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. dome...

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 &#181;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 &#176;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>
		</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 th...

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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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

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental studies. Like other aphids, A. pisum propagate rapidly via parthenogenetic viviparous reproduction, where the embryos develop within egg chambers in an assembly-line fashion in the ovariole. Previously we have established a robust platform of whole-mount in situ hybridization allowing detection of mRNA expression in the aphid embryos. For analyzing the expression of protein, though, established protocols for immunostaining the ovarioles of asexual viviparous aphids did not produce satisfactory results. Here we report conditions optimized for increasing tissue permeability and decreasing background staining, both of which were problems when applying established approaches. Optimizations include: (1) incubation of proteinase K (1 &#181;g/ml, 10 min), which was found essential for antibody penetration in mid- and late-stage aphid embryos; (2) replacement of normal goat serum/bovine serum albumin with a blocking reagent supplied by a Digoxigenin (DIG)-based buffer set and (3) application of methanol rather hydrogen peroxide (H2O2) for bleaching endogenous peroxidase; which significantly reduced the background staining in the aphid tissues. These critical conditions optimized for immunostaining will allow effective detection of gene products in the embryos of A. pisum and other aphids.

A protocol of whole-mount immunostaining on aphid embryos is presented, in which critical conditions for decreasing background staining and increasing tissue permeability are addressed. These conditions are specifically developed for effective detection of protein expression in the embryonic tissues of aphids.

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Collection of Post-mating Semen from the Female Reproductive Tract and Measurement of Semen Liquefaction in Mice

In mice, ejaculated semen is deposited in the uterus. After ejaculation, the semen changes consistency from gel-like to watery, a process called liquefaction. In this study, we show how to collect the post-ejaculated semen from the female reproductive tract in a mouse model. First, adult female mice in the estrus stage were housed in a male&#39;s cage overnight. The next morning, copulation was confirmed by the presence of copulatory plug at the vaginal opening. Female mice with copulatory plugs were euthanized, and each reproductive tract was collected as a whole (vagina, uterus, oviducts, ovaries), ensuring a closed system to contain the semen. The reproductive tract was placed in a 1.5 mL microcentrifuge tube, and the vagina was cut off to release the semen into the tube. To ensure maximum semen volume for analysis, toothless forceps were used to squeeze the uterine horns from ovarian end to vaginal end expelling remaining semen. The whole reproductive tract was then discarded. The semen-containing tube was briefly spun down. A 25 &#956;L capillary pipette was placed into the tube at a 180&#176; angle (parallel to the tube wall). The amount of time used to fill the capillary tube to the 25 &#956;L line was recorded. Semen from a proven male breeder usually takes approximately 60-180 s to fill a 25 &#956;L capillary tube. This semen collection technique can also be used in other downstream applications such as sperm imaging and motility analysis.

Semen liquefaction is required for sperm to be liberated from the seminal gel. This study provides the procedures for collecting semen from the female reproductive tract post-coitus, and measuring semen liquefaction time.

In mice, ejaculated semen is deposited in the uterus. After ejaculation, the semen changes consistency from gel-like to watery, a process called ...

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of cells or tissues in model organisms. These systems contain two components as separate transgenic lines. A driver line expresses a transcriptional activator under the control of tissue-specific promoters/enhancers, and a reporter/effector line harbors a target gene placed downstream to the binding site of the transcription activator. Animals harboring both components induce tissue-specific transactivation of a target gene expression. Precise spatiotemporal expression of the gene in targeted tissues is critical for unbiased interpretation of cell/gene activity. Therefore, developing a method for generating exclusive cell/tissue-specific driver lines is essential. Here we present a method to generate highly tissue-specific targeted expression system by employing a &#34;Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR-associated&#34; (CRISPR/Cas)-based genome editing technique. In this method, the endonuclease Cas9 is targeted by two chimeric guide RNAs (gRNA) to specific sites in the first coding exon of a gene in the Drosophila genome to create double-strand breaks (DSB). Subsequently, using an exogenous donor plasmid containing the transactivator sequence, the cell-autonomous repair machinery enables homology-directed repair (HDR) of the DSB, resulting in precise deletion and replacement of the exon with the transactivator sequence. The knocked-in transactivator is expressed exclusively in cells where the cis-regulatory elements of the replaced gene are functional. The detailed step-by-step protocol presented here for generating a binary transcriptional driver expressed in Drosophila fgf/branchless-producing epithelial/neuronal cells can be adopted for any gene- or tissue-specific expression.

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Here, we present a method for generating tissue-specific binary transcription systems in Drosophila by replacing the first coding exon of genes with transcription drivers. The CRISPR/Cas9-based method places a transactivator sequence under the endogenous regulation of a replaced gene, and consequently facilitates transctivator expression exclusively in gene-specific spatiotemporal patterns.

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

Brain Organoid Generation from Induced Pluripotent Stem Cells in Home-Made Mini Bioreactors

The iPSC-derived brain organoid is a promising technology for in vitro modeling the pathologies of the nervous system and drug screening. This technology has emerged recently. It is still in its infancy and has some limitations unsolved yet. The current protocols do not allow obtaining organoids to be consistent enough for drug discovery and preclinical studies. The maturation of organoids can take up to a year, pushing the researchers to launch multiple differentiation processes simultaneously. It imposes additional costs for the laboratory in terms of space and equipment. In addition, brain organoids often have a necrotic zone in the center, which suffers from nutrient and oxygen deficiency. Hence, most current protocols use a circulating system for culture medium to improve nutrition.
Meanwhile, there are no inexpensive dynamic systems or bioreactors for organoid cultivation. This paper describes a protocol for producing brain organoids in compact and inexpensive home-made mini bioreactors. This protocol allows obtaining high quality organoids in large quantities.

Here we describe a protocol for generating brain organoids from human induced pluripotent stem cells (iPSCs). To obtain brain organoids in large quantities and of high quality, we use home-made mini bioreactors.

The iPSC-derived brain organoid is a promising technology for in vitro modeling the pathologies of the nervous system and drug screening. This technology ...

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

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Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Technique to Target Microinjection to the Developing Xenopus Kidney

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Collection of Post-mating Semen from the Female Reproductive Tract and Measurement of Semen Liquefaction in Mice

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Brain Organoid Generation from Induced Pluripotent Stem Cells in Home-Made Mini Bioreactors