The overall goal of this procedure is to image cell behavior within the embryonic neural tube at high resolution for long periods of time. This is accomplished by first transecting the early neural tube with a construct driving expression of a fluorescent protein of choice to mark single cells. The next step is to make slices of the early embryo and mount them onto glass bottom dishes.
After recovery in an incubator, the embryo slices are imaged on a wide field microscope at regular intervals. Ultimately, this technique can be used to study cell behavior within the developing neuro epithelium over long periods of time with high spatial and temporal resolution. The main advantage of this technique over existing methods for imaging living tissue is that it allows us to observe individual cell behavior over long periods of time at high spatial and temporal resolution.
This method allows you to address key questions in the field of, of developmental neurobiology, such as the role of mitotic spindle orientation in sulfate choice, or how signaling dynamics might be regulating cell behavior Prior to electroporation of plasmids into the neural tube. Glass needles are prepared using a micro capillary puller under a dissecting microscope. Use fine forceps to break off the tip of the needle.
The end of the needle should be sharp enough to pierce the neural tube while not being so narrow that it impedes injection of the DNA solution. The X for this experiment have been incubated at 37 degrees Celsius for about 36 hours to hamburger Hamilton. Stage 10.
To begin this procedure, window and egg and place electrodes five millimeters apart on either side of the embryo, inject DNA into the neural tube. The DNA is colored with a small amount of fast green for visualization. Apply a current of 12 to 17 volts and 50 millisecond pulse length three times with 950 milliseconds between pulses.
Low concentrations of DNA and low electroporation voltages are used to achieve mosaic expression so that individual cells can be followed. Finally, cover the window in the eggshell with cellar tape, making sure it is sealed. Allow the embryos to recover for three to four hours or overnight at 37 degrees Celsius.
The collagen mixture and slice culture medium should be prepared about an hour before slicing the embryos. To prepare the collagen mixture at 100 microliters of 0.1%acetic acid solution to 300 microliters of type one collagen and vortex thoroughly, add 100 microliters of five times L 15, medium and vortex thoroughly. Again, the solution should turn yellow.
Next, add 15 to 20 microliters of 7.5%sodium bicarbonate, and vortex thoroughly. The solution should become slightly pink. Keep on ice.
This collagen mix should be prepared fresh each time. To prepare the slice culture medium at the following to 10 milliliters of neuro basal medium, B 27 supplement glut max and Gentamycin solution, place the medium in a 37 degree Celsius incubator, buffered with 5%carbon dioxide. Leave the top of the container loose to allow it to equilibrate with the carbon dioxide for at least an hour.
To begin this procedure, use a pair of small scissors to cut out the embryo. Remove the embryo from the egg with tweezers washing L 15 medium. Place the embryo in a tissue culture dish with a layer of syl guard at the bottom.
Pin the embryo out through the surrounding extra embryonic membranes so that the membranes are stretched taut. Using a micro knife, slice the embryo as straight as possible through the region of interest. For spinal cord slices should be between one to two somites thick leaf slices attached to the lateral tissue of the embryo so they are not lost while other embryos are being sliced.
A customized micro perpet tip is needed for transferring the spinal cord slices to the glass bottom dish. To prepare this, attach a 200 microliter tip to a P two or P 10 perman and cut off approximately one millimeter from the end of the tip. Code the inside of the tip with collagen by preparing one microliter of the collagen mix that was prepared earlier.
Leave for one to two minutes and then rinse with L 15 medium. This will prevent the tissue from sticking to the inside of the tip. Now detach a slice from the embryo using a micro knife and remove the slice from the dish using the P two or P 10 fitted With the 200 microliter tip.
Try to take up as little medium as possible. Vortex the collagen mix and drop five to eight microliters of it onto a polyol lysine coated glass bottom dish with a cover slip as the base, immediately put the embryo slice into the collagen and position it with a pair of fine forceps. LICs should be positioned so that the side to be imaged is flush with the cover slip.
The tissue should adhere to the polyol lysine coating on the cover slip. Repeat this until you have several slices on the cover slip. Usually six to nine slices are placed on one dish.
When all the slices are in place, add two to three microliters of L 15, medium to the earliest place collagen and cover the dish. Allow the collagen to set for 20 minutes. Once the collagen is set carefully add two milliliters of slice culture medium that has been equilibrated for at least an hour in 5%carbon dioxide and 37 degrees Celsius.
Care must be taken not to dislodge the collagen from the cover slip place in a 37 degree Celsius 5%carbon dioxide incubator and allow slices to recover for at least three hours before imaging. Imaging cell behavior within tissue for long periods of time at high temporal and spatial resolution is challenging. A combination of optimal culture conditions and the use of white field microscopy is essential for obtaining good results.
A delta vision core wide field microscope fitted with a weather station environmental chamber is used to image the slices. The chamber is constantly maintained at 37 degrees Celsius with a carbon dioxide perfusion apparatus to maintain the microscope stage at 5%carbon dioxide and 95%Air imaging is normally carried out using a 40 times 1.30 NA oil immersion lens and images are captured with a core snap HQ two called CCD camera Z Sections are captured every 1.5 microns through 45 microns of tissue exposure. Time should be kept as low as possible, for example, five to 50 milliseconds.
For each said section images are captured every seven minutes. Up to nine slices can be visited using the Delta vision systems. Precise point visiting function shown here is an example time-lapse sequence of a spinal cord progenitor cell transfected with a construct expressing GFP alpha Tubulin imaging was started on a spinal cord slice from a two day old HH stage 12 embryo.
During this early stage, neural progenitor cells undergo predominantly progenitor progenitor mode divisions during which the cells divide to generate two further cycling progenitor cells. This figure shows selected frames from the time-lapse sequence just shown. At two hours and 20 minutes, the cell divides with a cleavage plane that is perpendicular to the apical surface generating two daughter cells.
These two cells divide once again at 24 hours and 23 minutes and 25 hours and 54 minutes. This next time-lapse sequence is of a cell transfected with G-F-P-G-P-I marking the cell membrane. This cell undergoes a division during which the basal process splits in two indicated by the white arrows and is equally inherited by the daughter cells.
Selected frames from this time-lapse sequence show that the cell division occurs at zero hours and 49 minutes. After watching this video, you should have a good idea of how to prepare and image spinal cord slices. Remember, if you are new to this technique, you may struggle initially as there are a number of technically challenging steps that all need to come together for this to work.
