The aim of this procedure is to assess axon guidance defects in the chicken neural tube after downregulating gene expression in a cell type specific traceable manner. This is accomplished by preparing fertilized chicken eggs for innovo electroporation by cutting a window in the eggshell. The second step is to inject and electro operate DNA constructs encoding microRNAs into the neural tube.
Next, the electro rated spinal cords are dissected and fixed in flat mount open book preparations. The final step is to inject a fluorescent dye to visualize the axonal projections. Ultimately, fluorescent microscopy is used to show both the cells experiencing gene silencing and to assess the subsequent effects on commiss axon guidance.
The main advantage of this technique over existing methods is the cell type specific silencing of genes. To make the plasmids check the written protocol for instructions Incubate the eggs at 38.5 degrees Celsius and around 45%humidity until the desired stage of development is reached. To study Axon guidance of commissural neurons incubate the embryos for approximately three days until they reach hamburger and Hamilton Stage 17 to 18.
Once the desired stage of development is reached and before the protein of interest is accumulated within the embryo, remove the eggs from the incubator and place them in a stable horizontal position for 20 minutes to reposition the embryo on top of the yolk at the upper side of the egg. After the embryos have repositioned on top of the yolk, take the eggs out of the incubator and wipe them with 70%ethanol. Next place a strip of tape along the long axis of the egg and use a scalpel to make a small hole in the blunt end of the egg and another at the corner of the area to be windowed.
Then insert a syringe fitted with an 18 gauge needle into the hole at the blunt end of the egg at a 45 degree angle and aspirate approximately three milliliters of albumin while being careful to avoid damaging the yolk. Use small scissors held horizontally to cut the window into the egg without damaging the embryo. Then seal the window using the tape.
Finally, use a brush to apply melted paraffin wax to seal the hole at the blunt end of the egg and seal any cracks in the egg in the same manner. Then return the egg to the incubator. Repeat the process with all eggs to be electroporated.
Remove the tape from one of the windowed eggs and stage the embryo according to hamburger and Hamilton. Then use forceps to lift the extra embryonic membranes from the cordal half of the embryo in the region where major right and left viter line veins. Enter the trunk gently tear or cut the membranes with spring scissors and pour them towards the tail.
Now use gentle suction to load the DNA mixture to be electroporated into a glass micro capillary with attached tubing. Insert the micro capillary into the central canal of the neural tube just above the hind limbs and inject the DNA solution controlling the injection volume by mouth. The blue dye should spread from the tip of the tail up to the ventricle of the developing brain.
Add a few drops of sterile PBS on top of the embryo and place the electrodes parallel to the anterior posterior axis of the embryo without touching the embryo or any blood vessels. Then hold the electrodes steady and electro purate. Following the electroporation, remove the electrodes and quickly rinse with sterile water to remove denatured proteins from the egg white.
Finally, drop a little more sterile PBS onto the embryo and then reseal the egg with tape. As before, proper sealing is crucial to avoid dehydration of the embryo. Then return the embryo to the incubator until the desired stage of development is reached.
Once the embryo reaches Hamburger Hamilton, stage 25 to 26, use forceps to remove the embryo from the egg and place it in a Petri dish coated with sil guard elastomer. Use the forceps to remove the extra embryonic membranes and lay the embryo with the dorsal side down. Now use 0.2 millimeter insect pins to gently stretch out and pin the embryo through the neck and tail.
Then pin the limbs of the embryo, inserting the pins at an angle so that they do not interfere with the subsequent dissection. Illuminate the dish containing the embryo from below so that tissue density can be perceived. Use spring scissors to open the abdominal cavity and gentle scraping with forceps to remove the heart and internal organs.
When all organs have been removed completely, the segmented vertebrae and spinal cord will be visible. Next, rotate the embryo 180 degrees and use spring scissors to make a shallow cut through the vertebrae overlying the spinal cord at the neck. Then make two short longitudinal cuts through the vertebrae on either side of the spinal cord from the neck to the tail.
Use forceps to lift the flapper vertebrae away from the spinal cord and then peel off the vertebrae in a single strip towards the tail. Then gently stretch and re-pin the embryo through the tail and limbs. Now identify the spinal cord meninges, which should appear as a dark, dense line of tissue between the neural tube and the dorsal root ganglia, and then use a ster needle or fine microsurgical scalpel to cut the meninges longitudinally along each side of the cord from neck to tail.
The meninges should easily separate from the spinal cord due to the stretching of the pinned embryo. Finally, cut the spinal cord at the level of the wing bud and cordal to the limb bud. Then use forceps to lift out the spinal cord in a rostral cordal motion while ensuring that the cord remains immersed in the PBS First.
Spread the isolated spinal cord onto a spatula and transfer it to a fresh syl guard coated dish containing 4%para formaldehyde in PBS. Then carefully pin the spinal cord roly medially and quarterly with 0.1 millimeter insect pins to produce a flat mount preparation, incubating the paraldehyde for 30 minutes to one hour. Be careful not to over fix.
As this will increase background and reduce the efficiency of diod diffusion. Then carefully pour off the paraldehyde and replace with PBS. After preparing five milligrams per milliliter fast di in ethanol, break off the tip of a small diameter glass micro pipette attached to plastic tubing and draw the solution into the pipette iet.
Insert the filled needle into a dish of PBS and check that the dye doesn't leak from the needle. If it does, the diameter is too wide and a new needle should be prepared. Ensure that the open book preparation is illuminated from below and located denser longitudinal stripe of tissue located approximately one fifth of the width of the Hemi book From the lateral edge, the cell bodies of the commissural neurons are just dorsal to this dark line.
Insert the tip of the glass micro pipette into the tissue, and as the needle is withdrawn, puff a small amount of D eye using a mouth pipette working quickly repeat the procedure every 0.5 millimeters along the length of the open book. Once the injections are complete, use a transfer pipette to aspirate and discard. Any excess di failure to do so will result in high background.
Repeat the procedure for all of the open book preparations. Then place the dish containing the injected preparations of four degrees Celsius for approximately three days to allow the dye to spread along the axons. After this time mount the preparations according to the instructions in the written protocol.
Cross sections and open books were derived from hamburger Hamilton Stage 25 to 26 chicken embryos that were electro rated at Hamburger Hamilton Stage 18. As seen here in cross section, the beta acting promoter drives ubiquitous expression. The arrow indicates the dorsal ventral direction.
This image shows an example of fluorescent protein expression patterns obtained following electroporation of the plasmid vectors. Here ubiquitous expression from the beta actin promoter is seen in an open book preparation. The arrow indicates the rostral cordal direction.
This image shows a cross section after electroporation with a construct containing a math one enhancer. This drives expression in di one neurons. Here expression from the math one enhancer construct is seen in an open book preparation.
This cross section shows expression from the hawk, A one enhancer after bilateral electroporation of the construct. The hawk A one enhancer drives expression specifically in the floor plate. This open book preparation shows the expression pattern of the hawks a one construct in the floor plate.
This image shows a bright field image of the application of Dai injection sites. In an open book preparation, Dai was injected in a punctate pattern close to the lateral margin of the open book. On the electro rated side, this image shows visualization of GFP expression in the same preparation, and here Dai is visualized.
After three days of diffusion, commissure relax on trajectories should be able to be visualized under fluorescent microscopy. Normal axon trajectories will grow towards the floor plate, cross the floor plate, and then turn and grow. Ally as seen here.
Abnormal phenotypes arising from gene knockdown such as the one shown here can be compared to the archetypal trajectory. In the example, some axon stalling the floor plate or make erroneous turning decisions on the contralateral side. While attempting this procedure, it's important to be careful and patient throughout the fine injection and dissection steps.
These take some practice. After watching this video, you should have a good understanding of how to inject and ate RNAI plasmids to investigate gene function during neural tube development in a cell type specific manner.
