This work was supported by Shenzhen Science and Technology Program (Grant No. KQTD20180411143628272) and special funds for science technology innovation and industrial development of Shenzhen Dapeng New District (Grant No. PT202101-02).

1. Target design and in vitro synthesis of sgRNA
<ol>
	<li>Predict the structure of target genes of interest and determine the boundaries between exons and introns via bioinformatic analysis of the B. dorsalis genome (software applications used here are listed in the Table of Materials). 
	NOTE: BLAT20 was used to search potential gene loci in the genome. The high-quality RNA-seq reads (transcriptome) were aligned to the acquired gene loci using Hisat221. Samtools22 was used to generate the sorted bam files. The sorted bam files were input to Stringtie223 to provide the assemble transcripts. The assemble transcripts and gene loci information were combined by Transdecoder24. The results acquired from Transdecoder were visualized in IGV tools25 and the boundaries between exons and introns could be determined.</li>
	<li>Identify the suitable target regions within the candidate target gene site. The total length must be less than 750 bp for more convenient sequencing (Figure 1B). Design specific primers to amplify the target area from wild-type genomic DNA by PCR (Figure 1B) (Primers: F-primer: AACATTGAATATCTGGAATCAGGTAAACT, R-primer: CCTCATTGTTGATTAATTCCGACTTC). Clone the PCR products into a blunt end-vector26 and select 20 individual bacterial colonies for sequencing to determine how conserved the target region is in the laboratory insect populations.</li>
	<li>A typical target site contains a three-nucleotide sequence motif (NGG or CCN) and a 20 bp sequence adjacent to the NGG or CCN motifs (20 bp-NGG or CCN-20 bp). Blast the candidate target sites against the B. dorsalis genome and make sure the predicted efficiency is high enough and the off-targeting rate is low; several open-source software programs can automatically predict this. In this protocol, sgRNAcas9 -AI27 is used to select and evaluate the optimal targets. Details of use can be found in the manual for this software. 
	NOTE: Possible target sites closed to the 5&#39; UTR of the target gene and 1-2 Gs at the start of the 20 bp sequence are favored. In order to achieve a large deletion which will be identified by PCR followed by agarose gel electrophoresis, designing two targets28 separated by over 100 bp is recommended (Figure 1C).</li>
	<li>Use the commercially available gRNA synthesis kit to generate the designed sgRNA. Perform each step following the user&#39;s guide. Resuspend the gRNA product in nuclease-free water, quantify the concentration using a UV-Vis spectrophotometer, and store at -80 &#176;C prior to use (the concentration of successfully synthesized single gRNA [10 &#181;L] with the kit used here is about 4000-6000 ng/&#181;L, or even higher). 
	&#8203;NOTE: Although generating the mutant flies is the first step for every researcher, the appropriate negative control for downstream experiment/analysis is critical. Generating these controls alongside the mutants saves researchers time and effort. For example, set scrambled sgRNA as a negative control.</li>
</ol>
2. Embryo collection and preparation
<ol>
	<li>Place B. dorsalis pupae&#160;into plastic cages. Provide a mixture of sugar and yeast (1:1) as food along with a water source after the adults eclose (Figure 2A, B). The rearing conditions are 55% relative humidity (RH), 26.5 &#176;C, and a 14:10 L/D cycle(lights on at six in the morning, lights off at eight in the evening).</li>
	<li>Most adults reach sexual maturity 10 days after emergence. Provide a suitable environment to help adult flies mate as much as possible. Ideally, use a light stand with a light of 30-50 lux. This can improve the fecundity of females, therefore, improving the efficiency of embryo collection. 
	NOTE: Putting the adults (aged 5-6 days after emergence) in a dimly lit (&#60;100 lux) environment can promote mating. Generally, the peak of oviposition happens at 03:00 p.m. (14:10/L:D, lights on 06:00 a.m., lights off at 08:00 p.m.); to obtain enough embryos, placing oviposition chambers in the cages 30 min before 03:00 p.m. is recommended.</li>
	<li>Place a 200-mesh gauze in the oviposition chamber, 1-2 mm away from the chamber lid. This will help with obtaining as many embryos as possible. 
	NOTE: Avoid letting the embryos get soaked in orange juice or rubbed with gauze, as this can significantly decrease the embryos&#39; survival rate.</li>
	<li>Put a new oviposition chamber into the cage when the microinjection setup is ready. Collect the embryos every 10 min using a fine wet brush, then line them up on a self-made injection plate (plexiglass with a length of 55 mm, a width of 13.75 mm, and a height of 5 mm, A 45 mm x 5 mm x 0.3 mm shallow groove is opened in the middle to facilitate egg placement). If the embryos have high internal pressure, slight desiccation at &#60;10% RH for 10 min is optional. Dip the embryos into halocarbon oil during injection to avoid further desiccation (Figure 2C).</li>
</ol>
3. Microinjection of the embryo
<ol>
	<li>Prepare the glass injection needle using a micropipette puller. 
	NOTE: Setting the parameters following the user guide is important. In these protocols, different needle shapes can be made using the parameters suggested by the manual of the micropipette puller. The glass capillary must be as clean as possible to prevent dust from clogging the needle.</li>
	<li>Prepare the working solution. Mix the Cas9 protein and the corresponding sgRNA to the following working concentrations:&#160;sgRNA, 300 ng/&#181;L; Cas9 protein, 150 ng/&#181;L. Add 1 &#181;L of phenol red to the mixture to serve as a convenient way to mark injected embryos. Place the prepared mixture on ice to avoid degradation of the sgRNA. 
	NOTE: Use nuclease-free pipette tips and PCR tubes. The concentration of Cas9 protein needs to be less than or equal to 150 ng/&#181;L; higher concentrations are toxic and significantly decrease the embryo survival rate. Cas9 could also be delivered in DNA or mRNA format to achieve successful gene editing17,19. In this protocol, Cas9 protein is recommended since mRNAs are susceptible to degradation, and the expression of Cas9 protein from plasmid DNA takes time for transcription and translation.</li>
	<li>Set the parameters for the injector. The initial program is Pi-500 hPa, Ti-0.5 s, and PC-200 hPa. Adjust these parameters further as needed during microinjection.</li>
	<li>Add 3 &#181;L of the mixture into the injection needle. Avoid introducing air bubbles that can possibly clog the needle. Open the needle using a Microgrinder.</li>
	<li>Connect the needle to the micromanipulator according to the user guide (Figure 2E).</li>
	<li>Put the plate with lined-up embryos on the objective table. Adjust the position of the micropipette under a fine optical microscope, setting the micropipette and embryo on the same plane. Adjust the droplet volume by pressing the pedal; 1/10 the volume of embryos is recommended.</li>
	<li>Insert the needle tip into the posterior (vegetal pole) of the embryo. Deliver the mixtures into the embryo by pressing the pedal from the injector. If phenol red is added, a slightly reddish color is observed instantly. 
	&#8203;NOTE: A small volume of cytoplasmic backflow from the pinhole will not decrease the survival rate of the embryo. Injection of 200 embryos is enough for successful mutagenesis of one gene. Adding more halocarbon oil to immerse the embryos can prevent further desiccation. The injection needs to be performed before pole cell formation, which ensures every mutation can be efficiently inherited.</li>
</ol>
4. Post-injection insect rearing
<ol>
	<li>Put the injection plate with the injected embryos into an artificial climate chamber kept at 55% RH, 26.5 &#176;C, and a 14:10 L/D cycle(lights on at six in the morning, lights off at eight in the evening).</li>
	<li>After injection, the embryo formation usually takes 24 h to complete and 36 h to hatch. Use a fine-tip brush to transfer the larvae onto a prepared larval diet. Collect larvae three or four times daily until no more larvae hatch. Generally, larvae take 2-3 days to finish hatching and pupate within 7 days. 
	&#8203;NOTE: A maize-based diet was used to feed the larvae. The recipe contains 150 g of corn flour, 150 g of banana, 0.6 g of sodium benzoate, 30 g of yeast, 30 g of sucrose, 30 g of paper towel, 1.2 mL of hydrochloric acid, and 300 mL of water29. Minimize the time larvae are left in oil (&#60;2 h). Move the larvae onto the diet immediately after hatching as far as possible. This can significantly increase the survival and pupation rates. Do not provide too much diet (a diet with 2/3 of a 90 mm Petri dish is enough for 30 larvae); otherwise, it will become moldy and decrease the larval survival rate.</li>
	<li>Put the mature larvae into the wet sand to pupate. The pupation stage takes about 10 days. Move the pupae to plastic cages before eclosion begins.</li>
</ol>
5. Mutant screening
<ol>
	<li>A flow chart of the mutant screening is illustrated in Figure 3A. Cross G0 adults obtained by microinjection with wild-type adults to obtain heterozygous lines. Verify the type of mutations the offspring (G1) carry by genotyping.</li>
	<li>Self-cross G1 with the same mutant genotype to obtain homozygous lines in G2. If genotypes of the mutants in G1 are entirely different, cross the heterozygotes with wild-type flies. Homozygous lines are usually obtained in G3. Maintain two or three homozygous lines for subsequent phenotyping experiments.</li>
	<li>Use the genomic DNA from a fresh puparium or a single mid-leg of individual adults to perform genotyping. Design specific primers to amplify the target area; the primers must set at least 50 bps upstream and downstream of the target site. Refer to step 1.2 for the primer details. 
	NOTE: Since the amount of the DNA in a puparium is extremely low, commercially available DNA extraction kits are recommended.</li>
	<li>Perform PCR using the following cycling conditions: 98 &#176;C for 3 min, followed by 35 cycles of 98 &#176;C for 10 s, 15 s at an annealing temperature appropriate for the designed primers, 72 &#176;C for 35 s, followed by a final extension of 72 &#176;C for 10 min.</li>
	<li>Sequence the purified PCR products. When multiple overlapping peaks adjacent to the target site are detected (Figure 3B), successful mutagenesis has occurred. Sub-clone the purified PCR products into a blunt-end vector and select 10 individual bacterial colonies for sequencing to verify the genotype of the mutants. Maintain the lines with frameshift mutations and premature translation terminations (Figure 3C). 
	NOTE: There is no need to perform single clone sequencing to verify the genotype of G0 individuals. Just sequence the PCR products and maintain the individuals with obvious multiple overlap peaks adjacent to the target site. After the initial injection, selecting 10 embryos to extract genomic DNA and sequencing the PCR products based on the standards in step 5.3 is recommended. This can help to predict mutation rates in advance. In this study, sanger sequencing to determine the genotype was done by a sequencing company.</li>
</ol>

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H., 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=Knockout+of+juvenile+hormone+receptor,+Methoprene-tolerant,+induces+black+larval+phenotype+in+the+yellow+fever+mosquito,+Aedes+aegypti.">Knockout of juvenile hormone receptor, Methoprene-tolerant, induces black larval phenotype in the yellow fever mosquito, Aedes aegypti.</a> Proceedings of the National Academy of Sciences. 116 (43), 21501-21507 (2019).</li><li>Laohakieat, K., Isasawin, S., Thanaphum, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+transformer-2+and+fruitless+characterisation+with+developmental+expression+profiles+of+sex-determining+genes+in+Bactrocera+dorsalis+and+B.+correcta.">The transformer-2 and fruitless characterisation with developmental expression profiles of sex-determining genes in Bactrocera dorsalis and B. correcta.</a> Scientific Reports. 10 (1), 1-13 (2020).</li><li>Gabrieli, P., 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=Sperm-less+males+modulate+female+behaviour+in+Ceratitis+capitata+(Diptera:+Tephritidae).">Sperm-less males modulate female behaviour in Ceratitis capitata (Diptera: Tephritidae).</a> Insect Biochemistry and Molecular Biology. 79, 13-26 (2016).</li></ol>

The authors do not have any conflicts of interest.

The CRISPR/Cas9 system is the most widely used gene editing tool and has various applications, such as gene threpy30, crop breeding31, and basic studies of gene fuctions32. This system has already been applied for gene editing in various insect species and has served as an effective tool for functional gene studies in pests. The protocols we present here standardize the procedure of design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. Moreover, a troubleshooting table was added to summarize the potential problem from each step, and their solutions were provided to lighten the workload and improve efficiency (Table 2). This procedure provides a reliable way to edit genes of interest and improves the functional genomic studies in B. dorsalis.
First, target gene selection is critical to increasing the efficiency of the CRISPR/Cas9 system, and several open source software programs such as ChopChop33,34 (http://chopchop.cbu.uib.no/), sgRNAcas9 (V3.0)35, and CRISPR optimal target finder36 (http://targetfinder.flycrispr.neuro.brown.edu) can automatically generate targets and predict potential off-target rates. Since the high fecundity rate of&#160;B. dorsalis facilitates embryo collection, mutant rates can be predicted by sacrificing a portion of the embryos for DNA extraction and sequencing of the target region. If sequencing is not available in the lab, using T7 endonuclease I to predict mutation rates is also recommended37. Target sites with high genomic deletion rates can be selected through these three methods, and therefore, the efficiency of gene editing in B. dorsalis could be increased.
The development stage of the embryo determines whether mutations will be inherited. In order to obtain inherited mutations, gene editing must occur in germ cells. Generally, the mixture of sgRNA and Cas9 cannot be delivered into germ cells after pole cell formation has occurred. In B. dorsalis, the pole cell formation occurred at 3 h after egg laying38, whereas the protocol detailed here for B. dorsalis takes 30 min from embryo collection to the end of microinjection. During injection, nearly no pole cells are formed in the vegetal end of the embryo; therefore, gene mutations obtained in the G0 should be efficiently inherited. The time consumed by microinjection is also less than the method mentioned in previously published papers (generally 1-3 h)18,19.
It is important to minimize mechanical or chemical damage during the process of embryo collection and handling. Use a fine brush to gently line up embryos on the injection plate. The position of injection should reflect the work previously done in Drosophila (Methods - Nicolas Gompel&#39;s lab, http://gompel.org/methods) and&#160;Ceratitis capitata39. Inject embryos at the vegetal pole; never insert the needle into the animal pole. Prepare the needle following the micropipette puller guide; the opening of the needle should be as small as possible to avoid excessive cytoplasmic backflow. The method mentioned in published studies in B. dorsalis generally&#160;dechorionated the embryos with sodium hypochorite17,18,19; this could cause chemical damage and decrease the survival rates of the embryos. In this protocol, embryos are not dechorionated, and injection still works well. The chemical damage to the embryos could be minimal.
Post-injection insect rearing is very important. The standardized rearing protocol provided here can serve as a reference for rearing other fruit fly species, especially Bactrocera species. The embryos can be immersed in oil to avoid desiccation during injection using our self-made injection plate. Minimize the time larva spend in the oil (&#60;2 h) to achieve survival rates in the G0 above 50%.
A non-invasive method of genomic DNA extraction is recommended for mutant detection. For example, some lepidopteran researchers suggest that the exuviates of the final instar larvae could be used to extract genomic DNA for further genotyping. For B. dorsalis, one non-invasive method involves extracting genomic DNA from a fresh puparium. Injection of multiple sgRNAs targeting a marker gene and the gene of interest could also improve the identification of mutants. For example, co-injected sgRNAs targeting eye color (kmo) and juvenile hormone receptor (Met) can produce 75% of offspring with double mutations in mosquitos. Met mutants were preliminarily screened based on the eye color of larvae37. This should be evaluated in B. dorsalis in the future to further improve the efficacy of gene editing in this insect species by using the CRISPR/Cas9 system.
In conclusion, the CRSIPR/Cas9 system is a powerful tool in functional genomics in B. dorsalis. Our detailed protocols provide useful information to help researchers to achieve efficient embryo collection, ideal survival rates of larvae, and the desired editing efficiency. This could be a simple and quick way to help researchers to obtain mutagenesis in B. dorsalis. This technique could not be applied in the functional studies of lethal genes since the inherited mutations need successful cross-breedings. Future studies could focus on developing transgenic tools to express stage or tissue-specific CRISPR elements to break through this limitation.

The Oriental fruit fly, Bactrocera dorsalis, is a cosmopolitan insect pest species that causes damage to over 150 species of fruit crops, including guava, mango, Eugenia spp., Surinam cherry, citrus, loquat, and papaya1. The damage caused in Guangdong Province (China) alone is estimated at over 200 million yuans. Adult females insert their eggs beneath the skin of ripening or ripened fruits, causing decay and abscission of the fruit, which decreases fruit quality and overall yield of the crop2. Since adult fruit flies have great flight capacity and their larvae bore into the fruit skin, insecticides requiring direct contact with the pest perform poorly in the field. Additionally, the extensive use of insecticides has increased the resistance of B. dorsalis against various agricultural chemicals, making control of these damaging pests even more difficult3. Therefore, the development of effective and environmentally friendly pest management strategies is desperately needed.
Recently, with the development of molecular biological tools and high-throughput sequencing technologies, scientists are attempting to develop environmentally-friendly pest management strategies, such as RNAi, that target the functionality of important genes (molecular targets) of various insect pests. Genes that are critical to the survival and reproduction of the pest can be identified through functional gene studies and further serve as potential molecular targets for the improvement of specifically targeted and environmentally friendly pest management tools4. To adapt such strategies to Oriental fruit fly control, effective methods for functional gene research are needed.
The CRISPR/Cas (clustered regularly interspaced short palindromic repeats/CRISPR-associated) endonuclease system was initially discovered in bacteria and archaea and found to be an adaptive mechanism involved in the recognition and degradation of foreign intracellular DNA, such as that introduced by infecting bacteriophages5. In the type II CRISPR system, Cas9 endonuclease is guided by small associated RNAs (crRNA and tracrRNA) to cleave trespassing DNA6,7,8 and has become one of the most widely used tools for gene-editing to date9,10,11,12. Since the CRISPR/Cas9 system has several advantages, such as high efficiency of gene silencing and low cost, it has already been applied for gene editing in various insect species, including Aedes aegypti13,14, Locusta migratoria15, and Bombyx mori16. In B. dorsalis, genes related to body color, wing dimorphism, and sex determination have been successfully knocked out using CRISPR/Cas917,18,19. However, detailed procedures for CRISPR/Cas9 application in this insect remain incomplete. Moreover, some steps provided by researchers for B. dorsalis gene editing are also varied and in need of standardization. For example, the forms of Cas9 were different in published references17,18,19.
This paper provides a systematic method for mutagenesis of B. dorsalis using the CRISPR/Cas9 system, including the design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. This protocol will serve as a useful guide for generating mutant flies for researchers who are interested in the functional gene studies in B. dorsalis.

<table><tbody><tr><td>6x DNA Loading Buffer</td><td>TransGen Biotech</td><td>GH101-01</td><td></td></tr><tr><td>Artificial climate chamber</td><td>ShangHai BluePard</td><td>MGC-350P</td><td></td></tr><tr><td>AxyPrep Genomic DNA Mini-Extraction Kit</td><td>Axygen</td><td>AP-MN-MS-GDNA-250G</td><td></td></tr><tr><td>BLAT</td><td>NA</td><td>NA</td><td>For searching potential gene loci in the genome</td></tr><tr><td>Capillary Glass</td><td>WPI&amp;nbsp;</td><td>1B100F-4</td><td></td></tr><tr><td>Eppendorf InjectMan 4 micromanipulator</td><td>Eppendorf</td><td>InjectMan 4</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>Hisat2</td><td>NA</td><td>NA</td><td>For aligning the transcriptome to the acquired gene loci</td></tr><tr><td>IGV</td><td>NA</td><td>NA</td><td>For visualizing the results from Transdecoder</td></tr><tr><td>Microgrinder</td><td>NARISHIGE</td><td>EG-401</td><td></td></tr><tr><td>Olympus Microscope</td><td>Olympus Corporation</td><td>SZ2-ILST</td><td></td></tr><tr><td>pEASY-Blunt Cloning Kit</td><td>TransGen Biotech</td><td>CB101-02</td><td>https://www.transgenbiotech.com/data/upload/pdf/CB101_2022-07-14.pdf</td></tr><tr><td>Phenol red solution</td><td>Sigma-Aldrich</td><td>P0290-100ML</td><td></td></tr><tr><td>Pipette cookbook 2018 P-97 &amp;amp; P-1000 Micropipette Pullers</td><td>Instrument Company&amp;nbsp;</td><td></td><td>https://www.sutter.com/PDFs/cookbook.pdf</td></tr><tr><td>PrimeSTAR HS (Premix)</td><td>Takara Biomedical Technology</td><td>R040A</td><td></td></tr><tr><td>SAMtools</td><td>NA</td><td>NA</td><td>For generating the sorted bam files</td></tr><tr><td>sgRNAcas9-AI</td><td>NA</td><td>NA</td><td>sgRNA design&lt;br/&gt; http://123.57.239.141:8080/home</td></tr><tr><td>Sutter Micropipette Puller Sutter&amp;nbsp;</td><td>Instrument Company&amp;nbsp;</td><td>P-97</td><td></td></tr><tr><td>Trans2K DNA Marker</td><td>TransGen Biotech</td><td>BM101-02</td><td></td></tr><tr><td>Transdecoder</td><td>NA</td><td>NA</td><td>For combining the results of assemble transcripts and gene loci information&lt;br/&gt; https://github.com/TransDecoder/TransDecoder/releases/tag/TransDecoder-v5.5.0</td></tr><tr><td>TrueCut Cas9 Protein v2</td><td>Thermo Fisher Scientific</td><td>A36498</td><td></td></tr><tr><td>Ultra-trace biological detector</td><td>Thermo Fisher Scientific</td><td>Nanodrop 2000C</td><td></td></tr></tbody></table>

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.

This protocol presents detailed steps for the development of B. dorsalis mutants using CRISPR/Cas9 technology, including representative results from gDNA selection, collecting embryos and microinjection, insect maintenance, and mutant screening.
The example of the target site of the selected gene is located in the third exon (Figure 1C). This site is highly conserved, and a single band was detected by gel electrophoresis for the DNA template for synthetic gRNA (Figure 1D) and gRNA obtained by in vitro transcription (Figure 1E).
Injection into 200 freshly harvested eggs was performed as described in section 3. The embryos were maintained by following the protocols described in steps 4.1-4.3. As detected by sequencing the PCR products, 80% of the G0 individuals are mosaic mutants (Table 1). Here, mutants with 8 bp deletion which resulted in premature termination of amino acid translation in G1 were selected (Figure 3C). This should lead to changes in the corresponding functions of this gene product in B. dorsalis. The selected G1 were crossed with wild-type to obtain G2 heterozygotes. Self-cross the G2 heterozygotes and homozygotes were recovered in the next generation, demonstrating that this scheme is successful for developing B. dorsalis mutants and could be more widely applied in functional gene studies in this and closely related species.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64195/64195fig01.jpg" /> 
Figure 1: Preparation of sgRNA. (A) General procedures for sgRNA preparation. (B) Target area amplified by PCR from gDNA (100 bp). (C) Example of target gene structure and target cleavage site by Cas9. (D) PCR assembly of sgRNA (~100 bp). (E) Synthesis of sgRNA by in vitro transcription and purification. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64195/64195fig01large.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/64195/64195fig02.jpg" /> 
Figure 2: Embryo microinjection. (A) The process and method of injection. (B) Embryo collection device, laying eggs on gauze. (C) Line up embryos with water and cover with halocarbon oil. (D) Micropipette puller. (E) The whole setup of the microinjection system. Microscope (left), microinjector and micromanipulator (middle), and automatic air pump (right). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64195/64195fig02large.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/64195/64195fig03.jpg" /> 
Figure 3: Mutant Screening. (A) Mating of mutants and acquisition of homozygotes. (B) PCR products from the mutant genome which generates a distinct set of peaks at the target. (C) One of the mutation types and amino acid changes. Deletion mutations lead to premature termination of translation.&#160;Abbreviations: HE = heterozygotes, HO = homozygotes. Control: embryo injected with scrambled sgRNAs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64195/64195fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3" rowspan="2">Injection mix</td>
			<td rowspan="2">Injected embryos</td>
			<td rowspan="2">Hatched larvae</td>
			<td rowspan="2">Pupae</td>
			<td>Mosaic G0</td>
		</tr>
		<tr>
			<td>(Mosaic/Total adults)</td>
		</tr>
		<tr>
			<td colspan="3">BdorOrco_gRNA1 (300 ng/&#956;L)</td>
			<td rowspan="4">200</td>
			<td rowspan="4">83</td>
			<td rowspan="4">66</td>
			<td rowspan="4">49/60 (81%)</td>
		</tr>
		<tr>
			<td colspan="3">BdorOrco_gRNA2 (300 ng/&#956;L)</td>
		</tr>
		<tr>
			<td colspan="3">Cas9 (150 ng/&#956;L)&#160;</td>
		</tr>
		<tr>
			<td colspan="3">Phenol red (1 &#956;L)</td>
		</tr>
		<tr>
			<td colspan="3">BdorOrco_scrambled gRNA1 (300 ng/&#956;L)</td>
			<td rowspan="4">200</td>
			<td rowspan="4">78</td>
			<td rowspan="4">76</td>
			<td rowspan="4">0/70 (0%)</td>
		</tr>
		<tr>
			<td colspan="3">BdorOrco_scrambled gRNA2 (300 ng/&#956;L)</td>
		</tr>
		<tr>
			<td colspan="3">Cas9 (150 ng/&#956;L)</td>
		</tr>
		<tr>
			<td colspan="3">Phenol red (1 &#956;L)</td>
		</tr>
	</tbody>
</table>
Table 1: B. dorsalis survival and mutagenesis after microinjections.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Problem</td>
			<td>Potential cause(s)</td>
			<td colspan="2">Solutions</td>
		</tr>
		<tr>
			<td rowspan="2">Inefficient embryo collection</td>
			<td>1. Adults are in poor condition</td>
			<td colspan="2">1.Ensure the availability of food and water, especially the sugar content of the food.</td>
		</tr>
		<tr>
			<td>2. Insufficient mating</td>
			<td colspan="2">2. Mating in advance under 30-50 Lux dim conditions. It is best to select 12-15 days-old mated adults.</td>
		</tr>
		<tr>
			<td rowspan="3">Poor hatchability of eggs after injection</td>
			<td>1. Poor quality embryos</td>
			<td colspan="2">1. Pick fresh and plump eggs and gently brush them with a water-soaked brush.</td>
		</tr>
		<tr>
			<td>2. Needle does not fit</td>
			<td colspan="2">2. The tip of the needle is as small as possible to minimize damage to the egg during injection.</td>
		</tr>
		<tr>
			<td>3. Improper rearing of eggs after injection</td>
			<td colspan="2">3. Eggs need to be kept hydrated after injection to prevent dehydration from drying out. Eggs can also be transferred directly to food after injection.</td>
		</tr>
		<tr>
			<td rowspan="2">Low larval survival rate</td>
			<td>1. Moldy food is not suitable for larvae growth</td>
			<td colspan="2">1. For newly hatched larvae, add a small amount of food. Too much food&#160;will mold or dry out and affect the growth of the larvae.</td>
		</tr>
		<tr>
			<td>2. Newly hatched larvae soaked in oil for a long time</td>
			<td colspan="2">2.The newly hatched larvae need to be transferred to the larval food in time or the eggs should be picked directly into the food after injection.</td>
		</tr>
		<tr>
			<td rowspan="2">Adult mutation rate is low</td>
			<td>1. Low quality of gRNA</td>
			<td colspan="2">1. Rigorous experimental operation to synthesize high-quality gRNA to prevent degradation.</td>
		</tr>
		<tr>
			<td>2. Inefficient targets lead to off-targets</td>
			<td colspan="2">2. Screening high-efficiency targets through as our discussion mentioned.</td>
		</tr>
	</tbody>
</table>
Table 2: Possible problems with constructing mutants, potential causes, and solutions.

Watch this Scientific Journal Video about Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis at JoVE.com

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.

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 ...

protocols-for-crisprcas9-mutagenesis-oriental-fruit-fly-bactrocera

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2.2K Views.  Northeast Forestry University. 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.

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

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

Genetics

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 ...

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been harnessed by researchers to study important biologically relevant problems. The unparalleled power of the CRISPR/Cas genome editing method allows researchers to precisely edit any locus of their choosing, thereby facilitating an increased understanding of gene function. Several methods for editing the C. elegans genome by CRISPR/Cas9 have been described previously. Here, we discuss and demonstrate a method which utilizes in vitro assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. Specifically, in this article, we go through the step-by-step process of introducing premature stop codons into the C. elegans rbm-3.2 gene by homology-directed repair using this method of CRISPR/Cas9 editing. This relatively simple editing method can be used to study the function of any gene of interest and allows for the generation of homozygous-edited C. elegans by CRISPR/Cas9 editing&#160;in less than two weeks.

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The goal of this protocol is to enable seamless CRISPR/Cas9 editing of the C. elegans genome using assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. This protocol can be used to make a variety of genetic modifications in C. elegans including insertions, deletions, gene replacements and codon substitutions.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been ...

Indel Detection following CRISPR/Cas9 Mutagenesis using High-resolution Melt Analysis in the Mosquito Aedes aegypti

Mosquito gene editing has become routine in several laboratories with the establishment of systems such as transcription-activator-like effector nucleases (TALENs), zinc-finger nucleases (ZFNs), and homing endonucleases (HEs). More recently, clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) technology has offered an easier and cheaper alternative for precision genome engineering. Following nuclease action, DNA repair pathways will fix the broken DNA ends, often introducing indels. These out-of-frame mutations are then used for understanding gene function in the target organisms. A drawback, however, is that mutant individuals carry no dominant marker, making identification and tracking of mutant alleles challenging, especially at scales needed for many experiments. 
High-resolution melt analysis (HRMA) is a simple method to identify variations in nucleic acid sequences and utilizes PCR melting curves to detect such variations. This post-PCR analysis method uses fluorescent double-stranded DNA-binding dyes with instrumentation that has temperature ramp control data capture capability and is easily scaled to 96-well plate formats. Described here is a simple workflow using HRMA for the rapid detection of CRISPR/Cas9-induced indels and the establishment of mutant lines in the mosquito Ae. aegypti. Critically, all steps can be performed with a small amount of leg tissue and do not require sacrificing the organism, allowing genetic crosses or phenotyping assays to be performed after genotyping.

Indel Detection following CRISPR/Cas9 Mutagenesis using High-resolution Melt Analysis in the Mosquito Aedes aegypti

This article details a protocol for rapid identification of indels induced by CRISPR/Cas9 and selection of mutant lines in the mosquito Aedes aegypti using high-resolution melt analysis.

Mosquito gene editing has become routine in several laboratories with the establishment of systems such as transcription-activator-like effector nucleases ...

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.

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.

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

Behavior

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal experiments. When generating animal models, the electroporation of zygotes offers higher efficiency, simplicity, cost, and throughput as an alternative to the gold standard method of microinjection. Electroporation is also gentler, with higher viability, and reliably delivers Cas9/single-guide RNA (sgRNA) ribonucleoproteins (RNPs) into the zygotes of common laboratory mouse strains (e.g., C57BL/6J and C57BL/6N) that approaches 100% delivery efficiency. This technique enables insertion/deletion (indels) mutations, point mutations, the deletion of whole genes or exons, and small insertions in the range of 100-200 bp to insert LoxP or short tags like FLAG, HA, or V5. While constantly being improved, here we present the current state of CRISPR-EZ in a protocol that includes sgRNA production through in vitro transcription, embryo processing, RNP assembly, electroporation, and the genotyping of preimplantation embryos. A graduate-level researcher with minimal experience manipulating embryos can obtain genetically edited embryos in less than 1 week using this protocol. Here, we offer a straightforward, low-cost, efficient, high-capacity method that could be used with mouse embryos.

Here, we describe a simple technique intended for the efficient generation of genetically modified mice called CRISPR RNP Electroporation of Zygotes (CRISPR-EZ). This method delivers editing reagents by electroporation into embryos at an efficiency approaching 100%. This protocol is effective for point mutations, small genomic insertions, and deletions in mammalian embryos.

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal ...

Developmental Biology

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with an attractive off-target profile at both cellular and organismal levels. It is recently reported that the CRISPR/CasRx system can be used to achieve ubiquitous and tissue-specific gene transcript reduction in Drosophila melanogaster. This paper details the methods from the recent work, consisting of three parts: 1) ubiquitous&#160;in vivo endogenous RNA targeting using a two-component CasRx system; 2) ubiquitous in vivo exogenous RNA targeting using a three-component CasRx system; and 3) tissue-specific in vivo RNA targeting using a three-component CasRx system. The effects of RNA targeting observed include targeted gene specific phenotypic changes, targeted RNA transcript reduction, and occasional lethality phenotypes associated with high expression of CasRx protein and collateral activity. Overall, these results showed that the CasRx system is capable of target RNA transcript reduction at the organismal level in a programmable and efficient manner, demonstrating that in vivo transcriptome targeting, and engineering is feasible and lays the foundation for future in vivo CRISPR-based RNA targeting technologies.

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

This article outlines a detailed protocol for using the RNA-targeting Cas13D enzyme (RfxCas13D) in flies.

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with ...

Immunology and Infection

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, host-symbiont interactions, speciation, and venom synthesis. Despite the availability of several molecular tools, including CRISPR/Cas9, functional genetic studies are still limited in this organism. The major limitation of applying CRISPR/Cas9 technology in N. vitripennis stems from the challenges of embryonic microinjections. Injections of embryos are particularly difficult in this organism and in general in many parasitoid wasps, due to small embryo size and the requirement of a host pupa for embryonic development. To address these challenges, Cas9 ribonucleoprotein complex delivery into female&#160;ovaries by adult injection, rather than embryonic microinjection, was optimized, resulting in both somatic and heritable germline edits. The injection procedures were optimized in pupae and female wasps using either ReMOT Control (Receptor-Mediated Ovary Transduction of Cargo) or BAPC (Branched Amphiphilic Peptide Capsules). These methods are shown to be effective alternatives to embryo injection, enabling site-specific and heritable germline mutations.

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

Here, we describe methods for efficient pupal and adult injections in Nasonia vitripennis as accessible alternatives to embryo microinjection, enabling functional analysis of genes of interest using either RNA-silencing via RNA interference (RNAi) or gene knockout via CRISPR/Cas9 genome editing.

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome 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

Summary

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Chapters in this video

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

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

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

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

Indel Detection following CRISPR/Cas9 Mutagenesis using High-resolution Melt Analysis in the Mosquito Aedes aegypti

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

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

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