RNAi technologies are more environmentally friendly, as the technology provides greater specificity in pest targeting, while reducing the potential negative effects on ecosystems and leaving beneficial insects and other organisms unharmed in crop ecosystems. Consequently, the increase in native fauna improves the efficacy of biological control agents against pests and pathogens. A growing understanding of the ubiquitous nature of RNAi, along with evidence for efficient, non-transgenic, topical applications has already begun to garner support among organic and industry producers. Designing solutions to agricultural problems based upon the same mechanisms used in nature provides newer, safer solutions to pests and pathogens for all agricultural industries.
A phenomenon initially reported in plants  called the attention of the scientific community, leading to the discovery of a sophisticated mechanism of gene regulation and protection against invasive nucleic acids [2–4]. Furthermore, the mechanism described in plants, referred to as post-transcriptional gene silencing (PTGS), or virus-induced gene silencing (VIGS), had been described in the 1990s  and was often referred to as pathogen-derived resistance .
RNAi is a natural process of gene regulation and antiviral defense system of eukaryotic cells. RNAi is a mechanism that functions as a “gene silencer” by targeting specific RNA sequences. RNAi results in degradation, and in some situations, translation inhibition, resulting in a reduction or complete elimination of the expression of a targeted RNA . RNAi is also linked to suppressing gene expression at transcriptional level by directing epigenetic alterations on chromatin .
The basic RNAi process consists of the trigger molecule, a long endogenous or exogenous double-strand RNA (dsRNA) molecule that is expressed in, or introduced into, the cell, which is processed by Dicer, a ribonuclease III (RNase III) enzyme into small RNA duplexes of 21–23 nucleotides in length. These duplexes are separated with one strand (the guide strand), producing a protein complex known as the RNA-induced silencing complex (RISC). The RISC complex uses the specific sequence of the guide strand to determine potential target messenger RNAs (mRNA). Once bound to the mRNA, the guide strand directs a RISC-bound endonuclease [called “slicer”, an Argonaute (AGO) protein] to cleave the mRNA which has homology to the guide strand . Thus, the RISC complex can target the messenger RNA (mRNA), an invasive virus RNA, or a transposable element transcript (Please note that RNAi cannot eliminate transposons itself, but its transcripts.) . These components appear to be cosmopolitan in their distribution across the RNAi spectrum of the eukaryotic phyla. This implies that a common ancestor had a functional RNAi pathway , estimated to have occurred over a billion years or more ago .
The generation of virus-resistant transgenic plants, by expressing fragments of viral genomes, was the first demonstration of the beneficial use of the RNAi [13,14]. However, with the demonstration that ingestion of dsRNAs can robustly silence genes in
In this chapter, we outlined some aspects on the development of RNAi-based strategies to control insects, presenting some considerations and research steps that are important to be addressed.
2. RNAi-based products for agricultural management of insect pest
Advances in genomics and initiatives to sequence genomes of agriculturally important organisms created big breakthroughs within the entomology fields of study, including taxonomy, insect physiology, toxicology, immunology, pest management, and microbe–host interactions. This new paradigm affected how research could be conducted, to make discoveries and increase the speed by which these could be accomplished. With these breakthroughs, entomologists, pathologists, and biologists are rapidly advancing toward better, safer, and more specific pest and pathogen management. Development of gene-based methods was dependent upon having the knowledge to understand how the cells in a living organism respond to the threats from viral pathogens, how they regulate their own gene expression, and how they maintain these natural complex systems throughout their lives.
The initial genomes which were sequenced and annotated to elucidate these very complicated interactions from: the nematode –
The number of arthropod species which have had successful RNAi reports continues to increase, and this trend will carry agriculture into the future, covering five Classes, in four Subphyla of the Arthropoda phylum, which includes eight insect orders and over 30 insect species [16–17].
3. Pitfalls and solutions – Relevant considerations for development of RNAi in insects
Though the use of RNAi strategies to control a desired insect pest seems to be very straightforward; however, some issues should be taken into considerations: (1) for oral treatments the dsRNA must survive ingestion, then be absorbed by the epithelial cells, and depending upon the target translocated through the hemolymph to reach other tissues. In insects, the dsRNA is mainly introduced through feeding, but dermal application has already been reported to possibly bypass gut issues [18–21]; (2) once inside the body, the dsRNA must enter into the cell to activate the RNAi mechanism. In insects, cell uptake of dsRNA varies widely between species, because there are different mechanisms of systemic absorption and translocation of dsRNAs within and between cells, respectively, leading to differences in response and influencing the efficiency in silencing the target gene [17,22]; (3) once the RNAi mechanisms are triggered in a group of cells, it has been demonstrated that a systemic spread of silencing may also occur, so that other tissues / cells are also affected, which may increase the RNA effect. Successful studies have shown that dsRNA can circulate in the hemolymph, and cause a suppression of genes in tissues distant from initial entry sites in the insect gut, affecting cuticle formation, the nervous system, or ovaries[23–26]. However, in insects results can be highly variable and research efforts continue to elucidate the effects of systemic signaling.
In theory, any cellular mRNA can be inactivated in a precise and controlled manner. With this in mind, the use of the RNAi mechanism to manage an insect pest relies on the capacity to design the dsRNA. The sequence of the dsRNA provides specificity and the researcher must determine the active concentration needed to obtain the desired RNAi outcome; thus, proper design and evaluation of the dsRNA becomes critical.
Identification of vitally important (i.e., with high mortality) target genes of a particular insect is a crucial step toward development of RNAi-based control strategy. Thanks to world science development and increasing efforts of the research community, the identification of an essential target can be achieved by an extended literature search and analyses of available DNA/RNA sequence databases . Once identified, “candidate” target sequences are used to design potent “RNAi causing structures”. Then, the dsRNAs must be experimentally validated for functionality, specificity, and stability, toward the specific RNAi target of interest. Furthermore, the development of an efficient delivery system for “RNAi causing structures” is another key step. There are several methods available that include, but are not limited to: microinjection , soaking (for mosquito larvae, and nematodes) [29,30], and feeding (chewing and piercing-sucking insects) [17,17,27,31,32].
One approach to identify potential target genes which will function under field conditions is to perform bioassays that closely mimic the conditions in the field, for example, using a bioassay that mimics the feeding of a hemipteran insect acquiring the dsRNA during the natural feeding process performed on the crop plant. One problem with delivering dsRNA through feeding is that, depending on the bioassay, it may be difficult to measure the dose of dsRNA ingested, from the dose absorbed by the gut cells and the target cells .
Oral delivery of dsRNA through feeding can be performed by using artificial diet, detached plant parts (leaves, buds, roots), or intact plants [17,24,34,35]. Delivering dsRNA through the diet provides an easy procedure to screen large numbers of dsRNAs in insect larvae and adults [23,24,34,37]. In addition, it allows addressing different issues, such as effective length of dsRNA, determine regions of the gene to be target which may provide better suppression, and to determine the effective lethal concentration (LC50) [23,38].
Although oral feeding provides a more natural screening system, it is important to take into consideration that for some insect species from across all taxonomic orders, they may not provide an effective RNAi response when conducting oral feeding bioassays regardless of the dosage of dsRNA, as the dsRNA may not enter, or be detected in the insect’s body . In contrast, in these same insects when dsRNA was injected directly into the insect’s body, a potent RNAi response was observed [23,39,40]. Indeed, lack of positive results using feeding bioassays does not necessarily indicate that the insect is insensitive to RNAi, but in a majority of the situations, insects have nucleases in the saliva, in the midgut, or even in the hemolymph that degrades dsRNA before it can be absorbed by the cells [40–42].
Wynant et al.  discussed the interactions of enzymes and RNAi-causing dsRNAs in the alimentary tract and hemolymph of insects and other arthropods using oral delivery. The elucidation of the roles of microbes and host enzymes on RNAi efficacy across arthropod species continues to be a challenging field of research .
Where information about absorption of dsRNA or presence of nucleases is not available for a particular insect species, the use of reporter dsRNAs molecules are useful to clarify possible issues. It is essential that the dsRNAs sequence should not match with any insect’s transcript sequence. The dsRNA “movement” can be monitored (detected), or quantified in the insect’s body by RT-qPCR, showing that the insect has acquired the potent, fully functional, systemically spreading dsRNA during feeding (plant, diet, drop of water, etc.) (Figure 1).
When conducting a RT-PCR detection of reporter dsRNA after insect feeding, it is important to sample a tissue other than the gut such as the hemolymph, fat body, or ovary. Careful collection of tissues which are not in direct contact with the gut provides evidence that the dsRNA was truly absorbed by the cells, and you are not just detecting dsRNA just in the digestive tract.
With small insects, as the Asian Citrus Psyllid (ACP)
The reporter dsRNA is designed so that the sequence does not match with any known mRNA transcript in your insect. This is to avoid off target of other transcripts in the insect. Some commonly used dsRNAs which are used as negative controls in RNAi experiments are: green fluorescent protein (GFP), β-glucuronidase (GUS), and enhanced yellow fluorescent protein (EYFP) .
When designing RNAi experiments, important questions arise regarding the design of dsRNA, including: the length of the molecule and the region targeted within the mRNA. The minimal required length to achieve an RNAi effect will vary depending on insect species . For example, in
There is no consensus on the mRNA region that the dsRNA should match to (e.g., 5′ or 3′). For example, in the pea aphid,
In the context of field applications of RNAi for insect management, dsRNAs can be designed to be highly specific to both the target gene and the insect species. If desired, the RNAs can be designed to have a broader spectrum to affect several pest species. For example, RNAi strategies can be designed to remove one aphid species from a cropping system, or be designed to remove multiple aphid species from that same ecosystem [24,38].
4. Bioassays for dsRNA screening
For RNAi research attempting the development of a viable pest management product. It is of utmost importance to identify the best delivery mechanisms (i.e., topical sprays, baits, or transgenic plants) as early as possible; this will expedite the entire process and can cut years off of the development and commercialization timeline.
The example outlined below highlights RNAi bioassays directed toward two citrus insect pests, each one with different feeding behaviors: piercing-sucking plant-feeding (the Asian citrus psyllid
4.1. Bioassays for piercing-sucking insects
It is notable that liquid feeding bioassays (dsRNAs mixed in a liquid diet or a sucrose solution) frequently result in high mortality levels in the controls, and increased degradation of dsRNA in the solution due to bacterial or fungal contaminations [42,57]. In addition, these bioassays require significantly high dsRNA concentrations to achieve insect mortality. Concentrations up to 1µg/µL [58–60] cannot be reproduced inside plant vascular tissues.
Hemipteran pests in citrus (psyllids, leafhoppers, aphids, whiteflies) have piercing-sucking mouthparts that are inserted into the plant vascular system to feed. The development of an RNAi control strategy against these insects relies on effective delivery of the dsRNA through the vascular tissues.
Demonstration of the first dsRNA delivery into full-sized citrus trees and grapevines, without a delivery vector, expression vector, or transformation event was performed in 2008 . These results showed that two hemipteran insects, the xylem-feeding leafhopper (
Use of cut plant feeding bioassays for hemipteran pests enables the screening of a large number of dsRNAs molecules at a reduced cost of materials and time. The bioassay, can use leaf disks, whole leaf, new growth leaves and stem, or rooted cuttings, to absorb and deliver dsRNAs. In citrus, the “flush”, which are new growth foliar shoots, are collected from potted citrus seedlings grown in a glasshouse (USDA-ARS, Fort Pierce, FL). The leaves and stem material are about 7–8 cm long. The plant material is washed in 0.2% bleach water, for 10 min. Then the base of each stem is cut at a 45 degree angle while submerged in filtered water. The material is then placed into a 1.5 mL tube containing 0.5 mL water (Figure 2B and C). The dsRNA solution, 300 µL, is added to the water, the tube top is wrapped with plastic or Parafilm™ and placed under artificial lighting to stimulate absorption of dsRNA solution. The next day the tube is filled with water using a 26 gauge syringe needle and syringed filtered (0.45 µ). The treated cuttings are then placed into a cage and adult insects provided feeding access for 10 days (Figure 2D). The plant material can remain viable for up to 40 days on average. While most bioassays may terminate after eight to 10 days of observations for mortality, having a longer feeding access time enables observations on insect oviposition, egg viability, or nymph development.
Each dsRNA molecules has an optimal concentration. So each dsRNA molecule is evaluated across a range of total concentrations (i.e., 5, 20, 50, 100 nanograms/ tissue). The bioassay permits screening for synergistic effects of multiple dsRNAs and to screen a single dsRNA against multiple insect species. For example, the assay using citrus flush permits screening of dsRNAs designed against psyllids for off target effects in the citrus aphid (
4.2. Bioassays for chewing insects
For insects which are foliage feeders, the delivery of dsRNA can be achieved as a foliar topical spray. In this scenario, the dsRNAs are evaluated similarly as topical insecticides. The dsRNA solution is sprayed on leaves, and then fed to the insects. An example of the effectiveness of this approach was reported by Bolognesi  working with the coleopteran
Similar results were obtained while developing an RNAi strategy against the Diaprepes root weevil (DRW),
A test spray using only water established the volume needed to provide full coverage of leaf bouquets without excessive run off (Figure 3B). After the leaves have dried, they are caged with adult insects (Figure 3C). Freshly treated citrus leaf bouquets replaced previous bouquets every five to seven days for a 5-week period. The total amount of dsRNA to be sprayed over the leaf bouquet was determined by evaluating a range of concentrations in a pretest experiment for efficacy. The effects from RNAi in insects usually start to appear within 4 to 5 days post-ingestion, which suggests there may be a dose response . Since foliage feeding insects tend to eat a lot of leaf material each day, a low-dose spray may be able to deliver a significant amount of dsRNA.
5. Final considerations on RNAi applied to agriculture
Efficient delivery and increased stability of dsRNA need to be developed if non-transgenic, topically delivered, RNAi strategies are to be established. Increased stability and superior delivery into some insects can be achieved using nanoparticle-mediated RNAi [63–65], traditional crop improvement strategies, in which plants express hairpin dsRNAs, will continue to be a mainstay of agricultural approaches [63,65,66].
Transgenic plants have successfully used RNAi strategies to produce crops with improved virus resistance, increased nutrition and fiber content ; biotechnology companies are trying to move towards a faster, more natural process of topically applied RNAi. dsRNA molecules are part of naturally occurring processes in all living organisms. They exist in our foods, and our bodies . The short persistence time of dsRNA in the environment is demonstrated by the fact that analyses of soils and plant debris, treated with dsRNA have consistently shown rapid breakdown of dsRNAs within 2–3 days , also means less concerns about unintended contamination of water supplies, soils, or adverse air quality effects. Furthermore, since all living things have evolved to break down dsRNA and use the nucleic acids as cellular nutrients, this technology will be safer than conventional chemistries for those who apply RNAi products, or eat the produce [66–69].
RNAi technologies have greater specificity in pest targeting, which reduces negative impacts on crop ecosystems by leaving more insects and other organisms unharmed in the field. The increased fauna consequently improves the efficacy of pollination, and biological control agents that help suppress a broad range of pests. The increased understanding of the ubiquitous nature of RNAi, along with evidence of efficient topical application, has already begun to garner support for this technology among members of the organic grower’s communities, which desperately need a truly natural, innovative breakthrough, to manage many of the pests and pathogens which plague the organic industries.
5.1. Cost-effective methods for the mass production and formulation of dsRNA
Cost-efficient methods for mass production of vast amounts of dsRNA are being developed, and include bacterial, plant, and synthetic production . While small amounts of dsRNA can be easily produced in the laboratory for research purposes, commercially available kits are not a viable, cost-effective method for the production of large quantities of dsRNA . The costs associated with the commercialization and implementation of RNAi products are decreasing rapidly. The costs of dsRNA production have dropped from $500,000 USD for 40 g in 2008 to less than $4,000 USD for 40 g today.
5.2. Other applications
Future applications of RNAi and other gene-based targeting biotechnologies will add value to existing beneficial insects (pollinators, predators, parasitoids). A real-world example is a study conducted over several years in which an RNAi product designed to reduce Israeli acute paralysis virus replication was fed to honey bees. The treated bees had significantly greater survival and produced significantly more honey . RNAi strategies have also reduced honey bee parasites, like
Biotechnology has demonstrated the safe production of plants which are more nutritious, less toxic, more resistant to drought, and more efficient for biofuel production [66, 67]. RNAi has already been successfully used to produce crops which are virus- and drought-resistant . However, plants expressing dsRNAs while stable and safe take years to develop and millions of dollars to commercialize . Development of topically applied RNAi, which is a non-transgenic approach improves crop traits and provides a major step forward for environmentally sound crop management [66, 67].
As more insects and mites develop chemical resistance to one or more insecticides, now estimated to be over 500 species with resistance to one or more products , it is imperative that new types of pest control are developed. The public would like the world to be filled with environmentally friendly technologies, safe for human and animal consumption, technologies which are safe for use around animals and beneficial insects, safe for each type of ecosystem, forest, field, crop, or backyard, a technology that will not endanger water or food quality, a more natural solution, with a natural approach toward problem-solving. So enters RNA interference!
RNA interference, or gene silencing, is a way to reduce specific mRNAs so that a particular protein is either not made or it is reduced. The RNAi mechanism is a natural one which occurs in the cells of humans, animals, insects, and plants, and appears to have evolved as a primary defense system against virus replication . Andrew Fire and Craig Mello, won the Nobel Prize in 2006  for explaining how the RNAi mechanism is triggered, when a cell encounters double-stranded RNA, and how this could be used to benefit humanity. Humanity’s greatest discoveries have come from observing the natural world; while RNAi will not solve every problem, it certainly can help improve plant health, reduce insect pests and pathogens [17, 66, 67, 80]. Some of the benefits from developing RNAi as topically applied products are: 1) The rapid degradation of the molecules ensures low environmental risks. All cells have the capacity to degrade dsRNA, and the salvage pathways to recycle these bases and nucleosides to form new nucleotides. Thus, cells constantly breaking down DNA and RNA into recycled nucleotides . 2) Topical RNAi applications do not insert genes, so do not produce proteins. RNAi reduces the expression of the targeted proteins, a modulation effect of the natural system. 3) RNAi can be designed and tested faster (in about 2–3 years) than producing transgenic crops, which can take 10 to 20 years and cost hundreds of millions of dollars . 4) Finally, RNAi strategies as topical sprays would, for the first time, be able to remove one or two closely related insect species, while leaving all the other insects unharmed . The ability to design RNAi as highly specific pest control will finally provide relief to biological control agents and beneficial insects [17, 44, 73], significantly improving integrated pest management programs.
The advantages and promises from RNAi technology sound amazing. However, serious efforts in outreach and education are needed to better inform the different stake holders including the general public, and agricultural industry, leaders as well as decision makers in the regulatory and political communities to help expedite the release and adoption of RNAi products and technology.
Mention of trade names or commercial products herein is solely for the purpose of providing specific information and does not imply recommendation or endorsement, to the exclusion of other similar products or services by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer.
We thank Dr. Xiomara Sinisterra-Hunter, AgTec, LLC, Plant Biotechnology Consultant, Port St. Lucie, FL, for critical reviews of the manuscript, and Maria Gonzalez, Biological science technician, USDA, Fort Pierce, FL, for technical assistance.
Napoli C, Lemieux C, Jorgensen, R. Introduction of a chimeric chalcone synthase gene into petunia results in reversible co-suppression of homologous genes in trans. Plant Cell 1990;2:279–89. DOI: http://dx.doi.org/10.1105/tpc.2.4.279.
Voinnet O, Pinto YM, Baulcombe DC. Suppression of gene silencing: a general strategy used by diverse DNA and RNA viruses of plants. Proc Nat Acad Sci USA 1999;96:14147–52. DOI: 10.1073/pnas.96.24.14147.
Voinnet O. Induction and suppression of RNA silencing: insights from viral infections. Nat Rev Genetics 2005;6:206–20. DOI: 10.1038/nrg1555.
Sidahmed AM, Wilkie B. Endogenous antiviral mechanisms of RNA interference: a comparative biology perspective. Method Molecul Biol 2010;623:3–19. DOI:10.1007/978-1-60761-588-0_1.
Baulcombe DC. RNA as a target and an initiator of post-transcriptional gene silencing in trangenic plants .Plant Molecul Biol 1996;32:79–88. DOI 10.1007/BF00039378.
Molnar A, Melnyk C, Baulcombe DC. Silencing signals in plants: a long journey for small RNAs. Genome Biol 2011;12:215. DOI 10.1186/gb-2010-11-12-219.
Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 1998;391:806–11. DOI: 10.1038/35888.
Castel SE, Martienssen RA. RNA interference in the nucleus: roles for small RNAs in transcription, epigenetics and beyond. Nat Rev Genetics 2013;14:100–12. DOI:10.1038/nrg3355.
Tomari Y, Zamore PD. Perspective: machines for RNAi Genes Develop 2005;19:517–29. DOI: 10.1101/gad.1284105.
Buchon N, Vaury C. RNAi: a defensive RNA-silencing against viruses and transposable elements. Heredity 2006;96:195–202. DOI:10.1038/sj.hdy.6800789.
Cerutti H, Casas-Mollano, J.A. On the origin and functions of RNA-mediated silencing: from protists to man. Current Genetics 2006;50:81–99. DOI: 10.1007/s00294-006-0078-x.
Roger AJ, Hug LA. The origin and diversification of eukaryotes: problems with molecular phylogenetics and molecular clock estimation. Philosoph Transac Royal Soc B 2006;361:1039–1054. DOI: 10.1098/rstb.2006.1845.
Hamilton AJ, Baulcombe DC. A species of small antisense RNA in posttranscriptional gene silencing in plants. Science 1999;286:950–952. DOI: 10.1126/science.286.5441.950.
Bonfim K, Faria JC, Nogueira EO, Mendes EA, Aragão FJL. RNAi-mediated resistance to bean golden mosaic virus in genetically engineered common bean ( Phaseolus vugaris). Molecul Plant-Microbe Interact 2007;20:717–26. DOI: http://dx.doi.org/10.1094/MPMI-20-6-0717.
i5K Consortium. The i5K initiative: advancing arthropod genomics for knowledge, human health, agriculture, and the environment. J Heredity 2013;104:595–600. DOI: 10.1093/jhered/est050.
Zhang H, Li H, Miao X. Feasibility, limitation and possible solutions of RNAi-based technology for insect pest control. Insect Sci 2013:20:15–30. DOI 10.1111/j.1744-7917.2012.01513.x.
Zotti M, Smagghe G. RNAi technology for insect management and protection of beneficial insects from diseases: lessons, challenges and risk assessments. Neotropic Entomol 2015;44. DOI 10.1007/s13744-015-0291-8.
Pridgeon JW, Zhao L, Becnel JJ, Strickman DA, Clark GG, Linthicum KJ. Topically applied AaeIAP1double-stranded RNA kills female adults of Aedes aegypti. J Med Entomol 2008;45:414–20. DOI:http://dx.doi.org/10.1093/jmedent/45.3.414.
Wang Y, Zhang H, Li H, Miao X. Second-generation sequencing supply an effective way to screen RNAi targets in large scale for potential application in pest insect control. PLoS One 2011;6:e18644. DOI: 10.1371/journal.pone.0018644.
El-Shesheny I, Hajeri S, El-Hawary I, Gowda S, Killiny N. Silencing abnormal wing disc gene of the Asian citrus psyllid, Diaphorina citridisrupts adult wing development and increases nymph mortality. PLoS One 2013;8:e65392. DOI:10.1371/journal.pone.0065392.
Killiny N, Hajeri S, Tiwari S, Gowda S, Stelinski LL. Double-stranded RNA uptake through topical application, mediates silencing of five CYP4genes and suppresses insecticide resistance in Diaphorina citri. PLoS One 2014;9:e110536. DOI:10.1371/journal.pone.0110536.
Terenius O, Papanicolaou A, Garbutt JS, Eleftherianos I, Huvenne H, et al. RNA interference in Lepidoptera: an overview of successful and unsuccessful studies and implications for experimental design. J Insect Physiol 2011;57:231–245. DOI: 10.1016/j.jinsphys.2010.11.006.
Araujo RN, Santos A, Pinto FS, Gontijo NF, Lehane MJ, Pereira MH. RNA interference of the salivary gland nitrophorin 2 in the triatomine bug Rhodnius prolixus(Hemiptera: Reduviidae) by dsRNA ingestion or injection. Insect Biochem Molecul Biol 2006;36:683–93. ISSN 0965-1748; 0965-1748. DOI:10.1016/j.ibmb.2006.05.012
Whyard S, Singh AD, Wong S. Ingested double-stranded RNAs can act as species-specific insecticides. Insect Biochem Molecul Biol 2009;39:824–32. DOI: 10.1016/j.ibmb.2009.09.007.
Paim RM, Pereira MH, Di Ponzio R, Rodrigues JO, Guarneri AA, Gontijo NF, Araujo RN. Validation of reference genes for expression analysis in the salivary gland and the intestine of Rhodnius prolixus(Hemiptera, Reduviidae) under different experimental conditions by quantitative real-time PCR. BMC Res Notes 2012;5:128. DOI:10.1186/1756-0500-5-128.
Paim RM, Pereira MH, Araujo RN, Gontijo NF, Guarneri AA. The interaction between Trypanosoma rangeliand the nitrophorins in the salivary glands of the triatomine Rhodnius prolixus(Hemiptera; Reduviidae). Insect Biochem Molecul Biol 2013;43:229–36. DOI:10.1016/j.ibmb.2012.12.011.
Mao J, Zeng F. Feeding-based RNA interference of a gap gene is lethal to the pea aphid, Acyrthosiphon pisum. PLoS One 2012;7:e48718. DOI:10.1371/journal.pone.0048718.
Liu S, Ding Z, Zhang C, Yang B, Liu, Z. Gene knockdown by introthoracic injection of double-stranded RNA in the brown planthopper, Nilaparvata lugens. Insect Biochem Molecul Biol 2010;40:666–71. DOI: 10.1016/j.ibmb.2010.06.007.
Singh AD, Wong S, Ryan CP, Whyard S. Oral delivery of double-stranded RNA in larvae of the yellow fever mosquito, Aedes aegypti: implications for pest mosquito control .J Insect Sci 2013;13:69. Available from: http://www.insectscience.org/13.69.
Dutta TK, Banakar P, Rao U. The status of RNAi-based transgenic research in plant nematology. Front Microbiol 2015;5:1–7. DOI: 10.3389/fmicb.2014.00760.
Huvenne H, Smagghe G. Mechanisms of dsRNA uptake in insects and potential of RNAi for pest control: a review. J Insect Physiol 2010;56:227–35. DOI:10.1016/j.jinsphys.2009.10.004.
Pitino M, Coleman AD, Maffei ME, Ridout CJ, Hogenhout SA. Silencing of aphid genes by dsRNA feeding from plants. PLoS ONE 2011;6:e25709. DOI: 10.1371/journal.pone.0025709.
Li H, Guan R, Guo H, Miao X. New insights into an RNAi approach for plant defence against piercing-sucking and stem-borer insect pests. Plant Cell Environ 2015:38:1–9. DOI: 10.1111/pce.12546.
Baum JA, Roberts, J.K. Chapter Five- progress towards RNAi-mediated insect pest management. Adv Insect Physiol 2014;47:249–95. DOI: 10.1016/B978-0-12-800197-4.00005-1.
Zhu F, Xu JJ, Palli R, Ferguson J, Palli S.R. Ingested RNA: interference for managing the populations of the Colorado potato beetle, Leptinotarsa decemlineata. Pest Manage Sci 2011;67:175–82. DOI: 10.1002/ps.2048.
Turner CT, Davy MW, MacDiarmid RM, Plummer KM, Birch NP, Newcomb RD. RNA interference in the light brown apple moth, Epiphyas postvittana(Walker) induced by double-stranded RNA feeding. Insect Molecul Biol 2006;15:383–91. DOI:10.1111/j.1365-2583.2006.00656.x
Aronstein K, Oppert B, Lorenzen MD. RNAi in Agriculturally-Important Arthropods, RNA Processing, Prof. Paula Grabowski (Ed.), 2011, ISBN: 978-953-307-557-0, InTech, [Available from: http://www.intechopen.com/books/rna-processing/rnai-in-agriculturally-important-arthropods. Accessed 2015/08/25.
Bachman PM, Bolognesi R, Moar WJ, Mueller GM, Paradise MS, et al. Characterization of the spectrum of insecticidal activity of a double-stranded RNA with targeted activity against western corn rootworm ( Diabrotica virgifera virgiferaLeConte). Transgenic Res 2013;22:1207–22. DOI: 10.1007/s11248-013-9716-5.
Wynant N, Verlinden H, Breugelmans B, Simonet G, Vanden Broeck J. Tissue-dependence and sensitivity of the systemic RNA interference response in the desert locust, Schistocerca gregaria. Insect Biochem Molecul Biol 2012;42:911–7. DOI:10.1016/j.ibmb.2012.09.004.
Wynant N, Santos D, Verdonck R, Spit J, Van Wielendaele P, Vanden Broeck, J. Identification, functional characterization and phylogenetic analysis of double stranded RNA degrading enzymes present in the gut of the desert locust, Schistocerca gregaria. Insect Biochem Molecul Biol 2014;46:1–8.DOI: http://dx.DOI.org/10.1016/j.ibmb.2013.12.008.
Garbutt JS, Bellés X, Richards EH, Reynolds SE. Persistence of double-stranded RNA in insect hemolymph as a potential determiner of RNA interference success: evidence from Manduca sextaand Blattella germanica. J. Insect Physiology 2012. DOI: http://dx.doi.org/10.1016/j.jinsphys.2012.05.013.
Christiaens O, Swevers L, Smagghe G. DsRNA degradation in the pea aphid ( Acyrthosiphon pisum) associated with lack of response in RNAi feeding and injection assay. Peptides 2014;53:307–14. DOI: 10.1016/j.peptides.2013.12.014.
Sharma P, Sharma S, Maurya RK, De TD, Thomas T, Lata S, Singh N, Pandey KC, Valecha N, Dixti R. Salivary glands harbor more diverse microbial communities than gut in Anopheles culicificacies. Parasites Vectors 2014;7:235. DOI: 10.1186/1756-3305-7-235.
Scott JG, Michel K, Bartholomay LC, Siegfried BD, Hunter WB, Smagghe G, Zhu KY, Douglas AE. Towards the elements of successful insect RNAi. J Insect Physiol 2013;59:1212–21. http://dx.doi.org/10.1016/j.jinsphys.2013.08.014.
Bolognesi R, Ramaseshadri P, Anderson J, Bachman P, Clinton W, Flannagan R, Ilagan O, Lawrence C, Levine S, Moar W, et al. Characterizing the mechanism of action of double-stranded RNA activity against western corn rootworm ( Diabrotica virgifera virgiferaLeConte). PLoS One 2012;7: e47534. DOI:10.1371/journal.pone.0047534.
Miller SC, Miyata K, Brown SJ, Tomoyasu Y. Dissecting systemic RNA interference in the red flour beetle Tribolium castaneum: parameters affecting the efficiency of RNAi. PLoS One 2012;7:e47431. DOI:10.1371/journal.pone.0047431.
Wuriyanghan H, Rosa C, Falk BW. Oral delivery of double-stranded RNAs and siRNAs induces RNAi effects in the potato/tomato psyllid, Bactericerca cockerelli. PLoS One 2011;6:e27736. DOI: 10.1371/journal.pone.0027736.
Mutti NS, Park Y, Reese JC, Reeck GR. RNAi knockdown of a salivary transcript leading to lethality in the pea aphid, Acyrthosiphon pisum. J Insect Sci 2006;6:1–7. DOI:10.1673/031.006.3801.
Kumar P, Pandit SS, Baldwin I.T. Tobacco rattle virus vector: a rapid and transient means of silencing Manduca sextagenes by plant mediated RNA interference. PLoS One 2012;7:e31347. DOI:10.1371/journal.pone.0031347.
Noh MY, Beeman RW, Arakane Y. RNAi-based functional genomics in Tribolium castaneumand possible applications for controlling insect pests. Entomol Res 2012;42:1–10. DOI: 10.1111/j.1748-5967.2011.00437.x.
Hamilton MA. Further experiments on the artificial feeding of Myzus persicae(Sulz.) Annal Appl Biol 1935;32:243–58. DOI:10.1111/j.1744-7348.1935.tb07160.x.
Li J, Wang W-P, Wang M-Q, Ma W-H, Hua H-X. Advances in the use of the RNA interference technique in Hemiptera. Insect Sci 2013;20:31–9. DOI 10.1111/j.1744-7917.2012.01550.x.
Christiaens O, Smagghe G. The challenge of RNAi-mediated control of hemipterans. Curr Opin Insect Sci 2014;6:15–21. DOI: http://dx.doi.org/10.1016/j.cois.2014.09.012.
Hall DG, Shatters RG, Carpenter JE, Shapiro JP. Research toward an artificial diet for adult Asian Citrus Psyllid. Annal Entomol Soc Am 2010;03: 611–17. DOI:http://dx.doi.org/10.1603/AN10004.
Hall DG, Richardson ML, Ammar E-D, Halbert SE. Asian citrus psyllid, Diaphorina citri, vector of citrus huanglongbing disease. Entomol Experiment Applic 2012;146:207–23. DOI: 10.1111/eea.12025.
Hunter WB, Glick E, Paldi N, Bextine BR. Advances in RNA interference: dsRNA treatment in trees and grapevines for insect pest suppression. Southwest Entomol 2012;37:85–7. DOI: http://dx.doi.org/10.3958/059.037.0110.
Upadhyay SK, Chandrashekar K, Thakur N, Verma PC, Borgio JF, Singh PK, Tuli R. RNA interference for the control of whiteflies ( Bemisia tabaci) by oral route. J Biosci 2011;36:153–61. DOI:10.1007/s12038-011-9009-1.
Borgio JF. RNAi mediated gene knockdown in sucking and chewing insect pests. J Biopesticides 2010;3:386–93. Available from: http://www.jbiopest.com/users/lw8/efiles/francis_borgio.pdf.
Katoch R, Sethi A, Thakur N, Murdock LL. RNAi for insect control: current perspective and future challenges. Appl Biochem Biotechnol 2013;171:847–73. DOI:10.1007/s12010-013-0399-4.
Tomizawa M, Noda H. High mortality caused by high dose of dsRNA in the green rice leafhopper Nephotettix cincticeps(Hemiptera: Cicadellidae). Appl Entomol Zoo 2013;48:553–59. DOI:10.1007/s13355-013-0211-5.
Zhou X, Wheeler MM, Oi FM, Scharf ME. RNA interference in the termite Reticulitermes flavipesthrough ingestion of double-stranded RNA. Insect Biochem Molecul Biol 2008;38:805–15. DOI:10.1016/j.ibmb.2008.05.005.
Zha W, Peng X, Chen R, Du B, Zhu L, He, G. Knockdown of midgut genes by dsRNA-transgenic plant-mediated RNA interference in the Hemipteran insect Nilaparvata lugens. PLoS One 2011;6(5):e20504. DOI:10.1371/journal.pone.0020504.
Yu N, Christiaens O, Lui J, Niu J, Cappelle K, Caccia S, Huvenne H, Smagghe G. Delivery of dsRNA for RNAi in insects: an overview and future directions. Insect Sci 2013;20:4–40. Doi 10.1111/j.1744-7917.2012.01534.x.
Zhang X, Zhang J, Zhu KY. Chitosan/double-stranded RNA nanoparticle-mediated RNA interference to silence chitin synthase genes through larval feeding in the African malaria mosquito ( Anopheles gambiae). Insect Molecul Biol 2010;19:683–93. DOI:10.1111/j.1365-2583.2010.01029.x
Palli SR. RNA interference in Colorado potato beetle: steps toward development of dsRNA as a commercial insecticide. Curr Opin Insect Sci 2014;3. DOI: 10.1016/j.cois.2014.09.011.
Petrick JS, Brower-Toland B, Jackson A, Kier LD. Safety assessment of food and feed from biotechnology-derived crops employing RNA-mediated gene regulation to achieve desired traits: a scientific review. Regul Toxicol Pharmacol 2013;66:167–76. DOI: http://dx.doi.org/10.1016/j.yrtph.2013.03.008.
Koch A, Kogel K-H. New wind in the sails: improving the agronomic value of crop plants through RNAi-mediated gene silencing. Plant Biotechnol J 2014;12:821–31. DOI:10.1111/pbi.12226.
Dubelman S, Fischer J, Zapata F, Huizinga K, Jiang CJ, et al. Environmental fate of double-stranded RNA in agricultural soils. PLoS ONE 2014;9. DOI:10.1371/journal.pone.0093155.
Ivashuta SI, Petrick JS, Heisel SE, Zhang Y, Guo L, Reyholds TL, Rice JF, Allen E, Roberts JK. Endogenous small RNAs in grain: Semi-quantification and sequence homology to human and animal genes. Food Chem Toxicol 2009;47:353–60. DOI: 10.1016/j.fct.2008.11.025.
AgroRNA, South Korea, [Internet]. Available from: http://www.agrorna.com/. Accessed: 2015/08/25.
Nguyen TA, Fruehauf JH. transkingdomRNA interference ( tkRNAi): a novel method to induce therapeutic gene silencing. T-Cell Protocols. Method Molecul Biol 2009;514:27–34. Available from: http://link.springer.com/protocol/10.1007%2F978-1-60327-527-9_3.
APSE, LLC, [Internet]. Available from: http://www.apsellc.com/. Accessed: 2015/08/25.
Hunter W, Ellis J, VanEngelsdorp D, Hayes J, Westervelt D, Glick E, Paldi N. Large-scale field application of RNAi technology reducing Israeli acute paralysis virus disease in honey bees ( Apis mellifera, Hymenoptera: Apidae). PLoS Pathogens 2010;6:e1001160. DOI:10.1371/journal.ppat.1001160.
Campbell EM, Budge GE, Bowman AS. Gene-knockdown in the honey bee mite Varroa destructorby a non-invasive approach: studies on a glutathione S-transferase. Parasites Vectors 2010;3:73. DOI:10.1186/1756-3305-3-73.
Garbian Y, Maori E, Kalev H, Shafir S, Sela I. Bidirectional transfer of RNAi between honey bee and Varroa destructor: Varroagene silencing reduces Varroapopulation. PLoS Pathogens 2012;8(12):e1003035. DOI: 10.1371/journal.ppat.1003035.
Paldi N, Glick E, Oliva M, Zilberberg Y, Aubin L, Pettis JS, Chen YP, Evans JD. 2010. Effective gene silencing of a microsporidian parasite associated with honey bee ( Apis mellifera) colony declines. Appl Environ Microbiol 2010;76:5960–4. DOI:10.1128/AEM.01067-10.
Elzen GW, Hardee DD. United States Department of Agriculture–Agricultural Research Service, Research on managing insect resistance to insecticides. Pest Management Sci 2003;59:770–6.
Blair CD, Olson KE. The role of RNA interference (RNAi) in Arbovirus-vector interactions. Viruses 2015;7:820–43. DOI: 10.3390/v7020820.
The Nobel Prize in Physiology or Medicine 2006. Andrew Z. Fire and Craig C. Mello "For their discovery of RNA interference - gene silencing by double-stranded RNA". Available from: http://www.nobelprize.org/nobel_prizes/medicine/laureates/2006/. [Accessed: 2015-08-25].
Whyard S. Insecticidal RNA, the long and short of it. Science 2015;347:950–91. DOI: 10.1126/science.aaa7722.
Lane AN, Fan TW-M. Survey and summary. Regulation of mammalian nucleotide metabolism and biosynthesis. Nucleic Acids Res 2015;43:1–20. DOI: 10.1093/nar/gkv047.