This work was supported by National Natural Science Foundation of China (31725023, 31861133019 to GW, and 31171912 to CY).

1. Design of gene-specific primers and preparation of sgRNA
<ol>
	<li>Verify a conserved genomic region in the gene of interest through PCR amplification and sequencing analyses. Amplify the target gene from the genome DNA of H. armigera and distinguish the exons and introns. 
	NOTE: The sequence specificity of the guide site is necessary to avoid off-target gene editing. Search possible guide sites in the exons are close to the 5&#39; UTR of the gene. Then, it is important to make sure that the gene is completely nonfunctional. A summary of the flow path for the preparation of sgRNA is illustrated in Figure 1.</li>
	<li>Choose the sgRNA targets. Use the CRISPR online website CRISPOR (Version 4.97) (http://crispor.tefor.net/crispor.py)&#160;to search for possible guide sites in the exons close to the 5&#39; UTR of the gene. Input the exon sequence into the textbox and select the target genome to Helicoverpa armigera (Harm_1.0). Choose the protospacer adjacent motif (PAM) option of &#34;20 bp-NGG&#34; and leave the other settings on the default parameters according to the user manuals of the websites.</li>
	<li>Compare the predicted guide sequences from the software and choose the guide sequence with the highest predicted efficiency and fewest mismatches to improve the editing efficiency and reduce off-target editing. A 20 bp guide sequence containing one or two G on the 5&#39; UTR is recommended as it could increase the cutting efficiency. 
	NOTE: A pair of gRNAs across exon-regions are also recommended to obtain a large segment deletion, which simplifies mutant detection in later steps. Ensure the spacing distance between the two selected guide sequences is at least 100 bp. In this protocol, we choose the commonly used SpCas9 protein, which recognizes the NGG motif. According to the manufacturer&#39;s instruction, the guide sequence lacking G is also acceptable when choosing the T7 promoter because the promoter adds a G to the 5&#39; UTR of the sequence.</li>
	<li>Design forward and reverse oligonucleotides. Set the sequence order to 5&#39;-20 bp guide sequence-NGG-3&#39; and reverse complement the guide sequence. Add the T7 promoter sequence to the forward and reverse strand guide sequence, respectively according to the user guide of the gRNA synthesis kit. 
	NOTE: The PAM sequence NGG should be excluded from the oligonucleotide sequence.</li>
	<li>Generate the sgRNA using the gRNA synthesis kit. This process includes three steps: DNA template assembly, in vitro transcription, and purification of sgRNA (Figure 2). Perform each step in accordance with the user instructions.</li>
</ol>
2. Embryo preparation and collection
<ol>
	<li>Separate male and female pupae as described by Hongtao et al.34 and segregate them into two different net cages. After eclosion, feed them ~30 mL of 10% (w/v) white sugar solution in absorbent cotton in a Petri dish. 
	NOTE: 3 g of white sugar was dissolved in 30 mL of sterile water to prepare the 10% (w/v) white sugar solution.</li>
	<li>Select 50 healthy individuals from 3-day-old males and 2-day-old females, respectively, and mix them in a clean net cage. Place a piece of cotton containing 10% (w/v) sugar solution in the cage and keep the cotton moist. Cover the net cages with gauze and fix the gauze with a rubber band. Spray water onto the gauze to keep moist.</li>
	<li>Allow selected male and female moths in step 2 to mate completely and observe the egg-laying amount. 
	NOTE: The peak of oviposition of H. armigera appears after 9:00 p.m. Therefore, the time of the mating should be considered to ensure a sufficient number of eggs in the subsequent steps. At the peak of oviposition, replace the gauze with a black cloth and enable free oviposition for 30 min. Replace the black cloth every 30 min for collecting fresh eggs (Figure 3A).</li>
	<li>Cut the black cloth into irregularly shaped patches with a size of 3 mm approximately. Ensure more eggs on each patch. 
	NOTE: Avoid choosing wrinkled eggs as they are usually unfertilized.</li>
	<li>Paste double-sided tape onto a microscope slide (25 mm x 75 mm) (Figure 3B). Using forceps, paste the patches with eggs in a row on the surface of the double-sided tape. Press the margin of each patch to make sure they stick firmly to the tape. Collect 50-100 eggs per microscope slide (Figure 3C). 
	NOTE: The patches need to cover the full surface of the double-sided tape, otherwise the hatching larval will have difficulty crawling out.</li>
	<li>Before microinjection, keep the microscope slide on ice to delay the development of embryos.</li>
</ol>
3. Microinjection of embryos
<ol>
	<li>Prepare the needle by pulling a capillary glass using a micropipette puller (Figure 4A). Set Heat to 540, Pull to 80, Vel to 75, Time to 170, and Pressure to 450. Ground the needle tip using a micro grinder. The ideal needle shows a sharp-edged tip (Figure 4B).</li>
	<li>Preparation of injection solutions. Add 2 &#181;L of commercialized Cas9 protein (1 mg/mL) and sgRNA (300-500 ng/&#181;L final concentration) to RNase-free water in a PCR tube to obtain a 10 &#181;L volume mixture. The volume of sgRNA depends on its concentration. Mix well by pipetting and put it on ice. 
	NOTE: All the pipette tips and PCR tubes used in this step are RNase-free.</li>
	<li>Set the parameters of the electronic microinjector. Set the injection pressure (pi) to 1,500 hPa, the injection time (ti) to 0.1 s, and the compensation pressure (pc) to 30 hPa.</li>
	<li>Load 2 &#181;L of the mixture into a needle using a micro loader pipette tip. The residual air in the tip of the needle should be exhausted as much as possible.</li>
	<li>Connect the injection needle to a micromanipulator and ensure a tight connection between the two parts.</li>
	<li>Place a slide in a Petri dish (100 mm) and put them on the stage of the microscope (Figure 5A).</li>
	<li>Adjust the position of the needle tip under a light microscope until both the needle tip and embryos are visible under the microscope (Figure 5B).</li>
	<li>Adjust the volume of the droplet. Press the pedal and observe the liquid drop at the needle tip. Adjust the injection pressure of the microinjector until the volume of a liquid drop is about one-tenth of the volume of the embryos. 
	NOTE: The quality of the injection needle is vital for the survival rate of embryos.</li>
	<li>Carefully insert the needle tip into the top hemisphere of an embryo at a 45-degree angle (Figure 5C). Press the pedal to deliver the mixture into the embryo. The injection leads to a slight expansion of the embryo. Retract the needle immediately from the embryo and move the Petri dish with one hand until the next embryo is in proximity to the needle and inject the next embryo with the same procedure. 
	NOTE: The cytoplasmic outflow at the pinhole is acceptable. If the cytoplasmic leakage is too much, adjust the angle of the needle to a more severe angle until the fluid outflow is controlled.</li>
	<li>Inject at least 300 embryos to ensure a sufficient hatching amount. Cover the lid of the Petri dish(es) after injection. 
	&#8203;NOTE: The time from oviposition to injection of the 50-100 embryos is limited to 2 h. Most of the embryos are still at the one-cell stage within this time frame. In general, it is useful to repeat the oviposition procedure during embryo injection to promote efficiency.</li>
</ol>
4. Post-injection insect rearing
<ol>
	<li>Propagation of G0 embryos.
	<ol>
		<li>Incubate the embryos at 60% relative humidity and 28&#176;C in an artificial climate box with 14 h light/10 h dark.</li>
		<li>Check the development of embryos daily after injection. When the surface color of embryos has darkened, put artificial diet in the Petri dish around the microscope slide, and check the development of embryos every 12 h. 
		&#8203;NOTE: The artificial diet is prepared as described by Wu et al.35 and Jha et al.36.</li>
		<li>Prepare 24-well culture plates and fill each well to one-third of its volumetric capacity with artificial diet.</li>
		<li>Pick out hatching larvae (Figure 5D) using a small paintbrush and transfer them to the 24-well culture plate. Insert one larva per well to ensure that each larva has enough food to survive. 
		&#8203;NOTE: The larvae of H. armigera were reared individually in each well since they usually cannibalized each other.</li>
	</ol>
	</li>
	<li>G0 larvae rearing
	<ol>
		<li>Rear the larvae in the same conditions as the embryos.</li>
		<li>Check the hatched larvae every day. When larvae grow to the third instar stage, transfer each larva into a new glass dactylethrae, filling one-fifth of the volume with artificial diet.</li>
		<li>Approximately 12 d after incubation, the mature larvae should begin to pupate.</li>
		<li>The G0 mature pupae are distinguished based on sex and placed in separate cages before eclosion. The same-aged pupae of wild type are also prepared.</li>
	</ol>
	</li>
	<li>G0 adults rearing
	<ol>
		<li>Check the eclosion of both wild type and mutant pupae daily.</li>
		<li>Transfer the newly-eclosed G0 male adults and wild type female adults into a fresh net cage and make the ratio between G0 and wild type at approximately 1:1. Supply them with 10% (w/v) sugar solution dropped in cotton balls.</li>
		<li>Rear insects using the routine method above until the pupation of generation one (G1).</li>
		<li>Using a dactylethrae, transfer the newly-eclosed single pair of G1 adults into a plastic jar (13 cm x 12 cm x 12 cm) supplied with 10% (w/v) sugar solution. Cover each jar with gauze. Take about 50 pairs of G1 adults in total. 
		&#8203;NOTE: Eclosion of female moths predate that of male moths. In general, a 3-day-old male and a 2-day-old female are sexually mature and ready to mate. The newly-eclosed adults will not mate in their first light period without feeding.</li>
		<li>Collect the G1 adults in a plastic jar after they have laid eggs. Put each adult moth in a 1.5 mL centrifuge tube.</li>
	</ol>
	</li>
</ol>
5. Knock-out mutant detection
<ol>
	<li>Design a pair of primers spanning the predicted truncated site. The primers should be set at least 50 bp distance on either side (upstream and downstream) from the target site. 
	NOTE: The primers for the identification of the target sequence often cover large spans to amplify efficiently.</li>
	<li>Perform the PCR reaction using the genotyping primers with genome DNA extracted in section 1. Perform PCR cycling at 95 &#176;C for 20 s; 35 cycles of 95 &#176;C for 20 s, 55 &#176;C for 20 s, 72 &#176;C for 1 min; 72 &#176;C for 5 min, and 4 &#176;C on hold. Verify the reaction product via 1% (w/v) agarose gel. The selected pair of detection primers was confirmed by the quality of the PCR product. If the bands are evident and specific, the primers could be used for further mutant detection based on amplicon size.</li>
	<li>Screen for potential edited individuals. Remove a hind leg carefully using forceps and put each leg in a lysing matrix tube, respectively. Label the lysing matrix tube consistent with the number on the glass dactylethrae.</li>
	<li>Homogenize the hind leg using a tissue homogenizer. Set the speed to 6.0 m/s and the time to 60 s. Extract the genomic DNA of the homogenized sample using a commercial gDNA extraction kit according to the manufacturer&#39;s instructions.</li>
	<li>Amplify the gene segment using genotyping primers with the same PCR reaction conditions as described in step 2. Confirm the genotype by a gene sequencing service. Once G1 mutant genotypes (target the same site) contained in the same jar were detected, keep the G1 progeny and rename it as generation two (G2).</li>
	<li>Put G1 individuals of the same genotype in one net cage. Self-cross the G1 progenies and continue to screen using the same methods.</li>
	<li>Amplify the gene segment and confirm the genotype with the same procedure as outlined in step 2. Obtain G2 homozygous lines and maintain the knock-out mutant line.</li>
</ol>

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J., Anderson, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expansion+of+a+bitter+taste+receptor+family+in+a+polyphagous+insect+herbivore.">Expansion of a bitter taste receptor family in a polyphagous insect herbivore.</a> Science Reports. 6, 23666 (2016).</li><li>Gouin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+genomes+of+highly+polyphagous+lepidopteran+pests+(Spodoptera+frugiperda,+Noctuidae)+with+different+host-plant+ranges.">Two genomes of highly polyphagous lepidopteran pests (Spodoptera frugiperda, Noctuidae) with different host-plant ranges.</a> Scientific Reports. 7 (1), 1-12 (2017).</li><li>Wang, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+validation+of+cadherin+as+a+receptor+of+Bt+toxin+Cry1Ac+in+Helicoverpa+armigera+utilizing+the+CRISPR/Cas9+system.">Functional validation of cadherin as a receptor of Bt toxin Cry1Ac in Helicoverpa armigera utilizing the CRISPR/Cas9 system.</a> Insect Biochemistry and Molecular Biology. 76, 11-17 (2016).</li><li>Wang, J., Wang, H., Liu, S., Liu, L., Wu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9+mediated+genome+editing+of+Helicoverpa+armigera+with+mutations+of+an+ABC+transporter+gene+HaABCA2+confers+resistance+to+Bacillus+thuringiensis+Cry2A+toxins.">CRISPR/Cas9 mediated genome editing of Helicoverpa armigera with mutations of an ABC transporter gene HaABCA2 confers resistance to Bacillus thuringiensis Cry2A toxins.</a> Insect Biochemistry and Molecular biology. 87, 147 (2017).</li><li>Wang, J., Ma, H., Zhao, S., Huang, J., Wu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+redundancy+of+two+ABC+transporter+proteins+in+mediating+toxicity+of+Bacillus+thuringiensis+to+cotton+bollworm.">Functional redundancy of two ABC transporter proteins in mediating toxicity of Bacillus thuringiensis to cotton bollworm.</a> PLoS Pathogens. 16 (3), 1008427 (2020).</li><li>Jin, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dominant+point+mutation+in+a+tetraspanin+gene+associated+with+field-evolved+resistance+of+cotton+bollworm+to+transgenic+Bt+cotton.">Dominant point mutation in a tetraspanin gene associated with field-evolved resistance of cotton bollworm to transgenic Bt cotton.</a> Proceedings of the National Academy of Sciences. 115 (46), 11760-11765 (2018).</li><li>Hongtao, Z., Jianan, L., Lian, T., Yang, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sexing+of+Helicoverpa+assulta+and+Helicoverpa+armigera+pupae+based+on+machine+vision.">Sexing of Helicoverpa assulta and Helicoverpa armigera pupae based on machine vision.</a> Tobacco Sciene &amp; Technology. 53 (2), 21-26 (2019).</li><li>Wu, K., Gong, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+and+practical+artificial+diet+for+the+cotton+boll-worm.">A new and practical artificial diet for the cotton boll-worm.</a> Insect science. 4 (3), 277-282 (1997).</li><li>Jha, R. K., Chi, H., Li-Cheng, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Comparison+of+Artificial+Diet+and+Hybrid+Sweet+Corn+for+the+Rearing+of+Helicoverpa+armigera+(H&#252;bner)+(Lepidoptera:+Noctuidae)+Based+on+Life+Table+Characteristics.">A Comparison of Artificial Diet and Hybrid Sweet Corn for the Rearing of Helicoverpa armigera (H&#252;bner) (Lepidoptera: Noctuidae) Based on Life Table Characteristics.</a> Environmental Entomology. (1), 30 (2012).</li><li>Bassett, A. R., Tibbit, C., Ponting, C. P., Liu, J. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+efficient+targeted+mutagenesis+of+Drosophila+with+the+CRISPR/Cas9+system.">Highly efficient targeted mutagenesis of Drosophila with the CRISPR/Cas9 system.</a> Cell Reports. 4 (1), 220-228 (2013).</li><li>Zheng, M. -. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+non-destructive+method+of+genotyping+individual+insects+from+the+exuviate+of+final+instar+larvae,+or+puparia.">A non-destructive method of genotyping individual insects from the exuviate of final instar larvae, or puparia.</a> Chinese Journal of Applied Entomology. (2), 21 (2018).</li><li>Pashley, D. P., Hammond, A. M., Hardy, T. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reproductive+isolating+mechanisms+in+fall+armyworm+host+strains+(Lepidoptera:+Noctuidae).">Reproductive isolating mechanisms in fall armyworm host strains (Lepidoptera: Noctuidae).</a> Annals of The Entomological Society of America. 85 (4), 400-405 (1992).</li><li>Kamimura, M., Tatsuki, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diel+rhythms+of+calling+behavior+and+pheromone+production+of+oriental+tobacco+budworm+moth,+Helicoverpa+assulta(Lepidoptera:+Noctuidae).">Diel rhythms of calling behavior and pheromone production of oriental tobacco budworm moth, Helicoverpa assulta(Lepidoptera: Noctuidae).</a> Journal of Chemical Ecology. 19 (12), 2953-2963 (1993).</li><li>Edwards, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activity+rhythms+of+lepidopterous+defoliators:+ii.+Halisidota+argentata+pack.+(arctiidae),+and+nepytia+phantasmaria+stkr.+(GEOMETRIDAE).">Activity rhythms of lepidopterous defoliators: ii. Halisidota argentata pack. (arctiidae), and nepytia phantasmaria stkr. (GEOMETRIDAE).</a> Canadian Journal of Zoology. 42 (6), 939-958 (1964).</li></ol>

The authors do not have any conflicts of interest.

The application of the CRISPR/Cas9 system has provided powerful technical support for the analysis of gene function and interaction among various genes. The detailed protocol we present here demonstrates the generation of a homozygote mutant in H. armigera via CRISPR/Cas9 genome editing. This reliable procedure provides a straightforward way for directed gene mutagenesis in H. armigera.
The choice of CRISPR target sites could affect the mutagenesis efficiency<sup class="xref"...

The application of genome editing technology provides an efficient tool to achieve target-gene mutants in diverse species. The emergence of the clustered regularly interspaced short palindromic repeats (CRISPR)/associated protein 9 (Cas9) system provides a novel method to manipulate genomes1. The CRISPR/Cas9 system consists of a guide RNA (gRNA) and the Cas9 endonuclease2,3, while the gRNA can be further divided into two parts, a target complementary CRISPR RNA (crRNA) and a trans-activating crRNA (tracrRNA). The gRNA integrates with Cas9 endonuclease and forms a ribonucleoprotein (RNP). With the gRNA, Cas9 endonuclease can be directed to a specific site of the genome via base complementation. The RuvC and HNH domains of the Cas9 cleave the target site of the genome three bases before the protospacer-adjacent motif (PAM) sequence and create a double-strand break (DSB). The DNA cleavage can then be repaired through two mechanisms, non-homologous end joining (NHEJ) or homology-directed repair (HDR)4. Repair of the DSB introduces insertions or deletions as a way to inactivate the targeted gene, potentially causing a complete loss of gene function. Hence, the hereditable and specificity of the CRISPR/Cas9 system make it a robust method to characterize gene functions in vivo and analyze gene interactions5.
With numerous merits, the CRISPR/Cas9 system has been applied to various fields, including biomedicine6,7, gene therapy8,9, and agriculture10,11,12, and has been used for various biological systems including microorganisms13, plants14,15, nematodes16 , and mammals17. In invertebrates, many insect species have been subjected to CRISPR/Cas9 genome editing, such as the fruit fly Drosophila melanogaster and beyond18,19,20,21,22.
Helicoverpa armigera is one of the most destructive pests worldwide23, and damages numerous crops, including cotton, soybean, and sorghum24,25. With the development of sequencing technology, the genome of H. armigera, as well as that of a range of Lepidoptera insect species, have been sequenced completely26,27,28,29. A large number of resistance and olfactory receptor genes have been identified and characterized from these insects in recent years19,27,28,29. Some resistance-related genes have been identified in H. armigera, such as the genes encoding for cadherin30, an ATP-binding cassette transporter31,32, as well as HaTSPAN133. Knockout of these genes using CRISPR/Cas9 technology results in a high level of resistance to Bacillus thuringiensis (BT) toxin in susceptible strains. Also, Chang et al. (2017) knocked out a pheromone receptor, which validated its significant function in mating time regulation19. These reports suggest that CRISPR/Cas9 can act as an effective tool to study gene function in vivo in insect systems. However, a detailed procedure for CRISPR/Cas9 modification in insect systems remains incomplete, which limits its application range in insect functional genomics.
Here, we present a protocol for knocking out a functional gene in H. armigera using the CRISPR/Cas9 system. A detailed step-by-step protocol is provided, including the design and preparation of gene-specific primers for gRNA production, embryo collection, microinjection, insect rearing, and mutant identification. This protocol serves as a valuable reference to manipulate any functional genes in H. armigera and can be extended to other Lepidoptera species.

<table><tbody><tr><td>2kb DNA ladder</td><td>TransGen Biotech</td><td>BM101</td><td></td></tr><tr><td>Capillary Glass</td><td>World Precision Instrucments</td><td>504949</td><td>referred to as &quot;capillary glass&quot; in the protocol</td></tr><tr><td>Double Sided Tape</td><td>Minnesota Mining and Manufacturing Corporation</td><td>665</td><td></td></tr><tr><td>Eppendorf FemtoJet 4i Microinjector</td><td>Eppendorf Corporate</td><td>E5252000021</td><td></td></tr><tr><td>Eppendorf InjectMan 4 micromanipulator</td><td>Eppendorf Corporate</td><td>5192000051</td><td></td></tr><tr><td>Eppendorf Microloader Pipette Tips</td><td>Eppendorf Corporate</td><td>G2835241</td><td></td></tr><tr><td>GeneArt Precision gRNA Synthesis Kit</td><td>Thermo Fisher Scientific</td><td>A29377</td><td></td></tr><tr><td>Microscope Slide</td><td>Sail Brand</td><td>7105</td><td></td></tr><tr><td>Olympus Microscope</td><td>Olympus Corporation</td><td>SZX16</td><td></td></tr><tr><td>PrimeSTAR HS (Premix)</td><td>Takara Biomedical Technology</td><td>R040</td><td>used for mutant detection</td></tr><tr><td>Sutter Micropipette Puller</td><td>Sutter Instrument Company</td><td>P-1000</td><td></td></tr><tr><td>TIANamp Genomic DNA Kit</td><td>TIANGEN Corporate</td><td>DP304-03</td><td></td></tr><tr><td>TrueCut Cas9 Protein v2</td><td>Thermo Fisher Scientific</td><td>A36499</td><td></td></tr></tbody></table>

embryo microinjection and knockout mutant identification of crispr/cas9 genome-edited helicoverpa armigera (h&#252;bner)

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional genomics, and behavioral studies has made H. armigera a model species in Lepidoptera Noctuidae. To study the in vivo functions of and interactions between different genes, clustered regularly interspaced short palindromic repeats (CRISPR)/ associated protein 9 (Cas9) genome editing technology is a convenient and effective method used for performing functional genomic studies. In this study, we provide a step-by-step systematic method to complete gene knockout in H. armigera using the CRISPR/Cas9 system. The design and synthesis of guide RNA (gRNA) are described in detail. Then, the subsequent steps consisting of gene-specific primer design for guide RNA (gRNA) creation, embryo collection, microinjection, insect rearing, and mutant detection are summarized. Finally, troubleshooting advice and notes are provided to improve the efficiency of gene editing. Our method will serve as a reference for the application of CRISPR/Cas9 genome editing in H. armigera as well as other Lepidopteran moths.

This protocol provides detailed steps for obtaining gene knock-out lines of H. armigera using CRISPR/Cas9 technology. The representative results obtained by this protocol are summarized for gDNA selection, embryo collection and injection, insect rearing, and mutant detection.
In this study, the target site of our gene of interest was located in its second exon (Figure 2A). This site was hig...

Watch this Scientific Journal Video about Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera HÃ¼bner at JoVE.com

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera H&#252;bner

Presented here is a protocol of Helicoverpa armigera (H&#252;bner) embryo microinjection and knockout mutant identification created by CRISPR/Cas9 genome editing. Mutant insects enable further research of gene function and interaction among different genes in vivo.

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (H&#252;bner)

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional ...

embryo-microinjection-knockout-mutant-identification-crisprcas9

Research

JoVE Journal

Behavior

4.3K Views.  China University of Mining and Technology. Katoenbolworm is een van de meest destructieve plagen ter wereld. Genoombewerking met de CRISPR/Cas9-technologie maakt verder onderzoek naar genfunctie en interactie tussen verschillende genen in dit organisme mogelijk. De twee belangrijkste voordelen van deze techniek zijn dat het een hoge specificiteit heeft en herbewerkbaar is.Om de sgRNA-doelen te kiezen, gaat u naar de crispr online website om te zoeken naar mogelijke gidsplaatsen en de exonen in de buurt van de vijf belangrijkste...

Video: Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera Hübner

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Genetics

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, Eastern equine encephalitis, lymphatic filariasis, and Saint Louis encephalitis. Notably, avian malaria has played a major role in the extinction of numerous endemic island bird species, while WNV has become an important vector-borne disease in the United States. To gain further insight into C. quinquefasciatus biology and expand their genetic control toolbox, we need to develop more efficient and affordable methods for genome engineering in this species. However, some biological traits unique to Culex mosquitoes, particularly their egg rafts, have made it difficult to perform microinjection procedures required for genome engineering. To address these challenges, we have developed an optimized embryo microinjection protocol that focuses on mitigating the technical obstacles associated with the unique characteristics of Culex mosquitoes. These procedures demonstrate optimized methods for egg collection, egg raft separation and other handling procedures essential for successful microinjection in C. quinquefasciatus. When coupled with the CRISPR/Cas9 genome editing technology, these procedures allow us to achieve site-specific, efficient and heritable germline mutations, which are required to perform advanced genome engineering and develop genetic control technologies in this important, but currently understudied, disease vector.

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

This protocol describes microinjection procedures for Culex quinquefasciatus embryos that are optimized to work with CRISPR/Cas9 gene editing tools. This technique can efficiently generate site-specific, heritable, germline mutations that can be used for building genetic technologies in this understudied disease vector.

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, ...

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to visceral pathology. Deciphering the nature of the vector/parasite interactions is of primary importance for better understanding of Leishmania transmission to their hosts. Among the parameters controlling the sand fly vector competence (i.e. their ability to carry and transmit pathogens), parameters intrinsic to these insects were shown to play a key role. Insect immune response, for example, impacts sand fly vector competence to Leishmania. The study of such parameters has been limited by the lack of methods of gene expression modification adapted for use in these non-model organisms. Gene downregulation by small interfering RNA (siRNA) is possible, but in addition to being technically challenging, the silencing leads to only a partial loss of function, which cannot be transmitted from generation to generation. Targeted mutagenesis by CRISPR/Cas9 technology was recently adapted to the Phlebotomus papatasi sand fly. This technique leads to the generation of transmissible mutations in a specifically chosen locus, allowing to study the genes of interest. The CRISPR/Cas9 system relies on the induction of targeted double-strand DNA breaks, later repaired by either Non-Homologous End Joining (NHEJ) or by Homology Driven Repair (HDR). NHEJ consists of a simple closure of the break and frequently leads to small insertion/deletion events. In contrast, HDR uses the presence of a donor DNA molecule sharing homology with the target DNA as a template for repair. Here, we present a sand fly embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 using NHEJ, which is the only genome modification technique adapted to sand fly vectors to date.

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

This protocol details the steps of CRISPR/Cas9 targeted mutagenesis in sand flies: embryo collection, injection, insect rearing, and identification as well as selection of mutations of interest.

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to ...

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how genes regulate social traits, we have developed mutant ants via delivery of CRISPR complex into young embryos during their syncytial stage. Here, we provide a protocol of CRISPR-mediated mutagenesis in Harpegnathos saltator, a ponerine ant species that displays striking phenotypic plasticity. H. saltator ants are readily reared in a laboratory setting. Embryos are collected for microinjection with Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs) using home-made quartz needles. Post-injection embryos are reared outside the colony. Following emergence of the first larva, all embryos and larvae are transported to a nest box with a few nursing workers for further development. This protocol is suitable for inducing mutagenesis for analysis of caste-specific physiology and social behavior in ants, but may also be applied to a broader spectrum of hymenopterans and other insects.

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Many characteristics of insect eusociality rely on within-colony communication and division of labor. Genetic manipulation of key regulatory genes in ant embryos via microinjection and CRISPR-mediated mutagenesis provides insights into the nature of altruistic behavior in eusocial insects.

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how ...

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

The firebrat Thermobia domestica is an ametabolous, wingless species that is suitable for studying the developmental mechanisms of insects that led to their successful evolutionary radiation on the earth. The application of genetic tools such as genome editing is the key to understanding genetic changes that are responsible for evolutionary transitions in an Evo-Devo approach. In this article, we describe our current protocol for generating and maintaining mutant strains of T. domestica. We report a dry injection method, as an alternative to the reported wet injection method, that allows us to obtain stably high survival rates in injected embryos. We also report an optimized environment setting to mate adults and obtain subsequent generations with high efficiency. Our method underlines the importance of taking each species&#8217; unique biology into account for the successful application of genome editing methods to non-traditional model organisms. We predict that these genome editing protocols will help in implementing T. domestica as a laboratory model and to further accelerate the development and application of useful genetic tools in this species.

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

We provide a detailed protocol for rearing, microinjection of eggs and for efficient mating of the firebrat Thermobia domestica to generate and maintain mutant strains after genome editing.

The firebrat Thermobia domestica is an ametabolous, wingless species that is suitable for studying the developmental mechanisms of insects that led to ...

Developmental Biology

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although RNA interference (RNAi) has proved to be a powerful tool for studying WCR biology, it has its limitations. Specifically, RNAi itself is transient (i.e. does not result in long-term Mendelian inheritance of the associated phenotype), and it requires knowing the DNA sequence of the target gene. The latter can be limiting if the phenotype of interest is controlled by poorly conserved, or even novel genes, because identifying useful targets would be challenging, if not impossible. Therefore, the number of tools in WCR&#39;s genomic toolbox should be expanded by the development of methods that could be used to create stable mutant strains and enable sequence-independent surveys of the WCR genome. Herein, we detail the methods used to collect and microinject precellular WCR embryos with nucleic acids. While the protocols described herein are aimed at the creation of transgenic WCR, CRISPR/Cas9-genome editing could also be performed using the same protocols, with the only difference being the composition of the solution injected into the embryos.

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

Here we present protocols for collecting and microinjecting precellular western corn rootworm embryos for the purpose of performing functional-genomic assays such as germline transformation and CRISPR/Cas9-genome editing.

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although ...

Bioengineering

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

The corn planthopper, Peregrinus maidis, is&#160;a pest of maize and a vector of several maize viruses. Previously published methods describe the triggering of RNA interference (RNAi) in P. maidis through microinjection of double-stranded RNAs (dsRNAs) into nymphs and adults. Despite the power of RNAi, phenotypes generated via this technique are transient and lack long-term Mendelian inheritance. Therefore, the P. maidis toolbox needs to be expanded to include functional genomic tools that would enable the production of stable mutant strains, opening the door for researchers to bring new control methods to bear on this economically important pest. However, unlike the dsRNAs used for RNAi, the components used in CRISPR/Cas9-based genome editing and germline transformation do not easily cross cell membranes. As a result, plasmid DNAs, RNAs, and/or proteins must be microinjected into embryos before the embryo cellularizes, making the timing of injection a critical factor for success. To that end, an agarose-based egg-lay method was developed to allow embryos to be harvested from P. maidis females at relatively short intervals. Herein are provided detailed protocols for collecting and microinjecting precellular P. maidis embryos with CRISPR components (Cas9 nuclease that has been complexed with guide RNAs), and results of Cas9-based gene knockout of a P. maidis eye-color gene, white, are presented. Although these protocols describe CRISPR/Cas9-genome editing in P. maidis, they can also be used for producing transgenic P. maidis via germline transformation by simply changing the composition of the injection solution.

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

Herein are protocols for collecting and microinjecting precellular corn planthopper embryos for the purpose of modifying their genome via CRISPR/Cas9-based genome editing or for the addition of marked transposable elements through germline transformation.

The corn planthopper, Peregrinus maidis, is&#160;a pest of maize and a vector of several maize viruses. Previously published methods describe the ...

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by filarial nematodes such as canine heartworm and elephantasis. Recently, CRISPR/Cas9 genome editing has been used to induce site-directed mutations by injecting a Cas9 protein that has been complexed with a guide RNA (gRNA) into freshly laid embryos of several insect species, including mosquitoes that belong to the genera Anopheles and Aedes. Manipulating and injecting Culex mosquitoes is slightly more difficult as these mosquitoes lay their eggs upright in rafts rather than individually like other species of mosquitoes. Here we describe how to design gRNAs, complex them with Cas9 protein, induce female mosquitoes of Culex pipiens to lay eggs, and how to prepare and inject newly laid embryos for microinjection with Cas9/gRNA. We also describe how to rear and screen injected mosquitoes for the desired mutation. The representative results demonstrate that this technique can be used to induce site-directed mutations in the genome of Culex mosquitoes and, with slight modifications, can be used to generate null-mutants in other mosquito species as well.&#160;

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by ...

Biology

Construction of Homozygous Mutants of Migratory Locust Using CRISPR/Cas9 Technology

The migratory locust, Locusta migratoria, is not only one of the worldwide plague locusts that caused huge economic losses to human beings but also an important research model for insect metamorphosis. The CRISPR/Cas9 system can accurately locate at a specific DNA locus and cleave within the target site, efficiently introducing double-strand breaks to induce target gene knockout or integrate new gene fragments into the specific locus. CRISPR/Cas9-mediated genome editing is a powerful tool for addressing questions encountered in locust research as well as a promising technology for locust control. This study provides a systematic protocol for CRISPR/Cas9-mediated gene knockout with the complex of Cas9 protein and single guide RNAs (sgRNAs) in migratory locusts. The selection of target sites and design of sgRNA are described in detail, followed by in vitro synthesis and verification of the sgRNAs. Subsequent procedures include egg raft collection and tanned-egg separation to achieve successful microinjection with low mortality rate, egg culture,&#160;preliminary estimation of the mutation rate, locust breeding as well as detection, preservation, and passage of the mutants to ensure population stability of the edited locusts. This method can be used as a reference for CRISPR/Cas9 based gene editing applications in migratory locusts as well as in other insects.

This study provides a systematically optimized procedure of CRISPR/Cas9 ribonuclease-based construction of homozygous locust mutants as well as a detailed method for cryopreservation and resuscitation of the locust eggs.

The migratory locust, Locusta migratoria, is not only one of the worldwide plague locusts that caused huge economic losses to human beings but also an ...

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops worldwide. Since adult fruit flies have great flight capacity and females lay their eggs under the skins of fruit, insecticides requiring direct contact with the pest usually perform poorly in the field. With the development of molecular biological tools and high-throughput sequencing technology, many scientists are attempting to develop environmentally friendly pest management strategies. These include RNAi or gene editing-based pesticides that downregulate or silence genes (molecular targets), such as olfactory genes involved in searching behavior, in various insect pests. To adapt these strategies for Oriental fruit fly control, effective methods for functional gene research are needed. Genes with critical functions in the survival and reproduction of B. dorsalis serve as good molecular targets for gene knockdown and/or silencing. The CRISPR/Cas9 system is a reliable technique used for gene editing, especially in insects. This paper presents a systematic method for CRISPR/Cas9 mutagenesis of B. dorsalis, including the design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. These protocols will serve as a useful guide for generating mutant flies for researchers interested in functional gene studies in B. dorsalis.

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

This paper presents the step-by-step protocols for CRISPR/Cas9 mutagenesis of the Oriental fruit fly Bactrocera dorsalis. Detailed steps provided by this standardized protocol will serve as a useful guide for generating mutant flies for functional gene studies in B. dorsalis.

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops ...

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (Hübner)

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