The importance of honey bees to the world economy far surpasses their contribution in terms of honey production; they are responsible for up to 30% of the world' s food production through pollination of crops. Since fall 2006, honey bees in the U. S. have faced a serious population decline, due in part to a phenomenon called Colony Collapse Disorder (CCD), which is a disease syndrome that is likely caused by several factors. Data from an initial study in which investigators compared pathogens in honey bees affected by CCD suggested a putative role for Israeli Acute Paralysis Virus, IAPV. This is a single stranded RNA virus with no DNA stage placed taxonomically within the family Dicistroviridae. Although subsequent studies have failed to find IAPV in all CCD diagnosed colonies, IAPV has been shown to cause honey bee mortality. RNA interference technology (RNAi) has been used successfully to silence endogenous insect (including honey bee) genes both by injection and feeding. Moreover, RNAi was shown to prevent bees from succumbing to infection from IAPV under laboratory conditions. In the current study IAPV specific homologous dsRNA was used in the field, under natural beekeeping conditions in order to prevent mortality and improve the overall health of bees infected with IAPV. This controlled study included a total of 160 honey bee hives in two discrete climates, seasons and geographical locations (Florida and Pennsylvania). To our knowledge, this is the first successful large-scale real world use of RNAi for disease control.
The importance of honey bees as pollinators of crops to the global economy far surpasses their contributions in terms of honey production [1]. In all, 52 of the world' s 115 leading agricultural crops rely on honey bee pollination to some extent. These crops represent approximately 35% of the human diet [2]. Insect pollination, which is provided predominately by honey bees, is estimated to have a value of US$ 212 billion [3]. Honey bee populations have been decreasing globally in recent years [4]. Since fall 2006, honey bees overwintering in the U. S. A. have faced unusually high rates of mortality, in part because of a phenomenon now known as Colony Collapse Disorder (CCD) [5]. Several hypotheses have been offered to explain CCD and existing and emerging pathogens have been implicated either directly or indirectly [6]. Colonies affected by CCD are infected with larger numbers of pathogenic organisms than control colonies, yet no single pathogen was found associated with all affected colonies [7]. In another effort, researchers did find that single-stranded RNA viruses, specifically picorna-like viruses, occurred at elevated levels in CCD colonies. These elevated levels of viruses may interfere with gene transcription, thus reducing immune response competence and pesticide detoxification capabilities, subsequently leading to premature death of infected bees [8]. Honey bees are susceptible to a host of picorna-like viruses, including the closely related Acute Bee Paralysis Virus (ABPV), Kashmir Bee Virus (KBV), and Israeli Acute Paralysis Virus (IAPV) [9], [10]. The latter of these three viruses was identified as a good marker for CCD in initial studies, especially when found in association with the microsporidia Nosema sp. [6]. While IAPV is probably not the sole cause of CCD [7], its ability to cause increased mortality in honey bees has been established [11]. The process of post-transcriptional gene silencing is thought to be an evolutionarily-conserved cellular defense mechanism used to prevent the expression of foreign genes and is commonly shared by diverse flora and phyla [12]. The presence of long double-stranded RNAs in cells stimulates the activity of a ribonuclease III, Dicer, which is involved in the processing of the double stranded RNA (dsRNA) into short interfering RNAs (siRNAs). The RNAi response also features an endonuclease complex, commonly referred to as an RNA-induced silencing complex (RISC), which mediates cleavage of target ssRNA having sequence complementary to the antisense strand of the siRNA duplex. [13], [14], [15], [16]. In a variety of organisms, exogenously applied dsRNA or their siRNA derivatives, can be used to arrest, retard or even prevent a variety of pathogens. In some of these organisms, such as plants and the nematode C. elegans, an amplification stage follows the initiation stage of gene silencing, involving an RNA dependent RNA Polymerase (RdRp), which may lead subsequently to degradation of RNAs outside the initial dsRNA region of homology [17]. RNAi can spread from the initial site of dsRNA delivery, producing interference phenotypes throughout the treated animal. To serve as a preventive or curative strategy, amplification and systemic spread of the silencing signal are both paramount. In some invertebrates, including honey bees, a systemic interference defective (SID) gene encodes a transmembrane protein that is an important participator in the systemic RNAi pathway. Apparently, these SID1-like proteins channel dsRNAs between cells, enabling systemic spread of the silencing signal [18], [19]. Although a canonical invertebrate RNA dependent RNA Polymerase (RdRP) homologue has not yet been described, there is evidence that such RdRp activity may occur via other enzymes, leading to amplification of the silencing signal in insects [20]. IAPV specific dsRNA (Remebee-IAPV or herein Remebee-I) was used successfully to prevent bees from succumbing to infection from IAPV in small scale lab experiments whereas bees fed Green Fluorescent Protein (GFP) dsRNA and virus died in a manner similar to the IAPV fed control bees [12]. Although these results were exciting per-se, transferring RNAi from a well characterized and efficient tool in the lab and making it successful in preventing the adverse effects of virus infection in the field, remains notoriously difficult. We present the first large-scale real world successful use of RNAi for disease control. We attempted to determine if IAPV specific homologous dsRNA can be used to reduce impacts from IAPV infection in 160 honey bee hives in two discrete climates, seasons and geographical locations (Florida and Pennsylvania). To our knowledge, this is the first successful demonstration of the use of RNAi as a preventative treatment for an insect disease on such a large scale.
The field demonstration in FL was designed in a manner that permitted us to follow IAPV-infested bee colonies (some given Remebee-I and others not) for six weeks. One hundred standard colonies of honey bees were split into 5 groups with 20 colonies per group. Four groups were located within 100 m of one another (non-isolated) while a 5th group was isolated from the remaining four by at least 3. 2 km to measure any environmental effects due to location. Treatment allocations (20 colonies per treatment) were as follows: Treatment 1 – no treatment – non isolated Treatment 2 – Remebee-I only – non isolated Treatment 3 – Remebee-I+IAPV – non isolated Treatment 4 – IAPV only (fed in sugar water solution) – non isolated Treatment 5 – no treatment– isolated Honey (or net weight gain) is the ultimate proxy to the total active population of the hive. The non treated control produced the most honey in PA, but not in FL. In FL, colonies treated with Remebee-I+IAPV produced significantly more honey than colonies receiving IAPV alone (Figure 2, N = 40, p<0. 03). In PA, the difference between the weight at the start and the end of the experiment (4 months) shows that the non infected controls gained the most weight (mean gain = 23. 5kg), whereas Remebee-I+IAPV had gained slightly less (mean = 21kg). Both made significantly greater weight gains compared with the group receiving IAPV alone (mean = 16. 3kg) (Figure 3 F = 2. 7; df = 4. 92; P = 0. 034). Subsequent trials done under a similar protocol were repeated in the winter of 2009–10 in FL and in California (CA). Samples of bees were collected just before IAPV challenge and 2-weeks post treatment. Northern analysis was done with IAPV specific sequence probes corresponding with the Remebee-I sequence. The results of these are presented in detail in Supporting Information S2. High levels of discrete Dicer Remebee metabolites are evident in Remebee-I treated hives prior to IAPV challenge up to four weeks after a Remebee-I application. Non- Remebee-I treated bees are mostly negative, but a low signal was detected in some colonies. Subsequent to IAPV challenge, levels of siRNAs and IAPV metabolites are highly elevated in both Remebee-I and in non-Remebee-I treated hives, showing that production of dsRNA is a natural defense mechanism in bees against IAPV infection. Varroa levels are unaffected by treatment. The change in varroa prevalence on adult bees did not differ significantly between treatment groups in either the FL (F4,95 = 2. 39; P = 0. 056) or PA (F2,57 = 1. 03; P = 0. 3642) trials. Nosema levels: While the change in nosema levels in the FL trial was not significantly different between treatment groups (F4,95 = 0. 47; P = 0. 7586), a highly significant difference in nosema spore levels was detected at the end of the PA trial (F2,57 = 8. 62; P = 0. 0005) (Figure 4). Indeed, within the duration of the experiment, nosema spores levels increased in the IAPV treated group, yet went down in both Remebee-I+ IAPV and uninfected control colonies. A brief summary of findings is presented in Table 1.
The negative effects of IAPV on honey bee health and colony vigor is evidenced by lower honey weight gains (Figures 2 and 3). Environmental factors and terrain also influence the availability of forage and thus can seriously reduce or increase the effects of virus infection on a hive by altering foraging patterns. Large scale examination of the RNAi treatment under such varied conditions at two separate and diverse locations (i. e. FL and PA) was challenging. However, one would expect that 7–8 weeks after a young queen begins to oviposition prolifically (see Supporting Information S1 for brief overview), all hives would be overflowing with bees. This was evidently not the case in this situation, and the IAPV only group reversed in total population and bee/brood ratio. Thus, we observed relative de-population of the hive with probable greater loss of foragers in IAPV only infected hives. We conducted the trials described herein in spring and summer, whereas CCD is a mostly winter phenomenon [1], [7]. This may be the reason that we did not see many hives devastated by CCD. However, subsequent trials performed in the 2009–10 winter in Florida and California resulted in 40% and 60% collapse of hives, respectively (Hunter Wayne. and Oliver Randy. , personnel communication). Beyond cold weather providing additional stress on the bees, some of the difference may be attributed to the viruses' suppressors of gene silencing. In plants, these viral suppressors of gene silencing have more optimal enzymatic kinetic coefficients under cold temperatures in relation to the silencing enzymes, often leading to more acute virulence [21]. This could help explain the initiation and overall devastation of hives in the U. S. following winter cold snaps (Dennis VanEngelsdorp unpublished observations). Our hypothesis that Remebee-I would protect bees from IAPV infection was supported by multiple observations: First, in FL, the Remebee-I+ IAPV treated hives were the only colonies with significantly increasing numbers of bees during the study. In PA bees increased within all treatments with no significant differences. Some of the difference between the two trials could account for this difference in observations. In PA the virus was introduced twice into the colonies within a three day period (instead of once), and the total amount of virus introduced was thus much higher than the FL trial. The PA trial starter hives were weaker in strength, and had more collapses of hives across all treatments prior to infection than those starting the FL trial. Second, although bee to brood ratios started lower in Remebee-I treated hives it became stronger and remained so until the end of the trial. The change in the ratio is attributed to changes in the adult bee counts, since the capped brood coverage was the same between treated and non-treated bees, thus these hives also contained more adult foragers, which resulted in significantly more honey production over the IAPV only treatment (Figure 2,3). Furthermore, in subsequent trials, molecular evidence now proves that Remebee-I is active in the Remebee-I + IAPV treated groups, as determined by the presence of siRNA (see Supporting Information S2). The strong presence of siRNAs probably restricts the severity of the disease in the bees leading to a longer life-span and subsequently to an overall greater number of bees, with more foragers and consequently a greater yield of honey. It is interesting to note the natural occurrence of these siRNAs in bees receiving IAPV challenge. Presence of these siRNA in non- Remebee-I treated hives prior to infection may be a result of natural virus infection prior to the challenge, or by transcription of integrated viral sequences in the bee genome [22]. IAPV specific dsRNA (Remebee-I) was used successfully to prevent bees from succumbing to infection from IAPV. The results further demonstrate the possibility to produce targeted treatments for bee pathogenic diseases. These field results demonstrate the successful application of dsRNA as a viable treatment to solve a real world problem, which may further lead to concerted efforts to utilize this ubiquitous natural mechanism, RNAi, for the benefit of the bees, beekeepers, and hopefully to other applications in agriculture and veterinary health.
Essentially as described in [11]. Approximately 40 adult forager honey bees were collected from 10 colonies in a Florida bee yard (apiary) where CCD had been reported. Each bee was processed individually and tested using rtPCR for the presence of Israeli Acute Paralysis Virus (IAPV), genome – NC_009025; Acute Bee Paralysis Virus (ABPV) genome – NC_002548; Kashmir Bee Virus (KBV) genome – NC_004807; Black Queen Cell Virus (BQCV), genome-NC_003784; Deformed Wing Virus (DWV) genome NC_004830. All bees had more than one virus detected so inoculum was prepared from bees which tested positive only for IAPV+KBV by homogenizing the bees with glass beads in small amounts of 10 mM buffer phosphate, pH 7. 2 containing 0. 02% DETCA (Sigma-Aldrich Cat #22,868-0). Inoculum was prepared by passing the virus solution through a syringe filter, 0. 45 µm, to remove bacteria, after which ∼10 µl were administered by microinjection along the lateral side of the abdomen of ∼700 pupae using a Hamilton syringe with a 30Gx½ gauge sterile needle. Inoculated pupae were kept in petri dishes covered with slightly damp filter paper and maintained at 21–23°C for three days to permit virus replication. On the third day, batches of about 50 pupae were homogenized with glass beads. Small amounts of 10 mM buffer phosphate (pH 7. 2 contained 0. 02% DETCA, Sigma-Aldrich Cat #22,868-0) were added to the homogenates. The homogenates were collected in a beaker volume adjusted to ∼350 ml with buffer (see above) and mixed. Each sample was split into two 250 ml centrifuge tubes and centrifuged at 300×g (∼1,400 rpm) on a GSA rotor for 20 min. The supernatant (S1) was collected and kept at 4°C for 3 d. The pellet (P1) was recovered and saved at 4°C. Since some precipitation was noticed after 3 d, the supernatant was centrifuged again as before for 10 min to remove debris. Supernatant (S1) next was transferred to 12 ultracentrifuge tubes (about 26 ml/tube) (Beckman Cat #355618) and centrifuged for 4 h, 4°C, at 37,000 rpm (∼124,500×g) (Beckman Type 50. 2 Ti rotor, Beckman Optima L-70K Ultracentrifuge). After 4 h the supernatant (S2) was removed and saved. The pellet (P2) was resuspended in 10 mM phosphate buffer containing 4% Brij 58 (Aldrich Cat #388831) and 0. 4% Sodium deoxycholate (Sigma-Aldrich D6750): about 1 ml of buffer was used per tube. It was necessary to insert a spatula to help pellet into solution and this was followed by vortexing the suspension. The content from each tube was transferred to clean 50 ml centrifuge tubes. The process was repeated twice but only buffer phosphate was added the final time. Because the final solution was very thick, buffer was added to increase the final volume to ∼30 ml and this was mixed by inversion. The tube was centrifuged for 15 min at ∼10°C, 800×g (Beckman Coulter Allegra 25R), to remove debris. The pellet (P3) was saved at 4°C. (the pellet saved as a backup). The supernatant (S2) was transferred into two clean 50 ml tubes and 13. 2 g CsCl (Amresco Cat #0415) were added to each tube. To ensure the right CsCl concentration, 13. 2 g CsCl were added to ∼10 g sample; however the final volume was adjusted to 24 ml with buffer and gently mixed (up/down). The second tube was set by adding CsCl to the remaining sample. This final preparation was transferred to two ultracentrifuge tubes ∼25 ml (Beckman Cat #355618) and centrifuged at 37,000 rpm (∼124,500×g), 18°C for 24 h. After 24 h centrifugation, the tubes were removed carefully from the rotor and the whitish virus band collected by insertion of a needle attached to a syringe. Two more fractions were recovered for analyses: (1) the “liquid” part left after removing the virus band and (2) the “pellet” (P4) attached to the bottom of tube: Each fraction was transferred to dialysis tubes (Thomas Scientific Cat #3787-F42) and dialyzed overnight against nanopure filtered water followed by 3–4 additional changes in water the following day. After dialysis, content from the tubes was collected in 15 ml clean tubes and the volumes were measured. A subsample of 20 µl from each fraction was tested for virus presence. Adult bees were transferred to 1. 5 ml centrifuge tubes. Tri Reagent (Sigma Cat #T9424), was added and individual bees were homogenized in 0. 5 ml Tri reagent using disposable pestles and glass beads. Homogenates were frozen at −20°C if needed. Samples then were centrifuged 10 min at 12,000×g, at 4°C. The clear supernatant was transferred to a new tube and left at least 5 min at room temperature (RT). Next, 0. 2 ml chloroform was added and samples were shaken vigorously. This was followed by a10–15 min incubation at RT. Tubes were centrifuged 15 min at 12,000×g at 4°C. The colorless upper aqueous phase was transferred to a new tube and 0. 5 ml isopropanol was added. After mixing, samples were allowed to stand for 10 min then spun 10 min at 12,000×g at 4°C. The supernatant was removed and the pellet containing the RNA was washed with 1 ml 75% ethanol. After 5 min centrifugation at 7,500×g at 4°C, the RNA was allowed to dry (5–10 min) and reconstitute in ∼30 µl Nuclease free water (Qiagen). RNA concentrations were measured in a Nanodrop, ND-1000 Spectrophotometer. Samples were diluted in Nuclease free water. The field demonstration in FL was designed in a manner that permitted us to follow IAPV-infested bee colonies (some given Remebee-I and others not) for six weeks. One hundred standard colonies of honey bees were split into 5 groups with 20 colonies per group. Four groups were located within 100 m of one another (non-isolated) while a 5th group was isolated from the remaining four by at least 3. 2 km to measure any environmental effects due to location. Treatment allocations (20 colonies per treatment) were as follows: Treatment 1 – no treatment – non isolated Treatment 2 – Remebee-I only – non isolated Treatment 3 – Remebee-I+IAPV – non isolated Treatment 4 – IAPV only (fed in sugar water solution) – non isolated Treatment 5 – no treatment– isolated Colonies were equalized according to standard protocols prior to the beginning of the study (frames of bees/brood moved between colonies until populations leveled) and were managed optimally for honey production. Data collected at the beginning, middle, and end of the study included: frames of adult bees, cm2 brood, the presence of other bee maladies (nosema, varroa, and tracheal mites), bee activity, honey production and IAPV presence/absence and titer. The study lasted 6 weeks from the date of colony inoculation with IAPV and was replicated in PA with the following modifications: Only Treatment groups 1,3 and 4 were established and the trial lasted 12 weeks after inoculation to enable the bees to take advantage of a honeyflow (Tables 2 and 3). In PA the virus was introduced twice into the colonies within a three day period (instead of once as in FL), and the total amount of virus introduced was thus much higher than the FL trial. We calculated all test/sampling dates below from the date of the last treatment with Remebee-I. Controls 1,2, and 5, accounted for ‘within treatments’, a Remebee-I alone treatment to evaluate any potential detrimental effects to bees, and a distant control to measure environmental effects in the absence of IAPV. Colonies were equalized at the beginning of the studies, starting and ending colony strength parameters were compared using ANOVA recognizing treatment as the main effect (PROC GLM). Honey gains in treated colonies in FL and PA were compared identically. However, to compare colony size measures in the PA trial and for levels of nosema and varroa mites, a Before-After Control-Impact (BACI) design [33], [34] was used. A BACI design us a way of comparing data that are measured before treatment with data obtained after treatment. In general, it can be described as a repeated measures analysis of variance (ANOVA) which is performed using colonies as replicates and the covariance structure that best suits the data (PROC MIXED, SAS Institute). Each variable is measured at the start of the experiment to show existing conditions before treatment and then after a treatment. The analysis then looks at whether the change in variable measures was different between treatment groups. A repeated measures analysis of variance [34] was performed using colonies as replicates and an unstructured covariance structure was performed using SAS statistical software (PROC MIXED) [35]. Israeli Acute Paralysis Virus (IAPV) genome – NC_009025 (RefSec); Acute Bee Paralysis Virus (ABPV) genome – NC_002548 (RefSec); Kashmir Bee Virus (KBV) genome – NC_004807 (RefSec); Black Queen Cell Virus (BQCV), genome-NC_003784 (RefSec); Deformed Wing Virus (DWV) genome NC_004830 (RefSec); RNA dependent RNA Polymerase protein (C. elegans) – NP_492131 (RefSec); SID-1 protein (A. mellifera) – XP_395167 (RefSec); GFP nucleotide sequence – U87625 (GenBank).
High rates of honey bee mortality continue to threaten food security and apicultural industries worldwide. At least some of these losses are likely the result of viral infections. Application of RNAi technologies in the treatment and management of disease promises new solutions to disease problems through the naturally occurring biological processes of living organisms. This study applied a novel dsRNA product developed specifically with the aim of improving honey bee health. The results demonstrate the successful application of RNAi strategies to improve disease tolerance. Honey bees were fed a dsRNA product, Remebee-I, in the presence of the Israeli Acute Paralysis Virus. Treatment resulted in larger colony populations and thus increased honey production. We show that IAPV specific homologous dsRNA successfully curbed the negative effects of IAPV infection in 160 honey bee hives in two discrete climates, seasons and geographical locations (Florida and Pennsylvania). We provide the first successful demonstration of the use of RNAi as a preventative treatment for an insect disease on such a large scale.
biotechnology
molecular biology/post-translational regulation of gene expression
infectious diseases
microbiology/immunity to infections
pathology
virology/new therapies, including antivirals and immunotherapy
virology
immunology
microbiology
pathology/molecular pathology
molecular biology
genetics and genomics
cell biology/gene expression
Large-Scale Field Application of RNAi Technology Reducing Israeli Acute Paralysis Virus Disease in Honey Bees (Apis mellifera, Hymenoptera: Apidae)