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><tbody><tr><td>10x Microinjection Buffer</td><td>-</td><td>-</td><td>1 mM NaHPO&lt;sub&gt;4&lt;/sub&gt; 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 (ddH&lt;sub&gt;2&lt;/sub&gt;0)&amp;nbsp;</td><td>Mili-Q</td><td></td><td>In a wash bottle&amp;nbsp;</td></tr><tr><td>Dissecting microscope&amp;nbsp;</td><td>Leica&amp;nbsp;</td><td>Leica MZ12</td><td>For embryo alignment</td></tr><tr><td>Forceps&amp;nbsp;</td><td></td><td></td><td>No. 5 size&amp;nbsp;</td></tr><tr><td>Glass container&amp;nbsp;</td><td>Pyrex</td><td>No. 3140</td><td>125 x 65</td></tr><tr><td>Glass slide&amp;nbsp;</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 &amp;deg;C</td></tr><tr><td>KCl</td><td></td><td></td><td>50 mM</td></tr><tr><td>Latex dental film&amp;nbsp;</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&amp;nbsp;</td><td>Eppendorf&amp;nbsp;</td><td></td><td>20 &amp;micro;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&amp;nbsp;</td><td>Rainin&amp;nbsp;</td><td></td><td>20 &amp;micro;L</td></tr><tr><td>Micropipette puller&amp;nbsp;</td><td>Sutter Instrument</td><td>Sutter P-2000 micropipette puller</td><td></td></tr><tr><td>Microscope&amp;nbsp;</td><td>Leica</td><td>DM 1000 LED or M165 FC</td><td>For microinjection</td></tr><tr><td>Minimum fiber filter paper&amp;nbsp;</td><td>Fisher Brand</td><td>No. 05-714-4</td><td>Chromatography Paper, Thick&amp;nbsp;</td></tr><tr><td>Mosquitoes&amp;nbsp;</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>NaHPO&lt;sub&gt;4&lt;/sub&gt; buffer&amp;nbsp;</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>&amp;nbsp;3M</td></tr><tr><td>Quartz glass capillaries&amp;nbsp;</td><td>Sutter Instrument</td><td>No. QF100-70-10</td><td>With filament, 1 mm OD,&amp;nbsp; ID 0.7 10 cm length</td></tr><tr><td>Water PCR grade&amp;nbsp;</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

Universidad de Cartagena UDEC

Research

JoVE Journal

Genetics

4.5K 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

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. ...

Injection of An. stephensi Embryos to Generate Malaria-resistant Mosquitoes

The introduction of exogenous genes into the genomes of mosquitoes requires microinjection techniques tailored to the specific species of interest. This video protocol demonstrates a method used by the James laboratory to microinject DNA constructs into Anopheles stephensi embryos for the generation of transformed mosquitoes. Techniques for preparing microinjection needles, collecting and preparing embryos and performing the microinjection are illustrated.

Anopheles stephensi mosquitoes are vectors for malaria inhabiting India and throughout Asia. This video demonstrates the technique for performing microinjections of this species with transgenes that will confer resistance to the malaria to the mosquito. Much of the methodology demonstrated in this video is applicable to microinjection techniques of other mosquito species.

The introduction of exogenous genes into the genomes of mosquitoes requires microinjection techniques tailored to the specific species of interest. This ...

Biology

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 ...

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Mosquitoes are vectors for a diverse set of pathogens including arboviruses, protozoan parasites and nematodes. Investigation of transcripts and gene regulators that are expressed in tissues in which the mosquito host and pathogen interact, and in organs involved in reproduction are of great interest for strategies to reduce mosquito-borne disease transmission and disrupt egg development. A number of tools have been employed to study and validate the temporal and tissue-specific regulation of gene expression. Here, we describe protocols that have been developed to obtain spatial information, which enhances our understanding of where specific genes are expressed and their products accumulate. The protocol described has been used to validate expression and determine accumulation patterns of transcripts in tissues related to mosquito-borne pathogen transmission, such as female salivary glands, as well as subcellular compartments of ovaries and embryos, which relate to mosquito reproduction and development. 
 The following procedures represent an optimized methodology that improves the efficiency of various steps in the protocol without loss of target-specific hybridization signals. Guidelines for RNA probe preparation, dissection of soft tissues and the general procedure for fixation and hybridization are described in Part A, while steps specific for the collection, fixation, pre-hybridization and hybridization of mosquito embryos are detailed in Part B.

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Temporal and spatial gene expression analyses have a crucial role in functional genomics. Whole-mount hybridization in situ is useful for determining the localization of transcripts within tissues and subcellular compartments. Here we outline a hybridization in situ protocol with modifications for specific target tissues in mosquitoes.

Mosquitoes are vectors for a diverse set of pathogens including arboviruses, protozoan parasites and nematodes.  Investigation of transcripts and gene ...

Immunology and Infection

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, ...

Site-Directed &#966;C31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria

Functional genomic analysis and related strategies for genetic control of malaria rely on validated and reproducible methods to accurately modify the genome of Anopheles mosquitoes. Amongst these methods, the &#966;C31 system allows precise and stable site-directed integration of transgenes, or the substitution of integrated transgenic cassettes via recombinase-mediated cassette exchange (RMCE). This method relies on the action of the Streptomyces &#966;C31 bacteriophage integrase to catalyze recombination between two specific attachment sites designated attP (derived from the phage) and attB (derived from the host bacterium). The system uses one or two attP sites that have been integrated previously into the mosquito genome and attB site(s) in the donor template DNA. Here we illustrate how to stably modify the genome of attP-bearing Anopheles docking lines using two plasmids: an attB-tagged donor carrying the integration or exchange template and a helper plasmid encoding the &#966;C31 integrase. We report two representative results of &#966;C31-mediated site-directed modification: the single integration of a transgenic cassette in An. stephensi and RMCE in An. gambiae mosquitoes. &#966;C31-mediated genome manipulation offers the advantage of reproducible transgene expression from validated, fitness neutral genomic sites, allowing comparative qualitative and quantitative analyses of phenotypes. The site-directed nature of the integration also substantially simplifies the validation of the single insertion site and the mating scheme to obtain a stable transgenic line. These and other characteristics make the &#966;C31 system an essential component of the genetic toolkit for the transgenic manipulation of malaria mosquitoes and other insect vectors.

The protocol describes how to achieve site-directed modifications in the genome of Anopheles malaria mosquitoes using the &#966;C31 system. Modifications described include both the integration and the exchange of transgenic cassettes in the genome of attP-bearing docking lines.

Functional genomic analysis and related strategies for genetic control of malaria rely on validated and reproducible methods to accurately modify the ...

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is based on the injection of non-pathogenic particles into insects, as the particles will be detected by the immune system and will induce the production of immune effectors. We focus here on the stimulation of the melanization response in the mosquito Anopheles gambiae. The melanization response results in the encapsulation of foreign particles and parasites with a dark layer of melanin. To stimulate this response, mosquitoes are inoculated with beads in the thoracic cavity using microcapillary glass tubes. Then, after 24 hr, the mosquitoes are dissected to retrieve the beads. The degree of melanization of the bead is measured using image analysis software. Beads do not have the pathogenic effects of parasites, or their capacity to evade or suppress the immune response. These injections are a way to measure immune efficacy and the impact of immune stimulations on other life history traits, such as fecundity or longevity. It is not exactly the same as directly studying host-parasite interactions, but it is an interesting tool to study immunity and its evolutionary ecology.

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

Through inoculation with beads, the described technique enables the stimulation of the mosquito melanization response in the hemolymph circulating system. The amount of melanin covering the beads can be measured after dissection as a measure of the immune response.

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is ...

A Method for Microinjection of Patiria minata Zygotes

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and evolution of developmental processes. The sea star, Patiria miniata, is gaining prevalence as a model system for these types of studies which were previously performed almost exclusively in the sea urchins, Strongylocentrotus purpuratus and Lytechinus variegatus. An advantage of these model systems is the ease of producing modified embryos in which a particular gene is up or downregulated, labeling a group of cells, or introducing a reporter gene. A single microinjection method is capable of creating a wide variety of such modified embryos. Here, we present a method for obtaining gametes from P. miniata, producing zygotes, and introducing perturbing reagents via microinjection. Healthy morphant embryos are subsequently isolated for quantitative and qualitative studies of gene function. The availability of genome and transcriptome data for this organism has increased the types of studies that are performed and the ease of executing them.

A Method for Microinjection of Patiria minata Zygotes

Methods that produce morphant embryos are essential to study developmental mechanisms and gene regulatory networks. The sea star Patiria miniata is an emerging model system for these studies. Here we present a protocol for obtaining gametes, producing cultures of embryos, and rapid microinjection of zygotes from this species.

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and ...

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory for functional genomic studies. The ability to knockdown individual genes selectively via this reverse genetic technique has allowed many researchers to rapidly uncover the biological roles of numerous genes within many organisms, by evaluation of loss-of-function phenotypes. In the major human malaria vector Anopheles gambiae, the predominant method used to reduce the function of targeted genes involves injection of double-stranded (dsRNA) into the hemocoel of the adult mosquito. While this method has been successful, gene knockdown in adults excludes the functional assessment of genes that are expressed and potentially play roles during pre-adult stages, as well as genes that are expressed in limited numbers of cells in adult mosquitoes. We describe a method for the injection of Serine Protease Inhibitor 2 (SRPN2) dsRNA during the early pupal stage and validate SRPN2 protein knockdown by observing decreased target protein levels and the formation of melanotic pseudo-tumors in SRPN2 knockdown adult mosquitoes. This evident phenotype has been described previously for adult stage knockdown of SRPN2 function, and we have recapitulated this adult phenotype by SRPN2 knockdown initiated during pupal development. When used in conjunction with a dye-labeled dsRNA solution, this technique enables easy visualization by simple light microscopy of injection quality and distribution of dsRNA in the hemocoel.

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi) is an extremely valuable tool for uncovering gene function. However, the ability to target genes using RNAi during pre-adult stages is limited in the major human malaria vector Anopheles gambiae. We describe an RNAi protocol to reduce gene function via direct injection during pupal development.

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory ...

Mosquito Embryo Microinjection: A Technique to Deliver Exogenous DNA into Embryo Yolk of Anopheles Gambiae to Generate Germline Mutations

Source: Carballar-Lejaraz&#250;, R. et al, <a href="https://jove.unicartagenaproxy.elogim.com/v/62591">Microinjection Method for Anopheles gambiae Embryos.</a> J. Vis. Exp. (2021).
In this video, we demonstrate the delivery of gene drive construct into the embryo of Anopheles gambiae mosquito through embryo microinjection technique. This procedure allows for integration of&#160; delivered DNA construct, through homologous recombination, into the germline cells to generate transformed mosquito population.

Mikroiniekcja zarodka komara: technika dostarczania egzogennego DNA do żółtka zarodka Anopheles Gambiae w celu wytworzenia mutacji linii zarodkowej

Source: Carballar-Lejaraz&#250;, R. et al, Microinjection Method for Anopheles gambiae Embryos. J. Vis. Exp. (2021).
In this video, we demonstrate the ...

Microinjection Method for Anopheles gambiae Embryos

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Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

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

Site-Directed φC31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

A Method for Microinjection of Patiria minata Zygotes

RNAi Trigger Delivery into Anopheles gambiae Pupae

Mikroiniekcja zarodka komara: technika dostarczania egzogennego DNA do żółtka zarodka Anopheles Gambiae w celu wytworzenia mutacji linii zarodkowej