This work was supported in part by UCSD startup funds directed to O.S.A.

1. C. quinquefasciatus colony rearing
<ol>
	<li>Set up multiple colonies of C. quinquefasciatus adults in bug dorm cages. 
	NOTE: The colonies were provided by Dr. Laura Harrington at Cornell University20. Detailed protocols for rearing Culex mosquitoes can be found in other literature21.
	<ol>
		<li>Maintain the mosquitoes at 25 &#177; 1 &#730;C at 30% humidity with a 12:12 h day:night cycle.</li>
		<li>Provide 20% sugar solution ad libitum by introducing a sugar solution container with a wick into the cage or by saturating cotton balls with the sugar solution and placing it within the cage.</li>
	</ol>
	</li>
	<li>Allow mosquitoes to mate for at least 3 days prior to blood feeding (Figure 2A).</li>
</ol>
2. Collection of C. quinquefasciatus pre-blastoderm stage embryos
<ol>
	<li>After 3-6 days post eclosion, provide females with 1-2 mL of a citrated bovine blood-meal through a synthetic membrane and heated to approximately 40 &#176;C in a blood feeding system (Figure 2B). There are also alternative blood feeding protocols that can be used for Culex mosquitoes (e.g., see ref21). 
	NOTE: C. quinquefasciatus is known for a preference for avian blood. Some labs use anesthetized live birds or avian blood for their blood meal source21,22,23. However, colonies can be trained/selected to feed on bovine blood or small rodents if this feature is selected for over multiple generations.
	<ol>
		<li>Allow females to rest for a minimum of 3 days after blood feeding to undergo oogenesis and egg maturation before inducing oviposition for microinjection experiments.</li>
	</ol>
	</li>
	<li>On the day of embryonic microinjections, create an oviposition cup with organically infused oviposition water. Oviposition water should be made by fermenting rabbit feces (50 g/L), decomposing grass (4.5 g/L), or fish food (25 g/L) in water over the course of 5 or more days24,25.</li>
	<li>Place the oviposition cup into the cage and place the entire cage into a dark location (Figure 2C).</li>
	<li>After every 30 min, check the cup for egg rafts.
	<ol>
		<li>If egg rafts are present, collect the rafts by scooping them with a paintbrush and place them on a wet filter paper (Figure 2D,E).</li>
	</ol>
	</li>
</ol>
3. Alignment of C. quinquefasciatus pre-blastoderm stage embryos
<ol>
	<li>Separate the eggs from the rafts by pressing down on the raft and teasing apart the eggs individually using a fine tip paintbrush and forceps (Figure 2E).</li>
	<li>Align individual eggs on a thin strip of double-sided sticky tape placed across the top of a glass slide (Figure 2F).</li>
	<li>While aligning, try to point the anterior side of each egg in the same direction for easier accessibility. 
	NOTE: An alternative egg alignment method without double sided sticky tape can be found in a previously published Nasonia vitripennis embryo microinjection protocol26.</li>
	<li>Cover eggs with the halocarbon oil mix. 
	NOTE: Halocarbon mix can be prepared ahead of time by gently mixing two halocarbon reagents and water (9:1:20, halocarbon 700 : halocarbon 27 : Ultrapure water) and then incubating the mixture overnight at 25&#959;C to facilitate water saturation of the halocarbon oil.</li>
</ol>
4. Needle preparation for microinjections
<ol>
	<li>Generate the aluminosilicate capillary glass needles using a glass micropipette puller.
	<ol>
		<li>Place an aluminosilicate capillary glass into a needle puller, as per instructions from the needle puller&#8217;s user manual. 
		NOTE: Quartz and borosilicate capillary needles can also be used, but for this experiment, aluminosilicate is preferred because of its relative affordability and durability.</li>
		<li>Set Heat to 516, Velocity to 100, Delay to 70, Pull to 97, and Pressure to 500 on the needle puller.</li>
		<li>Activate the needle puller, as per the puller&#8217;s instructions and repeat as needed for additional needles. 
		NOTE: During the injection process, needles often become clogged or accidentally broken, therefore, pulling additional needles for a single experiment is highly recommended.</li>
	</ol>
	</li>
	<li>Bevel the needle tip by gently touching the tip of the pulled needle on the rotating diamond abrasive plate for around 10 s at a 50&#730; angle. 
	NOTE: Beveling the needle opens it to allow fluid to flow through while also creating a sharper tip for easier penetration into the embryo. An example of a proper needle can be seen in a previous article26.</li>
	<li>Store pulled and beveled needles by embedding them into lines of modeler&#8217;s clay in a Petri dish. 
	NOTE: To ensure the best quality for needle tips, needles should be freshly pulled and beveled as close to the time of injection as possible.</li>
</ol>
5. Loading the injection mixture
<ol>
	<li>Prepare the injection mixture consisting of genome modification reagents (e.g., 200 ng/&#956;L sgRNA and 200 ng/&#956;L Cas9 mixture), or preferred injection solutions and keep it on ice. 
	NOTE: This mix can be prepared while waiting for eggs to be laid. More details on Cas9 and sgRNA production and preparation for microinjection can be found in previous publications27,28,29,30.</li>
	<li>Load 2 &#956;L of injection mixture into the injection needle using a microloader tip.</li>
</ol>
6. Microinjection set up
<ol>
	<li>Place the filled injection needle into a micromanipulator set up linked to an electronic microinjector.</li>
	<li>Place the glass slide containing the aligned eggs on the stage of a compound microscope.</li>
	<li>Using the micromanipulator and compound microscope, align the needle to aim at the posterior end of the embryo at a 25-35&#730; angle.</li>
</ol>
7. Embryo microinjection (Figure 2G)
<ol>
	<li>Carefully insert the needle into the embryo and inject the mixture at a quantity of about 10% of the volume of the embryo (700-800 pL depending on the size of the eggs). 
	NOTE: When injecting, the egg should slightly swell, however, if too much fluid is injected, eggs may burst, or cytoplasmic fluid may leak out.</li>
	<li>Inject around 20 eggs at a time then stop and perform embryo recovery procedures. 
	NOTE: While injecting, there is a high chance of needles to either clog or break. If a clog occurs, which can be determined by a lack of injection fluid flowing through the needle, try either cleaning out the needle with the clean function on the electronic microinjector or try re-beveling the needle. If neither step works, quickly change to a new needle while ensuring the prepared eggs remain moistened.</li>
</ol>
8. Embryo recovery and hatching
<ol>
	<li>Leave the eggs undisturbed for at least 5 min. Within 20 min post-injection, carefully remove the halocarbon oil by brushing lightly with a clean paintbrush (Figure 2H).</li>
	<li>Lift the eggs gently using a paintbrush and place them into a cup of double-distilled water (Figure 2I). Take care to keep the eggs on the surface of the water.</li>
	<li>Check the eggs daily for 7 days for hatching (Figure 2J).</li>
	<li>Follow normal larval rearing procedures21.</li>
</ol>
9. Screening for genome modification
<ol>
	<li>Screen the injected mosquitoes for the mutant phenotypes using a stereoscope (Figure 2K).
	<ol>
		<li>Verify mutations that do not have an easily recognizable phenotype, by PCR amplification, T7 Endonuclease I assay, and sequencing by subcloning of the target region31.</li>
	</ol>
	</li>
	<li>Set up additional crosses between injected individuals to detect heritable mutations within the germline.</li>
</ol>

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With recent efforts to generate genetically engineered tools for mosquito vector control, there is a need to develop and optimize embryo microinjection protocols for common mosquito disease vectors. Although methods have been developed for Aedes and Anopheles mosquitoes, a protocol designed specifically for Culex has been minimally explored. In general mosquito embryo injection protocols can be divided into 3 general steps: 1) embryo collection and preparation, 2) embryo injection, and 3) post-injection recovery. In order to successfully generate mutants, all three steps must be optimized for the target species. This modified protocol for embryo microinjection is specific for successful genome engineering of Culex mosquitoes.
Maximizing embryo collection is a common bottleneck in embryo microinjection protocols. In order to increase the number of eggs collected in a short period of time, mosquitoes were restricted from an oviposition substrate for 2-3 days post blood meal. It is important to note that C. quinquefasciatus tend to prefer avian blood sources as opposed to mammalian sources or membrane feeding with mammalian blood. However, many researchers have difficulty acquiring a reliable source of live avians or avian blood for blood feeding, so laboratory stocks can be adapted to feed on more accessible blood sources. It is also crucial to use nutrient-rich water for the oviposition substrate as it provides oviposition cues for C. quinquefasciatus and most other Culex vectors. There are many methods to generate nutrient-rich water, which may need to be optimized between labs to ensure an appropriate number of eggs are available for microinjection. For example, while this method was the most successful with rabbit feces fermented in deionized water for 5 or more days, other researchers have more success with grass or fish food as the nutrient source and some even with distilled water21.
Since Culex mosquitoes lay groups of eggs in rafts, collecting eggs is fairly simple. Eggs can simply be collected by scooping the entire raft with a paintbrush. Egg separation is more complicated, but essential for successful injection. Eggs can be carefully separated from the raft by applying gentle downward pressure between eggs with paintbrush or forceps. With some practice, multiple users were reliably able to separate single eggs from egg rafts. After separating each egg, the eggs are aligned in the same orientation on double-sided sticky tape, which stabilizes the eggs during microinjection. Eggs are injected into the tapered posterior end and the addition of halocarbon oil keeps the eggs moist during manipulation.
To limit embryo damage during microinjection, injection needles need to be of appropriate strength and beveled at an appropriate angle. Aluminosilicate needles beveled for 10 seconds at a 50&#730; angle had the best results, but borosilicate and quartz needles may also work, even though they may be less durable and more expensive. In addition, proper needle beveling facilitates better flow of the Cas9 and sgRNA mixtures and creates a sharper point for easier penetration into the embryo. In many cases, beveling is also an efficient way to quickly fix clogged needles instead of spending the time and effort to replace clogged needles.
After injection, eggs should remain undisturbed for at least 5 minutes with the halocarbon oil being removed immediately afterward by gently brushing it away with a clean paintbrush. Following removal of the halocarbon oil, the injected eggs can be placed in water to hatch, which typically occurs within 3 days post-injection. With practice and sufficient numbers of eggs, this protocol can achieve consistent somatic and germline mutations in C. quinquefasciatus and is versatile enough that it should be easily adaptable to other Culex mosquitoes.

C. quinquefasciatus, commonly known as the southern house mosquito, is a competent vector of numerous pathogens including West Nile virus (WNV), Japanese encephalitis, Saint Louis encephalitis, and Eastern equine encephalitis. In particular, since it was first detected in New York in 1999, WNV has become a major vector-borne disease throughout the continental United States (US) with over 50,000 reported human cases resulting in around 2,300 deaths between 1999 and 20181 as well as over 4,500 reported equine cases between 2008-20192. In addition, at least 23 bird species found in North America have been impacted by WNV infections with at least 12 species classified as irrecoverable as a result of WNV3. The impact of WNV on human, equine, and avian populations is due to the opportunistic feeding behavior of its vectors. Typically, birds are the primary hosts for WNV, and humans and horses are incidental or dead-end hosts. Some pathogens vectored by C. quinquefasciatus only infect birds such as the avian malaria parasite, Plasmodium relictum. In Hawaii, C. quinquefasciatus is a principal vector of avian malaria and has caused the extinction of many native bird species4,5.
To control diseases transmitted by C. quinquefasciatus, researchers and vector control agencies have used commonly established mosquito population control tools such as insecticide application6, however, these methods are costly, not species-specific, and have limited effectiveness as resistance to insecticides is high in many C. quinquefasciatus populations6,7,8,9. Other control techniques, such as Wolbachia-based population control strategies have been developed in recent years10,11, but the fitness costs associated with Wolbachia infection limit the feasibility of this approach for this vector12. There are also genetic-based control methods that have been developed in other mosquito species such as Aedes aegypti13,14, Anopheles gambiae15 and Anopheles stephensi16, including the development of pathogen resistant mosquitoes17,18,19, that could also be developed for C. quinquefasciatus if the requisite genome engineering tools are developed for this species. However, C. quinquefasciatus biology differs greatly from other Aedes and Anopheles mosquito vectors which has made the development of similar genetic technologies difficult in this vector. With the advent of CRISPR-based genome engineering technologies, precise genome engineering has become increasingly trivial, affordable and adaptable and consequently has led to the development of novel genetic tools in a wide variety of species.
To generate mutations with CRISPR-based technologies, a mixture of Cas9 protein and synthetic guide RNA (sgRNA), complementary to the desired loci, is microinjected into pre-blastoderm stage embryos. Since C. quinquefasciatus females lay their eggs in groups attached in a floating raft structure (Figure 1), as opposed to ovipositing individual eggs, a trait of Aedes and Anopheles mosquitoes, embryo microinjections are increasingly complicated in this species. Culex larvae also emerge from the anterior side of each egg, which is in contact with the water surface (Figure 1), so egg orientation post manipulation is important in this species. Here we describe a detailed protocol designed for the microinjection of Cas9 protein and sgRNA into C. quinquefasciatus embryos. This protocol has been designed to accommodate traits unique to Culex biology in order to improve embryo survival and genome mutation rates through certain steps that are key for timely egg collection and egg survival.

<table><tbody><tr><td>9 oz clear plastic pet cup</td><td>Karat</td><td>C-KC9</td><td>Insect Rearing and Egg Collection</td></tr><tr><td>Aluminosilicate glass capillary tubing 1mm(outside diameter) X 0.58mm (inner diameter)</td><td>Sutter Instruments</td><td>BF100-58-10</td><td>Microinjection, borosilicate and quartz needles could also be used but we prefered aluminosilicate</td></tr><tr><td>Blood</td><td>Colorado Serum Company</td><td>31025</td><td>The colony took multiple generations to adapt to this blood source. Other blood sources are likely just as appropriate. See protocol notes on blood source selection.</td></tr><tr><td>Bugdorm</td><td>Bugdorm</td><td>4F2222</td><td>Insect Rearing Cage</td></tr><tr><td>Cas9 Protein with NLS</td><td>PNABio</td><td>CP01</td><td>Microinjection</td></tr><tr><td>Compound Microscope</td><td>Olympus BX41</td><td></td><td>Microinjection, for embryo injection</td></tr><tr><td>Diamond abrasive plate (0.7u to 2.0u tip sizes)</td><td>Sutter Instruments</td><td>104E</td><td>Microinjection, to be used with beveler</td></tr><tr><td>DNase/RNase-Free distilled Water</td><td>Invitrogen</td><td>10977-015</td><td>Microinjection</td></tr><tr><td>Double-sided Tape</td><td>Scotch</td><td>B084NVQGXD</td><td>Microinjection, embryo alignment</td></tr><tr><td>Femtojet 4x or 4i programmable microinjector</td><td>Eppendorf</td><td></td><td>Microinjection</td></tr><tr><td>Femtotips Microloader tips</td><td>Fisher Scientific</td><td>E5242956003</td><td>Microinjection</td></tr><tr><td>Filter Paper</td><td>Whatman</td><td>1001-090</td><td>Microinjection, embryo collection</td></tr><tr><td>Fine-tip paintbrush</td><td>ZEM</td><td>2595</td><td>Microinjection, embryo alignment</td></tr><tr><td>Halocarbon oil 27</td><td>Sigma-Aldrich</td><td>H8773</td><td>Microinjection, embryo alignment</td></tr><tr><td>Halocarbon oil 700</td><td>Sigma-Aldrich</td><td>H8898</td><td>Microinjection, embryo alignment</td></tr><tr><td>Hemotek</td><td>Hemotek</td><td>PS5</td><td>Line Maintenance</td></tr><tr><td>Microelectrode Beveler</td><td>Sutter Instruments</td><td>BV10</td><td>Microinjection, needle beveling</td></tr><tr><td>Micropipette Puller</td><td>Sutter Instruments</td><td>P-1000 or P-2000</td><td>Microinjection, needle pulling</td></tr><tr><td>Microscope Slides</td><td>Fisherbrand</td><td>12-550-A3</td><td>Microinjection, embryo alignment</td></tr><tr><td>Non-Drying Modeling Clay</td><td>Jovi</td><td>B0025Z71IM</td><td>Microinjection, needle storage</td></tr><tr><td>Stereo Microscope</td><td>Olympus</td><td>SZ51</td><td>Microinjection, for embryo alignment</td></tr><tr><td>Sugar</td><td>Domino</td><td></td><td>20% sugar solution for adult sugar source.</td></tr><tr><td>T7 Endonuclease I</td><td>NEB</td><td>M0302</td><td>Preparation of microinjection materials</td></tr><tr><td>TOPO TA Cloning Kit</td><td>ThermoFischer Scientific</td><td>K451020</td><td>Preparation of microinjection materials</td></tr><tr><td>Ultra-fine tip forceps</td><td>Fisher Scientific</td><td>16-100-121</td><td>Microinjection, embryo alignment</td></tr></tbody></table>

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.

In previously published experiments this method was used to successfully generate somatic and germline mutations of a gene critical for the development of dark eye pigmentation, white (CPIJ005542)30 (Table 1). CRISPR/Cas9 generated somatic mutations were scored by screening for loss of pigmentation in pupal stage eyes of injected individuals (G0). Somatic mutations were generally present as mosaic phenotypes where some but not all the cells have the mutant phenotype. For example, somatic mutations of the white gene resulted in a mixture of ommatidia with phenotypes that lack pigmentation and those with a wildtype pigmented phenotype30. When there was a germline mutation of the white gene, however, the next generation (G1) inherited the full white eyed phenotype. Germline mutation rates were determined by intercrossing mosaic G0 individuals and scoring for completely white eyed offspring (G1). These experiments resulted in a 64% to 82% embryo survival rate, as well as a 37% to 57% somatic mutagenesis rate and a &#62;61% germline mutagenesis rate (Table 1). By multiplexing sgRNAs to target multiple loci in the same gene, somatic and germline mutagenesis rates increased upwards to as much as 86% (Table 1). In addition, for many generations, viable homozygous stocks of the white mutants have been successfully kept in the lab.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61375/61375fig01.jpg" /> 
Figure 1: C. quinquefasciatus egg raft and single egg morphology. 
Dorsal (A), lateral (B), and ventral (C) views of C quinquefasciatus egg rafts. C. quinquefasciatus females lay their eggs with the more bulbous anterior side touching the surface of the water while the more pointed posterior away from the water&#8217;s surface (D). A- anterior; P- posterior <a href="https://www.jove.com/files/ftp_upload/61375/61375fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61375/61375fig2v2.jpg" /> 
Figure 2: Timeline for creating C. quinquefasciatus mutants by microinjection. 
Timeline of C. quinquefasciatus colony preparation and embryo collection for microinjection (A-D). Collected eggs are then separated (E) aligned in the same orientation and covered with halocarbon oil (F). The eggs are then injected (G) and allowed to rest for a minimum of 5 min before removing the halocarbon oil (H). Eggs are then transferred to a clean water source (I) for hatching (J) and screened for mutant phenotypes (K). <a href="https://www.jove.com/files/ftp_upload/61375/61375fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td colspan="3">Survival</td>
			<td colspan="3">Somatic mosaicism (G0)</td>
			<td>Germline mutagenesis (G1)</td>
		</tr>
		<tr>
			<td>sgRNA</td>
			<td>#injected</td>
			<td>&#9794;</td>
			<td>&#9792;</td>
			<td>Total (%)</td>
			<td>&#9794;</td>
			<td>&#9792;</td>
			<td>Total (%)</td>
			<td>G1 mutants (%)</td>
		</tr>
		<tr>
			<td>wsgRNA-1</td>
			<td>50</td>
			<td>15</td>
			<td>20</td>
			<td>35(70)</td>
			<td>7(47)</td>
			<td>13(65)</td>
			<td>20(57)</td>
			<td>128(69)</td>
		</tr>
		<tr>
			<td>wsgRNA-2</td>
			<td>50</td>
			<td>9</td>
			<td>32</td>
			<td>41(82)</td>
			<td>3(33)</td>
			<td>12(38)</td>
			<td>15(37)</td>
			<td>51(61)</td>
		</tr>
		<tr>
			<td>wsgRNA-3</td>
			<td>50</td>
			<td>17</td>
			<td>15</td>
			<td>32(64)</td>
			<td>7(41)</td>
			<td>8(53)</td>
			<td>15(47)</td>
			<td>157(72)</td>
		</tr>
		<tr>
			<td>wsgRNA-1/wsgRNA-2</td>
			<td>50</td>
			<td>7</td>
			<td>16</td>
			<td>23(46)</td>
			<td>5(71)</td>
			<td>12(75)</td>
			<td>17(74)</td>
			<td>123(79)</td>
		</tr>
		<tr>
			<td>wsgRNA-1/wsgRNA-3</td>
			<td>50</td>
			<td>13</td>
			<td>9</td>
			<td>22(44)</td>
			<td>10(77)</td>
			<td>6(67)</td>
			<td>16(73)</td>
			<td>72(81)</td>
		</tr>
		<tr>
			<td>wsgRNA-2/wsgRNA-3</td>
			<td>50</td>
			<td>17</td>
			<td>10</td>
			<td>27(54)</td>
			<td>13(76)</td>
			<td>8(80)</td>
			<td>21(78)</td>
			<td>101(85)</td>
		</tr>
		<tr>
			<td>wsgRNA-1/wsgRNA-2/wsgRNA-3</td>
			<td>50</td>
			<td>11</td>
			<td>10</td>
			<td>21(42)</td>
			<td>9(82)</td>
			<td>9(90)</td>
			<td>18(86)</td>
			<td>149(86)</td>
		</tr>
	</tbody>
</table>
Table 1: Survival and mutation rates of embryos injected with single and multiplexed gRNAs targeting white. The table is reprinted with permission from30

Watch this Scientific Journal Video about Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus at JoVE.com

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.

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, ...

embryo-microinjection-techniques-for-efficient-site-specific

Research

JoVE Journal

Genetics

5.5K Views.  University of California, San Diego. Dit protocol beschrijft micro-injectieprocedures voor Culex quinquefasciatus embryo's die zijn geoptimaliseerd om te werken met CRISPR/Cas9 genbewerkingstools. Deze techniek kan efficiënt site-specifieke, erfelijke, kiembaanmutaties genereren die kunnen worden gebruikt voor het bouwen van genetische technologieën in deze onderbelichte ziektevector.

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

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 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

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 ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

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

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

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

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

Developmental 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 ...

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 ...

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

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.

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.

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

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (&#62;1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily ...

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by filarial nematodes such as canine heartworm and elephantasis. Recently, CRISPR/Cas9 genome editing has been used to induce site-directed mutations by injecting a Cas9 protein that has been complexed with a guide RNA (gRNA) into freshly laid embryos of several insect species, including mosquitoes that belong to the genera Anopheles and Aedes. Manipulating and injecting Culex mosquitoes is slightly more difficult as these mosquitoes lay their eggs upright in rafts rather than individually like other species of mosquitoes. Here we describe how to design gRNAs, complex them with Cas9 protein, induce female mosquitoes of Culex pipiens to lay eggs, and how to prepare and inject newly laid embryos for microinjection with Cas9/gRNA. We also describe how to rear and screen injected mosquitoes for the desired mutation. The representative results demonstrate that this technique can be used to induce site-directed mutations in the genome of Culex mosquitoes and, with slight modifications, can be used to generate null-mutants in other mosquito species as well.&#160;

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by ...

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://www.jove.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.

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

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

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