This work was supported by funding from the National Institutes of Health (grants R21 NS45037 and R01 ES014901).

1. Preparation of Culture Medium (C2 medium)
<ol>
	<li>Add the following, in the order listed, to a sterile, disposable 500 ml Erlenmeyer flask:
	<table border="1">
		<tbody>
			<tr>
				<td>Amount </td>
				<td>Component </td>
				<td>Final Concentration</td>
			</tr>
			<tr>
				<td>190 ml</td>
				<td>DMEM (low glucose)</td>
				<td>N/A</td>
			</tr>
			<tr>
				<td>10 ml</td>
				<td>fatty acid free BSA (20mg/ml in DMEM)</td>
				<td>50 &mu;g/ml</td>
			</tr>
			<tr>
				<td>2.8 ml</td>
				<td>L-Glutamine</td>
				<td>1.4 mM</td>
			</tr>
			<tr>
				<td>4 ml</td>
				<td>insulin/transferrin/selenium(100x)</td>
				<td>10 &mu;g/ml insulin 5.5 &mu;g/ml transferrin 38.7 nM selenium</td>
			</tr>
			<tr>
				<td>0.4 ml</td>
				<td>NGF (125 &mu;g/ml)</td>
				<td>100 ng/ml</td>
			</tr>
			<tr>
				<td>200 ml</td>
				<td>Ham F-12</td>
				<td>N/A</td>
			</tr>
		</tbody>
	</table>
	</li>
	<li>Swirl gently to mix and aliquot 10 ml per tube in 17 x 100 sterile snap cap tubes.</li>
	<li>Store at -80 &deg;C</li>
	<li>Important notes:
	<ul type="disc">
		<li>Thaw NGF immediately prior to usage.</li>
		<li>Before aliquoting the NGF stock solution, precoat the interior walls of the tissue culture plastic serological pipet with protein to minimize sticking of NGF to the plastic. Easiest approach is to Pipette the DMEM mixture containing the BSA several times. Rinse the NGF tube several times with the DMEM mixture.</li>
		<li>Add F-12 last because free cysteine may destabilize disulfide bonds in the insulin.</li>
		<li>Don't freeze in volumes greater than 10 ml because CaPO4 crystals may form</li>
		<li>Do not freeze-thaw C2 medium more than 2 times.</li>
	</ul>
	</li>
</ol>
2. Preparation of Glass Coverslips
<ol>
	<li>We have found that only fine German glass coverslips work consistently in this protocol (see Table 1 for recommended source and catalog number).</li>
	<li>Add 75-100 ml of 70% nitric acid to 300 ml glass specimen bottles with polytetrafluoroethylene (PTFE or Teflon) lid liners (Fisher catalog #09-911-765).</li>
	<li>Wearing gloves, drop coverslips one by one into the nitric acid solution. Add enough coverslips to make a layer no more than 2-3 coverslips thick on the bottom of the jar.</li>
	<li>Gently shake the containers with the coverslips for 6-8 hours at room temperature on an orbital shaker set for no more than 185 rpm.</li>
	<li>Working in a fume hood, carefully decant the acid and dispose according to local regulations. Rinse coverslips once with ultra-pure tissue culture-grade water, then add 75-100 ml water to the container. Coverslips may be rinsed multiple times if the water becomes cloudy. Gently shake at room temperature overnight on an orbital shaker set for no more than 185 rpm.</li>
	<li>Repeat the above sequence of nitric acid/ tissue culture-grade water daily for a total of 3 days.</li>
	<li>Decant the final tissue culture-grade water and add enough acetone to barely cover the coverslips.</li>
	<li>Decant this acetone and dispose properly.</li>
	<li>Transfer coverslips to a 150 mm glass Petri dish. Add enough acetone to barely cover the coverslips. Ensure the coverslips are in a single layer in the dish.</li>
	<li>Allow the acetone to evaporate overnight in a fume hood.</li>
	<li>Bake coverslips in a drying oven at 180 &deg;C for 1.5 hours to sterilize.</li>
</ol>
3. Preparation of Coverslips for Culture
<ol>
	<li>The day before the dissection, add 1 glass coverslip (prepared as described above) to each well of a 24 well tissue culture plate using sterile technique. Add 200 &mu;l of sterile poly-D-lysine solution (100 mg/ml in 0.5 M Tris buffer, pH 8) to each well and store the plate at 4 &deg;C overnight.</li>
	<li>The day of the dissection, and ideally before starting the dissection, aspirate the poly-D-lysine solution from each well and wash each well 5 times with sterile tissue-culture grade water.</li>
	<li>Add 200 &mu;l low-glucose DMEM to each plate and store in 37 &deg;C incubator. Approximately 1 hour prior to plating cells, remove low-glucose DMEM and add 250 &mu;l C2 medium per well. Store plates at 37 &deg;C under 5% CO2 until cells are ready to be plated.</li>
</ol>
4. Dissection and Isolation of the Superior Cervical Ganglion
<ol>
	<li>Dissociated cultures of sympathetic neurons are usually prepared by dissociation of superior cervical ganglia (SCG) harvested from prenatal (embryonic days 19-21) rat pups because there are fewer glial cells. However, this same protocol can be used successfully with early postnatal (1 to 3 days old) rat pups or perinatal mouse pups.</li>
	<li>Euthanize pregnant rat via CO2 inhalation. Shave the abdomen and wash the skin with 70% ethanol. Remove the uterine horns under sterile conditions and place the uterine horns in a sterile 150 mm Petri dish. Avoid damaging the intestines or other internal organs of the dam during the dissection.</li>
	<li>Place the Petri dish with the pups onto a pan of ice to anesthetize the embryonic pups. Dissect the embryos free of the uterine horn and amniotic membranes. Cut the spinal cord of each pup along the midline under the shoulder to euthanize the pups. This also severs the vena cava, which reduces bleeding from the carotid artery during dissection of the SCG. Once dissected free of the uterine horn, place embryos in a sterile Leibovitz's L-15 medium (or any air-buffered medium) supplemented with penicillin and streptomycin.</li>
	<li>Place the embryos on their backs on a 150 mm Petri dish half-filled with Sylgard. Using sterile 20 gauge needles, pin the thorax and the head, gently hyperextending the neck, which facilitates dissection of the SCG.</li>
	<li>Make a midline incision along the neck at the level of the clavicles to reveal the submandibular glands. Remove the glands using fine forceps (no. 4 or no. 5 Dumont) to expose the muscle layer.</li>
	<li>Use the fine forceps to cut the sternocleidomastoid muscle near the clavicle. Then gently lift and incise the omohyoid muscle next to the trachea. This muscle is very thin, so avoid cutting too deeply to avoid tearing the underlying carotid artery and SCG. Once these muscle layers are removed, the carotid artery and the vagus nerve should be clearly visible.</li>
	<li>The SCG lies just below the bifurcation of the carotid artery. Using fine forceps (no. 4 or no. 5), blunt dissect tissue away from both sides of the carotid artery to expose the ganglion. Using 2 fine forceps, grasp the carotid artery just above and below the rostral and caudal ends of the SCG, respectively, to remove the section of the carotid artery lying above the SCG as well as the SCG itself. It is likely that the nodose ganglion, which lies alongside the SCG at this stage of development, will also be removed at this step. The nodose ganglia can be identified by its round shape and the large vagal nerve projecting from it. This lies along one of the branches of the carotid artery. In contrast, the SCG is almond shaped and lies directly under the bifurcation of the carotid artery. Place the dissected tissue into a sterile 35 mm culture dish containing L-15 medium and antibiotics. Remove the SCG from all pups before progressing to the next steps.</li>
	<li>Using fine forceps, gently separate the SCG from the carotid artery and any other tissues removed during the dissection (particularly the vagus nerve and nodose ganglion). Use fine forceps to remove the capsule from the ganglia. This greatly reduces the numbers of nonneuronal cells in culture. Transfer the ganglia to another sterile 35 mm dish containing L-15 medium.</li>
	<li>To dissociate the SCG, remove the L-15 medium and replace with 2 ml Collagenase (final concentration 1 mg/ml)/Dispase (final concentration 5 mg/ml) in calcium and magnesium-free Hank's balanced salt solution. Incubate at 37 &deg;C for 40 minutes.</li>
	<li>Transfer the SCG to a sterile 15 ml conical centrifuge tube and bring the volume up to 10 ml with L-15 supplemented with BSA (final concentration 1-2 mg/ml). Centrifuge at 200 X g at room temperature for 5 minutes. Discard the supernatant and wash once more with L-15 supplemented with BSA.</li>
	<li>After the second wash, remove all L-15 and resuspend the ganglia in ~2 ml of C2 medium. Triturate the cells using a fire-polished bent-tip pipette, make sure the pipette is coated with BSA/L15 before trituration to minimize sticking of cells to the glass. Triturate gently 3-4 times. Let the large clumps settle for about 1 minute and transfer the cell suspension to a 15 ml conical tube. Add more C2 media to the remaining SCG and triturate. After letting the clumps settle, transfer the cell suspension from the second trituration to that collected after the first trituration. Repeat this process several times until the ganglia are mostly dissociated. Discard any remaining clumps after the fourth trituration.</li>
	<li>For cells dissociated from one litter of rat pups (typically 12-16 pups), bring the total volume of medium in the cell suspension to 8-10 ml by adding additional C2 medium. Gently resuspend the cells using a pipette and remove a small aliquot to determine the cell density. Adjust the volume of the cell suspension to achieve 2X the cell density desired in the culture, and then, being sure to maintain the cells in suspension, aliquot 250 ml of cell suspension per well. For morphometric studies of dendritic growth, we typically plate cells at low density (~5 x 104 cells per well) in 24 well plates.</li>
	<li>Our laboratory incubates SCG cultures in glass desiccators rather than directly in a CO2 incubator. We do this because sympathetic neurons grown in serum-free medium perform better with a slightly higher CO2 concentration (approximately 6.5%). In addition we do not use antibiotics in our media since it inhibits neurite outgrowth and maintaining cultures in the desiccators seems to minimize spread of contamination if it develops in a subset of cultures. Using this system, we routinely maintain SCG cultures for up to 8 weeks. Once plated, SCG cultures are placed on the porcelain desiccator plate in glass desiccators containing sterile water in the bottom. Approximately 120 ml of CO2 is injected into the desiccator prior to sealing it shut and placing the entire desiccator into an incubator set for 35.5 &deg;C.</li>
</ol>
5. Feeding and Maintenance of Cultures
<ol>
	<li>Cultures are generally fed 3 times a week, with the first media exchange occurring 24 hours after plating. Using a sterile bent-tip pipette, gently withdraw about half of the media per well. Using a clean sterile bent-tip pipette, replace the volume of medium removed with fresh C2.</li>
	<li>To eliminate non-neuronal cells from the culture, the anti-mitotic agent cytosine-D-arabinofuranoside (ARA-C, final concentration 1-2 &mu;M) is typically added to the culture medium at the first media exchange (e.g., 24 hours after plating) for 48 hours. This concentration of ARA-C is usually sufficient to remove all non-neuronal cells. If additional ARA-C treatment is required, it is recommended that these be done every other feeding to minimize toxic effects on the neurons.</li>
</ol>
6. Inducing Dendritic Growth with Matrigel or BMP-7
<ol>
	<li>To induce dendritic growth, Matrigel or BMP-7 is added to cultures after non-neuronal cells have been eliminated and ARA-C levels in the cultures are significantly reduced. Typically, we add Matrigel or BMP-7 to the medium on day 5 in vitro, but dendritic growth can be induced by Matrigel or BMP-7 added at any time point in vitro. Matrigel or BMP-7 is added to the C2 medium and cultures are fed as described above.</li>
	<li>Matrigel and BMP-induced dendritic growth is time and concentration-dependent. For maximal effects, Matrigel is added to C2 at a final concentration of 50-75 &mu;g/ml; BMP-7 is added to the C2 medium at a final concentration of 50 ng/ml. Matrigel concentrations of greater than 100 &mu;g/ml or BMP-7 concentrations greater than 100 ng/ml have been noted to cause neuronal toxicity. Dendritic processes (as determined by morphological criteria) become apparent approximately 48 hours after initial exposure to Matrigel or BMP-7 at maximally effective concentrations. To induce robust dendritic growth, cultures are fed with BMP-7 containing medium at every feeding. Equivalent results have been obtained using BMPs 2, 4, 5 and 6 at comparable concentrations 20,21.</li>
</ol>
7. Immunocytochemical Analysis of BMP-7-induced Dendritic Growth
<ol>
	<li>When the desired extent of dendritic arborization has been achieved, cultures are fixed using 4% paraformaldehyde in 0.1 M phosphate buffer. The best fixation is achieved by replacing half the medium in the well with 4% paraformaldehyde and repeating this step 3-5 times over 10-15 minutes.</li>
	<li>Rinse cultures by replacing half the 4% paraformaldehyde with phosphate-buffered saline (PBS) and repeating this step three times over 10- 15 minutes.</li>
	<li>Remove all the PBS and add 200 &mu;l 0.1% Triton X-100 in PBS to each well for 5 minutes at room temperature to permeabilize the cells.</li>
	<li>Remove Triton solution by aspiration and add 200 &mu;l per well of blocking buffer solution (PBS supplemented with 3-5% BSA and 1% goat serum). Incubate for 20-30 minutes at room temperature.</li>
	<li>Remove the blocking buffer and add the primary antibody, microtubule-associated protein-2 (MAP2) diluted at 1:2000 - 1:5000 in blocking buffer. Incubate cultures for 1 hour at room temperature or overnight at 4 &deg;C.</li>
	<li>Remove the primary antibody solution, wash the cultures with PBS three times over 15 minutes at room temperature.</li>
	<li>Incubate the cultures with the secondary antibody diluted in PBS supplemented with 1% BSA for 1-2 hours at room temperature in the dark.</li>
	<li>Remove the secondary antibody solution and wash the cultures with PBS three times over 10-15 minutes at room temperature in the dark.</li>
	<li>Mount coverslips on glass slides cell side down in a drop of aqueous mounting medium at the appropriate pH for the fluorochrome tagged to the secondary antibody. Keep the slides at room temperature overnight to let the mounting medium dry. Store slides at 4 &deg;C until imaging.</li>
	<li>Allow the slides to equilibrate to room temp before imaging. Acquire fluorescent images using a phase-contrast microscope equipped for epifluorescence. Typically, the entire dendritic arbor of a single neuron can be captured using a 20X objective and appropriate filters.</li>
</ol>
8. Representative Results
Sympathetic neurons dissociated from the SCG of perinatal rats and grown in the absence of serum and ganglionic glial cells fail to extend MAP2 immunopositive processes (Figure 1), but rather, typically extend only a single axonal process 14,15. Exposure to BMP-7 (Figure 1) or Matrigel induces the formation of numerous processes that meet the morphological, biochemical and functional criteria of dendrites 17,18. The dendrite-promoting activity of either Matrigel or BMP-7 is concentration- and time-dependent 17,18,20. Maximal dendritic growth is observed using concentrations of Matrigel between 50 and 75 &mu;g/ml or BMP-7 between 30 and 100 ng/ml, and half-maximal effects are typically observed at BMP-7 concentrations of ~2 ng/ml. However, significant changes in dendritic growth can be detected with BMP-7 concentrations as low as 300 pg/ml. The dendritic response to either Matrigel or BMP-7 is relatively slow, with &lt;50% of the neurons forming a second process within 24 hours after exposure to either dendrite-promoting agent. However, within 3 days after the initial BMP-7 exposure, virtually all neurons respond to maximally effective Matrigel or BMP-7 concentrations. The number of dendrites per neurons continues to increase with continuous exposure to BMP-7, with most of the change occurring during the first 10 days of treatment. After 4 weeks, BMP-7-treated neurons typically have 6-8 primary dendrites that exhibit secondary, tertiary and even quaternary dendritic branches 17. This increase in dendritic arbor occurs in the absence of any significant change in axonal growth, and comparable dendritic responses can be elicited using similar concentrations of BMPs 2, 4, 5 or 6 20,21. Dendritic growth can also be elicited by co-culturing sympathetic neurons with endogenous ganglionic glial cells 15,22; however, the dendritic response is significantly slower under these conditions.
<img alt="Figure 1" src="/files/ftp_upload/3546/3546fig1.jpg" /> 
Figure 1. BMP-7 promotes dendritic growth in cultured sympathetic neurons. Non-neuronal cells were eliminated from SCG cultures by treatment with anti-mitotic for 48 hr beginning on day 2 in vitro. Beginning on day 5 in vitro, cultures were treated with either control medium (A) or medium supplemented with BMP-7 at 50 ng/ml (B). On day 11 in vitro (after 5 days of BMP treatment), cultures were immunostained with mAb against MAP2, a protein found primarily in dendrites and neuronal somata. As shown in representative fluorescence photomicrographs, neurons grown under control conditions lack dendrites as evidenced by the lack of MAP-2 immunopositive processes dendrites (A); in contrast, neurons exposed to BMP-7 (B) typically have several tapered MAP-2 immunopositive dendrites.

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Neurosci. 14, 203-215 (1996).</li><li>Lein, P., Johnson, M., Guo, X., Rueger, 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=Osteogenic+protein-1+induces+dendritic+growth+in+rat+sympathetic+neurons.">Osteogenic protein-1 induces dendritic growth in rat sympathetic neurons.</a> Neuron. 15, 597-605 (1995).</li><li>Lein, P. J., 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=Laminin+and+a+basement+membrane+extract+have+different+effects+on+axonal+and+dendritic+outgrowth+from+embryonic+rat+sympathetic+neurons+in+vitro.">Laminin and a basement membrane extract have different effects on axonal and dendritic outgrowth from embryonic rat sympathetic neurons in vitro.</a> 136, 330-345 (1989).</li><li>Higgins, D., Lein, P. J., Osterhout, D. J., Johnson, M. 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The authors have nothing to disclose.

The advantages of this model include: (a) the cultures are comprised of a homogenous population of sympathetic neurons devoid of other cell types 17,23; (b) the growth factor requirements of sympathetic neurons are well-established, which allows the use of defined medium, and a variety of defined media, and substrates work well for growing and maintaining sympathetic neurons 23; (c) neurons in these cultures respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of th...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments </td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td>Bellco</td>
			<td>1943-10012</td>
			<td>12 mm Round Coverslip #1 Thickness, German glass</td>
		</tr>
		<tr>
			<td>Bent tip pipets</td>
			<td>Bellco</td>
			<td>1273-50003</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Poly-D-lysine</td>
			<td>Sigma</td>
			<td>P0899-100MG</td>
			<td>Stock solution 1 mg/ml</td>
		</tr>
		<tr>
			<td>Distilled water</td>
			<td>Gibco</td>
			<td>15230</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Fine forceps Dumont no. 4 and 5</td>
			<td>Fine Science Tools (FST)</td>
			<td>11252-30 11241-30</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>20-gauge needles</td>
			<td>Becton Dickinson</td>
			<td>305176</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Leibovitz's L-15 medium</td>
			<td>Invitrogen</td>
			<td>11415</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Collagenase</td>
			<td>Worthington</td>
			<td>4176</td>
			<td>1 mg/ml</td>
		</tr>
		<tr>
			<td>Dispase</td>
			<td>Roche</td>
			<td>04942078001</td>
			<td>5 mg/ml</td>
		</tr>
		<tr>
			<td>Hank's balanced salt solution( -Ca,-Mg)</td>
			<td>Invitrogen</td>
			<td>14185-052</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>DMEM Low glucose</td>
			<td>Invitrogen</td>
			<td>11885</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Invitrogen</td>
			<td>25030</td>
			<td>20 mM</td>
		</tr>
		<tr>
			<td>Insulin/Transferrin/ Selenium</td>
			<td>Invitrogen</td>
			<td>51500-056</td>
			<td>100X</td>
		</tr>
		<tr>
			<td>Fatty-acid free BSA</td>
			<td>Calbiochem</td>
			<td>126609</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>NGF</td>
			<td>Harlan Bioproducts</td>
			<td>5017</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Ham F-12 nutrient medium (F-12)</td>
			<td>Invitrogen</td>
			<td>11765</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cytosine arabinoside</td>
			<td>Sigma</td>
			<td>C1768</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Matrigel</td>
			<td>BD Biosciences</td>
			<td>356234</td>
			<td>Avoid repeated freeze-thaw cycles</td>
		</tr>
		<tr>
			<td>BMP-7</td>
			<td>R&amp;D Systems Curis, Inc.</td>
			<td>345-BP BMP7-07H</td>
			<td>BMPs 4, 5 and 6 work equally as well as BMP-7 and are sometimes easier to obtain commercially</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma</td>
			<td>P6148</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher</td>
			<td>BP151-500</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mouse anti-rat MAP2 antibody</td>
			<td>Sternberg Monoclonals Inc.</td>
			<td>SMI52</td>
			<td>1/5000</td>
		</tr>
		<tr>
			<td>Alexa Fluor 546 goat-anti-mouse</td>
			<td>Invitrogen</td>
			<td>A11003</td>
			<td>1/10000</td>
		</tr>
		<tr>
			<td>Mounting media</td>
			<td>Prolong Antifade kit Molecular Probes</td>
			<td>P7481</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>
Table 1. Materials.

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.

Watch this Scientific Journal Video about Inducing Dendritic Growth in Cultured Sympathetic Neurons at JoVE.com

Inducing Dendritic Growth in Cultured Sympathetic Neurons

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 ...

inducing-dendritic-growth-in-cultured-sympathetic-neurons

Research

JoVE Journal

Neuroscience

12.7K Views.  University of California, Davis. Мы опишем протокол для использования костных морфогенетических белков-7 (BMP-7) или Матригель выборочно стимулировать рост дендритов в первичных симпатических нейронов в отрыве от верхних шейных ганглиев (SCG) перинатальной крысы.

Video: Inducing Dendritic Growth in Cultured Sympathetic Neurons

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.