In this video, we will be demonstrating the culturing of embryonic rats'superior cervical ganglia for morphological and proteomic analysis. Before beginning the dissection, prepare control and dissection media. Control medium contains F-12 and low glucose DMEM supplemented with an insulin-selenium-transferrin mixture, glutamine, nerve growth factor, and BSA.
Dissection medium contains L-15 supplemented with pen-strep and BSA. Refer to the manuscript for detailed protocols for media preparation. Preparation of plates for culturing neurons.
Dilute a one milligram per milliliter stock of poly-D-lysine to 100 micrograms per milliliter with sterile distilled water. For proteomic or genomic analysis, one to two days before the dissection, coat six-well plates with approximately two milliliters of sterile 100 grams per milliliter of poly-D-lysine. This is necessary to ensure cell adhesion to the well.
Wrap the plates with cling film and store the plates overnight at four degrees Celsius. On the day of the dissection, before the start of the dissection, remove the poly-D-lysine solution from the wells and rinse the wells five times with sterile distilled water, followed by once with low-glucose DMEM. During the enzymatic digestion of the ganglia, approximately an hour before plating the cells, aspirate the DMEM from the plates and replace it with 0.3 milliliters of control medium.
Store the plates at 35 degrees Celsius under 5%CO2 in a humidified chamber. In the hood, set up the following items for dissection. Four 50 milliliter sterile conical tubes with about 20 milliliters of dissection media in each tube.
Four sterile 35 millimeter dishes with 1.5 milliliters of dissection media. One sterile 50 milliliter conical tube with 20 milliliters of dissection media for centrifugation. One 15 milliliter sterile conical tube for centrifuging the ganglia, one sterile 10 milliliter tube for collecting the dissociated cells.
Use a dry beads sterilizer to sterilize a pair of fine forceps for at least one minute. All procedures performed in studies involving animals were approved by the Institutional Animal Care and Use Committee at Saint Mary's College of California. The animal care and use guidelines at Saint Mary's College of California were developed based on guidelines provided by the Office of Laboratory Animal Welfare at the National Institutes for Health.
Removal of E21 embryos from the pregnant rat. Note that the removal of the uterine horn can be performed outside of the hood if the surrounding area is thoroughly sterilized. Euthanize a pregnant rat with CO2 inhalation.
Shear the fur from the abdominal region and wipe the skin in the area with 70%alcohol to sterilize it. Using a fresh set of sterile scissors and forceps, cut through the skin and then the muscle to expose the uterine horns containing the embryos. Remove the uterine horn with the embryos, using a new set of scissors and forceps, taking care not to damage the intestines in the process.
Transfer the uterine horn with the embryos into 150 millimeter sterile Petri dish and transfer them into the hood. Using a new set of forceps and scissors, remove the embryos from the uterine horn and separate the embryos from the amniotic membranes and the placenta. To euthanize the embryos, cut the spinal cord of the embryos along the midline under the right arm.
This will reduce the bleeding from the carotid artery during removal of the SCG. Transfer these embryos into the prepared 50 milliliter conical tubes containing dissection media. Make sure the embryos are submerged in the media.
Each tube can hold up to three embryos. Isolation of the superior cervical ganglion from the embryo. Transfer one pup from the dissection media onto a sterile 150 millimeter Petri dish half-filled with solid substrate with its dorsal surface on the substrate.
Using three sterile 23 gauge needles, pin the pub to the dish with one needle under each arm and a third needle through the mouth to carefully hyperextend the neck. Cut through the skin in the neck region using sterile fine forceps to expose the salivary glands underneath. Remove these glands using fine forceps.
Locate the sternocleidomastoid and omohyoid muscles near the clavicle and trachea respectively. First, cut the transverse sternocleidomastoid muscles. And then carefully cut the thin omohyoid muscle using fine forceps.
Once these muscles are removed, the bifurcation in the carotid artery on the anterior end, will be visible on either side of the trachea with the SCG located under this fork in the carotid artery. Using closed forceps, gently lift the carotid artery to visualize the SCG. Using one forceps on either side of the SCG, pull out the carotid and transfer it to the prepared sterile 35 millimeter dishes.
This tissue will most likely contain the SCG with the carotid artery, the vagus nerve with the nodose ganglia, as well as other segments of muscle or fat in the area. Repeat the dissection process on the other side. Remove SCGs from all embryos before continuing with the remaining dissection steps outlined.
Distribute the isolated tissues between two of the 35 millimeter dishes to facilitate processing of the tissue samples. Post-processing of the SCG. To separate the SCG from the dissected tissue, first use fine forceps to remove any extraneous tissue, such as muscle or fat, taking care to avoid the area near the carotid bifurcation.
Once these tissues have been removed, two ganglia are visible. The nodose ganglion is smaller and circular while the SCG is almond shaped. Gently pull on the vagus nerve to separate the vagus nerve and the nodose ganglia from the carotid and then separate the SCG from the carotid artery.
Use fine forceps to remove the capsule that surrounds SCG. Transfer the SCG to a new 35 millimeter culture dish. Repeat this process with all of the dissected tissue samples.
Coat a sterilized, cotton-plugged glass pipette with dissection media to prevent the tissue from adhering to the pipette walls. Use the pipette to replace the dissection media with sterile two milliliters of collagenase type II, dispase type II in calcium and magnesium-free HBSS and incubate for 50 minutes at 37 degrees Celsius to help break down the tissues. Note that the incubation times may need to be optimized with different batches of collagenase/dispase and usually ranges from 40 minutes to an hour.
Following the incubation, transfer the SCGs and collagenase/dispase to a sterile 15 milliliter conical tube. Use the dissection media to rinse the plates and transfer the solution to the tube. Add enough dissection media to bring the volume to approximately 10 milliliters.
Centrifuge at 200 x g, 1, 000 to 1, 200 rpm for five minutes at room temperature to pellet the sample. Aspirate the supernatant, taking care to not dislodge the pellet. Resuspend the pellet with 10 milliliters of dissection media, repeat the centrifugation and discard the supernatant.
Replace with one to two milliliters of culture medium. Using a narrow-bore, bent-tip sterile Pasteur pipette, pre-coat it with culture medium, mechanically dissociate the clumps by gently triturating five to six times. Let the larger clumps settle for about one minute and transfer the supernatant cell suspension to a new 10 milliliter tube.
Repeat this process three more times with increasing force of trituration each time to ensure almost complete dissociation of the SCGs. Transfer the supernatant after each round of trituration to the 10 milliliter tube with the supernatant from the first trituration. Add enough culture media to bring the volume to eight to 10 milliliters.
Gently mix a cell suspension and quantify the cell density with a hemocytometer. Distribute the cell suspension into the wells at the appropriate cell density for the experiments. Mix the cell suspension continually during the plating process to ensure even distribution of cells into the wells.
For morphological analysis, plate the cells around 8, 000 cells per well in a 24-well plate. And for genomic and proteomic protocols, plate the cells as high as 30, 000 cells per well. Transfer the plates to a glass desiccator with sterile water at the bottom to create a humidified chamber.
Inject enough CO2, around 120 milliliters, to obtain a 5%CO2 environment in the desiccator, prior to sealing. Maintain the plates at 35.5 degrees Celsius. This is referred to as day zero in the protocol.
These plates can also be maintained in a regular 5%CO2 incubator. The method described above minimizes temperature and pH changes and also helps prevent cross-contamination. Maintenance of the cultured SCG neurons and treatments.
These pictures show the dissociated superior cervical ganglia cells on day zero, day one, and day five. The day zero image shows the cells upon plating. The cells plated on day zero contain a mixture of both neuronal and non-neuronal cells.
On day one, 24 hours after plating, carefully remove half of the culture media and replace with two micromolar Ara-C, cytosine D-arabinofuranoside, an anti-mitotic agent. Leave the treatment on the cells for 48 hours. Usually this length of treatment is sufficient to eliminate non-neuronal cells in the culture.
On day three, gently aspirate half the medium and replace with control medium. On day four, the cells are ready for experimental treatments. Feed cultures every other day with the appropriate medium.
Gently replacing half of the medium in the well with fresh medium. The picture on day five shows extensive axonal growth with no dendritic growth observed. To induce dendritic growth, neurons can be grown on Matrigel or treated with 10%serum or 50 nanograms per milliliter of bone morphogenetic protein-7.
Figure two shows changes in neuronal morphology of E21 rat SCG neurons following treatment with BMP-7. Representative phase contrast micrographs at 10 X magnification show the circular neuronal cell body with axons in control neurons in panel A, and neurons with multiple short dendrite-like processes when treated with BMP-7 shown by the arrows in panel B.Cultured SCG neurons following appropriate treatments can be used for morphological analysis using immunocytochemical staining for genomic analysis and for biochemical studies, such as for proteomic analyses. Refer to the manuscript for detailed protocols for immunostaining and sample preparation for proteomic analysis.
Figure three shows immunostaining from MAP-2 protein in cultured embryonic rats sympathetic neurons. Panels A through D show cultured SCG neurons treated with control media and panels E through H show neurons treated with BMP-7 at 50 nanograms per milliliter for five days. Representative micrographs showing immunocytochemical staining of the neurons with microtubule associated protein-2 in C and G, a nuclear stain in D and H with phase contrast micrographs in B and F, and emerge of the three channels in A and D.Immunocytochemical staining from microtubule associated protein-2 is present in the cell body and proximal axons of control neurons in panel C.And staining from MAP-2 protein in the cell body, and dendrites in BMP-7 treated neurons is in panel G.Here's a sample run of superior cervical ganglia treated with control media, which detected 1, 100 proteins.
Gene ontology using the Panther database found that most of the proteins were cytoplasmic. However, there were membrane-bound proteins and proteins at cellular junctions in the sample. This analysis also revealed that proteins that are involved in a wide array of neuronal signaling pathways were detected in the sample.
In conclusion, after watching this video, you should be able to isolate and culture rat embryonic superior cervical ganglion neurons for morphological and proteomic analysis. This work was funded by the faculty development fund and summer research program at Saint Mary's College of California.
