TO and TD were supported by JSPS KAKENHI grant numbers 19H02970 and 20H02999, respectively.

1. Maintenance of laboratory colonies
<ol>
	<li>For the maintenance of wildtype and mutant populations, use a large plastic container (460 mm x 360 mm x 170 mm) with regular artificial fish food, water in plastic cups with a ventilation hole on the top, a folded paper for hiding the insects, and layered cotton for laying eggs (Figure 2A). Keep all T. domestica cultures inside 37 &#176;C incubators and set the relative humidity (RH) inside each container to 60%&#8211;80%. 
	NOTE: Because T. domestica absorbs water vapor from the atmosphere11, a direct water supply is not needed. The appropriate RH is maintained due to the presence of water vapor from plastic cups containing a ventilation hole on the top or from lidless cups inside each container. There is no need to humidify inside an entire incubator which contains cultures. The approximate duration of each developmental stage under the condition described in this protocol is shown in Figure 1. The developmental speed could be adjusted by changing the temperature and/or RH12.</li>
	<li>Add food periodically. Add water before it dries up.</li>
	<li>Transfer the populations to a new clean container at least every 3 months given that adults stop laying eggs in a dirty environment and/or dense population (see Discussion).</li>
</ol>
2. Egg collection and microinjection
<ol>
	<li>Design a guide RNA (gRNA)
	<ol>
		<li>Design a gRNA sequence for each required target and BLAST the gRNA sequences against the genome assembly to check possible off-target recognition sites.</li>
		<li>Synthesize and purify the gRNAs according to the manufacturer&#8217;s instructions8. 
		NOTE: As an example, in the case of targeting the ATP-binding cassette transporter white gene, a 20 bp target sequence was designed and two synthesized DNA oligonucleotides 5&#180;-TAATACGACTCACTATAGTAAGTGTTGTGGGAC-3&#180; and 5&#180;-TTCTAGCTCTAAAACATCGGTCCCACAACACTTA-3&#180; were ordered. The DNA template was prepared by annealing these two oligonucleotides followed by PCR amplification and the gRNA is then in vitro transcribed from the template with a T7 RNA polymerase.</li>
	</ol>
	</li>
	<li>Prepare egg collection colonies
	<ol>
		<li>Transfer about 20 male and 20 female adults to a middle-sized container (200 mm x 150 mm x 90 mm) with food, water supply, a folded paper, and a small piece of layered cotton for egg laying (Figure 2B).</li>
		<li>Set up several colonies to obtain large number of staged embryos in a short time period to be used for genome editing. 
		NOTE: About 20&#8211;40 eggs are expected to be collected from a colony after 8 h at 37 &#176;C. It usually takes a few days for transferred adults to start laying eggs, possibly due to adaptation to a new environment.</li>
	</ol>
	</li>
	<li>On the day of injection, replace the cotton inside the containers with new ones.</li>
	<li>Place a 76 mm x 5 mm double-sided tape on a regular 76 mm x 26 mm glass slide.</li>
	<li>Eight hours later, collect the eggs from the layered cotton by separating the layers using forceps.</li>
	<li>Align the eggs on the double-sided tape using a wet paint brush and keep a 2 mm distance between the eggs. All eggs should be oriented so that the longitudinal axis of an egg faces the injection side (Figure 3A). Gently press down the eggs with a paint brush for firm holding during the injection. 
	NOTE: In this protocol, fungus grows fast in damaged injected eggs under wet and warm condition. It is important to keep the distance between the eggs to prevent cross contamination and expansion of fungus.</li>
	<li>Mix the gRNA and Cas9 protein to a final concentration of 100 ng/&#181;L and 500 ng/&#181;L, respectively. Use distilled water for dilution. Incubate the mix for 10 min at room temperature to promote ribonucleoprotein complex formation and then keep the mix on ice. 
	NOTE: Neutral red at a 1% final concentration could be added to the injection solution to monitor the injected amount.</li>
	<li>Load 2 &#181;L of the gRNA/Cas9 solution in a glass injection capillary with a microloader. Make sure there are no air bubbles in the solution before the injection. If necessary, tap the needle to remove the bubbles. 
	NOTE: In this experiment, a ready-made needle is used to obtain a fine needle tip (Figure 3B). A homemade needle with a similar shape could be used instead.</li>
	<li>Fix the glass injection capillary, previously loaded with the gRNA/Cas9 solution, to a holder equipped with a manipulator and connect the holder to an electronic microinjector.</li>
	<li>Optimize the shape of the needle tip by breaking it slightly with forceps to prevent clogging and getting better durability throughout a series of injections (an example of an appropriate needle is shown in Figure 3C). 
	NOTE: It is recommended to change a needle when it is clogged. It is possible to continue injecting with the same needle if it is broken again with forceps, but a wider tip leads to more egg damage and lowers their survival rate. As T. domestica eggs are soft and fragile, keeping a fine needle tip is key for having a high survival rate after injection.</li>
	<li>Insert the needle at the midpoint of the longitudinal axis of an egg and inject a slight amount of the solution (see Figure 3D&#8211;H for reference on the amount of injection). Adjust the configuration of the electronic microinjector during injection, depending on the shape of the needle tip. 
	NOTE: It is recommended to apply a constant pressure during injection, otherwise sticky egg contents can easily flow into the glass needle and may clog it. The solution may be to either inject with a short pressure pulse or with a constant pressure, depending on the amount of liquid injected. Keep in mind that when a needle tip has a wide opening, too much solution is injected into an egg, which causes lethality (Figure 3F&#8211;H). In that case, maintain a constant liquid flow by constant pressure, then insert the needle into an egg and pull it out immediately. If it causes too much overflow or the egg bursts, change the needle.</li>
	<li>Keep the injected eggs in a container with the appropriate size for the number of injected eggs (&#60;20 eggs: small dish; &#62;20 eggs: middle-sized container) with 60%&#8211;80% RH and 37 &#176;C.</li>
</ol>
3. Mating
<ol>
	<li>Check the injected eggs periodically and discard damaged eggs with forceps to avoid fungal growth (Figure 3I,J). In case too much fungus is growing on an egg, clean up the surface of the egg with 70% EtOH.</li>
	<li>Before hatching (approximately 10 days at 37 &#176;C after egg laying), dip the glass slide with the injected eggs into talcum powder to coat the surface of the double-sided tape. This will avoid the stacking of hatched nymphs. Transfer the powder-coated glass slides to a middle-sized container with food, water, and a folded paper for hiding of the insects.</li>
	<li>Remove the glass slides after the nymphs have hatched. Periodically supply food until they reach adulthood. 
	NOTE: It takes about 2.0&#8211;2.5 months for individuals to reach adulthood after they hatch (Figure 1A). The adult stage is judged based on a well-developed ovipositor in females (Figure 1B).</li>
	<li>To mate the individuals, transfer as many as needed wildtype female adults from a laboratory colony to the medium-sized container and incubate them for at least 14 days at 37 &#176;C to make sure they are virgin. 
	NOTE: It is not necessary to collect virgin females from a laboratory colony because adult T. domestica have a repeated cycle of molting and mating called &#8220;reproductive and molting cycle&#8221;, during which females throw away sperm with each molt and mate again in the next fertilization cycle13.</li>
	<li>Transfer either a male or a female G0 adult that developed from an injected egg and wildtype adult(s) to a small plastic dish (&#216; 100 mm x 40 mm) with food, a folded paper, and a small piece of cotton for laying G1 eggs (Figure 2C&#8217;; mating dish). Keep the mating dishes in a larger container with 60%&#8211;80% RH (Figure 2C). 
	NOTE: Multiple wildtype adults can be included in a dish to increase the chance of successful mating, although high success rates have been achieved with one-to-one pairing.</li>
</ol>
4. Genotyping
<ol>
	<li>Design PCR primer pairs for each gRNA to amplify a 100&#8211;200 bp product that includes the site targeted by the gRNA. BLAST each primer sequence against the genome assembly to check its specificity (an example is shown for the targeting of the white gene in Figure 4A).</li>
	<li>Check the germline transformation of G0 adults.
	<ol>
		<li>Five days after the eggs are laid, collect individual G1 eggs from each G0 adult pair into 0.2 mL tubes (one egg per tube; store collected samples at -20 &#176;C for a long-term storage). Separate the cotton layers to collect the eggs. 
		NOTE: It is recommended to genotype at least 12 G1 nymphs from each mating pair to evaluate the success of the germline transmission (see Representative Results).</li>
		<li>Add 15 &#181;L of a 0.25 mg/mL Proteinase K solution (dissolved in Tris-EDTA buffer) to each tube, briefly homogenize samples with toothpicks, and incubate at 55 &#176;C for 3 to 16 h.</li>
		<li>Inactivate the Proteinase K by placing the samples at 95 &#176;C for 10 min.</li>
		<li>Add 90 &#181;L of distilled water to each tube and mix well. Use 2 &#181;L of supernatant in a 10 &#181;L PCR reaction mix containing the primers designed in step 4.1. 
		NOTE: The use of a DNA polymerase optimized for crude templates is recommended to reach enough PCR amplification.</li>
		<li>To analyze the PCR products, perform an heteroduplex mobility assay (HMA) with a microchip electrophoresis system (Figure 3B; see Ohde et al., 2018)8. 
		NOTE: Mutations could be assessed with two alternative methods: (1) HMA with standard gel polymers, such as 8% polyacrylamide14; (2) digestion of PCR products with T7 endonuclease followed by agarose gel electrophoresis15.</li>
		<li>Keep only the G1 nymphs resulting from G0 adults that contain mutations in their germline and discard the others.</li>
	</ol>
	</li>
	<li>Individual genotyping of G1 nymphs/adults
	<ol>
		<li>Isolate G1 nymphs into 24-well plates with an aspirator or a paint brush. Place the 24-well plates in a larger container (e.g., the medium-sized container used in this protocol) with water supply as described above to keep a RH of 60%&#8211;80% (Figure 1D). Maintain the supply of artificial regular fish food (Figure 1D&#8217;). 
		NOTE: Although this step can be performed at any point of nymphal and adult stages, it is recommended to perform it after reaching adulthood and just before pairing (&#62;2.5 months after injection; Figure 1B) because it is easier to maintain firebrats in a large container. Individual rearing is required to track the genotype of each G1 nymph on the following steps. G1 nymphs from the same G0 adult can have different mutations.</li>
		<li>Pinch and pull cerci and the caudal filament from a nymph/adult using forceps and collect them into a 0.2 mL tube containing 50 &#181;L EtOH (store the collected samples at -20 &#176;C for a long-term storage). 
		NOTE: Tissue samples are collected in EtOH because it prevents the loss of these small samples due to static electricity. If one needs to stop the motion of insects, anesthetize nymph/adult on ice when taking tissue samples. Because T. domestica cannot survive after long-term cooling on ice, do not anesthetize them for more than a minute and immediately move them back to room temperature. Ablation of cerci and the caudal filament causes no increase of mortality.</li>
		<li>Place sample tubes with the lids open on a thermal block for 15 min at 70 &#176;C to evaporate the EtOH.</li>
		<li>Repeat steps 4.2.2&#8211;4.2.5 for genotyping.</li>
		<li>Submit the PCR products in which a mutant band pattern is observed to a standard Sanger sequencing service.</li>
		<li>Keep the G1 nymphs/adults with the desired mutations and discard the others (see Figure 4C for a representative sequencing result).</li>
	</ol>
	</li>
	<li>Cross the adults containing the desired mutations in a mating dish and obtain the next generations to establish a homozygous mutant strain.</li>
</ol>

<ol>
	<li>Peterson, M. D., Rogers, B. T., Popadi&#263;, A., Kaufman, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+embryonic+expression+pattern+of+labial,+posterior+homeotic+complex+genes+and+the+teashirt+homologue+in+an+apterygote+insect.">The embryonic expression pattern of labial, posterior homeotic complex genes and the teashirt homologue in an apterygote insect.</a> Development Genes and Evolution. 209, 77-90 (1999).</li><li>Farris, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+organization+of+the+mushroom+bodies+of+Thermobia+domestica+(Zygentoma,+Lepismatidae):+Insights+into+mushroom+body+evolution+from+a+basal+insect.">Developmental organization of the mushroom bodies of Thermobia domestica (Zygentoma, Lepismatidae): Insights into mushroom body evolution from a basal insect.</a> Evolution and Development. 7, 150-159 (2005).</li><li>Kamae, Y., Tanaka, F., Tomioka, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+cloning+and+functional+analysis+of+the+clock+genes,+Clock+and+cycle,+in+the+firebrat+Thermobia+domestica.">Molecular cloning and functional analysis of the clock genes, Clock and cycle, in the firebrat Thermobia domestica.</a> Journal of Insect Physiology. 56, 1291-1299 (2010).</li><li>Ohde, T., Masumoto, M., Yaginuma, T., Niimi, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+RNAi+analysis+in+the+firebrat,+Thermobia+domestica:+Distal-less+is+required+to+form+caudal+filament.">Embryonic RNAi analysis in the firebrat, Thermobia domestica: Distal-less is required to form caudal filament.</a> Journal of Insect Biotechnology and Sericology. 105, 99-105 (2009).</li><li>Ohde, T., Yaginuma, T., Niimi, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nymphal+RNAi+analysis+reveals+novel+function+of+scalloped+in+antenna,+cercus+and+caudal+filament+formation+in+the+firebrat,+Thermobia+domestica.">Nymphal RNAi analysis reveals novel function of scalloped in antenna, cercus and caudal filament formation in the firebrat, Thermobia domestica.</a> Journal of Insect Biotechnology and Sericology. 108, 101-108 (2012).</li><li>Bell&#233;s, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Beyond+Drosophila:+RNAi+in+vivo+and+functional+genomics+in+insects.">Beyond Drosophila: RNAi in vivo and functional genomics in insects.</a> Annual Review of Entomology. 55, 111-128 (2010).</li><li>Ohde, T., Takehana, Y., Shiotsuki, T., Niimi, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9-based+heritable+targeted+mutagenesis+in+Thermobia+domestica:+A+genetic+tool+in+an+apterygote+development+model+of+wing+evolution.">CRISPR/Cas9-based heritable targeted mutagenesis in Thermobia domestica: A genetic tool in an apterygote development model of wing evolution.</a> Arthropod Structure and Development. 47, 362-369 (2018).</li><li>Nji, C., 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=CRISPR/Cas9+in+insects:+Applications,+best+practices+and+biosafety+concerns.">CRISPR/Cas9 in insects: Applications, best practices and biosafety concerns.</a> Journal of Insect Physiology. 98, 245-257 (2017).</li><li>Brand, 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=The+origin+of+the+odorant+receptor+gene+family+in+insects.">The origin of the odorant receptor gene family in insects.</a> eLife. 7, 1-13 (2018).</li><li>Noble-Nesbitt, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Water+balance+in+the+firebrat,+Thermobia+domestica+(Packard).+Exchanges+of+water+with+the+atmosphere.">Water balance in the firebrat, Thermobia domestica (Packard). Exchanges of water with the atmosphere.</a> Journal of Experimental Biology. 50, 745-769 (1969).</li><li>Sweetman, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physical+ecology+of+the+firebrat,+Thermobia+domestica+(Packard).">Physical ecology of the firebrat, Thermobia domestica (Packard).</a> Ecological Monographs. 8, 285-311 (1938).</li><li>Nijhout, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Insect Hormones. , (1994).</li><li>Chen, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+detection,+quantification+and+enrichment+of+subtle+allelic+alterations.">Efficient detection, quantification and enrichment of subtle allelic alterations.</a> DNA Research. 19, 423-433 (2012).</li><li>Mashal, R. D., Koontz, J., Sklar, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+mutations+by+cleavage+of+DNA+heteroduplexes+with+bacteriophage+resolvases.">Detection of mutations by cleavage of DNA heteroduplexes with bacteriophage resolvases.</a> Nature Genetics. 9, 177-183 (1995).</li><li>Adams, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biological+notes+upon+the+firebrat,+Thermobia+domestica+Packard.">Biological notes upon the firebrat, Thermobia domestica Packard.</a> Journal of the New York Entomological Society. 41, 557-562 (1933).</li></ol>

The authors have nothing to disclose.

For the successful generation of the desired T. domestica mutant with CRISPR/Cas9, it is first important to collect a sufficient number of staged embryos for injection. For a constant collection of a sufficient number of T. domestica eggs, the key is to select an appropriate size of the container to have a lower population density because it would help the successful completion of a series of complex mating behaviors, which is repeated after every adult molt13. A male T. domestica adult transfers its sperm indirectly to a female via a spermatophore. Sweetman (1938) reported that it takes male adults 20 to 35 min from the initiation of the mating behavior to the placement of a spermatophore12. It is likely that disturbance of the interaction between a mating pair by other individuals prevents successful fertilization, which could happen more frequently in a dense environment.
Although eggs are collected 8 h after replacing the cotton inside a container to collect enough eggs in our protocol, the higher efficiency of genome editing may be achieved by collecting eggs and injecting them within a shorter time (e.g., 4 h after egg laying) when injected materials have more chance to be delivered to the large proportion of nuclei. Injection to a sufficient number of eggs within a shorter time could be done by (1) increasing the number or the size of containers if space allows, or (2) repeating the same procedure to obtain a sufficient number of injected eggs.
The site of injection is generally considered to be important for successful germline transformation in insects. T. domestica eggs are usually ellipsoidal in shape and contain a germ band at one pole of their longitudinal axis. It is recommended to inject the gRNA/Cas9 mixture at the midpoint of the egg longitudinal axis because it is hard to identify the pole where a germ band is formed in early embryos due to the variable shape of T. domestica egg. Although the gRNA/Cas9 solution is not directly injected to the site of the germ band formation, we have achieved germline transformation in as high as 39.1% of G0 adults8.
During the microinjection of eggs, it is important to use a fine needle tip to obtain a high survival rate, as it is the case for other animal models. We obtained a higher survival rate after injection with ready-made needles than when using homemade glass needles in T. domestica eggs. However, a similarly high survival rate could be achieved by replicating the fine needle shape with homemade needles (as shown in Figure 3B,C). According to our experience, the dry injection method results in a higher survival rate than the previously reported wet injection method8. We found that egg incubation in a dry environment is key for this improvement, taking advantage of the fact that T. domestica eggs and early nymphs are resistant to desiccation. This is in accordance with previous reports suggesting that this species prefers a dry environment particularly during early developmental stages and that a wet environment can even be harmful16. Instead of a plastic plate, an agarose gel can be used for a quick alignment of the eggs; however, transferring the injected eggs to a dry surface leads to a higher survival rate.
In conclusion, our method underlines the importance of taking the unique biology of T. domestica into account for a successful genome editing: keeping a sparse environment for collecting a sufficient number of eggs and a dry environment for having a higher survival rate of injected embryos. The basic protocols for injection, mating, and culture maintenance reported here are used not only for generating mutant strains with genome editing, but also for applications of other genetic tools such as RNAi and transgenesis and will help to understand the mechanisms underlying the early evolution of insects.

Thermobia domestica belongs to one of the most basal insect orders, Zygentoma, which retains an ancestral ametabolous and wingless life cycle. Such basal phylogenetic position and ancestral characteristics set this species as an attractive model for studying the mechanisms underlying the success of insects on Earth, which cover over 70% of the described animal species1. T. domestica has long been used mainly to study ancestral characteristics of insect physiology because of its suitable features as a laboratory model, such as a relatively short lifecycle (2.5&#8211;3.0 months from embryo to reproductive adult; Figure 1A) and an easy breeding. In the past three decades, its use has been expanded to investigate ancestral characteristics of various traits such as body plan, neural differentiation, and circadian rhythms2,3,4.
The application of advanced genetic tools in T. domestica could further accelerate such contributions in a wide research area. Successful RNA interference (RNAi)-mediated gene knockdown in embryos, nymphs, and adults has been reported in T. domestica4,5,6. The efficiency of systemic RNAi is still highly species-dependent&#8212;for example, it is generally high in coleoptera whereas it is low in the lepidoptera order7. The efficiency and duration of the RNAi knockdown in T. domestica is yet to be assessed. In addition to RNAi, we have previously reported a successful CRISPR/Cas9-mediated gene knockout in T. domestica8. The CRISPR/Cas system has been widely applied for genome editing in insects particularly for targeted gene knockout. Its use could be expanded for other applications such as gene reporter assay, cell lineage tracking, and manipulation of transcriptional activity by knocking-in exogenous constructs after the establishment of a protocol for delivering components of the CRISPR/Cas system into nuclei9. Combined with the published genome assembly10, the wide use and further development of the CRISPR/Cas-based genome editing in T. domestica would facilitate studies focusing on the evolutionary mechanisms behind the outstanding adaptive success of insects. Here, we describe a detailed protocol for embryo microinjection and for mating adult T. domestica to generate a mutant strain using CRISPR/Cas9. Considering this novel method, we discuss the importance of considering the unique biology of non-traditional model species for successful applications of these techniques.

<table><tbody><tr><td>24-well plate</td><td>Corning</td><td>83-3738</td><td></td></tr><tr><td>Alt-R S.p. HiFi Cas9 Nuclease V3, 100 &amp;micro;g</td><td>Integrated DNA Technologies</td><td>1081060</td><td></td></tr><tr><td>Anti-static cleaner</td><td>Hozan</td><td>Z-292</td><td>for removing static electricity from a 24-well plate</td></tr><tr><td>Barrier Box 20.7L</td><td>AS ONE</td><td>4-5606-01</td><td>Large container</td></tr><tr><td>FemtoJet 4i</td><td>Eppendorf</td><td>5252000013</td><td>Electronic microinjector</td></tr><tr><td>Femtotip II, injection capillary</td><td>Eppendorf</td><td>5242957000</td><td>Glass injection capillary</td></tr><tr><td>High Pack 2440mL</td><td>AS ONE</td><td>5-068-25</td><td>Middle-sized container</td></tr><tr><td>Incubator</td><td>Panasonic</td><td>MIR-554-PJ</td><td>for 37 &amp;deg;C incubation. No need to humidify inside the incubator.</td></tr><tr><td>KOD Fx Neo</td><td>Toyobo</td><td>KFX-101</td><td>PCR enzyme for genotyping. Optimized for an amplification from crude templates.</td></tr><tr><td>Magnetic stand</td><td>Narishige</td><td>GJ-8</td><td>for holding the micromanipulator</td></tr><tr><td>Microloader</td><td>Eppendorf</td><td>5242956003</td><td></td></tr><tr><td>Micromanipulator</td><td>Narishige</td><td>MM-3</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td>SZX12</td><td>for microinjection. More than 35X magnification is sufficient for the microinjection</td></tr><tr><td>MultiNA</td><td>Shimadzu</td><td>MCE-202</td><td>Microchip electrophoresis system</td></tr><tr><td>NiceTac</td><td>Nichiban</td><td>NW-5</td><td>Double-sided tape to place eggs on a glass slide</td></tr><tr><td>Paint brush (horse hair)</td><td>Pentel</td><td>ZBS1-0</td><td></td></tr><tr><td>Plant culture dish</td><td>SPL Life Sciences</td><td>310100</td><td>Mating dish and water supplies for a large and middle-sized containers</td></tr><tr><td>Proteinase K, recombinant, PCR Grade Lyophilizate from Pichia pastoris</td><td>Roche</td><td>3115836001</td><td></td></tr><tr><td>SZX 12 microscope</td><td>Olympus</td><td>SZX 12</td><td>More than 35X magnification is sufficient for the microinjection</td></tr><tr><td>Talcum powder</td><td>Maruishi</td><td>877113</td><td></td></tr><tr><td>Tetra Goldfish Gold Growth</td><td>Spectrum Brands</td><td></td><td>Artificial regular fish food</td></tr></tbody></table>

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.

In our hands, about 100 eggs can be well injected with a single injection capillary when it has the adequate tip (Figure 3C). Injection of gRNA/Cas9 ribonucleoprotein complex in embryos within the first 8 h after egg laying results in indels at the gRNA targeted site. This causes biallelic mutations in some cells of the injected generation (G0) and thus mutant mosaic phenotypes are usually obtained in G0. For example, when this protocol was used to inject a gRNA that is designed to target the white gene, 32.6% of G0 nymphs display partial loss of pigmentation in their compound eyes and dorsal regions (Figure 5)8.
Using the presently described dry injection method, when 80–120 eggs are injected the survival rate of the injected embryos is as high as 40%–60%. This is in contrast with the previous wet injection method, in which eggs are injected and maintained on an agarose plate, occasionally resulting in a survival rate of less than 10%.
Assessment of the germline transformation of G0 adults and mutated G1 individuals was done by genomic PCR followed by HMA. In HMA, wildtype and mutant alleles anneal in each possible combination, which typically results in four distinct bands on a gel electrophoresis (two homoduplexes and two heteroduplexes)14. In G1 samples, differential band patterns between wildtype and mutated samples are clearly distinguishable (Figure 4B). Germline transformation was found in 39.1% of the G0 adults when we targeted the white gene8. In our experience, the percentage of mutated individuals in G1 nymphs from a single G0 pair varies from 25% to 100%.
To evaluate the effect of our mating environment on the mating success, we crossed wildtype adults in a mating dish and obtained a success rate of 95.8% (23/24 pairs).
<img alt="Insect life cycle diagram with developmental stages; image of female insect for study of molting cycles." class="xfigimg" src="/files/ftp_upload/61885/61885fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: The lifecycle of T. domestica. (A) Approximate duration of each developmental stage in T. domestica. (B) Dorsal view of an adult female. Arrowhead indicates a well-developed ovipositor. <a href="https://www.jove.com/files/ftp_upload/61885/61885fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Laboratory insect-rearing setups labeled A-D with water, food boxes, cotton, monitors in containers." class="xfigimg" src="/files/ftp_upload/61885/61885fig02v3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Artificial environments used in this protocol. (A) A large container for laboratory colony, (B) a medium-size container for egg collection, (C) mating dishes with water supplies in a large container, (C’) a mating dish, (D) a 24-well plate with water supply in the medium-size container. Boxed region is magnified in (D’) to show a T. domestica individual. <a href="https://www.jove.com/files/ftp_upload/61885/61885fig02v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Microinjection technique in embryo samples; setup, needle preparation, and injection results analysis." class="xfigimg" src="/files/ftp_upload/61885/61885fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Dry injection of T. domestica eggs. (A) Eggs aligned on a glass slide. Black arrow indicates the point where a needle is inserted. (B) Shape of a glass needle tip used for injection. (C) The same glass needle before (top) and after (bottom) breaking the tip. Needles were filled with 1% neutral red. Arrowhead indicates the tip of the needle. Scale bar is 1 mm. (D) Uninjected egg. (E) A good example of an injection. Arrow indicates the point of injection. (F–H) The solution is overflowing from the injected site (F) or from the opposite side (G) of the injected egg; too much volume of injection caused a burst (H). Arrowhead indicates overflowed egg content. (I) Normally developed late embryo. Arrowhead indicates the colored compound eye. (J) Shrunken damaged egg 3 days after injection. <a href="https://www.jove.com/files/ftp_upload/61885/61885fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="CRISPR gene editing diagram with primers, gRNA, electrophoresis results, and sequencing data." class="xfigimg" src="/files/ftp_upload/61885/61885fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Genotyping of G1 individuals. (A) PCR primers are designed to amplify a 120 bp genomic region that includes the gRNA target site. (B) Multiple bands of homoduplex and heteroduplex DNAs are detected in mutated samples while single bands appear in unmutated samples. L: DNA ladder, U: unmutated samples, M: mutated samples. (C) Representative result of direct sequencing of PCR products from wildtype and heterozygous mutant samples. Sequences of wildtype and mutant (<img alt="Static equilibrium diagram with triangle symbol showing Δ for moment calculations." src="/files/ftp_upload/61885/61885equ01v2.jpg" data-altupdatedat="1747911167000" data-alt="Equation 1">) allele are shown on top. The forward primer shown in (A) was used as a sequencing primer. The sequence of a heterozygous mutant (bottom) is indicated by two overlapping sequences from the predicted cleavage site (arrow). <a href="https://www.jove.com/files/ftp_upload/61885/61885fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Insect morphology comparison, diagram showing anatomical differences in segments A and B." class="xfigimg" src="/files/ftp_upload/61885/61885fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Mosaic loss of pigmentations in compound eyes and in the dorsal region after targeting the white gene with gRNA/Cas9 protein injection. (A) Wildtype and (B) white gRNA/Cas9 protein-injected first instar nymphs. Partial loss of black and pink pigmentations in eyes (arrowhead) and in the dorsal region (arrow) are indicated. Scale bar is 200 µm. <a href="https://www.jove.com/files/ftp_upload/61885/61885fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica at JoVE.com

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.

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 ...

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Nanyang Technological University

Research

JoVE Journal

Developmental Biology

4.3K Views.  Kyoto University. 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.

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

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. This is due to its reproductive behavior: females damage the external surface of fruits and vegetables when they lay eggs and the hatched larvae feed on their pulp. Wild C. capitata populations are traditionally controlled through insecticide spraying and/or eco-friendly approaches, the most successful being the Sterile Insect Technique (SIT). The SIT relies on mass-rearing, radiation-based sterilization and field release of males that retain their capacity to mate but are not able to generate fertile progeny. The advent and the subsequent rapid development of biotechnological tools, together with the availability of the medfly genome sequence, has greatly boosted our understanding of the biology of this species. This favored the proliferation of new strategies for genome manipulation, which can be applied to population control.
In this context, embryo microinjection plays a dual role in expanding the toolbox for medfly control. The ability to interfere with the function of genes that regulate key biological processes, indeed, expands our understanding of the molecular machinery underlying medfly invasiveness. Furthermore, the ability to achieve germ-line transformation facilitates the production of multiple transgenic strains that can be tested for future field applications in novel SIT settings. Indeed, genetic manipulation can be used to confer desirable traits that can, for example, be used to monitor sterile male performance in the field, or that can result in early life-stage lethality. Here we describe a method to microinject nucleic acids into medfly embryos to achieve these two main goals.

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. ...

Genetics

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex determination, venom synthesis, and host-symbiont interactions, among others. A major limitation of working with this organism is the lack of effective protocols to perform directed genome modifications. An important part of genome modification is delivery of editing reagents, including CRISPR/Cas9 molecules, into embryos through microinjection. While microinjection is well established in many model organisms, this technique is particularly challenging to perform in N. vitripennis primarily due to its small embryo size, and the fact that embryonic development occurs entirely within a parasitized blowfly pupa. The following procedure overcomes these significant challenges while demonstrating a streamlined, visual procedure for effectively removing wasp embryos from parasitized host pupae, microinjecting them, and carefully transplanting them back into the host for continuation and completion of development. This protocol will strongly enhance the capability of research groups to perform advanced genome modifications in this organism.

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Microinjection of Nasonia vitripennis embryos is an essential method for generating heritable genome modifications. Described here is a detailed procedure for microinjection and transplantation of Nasonia vitripennis embryos, which will greatly facilitate future genome manipulation in this organism.

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex ...

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

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

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

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

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

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

Injecting Gryllus bimaculatus Eggs

Altering gene function in a developing organism is central to different kinds of experiments. While tremendously powerful genetic tools have been developed in traditional model systems, it is difficult to manipulate genes or messenger RNA (mRNA) in most other organisms. At the same time, evolutionary and comparative approaches rely on an exploration of gene function in many different species, necessitating the development and adaptation of techniques for manipulating expression outside currently genetically tractable species. This protocol describes a method for injecting reagents into cricket eggs to assay the effects of a given manipulation on embryonic or larval development. Instructions for how to collect and inject eggs with beveled needles are described. This relatively straightforward technique is flexible and potentially adaptable to other insects. One can gather and inject dozens of eggs in a single experiment, and survival rates for buffer-only injections improve with practice and can be as high as 80%. This technique will support several types of experimental approaches including injection of pharmacological agents, in vitro capped mRNA to express genes of interest, double-stranded RNA (dsRNA) to achieve RNA interference, use of clustered regularly interspaced short palindromic repeats (CRISPR) in concert with CRISPR-associated protein 9 (Cas9) reagents for genomic modification, and transposable elements to generate transient or stable transgenic lines.

Injecting Gryllus bimaculatus Eggs

Here we present a protocol to inject cricket eggs, a technique which serves as a foundational method in many experiments in the cricket, including, but not limited to, RNA interference and genomic manipulation.

Altering gene function in a developing organism is central to different kinds of experiments. While tremendously powerful genetic tools have been ...

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

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

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

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

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

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

Bioengineering

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

Biology

Hyperactive piggyBac Transposase-mediated Germline Transformation in the Fall Armyworm, Spodoptera frugiperda

Stable insertion of genetic cargo into insect genomes using transposable elements is a powerful tool for functional genomic studies and developing genetic pest management strategies. The most used transposable element in insect transformation is piggyBac, and piggyBac-based germline transformation has been successfully conducted in model insects. However, it is still challenging to employ this technology in non-model insects that include agricultural pests. This paper reports on germline transformation of a global agricultural pest, the fall armyworm (FAW), Spodoptera frugiperda, using the hyperactive piggyBac transposase (hyPBase).
In this work, the hyPBase mRNA was produced and used in place of helper plasmid in embryo microinjections. This change led to the successful generation of transgenic FAW. Furthermore, the methods of screening transgenic animals, PCR-based rapid detection of transgene insertion, and thermal asymmetric interlaced PCR (TAIL-PCR)-based determination of the integration site, are also described. Thus, this paper presents a protocol to produce transgenic FAW, which will facilitate piggyBac-based transgenesis in FAW and other lepidopteran insects.

Hyperactive piggyBac Transposase-mediated Germline Transformation in the Fall Armyworm, Spodoptera frugiperda

Successful germline transformation in the fall armyworm, Spodoptera frugiperda, was achieved using mRNA of hyperactive piggyBac transposase.

Stable insertion of genetic cargo into insect genomes using transposable elements is a powerful tool for functional genomic studies and developing genetic ...

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 ...

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

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Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

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

Injecting Gryllus bimaculatus Eggs

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

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

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Hyperactive piggyBac Transposase-mediated Germline Transformation in the Fall Armyworm, Spodoptera frugiperda

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