We are grateful to Drusilla Stillinger, Kiona Parker, Parrish Powell and Madeline Nottoli for mosquito husbandry. Funding was provided by the University of California, Irvine Malaria Initiative. AAJ is a Donald Bren Professor at the University of California, Irvine.

1. Preparing mosquitoes for microinjection
<ol>
	<li>Seed a cage13 (~5000 cm3) with ~100 male and 200-300 female 1-2 day adult post-eclosion mosquitoes and allow them to mate for 2 days.</li>
	<li>After the mating period, provide mosquitoes a blood meal using either 2 mL of blood with an artificial feeding device or live anesthetized animals depending on insectary practices14. The following day provide mosquitoes a second blood meal to ensure that all mated females have had an opportunity to feed and that partially-fed mosquitoes have become fully-engorged.</li>
	<li>Utilize the mosquitoes for embryo collections 2-4 days after the second blood meal. 
	&#8203;NOTE: Larval nutrition is important for adult robustness and reproductive fitness. See Benedict et al. (2020) for information about how to rear healthy insects14. No significant differences in egg laying have been observed when mosquitoes are fed on an artificial feeder or live anesthetized animals. Use whichever method is routinely used for Anopheles gambiae in the insectary.</li>
</ol>
2. Embryo preparation
<ol>
	<li>Use an aspirator to place 20-30 females in a transparent 50 mL conical tube that has been cut (with a band saw) to be open at both ends and is covered with latex dental film at one end and nylon mesh and filter paper at the other (Figure 1). 
	NOTE: The mosquitoes will deposit their eggs onto the filter paper and the nylon mesh will keep the filter paper securely in place. Mosquito eggs are cylindrical in shape, ~500 &#181;m in length, ~200 &#181;m in diameter at their widest point, and tapering at the anterior and posterior ends (Figure 2).</li>
	<li>Place the mosquito-filled tube in a small (60 mm x 15 mm) Petri dish filled with double-distilled water (ddH2O). Place the tube and dish in an incubator at 28 &#176;C for 45 min (Figure 3).</li>
	<li>Remove the tube from the incubator and insert the tube into an empty cage. Gently tap the tube to allow mosquitoes to fly out and remove the tube from the cage once all mosquitoes have flown out.</li>
	<li>When the tube is free of mosquitoes, unscrew the bottom ring, remove the nylon, and use forceps to carefully remove the filter paper with the eggs from the tube. Place the filter paper in a plastic Petri dish (100 mm x 15 mm) containing a layer of filter paper moistened with ddH2O.</li>
	<li>Observe the eggs. Eggs that have aged to the point where they are light gray in color (Figure 4) are ready for alignment.
	<ol>
		<li>If eggs are still white, return them to the incubator and check the color every 5 min. White eggs are fragile, rupture easily, and do not survive well after the injection process resulting in low hatching.</li>
		<li>Eggs that are dark gray or black have aged too much. Do not use them.</li>
	</ol>
	</li>
</ol>
3. Embryo alignment
<ol>
	<li>Cut a piece of nylon blotting membrane (2 cm x 1 cm) with a razor blade, making sure that the edge is neatly trimmed. 
	NOTE: If the edge is not straight, the eggs will not stay affixed to the membrane properly during the injection process.</li>
	<li>Put a membrane on a glass&#160;slide and cover the membrane with a piece of filter paper (2 cm x 2 cm), leaving ~1 mm of the membrane filter uncovered (Figure 5). 
	NOTE: Ensure microscope slides are clean before use and use clean forceps to manipulate the membrane filter.</li>
	<li>Wet the filter paper with deionized H2O as eggs will die if desiccated for prolonged periods of time. 
	NOTE: Do not put too much water on the paper because the embryos will move during the injection process, but make sure that there is enough to prevent the embryo from drying (Figure 6A, B). Excess water can be removed by absorbing it with a piece of filter paper.</li>
	<li>Gently transfer 30-50 embryos to the edge of the membrane with a paintbrush (size #0). Use the brush to align the eggs vertically along the membrane by gently rolling them over on the slide so that the ventral side (convex) is facing upward.</li>
	<li>Orient all the eggs in the same direction so that when the eggs are observed under the microinjection microscope, the posterior end (more pointed, Figure 6A, B) is in a down position and forms an angle of ~15&#176; with the needle (Figure 6C). Line the entire edge of the membrane with eggs (30-50) and place the slide under the microscope for injection. 
	NOTE: Choose the eggs carefully. Discard those that are white (too young or undeveloped) or dark gray (too old and will break the needle), and the abnormal ones (Figure 7). Make sure the filter paper remains moist at all times.</li>
</ol>
4. DNA preparation
<ol>
	<li>Purify plasmid DNAs for injection, at the minimum, with an endotoxin-free DNA plasmid maxiprep kit following the manufacturer&#39;s protocols.</li>
	<li>Resuspend the DNA in PCR-grade water and centrifuge at 17,100 x g in a refrigerated centrifuge for 30 min at 4 &#176;C. Then, transfer the supernatant&#160;to a new, clean 1.5 mL conical tube. Carefully micro-pipette to avoid transferring undesired particulate matter from the column.</li>
	<li>Perform a second precipitation of DNA using one-tenth volume of 3M sodium acetate and a two-fold volume of 100% ethanol.&#160;</li>
	<li>Resuspend the DNA in 300-500 &#181;L of PCR-grade water for a final plasmid concentration of 1000 ng/&#181;L. Mix the plasmid cocktail in the injection buffer for a final plasmid concentration of 300-400 ng/&#181;L and make 10-20 &#181;L aliquots. 
	&#8203;NOTE: DNA can be stored at -20 &#176;C. DNA microinjection aliquots can be stored at -20 &#176;C, or at -80 &#176;C if using RNA components. Do not exceed 800 ng/&#181;L of DNA in the injection mix because viscosity will cause the needles to clog.</li>
</ol>
5. Needle preparation
<ol>
	<li>Make the needles by pulling 10 cm quartz capillaries using a programmable micropipette puller.</li>
	<li>Examine the needle under microscope to ensure that the tip is formed well. Ensure that the needle setting on the programmable puller yields a tip that starts to taper ~0.5 mm from the terminal end and ends in a fine point (see Figure 6). Needles that break easily usually have too long of a taper. 
	NOTE: Conditions might differ depending on the needle puller (The authors will make available on request the settings for the programmable puller they use. Journal restrictions prevent us from citing a specific brand). Needles can be pulled by hand, but this requires a skill-set not available in most laboratories.</li>
</ol>
6. Embryo microinjection
<ol>
	<li>Use a micro-loader tip to fill the needles with 2 &#181;L of DNA mixture. Insert the needle into the needle holder and connect the automated pressure pump tubing (Figure 8).</li>
	<li>Important: Align the needle so that it makes an angle of 15&#176; with the plane of the slide (Figure 8).</li>
	<li>Open the needle tip by carefully touching the first egg and inject it by inserting the tip of the needle &#8804; 10 &#181;m in the posterior pole. A successful injection will lead to a small movement of the cytoplasm within the egg.</li>
	<li>Use the microscope coaxial stage controls to move to the next egg to continue injection.
	<ol>
		<li>To ensure that the needle tip remains open and has not clogged, press the Inject button before entering another embryo and visualize the small droplet at the opening of the needle.</li>
		<li>If the needle gets clogged, press the Clear button to clear the needle and repeat the droplet visualization test. Adjust the pressure as needed if the needle tip opening gets slightly bigger to ensure that the size of the droplet stays small.</li>
		<li>Inject ~40-50 eggs with one needle. 
		NOTE: Make sure the filter paper stays moist at all times. Keep sufficient back-pressure on the needle to keep it cleared. A needle that cannot be unclogged must be replaced with a new needle. Embryo placement, needle insertion and injection are made easier with microscopes that have coaxial controls for horizontal movement of the stage (Figure 9).</li>
	</ol>
	</li>
	<li>After injections are complete, rinse the eggs off into a glass container lined with filter paper and filled with 50 mL of deionized H2O (Figure 10). 
	NOTE: Embryos will start hatching 2 days post-injection and may take as long as 3-5 days. Hatched first instar larvae must be transferred immediately (check 2 times daily) to a clean container with water and food.</li>
</ol>

<ol>
	<li>Feramisco, J., Perona, R., Lacal, J. C., Lacal, J. C., Feramisco, J., Perona, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Needle+Microinjection:+A+Brief+History.">Needle Microinjection: A Brief History.</a> Microinjection. Methods and Tools in Biosciences and Medicine. , (1999).</li><li>Windbichler, N., 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+synthetic+homing+endonuclease-based+gene+drive+system+in+the+human+malaria+mosquito.">A synthetic homing endonuclease-based gene drive system in the human malaria mosquito.</a> Nature. 473 (7346), 212-215 (2011).</li><li>Meredith, J. M., 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=Site-specific+integration+and+expression+of+an+anti-malarial+gene+in+transgenic+Anopheles+gambiae+significantly+reduces+Plasmodium+infections.">Site-specific integration and expression of an anti-malarial gene in transgenic Anopheles gambiae significantly reduces Plasmodium infections.</a> PLoS One. 6 (1), 14587 (2011).</li><li>Hammond, A., 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+gene+drive+system+targeting+female+reproduction+in+the+malaria+mosquito+vector+Anopheles+gambiae.">CRISPR-Cas9 gene drive system targeting female reproduction in the malaria mosquito vector Anopheles gambiae.</a> Nature Biotechnology. 34 (1), 78-83 (2016).</li><li>Carballar-Lejarazu, R., 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=Next-generation+gene+drive+for+population+modification+of+the+malaria+vector+mosquito,+Anopheles+gambiae.">Next-generation gene drive for population modification of the malaria vector mosquito, Anopheles gambiae.</a> Proceedings of the National Academy of Sciences of the United States of America. 117 (37), 22805-22814 (2020).</li><li>Sim&#245;es, M. L., 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+Anopheles+FBN9+immune+factor+mediates+Plasmodium+species-specific+defense+through+transgenic+fat+body+expression.">The Anopheles FBN9 immune factor mediates Plasmodium species-specific defense through transgenic fat body expression.</a> Develomental &amp; Comparative Immunology. 67, 257-265 (2017).</li><li>Arik, A. 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=Increased+Akt+signaling+in+the+mosquito+fat+body+increases+adult+survivorship.">Increased Akt signaling in the mosquito fat body increases adult survivorship.</a> FASEB Journal. 4, 1404-1413 (2015).</li><li>Riabinina, O., 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=Organization+of+olfactory+centres+in+the+malaria+mosquito+Anopheles+gambiae.">Organization of olfactory centres in the malaria mosquito Anopheles gambiae.</a> Nature Communications. 7, 13010 (2016).</li><li>Kyrou, K., 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+CRISPR-Cas9+gene+drive+targeting+doublesex+causes+complete+population+suppression+in+caged+Anopheles+gambiae+mosquitoes.">A CRISPR-Cas9 gene drive targeting doublesex causes complete population suppression in caged Anopheles gambiae mosquitoes.</a> Nature Biotechnology. 36 (11), 1062-1066 (2018).</li><li>Dong, Y., Sim&#245;es, M. L., Dimopoulos, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Versatile+transgenic+multistage+effector-gene+combinations+for+Plasmodium+falciparum+suppression+in+Anopheles.">Versatile transgenic multistage effector-gene combinations for Plasmodium falciparum suppression in Anopheles.</a> Science Advances. 6 (20), (2020).</li><li>Grossman, G. L., 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=Germline+transformation+of+the+malaria+vector,+Anopheles+gambiae,+with+the+piggyBac+transposable+element.">Germline transformation of the malaria vector, Anopheles gambiae, with the piggyBac transposable element.</a> Insect Molecular Biology. 6, 597-604 (2001).</li><li>Benedict, M. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+in+Anopheles+research.+Chapter+3:+Specific+Anopheles+techniques.+3.1+Embryonic+Techniques.+3.1.1+Microinjection+methods+for+Anopheles+Embryos.">Methods in Anopheles research. Chapter 3: Specific Anopheles techniques. 3.1 Embryonic Techniques. 3.1.1 Microinjection methods for Anopheles Embryos.</a> BEI resources. , (2015).</li><li>Pham, T. B., 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=Experimental+population+modification+of+the+malaria+vector+mosquito,+Anopheles+stephensi.">Experimental population modification of the malaria vector mosquito, Anopheles stephensi.</a> PLoS Genetics. 15 (12), 1008440 (2019).</li><li>Benedict, M. Q., 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=Pragmatic+selection+of+larval+mosquito+diets+for+insectary+rearing+of+Anopheles+gambiae+and+Aedes+aegypti.">Pragmatic selection of larval mosquito diets for insectary rearing of Anopheles gambiae and Aedes aegypti.</a> PLoS One. 15 (3), 0221838 (2020).</li><li>Carballar-Lejaraz&#250;, R., James, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Population+modification+of+Anopheline+species+to+control+malaria+transmission.">Population modification of Anopheline species to control malaria transmission.</a> Pathogens and Global Health. 111 (8), 424-435 (2017).</li></ol>

The authors have nothing to disclose.

With the increased availability of precise and flexible genetic engineering technologies such as CRISPR/Cas9, transgenic organisms can be developed in a more straightforward and stable way than previously possible. These tools have allowed researchers to create transgenic strains of mosquito vectors that are very close to achieving the desired properties of either refractoriness to pathogens (population modification) or heritable sterility (population suppression). However, to develop the most safe and stable genetically...

Microinjection techniques have been used to experimentally manipulate organisms since the early 1900s1. Microinjection has been used to study both basic biological functions and/or introduce important changes in the biology of a desired organism. The microinjection technique has been of particular interest to vector biologists and has been widely used to manipulate vector genomes2-11. Transgenesis experiments in arthropod vectors often aim to make vectors less efficient at transmitting pathogens by either enacting changes that decrease a vector&#39;s fitness or increase refractoriness to the pathogens they transmit. Mosquitoes transmit a variety of human pathogens and have a significant impact on morbidity and mortality worldwide. The Anopheles genus of mosquitoes transmits the human malaria parasitic pathogens,&#160;Plasmodium spp.&#160;Genetic engineering experiments with Anopheles have aimed to better understand the biology and reduce the vectorial capacity of these mosquitoes in efforts to develop novel malaria elimination strategies.
The mosquito vectors that contribute the most malaria infections worldwide are in the Anopheles gambiae species complex. However, the majority of successful transgenesis experiments have been performed on the malaria vector of the Indian subcontinent, Anopheles stephensi. While plenty of laboratory-adapted Anopheles gambiae strains exist, the number of transgenic Anopheles gambiae spp. lines reported in the literature does not compare to that of Anopheles stephensi. It is thought that the Anopheles gambiae embryo is more difficult to inject and achieve successful transgenesis than Anopheles stephensi, although the reasons for these differences are unknown. This protocol describes a method that has been proven to be consistently successful in achieving transgenesis of Anopheles gambiae embryos via microinjection. The protocol is based on a method previously developed by Herv&#233;&#160;Bossin and Mark Benedict12 with some additional details and alterations added that have been found to increase the efficiency of transgenesis.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >10x Microinjection Buffer</td>
			<td >-</td>
			<td >-</td>
			<td >1 mM NaHPO4 buffer, pH 6.8, 50 mM KCl</td>
		</tr>
		<tr >
			<td >Blotting membrane (Zeta-Probe GT Genomic Tested Blotting Membrane)</td>
			<td >Bio-Rad</td>
			<td ></td>
			<td >Neatly and straightly cut into 2x1 cm piece</td>
		</tr>
		<tr >
			<td >Conical tubes 50 ml (disposable centrifuge tube, polypropylene)</td>
			<td >Fisher Brand</td>
			<td ></td>
			<td >Ends cut</td>
		</tr>
		<tr >
			<td >De-ionized or double-distilled water (ddH20)&#160;</td>
			<td >Mili-Q</td>
			<td ></td>
			<td >In a wash bottle&#160;</td>
		</tr>
		<tr >
			<td >Dissecting microscope&#160;</td>
			<td >Leica&#160;</td>
			<td >Leica MZ12</td>
			<td >For embryo alignment</td>
		</tr>
		<tr >
			<td >Forceps&#160;</td>
			<td ></td>
			<td ></td>
			<td >No. 5 size&#160;</td>
		</tr>
		<tr >
			<td >Glass container&#160;</td>
			<td >Pyrex</td>
			<td >No. 3140</td>
			<td >125 x 65</td>
		</tr>
		<tr >
			<td >Glass slide&#160;</td>
			<td >Fisher Brand</td>
			<td >No. 12-549-3</td>
			<td >75x26 mm</td>
		</tr>
		<tr >
			<td >Incubator</td>
			<td >Barnsted Lab-line</td>
			<td >Model No. 150</td>
			<td >28 &#176;C</td>
		</tr>
		<tr >
			<td >KCl</td>
			<td ></td>
			<td ></td>
			<td >50 mM</td>
		</tr>
		<tr >
			<td >Latex dental film&#160;</td>
			<td >Crosstex International</td>
			<td >No. 19302</td>
			<td ></td>
		</tr>
		<tr >
			<td >Microinjector</td>
			<td >Sutter Instrument</td>
			<td ></td>
			<td >XenoWorks Digital Microinjector</td>
		</tr>
		<tr >
			<td >Microloader Pipette tips&#160;</td>
			<td >Eppendorf&#160;</td>
			<td ></td>
			<td >20 &#181;L microloader epT.I.P.S.</td>
		</tr>
		<tr >
			<td >Micromanipulator</td>
			<td >Sutter Instrument</td>
			<td ></td>
			<td >XenoWorks Micromanipulator</td>
		</tr>
		<tr >
			<td >Micropipette&#160;</td>
			<td >Rainin&#160;</td>
			<td ></td>
			<td >20 &#181;L</td>
		</tr>
		<tr >
			<td >Micropipette puller&#160;</td>
			<td >Sutter Instrument</td>
			<td >Sutter P-2000 micropipette puller</td>
			<td ></td>
		</tr>
		<tr >
			<td >Microscope&#160;</td>
			<td >Leica</td>
			<td >DM 1000 LED or M165 FC</td>
			<td >For microinjection</td>
		</tr>
		<tr >
			<td >Minimum fiber filter paper&#160;</td>
			<td >Fisher Brand</td>
			<td >No. 05-714-4</td>
			<td >Chromatography Paper, Thick&#160;</td>
		</tr>
		<tr >
			<td >Mosquitoes&#160;</td>
			<td >MR4, BEI Resources</td>
			<td ></td>
			<td >Anopheles gambiae, mated adult females, blood-fed 4-5 days post-eclosion</td>
		</tr>
		<tr >
			<td >NaHPO4 buffer&#160;</td>
			<td ></td>
			<td ></td>
			<td >1 mM, ph 6.8</td>
		</tr>
		<tr >
			<td >Nylon mesh</td>
			<td ></td>
			<td ></td>
			<td ></td>
		</tr>
		<tr >
			<td >Paint brush</td>
			<td >Blick</td>
			<td >No. 05831-7040</td>
			<td >Fine, size 4/0</td>
		</tr>
		<tr >
			<td >Petri dish</td>
			<td ></td>
			<td ></td>
			<td >Plastic, (60x15 mm, 90x15 mm)</td>
		</tr>
		<tr >
			<td >Sodium acetate</td>
			<td ></td>
			<td ></td>
			<td >&#160;3M</td>
		</tr>
		<tr >
			<td >Quartz glass capillaries&#160;</td>
			<td >Sutter Instrument</td>
			<td >No. QF100-70-10</td>
			<td >With filament, 1 mm OD,&#160; ID 0.7 10 cm length</td>
		</tr>
		<tr >
			<td >Water PCR grade&#160;</td>
			<td >Roche</td>
			<td >No. 03315843001</td>
			<td ></td>
		</tr>
	</tbody>
</table>

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.

A representative example of the application of the microinjection protocol described can be found in Carballar-Lejaraz&#250;&#160;et al5. The intent here was to insert an autonomous gene-drive system into the germline of a laboratory strain, G3, of An. gambiae. The system was designed to target the cardinal ortholog locus (Agcd) on the third chromosome in this species, which encodes a heme peroxidase that catalyzes the conversion of 3-hydroxykynurenine to xanthommatin, t...

Watch this Scientific Journal Video about Microinjection Method for Anopheles gambiae Embryos at JoVE.com

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.

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 ...

microinjection-method-for-anopheles-gambiae-embryos

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4.4K Views.  University of California, Irvine. 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. 

Video: Microinjection Method for Anopheles gambiae Embryos

Laser-assisted Cytoplasmic Microinjection in Livestock Zygotes

Cytoplasmic microinjection into one-cell embryos is a very powerful technique. As an example, it enables the delivery of genome editing tools that can create genetic modifications that will be present in every cell of an adult organism. It can also be used to deliver siRNA, mRNAs or blocking antibodies to study gene function in preimplantation embryos. The conventional technique for microinjecting embryos used in rodents consists of a very thin micropipette that directly penetrates the plasma membrane when advanced into the embryo. When this technique is applied to livestock animals it usually results in low efficiency. This is mainly because in contrast to mice and rats, bovine, ovine, and porcine zygotes have a very dark cytoplasm and a highly elastic plasma membrane that makes visualization during injection and penetration of the plasma membrane hard to achieve. In this protocol, we describe a suitable microinjection method for the delivery of solutions into the cytoplasm of cattle zygotes that has proved to be successful for sheep and pig embryos as well. First, a laser is used to create a hole in the zona pellucida. Then a blunt-end glass micropipette is introduced through the hole and advanced until the tip of the needle reaches about 3/4 into the embryo. Then, the plasma membrane is broken by aspiration of cytoplasmic content inside the needle. Finally, the aspirated cytoplasmic content followed by the solution of interest is injected back into the embryonic cytoplasm. This protocol has been successfully used for the delivery of different solutions into bovine and ovine zygotes with 100% efficiency, minimal lysis, and normal blastocysts development rates.

This protocol shows how to perform cytoplasmic microinjection in farm animal zygotes. This technique can be used to deliver any solution into the one-cell embryo such as genome editing tools to generate knockout animals.

Cytoplasmic microinjection into one-cell embryos is a very powerful technique. As an example, it enables the delivery of genome editing tools that can ...

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

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

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

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

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

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of tissues, we have developed numerous modifications and improvements to our original fluorescent in situ hybridization (FISH) protocol. To facilitate throughput and cost effectiveness, all steps, from probe generation to signal detection, are performed using exon 96-well microtiter plates. Digoxygenin (DIG)-labelled antisense RNA probes are produced using either cDNA clones or genomic DNA as templates. After tissue fixation and permeabilization, probes are hybridized to transcripts of interest and then detected using a succession of anti-DIG antibody conjugated to biotin, streptavidin conjugated to horseradish peroxidase (HRP) and fluorescently conjugated tyramide, which in the presence of HRP, produces a highly reactive intermediate that binds to electron dense regions of immediately adjacent proteins. These amplification and localization steps produce a robust and highly localized signal that facilitates both cellular and subcellular transcript localization. The protocols provided have been optimized to produce highly specific signals in a variety of tissues and developmental stages. References are also provided for additional variations that allow the simultaneous detection of multiple transcripts, or transcripts and proteins, at the same time.

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

The described RNA in situ hybridization protocol allows the detection of RNA in whole Drosophila embryos or dissected tissues. Using 96-well microtiter plates and tyramide signal amplification, transcripts can be detected at high resolution, sensitivity, and throughput, and at a relatively low cost.

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of ...

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

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

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

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

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

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a protocol for performing the chromatin conformation capture technique in situ Hi-C on staged Drosophila melanogaster embryo populations. The result is a sequencing library that allows the mapping of all chromatin interactions that occur in the nucleus in a single experiment. Embryo sorting is done manually using a fluorescent stereo microscope and a transgenic fly line containing a nuclear marker. Using this technique, embryo populations from each nuclear division cycle, and with defined cell cycle status, can be obtained with very high purity. The protocol may also be adapted to sort older embryos beyond gastrulation. Sorted embryos are used as inputs for in situ Hi-C. All experiments, including sequencing library preparation, can be completed in five days. The protocol has low input requirements and works reliably using 20 blastoderm stage embryos as input material. The end result is a sequencing library for next generation sequencing. After sequencing, the data can be processed into genome-wide chromatin interaction maps that can be analyzed using a wide range of available tools to gain information about topologically associating domain (TAD) structure, chromatin loops, and chromatin compartments during Drosophila development.

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

This work describes a protocol for the generation of high resolution in situ Hi-C libraries from tightly staged pre-gastrulation Drosophila melanogaster embryos.

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a ...

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...

A Screening Method for Identification of Heterochromatin-Promoting Drugs Using Drosophila

Drosophila is an excellent model organism that can be used to screen compounds that might be useful for cancer therapy. The method described here is a cost-effective in vivo method to identify heterochromatin-promoting compounds by using Drosophila. The Drosophila&#39;s DX1 strain, having a variegated eye color phenotype that reflects the extents of heterochromatin formation, thereby providing a tool for a heterochromatin-promoting drug screen. In this screening method, eye variegation is quantified based on the surface area of red pigmentation occupying parts of the eye and is scored on a scale from 1 to 5. The screening method is straightforward and sensitive and allows for testing compounds in vivo. Drug screening using this method provides a fast and inexpensive way for identifying heterochromatin-promoting drugs that could have beneficial effects in cancer therapeutics. Identifying compounds that promote the formation of heterochromatin could also lead to the discovery of epigenetic mechanisms of cancer development.

A Screening Method for Identification of Heterochromatin-Promoting Drugs Using Drosophila

Drosophila is a widely used experimental model suitable for screening drugs with potential applications for cancer therapy. Here, we describe the use of Drosophila variegated eye color phenotypes as a method for screening small-molecule compounds that promote heterochromatin formation.

Drosophila is an excellent model organism that can be used to screen compounds that might be useful for cancer therapy. The method described here is a ...

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

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

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

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

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

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 ...