Culturing Rat Sympathetic Neurons from Embryonic Superior Cervical Ganglia for Morphological and Proteomic Analysis

All procedures performed in studies involving animals were approved by the Institutional Animal Care and Use Committee (IACUC) at Saint Mary’s College of California. The animal care and use guidelines at Saint Mary’s College were developed based on the guidelines provided by Office of Laboratory Animal Welfare at the National Institute for Health (https://olaw.nih.gov/sites/default/files/PHSPolicyLabAnimals.pdf and https://olaw.nih.gov/sites/default/files/Guide-for-the-Care-and-Use-of-Laboratory-Animals.pdf).

1. Preparation of culture media (also referred to as control medium)

1. Add the following ingredients, in the order listed below, to a sterile 500 mL flask: 190 mL of 1x low-glucose DMEM, 10 mL of 20 mg/mL fatty acid free bovine serum albumin(FAF-BSA) in DMEM, 2.8 mL of 200 mM L-glutamine, 4 mL of 100x insulin-selenium-transferrin mixture, 0.4 mL of 125 µg/mL nerve growth factor (in 0.2% inert protein stabilizer in DMEM), and 200 mL of Ham’s F-12 medium.
2. Make sure to coat the pipette with the medium containing FAF-BSA prior to the addition of protein-containing solutions such as insulin-selenium-transferrin and NGF to prevent sticking of proteins to the pipette.
3. Swirl the flask to mix the ingredients after each addition. Mix thoroughly with a 25 mL pipette after addition of F-12.
4. Aliquot the culture media in 10 mL aliquots and store at -80 °C. The aliquots can be stored for six months without loss of activity.

2. Preparation of plates for culturing neurons

1. Dilute a 1 mg/mL stock of poly-D-lysine (made in 0.5 M Tris buffer, pH 8) to 100 µg/mL with sterile distilled water.
2. For proteomic or genomic analysis, one to two days before the dissection, coat 6-well plates with approximately 2 mL of sterile 100 g/mL of poly-D-lysine (PDL). This is necessary to ensure cell adhesion to the well. Wrap the plates with cling film and store the plates overnight at 4 °C.
3. For morphological analysis and microscopy, place 12 mm² pre-treated German glass coverslips (which can be cleaned using nitric acid treatment as described previously¹⁸ or purchased) into each of the wells of a 24-well plate.
   1. Coat the coverslips with 0.3 mL of sterile 100 g/mL of poly-D-lysine (PDL). Store the plates overnight at 4 °C.
4. 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.
5. 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 mL of control media. Store the plates at 35.5 °C under 5% CO₂ in a humidified chamber.

3. Dissection setup

1. Preparation of 200 mL of media for dissection (referred to as Dissection media)
   1. Add 8 mL of 20 mg/mL fatty acid-free bovine serum albumin (FAF-BSA) in low glucose DMEM (with 1 mg/mL glucose) and 2 mL of 100x penicillin-streptomycin to 193 mL of sterile Leibovitz's L-15 medium (or any air-buffered medium)
2. In the hood, set up the following items for dissection: four 50 mL sterile conical tubes with about 20 mL of dissection media in each tube, four sterile 35 mm dishes with 1.5 mL of dissection media, one sterile 50 mL conical tube with 20 mL of dissection media for centrifugation, one sterile 15 mL conical tube for centrifuging the ganglia, and one sterile 10 mL tube for collecting the dissociated cells.
3. Use a dry bead sterilizer to sterilize a pair of fine forceps (no.4 or no. 5 forceps) for at least one minute.

4. Isolation of the superior cervical ganglia from embryonic rat pups

1. Removal of E21 embryos from the pregnant rat
   NOTE: Removal of the uterine horn can be performed outside of the hood if the surrounding area is thoroughly sterilized.
   1. Euthanize the pregnant rat using CO₂ inhalation. Shear the fur from the abdominal region and wipe the skin in the area with 70% alcohol to sterilize it.
   2. Using a fresh set of sterile scissors and forceps, cut through the skin and then the muscle layer to expose the uterine horns containing the embryos. Remove the uterine horns with the embryos using a new set of scissors and forceps, taking care not to damage the intestines in the process.
   3. Transfer the uterine horns with the embryos into a 150 mm² sterile Petri dish and transfer them into the hood.
   4. 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.
   5. To euthanize the embryos, cut the spinal cord of the embryos along the midline under the right arm. This will also reduce the bleeding from the carotid artery during the removal of the SCG.
   6. Transfer these embryos into the prepared 50 mL conical tubes containing dissection media. Make sure the embryos are submerged in the media. Each tube can hold up to 3 embryos.
2. Isolation of the superior cervical ganglion from the embryo
   1. Transfer one pup from the dissection media onto a sterile 150 mm² Petri dish half-filled with solid substrate (either paraffin wax or silicone polymer), with its dorsal surface on the substrate. Using three sterile 23 G needles, pin the pup to the dish with one needle under each arm and a third needle through the mouth to carefully hyperextend the neck.
   2. Cut through the skin in the neck region using sterile fine forceps (no. 4 or no. 5 forceps) to expose the salivary glands underneath. Remove these glands using fine forceps.
   3. 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.
   4. 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 mm² 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.
   5. Repeat the dissection process on the other side. Remove SCGs from all embryos before continuing with the remaining dissection steps outlined below. Distribute the isolated tissues between two of the 35 mm² dishes to facilitate processing of the tissue samples.
3. Post-processing of SCG
   1. To separate the SCG from the dissected tissue, first use fine forceps to remove any extraneous tissue, such as fat or muscle, taking care to avoid the area near the carotid bifurcation.
   2. 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 nodose ganglia from the carotid and then separate the SCG from the carotid artery.
   3. Use fine forceps to remove the capsule that surrounds the SCG. Transfer the SCG to a new 35 mm culture dish. Repeat the process with all the dissected tissue samples.
   4. 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 2 mL of Collagenase type II (1 mg/mL)/dispase type II (5 mg/mL) in calcium- and magnesium-free HBSS, and incubate for 50 min at 37 °C to help break down the tissues.
      NOTE: The incubation time may need to be optimized with different batches of Collagenase/Dispase and usually ranges from 40 min to an hour.
   5. During the incubation, aspirate the DMEM from the plates and replace it with 0.3 mL of control media. Store the plates at 35.5 °C under 5% CO₂ in a humidified chamber.
   6. Following the incubation, transfer the SCGs in collagenase/dispase to a sterile 15 mL conical tube. Use the dissection media to rinse the plates and transfer the solution to the tube. Add enough dissection media to bring up the volume to approximately 10 mL.
   7. Centrifuge at 200 x g (1000 – 1200 rpm) for 5 minutes at room temperature to pellet the sample. Aspirate the supernatant, taking care to not dislodge the pellet. Resuspend the pellet with 10 mL of dissection media. Repeat the centrifugation and discard the supernatant.
   8. Replace with 1 – 2 mL of culture medium. Using a narrow-bore, bent-tip sterile pasteur pipette (pre-coated with culture medium), mechanically dissociate the clumps by gently triturating 5-6 times. Let the larger clumps settle for about one minute. Transfer the supernatant cell suspension to a new 10 mL tube.
   9. Repeat this process 3 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 a 10 mL tube with the supernatant from the first trituration.
   10. Add enough culture media to bring up the volume to 8 – 10 mL. Gently mix the cell suspension and quantify the cell density with a hemocytometer.
   11. 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.
       NOTE: For morphological analysis, plate the cells around 8,000 cells/well in a 24 well plates and for genomic and proteomic protocols, plate the cells as high as 30,000 cells/well.
   12. Transfer the plates to a glass desiccator with sterile water at the bottom to create a humidified chamber. Inject enough CO₂ (around 120 mL) to obtain a 5% CO₂ environment in the desiccator, prior to sealing. Maintain the plates at 35.5 °C. This is referred to as Day 0 in the protocol.
       NOTE: 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.

5. Maintenance of the cultured SCG neurons and treatments

1. On Day 1 (24 hours after plating), carefully remove half of the culture media, and replace with 2 µM Ara-C (cytosine β-D-arabinofuranoside, an anti-mitotic agent). Leave the treatment on the cells for 48 h. Usually, this length of treatment is sufficient to eliminate non-neuronal cells in the culture.
2. On Day 3, gently aspirate half the medium and replace with control medium.
3. On Day 4, 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.
4. Cultured SCG neurons in serum-free medium extend only axons. If experiments require the presence of dendrites, treat cells with either 10% fetal calf serum, 75-100 µg/mL of basement membrane matrix extract or 50 ng/mL of bone morphogenetic protein-7. Dendritic growth is observed within 48 hours of treatment with elaborate dendritic arbor observed within a week of treatment.

6. Immunostaining cultured SCG neurons

1. Gradually replace the cell culture medium in the well with 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.2 in a fume hood.
   1. Remove half the culture media in the well and replace it gently with 4% paraformaldehyde. Then, repeat the process at least two more times, removing two-thirds of the media in the well each time and replacing with 4% paraformaldehyde. At the end of this process, the color of media in the well should have changed from pink to colorless.
   2. Once all the medium has been replaced by 4% paraformaldehyde, incubate the wells for 15 – 20 minutes at room temperature.
2. With a similar process as described above, gradually replace the 4% paraformaldehyde solution with phosphate-buffered saline (PBS), pH 7.2. Leave for 5 minutes. Rinse the wells two more times for 5 minutes each with PBS.
   1. Once the 4% paraformaldehyde is removed, move the plates to the benchtop for further processing. This step is a good stopping point where plates can be stored at 4 °C overnight.
3. Remove all the PBS. Add enough volume of 0.1% Triton X-100 in PBS to cover the cells. Leave it on the cultures for 5 minutes to permeabilize the neurons. The timing of this step is critical to ensure good staining while maintaining the integrity of the cells.
4. Carefully remove the Triton X-100 solution and rinse the wells three times for 5 minutes each time with PBS. Remove the PBS solution.
5. Add enough 5% BSA in PBS to cover the cells and incubate at room temperature for 20 minutes to reduce non-specific antibody binding.
6. Remove the blocking solution and replace it with a mouse monoclonal antibody against MAP-2 protein diluted in 5% BSA in PBS. Leave the primary antibody on the cells overnight at 4 °C.
7. Remove and save the primary antibody for reuse. The primary antibody can be reused at least twice without loss of detection intensity.
8. Rinse three times with enough PBS to cover the cells. Leave PBS on the cells for 10 minutes per rinse.
9. Remove the PBS solution and replace it with fluorescent tag-conjugated Goat anti-Mouse IgG secondary antibody at 1:1000 dilution in 5% BSA in PBS. Incubate for 2 hours in the dark at room temperature.
10. Remove the secondary antibody and replace it with PBS. Rinse the wells three times with PBS for 10 minutes per rinse. Rinse the wells one time with water to remove any salts.
11. Mount the coverslips onto glass slides containing a drop of aqueous mountant that is appropriate for fluorescently labeled samples. Store the slides at 4 °C, in a slide folder until ready for imaging.

7. Sample preparation for analysis of the proteome using liquid chromatography coupled with mass spectrometry

1. Lysis of cultured neurons
   1. Gently remove all the culture media in the wells. Replace it with cold, sterile calcium and magnesium-free phosphate-buffered saline (PBS), pH 7.4. Remove quickly and replace with PBS and let it sit for 5 min. This step is done to remove any proteins present in the culture media.
   2. Carefully remove all the PBS from all the wells for a particular treatment. Repeat the process one more time. Maintain plates over ice when lysing the cells to minimize neuronal damage and proteolysis.
   3. Add 100 – 150 µL of 50 mM ammonium bicarbonate, pH 7.5 (NH₄HCO₃) to one of the wells and scrape cells using a sterile cell scraper. Using a micropipette, transfer the liquid and repeat the scraping process with all the wells for a particular treatment. Examine the wells under the microscope after scraping to make sure that most of the cells have been removed.
      1. With the low number of neurons, use a limited volume to lyse the cells to ensure high enough concentration for proteomic analysis.
   4. Freeze the solution at -80 °C overnight to help with cell lysis. The lysates can be stored at this stage until ready for further processing.
   5. Once thawed, squirt the samples through a syringe with a 26 G or 28 G needle to mechanically lyse the cells.
   6. Sonicate the samples in a sonicating water bath two times for 10 minutes. Add ice to the water bath to prevent overheating and denaturation of proteins. Centrifuge for 5 minutes at 4 °C at 12,000 x g.
   7. Measure the protein concentration. Typical protein concentrations range from 0.4 – 1 µg/µL
2. Sample preparation and trypsin digestion for proteome analysis
   1. Transfer a maximum of 60 µL of the lysate or 50 µg of protein into a DNase-, RNase- and protease-free 1.5 mL tube. If the volume of the lysate is less than 60 µL, add enough 50 mM ammonium bicarbonate to make up the volume.
   2. Add 25 µL of 0.2% acid labile surfactant and vortex. Incubate the tube in a block heater at 80 °C for 15 minutes. Centrifuge the tube at 12,000 x g for 30 s.
   3. Add 2.5 µL of 100 mM dithiothreitol and vortex. This makes the protein more accessible for alkylation and digestion. Incubate the tube at 60 °C in a block heater for 30 minutes. Cool to room temperature and centrifuge.
   4. Add 2.5 µL of 300 mM iodoacetamide to the sample and vortex. This step helps to alkylate the cysteines and prevents them from reforming the disulfide bonds. Incubate the tubes at room temperature in the dark for 30 minutes.
   5. Add mass spectrometry grade trypsin (0.5 µg/µL) to the tube at a trypsin: protein ratio of 1:10. Digest the samples at 37 °C overnight.
   6. Add 10 µL of 5% trifluoroacetic acid (TFA) and vortex. Incubate the samples at 37 °C for 90 minutes. This step is necessary to hydrolyze the acid labile surfactant to prevent interference during mass spectrometry.
   7. Centrifuge the samples at 12,000 x g at 4 °C for 30 minutes and transfer supernatant to a chromatography certified clear glass vial with pre slit Teflon/Silicone septum caps. Samples can be stored at -20 °C prior to mass spectrometry analysis.
   8. Subject the sample vials to liquid chromatography coupled with high definition mass spectrometry.