كفاءة الإنتاج وتحديد كريسبر/Cas9-المنشأة الضربة القاضية الجينات في "النظام النموذجي" دانيو rerio

The overall goal of this procedure is to perform targeted genome editing using CRISPR/Cas9 technology in zebrafish to engineer gene knockouts in a robust, inexpensive fashion. This method can help answer key questions about the biological role of a gene in the context of a vertebrate model system such as how a gene contributes to developmental processes in higher order eukaryotes. With these simple robust procedures, laboratory personnel of all levels of experience including undergraduates can perform targeted genomic modifications in zebrafish.

Individuals new to this method should be able to follow the step by step instructions to identify optimal genomic targets for CRISPR mutagenesis, perform zebrafish embryo microinjection and identify modified alleles. Visual demonstration of this method is critical as this method is intended for undergraduates who may be unfamiliar with one or more of the necessary techniques. Guidelines for the design of template specific oligos for guide RNA production are provided in the text protocol.

To begin, suspend the Cas9 protein in buffer to generate a one milligram per milliliter solution. Store this solution in injection ready aliquots at minus 80 degrees Celsius for later use. Then synthesize the sgRNA with an in vitro transcription kit such as the one suggested in the materials of this article per the manufacturer's instructions.

Purify the synthesized sgRNA using an RNAse free ammonium acetate precipitation. Add 25 microliters of five molar ammonium acetate to the synthesized RNA and mix the solution by vortexing. Next, add 150 microliters of 200 proof nuclease free ethanol to the sample.

And place the reaction in a minus 80 degrees Celsius freezer for at least 20 minutes. Afterwards, centrifuge the samples at maximum speed. Use a pipette to remove the supernatant taking care not to disturb the RNA pellet.

Then add one milliliter of 70%ethanol and invert the tube several times to wash residual salt from the tube. Centrifuge again at maximum speed at four degrees Celsius for seven minutes. Use a P1000 pipette to remove most of the solution.

Next use a P200 pipette to remove as much of the remaining solution without disturbing the pellet. Taking care to avoid RNAse contamination, dry the pellet in a clean space until there is no liquid drops visible in the tube. Resuspend the pellet in 30 microliters of RNAse free water and use a spectrophotometer to quantify the product.

Aliquot the solution for long term storage at minus 80 degrees Celsius. Next mix 300 to 500 nanograms of sgRNA with an equal volume of 2X RNA gel loading dye. Then, using a P1000 pipette, clear each of the agarose gel wells of debris with running buffer several times.

Load the solutions into the agarose gel wells and run the gel at 10 volts per centimeter for sufficient time to separate the RNA bands on the gel. Time may vary per electrophoresis apparatus. In general, the dye front should be run to 2/3s the length of the gel.

Then use a nucleic acid stain to visualize the bands. Intact RNA appears as a single band such as in lane one and two shown here. Degraded RNA runs as a smear such as in lane three.

First, prepare the zebrafish subjects for breeding the night prior to performing injections as described in the text protocol. Then combine Cas9 protein and sgRNA in a two to one ratio. Incubate the solution at room temperature for five minutes while the Cas9 and sgRNA form a ribonucleoprotein complex.

Then add 0.5 microliters of 2.5%phenol red solutions in sufficient RNAse free water to achieve a final volume of five microliters. To prepare the injection needle, use a micropipette puller to pull a one millimeter glass capillary. Then cut the tip of the resulting glass needle with a razor blade to obtain an angled opening.

Place the needle in a micromanipulator attached to a microinjector. Using a light microscope, adjust the injection pressure until the needle consistently injects one nanometer of solution in the Petri dish. Before performing embryo microinjection, practice preparing needles and injecting control embryos using a dye only solution and record the survival after 24 hours of development.

You should be able to obtain great than 90%survival compared to an uninjected control. Next, select 10 to 15 embryos as a control population and place them in a separate labeled Petri dish. Using a transfer pipette, carefully line the remaining embryos up on a room temperature injection plate.

Under a dissection microscope, inject one nanoliter of the solution into the yolk sac of each embryo. Then return the injected embryos to a labeled Petri dish and cover them with 1X E3 media with methylene blue. Place the injected embryos in an incubator at 28 degrees Celsius for 24 hours.

Then inspect the injected embryos to remove dead or abnormally developing individuals. First, collect two sets of five 72 hour old embryos from the plates containing injected embryos and one set of five 72 hour old embryos from the uninjected control group into the microcentrifuge tubes. Gently pipette the media off of each embryo set and anesthetize them.

Remove the anesthetic after two minutes and add 45 microliters of 50 millimolar sodium hydroxide. Then incubate the samples at 95 degrees for 10 minutes. Next, remove the samples from the incubator and allow them to cool to room temperature.

Add five microliters of one molar pH tris hydrochloride and vortex the samples vigorously. Then centrifuge the solution at maximum speed at room temperature for three minutes. Transfer the supernatant which contains the genomic DNA to a new labeled tube and store the gDNA at minus 20 degrees Celsius.

Use the primers designed to amplify around the sgRNA target region as described in the text to amplify a PCR fragment for heteroduplex analysis. Use a PCR clean up kit to purify the products of the PCR reaction. Dilute the samples in 30 microliters of water and use a spectrophotometer to quantify the DNA.

Place the tubes containing the PCR amplicons in a floating rack in a boiling water bath. Next prepare a 15%polyacrylamide TBE gel using 30%polyacrylamide. After the gel sets, place it into an electrophoresis apparatus with TBE running buffer.

Then load 500 nanograms of the reannealed PCR products and load a control next to each set of sgRNA samples. Run the gel at 150 volts for two and a half hours or until the dye front is at the bottom of the gel. Grow up injected zebrafish embryos to adulthood that show heteroduplex bands in the HMA.

To interpret the HMA, compare the intensity of the homoduplex band from the PCR of the wild type uninjected control to the corresponding injected samples. If sufficient cutting has occurred the intensity of the band from the injected fish should be approximately 50%lower than the wild type control. To confirm the presence of indels in potential founder parent fish, anesthetize the fish in 0.004%tricaine and use a clean razor blade to remove 1/2 to 3/4 of the tail fin.

Then, place the tail in 45 microliters of 50 millimolar sodium hydroxide. Return the fish to a recovery tank and perform the gDNA extraction as previously described. Next, perform an HMA on the tail clip gDNA to determine if the individual was successfully modified by CRISPR injection.

Then breed the adults that exhibit heteroduplex bands in tail DNA to wild type fish. Select founder fish to generate F1 fish based on HMA analysis of pools. Finally, sequence the gDNA of the fish of interest to characterize the nature of the indel.

Following successful production and microinjection of CRISPR reagents, zebrafish embryos were analyzed for overt phenotypes and indel formation using HMA. Cas9 induced indel formation at tyrosinase results in loss of pigmentation and is easily scored by 48 hours post fertilization. Identification of CRISPR modified alleles that do not result in overt developmental phenotypes is performed using HMA.

Successful targeting of the gene of interest results in formation of heteroduplex bands and reduction of the intensity of the homoduplex band such as in lanes three and four. Don't forget that working with a vertebrate organism carries ethical responsibilities. This protocol describes strategies that minimize the number of zebrafish needed to obtain a knockout, a goal that should be taken into consideration while performing this procedure.

Once mastered, this technique can be used to quickly generate and identify zebrafish carrying CRISPR modified alleles that will be transmitted in the germline with high efficiency. Importantly, this technique has been optimized to be a low cost approach amenable to undergraduates and others with little prior experience. After watching this video, you should have a good understanding of how to easily design and implement a CRISPR based gene knockout approach using zebrafish.