Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

The overall goal of this procedure is to achieve high efficiency, plasmid, transfection of retinal cells to enable functional genetic studies in primary retinal cultures. This is accomplished by first collecting chicken embryos at stage 27 of the hamburger and Hamilton system. Appropriate staging is critical for achieving optimum efficiency.

The second step is to dissect the embryonic eyes and remove the retinal pigmented epithelium exposing the neural retina. Next, the retinal cup is placed in a chamber filled with plasmid solution. It is then subjected to electroporation under optimized conditions using easily constructed custom made electrodes.

The final steps are to remove the vitreous body and lens and dissociate the neural retina to single cells for low density primary culture. Ultimately, transfected cells can be visualized by the expression of reporter genes or immunohistochemistry, and the effective genetic manipulation can be assessed by evaluating the expression of downstream genes and or phenotypic cell changes. The main advantage of this technique over existing methods, such as lipofection or chemical transfection, is that it increases the rate of transfection of retinal cells by fivefold yielding efficiencies of 22 to 25%of the total cell population.

This method can help answer key questions in the field of photoreceptor biology, since it allows for the manipulation of gene expression in primary retinal cultures at a level of efficiency that can provide biologically meaningful insights. The implications of this technique extend beyond basic research as they can easily be adapted to high throughput technologies for drug development. Begin by incubating fertilized white leg horn chicken eggs at 37.5 degrees Celsius and 60%relative humidity for five days in an egg incubator.

Prior to the experiment, obtain a box filament normally used in micro pipette pulls. Use forceps to undo the square box and straighten the filament into a long strip. Then using a ruler as a guide, bend the filament into a square u u-shape, three millimeters long at the base and two millimeters high at one side with the other side, the remaining length of the filament use forceps to dip the electrode in 96%ethanol and briefly flame the electrode in order to sterilize it.

Then attach an electrical lead with a small alligator clip to the long arm of the electrode. Next wipe, clean the thick gold tipped electrode using 70%ethanol. Then prepare the electroporation chamber by cutting the lid off of a sterile 1.5 milliliter micro centrifuge tube using sterilized scissors and affixing it flat side down to a 100 millimeter Petri dish using tape.

Next, sterilize the microdisect tools by dipping the instrument tips in 96%ethanol and briefly flaming them. Also, wipe clean the dissection scope using 70%ethanol. Warm a bottle of sterile calcium and magnesium free Hanks balance salt solution in a 37 degree Celsius water bath.

Once warm, use it to fill a 100 millimeter sterile Petri dish three quarters of the way full. Next place the electroporation chamber under a clean dissection microscope. Then connect the electrodes to the electroporation apparatus, making sure the custom made electrode is connected to the negative pole, and the gold tipped electrode is connected to the positive pole of the machine.

Clean the eggs by wiping the exterior of the shells with 70%ethanol. Be cautious and maintain sterility throughout the rest of the protocol. To avoid carrying contamination into the cultures using big curved Maloney forceps, carefully open a hole in the eggshell by gently tapping on the rounded tip of an egg directly over the air chamber.

Pull out all of the shell pieces with another pair of forceps. Remove the membranes and collect the embryo by carefully scooping it out of the egg. Place the embryo in a Petri dish with the prewarm calcium and magnesium free Hanks Balance salt solution.

When an appropriate number of embryos have been collected, ensure they are all at the correct stage. According to the hamburger and Hamilton staging system for optimum efficiency, embryos must be at stage 27. Euthanize the embryos by decapitation using maloney forceps, and then use dumont tweezers to carefully and nucleate the eyes.

Transfer the eyes to a new Petri dish with calcium and magnesium free HBSS. Start at the posterior part of each eye close to the optic nerve head, and then introduce one tip of each pair of dumont tweezers between the neural retina and retinal pigmented epithelium. Working from the posterior part of the eye towards the front, use the tweezers to carefully dissect out the retinal pigmented epithelium.

Being cautious not to damage the neural retina. Set up the electro perter to deliver five square pulses of 15 volts at a 50 millisecond duration and an interval of 950 milliseconds. Next, fill the electroporation chamber with 120 microliters of plasmid solution at a concentration of 1.5 micrograms per microliter in PBS.

Then place the U-shaped negative electrode inside the micro centrifuge lid electroporation chamber. Using a pair of curved dumont tweezers. Transfer one retinal cup to the electroporation chamber and place it inside the U-shaped electrode.

Position the retina so that the posterior part of it is towards the bottom of the electrode and the lens is facing upwards. Next, place the anode so that it touches the anterior part of the eye next to the lens. Then use the electro raider's foot pedal to deliver five electric pulses.

When finished, transfer the electroporated retinal cup to a new Petri dish filled with calcium and magnesium free HBSS. Repeat the electroporation procedure with each additional retinal cup. Up to six retinal cups can be electroporated with the same plasmid solution without a significant decrease in transfection efficiency.

Next, use bond toothed forceps to gently remove the lens and vitreous body from each electroporated sample while holding the eye in place with the back of the curved dumont tweezers once removed, proceed to the dissociation and culture of the electroporated retinal cells using standard techniques. The retinal cells shown here were electroporated with a GFP expression, plasmid, dissociated, and cultured for four days. Fluorescent micrographs showed dappy stain nuclei and GFP expressing retinal cells.

Anti visin antibody staining was performed to identify photoreceptor cells. The parameters described in this protocol have been optimized to obtain transfect efficiencies that average 22%when compared to the total population after four days. In culture, this measure increases to an average of 25%efficiency when only the photoreceptor population is considered Considered once mastered.

This technique from embryo collection to electro operation can be done in less than 30 minutes for six eyes if it is performed properly. While attempting this procedure, it's important to remember that no more than three hours should pass between collection of the embryos and dis dissociated cell plating in order to obtain a good quality culture. This technique can be applied to gene overexpression or correlation, for example, using an RNAi system.

In addition, retina that have been transfected following this procedure can be used for organotypic retinal cultures.