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

Embryo microinjection techniques have paved the way for the development of numerous genetic based vector control tools. Although numerous insect species have protocols already well-developed, Culex quinquefasciatus remains relatively under-studied. This microinjection protocol has been specifically designed to accommodate the unique biological traits of Culex quinquefasciatus, including egg collection, egg raft separation, and post-injection procedures.

This method can likely be adapted to any Culex species of interest, and can be used to study gene function or to generate genetic based control technologies for Culex species. Begin by separating the eggs from the rafts. Use forceps or a paintbrush to press down on the raft and tease apart the individual eggs.

Align the eggs on a thin strip of double-sided sticky tape placed across the top of a glass slide, making an effort to point the anterior side of each egg in the same direction. Prepare the halocarbon mix by gently mixing halocarbon reagents with water, and incubating the mixture at 25 degrees Celsius overnight. Cover aligned eggs with the halocarbon oil mixture.

To generate needles for the microinjections, place an aluminosilicate capillary glass into a needle puller, following the manufacturer's instructions. Set the heat to 560, velocity to 100, delay to 70, pull to 97, and pressure to 500. Then activate the needle puller.

Gently touch the tip of the pulled needle on the rotating diamond abrasive plate for around 10 seconds at a 50 degree angle to bevel the needle tip. Embed pulled and beveled needles into lines of modelers'clay in a petri dish. Prepare the injection mixture consisting of genome modification reagents and store it on ice.

Use a microloader tip to load two microliters of injection mixture into the injection needle. Place the filled injection needle into a micromanipulator linked to an electronic microinjector and place the glass slide with the aligned eggs on the stage of a compound microscope. Using the micromanipulator, align the needle to aim at the posterior end of the embryo at a 25 to 35 degree angle.

Carefully insert the needle into the embryo and inject the mixture at a quantity of about 10%of the volume of the embryo. Inject 20 eggs at a time, then stop and perform embryo recovery procedures. Within 20 minutes post-injection, carefully remove the halocarbon oil from the eggs by brushing them lightly with a clean paintbrush.

Lift the eggs with the paintbrush and place them into a cup of double distilled water, taking care to keep the eggs on the surface. Over the next seven days, check the eggs daily for hatching and follow normal larval rearing procedures. Screen the injected mosquitoes for the mutant phenotypes using a stereoscope.

This method has been used to successfully generate somatic and germline mutations of a gene critical for the development of dark eye pigmentation. CRISPR-Cas9 generated somatic mutations were scored by screening for loss of pigmentation in the pupil stage eyes of injected individuals. Somatic mutations were generally present as mosaic phenotypes, where some but not all of the cells have the mutant phenotype.

Germline mutation rates were determined by intercrossing mosaic G zero individuals and scoring per completely white-eyed offspring. These experiments resulted in a 64 to 82%embryo survival rate, a 37 to 57%somatic mutagenesis rate, and a greater than 61%germline mutagenesis rate. By multiplexing sgRNAs to target multiple loci in the same gene, somatic and germline mutagenesis rates increased to as much as 86%In addition, for many generations, viable homozygous stocks of the white mutants have been successfully kept in the lab.

To optimize survival and mutagenesis rates, all materials, including halocarbon oil, injection fluid, and needles, should be properly prepared before attempting microinjections. This will allow for a more streamlined and focused microinjection process.