This work was supported by the National Natural Science Foundation of China (32070502, 31601697, 32072419 and the China Postdoctoral Science Foundation (2020M672205).

1. Target site selection and sgRNA design
<ol>
	<li>Collect as much sequence information as possible for the gene of interest through literature research and/or searching for the mRNA or coding DNA sequence (CDS) of the gene at NCBI and locust database24.</li>
	<li>Compare the sequence of the interested gene to its genomic DNA sequence to distinguish the exon and intron regions.</li>
	<li>Select a candidate region for target site design based on the research purpose. Design primer pairs to amplify the candidate region fragment and verify its wild-type sequence by sequencing analysis of the PCR product. 
	NOTE: There are different strategies for the selection of the candidate target region. For example, in most gene/protein function research, use the exon fragment close to its start codon as the candidate region to ensure that the whole protein/RNA function is lost after the genome editing (i.e., gene knock-out). While for the function analysis of a specific domain, select the candidate region at the beginning (or both ends) of the domain.</li>
	<li>Use online resources such as the E-CRISP Design25 to search for potential target sites in the candidate target region.
	<ol>
		<li>Select &#34;Drosophila melanogaster BDGP6&#34; (or any other insect) in the drop-down list of reference genomes and choose &#34;Input is FASTA sequence&#34; followed by pasting the sequence of target region into the textbox (in FASTA format).</li>
		<li>Start the application by pressing &#34;Start sgRNA search&#34; with &#34;medium&#34; and &#34;single design&#34; to get the possible target sites. Select 1-3 potential targets from the results based on their predicted scores and design the corresponding sgRNAs accordingly. 
		NOTE: There are many more online platforms for CRISPR design, such as CRISPOR, CHOPCHOP, CRISPRdirect, ZiFiT, and so on (see Table of Materials). Some of them have the function of specificity check for the species whose genomic sequence data is in their databases. However, the genomic sequence of locusts has not been found on any online website. It is better to use more than one online tool for the CRISPR design in locusts. Comprehensively, consider the results from different websites and select the sgRNA from the results on the principle of highest specificity, highest efficiency, and least mismatch rate (Figure 1A,B).</li>
	</ol>
	</li>
</ol>
2. Synthesis and verification of the sgRNA&#160;&#160;in vitro
<ol>
	<li>Synthesize the sgRNA using sgRNA synthesis kits according to the manufacturer&#180;s manual (Table of Materials). This procedure usually includes three steps: DNA template amplification, in vitro transcription, and sgRNA purification21. Dilute the synthesized sgRNA with nuclease-free water to a storage concentration of 300 ng/&#181;L for further use.</li>
	<li>Amplify and purify the target gene fragment to serve as the substrate of the in vitro cleavage assay of the CRISPR/Cas9 system. Perform these steps following the manufacturer&#39;s instructions.</li>
	<li>Dilute the purchased Cas9 protein to 300 ng/&#181;L with nuclease-free water. Incubate 1 &#181;L of sgRNA (300 ng/&#181;L) and 1 &#181;L of Cas9 protein (300 ng/&#181;L) with 200 ng of purified target fragment in the cleavage buffer at 37 &#176;C for 1 h (10 &#181;L of total volume; see Table 1). Estimate the efficiency of sgRNA induced Cas9 cleavage by agarose gel electrophoresis (Figure 1C). Select the sgRNAs with high activity for the following microinjection. 
	NOTE: The results of agarose gel electrophoresis can be converted to grayscale images with image processing software and the cleavage efficiency is estimated according to the grayscale of the bands of speculated sizes.</li>
</ol>
3. Microinjection and culture of the eggs
<ol>
	<li>Prepare the injection needles by pulling glass capillaries with a micropipette puller (Table of Materials). Set the parameters as follows: Heat to 588, Pull to 90, Velocity to 60, and Time to 40. Grind the tip of the injection needle with a micro grinder (Table of Materials). 
	NOTE: Ideal needle tips are open and sharp-edged (Figure 2A). It is highly recommended to prepare additional needles for the experiments because needles are sometimes blocked or accidentally broken during injections.</li>
	<li>Mix 1 &#181;L of Cas9 protein (300 ng/&#181;L) with 1 &#181;L of the verified sgRNA (300 ng/&#181;L) together to obtain the RNPs for injection. Add 8 &#181;L of RNase-free sterile water to make the final volume of the mixture to 10 &#181;L. Mix the solution thoroughly and keep it on ice. 
	NOTE: The final concentrations of the Cas9 protein and sgRNAs can be optimized according to the editing results. Avoid repetitive freezing and thawing of the prepared RNP solution and immediate use is recommended.</li>
	<li>Rear the male and female adult locusts together at 30 &#176;C with a 16 (light):8 (dark) photoperiod and supply them sufficient fresh wheat seedlings as food. Observe these locusts daily and put an oviposition pot (a plastic flowerpot or cup filled with wet sterile sand) into the rearing cage for oviposition once the locusts mated. 
	NOTE: Usually, 100 pairs of adult locusts reared in a rearing cage (40 cm x 40 cm x 40 cm) are enough to generate eggs for knocking out a single gene. Culture additional locust pairs if more eggs are needed.</li>
	<li>Collect the freshly-laid egg pods from the oviposition pot and wait for about 30 min for egg tanning. Gently isolate the eggs from the egg pods in water using a fine brush and wash them with sterile water three times. Keep the eggs in a Petri dish and add sterile water to keep them moist. 
	NOTE: Tanning of the newly-laid eggs can significantly improve the mutant efficiency21.</li>
	<li>Fill an injection needle with the prepared RNP solution and load it to the micromanipulator (Table of Materials).
	<ol>
		<li>Set the microinjection parameters as follows: 300 hPa of the injection pressure (pi), 0.5 s of the injection time (ti), and 25 hPa of the compensation pressure (pc).</li>
		<li>Press the pedal to evaluate the volume of the injected solution. Adjust the injection pressure and injection time to ensure that the injection volume is about 50-100 nL. 
		NOTE: Exhaust the residual air in the needle to make the injection solution continuous and controllable as much as possible. The connection of the needle and the micromanipulator need to be tight.</li>
	</ol>
	</li>
	<li>Arrange the sterile eggs regularly on the injection pad (Figure 2B,C) and place the pad on the object table of the microscope (Figure 3A). Adjust the magnification of the microscope until the eggs are observed. Adjust the microinjection needle under the microscope until the injection tip can be seen and position it near the egg to be injected.</li>
	<li>Start the injection at a suitable height and an angle of 30-45 degrees. Insert the tip into the egg carefully near the micropyles of the egg (Figure 3B) and press the pedal to accomplish the injection. Retract the needle quickly and move the injection pad for injection of the next egg. 
	NOTE: A slight expansion of the egg during the microinjection should be observed. A small amount of cytoplasmic leakage at the pinhole is acceptable. Replace the injection needle with a new one or adjust the injection angle if the outflow of fluid is too much.</li>
	<li>Transfer the injected eggs to a culture dish (a Petri dish with a piece of moist filter paper), and place them in an incubator at 30 &#176;C. 
	NOTE: It will take about 13-14 days till nymphs hatch from these injected eggs (on condition that mutation of the target gene does not affect the development of the embryos) (Figure 3C). Inject at least 100 eggs to ensure a sufficient hatching and eclosion amount for knocking out a specific gene.</li>
</ol>
4. Mutation rate estimation and the screening of mutants
<ol>
	<li>Check the development of the injected eggs every day after the injection. Transfer injected eggs to a new culture dish every 24 h in the first 5 days.</li>
	<li>On the 6th day (or later) after injection, randomly pick and transfer 10 eggs into a tube and adequately grind the eggs with two steel balls at 60 Hz for 6 min using a grinder (see Table of Materials). Resuspend the debris with 1 mL of PBS. Transfer 5 &#181;L of the mixture into 45 &#181;L of 50 mM NaOH and lyse at 95 &#176;C for 5 min. Add 5 &#181;L of 1 M Tris-HCl (pH=8.0) into the lysis system to terminate the alkaline lysis reaction. 
	NOTE: Eggs can be picked out for alkaline lysis on any day after the last transfer and multiple eggs can be used as one sample depending on their development situation (e.g., five 6-day-old embryos can be used as one sample, and two 10-day-old embryos can be used as one sample). Sometimes, it is not feasible to get the genomic DNA of eggs using the alkaline lysis method. Commercial genomic DNA extraction kits can be used as an alternative method for isolating the genomic DNA from the eggs.</li>
	<li>Take 1 &#181;L of the lysis product as the PCR template to amplify the target gene fragment (Table 2 and Table 3) and send the PCR products for sequencing. Compare the sequencing results with the wild-type sequence to preliminarily evaluate whether the CRISPR/Cas9 system cleaved the target gene in vivo (Figure 4A). Allow the rest of the eggs for subsequent development on the condition that indels are detected at the embryonic stage. 
	NOTE: In this way, the mutation rate can be preliminarily estimated at the embryo stage. If no indels are found at this step, prepare some new sgRNAs for the target gene and repeat the knock-out protocol from step 2.1.</li>
	<li>Transfer the hatched nymphs to a rearing cage and culture them as described in step 3.3. When the nymphs developed to the fifth instar stage, separate them with plastic culture cups (1 nymph/cup). 
	NOTE: It usually takes about 25-35 days for the nymphs to develop to their fifth instar and the most obvious phenotype is that the wing buds extend to the fourth or fifth abdominal segments. Moreover, it is recommended to record the phenotypes of dead nymphs to predict the relationship between the target gene and phenotypes in further research.</li>
	<li>Cut off about 2 mm length of the antennae with dissecting scissors and lyse it using the alkaline lysis method (step 4.2). Analyze the target gene fragment sequence of these nymphs as described above (step 4.3) to identify the G0 mutants (Figure 4B). 
	&#8203;NOTE: The antennae cut for lysis can be a little longer and grinding or mincing the antennae is helpful for lysis although the antennae can be directly digested. Individuals with multiple peaks near the target site in the sequencing result are identified as positive mutants and allowed for subsequent development and mating.</li>
</ol>
5. Establishment and passaging of mutant lines
<ol>
	<li>Perform cross breeding using the G0 mutants and wild-type locusts (Figure 4B). Collect the oocysts and incubate these oocysts separately at 30 &#176;C. Use 3-6 developed eggs in each oocyst to detect mutations as described in steps 4.2 and 4.3. Keep mutation-positive oocysts for the subsequent development and abandon the mutation-negative oocysts. 
	NOTE: Mutation rate estimation at the embryonic stage can greatly accelerate the screening of G1 mutants. Usually, 3-6 developed eggs can be mixed as one sample for PCR-mediated genotyping as described above.</li>
	<li>Rear the G1 nymphs as described in step 4.4. Cut off about 2 mm length of the antennae to perform PCR-based genotyping as described in step 4.5 for detecting G1 heterozygotes. Perform TA-cloning according to the manufacturer&#39;s instructions (see Table of Materials) to identify their mutations. Perform in-cross using G1 heterozygotes with the same mutations to obtain G2 nymphs. 
	NOTE: Sanger sequencing of the PCR products can only provide information for finding out the heterozygotes, but not enough information for identifying the exact mutations. Thus, TA cloning using the PCR products is required to clearly identify the mutations and can promote the establishment of stable mutant lines.</li>
	<li>Rear the G2 nymphs till their fifth instar. Use the PCR-based genotyping described in step 5.2 to identify the homozygotes and/or heterozygotes that are suitable for further research and stable passaging (Figure 4B). 
	&#8203;NOTE: Pay attention to avoid mixing of homozygotes and heterozygotes. It is recommended to perform PCR-based genotyping in every generation to confirm the homozygosity and/or heterozygosity of each population. The number of locusts used for this check depends on the population size. Moreover, the population of locusts can be expanded by the in-cross strategy.</li>
</ol>
6. Egg cryopreservation and resuscitation
<ol>
	<li>Wash the eggs to be cryopreserved with sterile water and incubate them in a culture dish as described above (step 3.8) for 5-6 days. Gather these eggs together in the culture dish and cover them with filter paper fragments. Wrap the entire culture dish with a paraffin film.</li>
	<li>Keep the dish at 25 &#176;C for 2 days followed by another 2 days at a relatively lower temperature (13-16 &#176;C). Then, transfer the dish to a 6 &#176;C refrigerator. Add water into it every 2 weeks to provide a moist environment for the eggs (Figure 5A). 
	NOTE: The embryos are speculated to be at the katatrepsis stage after being developed for 5-6 days at 30 &#176;C26. Gradient cooling is helpful for cryopreservation of eggs (Table 5).</li>
	<li>For resuscitating the cryopreserved eggs, take the Petri dish out from the refrigerator and keep it at 25 &#176;C for 2 days. Place these eggs in a 30 &#176;C incubator until the nymphs hatch (Figure 5B). 
	NOTE: It is necessary to put the cryopreserved eggs at 25 &#176;C before transferring them to a 30 &#176;C incubator. The embryos can be cryopreserved for at least five months although the hatching rate and eclosion rate can decrease because of the cryopreservation (Table 5).</li>
</ol>

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The authors declare that they have no conflicts of interest.

Locusts have been among the most devastating pests to agriculture since the civilization of human beings23. CRISPR/Cas9-based genome editing technology is a powerful tool for providing better knowledge of the biological mechanisms in locusts as well as a promising pest control strategy. Thus, it is of great benefit to develop an efficient and easy-to-use method of CRISPR/Cas9-mediated construction of homozygous locust mutants. Although some great works have been reported and provided some basic wo...

Gene editing technologies could be used to introduce insertions or deletions into a specific genome locus to artificially modify the target gene on purpose1. In the past years, CRISPR/Cas9 technology has developed rapidly and has a growing scope of applications in various fields of life sciences2,3,4,5,6. The CRISPR/Cas9 system was discovered back in 19877, and widely found in bacteria and archaea. Further research indicated that it was a prokaryotic adaptive immune system that depends on the RNA-guided nuclease Cas9 to fight against phages8. The artificially modified CRISPR/Cas9 system mainly consists of two components, a single guide RNA (sgRNA) and the Cas9 protein. The sgRNA is made up of a CRISPR RNA (crRNA) complementary to the target sequence and an auxiliary trans-activating crRNA (tracrRNA), which is relatively conserved. When the CRISPR/Cas9 system is activated, the sgRNA forms a ribonucleoprotein (RNP) with the Cas9 protein and guides Cas9 to its target site via the base pairing of RNA-DNA interactions. Then, the double-strand DNA can be cleaved by the Cas9 protein and as a result, the double-strand break (DSB) emerges near the protospacer adjacent motif (PAM) of the target site9,10,11,12. To mitigate the damage caused by the DSBs, cells would activate comprehensive DNA damage responses to efficiently detect the genomic damages and initiate the repair procedure. There are two distinct repair mechanisms in the cell: non-homologous end joining (NHEJ) and homology-directed repair (HDR). NHEJ is the most common repair pathway that can repair DNA double-strand breaks quickly and prevents cell apoptosis. However, it is error-prone because of leaving small fragments of insertions and deletions (indels) near the DSBs, which usually results in an open reading frame shift and thus can lead to gene knock-out. In contrast, homologous repairment is quite a rare event. On the condition that there is a repair template with sequences homologous to the context of the DSB, cells would occasionally repair the genomic break according to the nearby template. The result of HDR is that the DSB is precisely repaired. Especially, if there is an additional sequence between the homologous sequences in the template, they would be integrated into the genome through HDR, and in this way, the specific gene insertions could be realized13.
With the optimization and development of the sgRNA structures and Cas9 protein variants, the CRISPR/Cas9-based genetic editing system has also been successfully applied in research of insects, including but not limited to Drosophila melanogaster, Aedes aegypti, Bombyx mori, Helicoverpa Armigera, Plutella xylostella, and Locusta migratoria14,15,16,17,18,19. To the best of the authors&#39; knowledge, although RNPs consisting of the Cas9 protein and in vitro transcribed sgRNA have been used for locust genome editing20,21,22, a systematic and detailed protocol for CRISPR/Cas9 ribonuclease mediated construction of homozygous mutants of the migratory locust is still lacking.
The migratory locust is an important agricultural pest that has a global distribution and poses substantial threats to food production, being especially harmful to gramineous plants, such as wheat, maize, rice, and millet23. Gene function analysis based on genome editing technologies can provide novel targets and new strategies for the control of migratory locusts. This study proposes a detailed method for knocking out migratory locust genes via the CRISPR/Cas9 system, including the selection of target sites and design of sgRNAs, in vitro synthesis and verification of the sgRNAs, microinjection and culture of eggs, estimation of mutation rate at the embryonic stage, detection of mutants as well as passage and preservation of the mutants. This protocol could be used as a basal reference for manipulation of the vast majority of locust genes and can provide valuable references for genome editing of other insects.

<table><tbody><tr><td>10&amp;times;NEBuffer r3.1</td><td>New England Biolabs</td><td>B7030S</td><td>The buffer of &amp;nbsp;&lt;em&gt;in vitro&lt;/em&gt; Cas9 cleavage assays</td></tr><tr><td>2xEs Taq MasterMix (Dye)</td><td>Cwbio</td><td>CW0690</td><td>For gene amplification</td></tr><tr><td>2xPfu MasterMix (Dye)</td><td>Cwbio</td><td>CW0686</td><td>For gene amplification</td></tr><tr><td>CHOPCHOP</td><td></td><td></td><td>Online website for designing sgRNAs, http://chopchop.cbu.uib.no/.</td></tr><tr><td>CRISPOR</td><td></td><td></td><td>Online website for designing sgRNAs, http://crispor.org.</td></tr><tr><td>CRISPRdirect</td><td></td><td></td><td>Online website for designing sgRNAs, http://crispr.dbcls.jp/.</td></tr><tr><td>Electrophoresis power supply</td><td>LIUYI BIOLOGY</td><td>DYY-6D</td><td>Separation of nucleic acid molecules of different sizes</td></tr><tr><td>Eppendorf Tube</td><td>Eppendorf</td><td>30125177</td><td>For sample collection, etc.</td></tr><tr><td>Fine brushes</td><td>Annigoni</td><td>1235</td><td>For cleaning and isolating eggs. Purchased online.</td></tr><tr><td>Flaming/brown micropipette puller</td><td>Sutter Instrument</td><td>P97</td><td>For making the microinjection needles</td></tr><tr><td>Gel Extraction Kit</td><td>Cwbio</td><td>CW2302</td><td>DNA recovery and purification</td></tr><tr><td>Gel Imaging Analysis System</td><td>OLYMPUS</td><td>Gel Doc XR</td><td>Observe the electrophoresis results</td></tr><tr><td>GeneTouch Plus</td><td>Bioer</td><td>B-48DA</td><td>For gene amplification</td></tr><tr><td>Glass electrode capillary</td><td>Gairdner</td><td>GD-102</td><td>For making&amp;nbsp;injection needles with&amp;nbsp;a&amp;nbsp;micropipette Puller</td></tr><tr><td>Incubator</td><td>MEMMERT</td><td>INplus55</td><td>For migratory locust embryo culture</td></tr><tr><td>Metal bath</td><td>TIANGEN</td><td>AJ-800</td><td>For heating the sample</td></tr><tr><td>Micro autoinjector</td><td>Eppendorf</td><td>5253000068</td><td>Microinjection of embryos early in development</td></tr><tr><td>Micro centrifuge</td><td>Allsheng</td><td>MTV-1</td><td>Used for mixing reagents</td></tr><tr><td>Microgrinder</td><td>NARISHIGE</td><td>EG-401</td><td>To ground the tip of injection needle</td></tr><tr><td>Microloader</td><td>Eppendorf</td><td>5242956003</td><td>For loading solutions into the injection needles.</td></tr><tr><td>Micromanipulation system</td><td>Eppendorf</td><td>TransferMan 4r</td><td>An altinative manipulation system for microinjection</td></tr><tr><td>Microscope</td><td>cnoptec</td><td>SZ780</td><td>For microinjection</td></tr><tr><td>Motor-drive Manipulator</td><td>NARISHIGE</td><td>MM-94</td><td>For controling the position of the micropipette during the microinjection precedure</td></tr><tr><td>Multi-Sample Tissue Grinder</td><td>jingxin</td><td>Tissuelyser-64</td><td>Grind and homogenize the eggs</td></tr><tr><td>ovipisition pot</td><td>ChangShengYuanYi</td><td>CS-11</td><td>Filled with wet sterile sand for locust ovipositing in it. The oocysts are collected from this container. Purchased online.</td></tr><tr><td>Parafilm</td><td>ParafilmM</td><td>PM996</td><td>For wrapping the petri dishes.</td></tr><tr><td>pEASY-T3 Cloning Kit</td><td>TransGen Biotech</td><td>CT301-01</td><td>For TA cloning</td></tr><tr><td>Petri dish</td><td>NEST</td><td>752001</td><td>For culture and preservation of&amp;nbsp; the eggs.</td></tr><tr><td>Pipettor</td><td>Eppendorf</td><td>Research plus</td><td>For sample loading</td></tr><tr><td>plastic culture cup</td><td></td><td></td><td>For rearing locusts seperately and any plastic cup big enough (not less than 1000 mL in volume) will do. Purchased online.</td></tr><tr><td>Precision gRNA Synthesis Kit</td><td>Thermo</td><td>A29377</td><td>For sgRNA synthesis</td></tr><tr><td>Primer Premier</td><td>PREMIER Biosoft</td><td>Primer Premier 5.00</td><td>For primer design</td></tr><tr><td>SnapGene</td><td>Insightful Science</td><td>SnapGene&amp;reg;4.2.4</td><td>For analyzing&amp;nbsp; sequences</td></tr><tr><td>Steel balls</td><td>HuaXinGangQiu</td><td>HXGQ60</td><td>For sample grinding.Purchased online.</td></tr><tr><td>Tips</td><td>bioleaf</td><td>D781349</td><td>For sample loading</td></tr><tr><td>Trans DNA Marker II</td><td>TransGen Biotech</td><td>BM411-01</td><td>Used to determine gene size</td></tr><tr><td>TrueCut Cas9 Protein v2</td><td>Thermo</td><td>A36496</td><td>Cas9 protein</td></tr><tr><td>UniversalGen DNA Kit</td><td>Cwbio</td><td>CWY004</td><td>For genomic DNA extraction</td></tr><tr><td>VANNAS Scissors</td><td>Electron Microscopy Sciences</td><td>72932-01</td><td>For cutting off the antennae</td></tr><tr><td>Wheat</td><td></td><td></td><td>To generate wheat seedlings as the food for locusts. Bought from local farmers.</td></tr><tr><td>ZiFiT</td><td></td><td></td><td>Online website for designing sgRNAs, http://zifit.partners.org/ZiFiT/ChoiceMenu.aspx.</td></tr></tbody></table>

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 protocol contains the detailed steps for generating homozygous mutants of the migratory locusts with the RNP consisting of Cas9 protein and in vitro synthesized sgRNA. The following are some representative results of CRISPR/Cas9-mediated target gene knockout in locusts, including target selection, sgRNA&#160;synthesis and verification (Figure 1A), egg collection and injection, mutant screening and passaging, cryopreservation, and resuscitation of the homozygous eggs.
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Watch this Scientific Journal Video about Construction of Homozygous Mutants of Migratory Locust Using CRISPR/Cas9 Technology at JoVE.com

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

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 ...

construction-homozygous-mutants-migratory-locust-using-crisprcas9

Universidad de Cartagena UDEC

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Biology

2.0K Views.  Henan University. 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.

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

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the African clawed frog, Xenopus, a longstanding model organism for biomedical research. Traditional breeding schemes for creating homozygous mutant lines with CRISPR/Cas9-targeted mutagenesis have several time-consuming and laborious steps. To facilitate the creation of mutant embryos, particularly to overcome the obstacles associated with knocking out genes that are essential for embryogenesis, a new method called leapfrogging was developed. This technique leverages the robustness of Xenopus embryos to &#34;cut and paste&#34; embryological methods. Leapfrogging utilizes the transfer of primordial germ cells (PGCs) from efficiently-mutagenized donor embryos into PGC-ablated wildtype siblings. This method allows for the efficient mutation of essential genes by creating chimeric animals with wildtype somatic cells that carry a mutant germline. When two F0 animals carrying &#34;leapfrog transplants&#34; (i.e., mutant germ cells) are intercrossed, they produce homozygous, or compound heterozygous, null F1 embryos, thus saving a full generation time to obtain phenotypic data. Leapfrogging also provides a new approach for analyzing maternal effect genes, which are refractory to F0 phenotypic analysis following CRISPR/Cas9 mutagenesis. This manuscript details the method of leapfrogging, with special emphasis on how to successfully perform PGC transplantation.

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Genes essential for survival pose technical hurdles for creating mutant lines. Leapfrogging circumvents lethality by combining genome editing with primordial germ cell transplantation to create wild-type animals carrying germline mutations. Leapfrogging also permits the efficient generation of homozygous null mutants in the F1 generation. Here, the transplantation step is demonstrated.

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the ...

Genetics

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 ...

Neurocircuit Assays for Seizures in Epilepsy Mutants of Drosophila

Drosophila melanogaster is a useful tool for studying seizure like activity. A variety of mutants in which seizures can be induced through either physical shock or electrical stimulation is available for study of various aspects of seizure activity and behavior. All flies, including wild-type, will undergo seizure-like activity if stimulated at a high enough voltage. Seizure like activity is an all-or-nothing response and each genotype has a specific seizure threshold. The seizure threshold of a specific genotype of fly can be altered either by treatment with a drug or by genetic suppression or enhancement. The threshold is easily measured by electrophysiology. Seizure-like activity can be induced via high frequency electrical stimulation delivered directly to the brain and recorded through the dorsal longitudinal muscles (DLMs) in the thorax. The DLMs are innervated by part of the giant fiber system. Starting with low voltage, high frequency stimulation, and subsequently raising the voltage in small increments, the seizure threshold for a single fly can be measured.

Neurocircuit Assays for Seizures in Epilepsy Mutants of Drosophila

Using high frequency electrical stimulation, seizure-like activity can be induced in Drosophila. This activity is easily recorded from the giant fiber system.

Drosophila melanogaster is a useful tool for studying seizure like activity. A variety of mutants in which seizures can be induced through either physical ...

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 ...

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

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 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 ...

Bioengineering

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 ...

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 ...

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

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Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

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

Neurocircuit Assays for Seizures in Epilepsy Mutants of Drosophila

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

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

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

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

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

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

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