The overall goal of this protocol is to make targeted genomic changes in a variety of cells and organisms using CRISPR/Cas9 ribonucleoprotein complexes. This method can help address fundamental biological questions by giving the researchers the power to generate custom-designed insertions, deletions, and substitutions in otherwise unmodified genomes of model and non-model organisms. The main advantage of using ribonucleoprotein or RNP complexes instead of plasmid DNA is that efficiency is higher, and off target events are reduced.

The implications of this technique extend towards ex vivo therapies to fight things like cancer, HIV, or genetic disease. This is because genome editing can be used to correct harmful mutations or even enhance the ability of cells to fight off harmful diseases. So this method can provide insights into primary human cells, C.elegans, and Parhyale hawaiensis.

It can be applied to make genetic changes in hundreds of other systems including previously intractable organisms. Begin this procedure with design and preparation of guide RNA as described in the text protocol. Assemble an RNP complex by mixing a one to two times molar amount of guide RNA with 200 picomoles of Cas9 protein in a total volume of 10 microliters.

Very slowly, add concentrated Cas9 to the guide RNA for about 30 seconds, making quick circles with the pipette. Bringing the final Cas9 concentration to 20 micromolar. Next, add 10 microliters of RNP to each cuvette well.

After preparing HSPCs as described in the text protocol, count the cells with a hemocytometer and transfer 150, 000 to 200, 000 cells per cuvette to be electroporated to a centrifuge tube. Spin the tube at 100 times gravity for 10 minutes to pellet the cells. Aspirate the supernatant with a pipette or vacuum, removing any bubbles.

Gently resuspend the cells with 20 microliters of electroporation buffer per cuvette well. Then, add 20 microliters of the cells which contain 150, 000 to 200, 000 HSPCs to each cuvette well which already possess 10 microliters of the RNP and mix well by pipetting up and down without creating bubbles. Finally, electroporate the cuvettes after placing them in a Nucleofector.

For the HSPCs, use the pulse code ER-100. Begin with preparation of C.elegans in agarose pads as described in the text protocol. Place an agarose injection pad and cover slip onto a dissection scope.

Use a worm pick to lay a small track of Halocarbon oil along one edge of the pad. Then, use the worm pick coated in oil to lift several worms off the NG agar plate and into the track of oil. With a fine hair attached to a pipette, such as an eyelash or cat whisker, position the worms in parallel, gently pushing the worms into the agarose pad.

Until comfortable with the microinjection procedure only mount an inject one worm at a time. Once in position and attached to the pad, overlay the worms with another few drops of Halocarbon oil from the tip of the worm pick. Place the cover slip with the mounted worms onto the injection microscope.

Under a low magnification, position the worms perpendicular to the injection needle which has been filled with the RNP solution, as described in the text protocol. Switch to a high magnification and reposition the needle adjacent to the gonad arm, corresponding to the region near the nuclei in mid to late pachytene. Using the micromanipulator, move the needle against the worm, depressing the cuticle slightly.

Then, with one hand, tap the side of the microscope stage to jolt the needle through the cuticle. Depress the injection paddle or button, slowly fill the gonad arm with the injection mix and remove the needle. Repeat the step with the other gonad arm.

Once the worms are injected, remove the cover slip and agarose pad and place it under a dissecting microscope. Using a pulled capillary pipette, displace the oil from the worms by pipetting an M9 buffer over them. Perform this treatment to release the worms from the agar.

After 10 minutes, when the worms are thrashing around in the buffer, move them to an NG agar plate with OP50 bacteria using the pulled capillary pipette. Place the plate at 20 degrees Celsius for two to three hours until the worms have recovered and are moving around. Once recovered, individually transfer the worms to NG agar plates with OP50.

Then, transfer the plates to a 25 degree Celsius incubator. Collect the single cell Parhyale embryos at zero to four hours post fertilization by first anesthetizing gravid females with 0.02%clove oil and seawater. Gently scrape the embryos out of her ventral brood pouch using a flame pulled and rounded glass pipette and a dull pair of number three forceps.

Using compressed nitrogen, inject the Parhyale embryos under a dissecting microscope using a microinjector and a micromanipulator. Load 1.5 microliters of the injection mix into the back of a pulled capillary tube using a microloader pipette tip. After setting up the needle on the injection apparatus, break the tip of the needle using a pair of forceps under the dissecting scope.

Calibrate the delivered volume by injecting into Halocarbon oil 700 and measuring the diameter of the bubble. Cut a trough out of the curing agent using a razor blade. Fill it halfway with filter-sterilized seawater and line the Parhyale embryos up in the trough for injection.

Inject the embryos using the microinjection set-up, stabilizing each embryo with a pair of forceps during the injection. After injection, use a glass transfer pipette to transfer the embryos over to a fresh 60 millimeter culture dish filled halfway with filter-sterilized seawater before dissecting and fixing the embryos at various stages. Make dissection needles by threading a bent piece of tungsten wire approximately 0.5 inches in length into the end of an insulin needle.

Use a one milliliter syringe as the handle of the dissection needle and sharpen the needle in sodium hydroxide under a current. Fill one well of a three well glass dish halfway with a freshly-made solution of nine parts PEM buffer, one part 10X PBS, and one part 32%PFA. Place three to five embryos into the dish and poke a small hole into each embryo using a sharp tungsten needle to poke and a slightly dulled one to stabilize, allowing the yolk to flow out and the fixative to run in.

Using a pair of sharpened tungsten needles, gently tease away the outer two membranes surrounding the Parhyale embryo. Dissect them in fixative to make the embryos more robust but work quickly to keep the membrane from becoming fixed to the embryo, which makes membrane removal more difficult. Allow the embryos to fix for a total of 15 to 20 minutes for antibody staining, or 40 to 50 minutes for in situ hybridization.

Sometimes, results of an editing experiment are best assessed through an experimental assay. Representative results of both wild type T cells and CD25 knock out cells are shown here. Flow cytometry shows that cells that have not been edited with Cas9 RNP express CD25.

However, cells in which CD25 has been knocked out do not express the protein. For large scale perturbations, editing outcomes may have clear visual phenotypes. For example, wild type Parhyale hawaiensis have normal abdomens with swimming and anchor legs.

Disrupting the Abd-B gene replaces the abdominal swimming and anchor legs with jumping and walking legs, which are usually only associated with thorax. In addition to checking for phenotypes, experimenters should also analyze genotype to assess overall editing efficiency and to detect specific genetic changes. For single clones, this can be achieved through simple Sanger sequencing.

In mixed populations of cells, more advanced sequencing analysis is recommended. For example, TIDE can map all the different indels that have occurred in individuals cells within an RNP edited pool. When first attempting this procedure, try using one of the positive controls we've listed in table one.

Using a tried and true guide RNA can help you get a feel for editing with RNP before designing your own experiment.