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2014 - interactions in the Pacific white , Litopenaeus vannamei Duan Sriyotee Loy Iowa State University

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Host-virus interactions in the Pacific white shrimp, Litopenaeus vannamei

by

Duan Sriyotee Loy

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Veterinary Microbiology

Program of Study Committee: Lyric Bartholomay, Co-Major Professor Bradley Blitvich, Co-Major Professor D.L. Hank Harris Cathy Miller Michael Kimber

Iowa State University

Ames, Iowa

2014

Copyright © Duan Sriyotee Loy, 2014. All rights reserved.

ii

TABLE OF CONTENTS

CHAPTER 1: GENERAL INTRODUCTION...…………………………………………...1 Introduction…………………………………………………………………………………………1 Dissertation Organization…………………………………………………………………………3 Literature Review ...... 3 References ...... 23

CHAPTER 2: CHARACTERIZATION OF NEWLY REVEALED SEQUENCES IN THE INFECTIOUS MYONECROSIS VIRUS IN LITOPENAEUS VANNAMEI ...... 33 Abstract ...... 33 Introduction ...... 35 Materials and Methods ...... 37 Results and Discussion ...... 47 Acknowledgments ...... 53 References ...... 54

CHAPTER 3: VARIATION IN VIRULENCE IN GEOGRAPHICALLY DISTINCT ISOLATES OF INFECTIOUS MYONECROSIS VIRUS IN THE PACIFIC WHITE SHRIMP, LITOPENAEUS VANNAMEI ...... 71 Abstract ...... 71 Introduction ...... 72 Materials and Methods ...... 74 Results ...... 78 Discussion ...... 81 References ...... 84

CHAPTER 4: COMPARATIVE ANALYSIS OF SUPPRESSION OF IMMEDIATE EARLY, EARLY AND LATE GENES OF VIRUS ...... 96 Abstract ...... 96 Introduction ...... 97 Materials and Methods ...... 100 Results ...... 104 Discussion ...... 107 Acknowledgements ...... 111 References ...... 112

CHAPTER 5: DIRECT DELIVERY OF DOUBLE STRANDED RNA INTO THE FOREGUT LUMEN OF LITOPENAEUS VANNAMEI DEMONSTRATES PROTECTION AGAINST WHITE SPOT SYNDROME VIRUS AND INFECTIOUS MYONECROSIS VIRUS ...... 120 Abstract ...... 120 Introduction ...... 121 Materials and Methods ...... 124 Results ...... 128 Discussion ...... 130 Acknowledgements ...... 131 References ...... 132 iii

CHAPTER 6: GENERAL CONCLUSIONS ...... 141

ACKNOWLEDGEMENTS ...... 145

1

CHAPTER1: GENERAL INTRODUCTION

Introduction

The global population is predicted to expand from the current size to an estimated 7 billion to 9.1 billion by 2050. Aquatic food products, the fastest growing of the food-producing sectors, are predicted to supply more than 50% of this sector by 2015 (Stentiford et al. 2012). The industry continues to grow rapidly, particularly in the case of cultured marine shrimp. Indeed, cultured shrimp harvest was an estimated 3.5 million metric tons representing a sale value of $14.6 billion

USD in 2009 (Moss et al. 2012). In the early era of expansion

(1970s-1980s), was the major in culture species, and the industry relied on wild caught stocks to produce postlarvae (PL) and adult broodstock to supply seed for the industry (Moss et al. 2012). However, this is a high-risk practice because wild-caught animals can be carriers of diseases that then can be introduced into culture facilities. The industry largely has shifted from using wild seed stocks of P. monodon to using specific pathogen-free (SPF) domesticated lines of Litopenaeus vannamei, a North American species. L. vannamei was introduced to Asian crustacean fisheries in the late 1990s and soon became the dominant species in Asia and other shrimp farming regions (Flegel 2012, Lightner et al. 2012). Aquaculture practices have also changed from a traditional extensive farming to semi-intensive or intensive systems that are more efficient production systems. The adoption of L. vannamei in culture systems worldwide, along with effective biosecurity measures and intensive farming techniques, have resulted in 2 rapid growth of the global shrimp farming industry over the last four decades (Flegel

2012, Moss et al. 2012). This growth has been hampered only by pandemic viral diseases that have devastated harvests and yields; therefore, they have also greatly impacted the economics of the shrimp industry worldwide (Flegel 2012, Lightner et al. 2012). Thus, development of strategies to prevent and control viral diseases is of tremendous interest and importance for the sustainability of this industry.

The research described in this dissertation emphasizes two virus pathogens:

1) Infectious myonecrosis virus (IMNV) and 2) White spot syndrome virus (WSSV) that have caused more than $16 billion in losses combined since the first outbreaks

(Lightner et al. 2012). The first manuscript of this dissertation describes the characterization of newly revealed sequences in the IMNV genome. This paper provides evidence that additional sequences exist in the IMNV genome, resulting in an overall genome length that is 8226 base pairs (bp) as compared to the original description of 7561 bp. The second manuscript describes variation in virulence in geographically distinct isolates of IMNV from Brazil and Indonesia. The third manuscript describes the utilization of RNAi to silence WSSV by targeting early gene expression and that is more effective at suppressing virus replication than suppressing late genes. This result provides a new target antiviral for WSSV. The fourth manuscript presents the development of direct delivery methodology of double-stranded RNA (dsRNA) into the foregut lumen of L. vannamei. This manuscript indicates that delivery of dsRNA by reverse gavage elicits protection that is comparable to intramuscular injection methods, and provides evidence that shrimp 3 possess some mechanism to uptake dsRNA from the gut lumen. This method can be used as a proxy for possible per os vaccination trials.

Dissertation Organization

This dissertation is organized in manuscript format. Chapter 1 includes a brief introduction to a topic followed by a literature review with a general background of

IMNV and WSSV, current knowledge of RNA interference (RNAi), shrimp immunity and augmenting RNAi to control virus diseases. Chapter 2, 3, 4, and 5 are manuscripts describing the research and results obtained by the primary research author, Duan Loy, and co-authors. Chapter 6 provides general conclusions of the research, discussion of future work followed by an acknowledgements section.

Literature Review

History of Infectious myonecrosis virus (2002-2013)

An unknown disease outbreak described as “cotton shrimp disease” was reported in a Pacific white shrimp farm, L. vannamei, in northeastern Brazil in 2002

(Poulos et al. 2006). The infected shrimp were initially thought to be infected with a microsporidian parasite due to the gross signs of white abdominal muscle in the shrimp. Some of the animals were submitted to the Office International des

Epizooties (OIE) reference laboratory for shrimp diseases at the University of

Arizona. The samples were processed for histopathology and PCR/ RT-PCR and results showed that the shrimp samples were not infected with microsporidia or other known (Andrade 2009). The clinical signs of disease are white and opaque muscle, specifically at the distal abdominal segment and tail fan. Infected animals 4 are lethargic and have reduced feed consumption. The feed conversion ratio (FCR) in infected ponds rose from 1.5 to 4.4 and the survival rate decreased to as low as

21%. The disease progressed slowly throughout the culture season and the overall mortality rate ranged from 40-70%. In addition to L. vannamei, P. monodon,

Litopenaeus stylirostris and Farfantepenaeus subtilis are susceptible to IMNV infection (Tang et al. 2005, Coelho et al. 2009). IMNV transmission occurs horizontally by ingestion of infected tissue from dead or moribund shrimp.

The disease was then spread through shrimp farming areas across Brazil. At the end of 2005, it was estimated that shrimp farms in Brazil suffered $440 million

USD of economic losses as a result of this disease (Andrade et al. 2007). The etiologic agent was described and named Infectious myonecrosis virus (IMNV)

(Lightner 2004). Due to the significant impact that IMNV could have on disease free countries, IMNV was listed by the OIE as a pathogen and reportable disease in

2005. The OIE is an international organization that functions to inform governments of the emergence of disease and develop the control measurements for those diseases. Unfortunately, IMNV emerged in East Java in Indonesia in 2006. The of IMNV isolates from Indonesia and Brazil share 99.6% identity at the nucleotide (nt) level (Senapin et al. 2007). The translocation of IMNV was probably through unregulated transportation of live animals (Flegel 2006, Senapin et al. 2007,

Walker & Mohan 2009), and the virus subsequently spread to other regions of the country. Financial losses in these two endemic countries was estimated at more than $1 billion USD (Lightner et al. 2012). IMNV outbreaks in other countries have not yet been reported. Rumors of disease outbreaks in other Asian countries such 5 as Thailand and China have circulated periodically due to the similar clinical signs of white muscles (Senapin et al. 2011, Yan et al. 2013). However, white muscles can be caused by muscle cramp syndrome, stress factors, penaeid white tail disease

(Penaeus vannamei Nodavirus) and white tail disease of rosenbergii

(Nodavirus) (Tang et al. 2007, Bonami & Sri Widada 2011, Senapin et al. 2011).

IMNV disease has to be confirmed by an OIE approved, standard diagnostic techniques.

IMNV disease etiology and genome characterization

IMNV was classified in the member of family based on phylogenetic analysis of the RNA dependent RNA polymerase (RdRp) coding region; this revealed that IMNV is most closely related to Giardia lamblia virus

(GLV). Fungi and protozoa that are infected with , such as

Saccharomyces cerevisiae virus (ScV) and GLV, typically are asymptomatic and the virus is persistent. Most viruses in the family Totiviridae are transmitted to new cells only during division, sporogenesis and cell fusion (Poulos et al. 2006). GLV,

IMNV and Piscine myocarditis virus (PMCV) exhibit extracellular transmission, which is different from other viruses in family Totiviridae. IMNV and PMCV are unique amongst the genus because they infect penaeid shrimp and Atlantic salmon (Salmo salar L.), respectively (Haugland et al. 2011) as opposed to other viruses in Totiviridae family that infect fungi and protozoa (Tang et al. 2008). These two viruses cause significant pathological changes in muscle that can be observed by histopathology. IMNV replicates in the of muscle cells as demonstrated 6 by in situ hybridization (Poulos et al. 2006), resulting in gross signs that are easily observed as opaque muscle in the abdominal segments.

IMNV is non-segmented, non-enveloped, double-stranded RNA (dsRNA) and the virus particle is icosahedral with a 40 nm diameter. The initial report of IMNV genomes of both strains indicated the genome size 7561 bp (NC007915.2,

AY570982.2; EF061744.1), which is composed of ORF1 and ORF 2. The putative

ORF 1 (nt 136-4953) encodes a protein with a dsRNA-binding motif (DSRM) (nt 136-

315, 60 amino acids (aa)) and a major protein (MCP) (nt 2248-4953, 901 aa).

The putative ORF 2 (nt 5241- 7490) encodes RNA-dependent RNA polymerase

(RdRp) (nt 5241-7490, 749 aa) (Senapin et al. 2007). Additional studies revealed that 2A-like cleavage and a shifty heptamer of ORF1 produce polyprotein products; protein 1, protein 2 and protein 3 (Nibert 2007). The functions of these proteins remain unclear (Poulos et al. 2006, Nibert 2007). The study used cryomicroscopy and 3D imaging to further characterize the virion structure. The results demonstrate the protrusion fibers form five-fold axes that may play a role in facilitating extracellular transmission and pathogenesis of IMNV (Tang et al. 2008). A recent study showed that the laminin receptor (Lamr) is a proposed virus receptor because it interacts with the capsid protein of IMNV based on a yeast two-hybrid screen

(Busayarat et al. 2011). Further studies are needed to have a better understanding of the infection mechanisms of IMNV. The initial description of the IMNV genome included 7561 bp; however, our subsequent study revealed that the IMNV genome is larger in size (8226 bp) and therefore has an expanded protein coding capacity. The 7 addition of this novel sequence changes our understanding of the IMNV genome organization and provides novel insight into IMNV biology.

IMNV diagnostic methods

The clinical signs of white muscle in penaeid shrimp can be caused by infectious and non-infectious disease. A standardized diagnostic technique is necessary to differentiate from disease caused by stress, hypoxia or other infectious agents. The diagnostic methods approved by the OIE (2012), include histopathology, antibody-based antigen detection (Kunanopparat et al. 2011) and molecular techniques (quantitative real-time RT-PCR, nested RT-PCR and in-situ hybridization). Infected muscle tissue can be used to detect virus load by qRT-PCR

(Andrade et al. 2007). Histopathological lesions are evident in infected tissue, particularly in striated muscle (skeletal muscle and less frequently in cardiac muscle), connective tissue, hemocytes and lymphoid organ parenchymal cells. The histopathological changes appear as myonecrosis with some infiltration and accumulation of hemocytes (blood cells). Intracytoplasmic, dark basophilic inclusion bodies are seen within the muscles cells, hemocytes and connective tissue cells.

The lymphoid organ (LO) also shows significant histopathological changes. During

IMNV infection, the LO is hypertrophied and spheroids are commonly found (Poulos et al. 2006, Loy et al. 2012). In addition to laboratory diagnostics, a pond-side immunochromatographic test strip detection method has been developed to for

IMNV infection. This technique uses a monoclonal antibody specific to the capsid protein of IMNV. The IMNV strip test is 300-fold less sensitive than one-step RT- 8

PCR; however, it is inexpensive and user-friendly for farmers (Chaivisuthangkura et al. 2013).

IMNV disease prevention and control measurements

There are no methods to effectively prevent and control outbreaks of IMNV in the field. However, studies have shown that administering sequence-specific RNAi antiviral molecules result in significantly reduced pathology and virus load in infected animals (Loy et al. 2012). These data prompted us to put forth the argument that sequence-specific, optimized dsRNA functions as a vaccine to prevent infection and disease caused by IMNV (Bartholomay et al. 2012). Moreover, these dsRNA constructs can be utilized as a therapeutic antiviral treatment against IMNV when administered post-infection. Administration of 0.5 µg/shrimp of dsRNA194-275 48 hours following IMNV injection resulted in at least 50% survival compared to other treatment groups (Loy et al. 2013). Although these are promising and provocative results, exploiting RNAi as a disease control method in the field is in the developmental phase; thus the use of more basic prevention strategies to control and prevent is essential. These include use of Specific Pathogen Free

(SPF) animals, farm biosecurity and family-based selection to breed for disease resistance (Moss et al. 2012). Using SPF animals with biosecurity measures on farms is a more promising means to reduce the risk of IMNV outbreaks because breeding for IMNV resistance thus far has been minimally successful. Based on anecdotal evidence, selective breeding for IMNV resistance shows significant improvement in survival, from 3.2% in the F1 generation to 55.3% in F7 (White-

Noble et al. 2010). However, these data have not been published in the peer- 9 reviewed literature and the selection process was conducted using IMNV (Brazil)

(Lima et al. 2013) so studies are needed to determine if the IMNV resistant line is equally resistant to IMNV (Indonesia).

White spot syndrome virus: The most devastating virus to the shrimp industry

White Spot Disease (WSD) was first described in China and in 1992 and1993. Subsequently, the disease spread quickly throughout Asia including

Japan, Korea and Thailand (Chou et al. 1995, Wongteerasupaya et al. 2003, Walker

& Mohan 2009). WSD later emerged in the United States and was reported in South

Texas in 1995, probably as a result of imported frozen-bait shrimp products from

Asia (Hasson et al. 2006). Outbreaks of WSD were also found in other shrimp farming areas including South, North and , Europe and the Middle

East (Leu et al. 2009, Walker & Mohan 2009, Sanchez-Paz 2010, Lightner et al.

2012). To date, WSSV is the most contagious virus in shrimp aquaculture and it causes the highest economic losses of all infection shrimp diseases. The economic impact of WSD is estimated at more than $15 billion (Lightner et al. 2012). WSD is highly virulent and causes up to 100% mortality within 3-7 days of infection. White spots (ranging from 0.5-3.0 mm in diameter) can be observed on the carapace, appendages and epidermis of infected animals; however, these signs are not a definitive diagnostic because white spots also are caused by high alkalinity, bacterial infection and other environmental stressors (Leu et al. 2009). Other clinical signs are reddish-pinkish discoloration, lethargy and reduced feed intake. WSSV can infect a wide-host range of aquatic crustaceans including and that can serve as reservoir hosts in the natural environment. Other invertebrates such as copepods, 10 artemia and polychaetes also can serve as a mechanical vectors of WSSV infection

(Walker & Mohan 2009). Other WSSV hosts both natural and experimental infection has been reviewed by Sanchez-Paz (2010). WSSV has a wide range of tissue tropisms of infected shrimp including tissue of ectodermal (cuticular epidermis, foregut, hindgut, gills, nervous system) and mesodermal (lymphoid organs, connective tissue, antennal glands) origins. Virus replication occurs in the nucleus causing nuclear hypertrophy, nuclear dissolution and chromatin margination.

Infection results in an intranuclear eosinophilic Cowdry A-type inclusions, i.e., amorphous clear halos within the nuclear membrane (Leu et al. 2009).

White spot disease etiology

WSSV is a large (80-120x250-280 nm) rod to elliptical-shaped, double- stranded DNA, enveloped virus with unique tail-like appendages at one end. The virus genome is circular, approximately 300 kbp with at least 181 ORF. In initial reports, WSSV was classified as a non-occluded baculovirus, but later evidence showed that WSSV is a new virus (van Hulten et al. 2001) and within its own genus

(Whispovirus) and is not related to baculoviruses (Fauquet 2005). The WSSV genome has been completely sequenced from three WSSV isolates from China,

Thailand and Taiwan (accession no. AF332093, AF369029 and AF440570, respectively) which share 99.3% nt identity (Sanchez-Paz 2010). The virion is composed of at least 45 structural proteins that are arranged into three layers including the nucleocapsid, tegument and envelope. The nucleocapsid is a helical, baciliform structure that contains the dsDNA genome and nine proteins including, for example, VP15, a basic DNA-binding protein, and VP664 that forms a stack ring unit 11 in the tegument layers. The tegument layer is in-between the nucleocapsid and envelope and is composed of at least four structural proteins, including VP26. The envelope layers comprise at least 28 structural proteins such as VP19 and VP28.

WSSV genes are classified according to temporal gene expression in three phases including immediate-early (IE), early (E) and late (L) genes. Viral IE genes can be expressed independently during the early stage of infection, approximately 2 hours post-primary infection (Liu et al. 2005). Viral E genes usually encode regulatory proteins which are necessary for L gene expression (Han et al. 2007). Viral L genes are expressed in the late stage of infection, i.e., at least 16-24 hours post-infection

(Leu et al. 2009, Sanchez-Paz 2010).

There is a desperate need for measures to control and prevent outbreaks of

WSSV. In addition to SPF stocks and farm biosecurity, WSSV resistance breeding programs have been under development for decades with limited success (Lightner et al. 2012). Research efforts on developing vaccines to mitigate WSD have employed recombinant proteins, DNA- and RNAi-based strategies, all primarily targeting structural genes (Westenberg et al. 2005b, Vaseeharan 2006, Rout et al.

2007, Wu et al. 2007b, Attasart et al. 2009). We postulated that an RNAi strategy that targets immediate early or early genes may be more effective at shutting down virus replication earlier in the infection process and thereby reduce disease burden.

This was tested and described in Chapter 4.

Shrimp immunity

Immunity is a state in which the body is protected from pathogens and other toxic substances as a result of the coordinated effects of cells and molecules that 12 are involved in innate or adaptive immune responses. Generally speaking, innate immunity manifests as the first-line of defense mechanisms that acts immediately

(i.e., within minutes or hours) to destroy invading organisms in a non-specific manner. Innate immune responses do not rely on induction and expansion of antigen-specific lymphocytes as do adaptive responses, so do not directly generate canonical immunological memory, although innate immunity does impact adaptive immunity in organisms that have both innate and adaptive immunity (Maclachlan et al. 2011, Tizard 2013).

The innate defense system of arthopods, including crustaceans, is characterized as strictly innate in nature, with humoral and cellular effector arms.

Humoral responses include clotting and prophenoloxidase cascades, and antimicrobial . Shrimp have an open circulatory system that consists of plasma called hemolymph and blood cells called hemocytes. Hemocytes play an important role in the immune response of shrimp against infection (Jiravanichpaisal et al. 2006). Cellular immune responses include apoptosis, encapsulation, phagocytosis and melanization (Little et al. 2005, Tassanakajon et al. 2013). Shrimp utilize both humoral and cellular responses to combat pathogen infection (Bachere et al. 2004, Wang & Zhang 2008).

Innate immune responses in shrimp are initiated when pathogen-associated molecular patterns (PAMP) are recognized by pattern recognition receptors (PRR).

The interaction of these molecules triggers a number of complex responses that involves the production of antimicrobial peptides, including the enzymes and inhibitors that control the activity of these peptides (Tassanakajon et al. 2013). It is 13 generally believed that invertebrates lack a complex adaptive immune response due to the fact that there is no evidence of antigen-specific humoral compound production akin to antibodies in (Liu et al. 2009). Thus, there are no T cells, B cells, B cell receptors, or major histocompatibility complex molecules (MHC) in invertebrate immune systems (Little et al. 2005). An adaptive immune system is associated with immune memory, which will significantly enhance the ability to mitigate disease during the second and subsequent infections (Tizard 2013). There are data from shrimp and other that demonstrate that invertebrates possess some form of memory immune response but the precise underlying mechanisms are poorly understood (Little et al. 2005). Recently, evidence of a mechanism for a more specific immune response capacity has been discovered in crustaceans known as Down syndrome cell adhesion molecule (Dscam). Dscam, a member of the immunoglobulin super family (IgSF), that was identified and characterized in crustaceans including P. vannmei, P. monodon, Pacifastacus leniusculus and Eriocheir sinensis (Chinese mitten ) (Chou et al. 2009, Chou et al. 2011, Watthanasurorot et al. 2011, Wang et al. 2013). The Dscam molecule has hyper-variable region that can facilitate phagocytosis of the specific invading pathogens through alternative splicing by recognizing pathogens including bacteria

Escherichia coli, Streptococcus aureus, Vibrio alginolyticus and WSSV

(Watthanasurorot et al. 2011, Lin et al. 2013, Wang et al. 2013). Due to the extreme diversity of Dscam receptors, this could be a mechanism by which unique and specific responses are elicited to pathogens that have a tremendous impact on crustacean fisheries. 14

In innate immune system, Type I interferon (IFN) is a first line antiviral immune response. Interferons are glycoproteins secreted by virus-exposed cells that protect neighboring cells against viral infection. IFN-inducible genes include RNA dependent protein kinase R (PKR), the Matrix protein (Mx), oligoadenylate synthetase and IFN molecules themselves (Tizard 2013). Double- stranded RNA, which is not normally produced in a cell, but occurs as function of virus replication and formation of extensive secondary structure in virus RNA sequences, is a potent inducer of an IFN response (Robalino et al. 2004, Tizard

2013). The dsRNA is recognized by Toll-like receptor 3 (TLR3), PKR and retinoic acid-inducible gene I (RIG-I) (Tizard 2013). The consequences of dsRNA recognition include activation of the IFN system, initiation of apoptosis and inhibition of cellular protein synthesis to shut off virion production (Robalino et al. 2005). A Toll receptor has been identified in L. vannamei (lToll) (Yang et al. 2007), however the data show that lToll is not involved in dsRNA-induced antiviral immunity (Labreuche et al.

2009). Because invertebrates lack genes that are homologous to IFNs or the major effector molecules such as protein kinase R, it has long been assumed that invertebrates lack the ability to mount an antiviral response. However, with the discovery of the RNA interference (RNAi) in the , Caenorhabditis elegans

(Fire et al. 1998), it became clear that cells detect and internalize double-stranded

RNA (dsRNA) and then initiate intracellular gene-silencing events. Since then several lines of evidence have revealed that RNAi is, in fact, a robust antiviral immune response in invertebrates including fruit flies, mosquitoes, bees and shrimp

(Olson et al. 2008, Hunter et al. 2010, Bartholomay et al. 2012). 15

RNA interference mechanism and function

RNA interference or RNAi is a biological process that mediates gene silencing in a sequence-specific manner. The mechanism and the function of the biochemical molecules and events that are fundamental to a functional RNAi pathway have been extensively studied during the last decade. The phenomenon was first observed in the 1990s in flowering plants during an attempt to produce more purple coloration in pansies by introducing more transgene-encoding enzyme for flower pigmentation.

Surprisingly, the flowers turned white or variegated rather than more purple as expected. This phenomenon was called co-suppression of gene expression (Napoli et al. 1990) and later was reported as posttranscriptional gene silencing (PTGS).

Then, a related phenomenon was also observed in the , Neurospora crassa

(Pallavi et al. 2012). Andrew Fire and Craig Mello published their breakthrough study on the mechanism of RNAi in Caenorhabditis elegans in 1998 which showed that dsRNA that is introduced into worms can interfere with gene expression (Fire et al.

1998). This discovery won the Nobel Prize in Physiology or Medicine in 2006.

The RNAi pathway is initiated by long double-stranded RNA (dsRNA) that can be produced during , by integrated transposons, by the cell itself in the form of non-coding microRNA (miRNA), and by exogenous introduction of in vitro synthesized dsRNA. Following recognition, dsRNAs are processed into short dsRNA, siRNA or miRNA by Dicer enzymes. Dicer is an ATP-dependent RNase III- family enzyme that is conserved in plants, fungi, and animals including and invertebrates (Fire et al. 1998, Kim & Rossi 2008, Vijayendran et al. 2013).

R2D2 has two dsRNA binding domains that link the initiator and effector stages of 16 the RNAi pathway. Its function is to load Dicer/siRNA into RNA-induced silencing complex (RISC) complex. The passenger strand (sense strand) of the siRNA duplex must separate from the guide strand (anti-sense strand) before loading to RISC complex. Then, the passenger strand is degraded and the guide strand remains associated with RISC complex. The RISC complex is composed of an Argonaute protein (Ago) that is an endonuclease capable of degrading the complementary sequences of target mRNA into 21-22 fragments. Therefore, this functions as a post- transcriptional gene silencing mechanism and inhibits protein synthesis. In particular systems such as plants, worms and fungi, an RNA-dependent RNA polymerase

(RdRP) plays a critical role in maintaining and amplifying the silencing signal through synthesis of secondary dsRNA trigger molecules. An RdRp has not been identified in the genomes of either flies or humans (Olson et al. 2008, Pallavi et al. 2012). In the fruit fly, Drosophila melanogaster, the spread of systemic RNAi is mediated by endocytosis but the effect lasts only 5 days (Saleh et al. 2009). In penaeid shrimp,

Sid-1 protein (systemic interference defective protein), a membrane channel protein that functions to mediate the spreading of systemic RNAi, was identified by

Labreuche et al., (2010). The RNAi machinery characterized in penaeid shrimp also includes Dicer 1, Dicer 2 (Su et al. 2008, Yao et al. 2010, Chen et al. 2011),

Argonaute 1 and Argonaute 2 (Unajak et al. 2006, Dechklar et al. 2008, Labreuche et al. 2010). The presence of these key RNAi pathway components further supports the existence of RNAi in penaeid shrimp.

Three major classes of small regulatory have been characterized including: microRNA (miRNA), small interfering RNA (siRNA), and PIWI-interacting 17

RNA (piRNA). The function of each small RNA is relatively distinct but related proteins are required for small RNA biogenesis and function in all pathways. All three of these small RNA classes are involved in host-virus interactions. Small RNA can range from 18-32 nt in size and have distinct origins and functions. siRNA (21-

22 nt) or miRNA (18-25 nt) can be differentiated from each other based on their biogenesis or function (Cook & Blelloch 2013). In general, the miRNA binds target mRNA to inhibit and siRNA are thought to bind target mRNA and direct cleavage of a nascent transcript. The function of miRNA does not require that the target has a perfect match to inhibit translation. In contrast to miRNA, siRNA only binds to the target mRNA with identical sequence, which results in the cleavage of the transcript. piRNA are slightly longer (25-32 nt) and do not require Dicer for processing. The complete mechanism and function of producing piRNA remains unclear but they seem to play a role in the cellular defense against transposable elements and in antiviral responses in mosquitoes (Morazzani et al. 2012,

Vijayendran et al. 2013).

miRNAs are encoded by specific miRNA genes as a short hairpin primary- microRNA (pri-miRNAs) approximately 60-75 nt in length in the nucleus and are first processed by the proteins Drosha and Pasha to pre-miRNA. Then pri-miRNAs are exported to cytoplasm by Exportin 5 and processed into miRNA by Dicer1 (He &

Hannon 2004). The function of miRNA is to regulate gene expression by suppressing the mobilization of transposons that is important for the growth and development of organisms (Nykanen et al. 2001). miRNAs maintain genome stability by controlling transposons which are DNA sequences that can move around the 18 genome and cause deleterious effects. Transposons operate by copying their DNA to RNA and then reverse-transcribing back to DNA; the RNA form is usually double- stranded so can be recognized and targeted by RNA interference (Pallavi et al.

2012). miRNAs also are involved in regulation of larval development in C. elegans, growth control and apoptosis in D. melanogaster and mediating host-virus interactions (Vijayendran et al. 2013). miRNAs have been recently identified in the

WSSV genome using miRNA microarray and northern blot analyses. WSSV encodes 40 distinct viral miRNAs and engages RNAi pathway components including

Dicer1 and Drosha (He & Zhang 2012). The study showed that WSSV-miR-66 and

WSSV-miR-68 are key to the establishment of WSSV infection (He et al. 2014). This information supports the hypothesis that miRNA are involved in virus infection in arthropods and play a role in host-virus interactions.

siRNAs have been the focus of shrimp research in the last decade because compelling evidence shows that RNAi functions in antiviral immunity. In other arthropods including mosquitoes and Drosophila, the end products of RNAi or virus- derived small interfering RNAs (vsRNAs) are present in cell culture and host tissues during infection (Bartholomay et al. 2012). In our laboratory, a lymphoid organ transcriptome analysis revealed that vsRNAs (mostly 21-22 nt) are generated during

IMNV infection (unpublished data). Secondly, suppression of the RNAi pathway results in widespread virus replication and increases pathogenicity of disease. For example, suppressing RNAi pathway proteins such as Ago2, Dicer2 genes causes increasing viral titer of O’nyong-nyong virus and Sindbis virus in mosquitoes (Keene et al. 2004, Campbell et al. 2008, Bartholomay et al. 2012). In our laboratory, we 19 designed a study to suppress the RNAi pathway by targeting Ago2. Shrimp that received dsRNA to Ago2 showed mortality within 7 days post-injection without any virus infection (unpublished data). Labreuche et al. (2010) also reported this observation. Thus, the RNAi components may play a vital role in other physiological processes, and further research is needed to dissect the roles of these proteins.

Finally, several -specific virus genomes encode RNAi suppressors to counteract the host antiviral response. For instance, Cricket Paralysis (CrPV),

Drosophila C virus encoded 1A protein (Nayak et al. 2010) and Flock House virus

(FHV) encoded B2 protein (Chao et al. 2005). IMNV also may encode an RNAi suppressor because ORF1 encodes a protein with a dsRNA binding domain (Poulos et al. 2006, Bartholomay et al. 2012).

Augmenting RNAi to affect disease control

RNA interference (RNAi) technology was declared the “Scientific

Breakthrough of the Year” in 2002 in the journal Science because this is a tool that can be used to breakdown mRNA and thereby inhibit protein expression (Couzin

2002). RNAi provides a new and exciting tool for drug discovery with which putative drug targets can be suppressed and the biological effects observed (Leung &

Whittaker 2005). There are a number of pharmaceutical and biotechnology companies developing RNAi-based pharmaceuticals and vaccines. RNAi-based vaccines are arguably safer as compared to modified live vaccines (MLV) that have been used frequently to control viral diseases. MLV have the potential to mutate to a virulent form and could exacerbate disease in immunosuppressed animals

(Maclachlan et al. 2011). In contrast, RNAi-based vaccines synthesized from nucleic 20 acid (dsRNA) are more stable and can’t revert to a virulent form. In humans, RNAi technology has been used in clinical trials and provided promising results for treating several diseases such as ocular and retinal disorders, cancer, kidney disorders and hypercholesteremia (Davidson & McCray 2011).

RNAi was first studied in L. vannamei by Robalino and collaborators

(Robalino et al. 2004, Robalino et al. 2005). The discovery of this mechanism in shrimp was very exciting to shrimp researchers and the aquaculture industry because it provided a new potential tool to control virus disease outbreaks. In an

IMNV study conducted in our laboratory, a specific dsRNA corresponding to nucleotides from 95-475 (dsRNA95-475), at a dose of 2 µg/shrimp, provided 81.67% survival protection against IMNV challenge at 2 days post-vaccination. Those surviving animals were re-challenged with 100X higher dose of IMNV at 52 days post-initial challenge. The study was followed for an additional 40 days and the animals showed 94% survival (16/17 animals) (Loy et al. 2012). This evidence demonstrates some form of long-lasting protection. In a WSSV challenge study, shrimp were vaccinated with VP28 and VP26dsRNA and the survivors were re- challenged 3 times every 10 days post-initial infection with WSSV. The results showed 50% and 31% protection in VP28 and VP26dsRNA vaccinated groups, respectively (Mejía-Ruíz et al. 2011). Thus, these data may be the beginning of a paradigm shift in the understanding of the immune capacity of penaeid shrimp, i.e., shrimp have some form of memory immune response. This probably is not similar to memory mechanisms in vertebrates, with production of a clonal expansion of memory cells in to response to a specific pathogen infection. Further research is 21 needed to understand the underlying mechanisms of memory immune responses in invertebrates. The sequence-specific dsRNAs for WSSV and IMNV against viral diseases, which were generated in the studies described within this dissertation, could be used to interrogate the underlying mechanism of this long-term immunity.

The development of an RNAi-based vaccine for use in the crustacean fisheries industry would offer several advantages. Firstly, the nature of dsRNA molecules is that they are more stable and resistant to nuclease degradation compared to single-stranded (antisense) RNA molecules that also can be antiviral when introduced into cells (Bertrand et al. 2002). Secondly, RNAi-based vaccines can be produced in a cost-effective manner. For example, a dsRNA-based product was developed to successfully to control Israeli Acute Paralysis virus (IAPV), one of the major causes of Colony Collapse Disorder (CCD) in the honeybee, Apis mellifera. The dsRNA product, Remebee-I has been applied to honey beehives by mixing with sugar water and feeding to honey bee colonies. The results showed that there was higher honeybee production in groups that were treated with Remebee-I.

This was the first large-scale use of RNAi for disease control in insects (Hunter et al.

2010). Cost-effective and the large-scale production of dsRNA have been developed using transformed RNase-deficient Escherichia coli (E. coli). Production in an E. coli based fermentation system reduces the cost reduction to one-fourth that of a commercial in vitro transcription kit (Saksmerprome et al. 2009). That said, the cost of production of an RNAi-based vaccine may be more expensive depending on the delivery system of choice. The dose of dsRNA and the frequency of application is another factor that will affect the cost production of RNAi-based vaccines (Leung & 22

Whittaker 2005). However, our data both in IMNV and WSSV infection showed that targeting an early event of infection can provide the highest efficacy (using small dose of dsRNA) to inhibit virus replication (Loy et al. 2013 and Chapter 4) , would greatly reduce the amount of dsRNA needed and thus the cost of production of a dsRNA vaccine.

Oral vaccine delivery systems

A delivery system that is compatible with organisms in an aquatic habitat is the ultimate goal and primary hurdle for implementation of an RNAi-based vaccine for crustacean fisheries. Delivery systems for human vaccines include virus vector systems such as , , lentiviruses and adeno-associated viruses (AADV) and non-viral systems including, for example, -based systems

(Davidson & McCray 2011). In arthropods, RNAi technology is often implemented using injection because the specific dose and target site can be controlled; this method is impractical in field settings. In the case of the Remebee-I product, dsRNA is fed to the bees by mixing dsRNA with sugar water (Hunter et al. 2010). An RNAi- based vaccine for aquatic animals presents an additional challenge in that the vaccine is potentially significantly diluted in water. The ideal vaccine for shrimp would be an oral application that is user-friendly for hatchery workers or farmers to apply. The development of an RNAi-based oral vaccine has shown some promising results; for example, nanoparticles made of chitosan mixed with dsRNAVP28 and top-coated onto feed. After feeding the treated feed to the shrimp, 37% survival was observed in WSSV-infected animals in comparison to the control group (Sarathi et al. 2008a). In the same study, inactivated DE3 E. coli expressing dsRNAVP28 was 23 top-coated onto feed and provided 68% protection compared to 0% in untreated animals. Another study showed successful incorporation of bacteria-expressed dsRNA targeting Laem-Singh Virus (LSNV). LSNV is associated with monodon slow growth virus syndrome (MSGS) in Black tiger shrimp (P. monodon). The results showed the mean body weight (MBW) of shrimp that received feed with dsRNA for 9 weeks were higher and increased by 15% of biomass, which would account for

$4800 more yield per pond (Saksmerprome et al. 2013b). These results are promising; however consistent technology for delivery of vaccines through feed should be developed to reduce the requisite dose, and improve the stability and efficacy of the antiviral effect in hatchery or pond settings.

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33

CHAPTER 2: CHARACTERIZATION OF NEWLY REVEALED

SEQUENCES IN THE INFECTIOUS MYONECROSIS VIRUS GENOME

IN LITOPENAEUS VANNAMEI

A paper to be submitted to Journal of General Virology

Duan S. Loy1, Sijun Liu2, Mark A. Mogler3, J. Dustin Loy4, Bradley J. Blitvich1,

Lyric C. Bartholomay2

1Department of Veterinary Microbiology and Preventive Medicine, Iowa State University, Ames IA 50011, USA 2Department of Entomology, Iowa State University, Ames IA 50011, USA 3Harrisvaccines, Inc., Ames IA, 50010, USA 4School of Veterinary Medicine and Biomedical Sciences, University of Nebraska- Lincoln, Lincoln NE, 68583, USA

Abstract

Infectious myonecrosis virus (IMNV) causes significant economic losses in farmed shrimp, where associated mortality in ponds can reach 70%. To explore host-pathogen interactions, a next generation sequencing approach using lymphoid organ tissue from IMNV-infected shrimp, Litopenaeus vannamei, was conducted.

Preliminary deep sequencing assembly showed that there were at least an additional 639 base pairs (bp) at the 5’ terminus and 23 bp at the 3’ terminus as compared to the original description of the genome (7561 bp). Reverse transcriptase

PCR and Northern blot analysis confirmed the presence of novel sequence. Using 5’

Rapid Amplification of cDNA Ends (RACE), 4 additional nucleotides were discovered; 3’ RACE confirmed the presence of 22, rather than 23 bp of additional 34 sequence. Based on these data, the IMNV genome is 8226 bp in length1. Double- stranded RNA was used to suppress expression of the newly revealed sequence at the 5’ end of the IMNV genome in IMNV-infected L. vannamei. Suppression of a 376 bp region of the 5’UTR did not improve the survival rate of infected shrimp; in contrast, suppression of a 381 bp sequence that encompasses a portion of the coding region for ORF1 improved survival to 82.2% as compared to 2.2% survival in positive control animals. These studies reveal the critical nature of the new sequence to produce high titer infection, and associated disease and mortality, in infected shrimp.

1 The GenBank accession number associated with this sequence is KF836757 35

Introduction

A large number of pathogens threaten the shrimp aquaculture industry with a majority of these being viral in etiology. The total economic impact is significant, with losses in the billions of dollars (Lightner et al. 2012). The emergence of novel pathogens is inevitable because of the combined effects of high intensity farming, and a globalized aquaculture industry that exports and imports either live or unprocessed products and thereby rapidly disseminates pathogens worldwide. The emergence of Infectious myonecrosis virus (IMNV) is thought to have occurred via live animal movement from an endemic area to another intensive farming country

(Senapin et al. 2007). Clinically, the gross signs of IMNV-infected animals are lethargy and the appearance of white muscle starting from the tail to abdominal muscle followed by mortality in ponds that can reach 70% (Senapin et al. 2007).

IMNV was first reported in Brazil in 2003 after a disease outbreak occurred in 2002

(Poulos et al. 2006) and this was followed by an outbreak in Indonesia in 2006 which caused devastating economic losses (Senapin et al. 2007).

IMNV is non-segmented, doubled-stranded RNA virus with icosahedral symmetry and complex protrusions at the five-fold axis that are thought to be involved in virus entry (Tang et al. 2008). IMNV is classified in the family Totiviridae based on its genome organization and significant amino acid identity to Giardia

Lamblia virus (GLV) in the RNA-dependent RNA polymerase (RdRp) coding region

(Poulos et al. 2006). The IMNV genome encodes two open reading frames (ORFs).

ORF 1 is translated into a polyprotein that contains a predicted dsRNA binding domain and a major capsid protein (MCP) while ORF 2 encodes the RdRp. Another 36 protein (a MCP-RdRp fusion product) is generated as a result of a -1 frameshift within ORF1 (Poulos et al. 2006, Nibert 2007).

The IMNV genome was initially reported to be 7561 bp in length (Accession no. EF061744.2) (Senapin et al. 2007). Although the reported length of the IMNV genome was similar to that of other viruses in the family Totiviridae (5.0 - 7.0 kbp), the published IMNV sequence did not appear to have a predicted internal ribosomal entry site (IRES) in its 5’ (5’ UTR) that is characteristic of viruses in this family (Garlapati & Wang 2005). In addition, protein bands produced in purified IMNV protein preparations did not match the size of the proteins produced as a result of cleavage of the ORF1 polypeptide.

To explore shrimp-virus interactions, we undertook a next generation sequencing (NGS) approach using lymphoid organ (LO) tissues from L. vannamei.

The LO is located at the anterior end of the shrimp on the rostral-ventral side of hepatopancreas. One function of the LO is to remove pathogens from the hemolymph during circulation. Furthermore, the LO is a common site for virus replication and associated pathology (e.g., spheroids) to occur (Assavalapsakul et al. 2006, Escobedo-Bonilla et al. 2007). LO tissues express a significant amount of immune-related genes in normal and infected shrimp (Pongsomboon et al. 2008). It follows that significant transcriptional changes to this tissue would occur during active viral infection. We employed NGS to explore these transcriptional changes and to examine IMNV RNA production and breakdown during active IMNV infection.

Surprisingly, we identified virus genome sequence that extends significantly beyond 37 the original description. This additional sequence encodes a genomic element that is key to virus replication.

Materials and Methods

Shrimp culture

Specific Pathogen Free (SPF) L. vannamei were acquired from Shrimp

Improvement Systems (Islamorada, FL, USA) and maintained in a specific pathogen free colony in a biosecure facility at Iowa State University. Post-larvae were placed into one ton fiberglass tanks filled with artificial sea water. Water conditions were maintained at 25-27 °C, 30-35 parts per thousand (ppt) salinity, with constant airstone aeration. Each tank was maintained with a carbon filter and an oyster shell airlift biofilter. Water quality was measured according to ammonia and nitrite levels

(Nitriver3, Hach Company, Loveland, CO), and temperature and salinity were measured regularly. Shrimp were fed a commercial growout diet and cultured to different sizes for these studies.

IMNV (Indonesia) infected-lymphoid organs

The IMNV-infected tissues used for this study were collected during the IMNV outbreak in Indonesia. The virus inoculum was prepared as previously described

(Loy et al. 2012). Six shrimp (15-20 grams in size) were infected by intramuscular injection with IMNV (strain Indonesia). Virus stock was prepared and diluted in 1:100

2% NaCl (~2.11 x103 viral copy numbers µl -1 RNA) and filter-sterilized. Control shrimp were injected with 2% NaCl (viral diluent), as reported by Loy et al. 2012.

38

RNA sample and library preparation

LO were collected from IMNV-infected animals at 5 days post-infection or from normal control shrimp at the same time. Gross organ characteristics were confirmed via light microscope on a dissecting microscope. All LO were preserved in

RNAlater RNA Stabilization Reagent (Qiagen, Germantown, Maryland, USA) and stored at -80 °C. Samples were then transferred to 750 µl TRIzol LS (Invitrogen,

Carlsbad, California, USA) and processed for RNA extraction. Residual DNA was removed from the extracted RNA using the TURBO DNA-freeTM kit (Ambion, Foster

City, California, USA). RNA integrity was determination using an Agilent 2100 bioanalyzer. Total RNA from 5-6 individual animals was pooled and diluted in nuclease free water to achieve a concentration of 0.1 µg ml-1. cDNA libraries were prepared from pooled RNA samples using the TruSeq RNA Sample Prep Kit

(Illumina, San Diego, California, USA) following the manufacturer’s instructions.

Illumina sequencing, sequence assembly and sequencing data processing

cDNA libraries were subjected to 50 bp (single end) sequencing using the

HiSeq2000 platform (Illumina). Raw sequencing reads were obtained and trimmed to remove adaptor sequences, empty reads, and low quality sequences using the

FASTA-X toolkit (http://hannonlab.cshl.edu/fastx_toolkit/). These reads were assembled using Velvet (1.2.08) (Zerbino & Birney 2008). The performance of

Velvet assemblies is determined by hash length (k) and coverage cutoff (c). An iterative process was used to select the optimum k and c values to produce the best assembly (Liu et al. 2011). The parameters used for assembly of contigs were k=31 and multiple c (c=2~10). To identify contigs derived from IMNV sequences, the 39 resulting contig sets (>=100nt) were aligned against IMNV genomic sequences by running the local Basic Local Alignment Search Tool (BLAST) (Altschul et al. 1990).

To determine IMNV sequencing coverage, transcriptomic reads were clustered using the FASTA-X toolkit, and clustered reads were subsequently mapped to the IMNV genome using a custom-designed Perl program. The Perl script was designed to only permit mapping once to the reference sequence and n mismatched residues were allowed. The numbers of reads that mapped to each nucleotide position was used to calculate coverage (fold) of each nucleotide.

A biological sequence alignment editor (BioEdit v7.1.11)

(http://www.mbio.ncsu.edu/bioedit/page2.html) was used as a platform for basic sequence manipulations of IMNV (sequence alignment, protein translation, ORF prediction, etc). ClustalW2 was used for multiple sequences alignment (Larkin et al.

2007).

Identification of sequences at the 5’ and 3’ends of the IMNV genome by reverse-transcriptase PCR

Oligonucleotide primers were designed to amplify novel sequences at both the 5’ and 3’ ends of the IMNV genome (Table 1). Total RNA was extracted from shrimp inoculated with the Indonesia or Brazil strains of IMNV using TRIzol LS

(Invitrogen). One-step reverse transcriptase PCR (one-step RT-PCR kit, Qiagen) was performed using the following amplification parameters: 50 °C for 30 min, 95 °C for 15 min and 35 cycles of 95 °C for 15 s, 55 °C for 30 s and 72 °C for 45 s, followed by a final elongation at 72 °C for 5 min. RT-PCR products were cloned into the pCR®2.1 vector (TOPO TA Cloning® kit, Invitrogen) and transformed into 40 competent E. coli cells (OneShot®TOP10, Invitrogen). Cells were grown on Luria-

Bertani agar containing kanamycin (50 µg/ml). A subset of colonies was screened for the presence of the desired insert by PCR, and these inserts were then purified and sequenced with vector-specific primers (M13F and M13R) using a 3730x1 DNA sequencer (Applied Biosystems, Foster City, California, USA) at the ISU DNA facility.

Rapid Amplification cDNA Ends (RACE) PCR

Reverse transcription

Total RNA (1 µg) was mixed with 500µM of each dNTP and 25 µM of the appropriate reverse primer, heated at 70 °C for 10 min then chilled one ice for 1 min.

Heat-denatured RNA was reversed transcribed at 50 °C for 50 min using 0.2 units of superscript III reverse transcriptase (Invitrogen) in 1x first strand buffer (50 mM Tris-

HCl [pH 8.3], 75 mM KCl, 0.6 mM MgCl2), 0.02 M dithiothreitol (DTT) then incubated at 70 °C for 15 min. The resulting cDNAs were purified using PCR purification kit

(Purelink Quick PCR Purification Kit, Invitrogen).

Polymerase Chain Reaction

PCR amplifications were performed using 4 µl of cDNA, 1 unit of Platinum®

Taq DNA Polymerase High Fidelity (Invitrogen), 20 µM of each primer, 0.2 mM of each dNTP and 2 mM MgSO4, in 1 x High Fidelity PCR Buffer (600 mM Tris-SO4 [pH

8.9], 180 mM Ammonium Sulfate) using the following reaction conditions: 94 °C for 4 min, 35 cycles of 94 °C for 30 s, 50 °C for 45 s and 72 °C for 90 s followed by a final extension at 72 °C for 8 min. PCR products were examined by 1% agarose gel electrophoresis. 41

5’ RACE PCR

To identify the nucleotide sequences at the 5’ end of the IMNV genome, 5’

RACE was conducted. Total RNA was reverse transcribed using the appropriate primer (452R for IMNV [Indonesia] and 163R for IMNV [Brazil]), and the resulting cDNAs were purified using the Purelink Quick PCR Purification Kit (Invitrogen).

Oligo(dC) tails were added to the 3’ end of 2 µg of purified cDNA using 15 units of terminal deoxynucleotidyl transferase (Invitrogen), 1mM dCTP and 50 µg/ml of bovine serum albumin in 1x tailing buffer (100 mM potassium cacodylate [pH 7.2], 2 mM CoCl2, 0.2 mM DTT). Tailing reactions were performed at 37° C for 30 min and heat inactivated at 65 °C for 10 min. C-tailed products were purified then PCR amplified using a consensus forward primer specific to the C-tailed termini (C tail F) and one of several reverse primers specific to the IMNV cDNA sequences (428R for

IMNV [Indonesia] and 243R for IMNV [Brazil]). The resulting products were cloned and transformed into competent E. coli cells. Selected colonies were screened by

PCR using both vector-specific primers (M13F and M13R) and IMNV-specific primers (273F, 428R for IMNV [Indonesia] and 51F, 243R for IMNV [Brazil]). Select amplicons were purified (QIAquick PCR Purification Kit, Qiagen) and sequenced with an IMNV-specific reverse primer (374R for IMNV [Indonesia] and 304R for

IMNV [Brazil]).

3’ RACE PCR

To identify the nucleotide sequences at the 3’ end of the IMNV genome, synthetic poly (A) tails were added to 10 µg of total RNA using 6 units of poly(A) polymerase A (Ambion) in 1 x reaction buffer (40 mM Tris-HCl, pH 8.0, 10 mM 42

MgCl2, 2.5 mM MnCl2, 250 mM NaCl, 50 µg of bovine serum albumin/mL, and 1 mM

ATP). Tailing reactions were incubated at 37 °C for 1 hr and then placed in -20 °C for 2 min to inactivate the enzyme. Reaction products were purified by ethanol precipitation and poly (A)-tail enriched RNAs were reverse transcribed using a reverse primer specific to the poly(A) tail (3’ RT). Resulting cDNAs were then PCR amplified using the appropriate primers (IMNV7852F and 3’Rev). PCR products from the 3’ RACE were also cloned and screened with gene specific primers (IMNV7852F and 3’Rev) and plasmid primers (M13F and M13R) and then sequenced with 7945F for both IMNV strains.

Doubled-stranded RNA synthesis

Two dsRNAs, each approximately 381 bp in length, and corresponding to the newly identified IMNV genomic sequence were generated in the 639 bp of newly discovered sequence at the 5’ end. Primers for dsRNA synthesis were designed to amplify from position 26 to 312 nt (IMNV26T7F and IMNV401T7R) and 233 to 613 nt

(IMNV233T7F and IMNV613T7R). The aforementioned dsRNAs are nearly identical in size to dsRNA95-475 (with the nt range dictated by position on the original IMNV genome sequence) which has proven protective effect in suppressing IMNV replication. dsRNA95-475 was generated according to the parameters described in a previous report (Loy et al. 2012). Briefly, a PCR was performed using primers that contained the T7 promoter sequence at the 5’ end (Table1). Complementary

(cDNAs) were generated following the manufacturer’s instructions for ThermoScript

TM RT (Invitrogen) using total RNA extracted from IMNV-infected shrimp and forward primer IMNV26F (Table1). PCR was performed using PCR master mix (PuReTaq 43

Ready-To-Go PCR Beads, GE Healthcare, Pittsburgh, Pennsylvania, USA), cDNA template, and the primers containing T7 extensions (Table1) that were designed to specific regions of the newly revealed IMNV genome sequence. The reaction conditions were 95 °C for 4 min, followed by 35 cycles of 95 °C for 30 s, 55 °C for 30 s, 72 °C for 45 s and then a final elongation of 72 °C for 7 min. The resulting PCR products were confirmed to have a molecular weight approximately 381 bp by agarose gel electrophoresis. Nucleotides, enzymes and buffer were removed using the QIAquick PCR purification kit (Qiagen). PCR products were concentrated by vacuum concentrator (DNA120 SpeedVac, Thermo Scientific, Waltham,

Massachusetts) and 1 µg of purified PCR product was added to an in vitro transcription reaction. The synthesis reaction was incubated at 37 °C for 12-16 hours. The dsRNA synthesis product was then incubated for 45 min in a buffer solution containing RNase and DNase I. Following incubation, dsRNA was purified using the provided purification columns. The formation of dsRNA was confirmed by

1% agarose gel electrophoresis, and quantified by spectrophotometry (Nano

Drop2000, Thermo Scientific). The dsRNAs were stored in -20 °C until use and were then diluted in RNase free water to reach the desired concentration.

IMNV (Brazil) viral isolation

IMNV (Brazil) was kindly provided by the Aquaculture Pathology laboratory at the University of Arizona. Virus inoculum was clarified directly from frozen infected samples as previously described (Loy et al. 2012). Shrimp (7-10 gram) were injected intramuscularly (100 µL) with 1:100 dilution of IMNV (Brazil) (~7.2 x102 viral copy numbers µl -1 RNA) (Loy et al. 2012). Clinical signs of lethargy and white muscle 44 were observed at 3 days post-infection. Moribund shrimp with gross lesions were collected for RNA extraction using the Qiagen RNeasy Minikit according to the manufacturer’s instructions. RNA from shrimp infected with IMNV (Brazil) was tested by RT-PCR and molecular weight was confirmed on a 1% agarose gel stained with ethidium bromide.

Determination of novel dsRNA efficacy in vivo

Shrimp (3-5 gram) were divided into 5 groups of 10 shrimp with 3 replicates in each group (total 30 shrimp/group) and 10 shrimp for negative control. Tanks were maintained at 29-30 °C. Shrimp were injected intramuscularly with 5 µg/shrimp (100

µl) of dsRNA95-475, dsRNA26, dsRNA233 and dseGFP or 2% NaCl, and at 48 hours post-injection (p.i.) with 2% NaCl (negative control group) or IMNV 1:100 dilution of IMNV (Indonesia) that reach 95-100% mortality within two weeks (~2.11 x103 viral copy numbers µl -1 RNA) (Loy et al. 2012). Shrimp were fed and mortality observed twice per day. These studies were in triplicate. Tissue samples from moribund and dead shrimp were stored at -80 °C for further quantification of virus copy number.

Quantitative Reverse-Transcription PCR

A qRT-PCR assay for virus load was adapted from a previously described method (Andrade et al. 2007). RNA was extracted using the RNeasy Minikit

(Qiagen) and 3 µl of sample was added to each reaction of One Step RT-PCR

Master Mix (Qiagen). The one step RT-PCR reaction mix contained 5 x Master Mix

(5 µl), Enzyme mix (1 µl), 10 mM dNTP (1 µl), 20 µM of IMNV412F (0.3 µl), 20 µM

IMNV545R (0.3 µl) and 10 µM IMNV probe (0.3 µl) and RNase-free water (14.1 µl) 45 for a total volume of 25 µl. The qRT-PCR was performed with conditions from a previous report (Loy et al. 2012). Each reaction was run in duplicate on a BioRad

CFX96 Real Time PCR Detection System using a standard as described previously

(Andrade et al. 2007, Loy et al. 2012).

IMNV purification

IMNV was purified using a combination of methods adapted from Mello et al.

(2011) and Poulos et al. (2006). Briefly, one hundred 7-10 gram shrimp were infected with 1:100 dilution of IMNV (Indonesia) (Loy et al. 2012). Moribund shrimp were collected during a 2 week course of infection to obtain approximately 500 g of

IMNV tissue from acutely infected animals. Heads were removed to reduce the amount of lipid input into the virus purification process. Infected tissues were placed in 0.1 M PBS buffer, pH 7.5 plus 0.5% of Na2SO (w/v) at a ratio of 1:3 and homogenized in a blender. Additionally, 10% chloroform (v/v) was added into the homogenate and was then centrifuged at 8000 × g for 15 min to clarify the suspension. Following the first centrifugation, the supernatant was removed and centrifuged again to remove tissue debris. The supernatant was collected and precipitated by adding 10% polyethylene glycol (w/v) and 4% NaCl (w/v), stirred at

4 °C for 1 h, and centrifuged at 8000 × g, at 4 °C for 20 min (Mello et al. 2011). The pellet was then resuspended and loaded onto a 10-40% sucrose gradient and centrifuged at 70,000 × g for 2 h at 4 °C. A pellet was observed in the bottom of ultracentrifuge tube at the 40% sucrose gradient. The virus pellet was removed from the ultracentrifuge tube and diluted in the same buffer to be stored at -80°C. The presence of virus in the pellet was confirmed by RNA extraction and RT-PCR. 46

Purified virus was also injected intramuscularly into 7-10 gram shrimp to confirm the presence and viability of virus in the preparation.

Northern blot analysis

Northern blot analysis was performed using methods adapted from Current

Protocols in Molecular Biology (Brown et al. 2001). Briefly, total RNA was extracted from purified IMNV using Trizol LS (Invitrogen) and 5 µg was resolved on a 1% agarose glyoxal gel (NorthernMax-Gly, Life technologies). The gel was observed for presence of RNA via ethidium bromide staining in a UV light box prior to transfer.

Then RNA was nicked with 500 kilojoules UV light and transferred to a positively charged membrane (Whatman® Nytran™ Supercharge TurboBlotter, Sigma Aldrich) using a passive downhill transfer technique for 1 hour. Membranes were UV cross- linked at 1200 kilojoules and hybridized overnight ULTRAhyb® Ultrasensitive

Hybridization Buffer (Ambion) at 68 °C with Biotinylated RNA probes. Biotinylated

RNA probes were generated using BrightStar® Psoralen-Biotin Nonisotopic Labeling

Kit (Ambion) that were designed target to the original sequence of IMNV including 95 sense and 475 antisense probes and to the novel sequences at the 233 sense and

613 antisense probes (Table 1). PCR templates with T7 sequence at the 5’-ends were used to generate RNA in vitro transcripts (T7 RNA polymerase, Promega).

RNA transcripts were then labeled with biotin according to the manufacturer’s instruction to create biotinylated probes (BrightStar Biotin-Psoralen Kit). The hybridized membranes were developed following the manufacturer’s instruction and the signal was detected by chemiluminescence using the BrightStar BioDetect Kit

(Ambion). 47

Statistical analysis

Survival and qRT-PCR data were analyzed using one-way ANOVA followed by Tukey’s multiple comparison test (p<0.05) using JMP pro 10 software (SAS

Institute Inc.). The survival data and viral copy number were expressed as the mean

± standard error (SEM). Mean survival data were tested at the termination of experiment on day 14 post-infection and the mean log viral copy number was compared between groups.

Results and Discussion

IMNV sequencing analysis

A total of 43,567,120 trimmed reads of 50 bp were obtained from IMNV- infected shrimp and used for the sequence assembly. To identify IMNV sequence, each of the contig sets was aligned to the existing IMNV sequence (Accession No.

EF061744.1). Two IMNV contigs were identified, one that is 8044 bp in length and another one that is 8223 bp. Both contig8044 and contig8223 are significantly longer in size than that of the published IMNV genomic sequence (7561 bp). A sequence alignment revealed that the 663 bp of new, putative IMNV sequence is not within the limits of the known sequence but falls primarily upstream of the 5’end (639 bp) and, to a more limited extent, at the 3’end (23 bp) as shown in Fig. 1.

The new 5’-end and the known IMNV sequences have an equal depth of sequencing

IMNV sequences subjected to de novo assembly revealed possible novel sequences in the IMNV genome. To determine whether the new sequence fragments were represented with the same coverage as the published sequence, we 48 mapped the RNA-seq reads across the length of the genome (Fig. 2). Sequence coverage of the new sequences and the documented sequence are listed in Table 2.

Although there were significant variations of the sequencing depth at specific nucleotide locations in the IMNV genome, the average sequencing coverage between the new and known sequences was similar (Table 2), suggesting that the new sequences were equally represented in the cDNA library.

Confirmation of presence of sequence in IMNV Indonesia and Brazil

Deep sequencing results indicate that there are at least an additional 639 bp at the 5’-end and 23 bp at 3’-end when compared to the published IMNV genomic sequence. Several sets of primers were designed to confirm the presence of these novel IMNV sequences in Indonesia and Brazil isolates. Four sets of primers were used to confirm the presence of additional sequence at the 5’-end by RT-PCR including IMNV26F, IMNV312F, IMNV643R and IMNV1016R (Table 1). Previously published primers (95F, 475R) that amplify the known genome sequence were used as positive controls (Table 1). RT-PCR amplicons were subjected to gel electrophoresis and molecular weights were confirmed as predicted (Fig. 3, lane 1-

4). The largest amplicon (990 bp, lane 4, Fig. 3.) was cloned and sequenced and had 95% nucleotide identity compared to the published IMNV sequence in Genbank.

These four new primer pairs also were tested with IMNV (Brazil). RT-PCR results were identical to those for IMNV (Indonesia) (Fig. 3, lane 5-8) and sequenced PCR products had 100% and 99% identity to IMNV (Brazil) and IMNV (Indonesia), respectively.

Two reverse primers (IMNV8204R1, IMNV8204R2) were designed to confirm the present of additional sequence at the 3’ end of the IMNV genome (Table1). 49

Reverse primers successfully amplified the novel sequences in both IMNV strains (Fig. 4), confirming that these extra sequences at 3’ end are present, as revealed in the deep sequencing data. IMNV genome sized determination and Northern blot analysis

Gel electrophoresis was used to further confirm the size of the IMNV genome based on comparison to molecular weight markers. A distinct band of 8 kb was evident (Fig. 5). Total RNA from purified IMNV and non-infected shrimp were subjected to Northern analysis with RNA probes generated to the original sequences and novel sequences. A band of approximately 8-9 kb representing the IMNV genome was evident in IMNV RNA regardless of the RNA probes used, but not in uninfected animals (Fig. 6). These results further confirm that the novel sequence discovered by deep sequencing exists in the IMNV genome.

Analysis of the newly identified sequences at the 5’-end of the IMNV genome

The published IMNV sequence encodes two predicted ORFs. To clarify whether 639 bp of the new 5’- end sequences encodes any proteins, we translated sequence of Contig8223 in all six frames (Fig. 7). The translation results show that, like the published IMNV sequence, the new IMNV genome also encodes two ORFs.

The ORF2 (frame 3) is almost identical to the documented ORF2 sequences.

However, the predicted ORF1 (frame one) in Contg8223 is comprised of 1719 aa, which is 114 aa larger than the documented ORF1 (1605 aa). The predicted of the new ORF1 is at 342 bp upstream of the documented start codon. Fig. 8 shows the new N-terminal sequence and its alignment with the published IMNV

ORF1 sequence. 50

Examination of IMNV ORF sequence has revealed that the ORF1 contains 2

“2A-like” cleavage motifs (Nibert 2007), and another cleavage site that cleaves the C-terminus of ORF1 to release the MCP. These three cleavage motifs hydrolyze the ORF1 polyprotein into 4 products: the N-terminal peptide 1 (10 kDa), peptide 2 (31 kDa), predicted peptide 3 (36 kD) and the C-terminal 99 kDa MCP.

The ORF2 (frame 3) is translated by -1 ribosomal frame shifting within ORF1, and produces a 196 kDa MCP-RdRp fusion product (Poulos et al. 2006, Nibert 2007,

Tang et al. 2008). Hence, 5 peptide fragments should be expected. Our new N terminal sequence extends peptide 1 to a predicted 207 residue (23 kDa) protein.

Tang et al. showed that when purified IMNV virions were separated by SDS-PAGE and stained, 5 distinct protein bands could be observed. The 99 kDa and 196 kDa bands correspond to MCP and MCP/RdRp, respectively. The remaining 3 bands were between 25 kDa and 40 kDa (Tang et al. 2008). Therefore, it is likely that the

25 kD protein that was observed is peptide 1 and the other two bands are peptides 2 and 3.

5’ and 3’ RACE PCR confirm the presence of novel sequences in the IMNV genome

To confirm the presence of the IMNV sequences identified by NGS, 5’ and 3’

RACE were performed. Fourteen PCR products generated from the 5’ RACE of

IMNV (Indonesia) were sequenced and confirmed the presence of 639 bp identified by NGS, and revealed an additional 4 bp upstream. The additional 4 bp were present both in IMNV (Indonesia) and IMNV (Brazil). Thus, 5’ RACE PCR confirmed that the IMNV sequences previously published and available in Genbank 51

(NC_007915.2, AY570982.2, EF061744.1; 7561 bp) are truncated at the 5’ terminus by 643 bp.

Because the IMNV genome is not polyadenylated, a synthetic poly (A) tail was added to the 3’ end and then 3’ RACE experiments were performed to confirm the presence of additional sequence at the 3’ terminus of the genome. Ten PCR products were generated and sequenced using 3’ RACE on IMNV (Brazil). These experiments confirmed that an additional 22 bp are present at the distal 3’ end supporting the findings from the NGS analysis. We were unable to generate any positive clones by 3’ RACE with IMNV (Indonesia). However, we were able to amplify and sequence the 3’ end of IMNV (Indonesia) using (IMNV7852F) as the forward primer and (IMNV8210R2) as the reverse primer; the sequence of the latter primer corresponds to the distal 22 bp of the 3’ UTR of IMNV (Brazil) (Table1).

These data indicate that an additional 22 bp is present at the 3’ end of the genome of both virus strains. Therefore, using 5’ and 3’ RACE, we confirmed the presence of the nucleotide sequences identified by NGS at the 5’ and 3’ ends of the IMNV genome, and revealed the presence of another 4 bp at the distal 5’ end as shown in

Fig. 9. The full-length 8226 bp genomic sequence of IMNV (Indonesia) was submitted to Genbank (Accession No. KF836757).

Novel IMNV dsRNA showed protection against IMNV

In order to test the biological function of the novel sequences at the 5’ terminus of IMNV, two dsRNAs (designated dsRNA26 and dsRNA233) were designed and compared to dsRNA95-475, which targets the coding region of peptide

1. dsRNA95-475 was previously shown to provide 81.7% survival in animals infected 52 with IMNV at 30 days post-infection (Loy et al. 2012). Therefore, dsRNAs to the novel sequence were designed to have similar length to dsRNA95-475. Each shrimp was injected with 5 µg (100 µl) of either dsRNA26, dsRNA233, dsRNA95-475, dsRNAeGFP or 2% NaCl (sham-inoculation), held for 48 hours and then inoculated with IMNV (1:100). Negative control shrimp were also injected with 2% NaCl (virus diluent). In three separate trials, significant mortality was observed in shrimp exposed to dsRNA26, dsRNAeGFP or no with 3.3%, 2.2% and 2.2% mean survival, respectively. In contrast, shrimp that received dsRNA233 and dsRNA95-475 showed similar protection against IMNV with mean survival of 82.2% and 97.8%, respectively at 14 days post-infection as shown in Fig. 10. Mean survivals are significantly different from other groups (dsRNAeGFP, dsRNA26 and positive control) (p<0.001) using one-way ANOVA followed by Tukey’s multiple comparison test (JMP pro 10 software, SAS Institute Inc.). All negative control animals (non-challenged) survived during the full course of these experiments. Therefore, only dsRNA233, which encompasses ORF1 as compared to dsRNA26 which encompasses 5’ UTR sequence, showed protection against IMNV but the protection efficacy is similar to dsRNA95-475.

Novel IMNV dsRNA inhibition of IMNV replication

Viral load as measured by copy number was analyzed by qRT-PCR on muscle tissues from dead and survived animals in dsRNA26, dsRNAeGFP and positive control groups at 9-14 days post-infection and the dsRNA233 and dsRNA95-475 groups at 14 days post-infection. The results showed that dsRNA95-

475 and dsRNA233 can inhibit virus replication (mean 4.5 x 102 and 2.9 x 103 IMNV 53 copy numbers µl-1 RNA, respectively) and are significantly different from the positive control group (mean 1.1 x 105 IMNV copy numbers µl-1 RNA) (p= 0.013 and 0.016, respectively). By contrast, dsRNA26 and dsRNAeGFP-exposed shrimp did not show reduction of virus (mean 4.8 x 104 and 7.7 x 104 IMNV copy numbers µl-1 RNA, respectively) (Fig. 11). This is further evidence that dsRNA233 suppresses virus replication during IMNV infection.

Acknowledgments

The authors would like to thank Kara Jimenez and Zachary Flickinger for animal care and laboratory assistance, Shrimp Improvement System for supplying shrimp and Dr. Donald Lightner at the University of Arizona for providing IMNV

(Brazil). This work was financially supported by the National Science Foundation. 54

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Assavalapsakul, W., Smith, D. R. & Panyim, S. (2006). Identification and characterization of a Penaeus monodon lymphoid cell-expressed receptor for the yellow head virus. J Virol 80, 262-269.

Brown, T., Mackey, K. & Du, T. (2001). Analysis of RNA by Northern and Slot Blot Hybridization. In Current Protocols in Molecular Biology: John Wiley & Sons, Inc.

Escobedo-Bonilla, C. M., Wille, M., Alday Sanz, V., Sorgeloos, P., Pensaert, M. B. & Nauwynck, H. J. (2007). Pathogenesis of a Thai strain of white spot syndrome virus (WSSV) in juvenile, specific pathogen-free Litopenaeus vannamei. Dis Aquat Organ 74, 85-94.

Garlapati, S. & Wang, C. C. (2005). Structural elements in the 5'-untranslated region of transcript essential for internal entry site- mediated translation initiation. Eukaryot Cell 4, 742-754.

Larkin, M. A., Blackshields, G., Brown, N. P., Chenna, R., McGettigan, P. A., McWilliam, H., Valentin, F., Wallace, I. M., Wilm, A. & other authors (2007). Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947-2948.

Lightner, D. V., Redman, R. M., Pantoja, C. R., Tang, K. F., Noble, B. L., Schofield, P., Mohney, L. L., Nunan, L. M. & Navarro, S. A. (2012). Historic emergence, impact and current status of shrimp pathogens in the Americas. J Invertebr Pathol 110, 174-183.

Liu, S., Vijayendran, D. & Bonning, B. C. (2011). Next generation sequencing technologies for insect virus discovery. Viruses 3, 1849-1869.

Loy, J. D., Mogler, M. A., Loy, D. S., Janke, B., Kamrud, K., Scura, E. D., Harris, D. L. & Bartholomay, L. C. (2012). dsRNA provides sequence-dependent protection against infectious myonecrosis virus in Litopenaeus vannamei. J Gen Virol 93, 880-888. 55

Mello, M. V., Aragao, M. E., Torres-Franklin, M. L., Neto, J. M. & Guedes, M. I. (2011). Purification of infectious myonecrosis virus (IMNV) in species of marine shrimp Litopenaeus vannamei in the State of Ceara. J Virol Methods 177, 10-14.

Nibert, M. L. (2007). '2A-like' and 'shifty heptamer' motifs in penaeid shrimp infectious myonecrosis virus, a monosegmented double-stranded RNA virus. J Gen Virol 88, 1315-1318.

Pongsomboon, S., Wongpanya, R., Tang, S., Chalorsrikul, A. & Tassanakajon, A. (2008). Abundantly expressed transcripts in the lymphoid organ of the black tiger shrimp, Penaeus monodon, and their implication in immune function. Fish & Shellfish Immunology 25, 485-493.

Poulos, B. T., Tang, K. F., Pantoja, C. R., Bonami, J. R. & Lightner, D. V. (2006). Purification and characterization of infectious myonecrosis virus of penaeid shrimp. J Gen Virol 87, 987-996.

Senapin, S., Phewsaiya, K., Briggs, M. & Flegel, T. W. (2007). Outbreaks of infectious myonecrosis virus (IMNV) in Indonesia confirmed by genome sequencing and use of an alternative RT-PCR detection method. Aquaculture 266, 32-38.

Tang, J., Ochoa, W. F., Sinkovits, R. S., Poulos, B. T., Ghabrial, S. A., Lightner, D. V., Baker, T. S. & Nibert, M. L. (2008). Infectious myonecrosis virus has a totivirus-like, 120-subunit capsid, but with fiber complexes at the fivefold axes. Proc Natl Acad Sci U S A 105, 17526-17531.

Zerbino, D. R. & Birney, E. (2008). Velvet: Algorithms for de novo short read assembly using de Bruijn graphs. Genome Research 18, 821-829.

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A)

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 TCTACATCTGGCCAAGGAAAATCTGTTCGTAGGCAAGAACGAAGTGGCTATGGGGTTTGA 60 Contig8044 TCTACATCTGGCCAAGGAAAATCTGTTCGTAGGCAAGAACGAAGTGGCTATGGGGTTTGA 60

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 GGTTTAGCCGAAAGAGATCTCGACAGTCTTTAGTAGGATACACTTCTTACACTCCTGGGA 120 Contig8044 GGTTTAGCCGAAAGAGATCTCGACAGTCTTTAGTAGGATACACTTCTTACACTCCTGGGA 120

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 AAGAGAAACATTATCTGTCCAGCAATGATACACGGGTACGTCCTCCATTAGCTGTTGTTT 180 Contig8044 AAGAGAAACATTATCTGTCCAGCAATGATACACGGGTACGTCCTCCATTAGCTGTTGTTT 180

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 TCTTTTCCTCGGCAGGTCCTACAATGTCGGAGCGCAGGAACAGGACGAACCCACGAGTCG 240 Contig8044 TCTTTTCCTCGGCAGGTCCTACAATGTCGGAGCGCAGGAACAGGACGAACCCACGAGTCG 240

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 CATCCTTACGAGAAGTGCAGTAGGTTGGCGCTGTTGAGAGCGGGTGATCTGGGTAATGTC 300 Contig8044 CATCCTTACGAGAAGTGCAGTAGGTTGGCGCTGTTGAGAGCGGGTGATCTGGGTAATGTC 300

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 CGTCGTTACGGGTAGCATTACTTGGCGCATGCAGTGGGAAGTCATGGCCCACGTCACCTT 360 Contig8044 CGTCGTTACGGGTAGCATTACTTGGCGCATGCAGTGGGAAGTCATGGCCCACGTCACCTT 360

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 AACAAAAGACTAGTCCGACTGTAACCTCGGCGTATAGCGAAAGCGGTTAGATAACATTAA 420 Contig8044 AACAAAAGACTAGTCCGACTGTAACCTCGGCGTATAGCGAAAGCGGTTAGATAACATTAA 420

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 ATACCGGCTTGTATGCTAGAAACTGGACTGTCCATATACCATTGGATGTCTCAAACAAAG 480 Contig8044 ATACCGGCTTGTATGCTAGAAACTGGACTGTCCATATACCATTGGATGTCTCAAACAAAG 480

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 AATAAAGAACCTGTTGTTGAACAGCAAAATCAACAAACCTTCAACCAAAAAGACCACAAT 540 Contig8044 AATAAAGAACCTGTTGTTGAACAGCAAAATCAACAAACCTTCAACCAAAAAGACCACAAT 540

NC_007915.1 ------AY570982.1 ------EF061744.1 ------Contig8223 GCGCTAGTGATTGAACACAAAACTAGTGCAACAACGAGCGCCCAAAGCATTCCCGTTCGT 600 Contig8044 GCGCTAGTGATTGAACACAAAACTAGTGCAACAACGAGCGCCCAAAGCATTCCCGTTCGT 600 57

NC_007915.1 ------GGCAATTTCAACCTAATTCTA 21 AY570982.1 ------GGCAATTTCAACCTAATTCTA 21 EF061744.1 ------GGCAATTTCAACCTAATTCTA 21 Contig8223 CCTTCACGTAAGAAGGATGACTATGTGGACATGACACAGGGCAATTTCAACCTAATTCTA 660 Contig8044 CCTTCACGTAAGAAGGATGACTATGTGGACATGACACAGGGCAATTTCAACCTAATTCTA 660 *********************

B)

NC_007915.1 ATAAGATCGCCAAATTTATTTACCAAAAA-CAAAACAAGCAGAGAACATGATACTGCAAA 7460 AY570982.1 ATAAGATCGCCAAATTTATTTACCAAAAA-CAAAACAAGCAGAGAACATGATACTGCAAA 7460 EF061744.1 ATAAGATCGCCAAATTTATTTACCAAAAAACAAAACAAGCAGAGAACATGATACTGCAAA 7461 Contig8223 ATAAGATCGCCAAATTTATTTACCAAAAAACAAAACAAGCAGAGAACATGATACTGCAAA 8100 Contig8044 ATAA------8044 ****

NC_007915.1 GTAGTTTACAACAAATGTATAGGTACTAAATTAAGGGACCAATAAAGAAACTTCGAGTTT 7520 AY570982.1 GTAGTTTACAACAAATGTATAGGTACTAAATTAAGGGACCAATAAAGAAACTTCGAGTTT 7520 EF061744.1 GTAGTTTACAACAAATGTATAGGTACTAAATTAAGGGACCAATAAAGAAACTTCGAGTTT 7521 Contig8223 GTAGTTTACAACAAATGTATAGGTACTAAATTAAGGGACCAATAAAGAAACTTCGAGTTT 8160 Contig8044 ------

NC_007915.1 CCTATAACACATTCCCAGTTGGGTTTTGTGGCCAGCCATG------7560 AY570982.1 CCTATAACACATTCCCAGTTGGGTTTTGTGGCCAGCCATG------7560 EF061744.1 CCTATAACACATTCCCAGTTGGGTTTTGTGGCCAGCCATG------7561 Contig8223 CCTATAACACATTCCCAGTTGGGTTTTGTGGCCAGCCATGCGGTTGGCCCTAGGTTATAG 8220 Contig8044 ------

NC_007915.1 --- AY570982.1 --- EF061744.1 --- Contig8223 TCG 8223 Contig8044 ---

Fig. 1. Sequence alignment (partial) of Contig8044, Contig8223 and the published

IMNV sequences. The alignment was generated by ClustalW2 algorithm. A) 5’- end alignment of the sequences shows sequence differences between the contig8044, contig8023 and the published sequences (NC_007915.2, AY570982.2 and

EF061744.1). B) The 3’-end alignment of the contigs and IMNV sequences.

*Strictly conserved nucleotide sequences.

58

Fig. 2. Sequencing coverage of IMNV genomic sequences. The Illumina reads used for assembling the IMNV contigs (8223 bp) were mapped to the assembled IMNV sequence using a customer designed Perl program. No mismatched bases were allowed in the algorithm. The sequencing coverage was calculated by converting the numbers of mapped reads in the corresponding positions of the IMNV sequences to numbers of times that mapped to each base. Reads of the plus strand (blue) and minus strand (red) were calculated separately.

59

Fig. 3. RT-PCRs performed using four different pairs of primers specific to the 5’-end of the Indonesia (lanes 1-4) and Brazil (lanes 5-8) strains of IMNV confirmed the presence of the novel sequences. Primers specific to previously recognized IMNV sequence were included as positive controls (lane 9). M=Molecular weight marker.

These four new primers were also tested on RNA from specific free pathogen (SPF) shrimp and the RT-PCR results were negative for all new primers at a variety of cycling conditions (data not shown).

60

Fig. 4. RT-PCRs performed using two different pairs of primers specific to the 3’-end of the Indonesia (lanes 1-2) and Brazil (lanes 3-4) strains of IMNV confirmed the presence of the novel sequence. These primers were tested with SPF shrimp and showed no amplification of cDNA, indicating that sequences are present only in virus-infected tissues (data not shown).

61

Fig. 5. SPF Virus Negative shrimp RNA (lane 3) and IMNV viral RNA purified from shrimp tissues (lane 2) (2 µg) were subjected to electrophoresis on a denaturing

0.8% agarose glyoxal gel containing ethidium bromide. Two markers have been used to determine the molecular weight of RNA products, and IMNV genome including 0.5-9 kb Millennium Markers (Ambion) (lane 1) and 0.5-10 kb RNA ladder

(Invitrogen) (lane 4).

62

Fig. 6. RNA subjected to electrophoresis on a denaturing 1% agarose glyoxal gel that was loaded with 5 µg total RNA extracted from negative shrimp tissues to ensure probe specificity to virus RNA (U= uninfected tissue) and extracted from clarified IMNV infected shrimp tissues (I= infected tissue). The gel was then passively transferred to positively charged membrane (Sigma Aldrich) and hybridized overnight at 68 °C with different virus specific biotinlyated RNA probes generated that correspond to original sequences including 95F, 475R, and novel sequences including 275F and 613R. F indicates sense probe and R indicates antisense probe. The bars indicate the pairs of RNA gels and positively charged membranes. M is 0.5-9 kb Millennium Markers (Ambion).

63

1 TCT ACA TCT GGC CAA GGA AAA TCT GTT CGT AGG CAA GAA CGA AGT 45 1 S T S G Q G K S V R R Q E R S 15

46 GGC TAT GGG GTT TGA GGT TTA GCC GAA AGA GAT CTC GAC AGT CTT 90 16 G Y G V * G L A E R D L D S L 30

91 TAG TAG GAT ACA CTT CTT ACA CTC CTG GGA AAG AGA AAC ATT ATC 135 31 * * D T L L T L L G K R N I I 45

136 TGT CCA GCA ATG ATA CAC GGG TAC GTC CTC CAT TAG CTG TTG TTT 180 46 C P A M I H G Y V L H * L L F 60

181 TCT TTT CCT CGG CAG GTC CTA CAA TGT CGG AGC GCA GGA ACA GGA 225 61 S F P R Q V L Q C R S A G T G 75

226 CGA ACC CAC GAG TCG CAT CCT TAC GAG AAG TGC AGT AGG TTG GCG 270 76 R T H E S H P Y E K C S R L A 90

271 CTG TTG AGA GCG GGT GAT CTG GGT AAT GTC CGT CGT TAC GGG TAG 315 91 L L R A G D L G N V R R Y G * 105

316 CAT TAC TTG GCG CAT GCA GTG GGA AGT CAT GGC CCA CGT CAC CTT 360 106 H Y L A H A V G S H G P R H L 120

361 AAC AAA AGA CTA GTC CGA CTG TAA CCT CGG CGT ATA GCG AAA GCG 405 121 N K R L V R L * P R R I A K A 135

406 GTT AGA TAA CAT TAA ATA CCG GCT TGT ATG CTA GAA ACT GGA CTG 450 136 V R * H * I P A C M L E T G L 150

451 TCC ATA TAC CAT TGG ATG TCT CAA ACA AAG AAT AAA GAA CCT GTT 495 151 S I Y H W M S Q T K N K E P V 165

496 GTT GAA CAG CAA AAT CAA CAA ACC TTC AAC CAA AAA GAC CAC AAT 540 166 V E Q Q N Q Q T F N Q K D H N 180

541 GCG CTA GTG ATT GAA CAC AAA ACT AGT GCA ACA ACG AGC GCC CAA 585 181 A L V I E H K T S A T T S A Q 195

586 AGC ATT CCC GTT CGT CCT TCA CGT AAG AAG GAT GAC TAT GTG GAC 630 196 S I P V R P S R K K D D Y V D 210

631 ATG ACA CAG GGC AAT TTC AAC CTA ATT CTA AAA CTG TTA CCA ACA 675 211 M T Q G N F N L I L K L L P T 225

676 ATT TCT GAA CTA CAG AAG AGG CAT ATT CTT CAT CTA CTT CGA GAA 720 226 I S E L Q K R H I L H L L R E 240

721 GAA GTA GAA GGA AAG AAA GTT TGT TTC GTA AAG CGA GAA AAA CAG 765 241 E V E G K K V C F V K R E K Q 255

766 AAT CCA CTT ATG GCT ATA AAC GAA CTA GCC GTT AAA GTT GGA GAG 810 256 N P L M A I N E L A V K V G E 270

811 AAG CCT AAA TAC ACA TCG ACG AAA ACC GGA GCT GAC CAC ATT CCA 855 271 K P K Y T S T K T G A D H I P 285

856 AGC TGG ACT GTA TTG GTT GAG TTC GCA GGT TTT AGC GAA GCA GCG 900 286 S W T V L V E F A G F S E A A 300

901 ACA TGT GAC ACA GTT AAA AAC GCA AAA ATG ATT GCT GCT TAC AAA 945 301 T C D T V K N A K M I A A Y K 315

64

946 TTA GTT AAA AGA TTT TGT AAA TGG GAC CCA ACC TAC ATT GAA ATT 990 316 L V K R F C K W D P T Y I E I 330

991 TCT GAT TGT ATG CTG CCA CCT CCA GAC CTT ACA TCG TGC GGG GAC 1035 331 S D C M L P P P D L T S C G D 345

1036 GTT GAG AGT AAT CCT GGA CCT ATC ATA CAT AGC GTT GCA TTT GCA 1080 346 V E S N P G P I I H S V A F A 360

1081 AGA ACT GGT TCA GTA TGG ACA CCT GCC ACC TTT ACT TTC AAT ACT 1125 361 R T G S V W T P A T F T F N T 375

1126 ACA TCA TCC CCG GGT AGA CTG CAA GTA CAA ATG TCA TCC AGC GAC 1170 376 T S S P G R L Q V Q M S S S D 390

Fig. 7. Translation of the new 5’- sequences of IMNV. The new IMNV sequences were translated (six frames) using BioEdit program. The figure shows the 5’-end of the frame 1 translation. The first start codon (ATG) in frame one (italic M) locates at

433 bp, which starts a peptide sequences and merge the peptide sequence into the known ORF1 sequence started at 775 bp (bold M) of the new IMNV sequence.

65

NC_007915.1 ------AY570982.1 ------EF061744.1 ------New_ORF1 MLETGLSIYHWMSQTKNKEPVVEQQNQQTFNQKDHNALVIEHKTSATTSAQSIPVRPSRK 60

NC_007915.1 ------MAINEL 6 AY570982.1 ------MAINEL 6 EF061744.1 ------MAINEL 6 New_ORF1 KDDYVDMTQGNFNLILKLLPTISELQKRHILHLLREEVEGKKVCFVKREKQNPLMAINEL 120 Consensus ******

NC_007915.1 AVKVGKKPKYTSTKTGADHIPSWTVLVEFAGFSEAATCDTVKNAKMIAAYKLVKRFCKWD 66 AY570982.1 AVKVGKKPKYTSTKTGADHIPSWTVLVEFAGFSEAATCDTVKNAKMIAAYKLVKRFCKWD 66 EF061744.1 AVKVGKKPKYTSTKTGADHIPSWTVLVEFAGFSEAATCDTVKNAKMIAAYKLVKRFCKWD 66 New_ORF1 AVKVGEKPKYTSTKTGADHIPSWTVLVEFAGFSEAATCDTVKNAKMIAAYKLVKRFCKWD 180 Consensus *****:******************************************************

Fig. 8. Alignment of the new ORF1 and the published ORF1 protein sequences

(partial, N-terminus). The multiple amino acid sequence alignments show that the new ORF1 has additional 114 residues at the N-terminus compared to the published

ORF1 sequences.

66

Fig. 9. Genomic organization of IMNV (Indonesia). Previously published information indicates a genome length of 7561 bp (Accession No. AY570982.1, top). Newly revealed sequence information indicates the virus genome length is 8226 bp (Accession No. KF836757, bottom). The predicted putative ORF1 contains 1719 aa and thus is 114 aa longer than the previously reported sequences (1605 aa). Nucleotide positions indicate the following genome features; A is 5’UTR, B is

ORF1, C is intergenic region, D is ORF2 and E is 3’UTR.

67

100

90

80

70 SEM) ± 60 dsRNA 95-475 dsRNA 233 50 dsRNA 26 40 dsRNAeGFP 30 Positive control

Mean Mean Survival ( Negative control 20

10

0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Days post-IMNV infection (Days)

Fig.10. Shrimp survival post-administration with novel dsRNAs (dsRNA26 and dsRNA233) compared with previously reported dsRNA (dsRNA95-475) and heterologous dsRNA (dsRNAeGFP) post 48 hours IMNV infection. Y-axis indicates mean survival ± SEM (%) and X-axis indicates days post-IMNV infection. n=90 animals/treatment except negative control group; n=30 animals.

68

5

4

3 viral copies

10 2 Log

1

0

dseGFP dsRNA26 dsRNA233 dsRNA95-475 Positive control

Fig. 11. Confirmation of suppression of IMNV using muscle tissue by qRT-PCR analysis. X-axis is treatment and Y-axis is log10 viral genome copy number. Bars represent standard error within the sample. n=18 samples/treatment.

69

Table 1. Oligonucleotide primer sequences for RT-PCR, dsRNA synthesis and RACE PCR Primers Sequence 5’- 3’

RT-PCR to confirm the new IMNV genome IMNV26F TTCGTAGGCAAGAACGAAGTGGCT IMNV312F GTAGCATTACTTGGCGCATGCAGT IMNV643R ACTATGTGGACATGACACAGGGCA IMNV1016R TGATTGTATGCTGCCACCTCCAGA IMNV8204R1 CGACTATAACCTAGGGCCAACCGC IMNV8210R2 GACTATAACCTAGGGCC IMNV7852F TTGGACAAATGGTTCCCGGTCCAA

dsRNA synthesis IMNV26T7F TAATACGACTCACTATAGGGTTCGTAGGCAAGAACGAAGTGGCT IMNV401T7R TAATACGACTCACTATAGGGTTCGCTATACGCCGAGGTTACAGT IMNV233T7F TAATACGACTCACTATAGGGACGAGTCGCATCCTTACGAGAAGT IMNV613T7R TAATACGACTCACTATAGGGTCTTACGTGAAGGACGAACGGGAA

5’ RACE of IMNV (Indonesia) 273F ACGAGTCGCATCCTTACGAGAAG 452R AGACATCCAATGGTATATGGACAG 428R TCCAGTTTCTAGCATACAAGCCGG C tail F GACATCGAAAGGGGGGGGGGG 374R GCCGAGGTTACAGTCGGACTAG

5’ RACE of IMNV (Brazil) 51F GCTATGGGGTTTGAGGTTTAGCCG 163R GAGAACAACAGCTAATCGAGGACG 243R TACTGCACTTCTCGTAAGGATGCG 304R AAGTAATGCTACCCGTAACGACGG

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Primers Sequence 5’-3’

3’ RACE of Brazil and Indonesia strain 3’ RT GGCCACGCGTCGACTAGTACTTTTTTTTTTTTTTTTT 3’ Rev GGCCACGCGTCGACTAGTA 7945F GAAATTCCAACTGAACCAGCAGTT M13F GTAAAACGACGGCCAG M13R CAGGAAACAGCTATGAC

Table 2. Comparison of sequencing coverage between the new and published IMNV sequences

Sequencing coverage (fold) IMNV sequence Positive strand Negative Strand Total IMNV genome (8223 bp) 29.39 26.21 55.60 5’-new sequence (639 bp) 26.37 21.48 47.85 Published sequence (7561bp) 29.73 26.67 56.40

71

CHAPTER 3: VARIATION IN VIRULENCE IN GEOGRAPHICALLY

DISTINCT ISOLATES OF INFECTIOUS MYONECROSIS VIRUS IN

THE PACIFIC WHITE SHRIMP, LITOPENAEUS VANNAMEI

A paper to be submitted to Virus Research

Duan S. Loy1, Sijun Liu2, Mark A. Mogler3, J. Dustin Loy4, Bradley J. Blitvich1,

Lyric C. Bartholomay 2

1Department of Veterinary Microbiology and Preventive Medicine, Iowa State University, Ames IA 50011, USA 2Department of Entomology, Iowa State University, Ames IA 50011, USA 3Harrisvaccines, Inc., Ames IA, 50010, USA 4School of Veterinary Medicine and Biomedical Sciences, University of Nebraska- Lincoln, Lincoln NE, 68583, USA

Abstract

The emergence of new viral diseases can cause devastating losses in production animal systems including crustacean aquaculture. For example,

Infectious myonecrosis virus (IMNV) emerged in the last decade and is estimated to have caused losses in excess of $1 billion USD since the first outbreak in two endemic countries, Brazil and Indonesia. Recently, Next Generation Sequencing of

IMNV-infected Litopenaeus vannamei revealed the IMNV genome is 8226 bp in length as compared to 7561 bp in the initial description by additional of 639 bp at 5’ end and 22 bp at 3’end. Bioassays were conducted to compare the virulence of

IMNV isolates and viral replication was evaluated by qRT-PCR. Comparative 72 bioassay results showed that shrimp that were challenged with IMNV (Indonesia) had a survival rate of 13.3% that was significantly lower than 86.7% survival observed in shrimp infected with an equivalent dose of ~7.3 x102 viral copy numbers

µl-1 RNA of IMNV (Brazil) (p=0.0007). Animals challenged with ten times the virus inoculum of IMNV (Brazil) showed higher mortality than the lower dose group; however, even with this increased of dose, death caused by of IMNV (Brazil) did not approach the 100% mortality observed in shrimp infected with IMNV (Indonesia).

The complete genome sequences of both IMNV isolates were compared and shown to differ in 67 bp and 30 amino acid positions.

Introduction

Infectious myonecrosis virus (IMNV) emerged in 2002 in farmed Pacific white shrimp, L. vannamei, at Pernambuco, in the state of Piaui in northeastern Brazil and then spread to other regions of Brazil (Poulos et al. 2006, Andrade et al. 2007,

Walker & Mohan 2009). Several years after the emergence of IMNV in Brazil, IMNV was identified in shrimp by RT-PCR from samples collected in the Situbondo District of East Java in Indonesia in 2006 (Senapin et al. 2007, Walker & Mohan 2009), from which IMNV subsequently spread to other regions of the country. It is likely that the epizootic of IMNV in Indonesia occurred because of transportation and use of infected animals from Brazil; indeed IMNV (Brazil) and IMNV (Indonesia) are very closely related with genomes have 99.6% identity at nucleotide (nt) level (Poulos et al. 2006, Senapin et al. 2007). Clinical signs of IMNV infection include opaque to white muscles that start from the ventrum to dorsum of abdominal muscles and a mortality rate that reaches up to 70% in shrimp farms (Poulos et al. 2006, Senapin et 73 al. 2007). The consequence of the outbreak in these two countries is expected to cause financial losses of more than $1 billion USD (Lightner et al. 2012). Outbreaks of this disease thus far have not been reported in any other countries.

Andrade et al. (2007) reported 100% mortality at 52 days post-infection in a bioassay in which 1 gram animals were challenged with IMNV (Brazil) (~1.0 x 104

IMNV copy numbers µl-1 RNA). The infected animals began to experience increased mortality after 40 days post-infection. In contrast, Loy et al. (2012) demonstrated

100% mortality at 18 days post-infection in a bioassay in which 8-10 gram animals were challenged with IMNV (Indonesia) (~3.2 x 103 IMNV copy numbers µl-1 RNA).

The data indicated that even using the smaller animals as 1 gram challenged with the high viral titer, it took nearly two months to cause 100% mortality using IMNV

(Brazil) while IMNV (Indonesia) reached 100% mortality in shorter time period using bigger animals and lower viral titer. The virulence of these two IMNV isolates cannot be directly inferred from these data because the experiments were conducted in different laboratories with variability attributable to challenge techniques, animal source and qRT-PCR technique. We, therefore, conducted bioassays to directly compare the virulence of IMNV isolates.

IMNV is classified in the family Totiviridae because of its characteristic non- enveloped, double-stranded RNA (dsRNA) genome, and the virus particle with icosahedral symmetry that is 40 nm in diameter. The genome originally was reported to have 7561 base pairs (bp) (NC_007915.2, AY570982.2; EF061744.1; 7561 bp) with two non-overlapping open reading frames (ORFs): ORF 1 and ORF 2 (Poulos et al. 2006, Nibert 2007). We used Next Generation Sequencing (NGS) to 74 investigate IMNV infection in lymphoid organ tissue (Chapter 2). The sequence revealed that IMNV genome is larger than originally reported; the additional sequences include additions to the 5’ end (639 bp) and 3’ end (22 bp) such that the genome is 8226 bp in length. The extra sequence at 5’ end results in the expanding of the coding region of putative ORF1 of IMNV (Indonesia) by 114 amino acids. The novel sequences were confirmed with molecular techniques and the completed

IMNV genome was submitted to Genbank (IMNV (Indonesia), Accession No.

KF836757) (Loy et al., Chapter 2). IMNV genome sequences of both IMNV isolates were compared and revealed differences at both the nucleotides and amino acid level. The studies have shown that the mutation of virus can lead to the increasing of viral virulence or attenuation of virus (Cahour et al. 1995, Fayzulin & Frolov 2004,

Sirigulpanit et al. 2007). It is possible that differences in the coding and non-coding regions of IMNV isolates (Brazil and Indonesia) can affect virus virulence.

Materials and Methods

Animal Rearing

Specific Pathogen Free (SPF) juvenile L. vannamei were received from

Shrimp Improvement Systems (Islamorada, FL, USA) and maintained in a rearing facility at Iowa State University. Shrimp were placed in one ton fiberglass tanks filled with dechlorinated municipal water and Crystal Sea Marinemix (Marine Enterprises

International, Baltimore, Maryland). Water conditions were maintained at 25-27 °C, salinity 30-35 parts per thousand (ppt), with constant airstone aeration. Each tank was equipped with a carbon filter and an oyster shell airlift biofilter. Water quality was monitored weekly by measuring ammonia and nitrite levels (Nitriver3, Hach 75

Company, Loveland, CO). Shrimp were fed and monitored for twice a day with commercial growout diet and cultured to reach 3-7 grams for bioassays.

Bioassays

SPF shrimp sized 5-7 grams were divided into 3 groups of 10 shrimp with 3 replicates in each group and 10 shrimp for negative controls. Tanks were maintained at 29-30 °C. Shrimp were injected intramuscularly with IMNV (Brazil) or (Indonesia) diluted 1:100 (~7.3 x102 viral copy numbers µl-1 RNA). Shrimp were fed and mortality observed twice daily for 17 days post-infection. Another bioassay study was conducted with the smaller animals (3-5 grams) divided into groups (10 shrimp in 3 replicates) and 10 shrimp for negative controls. Shrimp were injected intramuscularly with IMNV. The experimental groups included IMNV (Brazil) and

(Indonesia) with ~7.3 x102 and ~7.3 x103 viral copy numbers µl-1 RNA and negative control group. Shrimp were monitored regularly until 21 days post-infection. These bioassays were repeated twice. Tissue samples were stored in -80 °C prior to qRT-

PCR testing.

Quantitative Reverse-Transcription PCR

Quantitative Reverse-Transcription PCR (qRT-PCR) was used to quantify virus titer according to protocol adapted from a previously described method

(Andrade et al. 2007). RNA was extracted from muscle tissue using the RNAeasy

Minikit (Qiagen) and 3 µl of sample (60 ng/µl) was added to each reaction of One

Step RT-PCR Master Mix (Qiagen). The one step RT-PCR reaction mixer contained

5 x Master Mix (5µl), Enzyme mix (1 µl), dNTP (1 µl), 20 uM of IMNV412F (0.3 µl),

20 uM IMNV545R (0.3 µl), 10 µM IMNV probe (0.3 µl) and RNase-free water (14.1 76

µl) for a total volume of 25 µl. Thermal cycling was performed with conditions from a previous report (Loy et al. 2012) using following conditions: 30 min at 48 °C and 10 min at 95 °C, followed by 15 s at 95 °C and 1 min at 60 °C for 35 cycles. Each reaction was run in duplicate on a BioRad CFX96 Real Time PCR Detection System using a standard curve as described previously (Andrade et al. 2007, Loy et al.

2012).

Sequencing full-length IMNV (Brazil)

The full-length IMNV (Brazil) genome was amplified by RT-PCR and sequenced as nine overlapping fragments using the primers listed in Table 1.

Complementary DNAs were generated using Superscript III reverse transcriptase

(Invitrogen, Carlsbad, CA, USA) following the manufacturer’s instructions. Briefly, an aliquot of total RNA (1–2 µg) was mixed with 500 µM of each dNTP and 25 ng of primer, and heated at 70 °C for 10 min. After briefly chilling on ice, samples were added to 200 units of Superscript III reverse transcriptase in 1x reaction buffer

(50 mM Tris-HCl [pH 8.3], 75 mM KCl, 3 mM MgCl2, 10 mM DTT) and incubated at

50 °C for 1 h, followed by 70 °C for 15 min. PCR amplifications were performed using 1 µl of cDNA template, 0.1 unit of Taq polymerase, 25 ng of each primer and each dNTP at 200 µM in 1x PCR buffer (20 mM Tris-HCl [pH 8.4], 50 mM KCl,

1.5 mM MgCl2). Reactions were performed as follows: 94°C for 3 min, 30 cycles of

94 °C for 30 sec, 50 °C for 45 sec and 72 °C for 3 min, followed by a final extension at 72 °C for 8 min. An aliquot of each PCR product was examined by 1% agarose gel electrophoresis, visualized with ethidium bromide and extracted using the

Purelink gel extraction kit (Invitrogen). Purified DNAs were sequenced in both 77 directions using a 3730x1 DNA sequencer (Applied Biosystems, Foster City, CA,

USA).

Statistical analysis

Survival data and viral copy number were expressed as the mean ± standard error. The mean survival data were tested at the end of experiment and the mean log viral copy number analyzed by qRT-PCR was compared between each group.

Survival data obtained from the experiments were analyzed using Student’s t-test.

One-way ANOVA followed by a Tukey’s multiple comparison test (p<0.05) was used to test the significance of mean survival and virus copy number. All statistical analysis was conducted using JMP 10 (SAS Institute Inc.).

78

Results

Sequence analysis of IMNV (Brazil and Indonesia)

When IMNV first was detected in Indonesia, a comparison of the genome as it was understood at the time (i.e., 7.5 kb in length) revealed 99.6% similarity between

IMNV (Brazil) and IMNV (Indonesia). More recently, our group described an additional 639 bp of sequence at the 5’end and 22 bp at the 3’ end of the IMNV

(Indonesia) genome, which was revealed in a high throughput sequencing effort of

IMNV-infected L. vannamei tissue (Chapter 2). In this study, we endeavored to generate full-length IMNV (Brazil) sequence using RT-PCR and discovered that the full-length genome of this strain also consists of 8226 bp (Accession No. KJ556923).

The full-length of IMNV genome differs from the sequences that were previously reported in Genbank (NC_007915.2, AY570982.2; EF061744.1; 7561 bp) as shown in Fig. 1 (A, B and C). The previously reported IMNV (Brazil and Indonesia) genome was described with two ORFs, ORF1 and ORF2. The first ORF (nt 136-4953) encodes a protein with a dsRNA-binding motif (DSRM) (nt 136-315, 60 amino acids

(aa)) and a major capsid protein (MCP) (nt 2248-4953, 901 aa). ORF 2 (nt 5241-

7490) encodes RNA-dependent RNA polymerase (RdRp) (nt 5241-7490, 749 aa)

(Poulos et al. 2006, Senapin et al. 2007). The recently revealed additional sequence of IMNV genome cause shifts in the nucleotide positions of key features in the genome as shown in Table 2. Most importantly, the extra sequence at the 5’ end results in a considerable increase in the predicted length of ORF1 in the IMNV genome. Originally, ORF1 was predicted to encode 1605 aa; with the addition of nucleotides at the 5’ end, IMNV (Indonesia) encodes 1719 aa and IMNV (Brazil) 79 encodes 1708 aa, increasing the coding region by 114 aa and 103 aa as depicted in

Fig. 1 (A, B and C). Surprisingly, these results suggest that IMNV (Indonesia) encodes 11 additional amino acids at the N-terminus as compared to IMNV (Brazil)

(Fig. 1B, C and D). Coding differences also are evident in the DSRM and the MCP.

ORF 2 encodes the RdRp and has 11 bp changes that result in 4 aa differences between isolates. All together, the two isolates differ by 67 bp and 30 aa as shown in

Table 2. These sequence differences could be key to the variation in the virulence observed in the field and in bioassays.

Variation in virulence of IMNV isolates

Shrimp (5-7 gram) were injected intramuscularly with one of the two isolates of IMNV. The virus inoculum was standardized by qRT-PCR to contain ~7.3 x 102

IMNV copy numbers µl-1 RNA (1:100 dilution) in both IMNV (Indonesia and Brazil) and study animals were observed for 17 days post-infection. The bioassay results showed that shrimp that were challenged with IMNV (Indonesia) had a survival rate of 13.3% which was significantly lower than the 86.7% survival rate in shrimp exposed to the Brazil strain (p=0.0007) as shown in Fig. 2. Another bioassay study was conducted by challenging smaller animals (3-5 grams) (n=60 animals/group for all groups except IMNV (Indonesia), n=30 animals/group for ~7.3 x 103 IMNV copies) with a ten-fold higher titer of IMNV (Brazil) (~7.3 x 103 IMNV copies) in order to determine if higher virus increases mortality in animals challenged with IMNV

(Brazil). The study was observed for a longer period for 21 days post-infection as shown in Fig. 3. Animals that were challenged with ~7.3 x 102 IMNV copy numbers

µl-1 RNA of IMNV (Brazil) showed significantly different mean survival (68.3%) as 80 compared to 5.0% survival in shrimp exposed to IMNV (Indonesia) (p=0.0004). A tenfold increase in the dose of virus with IMNV (Brazil) (~7.3 x 103 IMNV copy numbers µl-1 RNA) did not cause 100% mortality within 10 days post-infection as shown with and equivalent dose of IMNV (Indonesia) (~7.3 x 103 IMNV copy numbers µl-1 RNA). Thus, these bioassays confirm data from the literature that suggest that there are significant differences in virulence between IMNV isolates. In every experiment, exposure to IMNV (Indonesia) results in more rapid disease progression and higher mortality, and therefore higher virulence, than IMNV (Brazil).

Importantly, IMNV (Indonesia) produces100% mortality in L. vannamei within 10 days post-infection using our experimental paramaters; this level of mortality was never observed during infection with IMNV (Brazil). qRT-PCR analysis of virus load in IMNV-infected shrimp

IMNV virus load was analyzed by qRT-PCR using infected muscle tissue.

Although there are differences in the mortality between groups, the quantification of virus in infected animals showed that with either virus strain, dead animals have higher virus loads, with a mean viral titer of 2.02 x 106 IMNV copy numbers µl-1 RNA

(n=6 samples/group) as shown in Fig. 4. No statistically significant differences were observed between the viral load of dead animals between infections with IMNV

(Indonesia) or (Brazil) (p>0.05). Virus load was also assessed in animals infected with IMNV (Brazil) with at either of two doses to compare virus load between survivors and dead animals. The results indicated that the dead animals have higher virus loads titer than the survivors such that 1.04 x 105, and 1.25 x 106 viral copies were observed in survivors and dead animals, respectively (dose ~7.3 x 103 viral 81 copies); in animals exposed to ~7.3 x 102 viral copies, 1.92 x 104 and 2.70 x 106 viral copies were observed in survivors and dead animals, respectively). There was a statistically significant difference between survivors and dead animals in both viral concentration of IMNV (Brazil); ~7.3 x 103 viral copies (p=0.0028) and ~7.3 x 103 viral copies (p=0.005) as shown in Fig. 5. Thus, these data support bioassay data and reveal that virus replication is likely not impaired in IMNV (Brazil) and therefore not the primary cause of different virulence observed between the two strains.

Discussion

The study presented here highlights genotypic and phenotypic differences in

IMNV isolates from Brazil and Indonesia. IMNV emerged first in Brazil in 2002, and was detected in shrimp in Indonesia in 2006. The effect has been particularly devastating in Indonesia, which is home to the biggest shrimp farms in the world.

The mortality observed in challenge models of both viral isolates was confirmed by qRT-PCR to detect virus load in muscle tissue and showed a significant difference in virus titer of mortalities and survivors (Fig. 4 and Fig. 5). These bioassays confirm data from the literature that suggest that there are differences in virulence between

IMNV isolates. The genomes of IMNV (Brazil) and IMNV (Indonesia) contain 8 bp differences in the 5’ UTRs, the non-coding regions of the IMNV genome. During virus replication cycles, bind to the extreme 5’ end of the positive strand to initiate translation; however, if mutations occur in this region, this can impair protein synthesis and lead to attenuation of virus replication (Chatterjee & Pal 2009).

Secondary structures in the mRNA sequence of the 5’ UTR have been reported to be involved in the regulation of RNA replication and translation process of other RNA 82 viruses such as Hepatitis C virus, , and Sindbis virus (Cahour et al.

1995, Butrapet et al. 2000, Fayzulin & Frolov 2004, Shi ST 2006, Sirigulpanit et al.

2007). There are 8 bp substitutions in the 5’UTR between strains, which may account for the phenotypic differences (mortality) in virulence.

The comparison of both IMNV isolates also revealed position differences in the initial start codon of ORF 1 as depicted in Fig. 1 (D). In IMNV (Indonesia), the predicted start codon (AUG) initiates at the nucleotide position 437 while IMNV

(Brazil) AUG starts at the nucleotide position of 470 in which increases the length of putative ORF1 by 11 amino acids in IMNV (Indonesia). Mutations in 51 nt in the 5’

UTR of Sindbis virus (Togaviridae, Alphavirus genus) causes attenuation of viral replication in mosquito cells (Fayzulin & Frolov 2004). Moreover, the sequence comparison of both IMNV genomes showed that nucleotide polymorphisms affect the coding region in ORF 2 that encodes the RdRp. RdRp is key to virus replication so, clearly, mutations that occur in this region could affect the replicative capacity and therefore virulence of the virus (Gallei et al. 2006). Prior studies have shown that point mutations in the RdRp alter replication of (family

Flaviviridae, genus ) (Van Slyke et al. 2012). IMNV is an RNA virus that generally has a high rate of mutation associated with a lack a proof-reading ability

(Steinhauer et al. 1992, Drake 1993). IMNV originated in Brazil and then was translocated to Indonesia via the importing of live animals harboring the virus

(Senapin et al. 2007). During the movement of the disease from to

Asia, it is likely that genetic mutations occurred to enhance viral fitness in a new environment, for which there certainly is precedent in other shrimp viruses including 83

WSSV (Zwart et al. 2010); in fact, dramatic size differences in the White Spot

Syndrome Virus (WSSV) genome (293-312 kb) are involved in WSSV virulence from different countries (Zwart et al. 2010).

Bioassays were conducted in an attempt to evaluate differences in disease outcome and virus replication between IMNV isolates. The inoculation of L. vannamei with ~7.3 x 102 IMNV copies resulted in significant differences in mortality according to IMNV strain. Animals that were challenged with IMNV (Indonesia) showed significantly higher mortality rates than IMNV (Brazil) (Fig. 2). Another bioassay study was conducted with higher doses of virus and showed that animals that received a ten-fold higher dose of IMNV (Brazil) caused higher mortality than the lower dose group (~7.3 x 102 viral copies); infection with IMNV (Brazil) never caused100% mortality that is seen as a result of infection with IMNV (Indonesia)

(Fig. 3).

The data produced in this study did not directly evaluate the direct effect of specific IMNV mutations on pathogenesis; these naturally occurring mutations are the only available method to compare genomic differences because there is not an

IMNV infectious clone and there are no existing continuous shrimp cell lines to propagate the virus. Insect cell lines including Sf 9 (Spodoptera frugiperda cells) can be infected with WSSV (Liu et al. 2007) and a mosquito cell line, C6/36, (Aedes albopictus cells) can be infected with virus (TSV) (Dhar et al.

2004); however, there is no report of successful infection of IMNV in insect cell lines.

Primary cell lines from the hepatopancreas, hemocytes or lymphoid organs (Flegel

1998, George & Dhar 2010, George et al. 2011) might be an alternative method if 84 the primary cell cultures can be perpetuated and show cytopathic effects as a result of IMNV infection. Primary cell culture would facilitate more rigorous comparison of

IMNV strains.

Overall, the present study demonstrates the first direct comparison of IMNV virulence in geographically diverse isolates. This provides additional insight into

IMNV biology to include mutations that very likely have an effect on virulence.

References

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Butrapet, S., Huang, C. Y., Pierro, D. J., Bhamarapravati, N., Gubler, D. J. & Kinney, R. M. (2000). Attenuation markers of a candidate dengue type 2 vaccine virus, strain 16681 (PDK-53), are defined by mutations in the 5' noncoding region and nonstructural proteins 1 and 3. Journal of Virology 74, 3011-3019.

Cahour, A., Pletnev, A., Vazeille-Falcoz, M., Rosen, L. & Lai, C.-J. (1995). Growth-Restricted Dengue Virus Mutants Containing Deletions in the 5′ Noncoding Region of the RNA Genome. Virology 207, 68-76.

Chatterjee, S. & Pal, J. K. (2009). Role of 5'- and 3'-untranslated regions of mRNAs in human diseases. Biology of the cell / under the auspices of the European Cell Biology Organization 101, 251-262.

Dhar, A. K., Cowley, J. A., Hasson, K. W. & Walker, P. J. (2004). Genomic Organization, Biology, and Diagnosis of Taura Syndrome Virus and Yellowhead Virus of Penaeid Shrimp. In Advances in Virus Research, pp. 353-421: Academic Press.

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Drake, J. W. (1993). Rates of spontaneous mutation among RNA viruses. Proceedings of the National Academy of Sciences of the United States of America 90, 4171-4175.

Fayzulin, R. & Frolov, I. (2004). Changes of the secondary structure of the 5' end of the Sindbis virus genome inhibit virus growth in mosquito cells and lead to accumulation of adaptive mutations. Journal of Virology 78, 4953-4964.

Flegel, T. W. (1998). Advances in Shrimp Biotechnology: National Centre for Genetic Engineering and Biotechnology Thailand.

Gallei, A., Widauer, S., Thiel, H. J. & Becher, P. (2006). Mutations in the palm region of a plus-strand RNA virus polymerase result in attenuated phenotype. The Journal of General Virology 87, 3631-3636.

George, S. K. & Dhar, A. K. (2010). An improved method of cell culture system from eye stalk, hepatopancreas, muscle, ovary, and hemocytes of Penaeus vannamei. In Vitro Cellular & Developmental Biology Animal 46, 801-810.

George, S. K., Kaizer, K. N., Betz, Y. M. & Dhar, A. K. (2011). Multiplication of Taura syndrome virus in primary hemocyte culture of shrimp (Penaeus vannamei). Journal of Virological Methods 172, 54-59.

Lightner, D. V., Redman, R. M., Pantoja, C. R., Tang, K. F., Noble, B. L., Schofield, P., Mohney, L. L., Nunan, L. M. & Navarro, S. A. (2012). Historic emergence, impact and current status of shrimp pathogens in the Americas. Journal of Invertebrate Pathology 110, 174-183.

Liu, W. J., Chang, Y. S., Wang, A. H., Kou, G. H. & Lo, C. F. (2007). White spot syndrome virus annexes a shrimp STAT to enhance expression of the immediate-early gene ie1. Journal of Virology 81, 1461-1471.

Loy, J. D., Mogler, M. A., Loy, D. S., Janke, B., Kamrud, K., Scura, E. D., Harris, D. L. & Bartholomay, L. C. (2012). dsRNA provides sequence-dependent protection against infectious myonecrosis virus in Litopenaeus vannamei. The Journal of General Virology 93, 880-888.

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Nibert, M. L. (2007). '2A-like' and 'shifty heptamer' motifs in penaeid shrimp infectious myonecrosis virus, a monosegmented double-stranded RNA virus. The Journal of General Virology 88, 1315-1318.

Poulos, B. T., Tang, K. F., Pantoja, C. R., Bonami, J. R. & Lightner, D. V. (2006). Purification and characterization of infectious myonecrosis virus of penaeid shrimp. The Journal of General Virology 87, 987-996.

Senapin, S., Phewsaiya, K., Briggs, M. & Flegel, T. W. (2007). Outbreaks of infectious myonecrosis virus (IMNV) in Indonesia confirmed by genome sequencing and use of an alternative RT-PCR detection method. Aquaculture 266, 32-38.

Shi ST, L. M. (2006). HCV 5′ and 3′UTR: When Translation Meets Replication. In Hepatitis C Viruses: Genomes and Molecular Biology. Edited by T. SL. Norfolk (UK): Horizon Bioscience.

Sievers, F., Wilm, A., Dineen, D., Gibson, T. J., Karplus, K., Li, W., Lopez, R., McWilliam, H., Remmert, M., Soding, J., Thompson, J. D. & Higgins, D. G. (2011). Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Molecular Systems Biology 7, 539.

Sirigulpanit, W., Kinney, R. M. & Leardkamolkarn, V. (2007). Substitution or deletion mutations between nt 54 and 70 in the 5' non-coding region of dengue type 2 virus produce variable effects on virus viability. The Journal of General Virology 88, 1748-1752.

Steinhauer, D. A., Domingo, E. & Holland, J. J. (1992). Lack of evidence for proofreading mechanisms associated with an RNA virus polymerase. Gene 122, 281-288.

Van Slyke, G. A., Ciota, A. T., Willsey, G. G., Jaeger, J., Shi, P.-Y. & Kramer, L. D. (2012). Point mutations in the West Nile virus (; Flavivirus) RNA-dependent RNA polymerase alter viral fitness in a host-dependent manner in vitro and in vivo. Virology 427, 18-24.

Walker, P. J. & Mohan, C. V. (2009). Viral disease emergence in shrimp aquaculture: origins, impact and the effectiveness of health management strategies. Reviews in Aquaculture 1, 125-154. 87

Zwart, M. P., Dieu, B. T., Hemerik, L. & Vlak, J. M. (2010). Evolutionary trajectory of white spot syndrome virus (WSSV) genome shrinkage during spread in Asia. PloS one 5, e13400.

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Fig. 1. A schematic representation of IMNV genome organization (modified from

(Nibert 2007). A) Previously published information indicated a genome length of

7561 bp (Accession No. AY570982.2). B) Newly revealed sequence information from IMNV (Indonesia) indicates the virus genome length is 8226 bp (Accession No.

KF836757). C) Newly revealed sequence information of IMNV (Brazil) that generated by RT-PCR indicates the virus genome length is 8226 bp (Accession no.

KJ556923). Based on open reading frame (ORF) prediction, the putative ORF1 of

IMNV has 1719 and 1708 amino acids (aa) that is 114 aa and 103 aa longer than the previously reported sequences (1605 aa) (Indonesia and Brazil, respectively). 89

D) Multiple alignment of predicted amino acids sequences using the Clustal Omega program (Sievers et al. 2011). The comparison of both IMNV isolates revealed 11 additional amino acids in IMNV (Indonesia) at putative ORF 1. Nucleotide positions indicate the following genome features; A is 5’ UTR, B is ORF1, C is intergenic region, D is ORF2 and E is 3’ UTR.

100 90 80 70 60 Indonesia 50 Brazil 40 Negative control Survival (%) Survival 30 20 10 0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 Days post-IMNV infection (Days)

Fig. 2. Shrimp percent survival following the IMNV 1:100 challenge (~7.3 x102 IMNV copy number µl-1 RNA), n=30 animals/group.

90

100

90

80

70 Brazil 7.3E+02 60 Brazil 7.3E+03 Indonesia 7.3E+02 50 Indonesia 7.3E+03 40 Negative control Survival (%) Survival 30

20

10

0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

Days post-IMNV infection (Days)

Fig. 3. Shrimp percent survival following the IMNV infection (Brazil and Indonesia) with different dilution of 7.30E+02 and 7.30E+03, n=60 animals/group (except IMNV

(Indonesia) 7.30E+03, n=30 animals/group).

91

7

6

5

4

viral copies 3 10 Log 2

1

0

Brazil Brazil Indonesia 7.3E+03 Indonesia 7.3E+02 7.3E+03 7.3E+02

Fig. 4. Viral genome copies calculated by qRT-PCR in muscle tissue of dead animals post-IMNV infection (n=6 samples/ group). X-axis is treatment and Y-axis is log10 viral copies. Bars represent standard error within the sample. No significant difference was observed between groups of dead animals (p>0.05).

92

7

6

5

4 Survivors Mortalities viral copies 3 10

Log 2

1

0

Brazil Brazil 7.3E+03 7.3E+02

Fig. 5. Viral genome copies calculated by qRT-PCR in muscle tissue of dead and survival animals post-IMNV (Brazil) infection, n=12 samples/group (6 survived and 6 dead animals/group). X-axis is treatment and Y-axis is log10 viral copies. Bars represent standard error within the sample. There are significant differences between dead and surviving animals after infection with IMNV (Brazil) in both viral titer groups; 7.3E+03 (p=0.0028) and 7.3E+02 (p=0.005).

93

Table 1. Primers used for the RT-PCR amplification and nucleotide sequencing of the IMNV (Brazil) genome.

1Primers for PCR Nucleotide Sequence (5’ to 3’) and sequencing positions PCR-A-For 1-21 ATTTTCTACATCTGGCCAAGG PCR-A-Rev 1148-1169 TGGATGACATTTGTACTTGCAG Seq-1A 321-341 ATTACTTGGCGCATGCAGTGG Seq-2A 641-662 GAATTAGGTTGAAATTGCCCTG PCR-B-For 1041-1063 TTGAGAGTAATCCTGGACCTATC PCR-B-Rev 2084-2105 CTTCTCTAGTTAGAACTGGATC Seq-1B 1361-1380 GAGGAAGCTATAGAGGCAAC Seq-2B 1766-1787 CCTCAATTGCTGTCTTTAGTGG PCR-C-For 1921-1943 GGAACAGGTATCAATAACGCCAG PCR-C-Rev 2984-3006 GTCATATCAGCTAATACTCTTAG Seq-1C 2266-2288 ATCATCAAGCAACGATCACTATG Seq-2C 2339-2360 TCATTTCAAGTGGATCTTCTGG PCR-D-For 2804-2826 GTCAGTGACGTTACCTACACTAG PCR-D-Rev 3946-3946 TGTTGGTTGAATTACTACACGTG Seq-1D 3201-3222 TTGAAACACACATTACAGACAG Seq-2D 3474-3496 CTGTGTTCTTGAATCAAATGCAG PCR-E-For 3826-3849 CGAATCTACTGTACATATTCCAGG PCR-E-Rev 4937-4960 TGAATTATCAATTAGTGCTGGTTC Seq-1E 4241-4263 ACTGCATGTCTTGCAAACTTAGG Seq-2E 4518-4539 CTATCATATGCTGTGTCCCACG PCR-F-For 4881-4902 TGCCATGGCCAATGCCAGAAGG PCR-F-Rev 5943-5965 CTATTCCAAATACTAACATATAG Seq-1F 5361-5381 AGTGGGCACATGCTCAGAGAC Seq-2F 5466-5488 CTTGGGTGCCATCATTGCTGTGG

94

1Primers for PCR Nucleotide Sequence (5’ to 3’) and sequencing positions

PCR-G-For 5842-5864 ATACGTTTAATCGACCTTTGGCG PCR-G-Rev 6889-6911 CTCGTTATTCTCACTCCTGATGG Seq-1G 6242-6263 AAGTAATGACATGGTACAAAGG Seq-2G 6425-6447 ATCCCAAGCATAAACAATGTTGG PCR-H-For 6796-6820 GTCAATAAGATTGTTAATGCATATG PCR-H-Rev 7632-7655 GTCTGATTTCTACTCCATATGATC Seq-1H 7147-7169 GAGATATCAGTGACAGGAGTACG Seq-2H 7340-7360 AACTTTGTCTGACGTGTCTCG PCR-I-For 7226-7249 AACCTTGGAGTCCAAATGATGAAG PCR-I-Rev 8203-8226 GACTATAACCTAGGGCCAACCGCA Seq-1I 7559-7580 TGCAAACAGATGATATACCTGG

Seq-2I 7665-7686 ACGTACTGTGTGATCAGTATGG

1Primers A through I denote those used to RT-PCR amplify and/or sequence fragments 1 through 9, respectively; For denotes forward PCR primers; Rev denotes reverse PCR primers

95

Table 2. IMNV genomes comparison of geographically distinct isolates between

Brazil and Indonesia.

Indonesian isolate compared Region in Nucleotide positions to Brazilian isolate the genome Nucleotide Amino acid Indonesia Brazil differences differences

5' UTR 1- 436 1-469 8, Gap= 33 NA

ORF1 437-5596 470-5596 48 26

DSRM 779-958 779-958 2 1

MCP 2870-5596 2870-5596 22 6

Intergenic 5597-5883 5597-5883 NA NA

ORF2 5884-8133 5884-8133 11 4

RdRp 5884-8133 5884-8133 11 4

3'UTR 8134-8226 8134-8226 NA NA

96

CHAPTER 4: COMPARATIVE ANALYSIS OF SUPPRESSION OF

IMMEDIATE EARLY, EARLY AND LATE GENES OF WHITE SPOT

SYNDROME VIRUS

A paper to be submitted to Journal of Invertebrate Pathology

Duan S. Loy1, Tyasning Kroemer2, Lyric C. Bartholomay2

1Department of Veterinary Microbiology and Preventive Medicine, Iowa State University, Ames IA 50011, USA 2Department of Entomology Iowa State University, Ames IA 50011, USA

Abstract

White Spot Syndrome Virus (WSSV) is one of the most devastating disease agents to cultured shrimp. WSSV is a double-stranded DNA virus with a 300 kbp genome. WSSV gene expression occurs in a temporally regulated cascade that includes immediate-early (IE), early (E) and late (L) genes to facilitate host cell entry, initiation of replication and virus particle assembly. Recombinant proteins, RNA interference (RNAi) triggers and broad-spectrum immune activators have been explored as means to potentiate an antiviral response and thereby protect shrimp from White spot syndrome. Our studies and others have shown that double-stranded

RNA (dsRNA) molecules, specific to capsid protein-coding genes in the WSSV genome, provide significant protection from disease and mortality. We postulated that targeting immediate early or early genes could provide more protection against disease by shutting down virus replication more rapidly and thereby reducing the burden of infection on the host cell machinery. RNAi triggers were generated to 97 target an immediate early (wssv403) and an early (wssv477) as compared to a late gene (VP15). Animals that were subjected to injection with dsRNA477 or dsRNAVP15 showed high survival (95.5% and 90.0%, respectively) at 10 days post- infection. By contrast, dsRNA403 did not provide a significant protection against

WSSV infection. In addition, the viral copy number detected in dsRNA477 and dsRNAVP15-injected shrimp was lower than the control and dsRNAeGFP groups suggesting the inhibitory effects of virus replication by dsRNA. To carry this a step further, we conducted a minimum protective dose study. Shrimp that received a dose of 0.1 µg of dsRNA477 showed 74.0% survival post-infection. A dose ten times higher (1 µg) was necessary to achieve 64.4% survival using dsRNAVP15.

Introduction

In the last two decades, the global crustacean fisheries industry witnessed a dramatic increase in production (measured in millions of metric tons) as a result of widespread adoption of the Pacific white shrimp as the culture species of choice

(see Bondad-Reantaso et al. 2012). Epizootics of disease caused by virus infections are the primary obstacle to productivity and yield in these fisheries (Lightner 2011b).

White spot disease causes the majority of losses and remains the most significant infectious disease in shrimp production systems. WSSV outbreaks cause up to

100% mortality in cultured shrimp within 3-5 days.

White spot disease was first reported in shrimp farms in northern Taiwan in

1992 (Chou et al. 1995); within a few years, it spread to several shrimp farming countries (Lo et al. 1996, Wang et al. 1999). The spread of WSSV threatens the success and further development of shrimp aquaculture worldwide because it 98 causes tremendous economic losses, in fact an estimated $15 billion since 1992

(Lightner et al. 2012). Standard disease control measures for shrimp production include the use of Specific Pathogen Free (SPF) animals produced through a judicious quarantine procedure to guarantee disease-free status, selective breeding from families of animals that have desirable growth and disease resistance traits, and the rigorous use of hatchery and farm biosecurity measures (see Moss et al.

2012). Unfortunately, family-based selective breeding has had limited success in the case of resistance to WSSV because of limited heritability of disease resistance traits (Moss et al. 2012) and disease outbreaks continue to occur despite the availability of SPF stocks and use of on-farm biosecurity. As a result, there is a very real need to develop alternative disease control strategies for WSSV.

WSSV is large, enveloped virus with a circular double-stranded DNA (dsDNA) that has a genome size of approximately 300 kbp (van Hulten et al. 2001, Yang et al.

2001). The WSSV genome has at least 181 putative open reading frames (ORFs)

(van Hulten et al. 2001) and it is classified in the genus Whispovirus, family

Nimaviridae (Mayo 2002). WSSV gene expression occurs in a temporal cascade related to host cell entry, replication and capsid assembly. As such, these gene production clusters include immediate-early (IE), early (E) and late (L) genes. WSSV

IE genes can be expressed independently in the absence of viral proteins by utilizing host cell replicative machinery; expression of IE genes can be detected from 2 hours post-primary infection (Liu et al. 2005). Viral E genes usually encode for regulatory proteins that are necessary for L gene expression (Han et al. 2007). Viral L genes, including structural genes like capsid proteins are found in the late stages of 99 infection of cells, at least 16-24 hours post-infection (Leu et al. 2009, Sanchez-Paz

2010).

Significant efforts have been made to develop antiviral vaccines using different techniques such as inactivated WSSV vaccines, recombinant protein vaccines and DNA vaccines (Namikoshi et al. 2004, Vaseeharan 2006, Rout et al.

2007). However, currently there are no effective vaccines that are commercially available to control and mitigate WSSV outbreaks. Robalino et al. (2005) first reported the induction of an antiviral state by dsRNA in the shrimp Litopenaeus vannamei. Since then, RNA interference has been used extensively to demonstrate the function of genes in the WSSV genome (Robalino et al. 2004, Robalino et al.

2005, Westenberg et al. 2005a, Robalino et al. 2007, Xu et al. 2007, Raja et al.

2010, Sarathi et al. 2010, Mejía-Ruíz et al. 2011). Most of these studies targeted

WSSV structural genes including VP28, VP19, VP24 and VP15 (Westenberg et al.

2005a, Wu et al. 2007b, Attasart et al. 2009) that are expressed at the late stage of virus infection (Leu et al. 2009). We reasoned that another strategy that could be employed in order to improve the efficacy of RNAi for the purposes of developing a commercially viable WSSV vaccine would be to target early gene expression instead of structural genes; the idea being that suppressing these transcripts could mitigate some of the adverse effects of virus use of host cell machinery by interfering with the virus earlier in the infection cycle, and that these transcripts could be easier to target using RNAi because they would be in lower transcript abundance than Late, structural genes. Only one study has directly evaluated the efficacy of targets other than structural genes (Wu et al. 2007b, Attasart et al. 2009). Attasart et al. (2009) 100 developed RNAi targets to genes across the WSSV genome and reported promising results from suppression of several genes including the early gene, ribonucleotide reductase small subunit (rr2) gene, with or without PmRab 7 (cellular genes that involving in endocytosis of viral entry), against WSSV showed 95% survival in

Penaeus monodon, the Black tiger shrimp. Because the WSSV genome is so large, and potential for protection after RNAi-based suppression has been investigated for a fraction of the 181 total ORFs, there is little doubt that effective antiviral molecules against WSSV remain to be discovered. The purpose of our study was to compare the efficacy RNAi targets in the immediate early, early or late gene to provide protection from disease in WSSV infected Litopenaeus vannamei.

Materials and Methods

Shrimp Rearing

Specific pathogen free (SPF) L. vannamei were obtained from Shrimp

Improvement Systems (Islamorada, FL, USA) and reared in a bio-secure animal holding facility at Iowa State University. Shrimp were reared in 1000L tanks containing artificial seawater, an oystershell airlift biofilter, and an activated carbon filter.

Preparation of Virus Inoculum

Virus inoculum was prepared based on Hasson et al. (1995). Tissue from

WSSV infected animals was homogenized in sterile TN buffer (0.02 M Tris-HCl, 0.4

M NaCl, pH 7.4) and clarified through three centrifugation steps. Supernatant was transferred into a new tube after each centrifugation step: 14,000 × g for 30 min;

15,000 × g for 15 min; and 25,000 × g for 60 min (Hasson et al. 1995). The final 101 supernatant was diluted 1:10 with 2% saline and filtered through a 0.2 micron filter.

A 1000-fold dilution of the stock (~6.38x105 WSSV copies ul-1 DNA) was used as the viral challenge dose for the experiments.

Double-stranded RNA Synthesis

Template DNA for dsRNA production was extracted from virus infected shrimp using the DNeasy kit (Qiagen, Valencia, CA, USA) for a template of the dsRNA. Primers containing T7 promoter sites were designed to amplify PCR products that targeted VP15 (GenBank Accession No. DQ681072.1), wssv403

(GenBank Accession No. AF332093.1), and wssv477 (GenBank Accession No.

NC_003225.1). Primers used to produce the template for dsRNA targeting wssv403

(dsRNA403) were: 5′-TAA TAC GAC TCA CTA TAG GGG ACC AGT TCC AAC

CCA AGA A -3′ and 5′-TAA TAC GAC TCA CTA TAG GGC CTG TCA AAC GGG

ATA GCT G -3′. Primers used to synthesize the template for dsRNA targeting wssv477 (dsRNA477) were: 5′-TAA TAC GAC TCA CTA TAG GGA CCT TTG TCG

ACT CGT GGA A -3′ and 5′-TAA TAC GAC TCA CTA TAG GGG GGC ACA ACA

AAT CAC AAT G -3′. Primers used to amplify dsRNAVP15 were 5′- TAA TAC GAC

TCA CTA TAG GGA TGA CAA AAT ACC CCG AGA T-3′ and 5′- TAA TAC GAC

TCA CTA TAG GGT TAA CGC CTT GAC TTG CGG ACT C-3′. Primers used to amplify eGFP (enhanced Green Fluorescent Protein) to generate dsRNAeGFP were

5’-TAA TAC GAC TCA CTA TAG GGA GAA TGG TGA GCA AGG GCG AGG AGC

TGT-3’ and 5’-TAA TAC GAC TCA CTA TAG GGA GAT TAC TTG TAC AGC TCG

TCC ATG CCG-3’ (Loy et al. 2012). 102

PCR products were purified using the Gel/PCR DNA Fragments Extraction kit

(IBI Scientific, Peosta, IA, USA) and were sequenced prior to being used as template material for dsRNA synthesis. dsRNA was prepared using the Ambion

Megascript RNAi Kits (Life technologies, Grand Island, NY, USA) following the manufacturer’s instructions. 8 µl of PCR product was mixed with 2 µl ATP, 2 µl CTP,

2 µl, GTP, 2 µl UTP, 2 µl Buffer, and 2 µl of enzyme mix into a total of 20 µl per reaction. The mixture was incubated at 37 °C for 12 hours. dsRNA products were then incubated for 1 hr with DNase I and RNase, and purified through column centrifugation. dsRNA products were subjected to gel electrophoresis and quantified using a NanoDrop spectrophotometer (Thermo Scientific, Wilmington, DE, USA).

Bioassays

3-5 g SPF shrimp were stocked in tanks containing 200 L synthetic seawater and an oystershell airlift biofilter. Tanks were maintained at 28 ppt salinity and 28 °C.

Shrimp were divided into six experimental groups. Each tank contained 10 shrimp and a total of 30 shrimp for each treatment were used in each replicate of the experiments; the negative control group contained 10 shrimp. Following acclimation for 48 hours, each shrimp was injected intramuscularly with 2 µg dsRNA403, dsRNA477, dsRNAVP15, or a heterologous dsRNA control targeting eGFP. At 72 hours post-dsRNA injection, shrimp were challenged with WSSV including positive control group. Shrimp that were not subjected to any dsRNA treatments or WSSV challenge were used as a negative control. Mortality data were recorded for 10 days post-infection and dead shrimp were collected daily and stored in -80 °C freezer to be tested for the presence of WSSV by qPCR. 103

Minimal protective dose against WSSV

3-5 g SPF shrimp were divided into eight experimental groups including different doses of dsRNA, positive and negative control group. Each group was represented by three replicate tanks with 10 shrimp per replicate. Shrimp received different doses (2 µg, 1 µg and 0.1 µg) of dsRNA477 or dsRNAVP15 or were inoculated with 2% NaCl in the case of the positive control group. Following the dsRNA or 2% NaCl administration, shrimp were challenged with WSSV at 72 hours post-injection dsRNA. Shrimp were monitored for mortality twice a day and dead shrimp were removed daily and stored at -80 °C to be tested for the presence of

WSSV by qPCR. These studies were completed in triplicate.

Quantitative Real Time-PCR

DNA template was extracted from gills of an individual shrimp and homogenized using a QIAShedder column (Qiagen) followed by extraction with a

DNeasy blood and tissue kit (Qiagen). Moribund shrimp for each treatment were tested for the presence of WSSV. DNA template from each sample was eluted with

50 µl nuclease free water. Quantitative Real Time-PCR reaction was performed as described previously by Durand & Lightner (2002). The Quantitect Probe kit

(Qiagen) was used and a mixture reaction was prepared following manufacturer’s instructions. Briefly, 3 µl of DNA template was mixed with 12.5 µl of Master Mix, 0.5

µl of Taqman probe, 1 µl WSSV forward primer (20 µM), 1 µl WSSV reverse primer, and 7 µl nuclease free water for a total reaction of 25 µl. Primers used to detect the presence of WSSV were WSS1011F: 5′-TGG TCC CGT CCT CAT CTC AG-3′ and

WSS1079R: 5′-GCT GCC TTG CCG GAA ATT A-3′ and the TaqMan probe 104 sequences were 5′-FAM-AGC CAT GAA GAA TGC CGT CTA TCA CAC A-TAMRA-

3′ (Durand & Lightner 2002). This reaction was then run in two replicates on a

BioRad CFX96 Real Time PCR machine (BioRad, Hercules, CA, USA). Real time

PCR thermal cycling parameters included: 15 min at 95 °C followed by 40 cycles of

15 s at 95 °C, and 1 min at 60 °C. Virus copy number was calculated using CFX

Manager Software (BioRad) according to a standard curve.

Statistical analysis

The survival rate and the virus copy number data were analyzed by JMP 9 software (SAS Institute Inc. Cary, NC, USA). A One-way ANOVA, followed by a

Tukey’s multiple comparison test, was performed to find significant differences among treatment and control groups.

Results

Protection of shrimp mortality against WSSV

To examine the protective effects of dsRNA against WSSV infection, the mortality of shrimp injected with dsRNA specific to wssv403, wssv477, or VP15 was compared to the control groups after WSSV challenge (Fig.1). Shrimp injected only with WSSV (positive control group) showed 0% survival at 10 days post-challenge with WSSV. Shrimp that were injected with dsRNAeGFP also showed 0% survival at

10 days post-challenge. Shrimp injected with dsRNA403, dsRNA477 and dsRNAVP15 showed 61.1%, 95.5%, and 90.0% survival at 10 days post-challenge with WSSV, which was significantly different from the untreated positive controls and shrimp injected with dsRNAeGFP; dsRNA403 (p=0.0003), dsRNA477 (p<0.0001), and dsRNAVP15 (p<0.0001). The percent survival of dsRNA403 is significantly 105 lower from those of dsRNA477 and dsRNAVP15 groups at 10 days post-WSSV challenge (p=0.0069 and p=0.0286). No mortality was observed in the negative control group. Our data show that amongst the dsRNA groups, dsRNA477 and dsRNAVP15 provided the most effective protection against WSSV challenge.

Inhibition of WSSV replication

To investigate the suppression of virus replication by dsRNA, the copy number of virus in the dsRNA-injected shrimp was compared to that in control groups (dsRNAeGFP and positive control groups). WSSV was detected and quantified by using quantitative Real Time PCR according to Durand & Lightner

(2002). Virus loads in the positive control and dsRNAeGFP groups were 1.68x108 and 5.93x107 WSSV copy numbers µl-1 DNA, respectively, which was not significantly different (p=0.9580). Although the percent survival of dsRNA403 group is significantly higher than those of positive control (p<0.0001) at 10 days post

WSSV challenge, the viral copy number of dsRNA403 group (mean 7.53x106 WSSV copy numbers µl-1 DNA) was not significantly different from the positive control

(p=0.2754) and dsRNAeGFP group (p=0.6581). Amongst the dsRNA injected groups, dsRNA477 and dsRNAVP15 groups (mean 5.17 and 9.85x102 WSSV copy numbers µl-1 DNA, respectively) showed the lowest virus load as would be expected based on decreased mortality in these groups, which does represent a statistically difference from the virus copy number observed in the dsRNAeGFP group

(p<0.0001). The virus copy number of dsRNA477 group was not significantly different than the dsRNAVP15 group. 106

Minimal protective dose of dsRNA477 and dsRNAVP15 against WSSV

To test the possibility that suppression of an early stage gene requires less dsRNA trigger to control white spot syndrome, shrimp were subjected to different concentrations (2 µg, 1 µg and 0.1 µg/shrimp) of dsRNA477 and dsRNAVP15; these two dsRNAs were of particular interest based on initial results (above) that showed both significant protection from disease and significantly reduced virus loads according to qPCR. Animals were challenged at 72 hours post-injection with either dsRNA. The results demonstrate that shrimp that received dsRNA477 at doses of 2

µg, 1 µg and 0.1 µg/shrimp showed 97.0%, 96.0% and 74.0%, survival, respectively, all of which were significantly different (p<0.0001) from the positive control group which showed 1% survival (Fig. 3). Shrimp that received dsRNAVP15 at a dose of 2

µg, 1 µg/shrimp exhibited significantly different survival from the positive control group (p<0.0001) with average survival of 78.9% and 64.4%, respectively. However, the dsRNAVP15 group that was provided with 0.1 µg showed 8% survival which was not significantly difference from the positive control group (p=0.9954). Thus, a dose of dsRNA477 as low as 0.1 µg/shrimp provided significant protection from white spot disease, and ten times the amount was required to provide comparable protection using dsRNAVP15.

Muscle tissues were collected from the surviving or the dead animals at 7-10 days post-infection and assayed by qPCR to determine the effect of virus load as measured by virus copy number (Fig. 4). The data show that pre-exposure to either dsRNA477 (2 µg and 1 µg/shrimp, mean 2.3x101 and 7.15x101 WSSV copy numbers µl -1 DNA, respectively) or dsRNAVP15 (2 µg and 1 µg/shrimp, mean 107

2.58x101, 5.12x101 WSSV copy numbers µl-1 DNA, respectively) results in significantly fewer virus copies in muscle tissue of L. vannamei as compared to the control group, (mean 2.48x108 WSSV copy numbers µl-1 DNA) (p=0.002). In contrast, groups exposed to only 0.1 µg of either dsRNA477 or dsRNAVP15 had a mean of 3.32x102 WSSV copy numbers µl-1 DNA or 1.56x108 WSSV copy numbers

µl-1 DNA respectively, such that only the copy number in the group of dsRNA477 was significantly different from positive control group (p=0.002) as compared to dsRNAVP15 (p=0.6687). The data reveal that shrimp that received dsRNA477 and dsRNAVP15 at the two highest doses had high protection as measured by low virus copy numbers. This study also demonstrates that dsRNA477 can be provided with as little as 0.1 µg/shrimp.

Discussion

RNA interference (RNAi) is a highly conserved pathway for antiviral defense and gene regulation in plants, animals, and fungi. The RNAi pathway begins with the long double-stranded RNA that can be introduced exogenously or is produced as a function of virus replication. During an antiviral response in penaeid shrimp, the dsRNAs are processed by RNase-III-like enzymes (Dicer) into small RNA duplexes that function as small interfering RNA (siRNA) (Labreuche & Warr 2013). These duplexes are then unwound and one strand (the guide strand) is loaded into a protein complex (RNA-induced silencing complex or RISC) after the recruitment of

Argonaute proteins (Labreuche & Warr 2013). In the last step, the guide strand complexes with the messenger RNA (mRNA) target based on sequence identity between the mRNA and the guide strand to induce post-transcriptional gene 108 silencing (Labreuche & Warr 2013). In shrimp, different sets of Dicer and Argonaute- encoding proteins have been identified, thereby demonstrating the biochemical capability of these animals to carry out the RNAi pathway (Unajak et al. 2006,

Dechklar et al. 2008, Labreuche et al. 2010).

There are ample data to support the importance and robust nature of RNAi as an antiviral response in penaeid shrimp. In the absence of deliberate and systematic interrogation of the nuances of the RNAi pathway, there is a great deal known about

RNAi activation as a result of dsRNA injection or oral administration followed by virus challenge. Based on previous studies, dsRNA and siRNA specific for viral genes suppress virus replication for WSSV (Robalino et al. 2005, Xu et al. 2007,

Attasart et al. 2009, Sarathi et al. 2010, Mejía-Ruíz et al. 2011, Bartholomay et al.

2012, Loy et al. 2012), Yellow head virus (Tirasophon et al. 2005), IMNV (Loy et al.

2012) Taura syndrome virus (Tirasophon et al. 2005) and Penaeus monodon densovirus (PmDNV) (Attasart et al. 2010). Therefore, the applications of RNAi to control shrimp disease caused by viruses is an attractive approach to produce a commercial product to prevent the outbreak of many shrimp diseases in the farms.

Most WSSV early genes encode enzymes involved in virus replication and nucleotide metabolism (Sanchez-Paz 2010); the gene that was explored in this study, wssv477, is the only WSSV early gene that has been reported to possess a regulatory function (Han et al. 2007). In our study, the WSSV genes wssv403, wssv477, and VP15 were chosen as RNAi targets based on temporal expression associated with the stage of infection. The results demonstrated that injection of

RNAi triggers corresponding to wssv477 (early gene) and VP15 (late gene) showed 109 a significant reduction in mortality 10 days after WSSV challenge when compared to controls (Fig. 1). Wssv477 transcripts can be detected at increasing levels from 2 to

72 hours post-infection with expression of the corresponding protein product detected by 6 hours post-infection (Han et al. 2007, Wu et al. 2007a). Our results show that targeting the early genes, wssv477 indeed confers significant protection against WSSV. This result agrees with previously reported data that targeted the

WSSV early gene, ribonucleotide reductase small subunit (rr2) (Attasart et al. 2009).

The late gene, VP15, is one of the two major nucleocapsid proteins that comprise the WSSV nucleocapsid structure, and when targeted with RNAi provides protection against WSSV. VP15 was reported to have a clear preference for supercoiled DNA in lieu of double-stranded RNA, single-stranded RNA or single-stranded DNA

(Witteveldt et al. 2005). This suggests that VP15 is involved in the condensation and packaging of the genome in the nucleocapsid (Witteveldt et al. 2005, Leu et al.

2009). Both 477 and VP15 dsRNA-treated groups showed significant reduction in virus copy number as compared to the other groups (Fig. 2). The significant reduction in copy number may indicate the inhibition of viral replication in the shrimp by both dsRNA treatments. Thus, these data support the essential function of these two genes in virus replication.

Immediate early genes of DNA viruses like WSSV and the baculoviruses encode proteins that are important in activating the expression of viral early and late genes, altering host gene expression, and evading host immune responses

(Sanchez-Paz 2010). The data show that dsRNA that targets wssv403 (early gene) induced protection against WSSV, but the survival of dsRNA403 vaccinated animals 110 began to decline significantly 5 days post-WSSV challenge. Wssv403 encodes a protein with a RING-H2 finger motif that plays an important role in the ubiquitination pathway. However, there are three other proteins (wsv199, wsv249 and wsv222) that also predicted to have a similar RING-H2 finger motif (Leu & Lo 2011). Thus, knocking down only wssv403 might not be sufficient to inhibit viral replication because there are other proteins involved in the ubiquitination pathway that may serve as an alternative pathway for replication (Leu & Lo 2011). Ubiquitin proteins are commonly found in almost all tissue types of eukaryotic organisms (Pickart &

Eddins 2004).

The inhibitory effects of dsRNA targeting WSSV early, immediate early or late genes may not depend on whether the genes are expressed on the early stage or later stage of viral infection. Attasart (2009) suggested that the protective degree of dsRNA that targets a specific WSSV gene might be correlated with the gene function over the course of viral infection. The dsRNA that targeted immediate early genes ie1 and DNA pol did not show any viral inhibition, however, dsRNA that targeted immediate early gene ie3 and early gene rr2 showed a viral inhibition, making a temporal association with gene transcription and protection unclear (Attasart et al.

2009).

In a minimal protective dose study, shrimp were administered with different doses of dsRNA (2, 1 and 0.1 µg/shrimp) of dsRNA477 and dsRNAVP15, the data suggested that sequence specific of dsRNA of early gene, wssv477, can reduce the mortality from WSSV using as minimal dose as 0.1 µg/shrimp (Fig. 3). The qPCR also confirmed that using the dose of 0.1 µg/shrimp can inhibit viral replication and 111 significantly from positive control group (p=0.0234) (Fig. 4). However, the similar dose of dsRNAVP15 did not show significant protection against WSSV and a dose ten times higher (1 µg) was necessary to achieve 64.4% survival. Thus, the function of target genes may play a significant role in the degree of protection induced by antiviral RNA interference. The lower dose of dsRNA that can mitigate viral disease could contribute to the cost-effective of vaccine production resulting in benefit to the crustacean fisheries industry.

Antiviral molecule approaches to manage WSSV by using dsRNA is probably a safe approach to reduce the impact on non-target organisms, because dsRNA is developed specifically to target the WSSV transcripts. In addition, dsRNA reduces the ability of the virus to replicate as shown by the low virus copy number after the application of dsRNA in the shrimp when compared to the positive controls. This would ultimately reduce the amount of virus shed in the environment and reduce the ability for virus transmission in ponds. These advantages are certainly desirable for the development of potential control agents for WSSV. However, additional research is still needed to assess the environmental risk and food safety of the RNAi treatment.

Acknowledgements

The author would like to thank Kara Jamenez, Zach Flickinger and Kay

Kimpston-Burkgren for animal rearing and laboratory assistance and Shrimp

Improvement Systems (SIS) for providing shrimp. This work was supported by the

National Science Foundation.

112

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A B 100 100 90 90 80 80 70 70 60 60 50 50 dsRNA403 dsRNAeGFP 40 40 Positive control Survival (%) Survival 30 dsRNA477 (%) Survival 30 20 dsRNAVP15 20 Negative control 10 10 0 0 0 1 2 3 4 5 6 7 8 9 10 0 1 2 3 4 5 6 7 8 9 10 C 100 90 80 70 60 50 40

Survival (%) Survival 30 20 10 0

dsRNA403dsRNA477 dsRNAVP15dsRNAeGFP Positive controlNegative control

Fig. 1. Survival rate (%) of shrimp injected with dsRNAs (dsRNA403, dsRNA477, dsRNAVP15 or dsRNAeGFP) and challenged with WSSV at 72 hours after dsRNA injection. Each point represents the mean and standard error of values obtained from three replicates. A) Shrimp survival post administration of different dsRNAs; dsRNA477, dsRNA403 and dsRNAVP15. B) Shrimp survival post administration of dsRNAeGFP, positive controls and negative controls (injected with 2% NaCl). C)

Overall percent shrimp survival at the termination day of study (10 days post- infection) from three replicates. All treatment groups have 90 animals/treatment except negative control group has 30 animals/treatment from three replications. 117

10

8

6 viral copies

10 4 Log

2

0

dsRNA403 dsRNA477 dsRNAVP15 dsRNAeGFP Positive control

Fig. 2. Viral copy number data of each treatment (n=18 sample/treatment). DNA was extracted from each shrimp and quantitative Real Time PCR was performed to detect the presence of WSSV. Data shown in the graphs were means with the standard error bars from three replications.

118

A B

100 100 90 90 80 80 70 70 60 60 50 dsRNA477 2ug 50 dsRNAVP15 2ug 40 dsRNA477 1ug 40 dsRNAVP15 1ug 30 dsRNA477 0.1ug 30 dsRNAVP15 0.1ug 20 20 10 10 0 0 0 1 2 3 4 5 6 7 8 9 10 0 1 2 3 4 5 6 7 8 9 10

C D

Survival (%) Survival 100 100 90 90 80 80 70 70 60 60 50 Positive control 50 40 Negative control 40 30 30 20 20 10 10 0 0 0 1 2 3 4 5 6 7 8 9 10 - + dsRNAVP15 dsRNA477 0.1 1 2 0.1 1 2

Fig. 3. Survival rate (%) of shrimp that intramuscular injection with different doses of dsRNA477 and dsRNAVP15 (2, 1 and 0.1 µg/ shrimp) and challenged with WSSV at

72 hours post-dsRNA injection. Each point represents the mean and standard error of values obtained from three replicates with a total of 90 shrimp per treatment. A)

Shrimp survival post administration of various doses of dsRNA477. B) Shrimp survival post administration of various doses of dsRNAVP15. C) Survival of negative control group that injected with 2% NaCl and positive control post-WSSV infection.

D) Overall percent shrimp survival at the termination day of study (10 days post- infection) from three replicates. All treatment groups have 90 animals/treatment except the negative control group which has 30 animals/treatment from three replications. 119

10

8

6 viral copies 4 10 Log 2

0 dsRNA477 dsRNAVP15 + 2 1 0.1 2 1 0.1 (µg)

Fig. 4. Confirmation of suppression of WSSV replication by qPCR (n=6 samples/treatment). Muscle tissues were collected from shrimp that were administered with different doses of dsRNA (2, 1 and 0.1 µg/shrimp) of dsRNA477 and dsRNAVP15. Data shown in the graphs were means with the standard error bar of log10 of WSSV copies.

120

CHAPTER 5: DIRECT DELIVERY OF DOUBLE STRANDED RNA

INTO THE FOREGUT LUMEN OF LITOPENAEUS VANNAMEI

DEMONSTRATES PROTECTION AGAINST WHITE SPOT

SYNDROME VIRUS AND INFECTIOUS MYONECROSIS VIRUS

A paper to be submitted to Diseases of Aquatic Organisms

Duan S. Loy1, J. Dustin Loy3, Mark A. Mogler4, D.L. Hank Harris1, 4, 5, 6,

Lyric C. Bartholomay2

1Department of Veterinary Microbiology and Preventive Medicine, Iowa State University, Ames IA 50011, USA 2Department of Entomology, Iowa State University, Ames IA 50011, USA 3School of Veterinary Medicine and Biomedical Sciences, University of Nebraska- Lincoln, Lincoln NE 68583, USA 4Harrisvaccines, 1102 S. Hills Dr. Suite 101, Ames IA 50010, USA 5Department of Animal Science, 11 Kildee Hall, Iowa State University, Ames IA 50011, USA 6Department of Veterinary Diagnostic and Production Animal Medicine, Iowa State University, Ames IA 50011, USA

Abstract

Double-stranded RNA (dsRNA) is a potent trigger for induction of an antiviral

RNA interference (RNAi) response and has been shown to protect penaeid shrimp from infection with a wide range of DNA and RNA viruses. White Spot Syndrome

Virus (WSSV) and Infectious myonecrosis virus (IMNV) are two particular significant viral diseases that cause large economic losses in shrimp aquaculture. Vaccination 121 is a possible mean to prevent large-scale outbreaks. Intramuscular injection is routinely used to evaluate shrimp vaccine efficacy. As an alternative, reverse gavage can be utilized to directly deliver dsRNA into the foregut lumen of shrimp. We have developed RNAi triggers that effectively to protect shrimp against either WSSV or

IMNV. These dsRNAs were mixed with food coloring dye to facilitate the observation of dsRNA gavage into the gut. These experiments demonstrate that the introduction of specific anti-viral dsRNA via reverse gavage can elicit statistically significant

(p<0.05) protection against both IMNV and WSSV virus-induced mortality following infection. Delivery by this route elicits protection comparable to vaccination by intramuscular injection, and indicates that shrimp possess a mechanism to uptake dsRNA from the gut lumen. These data also demonstrate that viral gene specific dsRNAs significantly reduce viral replication in vivo.

Introduction

Infectious diseases caused by viruses are a major cause of economic losses in cultured shrimp, with cumulative losses of more than $15 billion USD (Lightner

2011a, Lightner et al. 2012). White Spot Syndrome Virus (WSSV), which first emerged in Southeast Asia in 1992 and has spread throughout shrimp culturing areas across the world (Lo et al. 1996, Flegel 1997, Wang et al. 1998, Lightner

2011a). Infectious myonecrosis virus (IMNV) emerged in the last decade in Brazil and spread to Indonesia, and it is expected to cause an economic loss of more than

$1 billion USD (Lightner et al. 2012). WSSV is a very large, enveloped, double- stranded DNA (dsDNA) virus. The viral genome size is around 300 kbp and it is classified in its own genus, Whispovirus, in the family, Nimaviridae. In contrast to 122

WSSV, IMNV is a small, non-enveloped, double-stranded RNA (dsRNA) virus with genome size around 8 kb that is classified in family Totiviridae (Poulos et al. 2006,

Loy et al, Chapter 2).

The clinical signs of disease cause by WSSV are white spots on the carapace, red to pink discoloration of the body, lethargy, abnormal swimming behavior, reduced feed consumption and ultimately death (Lo et al. 1996, Lo & Kou

1998, Mohan et al. 1998). IMNV-associated clinical signs of disease include white opaque abdominal muscle, red tail fans, lethargy and decreased feed consumption

(Poulos et al. 2006, Senapin et al. 2007). WSSV infected shrimp ponds have high mortality, up to 100%, within 3-10 days post-infection under ideal conditions. IMNV causes less mortality than WSSV, and ranges from 40-70% (Chou et al. 1995,

Poulos et al. 2006, Senapin et al. 2007). These two viruses can co-infect shrimp as was recently demonstrated in farms in Northeastern Brazil (Feijó et al. 2013).

Selective breeding and pathogen exclusion via biosecurity systems are used to reduce the impact of virus outbreaks in the shrimp farming industry. Shrimp vaccines have been developed using various modalities and delivery systems including formalin inactivated virions (Namikoshi et al. 2004), subunit protein vaccines (Vaseeharan et al. 2006), DNA vaccines (Rout et al. 2007) and dsRNA vaccines (see Bartholomay et al.2012). These vaccine studies have shown that protection and enhancement of immune responses in penaeid shrimp against virus infections is possible, and that shrimp immunity is more complex than previously understood (Johnson et al. 2008). RNA interference (RNAi) is undoubtedly key to the antiviral response as was first demonstrated by Robalino et al. (2004, 2005). To 123 test this, an intramuscular injection of in vitro synthesized dsRNA that targets specific WSSV genes provided protection against lethal challenge by intramuscular injection (Robalino et al. 2005). Other studies have shown that various vaccines provide significant protection against viral infection such as WSSV, IMNV,

Yellow head virus (YHV), Penaeus stylirostris densovirus (PstDNV) (Robalino et al.

2004, Robalino et al. 2005, Yodmuang et al. 2006, Robalino et al. 2007, Ho et al.

2011, Bartholomay et al. 2012, Loy et al. 2012). However, these studies used an injection route, which is impractical in the field.

An oral vaccine is the ideal for vaccine delivery system in aquatic animal production systems. However, one of the major impediments of developing an oral vaccine delivery system is that gut environment of animals may cause degradation or inactivation of nucleic acid vaccine components rendering them ineffective. Thus, an alternative reverse gavage method was utilized to directly deliver dsRNA into the foregut lumen of shrimp in order to determine if dsRNA targeted to WSSV and IMNV structural genes can establish protection against virus challenge. This technique allows us to test the efficacy of a dsRNA vaccine, at a specific dose, in the absence of uncontrolled variables that exist with per os delivery feeding; this includes the issue of dose based on feed uptake, heterogeneity of distribution of the vaccine on feed, release of vaccine from feed, and potential degradation of the vaccine as a result of mechanical processing of the feed through gastric sieve and digestive enzymes in the gut. Reverse gavage is a method that has been employed for disease challenge in penaeid shrimp; for example, Aranguren et al. used reverse gavage to challenge shrimp with the enteric bacterial pathogens, as necrotizing 124 hepatopancreatitis bacterium (NHP-B) (Aranguren et al. 2010). Similar methods have also been used successfully to infect mosquitoes with virus, e.g, Chikungunya virus (CHIKV), in Aedes albopictus (Nuckols et al. 2013). This methodology applied to mosquitoes allows researchers to infect animals with pathogens (e.g., a nematode) delivered directly to the gut, thereby bypassing physical damage that can be inflicted when the pathogen is taken up per os (Klowden 1981). This is the first time that reverse gavage has been used as methodology to test an antiviral in shrimp.

Materials and Methods

Animal rearing

Specific Pathogen Free (SPF) juvenile L. vannamei were received from

Shrimp Improvement Systems (Islamorada, FL, USA) and cultured in artificial seawater that was prepared using sea salt (Crystal Sea Marine Mix) at animal rearing facility on SPF units at Iowa State University. Water quality was maintained at 25 to 27 °C and the salinity at 30-35 parts per thousand (ppt). Other water qualities such as pH, ammonia and nitrite were monitored once a week using commercial available kits (Nitriver3, Hach Company, Loveland, CO). Shrimp was fed with the commercial feed (Zeigler) until sized 3-5 grams for the study.

Dose determination of WSSV and IMNV bioassay

Tanks were stocked with 20 SPF shrimp (3- 5 grams) in 8 x 50 L tanks and allowed to acclimate for 24 hours. Each tank contained artificial seawater with oyster shell biofilters. WSSV dilutions of virus stock were prepared at 1:103, 1:104 and 1:105 with 2% NaCl (WSSV stock ~ 6.8 x 106 WSSV copy numbers µl-1DNA). Shrimp were 125 challenged via intramuscular injection with 100 µl WSSV/ shrimp using the different dilutions. Mortality was observed for 8 days post-infection. The virus isolate used in this study was kindly provided by the Aquaculture Pathology laboratory, University of

Arizona. Virus stocks were purified from macerated shrimp tissue, clarified by centrifugation, and filtered through 0.2 micron filter (Hasson et al. 1995). Virus samples were stored in aliquots at -80°C for disease challenge experiments.

IMNV infected tissue were macerated and clarified and filtered following the method of (Loy et al. 2012). The virus was diluted with sterile 2% NaCl (viral diluent) into three different concentrations: 1:101, 1:102 and 1:103 (IMNV stock ~ 2.1 x 105

IMNV copy numbers µl-1 RNA). Animals in each group (n= 10) were injected intramuscularly with 100 µl of diluted virus and mortality was observed for 14 days.

Clinical signs of IMNV were observed at 4-5 days post-infection (p.i) and mortality was observed beginning on day 2 post-injection.

Doubled-stranded RNA synthesis

Double-stranded RNAs targeting VP19 envelope protein of WSSV (Robalino et al. 2005) and nucleotides 95-475 of IMNV genome (GenBank accession no.

EF061744) were generated by in vitro transcription by adding T7 promoter to the 5’ end of each primers sequence and using the Megascript RNAi (Ambion) according to the manufacturer’s protocol. Primers that were used for dsRNAVP19 synthesis were VP19T7F: 5’-TAA TAC GAC TCA CTA TAG GGC GAA GCT TGG CCA CCA

CGA CTA ACA CTC-3’ and VP19T7R: 5’-TAA TAC GAC TCA CTA TAG GGC GGA

GCT CCT GCC TCC TCT TGG GGT AAG AC-3’. Primers for dsRNA95-475 were generated according to a previous publication (Loy et al. 2012). Tracking dye was 126 prepared by mixing commercially available red food coloring (Tone’s, Ankeny IA) with nuclease free water in 1:103 dilution and filtered at 0.2 microns for sterility.

Then, either dsRNAVP19 or dsRNA95-475 were mixed with sterile filtered marking dye to facilitate observation of dsRNA introduction into the foregut and to ensure that the preparation did not perforate the gut lumen during the reverse gavage procedure.

Reverse gavage injection

SPF 3-5 gram juvenile L. vannamei were divided into 3 groups of 10 shrimp with 3 replicates in each group. A sham group (n=10) were injected with 2% NaCl.

Animals were acclimated for 48 hours and starved for 24 h prior to treatment.

Treatment groups received 2 µg/shrimp of VP19dsRNA either by reverse gavage

(RG) as shown in Fig. 2 or via intramuscular injection (IM) at the third abdominal segment from the telson. RG was performed using a tuberculin syringe, with a 27G x

½ CC needle (Becton, Dickinson and Company). A new syringe and needle were used to deliver 100 µl of dsRNA, mixed with tracking dye to the hindgut via the anus into each animal. The syringe cargo was transported (by positive pressure) through the midgut into the foregut (Felgenhauer, 1992), and care was taken to not puncture the gut epithelium. Successful delivery was indicated by visualizing the tracking dye translocating into the foregut. At 3 days post-dsRNAVP19 injection, shrimp in each treatment group, and positive control animals, were challenged with 100 µl with 0.2 micron filtered WSSV clarification diluted in 2% sterile saline at WSSV 1:104 dilution

(∼ 6.8 x 102 WSSV copy numbers µl-1 DNA). Mortality was observed for 21 days post-WSSV challenge. The WSSV study was repeated twice. The RG experiment 127 was then carried out using dsRNA95-475 at a dose of 2µg/shrimp and subsequent challenge with IMNV at 1:103 dilution (∼ 2.1 x 103 IMNV copy numbers µl-1 RNA) at 2 days post-injection was observed for 14 days post-IMNV challenge and the study was repeated three times. Water temperatures were maintained at 27-28 °C for

WSSV trials and at or above 30 °C for IMNV.

Quantitative PCR of WSSV and quantitative reverse-transcription PCR of IMNV

Dead and moribund shrimp were tested by quantitative PCR (qPCR) according to established protocols for the presence of WSSV (Durand & Lightner

2002) and IMNV (Andrade et al. 2007). For WSSV, qPCR was performed on a

BioRad CFX96 Real Time PCR Detection System. Template DNA was generated from gill tissues that were removed from shrimp using sterile forceps and scissors and nucleic acid was extracted using DNeasy Blood and Tissue kit (Qiagen) according to the manufacturer’s instructions. 10-50 ng (3 µl) of DNA template was added to 12.5 µl of PCR master mix (Quanti Tect Probe PCR Kit, Qiagen) containing

10 µl (20 µM) of each primer (WSSV1011F and 1079R) and 0.5 µl (10 µM) of WSSV

TaqMan probe and 7 µl of nuclease-free water for a total reaction volume of 25 µl.

This reaction was performed in duplicate with thermocycling conditions of 95 °C for

15 min, followed by 35 cycles of 95 °C for 15 s and 60 °C for 1 min. Virus copy number was calculated using CFX manager software (Bio-Rad) using a standard generated from a 69-bp insert plasmid as previously described (Durand & Lightner

2002) and performed 10 fold series dilutions. Quantitative reverse-transcription PCR

(qRT-PCR) for IMNV was performed using template RNA from muscle tissue of 128 infected animals and RT-PCR parameters according to the methods described previously (Loy et al. 2012).

Statistical analysis

Survival and virus copy number data obtained from the experiments were expressed as the mean ± standard error. The survival data was analyzed using one- way ANOVA followed by Tukey’s multiple comparison test. Survival data were collected until the termination of experiments on day 14 for IMNV trials and day 21 for WSSV trials. The mean log viral copy number was compared between each group using one-way ANOVA followed by Tukey’s multiple comparison test.

Statistical calculations were performed using JMP 10 (SAS Institute Inc).

Results

In vitro titration of WSSV and IMNV

The dose of virus given of WSSV and IMNV were established by titration in order to determine the challenge dose for the reverse gavage trials. SPF shrimp were stocked and challenged with the different dilution of WSSV and IMNV. Mortality was observed for 8 days p.i. for WSSV and 14 days p.i. for IMNV, the survival of

WSSV 1:103, 1:104 and 1:105 showed 5%, 30% and 100% survival, respectively.

The sham control also showed 100% survival. Thus, the dose that was utilized as

WSSV challenge model is 10000-fold dilution (1:104) of WSSV stock (~ 6.8 x 102

WSSV copy numbers µl-1 DNA) as shown in Fig. 1A. In the IMNV dose titration, shrimp exposed to each dilution (1:101, 1:102 and 1:103) showed 100% mortality at day 6, 9 and 13 p.i. The 1000-fold dilution (1:103) of IMNV stock (~ 2.1 x 103 IMNV copy numbers µl-1 RNA) was chosen for experiment as shown in Fig. 1B. 129

Survival of animals following dsRNAVP19 and dsRNA95-475 reverse gavage administration

Animals were subjected to vaccination by injection or RG then challenged with WSSV or IMNV using challenge model conditions. The groups that were vaccinated with dsRNAVP19 by reverse gavage, intramuscular injection and positive control group showed a survival rate of 98.1%, 98.3% and 5.0%, respectively, at the end of the experimental period (21 days p.i.) (Fig. 3). Survival of the groups vaccinated by RG and IM were significantly different from the positive control group at a significance level of p<0.0001. Similarly, dsRNA95-475 vaccinated animals showed significant protection (p<0.0001) against IMNV administered via RG and IM injection account for 81.0% and 77.0% survival, respectively, compared to the positive control group (2.4% survival) at the end of the experimental period (14 days p.i.) (Fig. 4).

Quantitative Polymerase Chain reaction

Quantitative real time polymerase chain reaction (qPCR) or Reverse transcriptase quantitative polymerase chain reaction (RT-qPCR) was performed on

RNA (IMNV) or DNA (WSSV) extracted from six shrimp tissue samples taken from animals randomly selected at termination from each treatment group, in order to quantify WSSV and IMNV copy number. The results indicated that the mean WSSV copies in RG and IM vaccinated group were both significantly reduced (p<0.0001) in comparison to the positive control group, accounting for 3.9 x 101, 3.0 x 101 and 4.0 x 106 WSSV copy numbers µl-1 DNA, respectively (Fig. 5A). This shows that vaccination by RG with dsRNAVP19 inhibits WSSV multiplication comparable to 130 vaccination by IM injection. In the IMNV trials, the mean IMNV viral copies of shrimp vaccinated with dsRNA95-475 via RG and IM are 4.7 x 103, 9.9 x 103 IMNV copy numbers µl-1 RNA, respectively. This was statistically significant different (p=0.0001) from the positive control group 4.6 x 106 IMNV copy numbers µl-1 RNA (Fig. 5B).

Discussion

In this study, we demonstrate for the first time the use of a reverse gavage system to deliver antiviral vaccines to penaeid shrimp. RNAi antiviral molecules have been proved to show protection against viral diseases. To fully explore the capacity of this methodology both DNA virus (WSSV) and RNA virus (IMNV) infection model were used. VP19 is a major envelope protein located in ORF 182 of the WSSV genome that contains two putative transmembrane domains (van Hulten et al. 2002). dsRNA95-475 is located in ORF1, and encodes a putative RNA-binding protein

(Poulos et al. 2006, Loy et al. 2012). These two viral gene targets were selected based on previous data that demonstrated the high survival of intramuscular vaccinated shrimp following subsequent viral challenge (Robalino et al. 2005, Loy et al. 2012). However, as injection administration is impractical to use in the field for shrimp farming, this study was undertaken as a proxy for per os vaccination, to demonstrate that delivery of dsRNA to gut epithelia could induce a protective response.

These experiments demonstrate that the introduction of dsRNA via reverse gavage can elicit statistically significant (p<0.05) protection against both IMNV and

WSSV virus-induced mortality. Delivery by this route elicits protection comparable to intramuscular injection that is a standard model method for vaccine delivery. These 131 data also demonstrate that gene-specific dsRNAs significantly reduce viral replication in vivo for both WSSV and IMNV. This provides evidence that dsRNA remains potent and effective after passing through the gut environment that could contain digestive enzymes or bacteria that can degrade the dsRNA. Previous work evaluating oral delivery of pellet feed coated with inactivated bacteria that expressed dsRNAVP28 showed 68% survival after challenge with WSSV, however it is unclear if the bacterial cells or some other factors provided protection (Sarathi et al. 2008b).

The oral delivery of DNA vector encoding expression of VP28 encapsulated in chitosan nanoparticles also showed 85% protection against WSSV challenged at 7 days post-feeding (Rajeshkumar et al. 2009). Recently, another oral application has been developed against Laem-Singh virus (LSNV) by incorporating bacterially expressed LSNV-dsRNA into feed, and it improves the growth of infected animals compared to controls (Saksmerprome et al. 2013a). Based on the information obtained from previous LSNV studies, delivering dsRNA into the gut by oral feeding is functional, reduces mortality and improves growth on peneaid shrimp. This work provides direct evidence that introduction of dsRNA sequences corresponding to viral pathogens to the shrimp foregut via reverse gavage can provide significant protection from virus-induced mortality. Further research should be conducted to maximize oral vaccine delivery with different delivery systems that are suited to a marine environment.

Acknowledgements

The authors would like to thank Kara Jamenez, Zach Flickinger, Kay

Kimpston-Burkgren for animal rearing and laboratory assistance, and Shrimp 132

Improvement Systems (SIS) for providing shrimp. This work was financially supported by United States Department of Agriculture, Small Business Innovation

Research Program (USDA SBIR) and the National Science Foundation.

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136

A 100

80 Sham 1:100000 60 1:10000 1:1000 40 Survival (%) Survival

20

0 0 1 2 3 4 5 6 7 8 Days post-WSSV infection (Days)

B 100

80 Sham 60 1:1000 1:100 40 1:10 Survival (%) Survival

20

0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 Days post-IMNV infection (Days)

Fig. 1 A) L. vannamei survival post challenge with serial dilutions of WSSV including

1:103, 1:104 and 1:105 or normal saline (sham) (WSSV stock ∼ 6.8 x 106 WSSV copy numbers µl-1 DNA). Mortality was observed for 8 days, n=20 animals/group.

B) L. vannamei survival post challenge with serial dilutions of IMNV including

1:101, 1:102 and 1:103 or sham (IMNV stock ∼ 2.1 x 105 IMNV copy numbers µl-1

RNA). Mortality was observed for 14 days, n=10 animals/group. 137

Fig. 2. Image of reverse gavage administration demonstrated with a tracking dye.

A) Shrimp were subjected to reverse gavage using a tuberculin syringe, 27Gauge x

½ CC to deliver 100 µl of tracking dye into the hindgut via the anus. B) In this experiment, tracking dye was used to ensure that the gut epithelium was not punctured; successful delivery was indicated by visualizing the tracking dye translocating into the foregut as seen in C.

138

100

90

80 IM 70 RG Positive control 60 Negative control 50

40 Survival (%) Survival 30

20

10

0 0 2 4 6 8 10 12 14 16 18 20

Days post-WSSV infection (Days)

Fig. 3. L.vannamei mean survival (%) following administration of dsRNAVP19 via

intramuscular injection (IM) or reverse gavage (RG). Animals were challenged

with WSSV at 72 hours post-vaccination and were observed for 21 days post-

infection. n=60 animals/treatment.

139

100

90

80

70

60 IM 50 RG 40 Positive control Survival (%) Survival 30 Negative control

20

10

0 0 2 4 6 8 10 12 14 Days post-IMNV infection (Days)

Fig. 4. L. vannamei mean survival (%) following administration of dsRNA95-475

via intramuscular injection (IM) or reverse gavage (RG). Animals were

challenged with IMNV at 48 hours post-vaccination and were observed for 14

days post-infection. n=90 animals/treatment.

140

A 9 8

7

6

5

viral copies 4 10 3 Log 2

1

0 dsRNAVP19 dsRNAVP19 Positive control (IM) (RG)

B 6

5

4

3 viral copies 10 2 Log

1

0 dsRNA95-475 dsRNA95-475 Positive control (IM) (RG)

Fig. 5. Confirmation of suppression of WSSV and IMNV viral replication by quantitative PCR. A) Gill tissue were used as template for WSSV viral copies, n=12 samples/group. B) Muscle tissue were used as template for IMNV viral copies detection, n=18 samples/group. All data are presented as mean ± standard error of the log10 of IMNV copies. 141

CHAPTER 6: GENERAL CONCLUSIONS

The research described in this dissertation demonstrates the interactions of the Pacific white shrimp, L. vannamei with two devastating viruses, WSSV and

IMNV, by utilizing RNA interference as tool to study this interaction. Viral diseases are the major cause tremendous negative impacts on the shrimp farming industry.

Efforts have been made to mitigate and control viral diseases. To understand the underlying mechanisms of host-virus interactions, the transcriptome of lymphoid organ from shrimp infected with IMNV was analyzed. Surprisingly, we found that

IMNV has additional sequences at both the 5’ (643 bp) and 3’ (22 bp) ends of the genome that results in the full-length IMNV genome sequence of 8226 bp as compared to the original description of 7561 bp. An RNAi silencing bioassay, in which we targeted the additional sequence at 5’ end for gene suppression revealed the critical nature of the new sequence to produce high titer infection and associated disease (Chapter 2). Outbreaks of IMNV have been reported in two countries, Brazil and Indonesia. Previous studies from different laboratories revealed differences in mortality rates in bioassays. Our studies confirmed that there is a significant difference in virulence between these geographically distinct isolates, with a higher virulence in IMNV (Indonesia). Even a ten-fold higher dose of IMNV (Brazil) did not equal that of shrimp exposed to IMNV (Indonesia). Genome sequencing revealed 67 nucleotides and 30 amino acid residues that are distinct between the two isolates

(Chapter 3). Further experimentation is needed to identify the specific nucleotide changes that are associated with the increased virulence phenotype in IMNV

(Indonesia). 142

WSSV is the most devastating virus in shrimp aquaculture that historically and currently causes the highest economic losses of all of the shrimp infectious diseases. Novel antiviral molecules against WSSV have been developed based on genes associated with the early stage of virus infection including immediate early

(dsRNA403), early (dsRNA477) and late genes (dsRNAVP15). dsRNA477 (early gene) showed a significant level of protection with the lowest dose of 0.1 µg/shrimp following the challenge with lethal dose of WSSV. In contrast, administration of dsRNAVP15 (late gene) at the same dose did not provide good protection. This provides evidence that gene function and the stage of expression may play an important role in selection of RNAi targets (Chapter 4). Furthermore, these data have important implications in context of field application of this technology, namely the small dose of dsRNA477 required to induce protection will reduce the cost of production for an antiviral vaccine that will benefit to shrimp production systems.

The goal of the aquaculture industry is to develop an oral vaccine that is suited to a marine environment. RNAi-based antiviral vaccines targeting viral genes on a sequence-specific level has been subject to intense research efforts for the last decade, and has provided some provocative and promising results for disease control. Development of an oral delivery system has shown some promise; however, a primary hurdle is a consistent vaccine delivery system. The hypothesis that directly delivering of dsRNA into the gut can provide protection against viral diseases has not been tested robustly. In our study, a new methodology leveraging reverse gavage to deliver dsRNA has been developed as shown in Chapter 5 and it can be used as a proxy for possible per os vaccination. We show that dsRNA delivered via 143 this method elicits protection against WSSV and IMNV that is comparable to the traditional intramuscular injection method. These results also indicate that dsRNA can be taken up through the gut epithelial cells. However, further research is needed to identify the underlying mechanism to dsRNA uptake into shrimp gut epithelial cells.

Future directions

Data based on in vivo bioassay studies described in this dissertation, show that shrimp utilize RNAi as antiviral mechanisms that can shut down virus replication in specific manner. To accomplish the goal of oral vaccine development, novel antiviral molecules that provide a high protection against viral diseases including dsRNA233 (IMNV) and dsRNA477 (WSSV) should be incorporated into feed using a delivery technology that can improve the stability and efficacy of nucleic acids in marine environment. Further study of the mechanisms and functions of RNAi pathway in penaeid shrimp are needed in order to advance our understanding in antiviral immunity in invertebrates. Shrimp immunity is a complex cellular system that data has shown that shrimp may possess some forms of memory immune response, however; further studies are needed to develop the model to study long- term memory immune response against viral diseases. Study of host-virus interactions has been hampered by a lack of shrimp genome sequencing for genetic manipulations and immortal cell lines that can be used as a tool for in vitro studies.

The research presented in this dissertation significantly advances knowledge of the host-virus interactions in the Pacific white shrimp and provides promising 144 results toward oral vaccine development. There is a rich landscape of research questions yet to be addressed from the fundamental mechanisms of RNAi pathway in penaeid shrimp to further advances vaccine development, all of which would prove useful in developing new shrimp disease control strategies.

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ACKNOWLEDGEMENTS

I would like to express my gratitude and appreciation to all those who gave me the possibility to become a PhD. Firstly, I would sincerely like to thank my co- major professor, Dr. Lyric Bartholomay for being the best mentor that I could ask for, and who gives me advice both personal and academic. I could never have made it to see the day that I am a PhD without her support. Through all the obstacles, she is always the best support and is always by my side. She not only showed me how to work but she showed how to work smartly and taught me how to be an independent researcher. She is a role model that always inspires and encourages me to be successful in both work and family. Thank you to her for the critical reading and editing of this dissertation.

I would like to thank Dr. Bradley Blitvich, who is my co-major professor, who also has supported me through graduate school. As he said when he tried to cheer me up on one of those bad days a few years ago that “one day when you look back, you will think it is funny and you will laugh at it”. This reminded me of an Aristotle quote “the roots of education are bitter, but the fruit is sweet”.

I would like to thank my other committee members that have provided assistance through graduate school. Dr. Hank Harris who gave me the best opportunity in my life when he accepted me to graduate school and guided my work in shrimp research. I’m so lucky to be his last graduate student. Dr. Cathy Miller, whose passion for molecular virology taught me that research can fail so many times but you need only one success to achieve great things. Dr. Michael Kimber, who provided excellent support with the great knowledge of RNA interference and gave 146 me a perspective of applied research. I also would like to thank Dr. Bruce Janke who was previously a committee member that supported me through graduate school and encouraged me to continue with shrimp research.

A special thank you goes to Dr. Edward Scura who gave me a constructive advice and an endless support for shrimp research projects.

I would like to thank you current and previous Harrisvaccines employees, Dr.

Mark Mogler, Dr. Ryan Vander Veen and their wives who are great friends that supported and assisted me in my research projects and gave a perspective for working in industry. Thanks to Dr. Kurt Kamrud for the guidance through graduate school and sharing his passion in science. I would like to thank Jill Gander whom I have learned so much of molecular techniques through her suggestions for troubleshooting. Thanks to Pamela Whitson, Kristen Summerfelt, James Wamboldt,

Patrick Jennings and Kara Jimenez who assisted and coordinated my research projects.

I would like to thank my past and current lab mates in the Bartholomay’ lab including Dr. Yashdeep Phanse, Dr. Supraja Puttamreddy, Dr. Tyasning Kroemer,

Lisa Fraser, Elizabeth Asque, Paul Airs and a great assistance from Brendan

Dunphy. Thanks to Dr. Sijun Liu and Dr. Diveena Vijayendran who gave me constructive advice for my research project on NGS.

All the research could not be successful without the animal care and laboratory assistance from past and current undergraduate researchers including

Kay Kimpston-Burkgren, Cara Fink, Kara Burrack, Zach Flickinger, Katie Schubert, 147

Katie Stueck, Johanna Degan and Alison Masters, Colton Schaben, Kaeli Flaska and Krista Thompson. It was a great experience to work with these students.

Thanks to my parents Kanokthip and Ian Gregory who always support me with love, patience and kindness. They believe in my ability to be a great scientist so that someday I can contribute my knowledge to the benefit of society. Thanks to my sister and her family, Daosai, Johnathan and Dan Evans for endless support. I wish to thank to the Loy family, Dr. Daniel and Laurie Loy and grandparents who always make me feel like Ames is my second home and provide support in many ways.

Thanks to Dr. Kenneth and Dr. Ratree Platt who are advisers for the Thai Student

Association (TSA) and all of the Thai friends that make me feel so at home. I would like to thank my professors, classmates (Vet 64), and friends from Kasetsart

University, Faculty of Veterinary Medicine, Thailand, who helped me develop the foundations of veterinary knowledge.

Finally, this dissertation would not have been possible without the love and the support of the most important person in my life, my beloved husband, Dr. John

Dustin Loy. I would like to thank him for being such an incredible scientist, researcher and professor. He has assisted my research projects in so many ways and always believes in me. I am so grateful that this world brought us together to share the long journey through graduate school together and beyond.