This research was funded by the National Natural Science Foundation of China (31870637, 31200487) and jointly funded by the Zhejiang Key Research Plan (2019C02024, LGN22C160004).

The study was approved by the council for animal experimentation of Zhejiang Agricultural &#38; Forestry University. The B. xylophilus isolate NXY61 was originally extracted from a diseased Pinus massoniana in the Ningbo area of Zhejiang province, China11.
1. Gene cloning
NOTE: See the Table of Materials for details about the primers used in this protocol.
<ol>
	<li>Collect nematodes.
	<ol>
		<li>Culture the B. xylophilus strain on the mycelia of Botrytis cinerea on Potato Dextrose Agar (PDA) plates at 25 &#176;C for 3-5 days.</li>
		<li>Collect the nematodes using the Bellman funnel method14.
		<ol>
			<li>Place a clamped rubber tube below a funnel and place two layers of filter paper in the mouth of the funnel. Transfer the fungal cultures to the funnel and add water to immerse the fungal mat. Wait for 2 h, then collect the nematodes.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Extract the total RNA from the nematodes using a total RNA extraction reagent (see the Table of Materials)11 according to the following steps.
	<ol>
		<li>Add 500 &#181;L of extraction reagent and 100 &#181;L of magnetic beads to a 2 mL centrifuge tube. Aspirate 20 &#181;L of the nematodes and transfer the sample to a grinder for grinding at 9,000 &#215; g for 30 s. Incubate for 5 min and then centrifuge for 10 min at 12,000 &#215; g and 4 &#176;C.</li>
		<li>Transfer the supernatant to a new centrifuge tube. Add 100 &#181;L of chloroform, cap the tube, and mix by inverting the tube several times. Incubate for 3 min and then centrifuge for 10 min at 12,000 &#215; g at 4 &#176;C.</li>
		<li>Transfer the supernatant to a new centrifuge tube. Add 250 &#181;L of isopropyl alcohol and vortex vigorously. Centrifuge at 12,000 &#215; g for 10 min.&#160;</li>
		<li>Discard the supernatant. Add 500 &#181;L of 75% ethanol to wash the RNA and then vortex the sample. Centrifuge it for 5 min at 12,000 &#215; g and 4 &#176;C.</li>
		<li>Air-dry the RNA pellet for 5 min.</li>
		<li>Resuspend the pellet in 30 &#181;L of RNase-free water.</li>
		<li>Calculate the RNA concentration using the formula: A260 &#215; dilution &#215; 40 = &#181;g RNA/mL. Calculate the A260/A280 ratio. 
		&#8203;NOTE: A ratio of ~2 is considered pure.</li>
	</ol>
	</li>
	<li>Perform reverse transcription of good quality RNA to obtain the cDNA template.
	<ol>
		<li>Design and use a pair of specific primers, ppm-1-F/R (see the Table of Materials), to amplify the partial coding sequence of the Bx-ppm-1 gene in B. xylophilus (GenBank accession number QTZ96795).</li>
		<li>Clone the ppm-1 gene sequences into the pGEM-Teasy vector containing the T7 promoter following a standard cloning protocol11.
		<ol>
			<li>Set up the PCR reactions as follows: 2 &#181;L of cDNA, 25 &#181;L of 2x Ex Taq Polymerase Premix, 2 &#181;L of each primer (10 pmol/l), and sterile distilled water to a final volume of 50 &#181;L.</li>
			<li>Perform the amplification procedure as follows: 5 min at 94 &#176;C; followed by 35 cycles of 30 s at 94 &#176;C, 30 s at 55 &#176;C, and 1 min at 72 &#176;C; and a final extension step at 72 &#176;C for 5 min.</li>
			<li>Clone the amplified products into a pGEM-T Easy vector for sequencing.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Synthesis of dsRNA
<ol>
	<li>Prepare the DNA template for dsRNA synthesis using PCR with primers designed to add T7 promoter sites at both ends. Add the T7 promoter sequence to the 5&#39; end of the primers.</li>
	<li>Use the plasmid containing the ppm-1 gene fragment (894 bp) as the template for PCR and recover the fragment containing the T7 promoter11. Use the PCR procedure and system described above.</li>
	<li>Use an in vitro transcription kit to synthesize dsRNA11.
	<ol>
		<li>Thaw the frozen reagents on ice.</li>
		<li>Add 2 &#181;L of 10x reaction buffer, 2 &#181;L of enzyme mix, and 1 &#181;g of DNA to a centrifuge tube. Add nuclease-free water to produce a standard 4 &#181;L reaction. Then, mix equal volumes of the four ribonucleotide solutions (ATP, CTP, GTP, and UTP) together and add 8 &#181;L of the mixture to the tube. Mix thoroughly and incubate at 37 &#176;C for 4 h.</li>
		<li>Add 1 &#181;L of DNase, mix well, and incubate for 15 min at 37 &#176;C.</li>
		<li>Stop the reaction and add 30 &#181;L of nuclease-free water and 30 &#181;L of LiCl precipitation solution to precipitate the RNA. Mix thoroughly. Incubate at -20 &#176;C overnight.</li>
		<li>Centrifuge for 15 min at 12,000 &#215; g and 4 &#176;C. Discard the supernatant.</li>
		<li>Add 1 mL of 75% ethanol to wash the RNA. Vortex the sample and centrifuge it for 10 min at 12,000 &#215; g and 4 &#176;C.</li>
		<li>Air-dry the RNA pellet for 3 min.</li>
		<li>Resuspend the pellet in 30 &#181;L of RNase-free water.</li>
		<li>Analyze the quality of the dsRNA using a spectrophotometer. Pipette 1 &#181;L of the dsRNA sample onto the measurement pedestal and set the wavelength to 340 nm. Visualize the products on a 1.0% agarose gel.</li>
	</ol>
	</li>
</ol>
3. RNAi by soaking
<ol>
	<li>Mix 4 &#181;L of 5x soaking buffer (0.05% gelatin, 5.5 mM KH2PO4, 2.1 mM NaCl, 4.7 mM NH4Cl, 3 mM spermidine) with the dsRNA and ddH2O to produce a total volume of 20 &#181;L and a final RNA concentration of 0.8 &#181;g/mL.</li>
	<li>Acquire J2 larvae.
	<ol>
		<li>Collect the nematodes from the fungal cultures and transfer them to a glass Petri dish 6 cm in diameter. Add 10 mL of water to the dish to ensure that the nematodes can swim freely. Keep the nematodes in the dish for 30 min and wait for the eggs to adhere to the bottom.</li>
		<li>Remove the water and nematodes carefully, making sure not to disturb the eggs. Repeat the steps until all the larvae and adults are removed, leaving only the eggs in the dish.</li>
		<li>Hatch the collected eggs for 24 h in the dark at 25 &#176;C to obtain J2 larvae. Collect the J2 larvae, place them in a tube, and wash them three times with ddH2O for the RNAi experiment15.</li>
		<li>Transfer the J2 larvae to the 2 mL tube containing the dsRNA solution and add resorcinol solution (wrapped in tinfoil and dissolved in water) to produce a final concentration of 1.0%. Incubate the larvae with centrifugation at 15&#160;&#215;&#160;g on a shaking table for 24 h at 25 &#176;C to ensure that the larvae effectively absorb the dsRNA.</li>
		<li>Soak the same quantity of nematodes in the soaking buffer without the dsRNA probe or with a GFP dsRNA probe as a control. Use the GFP gene (gfp, M62653.1) as a nonendogenous control and synthesize the dsRNA of gfp using gene-specific primers T7-GFP-F/R.</li>
	</ol>
	</li>
</ol>
4. qPCR detection
<ol>
	<li>Clean the J2 larvae with ddH2O, including those with target gene interference, GFP gene interference, and the undisturbed control group. Extract the total RNAs from each group using the method described above.</li>
	<li>Start the qPCR.
	<ol>
		<li>Design the q-ppm-1-F/R primers using the desired software (see the Table of Materials).</li>
		<li>Set up the qPCR reaction in 12 &#181;L containing 1 &#181;L of cDNA, 6 &#181;L of fluorescent premix, 0.4 &#181;L of each primer (10 pmol/l), and sterile distilled water.</li>
		<li>Perform qPCR as follows: 2 min at 95 &#176;C; followed by 40 cycles of 10 s at 95 &#176;C, 30 s at 55 &#176;C, and 1 min at 72 &#176;C.</li>
		<li>Use the actb gene (GenBank accession number EU100952) and tbb-2 gene (GenBank accession number MT769316),&#160;or other genes, as internal reference genes to evaluate the changes in gene expression level after RNAi15.</li>
	</ol>
	</li>
	<li>According to the cycle threshold (Ct) value and the dissolution curve, use the 2-&#916;&#916;Ct method to estimate the relative expression level of the target gene and verify the interference efficiency.
	<ol>
		<li>Subtract the Ct value of the internal reference gene of each sample from the Ct value of the target gene to obtain the &#916;Ct value. Then, subtract the &#916;Ct value of the interference group from the &#916;Ct value of the control group to obtain the &#916;&#916;Ct value. 
		&#8203;NOTE: A &#916;&#916;Ct value greater than 0 indicates that the interference is effective.</li>
	</ol>
	</li>
</ol>
5. Evaluate the body length of nematode adults following RNAi
<ol>
	<li>After RNAi, culture the J2 larvae until adulthood on B. cinerea lawns in PDA plates for 60 h at 25 &#176;C15.</li>
	<li>Collect the adults using the Bellman funnel method (see step 1.1.2.1)14.</li>
	<li>Acquire images of the adult nematodes under a microscope and use ImageJ software (or other measurement software) to measure the body lengths.
	<ol>
		<li>Measure the length using ImageJ by selecting Analyze | Set Scale. If the distance is known, enter the length value of the drawn straight line. Enter the unit of length.</li>
		<li>Check the Global check box (use this standard for all pictures), and click OK to select the length to be measured with a straight line on the image to be measured.</li>
		<li>Use the command Ctrl+M (measurement) to display the results and record them in the results window.</li>
	</ol>
	</li>
	<li>Take measurements of 50 male and 50 female nematodes for statistical analysis12.</li>
	<li>Analyze the data by calculating the mean and standard deviation for each sample. Compare the means of the samples from the different groups using the Student&#39;s t-test.</li>
</ol>

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No conflicts of interest were declared.

Although the life history and parasitic environment of B. xylophilus are different from those of other nematodes, there has been limited research on the molecular pathogenesis of this plant pathogen. Despite great progress made in the application of CRISPR/Cas9 genome editing technology in C. elegans and other nematodes, only RNAi technology applied to B. xylophilus has been published to date17. RNAi is one of the most powerful tools available to study the gene function of nematodes and has been widely used in research elucidating B. xylophilus gene function, signal transduction pathways, and gene therapy18,19,20. In contrast to the gene knockout approaches used in invertebrate models, RNAi-mediated knockdown of target genes in nematodes enables the rapid evaluation of gene function.
There are three ways to conduct RNAi in C. elegans: injection6, soaking21, and feeding6. Because J2 larvae only feed after infection, the soaking method is typically used for the RNAi of plant-parasitic nematodes. The key step in the soaking method is to add a nerve agent to stimulate the nematodes to feed. Urwin first used octopamine to stimulate the J2 of two oral cyst nematode species, Globodera pallida, and Heterodera glycines, resulting in the uptake of dsRNA from the soaking solution22. The same method has been successfully used to induce J2 of the root-knot nematode Meloidogyne incognita to absorb dsRNA 23. Resorcinol can also induce the uptake of dsRNA by J2 of&#160;M. incognita and may be more effective than octopamine for this nematode24. Furthermore, the addition of spermidine to the soaking buffer with a prolonged incubation time improved the efficiency of RNAi in nematodes25. After 24 h of soaking, dsRNA effectively enters B. xylophilus, thereby silencing the ppm-1 gene.
This protocol, therefore, provides a reference for the future study of the RNAi of other plant-parasitic nematodes with phagocytosis behavior. In addition, the suspension effect of the shaking table plays an important role in maximizing the advantage of the soaking method. This method can allow for the simultaneous treatment of a large number of nematodes. RNAi is easier to operate when focused on the target genes for which mutants cannot easily be obtained and for evaluating the effects at different points in development because nematodes can be soaked at any life stage. Thus, RNAi can be used to study the function of a specific gene at a specific developmental stage. Wang et al. analyzed the influence of MAPK on the fecundity of B. xylophilus and its important role in the growth and development of nematodes using dsRNA soaking26. Qiu et al. found that the migration speed and fecundity of B. xylophilus in pine trees decreased after downregulation of the Bxpel1 gene, which indicates that this gene is an important pathogenic factor of B. xylophilus27.
However, not all RNAi in nematodes works through dsRNA. Dulovic and Streit applied small interfering RNAs (siRNAs) rather than the longer dsRNAs to successfully interfere with Strongyloides ratti28. The limitation of using dsRNA for RNAi&#160;in Strongyloides may be related to the absence of genes such as rsd-6, sid-1, or sid-2 that are known to be involved in dsRNA uptake. RNAi is also associated with the disadvantages of a positional effect and temporary and incomplete knockout and has limited effectiveness for some genes and cell types (such as neurons)29. However, until a breakthrough in the transgenic technology of B. xylophilus is achieved, RNAi represents a relatively effective research strategy.
RNAi has shown great potential for crop protection. Using RNAi technology, transgenic potatoes capable of producing dsRNA that targets root-knot nematode genes have been bred to produce complete resistance to root-knot nematodes30. Expression of dsRNAs that target insect genes can provide crop protection in the absence of chemical insecticides and offers the additional advantage of not producing foreign proteins, with an almost unlimited number of target genes31. Therefore, accelerated research in the field of applied RNAi for pest control will provide better biosecurity for plant nematode control than transgenic methods or other chemical control methods.
In conclusion, this protocol describes the preparation of dsRNA of the ppm-1 gene to achieve RNAi by directly soaking the larvae of B. xylophilus in dsRNA solution. The interference effect was confirmed based on a significant reduction in the body size of the larvae after they developed into adults compared with that of the control, demonstrating that&#160;the ppm-1 gene exerts effects on the growth and development of B. xylophilus. This study provides a theoretical basis for further revealing the function of ppm-1 with additional practical value in guiding the biological control of this plant parasite. It is believed that with the further disclosure and improvement of the RNAi technology mechanism, its application in the field of B. xylophilus control will have broad prospects.

Plant-parasitic nematodes (PPNs) are a continuing threat to food security and forest ecosystems. They cause an estimated 100 billion USD in economic losses each year1, the most problematic of which are primarily root-knot nematodes, cyst nematodes, and pinewood nematodes. The pinewood nematode, Bursaphelenchus xylophilus, is a migratory, endoparasitic nematode, which is the causal pathogen of pine wilt disease2. It has caused great harm to pine forests worldwide3. Using the terminology of Van Megen et al.4, B. xylophilus is a member of the Parasitaphelenchidae and belongs to clade 10, whereas most other major plant parasites belong to clade 12.
As an independent and recently evolved plant parasite, B. xylophilus is an attractive model for comparative studies. To date, there has been substantial research on root-knot nematodes and cyst nematodes belonging to clade 12, which are obligate, sedentary endoparasites and are some of the most intensely studied nematodes. However, conducting further research in this important area comes with a major challenge: the function of parasitism genes is a research bottleneck. Functional studies generally include ectopic expression and knockdown/out experiments but rely on effective genetic transformation protocols for the nematode. As a result, reverse genetics in PPNs almost exclusively relies on gene silencing by RNAi.
RNAi, a mechanism widely present in eukaryotic cells, silences gene expression by introducing double-stranded RNA (dsRNA)5. To date, the posttranscriptional gene-silencing mechanism induced by dsRNA has been found in all studied eukaryotes, and RNAi technology, as a tool of functional genomics research and other applications, has developed rapidly in many organisms. Since the discovery of the RNAi machinery in Caenorhabditis elegans in 19986, RNAi techniques have become effective methods for identifying the gene function of nematodes and are proposed as a new way to effectively control pathogenic nematodes7.
RNAi is technically facile-soaking the juveniles in dsRNA can suffice; however, the efficacy and reproducibility of this approach vary widely with the nematode species and the target gene8. The silencing of 20 genes involved in the RNAi pathways of the root-knot nematode, Meloidogyne incognita, was investigated using long dsRNAs as triggers, resulting in diverse responses, including an increase and no change in the expression of some genes9. These results show that target genes may respond to RNAi knockdown differently, necessitating an exhaustive assessment of their suitability as targets for nematode control via RNAi. However, there is currently a paucity of research on the developmental and reproductive biology of B. xylophilus.
As a continuation of previous work10,11,12,13, we describe here a protocol for applying RNAi to study the function of&#160;the ppm-1 gene of B. xylophilus, including the synthesis of dsRNA, synthetic neurostimulant soaking, and quantitative polymerase chain reaction (qPCR) detection. The knowledge gained from this experimental approach will likely contribute markedly to understanding basic biological systems and preventing pine wilt disease.

<table><tbody><tr><td>Baermann funnel</td><td>n/a</td><td>n/a</td><td>to isolate nematodes</td></tr><tr><td>Beacon Designer 7.9</td><td>Shanghai kangyusheng information technology co.</td><td>n/a</td><td>to design qPCR primers</td></tr><tr><td>Botrytis cinerea</td><td>n/a</td><td>n/a</td><td>as food for nematodes</td></tr><tr><td>&lt;em&gt;Bursaphelenchus xylophilus&lt;/em&gt;</td><td>n/a</td><td>n/a</td><td>its number was NXY61 and was it was originally extracted from diseased&lt;br/&gt; Pinus massoniana in Ningbo, Zhejiang province, China.</td></tr><tr><td>constant temperature incubator</td><td>Shanghai Jing Hong Laboratory Instrument Co.</td><td>H1703544</td><td>to cultur nematodes</td></tr><tr><td>Electrophoresis apparatus</td><td>Bio-Rad Laboratories</td><td>1704466</td><td>to achieve electrophoretic analysis</td></tr><tr><td>Ethanol, 75%</td><td>Sinopharm Chemical Reagent Co.</td><td>80176961</td><td>to extract RNA</td></tr><tr><td>Ex Taq Polymerase Premix</td><td>Takara Bio Inc.</td><td>RR030A</td><td>for PCR</td></tr><tr><td>Ex Taq Polymerase Premix</td><td>Takara Bio Inc.</td><td>RR390A</td><td>for PCR</td></tr><tr><td>Gel imager</td><td>LongGene Scientific Instruments Co.</td><td>LG2020</td><td>to make nucleic acid bands visible</td></tr><tr><td>GraphPad Prism 8</td><td>GraphPad Prism</td><td>n/a</td><td>to analyze the data and make figurs</td></tr><tr><td>High Speed Centrifuge</td><td>Hangzhou Allsheng Instruments Co.</td><td>AS0813000</td><td>centrifug</td></tr><tr><td>High-flux tissue grinder</td><td>Bertin</td><td></td><td>to extract RNA</td></tr><tr><td>ImageJ software</td><td>National Institutes of Health</td><td>n/a</td><td>to measure the body lengths</td></tr><tr><td>isopropyl alcohol</td><td>Shanghai Aladdin Biochemical Technology Co.</td><td>L1909022</td><td>to extract RNA</td></tr><tr><td>Leica DM4B microscope</td><td>Leica Microsystems Inc.</td><td></td><td>to observe nematodes</td></tr><tr><td>magnetic beads</td><td>Aoran science technology co.</td><td>150010C</td><td>to extract RNA</td></tr><tr><td>MEGAscript T7 High Yield Transcription Kit</td><td>Thermo Fisher Scientific Inc.</td><td>AM1333</td><td>to synthesize dsRNA in vitro</td></tr><tr><td>NanoDrop ND-2000 spectrophotometer</td><td>Thermo Fisher Scientific Inc.</td><td>NanoDrop 2000/2000C</td><td>to analyze the quality of the dsRNA</td></tr><tr><td>PCR Amplifier</td><td>Bio-Rad Life Medical Products Co.</td><td>1851148</td><td>to amplify nucleic acid sequence</td></tr><tr><td>Petri dishes</td><td>n/a</td><td>n/a</td><td>to cultur nematodes</td></tr><tr><td>pGEM-T Easy vector</td><td>Promega Corporation</td><td>A1360</td><td>for cloning</td></tr><tr><td>Potato Dextrose Agar (Medium)</td><td>n/a</td><td>n/a</td><td>to cultur &lt;em&gt;Botrytis cinerea&lt;/em&gt;</td></tr><tr><td>Prime Script RT reagent Kit with gDNA Eraser</td><td>Takara Bio Inc.</td><td>RR047B</td><td>to synthetic cDNA</td></tr><tr><td>Primer Premier 5.0</td><td>PREMIER Biosoft</td><td>n/a</td><td>to design PCR primers</td></tr><tr><td>primers:ppm-1-F/R</td><td>Tsingke Biotechnology Co.</td><td>n/a</td><td>F: 5&#039;-GATGCGAAGTTGCCAATCATTCT -3&#039;; R: 5&#039;- CCAGATCCAGTCCACCATACACC -3</td></tr><tr><td>q-ppm-1-F/R</td><td>Tsingke Biotechnology Co.</td><td>n/a</td><td>F: 5&#039;-CATCCGAATGGCAATACAG-3&#039;; R: 5&#039;-ACTATCCTCAGCGTTAGC-3&#039;</td></tr><tr><td>Real-time thermal cycler&amp;nbsp;qTOWER 2.2</td><td>Analytique Jena Instruments (Beijing) Co.</td><td></td><td>for qPCR</td></tr><tr><td>shaking table</td><td>Shanghai Zhicheng analytical instrument manufacturing co.</td><td></td><td>to soak nematodes</td></tr><tr><td>stereoscopic microscope</td><td>Chongqing Optec Instrument Co.</td><td>1814120</td><td>to observe nematodes</td></tr><tr><td>T7-GFP-F/R</td><td>Tsingke Biotechnology Co.</td><td>n/a</td><td>F: 5&#039;-TAATACGACTCACTATAGGGAAA&lt;br/&gt; GGAGAAGAACTTTTCAC-3&#039;; R: 5&#039;-TAATACGACTCACTATAGGGCTG&lt;br/&gt; TTACAAACTCAAGAAGG-3&#039;</td></tr><tr><td>&amp;nbsp;T7 promoter</td><td>Tsingke Biotechnology Co.</td><td>n/a</td><td>TAATACGACTCACTATAGGG</td></tr><tr><td>Takara MiniBEST Agarose Gel DNA Extraction Kit</td><td>Takara Bio Inc.</td><td>9762</td><td>to recover DNA</td></tr><tr><td>TaKaRa TB Green Premix Ex Taq (Tli RNaseH Plus)</td><td>Takara Bio Inc.</td><td>RR820A</td><td>for qPCR</td></tr><tr><td>trichloroethane</td><td>Shanghai LingFeng Chemical Reagent Co.</td><td></td><td>to extract RNA</td></tr><tr><td>TRIzol Reagent</td><td>Thermo Fisher Scientific Inc.</td><td>15596026</td><td>total RNA extraction reagent,to extract RNA</td></tr></tbody></table>

application of rna interference in the pinewood nematode, bursaphelenchus xylophilus

The pinewood nematode, Bursaphelenchus xylophilus, is one of the most destructive invasive species worldwide, causing the wilting and eventual death of pine trees. Despite the recognition of their economic and environmental significance, it has thus far been impossible to study the detailed gene functions of plant-parasitic nematodes (PPNs) using conventional forward genetics and transgenic methods. However, as a reverse genetics technology, RNA interference (RNAi) facilitates the study of the functional genes of nematodes, including B. xylophilus. 
This paper outlines a new protocol for RNAi of the ppm-1 gene in B. xylophilus, which has been reported to play crucial roles in the development and reproduction of other pathogenic nematodes. For RNAi, the T7 promoter was linked to the 5&#8242;-terminal of the target fragment by polymerase chain reaction (PCR), and double-stranded RNA (dsRNA) was synthesized by in vitro transcription. Subsequently, dsRNA delivery was accomplished by soaking the nematodes in a dsRNA solution mixed with synthetic neurostimulants. Synchronized juveniles of B. xylophilus (approximately 20,000 individuals) were washed and soaked in dsRNA (0.8 &#181;g/mL) in the soaking buffer for 24 h in the dark at 25 &#176;C.
The same quantity of nematodes was placed in a soaking buffer without dsRNA as a control. Meanwhile, another identical quantity of nematodes was placed in a soaking buffer with green fluorescent protein (gfp) gene dsRNA as a control. After soaking, the expression level of the target transcripts was determined using real-time quantitative PCR. The effects of RNAi were then confirmed using microscopic observation of the phenotypes and a comparison of the body size of the adults among the groups. The current protocol can help advance research to better understand the functions of the genes of B. xylophilus and other parasitic nematodes toward developing control strategies through genetic engineering.

Analysis of ppm-1 expression of B. xylophilus after RNAi 
The relative expression level of the ppm-1 gene of B. xylophilus soaked with GFP dsRNA and that soaked with target gene dsRNA was 0.92 and 0.52, respectively (the ppm-1 gene expression level of the ddH2O-treated control group was set to 1) (Figure 1). Thus, exogenous dsRNA has no effect on the ppm-1 expression of B. xylophilus; however, ppm-1 dsRNA can effectively inhibit the expression of the target gene.
Effect of ppm-1 expression on growth and development of B. xylophilus 
After RNAi, the size of the adults markedly decreased (Figure 2), resulting in the SMA (small body size) mutant phenotype. Although RNAi-treated individuals developed to sexual maturity, their body length was substantially smaller than normal adults. Specifically, after RNAi in J2-stage nematodes, the mean body length of females and males was 544.61 &#181;m and 526.24 &#181;m, respectively. In contrast, the mean body length of females and males in the control group was 971.86 &#181;m and 912.31 &#181;m, respectively (Figure 3), representing significant differences (P = 0.0322).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63645/63645fig01.jpg" /> 
Figure 1: Expression of the ppm-1 gene following RNAi of Bursaphelenchus xylophilus. **P &#60; 0.001. Abbreviations: RNAi = RNA interference; GFP = green fluorescent protein. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63645/63645fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63645/63645fig02.jpg" /> 
Figure 2: Body length reduction of adults after the interference of ppm-1 in Bursaphelenchus xylophilus. Images of an adult male (A) and female (C) of the RNAi group. Images of an adult male (B) and female (D) of the control group. Scale bars = 50 &#181;m. Abbreviation: RNAi = RNA interference. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63645/63645fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63645/63645fig03.jpg" /> 
Figure 3: Quantification and statistical analysis of the body length of adults of Bursaphelenchus xylophilus after RNAi of the ppm-1 gene. (*P = 0.0322). Abbreviation: RNAi = RNA interference. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63645/63645fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Application of RNA Interference in the Pinewood Nematode, Bursaphelenchus xylophilus at JoVE.com

Application of RNA Interference in the Pinewood Nematode, Bursaphelenchus xylophilus

Here, we introduce a detailed soaking method of RNA interference in Bursaphelenchus xylophilus to facilitate the study of gene functions.

Application of RNA Interference in the Pinewood Nematode, Bursaphelenchus xylophilus

The pinewood nematode, Bursaphelenchus xylophilus, is one of the most destructive invasive species worldwide, causing the wilting and eventual death of ...

application-rna-interference-pinewood-nematode-bursaphelenchus

Nanyang Technological University

Research

JoVE Journal

Biology

2.4K Views.  Zhejiang Agricultural & Forestry University. RNAi is een effectieve methode geworden voor het identificeren van de functie van genen in nematoden en wordt voorgesteld als een nieuwe manier om pathogene nematoden effectief te bestrijden. RNAi is het weken van nematoden in dsRNA kan voldoende zijn voor de procedure. Verzamel om te beginnen de nematoden volgens de Baermann-trechtermethode.Plaats hiervoor een geklemde rubberen buis onder een trechter en plaats twee lagen filterpapier in de mond van de trechter. Breng de Botrytis cine...

Video: Application of RNA Interference in the Pinewood Nematode, Bursaphelenchus xylophilus

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

Genetics

Double-stranded RNA Oral Delivery Methods to Induce RNA Interference in Phloem and Plant-sap-feeding Hemipteran Insects

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran insects account for a number of economically substantial pests of plants that cause damage to crops by feeding on phloem sap. The brown marmorated stink bug (BMSB), Halyomorpha halys (Heteroptera: Pentatomidae) and the Asian citrus psyllid (ACP), Diaphorina citri Kuwayama (Hemiptera: Liviidae) are hemipteran insect pests introduced in North America, where they are an invasive agricultural pest of high-value specialty, row, and staple crops and citrus fruits, as well as a nuisance pest when they aggregate indoors. Insecticide resistance in many species has led to the development of alternate methods of pest management strategies. Double-stranded RNA (dsRNA)-mediated RNA interference (RNAi) is a gene silencing mechanism for functional genomic studies that has potential applications as a tool for the management of insect pests. Exogenously synthesized dsRNA or small interfering RNA (siRNA) can trigger highly efficient gene silencing through the degradation of endogenous RNA, which is homologous to that presented. Effective and environmental use of RNAi as molecular biopesticides for biocontrol of hemipteran insects requires the in vivo delivery of dsRNAs through feeding. Here we demonstrate methods for delivery of dsRNA to insects: loading of dsRNA into green beans by immersion, and absorbing of gene-specific dsRNA with oral delivery through ingestion. We have also outlined non-transgenic plant delivery approaches using foliar sprays, root drench, trunk injections as well as clay granules, all of which may be essential for sustained release of dsRNA. Efficient delivery by orally ingested dsRNA was confirmed as an effective dosage to induce a significant decrease in expression of targeted genes, such as juvenile hormone acid O-methyltransferase (JHAMT) and vitellogenin (Vg). These innovative methods represent strategies for delivery of dsRNA to use in crop protection and overcome environmental challenges for pest management.

This article demonstrates novel techniques developed for oral delivery of double-stranded RNA (dsRNA) through the vascular tissues of plants for RNA interference (RNAi) in phloem sap feeding insects.

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran ...

Environment

RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is applicable to a wide variety of organisms and is highly efficient in the species-rich order Coleoptera (beetles). Here, we summarize the necessary steps for developing this technique in a novel organism and illustrate its application to the different developmental stages of the aquatic diving beetle Thermonectus marmoratus. Target gene sequences can be obtained cost-effectively through the assembly of transcriptomes against a close relative with known genomics or de novo. Candidate gene cloning utilizes a specific cloning vector (the pCR4-TOPO plasmid), which allows the synthesis of double-stranded RNA (dsRNA) for any gene with the use of a single common primer. The synthesized dsRNA can be injected into either embryos for early developmental processes or larvae for later developmental processes. We then illustrate how RNAi can be injected into aquatic larvae using immobilization in agarose. To demonstrate the technique, we provide several examples of RNAi experiments, generating specific knockdowns with predicted phenotypes. Specifically, RNAi for the tanning gene laccase2 leads to cuticle lightening in both larvae and adults, and RNAi for the eye pigmentation gene white produces a lightening/lack of pigmentation in eye tubes. In addition, the knockdown of a key lens protein leads to larvae with optical deficiencies and a reduced ability to hunt prey. Combined, these results exemplify the power of RNAi as a tool for investigating both morphological patterning and behavioral traits in organisms with only transcriptomic databases.

RNA interference is a widely applicable, powerful technique for manipulating gene expression at specific developmental stages. Here, we describe the necessary steps for implementing this technique in the aquatic diving beetle Thermonectus marmoratus, from the acquisition of gene sequences to the knockdown of genes that affect structure or behavior.

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is ...

Electroporation-mediated RNA Interference Method in Odonata

Dragonflies and damselflies (order Odonata) represent one of the most ancestral insects with metamorphosis, in which they change their habitat, morphology, and behavior drastically from aquatic larvae to terrestrial/aerial adults without pupal stage. Odonata adults have a well-developed color vision and show a remarkable diversity in body colors and patterns across sexes, stages, and species. While many ecological and behavioral studies on Odonata have been conducted, molecular genetic studies have been scarce mainly due to the difficulty in applying gene functional analysis to Odonata. For instance, RNA interference (RNAi) is less effective in the Odonata, as reported in the Lepidoptera. To overcome this problem, we successfully established an RNAi method combined with in vivo electroporation. Here we provide a detailed protocol including a video of the electroporation-mediated RNAi method as follows: preparation of larvae, species identification, preparation of dsRNA/siRNA solution and injection needles, ice-cold anesthesia of larvae, dsRNA/siRNA injection, in vivo electroporation, and individual rearing until adult emergence. The electroporation-mediated RNAi method is applicable to both damselflies (suborder Zygoptera) and dragonflies (suborder Anisoptera). In this protocol, we present the methods for the blue-tailed damselfly Ischnura senegalensis (Coenagrionidae) as an example of damselfly species and the pied skimmer dragonfly Pseudothemis zonata (Libellulidae) as another example of dragonfly species. As representative examples, we show the results of RNAi targeting the melanin synthesis gene multicopper oxidase 2. This RNAi method will facilitate understanding of various gene functions involved in metamorphosis, morphogenesis, color pattern formation, and other biological features of Odonata. Moreover, this protocol may be generally applicable to non-model organisms in which RNAi is less effective in gene suppression due to the inefficiency and low penetrance.

We provide a detailed protocol for electroporation-mediated RNA interference in insects of the order Odonata (dragonflies and damselflies) using the blue-tailed damselfly (Ischnura senegalensis: Coenagironidae: Zygoptera) and the pied skimmer dragonfly (Pseudothemis zonata: Libellulidae: Anisoptera).

Dragonflies and damselflies (order Odonata) represent one of the most ancestral insects with metamorphosis, in which they change their habitat, ...

Developmental Biology

Applications of RNA Interference in American Cockroach

Cockroaches, a sanitary pest, are essential species in insect developmental and metamorphic studies due to their easy feeding and hemimetabolous characteristics. Altogether with well-annotated genome sequences, these advantages have made American cockroach, Periplaneta americana, an important hemimetabolous insect model. Limited by the shortage of knockout strategy, effective RNA interference (RNAi)-based gene knockdown becomes an indispensable technique in functional gene research of P. americana. The present protocol describes the RNAi operation techniques in P. americana. The protocol includes (1) selection of the P. americana at proper developmental stages, (2) preparation for the injection setting, (3) dsRNA injection, and (4) gene knockdown efficiency detection. RNAi is a powerful reverse genetic tool in P. americana. The majority of P. americana tissues are sensitive to extracellular dsRNA. Its simplicity allows researchers to quickly obtain dysfunctional phenotypes under one or multiple targeting dsRNA injections, enabling researchers to better use the P. americana for developmental and metamorphic studies.

The present protocol describes step-by-step guidelines for the RNAi operation techniques in P. americana.

Cockroaches, a sanitary pest, are essential species in insect developmental and metamorphic studies due to their easy feeding and hemimetabolous ...

Larval RNA Interference in the Red Flour Beetle, Tribolium castaneum

The red flour beetle, Tribolium&#160;castaneum, offers a repertoire of experimental tools for genetic and developmental studies, including a fully annotated genome sequence, transposon-based transgenesis, and effective RNA interference (RNAi). Among these advantages, RNAi-based gene knockdown techniques are at the core of Tribolium research. T. castaneum show a robust systemic RNAi response, making it possible to perform RNAi at any life stage by simply injecting double-stranded RNA (dsRNA) into the beetle&#8217;s body cavity.
In this report, we provide an overview of our larval RNAi technique in T. castaneum. The protocol includes (i) isolation of the proper stage of T. castaneum larvae for injection, (ii) preparation for the injection setting, and (iii) dsRNA injection. Larval RNAi is a simple, but powerful technique that provides us with quick access to loss-of-function phenotypes, including multiple gene knockdown phenotypes as well as a series of hypomorphic phenotypes. Since virtually all T. castaneum tissues are susceptible to extracellular dsRNA, the larval RNAi technique allows researchers to study a wide variety of tissues in diverse contexts, including the genetic basis of organismal responses to the outside environment. In addition, the simplicity of this technique stimulates more student involvement in research, making T. castaneum an ideal genetic system for use in a classroom setting.

Larval RNA Interference in the Red Flour Beetle, Tribolium castaneum

RNA interference (RNAi)-based gene knockdown techniques are at the core of Tribolium research. Here, we provide an overview of our larval RNAi technique in Tribolium castaneum. Larval RNAi is a simple, but powerful technique that provides quick access to loss-of-function phenotypes, allowing researchers to study gene functions in diverse contexts.

The red flour beetle, Tribolium&#160;castaneum, offers a repertoire of experimental tools for genetic and developmental studies, including a fully ...

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory for functional genomic studies. The ability to knockdown individual genes selectively via this reverse genetic technique has allowed many researchers to rapidly uncover the biological roles of numerous genes within many organisms, by evaluation of loss-of-function phenotypes. In the major human malaria vector Anopheles gambiae, the predominant method used to reduce the function of targeted genes involves injection of double-stranded (dsRNA) into the hemocoel of the adult mosquito. While this method has been successful, gene knockdown in adults excludes the functional assessment of genes that are expressed and potentially play roles during pre-adult stages, as well as genes that are expressed in limited numbers of cells in adult mosquitoes. We describe a method for the injection of Serine Protease Inhibitor 2 (SRPN2) dsRNA during the early pupal stage and validate SRPN2 protein knockdown by observing decreased target protein levels and the formation of melanotic pseudo-tumors in SRPN2 knockdown adult mosquitoes. This evident phenotype has been described previously for adult stage knockdown of SRPN2 function, and we have recapitulated this adult phenotype by SRPN2 knockdown initiated during pupal development. When used in conjunction with a dye-labeled dsRNA solution, this technique enables easy visualization by simple light microscopy of injection quality and distribution of dsRNA in the hemocoel.

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi) is an extremely valuable tool for uncovering gene function. However, the ability to target genes using RNAi during pre-adult stages is limited in the major human malaria vector Anopheles gambiae. We describe an RNAi protocol to reduce gene function via direct injection during pupal development.

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory ...

Practical Use of RNA Interference: Oral Delivery of Double-stranded RNA in Liposome Carriers for Cockroaches

RNA interference (RNAi) has been widely applied for uncovering the biological functions of numerous genes, and has been envisaged as a pest control tool operating by disruption of essential gene expression. Although different methods, such as injection, feeding, and soaking, have been reported for successful delivery of double-stranded RNA (dsRNA), the efficiency of RNAi through oral delivery of dsRNA is highly variable among different insect groups. The German cockroach, Blattella germanica, is highly sensitive to the injection of dsRNA, as shown by many studies published previously. The present study describes a method to demonstrate that the dsRNA encapsulated with liposome carriers is sufficient to retard the degradation of dsRNA by midgut juice. Notably, the continuous feeding of dsRNA encapsulated by liposomes significantly reduces the tubulin expression in the midgut, and led to the death of cockroaches. In conclusion, the formulation and utilization of dsRNA lipoplexes, which protect dsRNA against nucleases, could be a practical use of RNAi for insect pest control in the future.

This manuscript demonstrates the depletion of gene expression in the midgut of the German cockroach through oral ingestion of double-stranded RNA encapsulated in liposomes.

RNA interference (RNAi) has been widely applied for uncovering the biological functions of numerous genes, and has been envisaged as a pest control tool ...

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) administration has been accomplished primarily via injection, which requires significant time and is not suitable for field applications. To overcome these limitations, here we present a more efficient method for robust activation of RNAi by oral delivery of dsRNA to adult Anopheles gambiae. Long dsRNAs were produced in Escherichia coli strain HT115 (DE3), and a concentrated suspension of heat-killed dsRNA-containing bacteria in 10% sucrose was offered on cotton balls ad-libitum to adult mosquitoes. Cotton balls were replaced every 2 days for the duration of the treatment. Use of this method to target doublesex (a gene involved in sex differentiation) or fork head (which encodes a salivary gland transcription factor) resulted in reduced target gene expression and/or protein immunofluorescence signal, as measured by quantitative Real-Time PCR (qRT-PCR) or fluorescence confocal microscopy, respectively. Defects in salivary gland morphology were also observed. This highly flexible, user-friendly, low-cost, time-efficient method of dsRNA delivery could be broadly applicable to target genes important for insect vector physiology and beyond.

Effective Oral RNA Interference RNAi Administration to Adult Anopheles gambiae Mosquitoes

The oral administration of dsRNA produced by bacteria, a delivery method for RNA interference (RNAi) that is routinely used in Caenorhabditis elegans, was successfully applied here to adult mosquitoes. Our method allows for robust reverse genetics studies and transmission-blocking vector studies without the use of injection.

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) ...

Advancing RNAi Techniques in Trichogramma dendrolimi for Gene Function Analysis

RNA Interference in the Egg Parasitoid, Trichogramma dendrolimi Matsumura

The egg parasitoids, Trichogramma spp, are recognized as efficient biological control agents against various lepidopteran pests in agriculture and forests. The immature stages of Trichogramma offspring develop within the host egg, exhibiting remarkable diminutiveness (approximately 0.5 mm in adult length). RNA-interference (RNAi) methodology has emerged as a crucial tool for elucidating gene functions in numerous organisms. However, manipulating RNAi in certain small parasitoid species, such as Trichogramma, has generally posed significant challenges. In this study, we present an efficient RNAi method in Trichogramma denrolimi. The outlined procedure encompasses the acquisition and isolation of individual T. dendrolimi specimens from host eggs, the design and synthesis of double-stranded RNA (dsRNA), the in vitro transplantation and cultivation of T. dendrolimi pupae, the micro-injection of dsRNA, and the subsequent assessment of target gene knockdown through RT-qPCR analysis. This study furnishes a comprehensive, visually detailed procedure for conducting RNAi experiments in T. dendrolimi, thereby enabling researchers to investigate the gene regulation in this species. Furthermore, this methodology is adaptable for RNAi studies or micro-injections in other Trichogramma species with minor adjustments, rendering it a valuable reference for conducting RNAi experiments in other endoparasitic species.

Author Spotlight: Investigating Wolbachia-Induced Thelytokous Parthenogenesis and Genetic Toolkit Development Through RNA Interference

The manipulation of RNA interference (RNAi) presents a formidable challenge in many parasitoid species with diminutive size, such as Trichogramma wasps. This study delineated an efficient RNAi method in Trichogramma denrolimi. The present methodology provides a robust model for investigating gene regulation in Trichogramma wasps.

Application of RNA Interference in the Pinewood Nematode, Bursaphelenchus xylophilus

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