Biology
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Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis
Chapters
Summary September 13th, 2022
The present protocol describes a method for injecting tick embryos. Embryo injection is the preferred technique for genetic manipulation to generate transgenic lines.
Transcript
This is the first embryo injection protocol for a Chelicerata species. The ability to inject tick embryos will open up many lines of research, in particular, studies into tick gene function and tick pathogen interactions. This technique allows future work on functional studies of tick genes and provides a foundation for tick population control by using gene drive systems.
This method will facilitate the understanding of the interaction between ticks and the pathogens that they harbor on a molecular level. This will allow us to identify proteins that can be used for the development of vaccines for Lyme disease and other pathogens. While several research areas such as tick pathogen interactions and pathogen transmission to hosts by tick during feeding will benefit from this work.
More fundamental questions such as how do ticks evade immune response of host, and differences in DNA repair mechanisms can also be understood now. There are a few aspects of tick biology that need to be understood before carrying out this procedure. The high internal pressure of the embryo leads to bursting when touched by a needle.
It is very important to prepare embryos for injection. To begin, transfer females from four degree Celsius to a 27 degree Celsius incubator for the initiation of egg laying a week before injections. After three to four days when the females start laying eggs, remove any eggs using a fine tipped paintbrush while preparing the females for Gene's organ ablation.
To empty the gland, arrange the gravid tick on a glass slide under a microscope in a way that mouth parts are visible on both ventral and dorsal sides. Then carefully puncture the area behind the mouth parts on the dorsal side, and apply pressure on the ventral side using forceps. Place the edge of a lint-free wipe and remove the liquid wax coming out of the puncture site.
Alternatively, instead of emptying the gland by dissection, glues such as tissue adhesive can be used around the tick mouth parts. When the females begin laying eggs again, use a fine paintbrush to collect freshly deposited eggs and place them in a 1.5 milliliter micro centrifuge tube. Add approximately 200 microliters of 5%benzalkonium chloride water in the tube containing zero to 18 hours old eggs.
Gently swirl the eggs with the paintbrush for five minutes to avoid the eggs settling in the bottom of the tube, and remove the supernatant using a micro pipette, leaving the eggs behind in the tube. Add approximately 200 microliters of distilled water to the tube containing eggs. Swirl with a paintbrush and remove the water from the micro centrifuge tube using a micro pipette.
After washing with distilled water, add approximately 200 microliters of 5%sodium chloride and gently swirl with a paintbrush for five minutes. Then, remove the solution from the tube after five minutes, and wash the eggs twice with distilled water. Next, add approximately 100 microliters of 1%sodium chloride solution in the micro centrifuge tube containing eggs.
First, adhere two glass microscope slides together, leaving a gap of about 0.5 centimeters to support aligning eggs using a double-sided tape, and apply a piece of transparent film dressing on the double-sided tape in the gap between the slides. Then align eight to 10 eggs at a time on the slide setup using a paintbrush. Then place the slide with aligned eggs on the stage of a compound microscope and remove the 1%sodium chloride solution using a piece of lint-free wipe in which they are stored.
Attach the filled injection needle to a micro injector connected to a micro manipulator. Next, open the needle by gently rubbing it against the egg surface using a high pressure setting on the micro injector. After the needle is opened, reduce the pressure of the micro injector depending on the opening of the needle.
Then inject all the eggs on the slide at a 10 to 15 degree angle as quickly as possible, quickly press the foot pedal, and immediately move the slide to high humidity conditions in a Petri dish with a moist paper towel. After five to six hours of placing the slide containing injected embryos in a large Petri dish lined with moist filter paper, add a small drop of distilled water to the transparent film dressing with injected embryos attached. Then, gently displace the eggs using a paintbrush.
Next, transfer the eggs to a small Petri dish and submerge them in distilled water. Keep the eggs submerged in water, place the Petri dish in a six quart plastic box in an incubator at 27 degree Celsius at more than 90%relative humidity. When the larva begin to hatch from the injected embryos 21 to 25 days after injection, check them daily and transfer any hatched larvae to glass vials with a screen on top.
The results demonstrated that injecting the eggs early in embryogenesis 12 to 18 hours old, and aligning the longer axis of the egg perpendicular to the edge of the slide results in a higher survival rate of the injected eggs. Of all the injected eggs, up to 8.5%survived and larva hatched. Before beginning this procedure, it is important to remember steps 2.4, removal of the wax from the mother tick, and steps 3.1 through 3.4, softening of the chorion of the embryo with benzalkonium chloride, and desiccation of the embryo with sodium chloride solutions to enable successful embryo injection.
This method was developed primarily for tick gene editing, and this will allow us to do knockouts and knock-ins that will help us to understand tick gene function. This technique will permit the use of gene editing tools available for many other organisms to be applied to tick research. And this will accelerate the tick molecular biology research.
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