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The Pennsylvania State University The Graduate School College of Agricultural Sciences EXPLORING THE IMPACT OF THE ANTIVIRAL DRUG RIBAVIRIN ON RNA VIRUSES IN HONEY BEES AND THE PRESENCE OF RNA VIRUSES IN BEES IN BRAZIL A Thesis in Entomology by Michael Andrew Thompson Freiberg 2012 Michael Andrew Thompson Freiberg Submitted in Partial Fulfillment of the Requirements for the Degree of Master of Science May 2012 ii This thesis of Michael Andrew Thompson Freiberg was reviewed and approved* by the following: Diana Cox-Foster Professor of Entomology Thesis Advisor Ed Rajotte Professor of Entomology Christina Grozinger Associate Professor of Entomology Craig Cameron Paul Berg Professor of Biochemistry and Molecular Biology Eddie Holmes Professor of Biology Gary Felton Professor of Entomology Head of the Department of Entomology *Signatures are on file in the Graduate School. iii Abstract Honey bees, Apis mellifera L., are essential pollinators worldwide and have an enormous economic impact. Unfortunately, however, honey bee health is declining. Honey bee viruses represent a significant threat to honey bee health and are associated with a variety of pathologies such as dead brood, deformed wings, and general colony mortality. Several viruses have even been associated with the recent mass die-off of honey bees reported in North America called Colony Collapse Disorder. Despite the threat posed by honey bee viruses, there are no chemotherapeutic agents currently in use to treat these viruses and so treatment remains limited to management techniques which are largely ineffective. Beekeepers are thus often forced to destroy infected colonies in order to prevent the spread of viral diseases. I explored the broad-spectrum antiviral drug ribavirin for its potential as an antiviral treatment in honey bees. I present evidence that ribavirin was active against Deformed wing virus (DWV), Black queen cell virus (BQCV), and Sacbrood virus (SBV). I also explored the mutagenic effect of ribavirin on viral genomes. Although further study is required to determine if ribavirin is safe and can improve colony survival, this study serves as an important step in the development of an antiviral treatment for viral infections in honey bees and may also provide insight into the ecology and epidemiology of these viruses. In addition to this study on ribavirin, I explored the prevalence of honey bee viruses in Brazil, both in honey bees and native stingless bees (Apidae; Meliponini). In North America, honey bee viruses have been found in native bees such as the bumble bees (Apidae; Bombus) where they have been shown to cause disease. Transmission can occur through pollen between honey bees and native bees if both species are foraging on the same flowers. The potential for this cross-species transmission poses a health risk to native bees which are likely foraging on the same flowers as infected honey bees. I surveyed honey bee colonies and stingless bee colonies on the campus of the University of São Paulo in Ribeirão Preto, Brazil and present the first evidence of SBV in colonies in Brazil; however, I found no evidence of honey bee viruses infecting stingless bees even in those with colonies near infected honey bees. iv Table of Contents List of Figures .................................................................................................................... vi List of Tables ................................................................................................................... viii Acknowledgments............................................................................................................... x Chapter 1: 1. Thesis Introduction ....................................................................................... 1 The European Honey Bee, Apis mellifera L. ........................................................... 1 a. Summary of Honey Bee Biology, Colony Structure, and Lifestyle ....................1 b. Beekeeping ..........................................................................................................3 2. Non-Viral Pests and Pathogens of the Honey Bee................................................... 5 3. Viral Infections of the Honey Bee ........................................................................... 6 a. Overview of Honey Bee Viruses .........................................................................6 b. Detection of Honey Bee Viral Infections ............................................................8 Chapter 2: Exploring the Impact of the Antiviral Drug Ribavirin on RNA Viruses in Honey Bees ..................................................................................................................... 10 1. Abstract .................................................................................................................. 10 2. Introduction ............................................................................................................ 10 3. Materials and Methods ........................................................................................... 13 4. Results and discussions .......................................................................................... 18 5. Acknowledgments.................................................................................................. 49 Chapter 3: Exploring the Gypsy Moth Caterpillar, Lymantria dispar L., as a System to Test the Infectivity of a DWV Preparation ....................................................................... 50 1. Introduction ............................................................................................................ 50 2. Materials and Methods ........................................................................................... 51 3. Results and Discussion .......................................................................................... 52 Chapter 4: First Report of Sacbrood Virus in Honey Bee (Apis mellifera L.) Colonies in Brazil ..................................................................................................................... 55 v 1. Abstract .................................................................................................................. 55 2. Introduction ............................................................................................................ 55 3. Materials and Methods ........................................................................................... 57 4. Results and Discussion .......................................................................................... 58 5. Acknowledgments.................................................................................................. 59 Chapter 5: Prevalence of Honey Bee Viruses in Native Brazilian Stingless Bees (Apidae; Meliponini) ........................................................................................................ 60 1. Introduction ............................................................................................................ 60 2. Materials and Methods ........................................................................................... 61 3. Results and Discussion .......................................................................................... 62 Chapter 6: Conclusion and Summary of results ........................................................... 66 Appendix: Susceptibility of Encapsidated Viruses to RNase Degradation ...................... 68 1. Materials and Methods ........................................................................................... 68 2. Results and Discussion .......................................................................................... 69 References ......................................................................................................................... 72 vi List of Figures Figure 1: The percent mortality of caged bees fed with 10-ml of sugar water with different concentrations of cordycepin. Each line represents a different cage with its corresponding concentration of cordycepin. Percent mortality is the percent of the total bees that had died at the corresponding time point after treatment began. ........................21 Figure 2: Timeline showing the treatment of ribavirin and other important observations of the four colonies (T1, T2, C1, and C2) from June 2010 to fall 2011 . Asterisks (*) indicate the points in time where colonies were sample and tested for viruses. ................25 Figure 3: RT-PCR analysis for actin mRNA on 8 bees per hive collected at four different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ......29 Figure 4: RT-PCR analysis for DWV on 8 bees per hive collected at three different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ......30 Figure 5: RT-PCR analysis for BQCV on 8 bees per hive collected at four different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ......31 Figure 6: RT-PCR analysis for SBV on 8 bees per hive collected at two different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................31 Figure 7: RT-PCR analysis for IAPV on 8 bees per hive collected at four different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................32 Figure 8: DWV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ...................................34 Figure 9: BQCV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................35 Figure 10: SBV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ...................................36 Figure 11: IAPV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................37 Figure 12: Ion Torrent sequencing coverage across the genome of LSV-2 for RNA extracted from bees from colony T2 (treated colony) collected June 14, 2010 (pretreatment). ....................................................................................................................42 vii Figure 13: Ion Torrent sequencing coverage across the genome of DWV for RNA extracted from bees from colony T2 (treated colony) collected June 14, 2010 (pretreatment). ....................................................................................................................43 Figure 14: Ion Torrent sequencing coverage across the genome of BQCV for RNA extracted from bees from colony T2 (treated colony) collected June 14, 2010 (pretreatment). ....................................................................................................................44 Figure 15: Ion Torrent sequencing coverage across the genome of BQCV for RNA extracted from bees from colony T2 (treated colony) collected July, 7 2010. ..................45 Figure 16: The percent of nucleotides differing from the consensus sequence over a 95bp region of the BQCV genome in the 3’ untranslated region is shown for bees sequenced from the June 14, 2010 collection of T2 (treated colony). This region from the 8435bp loci to the 8530bp loci represents the only region with coverage of greater than 1000bp for the BQCV genome in this sample. ...............................................................................46 Figure 17: DWV RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. ....53 Figure 18: DWV positive strand-specific RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. ............................................................................................53 Figure 19: DWV negative strand-specific RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. ............................................................................................54 Figure 20: Susceptibility of actin mRNA and several honey bee Picornavirles viruses to ribonuclease A and deoxyribonuclease I digestion. ...........................................................70 viii List of Tables Table 1: Summary of the primers used in these experiments for detection of viral genomes using RT-PCR. ....................................................................................................16 Table 2: Summary of the thermal protocols used to amplify cDNA with viral specific primers. ..............................................................................................................................16 Table 3: Summary of the primers used for qPCR viral detection. ....................................17 Table 4: Cage bees were fed 10-ml of sugar water with varying concentrations of cordycepin. The number of dead bees at specific time intervals post-treatment in each cage is reported here. The remaining number represents the number of live bees left in each cage 108 hours after the start of treatment. The total number is the number of bees in each cage at the beginning of the experiment. ...............................................................20 Table 5: Number of dead bees found in ribavirin-treated cages at specific times. Four cages of approximately 200 bees each were prepared and fed 10-ml of sugar water with dissolved ribavirin at different concentrations. Live bees were removed at regular intervals from each cage for testing and the number of dead bees was counted at regular intervals. .............................................................................................................................22 Table 6: Caged bees were treated with ribavirin in sugar water at varying concentrations and a sample of five bees from each cage was removed at successive time points. RNA was extracted from five pooled bees or five individual bees from each cage and timepoint. RNA was tested for Actin, DWV, and IAPV using a RT-PCR assay. ............23 Table 7: Summary of the feeding and ribavirin treatment schedule over the summer 2010. Four colonies (T1, T2, C1, and C2) were set up in an isolated apiary and given sugar water with ribavirin (T1 and T2) or plain sugar water (C1 and C2). .......................26 Table 8: Prevalence of RNA viruses in colonies sampled over the course of treatment with ribavirin. Numbers given indicate how many bees out of eight tested positive for the virus via virus-specific RT-PCR. Ribavirin-sugar water was given to T1 and T2; C1 and C2 were only given sugar water.........................................................................................28 Table 9: Summary of PCR products submitted for Ion Torrent sequencing. ...................48 Table 10: Summary of results from RT-PCR analysis for viruses in 10 colonies at the University of São Paulo in Ribeirão Preto, São Paulo, Brazil. Ten pooled honey bees ix from 10 individual colonies were tested for DWV, IAPV, SBV, BQCV, ABPV, CBPV, and KBV. ...........................................................................................................................62 Table 11: A variety of species of stingless bees from three different meliponiaries were collected and tested for DWV, IAPV, SBV, and KBV. ....................................................64 Table 12: Concentration of RNA extracted from viral homogenate digested with RNase A and DNase I for varying amounts of time. RNA was dissolved in a total of 20µl of RNase free water. ...............................................................................................................70 x Acknowledgments I would like to thank Penn State University and the entire Department of Entomology for the support that they have provided me over the last few years; it has been a pleasure to work among such great scientists. I would like to especially thank fellow graduate students Abby Levitt and Rajwinder Singh and laboratory technicians Amanda Fisher and Monica Kalkstein for invaluable guidance, support, and technical knowledge that they have provided me. I would like to the Mr. Jim Bobb for providing the colonies used in the ribavirin field experiment and Mrs. Harriett Cox for allowing the use of her land for the apiary in Port Matilda, PA. The work in Brazil would not have been as successful or enjoyable without the help and guidance of Dr. David de Jong and numerous other faculty and students at the genetics department of the University of São Paulo in Ribeirão Preto, Brazil. I would like to thank them for their hospitality, guidance, and patience while I was there in June and July 2011. Funding for this project was provided through the Pennsylvania Pollinator Research Grant, which was supported by the Pennsylvania Department of Agriculture, the Pennsylvania Beekeeper’s Association, the Center for Pollinator Research, and the Montgomery County Beekeepers Association. Funding for travel and living expenses while I was in Brazil was provided through the Penn State International Program in Brazil Grant. I would especially like to thank my advisor Dr. Diana Cox-Foster who assisted in countless edits of this thesis and has been a tremendous mentor and teacher over the course of my undergraduate and graduate career at Penn State. Finally, I would like to thank the additional members of my Master’s committee: Dr. Craig Cameron, Dr. Eddie Holmes, Dr. Christina Grozinger, and Dr. Ed Rajotte who provided invaluable feedback and advice over the course of this work. 1 Chapter 1: Thesis Introduction 1. The European Honey Bee, Apis mellifera L. The western honey bee, Apis mellifera L., is a eusocial insect in the order Hymenoptera. It is unique as one of the few domesticated insects and is extremely important to agriculture worldwide. The honey bee produces many commercial products such as honey, wax, and propolis; however, it is most economically important as a pollinator and its impact has been estimated to be in excess of $14 billon in the United States alone (Morse & Calderone, 2000). In addition to being important to agriculture, the honey bee is one of the most well studied insects. Much is known about honey bee biology and natural history and its genome was one of the first insect genomes to be sequenced (The Honeybee Genome Sequencing Consortium, 2006). This makes the honey bee an ideal model organism for a variety of studies including those on pathogen ecology and the evolution of social insects. a. Summary of Honey Bee Biology, Colony Structure, and Lifestyle A honey bee colony builds its hive in a hollow, enclosed cavity such as a hollow tree trunk or a man-made hive designed specifically for keeping bees and there are two primary building materials used: wax and propolis. Wax is secreted from a special gland on the worker bee and is used to build the comb where honey, pollen, and developing brood are kept while propolis is a sticky, resinous plant material used as a sealant within the hive. One colony can contain up to 60,000 individuals; although, the actual population varies depending on the time of year and strength of the colony. This superorganism consists of individuals in one of three different castes: the fertile female gyne (queen), the sterile female (worker), and the fertile male (drone). These castes can be differentiated by morphological features and have distinct roles within the colony. The queen is the only fertile female in the colony. She can be identified by her elongated, wasp-like abdomen and is about twice the size of the worker bees. A normal colony has only one and she is responsible for laying all eggs in the colony, up to 2,000 per day. A queen is reared from a normal female egg but is fed royal jelly which contains 2 a specific factor, royalactin, that triggers the development of female larvae into a queen (Kamakura, 2011). After the virgin queen emerges, she goes on a series of mating flights to collect semen from drones; a queen can mate with multiple drones with an average of 12 (Tarpy & Nielsen, 2002). Once she returns to the hive and starts laying eggs, she cannot mate again and must rely on the sperm that she has stored in an organ called the spermatheca for the duration of her life. With each egg that is laid, she has the option of fertilizing it so that it will develop into a diploid female or laying an unfertilized egg which will develop into a haploid male (Mackensen, 1951). The average queen lives longer than a worker or drone bee, often three to four years, and will be replaced when the colony recognizes certain signals that she has become unproductive (Butler, 1957). The worker bees are sterile females and constitute the majority of the total bees in a colony. Worker bees develop from fertilized, diploid eggs and can live for up to six weeks in the summer and six months in the winter. Most of the tasks associated with maintaining a colony are performed by the workers such as: producing wax, caring for brood, cleaning the hive, defending the hive, storing nectar, feeding and caring for the queen, and foraging for nectar and pollen. A distinct division of labor among the worker bees is often observed; however, the role of one individual bee often changes at different life stages. In general, bees progress over their life cycle from tasks inside the hive such as rearing brood to tasks outside the hive such as foraging for nectar with older bees generally performing the latter tasks. This age-based division of labor is, however, highly variable depending on the needs of the hive and is often difficult to predict (Ribbands, 1952; Rösch, 1925). A worker bee cannot lay fertilized eggs but can lay unfertilized eggs under certain conditions—typically only in a chronically queenless colony—which trigger the ovaries of some worker bees to develop allowing them to lay unfertilized eggs that will ultimately develop into drones. Drones are haploid males, which develop from unfertilized eggs. They are larger than worker bees but smaller than queens, and unlike both workers and queens, they lack a stinger. Their primary purpose is to mate with a queen; although, they do not usually mate with queens from their own colony. During the early afternoon, drones can be found in drone congregation areas (Zmarlicki & Morse, 1963) where they are available to mate with virgin queens. The mating act itself occurs in flight and the drone immediately 3 dies after releasing its semen. The ratio of workers to drones varies throughout the year as there are more drones in the spring and summer and fewer drones in the winter when it is less likely that a colony will need to mate a new queen. All of these individuals function together at the colony level as one superorganism and rely on a complex array of chemical and visual cues to communicate with each other. Examples include queen mandibular pheromone (QMP) which is released by the queen and communicates her presence to the rest of the colony, and the waggle dancing which allows foragers to communicate the location of a food source to other bees (Frisch, 1967). Colonies multiply by a process called swarming during which the current queen leaves the hive with a portion of the workers. The swarm finds a new place to build a hive and the bees in the original hive rear a new queen. It is unclear exactly what causes swarms to occur but it is probable that overcrowding and abundance of food are both important triggers for swarming behavior. Nectar and pollen are the primary nutrition sources for honey bees. Pollen serves as a protein source while nectar serves as a carbohydrate source. The primary source of nutrition of an individual bee differs with the age of the worker and task that the worker is performing; nurse bees consume more pollen in order to be able to synthesize royal jelly for the brood (Crailsheim et al., 1992) while foragers consume much more nectar in order to be able to keep up with the energetic demands of flight. Honey bees also have several methods of storing food for later use. Bees store nectar in the form of honey by dehydrating and storing it in a comb with a wax cap. The dehydrated honey is more energetically dense than nectar and has important antimicrobial properties. Pollen is stored in the form of bee bread which is a mixture of pollen, honey, and enzymes and is modified by microorganisms such as yeast (Gilliam, 1979) and lactic acid bacteria (Vasquez & Olofsson, 2009). b. Beekeeping Humans have used honey from honey bees since ancient times. While it is likely that many hunter-gatherer societies in Europe, Africa, and Asia collected honey from wild colonies, some of the earliest evidence of the domestication of honey bees comes from ancient Egyptian tombs and dates as early as the 7th century BC. Initially, honey bees 4 were likely kept in wooden skeps or other hollow containers. In order to extract honey, the beekeeper removed everything from the hive including the bees, wax, and honey. The obvious disadvantage of this method was that the colony was killed in this process. Today, there are several hive designs commonly used which improve upon older designs but the most common in Europe and the United States is the movable comb hive invented by Lorenzo Langstroth which he described in his landmark book, The Hive and HoneyBee (1853). His innovative hive design not only allowed the beekeeper to be able to inspect his colonies but also allowed him to be able to remove honey from the hive without sacrificing the colony. This transformed beekeeping into a sustainable practice and made it more feasible to selectively breed honey bees since it was now possible for a colony to produce honey for the beekeeper and be kept for multiple years. Because of the numerous benefits of beekeeping, the practice has spread throughout the world including to the Americas where there were originally no native Apis species prior to the arrival of Europeans. While the dominant domesticated honey bee worldwide is the western honey bee, Apis mellifera, the eastern honey bee, Apis cerana F. is commonly kept in much of Asia. A number of recognized subspecies of both the western and eastern honey bees are distributed throughout the world, as well, and are often adapted or have been bred for the specific climate in which they live. An important example of this is the Africanized honey bee, Apis mellifera scutellata Lepeletier, which gradually become the dominate species of managed honey bee in South America, Central America, and parts of North America after it was accidently introduced into southern Brazil in 1956 (Kerr, 1957). Despite the efforts of beekeepers, the Africanized honeybee has spread around the tropical Americas, replacing the European subspecies, largely because of its improved fitness in those climates (Schneider et al., 2004; Spivak, 1992). Today, honey bees are kept for both honey production and pollination. While honey is the most visible and direct product of honey bees, the economic value of honey is dwarfed by honey bee pollination which has been estimated at a value to agriculture in the United States alone in excess of $14 billion (Morse & Calderone, 2000). Honey bees are essential pollinators for many crops including almonds, apples, citrus fruit, and alfalfa and there are many large commercial beekeeping operations that move colonies around the United States in order to pollinate various crops. One example is the California 5 almond pollination which is the largest managed pollination event in the world and involves approximately half of all managed honey bee colonies in the United States (Runckel et al., 2011). Thus, honey bees and the practice of beekeeping are extremely important to agriculture and their study and preservation is essential not only to the global economy but also to the global food supply. 2. Non-Viral Pests and Pathogens of the Honey Bee The health of honey bees is declining around the world and especially in Europe and North America (Potts et al., 2010; VanEngelsdorp & Meixner, 2010). Although this is likely caused by numerous factors, pests and pathogens are a definite contributor to this decline and represent a significant threat to honey bee health. Because of the global importance of honey bees to agriculture, it is vital to be able to understand and control these pests. Honey bee pests and pathogens come from a wide range of taxa and include vertebrates, arthropods, bacteria, fungi, and viruses. Although the prevalence of specific pests varies around the world, there are many which are common for honey bees everywhere. The most problematic vertebrate pests include bears (Ursus species) and skunks (Mephitis species and Spilogale species) which are capable of completely destroying a hive to eat the honey and brood inside, and also house mice (Mus musculus L.) which often seek shelter inside the hive (O’Brien & Marsh, 1990). Important insect pests include the small hive beetle, Aethina tumida Murray, and the wax moth, Galleria mellonella L., both of which can destroy the comb structure of a hive. In addition, a number of insect predators are known to steal honey from honey bee colonies: especially wasps (Hymenoptera: Vespidae), ants (Hymenoptera: Formicidae), and honey bees from other colonies. The fugal pathogens Nosema apis and Nosema ceranae can both cause nonspecific disease symptoms in honey bees and the fungus, Ascosphaera apis, causes chalkbrood disease which results in dead larvae with a “chalky” appearance. The most serious bacterial pathogen of honey bees is Paenibacillus larvae which causes American foulbrood disease. The most significant parasite of honey bees worldwide and in North America is the Varroa mite, Varroa destructor (Anderson & Trueman, 2000). This external parasite 6 plagues honey bees by feeding on the hemolymph of prepupae, pupae, and adult bees and in doing so weakens them and decreases their lifespan. It is also likely that Varroa infestations can make a colony more susceptible to infection by viral pathogens, both by weakening the bee’s immune system and by vectoring the virus itself (Shen et al., 2005a, b; Yang & Cox-Foster, 2005). There are some strains of honey bees which are particularly resistant to Varroa mite infestation and display “hygienic behavior” where bees can sense and remove parasitized brood from the colony in order to control an infestation (Arthi et al., 2000). In addition, it has been proposed that the increased resistance to Varroa mites shown by Africanized honey bees has contributed to, a least in part, their success over European honey bees in the tropical Americas (Guzman-Novoa et al., 1999; Mondragon et al., 2005). 3. Viral Infections of the Honey Bee a. Overview of Honey Bee Viruses There are over twenty known viruses that infect honey bees and new viruses are regularly being discovered (Allen & Ball, 1996; Bromenshenk et al., 2010; Runckel et al., 2011). Most significant viral pathogens are positive, single-stranded RNA viruses in the order Picronovirales and family Iflaviridae or Dicistroviridae. Those in the family Iflaviridae have one open reading frame and include Deformed wing virus (DWV) and Sacbrood virus (SBV). Those in the family Dicistroviridae have two open reading frames and include Black queen cell virus (BQCV), Acute bee paralysis virus (ABPV), Kashmir bee virus (KBV), and Israeli acute paralysis virus (IAPV). An additional RNA virus of pathological importance is Chronic bee paralysis virus (CBPV) which has a different genomic structure and remains unclassified (Olivier et al., 2008). Related to CBPV are another set of RNA viruses called Lake Sinai Virus strains 1 and 2 (LSV-1, LSV-2) which were recently discovered by deep sequencing (Runckel et al., 2011); although, the pathologies, if any, associated with these viruses are unknown. Also of note are a family of DNA viruses invertebrate iridovirus (Iridoviridae), at least one of which is known to infect honeybees (Bailey et al., 1976). 7 Viruses represent a major threat to honey bee health and cause a variety of pathologies. The most common honey bee viruses in North America are DWV, SBV, BQCV, and KBV (Chen & Siede, 2007; Welch et al., 2009). DWV causes bees to develop with deformed wings unusable for flight. SBV causes Sacbrood disease, a condition where larvae fail to pupate and die (White, 1913). BQCV is associated with a blackening of cells containing infected pupae (Bailey & Woods, 1977). KBV does not have clearly defined symptoms but is associated with honey bee mortality (Chen & Siede, 2007). Recently, viruses have also been implicated as a potential cause of the newly described honey bee disappearance called colony collapse disorder (CCD) observed in North America since 2007 (Vanengelsdorp et al., 2009). The newly discovered IAPV was shown to be strongly correlated with collapsed colonies in a metagenomic study (Cox-Foster et al., 2007). In addition, an invertebrate iridovirus has been proposed as a possible cause of CCD (Bromenshenk et al., 2010) although the evidence for this association remains minimal and disputed (Foster, 2011; Knudsen & Chalkley, 2011; Tokarz et al., 2011). A common feature of honey bee RNA viruses and indeed many insect RNA viruses is that they usually persist as latent, unapparent infections but can replicate rapidly and cause severe disease under certain conditions such as stress induced by a severe Varroa mite infection (Martin, 2001). An example of this is DWV which causes honey bees to develop with ragged wings that are unusable for flight. While this virus is common and may be detectable year round within a colony, the symptom is most often seen when the colony is heavily co-infected with Varroa mites during the late summer and fall. Viruses have been found in bees at all life stages including eggs, larvae, pupae, and adults and are known to be transmitted in a variety of ways (Chen & Siede, 2007). The Varroa mite is an important vector and has been shown to be able to transmit DWV, IAPV, and KBV (Bowen-Walker et al., 1999; Chen et al., 2004; Di Prisco et al., 2011). In addition to mites, pollen may also be a vector of honey bee viruses (Singh et al., 2010) providing a way for two insects foraging on the same flower to infect each other. Viruses can also be transmitted vertically from queens and drones to their offspring (Yue et al., 2007). Viruses have also been found in pollen, royal jelly, honey (Chen et al., 2006b; Shen et al., 2005c), and bee feces (Chen et al., 2006a; Hung, 2000) which suggests that 8 they can also be transmitted horizontally between bees in the colony. Inter-taxa transmission is also possible with some honey bee viruses such as DWV which is also able to infect and cause disease in bumble bees (Genersch et al., 2006; Singh et al., 2010). b. Detection of Honey Bee Viral Infections While viral infections can sometimes be diagnosed by their symptoms, most infections remain asymptomatic. This creates a significant challenge for scientists who study and monitor honey bee viruses. Infections cannot be conclusively diagnosed by the beekeeper but instead require molecular methods for accurate detection. Broadly, there are two categories of molecular methods used to detect specific viruses: immunological methods and genome-based methods. Immunological detection methods use viralspecific antibodies to detect unique epitopes of viral proteins. The most commonly used immunological assays include: the enzyme-linked immunosorbent assay (ELISA) and immunodiffusion. Genomic methods, in contrast, derive their specificity by detecting the unique sequence of the viral genome and include: polymerase chain reaction (PCR), reverse-transcriptase polymerase chain reaction (RT-PCR), quantitative polymerase chain reaction (qPCR), microarrays, and viral genomic sequencing. While immunological methods are still in use today for detection of honey bee viruses, they have been largely replaced by the use of genomic methods, specifically RTPCR, which is typically faster, less expensive, and more sensitive than immunological methods (Benjeddou et al., 2001; Chen & Siede, 2007; Genersch, 2005; Grabensteiner et al., 2001; Shen et al., 2005a). RT-PCR consists of several steps. First, viral RNA must be extracted from the honey bee. Next, the RNA is converted into a complimentary DNA (cDNA) library using reverse transcriptase. Finally, the cDNA is amplified by PCR with viral-specific primers; amplification suggests the presence of the viral genome in the original RNA sample. Amplification can be detected by electrophoresis of the PCR product or by monitoring the PCR reaction by real-time PCR (qPCR). Since non-specific amplification can occur, it is often necessary to sequence the PCR product in order to confirm that it is virally derived. In these experiments I used RT-PCR and qPCR for detection of both host-derived and viral-derived RNA. 9 It should also be noted that several controls should be used for the RT-PCR reaction. A negative control reaction, with no cDNA added, is run to check for contamination in any of the reagents. A positive control reaction is run by adding a small amount of the target sequence, in order to verify the viability of the PCR reaction mix and primers. A PCR reaction with extracted RNA, rather than cDNA, can be run to check for DNA contamination of the RNA sample. In addition, an endogenous control gene, such as actin, can be used. Since actin mRNA is transcribed in all honey bee cells, actin can be amplified in order to verify a successful RNA extraction and cDNA reaction and allows for the normalization of differences of RNA extractions and cDNA reactions among different qPCR reactions. 10 Chapter 2: Exploring the Impact of the Antiviral Drug Ribavirin on RNA Viruses in Honey Bees 1. Abstract Honey bee viruses represent a significant threat to honey bee health and are associated with a variety of pathologies such as dead brood, deformed wings, and general colony mortality. It has also been suggested that Colony Collapse Disorder, the recent mass-dieoff of honeybees in the North America, is caused by a virus. Despite the threat posed by honey bee viruses, there are no chemotherapeutic agents currently in use to treat these viruses and so treatment remains limited to management techniques which are largely ineffective. Beekeepers are thus often forced to destroy infected colonies in order to prevent the spread of viral diseases. I explored the broad-spectrum antiviral drug ribavirin for its potential as an antiviral treatment in honeybees. I present evidence that ribavirin may be active against Deformed wing virus (DWV), Black queen cell virus (BQCV), and Sacbrood virus (SBV). I also explored the mutagenic effect of ribavirin on viral genomes. Although further study is required to determine if ribavirin is safe and can improve colony survival, this study serves as an important step in the development of an antiviral treatment for viral infections in honey bees and may also provide insight into the ecology and epidemiology of these viruses. 2. Introduction The western honey bee, Apis mellifera L., is an essential pollinator for numerous crops and has an enormous economic impact (Klein et al., 2007; Morse & Calderone, 2000). Unfortunately, however, honey bee health is declining worldwide (Potts et al., 2010; VanEngelsdorp & Meixner, 2010). This issue of honey bee decline is complex and likely has multiple causes however viruses are an important contributing factor because they are linked to numerous honey bee diseases with a variety of pathologies (Chen & Siede, 2007). For example, Israeli acute paralysis virus (IAPV) has been associated with the recent mass die-off of honey bees in North America called colony collapse disorder (Cox-Foster et al., 2007). Despite the threat posed by viruses, there is little a beekeeper can do to treat them: treatment is largely limited to management techniques such as the 11 removal of infected brood. Consequently, beekeepers are often forced to destroy chronically infected colonies to prevent them from infecting others. This practice, however, limits the productivity and sustainability of beekeeping operations and increases the overall cost by decreasing yield of honey bee products and pollination services. Development of an antiviral treatment is therefore necessary and would provide an essential tool for beekeepers. Some antiviral treatments have been explored such as Remebee (Beeologics), an RNAi based treatment for IAPV, which is currently undergoing FDA and EPA trials (Hunter et al., 2010). While RNAi technology shows promise, it remains to be seen if this can be successfully implemented in the field as a cost-effective way to control the viruses that plague honey bees. The ideal viral treatment would be inexpensive, safe, readily available, and active against a wide range of honey bee viral pathogens. Here I explore the broad-spectrum antiviral drug ribavirin for the potential to serve as an antiviral treatment in honey bees. Ribavirin (1-β-D-ribofuranosyl-1,2,4-triazole-3-carboxamide) was chosen for this study because of its broad-spectrum antiviral properties. It is active against a wide range of viral taxa including both DNA and RNA viruses (Sidwell et al., 1972). The drug is used clinically in humans to treat hepatitis C virus (Davis et al., 1998), respiratory syncytial virus (Wyde, 1998), and lassa fever virus (McCormick et al., 1986). Promisingly, it is active against foot-and-mouth disease virus (De La Torre et al., 1987), human rhinovirus, and poliovirus (Anderson et al., 1992), which are all in the same order, Picornavirales, as many common RNA viruses infecting honey bees. In addition, the generic status of the drug means that it is relatively inexpensive. The mechanism of action of ribavirin is best understood for RNA viruses. Ribavirin is a nucleoside analogue which has considerable structural similarity to purine RNA nucleotides, a property which allows the drug to interfere with several aspects of RNA production. The active form of ribavirin is ribavirin triphosphate (RTP) (Streeter et al., 1973) and the drug is triphosphorylated in the cell by cellular deoxyadenosine kinases (Streeter et al., 1974). RTP is a powerful competitive inhibitor of the enzyme inosine-5'monophosphate (IMP) dehydrogenase which catalyzes a key step in the production of guanosine monophosphate (GMP) (Streeter et al., 1973). Consequently, treatment with ribavirin results in a reduced intercellular concentration of both GMP and its 12 triphosphorylated form, guanosine triphosphate (GTP). A cellular deficiency of GTP can partially explain the antiviral activity and clinical side effects of ribavirin since GTP is essential to cellular and viral processes including protein and RNA synthesis. While this is true, it does not completely explain the antiviral effects of ribavirin (Gilbert & Knight, 1986). At least four additional mechanisms of action of ribavirin have also been proposed (Graci & Cameron, 2006) however the most well studied is that of lethal mutagenesis. Crotty et al. (2000) demonstrated that cellular RTP is incorporated into replicating poliovirus by the virally encoded RNA-dependent RNA polymerase. The ribavirin base can pair equally well with either cytidine or uridine. In this way, ribavirin takes the place of either guanosine or adenosine in the actively replicating viral RNA strand. When the viral genome with incorporated ribavirin is recopied, either cytosine or uracil forms a base pair with the ribavirin and is added to the elongating RNA. Thus, C-to-U and G-toA transition mutations are induced by ribavirin (Crotty et al., 2000). This antiviral strategy of “lethal mutagenesis” relies on the deleterious effects of the drug-induced mutations (Bull et al., 2007). RNA viruses—which have naturally high mutation rates—produce fewer viable progeny with the increased mutation rate induced by the drug. With fewer viable progeny, the infection within a host becomes less severe, potentially allowing the host to better control or eliminate the infection. If the mutation rate reaches a certain “extinction threshold” the infection is no longer able to produce enough viral progeny to sustain itself and the population will go extinct within the host (Bull et al., 2007). Lethal mutagenesis is a mechanism particularly relevant to honey bees since the most important viral pathogens for honey bees are positive, single-stranded RNA viruses in the order Picornavirales and family Iflaviridae or Dicistroviridae. Those in the family Iflaviridae have one open reading frame and include Deformed wing virus (DWV) and Sacbrood virus (SBV). Those in the family Dicistroviridae have two open reading frames and include Black queen cell virus (BQCV), Acute bee paralysis virus (ABPV), Kashmir bee virus (KBV), and Israeli acute paralysis virus (IAPV). An additional RNA virus of pathological importance is Chronic bee paralysis virus (CBPV) which has a different genomic structure and remains unclassified (Olivier et al., 2008). Related to CBPV are 13 another set of RNA viruses called Lake Sinai Virus strains 1 and 2 (LSV-1, LSV-2) which were recently discovered by deep sequencing of migratory colonies (Runckel et al., 2011), although the pathologies associated with these viruses are unknown. The most common honey bee viruses in North America are DWV, SBV, BQCV, and KBV (Chen & Siede, 2007; Welch et al., 2009). In this study, I explore ribavirin for use as a potential antiviral treatment in honey bees by preforming preliminary tests of the drug on both caged bees and live colonies. I address several key questions: (i) What is an effective dose of ribavirin? (ii) Is ribavirin safe for honey bees? (iii) Can ribavirin be used to treat viral infections in honey bees? (iv) What are the effects of ribavirin on the viral genome? Here, I present preliminary evidence that ribavirin is active against several important honey bee viruses including SBV, DWV, and BQCV. 3. Materials and Methods Collection of Bees for Cage Studies A sample of worker bees was collected from a queenless colony kept in a greenhouse which was known to be infected with several viruses. Bees were put into a cold room at 4°C for ten minutes and then separated into mesh cages. Mesh cages with bees were then put in a dark incubator at 35°C with a pan of water to maintain humidity. Bees were given an excess of 50% sucrose solution. Feeding Cordycepin and Ribavirin to Caged Bees A 10-ml solution of 50% sucrose with varying concentrations of cordycepin, a nucleoside analogue used as a positive control, or ribavirin was given to each cage with each cage receiving a different concentration of either cordycepin or ribavirin. Bees were kept in an incubator with a water pan near the fan to maintain humidity. The incubator was kept at a constant temperature of 35°C with no interior lighting. 14 Collection of Samples from Caged Bees Live or dead bees from each cage were collected in a 15 ml sterile screw-top conical tube at successive time points. Bees were then immediately frozen at -80°C for further analysis. Preparation of Colonies Four honey bee nucleus colonies were prepared in an isolated apiary in Port Matilda, Centre County, Pennsylvania at N 40° 47.588 W 78° 4.642 during the early summer 2010. Each of the four colonies was prepared by splitting one parent colony so that all bees and all four queens were produced from the original colony. Each nucleus colony was put in two standard five frame deep wooden hive boxes on top of each other. In this arrangement, the bottom box contained five deep frames while the top box was empty allowing for the placement of a feeder. The four hives were set next to each other, about 0.5 meters apart. The hives were each placed on top of individual cinderblocks with the entrance angled slightly downward and located inside an electric fence in order to prevent attacks by vertebrate pests such as bears. An index card with a unique design was placed above the entrance of each colony. Feeding and Ribavirin Treatment of Colonies A 50% (v/v) sucrose syrup was prepared by mixing food grade cane sugar (sucrose) with an equal volume of tap water. The syrup was then poured into a feeder which was placed inside the hive in the empty box on top of the frames of the box below it. For colonies which received ribavirin treatment, the appropriate amount of ribavirin was first dissolved in distilled water and then added to and mixed with the syrup before treating the hive. Over the course of the summer, two of the colonies were treated with ribavirin while the remaining control colonies received only sugar water. Collection of Samples from Colonies Nurse worker bees were collected by scooping bees from brood comb into sterile 15ml plastic conical screw top tubes. All samples of bees were stored on dry ice until they were transferred to a -80°C freezer where they were stored for further analysis. 15 RNA Extraction Bees were sorted on dry ice and homogenized, either individually or in pools, in 500µl of TRIzol reagent (Invitrogen). RNA was extracted in accordance with the manufacturer’s instructions and suspended in 20µl of RNase-free water (Promega). The concentration of RNA was measured using a SpectraMax 250 spectrophotometer (Molecular Devices), a NanoDrop 2000 spectrophotometer (Thermo Scientific), or the Qubit Fluorometer (Invitrogen) with the RNA protocol. First-Strand cDNA Library Synthesis A complimentary DNA library was made from 5µg total RNA using M-MLV reverse transcriptase (Promega). The cDNA reaction was primed using random primers (Promega, Cat# C1181). Viral Specific PCR Reaction GoTaq green master mix (Promega) was used to amplify cDNA with viral specific primers. Individual reactions were prepared with 1µl of cDNA and forward and reverse primers each at a concentration of 0.4µM. Reactions were subjected to one of three thermal protocols (Table 1) depending on the annealing temperature of the primer. Primer sequences and corresponding PCR temperature protocol are summarized in (Table 2). 16 Table 1: Summary of the primers used in these experiments for detection of viral genomes using RT-PCR. Virus IAPV DWV DWV KBV BQCV SBV ABPV LSV-2 Gene Product (bp) Forward (5'-3') Reverse (5'-3') Thermal Protocol Reference Honeybee Actin 514 ATGAAGATCCTTACAGAAAG TCTTGTTTAGAGATCCACAT PCR1 Shen et al. 2005c Bumble Bee Actin 218 GGAGAAACTTTGTTACGTCGCC CGCACTTCATGATCGAGTTG PCR1 Singh et al. 2010 18s Ribosomal RNA 470 GCCAGCGATCCGCCGAAGTT GCGTGCGGCCCAGAACATCT PCR3 This Thesis Singh et al. 2010 Capsid 840 GGTCCAAACCTCGAAATCAA TTGGTCCGGATGTTAATGGT PCR2 Singh et al. 2010 Capsid (VP1a) 424 CTCGTCATTTTGTCCCGACT TGCAAAGATGCTGTCAAACC PCR2 Capsid (VP1b) 651 GGCGTGGTTCATTAGAATATAGG AAGCAGATCCCCACCTAAAAA PCR2 Singh et al. 2011 Capsid 625 TGTTTGTGGCAATCCAGCTA TACGTCTTCTGCCCATTTCC PCR3 Singh et al. 2012 Capsid 792 TGGCAACCTAGCCATTTAGC GGTAGTGGGAGCTGACCAAA PCR3 Benjeddou et al. 2001 Capsid 210 CACTCAACTTACACAAAAAC CATTAACTACTCTCACTTTC PCR1 Shen et al. 2005c Benjeddou et al. 2001 RNA Polymerase 900 TTATGTGTCCAGAGACTGTATCCA GCTCCTATTGCTCGGTTTTTCGGT PCR3 Runckel et al. 2011 658 CGGCTGGTCTAGCGTGGCTG TGGCAAGCTGTGACGAATCCCT PCR3 Table 2: Summary of the thermal protocols used to amplify cDNA with viral specific primers. PCR 1 1 Hold Temp. (°C) Time PCR 2 94 8 min 1 Hold 35 Cycles 1 Hold 94 51.5 72 72 55 sec 55 sec 1:25 sec 10 min 38 Cycles 1 Hold Temp. (°C) 94 94 55 72 72 Time 8 min 1 min 1 min 1:15 sec 10 min PCR 3 1 Hold 35 Cycles 1 Hold Temp. (°C) 94 94 57.5 72 72 Time 8 min 55 sec 55 sec 1:25 sec 10 min 17 Visualization of PCR Product PCR products were loaded on a 1.5% agarose gel with a 100bp DNA ladder and separated by electrophoresis. Viral Specific Quantitative PCR Reaction TaqMan master mix (Applied Biosystems) was used to amplify cDNA with actin and viral specific quantitative PCR primers (Table 3). Individual reactions were prepared with 1µl of cDNA with forward and reverse primers each at a concentration of 0.1µM in a total reaction volume of 20µl. Reactions were run on a 96-well Fast 7500 qPCR machine (Applied Biosystems). The disassociation curves were verified and the Ct values were calculated using the Fast 7500 Software. Table 3: Summary of the primers used for qPCR viral detection. Virus or Gene Forward (5'-3') Reverse (5'-3') Reference Actin ATGCCAACACTGTCCTTTCTGG GACCCACCAATCCATACGGA Yang et al. 2005 DWV GACAAAATGACGAGGAGATTGTT CAACTACCTGTAATGTCGTCGTGTT Yang et al. 2006 IAPV CGAACTTGGTGACTTGAAGG GCATCAGTCGTCTTCCAGGT Cox-Foster et al. 2007 BQCV GGTGCGGGAGATGATATGGA GCCGTGTGAGATGCATGAATAC Chantawannakul et al. 2005 Runckel et al. 2011 LSV-2 CGTGCTGAGGCCACGGTTGT CCGGTGTCGATCTCGCGGAC Enrichment for RNA Viruses by Filtration and Nuclease Digestion Six bees were homogenized in a total of 2ml of sterile bee Ringer’s solution (155mM NaCl, 3mM KCl, 2mM CaCl2). Homogenate was centrifuged and the supernatant was filtered through a sterile 0.2µm cellulose filter to remove bacteria. Four hundred microliters of filtrate was digested with 50µl each of RNase A (10mg/ml, Affymetrix) and DNase I (1mg/ml, Affymetrix) for 45 minutes at 37°C to digest unencapsidated RNA and genomic DNA. RNA was then extracted from digested filtrate using 3.5ml of TRIzol reagent (Invitrogen). RNA pellet was resuspended in 19µl of RNase-free water and 1µl of SUPERase·In RNase inhibitor (Ambion) and stored at 80°C. Random PCR Amplification of cDNA for Ion Torrent Sequencing First-strand cDNA synthesis was performed using 2µl of each RNA sample and Superscript III reverse-transcriptase (Invitrogen) according to the manufacturer’s 18 instructions. The reaction was primed with tagged random septamers at 0.2µM (5’GCCGGAGCTCTGCAGATATCNNNNNNN-3’) and tagged oligo-dT primers at 0.2µM (5’-GCCGGAGCTCTGCAGATATCNNNNNNN-3’) (Allander et al., 2005; Blomström et al., 2010). Second-strand cDNA synthesis was performed by incubating the sample at 37°C for 1 hour with 1µl of the Klenow fragment (New England Biolabs). The cDNA (10µl) was amplified using Platinum pfx DNA polymerase (Invitrogen) the tag primer (5’-GCCGGAGCTCTGCAGATATC-3’) at a concentration of 0.6µM with the following thermal protocol: an initial denaturation at 94°C for 8 minutes; 40 cycles of 94°C for 30 seconds, 58°C for 1 minute, 68°C for 2 minutes; a final elongation at 68°C for 10 minutes. Ion Torrent Sequencing Samples were digested with EcoRV (New England Biolabs) and RNase A (Affymetrix) for 1 hour at 37°C to remove the PCR tag and excess RNA. Digested samples were purified using the QIAquick PCR purification kit (Qiagen). Samples were then sent to the Penn State Genomics Core Facility – University Park, PA for library preparation and Ion Torrent sequencing. 4. Results and discussions What is an effective dose of ribavirin? In order to estimate an effective dose of ribavirin, the chain-terminating nucleoside cordycepin (3’-deoxyadenosine) was given to honey bees as a positive control. Cordycepin is a toxin originally isolated from Cordyceps, a genus of entomopathogenic fungi (Cunningham et al., 1950). Cordycepin is triphosphorylated by cellular kinases and then incorporated into replicating mRNA in place of adenosine by cellular RNA polymerases. Since cordycepin lacks the 3’-hydroxyl group present on the adenosine nucleoside, the incorporated cordycepin cannot accept an additional nucleoside and mRNA synthesis is thus terminated (Müller et al., 1977). While this mechanism of toxicity in cordycepin is different than the mechanism of action of ribavirin—cordycepin 19 is a chain terminating nucleoside while ribavirin is a mutagen—their mechanisms share a fundamental similarity. They both work by being improperly incorporated into replicating RNA. The toxic effect of cordycepin, mortality, is much easier to observe than the antiviral effects of ribavirin and this feature made cordycepin a good a positive control to estimate an effective dose of ribavirin. While it is not possible to know exactly how the dosage and effect ratio compares between ribavirin and cordycepin, this at least provides a reference point toward estimating an effective dose of ribavirin. Four cages of approximately 100 bees each were prepared. Each cage was given a 10-ml solution of sugar water with cordycepin at one of four concentrations: 0, 0.4mM, 1.0mM, or 1.3mM. These doses were chosen based on active doses of ribavirin in tissue culture (Crotty et al., 2001; De La Torre et al., 1987). The mortality in each cage was measured daily. Comparable mortality was seen in all treatments after a period of 24 hours, suggesting that cordycepin needs at least 24 hours to show an effect. A considerable difference in mortality between treated and untreated bees was seen after 48 hours with the two highest treatments having mortality rates of 53% and 49% while the untreated bees had a mortality rate of only 11%. Table 4 summarizes the number of bees found dead at each time point for each of the four different cordycepin concentrations. Figure 1 depicts the percent of total bees dead at each time point. By the end of the experiment, over 90% of bees in the two highest treatments had died while fewer than 50% of the non-treated bees had died. The lowest cordycepin treatment, 0.4mM, also showed an increase in mortality rate over the non-treatment control, although it was not as large as the mortality seen in the higher treatments suggesting that toxic levels had not been reached in this dose. The significant increase in mortality by cordycepin treatment suggested that ribavirin would also be effective at these dosages and may provide a good reference concentration for the treatment of honey bees with ribavirin. 20 Table 4: Cage bees were fed 10-ml of sugar water with varying concentrations of cordycepin. The number of dead bees at specific time intervals post-treatment in each cage is reported here. The remaining number represents the number of live bees left in each cage 108 hours after the start of treatment. The total number is the number of bees in each cage at the beginning of the experiment. Concentration of Cordycepin (mM) Time (hours) 0.0 0.4 1.0 1.3 24 4 9 13 5 48 10 11 33 35 72 6 16 25 24 96 7 10 7 11 108 11 9 4 3 Remaining 49 42 5 3 Total 87 97 87 81 21 100% 90% 80% Percent Mortality 70% 60% 1.3mM 50% 1.0mM 0.4mM 40% 0 30% 20% 10% 0% 20 40 60 80 Time post-treatmtnet (hr) 100 120 Figure 1: The percent mortality of caged bees fed with 10-ml of sugar water with different concentrations of cordycepin. Each line represents a different cage with its corresponding concentration of cordycepin. Percent mortality is the percent of the total bees that had died at the corresponding time point after treatment began. Is ribavirin safe for honey bees? The next step of this study was to examine the safety of ribavirin on caged bees. Four cages of approximately 200 bees each were prepared from bees in a colony known to be infected with several viruses. After one day in the incubator, each cage was given a 10ml solution of sugar water with dissolved ribavirin at varying concentrations. Three of the cages were given ribavirin at either 0.4mM, 1.0mM, 1.3mM as experimental groups and the fourth cage received only sugar water as a negative control. All cages were also given excess distilled water. After all of the initial sugar water treatment was consumed, the cages were then given an excess of sugar water without dissolved ribavirin. 22 Five live bees from each cage were collected at successive time points. Each cage was sampled three times per day for the first two days of the experiment, twice per day for the next two days of the experiment and then once per day for the next 12 days after which the experiment was terminated. Dead bees were removed and counted at 54 hours and 12 days after the beginning of the treatment (Table 5). Table 5: Number of dead bees found in ribavirin-treated cages at specific times. Four cages of approximately 200 bees each were prepared and fed 10-ml of sugar water with dissolved ribavirin at different concentrations. Live bees were removed at regular intervals from each cage for testing and the number of dead bees was counted at regular intervals. Ribavirin (mM) Time (hours) 0.0 0.4 54 Hrs 6 3 12 Days 54 34 % Mortality 30% 19% 1.0 3 55 29% 1.3 7 67 37% RNA was extracted from five individual bees from each cage from the zero time point collection and the 144 hour post-treatment collection. RNA was extracted from five pooled bees from each cage for each of the 24 hour, 48 hour, 66 hour, and 90 hour posttreatment collections. All RNA extractions were tested by RT-PCR for DWV and IAPV. No significant difference in health or mortality rates (Table 5) was observed among the four treatments or controls suggesting that ribavirin is safe at this dosage. All bees or pools of bees tested positive for DWV while the prevalence of IAPV was minimal and the prevalence of either virus did not appear to change over the course of the experiment. (Table 6). 23 Table 6: Caged bees were treated with ribavirin in sugar water at varying concentrations and a sample of five bees from each cage was removed at successive time points. RNA was extracted from five pooled bees or five individual bees from each cage and timepoint. RNA was tested for Actin, DWV, and IAPV using a RT-PCR assay. [Ribavirin] (mM) 0.0 0.4 1.0 1.3 ∗ Time Post Treatment (hr.) Samples * % DWVVp1a DWV-Vp1a Presence 0 5 100 + 24 1 + 48 1 + 66 1 + 90 1 + 114 5 100 + 0 5 100 + 24 1 + 48 1 + 66 1 + 90 1 + 114 5 100 + 0 5 100 + 24 1 + 48 1 + 66 1 + 90 1 + 114 5 100 + 0 5 100 + 24 1 + 48 1 + 66 1 + 90 1 + 114 5 100 + Samples were extracted as either 5 individual bees (N=5) or a pool of 5 bees (N=1) IAPV was also determined for all samples and found to be missing from all samples except for a single bee at 114 hrs in the 1.3 mm Ribavirin treatment 24 While this result suggested that ribavirin was not effective at clearing a DWV infection within a honey bee, it does not indicate that the drug is ineffective or useless against DWV or any other virus. With the additional deleterious mutations induced by the drug, the virus produces fewer infectious genomes (Bull et al., 2007). Thus, the question of whether the virus remains infectious in the host becomes more relevant than whether the virus remains detectable in the host. In honeybees, a treatment which prevents transmission between individuals in the colony may be sufficient at clearing a viral infection since horizontal transmission between bees within a colony is an important viral transmission route and thus probably contributes to sustaining a viral infection within a colony over successive generations. Even if the ribavirin treatment is capable of rendering DWV uninfectious without completely clearing the virus from the bees, DWV may still be detectable with the RT-PCR assay used in this experiment. Thus, the next step in this study is to treat infected colonies with ribavirin in order to determine whether the drug has an impact on RNA viruses within a colony. Can ribavirin be used to treat viral infections in honey bees? In early June of 2010, four honey bee colonies (T1, T2, C1, C2) were set up in an isolated apiary in Port Matilda, Centre County, Pennsylvania. Near the apiary was a small stream and ample floral resources. This apiary was set up away from other honey bee colonies in order to minimize the potential for ribavirin-contaminated honey to be robbed by other colonies and also minimize the potential for other colonies to infect the experimental colonies. To my knowledge, there were no other apiaries within one mile of this apiary. Each of the four colonies was prepared by splitting one parent colony so that all bees and all four queens were progeny of the queen of the parent colony: this was done in order to minimize genetic diversity among the colonies and to ensure that each colony had the same initial viruses. An index card with a unique design was placed above the entrance of each colony so that honey bees would be better able to identify their own colony in order to minimize drifting. Two colonies (T1 and T2) were chosen as experimental colonies that were given the same ribavirin treatment over the summer in the form of ribavirin dissolved in sugar 25 water. The remaining two colonies (C1 and C2) were chosen as control and given only sugar water without ribavirin. Treatment began on June 30, 2010 and continued until September 19, 2010 (Figure 2). The dose of ribavirin was chosen based on the amount of cordycepin that was shown to cause mortality in caged bees. This number was then extrapolated for the estimated number of bees in each colony. The weekly dose was increased (Table 7) over the summer as the colonies strengthened. Each of the four colonies consumed most of the sugar water or sugar water with ribavirin that they were given and in fact consumed the entire amount of sugar water with ribavirin or plain sugar water that they were given for every feeding after July 7, 2010 (Table 7). During this time, the bees were storing honey, so it is possible that the ribavirin-sugar water solution was stored instead of being immediately consumed. Figure 2: Timeline showing the treatment of ribavirin and other important observations of the four colonies (T1, T2, C1, and C2) from June 2010 to fall 2011 . Asterisks (*) indicate the points in time where colonies were sample and tested for viruses. 26 Table 7: Summary of the feeding and ribavirin treatment schedule over the summer 2010. Four colonies (T1, T2, C1, and C2) were set up in an isolated apiary and given sugar water with ribavirin (T1 and T2) or plain sugar water (C1 and C2). Date 6/10/2010 6/14/2010 6/18/2010 6/30/2010 7/2/2010 7/7/2010 7/9/2010 7/13/2010 7/15/2010 7/20/2010 7/23/2010 7/28/2010 8/2/2010 8/5/2010 8/10/2010 8/15/2010 8/24/2010 8/28/2010 9/5/2010 9/9/2010 9/11/2010 9/19/2010 10/3/2010 Amount of 50% (v/v) Sugar Syrup Given (L) Ribavirin Treatment (mg) T1 T2 C1 C2 T1 T2 C1 3.8 3.8 3.8 3.8 0 0 0 1.9 1.9 1.9 1.9 0 0 0 1.9 1.9 1.9 1.9 0 0 0 1.0 1.0 1.0 1.0 70 70 0 1.0 1.0 1.0 1.0 70 70 0 1.0 1.0 1.0 1.0 70 70 0 1.0 1.0 1.0 1.0 70 70 0 1.0 1.0 1.0 1.0 70 70 0 3.0 3.0 3.0 3.0 210 210 0 3.0 3.0 3.0 3.0 210 210 0 3.0 3.0 3.0 3.0 210 210 0 3.0 3.0 3.0 3.0 210 210 0 3.0 3.0 3.0 3.0 210 210 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 840 840 0 3.0 0.0 3.0 0.0 840 0 0 3.0 3.0 3.0 3.0 840 840 0 3.0 3.0 3.0 3.0 0 0 0 C2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 27 Over the course of summer 2010, the four colonies were generally healthy and flourished, although colonies T1 and C1 were noticeably stronger than T2 and C2. All colonies were observed to have healthy, laying queens at each collection point throughout the summer with the exception of C2 which did not have eggs or a queen when it was checked on September 11, 2010 or September 19, 2010 but was presumably able to requeen because a queen and eggs were found when it was checked again on November 3, 2010. The colonies expanded over the course of the summer and when necessary, an additional 5-frame deep nucleus box was added to the top of the existing one to provide additional room in which the colony could expand. By the end of the summer, both T1 and C1 had a total of three nucleus boxes with full frames, plus one empty box for the feeder while T2 and C2 had only two nucleus boxes full with frames plus empty box for the feeder. At the end of the summer, all four colonies were transferred into ten-frame deep bottom supers for the winter. Colonies T1 and C1 were transferred into 2 ten-frame boxes while T2 and C2 were transferred into one. All four colonies survived the winter and were alive and active in the spring of 2011, at least until May of that year. The colonies were not monitored closely, fed or given ribavirin over the summer 2011 and by fall of 2011 there were no bees in T1 while C1, T2, and C2 were active. The cause of the colony demise in T1 is not known. A sample of bees was collected weekly over the summer 2010 and several additional samples were collected in the spring and summer of 2011. Collections from six different time points were analyzed for viral infections using a reverse-transcriptase PCR (RTPCR) reaction to detect honey bee actin mRNA (Figure 3) and six different honey bee RNA viruses: DWV(Figure 4), KBV, BQCV(Figure 5), SBV(Figure 6), IAPV(Figure 7), and ABPV. A total of eight bees from each colony and time point were analyzed. A summary of results is available in Table 8. 28 Table 8: Prevalence of RNA viruses in colonies sampled over the course of treatment with ribavirin. Numbers given indicate how many bees out of eight tested positive for the virus via virus-specific RT-PCR. Ribavirin-sugar water was given to T1 and T2; C1 and C2 were only given sugar water. VIRUS DATE DWV 6/14/10 7/28/10 9/19/10 4/4/11 5/21/11 COLONY T1 7/8 88% 8/8 100% 8/8 100% 6/8 75% 7/8 88% BQCV 6/14/10 7/28/10 9/19/10 4/4/11 5/21/11 7/8 88% 8/8 100% 7/8 88% 2/8 25% 5/8 63% 8/8 100% 7/7 100% 7/8 88% 7/8 88% 7/8 88% 8/8 100% 7/8 88% 7/7 100% 2/8 25% 8/8 100% 7/8 6/8 4/8 7/8 7/8 88% 75% 50% 88% 88% SBV 6/14/10 8/8 100% 4/4/11 1/8 13% 5/21/11 5/8 63% 8/8 100% 7/8 88% 8/8 100% 8/8 100% 2/8 25% 8/8 100% 7/8 0/8 7/8 88% 0% 88% IAPV 6/14/10 7/28/10 9/19/10 4/4/11 5/21/11 7/8 0/8 0/8 0/8 3/8 88% 0% 0% 0% 38% 8/8 100% 0/7 0% 0/8 0% 0/8 0% 0/8 0% 8/8 100% 0/8 0% 0/7 0% 0/8 0% 2/8 25% 6/8 0/8 0/8 0/8 0/8 75% 0% 0% 0% 0% KBV 6/14/10 0/8 0% 0/8 0% 1/8 13% 0/8 0% ABPV 6/14/10 0/8 4/4/11 0/8 5/21/11 0/8 0% 0% 0% 0/8 0/8 0/8 0% 0% 0% 0/8 0/8 0/8 0% 0% 0% 0/8 0/8 0/8 0% 0% 0% 8/8 7/7 8/8 8/8 8/8 C1 100% 100% 100% 100% 100% 8/8 8/8 7/7 8/8 8/8 T2 100% 100% 88% 100% 100% 8/8 8/8 6/8 7/8 6/8 C2 100% 100% 75% 88% 75% 29 Figure 3: RT-PCR analysis for actin mRNA on 8 bees per hive collected at four different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 30 Figure 4: RT-PCR analysis for DWV on 8 bees per hive collected at three different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 31 Figure 5: RT-PCR analysis for BQCV on 8 bees per hive collected at four different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. Figure 6: RT-PCR analysis for SBV on 8 bees per hive collected at two different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 32 Figure 7: RT-PCR analysis for IAPV on 8 bees per hive collected at four different times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 33 Next, in order to quantify viral titer, pools of six bees were tested for viral titer relative to actin using qPCR. Six bees from each colony and each time point were homogenized together and filtered to remove bacteria. RNA was then extracted from the viral homogenate and a cDNA library was made. The relative viral titer was then quantified using primers for Actin, DWV, BQCV, SBV, and IAPV by qPCR (Table 3). 34 10 T1 (Treatment) T2 (Treatment) C1 (Control) C2 (Control) DWV Viral Titer (2-ΔCt) 1 0.1 0.01 0.001 0.0001 0.00001 0.000001 Collection Date Figure 8: DWV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 35 1 T1 (Treatment) T2 (Treatment) C1 (Control) C2 (Control) BQCV Viral Titer (2-ΔCt) 0.1 0.01 0.001 0.0001 Collection Date Figure 9: BQCV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 36 1 0.1 0.01 T1 (Treatment) T2 (Treatment) C1 (Control) C2 (Control) SBV Viral Titer (2-ΔCt) 0.001 0.0001 0.00001 0.000001 0.0000001 1E-08 Collection Date Figure 10: SBV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 37 1 IAPV Viral Titer (2-ΔCt) 0.1 0.01 T1 (Treatment) T2 (Treatment) C1 (Control) C2 (Control) 0.001 0.0001 0.00001 0.000001 0.0000001 Collection Date Figure 11: IAPV viral titer relative to actin in four colonies at four time points. Treatment colonies were T1 and T2 while control colonies were C1 and C2. 38 All four colonies were tested for DWV, IAPV, BQCV, SBV, and ABPV from the pre-treatment collection (June 14, 2011) and had the same prevalence of these viruses at the beginning of the experiment. They tested positive for DWV, IAPV, BQCV and SBV (Table 8). Only one colony, T2, had one bee that tested positive for KBV suggesting that this infection was probably present but only at very low levels. This finding, that all colonies had the same initial set of viruses at the beginning of the experiment, was expected since the four colonies were split from one parent colony. Over the course of the summer, the prevalence of some viral infections did change, specifically for IAPV, which was detected in all four colonies at the beginning of the summer but not detected in any of the subsequent collections. While viral prevalence did change between time points, it remained consistent among the four colonies regardless of whether they were control colonies or ribavirin treated colonies. Thus, no effect of the antiviral treatment was measurable using the RT-PCR assay described above for any of the collections over the summer of 2010. Interestingly, there was a difference in the prevalence of the virus between the treatment and control colonies for DWV (Figure 4), BQCV (Figure 5), and SBV (Figure 6) in bees collected on April 4, 2011. The prevalence and intensity of the viral bands for these three viruses was greater in the untreated colonies vs. the treated colonies suggesting that ribavirin had an impact on these viruses. This was also observed in the RT-qPCR data for SBV (Figure 10) that saw a much greater increase in SBV between the September 2010 and April 2011 in the untreated colonies compared to the treated colonies although additional replications will need to be done for this data to be significant. It was puzzling, however, that this impact of the drug was not seen until April of 2011, despite the fact that the final treatment with ribavirin was in September of 2010. I propose several hypotheses for further study to explain this result. First, it may be possible that while the drug was treating the viruses, treated bees were constantly becoming re-infected from the environment or other colonies over the course of treatment. Singh et al. (2010) demonstrated that honey bees can become infected with RNA viruses simply by foraging on the same flowers as other infected honey bees or other infected hymenopteran pollinators. Alternately, colonies could have become re- 39 infected through direct contact with infected bees from other colonies such as though robbing or bees which entered the wrong hive. This is more likely to occur during the summer than the winter because bees are more active in the summer. Supporting this hypothesis is the observation that prevalence of these viruses went up in May of 2011 (data not shown). Second, I suggest that while the bees were taking the sugar water with ribavirin from the feeder, it may not actually have been consumed by the bees but instead stored for the winter. In this scenario, it was not until the bees began consuming their winter stores with ribavirin that they were actually receiving the ribavirin treatment. A deeper understanding of the stability of ribavirin in sugar water will be necessary to understand the validity of this hypothesis. Additionally, bees in the winter have a longer lifespan so they would have been exposed to ribavirin for a longer period of time than summer bees. Finally, I suggest that ribavirin was an effective treatment in these bees but was unable to completely clear the infections from treated bees but did decrease the transmissibility of the virus. It was not until new bees were reared in the colony that the infection was cleared from the colony since the drug treatment prevented the new bees from becoming infected. An analysis of the sequence variation will provide more insight into these hypotheses and help to understanding of the impact of ribavirin. Is ribavirin causing a detectable level of mutations in treated viruses consistent with the proposed “lethal mutagenesis” mechanism of the drug? Ribavirin is a mutagen of RNA viruses (Cuevas et al., 2009). To explore this mechanism of action of ribavirin and determine if ribavirin has an impact in our system, I examined the intrahost diversity of viral genomes in treated and untreated bees. I hypothesized that the mutagenic properties of ribavirin will increase the intrahost diversity of viral sequences within the host. Thus, viral genomes in treated colonies will show an increase in intrahost diversity of compared to control colonies. Recent advances in next-generation sequencing (NGS) technology such as 454pyrosequencing, Illumnia, SOLiD, and Ion Torrent provide powerful tools for determining intrahost diversity. Determining intrahost diversity has previously required cloning and sequencing many viral genome fragments (Cuevas et al., 2009; Hoelzer et al., 2010; Murcia et al., 2010); however, it has now been done successfully using NGS 40 (Hoelzer et al., 2010). NGS is faster, more cost-effective per nucleotide sequenced, and is less labor intensive than traditional cloning and Sanger sequencing. Here I explored the Ion Torrent PGM platform (Rothberg et al., 2011) of NGS to estimate intrahost diversity in viral genomes. I first performed a deep sequencing experiment of the transcriptome. Six bees from each of four successive collections from T1 (treated colony) were homogenized together and filtered to remove bacteria. RNA was extracted from each homogenate and a cDNA library prepared from each RNA sample. Each of the four samples was given a unique barcode during the library preparation and the samples were run together on the Ion Torrent PGM sequencer. A total of 4.1 million reads were produced with an average read length of slightly fewer than 200 base pairs. The four datasets were mapped to the genomes of DWV, BQCV, IAPV, SBV, LSV-2, CBPV and honeybee ribosomal RNA using the Bowtie short read aligner (Trapnell et al., 2009) on the Galaxy platform (Blankenberg et al., 2010; Giardine et al., 2005; Goecks et al., 2010). Only five reads aligned from one sample with any viral genome, SBV. In fact, the majority of reads were ribosomal RNA. This suggests that the prevalence of viral RNA relative to ribosomal RNA was much too low for this deep sequencing method to yield useful results. Next, I sought to develop a technique to enrich the homogenate for viral RNA so I explored a method to use RNase A to digest ribosomal RNA. RNase A is an exonuclease specific to single-stranded RNA. Encapsidated viral RNA should be resistant to RNase A degradation because the protein capsid can protect the RNA genome from RNase A degradation. In contrast, free mRNA and ribosomal RNA should be susceptible to degradation by RNase A. This method has been used successfully for metagenomic studies using deep sequencing to identify novel viral pathogens (Blomström et al., 2010). To enrich the preparation for viral RNA, six bees from each of four successive collections from T2 (treated colony) were homogenized and filtered to remove bacteria. The filtrate was then digested with RNase A to remove ribosomal RNA and DNase I to remove DNA. The RNA was extracted from each homogenate and a cDNA library was prepared from each RNA sample using tagged random and oligo-dT primers for the cDNA reaction. The cDNA was then amplified using PCR with the tag primer. Each PCR product was given a unique barcoding sequence during the library preparation and 41 all four samples were run on the Ion Torrent PGM sequencer. The four datasets were mapped to the genomes of DWV, BQCV, IAPV, SBV, LSV-2, CBPV, and honeybee ribosomal RNA using the Bowtie short read aligner (Trapnell et al., 2009) on the Galaxy platform (Blankenberg et al., 2010; Giardine et al., 2005; Goecks et al., 2010). Again, ribosomal RNA sequences dominated the reads; although, many reads did map to viral sequences of LSV-2 (Figure 12), DWV (Figure 13) and BQCV (Figure 14 and Figure 15) and the number of reads for each virus reflected the titer of virus present in the bees at various time points. There was, however, insufficient coverage to use this data to determine intrahost diversity. For DWV and BQCV, reads tended to be biased toward the 3’-end of the genome, possibly because of the tagged oligo-dTs used to prime the cDNA reaction or because of digestion of the 5’-end by the RNase A. This deep-sequencing experiment did reveal some interesting results. First, it showed the presence of the recently discovered virus LSV-2 (Runckel et al., 2011) in one of the samples from T2 collected on June 14, 2010 that I would not have searched for otherwise. The presence of LSV-2 in this sample was later confirmed by RT-PCR and qPCR. I then tested all four colonies from collections at four different time points (June 14, 2010; July 7, 2010; September 19, 2010; and April 10, 2011) using RT-PCR and qPCR; however, LSV-2 was only found in the Jun 14, 2010 collection of T2. Also of note was a mutation analysis of BQCV. This revealed a region in the 3’-untranslated region of the viral genome (Leat et al., 2000) from position 8438 to 8444 with a high incidence of mutations (Figure 16). 42 9000 Sequencing Coverage (Nucleotides) 8000 7000 6000 5000 4000 3000 2000 1000 0 0 1000 2000 3000 4000 Base Position (LSV-2 Partial Genome) 5000 Figure 12: Ion Torrent sequencing coverage across the genome of LSV-2 for RNA extracted from bees from colony T2 (treated colony) collected June 14, 2010 (pretreatment). 6000 43 140 Sequencing Coverage (Nucleotides) 120 100 80 60 40 20 0 0 2000 4000 6000 8000 Base Position (DWV Genome) 10000 Figure 13: Ion Torrent sequencing coverage across the genome of DWV for RNA extracted from bees from colony T2 (treated colony) collected June 14, 2010 (pretreatment). 12000 44 5000 Sequencing Coverage (Nucleotides) 4500 4000 3500 3000 2500 2000 1500 1000 500 0 0 1000 2000 3000 4000 5000 6000 Base Position (BQCV Genome) 7000 8000 Figure 14: Ion Torrent sequencing coverage across the genome of BQCV for RNA extracted from bees from colony T2 (treated colony) collected June 14, 2010 (pretreatment). 9000 45 140 120 Sequencing Coverage (Nucleotides) 100 80 60 40 20 0 0 1000 2000 3000 4000 5000 6000 Base Position (BQCV Genome) 7000 8000 Figure 15: Ion Torrent sequencing coverage across the genome of BQCV for RNA extracted from bees from colony T2 (treated colony) collected July, 7 2010. 9000 46 4.00% Total Mutations per Total Coverage 3.50% 3.00% 2.50% 2.00% 1.50% 1.00% 0.50% 0.00% 8420 8440 8460 8480 8500 Base Position (BQCV Genome) 8520 8540 Figure 16: The percent of nucleotides differing from the consensus sequence over a 95bp region of the BQCV genome in the 3’ untranslated region is shown for bees sequenced from the June 14, 2010 collection of T2 (treated colony). This region from the 8435bp loci to the 8530bp loci represents the only region with coverage of greater than 1000bp for the BQCV genome in this sample. 47 Since neither of the previous techniques provided sufficient data for analysis of intrahost diversity, I resorted to sequencing specific PCR products amplified from RNA from homogenized samples. Samples of six bees were homogenized and filtered to remove bacteria. RNA was extracted and cDNA was prepared using random primers. The cDNA library was amplified using primers for DWV or BQCV with either GoTaq green master mix or Platinum Pfx DNA polymerase and purified. Platinum Pfx polymerase is preferred because it has a smaller error rate compared to GoTaq, however some samples did not amplify with Platinum Pfx so GoTaq was used instead. PCR products from both control colonies and both treatment colonies were used so that the inherent error rate of the sequencing technology could be distinguished from mutations caused by ribavirin. A total of 32 PCR products were sent for sequencing on two independent Ion Torrent runs. Table 9 summarizes the samples that were sent for sequencing. At the time of this writing, sequencing results had been returned by the Penn State Genomics Core Facility and were being analyzed. 48 Table 9: Summary of PCR products submitted for Ion Torrent sequencing. Sample Number Run 1 Run 2 RNA number 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 Colony Collection Date 6/14/2010 346 T1 347 T1 7/7/2010 349 C1 6/14/2010 7/7/2010 350 C1 351 C1 9/19/2010 353 T2 6/14/2010 354 T2 7/7/2010 355 T2 9/11/2010 356 T2 4/10/2011 357 C2 6/14/2010 7/7/2010 358 C2 359 C2 9/11/2010 4/10/2011 352 C1 4/10/2011 360 C2 6/14/2010 357 C2 358 C2 7/7/2010 6/14/2010 345 T1 7/7/2010 346 T1 6/14/2010 353 T2 354 T2 7/7/2010 355 T2 9/11/2010 4/10/2011 356 T2 345 T1 6/14/2010 346 T1 7/7/2010 9/19/2010 347 T1 4/10/2011 348 T1 349 C1 6/14/2010 350 C1 7/7/2010 351 C1 9/19/2010 352 C1 4/10/2011 353 T2 6/14/2010 7/7/2010 354 T2 Primer DNA Polymerase DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab DWV-vp1ab BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV BQCV Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx GoTaq GoTaq Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx Platinum pfx GoTaq GoTaq GoTaq GoTaq GoTaq GoTaq GoTaq GoTaq GoTaq GoTaq Summary of results and ongoing Research Here, I have explored the impacts of the antiviral drug ribavirin on RNA viruses in honey bees. I have presented some evidence that ribavirin may be an effective treatment for honey bee RNA viruses and the results of the most recent sequencing run will help to elucidate the effect of ribavirin on the genome of replicating RNA viruses. While this work is an important step in the development of an antiviral drug for honey bees, much work remains before ribavirin can be used by beekeepers as an antiviral drug. Further 49 research is needed to determine if the drug has an effect on survivorship of the colony, whether the drug is safe for the environment, and whether there is a risk for the evolution of ribavirin-resistant viral strains. In addition, the use of ribavirin will probably need to be strictly limited to colonies which will not produce honey for human consumption since ribavirin is known to be a teratogen in some animal models (Johnson, 1990) and could have potentially devastating effects on a developing fetus if it were unintentionally consumed by a pregnant women. Before publication of this study, the RNA samples extracted from 8 individual bees from each of the four colonies at four different time points will be quantified using qPCR. In addition, the results of pending Ion Torrent sequencing runs will be analyzed. 5. Acknowledgments I would like to thank Jim Bobb for graciously providing and assisting with the preparation of colonies used in this experiment and Harriet Cox for the use of her land for the apiaries in the field studies. I appreciate the help of Deborah Grove and Dan Hannon at the Penn State Genomics Core Facility - University Park, PA. This work was funded by the Pennsylvania Pollinator Research Grant, which was supported by the Pennsylvania Department of Agriculture, the Pennsylvania State Beekeepers Association, the Montgomery County Beekeepers’ Association, and the Center for Pollinator Research. 50 Chapter 3: Exploring the Gypsy Moth Caterpillar, Lymantria dispar L., as a System to Test the Infectivity of a DWV Preparation 1. Introduction Results of the cage study in the previous chapter suggested that ribavirin was ineffective at completely clearing an infection of DWV since it was still detectable in treated bees. Under the lethal mutagenesis model for ribavirin, the drug causes mutations in the replicating virus. With the additional deleterious mutations induced by the drug, the virus is unable to produce as many infectious genomes (Bull et al., 2007). Thus, the question of whether the virus remains infectious in the host becomes potentially more relevant than whether the virus remains detectable in the host. In honeybees a treatment which makes the virus less virulent and pathogenic or less transmissible may potentially be sufficient since bee to bee transmission within the colony may be an important viral transmission mechanism and thus potentially contributes to sustaining a viral infection within a colony. The goal of this experiment was to design and investigate a system to test a DWV preparation for its infectivity. It was necessary to select a non-honey bee system since neither DWV-free bees nor honey bee tissue culture cell lines are available and in this study, I explore the gypsy moth caterpillar, Lymantria dispar L. (Lepidoptera; Lymantriidae), as a system to test the infectivity of a DWV preparation. The gypsy moth caterpillar was a convenient system to try because it was readily available in our department. Although DWV is a honey bee virus, it has been detected in other insects (Genersch et al., 2006; Singh et al., 2010). In addition, previous studies from our lab provided some evidence that DWV is capable of infecting gypsy moth cells in tissue culture; however, from this study I conclude that DWV does not replicate in intact caterpillars. 51 2. Materials and Methods Preparation of DWV homogenate Four bees were cut in half on dry ice; one half was used to confirm the presence of DWV using RT-PCR while the other half was stored at -80°C for use in preparation of the homogenate. The bee halves were homogenized in 2 ml of Grace’s insect media (Sigma), passed through a sterile, 0.2 µm filter, and frozen at -80°C. Injection of gypsy moth caterpillars Newly molted fourth-instar Lymantria dispar L. larva were injected with either 0.5 µl, 1.0 µl or 5.0 µl of either Grace’s insect media or the DWV homogenate. Injections were performed under a stereo dissecting microscope with a sterile 32-guage sharp needle which was inserted at the base of the second proleg. Four caterpillars were injected with each combination of volume and either grace’s media or DWV homogenate; two caterpillars from each combination were immediately frozen at -80°C while the remaining two were reared for five days. Rearing of injected caterpillars The remaining caterpillars, two from each treatment combination, were each put in an individual cup with artificial gypsy moth diet and incubated at 37°C for five days. After five days, caterpillars were frozen at -80°C for further analysis. RNA extraction RNA was extracted from each individual caterpillar by grinding the sample in 500µl of TRIzol reagent (Invitrogen) and extracting the RNA in accordance with the manufacturer’s instructions. RNA library and Strand-specific cDNA synthesis Three different sets of complimentary DNA from each extraction were each made from 5µg total RNA using M-MLV reverse transcriptase (Promega). One set of cDNA reactions was primed using random primers (Promega). The two remaining sets of 52 cDNA reactions were primed using either forward or reverse DWV primers (Table 1) at a concentration of 0.75µM each. DWV specific PCR reaction GoTaq green master mix (Promega) was used to amplify each of the three cDNA with primers for DWV. Individual reactions were prepared with 1µl of cDNA and forward and reverse primers each at a concentration of 0.4µM. Thermal protocols are summarized in (Table 2). Primer sequences and corresponding thermal protocol are summarized in (Table 2). Visualization of PCR product PCR products were loaded on a 1.5% agarose gel with a 100bp DNA ladder and separated by electrophoresis. 3. Results and Discussion The concept behind this technique is based on the ability to detect the anti-sense replicative intermediate of DWV. This is an indicator of viral replication since an encapsidated virion carries the positive-sense genome; however, only an actively replicating infection has the negative-strand replicative intermediate. If just DWV virions are present, then only the positive strand will be detectable in a strand-specific RT-PCR assay. If, however, the virus is actively replicating then both the positive and negative-sense strands will be detectable (Boncristiani et al., 2009). Gypsy moth caterpillars were injected with varying amounts of either Grace’s insect medium (control) or a DWV homogenate (experimental). Half of the caterpillars were frozen immediately while the other half were reared for five days and then frozen. All gypsy moth caterpillars that were not frozen immediately after injection survived for the entire five days before they were frozen. A strand-specific RT-PCR assay was used to detect the positive and negative strand of DWV in each caterpillar. Complimentary DNA primed with random primers was used to detect the presence of DWV in the gypsy moths (Figure 17). Complimentary DNA primed with the DWV reverse primer was used to 53 detect the positive strand of the virus (Figure 18). Complimentary DNA primed with the DWV forward primer was used to detect the negative strand of the virus (Figure 19). Figure 17: DWV RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. Figure 18: DWV positive strand-specific RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. 54 Figure 19: DWV negative strand-specific RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. The positive strand and negative strand of DWV were both present in caterpillars injected with the viral preparation and immediately frozen. This indicates that the virus was actively replicating in the honey bees used to make the viral preparation. After the five day incubation period, however, only the positive strand was detectable suggesting that DWV was probably not actively replicating in this system. Since the encapsidated positive-sense genome is much more resistant to degradation by host RNases than the free negative-strand replicative intermediate, it is much more likely that the positive strand would survive the five day incubation period within the host than the negative strand. Since only the positive strand was detectable after the five day incubation period, it is likely that DWV was not actively replicating in this system. This suggests that DWV was not capable of replicating to high levels over extended time in intact gypsy moths, although it cannot be ruled out that the homogenate itself was not infectious, perhaps due to changes in the virus induced by the freeze-thaw cycle. In addition, the initial period following infection needs to be evaluated to determine if the virus is replicating in the first few hours. Further experiments could include using a fresh homogenate which was not frozen and in which only encapsidated virus is injected. In addition, this experiment could be repeated on gypsy moth cell culture lines which may lack the host defenses of the caterpillar. 55 Chapter 4: First Report of Sacbrood Virus in Honey Bee (Apis mellifera L.) Colonies in Brazil 1. Abstract Sacbrood disease is an affliction of honey bees (Apis mellifera L.) characterized by diseased brood which fails to pupate and dies and this disease represents a significant health threat to honey bee colonies. The disease is caused by Sacbrood virus (SBV), a positive, single-stranded RNA virus in the order Picornavirales which has been found across the world and recently in some countries in South America. I report the first evidence of SBV in honey bee colonies in Brazil. SBV was detected using ReverseTranscriptase Polymerase Chain Reaction (RT-PCR) and confirmed with Sanger sequencing to verify the identity of the PCR product. 2. Introduction The honey bee, Apis mellifera L., is extremely important to agriculture worldwide, not only for its honey production but also for its vital role as a pollinator. In Brazil, honey bee health is declining (Teixeira et al., 2008) and although this is mild compared to the recent die-off’s reported in Europe and North America (Vanengelsdorp et al., 2009) it is still of great concern considering the economic importance of honey bees to Brazilian agriculture. This difference may be related to the fact that the African subspecies, Apis mellifera scutellata, which was introduced to Brazil in 1956 (Kerr, 1957) is now the dominant type of honey bee found in South America. The Africanized subspecies seems to show an increased resistance to varroa mites (Guzman-Novoa et al., 1999; Mondragon et al., 2005). Honey bee health is influenced by numerous factors including parasites, pathogens, pesticides, and nutrition. Viral pathogens are often associated with disease and have even been associated with the recent honey bee die-offs observed in the United States (CoxFoster et al., 2007). At least 22 different viruses are known to infect the honey bee with the best studied being positive sense single-stranded RNA ((+)ssRNA) viruses in the order Picornavirales (Chen & Siede, 2007; Runckel et al., 2011). While many of these viruses can cause 56 disease, they frequently exist in the colony as latent infections that can multiply rapidly under certain conditions such as when the colony is compromised by some other stress such as an infestation with the Varroa mite, Varroa destructor Anderson & Trueman, which can spread the virus (Genersch & Aubert, 2010) and impair the host immune function (Yang & Cox-Foster, 2005). This presents an additional challenge to the diagnosis and monitoring of RNA viral infections in honey bees because molecular techniques are thus required to confirm the presence of the virus rather than visible symptoms observable by the beekeeper. In the past, RNA viruses were detected by immunological methods; however, Reverse-Transcriptase Polymerase Chain Reaction has largely replaced immunological methods as the primary way to detect RNA viruses in honey bees because of improved accuracy and sensitivity as well as economic benefits (Benjeddou et al., 2001; Chen & Siede, 2007; Genersch, 2005; Grabensteiner et al., 2001; Shen et al., 2005a). Many Picornavirales viruses are known to cause disease in honey bees including the Iflavirus Sacbrood Virus (SBV), which is the etiological agent of Sacbrood disease in A. mellifera (Bailey et al., 1964). Sacbrood disease is an important malady of honey bees and is characterized by larvae which fail to pupate and turn pale yellow and eventually dark brown before they die. The virus was first isolated and characterized by Bailey et al. (1964) although the disease itself was first described in 1913 (White) and attributed to a viral infection in 1917 (White). Since the disease was first characterized, it has been found throughout Europe, Africa, Asia, North America, and recently South America where it has been found in both Uruguay(Antúnez et al., 2006) and Argentina (Reynaldi et al., 2010). I report the detection, using RT-PCR, of SBV in managed A. mellifera colonies in the state of São Paulo, Brazil. While a number of other Picornavirales have been found in colonies in Brazil including Deformed wing virus (DWV), Acute bee paralysis virus (ABPV), and Black queen cell virus (BQCV) (Teixeira et al., 2008), this represents the first detection of SBV in Brazilian A. mellifera colonies. 57 3. Materials and Methods Collection of Bees All bees were collected from apiaries on the University of São Paulo campus in Ribeirão Preto, Brazil. Foraging workers were sampled by collection of bees entering the hive while nurse bees were sampled by collection of bees directly on the brood. Brood was sampled by careful removal with tweezers. Samples were stored at -80°C until being processed. RNA Extraction RNA was then extracted using the TRIzol reagent (Invitrogen) according to the manufacturer’s directions. Honey bee RNA was extracted from a pool of 10 bees homogenized together, from individual bees, or from individual brood. The RNA pellet was precipitated with isopropyl alcohol overnight at -20°C. The pellet was then washed with 75% ethanol, resuspended in 20µl of nuclease-free water and stored at -80°C for further analysis. The concentration of RNA in each sample was measured using a spectrophotometer (NanoDrop ND-1000). Reverse Transcriptase PCR First-strand reverse-transcriptase cDNA synthesis was performed on extracted RNA samples using 5µg of total RNA and superscript II (Invitrogen) according to the manufacturer’s instructions. The RNA template was primed using an oligonucliotidedeoxythymidine (oligo-dT) primer. Viral Specific PCR Honey bee cDNA was amplified with SBV primers (SBV-F 5’CACTCAACTTACACAAAAAC-3’; SBV-R 5’-CATTAACTACTCTCACTTTC-3’) specific to a 210bp section in the capsid region of SBV as described by Shen et al. (Shen et al., 2005c). Samples were amplified with Taq DNA polymerase (Promega) with each primer at a concentration of 0.4µM. Reactions were carried out on a thermal protocol consisting of an initial incubation at 94°C for 8 minutes followed by 35 cycles of 94°C 58 for 55 seconds, 51.5°C for 55 seconds, 72°C for 85 seconds with a final incubation at 72°C for 10 minutes. Gel Electrophoresis DNA samples were visualized on a 1.5% agarose gel and compared to a standard 100bp molecular weight DNA ladder. Sequencing PCR products were precipitated with isopropyl alcohol and resuspended in 10µl of nuclease-free water. Samples were prepared for sequencing using the BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) and a GeneAmp PCR System 9600 (Applied Biosystems) in accordance with the manufacturer’s instructions. Samples were then sequenced using an ABI Prism 310 Genetic Analyzer (Applied Biosystems). 4. Results and Discussion An initial survey of bees from 10 colonies in an apiary on the University of São Paulo campus in Ribeirão Preto was done by pooling 10 forager bees from each colony and testing them for SBV using the RT-PCR assay described above. Of the 10 colonies surveyed, one had a positive band of approximately 210bp, suggesting the presence of SBV in this colony. This colony was retested for SBV using the RT-PCR assay on a total of 8 individual forager bees; 3 of the 8 bees tested using the RT-PCR assay were positive for SBV. While the initial colony with SBV was observed to be healthy, an additional colony was tested in a nearby apiary on the campus that was exhibiting symptoms suggestive of SBV. From this colony, eight larvae and eight nurse bees were tested for SBV using the RT-PCR assay described above. Of the larvae tested, 1 of 8 tested positive for SBV. Of the brood nurse bees tested, 4 of 8 tested positive for SBV. The PCR product of each reaction was sequenced and aligned to the SBV complete genome (GenBank accession number AF092924.1) (Ghosh et al., 1999). This represents the first detection of SBV in Brazil, although it is not surprising that SBV has been found there considering its worldwide incidence and the recent findings in 59 neighboring countries (Antunez et al., 2006). With global trade and travel occurring at unprecedented rates, the global spread of viruses and other pathogens can occur faster than ever before. It is thus extremely important to monitor the global spread of pathogens including those that infect honey bees. For SBV in particular, further and more extensive studies will be vital to understanding how the virus spreads as well as the prevalence and impact of the virus around Southern Brazil and throughout the entire country. 5. Acknowledgments I would like to thank the Penn State International Graduate Program in Brazil for providing travel expenses and Dra. Zilá Simões at the University of São Paulo, Ribeirão Preto campus for providing supplies to do the molecular analysis. 60 Chapter 5: Prevalence of Honey Bee Viruses in Native Brazilian Stingless Bees (Apidae; Meliponini) 1. Introduction Stingless bees (Apidae; Meliponini) are a tribe of eusocial bees found throughout the tropics including South and Central America, Australia, Africa, and Asia. There are over 500 species and many are important natural pollinators for both wild fauna and agricultural crops (Slaa et al., 2006). In Brazil and other countries, many techniques of meliponiculture, the management of native bees, have been developed in order to use native bees as managed pollinators (Roubic et al., 1987). One advantage of keeping stingless bees is that they lack stingers, making them easier to manage, although many species display other defense mechanisms such as flying toward and intruder or biting it and injecting formic acid (Roubic et al., 1987). In addition, several species of stingless bees can produce a small amount of honey which can be sold for a premium in Brazil and around the world because of its unique taste and suggested medicinal properties. There have been a few diseases reported in stingless bees (Nogueira-Neto, 1997); however, they are poorly studied and almost nothing is known about their causes (NunesSilva et al., 2009). One potential source of diseases in stingless bees is from honey bees. In North America, some viruses traditionally characterized as honey bee viruses are known to infect species of native bumble bees and can even cause pathology similar to that caused by these viruses in honey bees (Genersch et al., 2006). Viruses can also be transferred between bee species through pollen (Singh et al., 2010) putting native bees at risk for honey bee viruses when they forage on the same flowers as honey bees. This is an additional unintended impact that this non-native species may have on native bees. It is unknown whether honey bee viruses can also infect stingless bees, but it may be possible considering the inter-taxa range already documented in honey bee viruses. Given that stingless bees are important insects for both natural and agricultural ecosystems, it is necessary to explore the potential impact that honeybee viruses may have on stingless bees. In this experiment, I search for viruses in managed stingless bee colonies at three sites—which varied in their proximity to managed honey bee colonies— in the State of São Paulo, Brazil. 61 2. Materials and Methods Collection of Bees Foraging worker honey bees and foraging native bees were sampled by collecting bees entering the hive. Samples were stored at -80°C until being processed. RNA Extraction RNA was extracted using the TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. The RNA pellet was precipitated with isopropyl alcohol overnight at -20°C. The pellet was then washed with 75% ethanol, resuspended in 20µl of nuclease-free water and stored at -80°C for further analysis. The concentration of RNA in each sample was measured using a NanoDrop 1000 spectrophotometer (Thermo Scientific). Reverse Transcriptase PCR First-strand reverse-transcriptase cDNA synthesis was performed on extracted RNA samples using 5µg of total RNA and superscript II (Invitrogen) according to the manufacturer’s instructions. The RNA template was primed using an oligonucliotidedeoxythymidine (oligo-dT) primer. Viral Specific PCR One microliter of cDNA used for each PCR reaction. Samples were amplified with Taq DNA polymerase (Promega) with each primer at a concentration of 0.4µM. Viral specific primers are summarized in Table 1 and corresponding thermal protocols are summarized in Table 2. Gel Electrophoresis DNA samples were visualized on a 1.5% agarose gel and compared to a standard 100bp molecular weight DNA ladder. 62 3. Results and Discussion Ten honeybee colonies at the University of São Paulo in Ribeirão Preto, São Paulo, Brazil were tested for a variety of honey bee viruses in order to first determine what viruses might be present in honey bee colonies on the campus. A sample of ten forager bees was collected from ten different colonies in the apiary on campus. RNA was extracted for the ten bees together and tested using the RT-PCR assay for DWV, IAPV, SBV, BQCV ABPV, CBPV, and KBV. Of the ten colonies, all tested positive for DWV, one tested positive for IAPV, one tested positive for SBV, and three tested positive for KBV. Results are summarized in Table 10. Table 10: Summary of results from RT-PCR analysis for viruses in 10 colonies at the University of São Paulo in Ribeirão Preto, São Paulo, Brazil. Ten pooled honey bees from 10 individual colonies were tested for DWV, IAPV, SBV, BQCV, ABPV, CBPV, and KBV. Colony 1 2 3 4 5 6 7 8 9 10 Genus Apis Apis Apis Apis Apis Apis Apis Apis Apis Apis Species Actin mellifera + mellifera + mellifera + mellifera + mellifera + mellifera + mellifera + mellifera + mellifera + mellifera + DWV-vp1a IAPV SBV BQCV ABPV CBPV KBV + + + + + + + + + + + + + + + Since honey bees may be a source of viruses in native bees, three meliponiaries (places with managed stingless bee colonies) were chosen based on their proximity to honey bee colonies. Meliponiary 1 was located on the campus directly adjacent to the apiary where the original tested honeybees were kept. Meliponiary 2 was a located approximately 500m from the original apiary on campus and was separated by a forests and a lake 100m wide. Meliponiary 3 was a on a farm in São Simão, São Paulo, Brazil with no managed honeybee colonies within foraging distance. A sample of ten bees was collected from ten to twelve different colonies in each meliponiary; genus and species of 63 bees are summarized in Table 11. RNA was extracted for the ten bees together and tested for DWV, IAPV, SBV, and KBV using the RT-PCR assay; these viruses were chosen because they were first found in the tested honey bee colonies. 64 Table 11: A variety of species of stingless bees from three different meliponiaries were collected and tested for DWV, IAPV, SBV, and KBV. Colony Number Meliponiary 1 1 2 3 4 5 6 7 8 9 10 Meliponiary 2 11 12 13 14 15 16 17 18 19 20 Meliponiary 3 21 22 23 24 25 26 27 28 29 30 31 32 Genus Tetragona Tetragona Frieseomelitta Scaptotrigona Partamona Melipona Melipona Nannotrigona Leurotrigona Plebeia Melipona Melipona Scaptotrigona Scaptotrigona Nannotrigona Plebeia Tetragonisca Tetragona Partamona Frieseomelitta Frieseomelitta Plebeia Marmelita Scaptotrigona Scaptotrigona Tetragonisca Frieseomelitta Melipona Melipona Melipona Nannotrigona Scaptotrigona Species clavipes angustula varia bipunctata helleri quadrifasciata marginata testaceicornis muelleri remota scutellaris quadrifasciata bipunctata depilis testaceicornis droryana angustula clavipes helleri varia polistica polysticta angustula preta scutellaris quadrifasciata testaceicornis - 65 These native bees were tested for honey bee viruses and no evidence of these viruses was found in any colonies. While this preliminary result is hopeful, more work needs to be done to understand whether honey bee viruses or other pathogens are infecting native bees. This study has several limitations that should be addressed in future studies. First, the sampling was severely limited in scope. Future studies should involve sampling from more meliponiaries and wild stingless bee colonies from across Brazil and in other countries were stingless bees are found. Second, the RT-PCR assay used to detect viruses is limiting. It is unknown what the minimum infection threshold is for detecting viruses is, but it is possible that a non-severe infection could potentially go undetected with the RT-PCR assay. Also, since the RT-PCR assay is sequence-specific, mutations in the viral genome—which may have been necessary for the cross-species transmission of viruses—could make the genome undetectable using the same RT-PCR primers if the mutations occurred at sites where the PCR primers normally bind. 66 Chapter 6: Conclusion and Summary of results Unfortunately, honey bee health is threatened. Both managed and wild colonies are dying in many places around the word, especially in North America and Europe (Potts et al., 2010). Although it is not clear what is causing this decline, it is likely due to a variety of factors which may include: pesticides, pathogens, human activities, and management techniques. More research will be necessary to understand this problem and to work toward a solution to help save honey bees. One cause of honey bee disease is from a variety of single-stranded RNA viruses in the order Picornavirales. These can cause a variety of diseases ranging from specific symptoms such as deformed wings to nonspecific honey bee mortality. Because of the threat of viruses, the development of a treatment for honey bee viruses has the potential to help beekeepers around the world. In addition, the lack of a virus-free bee makes the study of viral transmission in honey bees difficult; a drug capable of clearing viral infections in honey bees could help researchers who study these viruses. In this thesis, I explored the drug ribavirin as a potential antiviral agent in honeybees. I first tested the drug by feeding it to caged honey bees and find evidence to suggest that ribavirin does not induce mortality in honey bees at potentially therapeutic doses. I next tested the drug on actual colonies by feeding the drug to bees in a sugar water solution. I found that ribavirin does seem to have an impact on viral prevalence; although, this impact was not detectible until the spring after bees were treated with ribavirin. Finally, I explored several techniques to determine the impact of ribavirin on the intrahost diversity of viral genomes within honey bees. While these results are promising, it will be a long time before ribavirin can be used as a chemotherapeutic agent by beekeepers. A myriad of questions remain and further studies should focus on whether drug treatment has an effect on survivorship of the colony, whether the drug treatment poses a risk of contaminating honey or other bee products that may be consumed by humans, whether the drug is safe for the environment, and whether the evolution of ribavirin-resistant viral strains is possible. Given that ribavirin is a teratogen in some animal models and probably humans as well, it is unlikely that it would be approved for use by beekeepers; other antivirals may offer greater safety. 67 For researchers who can maintain isolated colonies or cages of bees, this treatment does offer the promise of being able to eliminate virus from those bees to allow for virus-free bees. In the next part of this thesis, I searched for honey bee viruses in managed stingless bee colonies in Brazil. Transmission of honey bee viruses to other hymenopteran pollinators has been documented and represents an unintended negative consequence to native pollinators from honey bees which are not native to the Americas. While I found no evidence of honey bee viruses in the native bees that I tested, a more in-depth study is necessary to fully understand what risk honey bee viruses have on native stingless bees in Brazil and throughout the tropics. Honey bees are unique because they are one of very few insects that have been successfully domesticated. Humans have used honey from honey bees for thousands of years because it has traditionally been one of the only available sweeteners. In recent times, the role of honey bees as managed pollinators has become vital to agriculture and the global food supply because they are often only insect capable of effectively pollinating large-scale monocultures common to modern agriculture in developed countries. The impact of honey bees is evident in many aspects of culture around the world from simple works of art to multi-million dollar movies (Hickner & Smith, 2007). In short, honey bees are extremely important to humans. Thus, research that helps to understand and help honey bee health is extremely important and worthwhile. It is my sincere hope that this work has contributed to our understanding of viruses in honey bees and can be developed further to provide additional tools to help save this invaluable insect. 68 Appendix: Susceptibility of Encapsidated Viruses to RNase Degradation 1. Materials and Methods Preparation of Viral Homogenate Eighteen bees were homogenized in a total of 6mL of sterile Bee Ringer’s Solution (155mM NaCl, 3mM KCl, 2mM CaCl2). Bees used were taken from a colony known to be infected with several viruses and were stored at -80°C prior to use in this experiment. Homogenate was centrifuged and the supernatant was filtered through a sterile 0.2µm cellulose filter. Nuclease Digestion Five aliquots of 400µl filtered homogenate were prepared. TRIzol reagent (Invitrogen) (2.5ml) was immediately added to the zero time point. Fifty microliters each of RNase A (10mg/ml, Affymetrix) and DNase I (1mg/ml, Affymetrix) were added to each aliquot. Each aliquot was incubated at 37°C for a varying amount of time: 15 minutes, 60 minutes, 90 minutes, and 120 minutes after which the enzymatic reaction was stopped by the addition of 2.5ml of TRIzol reagent. RNA Extraction RNA was extracted from digested homogenate with TRIzol reagent according to the manufacturer’s instructions. The resulting RNA pellet was then resuspended in 20µl of RNase-free water. The RNA concentration of each sample was measured using a NanoDrop 2000 (Thermo Scientific). RNA was stored at -80°C for further analysis. Complimentary DNA Library Preparation First-strand cDNA synthesis was performed using 2µl of each RNA sample and Superscript III reverse-transcriptase (Invitrogen) according to the manufacturer’s instructions. The reaction was primed with tagged random septamers at 0.2µM (5’GCCGGAGCTCTGCAGATATCNNNNNNN-3’) and tagged oligo-dT primers at 0.2µM (5’-GCCGGAGCTCTGCAGATATCTTTTTTT-3’) (Allander et al., 2005; Blomström et 69 al., 2010). Second-strand cDNA synthesis was performed by incubating the sample at 37°C for 1 hour with 1µl of the Klenow fragment (New England Biolabs). PCR Amplification GoTaq green master mix (Promega) was used to amplify cDNA with primers specific to Actin and DWV, IAPB, SBV, and BQCV. Individual reactions were prepared with 1µl of cDNA and forward and reverse primers, each at a concentration of 0.4µM. Reactions were subjected to one of three thermal (Table 2) depending on the annealing temperature of the primer. Primer sequences and corresponding thermal protocol are summarized in Table 1. Visualization of PCR product PCR products were loaded on a 1.5% agarose gel with a 100bp DNA ladder and separated by electrophoresis. 2. Results and Discussion This technique was being explored as a method to enrich an RNA preparation for viral RNA while removing the excess mRNA and ribosomal RNA. It has been used successfully for metagenomic studies using deep sequencing to identify novel viral pathogens (Blomström et al., 2010). RNase A is an exonuclease specific to singlestranded RNA. Encapsidated viral RNA should be resistant to RNase A degradation because the protein capsid can protect the RNA genome from RNase A degradation. In contrast, free mRNA and ribosomal RNA is susceptible to degradation by RNase A. A honey bee homogenate was prepared and passed through a 0.2µm cellulose filter to remove bacteria and other contaminates. The homogenate was then divided into aliquots and digested with DNase I (to remove genomic DNA) and RNase A (to remove mRNA and rRNA) for varying amounts of time. Aliquots from the same initial homogenate were used to ensure that each sample had the same viruses at the same concentrations before digestion. The RNA was then extracted from each digested homogenate and the concentration of each RNA sample was measured on a spectrophotometer (Table 12). RNA was then tested for the presence of viral RNA using an RT-PCR assay (Figure 20). 70 Table 12: Concentration of RNA extracted from viral homogenate digested with RNase A and DNase I for varying amounts of time. RNA was dissolved in a total of 20µl of RNase free water. Digestion Time (min) RNA Concentration (ng/µl) 0 1422.7 15 611 60 92 90 138.9 120 99.4 Figure 20: Susceptibility of actin mRNA and several honey bee Picornavirles viruses to ribonuclease A and deoxyribonuclease I digestion. As expected, the concentration of RNA extracted generally decreased with a longer digestion time. It decreased between the 0 and 15 minute digestion and between the 15 and 60 minute digestion but remained similar for subsequent digestion times (Table 12). 71 This suggests that the nucleases are effectively digesting some but not all of the RNA and some RNA is resistant to digestion. Not surprisingly, Actin mRNA was found in the undigested homogenate but was not present in the digested homogenate. DWV, SBV, and BQCV were all found in the undigested samples, however, only IAPV and DWV were found in the digested samples suggesting that DWV and IAPV are resistant to RNase A digestion while SBV and BQCV are not. It was also observed that the smallest amount of PCR product for both DWV and IAPV was found in the 120 minute digestion, suggesting that viral RNA was present in a lower amount in this sample compared to less digested samples. This is likely because these viruses are not completely resistant to RNase A digestion. IAPV was only found in digested samples suggesting that the concentration of IAPV relative to other RNA in the undigested samples is much lower than in digested samples and that the nuclease digestion enriched for this virus and increased its relative concentration above a certain detectability threshold. It is unknown why these viruses have different susceptibilities to RNase A digestion. One hypothesis is simply that the amount of virus present in the homogenate was much greater for DWV and IAPV so they required a longer digestion time to be completely removed. Another hypothesis is that the capsids differ in permeability to RNase or stability which could be altered by the specific buffer composition. A third hypothesis is that DWV and IAPV contain a specific factor which makes them resistant to RNase A digestion. For example, Yang and colleagues (2004) demonstrated that a specific protease-sensitive factor protects newly synthesized Hepatitis C viral RNA from nuclease degradation and it is possible that such a factor could exist for DWV or IAPV as well. A fourth hypothesis is that DWV and IAPV are localized in a cellular vesicle or other protected cellular structure. The vesicle, not the capsid, was shielding the genome from RNase A. 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