The work was supported by the Scientific Research Fund for High-Level Talents, University of South China.

1. Preparation of plasmids
<ol>
	<li>Isolate plasmids from 4 mL overnight bacterial cultures using a plasmid miniprep kit. Elute with 40 &#181;L of elution buffer. 
	NOTE: Isolation of plasmids using a plasmid midi-prep kit is not necessary. A plasmid miniprep kit can fully meet the experimental requirements for embryo microinjection. In this protocol, the prepared UAS-cDNA/ORF plasmids contain ten copies of UAS, an Hsp70 minimal promoter, an attB site, a mini-white gene, and a gene of interest.</li>
	<li>Determine the plasmid DNA concentration using a spectrophotometer and store it at -80 &#176;C freezer. 
	NOTE: Plasmid concentration is not critical for successful embryo microinjection, as plasmid concentrations ranging from 20 ng/&#181;L to 1683 ng/&#181;L work well for transgenesis in Drosophila (Supplementary Table 1).</li>
	<li>Just before microinjection, thaw plasmids and centrifuge at 13000 x g for 5 min.</li>
	<li>Transfer 10 &#181;L of each plasmid from the upper layer of the liquid into a new 1.5 mL tube.</li>
</ol>
2. Pulling needles
<ol>
	<li>Turn on a needle puller with selected settings (Heat: 500, Pull: 50, Vel: 60, Delay: 90, Pressure: 200, Ramp Test: 510).</li>
	<li>Insert a borosilicate glass capillary (outer diameter 1.0 mm, inner diameter 0.75 mm, 10 cm length) into the needle puller and close the lid.</li>
	<li>Press the Pull button. Each glass capillary makes two needles.</li>
</ol>
3. Grinding needles (Figure 1)
<ol>
	<li>Use a needle grinder mounted with an appropriate abrasive plate (for 0.7-2.0 &#181;m tip sizes) of a needle grinder.</li>
	<li>Wet the cloth (reference wire) with grinding solution (100 mL of distilled water with 0.9 g of NaCl, 1 mL of wetting agent) and fix the cloth.</li>
	<li>Turn on the needle grinder and keep the grinding plate rotating.</li>
	<li>Connect a needle to a foot air pump with a pressure gauge via a silicone capillary tube, and then mount the needle onto a needle holder of the needle grinder. Adjust the angle of the needle relative to the grinding plate to 35&#176; (Figure 1A, B).</li>
	<li>Adjust the position of the needle tip just above the grinding plate surface by using a coarse control knob under the microscope (Figure 1C). 
	NOTE: The pulled needles obtained in section 2 have a 6-8 mm taper and a less than 1 &#181;m tip (Figure 1D), as is described in the user manual of the puller.</li>
	<li>Keep the inner pressure of the needle at 40 psi using the foot air pump. Adjust the position of the needle tip by using a fine control knob under the microscope, and make the tip of the needle touch the grinding plate.</li>
	<li>Grind for 3-5&#160;s. When tiny air bubbles come out of the needle tip, unload the needle (Figure 1E). 
	NOTE: In the process of grinding, using an air pump prevents the grinding solution from siphoning.</li>
	<li>Put the ground needles into a storage case. 
	NOTE: Use freshly ground needles as much as possible, though the ground needles can be kept in a storage case for at least a few weeks before microinjection.</li>
</ol>
4. Preparing pipettes for loading plasmids
<ol>
	<li>Heat a 200 &#181;L pipette with the outer flame of an alcohol burner.</li>
	<li>Once it starts to melt, stretch the pipette.</li>
	<li>Cut off the sealed end of the stretched pipette with scissors, and the stretched pipette is then ready to load DNA into a ground needle. 
	NOTE: Alternatively, extremely long and fine tips for filling ground needles are commercially available.</li>
	<li>Place these pipettes in an autoclaved plastic box.</li>
</ol>
5. Collecting embryos
<ol>
	<li>Collect flies (vas-phiC31; + ; attP2) expressing phiC31 integrase and bearing the attP2 docking site 3 days after they are hatched from 50 vials and transfer them to a cage (&#966; 90 mm x 150 mm). Add a little fresh yeast paste to the center of each grape agar plate18.</li>
	<li>Prepare 2-4 cages of flies. Keep the cages and plates at 25 &#176;C and change the plates every day for 2-4 days. 
	NOTE: This study combined the vas-phiC31 transgene (Stock # 36313, Bloomington Drosophila Stock Center) on the first chromosome and the attP2 transgene (Stock # 36313, Bloomington Drosophila Stock Center) on the third chromosome. The resulting stock (vas-phiC31; + ; attP2) is used in this protocol. Yeast paste needs to be prepared daily, and fresh yeast paste must be fed to the flies. The replaced cages need to be cleared and dried to get rid of the remaining embryos in the cages.</li>
	<li>On the day of microinjection, change the cages and plates in the morning. Then, change the plates every 30 min and collect embryos from these plates. Change all the cages and plates at room temperature (RT) and afterward keep the cages and plates at 25 &#176;C. 
	NOTE: Some protocols prefer to incubate the cages and plates at 18 &#176;C for egg laying. It depends on the number of embryos one needs for microinjection.</li>
</ol>
6. Dechorionating embryos
<ol>
	<li>Prepare 30% bleach by mixing 60 mL of bleach (containing active chlorine content of 34-46 g/L) and 140 mL of double distilled water.</li>
	<li>Remove the remaining flies on the grape juice agar plates with a brush or forceps, and rinse embryos into a cell strainer with double distilled water.</li>
	<li>Dry the cell strainer containing the embryos with tissue paper.</li>
	<li>Place it in a Petri dish containing 30% bleach, and swirl the cell strainer by hand or a horizontal shaker for 1.5 min at 60 rpm. 
	NOTE: Rinsing is a critical step. If the rinsing time is not long enough, the shell of an embryo will not come off completely. If the rinsing time is too long, it will cause the embryo to become soft and break easily. The cell strainer should be completely submerged in the 30% bleach. Wear a pair of rubber gloves and a lab coat to avoid injuries caused by bleach.</li>
	<li>Rinse embryos for 2 min with double distilled water stored in a laboratory plastic wash bottle to remove the residual bleach. Pick the floating embryos for the line-up.</li>
</ol>
7. Lining up embryos
<ol>
	<li>Cut the solid grape juice agar into long strips using a blade. 
	NOTE: Use purple grape juice to make it easier to line up white embryos.</li>
	<li>Transfer the embryos onto a long strip using a paintbrush.</li>
	<li>Line up around 60 embryos with a dissecting needle along the edge of a long strip with their posteriors facing the outside of the strip. Line up around 60 embryos within 5 min. 
	NOTE: The micropyle is a cone-shaped protrusion at the anterior pole of the embryo, through which the sperm enters to fertilize the oocyte (Figure 2A). Therefore, based on the cone-shaped protrusion at the anterior pole of the embryo, it is easy to determine the posterior of the dechorionated embryo.</li>
	<li>Place an 18 mm double-sided tape lengthwise along the edge of a coverslip (18 mm x 18 mm) and remove the paper backing. Gently press the coverslip onto the aligned embryos. Ensure the embryos are glued to the center of the tape.</li>
</ol>
8. Desiccating embryos
<ol>
	<li>Place the coverslip with the lined-up embryos in a box containing desiccant. The drying time varies depending on temperature and humidity. 
	NOTE: The drying time is critical for microinjection and needs to be experimentally determined. The drying time is usually 9-10 min in summer and 17-18 min in winter.</li>
	<li>Cover the embryos with halocarbon oil. 
	NOTE: Do not drop too much oil onto the embryos, as the extra oil may overflow the adhesive. The volume ratio of halocarbon oil 700 and halocarbon oil 27 is 1:3.</li>
</ol>
9. Microinjecting embryos
<ol>
	<li>Load 2 &#181;L of DNA into the needle by using the stretched pipette. Ensure that there are no bubbles in the needle. Place the needle into the needle holder of the micromanipulator.</li>
	<li>Place the aligned embryos covered with halocarbon oil under an inverted microscope. Ensure that the posterior ends of the embryos face the needle.</li>
	<li>Use the micromanipulator to bring the needle close to the embryos. Clear the needle. Ensure that a drop of liquid coming out of the needle is visible. Starting from 300 hPa injection pressure and 20 hPa equilibrium pressure, adjust the pressure of injection and the pressure of equilibrium to ensure the optimal size of the drop of liquid.</li>
	<li>Use the micromanipulator to insert the needle into the posterior ends of the embryos, and use the foot control connected to the microinjector to inject the DNA (Figure 2). If a small halo is visible, DNA is successfully injected into the embryo. 
	NOTE: Select the multinucleate embryos for injection and kill the older embryos using the &#34;clean&#34; button. If a large amount of cytoplasm leaks after injection, embryos are under-dried. If only a small amount of or no amount of cytoplasm leaks after injection, injections are considered successful. If more than one-third of the embryos leak cytoplasm after injection, extend the drying time by 1-2 min. If more than one-third of the embryos are over-dried, shorten the drying time by 1-2 min.</li>
	<li>Remove the needle from the embryo. Move the needle to the posterior end of the next embryo. Repeat the above steps.</li>
	<li>Put the injected embryos into an agar plate. Place the plate in a room at 18-20 &#176;C. On the next day, transfer the plate to an incubator at a temperature of 25 &#176;C and a humidity of 50%-60%, and incubate for 1 day. 
	NOTE: After injection, keep embryos at a temperature of 18-20 &#176;C to slow down their development, facilitate the integration of plasmid DNA into the Drosophila genome, and increase the survival rate of the injected embryos.</li>
</ol>
10. Collecting larvae
<ol>
	<li>Use a dissecting needle to scrape and loosen the food in a vial. Supply the food with a few drops of distilled water. 
	NOTE: Allow the distilled water to be absorbed into the food to prevent larvae from drowning.</li>
	<li>Take out the plates from the incubator.</li>
	<li>Collect the hatched larvae with a paintbrush under a stereo microscope and transfer them to the vial.</li>
	<li>Place the vial into an incubator at a temperature of 25 &#176;C and a humidity of 50%-60%.</li>
</ol>
11. Obtaining transgenic flies
<ol>
	<li>Collect the hatched flies after the vial is incubated at 25 &#176;C for 10 days.</li>
	<li>Cross a single male fly to three virgin female flies of the TM2/TM6C Sb1 Tb1 balancers, and cross a single female fly to three male flies of the TM2/TM6C Sb1 Tb1 balancers.</li>
	<li>Place the crosses into an incubator at a temperature of 25 &#176;C and a humidity of 50%-60%.</li>
	<li>Screen the progeny for flies with orange eyes. These are the transgenic flies (Figure 3A-C). 
	NOTE: Flies bearing a copy of transgene at the attP2 docking site have orange eyes.</li>
	<li>Cross a single orange-eyed male fly to three virgin female flies of the TM2/TM6C Sb1 Tb1 balancers to establish a stock. 
	NOTE: More than one generation of crosses need to be set up in order to cross out the phiC31 transgene on the first chromosome.</li>
</ol>

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

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	<li>Rubin, G. M., Spradling, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+transformation+of+Drosophila+with+transposable+element+vectors.">Genetic transformation of Drosophila with transposable element vectors.</a> Science. 218 (4570), 348-353 (1982).</li><li>Venken, K. J., Bellen, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transgenesis+upgrades+for+Drosophila+melanogaster.">Transgenesis upgrades for Drosophila melanogaster.</a> Development. 134 (20), 3571-3584 (2007).</li><li>Groth, A. C., Fish, M., Nusse, R., Calos, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Construction+of+transgenic+Drosophila+by+using+the+site-specific+integrase+from+phage+phic31.">Construction of transgenic Drosophila by using the site-specific integrase from phage phic31.</a> Genetics. 166 (4), 1775-1782 (2004).</li><li>Bischof, J., Maeda, R. K., Hediger, M., Karch, F., Basler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optimized+transgenesis+system+for+Drosophila+using+germ-line-specific+phic31+integrases.">An optimized transgenesis system for Drosophila using germ-line-specific phic31 integrases.</a> Proc Natl Acad Sci U S A. 104 (9), 3312-3317 (2007).</li><li>Markstein, M., Pitsouli, C., Villalta, C., Celniker, S. E., Perrimon, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exploiting+position+effects+and+the+gypsy+retrovirus+insulator+to+engineer+precisely+expressed+transgenes.">Exploiting position effects and the gypsy retrovirus insulator to engineer precisely expressed transgenes.</a> Nat Genet. 40 (4), 476-483 (2008).</li><li>Bateman, J. R., Lee, A. M., Wu, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Site-specific+transformation+of+Drosophila+via+phic31+integrase-mediated+cassette+exchange.">Site-specific transformation of Drosophila via phic31 integrase-mediated cassette exchange.</a> Genetics. 173 (2), 769-777 (2006).</li><li>Venken, K. J., He, Y., Hoskins, R. A., Bellen, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=P[acman]:+A+BAC+transgenic+platform+for+targeted+insertion+of+large+DNA+fragments+in+D.+melanogaster.">P[acman]: A BAC transgenic platform for targeted insertion of large DNA fragments in D. melanogaster.</a> Science. 314 (5806), 1747-1751 (2006).</li><li>Szabad, J., Bellen, H. J., Venken, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+assay+to+detect+in+vivo+Y+chromosome+loss+in+Drosophila+wing+disc+cells.">An assay to detect in vivo Y chromosome loss in Drosophila wing disc cells.</a> G3. 2 (9), 1095-1102 (2012).</li><li>Ringrose, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transgenesis+in+Drosophila+melanogaster.">Transgenesis in Drosophila melanogaster.</a> Methods Mol Biol. 561, 3-19 (2009).</li><li>Van Der Graaf, K., Srivastav, S., Singh, P., Mcnew, J. A., Stern, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+melanogaster+attp40+docking+site+and+derivatives+are+insertion+mutations+of+msp-300.">The Drosophila melanogaster attp40 docking site and derivatives are insertion mutations of msp-300.</a> PLoS One. 17 (12), e0278598 (2022).</li><li>Duan, Q., Estrella, R., Carson, A., Chen, Y., Volkan, P. 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+effect+of+Drosophila+attp40+background+on+the+glomerular+organization+of+or47b+olfactory+receptor+neurons.">The effect of Drosophila attp40 background on the glomerular organization of or47b olfactory receptor neurons.</a> G3. 13 (4), 022 (2023).</li><li>Dietzl, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+genome-wide+transgenic+RNAi+library+for+conditional+gene+inactivation+in+Drosophila.">A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila.</a> Nature. 448 (7150), 151-156 (2007).</li><li>Pfeiffer, B. D., 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=Tools+for+neuroanatomy+and+neurogenetics+in+Drosophila.">Tools for neuroanatomy and neurogenetics in Drosophila.</a> Proc Natl Acad Sci U S A. 105 (28), 9715-9720 (2008).</li><li>Bellen, H. 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=The+Drosophila+gene+disruption+project:+Progress+using+transposons+with+distinctive+site+specificities.">The Drosophila gene disruption project: Progress using transposons with distinctive site specificities.</a> Genetics. 188 (3), 731-743 (2011).</li><li>Zirin, 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=Large-scale+transgenic+drosophila+resource+collections+for+loss-+and+gain-of-function+studies.">Large-scale transgenic drosophila resource collections for loss- and gain-of-function studies.</a> Genetics. 214 (4), 755-767 (2020).</li><li>Wei, P., Xue, W., Zhao, Y., Ning, G., Wang, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR-based+modular+assembly+of+a+UAS-cDNA/ORF+plasmid+library+for+more+than+5500+Drosophila+genes+conserved+in+humans.">CRISPR-based modular assembly of a UAS-cDNA/ORF plasmid library for more than 5500 Drosophila genes conserved in humans.</a> Genome Res. 30 (1), 95-106 (2020).</li><li>Fish, M. P., Groth, A. C., Calos, M. P., Nusse, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Creating+transgenic+Drosophila+by+microinjecting+the+site-specific+phic31+integrase+mRNA+and+a+transgene-containing+donor+plasmid.">Creating transgenic Drosophila by microinjecting the site-specific phic31 integrase mRNA and a transgene-containing donor plasmid.</a> Nat Protoc. 2 (10), 2325-2331 (2007).</li><li>Fujioka, M., Jaynes, J. B., Bejsovec, A., Weir, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production+of+transgenic+Drosophila.">Production of transgenic Drosophila.</a> Methods Mol Biol. 136, 353-363 (2000).</li><li>Loncar, D., Singer, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+membrane+formation+during+the+cellularization+of+the+syncytial+blastoderm+of+Drosophila.">Cell membrane formation during the cellularization of the syncytial blastoderm of Drosophila.</a> Proc Natl Acad Sci U S A. 92 (6), 2199-2203 (1995).</li><li>Carmo, C., Araujo, M., Oliveira, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+techniques+in+fly+embryos+to+study+the+function+and+dynamics+of+SMC+complexes.">Microinjection techniques in fly embryos to study the function and dynamics of SMC complexes.</a> Methods Mol Biol. 2004, 251-268 (2019).</li></ol>

The authors have nothing to disclose.

Here, we present a protocol for embryo microinjection for transgenesis in Drosophila. For phiC31-mediated site-specific transgenesis, we injected 120-140 embryos for each plasmid and obtained at least one transgenic line for around 85% of plasmids (Supplementary Table 2). In our experience, plasmid DNA isolated by a plasmid miniprep kit suffices for transgenesis in Drosophila. Plasmid DNA concentrations ranging from 20 ng/&#181;L to 1683 ng/&#181;L might have no apparent effect on the s...

The fruit fly Drosophila melanogaster is extremely amenable to genetic manipulation and genetic analysis. Transgenic fruit flies are widely used in biological research. Since it was developed in the early 1980s, P-element transposon-mediated transgenesis has been indispensable for Drosophila research1. In some scenarios, other transposons, such as piggyBac and Minos have been used for transgenesis in Drosophila as well2. Transgenes via transposon are randomly inserted into the Drosophila genome, and the expression levels of transgenes at different genomic loci may vary due to position effects and thereby can not be compared2. These drawbacks were overcome by the phiC31-mediated site-specific transgenesis3. The bacteriophage&#160;phiC31 gene encodes an integrase that mediates sequence-specific recombination between the attB and attP sites in Drosophila3. Many attP docking sites have been characterized and are available in the Bloomington Drosophila Stock Center3,4,5,6,7,8,9.
The phiC31 transgenesis system for Drosophila has been widely used by the fly community. Among the attP docking sites, attP18 on the X chromosome, attP40 on the second chromosome, and attP2 on the third chromosome are usually the preferred sites for transgene integration on each chromosome because transgenes at the three sites show high levels of inducible expression while having low levels of basal expression5. Recently, however, van der Graaf and colleagues found that the attP40 site, close to the Msp300 gene locus, and transgenes integrated at attP40 cause larval muscle nuclear clustering in Drosophila10. In another recent study, Duan and colleagues found that the homozygous attP40 chromosome disrupts the normal glomerular organization of the Or47b olfactory receptor neuron class in Drosophila11. These findings suggest that rigorous controls should be included when designing experiments and interpreting data.
Drosophila is an ideal model organism for in vivo interrogation of gene function due to its short life cycle, low cost, and easy maintenance. Genome-wide genetic screening relies on the construction of genome-wide transgenic libraries in Drosophila12,13,14,15. We previously generated 5551 UAS-cDNA/ORF constructs based on the binary GAL4/upstream activating sequence (GAL4/UAS) system, covering 83% of the Drosophila genes conserved in humans in the Drosophila Genomics Resource Center (DGRC) Gold Collection16. The UAS-cDNA/ORF plasmids can be used for the creation of a transgenic UAS-cDNA/ORF library in Drosophila. Efficient embryo microinjection technology can accelerate the creation of transgenic fly libraries.
For embryo microinjection, the needle tip is normally broken against a coverslip17. Thereby, needles frequently become clogged or cause leakage of large amounts of cytoplasm, and when these issues arise, new needles must be employed. Although beveling needles can enhance microinjection efficiency, the grinding solution enters the beveled tip during the grinding process. The grinding solution in the beveled tip does not quickly evaporate naturally; therefore, needles can not be utilized for microinjection directly after beveling. These factors reduce the efficiency of embryo microinjection procedures. Here, using the phiC31-mediated site-specific transgenesis in Drosophila as an example, we present a protocol for embryo microinjection for transgenesis in Drosophila.&#160;To prevent the grinding solution from entering the needle, we utilize an air pump to exert air pressure within the needle, thereby preventing the grinding solution from entering the beveled tip. Beveled needles are ready for sample loading and microinjection directly after beveling.

<table><tbody><tr><td>100 &amp;mu;m Cell Strainer</td><td>NEST</td><td>258367</td><td></td></tr><tr><td>3M double-sided adhesive</td><td>3M</td><td>415</td><td></td></tr><tr><td>Borosilicate glass</td><td>SUTTER</td><td>B100-75-10</td><td></td></tr><tr><td>Diamond abrasive plate</td><td>SUTTER</td><td>104E</td><td></td></tr><tr><td>FLAMING/BROWN Micropipette Puller</td><td>SUTTER</td><td>P-1000</td><td></td></tr><tr><td>Foot air pump with pressure gauge</td><td>Shenfeng</td><td>SF8705D</td><td></td></tr><tr><td>Halocarbon oil 27</td><td>Sigma</td><td>H8773</td><td></td></tr><tr><td>Halocarbon oil 700</td><td>Sigma</td><td>H8898</td><td></td></tr><tr><td>Inverted microscope</td><td>Nikon</td><td>ECLIPSE Ts2R</td><td></td></tr><tr><td>Microinjection pump</td><td>eppendorf</td><td>FemtoJet 4i</td><td></td></tr><tr><td>Micromanipulator</td><td>eppendorf</td><td>TransferMan 4r</td><td></td></tr><tr><td>Micropipette Beveler</td><td>SUTTER</td><td>BV-10</td><td></td></tr><tr><td>Microscope cover glasses (18 mm x 18 mm)</td><td>CITOLAS</td><td>10211818C</td><td></td></tr><tr><td>Microscope slides (25 mm x 75 mm)</td><td>CITOLAS</td><td>188105W</td><td></td></tr><tr><td>Petri dish (90 mm x 15 mm)</td><td>LAIBOER</td><td>4190152</td><td></td></tr><tr><td>Photo-Flo 200</td><td>Kodak</td><td>1026269</td><td></td></tr><tr><td>QIAprep Spin Miniprep Kit</td><td>Qiagen</td><td>27106</td><td></td></tr><tr><td>Stereo Microscope</td><td>Nikon</td><td>SMZ745</td><td></td></tr></tbody></table>

embryo microinjection for transgenesis in drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the construction of transgenic flies. Microinjection requires some types of equipment, including a microinjector, a micromanipulator, an inverted microscope, and a stereo microscope.&#160;Plasmids isolated with a plasmid miniprep kit are qualified for microinjection. Embryos at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm, are subjected to microinjection. A cell strainer eases the process of dechorionating embryos. The optimal time for dechorionation and desiccation of embryos needs to be determined experimentally. To increase the efficiency of embryo microinjection, needles prepared by a puller need to be beveled by a needle grinder. In the process of grinding needles, we utilize a foot air pump with a pressure gauge to avoid the capillary effect of the needle tip. We routinely inject 120-140 embryos for each plasmid and obtain at least one transgenic line for around 85% of plasmids. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and&#160;presents a detailed protocol for embryo microinjection for transgenesis in Drosophila.

The tip of an injection needle is beveled by a needle grinder (Figure 1). One can bevel 50-60 needles in one hour. DNA is microinjected into the posterior of an embryo at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm (Figure 2A). The posterior of an embryo can be readily located based on the micropyle at the anterior of the embryo (Figure 2A). Embryos at the cellular blastoderm and gastrulat...

Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection,&#160;a crucial step for creating transgenic flies.

Embryo Microinjection for Transgenesis in Drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the ...

embryo-microinjection-for-transgenesis-in-drosophila

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1.7K Views.  University of South China. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection, a crucial step for creating transgenic flies.

Video: Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

Biology

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

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

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

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

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

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

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

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

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

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

Microinjection Method for Anopheles gambiae Embryos

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA fragments encoding genes of interest as well as favorable traits into the insect germline in a stable and heritable manner. The resulting transgenic strains can be studied for phenotypic changes resulting from the expression of the integrated DNA to answer basic questions or used in practical applications. Although the technology is straightforward, it requires of the investigator patience and practice to achieve a level of skill that maximizes efficiency. Shown here is a method for microinjection of embryos of the African malaria mosquito, Anopheles gambiae. The objective is to deliver by microinjection exogenous DNA to the embryo so that it can be taken up in the developing germline (pole) cells. Expression from the injected DNA of transposases, integrases, recombinases, or other nucleases (for example CRISPR-associated proteins, Cas) can trigger events that lead to its covalent insertion into chromosomes. Transgenic An. gambiae generated from these technologies have been used for basic studies of immune system components, genes involved in blood-feeding, and elements of the olfactory system. In addition, these techniques have been used to produce An. gambiae strains with traits that may help control the transmission of malaria parasites.

Microinjection Method for Anopheles gambiae Embryos

Microinjection techniques are essential to introduce exogenous genes into the genomes of mosquitoes. This protocol explains a method used by the James laboratory to microinject DNA constructs into Anopheles gambiae embryos to generate transformed mosquitoes.

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

Developmental Biology

RNAi Interference by dsRNA Injection into Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements and tools for genetic manipulation have become available in Drosophila 2. Soon after the initial discovery by Frie and Mello 3 that double stranded RNA can be used to knockdown the activity of individual genes in Caenorhabditis elegans, RNA interference (RNAi) was shown to provide a powerful reverse genetic approach to analyze gene functions in Drosophila organ development 4, 5.
Many organs, including lung, kidney, liver, and vascular system, are composed of branched tubular networks that transport vital fluids or gases 6, 7. The analysis of Drosophila tracheal formation provides an excellent model system to study the morphogenesis of other tubular organs 8. The Berkeley Drosophila genome project has revealed hundreds of genes that are expressed in the tracheal system. To study the molecular and cellular mechanism of tube formation, the challenge is to understand the roles of these genes in tracheal development. Here, we described a detailed method of dsRNA injection into Drosophila embryo to knockdown individual gene expression. We successfully knocked down endogenous dysfusion(dys) gene expression by dsRNA injection. Dys is a bHLH-PAS protein expressed in tracheal fusion cells, and it is required for tracheal branch fusion 9, 10. dys-RNAi completely eliminated dys expression and resulted in tracheal fusion defect. This relatively simple method provides a tool to identify genes requried for tissure and organ development in Drosophila.

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNA interference has been proven very effective to analyze gene function in Drosophila tracheal development. A detailed protocol used by Jiang lab to inject dsRNA into fly embryos to knockdown gene expression is illustrated. This technique has the potential for screening genes required for tissue and organ development in Drosophila.

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements ...

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

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

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

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

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

Bioengineering

Microinjection of Drosophila Nurse Cells: A Method of Intracellular Delivery

Source: Catrina, I. E., et al. <a href="https://www.jove.com/video/58545/" target="_blank">Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers</a>. J. Vis. Exp. (2019).
Due to their large size and relatively simple organization, Drosophila nurse cells are ideally suited for live cell imaging studies. This video describes a microinjection method for exogenous compounds delivery into the cells, and the featured protocol demonstrates the procedure with fluorescent oligonucleotide probes called molecular beacons (MBs) used to visualize endogenous mRNA transcripts.

Source: Catrina, I. E., et al. Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers. J. Vis. Exp. (2019).
Due to their ...

JoVE Encyclopedia of Experiments

Drosophila melanogaster (fruit fly)

Embryo

Microinjection of Live Drosophila Embryos: Early Delivery of Reagents to the Developing Embryo

Source: Brust-Mascher, I., Scholey, J. M. <a href="https://www.jove.com/video/1382/" target="_blank">Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo</a>. J. Vis. Exp. (2009).
This video describes how to perform microinjections that deliver reagents into live Drosophila embryos. Researchers can use this method to introduce exogenous compounds like nucleic acids and proteins during the early stages of embryo development. The featured protocol demonstrates how to set up and perform the procedure for injection of any soluble reagent&#8211;here, a fluorescent protein for live imaging experiments.

Source: Brust-Mascher, I., Scholey, J. M. Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo. J. Vis. Exp. ...

Embryo Microinjection for Transgenesis in Drosophila

Summary

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Chapters in this video

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

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

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

Microinjection Method for Anopheles gambiae Embryos

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

RNAi Interference by dsRNA Injection into Drosophila Embryos

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

Microinjection of Drosophila Nurse Cells: A Method of Intracellular Delivery

Microinjection of Live Drosophila Embryos: Early Delivery of Reagents to the Developing Embryo