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, ...

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5.3K Views.  University of California, San Diego. 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.

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

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. ...

Tools to Study the Role of Architectural Protein HMGB1 in the Processing of Helix Distorting, Site-specific DNA Interstrand Crosslinks

High mobility group box 1 (HMGB1) protein is a non-histone architectural protein that is involved in regulating many important functions in the genome, such as transcription, DNA replication, and DNA repair. HMGB1 binds to structurally distorted DNA with higher affinity than to canonical B-DNA. For example, we found that HMGB1 binds to DNA interstrand crosslinks (ICLs), which covalently link the two strands of the DNA, cause distortion of the helix, and if left unrepaired can cause cell death. Due to their cytotoxic potential, several ICL-inducing agents are currently used as chemotherapeutic agents in the clinic. While ICL-forming agents show preferences for certain base sequences (e.g., 5&#39;-TA-3&#39; is the preferred crosslinking site for psoralen), they largely induce DNA damage in an indiscriminate fashion. However, by covalently coupling the ICL-inducing agent to a triplex-forming oligonucleotide (TFO), which binds to DNA in a sequence-specific manner, targeted DNA damage can be achieved. Here, we use a TFO covalently conjugated on the 5&#39; end to a 4&#39;-hydroxymethyl-4,5&#39;,8-trimethylpsoralen (HMT) psoralen to generate a site-specific ICL on a mutation-reporter plasmid to use as a tool to study the architectural modification, processing, and repair of complex DNA lesions by HMGB1 in human cells. We describe experimental techniques to prepare TFO-directed ICLs on reporter plasmids, and to interrogate the association of HMGB1 with the TFO-directed ICLs in a cellular context using chromatin immunoprecipitation assays. In addition, we describe DNA supercoiling assays to assess specific architectural modification of the damaged DNA by measuring the amount of superhelical turns introduced on the psoralen-crosslinked plasmid by HMGB1. These techniques can be used to study the roles of other proteins involved in the processing and repair of TFO-directed ICLs or other targeted DNA damage in any cell line of interest.

Targeted DNA damage can be achieved by tethering a DNA damaging agent to a triplex-forming oligonucleotide (TFO). Using modified TFOs, DNA damage-specific protein association, and DNA topology modification can be studied in human cells by the utilization of modified chromatin immunoprecipitation assays and DNA supercoiling assays described herein.

High mobility group box 1 (HMGB1) protein is a non-histone architectural protein that is involved in regulating many important functions in the genome, ...

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most mutagenesis-based screens, researchers have relied on the use of toxic chemicals, carcinogens, or irradiation, which requires designated equipment, safety setup, and/or disposal of hazardous materials. We have developed a simple approach to induce heritable mutations in C. elegans using germline-expressed histone-miniSOG, a light-inducible potent generator of reactive oxygen species. This mutagenesis method is free of toxic chemicals and requires minimal laboratory safety and waste management. The induced DNA modifications include single-nucleotide changes and small deletions, and complement those caused by classical chemical mutagenesis. This methodology can also be used to induce integration of extrachromosomal transgenes. Here, we provide the details of the LED setup and protocols for standard mutagenesis and transgene integration.

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Genetically-encoded histone-miniSOG induces genome-wide heritable mutations in a blue&#160;light-dependent manner. This mutagenesis method is simple, fast, free of toxic chemicals, and well-suited for forward genetic screening and transgene integration.

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most ...

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

We describe a PCR- and homologous recombination-based system for generating targeted mutations in histone genes in budding yeast cells. The resulting mutant alleles reside at their endogenous genomic sites and no exogenous DNA sequences are left in the genome following the procedure. Since in haploid yeast cells each of the four core histone proteins is encoded by two non-allelic genes with highly homologous open reading frames (ORFs), targeting mutagenesis specifically to one of two genes encoding a particular histone protein can be problematic. The strategy we describe here bypasses this problem by utilizing sequences outside, rather than within, the ORF of the target genes for the homologous recombination step. Another feature of this system is that the regions of DNA driving the homologous recombination steps can be made to be very extensive, thus increasing the likelihood of successful integration events. These features make this strategy particularly well-suited for histone gene mutagenesis, but can also be adapted for mutagenesis of other genes in the yeast genome.

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

A strategy for generating mutations in histone genes at their endogenous location in Saccharomyces cerevisiae is presented.

We describe a PCR- and homologous recombination-based system for generating targeted mutations in histone genes in budding yeast cells. The resulting ...

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 ...

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 ...

Gene-targeted Random Mutagenesis to Select Heterochromatin-destabilizing Proteasome Mutants in Fission Yeast

Random mutagenesis of a target gene is commonly used to identify mutations that yield the desired phenotype. Of the methods that may be used to achieve random mutagenesis, error-prone PCR is a convenient and efficient strategy for generating a diverse pool of mutants (i.e., a mutant library). Error-prone PCR is the method of choice when a researcher seeks to mutate a pre-defined region, such as the coding region of a gene while leaving other genomic regions unaffected. After the mutant library is amplified by error-prone PCR, it must be cloned into a suitable plasmid. The size of the library generated by error-prone PCR is constrained by the efficiency of the cloning step. However, in the fission yeast, Schizosaccharomyces pombe, the cloning step can be replaced by the use of a highly efficient one-step fusion PCR to generate constructs for transformation. Mutants of desired phenotypes may then be selected using appropriate reporters. Here, we describe this strategy in detail, taking as an example, a reporter inserted at centromeric heterochromatin.

This article describes a detailed methodology for random mutagenesis of a target gene in fission yeast. As an example, we target rpt4+, which encodes a subunit of the 19S proteasome, and screen for mutations that destabilize heterochromatin.

Random mutagenesis of a target gene is commonly used to identify mutations that yield the desired phenotype. Of the methods that may be used to achieve ...

Universal and Efficient Electroporation Protocol for Genetic Engineering of Gastrointestinal Organoids

Electroporation is a common method for transfection with different kinds of molecules by electrical permeabilization of the plasma membrane. With the increasing use of organoids as a culturing method for primary patient material in the last years, efficient transfer methods of components for genetic engineering in this 3D culture system are in need. Especially for organoids, the efficiency of genetic manipulations depends on a successful transfection. Thus, this protocol was developed to facilitate the electroporation of organoids and to prove its universal functionality in different entities. Human colorectal, pancreatic, hepatic and gastric cancer organoids were successfully electroporated with small and large plasmids in comparison. Based on GFP encoding vectors, the transfection efficiency was determined by FACS. No extensive preparation of the cells or special, cost-intensive electroporation buffers are necessary, and the protocol can be performed within one day.

This protocol describes an efficient electroporation method for the transfection of four different gastrointestinal organoid entities with larger plasmids (to the extent of 10 kB). It can be performed within one day and does not need extensive preparation or special, cost-intensive electroporation buffers.

Electroporation is a common method for transfection with different kinds of molecules by electrical permeabilization of the plasma membrane. With the ...

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 ...