In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

In utero, retinal electroporation begins with an abdominal incision on an anesthetized E 13.5 to 14.5 stage dam to expose the embryonic chain. DNA is injected into one retina per embryo and heads are electroporated causing DNA to be taken up by RGC progenitors. At E 18.5 embryos are perfused and GFP positive.

Retina and visual projections are analyzed for X vivo retinal electroporation. E 13.5 to 14.5 heads are pinned dorsal side up in a dish and DNA is injected into the peripheral dorsal region of both retina, followed by electroporation. Whole retina are cultured for 40 hours, followed by plating GFP positive retinal explants for in vitro culture assays.

Hi, I’m Tim Petros from the laboratory of Dr.Carol Mason in the Department of Pathology and cell biology and neuroscience at Columbia University. Today we will show you how to introduce genes into embryonic mirroring retina by utilizing in utero and ex vivo retinal electroporation. We use this procedure in our lab to study how manipulating gene expression and retinal ganglion cells alters their projection pathway specifically at the optic chiasm.

Okay, let’s get started. Before attempting in utero retinal electroporation, proper preparation is essential. Please see our written protocol for detailed information regarding preparing the anesthetic sterilization of tools and instrumentation, preparing the DNA solutions for injection and antibiotic anti mycotic solutions for after the surgery, prepping the surgical area and warming the proper solutions.

Fine tipped micro pipettes should be prepared prior to the surgery using a micro electrode puller. Break the tip to make an angled point for in utero electroporation First inject 0.15 milliliters of the anesthetic mix interperitoneal into an E 13.5 to E 14.5 stage dam. It should take three to seven minutes until the dam is fully under anesthetic to test if the mouse is anesthetized.

Pinch the hind paw and check for a lack of response during the interval where the animal is succumbing to anesthetic. Cut a small piece of and pipette five microliters of the DNA solution onto the param. Bend a metal wire plunger at a 90 degree angle and insert it into a pulled micro pipette using the plunger aspirate the DNA solution into the micro pipet.

Now surgically expose the mouse embryos. Please see the JoVE protocol by Wal Inus et al for the proper technique. Now we’re ready for the electroporation.

The first step is to orient the retina properly. Then we’ll inject DNA into the retina and then we will electro parade the head. Okay, let’s get started.

Okay, the first step is to orient the retina so that it is facing up like this one. Then with the micro pipette, we will pierce the amniotic wall and ideally inject the DNA directly into the retina. Once you have pierced through the uterine wall, you will depress the plunger, which will expel the DNA into the retina and pull back on the plunger.

As such, once you’ve injected the DNA, you should see the green dye filling the amniotic fluid and also the retina. Okay, now that we’ve done the injection, we’re ready for the electroporation. So to electro pray, we’re gonna place the electroporation paddles around the embryo with the positive paddle adjacent to the retina that was electro paraded.

And when ready, a depressive foot paddle and that will deliver current to the embryo. You see bubbles forming on the negative charge pan paddle and that shows that the electroporation was a success. And you’ll repeat this step for all of the injected embryos.

So that is how you electro parade DNA into one embryonic retina. You can repeat this procedure up to as many embryos as you’d like. Keeping in mind that the more embryos you electro parade, the greater chance that the mother will abort the embryos.

After electro parading the embryos, use your fingers to massage the embryonic chain back into the dam once the embryos are back in. Pour about two milliliters of the antibiotic anti mycotic solution into the abdominal cavity. Use the hemostat and forceps to suture the peritoneum starting with a double knot on the anterior portion of the incision and continue in a simple continuous suture pattern to the posterior portion of the incision.

Use the final loop to tie off the suture and trim excess suture. Finally, staple the incision closed. To do this, lift the skin at the anterior portion of the incision with blunt forceps.

Be careful to separate the skin from the peritoneum. Next, place the teeth of the stapler around the skin and to depress the stapler, continue stapling the incision closed to the posterior end, which usually requires five to seven staples. Once finished, stapling, wipe the wound around the staples with an alcohol pad.

Let the dam recover on the heating pad and then place her in a new cage. When she begins to twitch and roll over, it usually takes between 15 to 90 minutes for her to wake up to minimize pain and discomfort. Mothers receive buprenorphine upon awakening and at later time points.

If discomfort continues to prevent dehydration, inject the dam with one milliliter saline subcutaneously about two to four hours post-surgery. After 24 hours post-surgery, the dam should regain normal behavior and be drinking and eating regularly. We will harvest the electroporated retina at E 18.5 for analysis.

Alternatively, you can let the dam give birth and harvest the retina at postnatal ages for analysis at later ages. After anesthetizing the animal, use scissors to cut the skin and peritoneum underneath the staples. To expose the abdominal cavity and uterine horns, remove an electroporated embryo and pin the embryo through the limbs to a dissecting dish ventral side up cut through the abdominal skin and peritoneum and cut through the lateral portions of the rib cage to expose the heart.

Next, pierce the left ventricle with the perfusion needle and cut the right atrium with the micro scissors. Now start the perfusion and allow PFA to flow for about 60 to 90 seconds. If done correctly, blood should exit the right atrium and the embryo should become pale.

Once finished with the perfusion, decapitate the embryo and collect heads in 4%PF in a 15 milliliter conical vial. Repeat this procedure for all electroporated embryos and all dams. Once finished collecting all the embryos, sacrifice the dam via cervical dislocation.

Keep heads in 4%PFA overnight at four degrees Celsius and then wash in PBS for several days after washing with PBS, you are now ready to examine retina for successful GFP expression. First pin the head in a dissection dish with a retina facing up and immersed in PBS. Remove skin from around the retina to reveal the retinal pigment epithelium or RPE using a 26 and a half gauge needle.

Puncture the ventral retina at the lens RPE junction. Next, insert one edge of the micro scissors into this hole and make a vertical incision through the ventral retina to the optic disc. This will allow you to orient the retina when you take it out.

Separate the RPE from the retina by grabbing the RPE at the ventral incision with one pair of forceps. And use another pair of forceps to peel the RPE away from the retina, specifically at the RPE lens junction where the RPE is firmly attached. After removing the RPE, grasp the lens with forceps and pull up and away.

To remove the lens, carefully remove the retina by placing forceps underneath the retina and pinch the optic nerve head to free the retina. Finally transfer the retina to A PBS filled. Well be sure to keep track of which retina came from, which heads repeat these steps for the contralateral retina and for all other electroporated embryos, use a fluorescent dissecting microscope to scan all retina for GFP positive retina, which would indicate that the electroporation was successful in this embryo and these retina and corresponding brain can be processed for further analysis.

The ex vivo retinal electroporation procedure requires several additional preparatory steps that are distinct from in utero electroporation. Please see the written protocol for detailed information on preparation and oxygenation of serum free medium and preparation of culture dishes and border assays. After anesthetizing the animal, the abdominal cavity is opened as previously demonstrated and the entire embryonic chain is removed and placed in D-M-E-M-F 12 on ice.

The dam should then be euthanized using cervical dislocation. Use scissors to cut through the uterine wall, then remove embryos from the amniotic sac and collect in D-M-E-M-F 12 and keep on ice decapitate embryos and keep in DMEA medium on ice bodies can be discarded. Next, use a mouth pipette or a plunger pipette and fill a micro pipette with the desired amount of DNA solution.

Then transfer one. Head to a dissecting dish with PBS and pin the head dorsal side up. Move to the dissecting scope for the injections.

Okay, now we’re ready to electro prate. I’m going to inject the DNA into the peripheral dorsal of both retina and then electro prate the head. Okay, here I go.

The first step is to remove the skin from above the retina. To do that, we’ll use the tweezers just to peel away the skin. Now we can see the dorsal portion of both retina.

Next I’m gonna to inject DNA just underneath the retinal pigment epithelium layer in both retinas. To do so, I’m gonna hold the electrode at a almost horizontal position and pierce through the RPE on the most peripheral dorsal part of the retina. You should see the green dye fill the retinal space and leak out into the PBS solution.

Okay, now it’s time for the electroporation. First you unpin the head. Now we’re gonna electro prey the paddles and we want the positive paddle to be on the ventral side of the head.

So we’ll place the head within the paddles with the positive paddle on the ventral surface and we’ll apply pressure to keep the paddle steady like so. And now we’ll electro parade. You can see the bubbles forming on the paddle and on the top of the head.

And that’s it. Now we’ll move the head to a dish with median. So that’s it for the ex vivo retinal electroporation.

Repeat this procedure for as many embryos and DNA conditions as you like. Now we’re gonna get ready to culture the retina. After completing all electro preparations, add about 1.5 milliliters of oxygenated SFM solution to a four well dish and keep it on ice.

Have one well for each different DNA solution injected. Next, transfer one electroporated. Head to a dissecting dish with D-M-E-M-F 12 with one retina facing up on ventral retina or the side opposite the target region.

Use fine forceps to pinch and tear a hole in the RPE. Use this hole as a starting point to peel back RPE from around the retina. Do not cut or damage the eye as this will cause the retina to collapse onto itself during incubation.

After removing the RPE use forceps to pop out the retina by pinching the optic nerve at the base of the retina leave lens intact. Transfer the retina to the oxygenated SFM in the four well dish flip. Head over and repeat the retina removal for the contralateral eye.

Complete this step for all electroporated heads and incubate the retina in the oxygenated SFM at 37 degrees Celsius for 40 to 48 hours. After incubation of whole retina transfer retina from one well of oxygenated SFM into a Petri dish lid with D-M-E-M-F 12. Using a fluorescent dissecting scope, choose one retina and visualize GFP positive expressing portion of retina.

Next, using a micro scalpel and forceps, dissect and discard the non GFP positive portion of retina so that only A GFP positive chunk of retina remains. It is helpful to continually switch between fluorescent and regular light throughout the dissection. To specifically select the GFP positive retina transfer the GFP positive piece of retina to a well with G-M-E-M-F 12.

Repeat this step for all retina from one Well, for each DNA condition, use a new Petri dish and D-M-E-M-F 12 medium to dissect these GFP positive pieces of retina into smaller retinal explan for plating. Use a dissecting scope to cut the large piece of GFP positive retina into smaller explan with a micro scalpel. The explan should be about 200 to 300 micrometers length and width.

One GFP positive chunk of retina should produce about four to 10 GFP positive retinal explan Collect all explan in a well with D-M-E-M-F 12. Repeat this step for all GFP positive retinal regions that are harvested. After preparing all GFP positive retinal explan add 250 microliters of cold SFM plus 0.4%methylcellulose to laminate coated culture dishes.

Next, transfer four to eight GFP positive explan to the culture dish and use the forceps to gently arrange the X explan into a polygon with x explan located halfway between the center and the edge of the dish. Determine which side of the X explan is the RGC layer. This can be done by looking for RPE crystals, which can sometimes remain attached to the outer retinal layer indicating that the opposite side is the RGC layer.

The RGC side also usually has some cellular debris that can aid in the proper explanation with the RGC layer down. Use forceps to firmly depress the center of the explan so that it will adhere to the glass cover slip on the base of the culture dish. After plating all X explan on a culture dish transfer to a 37 degrees Celsius incubator.

Retinal X explants display abundant axon growth after 24 hours and can be used for numerous in vitro assays, then fixed with 4%PFA for 30 minutes. And immuno stain for the border assay. Transfer 4G FP positive explan to a dish that has been prepared for the border assay.

Arrange the X explants in a straight line that approximates the fluorescent border with your substrate of interest. Under a fluorescent dissecting microscope, use forceps to plate the EXPLAN so that they are about 50 to 150 micrometers from the fluorescent border. Alternating between the two fluorescent channels to visualize the green explan and the red border.

Next incubate dishes at 37 degrees Celsius for 24 hours, allowing ample time for RGC axons to reach the border substrate. X explan can then be fixed in 4%PFA for 30 minutes followed by immuno staining. Following in utero retinal electroporation.

Retinal hole mounts can be prepared to visualize GFP positive R GCs in the retina and semi intact. Visual systems can be prepared to visualize GFP positive axons throughout the optic nerve optic chiasm and optic tract. Alternatively, frontal cryo sections can be made through the retina and RGC pathway to visualize and analyze GFP positive RGC cell bodies in the retina and axons in the optic nerve chiasm and tract.

All retinal axons are visualized with neurofilament staining. In addition in utero, electroporation can be used to study RGC axon targeting in the lateral gen nucleus LGN by harvesting pups at P four to P 10. At P eight GFP positive axons are visible in the LGN with CTB LOR 5 94 red channel injected into the ipsilateral retina and CT bor 6 47 Blue Channel into the contralateral retina to label the entire RGC projection following ex vivo electroporation and plating of retinal implants.

A subset of RGC axons RGFP positive and can clearly be visualized both at lower and higher power as seen in the top and bottom images respectively. Response of GFP positive axons can be analyzed by plating XS adjacent to borders of various substrates shown here in blue. We’ve just shown you how to introduce genes into embryonic mirroring retina by utilizing in utero and ex vivo retinal electroporation.

While we have focused our analysis on axon guidance of the optic chiasm, this technique could be beneficial for those studying RGC differentiation, dendritic morphology, and electrophysiological properties of retinal ganglion cells. When doing this procedure, it’s important to remember to minimize the damage to the retina when injecting the DNA and to always keep the embryos hydrated with PBS throughout the experiment. So that’s it.

Thanks for watching and good luck with your experiments.