L'imagerie confocale temporelle des neurones migrateurs dans la culture de tranches organotypiques du cerveau embryonnaire de souris en utilisant

The overall goal of this procedure is to directly observe radially migrating neurons in organotypic slice culture prepared from electroporated embryonic brain by time-lapse confocal microscopy. This method can help study key aspects of neocortex development. It allows investigation of the molecular mechanisms involved in neuron polarization and the radial migration of neurons to their final position within the brain.

The main advantage of this technique is the dynamic properties of migrating neurons can be analyzed including speed profiles, average migration speed as well changes of migratory direction. Begin by placing a properly anesthetized pregnant mouse in the supine position on a warming plate. Check for the absence of the pedal reflex.

And then, carefully cover the eyes with petroleum jelly to prevent them from drying. Next, gently spread out the limbs and fix them to the warming plate with surgical tape. Sterilize the abdomen by swabbing with 70%ethanol followed by iodine solution, then place sterile gauze, in which a cut has been made for the abdominal incision, over the abdomen.

Moisten the gauze with bacteriostatic sodium chloride solution. After reassessing the development of surgical anesthesia, by loss of the pedal reflex, use serrated micro Adson forceps and angled fine scissors to cut the skin approximately 1.5 centimeters along the midline of the abdomen. Then, cut the underlying muscle along the linea alba.

Grasp the uterine horn between embryos with ring forceps and gently place it onto the moistened gauze without perturbing the placentas or supply vessels. Then, carefully position an embryo to give a clear view of the lateral ventricle which presents as a crescent-shaped shadow parallel to the sagittal sinus. The injection site lies in the middle of a line between the pigmented eye and the confluence of sinuses where the sagittal sinus branches to two transfer sinuses.

Once identified, push the microinjection needle through the uterine wall and into the lateral ventricle. Next, use a microinjector operated with a foot pedal to inject one to two microliters of DNA solution with five to 10 pulses. Successful injection can be monitored by the colored DNA solution which should fill the ventricular system.

Then, place tweezers-type electrodes so that the positive terminal is on the same side as the injected ventricle and the negative terminal is on the opposite side of the injected ventricle below the ear of the embryo's head. Moisten the electroporation site with a few drops of bacteriostatic sodium chloride solution and apply five electrical current pulses of 50-millisecond duration with 950-millisecond intervals. Bubbles at the negative electrode indicate electrical current.

60 to 90 milliamperes are usually sufficient for successful electroporation. Lower currents will not efficiently transfect neurons whereas higher currents may lead to embryonic death. After repeating the procedure for every embryo of the first uterine horn, gently place it back into the abdominal cavity.

After suturing the muscle layer and skin, carefully disinfect with iodine solution and gently remove the petroleum jelly from the eyes using cellulose swabs. Place the mouse under an infrared lamp until it wakes up. After euthanizing the pregnant female using an approved method, remove the uterus containing the embryos and place it into a 10-centimeter Petri dish with ice-cold complete HBSS.

From this point onward, keep all solutions, embryos and brains on ice. While working under a stereomicroscope, use fine-tipped forceps and a pair of Vannas Tubingen spring scissors to separate each embryo from the uterus. Transfer the embryos to another Petri dish containing ice-cold complete HBSS.

Make an incision at the level of the brainstem and cut along the sagittal midline. Peel away the skin and cartilage covering the brain. Then, cut off the brainstem right behind the hemispheres and remove the brain from the skull.

Transfer the brain with a micro spoon spatula to a 12-well plate containing ice-cold complete HBSS. Collect all brains from the litter in the 12-well plate. Then, use a fluorescent stereomicroscope to screen the brains in the 12-well plate for brightness of florescence as well as the size of the electroporated region.

Select two to four brains with bright fluorescent signals and the presumptive somatosensory cortex for further processing. Then, pour molten 3%low-melting-point agarose solution into a peel-away disposable embedding mold. Use a micro spoon spatula to remove the brain from the 12-well plate and carefully drain excess complete HBSS from around the brain using fine tissue paper.

Place the brain gently into the agarose solution. Then, use a 20-gauge needle to push it to the bottom of the mold and carefully adjust its position. For coronal sections, orient the brain with the olfactory bulbs pointing upwards.

Keep the mold on ice until the agarose solution is solidified then proceed to sectioning the brain. Use a clean razor blade to trim away excess agarose around the brain leaving approximately one millimeter of agarose on all sides except for the olfactory bulbs where two to three millimeters of agarose should be left. After applying a small amount of cyanoacrylate glue to a specimen, fix the trimmed agarose block with the olfactory bulbs pointing downwards.

Transfer the specimen stage to the slicing chamber of a vibrating blade microtome containing ice-cold complete HBSS and position the brain with its dorsal side towards the blade. Cut 250 micron-thick brain slices containing the electroporated region with a low to moderate amplitude and a very slow sectioning speed. Usually four to six slices with bright fluorescent signal are obtained from each successfully electroporated brain.

Grasp the agarose rim of one section with fine-tipped forceps and pull the slice onto a bent micro spoon spatula. In this manner, carefully transfer the brain slices to a six-well plate containing ice-cold complete HBSS. Moisten laminin-coated membrane inserts with 100 microliters of complete HBSS.

Then, carefully transfer the slices onto the membranes using the bent micro spoon spatula and forceps. Gently push the slice from the spatula onto the membrane, then position using forceps. Continue to transfer the remaining slices.

Up to five slices can be placed on one membrane insert. Remove excess complete HBSS with a pipette. Then, with the help of forceps, carefully place the membrane inserts with the slices into a six-well plate containing 1.8 milliliters of sliced culture medium.

Select a brain slice for imaging by viewing the slices through an inverted microscope. Choose a slice with bright single neurons in the upper SVZ migrating radially towards the pile surface of the slice. The presence of fine fluorescent processes of radial glial cells that span the entire conjugate plate indicates an intact radial glial scaffold which is used by migrating conjugate neurons.

Transfer the membrane insert with a selected slice into a 50-millimeter-diameter glass-bottomed dish containing two milliliters of slice culture medium. Place the dish into the climate chamber of the confocal microscope. Set the resolution to 512 by 512 pixels.

Increase the scan speed from 400 hertz to 700 hertz to increase the frame rate from about 1.4 to about 2.5 frames per second. Use no more than two times averaging. Define a Z-stack through the electroporated region with the step size of 1.5 microns.

Start the time-lapse series by taking a Z-stack every 30 minutes. The settings described allow for sufficient resolution and brightness of the image while keeping photo damage low during acquisition. This movie shows E16.5 cortical neurons migrating from the ventricular subventrical zones to the cortical plate.

In the Bcl11a floxed conditional transgenic after electroporation of the IRES-GFP control plasmid on embryonic day 14.5. The movie is comprised of 108 frames at the rate of five frames per second. The scale bar represents 100 microns.

Electroporation of a DNA plasmid vector containing Cre-IRES-GFP affected cortical neuron migration in conditional Bcl11a floxed brains. As seen here, very few neurons move past the intermediate zone to reach the cortical plate. This movie shows the polarization of cortical neurons from a GFP-labeled control on the left compared to neurons from a Bcl11a mutant on the right.

These animated traces of representative control in Bcl11a mutant neurons were obtained from time-lapse series of E16.5 slice cultures with one-hour-interval resolution. Again, data from the control brain is shown on the left and data from the Cre-IRES-GFP electroporated brain is shown on the right. As seen from this collection of representative traces from control and Bcl11a mutant slice cultures, the Bcl11a mutant neurons frequently undergo repetitive phases of reduced migration speed and randomly changed orientation as marked by the red arrowheads.

Speed profiles can be calculated from traces of migrating neurons. As seen here, a greater proportion of Bcl11a mutant neurons have slower migration speeds compared to controls. The deviation angle may be calculated from the trace data.

As seen from this histogram, the Bcl11a mutant neurons represented by the black bars exhibit greater angles of deviation compared to controls. Once mastered, this technique can be done in less than two hours each for the in utero electroporation and the preparation of organotypic brain slice culture. This procedure can easily be adapted to perform fractional analysis of genes of interest in radially migrating cortical neurons by gain and loss of function as well as vascular experiments.

After watching this video, you should have a good understanding of how to directly observe radially migrating neurons in organotypic slice culture prepared from electroporated embryonic brains by time-lapse confocal microscopy.