Source: KANSAS STATE UNIV submitted to NRP
BREAKING DOWN THE BARRIER: RATIONAL DESIGN OF EFFECTIVE RNAI FOR INSECT PEST MANAGEMENT
Sponsoring Institution
National Institute of Food and Agriculture
Project Status
ACTIVE
Funding Source
AFRI COMPETITIVE GRANT
Reporting Frequency
Annual
Accession No.
1029856
Grant No.
2023-67013-39166
Cumulative Award Amt.
$714,447.00
Proposal No.
2022-08801
Multistate No.
(N/A)
Project Start Date
Apr 1, 2023
Project End Date
Mar 31, 2026
Grant Year
2023
Program Code
[A1112]- Pests and Beneficial Species in Agricultural Production Systems
Recipient Organization
KANSAS STATE UNIV
(N/A)
MANHATTAN,KS 66506
Performing Department
(N/A)
Non Technical Summary
RNA interference (RNAi), a naturally occurring process in most eukaryotic organisms, has shown enormous potentials for species-specific management of insect pests. However, developing RNAi-based pest management strategies has been challenged by low or lack of RNAi efficiency of small interfering RNA (siRNA) delivered into insects. The overall goal of this project is to develop siRNA-based sprayable formulations and baits through rational design of effective siRNA constructs and development of new strategies to overcome the major challenges toward the widespread application of siRNA for insect pest management. We hypothesize that low or lack of RNAi efficiency of siRNA is caused by the application of ineffective synthetic siRNA, rapid degradation of siRNA by exoribonucleases (exoRNases), low cellular uptake of siRNA, and endosomal entrapment of siRNA in insects. To test these hypotheses, we plan to develop novel strategies for designing highly effective siRNA constructs based on the abundance of the siRNA antisense strands generated by insect species-specific Dicer-2. We will then use exoRNase inhibitors, exoRNase-resistant siRNA constructs, and siRNA mixtures to protect siRNA from exoRNase-mediated degradation. Finally, we will develop siRNA nanoparticle-based delivery strategies to enhance cellular uptake of siRNA, and test endosomal escape enhancing and endosome-disrupting agents to promote endosomal release of siRNA. This project will advance our knowledge on the mechanisms affecting RNAi efficiency of siRNA in insects and develop new strategies to overcome the major obstacles toward successful application of siRNA for insect pest management. This project addresses Program Area Priority (A1112): Pests and Beneficial Species in Agricultural Production Systems.
Animal Health Component
0%
Research Effort Categories
Basic
100%
Applied
0%
Developmental
0%
Classification

Knowledge Area (KA)Subject of Investigation (SOI)Field of Science (FOS)Percent
21131101130100%
Knowledge Area
211 - Insects, Mites, and Other Arthropods Affecting Plants;

Subject Of Investigation
3110 - Insects;

Field Of Science
1130 - Entomology and acarology;
Goals / Objectives
Arthropod pests alone destroy an estimated 18-20% of the crop production worldwide resulting in an economic loss of $470 billion US dollar annually. RNA interference (RNAi) has shown enormous potentials as a novel approach for species-specific management of insect pests. RNAi is environmentally friendly and can be designed to control specific pest species. Significant progresses have been made by developing transgenic crops expressing insect-specific double-stranded RNA (dsRNA). However, there has been great interest in developing non-transgenic RNAi approaches for insect pest management, including small interfering RNA (siRNA)-based sprayable formulations and baits.There are several distinct advantages of using siRNA for insect pest management: a) the production of siRNA can be fully scalable with a low cost; b) the structure of siRNA can be modified to improve RNAi efficiency; and c) siRNA bypasses the Dicer-mediated dicing step, which allows escaping potential RNAi resistance conferred by Dicers (e.g., mutations) in insects. However, the use of siRNA for insect pest management is currently limited by many factors. A long-standing challenge is that siRNA delivered into insects is often not effective to trigger RNAi responses, which becomes a roadblock toward successful applications of RNAi technology for insect pest management. The overall goal of this research is to develop siRNA-based sprayable formulations and baits through rational designs of effective siRNA constructs and development of novel strategies to overcome the obstacles toward widespread applications of siRNA for insect pest management. Our specific objectives of this project are to:1) Develop novel strategies for designing siRNA constructs for effective RNAi. Background: Dicer-2 cleaves dsRNA to generate species-specific siRNA populations with different abundances and RNAi efficiencies in different insect species. Supporting Objectives: a) produce insect species-specific recombinant Dicer-2, b) generate siRNA population from each dsRNA by each recombinant Dicer-2, c) sequence each siRNA population and map siRNAs to each dsRNA to identify most abundant antisense strands, d) design highly effective siRNA constructs, and e) determine RNAi efficiency of siRNA constructs in insects.2) Improve RNAi efficiency by protecting siRNA from degradation by RNases. Background: The degradation of dsRNA or siRNA by RNases (e.g., dsRNases) is a major factor causing low RNAi efficiency in insects. Supporting Objectives: a) co-deliver siRNA with naturally occurring RNase inhibitor; b) improve siRNA stability using structurally modified, dsRNase-resistant siRNA constructs; and c) enhance RNAi efficiency using RNase-resistant siRNA mixtures against both target and RNase genes simultaneously.3) Enhance RNAi efficiency by promoting cellular uptake and endosomal release of siRNA. Background: RNAi efficiency is directly related to the amount of siRNA internalized by cells and the amount of siRNA released from the endosomes. Supporting Objectives: a) facilitate cellular uptake of siRNA using polymer nanocarriers, b) prevent endosomal entrapment of siRNA using endosomolytic (endosome lysing) agents, and c) promote endosomal release of siRNA using endosome-disrupting nanoparticles.
Project Methods
Objective 1. Develop novel strategies for designing siRNA constructs for effective RNAiWe hypothesize that RNAi efficiency induced by siRNA is largely determined by the abundance of the antisense strand of a siRNA duplex because only the antisense siRNA strand loaded onto the RNA-induced silencing complex (RISC) is responsible for inducing the RNAi effect in the siRNA-mediated RNAi pathway. Therefore, the abundance of the siRNA antisense strands generated from a long dsRNA should be considered as an important criterion for designing effective siRNA constructs. To test this hypothesis, we will use the red flour beetle (RFB, Tribolium castaneum) representing coleopterans with robust RNAi response and the diamondback moth (DBM, Plutella xylostella) representing lepidopterans with poor RNAi efficiency in our study. We plan to use two insect essential genes including CHS2 encoding chitin synthase 2 and CHT10 encoding chitinase 10.First, we will produce recombinant Dicer-2 from each insect species using Escherichia coli BL21 (DE3) cells. Recombinant Dicer-2 in all fractions eluted from the column will be analyzed using SDS-polyacrylamide gel electrophoresis. Second, we will generate siRNA population from dsRNA by recombinant Dicer-2. Four dsRNAs including two (dsCHS2, dsCHT10) for each insect species will be synthesized using a dsRNA synthesis kit. Each dsRNA will be subjected to in vitro cleavage by species-specific recombinant Dicer-2. After each siRNA population generated by each recombinant Dicer-2 is sequenced, the sequences of each siRNA population will then be mapped to each dsRNA sequence and the dsRNA cleavage sites will be determined.Based on the sequences of the antisense strands mapped to each dsRNA, single-stranded RNA (ssRNA) oligos for both the antisense strand and its complementary sense strand will be synthesized. By annealing the sense and antisense oligos, we will produce 10 siRNA constructs for each target gene for each insect species. These siRNA constructs are expected to generate high abundant antisense strands against each target gene when delivered into insects, which is critical for high RNAi efficiency. We will then treat the fourth-instar larvae of RFB and the third-instar larvae of DBM by injecting each siRNA construct at each of three doses. For feeding-based bioassays using an artificial diet for DBM larvae, we will mix each siRNA construct at each of three concentrations to the artificial diet. For feeding-based bioassay using cabbage leaf disc and seedlings, we will treat the surface of the leaf discs and seedlings with each siRNA construct at each of three concentrations. Each treatment and control will be repeated four times. The suppression level of each target gene will be examined using reverse transcription quantitative PCR (RT-qPCR) and the effect of RNAi on insect survivorship or development will be examined at different time points. The data will be analyzed using ANOVA followed by Tukey's HSD multiple comparison test.Objective 2. Improve RNAi efficiency by protecting siRNA from degradation by exoRNasesWe hypothesize that rapid degradation of siRNA delivered into insects by exoRNases is a key mechanism causing low RNAi efficiency, and therefore, protecting siRNA from exoRNase-mediated degradation can significantly improve RNAi efficiency. To test this hypothesis, we will evaluate three strategies to enhance RNAi efficiency of siRNA by using a naturally occurring exoRNase inhibitor; structurally modified, RNase-resistant siRNA; and exoRNase-resistant siRNA mixtures against both target and RNase genes simultaneously. First, we will extract hemolymph and gut contents from DBM larvae, and incubate each siRNA construct in the presence or absence of adenosine 3', 5'-bisphosphate (PAP), a known inhibitor of exoRNases, at each of three concentrations. The remaining un-degraded siRNA after the incubations will be quantified using NanoPhotometer and visualized by gel electrophoresis. For in vivo study, we will compare RNAi efficiency of each of three siRNA constructs with and without PAP against each of the two target genes (CHS2, CHT10) by injection in RFB and DBM larvae and by feeding using artificial diet, leaf-discs, and cabbage seedlings in DBM larvae as described in Objective 1.Second, we will select three siRNA constructs showing high RNAi efficiency and structurally modify these siRNA constructs by introducing a dTdT overhang at each 3' end of a siRNA duplex. To accomplish this, ssRNA oligos for both the antisense strand and its complementary sense strand will be synthesized using the custom ssRNA synthesis service. By annealing the 3'dTdT sense and 3'dTdT antisense oligos, we will obtain structurally modified siRNA constructs. To examine if the structurally modified siRNA constructs can enhance their stability to the 3'-exoRNase-mediated degradation and improve their RNAi efficiency against their corresponding target genes, we will perform both the in vitro and in vivo studies as described above. Finally, we will combine the two strategies, 3'dTdT RNase-resistant siRNA and siRNA mixture simultaneously targeting a target and dsRNase genes, to test their effectiveness. The RNAi efficiency of the RNase-resistant siRNA mixtures will be examined against each of the two target genes (CHS2 and CHT10) using the injection-based RNAi approach for RFB and DBM larvae. Additional RNA bioassays will be conducted using the three feeding-based approaches, including artificial diet, cabbage leaf disc, and cabbage seedlings treated with the RNase-resistant siRNA mixtures for DBM larvae as described in Objective 1.Objective 3. Enhance RNAi efficiency by promoting cellular uptake and endosomal release of siRNAWe hypothesize that low or lack of RNAi efficiency of siRNA is further caused by limited cellular uptake of siRNA and the entrapment of siRNA in endosomes in insects. First, we will generate siRNA-based nanoparticles using chitosan and its derivatives (galactochitosan, glycol chitosan, methylglycol chitosan) as polymer nanocarriers. The formation of siRNA-based nanoparticles will be confirmed and the size of the nanoparticles will be measured using the ZetaView instrument. The RNAi efficiency of the siRNA-based nanoparticles will be examined against each of the two target genes by the injection-based approach for RFB and DBM larvae, and the three feeding-based approaches for DBM larvae. Second, we will choose at least three concentrations of chloroquine (an endosomal escape enhancing agent) and co-deliver each structurally modified, RNase-resistant siRNA construct against each of two target genes using both the in vitro and in vivo approaches as described in Objective 1.Finally, we will use endosome-disrupting nanoparticles to promote endosomal release of siRNA. Specifically, we will generate polyamidoamine (PAMAM) dendrimer-siRNA endosome-disrupting nanoparticles according to previously reported procedures. Both the siRNA binding ability of PAMAM dendrimers and the formation of nanoparticle will be evaluated based on the size shifting of siRNA using agarose gel electrophoresis. The size of the nanoparticles will be measured using the ZetaView instrument. Assays of endosomal release of siRNA will be carried out using midgut tissue culture in DBM larvae as endosomal entrapment of dsRNA has been reported in lepidopteran insect cells. To compare RNAi efficiency between the naked siRNA (control) and PAMAM dendrimer-siRNA nanoparticles (treatment), we will perform RNAi bioassays using the injection-based approach for RFB and DBM larvae, and the three feeding-based approaches for DBM larvae as described in Objective 1. Relative expression levels of a target gene in the treatment and control will be compared using RT-qPCR and various RNAi phenotypes including insect mortality and developmental disorders will also be examined and statistically analyzed as described in Objective 1.

Progress 04/01/24 to 03/31/25

Outputs
Target Audience:The primary target audiences of this study are researchers interested in RNA interference (RNAi), insect molecular biology, and insect pest management. Because RNAi has been shown to have its great potential for pest management, industries and pesticide regulatory agencies involved in pest management may also be interested in our findings. Changes/Problems:Our research progress has been delayed due to challenges in hiring a highly qualified postdoctoral fellow for the project. We initially identified and extended offers to two well-trained recent Ph.D. graduates in the U.S. at different times, but both ultimately declined due to other employment opportunities. Subsequently, we offered the position to an international researcher, one of our top three candidates with extensive experience in relevant research areas. Unfortunately, there has been a significant delay in the visa process. If the researcher is ultimately unable to join the project, we will initiate a new recruitment process. We remain committed to catching up on the research timeline and plan to request a one-year no-cost extension during the third year of the project. What opportunities for training and professional development has the project provided?This project supported the training of one Ph.D. graduate student and one undergraduate research assistant during its second year. The graduate student made excellent progress in her Ph.D. program and successfully presented her dissertation research proposal seminar in the department. She was awarded the 2025 Lambley Family Scholarship by the Department of Entomology at K-State. In addition, she was selected as a 2024 ESA Science Policy Fellow, a prestigious two-year program that trains entomologists in policy advocacy. In Spring 2025, she had the opportunity to meet with Congressional offices on Capitol Hill, as well as representatives from federal agencies and non-governmental organizations, to discuss her research. The undergraduate research assistant involved in the project received the 2025 Most Promising Student Award from the Division of Biology at K-State. How have the results been disseminated to communities of interest?Relevant research findings were disseminated to scientific communities through oral and poster presentations at international, national, and regional conferences. Notably, results from this project were presented at the national meeting of the Entomological Society of America, held November 10-13, 2024. What do you plan to do during the next reporting period to accomplish the goals?Due to delays in recruiting a highly qualified postdoctoral fellow for the project, we will be working diligently to catch up on our research progress. Our efforts will focus on optimizing RNAi bioassay techniques, including the selection of HRP sequences and dosages, to enhance RNAi efficiency in diamondback moth larvae. We will profile siRNA populations in two insect species, the red flour beetle and the diamondback moth, by incubating species-specific long dsRNA targeting the chitinase 10 gene with gut tissues from each species. The resulting siRNAs will be sequenced and mapped to the corresponding long dsRNA to identify the most abundant siRNA sequences. We will then design and synthesize species-specific siRNAs and HRPs, incorporate them into LDH nanoparticles, and perform RNAi bioassays via oral delivery for each insect species.

Impacts
What was accomplished under these goals? We optimized a protocol to generate hairpin RNA polymer (HRP) targeting the chitin synthase 1 gene (CHS1), which plays a key role in the chitin biosynthetic pathway, in the diamondback moth, using a rolling circle transcription (RCT) platform. The optimized protocol simplifies the experimental procedure and yields more consistent HRP production. The resulting HRP ranged in size from <100 to 300 nucleotides, smaller than expected. This discrepancy may be due to the hairpin structure, which likely causes faster migration than linear RNA of the same length on agarose gels. These findings may also indicate that RCT synthesis inherently limits the length of HRP that can be generated. We evaluated galactochitosan as a potential nanocarrier for delivering CHS1 HRP. Galactochitosan is a chemically modified chitosan polymer synthesized by our research collaborators in the Department of Chemistry at our university and has been successfully used to deliver double-stranded RNA (dsRNA) to adult western corn rootworms via feeding. However, in this study, no visible nanoparticles were produced when galactochitosan and CHS1 HRP were combined using our previously established protocol for dsRNA. While the exact reason remains unclear, this outcome is likely due to an unsuitable ratio between galactochitosan and HRP, as the binding capacity of galactochitosan to HRP may differ from that to dsRNA. Further investigation is needed to identify the optimal ratio and enable the use of galactochitosan as a nanocarrier for HRP delivery. To explore more advanced nanocarriers for HRP delivery in insects, we adapted and optimized a protocol for synthesizing layered double hydroxide (LDH) nanosheets as an alternative delivery system. LDH nanosheets are positively charged, two-dimensional nanomaterials that serve as effective carriers for delivering dsRNA. Their layered structure allows for the intercalation or surface adsorption of negatively charged dsRNA molecules, protecting them from degradation and enhancing their cellular uptake. LDH was synthesized using magnesium nitrate and aluminum nitrate solutions in a Teflon-lined reaction chamber at 100?°C overnight. The resulting LDH slurry was washed with deionized water and resuspended in deionized water using ultrasonication. To determine the optimal HRP-to-LDH ratio, we tested 10 different LDH:HRP ratios (from 1:1 to 1:10) by incubating the mixtures at 37?°C for 1 hour, followed by evaluation of unbound HRP. Our results identified 1:8 as the optimal LDH:HRP ratio, at which 92.2% of HRP was incorporated into LDH, demonstrating the high binding capacity of LDH for HRP. This ratio now serves as a guideline for the efficient generation of HRP/LDH nanoparticles for this project. Degradation of interfering RNA in the insect gut following oral delivery or in the hemolymph following injection is one of the major factors contributing to low RNAi efficiency in insects, particularly in lepidopterans. To assess whether LDH can protect HRP and dsRNA from degradation in insect tissues, we compared the stability of naked HRP and dsRNA versus their LDH-incorporated forms after 24-hour incubations with hemolymph and midgut solutions dissected from fourth-instar diamondback moth larvae. Both naked HRP and dsRNA were degraded under these conditions, while their LDH-bound counterparts remained intact. These results indicate that LDH can effectively protect interfering RNA from degradation in the insect hemolymph and midgut. We further conducted feeding-based bioassays using cabbage leaf discs treated with either naked CHS1 HRP or CHS1 HRP/LDH nanoparticles to target CHS1 in third-instar larvae of the diamondback moth under laboratory conditions. Although the differences were not statistically significant, a clear trend of increased larval mortality was observed in all CHS1 HRP/LDH-treated groups. Specifically, larvae that were fed CHS1 HRP/LDH nanoparticles showed a 16.7% increase in mortality compared to those fed GFP HRP/LDH control nanoparticles. Similarly, larvae fed cabbage leaf discs treated with IAP1 HRP/LDH nanoparticles, targeting the inhibitor of apoptosis 1 gene (IAP1) involved in apoptosis regulation, exhibited a 20% increase in mortality relative to the control group. These promising findings support further optimization of RNAi bioassay techniques, including the selection of HRP sequences and dosages, to enhance RNAi efficiency in diamondback moth larvae.

Publications

  • Type: Peer Reviewed Journal Articles Status: Accepted Year Published: 2025 Citation: Liu Y., Zhang J., Li S., Chai L., Chang B. H., Malak M., Wakil A. E., Moussian B., Zhao Z., Zeng Z., Zhu K. Y., Zhang J. 2025. Chitosan nanoparticle-mediated delivery of dsRNA for enhancing RNAi efficiency in Locusta migratoria. Pest Manag. Sci. (in press, https://doi.org/10.1002/ps.8880).
  • Type: Peer Reviewed Journal Articles Status: Published Year Published: 2025 Citation: Kong L., Shen W., Zhang S., Wang X., Xu J., Yuan X., Xu Z., Zhu K. Y. 2025. Development and evaluation of RNA microsphere-based RNAi approaches for managing the striped flea beetle (Phyllotreta striolata), a globally destructive pest of Cruciferae crops. Pest Manag. Sci. 81: 1529-1538 (https://doi.org/10.1002/ps.8557).
  • Type: Peer Reviewed Journal Articles Status: Published Year Published: 2025 Citation: Soumaila Issa M., Johnson R., Park Y., Zhu K. Y. 2024. Functional roles of five cytochrome P450 transcripts in the susceptibility of the yellow fever mosquito to pyrethroids revealed by RNAi coupled with insecticide bioassay. Arch. Insect Biochem. Physiol. 117: e70013 (https://doi.org/10.1002/arch.70013).
  • Type: Book Chapters Status: Published Year Published: 2025 Citation: Liu W., Liu X., Zhao X., Zhu K. Y., Zhang J. 2025. RNAi in the migratory locust: Functional studies of the genes in the formation and development of cuticle. In: Smagghe G., Palli S. R. and Swevers L. [eds.], RNA interference in Agriculture: Basic Science to Applications, pp. 249-268. Springer Nature, Switzerland AG (ISBN: 978-3-031-81548-5)
  • Type: Book Chapters Status: Published Year Published: 2025 Citation: Wang Y., Zhang J., Song H., Shi X., Zhu K. Y., Zhang J. 2025. The migratory locust as a model for studying the mechanisms of RNAi. In: Smagghe G., Palli S. R. and Swevers L. [eds.], RNA interference in Agriculture: Basic Science to Applications, pp. 303-319. Springer Nature, Switzerland AG (ISBN: 978-3-031-81548-5, https://doi.org/10.1007/978-3-031-81549-2).
  • Type: Book Chapters Status: Published Year Published: 2025 Citation: Cooper A. M. W., Silver K., Zhu K. Y. 2024. Chapter 17: RNA interference. pp. 163-175. In: Liu D. [ed.], Handbook of Molecular Biotechnology. CRC Press, Boca Raton, FL, USA. (ISBN: 9780367517878, https://doi.org/10.1201/9781003055211).
  • Type: Conference Papers and Presentations Status: Other Year Published: 2025 Citation: Maille J., Reed J., Morrison W., Sixbury N., Rust W., Kessler-Mathieu M., Brabec D., Zhu K. Y., Scully E., Indian meal moth flight activity in response to food and pheromonal cues. Student 10-Min Paper Competition presented by JM at the 2025 North Central Branch Meeting, Lincoln, NE. April 12-16, 2025. Quellhorst H., Sakka M., Baliota G., Athanassiou C., Zhu K. Y., Morrison W., Amorphous silica dust for use against Sitophilus granarius and Rhyzopertha dominica after harvest. Ten-Min Paper presented by HQ at the 2025 North Central Branch Meeting, Lincoln, NE. April 12-16, 2025. Gerken A., Juan R. S., Maille J., Scully E., Zhu K. Y., Abshire J., Morrison W., Using AI detection for better monitoring of stored product insect pests. Symposium Presentation by AG at the 2025 North Central Branch Meeting, Lincoln, NE. April 12-16, 2025 (INVITED). Boddepalli A., Zhu K. Y., McKay T., Starkus L., Scheff D., Efficacy of contact insecticides applied to wood and metal surfaces for controlling stored product insects in shipping containers. Student Poster Competition presented by AB at the 2025 North Central Branch Meeting, Lincoln, NE. April 12-16, 2025. Singh R., Castaldi J., Zhu K. Y., Smolensky D., Scully E., Impact of sorghum polyphenolics on insecticidal susceptibility of lesser grain borer, Rhyzopertha dominica (Coleoptera: Bostrichidae). Student 10-Min Paper Competition presented by RS at the 2025 North Central Branch Meeting, Lincoln, NE. April 12-16, 2025. Maille J. M., Morrison W. R., Zhu K. Y., Scully E. D., Flying under the radar Using flight mills to improve Indian meal moth management. 3MT, Kansas State University, Manhattan, KS. Feb. 05, 2025.


Progress 04/01/23 to 03/31/24

Outputs
Target Audience:The primary target audiences of this study are the researchers interested in RNA interference (RNAi), insect molecular biology, and insect pest management. Because RNAi has been shown to have its great potential for pest management, industries and pesticide regulatory agencies involved in pest management may also be interested in our findings. Changes/Problems:Our research progress has been delayed due to the process of hiring a highly qualified postdoctoral fellow for the project. We identified and offered the position to two well-trained recent Ph.D. graduates in the U.S. at two different time points, but both finally declined our offers due to their other employment opportunities. We then made an offer to an international researcher (one of the three candidates) with extensive research experience in the related areas. However, the researcher experienced a significant delay in obtaining his visa. However, we are optimistic that he will start to work within a few weeks. We will quickly catch up with the research progress during the second year of the project and plan to request a no-cost extension for one year in the third year of the project. What opportunities for training and professional development has the project provided?This project allowed for the training of one Ph.D. graduate student and two undergraduate students in the first year of the project. The graduate student made excellent progress with her Ph.D. program. In presenting her research from this project, she won the third-place award in the Ph.D. graduate student poster competition at the Entomological Society of America North Central Branch Conference (March 24-27, 2024). The graduate student was also selected by the Graduate School as one of three students to represent K-State for attending the AAAS Catalyzing Advocacy in Science and Engineering (CASE) workshop in Washington, DC (April 2024). The two undergraduate students completed their undergraduate research experience (URE) by participating in this project. Each student made a poster presentation at the 2023 Entomology Undergraduate Research Symposium at K-State (December 8, 2023). How have the results been disseminated to communities of interest?Relevant research was disseminated to scientific communities of interest through oral and poster presentations at international, national, and regional conferences. The PD was invited as a keynote speaker to present RNAi research at the Second International Molecular Plant Protection Congress in Turkey (May 15-18, 2023). Results from this project were also presented at the national meeting of the Entomological Society of America (Nov. 5-8, 2023), and at the 79th annual meeting of the Entomological Society of America North Central Branch by the PD (Mar. 24-27, 2024). Through the Undergraduate Research Experience (URE) program in the Department of Entomology at K-State, two undergraduate students completed their research, and each made a poster presentation at the 2023 Entomology Undergraduate Research Symposium at K-State (December 8, 2023). What do you plan to do during the next reporting period to accomplish the goals?Due to a delay in recruiting a highly qualified postdoctoral fellow for the project, we will be working very hard to catch up with our research progress. We will generate recombinant Dicer-2 protein from each of the two insect species (the red flour beetle, the diamondback moth), profile siRNA populations using recombinant Dicer-2 proteins, sequence and map the siRNA against long dsRNA for each insect species, design and produce insect species-specific siRNA constructs, and perform RNAi bioassay for each siRNA construct to determine their efficacy against the target genes in each insect species. Further, we will continue to optimize the synthesis of RNA microsphere (RMS) which contains the repeated hairpin RNA units and evaluate the gene silencing efficiency of RMS against the diamondback moth larvae by feeding. We will start to write manuscripts based on our results from this project.

Impacts
What was accomplished under these goals? The progress of our research in the first year was delayed due to the challenge of recruiting highly qualified research personnel including a graduate student and a postdoctoral research fellow for the project. After long searches, a highly experienced graduate student was recruited and started to work on the project in July 2023 whereas a highly qualified postdoctoral fellow was also recruited and will start his research shortly. With the help of these highly experienced researchers, we would be able to make rapid progress during the remaining funding period. Our research progresses for the first year are highlighted as follows: We have successfully established a laboratory colony of the diamondback moth using artificial diet. The egg masses were placed into Styrofoam cups for them to hatch and for the larvae to develop on the diet. After the larvae developed into a desirable stage, they were collected for laboratory studies. To maintain the diamondback moth colony in the laboratory, a certain number of the diamondback moth larvae were allowed to develop into pupae. The rearing cups with the pupae were placed in a rearing cage in a growth chamber. The adults were fed with 5% sugar solution and allowed to lay their eggs on aluminum foil treated with cabbage leaf juice. The egg strips were then used to produce the larvae. We further developed a feeding-based RNAi bioassay method using cabbage leaf discs for the diamondback moth larvae. We found that a method using 10 third-instar larvae per glass vial (19 mm in diameter and 51 mm in height) containing one cabbage leaf disc (11 mm in diameter) was most appropriate for the RNAi bioassay. We identified candidate RNAi target genes encoding chitin synthase 1 (CHS1), a key enzyme involved in the biosynthesis of new chitin for insect exoskeleton, and chitinase 10 (CHT10), an enzyme involved in the degradation of old chitin in insect midgut and exoskeleton, from both the red flour beetle and the diamondback moth. Based on the identified cDNA sequences, we designed three pairs of PCR primers for each gene and for each of the two insect species. The feasibility of using these primer sets to amplify approximately 500 bp cDNA fragments were validated by PCR. The alignments of their deduced protein sequences with those identified in other insect species using Neighbor-joining methos in Mega X revealed the identities of these sequences for the synthesis of double-stranded RNA (dsRNA). To address the challenges associated with the low RNAi efficiency in insects, particularly in lepidopteran insect pests, and with the complicated procedures to produce the conventional RNAi nanoparticles, we used a new technique known as rolling circle transcription (RCT) to generate RNA microspheres (RMS) to suppress the expression of CHS1 and CHT10 in the diamondback moth larvae by feeding. The RCT technology relies on the transcription based on a circular single-stranded DNA (ssDNA) template through the action of T7 RNA polymerase to produce long, single-stranded RNA containing multiple hairpin RNA unit repeats. The resulting RNA molecule forms a stable microsphere structure without a need for additional nanomaterials. The structures of the RMS can protect RNA from degradation during siRNA delivery and be successfully transported into the cytoplasm of the cells. Using siRNA design software, we designed several 75-base long ssDNA templates for RMS synthesis. In addition, we tried to optimize the synthesis methods to improve the RMS yield. Our initial characterization of RMS using a scanning electron microscope showed the size of approximately 180 micrometers in diameter. We also included experiments for comparisons between the long doubled-stranded RNA (dsRNA) and the RMS targeting the same genes in the diamondback moth larvae. We found that there was significant knockdown of CHT10 gene 3 days after treatment with the CHT10 dsRNA and significant knockdown of CHS1 gene 7 days after treatment with the CHS1 dsRNA. These results suggest that the timeline of suppression could vary by target gene. When the diamondback moth larvae were fed the RMS to target CHS1, we found a significantly increased larval mortality as compared with that of the control larvae fed the GFP RMS on day 3. However, we would need to adjust the sampling timepoints and bioassay methods to avoid the larval mortality in the control which might have confounded our results.

Publications

  • Type: Journal Articles Status: Published Year Published: 2024 Citation: Zhang F., Zhang Y. C., Yu Z. T., Zeng B., Sun H., Xie Y. Q., Zhu K. Y., Gao C. F. 2024. The G932C mutation of chitin synthase 1 gene (CHS1) mediates buprofezin resistance as confirmed by CRISPR/Cas9-mediated knock-in approach in the brown planthopper, Nilaparvata lugens. Pestic. Biochem. Physiol. 202: 105953 (https://doi.org/10.1016/j.pestbp.2024.105953). Zhao Y., Liu W., Zhao X., Yu Z., Guo H., Yang Y., Merzendorfer H., Zhu K. Y., Zhang J. 2024. Low-density lipoprotein receptor-related protein 2 (LRP2) is required for lipid export in the midgut of the migratory locust, Locusta migratoria. J. Integr. Agr. 23: 1618-1633 (https://doi.org/10.1016/j.jia.2023.07.027). Zhu Y., Kong L., Wang X., Xu J., Qian X., Yang Y., Xu Z., Zhu K. Y. 2023. Rolling circle transcription: A new system to produce RNA microspheres for improving RNAi efficiency in an agriculturally important lepidopteran pest (Mythimna separate). Pestic. Biochem. Physiol. 197: 105680 (https://doi.org/10.1016/j.pestbp.2023.105680). Wang Y., Li H., Liu X., Gao L., Fan Y., Zhu K. Y., Zhang J. 2023. Three alternative splicing variants of Loquacious play different roles in miRNA- and siRNA-mediated RNAi pathways in Locusta migratoria. RNA Biol. 20: 323-333 (https://doi.org/10.1080/15476286.2023.2223484). Zhang Y.-C., Gao Y., Ye W.-N., Peng Y.-X., Zhu K. Y., Gao C.-F. 2023. CRISPR/Cas9-mediated knockout of NlCYP6CS1 gene reveals its role in detoxification of insecticides in Nilaparvata lugens (Hemiptera: Delphacidae). Pest Manag. Sci. 79: 2239-2246 (https://doi.org/10.1002/ps.7404). Han P., Chen D., Fan J., Zhang J., Jiang S., Zhu K. Y., Zhang J. 2023. Genetically engineered Metarhizium anisopliae expressing dsRNA of Apolipophorin-D exhibits enhanced insecticidal virulence against Locusta migratoria. Entomol. Gen. 43: 167-175 (10.1127/entomologia/2023/1772). Xiao D., Yao J., Gao X., Zhu K. Y. 2023. Clathrin-dependent endocytosis plays a critical role in larval and pupal development, and female oocyte production in the red flour beetle (Tribolium castaneum). Pest Manag. Sci. 79: 1731-1742 (https://doi.org/10.1002/ps.7348). Zhao Y., Liu W., Zhao X., Yu Z., Guo H., Yang Y., Moussian B., Zhu K. Y., Zhang J. 2023. Lipophorin receptor is required for the accumulations of cuticular hydrocarbons and ovarian neutral lipids in Locusta migratoria. Inte. J. Biol. Macromol. 236: 123746 (https://doi.org/10.1016/j.ijbiomac.2023.123746). Fatehi S., Aikins M., Phillips T. W., Brown S., Zhu K. Y., Scully E. D., Park Y. 2023. Characterization of Iflavirus in the red flour beetle, Tribolium castaneum (Coleoptera: Tenebrionidae). Insects 14: 220 (https://www.mdpi.com/2075-4450/14/3/220). Zeng B., Chen F. R., Liu Y. T., Guo D., Zhang Y. J., Feng Z. R., Wang L. X., Vontas J., Wu S. F., Zhu K. Y., Gao C. F. 2023. A chitin synthase mutation confers widespread resistance to buprofezin, a chitin synthesis inhibitor, in the brown planthopper, Nilaparvata lugens. J. Pest Sci. 96: 819-832 (https://doi.org/10.1007/s10340-022-01538-9).
  • Type: Book Chapters Status: Accepted Year Published: 2024 Citation: Wang Y., Zhang J., Song H., Shi X., Zhu K. Y., Zhang J. 2024. Chapter A: The migratory locust as a model for studying the mechanisms of RNA interference. RNAi Book (in press). Liu W., Liu X., Zhao X., Zhu K. Y., Zhang J. 2024. Chapter B: RNA interference in the migratory locust: Functional studies of the genes in the formation and development of cuticles. RNAi Book (in press). Zhang J., Zhu K. Y. et al., 2024. Development of Insect Cuticle and Pest Management. Science Press, Beijing, China (In Chinese). pp. 254. (ISBN: 9787030775757) Cooper A. M. W., Silver K., Zhu K. Y. 2022. Chapter 19: RNA interference. pp. xx-xx. In: Liu D. [ed.], Handbook of Molecular Biotechnology. CRC Press, Boca Raton, FL, USA (in press).
  • Type: Conference Papers and Presentations Status: Published Year Published: 2024 Citation: Johnson R., Zhu K. Y., Comparing the RNAi efficiency of long double stranded RNA and RNA microspheres in suppressing gene expression in the diamondback moth, Plutella xylostella. Poster (D37) presented by RJ at the 79th Annual Meeting of the Entomological Society of America North Central Branch, Fort Collins, CO. Mar. 24-27, 2024. Zhu K. Y., RNAi-based insect pest management: Promises, challenges, and opportunities. College of Biological Science and Technology, Taiyuan Normal University, Jinzhong City, Shanxi Province, China. Dec. 29, 2023 (INVITED). Zhu K. Y., RNAi-based insect pest management: Promises, challenges, and opportunities. College of Plant Protection, Nanjing Agricultural University, Nanjing, China. Dec. 19, 2023 (INVITED). Zhu K. Y., RNAi-based insect pest management: Promises, challenges, and opportunities. NeoAgro, Zhejiang Academy of Agricultural Sciences, Hangzhou, China. Dec. 15, 2023 (INVITED). Gibson J., Johnson R., Zhu K. Y., Efficacy of RNAi delivered via artificial diet on the diamondback moth, Plutella xylostella. Poster presented by JG at the 2023 Entomology Undergraduate Research Symposium at Kansas State University, Manhattan, KS. Dec. 8, 2023. Hulse K., Johnson R., Zhu K. Y., Feeding-based RNAi against chitin-related genes in the diamondback moth, Plutella xylostella. Poster presented by KH at the 2023 Entomology Undergraduate Research Symposium at Kansas State University, Manhattan, KS. Dec. 8, 2023. Zhu K. Y., Implication of a potential mechanism of resistance to RNAi-based biopesticides in insects. Section Symposium, Entomology 2023: National Meeting of the Entomological Society of America, National Harbor, MD. Nov. 5-8, 2023 (INVITED). Zhu K. Y., Zhang J., Breaking down the barriers: Strategies to enhance RNAi efficiency in insects. Presented by KYZ at the Second International Molecular Plant Protection Congress, Orhangazi-Bursa, Turkey. May 15-18, 2023 (INVITED KEYNOTE SPEECH). Zhu K. Y., Breaking down the barriers: Strategies to enhance RNAi efficiency in insects. Insect Group Meeting, Department of Biochemistry and Biophysics, Kansas State University, Manhattan, KS. Jan. 27, 2023.