: [The Open Veterinary Science Journal][1], copyright 2008...... 33

Figure 1.5: Genome organization of EHV-1. The UL region is flanked by TRL and IRL sequences. The US region is flanked by TRS and IRS sequences. The UL region contains ORFs 1-63 and the US region contains ORFs 68-76. The TRS and IRS regions contain copies of ORFs 64-67 [3; 4]...... 34

Figure 2.1: Virus growth curves of EHV-1 (T262), EHV-1, EHV-4 (T445) and EAV in EECs ...... 84 Figure 2.2: IFNα production by EECs infected with EHV-1 (T262 and T953), EHV4 (T445) and EAV. * indicates statistically significant increase from mock at

12hpi (p≤0.05). #* indicates the increase in IFNα production from 6 to 12 hpi is statistically significant (p ≤0.05) ...... 84 Figure 2.3: IL17 production by EECs infected with EHV-1 (T262 and T953), EHV4 (T445) and EAV. The increase in IL17 production from 6 to 12 hpi is statistically not significant (p ≥0.05)...... … 85 Figure 2.4: CXCL10 production by EECs infected with EHV-1 (T262 and T953), EHV-4 (T445) and EAV. # indicates statistically significant increase from mock at 6hpi (p≤0.05). * indicates statistically significant increase from mock at 12hpi (p≤0.05). #* indicates statistically significant increase from 6 to 12 hpi (p≤0.05)...... 85 Figure 2.5: Flow cytometry results of the percentage of virus infected EECs when compared to uninfected EECs (A). EHV-1 (T262) infected EECs (B) with a 41.21% infection rate, EHV-1 infected EECs (C) with a 29.35% infection rate and EAV infected EECs (D) with a 45.43 % infection rate...... 86 Figure 3.1: Heat map of the gene expression of EHV-1, EAV and mock infected EECs...... 119 Figure 3.2: Principal components analysis of genes of EHV-1, EAV and mock infected EECs at both 6 and 12 hpi timepoints. The clustering of the genes of EAV infected EECs at the 6 hpi timepoint shows significant variance. No such variance observed in the clustering of EHV-1 or mock infected EECs. .. 120 Figure 3.3: Differential gene expression of EHV-1 infected EECs when compared with mock infected EECs at the 6 (A) and 12 (B) hpi timepoints and when compared among 6 and 12 hpi timepoints (C). Differential gene expression of EAV infected EECs when compared with mock infected EECs at the 6 (D) and 12 (E) hpi timepoints and when compared among 6 and 12 hpi timepoints (F)...... 121 Figure 3.4: Venn diagram illustrating the distribution of differentially expressed genes in EECs infected with; EAV at 6 and 12 hpi timepoints (A), EHV-1 at 6 and 12 hpi timepoints (B), EAV and EHV-1 at the 6 hpi timepoint (C) and EAV and EHV-1 at the 12 hpi timepoint (D)...... 122 Figure 4.1: Transcriptome results (FPKM values) of toll-like receptor expression in EHV- 1 and mock infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. * indicates a statistically significant increase from mock infected EECs (p≤0.05). ... 138 Figure 4.2: Transcriptome results (FPKM values) of toll-like receptor expression in EAV and mock infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. * indicates a statistically significant increase from mock infected EECs (p≤0.05) .... .138 Figure 4.3: Relative quantification values of toll-like receptor expression in EHV-1 (A) and EAV infected EECs. The red line marks the basal level of expression

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(RQ=1). * indicates a statistically significant increase from mock infected EECs (p≤0.05)...... 139 Figure 4.4: Relative quantification values of toll-like receptor expression in TLR 3 siRNA transfected and EHV-1 infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. The red line marks the basal level of expression (RQ=1). * indicates a statistically significant decrease in gene expression from mock infected EECs (p≤0.05)...... 139 Figure 3.5: Relative quantification values of toll-like receptor expression in TLR 3 siRNA transfected and EAV infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. The red line marks the basal level of expression (RQ=1). * indicates a statistically significant decrease in gene expression from mock infected EECs (p≤0.05)...... 139 Figure 5.1 Manhattan plot of the -log10 P values for correlation of SNPs with EHM based on recessive model genome-wide association (GWAS) study using 94 horses with 382,529 markers were analyzed using the Ecab 3.0 data. No significance was observed after adjusting for Bonferroni corrections ...... 148 Figure 5.2 Genome plot (Ecab 3.0) of -log10 P values for association test of runs of homozygosity (ROH; minimum 5 samples with minimum homozygosity length of 400 kb with at least 60 SNPs) Association test. No significant associations were found for any ROH...... 151

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LIST OF ABBREVIATIONS

A.H.C.F. - American Horse Council Foundation

B.C. – Before Christ

CAV – Cell associated viremia

CCKR – Cholecystokinin receptor

CNS - Central nervous system

CTL - Cytotoxic T-lymphocytes

DAMPs – Damage associated molecular patterns

EAdV-1 - Equine Adenovirus type-1

EAV – Equine arteritis virus

EECs – Equine endothelial cells

EGF - Epidermal growth factor

EHM – Equine herpesvirus myeloencephalopathy

EHV-1 – Equine herpesvirus-1

EHV-2 – Equine herpesvirus 2

EHV-4 – Equine herpesvirus-4

EHV-5 - Equine herpesvirus-5

EIV – Equine influenza virus

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ERV - Equine rhinitis virus

ETIF - EHV-1 α-gene transinducing factor

GWAS – Genome-wide association study hpi – Hours post infection

HSV-1 - Herpes simplex virus-1

ICTV - International Committee for the Taxonomy of Viruses

IE – Immediate early gene

INM – Inner nuclear membrane kb - kilobases

LATs - Latency associated transcripts

MHC-1 - Major histocompatibility complex class I

N- Nucleocapsid protein

NPC - Nuclear pore complex nsps – Non-structural proteins

ORF - open reading frame

PAMPs – Pathogen associated molecular patterns

PBMC - Peripheral blood mononuclear cells

PCP - Papainlike cysteine protease

PDGF - Platelet-derived growth factor

PRRSV - Porcine reproductive and respiratory syndrome virus

RdRp - RNA dependent RNA polymerase

RFS - Ribosomal frameshift site

ROCK-1 - Rho associated coiled-coil kinase 1

RTC - Replication/transcription complex

TLRs – Toll-like receptors

U.S. - United States

URT - Upper respiratory tract

VBS – Virulent Bucyrus strain

VZV - varicella zoster virus

Wnt - Wingless-integrated

WNV – West Nile virus

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CHAPTER ONE Literature Review 1.0.Introduction

1.1 History of horses

Fossil records of horses (family Equidae) dates back to 55 million years [7]. However, it is believed that at the end of the Pleistocene period around 11,000 years ago, wild horses of the New World went extinct but the Old-World wild horses and their relatives (zebras, onagers and asses) survived. Horses were re-introduced to the New World during the 16th century by Spanish explorers [8]. A key aspect in the progression of the human civilization is the domestication of plants and animals. Among the animal species domesticated by humans throughout the course of history, the dog and the horse hold major importance since they were not bred mainly as a food source [9; 10]. Throughout history, the main usage of horses was for transportation and warfare [9]. The common consensus is that the first instance of horse domestication took place between the 5th and 4th millennium BC in the Eurasian steppes [11]. Arguably, these domesticated horses descended from either

Tarpan or Przewalski’s horses [8]. Out of these two truly wild horse groups the Tarpans became extinct in the wild in the late 1800s but the Przewalski’s horses are still extant in the wild in the steppe regions of east Asia [8]. The domestication of horses became a popular and common practice among ancient human civilizations of the Old-World.

Evidence for the close association between horses and Old-World human cultures can be identified in the linguistics of proto-Indo-European cultures where the words for “horse” and “foal” were part of their vocabulary. Furthermore, petroglyphs from the bronze age, such as the ones found in central Asia depicts horses and horse drawn chariots demonstrating the importance of horses in human civilizations. Likewise, the 22-century

old Qin “terracotta” army discovered in China contains sculptures of horses bearing similarities to the Przewalki’s horses. This spectacular find indicates the importance and the usage of horses in Chinese culture. Therefore, since the second millennium, utilization of horses for draught work, transportation, sports and in warfare was wise spread in ancient

Asian and European cultures and since then until the industrial revolution horses have contributed immensely to the progression of the human civilization.

1.2. Equine Industry of the United States of America

The equine industry of the United States (U.S.) is a large, economically diverse industry with a significant contribution to the US economy. According to the results of the

2017 economic impact study carried out by the American Horse Council Foundation

(AHCF), the U.S. equine industry has generated approximately $122 billion in total economic impact which is an increase from $102 billion since 2005. The U.S. equine industry contributes around $50 billion in direct economic impact to the U.S. economy. It amounts to a direct employment impact of 988,394 jobs and a total employment impact of

1.7 million jobs. Furthermore, it contributes approximately $38 billion in direct wages, salaries, and benefits. Approximately there are around 7.2 million horses in the U.S., and they are being used for recreation, showing, racing and as draught horses. The states of

Texas, California and Florida occupy the first three spots respectively as the states with the greatest number of horses in the U.S. Moreover, around 32 million acres of land is owned by the U.S. horse industry while approximately 49 million acres of land is leased for horse- related uses (https://www.horsecouncil.org/resources/economics/). Kentucky is considered as horse country and the city of Lexington is considered as the horse capitol of the world.

The horse population in Kentucky is approximately 242,400 and it boasts $23.4 billion in

equine and equine-related assets. Furthermore, the state of Kentucky is associated with

35,000 equine operations and 1.1 million acres devoted to the equine industry

(https://www.horseproperties.net/state/kentucky).

1.3. Impact of infectious diseases on the equine industry

Horses in the U.S. are affected by a vast array of infectious agents including viral, bacterial, fungal and protozoal infectious agents. These infectious agents have the capacity to causes disease conditions in horses ranging from mild diseases which the horse recover fairly quickly to fatal conditions that may spread rapidly among horse populations and develop into outbreaks [12]. A direct consequence of infectious diseases in horses is the detrimental effect it can have on the U.S. equine industry, which is estimated to be around

$122 billion as well as the U.S. economy as a whole

(https://www.horsecouncil.org/resources/economics/). Infectious diseases such as West

Nile, Potomac Horse Fever, and equine herpesvirus myeloencephalopathy (EHM) imparts economic losses to the equine industry by causing fatalities in adult horses [13-16].

Abortions induced by infectious agents such as the equine herpesvirus-1 (EHV-1) and the equine arteritis virus (EAV) imparts severe economic losses by causing the elimination of valuable progeny as well as reproductive complications following infection [5; 6; 17-19].

Respiratory disease conditions caused by infectious agents such as EHV-1, EHV-4, EAV, and the equine influenza virus reduces the performance of race and show horses which in turn affects the economy of the equine industry [5; 6; 19; 20]. EHV-1 and EAV are two of the most prominent viruses causing reproductive and respiratory disease conditions in horses around the world. Collectively, these two viruses are a significant threat to the U.S. and the global equine industry as well as the focus of this dissertation.

1.4. Viruses that cause respiratory and reproductive diseases of the horse

As mentioned above, acute and chronic respiratory and reproductive diseases resulting from contagious infectious agents have a significant negative effect on the $122 billion equine industry of the U.S. (https://www.horsecouncil.org/resources/economics/)

[21]. The most prominent viral respiratory infectious agents of horses in North America include equine influenza virus (EIV), equine rhinovirus types 1 (ERhV1) and type 2

(ERhV-2), equine herpesvirus type-5 (EHV-5), equine herpesvirus type-4 (EHV-4), equine herpesvirus type-1 (EHV-1), equine herpesvirus type 2 (EHV-2), equine arteritis virus

(EAV), and the equine adenovirus while EHV-1, EHV-3 and EAV remains as major virus infections of the horse that results in reproductive consequences [21].

1.4.1. Equine Influenza virus (EIV)

Influenza viruses belong to the family Orthomyxoviridae and the horse is considered a “dead-end” host for influenza A viruses. Being a member of the influenza A viruses, equine influenza viruses are considered to be relatively stable in their natural host the horse [22]. Recent EIV outbreaks have been caused by the subtype H3N8 which was first isolated in 1963. However, prior to the emergence of this strain, the now extinct H7N7 subtype was considered the major cause of epidemics [23]. Horses of all ages are susceptible to equine influenza infections, but foals have a higher resistance towards infections due to the presence of maternal antibodies [20; 22]. The clinical signs of equine influenza usually include pyrexia, anorexia, nasal discharge and cough [22]. Equine influenza infections are rarely lethal but can be accompanied with complications such as secondary bacterial pneumonia, myositis, myocarditis, and limb edema [24]. Since 2000,

several major equine influenza outbreaks have taken place in the United Kingdom, South

Africa, India, Japan and Australia [22].

1.4.2. Equine rhinitis virus (ERV)

Equine Rhinitis virus (ERV) infections are a major cause of acute respiratory diseases in horses all around the world. ERVs belong to the family Picornaviridae and genus Aphthovirus. The seropositive nature of horses to ERVs vary depending on their age with horses over 12 months of age being more seropositive to the virus. The clinical signs of Equine rhinitis A (ERAV) and B (ERBV) viruses include fever, anorexia, nasal discharge, coughing, pharyngitis, and lymphadenitis of the head and neck and viremia [25].

Moreover, subclinical ERAV and ERBV infections may occur as well. Persistent virus shedding in urine and feces may occur following ERAV infections. If left untreated, ERAV and ERBV infections can lead to secondary bacterial infections of the upper respiratory tract. [26].

1.4.3. Equine arteritis virus (EAV)

Similar to EHV-1, EAV too is one of the most important viruses causing reproductive and respiratory diseases in horses and has the capacity to severely impact the

U.S. and the global equine industry [5; 6; 19; 27]. EAV belongs to the order Arteriviridae and genus Alphaarterivirus (https://talk.ictvonline.org/taxonomy/) and induces a flu-like syndrome as well as abortions. Furthermore, EAV establishes persistent infection in the reproductive tract of sexually matured stallions [5; 6; 19; 27]. The pathogenesis and disease conditions caused by EHV-1 is discussed in more detail in subsequent chapters.

1.4.4. Equine herpesvirus-1

EHV-1 belongs to the subfamily Alphaherpesvirinae and genus Varicellovirus [28].

It is one of the most important viruses causing reproductive and respiratory diseases in horses and has the capacity to severely impact the U.S. and the global equine industry [15;

17; 18]. The respiratory disease caused by EHV-1 is similar to the disease condition caused by EHV-4. But unlike EHV-4, EHV-1 can also induce abortions and equine herpesvirus myeloencephalopathy (EHM) [15; 17; 18]. Furthermore, EHV-1 establishes latent infections in the sensory nerves of the trigeminal ganglia and lymphocytes of the peripheral circulation as well as in lymph nodes [29-31] of infected horses. The pathogenesis and disease conditions caused by EHV-1 is discussed in more detail in subsequent chapters.

1.4.5. Equine herpesvirus-2 (EHV-2)

EHV-2 is a slow growing virus in the subfamily Gammaherpesvirinae and the genus Percavirus [28]. It is ubiquitous among horse populations and has been isolated from horse populations from many different countries including United Kingdom, Japan,

Australia, New Zealand, Switzerland, Germany, United States, Canada, Hungary and

Poland [32]. The significance of EHV-2 as an equine upper respiratory tract pathogen has not been studied extensively. However, several studies have implicated EHV-2 with upper respiratory tract disease, inappetence, lymphadenopathy, immunosuppression, keratoconjunctivitis, general malaise and poor performance [32]. A study done in

Argentina has revealed that 79.7% of horses were sero-positive for EHV-2 while a similar study has revealed that 67% of horses displaying signs of clinical respiratory diseases were positive for EHV-2 [32]. Furthermore, an experimental infection study has shown that

unvaccinated adult horses as well as foals develop respiratory symptoms following infection with EHV-2 [33].

1.4.6. Equine herpesvirus-4 (EHV-4)

Equine herpesvirus-4 belongs to the subfamily Alphaherpesvirinae and genus

Varicellovirus [28]. It is closely related to EHV-1 which belongs to the same genus sharing a nucleotide similarity between 55 - 84% and an amino acid similarity between 55 – 96% with EHV-1 [3; 4]. EHV-4 is one of the most important herpesviruses infecting horse populations around the world as well as a major cause of respiratory diseases in horses [17;

18]. It imparts severe economic losses to the global equine industry through the respiratory illness it causes as well as due to the lost time for training and performance [34].

Furthermore, it is reported that on rare occasion EHV-4 may cause isolated incidents of abortions [4]. Serological studies from around the world has attested to the relatively high prevalence of EHV-4 among horse populations [18; 35]. The pathogenesis of EHV-4 is quite different than that of EHV-1 and is most often limited to the upper respiratory tract resulting in mild rhinopneumonitis [17; 18; 36]. Even though rare, abortions due to EHV-

4 infections have been reported in the United States (US) and in the United Kingdom [18].

Studies have shown that EHV-4 can also infect and replicate in endothelial cells of local capillaries of lymph nodes and pulmonary vessels, and therefore, it has been suggested that the pathogenesis of EHV-4 abortions might be following a vasculitis-based mechanism similar to that of EHV-1 [37]. However, even though EHV-4 incidences are relatively higher than EHV-1 in Kentucky, abortions due to EHV-4 in the US are considerably low

(less than 1%) [38]. Interestingly, in Japan, it is believed that prior to 1967, only EHV-4 strains were responsible for abortions and respiratory diseases in horses. Clinical signs of

EHV-4 induced respiratory disease include pyrexia, anorexia, nasal and ocular discharge.

Secondary bacterial infections in the upper respiratory tract may be a contributory factor in the development of rhinopneumonitis [17; 18].

1.4.7. Equine herpesvirus-5 (EHV-5)

EHV-5 is a member of the subfamily Gammaherpesvirinae and the genus

Percavirus [28]. Interestingly, studies done on EHV-2 lead to the discovery of EHV-5 from horses in Australia with upper respiratory tract disease [39]. Following the initial discovery of EHV-5, several countries including New Zealand, Switzerland, Argentina and Germany has reported EHV-5 infections [40]. Apart from the upper respiratory tract disease caused by EHV-5, it is also implicated in the development of equine multinodular pulmonary fibrosis (EMPF), which causes pulmonary fibrosis in adult horses [41].

1.4.8. Equine Adenovirus type-1 (EAdV-1)

EAdV-1 is a nonenveloped, icosahedral, double-stranded DNA viruses of the family Adenoviridae and genus Mastadenovirus (https://talk.ictvonline.org/ictv- reports/ictv_9th_report/dsdna-viruses-2011/w/dsdna_viruses/93/adenoviridae). EAdVs were first isolated in the U.S. in 1969 and since then, two EAdV serotypes have been identified (EAdV-1 and EAdV-2) [42]. EAdV-1 is a major causative agent of upper respiratory tract infections in horses while EAdV-2 is associated with gastrointestinal tract infections in horses [42]. EAdV-1 has a worldwide prevalence and has been isolated frequently from nasopharyngeal secretions from foals and adult horses with or without respiratory disease [43]. In particular, several reports have indicated that EAdV-1 infections cause severe respiratory tract infections in Thoroughbred and Arabian foals

which can become life threatening [42; 43]. Fatal pneumonia has been reported in Arabian foals and Fell pony foals with severe combined immunodeficiency. However, EAdV-1 induced pneumonia has been reported in immunocompetent foals as well [42; 43].

1.5.Viruses that cause neurological conditions in horses

1.5.1. West Nile Virus (WNV)

West Nile virus was initially isolated in 1937 from the West Nile district of Uganda

[44]. WNV belongs to the family Flaviviridae and genus Flavivirus. It is an arbovirus transmitted by mosquitos and usually maintained in a bird-mosquito-bird transmission cycle [45]. However, occasional WNV transmission occur to other vertebrates such as horses and humans who are considered as “dead-end” hosts. These “dead-end” hosts do not develop sufficient viremia to re-infect mosquito species to sustain infectivity cycles.

Within the U.S., only a few species of Culex mosquito drive the transmission of the virus even though the virus has been detected in 65 different mosquito species and 326 bird species [44; 45]. WNV infected horses, in most cases, do not display a clinical illness other than fever. However, during recent WNV outbreaks, around 10% of infected horses displayed neurological disorders. The most common symptoms in horses include ataxia, paresis or paralysis of the limbs, recumbency, skin fasciculations, muscle tremors and rigidity, hyperesthesia and even aggressiveness [44].

1.5.2. Equine Rabies

Rabies belongs to the family Rhabdoviridae and genus Lyssavirus and it is an uncommon neurological disease of horses [46; 47]. However, human and animal rabies remains a notifiable condition in the U.S. since 1944 [46]. In the U.S., equine rabies is not a significant cause of equine deaths as it is in Central and South America. In 2017, 13 rabid horses and donkeys have been reported in the U.S. which is a significant decrease from the

23 reported in 2016. However, England, New Zealand, Hawaii, Australia, and some

Caribbean and Pacific Islands are reported as free from the disease. Most often, rabies gets transmitted to horses via a bite from a rabid dog although transmission due to encounters with rabid wild animals have also been reported. In North America, racoons, foxes and skunks are the main reservoir for rabies although bats have also been reported to transmit rabies to horses. Rabies is almost always fatal once clinical signs develop and in horses these symptoms include neurological signs, behavioral changes, lameness and colic.

Unlike in dogs, the furious form of rabies, which makes rabid animals aggressive, is not common in horses but the dumb form is. However, most cases of equine rabies exhibits fever, hyperesthesia, ataxia followed by recumbency and loss of anal sphincter tone and death is often due to cardiorespiratory failure [46; 47].

1.5.3. Equine herpesvirus myeloencephalopathy (EHM)

As mentioned above, EHV-1 is one of the most important equine herpesviruses that poses a significant threat to the U.S. and the global equine industry. One of the pathological manifestations of EHV-1 infection is EHM, which is a rare and fatal neurological condition that has a a 15-50% case-fatality rate [48; 49]. In Recent years, the occurrence of EHM has increased dramatically with outbreaks reported in the United States, South Africa, New

Zealand and several European nations, including the United Kingdom. Susceptibility to

EHM appears to be determined by several host factors including age, physical condition, immune status and whether the infection is a primary exposure [48; 50]. Furthermore, there is some evidence to suggest that a single point mutation in the DNA polymerase gene of some EHV-1 strains increase their ability to induce EHM following infection [51]. In most cases, horses that develop EHM display neurological signs which include dysmetria, ataxia and paresis of limbs, toe dragging, hypotonia of the tail and recumbency [15; 52].

1.6. Persistent viral infections

During persistent infections, the infected virus is not cleared and remains in cells of the infected host. Silent as well as productive infections are associated with persistent virus infections in the absence of host cell death or even extensive cell damage. Persistent virus- host interaction maybe of three types defined as latent, chronic and slow infections [53].

Several host organ systems are targeted by viruses that has the capacity to establish persistent infections [53]. Herpes viruses such as the Herpes simplex-1 (HSV-1), EHV-1 and varicella-zoster viruses establishes lifelong latent infections in the host nervous system.

HSV-1 and varicella-zoster viruses establishes latent infections in the dorsal root ganglia while EHV-1 utilizes the sensory nerves of the trigeminal ganglia, lymphocytes of the peripheral circulation as well as in lymph nodes. [29-31; 54]. Following reactivation, infectious virus particles travel along peripheral nerves to body surface and may initiate lesion (cold sore or zoster) [53; 54]. However, some viruses choose to persist in the central nervous system (CNS). Such sites of virus persistence can be regarded as a “dead-end” due to the little to no opportunity the virus has in shedding infectious virus particles to the outside. One reason why some viruses choose the CNS to establish persistent infections

maybe because host anti-viral mechanisms have restricted access to the CNS and therefore, persistent infection can be easily established [54]. Subacute sclerosing panencephalitis caused by the measles virus, progressive multifocal leukoencephalopathy caused by the human polyomavirus 2 establish persistent infection in the CNS [54; 55].

There is evidence to suggest the filoviruses such as the Ebola virus persists in the reproductive tract of infected individuals. However, in the case of the Ebola virus the host immune system clears virus material over time which may take from a few months to years

[55]. In more recent years, it was reported that individuals infected with the Zika virus may shed virus particles in semen for about 80 days while viral nucleic acids maybe detected for a period up to 6 months [56; 57]. Similarly, EAV persists in the male reproductive tract as well and virus infected stallions may shed the virus for a few months to lifelong [5; 6;

19].

Some viruses choose the renal system of the host to establish persist infection which enables them to readily spread to the outside when reactivated. The human cytomegalovirus utilizes the tubular epithelial cells in the kidney to establish persistent infection [54]. In addition, the renal epithelium is utilized by several other polyomaviruses such as the simian virus 40 (SV40), the K virus of mice, and the JC and BK viruses of humans to establish persistent infection [58].

The lymphoreticular system is another host system targeted by viruses to establish persistent infection. The Epsteen-Bar virus targets memory B cells which are long lived immune cells to establish persistent infection. Furthermore, the human immunodeficiency virus utilizes resting CD4+ T cells to maintain a dormant reservoir [54; 58].

The murine leukemia virus and the mouse mammary tumor virus which are retrovirus integrates its genome into the hematopoietic progenitor cells and mammary epithelial stem cells respectively. Since these viruses infects and integrates the viral genome into host stem cells the virus can persist for an indefinite period, providing a continuous supply of infectious virions [54; 58].

Interestingly, the bovine viral diarrhea virus (BVDV) establishes persistent infection in the fetus of cattle. BVDV infected bovine fetuses develop immunocompetence towards

BVDV which helps the virus to evade the host immune response and maintain persistent infection [59].

1.6.1. EAV persistent infection

Following infection via the respiratory or venereal routs, EAV establishes persistent infection in the reproductive tract of stallions which results in the continuous shedding of the virus in semen [5; 6; 19]. Furthermore, this establishment of persistent infection is androgen dependent [60]. In some cases, virus shedding of these persistently infected carrier stallions may stop within a few weeks or months post infection. However, in 10-70% of infected stallions, virus shedding may persist for many years to lifelong [5;

6; 19; 27; 61]. Carrier stallions do not display any clinical signs of equine viral arteritis

(EVA) or a decrease in reproductive capacity [27; 62; 63]. Furthermore, studies have demonstrated that there is a correlation between the EAV carrier status of stallions and the susceptibility of a subpopulation of CD3+ T cells of those stallions to in-vitro EAV infection. Furthermore, the same studies demonstrated that this correlation has a strong association with the CXCL16 gene on equine chromosome 11 [64; 65].

Interestingly, the virus persists in the male reproductive tract despite the presence of very high neutralizing antibodies in the serum [27; 62; 63]. Studies have demonstrated that there is a negative correlation between the increase in serum neutralizing antibody titers and the host viremia titers suggesting neutralizing antibodies play a role in virus clearance, except in the case of carrier stallions [66]. Even though the mechanisms utilized by EAV to evade the host immune responses has not been fully identified, several previous studies have demonstrated the importance of host factors in the development EAV persistent infections in stallions. Recent work has demonstrated that establishment of long- term persistent infection correlates with the in vitro susceptibility of a subpopulation of

CD3+ T lymphocytes to EAV infection [64; 65; 67-69]. As such, stallions with the susceptible phenotype for CD3+ T lymphocytes are at a greater risk of becoming long-term persistently infected carriers compared to stallions with the resistant phenotype. It has been demonstrated through a genome-wide association study that these susceptible/resistance phenotypes are associated with the CXCL16 gene located in equine chromosome 11 [67].

Subsequent studies have further demonstrated that the CXCL16 gene consists of two allelic variants and the stallions with the susceptible phenotype (CXCL16S) establishment of long- term persistence whereas stallions with the resistant phenotype (CXCL16R) is associated with early viral clearance [70]. Interestingly, the CXCL16S isoform functions as an entry receptor for EAV while the CXCL16R isoform does not [69; 70]. Furthermore, Carossino et.al. (2018) has demonstrated the microRNA eca-mir-128 present within seminal exosomes, which is also a putative regulator of CXCL16, is downregulated in horses with persistent EAV infections. The downregulation of this microRNA promotes the upregulation of CXCL16 at anatomical sites where EAV establishes persistent infection

[71]. Taken together, CXCL16 and eca-mir-128 are important host factors that play significant roles in the complex mechanism utilized by EAV to establish persistent infection in stallions [72].

It has been demonstrated that the ampullae is the primary sites of EAV persistent infection [73; 74]. Within these glands, EAV shows a strong tropism towards vimentin- positive stromal cells of the lamina propria as well as T (CD2+, CD3+, CD5+, and CD8+) and CD21+ B lymphocytes [74]. Moreover, transcriptome analysis of the accessory sex gland ampullae from stallions with persistent EAV infections has demonstrated CD8+ T lymphocyte infiltrations suggesting a predominant CD8+ T lymphocyte response [73].

However, EAV infected cells usually localize within inflammatory infiltrates suggesting

EAV has the capacity to replicate regardless of this host anti-viral response [74].

Transcriptome analysis of the ampullae of persistently infected stallions further revealed the upregulation of inhibitory receptor molecules such as PDCD1 and transcriptional factors EOMES, NFATC2 and PRDM1 which maybe resulting in CD8+ T lymphocyte dysfunction which can hinder the virus clearance process by these lymphocytes [73].

Furthermore, T regulatory cells, which have been implicated in the suppression of local immune responses following virus infections, have also been detected among inflammatory infiltrates. These factors may contribute to the local immunosuppression in the ampullae and other accessory sex glands which may promote viral persistence [74].

Furthermore, only a small fraction of the CD8+ T lymphocytes infiltrates express granzyme

B or interferon gamma which indicates a local suppression of the CD8+ T cell mediated immune response at these accessory sex glands [74]. Additionally, studies have demonstrated that in persistently infected stallions EAV undergoes antigenic drift that may

contribute to immune evasion [74]. The importance of androgens in the establishment of

EAV persistent infection is evident by the fact that the only successful method of clearing the virus in persistently infected stallions is through castration [60; 74]. The local immunosuppression induced by androgens maybe another factor influencing the establishment of EAV persistence. As such all these factors point towards a multifaceted mechanism at play during the development of EAV persistence [74].

1.6.2. EHV-1 latent life cycle

Herpesviruses are among the most successful viral pathogens affecting both humans and animals and one key reason for their success is their ability to establish latent infections and periodically reactivate [75]. As mentioned before, the order Herpesvirales has 3 subfamilies and all these subfamilies include viruses that can establish latency. Following the initial lytic cycle of infection in horses, EHV-1 establishes latent infection in about

60% of naturally infected horses [29]. During this time the infected animals show no apparent clinical signs, virus shedding or viremia [1]. In HSV-1, latency is established in sensory neurons innervating the site of primary infection [76]. Many different studies has looked into elucidating the specific site of EHV-1 latency and they have reported the sensory nerves of the trigeminal ganglia and lymphocytes of the peripheral circulation as well as in lymph nodes as possible candidate sites [29-31]. Furthermore, reactivation of latently infected EHV-1 from leukocytes has been demonstrated but not from the trigeminal ganglia [29]. The reasons and the mechanisms that drive herpesviruses towards latency is not fully understood. However, the prevailing consensus regarding latency is that

(a) the failure to initiate the expression of viral IE genes promote latency; (b) The herpesvirus genome exists in a non-replicating “endless” state during latency; (c) only the

latency associated transcripts (LATs) are transcribed during latency; (d) cellular signals

that can activate the expression of viral IE genes induce herpesvirus reactivation [76]. With

these characteristics, the latent herpesvirus particles escape the immune system and the

host becomes a life-long carrier of the virus [1].

1.7. Equine arteritis virus

1.7.1. EAV classification

The equine arteritis virus (EAV) is a positive sense single stranded RNA virus in the order Nidovirales. The order Nidovirales is a relatively large group of positive sense

RNA viruses [77]. According to the previous classification, the order Nidovirales consists of three families: Coronaviridae, Arteriviridae, and Roniviridae. The family Arteriviridae contained the single genus Arterivirus to which EAV belonged [78]. However, according to the latest ICTV classifications, the order Nidovirales contains 7 sub orders, 9 families,

20 subfamilies, 33 genera, 59 sub genera and 88 virus species. Within this large collection of RNA viruses, EAV belongs to the suborder Arnidovirineae, family Arteriviridae, subfamily Equarterivirinae, and the genus Alphaarterivirus

(https://talk.ictvonline.org/taxonomy/). In contrast to the family Herpesviridae, virion morphology of the nidoviruses do not share significant similarities. Furthermore, they differ in their host specificity and genome size as well. The basis for classifying viruses under the order Nidovirales lies with the similarities in genome organization, mechanisms of gene expression, and in sharing a common ancestry for the key replicase gene [77-81].

1.7.2. EAV virion architecture

EAV is the prototype virus of the family Arteriviridae and the causative agent of equine viral arteritis (EVA) [5; 6; 19; 77]. Arteriviruses are enveloped, spherical virus particles containing glycoprotein projections on their envelope. The EAV virion has a spherical morphology with a 40 to 60 nm diameter [82]. The genome size and the number of encoded genes vary considerably among the members of the order Nidovirales [78; 80; 83].

Arteriviruses have relatively smaller genomes when compared to other members of the

order Nidovirales and the genome size of EAV varies between 12,704 and 12,731 kilobases

(kb) between different viral strains (kb) [5; 6; 19; 84].

Figure 1.2: Virion architecture of EAV consists of a nucleocapsid and 7 envelope proteins. Major envelope proteins (GP5 and M) forms dimers while 3 of the 5 minor envelope proteins (GP2, GP3 and GP4) forms trimers. E and ORF5a are also minor envelope proteins. Adapted and reproduced with permission from Elsevier: [Vet Clin North Am Equine Pract.][5], copyright 2014.

The EAV genome is encapsidated by an icosahedral core composed of the nucleocapsid protein and this core particle is wrapped around by a smooth lipid bilayer envelope lacking large projections. However, it contains 7 envelope proteins (Figure 1.2) [5; 6; 80; 82].

1.7.3. EAV genome organization

EAV has at least 10 open reading frames (ORFs) encoding for 13 non-structural proteins

(nsps) and 8 structural proteins (Figure 1.3) [5; 6; 80; 82]. The EAV genome, as it is with all the members of the family Arteriviridae has a capped structure at its 5' end and a polyadenylated (poly A) tail at the 3' end. In addition to that, the 5' and the 3' ends of the genome contains untranslated regions (UTRs).

The 5' proximal three-quarters of the genome is occupied ORF1a and ORF1b which are the two largest ORFs in the EAV genome (Figure 1.3) [5; 6; 82; 84]. ORF1a and ORF1b are connected to each other with a -1 ribosomal frameshift site (RFS) and they encode for two replicase polyproteins which gets extensively processed post-translation by 3 viral proteases (nsp1, 2 and 4) to produce at least 13 nsps (nsp1-12, including nsp7 α/β) [5; 6;

82; 84]. The 8 structural proteins of EAV are encoded by a nested set of segmented mRNAs located at the 3' proximal quarter of its genome and not from its genomic RNA (Figure 1.3)

[5; 6; 82; 84]. The 8 structural proteins of EAV include 7 envelope proteins (E, GP2, GP3,

GP4, ORF5a protein, GP5 and M) and the nucleocapsid protein (N) [5; 6; 82; 84]. Out of the 7 envelope proteins 3 (GP2, GP3 and GP4) are considered minor envelope proteins and

they form a heterotrimer while the major envelope proteins, M and GP5 form disulfide- linked heterodimers [5; 6].

1.7.4. EAV viral proteins

Compared to EHV-1, EAV is a relatively small virus with a smaller genome [3-6].

While the EHV-1 genome contains 76 genes coding for a total of 77 viral proteins, the 10

ORFs of EAV codes for a mere 21 viral proteins [1; 3; 5; 6; 28]. Proteins encoded by the

EAV genome is classified into two groups as structural and non-structural proteins. EAV structural proteins consists of the 7 envelope proteins E, GP2, GP3, GP4, ORF5a protein,

GP5, and M encoded by ORFs 2a, 2b, 3-4, 5a, 5b, and 6, and a nucleocapsid protein (N) encoded by ORF7. The 3' proximal quarter of the EAV genome contains the ORFs that code for the structural proteins (Figure 1.4). The minor envelope proteins GP2, GP3 and

GP4 form heterotrimers while the major envelope proteins GP5, and M form a disulfide- linked heterodimer in the EAV particle (Figure 1.3). The structural proteins of EAV takes part in the formation of the EAV virion [5; 6; 82; 84].

EAV non-structural proteins (nsps) are synthesized by the autoproteolysis of the large polyproteins pp1a and pp1ab which are the products of ORF1a and ORF1ab respectively [85]. ORF1a and ORF1b together is known as the replicase gene and it occupies 75% of the 5’ end of the EAV genome. Translation of replicase gene results in the production of pp1a (1728 aa) and pp1ab (3175 aa) which gets auto-catalytically cleaved at 11 sites by 3 internal proteinases (nsp1, nsp2, and nsp4) resulting in the production of the EAV non-structural proteins (nsps) 1-12 [86; 87]. Expression of pp1ab occurs through a -1 ribosomal frameshift mechanism in the overlapping region of ORF1a and ORF1b

(Figure 1.5) [80].

1.8. Equine viral arteritis

Equine viral arteritis (EVA) is a respiratory and reproductive disease of the family

Equidae caused by EAV [5; 6; 19; 27; 88-90]. It was first isolated in 1953 from a fetal lung following a respiratory disease outbreak which also accompanied abortions [91; 92]. Since then EAV outbreaks have been reported from countries [61; 93-95]. Furthermore, serological studies done using different horse populations around the world has demonstrated that EAV has a worldwide prevalence [61]. Horses in almost every country in the European continent is seropositive to EAV. However, the percentage of seroconverted horses in European countries vary between 6 and 90% [96-100].

Furthermore, the seropositivity for EAV among different breeds of horses such as the

European Haflingers, Norikers, Warmbloods, and Lipizzans and the American

Standardbreds were between 50 and 94% [95; 99]. The seropositivity for EAV among

American Quarter horses and thoroughbreds were around 0.6% and <5,4% respectively

[61; 95; 101]. This indicates the seroprevalence for EAV varies among different countries and breeds [19]. Only a hand full of countries such as Iceland, Japan, and New Zealand are free from the disease [27; 102]. Interestingly, the seroprevalence for EAV increases with the age of the horse suggesting a repeated exposer to EAV as the horses age [61; 100].

Recent studies have demonstrated that the difference of seroprevalence among horse breeds have a genetic origin. Among stallions, the trait of becoming long term carriers or clearing

EAV shortly after infection is linked to the in-vitro susceptibility of a subset of CD3+ T lymphocytes to EAV infection. As such, stallions with this subset of susceptible CD3+ T lymphocytes are at a higher risk of becoming long term carriers [67; 68; 103].

Transmission of EAV occurs via two main routs which are the venereal and respiratory routs [61; 92]. During the acute phase of infection, horses shed the virus in nasal secretions

7-14 days post infection [104]. Aerosolization of virus particles in respiratory secretions facilitate horizontal transmission of the virus from infected to non-infected horses [19].

Aerosolized virus particles originating from urine and other bodily secretions, as well as from aborted fetuses and fetal membranes from infected horses can also transmit the virus via the respiratory rout [98; 99; 105]. In addition, the virus can be present in the feces of acutely infected animals up to 8 days post infection [106] as well as in the female reproductive tract for a short time following infection [105]. Transmission of EAV via the venereal rout occurs with semen from acutely or chronically infected stallions during natural or artificial breeding [63]. Virus shedding via the venereal rout can occur during the acute phase of the infection or when an acutely infected stallion becomes a carrier animal. It is estimated that around 10-70% of the acutely infected stallions become carriers that continuously shed the virus in their semen ranging from several weeks (short term carriers) to life long (long term carriers) [61; 63]. The establishment of persistent infection in stallions by EAV is similar to the establishment of latent infections by EHV-1 in that it allows the virus to establish a natural reservoir that is responsible for the survival of the virus as well as its perpetuation and evolution [61; 62; 107]

1.8.1. Clinical signs

Following EAV infection via the respiratory rout, the initial virus replication takes place in alveolar macrophages and bronchiolar epithelial cells within 24 hours post- infection (hpi) [108]. Within the first 48 after infection the virus drains into regional lymph nodes, especially the bronchial lymph nodes [109]. This is followed with virus shedding in

nasal secretions and typically lasts up to 7-21 days [109]. Once inside the lymph nodes the virus infects Peripheral blood mononuclear cells (PBMCs) and initiates cell associated viremia (CAV) within 72 hpi which may last up to 3-19 days post infection. Development of CAV is essential for the pathogenesis of the virus since it allows the virus to spread throughout the body and replicates in 2ry replication sites such as the vascular smooth muscle cells and endothelial cells, and cause systemic panvasculitis [6; 88; 110].

Furthermore, the characteristic histological lesion associated with EVA is severe necrotizing panvasculitis of small vessel. Therefore, abortions, which is one of the main reproductive consequences of EAV infections is a result of the vasculitis and thrombosis induced by EAV in the myometrial blood vessels of the pregnant uterus [88].

The severity of the clinical signs displayed during the acute phase of EAV infection varies according to the age, genetics and the physical condition of the horses as well as the viral strain, challenge dose and the route of infection [111; 112]. The horse adapted, highly virulent Bucyrus strain of EAV causes high mortality in healthy mature horse. However, it is a laboratory derived experimental virus variant with an extreme form of pathology that is not seen in EAV field isolates. The field isolates of EAV can be categorized into three groups according to the severity of clinical signs and the following disease manifestations.

As such, strains such as EAV KY84 and EAV KY77 cause a severe and a moderate disease of EVA respectively. Strains such as EAV SWZ64 and EAV AUT68 cause a mild form of

EVA and EAV KY63, and EAV CA95 infections are asymptomatic [113-115].

1.8.2. Acute infection

Most acute EAV infections are subclinical or produce barely detectible symptoms [98].

Usually it is the very young, old and the stressed horses that develop a clinically apparent

illness following acute infection with EAV [98]. Clinical signs usually develop 3 to 14 days following exposure to virus and may last up to 5 to 9 days [98]. Acutely infected horses initially show a flu-like illness followed with fever up to 41°C, depression, anorexia limb and ventral edema, periorbital edema, conjunctivitis, lacrimation, nasal discharge, and urticaria [27; 92]. Onset of fever is accompanied with leucopenia involving both neutrophils and lymphocytes [116]. In fact fever and leukopenia are considered the two most consistent clinical signs [92; 117]. Severe clinical signs are usually seen with foals and neonates and these include respiratory distress, colic, and diarrhea and may result in mortality [112; 118]. However, other than with foals and neonates, infections with EAV field isolates are rarely fatal to adult horses [61; 114]. As mentioned earlier, the virus can establish persistent infections in the reproductive tract of the stallion but in geldings and mares, the virus gets cleared within about a month after infection [61].

1.8.3. Reproductive consequences

Abortions in pregnant mares is one of the main reproductive consequences of EAV infections. A pregnant mare may abort starting from 3 months to over 10 months into gestation either with or without displaying any clinical signs. Furthermore, EAV induced abortions may occur at the end of the acute phase or during the convalescent phase of infection [61; 119; 120]. The incidence of abortions during natural EAV outbreaks range from <10% to 71% and this could be due to the differences in the abortigenic potential of different EAV strains [19; 61; 91; 121]. The aborted fetuses, in most cases are partially autolyzed but may be expelled fresh as well [61; 120; 122]. EAV infections in the late gestation period may result in a premature delivery of a weak foal that may not survive or a foal that develops the clinical disease within a short period of time [112].

EAV infections in stallions may result in a period of subfertility during the acute phase of the infection ranging from 6 to 11 weeks after infection. This sub-fertile period is usually associated with decreased sperm motility, concentration and percentage of morphologically normal sperm cells as well as a decrease in libido [123]. Furthermore, 10–

70% of EAV infected stallions and sexually mature colts become carriers of the virus by establishing persistent infection and continuously shed the virus in their semen ranging from several weeks (short term carriers) to months, years or lifelong (long term carriers) that shows no changes in their semen quality [19; 61; 63; 124]. There is evidence to suggest that the carrier state is androgen-dependent and this explains the lack of it in mares geldings or prepubertal colts [61; 125]. Virus shedding in semen usually starts around 5 days post infection and the sperm-rich fraction of semen contains the virus [63; 123].

1.8.4. EAV vaccination and management

Several studies have investigated the use of antiviral compounds against EAV infections but none of them have been proven to be successful in treating acute EVA [126;

127]. As such vaccination and proper management applications are one of the best methods to prevent and control this disease. Currently, in the US and Canada, a modified live vaccine (MLV) is being used against EAV (ARVAC®) [62]. However, this MLV vaccine is not licensed in European countries and Japan. An inactivated EAV vaccine (Artervac®) is used in several European countries instead of the MLV vaccine. Compared to the MLV vaccine, Artervac® has a reduced immunogenicity and needs to be re-vaccinated semi- annually. Furthermore, its efficacy in preventing clinical EVA is not well characterized

[19; 98].

The intra-muscular MLV vaccine has been in use in the US since 1985 and has been proven to be safe as well as induce protective immunity against EVA [105; 128; 129].

Studies have shown that the vaccine elicits the production of neutralizing antibodies 5-8 days post vaccination and this antibody titer peaks 7-14 days post-vaccination [105; 128-

130]. The protective antibody response may last up to 2 years and annual revaccination is recommended to maintain the protective level of neutralizing antibodies titers which is estimated to be 1:64 [19; 131; 132]. Even though vaccination with the MLV can prevent clinical EVA as well as prevent virus shedding in semen as well as persistent infection, it does not prevent re-infection and therefore, it is possible for vaccinated horses to display viremia or shed the virus is nasal secretions if they get exposed to and re-infected with

EAV post vaccination [61; 105; 130].

It is not recommended to vaccinate pregnant mares with the MLV vaccine especially during the last 2 months of gestation since there is a high chance for abortions [133].

Furthermore, it is not recommended to vaccinate dams with foals <6 weeks of age. Foals should be vaccinated only when they are 6 months of age or older. This is to prevent the neutralization of the MLV vaccine by maternal antibodies against EAV. Moreover, vaccination of foals before the onset of puberty is highly recommended to prevent them from becoming carriers of the virus [19; 98].

Management practices to control and prevent EVA are aimed at preventing the spread of EAV through Identification of carrier stallions, the quarantine of new arrivals, the segregation of pregnant mares and foals from other horses, implementation of biosecurity measures, and enforcement of vaccination programs [19; 101]. It is recommended to assess a stallion’s vaccination history, serological status and the virological status of their semen

prior to breeding or introduction into a new farm in order to prevent new EAV outbreaks.

Furthermore, 60 days prior to breeding, stallions should be assessed for the presence of neutralizing antibodies against EAV. Virological assessment of semen should be performed in unvaccinated yet seropositive stallions in order to determine their infection status and potential carrier state. If a stallion is confirmed as a carrier it should be kept isolated from other non-infected stallions, however, it can be bred with a seropositive mare

[105]. Furthermore, it is highly recommended to vaccinate horses that frequently travel to competitions or come into close contact with other horses on a regular basis.

During EAV outbreaks, the state veterinarian should be immediately notified, and the premises placed under strict quarantine. Horses with symptoms or in close contact with affected horses should be isolated and all animal movements should be suspended

(http://www.aaep.org/info/guidelines). Furthermore, breeding programs at the affected premises should be put on hold and at-risk horses should be vaccinated immediately.

Following quarantine, it is important to decontaminate stalls and equipment on the affected premises with any disinfectant. Usually disinfectants containing phenolic, chlorine, iodine, and quaternary ammonium compounds are adequate to inactivate EAV [5; 19].

1.9. Host immune response against EAV

1.9.1. Innate immune response

As mentioned earlier, EAV infects horses in two main ways, either via the respiratory rout or the venereal rout [61; 92]. Therefore, during the infection process, the

EAV particles first encounter the mucosal lining of the upper respiratory tract or the reproductive tract and the host mucosal immune response acts as the first line of defense against EAV. The host innate immune response against EAV has not been extensively

studied, however, the available information provide insights in to the interactions between the virus and the innate immune system [19].

EAV is an endotheliotropic virus and during its pathogenesis it mainly infects equine endothelial cells (EECs) and PBMCs [6; 88; 110]. Infection of these cells induce the production of various anti-viral compounds as well as proinflammatory cytokines and chemokines. One of the major anti-viral compounds produced by the triggering of the innate immune system are the type I (IFNα and IFNβ) interferons [134-136]. Binding of

IFN molecules with their cell surface receptors triggers a cascade of signaling events through the Janus kinase signal transducer and activator of transcription (JAK-STAT) pathway which induces the transcription of hundreds of IFN regulated genes (IRGs) [137].

Investigations into the host interferon response induced by EAV has shown that in EECs,

EAV suppresses the production of IFNβ which is one arm of the type I interferon system

[138]. The same study demonstrated that nsp1, nsp2, and nsp11 of EAV exerts strong inhibition on the IFNβ promoter and that nsp1 interferes with the host IRF-3 and signaling via NF-휅B to inhibit IFNβ production [138]. Similar to EAV, PRRSV which is another member of the order Nidovirales is also known to elicit an attenuated type I interferon response. Upregulation of TNF-α, IL-1β, IL-6, and IL-8 mRNA in EECs have been demonstrated following in-vitro infection of virulent and avirulent strains of EAV.

Furthermore, the secretion of biologically active TNF-α has also been detected in EAV infected EECs [139]. Studies have demonstrated that the virulent Bucyrus strain (VBS) of

EAV could in-vitro infect CD3+ T lymphocytes of only some horses [103]. In-vivo infection of horses belonging to both the susceptible and resistant phenotypes have demonstrated that horses in both groups upregulated the mRNA production of IL-1β, TNF-

α, IL-8 and IFNɣ following infection with EAV. However, IFNα and IL-10 gene expression was observed only in horses with the resistant phenotype [64].

Following the respiratory rout of infection, EAV infects alveolar macrophages

(AMΦ) but in-vitro studies have shown that EAV is unable to replicate in them. However,

EAV is capable of both infecting and replicating in blood-derived equine macrophages

(BMΦ) [139]. In-vitro infection of both AMΦ and BMΦ by virulent and avirulent strains of EAV stimulate the mRNA production from TNF-α, IL-1β, IL-6, and IL-8 genes as well as the production of biologically active TNF-α secretion [139].

1.9.2. Humoral immune response

The host humoral immune response against EAV is characterized by the production of complement-fixing (CF) and virus-specific neutralizing (VN) antibodies [129; 140].

Several studies have demonstrated that the EAV structural proteins GP5, M and N as well as the nsps nsp2, nsp4, nsp5 and nsp12 contribute to induce the humoral response in the host [141; 142]. However, several studies have demonstrated that the major neutralization sites for EAV-specific antibodies are located on the GP5 protein and these include 4 major neutralization sites [143; 144]. Similarly, Chirnside et. al. (1995) identified another epitope on the GP5 protein [145]. Moreover, another study done by Glaser et.al. (1995) demonstrated that mutations on the GP5 neutralization sites render EAV resistant to neutralization with VN antibodies [146].

Both CF and VN antibodies appear between 7-14 days post infection. CF antibody titers peaks around 2-3 weeks and declines around 8 months post infection while VN antibody titers peak around 2-4 months post infection and may persist for 3 years or more

[104; 134; 140; 147]. Avirulent strains of EAV such as the MLV strain is shown to induce a stronger CF antibody response in the host than virulent EAV strains [140]. The induction of CF antibodies is important because EAV neutralization is thought to be complement dependent and the heat inactivation of host sera decreases its capacity to inactivate EAV.

Furthermore, the IgG fraction of late antiserum is responsible for EAV neutralization [148;

149]. A recent study done by Carossino et.al. (2018) has demonstrated that following EAV infection a short-lived IgM response is initially triggered followed by a long-lasting IgG1 response in infected horses. The appearance of this humoral response correlates with the reduction of viremia and subsequent viral clearance except in the case of carrier stallions

[66; 74]. Furthermore, EAV infections also triggers an IgG1 and IgG4/7 antibody response in the seminal plasma [74].

Seropositive dams transfer their EAV-specific VN antibodies to foals via colostrum and they protect the foal from EAV infections during the first few months of their life. The fact that maternal antibodies are not detected in the serum of foals prior to nursing suggests that the transfer of the EAV-specific passive immunity is through colostrum and not via the placenta in-utero [150; 151]. As mentioned earlier, the maternal antibodies against

EAV remains in the sera of foals up to 6 months and foals should be vaccinated during the time of weaning to maintain this EAV-specific immunity [19; 98; 129]. The half-life of

EAV-specific maternal antibodies in serum of foals is 32 days [150; 151].

1.9.3. Cell mediated immune response

Cell mediated immunity, similar to the humoral immunity plays a key role during infections. A study done by Castillo-Olivares et al has demonstrated the presence of an

EAV-specific cytotoxic T-lymphocyte (CTL) responses in experimentally infected ponies.

The study shows that the cytotoxicity displayed by EAV stimulated PBMCs was virus specific and was mediated by CD8+ T lymphocytes. Furthermore, this CTL response lasted for period between 4 months to more than 1-year post-infection [152].

1.10 Equine herpesvirus-1 (EHV-1)

1.10.1 EHV-1 classification

With the isolation of more than 80 different herpesviruses from different animal species during the past 50 years necessitated a scientific classification for the Herpesviridae family.

The first attempt of organized scientific classification of the herpesviruses came in 1971 by ICTV [153]. However, this classification was updated in 2009 to form the currently used ICTV classification of herpesviruses [28]. During the 2009 update, the former family

Herpesviridae was split into 3 families and they were incorporated into the new order

Herpesvirales [28]. The three new families created were the family Herpesviridae which contains the mammal, bird and reptile herpesviruses, family Alloherpesviridae which contains the fish and frog herpesviruses and the family Malacoherpesviridae which contains a bivalve herpesvirus [28]. The family Herpesviridae is divided into 3 sub families; namely Alphaherpesvirinae, Betaherpesvirinae and Gammaherpesvirinae and the two equine herpesviruses used in this study, EHV-1 and the equine herpesvirus-4 (EHV-

4) are included in the subfamily Alphaherpesvirinae [28].

1.10.2 EHV-1 virion architecture

Members of the family Herpesviridae share similar virion structural features. The virion of the members of this family has a spherical morphology and it contains four major components which are the capsid, the core, the tegument and the envelope [28; 153; 154].

The core contains the genetic material of the virus which is a double stranded deoxyribonucleic acid (DNA) molecule of 125-290 kbp arranged into unique long and short regions with internal and terminal repeats. It is a fibrillar spool in which the viral

DNA is wrapped around [17; 18; 28; 154]. Encasing the core is an icosahedral capsid which is composed of 12 pentameric and 150 hexameric capsomeres. The external diameter of this 162 capsomere capsid is 200 nm. The capsid of the Herpesviridae family members is surrounded by an asymmetrically arranged proteinaceous matrix containing over 30 viral proteins called the tegument [28; 153-155]. Surrounding this entire structure is an envelope or a lipid bilayer which contains viral and some host glycoproteins embedded in it (Figure

1.4) [28; 153-155].

Figure 1.4: Virion architecture of EHV-1 consists of a core containing viral genomic DNA, a capsid, tegument and an envelope with 12 embedded glycoproteins. Adapted and reproduced with permission from Bentham Open: [The Open Veterinary Science Journal][1], copyright 2008.

1.10.3 EHV-1 genome organization

EHV-1 together with its closely related cousin EHV-4 are two of the most important herpesviruses affecting horse populations worldwide and up until 1981, both EHV-1 and

EHV-4 were considered two subtypes of the same virus [156]. However, the complete genome sequencing of EHV-1 and EHV-4 paved the way in separating them as two different virus species [3; 4]. Just as all the other herpesviruses, EHV-1 has a linear dsDNA genome encapsidated by a thick walled, spherical to pleomorphic icosahedral capsid of about 120 nm in diameter and 162 capsomeres [28; 154; 155]. The genome of EHV-1 is approximately 150,224 bp in size [3; 4] and can be divided into a unique long (UL) region

(112,871 bp) and a unique short (US) region (11,861 bp) which are covalently linked to each other. The UL region is flanked by two inverted repeat sequences of 32 bp in EHV-1 called the internal repeat (IRL) and the terminal repeat (TRL) [3; 4; 17; 28].

Figure 1.5: Genome organization of EHV-1. The UL region is flanked by TRL and IRL sequences. The US region is flanked by TRS and IRS sequences. The UL region contains

ORFs 1-63 and the US region contains ORFs 68-76. The TRS and IRS regions contain copies of ORFs 64-67 [3; 4].

Similarly, the US region is also flanked by two inverted repeat sequences of 12,714 bp called internal repeat (IRS) and the terminal repeat (TRS) (Figure 1.5) [3; 4]. EHV-1 genome consists of 80 ORFs and they code for at least 76 individual genes of which four

(genes 64, 65, 66 and 67) have been duplicated [3; 4; 17]. The average G+C content of the

EHV-1 genome is 56.7% [3; 4]. Furthermore, similar to other herpesviruses, the genome of EHV-1 contains 20 sets of tandemly reiterated DNA sequences in which 8 are located in the UL region, 2 in the US region and 5 each in the TRS and IRS regions. Interestingly, most of these sequences are in non-coding regions of the genome. Even though, EHV-1 and EHV-4 has some very close similarities in genome arrangement and composition they share with their alphaherpesvirus cousins, EHV-1 is assumed to contain five distinct genes encoded by ORF 1, ORF 2, ORF 67, ORF 71 and ORF 75 which have not been found in any other herpesviruses to date.

1.10.4. EHV-1 viral proteins

As mentioned previously, the components of an EHV-1 virion includes a nucleocapsid core, an icosahedral capsid, the tegument and a lipid envelope [28; 154].

Several EHV-1 viral proteins contribute to the formation of the nucleocapsid core and among them are the gene products of EHV-1 genes 22, 25, 35, 42, 43 and 56. These genes encode for “capsid triplex subunit 1”, “capsid protein”, “capsid maturation protease”,

“major capsid protein”, “capsid triplex subunit 2” and the “capsid portal protein” respectively. The function of the capsid portal protein is the encapsidation of viral DNA while the major contribution of the other nucleocapsid core proteins is in capsid morphogenesis.

The gene products of the genes 8, 11, 12, 23, 40, 46, 48 and 51 which are the

“tegument protein UL51”, “tegument protein VP22”, “transactivating tegument protein

VP16”, “tegument protein UL37”, “tegument protein UL21”, “tegument protein UL16”,

“tegument protein UL14” and the “myristylated tegument protein” respectively, contribute in the virion morphogenesis of EHV-1. The gene products of the EHV-1 genes 13 and 14

which are the “tegument protein VP13/14” and “tegument protein VP11/12” take part in gene regulation by modulating the gene expression of VP16. Gene products of the genes

15, 19, 45, and 49 which are the “membrane protein UL45”, “Tegument host shut-off protein”, “DNA packaging tegument protein UL17” and the “Tegument serine/threonine protein kinase” takes part in the membrane fusion of the virion, host mRNA degradation,

DNA encapsidation and protein phosphorylation respectively.

The EHV-1 virion is surrounded by a lipid envelope with 12 glycoprotein projections. The 12 glycoproteins of EHV-1 are conserved in all other alpha herpesviruses as well and therefore, named according to the nomenclature adopted for the prototype member of this family which is HSV-1 [1]. These envelope glycoproteins play a major role in inducing an immune response from the host following infection as well as in virus adsorption, penetration, and cell-to-cell spread [1; 157]. The 12 EHV-1 glycoproteins are encoded by the genes 6, 10, 16, 33, 39, 52, 62, 70, 71, 72, 73 and 74 and termed gK, gN, gC, gB, gH, gM, gL, gG, gp2, gD, gI and gE respectively [1]. EHV-1 gC is essential for host cell binding of the virus as it assists in binding heparan sulphate moieties on the host cell surface as well as virus egression from host cells [1]. However, most of the EHV-1 glycoproteins play roles in cell to cell spread and these include gB, gD, gE, gI, gK and gM

[1]. The glycoproteins gD and gM is also important in entry into the host cell [1]. Apart from these functions, gC, gE, gG, gI, gM and gp2 are important in the virulence of EHV-

1 while gB, gD, gH, gL and gK play key roles in EHV-1 replication [158]. The glycoproteins gp2, gC, gB, gD and gM represent the most glycosylated proteins in the

EHV-1 virion and they function as immunogens inducing a humoral as well as a cell mediated immune response from the host [1; 157]. The fact that sera obtained from rabbits

and mice injected with purified EHV-1 virions contain polyclonal antibodies directed at this set of glycoproteins demonstrate their importance in stimulating a host immune response [157].

1.11. EHV-1 infection in horses and diseases

1.11.1. EHV-1 respiratory disease

EHV-1 infections are one of the major causes of acute viral upper respiratory tract

(URT) diseases in horses. Furthermore, it has been suggested that the viruses isolated from respiratory infections differ in antigenicity, host range, plaque size and growth rate in-vitro and in-vivo from the abortigenic isolates [159]. In the absence of mucosal antibodies, 1ry infection of EHV-1 takes place at the upper respiratory tract epithelium in horses, though, transmission is also possible through the conjunctival epithelium upon direct contact with aerosols containing the virus [160; 161]. The incubation period of the respiratory form of

EHV-1 infections vary depending on the strain of virus, infecting dosage and immune condition of the host, but in most cases its between 1-3 days [162]. Experimental intra- nasal infections done on ponies have revealed that infectious virus particles and virus antigen expression can be detected in the nasal and nasopharyngeal epithelium of infected animals as early as 12 hpi. The URT infections by EHV-1 results in the erosion of the nasal and nasopharyngeal epithelial cell around 4-7 days post infection and during this time the infected animals also shed the virus from epithelial surfaces. The erosion of epithelial cells expose the underlying lamina propria to which the virus readily spreads and infects the leukocytes and endothelial cells of the blood and lymphatic vessels around 4 days post infection [163]. Once inside the circulatory system, the virus amplifies itself by going through the lytic cycle of replication which results in cell-associated viremia and the spread

of the virus to its secondary sites of replication such as the pregnant uterus, CNS, testes and endocrine organs [15; 17; 18; 163-165]. The cell-associated viremia is a critical development in EHV-1 induced abortions and EHM and can develops as early as 3 days post infection and last up to 22 days [163].

The respiratory form of EHV-1 infections usually results in rhinopharyngitis and tracheobronchitis [17; 18; 166]. In young horses, the respiratory form of EHV-1 usually results in depression, fever, anorexia, coughing, and nasal discharge. However, neutropenia, lymphopenia, and progressive lymphadenopathy may also be visible [162].

The nasal discharge appears serous for the first few days of infection and may become mucopurulent with development of 2ry bacterial infections. The infected horses may develop bronchopneumonia when the virus infects the lower part of the respiratory tract

[1]. Virus clearance is takes about 1-3 weeks, however, bronchial hypersensitivity, chronic obstructive pulmonary disease syndrome or compromised athletic performance may still persist and this condition is called “poor performance syndrome” [162; 167].

1.11.2. Abortion and neonatal diseases

With highly virulent EHV-1 strains, endothelial cell infection, vasculitis and thrombosis in the tale pregnant uterus is critical for abortions to occur [18; 168]. Following experimental intra-nasal infection, endothelial infection of the blood vessels in the pregnant uterus has been detected as early as 6 days post infection and it gets widespread over 9-13 days post infection, and as mentioned before, the development of cell-associated viremia is critical for this [163; 168]. The interaction between viremic leukocytes and endothelial cells appears to be tissue selective suggesting the micro-environmental factors such as cytokines and hormones may influence endothelial cell activation and the subsequent

interaction with leukocytes to facilitate virus transfer [1; 18; 163; 168]. The principal determinant of abortion is the degree of thrombosis in the pregnant uterus. Endothelial cell infection in blood vessels of the late pregnant uterus leads to inflammation and thrombosis.

Severe and wide-spread thrombosis results in rapid placental thrombo-ischemic necrosis leading to the expulsion of a fresh virus negative fetus. Less severe and less wide-spread thrombosis do not cause rapid placental separation providing more time for the virus to leak into and infect the fetus via the placenta [163; 169; 170]. In some instances, the fetuses may be born alive after virus-induced expulsion but develops interstitial pneumonia shortly after birth [17; 18]. Early pregnancy abortions are rare with EHV-1 infections. several possible reasons maybe at play making the late gravis uterus more susceptible for abortions by EHV-1. One such reason could be that during mid to late pregnancy, the placenta releases progesterone which causes local immunosuppression. Furthermore, the development of individually vascularized micro cotyledons in the placenta makes these micro cotyledons susceptible to single vessel thrombi infarction [17; 18; 163; 168; 171].

1.11.3. Equine herpesvirus myeloencephalopathy (EHM)

EHV-1 infections usually result in mild respiratory diseases in young horses, neonatal foal death and abortion storms in pregnant mares and establish lifelong latent infection in horses which allows viral perpetuation in equine populations [18; 172]. Interestingly, some of the EHV-1 strains can cause a fatal neurological disease called equine herpes myeloencephalopathy (EHM) [15; 18]. The association between EHV-1 and a severe form of myeloencephalopathy was first discovered in Norway in 1966 [173]. EHM has a 15-

50% case-fatality rate and poses a significant threat to the $102 billion equine industry in the United States. Unfortunately, the occurrence of EHM has increased dramatically over

the past fifteen years, with extensive outbreaks reported in the United States, South Africa,

New Zealand and several European nations, including the United Kingdom [14; 52; 164;

165; 174; 175]. Similar to EHV-1, other alpha herpesviruses such as HSV-1 and VZV too induces a neurological disease form. However, unlike HSV-1 or VZV, EHV-1 does not have a neurotropism [176; 177]. EHM is a result of vasculitis and thrombosis caused by

EHV-1 following infection of the endothelial cells of the vasculature of the CNS [15; 52;

164; 165; 175; 178]. (8). It has been demonstrated that following experimental EHV-1 infection in foals, the initial viral replication takes place in the epithelial cells and local lymph nodes of the upper respiratory tract [18; 166]. which result in leukocyte-associated viremia [18; 166; 179]. Subsequently, the virus infects the endothelium of the vasculature specifically located in the CNS and the uterus resulting in the development of an inflammatory process that leads to vascular damage (vasculitis and micro thrombosis) promoting the development of EHM and abortions [15; 18; 166].

The ability to establish EHM by EHV-1 strains is linked to a single nucleotide polymorphism (SNP) in the DNA polymerase gene (open reading frame 30 [ORF 30]).

This relationship was established by analyzing a panel of 131 EHV-1 field isolates [51].

As such, EHV-1 strains that has a Guanine (G) residue at the nucleotide position 2254 in the open reading frame 30 (ORF 30) are considered neuropathogenic while strains that has an Adenine (A) residue at this position are considered non-neuropathogenic [51]. This SNP results in the replacement of Asparagine (N) with Aspartic acid (D) at the amino acid position 752 in the DNA polymerase gene [51]. As such, it highlights a genetic basis associated with the virus in establishing EHM. However, this SNP of the pathogen’s genome cannot explain all the EHM cases resulting from EHV-1 infections since 16-24%

of the EHM cases are caused by EHV-1 strains that do not have this mutation on the DNA pol gene [175]. Therefore, it can be hypothesized that the development of EHM is multifactorial.

Clinically, horses with EHM display ataxia, paresis, swelling of hind limbs and urinary incontinence [52; 165; 180] while histopathological examinations often reveal perivascular cuffing in some areas of the central nervous system (CNS) together with axonal swelling and/or malacia, areas of microscopic hemorrhage, vasculitis and thrombosis [52; 178; 180].

EHM has no specific treatment and the prognosis is directly related to the severity of the neurological signs exhibited by affected horses [165]. Supportive medical care provided to such horses focus on treatments to reduce CNS inflammation due to vasculitis.

Corticosteroids such as prednisolone and dexamethasone, NSAIDS such as Flunixin megalumine and Dimethyl sulfoxide is often used as treatments to counter CNS inflammation together with Acyclovir [165]. Since 1941 many different vaccines have been developed to protect horses from EHV-1 infections as well as disease manifestations

[172]. However, these vaccines have not been successful since their protective efficacy is largely unknown or unsatisfactory [17]. Currently there are inactivated as well as modified live vaccines (MLV) available against EHV-1 but none of them seem to be able to prevent the development of EHM.

1.11.4. Vaccination and management

Vaccination, together with management measures remains as one of the best measures to control EHV-1 infections [1]. Vaccines against EHV-1 infections were first introduced in the 1950s and since then the main types of vaccines commercially available against

EHV-1 infections have been either inactivated whole virus or subunit vaccines [1]. The

earliest attempt at producing a vaccine against EHV-1 resulted in the development of a killed EHV-1 vaccine prepared using infected tissues from foals or hamsters. This was followed with the development of a partially purified, formaldehyde inactivated EHV-1 vaccine [181-183]. Likewise, inactivated whole EHV-1 vaccines are widely used during current times as well. Duvaxyn EHV-1/4 is an inactivated whole EHV-1 and -4 vaccine currently being used. This vaccine has been shown to induce both virus neutralizing as well as complement fixing antibodies in foals and mares [1]. Vaccination with Duvaxyn EHV-

1/4 significantly reduced the clinical signs of disease and the duration of both cell- associated viremia and virus shedding in foals and virus shedding was also reduced in vaccinated mares when compared with control ponies [1] upon experimental infection with

EHV-1. However, Response of foals and mares to inactivated EHV-1 vaccines in the field conditions have not been very efficient. A study carried out in Australia has demonstrated that the seroconversion of foals and mares vaccinated with inactivated whole EHV-1 vaccines are 50% and less than 30% respectively [184].

Apart from inactivated whole EHV-1 vaccines, the only other type of vaccine commercially available in the market are modified live vaccines (MLV) [17]. The vaccine

Rhinomune is an MLV licensed for the use in US and Europe (as Prevaccinol in Europe)

[1; 17]. Rhinomune has been demonstrated to have an excellent safety record and studies have demonstrated that this vaccine can provide protection against EHM under experimental conditions and against abortions in field conditions [185; 186]. Foals are usually protected against EHV-1 infections until 5-6 months of age due to the transfer of maternal antibodies. Therefore, during the time of weaning, it is recommended for foals to be vaccinated against EHV-1 followed with booster doses at every 3-6 months to prevent

the respiratory form of disease [187]. It is also recommended to vaccinate pregnant mares to reduce the risk of EHV-1 induced abortions [187].

Good managemental practices coupled with proper vaccination has shown to reduce the risk of EHV-1 induced abortions by about 75% [15]. One important aspect of proper management is to segregate the horse population in a farm to distinct categories such as weanlings, yearlings and new arrivals [187]. Horses arriving newly should be kept in quarantine or isolation for a period of 21 days before introducing to the herd. Since stress can reactivate the latently infected EHV-1 as well as immunosuppress animals making them more susceptible to various forms of pathogenic infections including EHV-1, it is important to control factors that cause stress in horses. Over-crowding, poor nutritional conditions, heavy parasite infestations, disruption of established social groups or exposure to inclement weather are some of the controllable factors that can induce stress in horses

[187]. Proper disinfection practices on tools and utensils as well as proper hygienic practices must also be followed to prevent EHV-1 infections and the spread of EHV-1 infection during outbreaks [188].

1.12. Host immune response against EHV-1

1.12.1. Innate immune response

EHV-1 initially infects the respiratory mucosal surface and the epithelium of the upper respiratory tract and as such this is the initial site of immune resistance against the virus [189] [190]. A recent study by Hussey et.al 2014 using primary equine respiratory epithelial cells has investigated the innate immune response against EHV-1 infections

[190]. The study revealed that upon infection the gene expression of the proinflammatory

cytokines and chemokines IFNα, Ilβ, TNFα, IL8 and IL6 was upregulated whereas the gene expressions of IFNβ, GM-CSF, MCP-1 and IL10 did not show a significant upregulation

[190]. Furthermore, the same study demonstrated that in-vitro EHV-1 infected equine respiratory epithelial cells suppress the secretion of IFNɣ. Even though the gene expressions of IFNα and TNFα was upregulated following EHV-1 infections, the secretion of these cytokines from equine respiratory epithelial cells was not detected. Furthermore, the secretion of IL-1, IL-17, IL-4, and IL-10 was also not detected [191]. However, a study done by Edington et al. 1986 has demonstrated an increased production of IFNα in nasal secretions as well as in serum following in-vivo EHV-1 infection of Welsh mares and ponies [192].

Interestingly, Hussey et.al 2014 revealed that EHV-1 infections of equine respiratory epithelial cells suppress MHC-1 and MHC-II expression. MHC-I and MHC-II molecules are antigen presenting cells and they are essential molecules in bridging the innate and the adaptive immune responses [193]. In the case of EHV-1 infections, the downregulation of MHC-1 molecules are of importance since they are responsible for the activation of CD8+ cytotoxic T-lymphocytes (CTLs) which plays a key role in EHV-1 clearance [194; 195]. As such, the downregulation of MHC-1 may play as a survival tactic by the virus to delay an EHV-1-specific immune response. Furthermore, the downregulation of MCH-I molecules by EHV-1 had been reported in several previous studies which identified it as an entry receptor for EHV-1 [196-198]. There is also enough evidence to conclude the secretions of equine respiratory epithelial cells which includes the proinflammatory cytokines and chemokines induces the chemotaxis of monocytes towards the site of infection and the EHV-1 pUL56 plays a role in suppressing this

chemotaxis[190; 199]. However, Edington et al. 1986 has shown that the phagocytic function and the killing efficiency of monocytes in EHV-1 infected mares and yearlings was reduced [192]. Furthermore, PBMCs in-vitro infected with EHV-1 has been shown to upregulate the gene expression of type I interferons (IFNα and IFNβ), TNFα, and IL8 while

TGFβ expression was downregulated.

1.12.2. Humoral immune response

As mentioned earlier, EHV-1 establishes latent infections in about 60% of infected horses [29]. Primary infection with EHV-1 is known to induce a protective yet short lived immune response, making hosts susceptible for reinfection within 3-4 months of the initial infection [200]. However, horses suffering multiple reinfection incidences gradually develop an immunity capable of preventing clinical signs of re-infection [201]. The mucosal humoral response against EHV-1 is an important aspect in EHV-1 pathogenesis since it is the first site of contact during natural EHV-1 infections. Experimental infection of foals has demonstrated the production of EHV-1-specific mucosal antibodies which was absent following vaccination. Furthermore, these EHV-1-specific mucosal antibodies were predominantly of the IgA isotype and also was the most long lasting. Furthermore, the same study demonstrated that mucosal IgA has virus neutralizing capability and may play an important role in reducing virus shedding. For brief moments during the acute phase of infection EHV-1-specific IgGa and IgGb were in excess of IgA but these IgG isotypes did not have any virus neutralizing capability [202].

There is evidence to suggest that the immunity bestowed on the host following an

EHV-1 induced abortion incidence is more durable than the immunity gained by contracting the EHV-1 induced respiratory disease [200]. Natural as well as experimental

EHV-1 infections of horses stimulate the production of both VN and CF antibodies starting around 2 weeks post infection by which time the CF antibody titer showed a peak. The VN antibodies, however, required another week to acquire peak titers [202-204]. Antibodies against EHV-1 targets the envelope glycoproteins gB, gC, gD, gH and gp2 while gC and gD glycoproteins contain neutralizing epitopes. EHV-1 specific VN antibodies lasts up to a year and is considered more protective against reinfection than CF antibodies which lasts only up to 10 weeks [204-208]. However, several studies have demonstrated that high titers of serum VN antibodies are more protective against the respiratory disease induced by

EHV-1 than abortions or EHM [209-211].

1.12.3. Cell mediated immune response

Cytotoxic activity against virus infected host cells is mainly carried out by CD8+ T lymphocytes and this is true in the case of EHV-1 infections as well [193-195; 212].

Following infection, EHV-1 internalizes itself into host cells and sequentially works its way into developing CAV which may last up to 3 weeks [213]. This intra-cellular nature of EHV-1, as with all viruses helps it in evading destruction via VN and CF antibodies. As such, the CLT response generated against EHV-1 is essential for virus clearance [163; 195].

Allen et al 1995 has demonstrated that the EHV-1-specific CTL response is carried out mainly by MHC-1 restricted CD8+ T lymphocytes and the maximum lysis of virus infected target cells through this CTL activity happens between 2-3 weeks post infection.

Furthermore, this generated EHV-1-specific CTL response may persist up to a year [212].

Other Experimental infection studies have demonstrated that ponies with repeated EHV-1 exposure contains a higher number of EHV-1-specific CTLs in their circulation and exhibited significantly reduced or no clinical and virological signs following infection

when compared to naive ponies with little to no EHV-1-specific CTLs in their circulation and displayed a more severe form of the infection [195; 213]. Furthermore, studies have also shown that following EHV-1 infection, the number of CD8+ T lymphocytes increase in lungs as well as in the peripheral circulation of mares significantly [214; 215].

A study done by Kydd et al 2003 using experimental infection of pregnant mares with EHV-1 has demonstrated that mares with EHV-1-specific CTLs in the circulation immediately before and approximately 1 month after experimental infection did not abort whereas the mares that did not have circulating EHV-1-specific CTLs did [195]. This protection granted by EHV-1-specific CTLs against EHV-1 induced abortions maybe through the prevention of CAV which is essential for the virus to reach the pregnant uterus after initial infection of the upper respiratory tract [195]. Findings from these studies indicate that the rise and recruitment of CD8+ T lymphocytes coincide with the recovery from EHV-1 infections and hence might play an important role in clearing the virus [195].

Dendritic cells transfected with the EHV-1 IE protein has shown to increase the EHV-1- specific CTL activity in-vitro indicating a possible epitope that might be useful in the development of an EHV-1 specific vaccine. Furthermore, dendritic cells transfected with the EHV-1 glycoproteins gC, gD, gI and gL has also shown increased EHV-1-specific CTL activity in individual horses suggesting these proteins may also act as inducers of a CTL response against EHV-1 [216].

1.13. Inflammation and its mediators

The term inflammation is derived from the Latin word “inflammare” which means

“to burn” and it is one of the most important processes utilized by host cells against injury or infections [217]. During the inflammation process a series of highly organized events

takes place involving cellular and vascular events as well as specific humoral secretions

[217]. These organized events induce a multitude of cellular and vascular pathways that induces vasodilation, migration of white blood cells, capture and adhesion of inflammatory cells and the secretion of mediators and signaling molecules which contribute to the development of inflammation [217]. Inflammation is characterized as acute and chronic.

During the acute phase of inflammation increase in blood flow to the affected tissue, vascular permeability as well as accumulation of fluid, leukocytes, and inflammatory mediators such as cytokines is observed. The chronic phase of inflammation is accompanied with the development of specific adaptive (humoral and cellular) immune responses to the infection [218].

1.13.1. Cytokines

Cytokines are one of the most important mediators of acute and chronic inflammation [217; 219]. This group of relatively small signaling proteins act on a variety of host cells at picomolar or nanomolar concentrations to impart their inflammatory effects

[219; 220]. A total of 44 chemokines and 23 chemokine receptors and over 50 cytokines have been identified in humans to date [221]. Cytokines can be divided as either pro or anti-inflammatory, based on their structural homology or receptor utilization [220]. Many host cells has the capacity to secrete cytokines including stromal, fibroblasts, and endothelial cells and the binding of a particular cytokine with its receptor triggers a signaling cascade that induces many events associated with inflammation such as adhesion, phagocytosis, and apoptosis [217; 220]. The most prominent proinflammatory cytokines are TNFα, IL-1β, IL6 and type I interferons while IL12, IL23 and IL10 have anti- inflammatory properties [219].

An interesting as well as an important group of cytokines are the interferons which are a subset of the class 2 α-helical cytokines. Interferons can be subdivided into two classes as type I and type II interferons and members of both classes play important roles in providing protection against invading pathogens [222]. In particular, IFNα and IFNβ of the type I interferon group imparts antiviral effects on a variety of cells and prevents the replication and the spread of viruses. Furthermore, type I IFNs stimulate the cytotoxic activity of T-cells, natural killer cells, monocytes and macrophages which facilitates the clearance of virus infected cells [222].

1.13.2. Chemokines

Together with cytokines, chemokines play an important role in the inflammatory process. The attraction of inflammatory cells towards sites of injury or infection is an integral part of inflammation which is mediated by chemokines [219; 220; 223]. Around

40 chemokines that play roles in inflammation has been identified up to date. Chemokines are around 8-10 kDa in size and they share an amino acid homology between 20-70%.

Chemokines are divided into families based on relative position of their cysteine residues

[223; 224]. Based on this classification there are 4 subfamilies of chemokines but only two have been extensively characterized [223; 224]. The α-chemokines, based on the presence of a glutamic acid–leucine–arginine near the N terminal can be chemo attractive to neutrophils or lymphocytes [223; 224]. In contrast, β- chemokines are not chemo attractive to neutrophils. Instead they act on monocytes, eosinophils and basophils [223; 224].

Chemokines impart their chemo attractive action on immune cells by binding with specific

G-protein–coupled cell-surface receptors [225]. These receptors are constitutively expressed on some cell types whereas they are inducible on some cell types [223]. A wide

variety of injury and disease conditions require the activation and accumulation of inflammatory cells and this requires the secretion of chemokines [224]. The different subgroups of chemokines that are being secreted during a tissue injury or an infection determines, in part, the inflammatory infiltrates. The ability to control the movement of inflammatory cells by chemokines present novel opportunities to develop therapeutics that may be effective in controlling and managing inflammatory disease conditions [223; 224].

1.13.3. Toll-like receptors

The selective pressure imposed on multi-cellular organisms by pathogens has advanced the evolution of the immune system [226]. Traditionally, the mammalian immune system has been categorized in two arms: The innate immune system and the adaptive immune system [227]. The adaptive immune system utilizes antigen receptors present on effector cells such as T and B lymphocytes to detect and mount an immune response against foreign antigens [228]. The innate immune was first described over a century ago and it is the first line of defense against invading pathogens [226; 228; 229].

The innate immune system senses these invaders through pathogen associated molecular patterns (PAMPs) or damage associated molecular patterns (DAMPs) present on them

[226; 229]. The host cells uses a specialized group of structurally diverse cell surface and intra-cellular receptors known as pattern recognition receptors (PRRs) to sense these

PAMPs or DAMPs and one such extensively studied group of PRRs are the Toll-like receptors (TLRs) [229].

The discovery of Toll receptors were first done in Drosophila as a receptor involved in the establishment of dorso-ventral polarity in the developing embryo [230]. However, later it was discovered that the absence of Toll receptors made Drosophila susceptible to

fungal infections [231; 232] suggesting its importance in the host immune response. In mammals, 12 different homologs have been identified for the Drosophila Toll receptors and they play crucial roles in the host immune response [233; 234]. Mammalian TLRs can be structurally characterized based on the presence of an extra-cellular a leucine rich repeat

(LRR) domain and an intra-cellular TIR domain [228]. Human TLRs can be grouped into

5 sub families based on their amino acid sequences: the TLR2, TLR3, TLR4, TLR5, and

TLR9 subfamilies. The TLR2 subfamily is composed of TLR1, TLR2, TLR6, and TLR10 while the TLR9 subfamily is composed of TLR7, TLR8, and TLR9 [228]. Furthermore, it has been demonstrated that different TLRs recognizes different categories of ligands. In general, TLR2 recognizes peptidoglycan, bacterial lipoproteins, and zymosan, TLR3 detects double-stranded RNA, TLR4 can detect lipopolysaccharide and heat-shock proteins, TLR5 is associated with flagellin detection, TLRs 7 and 8 can detect single- stranded RNA and TLR9 is involved with the detection of CpG motifs of unmethylated bacterial DNA [235].

1.13.3.1 Equine Toll-like receptors

As mentioned earlier, mammalian cells express 12 different TLRs [233; 234]. However, in most cases a particular cell type may not express all 12 TLRs. There isn’t much data on the expression of different TLRs in equine tissues. Only a handful of studies have reported the expression of different TLRs in equine cells [234; 236-239]. A study done by Gornik et. al

(2011) has demonstrated the expression of TLR 2, 3, 4, 6 and 9 mRNA in equine corneal, conjunctival and limbal tissue [237]. A study done by Berndt et. al. (2006) has demonstrated a correlation between TLR4 and IL8 mRNA expression in bronchial epithelial cells of horses with recurrent airway obstruction due to neutrophilic airway

inflammation [236]. Furthermore, LPS treated equine lung tissue has been shown to increase the expression of TLR2 and TLR4 mRNA [239].

The expression of TLR9 in equine pulmonary intravascular macrophages,

Neutrophils in pulmonary capillaries and type-II epithelial cells of alveolar septa by

Schneberger et. al. (2009) [238]. Furthermore, the same study demonstrated the expression of TLR9 in pulmonary vascular endothelial cells, airway epithelial cells and in alveolar macrophages [238]. Another study using equine tissue samples from the kidney, jejunum, liver, spleen and mesenteric lymph nodes detected TLR transcripts for TLRs 1-4, -9 6 and

TLR10 [240]. However, not much data is available on the expression of TLRs in equine endothelial cells and this is one aspect which has been addressed by chapter 3 of this thesis.

Moreover, a study done by Soboll-Hussey et.al. (2014) has demonstrated in-vitro EHV-1 infection of equine respiratory epithelial cells result in the increased expression of TLR3 and TLR9 mRNA [190]. Furthermore, the same study demonstrated the upregulation of

INFα, IFNβ, IL1, IL6, TNFα, GM-CSF, MCP-1 and IL8 mRNA suggesting an involvement in TLRs 3 and 9 in upregulating these cytokines and chemokines [190].

1.14. Genomics in infectious diseases

With the advances made in sequencing since 1995, the global analysis of genomes, epigenomes, transcriptome, proteome and the metabolome with regards to various diseases conditions has advanced the knowledge of disease pathology as well as disease diagnostics.

Furthermore, it has shed light into the different ways a host responds to infections [241;

242]. One advantage of genomics information is that it offers the opportunity for personalized medicine in clinical and public health settings [243]. Such efforts are now

being utilized to control and prevent diseases such as cancers, heart disease and neurodegenerative diseases as well as in the management of infectious diseases and epidemics [243]. As such, genomic analysis is fast becoming an indispensable tool in therapeutics and diagnostics.

1.14.1. Transcriptome in infectious diseases

The transcriptome is defined as the complete set of transcripts in a cell, and their quantity, for a specific developmental stage or physiological condition [244]. With the advances made in sequencing, investigating the transcriptome has become an essential aspect in understanding the functional elements of the genome as well as the molecular constituents of cells that play key roles during disease progression [244].

To determine the transcriptomes of various samples multiple technologies have been utilized thus far. Theses including hybridization-or sequence-based approaches [244].

Hybridization-based approaches usually involve the incubation of fluorescently labelled cDNA with custom-made microarrays or commercial high-density oligo microarrays.

Hybridization-based approaches are high throughput and relatively inexpensive however they require an existing knowledge of gene sequences of the target organism. Furthermore, this method is inefficient at accurately quantifying low or high levels of gene expression

[244; 245]. Tag-based sequencing methods such as serial analysis of gene expression and cap analysis of gene expression were utilized to overcome the limitations of hybridization- based approaches. Indeed, the tag-based methods were much more accurate in quantifying gene expression microarrays. However, these methods were relatively expensive while requiring laborious methodology and relatively large input RNA amounts [244; 245]. The recent technological advancements in high-throughput DNA sequencing paved the way for

the development of a novel method for mapping and quantifying transcriptomes. This method, named RNASeq (RNA sequencing), is both qualitative and quantitative and is capable of accurately measuring both low as well as high levels of gene expression [244;

245]. Furthermore, it can be used to quantify the expression levels of isoforms [244; 245].

Precision or personalized medicine is a new and emerging area in modern medicine that provide personalized diagnosis, treatment, and prevention of disease conditions by taking into account an individual’s genetic information, environment, and lifestyle [246].

Since the genetics and therefore, the gene expression varies considerably among individuals, the response towards infections and injuries may also show considerable variations among different individuals. These differences in gene expressions can be quantified and analyzed with RNAseq which may provide valuable information in diagnosing, controlling as well as preventing disease conditions taking into account individual variations among patients [247].

Infectious diseases are one of the main causes of morbidity and mortality worldwide amassing up to approximately 15 million of 57 million (over 25%) annual deaths [241]. Newly emerging as well as re-emerging pathogens contribute to a significant portion of these deaths while the frequent use of antibiotics is driving the emergence of antibiotic resistant bacterial strains that pose a grave public health risk. Furthermore, with regards to conventional anti-viral therapies, the repertoire of druggable viral proteins are extremely limited and also promotes the development of resistance [241]. Due to these drawbacks in conventional treatments against pathogens, modern drug discovery strategies are focused on identifying and manipulating host-oriented molecules that play key roles in

host-pathogen interactions during disease progression. As such, transcriptomics is an indispensable tool in the future of drug discovery and therapeutics [241; 248].

1.14.2. Genome-wide association studies (GWAS) in infectious diseases

Genome-wide association studies (GWAS) is utilized to screen hundreds of thousands to millions of genetic variants across the genomes of many individuals to identify genotype–phenotype associations. This genomics approach, which share similarities with the transcriptomics-based approach, has revolutionized field of complex disease genetics over the past decade [249]. The first publication of a GWAS was in 2005 for age-related macular degeneration and since then over 50,000 GWAS studies have reported significance between genetic variants and common diseases traits [249; 250]. In more detail, GWAS has been successfully utilized to identify risk loci for a vast number of complex disease phenotypes such as anorexia nervosa, major depressive disorder, cancers and diabetes mellitus, among others [249; 251-253]. Furthermore, the identification of

SNPs on genes which implicates risk loci often leads to the discovery of genes of unidentified functions or of previously unsuspected relevance. Additionally, it also leads to the discovery of novel biological pathways and mechanisms underlying disease phenotypes [249; 250; 254]. Indeed, a GWAS done in Dr. U.B.R. Balasuriya’s lab has also demonstrated significance between the development of persistent EAV infection in stallions and the presence of the CXCL16s receptor variant. Furthermore, this study was the first GWAS study done to investigate genotype–phenotype associations in horses [64;

65; 67].

As such, GWAS is an important tool for the identification of novel mutations in genes in the form of single nucleotide polymorphisms (SNPs) which may predispose

individuals towards developing diseases. It is also an important tool in clinical care as it helps in identifying novel drug targets and disease biomarkers which, together with transcriptomics techniques has the potential to advance personalized medicine [249].

1.15. Scope and outline of dissertation

EHV-1 is a double-stranded DNA virus whereas EAV is a positive sense, single- stranded RNA virus. Therefore, these two viruses are genetically very different from one another. However, they share some interesting similarities in their pathology and disease outcomes. EHV-1, similar to most alpha herpesviruses, establishes latent infection following primary infections while EAV establishes persistent infection in the male reproductive tract of the horse. Furthermore, following primary infection, the development of CAV is an essential step for both EHV-1 and EAV to spread from its 1ry site of infection to secondary replication sites. Characteristic to both these viruses is their endotheliotropic nature and thus, following CAV, they infect and replicates in equine endothelial cells resulting in vasculitis. Induction of vasculitis in the blood vessels of the pregnant uterus is the underlying reason for abortions in pregnant mares following EHV-1 or EAV infections.

Moreover, EHV-1 infections can infect the endothelial cells supplying the CNS and the resulting vasculitis induces EHM in affected horses, which is the neurological form of

EHV-1. As such, vasculitis plays an important role in the pathogenesis and disease outcomes of these two viruses.

The main objective of this thesis was to investigate the innate immune response of

EECs that contributes to the development of vasculitis following infection with EHV-1 and

EAV in-vitro as well as to compare the innate immune responses induced by the two virus strains. In particular, we investigated the involvement of the pro-inflammatory cytokines,

chemokines and Toll-like receptors as well as the biological pathways that may play a role in the induction of vasculitis. The central hypothesis of this study was whether in-vitro infection of EECs with EHV-1 and EAV virus strains will induce similar combinations of pro-inflammatory cytokines, chemokines and Toll-like receptors.

Vasculitis is the inflammation of the walls of the blood vessels and a vast number of mechanisms contribute to the development of vasculitis [255]. However, the production of pro-inflammatory cytokines and chemokines by immune and endothelial cells play a significant role in the development of vasculitis [255]. Since the upregulation of proinflammatory cytokines and chemokines occur frequently in vasculitis syndromes [255;

256], understanding the pro-inflammatory cytokine and chemokine profile of EECs infected with EHV-1 and EAV is important in determining the pathogenesis of these two viruses. The characterization of the innate immune response of EECs against EHV-1 and

EAV has not been done extensively, in particular the pro-inflammatory cytokine and chemokine response towards viral infections. As such the second chapter of this thesis was focused on elucidating the gene expression as well as the secretion of pro-inflammatory cytokines and chemokines from virus infected EECs to determine their contribution in promoting vasculitis.

The proinflammatory cytokine and chemokine response by endothelial cells is part of the innate immune response and it mediates this response by sensing pathogen associated molecular patterns (PAMPs) or damage associated molecular patterns (DAMPs) [226;

229]. Toll-like receptors (TLRs) sense PAMPs and DAMPs and initiate an innate immune response via proinflammatory cytokines and chemokines. However, no extensive study has been carried out to definitively confirm the full repertoire of TLRs expressed by EECs or

to determine the involvement of TLRs in inducing an inflammatory response from EECs following infection with EHV-1 or EAV. Therefore, chapter 3 of this thesis was focused on filling this gap of information on EHV-1 and EAV pathogenesis in EECs.

Chapter 4 of this thesis utilized RNA-seq to further investigate the transcriptome of

EHV-1 and EAV infected EECs. The differential expression of host genes following infection with EHV-1 and EAV provides insights into the biological pathways that are affected during the infection process.

Chapter 5 of this thesis was focused on identifying host genetic factors associated with EHV-1 induced myeloencephalopathy (EHM) in horses. The development of EHM in horses following EHV-1 infections has been linked with the neuropathogenic phenotype of EHV-1 which is a result of a single nucleotide substitution within ORF30 [51]. Since the development of EHM in horses cannot be explained solely by this factor, we investigated the involvement of host genetic factors influencing EHM using a genome- wide association study (GWAS).

CHAPTER TWO

COMPARISON OF THE PROINFLAMMATORY GENE MODULATION IN

EQUINE ENDOTHELIAL CELLS BY EQUID HERPESVIRUS-1, EQUID

HERPESVIRUS-4 AND THE EQUINE ARTERITIS VIRUS

2.1. SUMMARY

Equid herpesvirus-1 (EHV-1) and equid herpesvirus-4 (EHV-4) are closely related

DNA viruses while EAV is an RNA virus. All three viruses are endotheliotropic viruses of the horse with a global presence and having a significant economic impact on the equine industry. Even though genetically very different, all these viruses share close similarities in their disease outcomes. The infection of endothelial cells with these viruses results in vasculitis that is the underlying etiology for the disease conditions caused by these three viruses. Here we in-vitro infected EECs with either the two EHV-1 strains (T953 and

T262), EHV-4 (T445) or EAV and utilized RT-qPCR, Luminex and flow cytometry to investigate the proinflammatory cytokine and chemokine profile. In-vitro infection of

EECs with the two EHV-1 strains (T953 and T262) and EAV produced higher viral titers when compared with EHV-4 (T445). However, based on relative gene expression and

Luminex findings the proinflammatory cytokine and chemokine response of EECs induced by EAV is relatively less than the corresponding values for EECs infected with the three herpesviruses. Furthermore, EAV downregulates the type 1 interferon response in EECs by comparison with the EECs infected with the three herpesviruses.

Key words: EHV-1, EAV, RT-qPCR, Luminex, Flow cytometry, IFNα,

2.2. INTRODUCTION

Equid herpesvirus–1 (EHV-1) and Equid herpesvirus–4 (EHV-4) are double stranded DNA viruses of the subfamily Alphaherpesvirinae and genus Varicellovirus [28] that are endemic in horse populations in most continents [18]. Furthermore, these two viruses have striking similarities in nucleotide (55-84%) and amino acid (55-96%) sequences [257; 258]. Equine arteritis virus (EAV) is a positive sense, single stranded RNA virus belonging to the order Arteriviridae and genus Alphaarterivirus

(https://talk.ictvonline.org/taxonomy/)[259] and it also has a worldwide distribution [5; 6].

Collectively, these viruses are a significant threat to the global equine industry, as well as the $122 billion US equine industry (https://www.horsecouncil.org/resources/economics/).

Even though reproductive consequences following EHV-4 infections are relatively rare, all three viruses are associated with respiratory and reproductive diseases in horses and have a tropism for equine endothelial cells (EECs) of the small blood vessels in horses

(endotheliotropic) [5; 6; 17; 18; 88]. Infection of EECs with these viruses results in cell injury which leads to thrombosis and vasculitis of small blood vessels [5; 6; 17; 19; 52;

170]. Furthermore, some EHV-1 strains [neuropathogenic (e.g. T953 strain)] are capable of infecting endothelial cells of small blood vessels which supply the central nervous system (CNS) causing vascular thrombosis and vasculitis leading to a frequently fatal neurological disease called equine herpesvirus myeloencephalopathy (EHM) [15; 18; 52;

178]. Recently, EHM outbreaks have been reported in an increasing number of countries including the USA, prompting the USDA to designate it as an emerging disease [175].

However, most of the strains of EHV-1 [non-neuropathogenic (e.g. T262 strain)] and occasionally EHV-4 strains cause thrombosis and vasculitis in small vessels of the late

pregnant uterus resulting in abortion [18; 260]. Thus, the underlining etiology for the disease conditions caused by all three viruses used in this study is the viral vasculitis of the

CNS and the late gravid uterus.

EAV was first isolated from the lung of an aborted fetus in a standardbred breeding farm near Bucyrus, Ohio in 1953 [121]. Since then, EAV outbreaks have occurred in the

US and other parts of the world causing direct and indirect economic losses due to abortion in pregnant mares, increased mortality in young foals and establishment of persistent infection in stallions [27; 89; 90]. In contrast to neuropathologic EHV-1 strains, EAV infections do not cause neurological clinical signs but similar to non-neuropathogenic

EHV-1 and some EHV-4 strains, they can cause abortion through endothelial cell injury which leads to severe necrotizing panvasculitis and increased vascular permeability [5; 19].

Therefore, the main factor associated with the pathological outcomes of these virus infections is vasculitis. Vasculitis is the inflammation of the walls of the blood vessels and a vast number of mechanisms contribute to its development [255]. However, the production of cytokines and expression of adhesions molecules by immune and endothelial cells play a significant role in the development of vasculitis and thrombosis of the small blood vessels [255]. Since the up regulation of proinflammatory cytokines and chemokines occurs frequently in vasculitis syndromes [255; 256], understanding the proinflammatory cytokine and chemokine profile of equine endothelial cells (EECs) infected with EHV-1,

EHV-1 (T262), EHV-4 (T445) and EAV is important in contributing to the understanding of the pathogenesis of vasculitis caused by each of these viruses. Furthermore, even though genetically very different, all four viruses share some close similarities in their disease outcomes and pathogenesis and the fact that vasculitis is a major factor in their

pathogenesis led us hypothesize that the proinflammatory cytokine and chemokine profile of EECs infected with these viruses should also have some close similarities. Several studies have explored the cytokine and chemokine gene expression of immune cells, EECs and also the serum cytokine levels to EAV [64; 261] and EHV-1 and -4 infections [191;

262; 263]. The primary objective of this study however, was to compare the proinflammatory cytokine and chemokine gene expression as well as protein secretion by

EECs following infection with these three endotheliotropic viruses in order to establish whether their similarities in disease outcomes and pathogenesis would be reflected in gene and protein expression in infected EECs. As such, we hypothesized that the proinflammatory cytokine and chemokine gene expression as well as secretion of these virus infected EECs would show similar combinations.

2.3. MATERIALS AND METHODS

2.3.1 Cell lines

Equine endothelial cells (EECs) were derived from the pulmonary artery of a horse as previously been described [264]. The EECs were cultured in Dulbecco’s Modified

Eagle’s Medium (DMEM, Corning, New York, NY) with 10% bovine calf serum (BCS,

Hyclone Laboratories, Inc., Logan, UT), 100 U/mL penicillin–streptomycin, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids and 200 mM L-glutamine (Thermo Fisher

Scientific, Waltham, MA) in a humidified incubator at 37 °C with 5% CO2. All experiments were performed in EECs between passage numbers 15-20.

2.3.2 Viruses

Four viruses were used in this study. The horse adapted virulent Bucyrus strain of

EAV (VBS; VR796; ATCC), the neuropathogenic T953 (Findlay) strain [14] and the abortigenic T262 strain [265] of equine herpesvirus 1 and the T445 strain of EHV-4. The

EHV-1 and EHV-4 viruses were kindly provided by the late Dr. George Allen, Maxwell

H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky. All viruses were propagated in EECs to generate high titer working stocks. Briefly, triple decker cell culture flasks (Nunc, Rochester, NY) containing EEC monolayers were infected with each of the four viruses. The flasks were frozen at -80°C when a 100% cytopathic effect (CPE) was observed. Cell lysates were clarified by centrifugation (2000 x g) at 4 °C for 15 min, followed by ultracentrifugation (Beckman Coulter, Miami, FL) at 121,600 x g for EAV and 100,000 x g for EHVs through a 20% sucrose cushion in NET buffer (150 mM NaCl, 5 mM EDTA, 50 mM Tris-HCl, pH 7.5) at 4 °C for 4 h to pellet the virus.

Purified preparations of each virus strain were resuspended in Dulbecco’s phosphate buffered saline (DPBS, Thermo Fisher Scientific, Waltham, MA), briefly sonicated and 50

µL aliquots were frozen at -80 °C until further use. Virus stocks were titrated by standard plaque assay in EECs, and titers were expressed as the number of plaque forming units

(PFU) per milliliter (PFU/mL).

2.3.3 Antibodies

To determine the presence of viral antigens in in-vitro infected EECs, the following monoclonal antibodies (mAbs) were used. To detect EAV antigens, mAb 12A4 against

EAV non-structural protein 1 (nsp-1) [266] was conjugated with Alexa Fluor 488 (AF-488)

[267]. To detect EHV-1 antigens in T953 and T262 infected cells, the mAb 14H7 against

EHV-1 glycoprotein 13 (gp13) was used [268; 269]. A Melon Gel IgG Spin Purification kit (Pierce, Rockford, IL) was used to purify mouse ascitic fluid containing mAb 14H7.

Purified IgG was directly conjugated to AF-647 (Thermo Fisher Scientific, Waltham, MA) following the manufacturer’s protocol. Briefly, purified IgG from mAb 14H7 was diluted with phosphate-buffered saline (PBS, pH 7.4) to a concentration of 2 mg/mL. The diluted

IgG solution was incubated with AF-647 reactive dye for 1 h at room temperature in the dark with slow stirring. The reaction mixture was loaded onto an exclusion purification resin column for separation of labeled antibodies from unincorporated dye. Fractions of

AF-647 labeled antibodies were collected and stored at 4 °C. To detect nonspecific binding, an AF-647 conjugated mouse IgG2a [270] isotype control was used (BD Biosciences, San

Jose, CA).

2.3.4 Viral growth curves

The one step growth curves in EECs for each virus strain used in this study in EECs were plotted over a period of 24 hours post infection (hpi) according to a standard laboratory protocol. Briefly, EECs were plated in 6 well plates (3×106 cells per well) and left overnight in a humidified incubator at 37 °C in the presence of 5% CO2 for the cells to attach and form a cell monolayer. Following this, the media was aspirated, and the monolayer was washed once with 1 mL of DMEM (Corning, New York, NY) without

BCS. The EEC monolayer was then infected with 200 µL of Minimum Essential Media

(MEM, Corning, New York, NY) containing virus strains (EHV-1 [T262 or T953], EHV-

4 [T445], or EAV) at a multiplicity of infection (MOI) of 2 and incubated in a humidified incubator at 37 °C in the presence of 5% CO2 for 1 h. After 1 hour the medium containing the virus was removed and the EEC monolayer was washed three times with pre- warmed

(37 °C) PBS. Following this, pre-warmed (37 °C) DMEM (Corning, New York, NY) with

10% BCS (Hyclone Laboratories, Inc., Logan, UT), 100 U/mL penicillin–streptomycin, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids and 200 mM L-glutamine

(Thermo Fisher Scientific, Waltham, MA) was added to the wells and the plates incubated in a humidified incubator at 37 °C in the presence of 5% CO2. Tissue culture fluid (TCF) supernatant from the virus infected EEC monolayers were collected at time points 3, 6, 9,

12, 18 and 24 hpi. The collected TCF was used to plot the growth curves of the virus strains using the Reed and Muench 50% end point (TCID50) method.

2.3.5 RNA extraction and quantitative real-time RT-PCR (RT-qPCR)

EECs were seeded in 6 well plates (3×106 cells per well) and left overnight in a humidified incubator at 37 °C in the presence of 5% CO2 for the cells to attach and form a cell monolayer. Following this, monolayers were infected with the four virus strains as described above. Upon completion of infection, the virus containing MEM (Corning, New

York, NY) was aspirated and pre-warmed (37 °C) DMEM (Corning, New York, NY) with

10% BCS (Hyclone Laboratories, Inc., Logan, UT), 100 U/mL penicillin–streptomycin, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids and 200 mM L-glutamine

(Thermo Fisher Scientific, Waltham, MA) was added into each well and re-incubated until

6 or 12 hpi. EECs grown in identical conditions absent virus infection was used as the mock control. TCF supernatants from each of the virus and mock infected EEC monolayers were collected and stored at -80 °C to be used later for Luminex assay (Millipore sigma,

Burlington, MA) analysis. RT-qPCR was performed on total RNA extracted from EHV-1,

EHV-1 (T262), EHV-4 (T445) and EAV infected EECs at 6 and 12 hpi using the miRNeasy mini kit (Qiagen, Hilden, Germany). Total RNA from uninfected EECs cultured under

identical conditions was used as the reference sample. Any contaminating genomic DNA present was removed by performing DNase I digestion (RNase free DNase set, Qiagen,

Hilden, Germany). The total RNA quality and quantity was assessed using a Bioanalyzer

(Agilent technologies, Santa Clara, CA). One micro gram of total RNA was reverse transcribed to generate cDNA using a High capacity reverse transcription kit (Thermo

Fisher Scientific, Waltham, MA). Real-Time PCR amplification was carried out using 1:1 diluted cDNA to assess the relative expression of different proinflammatory cytokine and chemokine genes using commercially available and custom made equine TaqMan™ primers and probes (Thermo Fisher Scientific, Waltham, MA) in a 7500 Fast Real-Time

PCR system (Thermo Fisher Scientific, Waltham, MA). The panel of proinflammatory cytokine and chemokine genes investigated in this study are listed in Table 2.3. All reactions were performed in triplicate. Equine β- glucuronidase gene was used as an endogenous control to normalize the quantitative RT-qPCR data as has been described in previous studies [64]. PCR efficiency for all reactions was assessed using LinRegPCR

[271]. For relative quantification, fold changes in gene expression were calculated using the ΔΔCT method [272].

2.3.6 Cytokine and chemokine multiplex assay

To screen for the proinflammatory cytokines and chemokines secreted by EECs infected with the viruses used in this study, Luminex technology based equine multiplex cytokine assays were carried out. Analysis of IFNα, IFNɣ, IL17, IL4 and IL10 content was performed using a horse cytokine 5 plex assay at the Animal Health Diagnostic Center at the University of Cornell, Ithaca, NY as described by Wagner et. al. 2009 [273]. All the other cytokine (IL1α, IL1β, IL12, IL18, TNFα, GM-CSF and G-CSF) and chemokine (IL8,

CCL2, CCL5, CXCL10, Eotaxin and Fractalkine) protein analyses were carried out using the MILLIPLEX MAP Equine Cytokine/Chemokine Magnetic Bead Panel - Immunology

Multiplex Assay (Millipore sigma, Burlington, MA) in conjunction with the Luminex

200™ System (Luminex Corporation, Austin, TX). TCF supernatant collected previously were used for the analysis. In brief, 25 µL of the TCF supernatants collected from virus infected EEC cultures at 6 and 12 hpi were treated with the primary antibody cocktail

(IL1α, IL1β, IL8, IL12, IL18, CXCL10, CCL2, CCL5, Eotaxin, Fractalkine, TNFα, G-CSF and GM-CSF) according to the manufacturer’s protocol. The treated cell culture supernatants were loaded into a 96 well plate provided with the kit and incubated overnight at 4 °C on a shaker. Following incubation, the 1ry antibody cocktail is removed and the detection antibody was added and incubated with agitation on a plate shaker for 1 hour at room temperature. Streptavidin-Phycoerythrin was added to each well containing the detection antibody followed with a 30-minute incubation and washing. The plate was read using a Luminex 200™ System (Luminex Corporation, Austin, TX).

2.3.7 Flow cytometry analysis of EECs

In order to make certain that the difference in gene expression levels and cytokine production by EHV-1 and EAV infected EECs was not due to the differences in the infection levels of EECs, flow cytometry was performed. EECs were seeded in 6 well plates (3×106 cells per well) and left overnight in a humidified incubator at 37 °C in the presence of 5% CO2 for the cells to attach and form a cell monolayer. Following this, the monolayer was infected with either EHV-1 (T953 or T262) or EAV at a MOI of 2 as described above. Upon the completion of infection, the virus containing MEM (Corning,

New York, NY) was aspirated and pre-warmed (37 °C) DMEM (Corning, New York, NY)

with 10% BCS (Hyclone Laboratories, Inc., Logan, UT), 100 U/mL penicillin– streptomycin, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids and 200 mM L- glutamine (Thermo Fisher Scientific, Waltham, MA) was added into each well and re- incubated until a 12 hpi timepoint was reached. After 12 h of incubation, EECs were harvested from plates using a cell scraper. Up to 1×106 cells were incubated at room temperature in 200 µL of 2% paraformaldehyde for 30 minutes to fix the cells.

Permeabilization of EECs was carried out by incubating EECs using 200 µL of 1% saponin at room temperature for 5 minutes. Following permeabilization, EHV-1 (T953 and T262) infected EECs were incubated with 14H7-AF647 mouse mAb and EAV infected EECs were incubated with 12A4-AF488 mouse mAb for 30 minutes at room temperature. After washing, cells were resuspended in flow buffer (PBS supplemented with 10% normal goat serum and 0.1% sodium azide) and acquisition was performed using a BD LSR II flow cytometer (Becton Dickinson, Franklin Lakes, NJ). Data were analyzed using the BD

CellQuest™ pro software (Becton Dickinson, Franklin Lakes, NJ).

2.3.8. Statistical analysis

The statistical analysis of the virus growth curves was performed using the natural log values (Ln) of the virus titers employing t tests. For RT-qPCR analysis, ANOVA was performed using the means of fold changes. Statistical analysis of Luminex data was performed using t tests as well. The significant level (p-value) for all experiments was set at 0.05 for all tests. Statistical analysis was performed using the JMP pro 13 software

(JMP®, Version 13 SAS Institute Inc., Cary, NC, 1989-2019).

2.4. RESULTS

2.4.1 Virus growth curve comparison

First, the production of infectious progeny virus in EECs were analyzed using two

EHV-1 strains (T953 and T262), EHV-4 (T445) strain and EAV over a period of 24 hours.

The growth curves of the virus strains were plotted using the Reed and Muench 50% endpoint (TCID50) method [274].

The growth patterns of the three herpesviruses were similar. The titers of the three herpesviruses shows a sharp increase from 12 to 18 hpi timepoints indicating the viruses might be completing one infectious cycle in EECs between these two timepoints (Figure

2.1). Furthermore, the titer at the 18 hpi timepoint was statistically significantly higher than the titer at the 12 hpi timepoint for the two EHV-1 strains (T953 and T262) but not for

EHV-4 (T445) (Tables 2.1 and 2.2). At the end of 24 h, of the three herpesviruses, the highest virus titer was achieved with EHV-1 (T262) followed by EHV-1 (Table 2.1).

However, the virus titers of these two viruses at the 24 hpi timepoint were not statistically significant (Table 2.2). EHV-4 (T445) achieved the lowest virus titer after 24 hpi and the final titer was statistically significantly lower than all the other virus titers at the same timepoint (Tables 2.1 and 2.2).

Compared to the three herpesviruses, EAV had a different growth pattern (Figure

2.1). In the case of the growth curve of EAV the virus titer increases steeply from 6 to 9 hpi timepoints indicating the virus might be completing one infectious cycle in EECs between these two timepoints (Figure 2.1). Furthermore, the titer at the 9 hpi timepoint was significantly higher than the titer at the 6 hpi timepoint (Table 2.1). Based on the virus

growth curves, it can be concluded that the two EHV-1 strains (T953 and T262) and EAV elicited a similar ability to replicate in EECs when compared with EHV-4 (T445) which generated a lower virus titer in EECs.

2.4.2 Inflammatory cytokine and chemokine gene expression of EECs following virus infection

We explored the gene expression profile of eight proinflammatory cytokines (IL6,

IL17, IL18, TNFα, IFNα, IFNω, IFNɣ and GM-CSF) and eight proinflammatory chemokines (IL8, CXCL9, CXCL10, CXCL11, CXCL16, CCL3, CCL4 and CCL5) in virus infected EECs at 6 and 12 hpi timepoint using RT-qPCR. Gene expression values from 3 independent biological replicates are reported for the 4 virus strains used in this study. The relative quantification values of these genes in EECs infected with each virus are provided in Table 2.3. The gene expression levels of 4 proinflammatory cytokines

(IL18, GM-CSF, CXCL16 and IFNɣ) and 2 proinflammatory chemokines (IL8 and CCL3) were upregulated less than 5-fold at both 6 and 12 hpi timepoints in EECs infected with these four viruses. All other genes showed a 5-fold or a greater gene expression, at least at one timepoint (6 or 12 hpi).

Differences in gene expression in virus infected EECs between 6 and 12 hpi timepoints was investigated. In EHV-1 infected EECs, three cytokines (IL6, IL17c, IFNα) and one chemokine (CCL5) showed a statistically significant increase in gene expression from 6 to 12 hpi (Table 2.3). In EHV-1 (T262) and EHV-4 (T445) infected EECs, the gene expression of only IL17c increased significantly from 6 to 12 hpi (Table 2.3). In EAV infected EECs, three cytokines (IL17c, IFNα, IFNω) and one chemokine (CCL4) increased significantly from 6 to 12 hpi (Table 2.3).

Any differences in gene expression in EECs infected with the four virus strains were compared with each other. As mentioned previously, a 5-fold or greater change in gene expression was observed in the case of proinflammatory cytokines IL6, IL17, TNFα,

IFNα, IFNω and the proinflammatory chemokines CXCL9, CXCL10, CXCL11, CCL4 and

CCL5. Interestingly, the gene expression levels of these proinflammatory cytokines and chemokines in infected EECs were similar among the three herpesvirus and were not significantly different from one another. When compared with the gene expression levels of the EAV infected EECs however, the three herpesvirus infected EECs had a higher gene expression for all of the cytokines and chemokines and some of them were significantly different.

When considering the EEC gene expression levels of EHV-4 (T445) vs EAV, at the 6 hpi timepoint, IL6, IFNα and CXCL11were significantly higher in the case of EHV-

4 (T445) infected EECs (Table 2.4). At the 12 hpi timepoint, IL17, CXCL10, CXCL11 and

CCL5 gene expression were significantly higher in EHV-4 (T445) infected EECs (Table

2.4). When considering the EEC gene expression levels of EHV-1 (T262) vs EAV, at the

6 hpi timepoint, CXCL11 and IFNω gene expressions were significantly higher in EHV-1

(T262) infected EECs (Table 2.5). At the 12 hpi timepoint, IFNα gene expressions was significantly higher in EHV-1 (T262) infected EECs (Table 2.5). When considering the

EEC gene expression levels of EHV-1 vs EAV, at the 6 hpi timepoint, CXCL11, IFNα and

CCL4 gene expressions were significantly higher in EHV-1 infected EECs (Table 2.6). At the 12 hpi timepoint, IL6 gene expressions was significantly higher in EHV-1 infected

EECs (Table 2.6). As such, the gene expression data indicates that the three herpesviruses induce a higher proinflammatory response in EECs when compared with EAV.

During our analysis of virus growth curves in EECs we observed that, out of these

4 virus strains, EHV-4 (T445) achieved the lowest virus titer during replication in EECs.

However, during our analysis of the cellular response towards virus infection in terms of proinflammatory cytokine and chemokine gene expression, we do not see a statistically significant difference between EHV-4 (T445) and its two closely related EHV-1 cousins

(T953 and T262). This indicates that EECs can mount a defense response against EHV-4

(T445) infection in the same magnitude as in EHV-1 strains. Interestingly, EAV shows the lowest fold changes for proinflammatory cytokine and chemokine gene expression when compared with the three herpesviruses indicating that compared to the three herpesviruses used in this study, EAV suppresses the proinflammatory response of EECs.

2.4.3 Inflammatory cytokine and chemokine production by EECs against virus infection

Following RT-qPCR analysis of virus infected EECs, we observed that compared to the three herpesviruses EAV suppresses the proinflammatory gene response in EECs.

Here, we investigated if this suppression at the mRNA level is also present at the protein level. We analyzed the levels of proinflammatory cytokines and chemokines produced by virus infected EECs using a Luminex assay. A total of 18 analytes were analyzed using the

TCF supernatants collected from EEC monolayers infected with the four virus strains used in this study at both 6 and 12 hpi timepoints. TCF supernatant from uninfected EEC monolayers was used as the negative control. Here we present data from 3 independent experiments. Out of the 18 cytokines and chemokines analyzed, 15 (IL1α, IL1β, IL4, IL8,

IL10, IL12, IL18, MCP1, CCL5, IFNɣ, Eotaxin, Fractalkine, TNFα, G-CSF and GM-CSF) did not show any detectable amounts at either of the two timepoints for any of the virus

infected EEC cultures. Out of these analytes, IL8, IL18, GM-CSF and IFNɣ showed very low levels of gene expression in RT-qPCR analysis at both 6 and 12 hpi timepoints as well suggesting that in vitro infection of EECs by these 4 virus strains do not influence their production at the mRNA as well as protein levels. CCL5 and TNFα showed gene expression levels over 5 folds in infected EECs in RT-qPCR analysis but their presence was not detected in the TCF supernatants of infected EECs suggesting that in vitro infection of EECs by these 4 virus strains induce their mRNA expression but not the secreted proteins. Only three analytes (IL17, CXCL10 and IFNα) were present in detectable amounts in the TCF supernatants of infected EECs.

At the 6 hpi timepoint, only the TCF supernatants from EEC cultures infected with the three herpesvirus strains produced detectable levels of IFNα but didn’t show a statistically significant difference from the negative control (Figure 2.2). At the 12 hpi timepoint, IFNα levels produced by EECs infected with the three herpesviruses increased and this increase from 6 to 12 hpi was statistically significant for each of the three herpesviruses (Figure 2.2). However, the IFNα levels produced by the three herpesviruses at the 12 hpi timepoint were not significantly different from one another. In contrast, at both 6 and 12 hpi timepoints, the TCF supernatants from EAV infected EECs did not contain any detectable amounts of IFNα (Figure 2.2). This indicates that in vitro infection of EECs with EHV-1 and EHV-4 strains induce the secretion of IFNα while EAV suppresses IFNα production from EECs.

At both 6 and 12 hpi timepoints, IL17 was detected in the TCF of EECs infected with all 4 virus strains (Figure 2.3). However, at both these timepoints, the levels of IL17 was not significantly different from the negative control or between each other indicating

that in vitro infection of EECs by these 4 virus strains does not induce IL17 secretion

(Figure 2.3).

At the 6 hpi timepoint, CXCL10 levels produced by the two EHV-1 strains (T953 and T262) was significantly higher than the level produced by EAV infected EECs (Figure

2.4). CXCL10 levels produced by EHV-4 (T445) was higher though not statistically different from what was produced by EAV infected EECs (Figure 2.4). At the 12 hpi timepoint, the CXCL10 levels produced by EECs infected with the 3 herpesviruses were significantly higher than the negative control. However, the increase from 6 to 12 hpi was statistically significant only for EHV-4 (T445) infected EECs (Figure 2.4). Interestingly,

CXCL10 produced by EAV infected EECs was not statistically different from the negative control at both 6 and 12 hpi timepoints. Furthermore, at the 12 hpi timepoint, The CXCL10 produced by EHV-1 infected EECs was significantly higher than EAV infected EECs.

EHV-1 (T262) and EHV-4 (T445) infected EECs produced CXCL10 levels higher than

EAV infected EECs at 12 hpi even though no statistical significance was observed among them.

2.4.4 Flow cytometry

Based on the results of the RT-qPCR and Luminex assays the proinflammatory cytokine and chemokine production by the EAV infected EECs is lower than that of the herpesviruses. Therefore, in order to make sure this reduction is not due to reduced EEC infection levels by EAV we performed flow cytometry analysis of virus infected EECs.

EECs were in-vitro infected with EHV-1 (T262), EHV-1 and EAV at MOI of 2 and incubated for 12 h. Following incubation, cells were stained with mAbs against EHV-1 gp13 and EAV nsp-1. Flow cytometry analysis indicated that on average, EAV percentage

of infected EEC was 38.91 whereas the average for EHV-1 and EHV-1 (T262) was 29.35 and 38.07 respectively (Figure 2.5). This would suggest that the reduced proinflammatory cytokine, chemokine and IFN gene expression as well as production by EAV infected

EECs was not due to reduced EAV infection and confirms that compared with the herpesviruses, EAV suppresses the inflammatory response elicited by endothelial cells following infection.

In conclusion, the proinflammatory cytokine and chemokine gene expression and secretion in EAV infected EECs appear to be lower when compared with the three herpesvirus infected EECs.

2.5 DISCUSSION

This study explored the growth characteristics and the proinflammatory cytokine, chemokine and interferon gene expression and production in EECs following infection with EHV-1, EHV-4 and EAV. All three viruses are endotheliotropic, meaning they target endothelial cells during infection resulting in vasculitis which is the underlining etiology for the disease conditions caused by these three viruses [5; 6; 19; 52; 170]. A characteristic of vasculitides is the dysregulated production of cytokines by immune and endothelial cells which often leads to inflammation of blood vessels followed by ischemia that results in damage to the organ being supplied with [255; 256]. This study utilized two EHV-1 strains

(T953 and T262). EHV-1 is a prototypic neuropathogenic strain with an ORF30 G2254 genotype whereas the T262 strain of EHV-1 is a non-neuropathogenic yet abortogenic strain with an ORF30 A2254 genotype. This study also included EHV-4 (T445) which is very closely related to EHV-1 and the virulent Bucyrus strain of EAV which is a highly pathogenic and endotheliotropic virus of horses.

Results of the study demonstrated the production of progeny virus by EECs followed a similar pattern for the three EHV strains was different for EAV. This should not surprising since EHV-1 and -4 strains are closely related DNA viruses whereas EAV is an RNA virus. However, as demonstrated by the virus growth curves, the amount of virus particles remaining in EHV-1 (T262) and EAV infected EECs following 3 washes is relatively higher than in EHV-1 and EHV-4 (T445) infected EEC monolayers. One possible reason for this could be that 3 washes may not be sufficient to remove excess virus particles.

During natural infection, EHV-1 strains can cause abortion and occasionally EHM by inducing viral vasculitis. However, such disease manifestations are rare following natural EHV-4 infection, which is primarily regarded as a respiratory pathogen. This is because leukocyte associated viremia is not common following EHV-4 infections [178;

275] and in cases where viremia is detected following EHV-4 infections, viral DNA remains in low amounts in PBMCs and only for a short period of time [276]. These factors might be preventing the successful transfer of EHV-4 from its primary sites of infection to endothelial cells of the pregnant uterus or the CNS where it can infect and induce viral vasculitis. Studies including this one have demonstrated that EHV-4 is capable of infecting

EECs at least in-vitro [277]. Furthermore, this study demonstrates that the cellular response of EECs towards EHV-1 and EHV-4 infections, for the most part, remains the same with regards to the proinflammatory gene expression and the production of proinflammatory cytokines and chemokines. Therefore, it is reasonable to assume that if EHV-4 strains infect endothelial cells in vivo the vasculitis process and the following disease manifestations in the form of abortions or EHM might be the same as in EHV-1 strains.

The results of this study indicate that the lack of such disease manifestations is most probably due to the virus not being transported to endothelial cells of blood vessels of the pregnant uterus and the CNS rather than EHV-4 strains not being able to infect endothelial cells and induce an inflammatory response. Therefore, the results of this study strongly point towards inefficient leukocyte associated viremia as a major factor in the mild disease outcomes associated with EHV-4 infections.

Results from this study indicate that compared to the three herpesviruses infected

EECs, EAV infected EECs elicit a relatively lower proinflammatory response. The secretion of CXCL10, which is a proinflammatory chemokine, by EAV infected EECs was significantly lower when compared with the three herpesviruses infected EECs. CXCL10 activates CXCR3 receptors which are predominantly expressed on T and B lymphocytes, natural killer cells, dendritic cells and macrophages which induces chemotaxis [278-280].

As such CXCL10 plays an important role in inflammation and protection against virus infections. Therefore, reduced production of CXCL10 relative to herpesvirus infected

EECs suggests a suppressed proinflammatory response in EAV infected EECs.

Furthermore, CXCL10 aids herpes simplex virus type 2 infections by stimulating virus replication [281]. Therefore, the higher CXCL10 production in EHV infected EECs may be an adaptation by the EHVs to facilitate their replication in host cells.

The production of IFNα, was undetected at both 6 and 12 hpi timepoints in EAV infected EECs. Flow cytometry analysis showed that at 2 MOI in-vitro infection, the percentage of EAV infected EECs are higher than EHV-1 (T953 or T262) infected EECs ruling out lower EEC infection by EAV as the cause for the absence of an IFNα response.

Go et al. (2014) has shown that EAV suppresses IFNβ production in EECs due to the

interference caused by EAV nsp1 on IRF3 and NF-휅B [138]. As such, the observation made by Go et al. (2014) taken together with the relative gene expression and Luminex data from this study makes a strong case for the same interference mechanism by EAV nsp1 to be at play in IFNα inhibition in EAV infected EECs.

Go et al. (2014) has shown that EAV suppresses IFNβ production in infected EECs.

Here, we show that EAV suppresses IFNα production as well, indicating the suppression of the type I interferon pathway in EECs. The porcine reproductive and respiratory syndrome virus, which is another member of the family Arteriviridae, is also known to suppress the type I interferon system [136; 282]. Being a component of the innate immune system, type I (IFNα and IFNβ) and type II (IFNɣ) interferons are essential for the host antiviral response towards invading viruses [135; 283]. The results of this study have also shown that EECs do not express IFNɣ mRNA or secrete IFNɣ following infection by any of the four viruses used in this study indicating that EECs may not be a source for IFNɣ production during viral infections. Sarkar et. al. (2015) has shown that T953 strain of EHV-

1 suppress IFNβ production from EECs. However, this study indicates that the production of IFNα remains intact in EHV-1 and EHV-4 infected EECs. In accordance, other studies have demonstrated the induction of IFNα from EHV-1 infected PBMCs [284] and airway epithelial cells [285; 286] suggesting that the EHV-1 suppression of the type 1 interferon system may be limited to just IFNβ. During EHV-1 infections, the lesions are restricted to the uterus, CNS and the upper respiratory tract with no evidence of systemic lesions such as pan vasculitis observed during EAV infections. The fact that at least one arm of the type

I interferon system remaining intact during EHV-1 infections maybe a contributing factor in preventing systemic lesions whereas the complete suppression of the type I interferon

system in EECs during EAV infections maybe contributing to the much more severe systemic vascular lesions observed during EAV infections.

Table 2.1: Average virus titers of the three EHV-1 strains and EAV used in this study at different timepoints. These values were obtained from 3 independent experiments. Virus 3 hpi 6 hpi 9 hpi 12 hpi 18 hpi 24 hpi EHV-1 102.6 ± 102.6 ± 102.8 ± 103.6 ± 106 ± 106 ± (T262) 100.1 100.1 100.38 100.1 100.5 100.6

EHV-1 100.5 ± 100.9 ± 101.8 ± 102.4 ± 104.75 ± 105.25 ± 100.9 100.9 100.7 100.6 100.5 100.4

EHV-4 100.3 ± 100.3 ± 100.9 ± 100.9 ± 102.25 ± 102.9 ± (T445) 100.6 100.6 100.9 100.9 100.5 100.5

EAV 102.25 ± 102.1 ± 104.5 ± 105.6 ± 105.6 ± 105.8 ± 100.4 100.3 100.25 100.1 100.1 100.4

Table 2.2: The p values of statistically significant virus titers (p<0.05) among different virus strains (three EHV-1 strains and EAV) at 3-24 hpi. Statistical analysis was done using the natural log values of the log 10 TCID50 values obtained from cell culture supernatants. NS indicates timepoints with no statistical significance

Timepoint VBS v VBS v VBS v T262 v T262 v T953 v (hpi) T445 T262 T953 T445 T953 T445 3 p=0.014 NS p=0.023 p=0.006 p=0.09 NS 6 p=0.02 NS NS p=0.004 p=0.024 NS 9 p=0.0004 p=0.004 p=0.003 p=0.02 NS NS 12 p<0.0001 p=0.008 p=0.0004 p=0.002 NS p=0.04 18 p<0.0001 NS NS p<0.0001 p=0.03 p=0.0005 24 p=0.0005 NS NS p=0.0003 NS p=0.002

Table 2.3: Average fold change in gene expression ± SD in EECs following EHV-1 (T953 and T262), EHV-4 (T445) and EAV. This data is from three independent biological replicates for each of the virus strains. Bold texts show where the increase of fold change from 6 to 12 hpi was statistically significant (p≤0.05) Virus EHV-1 (T262) EHV-1 EHV-4 (T445) EAV Gene 6hpi SD 12hpi SD 6hpi SD 12hpi SD 6hpi SD 12hpi SD 6hpi SD 12hpi SD IL6 19.4 18.7 28 25.9 14.2 14.7 31.2 31.5 15.6 9.4 16.7 11.2 3.1 3.3 5.8 7.0 IL-8 1.7 0.2 3.8 1.3 2.4 1.1 1.8 2.3 1.6 0.2 1.4 0.9 0.2 0.2 1.5 0.6 IL-17 1.7 0.5 13.8 13.9 1.7 0.6 5.2 1.4 2.0 0.7 7.2 2.1 1.3 0.5 4.1 0.9 IL-18 1.6 0.2 1.3 1.2 0.7 0.4 0.3 0.0 1.6 1.0 1.4 1.3 1.8 0.3 0.8 0.0 CXCL9 156.3 259.6 230.7 196.3 172.6 282.6 182.4 283.6 173.8 164.7 753.3 692.9 38.4 54.0 154.9 22.1 CXCL10 111 167.4 101.5 71.9 137.2 193.2 219.7 184.6 1075.6 1560.7 2011.5 3250.5 52.7 51.8 54.2 68.7

82 CXCL11 130.5 174.7 345.5 304.7 204.3 292.4 265.7 377.9 561.5 417.7 622.2 216.4 14.5 21.7 115.2 38.1

TNFα 11.3 13.1 112.8 124.5 4.9 4.8 34.1 26.4 8.1 9.3 18.0 12.3 1.2 1.0 25.2 14.8 IFNα 96.1 84.1 251.6 207 129.6 114.8 404.6 343.5 198.1 275.6 517.5 575.2 6.9 5.3 57.3 43.8 IFNω 57.3 49 115.7 72.8 87.4 37.5 169.5 135.7 138.7 120 48.6 32.2 20.1 17.2 129.3 118.4 IFNɣ 1.7 0.0 0.7 1.0 0.8 1.2 0.0 0.0 0.8 1.2 1.0 1.4 0.6 0.8 1.6 0.5 GM- 2.9 2.9 4.5 1.0 1.5 0.4 3.3 2.0 1.3 0.9 1.6 0.7 0.7 0.4 4.4 2.3 CSF CCL3 2.4 1.1 4.1 1.5 1.9 0.3 3.6 0.7 2.1 0.4 1.5 0.4 0.6 0.1 0.4 0.3 CCL4 11.7 9.3 66.1 38.2 12.9 9.7 57.3 57.6 15.0 18.4 42.3 14.0 1.6 0.2 12.5 8.5 CCL5 22.8 24.4 151.7 135.3 20.6 20.7 163.6 195.1 82.9 70.7 1488.5 1953.8 3.7 3.7 8.5 1.2 CXCL16 1.4 0.7 2.2 0.9 1.3 0.7 2.0 0.5 1.1 0.2 1.5 0.3 0.7 0.2 1.6 0.2

CXCR6 1.6 0.0 1.1 0.2 1.8 0.4 1.2 0.6 1.7 0.4 1.3 0.3 1.1 1.1 1.0 0.9

Table 2.4: Average gene expressions that are significantly different between EHV-4 (T445) and EAV at 6 and 12 hpi. Bold texts indicate the higher fold change. Gene 6hpi 12hpi EHV-4 EAV p value EHV-4 EAV p value (T445) (T445) IL6 15.6 3.1 0.025 - - - CXCL11 561.5 14.5 0.036 622.2 115.2 0.007 IFNα 198.1 6.9 0.049 - - - IL17 - - - 7.2 4.1 0.038 CXCL10 - - - 2011.5 54.2 0.033 CCL5 - - - 1488.5 8.5 0.025

Table 2.5: Average gene expressions that are significantly different between EHV-1 (T262) and EAV at 6 and 12 hpi. Bold texts indicate the higher fold change. Gene 6hpi 12hpi EHV-1 EAV p value EHV-1 EAV p value (T262) (T262) CXCL11 130.5 14.5 0.041 - - - IFNω 57.3 20.1 0.025 - - - IFNα 251.6 57.3 0.004

Table 2.6: Average gene expressions that are significantly different between EHV-1 and EAV at 6 and 12 hpi. Bold texts indicate the higher fold change. Gene 6hpi 12hpi EHV-1 EAV p value EHV-1 EAV p value CXCL11 204.3 14.5 0.022 - - - IFNα 129.6 6.9 0.02 - - - CCL4 12.9 1.6 0.028 - - - IL6 - - - 31.2 5.8 0.014

Figure 2.1: Virus growth curves of EHV-1 (T262), EHV-1, EHV-4 (T445) and EAV in EECs.

Figure 2.2: IFNα production by EECs infected with EHV-1 (T262 and T953), EHV-4 (T445) and EAV. * indicates statistically significant increase from mock at 12hpi (p≤0.05). #* indicates the increase in IFNα production from 6 to 12 hpi is statistically significant (p ≤0.05)

Figure 2.3: IL17 production by EECs infected with EHV-1 (T262 and T953), EHV -4 (T445) and EAV. The increase in IL17 production from 6 to 12 hpi is statistically not significant (p ≥0.05)

Figure 2.4: CXCL10 production by EECs infected with EHV-1 (T262 and T953), EHV-4 (T445) and EAV. # indicates statistically significant increase from mock at 6hpi (p≤0.05). * indicates statistically significant increase from mock at 12hpi (p≤0.05). #* indicates statistically significant increase from 6 to 12 hpi (p≤0.05).

Figure 2.5: Flow cytometry results of the percentage of virus infected EECs when compared to uninfected EECs (A). EHV-1 (T262) infected EECs (B) with a 41.21% infection rate, EHV-1 infected EECs (C) with a 29.35% infection rate and EAV infected EECs (D) with a 45.43 % infection rate.

CHAPTER THREE

RNA SEQUENCE ANALYSIS OF EQUINE ENDOTHELIAL CELLS

FOLLOWING IN VITRO INFECTION WITH EQUINE HERPESVIRUS-1 AND

EQUINE ARTERITIS VIRUS STRAINS

3.1. SUMMARY

Equine herpesvirus-1 is double stranded DNA virus in the genus Varicellovirus of the subfamily Alphaherpesvirinae. Equine arteritis virus is a positive sense single stranded

RNA virus in the order Arteriviridae and genus Alphaarterivirus

(https://talk.ictvonline.org/taxonomy/). Both these viruses are endotheliotropic and pose a significant threat to the $102 billion US equine industry. In the previous chapter we investigated the gene expression of proinflammatory cytokines and chemokines in EHV-

1, EHV-4 and EAV infected EECs. The results of that study indicated that the expression of proinflammatory cytokine and chemokine genes are higher in EHV-1 infected EECs compared to EAV infected EECs, and also that EAV downregulates the expression of IFNα in EECs following infection. In this study we investigated the transcriptome of EHV-1 and

EAV infected EECs at 6 and 12 hpi timepoints using RNA-Seq. The differentially expressed genes were used in combination with gene ontology software to reveal a complex pattern of gene regulation and cellular pathways related to cellular immune, inflammatory and apoptotic responses. The results revealed clusters of differentially expressed genes in

EHV-1 and EAV infected EECs. Furthermore, the “gonadotrophin hormone releasing receptor pathway”, “CCKR signaling map” and the “Inflammation mediated by cytokine and chemokine pathway” was differentially expressed frequently in EECs infected with both EHV-1 and EAV.

Key words: RNA-seq, EHV-1, EAV, EECs, PANTHER, Gene ontology

3.2. INTRODUCTION

Equine herpesvirus-1 (EHV-1) and the equine arteritis virus (EAV) are two endotheliotropic equine viruses that pose a significant threat to the $102 billion US equine industry. EHV-1 is a double stranded DNA virus in the genus Varicellovirus of the subfamily Alphaherpesvirinae [18; 28]. Its type D DNA genome is about 150 kb with about

57% G+C content [18; 287]. The genome of EHV-1 is arranged into a long unique region covalently linked to a short unique region which is flanked by internal and terminal repeat sequences [288; 289]. Furthermore, EHV-1 strains have 80 open reading frames (ORFs) containing 76 genes capable of coding for 77 different viral proteins [18]. EAV is a positive sense single stranded RNA virus in the genus Equarterivirus of the family Arteriviridae

[290]. The genome of EAV strains include a 5’ leader sequence and at least 10 ORFs which encodes for 13 non-structural and 8 structural proteins (7 envelope proteins and 1 nucleocapsid protein). The size of the EAV genome varies between 12,704 and 12,731 bp among different viral strains [5; 6; 291]. In horses, these two viruses are associated with respiratory and reproductive diseases. Upon infection, both EHV-1 and EAV strains cause mild upper respiratory tract infections and abortions in pregnant mares [5; 6; 17-19].

However, neuropathogenic strains of EHV-1, such as the T953 (Findlay) strain [14], can cause a rare but fatal neurologic condition in horses called equine herpes myeloencephalopathy (EHM) [15; 18; 52; 178]. A single nucleotide polymorphism (SNP) in the DNA polymerase gene (ORF30) of neuropathogenic EHV-1 strains has been linked with the development of EHM in infected horses [51]. EHV-1 strains with a Guanine (G) residue at the nucleotide position 2254 in the ORF 30 are categorized as neuropathogenic

while strains that have an Adenine (A) residue at this position are considered non- neuropathogenic [51]. However, EAV is not linked with the development of any neurological diseases.

As mentioned earlier, these two viruses are endotheliotropic, meaning endothelial cells are their main target. Several studies have investigated the gene expression of endothelial cells following in-vitro infection by these two viruses [138; 263; 292]. Gene expression studies of equine endothelial cells (EECs) infected with EAV and EHV-1 has shown that both these viruses suppress the production of IFNβ [138; 292]. Furthermore, a study done by our group has demonstrated that EAV suppresses IFNα production in infected EECs but not EHV-1 and the upregulation of proinflammatory cytokines and chemokines in EHV-1 infected EECs is relatively higher than EAV infected EECs

(mentioned in chapter 2). Furthermore, PCR array studies done on EECs infected with

EHV-1 have demonstrated the upregulation of proinflammatory cytokines [263].

Therefore, we wanted to further investigate the interferon and proinflammatory cytokine gene expression and production in EECs by these two virus strains. However, no study has been done to extensively investigate the transcriptome changes induced by EHV-1 and

EAV in infected EECs and to elucidate the host biological pathways most affected by the infection of these two virus strains. Therefore, the purpose of this study was to elucidate the transcriptome changes in EECs following infection with either EHV-1 or EAV.

In this study, we utilized RNA-seq to further investigate the transcriptome of EHV-

1 and EAV infected EECs. The complete set of transcripts in a cell and its quantity is referred to as the transcriptome and understanding the transcriptome is important in understanding disease conditions [293]. Most of the widely used methods available to

investigate the cellular transcriptome profile are based on hybridization or sequence-based approaches [293]. Hybridization-based approaches utilize the incubation of fluorescently labelled cDNA with custom-made microarrays or commercial high-density oligo microarrays [293]. However, the use of hybridization-based approaches are limited by the existing knowledge about genome sequences and high background levels caused by cross- hybridization [294; 295], and it is difficult to compare the expression levels among different experiments [293]. However, with the development of high-throughput DNA sequencing, mapping and quantifying transcriptomes have become much efficient and accurate. Termed RNA-Seq (RNA sequencing), this approach utilizes recently developed deep sequencing technologies [293]. Compared to other methods, RNA-seq has several advantages since it allows the investigation of all unique sequences of the genome and it is a method that is very sensitive and offers a large dynamic range. This makes RNA-seq the better method to detect and quantify levels of RNAs expressed at very low levels over microarrays [296].

3.3. MATERIALS AND METHODS

3.3.1. Cells

Endothelial cells derived from the pulmonary artery of the horse (EECs) [264] were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM, Corning, New York,

NY) with 10% bovine calf serum (BCS, Hyclone Laboratories, Inc., Logan, UT), 100

U/mL penicillin–streptomycin, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids and 200 mM l-glutamine (Life Technologies, Carlsbad, CA) in a humidified incubator at

37 °C with 5% CO2. All experiments were performed in EECs with passage numbers 15-

20.

3.3.2. Viruses and purification

The horse adapted virulent Bucyrus strain of EAV (VBS, VR-796; ATCC) and the neuropathogenic T953 (Findlay) strain (Henninger et al., 2007) were used in this study.

The EHV-1 strain was kindly provided by the late Dr. George Allen at the Maxwell H.

Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky. All viruses were propagated in EECs to generate high titer working stocks. Briefly, triple decker cell culture flasks (Nunc, Rochester, NY) containing EECs were infected with each of the above-mentioned viruses. The flasks were frozen at -80 °C when a 100% cytopathic effect

(CPE) was observed. Cell lysates were clarified by centrifugation (2000 x g) at 4 °C for 15 min, followed by ultracentrifugation (Beckman Coulter, Miami, FL) at 121,600 x g through a 20% sucrose cushion in NET buffer (150 mM NaCl, 5 mM EDTA, 50 mM Tris-HCl, pH

7.5) at 4 °C for 4 h to pellet the virus. Purified preparations of each virus strain were resuspended in phosphate-buffered saline (PBS; pH 7.4) and frozen at -80 °C. Virus stocks were titrated by standard plaque assay in EECs, and titers were expressed as the number of

Plaque forming units (PFU) per milliliter (38).

3.3.3 In-vitro infection

EECs (Passage 15-20) were plated in 6 well plates (3×106 cells per well) with

Dulbecco’s Modified Eagle’s Medium as mentioned above and left overnight in a humidified incubator at 37 °C with 5% CO2 for the cells to attach to the bottom of the wells and form a monolayer. After overnight incubation, the media was aspirated, and the EEC monolayer was washed once with 1 mL of Dulbecco’s Modified Eagle’s Medium without

bovine calf serum. The cell monolayer was then infected with 200 µL of Minimum

Essential Media (MEM, Corning, New York, NY) containing either EHV-1 or EAV at a multiplicity of infection (MOI) of 2 and incubated in a humidified incubator at 37 °C with

5% CO2 for 1 hour to complete the infection of EECs. Following the 1 h incubation period, the media containing the virus was removed and the EEC monolayer was washed three times with warm PBS and pre-warmed Dulbecco’s Modified Eagle’s Medium (DMEM,

Corning, New York, NY) with 10% bovine calf serum (BCS, Hyclone Laboratories, Inc.,

Logan, UT), 100 U/mL penicillin–streptomycin, 1 mM sodium pyruvate, 0.1mM nonessential amino acids and 200 mM L-glutamine (Life Technologies, Carlsbad, CA) was added to the wells and was incubated in a humidified incubator at 37 °C with 5% CO2.

Cells were harvested at 6- and 12-hours post infection (hpi) timepoints for total RNA was extraction. Total RNA from uninfected EECs cultured under identical conditions mentioned above was used as the negative control.

3.3.4. Total RNA extraction

EHV-1, EAV or mock infected EECs were used for total RNA extraction following

6 and 12 hpi. Total RNA was extracted using the miRNeasy mini kit (Qiagen, Hilden,

Germany). Total RNA was extracted from three biological replicates for every sample at every timepoint. Any contaminating genomic DNA present was removed by performing

DNase I digestion (RNase free DNase set, Qiagen, Hilden, Germany). The total RNA quality and quantity was assessed using a Bioanalyzer (Agilent technologies, Santa Clara,

CA). All samples had an RNA integrity number > 8.0.

3.3.5. Sample submission

A minimum of 300 ng of total RNA was submitted to Exiqon (Exiqon, South

Korea) for whole transcriptome sequencing with single end sequencing with 75 bp read lengths and 30 million read counts. Library preparation was performed using the TruSeq

Stranded mRNA Sample Prep Kit (Illumina, San Diego, CA, USA) per manufacturer’s instructions. The adapter for read 1 was;

AGATCGGAAGAGCACACGTCTGAACTCCAGTCACNNNNNNATCTCGTATGCC

GTCTTCTGCTTG, with NNNNNN being the index sequence.

The read 2 adapter was;

AGATCGGAAGAGCGTCGTGTAGGGAAAGAGTGTAGATCTCGGTGGTCGCCGT

ATCAT. The libraries were quantified by qPCR. Sequencing was performed on a HiSeq

4000 (Illumina) using a HiSeq 4000 sequencing kit version 1, generating 150 bp single end reads. FASTQ files were generated and demultiplexed using bcl2fastq v2.17.1.14

Conversion Software (Illumina).

3.3.6. Data analysis:

For RNA-seq data analysis, the reads were initially trimmed for adapters and quality using TrimGalore Version 0.4.4 (Babraham Bioinformatics; www.bioinformatics.babraham.ac.uk), then trimmed reads were aligned to the horse reference genome (EquCab3.0) using STAR (Release 2.5.2b) [32], and annotated to the equine reference transcriptome available in NCBI database using Cufflinks (Release 2.2.1; http://cole-trapnell-lab.github.io/cufflinks/) [33]. Differential expression analysis was performed on the normalized read count (FPKM; fragments per kilobase of exon per million mapped reads) [34]. we used Cuffdiff 2.2.1 to calculate differently expressed genes

(DEG) between samples from the mock and virus infected groups [297]. Significance level was set at FDR-adjusted p value of the test statistic < 0.1 using a Benjamini-Hochberg correction.

For all data analyses, separate databases were created for EHV and EAV samples.

Differentially expressed genes were determined within tissue. Principal components analysis was performed on all genes included in either database to verify clustering. Initial identification of differentially expressed genes was performed using one-way ANOVA, using the Benjamini-Hochberg correction for false discovery rate (FDR P < 0.1).

PANTHER (version 13.1, http://www.pantherdb.org/) was used to annotate DEG in relation to biological process, molecular function, cellular component and pathways [298].

3.4. RESULTS

RNA-seq data obtained for this study came from a total of 18 libraries sequenced from EECs. These were EHV-1 infected EECs at 6 (3 samples) and 12 (3 samples) hpi timepoints, EAV infected EECs at 6 (3 samples) and 12 (3 samples) hpi timepoints and mock infected EECs at 6 (3 samples) and 12 (3 samples) hpi timepoints.

3.4.1 Gene clustering and arrangement

A heatmap with dendrograms and a principal component analysis (PCA) was done to identify the arrangement and clustering of differentially expressed genes of the EHV-1 infected EECs at 6 and 12 hpi timepoints, EAV infected EECs at 6 and 12 hpi timepoints and mock infected EECs at 6 and 12 hpi timepoints (Figures 4.1 and 4.2). Based on the heatmap and the PCA the 3 RNA seq libraries generated from the 3 replicates of EHV-1 infected EECs at the 6 and 12 hpi timepoints clustered together and did not show a

significant variance (Figures 4.1 and 4.2). With EAV infected EECs, the RNA seq libraries generated by the 3 replicates at the 12 hpi timepoint clustered together but did not at the 6 hpi timepoint and showed a significant variance among the replicates (Figures 4.1 and 4.2).

When considering the differential gene expression of EHV-1 infected EECs, at the

6 hpi timepoint majority of the genes were upregulated compared to mock infected EECs

(Figure 4.3A). At the 12 hpi timepoint, however, only about 50% of the genes were upregulated in EHV-1 infected EECs (Figure 4.3B). Furthermore, between 6 and 12 hpi timepoints, most of the genes were upregulated at the 6 hpi timepoint in EHV-1 infected

EECs (Figure 4.3C). When considering the differential gene expression of EAV infected

EECs, similar to EHV-1 infected EECs, the majority of the differentially expressed genes were upregulated at the 6 hpi timepoint when compared with mock infected EECs (Figure

4.3D). At the 12 hpi timepoint, most of the genes that were differentially expressed were downregulated in EAV infected EECs when compared with mock infected EECs (Figure

4.3E). Furthermore, between 6 and 12 hpi timepoints, similar to EHV-1 infected EECs, most of the genes were upregulated at the 6 hpi timepoint in EAV infected EECs (Figure

4.3F).

3.4.2 Differential expression of genes and pathways in EAV vs Mock-infected EECs

The PANTHER 14.1 database (http://www.pantherdb.org/) was used to map the differentially expressed genes in EECs to canonical pathways. When considering the EAV and mock-infected EECs, at the 6 hpi timepoint, 544 genes showed statistically significant differential expression (q≤0.05). Out of those, 367 genes were upregulated while 177 genes were downregulated in EAV infected EECs compared to the mock infected. Out of the 544 differentially expressed genes, 466 could be mapped to the PANTHER 14.1 database to

assign gene ontology. The top five canonical pathways differentially expressed in EAV infected EECs when compared to mock EECs at the 6 hpi timepoint included the

“Cholecystokinin receptor (CCKR) signaling map”, “Inflammation mediated by chemokine and cytokine signaling pathway” and the “Wingless-integrated (Wnt) signaling pathway” with 10 differentially expressed genes each, and the “Epidermal growth factor

(EGF) receptor signaling pathway” and the “Gonadotropin-releasing hormone receptor pathway” with 9 differentially expressed genes each” (Table 4.1). The 10 genes differentially expressed in the “Inflammation mediated by chemokine and cytokine signaling pathway” were all upregulated. Therefore, the “Inflammation mediated by chemokine and cytokine signaling pathway” appears to be predominantly upregulated in

EAV infected EECs when compared to mock EECs at the 6 hpi timepoint. The genes upregulated in the “Inflammation mediated by chemokine and cytokine signaling pathway” included CCL13 and CCL26 (Table 4.1) which play roles in T cell monocyte and macrophage Chemoattraction. Also upregulated in the same pathway was NFKB2 which functions as an inhibitor of NF-κB and aids in controlling the inflammatory response [299;

300] and PREX1, PLA2G4A and GNG4 which are associated with anti-inflammatory effects (Table 3.1) [301-303]. The “Wnt signaling pathway” is known to stimulate the expression of many inflammatory molecules during bacterial infections suggesting it plays a role during the host inflammatory response during infections [304]. Furthermore, increased Wnt pathway signaling is associated with inflammatory disorders [304-306].

Among the upregulated genes in this pathway is WNT9A which is a gene usually upregulated during endotoxemia driven inflammation and inflammatory bowel disease

(Table 4.1) [304; 307]. Out of the 10 genes in the “CCKR signaling map” 5 genes each

were up as well as down regulated. Activation of the “CCKR signaling map” is usually associated with anti-apoptotic and inflammatory effects [308]. The “EGF receptor signaling pathway” is associated with virus entry into cells and inflammation [309-311].

Out of the 9 differentially expressed genes in the “EGF receptor signaling pathway” 3 were upregulated and 6 were downregulated while the “Gonadotropin-releasing hormone receptor pathway” had 2 upregulated genes while 7 were downregulated indicating these two pathways were predominantly downregulated at this timepoint. Therefore, at the 6 hpi timepoint, the top canonical pathways showing differential gene expression are linked to the host inflammatory and immune responses.

At the 12 hpi timepoint between EAV and mock-infected EECs, 3109 genes showed statistically significant differential expression (q≤0.05) and out of those, 1621 genes were upregulated while 1488 genes were downregulated in EAV infected EECs compared to the mock-infected. Out of the 3109 differentially expressed genes, 2675 could be mapped to the PANTHER 14.1 database to assign gene ontology. Among the top 5 canonical pathways differentially expressed at this timepoint were the “Gonadotropin- releasing hormone receptor pathway”, “CCKR signaling map” and the “Inflammation mediated by chemokine and cytokine signaling pathway” and they were differentially expressed at the 6 hpi timepoint as well (Table 4.2).

However, at the 12 hpi timepoint more genes were differentially expressed in these pathways compared to the 6 hpi timepoint (Table 4.2). For the “Gonadotropin-releasing hormone receptor pathway” 51 genes were differentially expressed and out of them 37 were upregulated while 14 were downregulated. The upregulated genes in this pathway includes MAP3K2, MAP3K5, MAP3K11, MAP3K14, MAP2K3 and MAPK8 (Table 4.2)

which are associated with the mitogen-activated protein kinases (MAPK) signaling cascade. MAPK is associated with promoting inflammation and apoptosis [312; 313].

“CCKR signaling map” pathway had 39 differentially expressed genes out of which 30 were upregulated and 9 were downregulated. As mentioned before, the “CCKR signaling map” is associated inflammatory and anti-apoptotic effects [308]. The “Inflammation mediated by chemokine and cytokine signaling pathway” had 36 differentially expressed genes and 25 of them were upregulated while 11 were downregulated. Among the important genes upregulated at this timepoint is JAK2, STAT1, STAT3 and STAT6. These genes are associated with the Janus kinase/signal transducer and activator of transcription

(Jak/STAT) signaling pathway [314] which promotes inflammation during diseases [315].

The “Platelet-derived growth factor (PDGF) signaling pathway” and the “Apoptosis signaling pathway” were also among the top five pathways differentially expressed in EAV vs mock infected EECs at the 12 hpi timepoint. The “PDGF signaling pathway” had 44 differentially expressed genes out of which 33 were upregulated and 11 were downregulated. Vascular endothelial cells are one of the main sites expressing the PDGF receptors (PDGFRs) which initiates the downstream “PDGF signaling pathway” which is involved with vascular inflammation and chemotactic activity [316; 317]. Among the upregulated genes in this pathway is PDGFB which is known to be expressed in endothelial cells and encodes the B chain of PDGF (Table 3.2) [318; 319]. The “Apoptosis signaling pathway” which plays a role in cell death is a cellular defense mechanism against virus infections to prevent the spread of progeny virus [320]. This pathway had 34 differentially expressed genes and out of them 31 were upregulated while 3 were downregulated. EAV promotes cellular apoptosis via caspase-8 and caspase-9 [321] and at this timepoint the

CASP8 and CASP10 genes were upregulated in EAV infected EECs compared to mock infected cells (Table 4.2). Furthermore, MAPK8 (Table 4.2) was also among the upregulated genes associated with this pathway known to promote apoptosis [312] confirming the involvement of caspase genes in promoting apoptosis as well as the triggering of apoptosis in EECs by EAV. Therefore, at the 12 hpi timepoint the top 5 pathways differentially expressed in EAV infected EECs compared to mock infected EECs were associated with the host inflammatory, immune and apoptotic responses. This was similar to the 6 hpi timepoint and these differentially expressed pathways in EECs correlates with EAV pathogenesis.

3.4.3 Differential expression of genes and pathways in EAV 6 hpi vs EAV 12 hpi in

EECs

As mentioned above, 544 and 3109 genes were differentially expressed in EAV infected EECs at the 6 and 12 hpi timepoints respectively when compared with mock- infected EECs. Out of these differentially expressed genes 240 (7%) was differentially expressed at both 6 and 12 hpi timepoints while 304 and 2869 genes were exclusively differentially expressed at the 6 and 12 hpi timepoints respectively (Figure 4.4A). Out of the 240 differentially expressed genes common to both 6 and 12 hpi timepoints, 213 could be mapped to the PANTHER 14.1 database to assign gene ontology. The top 5 differentially expressed pathways associated with these genes were “CCKR signaling map”, “EGF receptor signaling pathway”, “PDGF signaling pathway” “Apoptosis signaling pathway” and the “Gonadotrophin-releasing hormone receptor pathway” (Table

4.3A). Therefore, at both 6 and 12 hpi timepoints, cellular pathways differentially

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expressed in EECs infected EAV are associated with host inflammatory, immune and apoptotic responses.

3.4.4 Differential expression of genes and pathways in EHV-1 vs Mock-infected EECs

When considering the EHV-1 and mock-infected EECs, at the 6 hpi timepoint 4737 genes showed statistically significant differential expression (q≤0.05). Out of those, 4003 genes were upregulated while 734 genes were downregulated in EHV-1 infected EECs compared to the mock-infected. Out of the 4737 differentially expressed genes, 4196 could be mapped to the PANTHER 14.1 database to assign gene ontology. The top five canonical pathways differentially expressed in EHV-1 infected EECs when compared to mock EECs at the 6 hpi timepoint included the “Gonadotropin-releasing hormone receptor pathway” with 59 differentially expressed genes out of which 41 were upregulated and 18 were downregulated (Table 4.3B). The upregulated genes in this pathway included MAP4K5,

MAP3K5, MAP2K1, MAPK11 and MAPK14 (Table 4.3B) and as mentioned before, these genes are associated with the MAPK signaling cascade which is associated with promoting inflammation and apoptosis [312; 313]. The “CCKR signaling map” had 50 differentially expressed genes out of which 35 were upregulated and 15 were downregulated (Table

4.3B). Activation of the “CCKR signaling map” is associated inflammatory and anti- apoptotic effects [308]. The “Integrin signaling pathway” had 48 differentially expressed genes and out of those 30 were upregulated while 18 were downregulated (Table 4.3B).

Integrins are cell surface receptors critical for cell to cell and cell to extra cellular matrix communication. The Herpesviridae family is known to utilize integrins as entry receptors

[322]. However, a study done by Van de Walle et al (2008) has shown that integrins might not be utilized by EHV-1 during infection of EECs. In the transcriptome data we obtained

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for EHV-1 infected EECs, at the 6 hpi timepoint, only the α integrin subunit ITGAE was upregulated while the β subunits ITGB3 and ITGB4 were downregulated suggesting a lesser involvement of integrins during infection of EECs by EHV-1 as mentioned by Van de Walle et al (2008). Furthermore, several studies have indicated the importance of MHC class I molecule as an entry receptor for EHV-1 infections in several cell lines [196; 323].

At the 6 hpi timepoint, transcriptome data indicates the upregulation of EQMHCC1 which encodes for the equine MHC class I heavy chain in EHV-1 infected EECs which bolsters the observation of the involvement of MHC class I molecules in EHV-1 entry into EECs.

Furthermore, the cell adhesion molecules ICAM-1 and VCAM-1 were also upregulated in

EHV-1 infected EECs at the 6 hpi timepoint. The upregulation of these molecules are important for the firm adhesion of leukocytes to the endothelium [324] which may facilitate the transfer of EHV-1 virus particles from EHV-1 infected leukocytes to endothelial cells

[17] and may also play a role in the development of thrombosis which is an important factors associated with EHV-1 pathogenesis [17; 18; 52; 170]. The “Inflammation mediated by chemokine and cytokine signaling pathway” had 47 differentially expressed genes and out of those 38 were upregulated and 9 were downregulated (Table 4.3B).

Upregulated genes included IL6, IL18, IL15, IL11, CCL13, CCL26 and CXCL8, (Table

4.3B) which play important roles in acute inflammation, immunity against infections, T cell, monocyte and macrophage chemoattraction as well as promote systemic vasculitis diseases such as the Churg–Strauss Syndrome [325-329]. Finally, the “Wnt signaling pathway” had 46 differentially expressed genes with 22 of them upregulated while 24 being downregulated (Table 4.3B) and as mentioned before, increased Wnt pathway signaling is associated with inflammatory disorders [304-306]. Therefore, similar to EAV infected

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EECs, EHV-1 infected EECs differentially express cellular pathways associated with host inflammatory and immune responses

At the 12 hpi timepoint between EHV-1 and mock-infected EECs 2503 genes showed statistically significant differential expression (q≤0.05) and out of those, 1046 genes were upregulated while 1457 genes were downregulated in EAV infected EECs compared to the mock-infected. Out of the 2503 differentially expressed genes, 2109 could be mapped to the PANTHER 14.1 database to assign gene ontology. The top five canonical pathways differentially expressed in EHV-1 infected EECs when compared to mock EECs at the 12 hpi timepoint included the following: The “Integrin signaling pathway” had 47 differentially expressed genes. Out of those 47 genes, 8 were upregulated and 39 were down regulated (Table 3.4B). Similar to the 6 hpi timepoint the integrin subunit genes were among the downregulated genes. These included the integrin α subunit genes ITGA2,

ITGA5, ITGA11, and ITGAX, and the integrin β subunit genes ITGB3, ITGB4, and ITGB5

(Table 3.4B), indicating a lesser involvement of integrins during infection of EECs by

EHV-1 as mentioned by previous studies [330]. Furthermore, at the 12 hpi timepoint the cell adhesion molecules ICAM-1 and VCAM-1 were among the upregulated genes just as it was at the 6 hpi timepoint. These molecules play important roles in the firm adhesion of leukocytes to the endothelium which may facilitate the transfer of EVH-1 particles from infected leukocytes to the endothelium and may also contribute to the development of thrombosis [17; 18; 52; 170; 324]. Similar to the 6 hpi timepoint, the “Gonadotropin- releasing hormone receptor pathway” was among the top 5 differentially expressed pathways at the 12 hpi timepoint (Table 4.4B). This pathway had 46 differentially expressed genes out of which 16 were upregulated and 30 were downregulated. The

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upregulated genes in this pathway included MAP3K5, MAPK4, MAPK6 and MAPK11 which are associated with the MAPK signaling cascade that promotes inflammation and apoptosis [312; 313]. The “Angiogenesis” pathway had 35 differentially expressed genes out of which 8 were upregulated and 27 were downregulated (Table 4.4B). Based on the amounts of upregulated genes in these three pathways, we can conclude that these three pathways are downregulated in EHV-1 infected EECs compared to uninfected EECs at the

12 hpi timepoint. The “Inflammation mediated by chemokine and cytokine signaling pathway” and the “CCKR signaling map” was also among the top 5 pathways (Table 4.4B).

The “Inflammation mediated by chemokine and cytokine signaling pathway” had 44 differentially expressed genes and 23 of them were upregulated while 21 were downregulated. Among the upregulated genes were CCL2, CCL5, CCL13, CCL20, CXCL8 and IL6 which play important roles in acute inflammation, immunity against infections, T cell monocyte and macrophage chemoattraction as well as be involved in systemic vasculitis diseases such as the Churg–Strauss Syndrome [325; 326; 328; 331]. The “CCKR signaling map” had 37 differentially expressed genes out of which 21 were upregulated and 16 were downregulated. Therefore, at the 12 hpi timepoint as well, EECs infected with

EHV-1 differentially express cellular pathways associated with host immune and inflammatory responses.

3.4.5 Differential expression of genes and pathways in EHV-1 6 hpi vs EHV-1 12 hpi in EECs

As mentioned above, 4737 and 2503 genes were differentially expressed in EHV-

1 infected EECs at the 6 and 12 hpi timepoints respectively when compared with mock- infected EECs. Out of these differentially expressed genes, 1315 (22.2%) were common to

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both 6 and 12 hpi timepoints while 3422 and 1188 genes were exclusively differentially expressed at the 6 and 12 hpi timepoints respectively (Figure 4.4B). Out of the 1315 differentially expressed genes common to both 6 and 12 hpi timepoints, 1076 could be mapped to the PANTHER 14.1 database to assign gene ontology. The top 5 differentially expressed pathways associated with these genes were “Gonadotrophin-releasing hormone receptor pathway”, “Integrin signaling pathway”, “CCKR signaling map”, “Inflammation mediated by chemokine and cytokine signaling pathway”, and the “Angiogenesis pathway”

(Table 4.5A). Therefore, in EHV-1 infected EECs genes that were commonly differentially expressed were associated with host immune and inflammatory responses.

3.4.6 Differential expression of genes, pathways and biological functions in EHV-1 vs

EAV infected EECs

When considering the differentially expressed genes in EHV-1 and EAV infected

EECs, at the 6 hpi timepoint, 518 genes (10.9%) were common while 4217 and 26 genes were exclusively differentially expressed in EHV-1 and EAV infected EECs respectively

(Figure 4.4C). Out of the 518 common genes, 443 could be mapped to the PANTHER 14.1 database to assign gene ontology. The top 5 differentially expressed pathways associated with these genes were the “Inflammation mediated by chemokine and cytokine signaling pathway”, “CCKR signaling map”, “Wnt signaling pathway”, “EGF receptor signaling pathway” and the “Gonadotrophin-releasing hormone receptor pathway” (Table 4.5A) suggesting the host cellular pathways associated with immune and inflammatory responses are the top pathways differentially expressed in EECs following infection with these two viruses.

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At the 12 hpi timepoint, out of the differentially expressed genes in EHV-1 and

EAV infected EECs, 910 genes (19.4%) were common while 1593 and 2199 genes were exclusively differentially expressed in EHV-1 and EAV infected EECs respectively

(Figure 4.4D). Out of the 910 common genes, 785 could be mapped to the PANTHER 14.1 database to assign gene ontology. The top 5 differentially expressed pathways associated with these genes were the “Gonadotrophin-releasing hormone receptor pathway”, “CCKR signaling map”, “Apoptosis signaling pathway”, “Inflammation mediated by chemokine and cytokine signaling pathway” and the “Integrin signaling pathway” (Table 4.5B).

3.5. DISCUSSION

This study elucidates the transcriptome of EECs infected with EHV-1 and EAV.

3.5.1 Gene clustering and arrangement

According to the heat map and the PCA, the differentially expressed genes in EHV-

1, infected EECs at the 6 and 12 hpi timepoint were clustered together indicating no significant differences in the differential gene expression among the replicates. In EAV infected EECs, at the 12 hpi timepoint, the differentially expressed genes were clustered together indicating no significant differences among the replicates. Two out of the three replicates of EAV infected EECs at the 6 hpi timepoint clustered together but one replicate did not, even though virus from the same EAV stock and EECs with the same passage number was used throughout the experiment. Several reasons may have contributed to this including differences in virus infection efficiency of EECs to the single ended sequencing.

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3.5.2Differential gene expression and cellular pathways

When considering the differential gene expression in EHV-1 and EAV infected

EECs, more genes were upregulated at the 6 hpi timepoint when compared with the 12 hpi timepoint. Virus infections of mammalian cells activates a myriad of gene expression pathways that may assist the host cells in overcoming invading pathogens [332].

Furthermore, it has been demonstrated that with the herpes simplex type I (HSV-1) virus infections, a vast number of host genes get transcriptionally activated even though they do not get translated [333]. These observations fit well with the activation of the transcriptional machinery at the 6 hpi timepoint in EHV-1 infected EECs. However, the innate immune system of host cells also has a tendency to shut down cellular protein synthesis as a measure to control viral replication [334-336] and members of many different virus families too, follow the same trend in shutting down host gene expression and mRNA translation while promoting the translation of viral mRNAs [335]. It has been suggested that the downregulation of host gene transcription by viruses is not merely to acquire more cellular resources for viral biogenesis but to inhibit the host anti-viral response [335]. Therefore, downregulation of the host gene expression in order to circumvent the host antiviral response can be taken as an important factor in viral pathogenesis [335]. A previous study done by Sarkar et al. (2016) has shown that EHV-1 can downregulate the production of IFNβ in infected EECs and studies done by Go et al.

(2014) taken together with the results obtained during the course of this study indicates

EAV maybe downregulating the type I IFN response (IFNα and IFNβ) altogether in infected EECs. The type I interferon system is an essential component of the innate immune system providing protection against invading viruses [135; 136]. As such, the

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downregulation of the host gene expression of EAV and EHV-1 infected EECs at the 12 hpi timepoint appears to be a direct consequence of the mechanism taken by some virus families to downregulate the host anti-viral response. Furthermore, the porcine reproductive and respiratory syndrome virus, which is another member of the family

Arteriviridae, is also known to suppress the type I interferon system [136; 337] suggesting that the suppression of the host antiviral response maybe a common mechanism utilized by members of the Arteriviridae family.

When considering the differentially expressed genes and pathways in EAV infected

EECs, at the 6 hpi timepoint, the “CCKR signaling map”, “Inflammation mediated by chemokine and cytokine signaling pathway” and the “Gonadotropin-releasing hormone receptor pathway” were differentially expressed at both 6 and 12 hpi timepoints. In a previous study, we demonstrated that the infection of EECs with EAV induces the expression of proinflammatory cytokine and chemokine mRNA. The upregulation of the

“Inflammation mediated by chemokine and cytokine signaling pathway” at both 6 and 12 hpi timepoints in EAV infected EECs confirms these results and indicates an active role played by proinflammatory cytokines and chemokines during EAV infection of EECs. At the 12 hpi timepoint, the “apoptosis signaling pathway” is differentially expressed in EAV infected EECs when compared to uninfected EECs. Caspase-8 and caspase-9 genes have been shown to be involved in the induction of apoptosis during EAV infections [338]. The transcriptome data of EAV infected EECs obtained during this study shows that Caspase-

8 gene is upregulated confirming previous studies. However, we did not see a differential expression on the caspase-9 gene. In addition to caspase-8, caspase-10 and MAPK8 genes were also upregulated at the 12 hpi timepoint suggesting these two apoptosis promoting

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genes may also play a role in inducing apoptosis in EAV infected EECs [312]. EAV infections in horses results in abortions in pregnant mares and even fatalities due to endothelial cell injury and increased vascular permeability caused by severe necrotizing panvasculitis [339; 340]. As such, vasculitis plays major roles in the pathological outcomes associated with EAV infections. Vasculitis is the inflammation of the walls of the blood vessels which can be influenced by many factors. The transcriptome results at the 6 hpi timepoint for EAV VBA infected EECs suggest the upregulation of the host inflammatory and immune responses when compared to uninfected EECs. Interestingly, the

“Gonadotropin-releasing hormone receptor pathway” was differentially expressed in EAV and EHV-1 infected EECs. Signaling by this pathway is necessary for the normal secretion of the gonadotropin-releasing hormone (GnRH) and the gonadotropins LH and FSH. Even though GnRH, LH and FSH are proinflammatory by nature [341], it is unlikely that activation of the “Gonadotropin-releasing hormone receptor pathway” in endothelial cells will have an effect on inducing the release of these hormones by the pituitary gland.

However, the upregulated genes in this pathway such as MAP3K2, MAP3K5, MAP3K11,

MAP3K14, MAP2K3 and MAPK8 are associated with the mitogen-activated protein kinases (MAPK) signaling cascade which is associated with promoting inflammation and apoptosis [312; 313].

Similar to EAV infected EECs, the “Inflammation mediated by chemokine and cytokine signaling pathway” in EHV-1 infected EECs is differentially expressed at both 6 and 12 hpi timepoints. Vasculitis is one of the important underlying causes in the pathogenesis of these two viruses and pro inflammatory cytokines and chemokines produced by endothelial cells and PBMCs play a key role in inducing vasculitis. The

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upregulation of the “Inflammation mediated by chemokine and cytokine signaling pathway” in both EAV and EHV-1 infected EECs suggest that this pathway plays a key role in promoting vasculitis following infection by these two viruses.

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Table 3.1: List of the top 5 pathways and the differentially expressed genes in EAV 6 hpi vs mock 6 hpi comparison

EAV 6 hpi vs. mock 6 hpi PANTHER pathways Number Genes of genes CCKR signaling map 10 PIK3R1, ITPR1, PRKD1, MEF2D, FOXO1, ODC1, SERPINE1, EGR1, HBEGF, PLA2G4A

Inflammation mediated by 10 ITPR1, GNG10, PREX1, CCL13, RGS17, CCL26, NFKB2, ARPC5,

chemokine and cytokine PLA2G4A, GNG4

signaling pathway

Wnt signaling pathway 10 ITPR1, GNG10, WNT9A, EP400, SRCAP, PPP2R5E, ARID1A, EP300, 111 CREBB, GNG4

EGF receptor signaling 9 CBLB, PRKD1, STAT4, STAT5A, PPP2R5E, PHLDB2, HBEGF, GAB1, pathway SPRY4

Gonadotropin-releasing 9 PIK3R1, ITPR1, IGF1R, BMP2, NR5A1, EGR1, EP300, CREBBP, PLA2G4A hormone receptor pathway

Table 3.2: List of the top 5 pathways and the differentially expressed genes in EAV 12 hpi vs mock 12 hpi

EAV 12 hpi vs. mock 12 hpi PANTHER Number Genes pathways of genes

Gonadotropin- 51 HSPA1A, ACVR2B, FOSB, CREB1, MAPK8, BMP2, IRS2, BMPR2, TGFBR3, GNB3,

releasing hormone ID2, VCL, ATF3, SMAD3, TGIF1, FOS, SP1, MAP3K8, ELK1, PRKAG2, NR4A1, ISL1,

receptor pathway ANXA5, EGR1, MAP3K11, MAP3K14, ITPR2, JUND, SDF4, MAP3K5, PLA2G4A,

STAT3, JUN, JUNB, MAP3K9, SRF, ID3, INHBB, MAP2K3, MAPK14, BMP4, PER1,

PIK3R1, ITPR1, MAP3K2, GATA2, SLC2A1, YBX3, ADIPOR1

PDGF signaling 44 FLI1, RPS6KC1, MAPK8, ELF1, STARD8, MAPKAPK2, ARHGAP5, EPHB2, STAT4, pathway FOS, RAB11B, STAT5A, MTOR, SHC1, ELK1, PDGFB, NINL, ERF, CDK2, OPHN1, NRAS, JAK2, GABPA, MKNK2, MYCBP, RPS6KB2, ITPR2, MYC, GAB1, STAT3,

112 PDGFRA, ARHGAP12, JAK1, JUN, SRF, STAT6, RASA2, MAPK6, VAV2, PIK3R1, ITPR1, MAP3K2, STAT1, DLC1

CCKR signaling 39 CREB1, MAPK8, BCAR1, CXCL1, PTEN, BIRC3, FOS, SP1, FOXO1, SHC1, FOXO3, map ELK1, YES1, NR4A1, CXCL8, ODC1, MCL1, SERPINE1, JAK2, EGR1, IER3, PLAU, MAP3K11, MAP3K14, KLF4, SNAI1, PTGS2, HBEGF, MYC, RGS2, CLU, PLA2G4A, STAT3, JUN, SRF, CXCL2, MAPK14, PIK3R1, ITPR1 Inflammation 36 GNG12, RELB, RAC3, PLCL2, ACTB, GNB3, PTEN, SHC1, KRAS, CXCL8, PLCD1, mediated by NRAS, JAK2, NFKB2, ACTG1, PTGS2, ITPR2, JUND, MYH3, REL, PLA2G4A, chemokine and STAT3, PLCD4, JUN, JUNB, NFKBIE, STAT6, PLA2G4B, CCL3, PLCL1, MYLK3, cytokine signaling ITPR1, STAT1, MYH10, NFKB1, ITGA2 pathway Apoptosis signaling 34 HSPA1A, CREB1, MAPK8, RELB, FADD, EIF2AK2, HSPA8, HSPA6, ATF4, ATF3, pathway BCL2L11, BCL2L2, AIFM1, BIRC3, FOS, PRKRA, HSPA14, TNFRSF10B, TRAF2, MCL1, BIK, BAG3, NFKB2, MAP3K14, TNFRSF1B, REL, MAP3K5, CASP10, APAF1, JUN, CASP8, MAP2K3, NFKB1, HSPA5, TNFRSF10B, TRAF2, MCL1, BIK, BAG3, NFKB2, MAP3K14, TNFRSF1B, REL, MAP3K5, CASP10, APAF1, JUN, CASP8, MAP2K3, NFKB1, HSPA5

Table 3.3: List of the top 5 pathways and the differentially expressed genes in EAV 6 hpi vs EAV 12 hpi (A) and EHV 6 hpi vs mock 6 hpi (B) comparison

A. EAV 6 hpi vs. EAV 12 hpi PANTHER pathways Number Genes of genes CCKR signaling map 8 PIK3R1, ITPR1, FOXO1, ODC1, SERPINE1, EGR1, HBEGF, PLA2G4A EGF receptor signaling 7 CBLB, STAT4, STAT5A, PPP2R5E, PHLDB2, HBEGF, GAB1

pathway

PDGF signaling pathway 6 PIK3R1, ITPR1, DLC1, STAT4, STAT5A, GAB1 Apoptosis signaling pathway 5 HSPA5, HSPA8, HSPA6, NFKB2, TNFRSF1B Gonadotropin-releasing 5 PIK3R1, ITPR1, BMP2, EGR1, PLA2G4A hormone receptor pathway

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B. EHV 6 hpi vs. mock 6 hpi PANTHER pathways Number Genes of genes Gonadotropin-releasing 59 HSPA1A, KSR1, BMP2, IRS2, BMPR2, GNAI3, PRKAG1, ITPR3, ATF3, TGIF1, hormone receptor pathway NR5A1, RAP1B, NR4A1, ISL1, MMP14, EGR1, SMAD9, CREBBP, GUCY1B1, PXN, ESRRA, NFATC2, NAB1, CDC42, JUNB, FST, ID3, BMP4, MAP4K5, ITPR1, SMAD5, GATA2, IGF1R, MAP2K1, TCF7L1, DUSP1, GNB4, ADIPOR1, CTNNB1, MAPK11, GNB5, FOS, SP1, HRAS, PRKAB2, SNRPN, NAB2, PRKCA, PPP3CA, EP300, TUBA1A, SOS1, SDF4, MAP3K5, PLA2G4A, PKNOX1, POU2F1, MAPK14, CAV1

CCKR signaling map 50 MEF2D, CREM, CALM1, BCAR1, TACR1, CPE, ACAT1, BIRC3, FOXO1,

MOB1A, ELK4, NR4A1, ODC1, MCL1, SERPINE1, EGR1, CRK, HBEGF, RGS2, PXN, NFATC2, CDC42, PRKD2, PLCG1, AP2M1, PRKACB, ITPR1, PRKD1, EIF4E, MAP2K1, CTNNB1, SLC18A2, FOS, SP1, PRKACA, YES1, Table 3.3 continued… CXCL8, RHEB, RPS6, HDAC7, PRKCH, PRKCA, PPP3CA, KLF4, SOS1, PLA2G4A, PDPK1, PAK1, MAPK14, TPCN1

B. EHV-1 6 hpi vs. mock 6 hpi PANTHER pathways Number Genes of genes Integrin signaling pathway 48 COL8A1, BCAR1, ARF1, RAP2B, LAMA5, PIK3C3, RND3, LIMS1, RAP1B, ITGB4, COL4A2, ITGAE, NRAS, COL5A2, LAMA4, RAP2C, ARF6, CRK, ACTN4, ARPC3, ARPC1A, COL18A1, PXN, COL2A1, CDC42, RAP2A, RHOC, MAPK6, ASAP1, ABL1, MAP2K1, NTN4, ILK, COL6A2, COL5A1, PIK3C2B, ARL1, LAMB1, ARPC1B, HRAS, ARPC5, SOS1, ITGB3, MAP3K5, COL1A1, ARFGAP1, CAV1, PIK3R2

Inflammation mediated by 47 PREX1, GNAI3, ITPR3, IFNGR2, TYK2, AKT3, CAMK2D, MYH9, RGS17, chemokine and cytokine NRAS, IL10RB, RRAS2, REL, GNG4, ARPC3, ARPC1A, NFATC2, CDC42, 114 signaling pathway IL15, JUNB, RHOC, PLCG1, PRKACB, CHUK, SOCS5, ITPR1, GNG10, STAT1,

MYH10, COL6A2, CYTH2, CCL13, RGS4, PRKACA, CXCL8, ARPC1B, CCL26, NFKB2, ARPC5, IL6, GNG5, SOS1, VWF, PLA2G4A, STAT6, PDPK1, PAK1

Wnt signaling pathway 46 PPP2R5A, SMARCC1, SMARCA5, DVL3, ITPR3, HDAC3, SMARCE1, CDH5, SMARCD2, ADSS, FBXW11, LRP5, ARID1B, SMAD9, CREBBP, TLE2, GNG4, NFATC2, APC, PPP3CB, FAT1, ITPR1, SMAD5, GNG10, FZD6, PPP2CB, BCL9, TCF7L1, GNB4, CTNNB1, PPP3CC, EP400, SRCAP, SMARCC2, B4GALT7, PPP2R5E, ARID1A, PRKCH, TCF7L2, PRKCA, PPP3CA, EP300, GNG5, DCHS1, SMARCA4, CTNNAL1

Table 3.4: List of the top 5 pathways and the differentially expressed genes in EHV 12 hpi vs mock 12 hpi (A) and EHV 6 hpi vs EHV 12 hpi (B) comparison.

A. EHV-1 12 hpi vs. mock 12 hpi PANTHER Number of Genes pathways genes Integrin signaling 47 FLNB, ITGA5, PIK3R2, COL8A1, DOCK1, COL6A2, MAP3K3, COL5A1, ACTB, pathway COL14A1, ITGA2, LAMA5, RND1, RND3, PIK3C2B, RHOA, RAP1B, ITGB4, COL16A1, COL4A2, LAMB2, ITGB5, LAMB1, HRAS, LAMC2, COL12A1, COL5A2, ARPC5, ITGAX, ARF6, PARVA, SOS1, COL6A1, ACTN4, ITGB3, TLN1, MAP3K5, COL18A1, PXN, COL1A1, LAMC1, COL1A2, RHOB, MAPK6, PTK2B, ASAP1, ITGA11

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Gonadotropin- 46 BMPR1A, ACVR2B, FOSB, GATA2, NCOA3, IGF1R, SLC2A1, KSR1, IRS2, releasing hormone MAP3K3, DUSP1, GNAI2, BMPR2, TGFBR3, ID2, ATF3, MAPK11, FOS, SP1, receptor pathway RAP1B, NR4A1, HRAS, MAP4K2, MMP14, MAP2K7, ANXA5, EGR1, PBX1, SMAD9, PRKCA, EP300, CREBBP, JUND, SOS1, EIF2A, MAP3K5, PXN, GNB2, JUNB, MAP3K9, SRF, FST, POU2F1, PTK2B, EIF2AK3, BMP4

Inflammation 44 GNG12, GNG10, CCL5, RELB, STAT1, NFKBIA, MYH10, COL6A2, GNAI2, ACTB, mediated by COL14A1, ITGA2, IFNGR2, TYK2, NFAT5, AKT3, CCL13, RHOA, INPPL1, chemokine and PRKACA, MYH9, CXCL8, PLCD1, COL12A1, RGS17, JAK2, PAK2, NFKB2, cytokine signaling ARPC5, IL6, RRAS2, PTGS2, JUND, SOS1, VWF, COL6A1, CCL2, CCL20, REL, pathway ROCK1, JUNB, NFKBIE, PDPK1, PTK2B

CCKR signaling 37 PRKD1, MEF2D, CREM, HSPB1, NFKBIA, BIRC2, BIRC3, RHOA, FOS, SP1, map FOXO1, FOXO3, PRKACA, TPST1, NR4A1, CXCL8, ODC1, MCL1, SERPINE1, JAK2, RPS6, EGR1, IER3, HDAC7, PRKCA, KLF4, SNAI1, PTGS2, HBEGF, SOS1, CLU, ROCK1, PXN, SRF, PRKD2, PDPK1, PTK2B

Angiogenesis 35 PIK3R2, PRKD1, HSPB1, EFNB2, SPHK2, STAT1, PRR5, DVL3, NOTCH1, signaling pathway TGFB1I1, PIK3C2B, AKT3, RHOA, FOS, BIRC5, JAG1, HRAS, PDGFRB, ARHGAP1, PAK2, PRKCA, CRYAB, F3, SOS1, PXN, PDGFRA, PTPRB, PRKD2, APC, NOS3, VEGFA, NOTCH2, RHOB, MAPK6, JAG2

Table 3.4 continued…

B. EHV-1 6 hpi vs. EHV 12 hpi

PANTHER pathways Number Genes of genes Gonadotropin-releasing 26 GATA2, IGF1R, KSR1, IRS2, DUSP1, BMPR2, ATF3, MAPK11, FOS, SP1, hormone receptor RAP1B, NR4A1, HRAS, MMP14, EGR1, SMAD9, PRKCA, EP300, CREBBP,

116 pathway SOS1, MAP3K5, PXN, JUNB, FST, POU2F1, BMP4

Integrin signaling 24 PIK3R2, COL8A1, COL6A2, COL5A1, LAMA5, RND3, PIK3C2B, RAP1B, ITGB4, pathway COL4A2, LAMB1, HRAS, COL5A2, ARPC5, ARF6, SOS1, ACTN4, ITGB3, MAP3K5, COL18A1, PXN, COL1A1, MAPK6, ASAP1

CCKR signaling map 22 PRKD1, MEF2D, CREM, BIRC3, FOS, SP1, FOXO1, PRKACA, NR4A1, CXCL8, ODC1, MCL1, SERPINE1, RPS6, EGR1, HDAC7, PRKCA, KLF4, HBEGF, SOS1, PXN, PDPK1

Inflammation mediated 21 GNG10, STAT1, MYH10, COL6A2, IFNGR2, TYK2, AKT3, CCL13, PRKACA, by chemokine and MYH9, CXCL8, RGS17, NFKB2, ARPC5, IL6, RRAS2, SOS1, VWF, REL, JUNB, cytokine signaling PDPK1 pathway

Table 3.5: List of the top 5 pathways and the differentially expressed genes in EHV 6 hpi vs EAV 6 hpi (A) and EHV 12 hpi vs EAV 12 hpi (B) comparison.

A. EHV-1 6 hpi vs. EAV 6 hpi PANTHER pathways Number Genes of genes Inflammation mediated by 10 ITPR1, GNG10, PREX1, CCL13, RGS17, CCL26, NFKB2, ARPC5, PLA2G4A, GNG4 chemokine and cytokine signaling pathway CCKR signaling map 9 ITPR1, PRKD1, MEF2D, FOXO1, ODC1, SERPINE1, EGR1, HBEGF, PLA2G4A

Wnt signaling pathway 9 ITPR1, GNG10, EP400, SRCAP, PPP2R5E, ARID1A, EP300, CREBBP, GNG4

117 EGF receptor pathway 8 CBLB, PRKD1, STAT4, STAT5A, PPP2R5E, HBEGF, GAB1, SPRY4

Gonadotropin-releasing 8 ITPR1, IGF1R, BMP2, NR5A1, EGR1, EP300, CREBBP, PLA2G4A hormone receptor pathway

B. EHV-1 12 hpi vs. EAV 12 hpi PANTHER pathways Number Genes of genes Gonadotropin-releasing hormone 21 ACVR2B, FOSB, GATA2, SLC2A1, IRS2, DUSP1, BMPR2, TGFBR3, ID2, receptor pathway ATF3, FOS, SP1, NR4A1, ANXA5, EGR1, JUND, MAP3K5, JUNB, MAP3K9, SRF, BMP4 CCKR signaling map 19 BIRC3, FOS, SP1, FOXO1, FOXO3, NR4A1, CXCL8, ODC1, MCL1, SERPINE1, JAK2, EGR1, IER3, KLF4, SNAI1, PTGS2, HBEGF, CLU, SRF

Apoptosis signaling pathway 15 RELB, EIF2AK2, HSPA8, HSPA6, ATF4, ATF3, BCL2L11, BIRC3, FOS, HSPA14, MCL1, BIK, NFKB2, REL, MAP3K5

Inflammation mediated by 15 GNG12, RELB, STAT1, MYH10, ACTB, ITGA2, CXCL8, PLCD1, JAK2,

118 chemokine and cytokine signaling NFKB2, PTGS2, JUND, REL, JUNB, NFKBIE

pathway Integrin signaling pathway 14 COL5A1, ACTB, ITGA2, RND1, RND3, COL4A2, LAMB2, COL5A2, ITGAX, ITGB3, MAP3K5, COL18A1, LAMC1, MAPK6

Figure 3.1: Heatmap of the gene expression of EHV-1, EAV and mock infected EECs.

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Figure 3.2: Principal components analysis of genes of EHV-1, EAV and mock infected EECs at both 6 and 12 hpi timepoints. The clustering of the genes of EAV infected EECs at the 6 hpi timepoint shows significant variance. No such variance observed in the clustering of EHV-1 or mock infected EECs.

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- Upregulated genes - Downregulated genes - Upregulated genes - Downregulated genes

Upregulated Downregulated

Figure 3.3: Differential gene expression of EHV-1 infected EECs when compared with mock infected EECs at the 6 (A) and 12 (B) hpi timepoints and when compared among 6 and 12 hpi timepoints (C). Differential gene expression of EAV infected EECs when compared with mock infected EECs at the 6 (D) and 12 (E) hpi timepoints and when compared among 6 and 12 hpi timepoints (F).

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Figure 3.4: Venn diagram illustrating the distribution of differentially expressed genes in EECs infected with; EAV at 6 and 12 hpi timepoints (A), EHV-1 at 6 and 12 hpi timepoints (B), EAV and EHV-1 at the 6 hpi timepoint (C) and EAV and EHV-1 at the 12 hpi timepoint (D).

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CHAPTER FOUR

INVOLVEMENT OF TOLL-LIKE RECEPTORS IN THE INNATE IMMUNE RESPONSE OF EQUINE ENDOTHELIAL CELLS FOLLOWING EHV-1 AND EAV INFECTIONS

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4.1. SUMMARY

Equid herpesvirus–1 (EHV-1) is a double-stranded DNA virus whereas equine arteritis virus (EAV) is a positive sense, single stranded RNA virus. Both these viruses have a worldwide distribution and collectively, are a significant threat to the global and US equine industry. One of the most important pathological conditions underlying the disease manifestations associated with EAV and EHV-1 infections is vasculitis. The vasculitis induced by EHV-1 is restricted to either the vasculature of the pregnant uterus or the central nervous system while in the case of EAV it can be systemic. In the 2nd chapter proinflammatory cytokine and chemokine response of EHV-1 and EAV infected EECs were investigated and the expression of proinflammatory cytokine and chemokine genes was found to be higher in EHV-1 infected EECs compared to EAV infected EECs, furthermore, EAV downregulates the expression of IFNα in EECs following infection suggesting that toll-like receptors (TLRs) are involved in the induction of the proinflammatory response in the infected EECs. Here we report an active role played by

TLR 3 in inducing the proinflammatory response in EHV-1 infected EECs at early stages and also that TLR 3 is not the main pattern recognition receptor in inducing a proinflammatory response in EAV infected EECs.

Keywords: EHV-1, EECs, TLRs, TLR3, Horses, siRNA, Electrophoresis

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4.2. INTRODUCTION

Equid herpesvirus–1 (EHV-1) is a DNA viruses of the subfamily

Alphaherpesvirinae and genus Varicellovirus [28]. Equine arteritis virus (EAV) is a positive sense, single stranded RNA virus belonging to the order Arteriviridae and genus

Alphaarterivirus (https://talk.ictvonline.org/taxonomy/)[290]. Both these viruses have a worldwide distribution [5; 6] and collectively, are a significant threat to the global and US equine industry. These two genetically very different viruses share some interesting similarities in their disease manifestation and clinical outcomes (establishment of latent or persistent infections[5; 6; 342], respiratory and/or reproductive tract diseases[5; 6; 15; 17-

19; 27; 34; 38; 88; 89; 169] and abortions in pregnant mares[5; 6; 17; 19; 27; 38; 88; 89;

166; 198; 343; 344]). One of the most important pathological conditions underlying the disease manifestations associated with EAV and EHV-1 infections is viral vasculitis [5; 6;

17; 19; 52; 170]. In the case of EHV-1 infections, the viral vasculitis is restricted to either the vasculature of the pregnant uterus or the central nervous system (CNS) resulting in abortions and a neurological condition termed equine herpesvirus myeloencephalopathy

(EHM) [15; 17; 18; 52; 170; 345]. Viral vasculitis induced by EAV infection affects the pregnant uterus which can result in abortion [5; 6; 19; 88]. However, it can also become systemic (Systemic panvasculitis) leading to fatalities [5; 6; 19; 89; 261]. Yet, in contrast to EHV-1 infections, EAV infections do not induce any form of neurological disease.

In a previous study, we investigated the cytokine response of EECs following infection with EHV-1, EHV-4 and EAV. The results of that study indicated that both EHV-

1 and EAV infected EECs upregulate the gene expression of proinflammatory cytokines

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and chemokines. However, the expression levels of these genes were relatively higher in

EHV-1 infected EECs when compared with EAV infected EECs (Unpublished data).

Following these results, whole transcriptome sequencing of EHV-1 and EAV infected

EECs was conducted. The results indicated the differential expression of the “Inflammation mediated by cytokine and chemokine” pathway as well as other pathways associated with the host immune, inflammatory and apoptotic responses in EECs following infection by

EHV-1 and EAV.

Inflammation is a coordinated cellular response evolved over time that can result in positive or sometimes negative effects for the host [227]. The cellular inflammatory response is advantageous to the host during infections or cellular damage but becomes disadvantageous when directed at a wrong stimulus or when reacting in excess [227]. The immune system consists of two arms, the innate and the adaptive immune systems [226;

229]. The innate immune system is the first line of defense against invading pathogens and it senses the presence of invaders through pathogen associated molecular patterns (PAMPs) or damage associated molecular patterns (DAMPs) [226; 229]. Host cells use a specialized group of structurally diverse cell surface and intra-cellular receptors known as pattern recognition receptors (PRRs) to sense these PAMPs or DAMPs and one such extensively studied group of PRRs are the Toll-like receptors (TLRs) [229]. TLRs sense PAMPs and

DAMPs and initiate an immune and inflammatory response via proinflammatory cytokines and chemokines. Virus replication in endothelial cells causes cell injury and virus infections that damage blood vessels are known to induce vasculitis [346]. Upon sensing

PAMPs and DAMPs, the innate immune system triggers an inflammatory response and this is mediated by proinflammatory cytokines and chemokines [226]. As such, TLRs are

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important in the initiation of the host inflammatory response and to this point, no investigation has been carried out to elucidate the TLRs involved with the inflammatory response induced by EHV-1 and EAV in EECs. The purpose of this study was to elucidate the involvement of TLRs in the immune and inflammatory response against EHV-1 and

EAV infections in EECs.

4.3 MATERIALS AND METHODS

4.3.1 Transcriptome data analysis

For RNA-seq data analysis, the reads were initially trimmed for adapters and quality using TrimGalore Version 0.4.4 (Babraham Bioinformatics; www.bioinformatics.babraham.ac.uk), then trimmed reads were aligned to the horse reference genome (EquCab3.0) using STAR (Release 2.5.2b) [32], and annotated to the equine reference transcriptome available in NCBI database using Cufflinks (Release 2.2.1; http://cole-trapnell-lab.github.io/cufflinks/) [33]. Differential expression analysis was performed on the normalized read count (FPKM; fragments per kilobase of exon per million mapped reads) [34]. We used Cuffdiff 2.2.1 (Trapnell, Roberts et al. 2012) to calculate differentially expressed genes (DEG) between samples from mock and virus infected groups. Significance level was set at FDR-adjusted p value of the test statistic <

0.1 using a Benjamini- Hochberg correction. For all data analyses, separate databases were created for EHV and EAV samples. Differentially expressed genes were determined within tissue. Initial identification of differentially expressed genes was performed using one- way ANOVA, using the Benjamini-Hochberg correction for false discovery rate (FDR P <

0.1).

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4.3.2 Cells

Equine endothelial cells (EECs) were derived from the pulmonary artery of a horse as previously described [264]. The EECs were cultured in Dulbecco’s Modified Eagle’s

Medium (DMEM, Corning, New York, NY) with 10% bovine calf serum (BCS, Hyclone

Laboratories, Inc., Logan, UT), 100 U/mL penicillin–streptomycin, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids and 200 mM L-glutamine (Thermo Fisher

Scientific, Waltham, MA) in a humidified incubator at 37 °C with 5% CO2. All experiments were performed in EECs between passage numbers 15-20.

4.3.3 Viruses

Two viruses were used in this study. The horse adapted virulent Bucyrus strain of

EAV (VBS; VR796; ATCC) and the neuropathogenic EHV-1 (Findlay) strain [14]. The

EHV-1 virus was kindly provided by the late Dr. George Allen, Maxwell H. Gluck Equine

Research Center, University of Kentucky, Lexington, Kentucky. All viruses were propagated in EECs to generate high titer working stocks. Briefly, triple-decker cell culture flasks (Nunc, Rochester, NY) containing EEC monolayers were infected with each of the four viruses. The flasks were frozen at -80°C when a 100% cytopathic effect (CPE) was observed. Cell lysates were clarified by centrifugation (2000 x g) at 4 °C for 15 min, followed by ultracentrifugation (Beckman Coulter, Miami, FL) at 121,600 x g for EAV and 100,000 x g for EHVs through a 20% sucrose cushion in NET buffer (150 mM NaCl,

5 mM EDTA, 50 mM Tris-HCl, pH 7.5) at 4°C for 4 h to pellet the virus. Purified preparations of each virus strain were resuspended in Dulbecco’s phosphate buffered saline

(DPBS, Thermo Fisher Scientific, Waltham, MA), briefly sonicated and 50 µL aliquots were frozen at -80 °C until further use. Virus stocks were titrated by standard plaque assay

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in EECs, and titers were expressed as the number of plaque forming units (PFU) per milliliter (PFU/mL).

4.3.4 siRNA transfection

EECs (Passage 15-20) were used for TLR3 siRNA transfection. In brief, 1×105 cells were transfected with either 0.5 pmol of TLR3 siRNA or negative control using the Lonza

4D nucleofector X unit system (Basel, Switzerland) following the manufacturers protocol.

The sequence of the TLR3 siRNA is; sense: 5’ AGAUCAAACUCUCAAGUUUtt 3’ antisense: 5’ AAACUUGAGAGUUUGAUCUtg 3’ (Silencer™ Select, Thermo Fisher

Scientific, Waltham, MA). The negative control siRNA used was a commercially available product (MISSION® siRNA Universal Negative Control, Sigma-Aldrich, St. Louis, MO).

4.3.5 Virus infection, RNA extraction and cDNA generation

siRNA transfected EECs (Passage 15-20) were seeded in 24 well plates (1×105 cells per well) and incubated for 24 hours. At 24 hours post transfection (hpt) timepoint, the cells were infected with either EHV-1 or EAV virus strains at a MOI of 2 and incubated for 6 and 12 hours. Following 6 and 12 hours of infection with virus strains total RNA was extracted using the miRNeasy mini kit (Qiagen, Hilden, Germany). Any contaminating genomic DNA present was removed by performing DNase I digestion (RNase free DNase set, Qiagen, Hilden, Germany). The total RNA quality and quantity was assessed by

OD260/OD280 measurement using a NanoDrop (Thermo Scientific, Wilmington, DE). 0.5 micrograms of total RNA were reverse transcribed to generate cDNA using the High capacity reverse transcription kit (Applied Biosystems, Foster city, CA).

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4.3.6 RT-qPCR:

Real-Time PCR amplification was carried out using 1:1 diluted cDNA to assess the relative expression of different pro-inflammatory cytokine and chemokine genes using commercially available TaqMan™ primers and probes (Thermo Fisher Scientific,

Waltham, MA) in a 7500 Fast Real-Time PCR system (Thermo Fisher Scientific, Waltham,

MA). The panel of TLR genes investigated in this study were TLRs 1, 2, 3, 4, 7, 8 and 9.

The proinflammatory cytokine and chemokine panel investigated was IL6, IL17, CXCL10 and IFNα. All reactions were performed in triplicate. EECs grown in identical conditions absent virus infection was used as the mock control. Equine β- glucuronidase gene was used as the endogenous control to normalize the quantitative RT-qPCR data as has been described in previous studies [64]. PCR efficiency for all reactions was assessed using

LinRegPCR [271]. For relative quantification, fold changes in gene expression were calculated using the ΔΔCT method [272].

4.3.7. Statistical analysis

The statistical analysis of RT-qPCR analysis was performed using the means of fold changes employing t tests. The significant level (p-value) was set at 0.05 for all tests.

Statistical analysis was performed using the JMP pro 13 software (JMP®, Version 13 SAS

Institute Inc., Cary, NC, 1989-2019).

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4.4. RESULTS

4.4.1 Transcriptome profile of TLR expression

As mentioned in chapter 3, the RNA-seq data obtained for this study came from a total of 18 libraries sequenced from EECs. Transcriptome data obtained from EHV-1, EAV and mock infected EEC libraries was analyzed for TLR expression. Transcriptome data from EHV-1 infected EECs showed that at the 6 hpi timepoint there was no detectable expression of TLRs 2 and 4 in either EHV-1 or mock infected EECs (Figure 3.1A). TLRs

1, 3, 7, 8 and 9 were expressed in both EHV-1 and mock infected EECs but did not show any significant differences from each other (Figure 3.1A). Only the expression of TLR 3 in EHV-1 infected EECs was statistically significantly increased from mock infected EECs at the 6 hpi timepoint (Figure 3.1A). Similar to the 6 hpi timepoint, the expression of TLRs

2 and 4 was not detected in EHV-1 infected or mock infected EECs at the 12 hpi timepoint

(Figure 3.1B). The expression of the TLRs 1, 3, 8, and 9 was detected in EHV-1 infected

EECs, but no significant differences from mock was observed (Figure 3.1B). The expression of TLRs 7 and 9 was not detected in EHV-1 infected EECs but was detected in mock infected EECs at the 12 hpi timepoint, but this downregulation in expression was not statistically significant (Figure 3.1B). similar to findings at the 6 hpi timepoint, TLR 3 expression was detected in EHV-1 infected EECs at the 12 hpi timepoint and it was significantly higher when compared with mock infected EECs.

In EAV infected EECs at the 6 hpi timepoint, transcriptome data revealed that TLRs

2 and 4 was not expressed either in EAV or mock infected EECs (Figure 3.2A). Moreover,

TLRs 1, 3, 7, 8 and 9 were expressed in both EAV and mock infected EECs but did not show any significant differences from each other (Figure 3.2A). Only TLR 3 expression in

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EAV infected EECs was significantly increased from mock infected EECs at the 6 hpi timepoint (Figure 3.2A).

Similarly, at the 12 hpi timepoint, TLRs 2 and 4 were not expressed in either EAV or mock infected EECs (Figure 3.2B). The expression of the TLRs 1, 3, 7, 8 and 9 were expressed in both EAV and mock infected EECs, neither differed significantly from each other (Figure 3.2B). Only TLR 3 expression in EAV infected EECs significantly increased from mock infected EECs at the 6 hpi timepoint (Figure 3.2B).

4.4.2 RT-qPCR validation of transcriptome results

In EHV-1 infected EECs gene expression was detected for TLRs 1, 2, 3, 4, 7, 8 and

9 at both 6 and 12 hpi timepoints (Figure 3.3A). The gene expression of the TLRs 1, 2, 4,

8 and 9 did not show a statistically significant difference at both 6 and 12 hpi timepoints when compared with mock infected EECs (Figure 3.3A). The gene expression level of TLR

7 was significantly upregulated in EHV-1 infected EECs at the 6 hpi timepoint but was not significant at the 12 hpi timepoint (Figure 3.3A). The gene expression level of TLR 3 was significantly upregulated at both 6 and 12 hpi timepoints in EHV-1 infected EECs compared to the mock infected EECs (Figure 3.3A). Furthermore, gene expression significantly increased from 6 to 12 hpi timepoints suggesting EHV-1 infection of EECs promote the expression of the TLR 3 gene (Figure 3.3A). Furthermore, the RT-qPCR data validates the transcriptome results for TLR 3 gene expression in EHV-1 infected EECs.

Similar to EHV-1 infected EECs, gene expression was detected for TLRs 1, 2, 3,

4, 7, 8 and 9 at both 6 and 12 hpi timepoints in EAV infected EECs (Figure 3.3B). The gene expression of the TLRs 1, 2, 3, 4, and 8 was not significantly different at both 6 and

12 hpi timepoints when compared with mock infected EECs (Figure 3.3B). The gene

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expression of TLR 7 was significantly upregulated in EAV infected EECs at both 6 and 12 hpi timepoints (Figure 3.3B). However, the gene expression level was lower at the 12 hpi timepoint compared to the 6 hpi timepoint. The gene expression of TLR 9 was significantly upregulated in EAV infected EECs at both 6 and 12 hpi timepoints (Figure 3.3B). These

RT-qPCR results suggest that TLRs 7 and 9 are differentially expressed in EAV infected

EECs. Furthermore, the data does not confirm the transcriptome results for TLR 3 expression in EAV infected EECs.

4.4.3 Involvement of TLR 3 in the innate immune response against EHV-1 and EAV infections

In order to investigate the involvement of TLR 3 in promoting the inflammatory response in EHV-1 and EAV infected EECs, we used siRNA sequences generated against equine TLR 3 mRNA. The downregulation of equine TLR 3 and the proinflammatory cytokine (IL6, IL17, CXCL10 and IFNα) mRNA was assessed using RT-qPCR.

The siRNA sequence generated against equine TLR 3 downregulated equine TLR

3 mRNA expression significantly at the 6 hpi timepoint (Figure 3.4A). Furthermore, parallel to the downregulation of TLR 3 mRNA, the IFNα mRNA level was significantly downregulated compared to mock infected EECs. This suggests that TLR 3 plays a role in

IFNα production in EHV-1 infected EECs at the 6 hpi timepoint (Figure 3.4A). However, the gene expression levels of IL6, IL17 and CXCL10 was not significantly downregulated, suggesting the expression of these proinflammatory cytokines and chemokines are not induced by TLR 3 (Figure 3.4A). Similarly, at the 12 hpi timepoint, the siRNA sequence generated against equine TLR 3 downregulated equine TLR 3 mRNA expression significantly (Figure 4.4B). I n contrast to the results obtained at the 6 hpi timepoint

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however, none of the proinflammatory cytokines and chemokines used in the experiment was significantly downregulated, suggesting that at the 12 hpi timepoint the importance of

TLR 3 in inducing an inflammatory response in EHV-1 infected EECs is reduced (Figure

3.4B).

The same experiment was performed using EAV infected EECs to investigate the involvement of TLR 3 in inducing an inflammatory response in infected cells at both 6 and

12 hpi timepoints. Similar to the observations in EHV-1 infected EECs, the siRNA sequence generated against equine TLR 3 downregulated equine TLR 3 mRNA expression significantly in EAV infected EECs at both 6 and 12 hpi timepoints (Figure 3.5 A&B).

However, the downregulation of TLR 3 mRNA did not induce the downregulation of any of the proinflammatory cytokines and chemokines investigated in this study, suggesting that TLR 3 is of lesser importance in inducing an inflammatory response in EAV infected

EECs compared to EHV-1 infected EECs (Figure 3.4B).

4.5 DISCUSSION

Eleven known TLRs have been identified in mammalian cells and the expression of these TLRs are cell type dependent [347]. To date, studies have reported the expression of TLR 4 and 2 in equine lungs and bronchial epithelial cells [236; 239], TLR 9 in equine lungs, spleen, lymph nodes, and peripheral blood leukocytes [234; 238] and TLRs 2, 3, 4,

6 and 9 in equine cornea, limbus, and the conjunctiva [237]. Here, we report the expression of TLRs 1, 2, 3, 4, 7, 8 and 9 in equine endothelial cells. The transcriptome results of this study indicated gene expression of TLRs 1, 3, 7, 8 and 9 in EECs but did not detect any gene expression for TLRs 2 and 4. However, RT-qPCR results indicated gene expression

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for TLRs 2 and 4 as well, indicating that EECs express mRNA for at least TLRs 1, 2, 3, 4,

7, 8 and 9. This is the first study to investigate the expression of TLRs in equine pulmonary artery endothelial cells. Furthermore, in the transcriptome profiles of infected and uninfected EECs, no expression was detected for TLRs 2, 4 and 5. However, RT-qPCR results indicated the presence of gene expression for these TLRs as well. This could be due to the higher sensitivity of RT-qPCR in detecting gene expression with low copy numbers compared to Next-generation sequencing and microarrays.

The transcriptome and RT-qPCR results of EHV-1 infected EECs indicate the upregulation of TLR 3 gene expression at both 6 and 12 hpi timepoints. Furthermore, the downregulation of TLR 3 mRNA using siRNA induced a downregulation of IFNα mRNA at the 6 hpi timepoint in EHV-1 infected EECs suggesting TLR 3 has a role to play in induction of IFNα during EHV-1 infection of EECs. TLR 3, together with TLRs 7, 8 and

9 are located intracellularly in endosomal membranes and they sense nucleic acids of invading pathogens [348; 349]. TLR 3 specifically recognizes double-stranded RNA which is an intermediate produced during the replication of most viruses including herpesviruses

[348]. Several studies have shown the importance of TLR 3 in sensing infections by herpes simplex type 1 virus [234; 350]. Therefore, this study indicates that just as it is in herpes simplex viruses, TLR 3 is important for the sensing of EHV-1 and for the induction of

IFNα in EECs at the early stages of infection. This is the first study to demonstrate the importance of TLR 3 in the infection of EECs by EHV-1 strains. However, at the 12 hpi timepoint, the downregulation of TLR 3 mRNA did not induce IFNα mRNA downregulation indicating that the importance of TLR 3 in inducing IFNα during the initial stages of infection has become redundant. Recent studies have demonstrated that the capsid

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of DNA viruses such as HSV and adenoviruses to become ubiquitinated and degraded in the cytoplasm by proteasomal pathways [351; 352]. This proteasomal degradation of the capsid maybe a mechanism utilized by DNA viruses which replicate in the cytoplasm of host cells to avoid capsid sensing as well as to release the viral genomic DNA into the cytoplasm [353], what it also does is expose the viral DNA to innate DNA sensors such as

DAI, RNA polymerase III, RIG-I and STING [353-358]. These cytosolic DNA sensors are capable of inducing a type I IFN response when exposed to DNA in the cytoplasm [353;

355-357]. This mechanism may be able to compensate for the downregulation in TLR 3 and induce the production of IFNα.

One thing that needs to be mentioned here is the downregulation of TLR3 mRNA brought about by siRNA is relatively low when compared with other such RNA silencing experiments [359; 360]. This could be because not all cells get transfected with the anti-

TLR 3 siRNA during the siRNA transfection process. Transfection with fluorescently labelled siRNA followed by cell sorting will aid in acquiring a uniformly transfected cell population with a relatively higher downregulation of TLR 3 mRNA and increase the sensitivity of the experiment.

It is well established that up or downregulation of mRNA doesn’t always reflect in protein translation and the subsequent expression of a protein. As such, TLR 3 mRNA downregulation demonstrated by RT-qPCR alone is not sufficient to confirm the downregulation of TLR 3 protein expression in EECs. In most experiments this confirmation of protein downregulation can be made with the use of western blots.

However, it proved a difficult task in this situation as we ran into difficulty finding a monoclonal or a polyclonal anti-equine TLR 3 antibody.

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The downregulation of TLR 3 mRNA in EAV infected EECs did not reduce the expression of any of the cytokine and chemokine genes analyzed during this study indicating that the induction of the proinflammatory response in EAV infected EECs may not be coming from TLR 3 but from a different TLR.

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Figure 4.1: Transcriptome results (FPKM values) of TLR expression in EHV-1 and mock infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. * indicates a statistically significant increase from mock infected EECs (p≤0.05).

Figure 4.2: Transcriptome results (FPKM values) of TLR expression in EAV and mock infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. * indicates a statistically significant increase from mock infected EECs (p≤0.05).

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Figure 4.3: Relative quantification values of TLR in EHV-1 (A) and EAV infected EECs. The red line marks the basal level of expression (RQ=1). * indicates a statistically significant increase from mock infected EECs (p≤0.05).

A 4 B 6 hpi 1.4 12 hpi 3.5 1.2 3 * 1 2.5 0.8 2

0.6 RQ values RQ 1.5 RQ values 1 0.4 * * 0.5 0.2

0 0 IL6 IL17 CXCL10 IFNa TLR3 IL6 IL17 CXCL10 IFNa TLR3 Groups Groups

Figure 4.4: Relative quantification values of TLR expression in TLR 3 siRNA transfected and EHV-1 infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. The red line marks the basal level of expression (RQ=1). * indicates a statistically significant decrease in gene expression from negative control transfected EECs (p≤0.05).

2.5 35 A 6 hpi B 12 hpi

30 2 25

1.5 20

1 RQvalues values RQ * * 10

0.5 5 * 0 0 IL6 IL17 CXCL10 IFNa TLR3 IL6 IL17 CXCL10 IFNa TLR3 Groups Groups

Figure 4.5: Relative quantification values of toll-like receptor expression in TLR 3 siRNA transfected and EAV infected EECs at 6 hpi (A) and 12 hpi (B) timepoints. The red line marks the basal level of expression (RQ=1). * indicates a statistically significant decrease in gene expression from negative control transfected EECs (p≤0.05).

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CHAPTER FIVE

GENOME-WIDE ASSOCIATION STUDY (GWAS) TO IDENTIFY HOST

GENETIC FACTORS ASSOCIATED WITH EQUID HERPESVIRUS TYPE-1

(EHV-1) INDUCED MYELOENCEPHALOPATHY (EHM) IN HORSES

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5.1. SUMMARY

Equid herpesvirus (EHV-1) infections in horses can lead to equine herpesvirus myeloencephalopathy (EHM), characterized by neurological clinical signs. We investigated the importance of the host genetic background in the development EHM using a genome-wide association study (GWAS). Genomic DNA from 28 horses with neurologic signs due to EHV-1 infection (EHM) and 67 control horses without any neurologic signs but infected with EHV-1 were included in this study. DNA samples were tested with the

Affymetrix Axiom 670K SNP array to identify single nucleotide polymorphisms (SNPs) associated with EHM. Results from this study did not identify significant SNPs or haplotype associations for the development of EHM following EHV-1 infections and did not corroborate with a previously published study. The results exclude the involvement of a recessive genetic factor in the susceptibility to develop EHM but do not have the power to exclude the involvement of other, complex host genetic factors.

Key words: EHV-1, EHM, GWAS, Horses, SNPs

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5.2. INTRODUCTION

Equid herpesvirus 1 (EHV-1) is a double-stranded DNA virus in the family

Herpesviridaein the genus Varicellovirus [28]. EHV-1 has a global distribution and at some time during their lives almost all domesticated horses will become infected with the virus resulting in a wide spectrum of clinical signs including an acute upper respiratory tract disease, abortion, neonatal death, retinopathy and severe neurological disease (also known as equine herpesvirus myeloencephalopathy [EHM]) [18; 172]. This neurologic form has a 15-50% case-fatality rate and poses a significant threat to the $102 billion equine industry in the United States [48; 49]. Unfortunately, the occurrence of EHM has increased dramatically over the past fifteen years, with extensive outbreaks reported in the United

States, South Africa, New Zealand and several European nations, including the United

Kingdom. Susceptibility to EHM appears to be determined by several host factors including age, physical condition, immune status and whether the infection is a primary exposure [48; 50]. It has also been reported that the neuropathogenic phenotype of EHV-1 can result from a single nucleotide substitution within ORF30 (A→G2254 [amino acid

N→D752] ) in the viral DNA polymerase [51]. However, data recently generated in our laboratory and other laboratories have demonstrated this interpretation is probably overly simplistic and the development of EHM cannot be solely explained by studying the viral determinants of virulence [361]. An experimental infection study with a neuropathogenic

EHV-1 strain (Ab4) resulted in 0-35% EHM cases [185]; similar rates have been reported for natural outbreaks in the US and Europe (3-45%) [362]. These observations suggest that host genetics may play an important part in the development of EHM. Thus, the rationale for this research is to understand the host genetic factors conferring the risk of developing

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EHM following infection with EHV-1. A recent genome-wide association study (GWAS) study suggested an association of a single nucleotide polymorphism

5.3. MATERIALS AND METHODS

5.3.1 Animals:

We tested 95 DNA samples from horses that had EHM (n=28) and horses that were infected with EHV-1 but did not develop EHM (n=67). The DNA was extracted from nasal swabs or whole blood samples that were received fresh or had been archived at -80°C.

These samples were obtained from several EHV-1 outbreaks in the US during the time period of 2013-2018.

5.3.2 Clinical Characterization:

The case definition of EHM for the selection of these samples involved horses that had clinical signs of neurologic disease consistent with EHM. The neurological signs included dysmetria, ataxia and paresis of limbs, toe dragging, hypotonia of the tail and recumbency. Licensed veterinarians performed the clinical diagnosis of EHM. The horses included in the control group showed signs of EHV-1 infection (fever, swollen legs and lymph nodes, nasal discharge and abortions) without any neurological signs.

5.3.3 Genome Wide Association Studies:

Genomic DNA was extracted from samples using the QIAamp® DNA blood mini kit (Qiagen, Hilden, Germany) following the manufacturer’s instructions. Extracted genomic DNA was stored at -80°C until further use. Genotyping was carried out using the

Axiom® Equine Genotyping Array (Affymetrix, Santa Clara, CA, USA) which contains a total of 670,796 markers (Neogen Corporation-GeneSeek Operations, Lincoln, NE, USA).

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The SNP data analysis was done using the Golden Helix SNP & Variation Suite 8.8.3

(SVS, Bozeman, MT, USA). Affymetrix quality control for the samples validated 582,598

SNPs. The call rate for one horse was less than 80% and it was dropped from the analyses.

The remaining 94 horses had call rates higher than 95%.

SNP data were removed for call rates below 95%, greater than two alleles or minor allele frequency less than 0.05 leaving 382,529 SNPs for analyses. Further analysis was done with only Thoroughbreds, Paints and Quarter Horses resulting in 55 samples (17 cases and 38 controls). Filtering left 369,064 SNPs for subsequent analyses. The Genotype

Association Tests (Basic, Genotype, Additive, Dominant and Recessive), Haplotype

Association Tests (moving window of 6 SNPs). In addition, tests were conducted for Runs of Homozygosity (ROH) based on regions of varying minimum sizes, numbers of samples and number of SNPs, allowing for 1 heterozygous position in the run. Probabilities were calculated using chi-square or correlation tests. To control for repeated analyses, probability values were corrected using the Bonferroni correction.

5.4. RESULTS

5.4.1 Association Tests

None of the association tests (Basic, Genotypic, Additive, Dominant and

Recessive) identified SNPs with significant differences in distribution when comparing cases and controls. Figure 5.1 shows the Manhattan Plot for the investigation of the recessive mode of inheritance. Investigations for haplotype associations were conducted using sliding windows of 6 SNPs and, again, no associations were found. The distribution of genotypes did not indicate an influence by population substructure; however we did

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repeat the association tests using a subset of Thoroughbred, Paint and Quarter horses based on the close relationship among those breeds Again, no significant associations (using

Bonferroni corrections) were uncovered for the genotype or haplotype association tests.

5.4.2 Runs of Homozygosity (ROH):

Traits with recessive modes of inheritance may be missed based on recessive genotypic studies due to absence of individual SNPs which distinguish cases and controls.

Therefor analyses were conducted for runs of homozygosity. The conditions used for this study ranged from 20 samples to 5 samples required to have lengths of homozygosity from

400kb to 1000kb with an expectation of 6- 10 SNPs per kb. None of these showed statistical association with EHM. A Manhattan plot for a test of recessive association this data is shown in Figure 5.2. Conditions for this figure were requirements for minimum of

400kb runs with no more than 1 heterozygote found among 5 of the 94 horses with at least

60 SNPs. The assay identified 307 runs distributed throughout the genome with none achieving statistical significance.

5.4.3 Testing BIEC2-946397:

The single SNP identified as having a statistically significant association with EHM by Brosnahan et al., (2018) was BIEC2-946397. This SNP was included in the Affymetrix array used in this study. The locus was polymorphic in this assay with two alleles, G and

A. However, the level of polymorphism was low with only 5 horses being heterozygous for the two alleles, 2 cases and 3 controls.

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5.5. DISCUSSION

The sporadic nature for the occurrence of EHM among horses previously infected with EHV-1 led to the speculation that some host factor may play a role in the occurrence of EHM. Brosnahan et al. (2018) reported an association between a SNP and the occurrence of EHM in a manner suggesting a recessive mode of inheritance. In this current study we examined 382,529 SNPs using a population of 94 horses (28 cases and 66 controls). In contrast to the report by Brosnahan et al. (2018), (129 horses with 37,173

SNPs) we did not find evidence for a genetic association with EHM. The power of our test was sufficient to reject a hypothesis of genetic effect by recessive alleles. We did not see any evidence for trends in the data identifying other possible modes of inheritance although we would not have sufficient power to detect complex genetic causes based on multiple loci.

Our assay included the SNP identified in the previous work by Brosnahan et al.

(2018), BIEC2- 946397. The marker exhibited a low level of variation in our study and there were no statistical differences between the cases and controls for the presence of variants. While most SNPs test the same regardless of assay used, some SNPS do not. We did not use other assays to assess the variation for the SNP and it is possible that the

Affymetrix assay was not reliable for this SNP. Brosnahan and co-workers confirmed their tests with a second assay, so it is more likely that our results under-represent the variation for this SNP. Nevertheless, we did not find any evidence with other SNPs in the region, or with runs of homozygosity implicating genes on chromosome 6 (Figures 1 and 2).

The difference between the two studies may have several explanations. The test is statistical, and it may be that either of the reports failed, either in detecting or not detecting

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the association. There were also a few differences in the nature of the two studies that could have had an impact. In this study all cases came from natural infections while Brosnahan and co-workers (2018) used cases from both experimental and natural infections. Other differences could be related to the genetic background of the horses. In order to obtain enough horses for the GWAS study, we included horses of diverse breeds (Supplemental

Table 1). We do not know what breeds were used in the other study, but such differences also could have an impact on the statistical evaluations.

In conclusion, based on the results of this study, we do not see any evidence for a recessive allele being determinative for the development of EHM following EHV-1 infection for any genetic locus. More complex host-pathogen interactions are possible, and discovery will require testing a larger number of horses with EHM.

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Table 5.1: List of case and control samples used in this study indicating their geographical location, breed and case/control classification. Samples were generously provided for this study by Anoka Equine Veterinary Services, MN, Equine Diagnostic Solutions, KY, Virginia Department of Agriculture and Consumer Services, VA, Alpine Veterinary Hospital, CO, Animal Health Clinic, ID, Eastern Lancaster Veterinary Clinic, PA, Oklahoma State University-Department of Veterinary Clinical Sciences, OK, Veterinary Integrative Performance Services, IA, and Dr. Nicola Pusterla School of Veterinary Medicine, University of California, Davis, CA. Sample Location Case/Control Breed No. 1 Virginia Case Warmblood 2 Virginia Case Thoroughbred 3 Virginia Case Quarter Horse 4 Virginia Case Thoroughbred 5 Virginia Case Thoroughbred 6 California Case Warmblood 7 California Case Warmblood 8 California Case Quarter Horse 9 California Case Quarter Horse 10 California Case Thoroughbred 11 California Case Thoroughbred 12 California Case Quarter Horse 13 California Case Warmblood 14 California Case Thoroughbred 15 California Case Paint 16 California Case Thoroughbred 17 California Case Quarter Horse 18 California Case Thoroughbred 19 Kentucky Case Thoroughbred 20 Kentucky Case Thoroughbred 21 Minnesota Case Quarter Horse 22 Minnesota Case Quarter Horse 23 Minnesota Case Quarter Horse 24 Minnesota Case Not mentioned 25 Iowa Case Not mentioned 26 Idaho Case Thoroughbred 27 Pennsylvania Case Not mentioned 28 Wisconsin Case Not mentioned 29 Minnesota Control Not mentioned 30 Minnesota Control Not mentioned 31 Minnesota Control Not mentioned 32 Minnesota Control Not mentioned 33 Minnesota Control Not mentioned 34 Minnesota Control Not mentioned 35 Minnesota Control Not mentioned 36 Minnesota Control Not mentioned

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Table 5.1 continued... 37 Minnesota Control Quarter Horse 38 Minnesota Control Quarter Horse 39 Minnesota Control Quarter Horse 40 Colorado Control Not mentioned 41 Virginia Control Thoroughbred 42 Virginia Control Dutch Warmblood 43 Virginia Control Draft cross 44 California Control Quarter Horse 45 California Control Quarter Horse 46 California Control Quarter Horse 47 California Control Quarter Horse 48 California Control Quarter Horse 49 California Control Quarter Horse 50 California Control Quarter Horse 51 California Control Quarter Horse 52 California Control Quarter Horse 53 California Control Quarter Horse 54 California Control Quarter Horse 55 California Control Quarter Horse 56 California Control Crossbred 57 California Control Warmblood 58 California Control Quarter Horse 59 California Control Mustang 60 California Control Thoroughbred 61 California Control Crossbred 62 California Control Crossbred 63 California Control Quarter Horse 64 California Control Lusitano 65 California Control Warmblood 66 California Control Warmblood 67 California Control Not mentioned 68 California Control Pony 69 California Control Paint 70 California Control Crossbred 71 California Control Welsh pony 72 California Control Welsh pony 73 California Control Welsh pony 74 California Control Quarter Horse 75 California Control Quarter Horse 76 California Control Pony 77 California Control Thoroughbred 78 California Control Warmblood 79 Kentucky Control Thoroughbred 80 Kentucky Control Thoroughbred

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Table 5.1 continued…. 81 Kentucky Control Thoroughbred 82 Kentucky Control Thoroughbred 83 Kentucky Control Thoroughbred 84 Kentucky Control Thoroughbred 85 Kentucky Control Thoroughbred 86 Kentucky Control Thoroughbred 87 Kentucky Control Thoroughbred 88 Kentucky Control Thoroughbred 89 Kentucky Control Thoroughbred 90 Kentucky Control Thoroughbred 91 Kentucky Control Thoroughbred 92 Iowa Control Not mentioned 93 Iowa Control Not mentioned 94 Iowa Control Not mentioned 95 Oklahoma Control Paint

(SNP) in horse chromosome 6 (ECA6) with the occurrence of EHM [363]. In the study described below, we were unable to find this association on ECA6, or any other association.

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Figure 5.1: Manhattan plot of the -log10 P values for correlation of SNPs with EHM based on recessive model genome-wide association (GWAS) study using 94 horses with 382,529 markers were analyzed using the Ecab 3.0 data. No significance was observed after adjusting for Bonferroni corrections.

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Figure 5.2: Genome plot (Ecab 3.0) of -log10 P values for association test of runs of homozygosity (ROH; minimum 5 samples with minimum homozygosity length of 400 kb with at least 60 SNPs) Association test. No significant associations were found for any ROH.

CHAPTER SIX GENERAL DISCUSSION

EHV-1, EHV-4 and EAV are among the most important viruses infecting horse populations around the world and pose a significant threat to the global as well as the U.S. equine industry which is estimated to be approximately 122 billion dollars

(https://www.horsecouncil.org/resources/economics/). Characteristic to EHV-1, EHV-4 and EAV is their endotheliotropic nature which plays a major role in inducing vasculitis.

Induction of vasculitis in the blood vessels of the pregnant uterus is the underlying reason for abortions in pregnant mares following EHV-1 or EAV infections [5; 6; 17-19; 61].

EHV-4 is primarily regarded as a respiratory pathogen of equids yet on rare occasions

EHV-4 infect and replicate in endothelial cells of local capillaries of lymph nodes and pulmonary vessels. Vasculitis is the inflammation of the walls of the blood vessels and several factors including the production of proinflammatory cytokines and chemokines by immune and endothelial cells play significant roles [255]. However, the proinflammatory cytokine and chemokine response mounted by EECs in response to EHV-1, EHV-4 and

EAV infections have not been extensively characterized.

The proinflammatory cytokine and chemokine profile of virus infected EECs revealed the gene expression levels of IL18, GM-CSF, CXCL16, IFNɣ (proinflammatory cytokines) and IL8 and CCL3 (proinflammatory chemokines) were expressed less than two-fold in most instances (Table 2.3) suggesting that, at least in the case of virus infections, the expression of these genes were not being induced in EECs. Furthermore, It is well established that EHV-1 and EAV infected PBMCs upregulate IFNɣ gene expression [64;

364; 365]. However, IFNɣ gene expression or secretion was not detected in EECs

152

following infection with EHV-1, EHV-4 or EAV as demonstrated by RT-qPCR and

Luminex data suggesting EECs may not be a good source of IFNɣ following virus infections (Table 2.3). The gene expression of the rest of the proinflammatory cytokines and chemokines (IL6, IL17, CXCL9, CXCL10, CXCL11, TNFα, IFNα, IFNω, CCL4 and

CCL5) analyzed during this study were upregulated 5-folds or higher in virus infected

EECs (Table 2.3). Interestingly, it was observed that the relative gene expression as well as the secretion of these proinflammatory cytokines and chemokines were relatively lower in EECs infected with EAV, suggesting a relatively lower induction of the host inflammatory response by EAV compared to EHV-1 and EAV infected EECs (Table 2.3).

Moreover, as is evident from the RT-qPCR and Luminex data, EAV infected EECs induces a lower gene expression as well as a lower secretion of CXCL10 and IFNα relative to EHV-1 and EHV-4 infected EECs (Table 2.3, Figures 2.2 & 2.4). Previous studies done in our lab has unequivocally demonstrated EAV suppresses the production of IFNβ secretion from EECs and this is brought about by the action of EAV nsp1 on IRF3 and NF-

휅B [138]. Therefore, the relatively lower IFNα gene expression as well as secretion might be a result of the same suppression mechanism since IFNα, similar to IFNβ, is also stimulated by IRF3 and NF-휅B [222]. As such, the data from this study together with the study previously done in our lab by Go et.al. (2012) [138] makes a strong case for the reduction of type I interferon ( IFNα and IFNβ) gene expression as well as their secretion from EECs following EAV infection. Moreover, EAV infected EECs has a relatively lower gene expression for CXCL10 which is a proinflammatory chemokine that induces T and B lymphocytes, natural killer cells, dendritic cells and macrophages [278-280].

153

Interestingly, as demonstrated by the RT-qPCR and Luminex results, the proinflammatory cytokine and chemokine gene expression and secretion from EECs following EHV-1 and EHV-4 infections, for the most part, did not show significant differences (Table 2.3, Figures 2.2 & 2.4). Furthermore, the growth curves for EHV-1 and

EHV-4 in EECs followed a similar pattern as well (Figure 2.1). During natural infection,

EHV-1 strains cause abortions and occasionally EHM by inducing vasculitis in the pregnant uterus and the CNS. However, such disease manifestations are rare following natural EHV-4 infections due to the absence of CAV [178; 275]. On instances CAV is detected following EHV-4 infections, viral DNA remains in low amounts in PBMCs and only for a short period of time [276]. This appears to be a major limitation in EHV-4 pathology that prevents the efficient transfer of virus particles from its primary sites of infection to endothelial cells of the pregnant uterus or the CNS where it can infect and induce viral vasculitis. Studies including this one have demonstrated that EHV-4 is capable of infecting and replicating in EECs in-vitro and occasionally in-vivo [37; 277].

Furthermore, this study demonstrates that the cellular response of EECs towards EHV-1 and EHV-4 infections, for the most part, remains the same with regards to the proinflammatory cytokine and chemokine gene expression as well as secretion (Table 2.3,

Figures 2.2 & 2.4). Therefore, it is reasonable to assume that if EHV-4 strains infected endothelial cells in-vivo, the vasculitis process and the following disease manifestations in the form of abortions or EHM might be the same as with EHV-1 strains. The results of this study indicate the lack of such disease manifestations is probably due to the inefficient transportation of the virus to endothelial cells of blood vessels of the pregnant uterus and the CNS rather than EHV-4 strains not being able to infect endothelial. Therefore, the

154

results of this study strongly point towards inefficient leukocyte associated viremia as a major reason for the mild disease outcomes associated with EHV-4 infections.

Global transcriptomic analysis of EAV and EHV-1 infected cells provided valuable insights about the molecular pathways and genes associated with the host response against

EHV-1 and EAV infections. The pathway analysis revealed that EECs infected with both

EHV-1 and EAV differentially expressed the CCKR signaling map”, “Inflammation mediated by chemokine and cytokine signaling pathway” and the “Gonadotropin-releasing hormone receptor pathway” during the first 12 h of infection (Tables 4.1, 4.2, 4.3, 4.4A&B and 4.5A&B). These 3 pathways are associated with the host Inflammatory responses against infections [308; 325; 326]. As such, the data from this study reveals both viruses stimulate pathways that contribute to the local inflammatory response in EECs. This, together with the relative gene expression and Luminex data indicates the molecular pathways as well as the proinflammatory cytokines and chemokines that play key roles in inducing vasculitis which is the major pathological condition that drives the disease outcomes associated with both EHV-1 and EAV infections.

Global transcriptome analysis together with RT-qPCR revealed the upregulation of

TLR3 mRNA in EECs following EHV-1 infection suggesting the involvement of TLR3 in the induction of the inflammatory response associated with EHV-1 infections (Figure 4.1). interestingly, siRNA silencing of TLR3 in EECs revealed that TLR3 plays a key role in the upregulation of IFNα mRNA during the first 6 h of EEC infection. However, no such involvement for TLR3 following EAV infection of EECs was made (Figure 4.3A&B).

The GWAS analysis of horses with EHM was done in order to investigate host genetic factors that may play a role in EHM development following EHV-1 infection. In

155

this current study we examined 382,529 SNPs using a population of 94 horses (28 cases and 66 controls), however, we did not find evidence for a host genetic association with

EHM development following EHV-1 infections (Table 5.1, Figures 5.1&5.2). The power of our test was sufficient to reject a hypothesis of genetic effect by recessive alleles.

Furthermore, we did not see any evidence for trends in the data identifying other possible modes of inheritance (Figure 5.2). However, the power of our test was not sufficient to detect complex genetic causes involving multiple loci. One of the major limitations to this test was the lower sample size and this may have affected the detection of genetic loci associated with EHM development. Furthermore, we failed to corroborate the results obtained by Brosnahan et al. (2018) that reported an association between a SNP and the occurrence of EHM in a manner suggesting a recessive mode of inheritance [363].

Future directions

The results presented here provide novel insights about the modulation of the innate immune response of EECs against EHV-1 and EAV infections. In particular, our data taken together with the observations of Go et.al. (2012), makes a strong case for the suppression of type I interferon (IFNα and IFNβ) gene expression as well as their secretion from EECs following EAV infection (Table 2.3 & Figure 2.2). Furthermore, the secretion of CXCL10 as well as the gene expression of proinflammatory cytokines and chemokines were also relatively lower in EAV infected EECs when compared with EHV-1 infected EECs (Table

2.3 & Figure 2.4). However, the mechanism utilized by EAV to influence this relatively lower proinflammatory response from EECs remains to be further elucidated. Go et.al.

(2012) has demonstrated that EAV nsp1 influences the suppression of IFNβ production from EAV infected EECs by interfering with the action of IRF3 and NF-휅B [138].

156

Therefore, further investigations are warranted to elucidate if EAV nsp1 is responsible for the lower proinflammatory response as well as the mechanism that bring about this lower inflammatory response.

Data from this study also demonstrated the involvement of TLR 3 in inducing IFNα from EHV-1 infected EECs during the first 6 h of infection. However, TLR 3 was not involved in IFNα stimulation after the initial stages of infection even though IFNα gene expression as well as secretion was observed (Figure 3.3A&B). However, what molecules and pathways are involved in this process remains to be elucidated and warrants further investigations. Furthermore, TLR 3 was not deemed to be involved in the proinflammatory response induced by EAV in EECs. Therefore, it is important to further investigate the role played by TLRs during EAV infection of EECs.

The transcriptomic analysis of EHV-1 and EAV infected EECs displayed distinct clusters of up and downregulated genes as well as pathways (Tables 4.1, 4.2, 4.3, 4.4A&B and 4.5A&B and Figure 4.1). Among others, the “gonadotrophin hormone releasing receptor pathway”, “CCKR signaling map” and the “Inflammation mediated by cytokine and chemokine pathway” was differentially expressed frequently in EECs infected with both EHV-1 and EAV (Tables 4.1, 4.2, 4.3, 4.4A&B and 4.5A&B). Since this study was only focused on elucidating these differentially expressed genes and pathways in-silico, further work is required to confirm these results with functional studies.

The GWAS study investigated the involvement of host genetic factors in the development of EHV-1 induced EHM but found no evidence for a genetic association

(Figure 5.1). However, the biggest limitation in that study was the sample size (94 samples). Even though the power of our test was sufficient to reject a hypothesis of genetic

157 effect by recessive alleles it wasn’t sufficient to detect complex genetic causes based on multiple loci. Therefore, a repeated experiment with a higher sample number has the potential to reveal more promising results.

158 Appendix 1 EXPERIMENTAL METHODS

TLR siRNA Transfection Method to EECs siRNA stock solution preparation 1. Dilute 5 nmols of siRNA in 100 µL of nuclease free water to make a 50 µM stock solution. Store the stock solution in -20 °C until further use. Prepare fresh working solutions from the stock solution for siRNA transfection. siRNA transfection into EECs 1. Prewarm 24 well cell culture plates with complete DMEM media 2. Harvest cells by trypsinizing (1×105 cells/well) and centrifuge at 90g for 10 minutes 3. Remove the supernatant and resuspend the cell pellet in 4DNucleofector™ Solution. 4. Prepare master mixes (4DNucleofector™ Solution+cells+siRNA) by dividing cell suspension according to number of substrates 5. Add required amount of substrates to each aliquot 6. Transfer master mixes into the Nucleocuvette™ Vessels 7. Place the Nucleocuvette™ Vessels inside the Lonza 4D nucleofection unit. 8. Use the program for BHK21 to electroporate cells 9. Remove the Nucleocuvette™ Vessels from the Lonza 4D nucleofection unit 10. Mix 500 µL of prewarmed media with the electroporated cells in Nucleocuvette™ Vessels and transfer the cells into 24 well cell culture plates 11. Incubate the cell culture plates for 24 h in a 37 °C incubator in the presence of 5% CO2. In-vitro infection of siRNA transfected EECs 1. Prepare fresh working stocks of virus by diluting concentrated stocks in MEM. All in-vitro infections are carried out at 2 MOI. 2. Remove cell culture media from siRNA transfected cell containing 24 cell culture plates by aspiration 3. Add the virus containing MEM onto cell monolayers and incubate the plates for 1 h in a 37 °C incubator in the presence of 5% CO2. 4. After 1 h remove the MEM medium containing the virus through aspiration and add 1 mL of fresh complete DMEM medium to each well. 5. Incubate the cell culture plates for 6 and 12 h in a 37 °C incubator in the presence of 5% CO2.

159 Luminex Assay for Cell Culture Supernatant Preparation of antibody-immobilized beads 1. Sonicate antibody-bead vials for 30 seconds followed by vortexing for 1 minute. Add 60 μL from each antibody-bead vial to the mixing bottle and bring final volume to 3.0 mL with bead diluent. Vortex the final bead mix thoroughly. Preparation of quality controls 1. Reconstitute quality controls 1 and 2 by adding 250 μL of deionized water. Invert the vial several times to mix. Preparation of wash buffer 1. Dilute 60 mL of 10X wash buffer with 540 mL deionized water. Preparation of equine cytokine standard 1. Reconstitute the equine cytokine standard with 250 μL deionized water. Invert and mix the vial several times. Vortex the vial for 10 seconds. Preparation of samples 1. Thaw samples on ice and centrifuge the sample to remove debris. Luminex assay for cell culture supernatant 1. Add 200 μL of wash buffer into each well of the plate. Seal the plate and mix on a plate shaker for 10 minutes at room temperature. 2. Invert the plate and decant the wash buffer. 3. Add 25 μL of each standard and control into the appropriate wells. 4. Add 25 μL of assay buffer to the sample wells. 5. Add 25 μL of complete DMEM to the background, standards, and control wells. 6. Add 25 μL from each sample to appropriate wells. 7. Add 25 μL of mixed beads to each well. 8. Seal the plate with a plate sealer and wrap the plate with foil. Incubate with agitation on a plate shaker overnight (16-18 hours) at 2-8°C. 9. Following incubation, remove well contents and wash the plate 3 times with the wash buffer. 10. Add 25 μL from the detection antibodies into each well. 11. Seal the plate with a plate sealer and cover with foil. Incubate with agitation on a plate shaker for 1 hour at room temperature. 12. Add 25 μL of Streptavidin-Phycoerythrin to each well containing the detection antibody mix. 13. Seal the plate with a plate sealer and cover with foil. Incubate with agitation on a plate shaker for 30 minutes at room temperature. 14. Gently remove the well contents and wash the plate 3 times with wash buffer.

160 15. Add 150 μL of sheath fluid into all the wells. Resuspend the beads on a plate shaker for 5 minutes. 16. Run the plate on a Luminex® 200™ machine. Transcriptome Analysis of Virus Infected EECs In-vitro infection of EECs with EHV-1 and EAV 1. Seed 1×106 EECs in 6 well cell culture plates and incubate in a 37 °C incubator in the presence of 5% CO2 to form cell monolayers. 2. Calculate the volume of virus needed for a MOI of 2. Mix the virus with MEM medium and add onto cell monolayers. Adsorb for 1 h at 37 °C in the presence of 5% CO2. 3. After 1 h, remove the virus containing medium and add fresh complete DMEM medium to cell monolayers. Total RNA extraction 1. Total RNA was extracted from EHV-1 and EAV infected EECs at 6 and 12 hpi timepoints using the miRNeasy mini kit (Qiagen, Hilden, Germany) following the manufacturers protocol. 2. Extracted RNA was quantified using a bioanalyzer. 3. 300 ng of total RNA was submitted for whole transcriptome sequencing. Data analysis 1. Reads were initially trimmed for adapters and quality using TrimGalore Version 4.4. 2. Trimmed reads were aligned to the horse reference genome (EquCab3.0) using STAR (Release 2.5.2b). 3. Cufflinks (Release 2.2.1) was used to annotate to the equine reference transcriptome available in NCBI database. 4. Cuffdiff 2.2.1 was used to calculate differently expressed genes between samples from the mock and virus infected groups.

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194 VITA

Saranajith Wangisa Dunuwille

PERSONAL INFORMATION

Place of birth: Peradeniya, Sri Lanka

EDUCATION

2014 Master of Science (M.Sc.).

University of Peradeniya, Post Graduate Institute of Science, Peradeniya, Sri Lanka.

2011 Bachelor’s in Veterinary Science (B.V.Sc.).

University of Peradeniya, Faculty of Veterinary Medicine and Animal Science,

Peradeniya, Sri Lanka.

PROFESSIONAL EXPERIENCE

2014 – 2019 Graduate Research Assistant, Molecular Virology Laboratory, Department of

Veterinary Science, Maxwell H. Gluck Equine Research Center, University

of Kentucky, Lexington, KY, USA

2013–2014 Graduate Research Assistant, Department of Molecular Biology and

Biotechnology, University of Peradeniya

2011 – Veterinary Surgeon, Sri Lanka.

AWARDS AND FELLOWSHIPS

1. Geoffrey C. Hughes Fellowship, Maxwell H. Gluck Equine Research Center.

2014 – 2019

2. Department of Veterinary Science Travel Award to Attend the 37th Annual

meeting of the American Society for Virology. (2018)

195

3. 3rd Place Presentation at the Gluck Equine Research Center 3MT

competition. (2018)

4. University of Kentucky, Graduate School Travel Award to Attend the 36th

Annual meeting of the American Society for Virology. (2017).

5. 3rd Place presentation at the Gluck Equine Research Center 3MT competition

(2017)

PUBLICATIONS IN PREPARATION

1. Wangisa M.B. Dunuwille., Navid Y. Mashouf., Udeni B.R. Balasuriya., Nicola Pusterla., Ernest Bailey. Genome Wide Association Study of Horses with Equine Herpesvirus Myeloencephalopathy to Identify Single Nucleotide Polymorphisms in the Equine Genome. Equine veterinary journal, 2019

ABSTRACTS AT NATIONAL AND INTERNATIONAL MEETINGS

1. Wangisa Dunuwille, Sanjay Sarkar, Annet Kyomuhangi, R. Frank Cook, and Udeni B. R. Balasuriya (2017), Expression of Cell Surface Inflammatory Markers, Inflammatory Cytokines and Chemokines in Equine Endothelial Cells Following Infection with Equine Herpesvirus-1 (EHV-1) and Equine Herpesvirus-4 (EHV-4). 36th Annual meeting of the American Society for Virology. University of Wisconsin-Madison, WI. USA

2. Wangisa Dunuwille, R. Frank Cook, and Udeni B. R. Balasuriya (2018), Equid alphaherpesvirus 1 and Equine Arteritis Virus Induces Proinflammatory Cytokine and Chemokine Gene Expression Via Two Different Toll-like Receptors. 37th Annual meeting of the American Society for Virology. University of Maryland in College Park, MD. USA

3. Dunuwille, S.W.M.B., Premachandra, T.N., Rajapaksha, P., Rajapakse, S., Jayawardane, B.C., Himali, S.M.C., Kodithuwakku, S., and Sooriyapathirana, S.D.S.S., (2014). Development of a PCR based genotyping procedure to identify the adulterations to chicken and wild boar meat in sri lanka. Proceedings of the Peradeniya Univ. International Research Sessions, Sri Lanka, 18, 203

4. Dunuwille, S.W.M.B., Premachandra, T.N., Rajapaksha, P., and Sooriyapathirana, S.D.S.S., (2013). Development of a PCR based genotyping procedure to identify the adulterations to beef in Sri Lankan meat market. Proceedings of the Research Symposium of Uva Wellassa University, 7-9.

196