Induction of Protein Deletion Through In Utero Electroporation to Define Deficits in Neuronal Migration in Transgenic Models

The overall goal of the following experiment is to analyze defects in neural development and migration in transgenic mouse models using in utero electroporation of various regions of the embryonic forebrain. This is achieved by injecting a plasmid carrying genes for Cree and a reporter protein like GFP into the ventricle of an embryonic brain and applying an electric current to the head to draw that plasmid into the cell lining of the ventricle. As a second step, the embryos are returned to the dam's abdominal cavity to allow the embryos to continue developing while crease expressed and mediates gene excision specifically in the electroporated neurons.

Next embryos are collected and the brains are fixed for analysis in order to determine the effect of CRE mediated gene excision and subsequent protein knockdown in the electroporated neurons results are obtained that show how excision of the gene of interest regulates neural development and migration in a cell specific manner using immunohisto chemistry. The main advantage of this technique over existing methods such as tissue specific transgenic CRE lines, is that electroporation of Cree excises the gene of interest only from a small population of neurons in a region of the brain chosen by the researcher, this greatly simplifies results and facilitates phenotype analysis. Visual demonstration of this technique is critical because the surgical steps can be very difficult to learn, and the embryos at this stage are very delicate and easy to inadvertently damage.

Before the surgery, dilute a single plasmid to two micrograms per microliter in sterile water with one microliter of 0.1%weight per volume fast green dye per 10 or 20 microliters of diluted plasmid solution. Next, pull the glass micro capillary tubes on a standard micro pipette polar at a 62 degree Celsius single step. Then use the micro loader pipette tips to back load the pulled needles with plasmid solution.

Do not fill all the way into the tip of the needle, as this will cause the needle to become clogged. In this procedure, make an incision in the skin over the abdomen between the lower tets of an anesthetized pregnant mouse. Pull the skin away from the underlying muscle layer by tearing the connective tissue.

Then cut through the peritoneum into the abdominal cavity along the linear elbow. Insert the wound retractor to pull the edges apart. Then remove the uterus from the abdominal cavity and lay it out as shown.

If different pups are to be injected with different plasmids, count the pups on each side and decide which ones and how many to inject with each plasmid return one side of the uterus back into the abdomen to keep the embryos warm and lubricated immediately moisten the exposed uterus with warm saline. Next, put a loaded needle in the grip head of the injection wand. Carefully cut the tip with a pair of small surgical scissors or fine forceps in such a way that the needle stays as long as possible while still allowing fluid to pass through.

Then adjust the length of injection and the injection pressure on the micro injector so that a small yet easily visible droplet of 0.1 to 0.2 microliters appears on the tip of the needle. When the foot paddle is pressed, keep the volume of this droplet as consistent as possible between different needles. Now select an embryo and gently position it on its back with the head tilted upward.

Once the embryo is in place, locate the ventricle, which presents as a crescent shaped shadow parallel to the sagittal sinus. Touch the uterus above the ventricle at a slight angle with the tip of the needle and guide the needle through the uterine wall. Then push the needle through the cortex into the ventricle.

Next, pump the foot paddle to inject the plasmid solution. Continue to pump until a green area is visible and conforms to the arc shape of the ventricle. In young embryos, the dye will often diffuse into the contralateral ventricle.

Inject about three or four embryos altogether and keep all the exposed embryos moist with saline throughout the procedure. Starting with the first embryo injected, place the paddles such that the positive electrode will draw the plasmid into the desired region of the brain. Angle the negative electrode so that the shortest path between the electrodes passes directly through the structure targeted for electroporation.

Once the paddles are in place, apply the shock. Depending on the age and size of the embryos, use five pulses of 35 to 50 volts, five millisecond pulse length, and 950 millisecond pulse interval. For electroporation, repeat with all the injected embryos.

Afterward, return both sides of the uterus to the mouse. Be careful to keep the surface moist with saline and to not apply too much pressure to prevent trauma to the amniotic sax. Next, suture the muscle layer using a simple continuous stitch.

Then staple the skin closed. Now prepare four degrees Celsius filtered 4%PFA. After euthanizing the animal with an intraperitoneal injection of a lethal dose of sodium pentobarbital, cut open the abdomen.

Then pull both sides of the uterus out and lay them out as when the pups were initially injected. If the pups were injected with different plasmids, be sure to locate each pup noting any dead pups, and determine which ones were injected with control and experimental plasmids. Before removing the uterus from the female, remove each pup from the uterus and amniotic sack individually.

Decapitate with sharp scissors or a razor blade, and place the head in PBS. In the case of transgenic mice, collect tissue samples for genotyping and keep each head separate, such as in a 12 well plate under a dissecting scope, remove the brain from the skull, all being careful not to nick the cortex or tear the midline. Using a fluorescent microscope check to see if the brains have a GFP positive patch indicating that they were electroporated successfully.

Afterward, fix the brains in PFA at four degrees Celsius overnight. Now, heat citric acid where the pH was previously set to six in a staining chamber in a microwave or water bath until it reaches 75 to 80 degrees Celsius. Then put the slides in the staining chamber for 10 to 15 minutes.

Afterward, wash the slides in PB S3 times for three minutes each time at room temperature. Apply the primary antibody and proceed with immuno staining according to a standard protocol. Then validate CRE mediated excision by co labeling for GFP and the gene of interest to confirm knockdown.

This figure shows the confirmation of CRE mediad excision by cos staining for a downstream target of the deleted gene. RB Flocked, P 1 0 7 knockout P one 30 flocked embryos were electroporated with either the empty PCI two plasmid or the PCI two Cree plasmid and stained for GFP and KI 67. A marker of cell proliferation.

Double stained cells were seen in neurons, electroporated with Cree, but not in neurons. Electroporated with psig. Two alone.

Excision of RB was also confirmed by performing PCR with genotyping primers on DNA extracted from the tissue sections removed from the slides. The upper band represents the F flox allele and the lower band represents the wild type allele. The arrow indicates the excised allele in the electroporated side of the brain.

Only CRE mediated excision can also be confirmed in vitro using plasmids with flocked stop codons. Upstream of RFP such as NL DS Red RFP is only expressed in he EK cells when cot transfected with Cree GFP in HEK 2 93 cells indicating that excision was successful Following this technique. Other methods such as cell culture, immunohistochemistry or electrophysiology can be used in order to answer additional questions regarding the development and function of individual neurons following c mediad gene excision.

After watching this technique, you can see how in utero electroporation can be used to target the different regions of the brain, and this can really help understand some of those complex phenotypes that we have with different transgenic animals.