This work was supported by the Faculty Development Fund and Summer Research Program grant at Saint Mary&#8217;s College of California. The authors would also like to thank Dr. Pamela Lein at University of California at Davis and Dr. Anthony Iavarone at UC Berkeley Mass spectrometry facility for their advice during the development of these protocols. The authors would also like to thank Haley Nelson in the Office of College Communications at Saint Mary&#8217;s College of California for her help with video production and editing.

All procedures performed in studies involving animals were approved by the Institutional Animal Care and Use Committee (IACUC) at Saint Mary&#8217;s College of California. The animal care and use guidelines at Saint Mary&#8217;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)
<ol>
	<li>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 &#181;g/mL nerve growth factor (in 0.2% inert protein stabilizer in DMEM), and 200 mL of Ham&#8217;s F-12 medium.</li>
	<li>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.</li>
	<li>Swirl the flask to mix the ingredients after each addition. Mix thoroughly with a 25 mL pipette after addition of F-12.</li>
	<li>Aliquot the culture media in 10 mL aliquots and store at -80 &#176;C. The aliquots can be stored for six months without loss of activity.</li>
</ol>
2. Preparation of plates for culturing neurons
<ol>
	<li>Dilute a 1 mg/mL stock of poly-D-lysine (made in 0.5 M Tris buffer, pH 8) to 100 &#181;g/mL with sterile distilled water.</li>
	<li>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 &#176;C.</li>
	<li>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.
	<ol>
		<li>Coat the coverslips with 0.3 mL of sterile 100 g/mL of poly-D-lysine (PDL). Store the plates overnight at 4 &#176;C.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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 &#176;C under 5% CO2 in a humidified chamber.</li>
</ol>
3. Dissection setup
<ol>
	<li>Preparation of 200 mL of media for dissection (referred to as Dissection media)
	<ol>
		<li>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&#39;s L-15 medium (or any air-buffered medium)</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Use a dry bead sterilizer to sterilize a pair of fine forceps (no.4 or no. 5 forceps) for at least one minute.</li>
</ol>
4. Isolation of the superior cervical ganglia from embryonic rat pups
<ol>
	<li>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.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Transfer the uterine horns with the embryos into a 150 mm2 sterile Petri dish and transfer them into the hood.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Isolation of the superior cervical ganglion from the embryo
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Post-processing of SCG
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#176;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.</li>
		<li>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 &#176;C under 5% CO2 in a humidified chamber.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#176;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.</li>
	</ol>
	</li>
</ol>
5. Maintenance of the cultured SCG neurons and treatments
<ol>
	<li>On Day 1 (24 hours after plating), carefully remove half of the culture media, and replace with 2 &#181;M Ara-C (cytosine &#946;-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.</li>
	<li>On Day 3, gently aspirate half the medium and replace with control medium.</li>
	<li>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.</li>
	<li>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 &#181;g/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.</li>
</ol>
6. Immunostaining cultured SCG neurons
<ol>
	<li>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.
	<ol>
		<li>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.</li>
		<li>Once all the medium has been replaced by 4% paraformaldehyde, incubate the wells for 15 - 20 minutes at room temperature.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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 &#176;C overnight.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Carefully remove the Triton X-100 solution and rinse the wells three times for 5 minutes each time with PBS. Remove the PBS solution.</li>
	<li>Add enough 5% BSA in PBS to cover the cells and incubate at room temperature for 20 minutes to reduce non-specific antibody binding.</li>
	<li>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 &#176;C.</li>
	<li>Remove and save the primary antibody for reuse. The primary antibody can be reused at least twice without loss of detection intensity.</li>
	<li>Rinse three times with enough PBS to cover the cells. Leave PBS on the cells for 10 minutes per rinse.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Mount the coverslips onto glass slides containing a drop of aqueous mountant that is appropriate for fluorescently labeled samples. Store the slides at 4 &#176;C, in a slide folder until ready for imaging.</li>
</ol>
7. Sample preparation for analysis of the proteome using liquid chromatography coupled with mass spectrometry
<ol>
	<li>Lysis of cultured neurons
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Add 100 - 150 &#181;L 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.
		<ol>
			<li>With the low number of neurons, use a limited volume to lyse the cells to ensure high enough concentration for proteomic analysis.</li>
		</ol>
		</li>
		<li>Freeze the solution at -80 &#176;C overnight to help with cell lysis. The lysates can be stored at this stage until ready for further processing.</li>
		<li>Once thawed, squirt the samples through a syringe with a 26 G or 28 G needle to mechanically lyse the cells.</li>
		<li>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 &#176;C at 12,000 x g.</li>
		<li>Measure the protein concentration. Typical protein concentrations range from 0.4 - 1 &#181;g/&#181;L</li>
	</ol>
	</li>
	<li>Sample preparation and trypsin digestion for proteome analysis
	<ol>
		<li>Transfer a maximum of 60 &#181;L of the lysate or 50 &#181;g of protein into a DNase-, RNase- and protease-free 1.5 mL tube. If the volume of the lysate is less than 60 &#181;L, add enough 50 mM ammonium bicarbonate to make up the volume.</li>
		<li>Add 25 &#181;L of 0.2% acid labile surfactant and vortex. Incubate the tube in a block heater at 80 &#176;C for 15 minutes. Centrifuge the tube at 12,000 x g for 30 s.</li>
		<li>Add 2.5 &#181;L of 100 mM dithiothreitol and vortex. This makes the protein more accessible for alkylation and digestion. Incubate the tube at 60 &#176;C in a block heater for 30 minutes. Cool to room temperature and centrifuge.</li>
		<li>Add 2.5 &#181;L 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.</li>
		<li>Add mass spectrometry grade trypsin (0.5 &#181;g/&#181;L) to the tube at a trypsin: protein ratio of 1:10. Digest the samples at 37 &#176;C overnight.</li>
		<li>Add 10 &#181;L of 5% trifluoroacetic acid (TFA) and vortex. Incubate the samples at 37 &#176;C for 90 minutes. This step is necessary to hydrolyze the acid labile surfactant to prevent interference during mass spectrometry.</li>
		<li>Centrifuge the samples at 12,000 x g at 4 &#176;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 &#176;C prior to mass spectrometry analysis.</li>
		<li>Subject the sample vials to liquid chromatography coupled with high definition mass spectrometry.</li>
	</ol>
	</li>
</ol>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BMP7-induced+dendritic+growth+in+sympathetic+neurons+requires+p75+NTR+signaling.">BMP7-induced dendritic growth in sympathetic neurons requires p75 NTR signaling.</a> Developmental Neurobiology. 76 (9), 1003-1013 (2016).</li><li>Lein, P. J., Fryer, A. D., Higgins, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+Culture:+Autonomic+and+Enteric+Neurons.">Cell Culture: Autonomic and Enteric Neurons.</a> Encyclopedia of Neuroscience. , 625-632 (2009).</li><li>Neto, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compartmentalized+Microfluidic+Platforms:+The+Unrivaled+Breakthrough+of+In+vitro+Tools+for+Neurobiological+Research.">Compartmentalized Microfluidic Platforms: The Unrivaled Breakthrough of In vitro Tools for Neurobiological Research.</a> Journal of Neuroscience. 36 (46), 11573-11584 (2016).</li><li>Sleigh, J. N., Weir, G. A., Schiavo, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple,+step-by-step+dissection+protocol+for+the+rapid+isolation+of+mouse+dorsal+root+ganglia.">A simple, step-by-step dissection protocol for the rapid isolation of mouse dorsal root ganglia.</a> BMC Research Notes. 9 (1), 82 (2016).</li><li>Conrad, R., Jablonka, S., Sczepan, T., Sendtner, M., Wiese, S., Klausmeyer, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lectin-based+isolation+and+culture+of+mouse+embryonic+motoneurons.">Lectin-based isolation and culture of mouse embryonic motoneurons.</a> Journal of Visualized Experiments. (55), e3200 (2011).</li><li>Takeuchi, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfabricated+device+for+co-culture+of+sympathetic+neuron+and+iPS-derived+cardiomyocytes.">Microfabricated device for co-culture of sympathetic neuron and iPS-derived cardiomyocytes.</a> Proceedings of the Annual International Conference of the IEEE Engineering in Medicine and Biology Society, EMBS. 2013, 3817-3820 (2013).</li><li>Takeuchi, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+semi-separated+co-culture+system+of+sympathetic+neuron+and+cardiomyocyte.">Development of semi-separated co-culture system of sympathetic neuron and cardiomyocyte.</a> Proceedings of the 31st Annual International Conference of the IEEE Engineering in Medicine and Biology Society: Engineering the Future of Biomedicine, EMBC 2009. , 1832-1835 (2009).</li><li>Chandrasekaran, V., Lea, C., Sosa, J. C., Higgins, D., Lein, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reactive+oxygen+species+are+involved+in+BMP-induced+dendritic+growth+in+cultured+rat+sympathetic+neurons.">Reactive oxygen species are involved in BMP-induced dendritic growth in cultured rat sympathetic neurons.</a> Molecular and Cellular Neuroscience. 67, (2015).</li><li>Kim, W. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Statins+decrease+dendritic+arborization+in+rat+sympathetic+neurons+by+blocking+RhoA+activation.">Statins decrease dendritic arborization in rat sympathetic neurons by blocking RhoA activation.</a> Journal of Neurochemistry. 108 (4), 1057-1071 (2009).</li><li>Dalby, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advanced+transfection+with+Lipofectamine+2000+reagent:+Primary+neurons,+siRNA,+and+high-throughput+applications.">Advanced transfection with Lipofectamine 2000 reagent: Primary neurons, siRNA, and high-throughput applications.</a> Methods. 33 (2), 95-103 (2004).</li><li>Szpara, M. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+gene+expression+during+neurite+outgrowth+and+regeneration.">Analysis of gene expression during neurite outgrowth and regeneration.</a> BMC Neuroscience. 8 (1), 100 (2007).</li><li>Pop, C., Mogosan, C., Loghin, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+rapigest+efficacy+for+the+digestion+of+proteins+from+cell+cultures+and+heart+tissue.">Evaluation of rapigest efficacy for the digestion of proteins from cell cultures and heart tissue.</a> Clujul Medical. 87 (4), 5 (2014).</li><li>Vit, O., Petrak, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integral+membrane+proteins+in+proteomics.+How+to+break+open+the+black+box.">Integral membrane proteins in proteomics. How to break open the black box.</a> Journal of Proteomics. 153, 8-20 (2017).</li></ol>

The authors have nothing to disclose.

This paper describes the protocols for culturing sympathetic neurons from superior cervical ganglia of embryonic rat pups. The advantages of using this model system are that it is possible to obtain a homogeneous population of neurons providing a similar response to growth factors, and since the growth factor requirements for these neurons has been well -characterized, it is possible to grow these neurons in vitro in defined media, under serum-free conditions10. Although the protocol describes the...

Sympathetic neurons derived from embryonic superior cervical ganglia (SCG) have been widely used as a primary neuronal culture system for studying many aspects of neuronal development including growth factor dependence, neuron-target interactions, neurotransmitter signaling, axonal growth, dendrite development and plasticity, synaptogenesis and signaling mechanisms underlying nerve-target/neuron-glia interactions1,2,3,4,5,6,7,8,9. Despite their small size (around 10000 neurons/ganglia), there are three main reasons for the development and extensive use of this culture system are i) being the first ganglia in the sympathetic chain, they are larger, and therefore easier to isolate, than the rest of the sympathetic ganglia10; ii) unlike central neurons, the neurons in the SCG are fairly homogeneous with all the neurons being derived from the neural crest, having a similar size, dependence on nerve growth factor and being nor-adrenergic. This makes them a valuable model for morphological and genomic studies10,11 and iii) these neurons can be maintained in a defined serum-free medium containing nerve growth factor for over a month10,12. Perinatal SCG neurons have been extensively used for studying the mechanisms underlying the initiation and maintenance of dendrites2. This is mainly because, although SCG neurons have an extensive dendritic arbor in vivo, they do not extend dendrites in vitro in the absence of serum but can be induced to grow dendrites in the presence of certain growth factors such as bone morphogenetic proteins2,12,13.
This paper describes the protocol for isolating and culturing embryonic rat SCG neurons. Over the past 50 years, primary neuronal cultures from the SCG have been mainly used for morphological studies with a limited number of studies examining the large-scale genomic or proteomic changes. This is mainly due to small tissue size resulting in the isolation of low amounts of DNA or protein, which makes it difficult to perform genomic and proteomic analyses on these neurons. However, in recent years, increased detection sensitivity has enabled development of methods to examine the genome, miRNome and proteome in the SCG neurons during dendritic growth development14,15,16,17. This paper will also describe the method for morphological analysis of these neurons using immunocytochemistry and a protocol to obtain neuronal protein extracts for mass spectrometric analysis.

<table>
	<tbody>
		<tr>
			<td>2D nanoACQUITY</td>
			<td>Waters Corporation</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>9830</td>
			<td></td>
		</tr>
		<tr>
			<td>BMP-7</td>
			<td>R&#38;D Systems</td>
			<td>354-BP</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Alumin</td>
			<td>Sigma-Aldrich</td>
			<td>5470</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell scraper</td>
			<td>Corning</td>
			<td>CLS-3010</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase</td>
			<td>Worthington Biochemical</td>
			<td>4176</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Costar or Nunc Flat bottomed Cell culture plates</td>
			<td>Fisher Scientific</td>
			<td>07-200, 140675, 142475</td>
			<td></td>
		</tr>
		<tr>
			<td>Cytosine- &#946;- D-arabinofuranoside</td>
			<td>Sigma-Aldrich</td>
			<td>C1768</td>
			<td></td>
		</tr>
		<tr>
			<td>D-phosphate buffered saline (Calcium and magnesium free)</td>
			<td>ATCC</td>
			<td>30-2200</td>
			<td></td>
		</tr>
		<tr>
			<td>Dispase II</td>
			<td>Roche</td>
			<td>4942078001</td>
			<td></td>
		</tr>
		<tr>
			<td>Distilled Water</td>
			<td>Thermo Fisher Scientific</td>
			<td>15230</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol</td>
			<td>Sigma-Aldrich</td>
			<td>D0632</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM - Low glucose + Glutamine, + sodium pyruvate</td>
			<td>Thermo Fisher Scientific</td>
			<td>11885</td>
			<td></td>
		</tr>
		<tr>
			<td>Fatty Acid Free BSA</td>
			<td>Calbiochem</td>
			<td>126609</td>
			<td>20 mg/mL stock in low glucose DMEM</td>
		</tr>
		<tr>
			<td>Fine forceps Dumont no.4 and no.5</td>
			<td>Ted Pella Inc</td>
			<td>5621, 5622</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps and Scissors for Dissection</td>
			<td>Ted Pella Inc</td>
			<td>1328, 1329, 5002</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverlips - 12mm</td>
			<td>Neuvitro Corporation</td>
			<td>GG-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat-Anti Mouse IgG Alexa 488 conjugated</td>
			<td>Thermo Fisher Scientific</td>
			<td>A32723</td>
			<td></td>
		</tr>
		<tr>
			<td>Ham&#39;s F-12 Nutrient Mix</td>
			<td>Thermo Fisher Scientific</td>
			<td>11765</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#39;s balanced salt soltion (Calcium and Magnesium free)</td>
			<td>Thermo Fisher Scientific</td>
			<td>14185</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin-Selenium-Transferrin (100X)</td>
			<td>Thermo Fisher Scientific</td>
			<td>41400-045</td>
			<td></td>
		</tr>
		<tr>
			<td>Iodoacetamide</td>
			<td>Sigma-Aldrich</td>
			<td>A3221</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Thermo Fisher Scientific</td>
			<td>25030</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz L-15 medium</td>
			<td>Thermo Fisher Scientific</td>
			<td>11415064</td>
			<td></td>
		</tr>
		<tr>
			<td>Mounting media for glass coverslips</td>
			<td>Thermo Fisher Scientific</td>
			<td>P36931, P36934</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse-anti- MAP2 antibody (SMI-52)</td>
			<td>BioLegend</td>
			<td>SMI 52</td>
			<td></td>
		</tr>
		<tr>
			<td>Nerve growth factor</td>
			<td>Envigo Bioproducts (formerly Harlan Bioproducts)</td>
			<td>BT5017</td>
			<td>Stock 125 &#956;g/mL in 0.2% Prionex in DMEM</td>
		</tr>
		<tr>
			<td>Paraformaldehye</td>
			<td>Spectrum Chemicals</td>
			<td>P1010</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (100X)</td>
			<td>Thermo Fisher Scientific</td>
			<td>15140</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-D-Lysine</td>
			<td>Sigma-Aldrich</td>
			<td>P0899</td>
			<td></td>
		</tr>
		<tr>
			<td>Prionex</td>
			<td>Millipore</td>
			<td>529600</td>
			<td>10% solution, 100 mL</td>
		</tr>
		<tr>
			<td>RapiGest SF</td>
			<td>Waters Corporation</td>
			<td>186001861</td>
			<td>5 X 1 mg</td>
		</tr>
		<tr>
			<td>Synapt G2 High Definition Mass Spectrometry</td>
			<td>Waters Corporation</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trifluoro acetic acid - Sequencing grade</td>
			<td>Thermo Fisher Scientific</td>
			<td>28904</td>
			<td>10 X 1 mL</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>X100</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Promega or NEB</td>
			<td>V511A, P8101S</td>
			<td>100 &#956;g or 5 X 20 mg</td>
		</tr>
		<tr>
			<td>Waters Total recovery vials</td>
			<td>Waters Corporation</td>
			<td>186000385c</td>
			<td></td>
		</tr>
	</tbody>
</table>

culturing rat sympathetic neurons from embryonic superior cervical ganglia for morphological and proteomic analysis

Sympathetic neurons from the embryonic rat superior cervical ganglia (SCG) have been used as an in vitro model system for peripheral neurons to study axonal growth, axonal trafficking, synaptogenesis, dendritic growth, dendritic plasticity and nerve-target interactions in co-culture systems. This protocol describes the isolation and dissociation of neurons from the superior cervical ganglia of E21 rat embryos, followed by the preparation and maintenance of pure neuronal cultures in serum-free medium. Since neurons do not adhere to uncoated plastic, neurons will be cultured on either 12 mm glass coverslips or 6-well plates coated with poly-D-lysine. Following treatment with an antimitotic agent (Ara-C, cytosine &#946;-D-arabinofuranoside), this protocol generates healthy neuronal cultures with less than 5% non-neuronal cells, which can be maintained for over a month in vitro. Although embryonic rat SCG neurons are multipolar with 5-8 dendrites in vivo; under serum-free conditions, these neurons extend only a single axon in culture and continue to be unipolar for the duration of the culture. However, these neurons can be induced to extend dendrites in the presence of basement membrane extract, bone morphogenetic proteins (BMPs), or 10% fetal calf serum. These homogenous neuronal cultures can be used for immunocytochemical staining and for biochemical studies. This paper also describes optimized protocol for immunocytochemical staining for microtubule associated protein-2 (MAP-2) in these neurons and for the preparation of neuronal extracts for mass spectrometry.

Isolating and maintaining neuronal cultures of embryonic SCG neurons 
Dissociated cells from the rat embryonic SCG were plated in a poly-D-lysine coated plate or coverslip and maintained in serum free culture media containing b-nerve growth factor. The dissociated cells containing a mixture of neurons and glial cells look circular upon plating (Figure 1A). Within 24 hours of plating, the neurons extend small axonal processes with glial cells flattening and appearing pha...

Watch this Scientific Journal Video about Culturing Rat Sympathetic Neurons from Embryonic Superior Cervical Ganglia for Morphological and Proteomic Analysis at JoVE.com

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

This paper describes the isolation and culturing of embryonic rat sympathetic neurons from the superior cervical ganglia. It also provides detailed protocols for immunocytochemical staining and for preparing neuronal extracts for mass spectrometric analysis.

Sympathetic neurons from the embryonic rat superior cervical ganglia (SCG) have been used as an in vitro model system for peripheral neurons to study ...

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7.4K Views.  Saint Mary's College of California. This paper describes the isolation and culturing of embryonic rat sympathetic neurons from the superior cervical ganglia. It also provides detailed protocols for immunocytochemical staining and for preparing neuronal extracts for mass spectrometric analysis.

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

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2 or record extracellular neuronal activity during a movement.3,4 While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo movement thus allowing substantial insight into the role of the neuron in sensorimotor function.

A technique is described to quantify the in vivo physiological response of mammalian neurons during movement and correlate the physiology of the neuron with neuronal morphology, neurochemical phenotype and synaptic microcircuitry.

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor ...

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after injury (Teng et al., 2006). It is recognized that transcriptional programs essential for neurite and axonal outgrowth are 
reactivated upon injury in the PNS (Makwana et al., 2005). However the tools available to analyze neuronal gene regulation in vivo are limited and 
often challenging.
The dorsal root ganglia (DRG) offer an excellent injury model system because both the CNS and PNS are innervated by a 
bifurcated axon originating from the same soma. The ganglia represent a discrete collection of cell bodies where all transcriptional events occur, 
and thus provide a clearly defined region of transcriptional activity that can be easily and reproducibly removed from the animal. Injury of nerve 
fibers in the PNS (e.g. sciatic nerve), where axonal regeneration does occur, should reveal a set of transcriptional programs that are distinct from 
those responding to a similar injury in the CNS, where regeneration does not take place (e.g. spinal cord). Sites for transcription factor binding, 
histone and DNA modification resulting from injury to either PNS or CNS can be characterized using chromatin immunoprecipitation (ChIP). 
Here, we describe a ChIP protocol using fixed mouse DRG tissue following axonal injury. This powerful combination provides a means for characterizing the pro-regeneration chromatin environment necessary for promoting axonal regeneration.

We present a method for chromatin immunoprecipitation from dorsal root ganglia tissue following axonal injury. The approach can be used to identify specific transcription factor binding sites and epigenetic modification of histone and DNA important for the regeneration of injured axons in both the peripheral and central nervous system.

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after ...

Cerebrovascular Casting of the Adult Mouse for 3D Imaging and Morphological Analysis

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). Animal models are necessary for studying the etiopathology and potential therapies of cerebrovascular diseases. Imaging the vasculature in large animals is relatively easy. However, developing vessel imaging methods of murine brain disease models is desirable due to the cost and availability of genetically-modified mouse lines. Imaging the murine cerebral vascular tree is a challenge. In humans and larger animals, the gold standard for assessing the angioarchitecture at the macrovascular (conductance) level is x-ray catheter contrast-based angiography, a method not suited for small rodents.
In this article, we present a method of cerebrovascular casting that produces a durable skeleton of the entire vascular bed, including arteries, veins, and capillaries that may be analyzed using many different modalities. Complete casting of the microvessels of the mouse cerebrovasculature can be difficult; however, these challenges are addressed in this step-by-step protocol. Through intracardial perfusion of the vascular casting material, all vessels of the body are casted. The brain can then be removed and clarified using the organic solvent methyl salicylate. Three dimensional imaging of the brain blood vessels can be visualized simply and inexpensively with any conventional brightfield microscope or dissecting microscope. The casted cerebrovasculature can also be imaged and quantified using micro-computed tomography (micro-CT)1. In addition, after being imaged, the casted brain can be embedded in paraffin for histological analysis.
The benefit of this vascular casting method as compared to other techniques is its broad adaptation to various analytic tools, including brightfield microscopic analysis, CT scanning due to the radiopaque characteristic of the material, as well as histological and immunohistochemical analysis. This efficient use of tissue can save animal usage and reduce costs. We have recently demonstrated application of this method to visualize the irregular blood vessels in a mouse model of adult BAVM at a microscopic level2, and provide additional images of the malformed vessels imaged by micro-CT scan. Although this method has drawbacks and may not be ideal for all types of analyses, it is a simple, practical technique that can be easily learned and widely applied to vascular casting of blood vessels throughout the body.

In this article, we present a simple, practical technique for cerebrovascular casting that is easy to perform and can be utilized to image the vascular tree of the adult mouse brain.

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Production and Isolation of Axons from Sensory Neurons for Biochemical Analysis Using Porous Filters

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is required to shape functional neuronal circuits. The protocol described here explains how to use cell culture inserts bearing a porous membrane (filter) to obtain large amounts of pure axonal preparations suitable for examination by conventional biochemical or immunocytochemical techniques. The functionality of these filter inserts will be demonstrated with models of developmental pruning and Wallerian degeneration, using explants of embryonic dorsal root ganglion. Axonal integrity and function is compromised in a wide variety of neurodegenerative pathologies. Indeed, it is now clear that axonal dysfunction appears much earlier in the course of the disease than neuronal soma loss in several neurodegenerative diseases, indicating that axonal-specific processes are primarily targeted in these disorders. By obtaining pure axonal samples for analysis by molecular and biochemical techniques, this technique has the potential to shed new light into mechanisms regulating the physiology and pathophysiology of axons. This in turn will have an impact in our understanding of the processes that drive degenerative diseases of the nervous system.

This article describes a neuronal culture system that can be used to obtain large quantities of pure axonal samples for biochemical and immunocytochemical analyses. This technique will allow an improved understanding of normal axonal physiology and signaling pathways leading to neurodegeneration.

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is ...

A Method of Nodose Ganglia Injection in Sprague-Dawley Rat

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located in the organs of the abdomen and thorax. The vagus nerve communicates information from stimuli such as heart rate, blood pressure, bronchopulmonary irritation, and gastrointestinal distension to the nucleus of solitary tract of the medulla. The cell bodies of the vagus nerve are located in the nodose and petrosal ganglia, of which the majority are located in the former. The nodose ganglia contain a wealth of receptors for amino acids, monoamines, neuropeptides, and other neurochemicals that can modify afferent vagus nerve activity. Modifying vagal afferents through systemic peripheral drug treatments targeted at the receptors on nodose ganglia has the potential of treating diseases such as sleep apnea, gastroesophageal reflux disease, or chronic cough. The protocol here describes a method of injection neurochemicals directly into the nodose ganglion. Injecting neurochemicals directly into the nodose ganglia allows study of effects solely on cell bodies that modulate afferent nerve activity, and prevents the complication of involving the central nervous system as seen in systemic neurochemical treatment. Using readily available and inexpensive equipment, intranodose ganglia injections are easily done in anesthetized Sprague-Dawley rats.

Afferent vagal signaling transmits important information to central nervous system from receptors located in organs of the abdomen and thorax. The nodose ganglia of vagus nerves contain many types of receptors that modulate vagal activity. This protocol describes a method of local injections of neurochemicals into the nodose ganglia.

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located ...

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and morphological levels. Compound muscle action potential (CMAP) recording following phrenic nerve stimulation can be used to quantitatively assess functional diaphragm innervation by PhMNs of the cervical spinal cord in vivo in anesthetized rats and mice. Because CMAPs represent simultaneous recording of all myofibers of the whole hemi-diaphragm, it is useful to also examine the phenotypes of individual motor axons and myofibers at the diaphragm NMJ in order to track disease- and therapy-relevant morphological changes such as partial and complete denervation, regenerative sprouting and reinnervation. This can be accomplished via whole-mount immunohistochemistry (IHC) of the diaphragm, followed by detailed morphological assessment of individual NMJs throughout the muscle. Combining CMAPs and NMJ analysis provides a powerful approach for quantitatively studying diaphragmatic innervation in rodent models of CNS and PNS disease.

Compound muscle action potential recording quantitatively assesses functional diaphragm innervation by phrenic motor neurons. Whole-mount diaphragm immunohistochemistry assesses morphological innervation at individual neuromuscular junctions. The goal of this protocol is to demonstrate how these two powerful methodologies can be used in various rodent models of spinal cord disease.

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen films. Favorable clinical outcomes have been achieved when using these tubes for the treatment of damaged inferior alveolar nerve (IAN). A critical factor for the successful nerve regeneration using PGA-C tubes is blood supply to the surrounding tissue. Cervical sympathetic ganglion block (CSGB) creates a sympathetic blockade in the head and neck region thus increasing blood flow in the area. To ensure an adequate effect, the blockade must be administered with local anesthetics one to two times a day for several consecutive weeks; this poses a challenge when creating animal models for investigating this technique. To address this limitation, we developed an ethanol-induced CSGB in a canine model of long-term increase in blood flow in the orofacial region. We examined whether IAN regeneration via PGA-C tube implantation can be enhanced by this model. Fourteen Beagles were each implanted with a PGA-C tube across a 10-mm gap in the left IAN. The IAN is located within the mandibular canal surrounded by bone, therefore we chose piezoelectric surgery, consisting of ultrasonic waves, for bone processing, in order to minimize the risk of nerve and vessel injury. A good surgical outcome was obtained with this approach. A week after surgery, seven of these dogs were subjected to left CSGB by injection of ethanol. Ethanol-induced CSGB resulted in improved nerve regeneration, suggesting that the increased blood flow effectively promotes nerve regeneration in IAN defects. This canine model can contribute to further research on the long-term effects of CSGB.

We evaluated the effect of cervical sympathetic ganglion block on nerve repair using artificial nerve conduits. Male beagle dogs were each implanted with an artificial nerve across a 10-mm gap in the left inferior alveolar nerve; left cervical sympathetic ganglion was blocked by injecting 99.5% ethanol via lateral thoracotomy.

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen ...