Double in utero electroporation is a powerful method for studying the important aspects of mammalian corticogenesis such as the interaction between neuron cells, generated at different locations and developmental ages. The main advantage of this technique is the ability to examine unperturbed interactions among these different cell populations in a quick, detailed, and inexpensive manner. One striking implication of our method is that it facilitates a better understanding of how similar our wiring is altered in neuronal disorders like autism and schizophrenia.
This study is not restricted to the neocortex but it can also be employed to analyze cell to cell interactions, in exocortico areas such as the sepalium or cerebellum. One of the most challenging steps is injection of DNA into the lateral ventricles of early embryos. The use of a bright light source is essential for successful injections.
For each surgery first prepare a 10 microliter solution containing one microliter of fast green dye, a one microgram per microliter plasmid DNA solution of interest for each plasmid, and the appropriate volume on Endotoxin free TE buffer. And preload a glass microcapillary pipette with the appropriate prepared DNA solution. For the first in utero electroporation surgery, apply ointment to the eyes of an appropriately timed anesthetized pregnant mouse, and use a razor to shave the abdomen.
Disinfect the exposed skin two times with sequential 70%ethanol and iodine wipes before proceeding with the incision on the right side of the abdomen. Once the uterus is accessible use ring forceps to carefully extract the organ. Using a mouth controlled aspirator tube, inject approximately 0.5 microliters of solution into the lateral ventricle of the left hemisphere of E11.5 embryos for strategy A.Or into the lateral ventricle of the right hemisphere of E13.5 embryos for strategy B, until the fast green dye is visible inside the ventricle.
When all of the solution has been injected, place forceps type platinum electrodes laterally around of the head of the injected embryos, with the positive pole oriented to the medial wall to target the cortical hem for strategy A, or oriented to the lateral cortex for strategy B, taking care to avoid the heart and placenta. Use a square-wave electroporater to apply the appropriate sequence of electric pulses for each embryonic age, as indicated in the table. When all of the embryos have been injected and electroporated, use forceps to carefully return the uterus to the abdominal cavity, and fill the abdomen with warm saline.
Using a needle 6-0 suture, close the abdominal wall, and carefully close the skin incision. Then return the animal to its home cage with monitoring until full recovery. Two days after the initial surgery, confirm that the mouse is exhibiting normal behavior, and presents no signs of pain or distress.
Before preparing the animal for the second in utero electroporation as just demonstrated. Inject 0.5 microliters of the desired DNA solution into the left lateral brain ventricle of E13.5 embryos for strategy A, and into the left lateral brain ventricle of E15.5 embryos for strategy B.After each injection, place the electrodes with the positive pole oriented to the lateral cortex of the injected hemisphere for strategy A and strategy B.And electroporate embryos as indicated in the table. When the electroporation is finished, return the uterus to the abdominal cavity, and complete the surgery and post-surgical care as demonstrated.
When implementing strategy A, four days after the second electroporation, harvest the uterus of the embryonic day 17.5 pregnant female mouse into a petri dish of PBS on ice. And use tweezers to transfer the embryos into a new dish of PBS under a dissecting microscope. Using forceps, carefully grasp the head of each embryo to facilitate removal of the skin over the head and skull.
Use forceps or a spatula to lift out the exposed brains. Then use spoon to transfer each brain into individual wells of a 48 well plate containing around 600 microliters of fixative solution per well. When all of the brains have been collected, fix the brains at four degrees Celsius overnight on an orbital shaker, followed by three ten minute washes in PBS.
After the last wash, embed the brains in 4%low melting point agarose in PBS for about ten minutes. After the agarose has solidified, use cyanoacrylate glue to secure each brain to a vibratome tissue holder olfactory bulb side up. Place the holder inside the vibratome container, and fill the container with PBS.
Then use the vibratome and a brush to obtain 100 micrometer thick coronal serial sections. Transferring up to seven sections per well and six wells per embryo, in a new 48 well plate containing 600 microliters per well of fresh PBS, supplemented with anti-fungal preservatives as they are collected. When all of the sections have been acquired, use a fine brush to mount each section onto a glass microscope slide, and cover them with a glass coverslip for assessment of the electroporation efficacy on an upright epifluorescence microscope.
When implementing strategy B, on postnatal day 15, after confirming a lack of response to topinch, secure the mouse in the supine position, and use scissors to make a midline skin incision to expose the ribcage and diaphragm, and cut the diaphragm to gain access to the heart. Using fine scissors, make an incision in the right atrium, and use a 25 gage needle connected to a flexible tube to a peristaltic perfusion pump to penetrate the left ventricle. When the needle is in place, deliver at least 25 milliliters of 4%paraformaldehyde by transcardial perfusion, at a constant flux at 5.5 milliliters per minute.
After approximately five minutes, stop the perfusion, and use scissors to remove the skin over the head. Remove the skull, and use a spatula to remove the brain tissue. Place the brain into one well of a 24 well plate containing 1.5 milliliters of fresh 4%paraformaldehyde, for an overnight incubation at four degrees Celsius on an orbital shaker.
Then acquire 40 micrometer sections of the postnatal brains as just demonstrated. For embryonic and postnatal brain slice imaging, place the slides containing the mounted brain sections of interest onto a microscope slide holder. Introduce the slider holder in the confocal microscope, and select the appropriate channel for the fluorofores used in the experiment.
Perform sample scanning to obtain map images of each brain section at two different wavelengths for a general view of the double electroporation output. Once finished, select the 10x lens, in the multi-area Z-stack time lapse observation mode. In each of the select XY regions, set the appropriate imaging perimeters as well as the depth of the scanning along the Z axis according to the planes of the sample, in which the fluorescence is visible.
Obtain low magnification images of all of the chosen regions, and export the images from OIF to TIFF format in the microscope viewer software. Then select the 60x lens, and capture high magnification images as just demonstrated, to observe the cell-to-cell interactions at a more detailed level. The temporal gap between the double in utero electroporation surgeries allows CAG toresial cells targeted on embryonic day 11.5 in one of its places of origin to reach the marginal zone of the lateral neocortex including the somatosensory area, in time to establish contacts with cortical projection neurons, labeled on embryonic day 13.5.
The leading processes of projection neurons expressing enhanced GFP profusely uprise in the marginal zone of the cortex, and intermingle with the processes of M-cherry expressing cataretzial cells. In utero electroporation at embryonic day 13.5, using a BFP expressing plasmid facilitates the labeling of projection neurons across different cortical layers. A second electroporation in the contralateral side of embryonic day 15.5 targets the upper layer callosal projection neurons subpopulation, but not the lower layer projection neurons that develop at earlier stages.
Upper layer targeted neurons extend their axons to the contralateral hemisphere, with the characteristic arborization pattern. High magnification allows a more detailed visualization of callosal axon innervation of the targeted projection neurons within the contralateral hemisphere. When attempting this protocol, it is important to plan electroporation timing, the hemisphere to inject, and the position of the electrodes according to the cell populations to be targeted.
Following this procedure all the methods, such as electroporation is the interesting one. For both cell types can be performed to study possible circuit alterations in vivo. Using this technique, it is possible to study fine interactions between two different targeted cells.
Including new science information in vivo, by the electroporation of fluorescent synaptic proteins.
