All procedures were approved by the University of Wisconsin Committee on Animal Care and were in accordance with NIH guidelines. We thank Dr. Katherine Kalil for the generous use of her Nucleofector device. We also thank members of the Dent lab for comments on the protocol. This work was supported by grants NIH R01-NS064014, Dana Foundation and Whitehall Foundation to E.W.D.
Christopher Viesselmann, Jason Ballweg and Derek Lumbard contributed equally to this paper.

1. 	Preparation of Coverslips and Chambers
<ol>
<li>Preparation of clean coverslips and chambers is essential for healthy cultures. Shortcuts should not be taken at any of these steps.</li>
<li>Wash the coverslips (12mm or 22mm round, German glass - Carolina Assistant Brand) overnight in concentrated nitric acid (HNO3) in a dedicated glass jar or beaker.</li>
<li>Remove the coverslips from nitric acid and wash extensively (5-7x) in deionized water. </li>
<li>Separate the coverslips and dry in a laminar flow hood or biosafety cabinet. When dry, sterilize them with UV light for 30 minutes. Place sterilized coverslips into a sterile petri dishes for storage. If plating neurons on 12mm coverslips directly place coverslips in a sterile 35mm dish and proceed to Section 2. </li>
<li>Imaging chambers are constructed by drilling a 15mm hole in the bottom of 35mm petri dishes (remove all burrs) and attaching the cleaned coverslips with a 3:1 mixture of paraffin and petroleum jelly.</li>
<li>Melt the paraffin/petroleum jelly mixture in a conical tube inside a boiling water bath. Use a small paint brush and coat the underside of the dish around the 15mm hole. Make sure to keep stirring the paraffin/petroleum jelly mixture as it will separate. This usually results in chambers glued together with a higher concentration of petroleum jelly, which will become viscous when dishes are placed in the incubator, resulting in coverslip detachment. The remaining paraffin/petroleum jelly can be stored at room temperature. </li>
<li>Place the dish upside down on a flat tray and place the coverslip over the hole. Heat in 80&deg;C oven until paraffin mixture is melted (~10 minutes). Remove dishes onto a flat surface and let the paraffin mixture set. </li>
<li>Turn the dishes over and sterilize both the insides of the covers and the bottoms of chambers with UV light.</li>
<li>Coat coverslips or glass regions of chambers with 1.0mg/mL Poly-d-lysine (30kDa) in borate buffer (0.1M sodium borate, pH 8.5) for one hour. Rinse 3-5 times with copious amounts of tissue culture grade deionized water. Make sure to remove all traces of borate buffer. Dry and use immediately or store chambers/coverslips for later use. We usually use cleaned coverslips within one month of preparation.</li>
</ol>
2. 	Preparation of Neuronal Dissection and Culture Medium
<ol>
<li>Prepare the dissection medium (DM) by adding appropriate amounts of 10x HBSS and 100X HEPES to tissue culture grade water. Store at 4&deg;C. Keep on ice during the dissection.</li>
<li>The day before the dissection prepare the plating medium (PM) and serum-free medium (SFM). PM consists of Neurobasal Medium, B27 supplement, 2 mM glutamine, 0.3% glucose, 37.5 mM NaCl and 5% fetal bovine serum (FBS). SFM consists of Neurobasal Medium, B27 supplement, 2 mM glutamine, 0.3% glucose and 37.5 mM NaCl.</li>
<li>Make only enough for the dissection and store in a tissue culture incubator overnight with the cap slightly ajar so that the temperature and CO2 content of the medium equilibrates. We add extra glucose and increase the osmolality to approximately 310mOsm with NaCl. We find the cultures do better at a more physiological osmolality (Neurobasal osmolality is typically 205-245mOsm).</li>
</ol>
3. 	Cortical Glial Feeder Layer Preparation for Long-term Cultures 
<ol>
<li>If long term cultures are to be prepared, perform this part of the protocol two to three weeks before continuing with the cortical or hippocampal dissections.</li>
<li>Prepare glial medium (GM) with MEM, 0.3% glucose, penicillin/streptomycin and 10% horse serum.</li>
<li>Euthanize P1-P3 mouse pups by cooling on ice for 5 minutes. Remove each pup from the ice and spray with 70% ethanol. Quickly decapitate with scissors. Remove the entire brain to a dish containing cold DM (step 2.1).</li>
<li>Remove the two cerebral hemispheres and meninges. Excise the neocortex and remove to a new dish containing no media. Prepare cortices from 4 brains total.</li>
<li>Mince the cortices with a clean, sterile razor blade as fine as possible and remove chopped tissue with a plastic pipette to a 50 mL conical tube containing 12 mL of cold DM. Add trypsin and DNase to final concentrations of 0.25% (1.5 mL) and 0.1% (1.5 mL), respectively. Incubate in a 37&deg;C water bath for 10 minutes with intermittent swirling.</li>
<li>Remove the tube containing the cortical tissue and clean thoroughly with 70% ethanol before bringing into the tissue culture hood. Pipet cortical tissue up and down with a 10 mL pipette approximately 10-15 times, or until most chunks disappear.</li>
<li>Return the tube to 37&deg;C in a water bath for another 10 minutes with intermittent swirling.</li>
<li>Thoroughly clean the tube with 70% ethanol and bring it back to the tissue culture hood. Pipet the cortical tissue up and down with a 5 mL pipette approximately 10-15 times, or until the chunks disappear.</li>
<li>Add 15 mL of warm GM and centrifuge at 200xg (1000rpm) for 10 minutes.</li>
<li>Discard supernatant, resuspend the pelleted cells in 20 mL of fresh GM and count with a hemocytometer. Plate 5-7.5x106 cells in 15 mL of GM per 75cm2 flask.</li>
<li>After one day and every 2-3 subsequent days in culture, dislodge loose cells by knocking the flask against your hand. Remove the medium along with any dislodged cells and replace with 15 mL of fresh GM.</li>
<li>Glia can be harvested after 1-2 weeks of growth in the flasks, when they are about 70-100% confluent. To prepare individual coverslips coated with glia, place 6 nitric acid cleaned and sterilized 25mm round coverslips in a 10cm dish, and place 3 dots of the 3:1 mixture of paraffin/petroleum jelly onto each coverslip in a triangular pattern with a small paint brush. Treat open dishes with UV light for 30 minutes. Coat the coverslips with 0.1 mg/ mL Poly-d-lysine (30kDa) in borate buffer for one hour, then wash extensively (3-5x) with sterile tissue culture grade deionized water and let dry.</li>
<li>Remove the glial-containing flask from incubator, discard the medium and rinse with 5 mL of pre-warmed trypsin/EDTA solution. Remove the trypsin/EDTA solution from the flask and pipette 3 mL of fresh pre-warmed trypsin/EDTA into the flask. Incubate the flask for 1 minute at 37&deg;C before adding 5 mL of GM to stop the trypsinization.</li>
<li>Remove the glia from flask by repeated pipetting 10-15 times, and then transfer the media to a 15 mL conical tube. Centrifuge at 200xg (1000rpm) for 8 minutes. Remove the supernatant and add 10 mL of GM, count cells, and plate 5x105 cells in 12.5 mL of GM per 10cm dish containing the coverslips.</li>
<li>Exchange the medium with fresh pre-warmed GM every 2-3 days. The day before the neuron dissection, remove the GM and replace with SFM (section 2.2). Use this glial-conditioned SFM in step 4.12 when flooding cortical or hippocampal cultures.</li>
</ol>
4. 	Cortical And/or Hippocampal Dissection and Electroporation
<ol>
<li>Remove appropriate amounts of nucleofection solutions (Lonza), combine, and warm to room temperature before starting the dissection. Since Nucleofection solution has a limited lifetime when combined, we only combine the needed amount for each preparation (100 &mu;L per transfection). </li>
<li>Euthanize a pregnant mouse at E15.5 with CO2 (day of plug is E0.5) and remove the uterus to a 10cm petri dish. Remove fetuses and decapitate into cold DM (section 2.1).</li>
<li>Remove the entire brain into a separate dish of cold DM and with a bent tungsten needle, remove both neocortices. Remove the meninges with microforceps and place cortices into new dish of cold DM. With a small iris or Wecker scissors, remove the cortex or hippocampus and place in a 1.5 mL Eppendorf tube filled with 1.0 mL of cold DM. Keep this tube on ice.</li>
<li>After dissecting all cortices or hippocampi, add 110 &mu;L of 2.5% trypsin to the Eppendorf tube containing the tissue and place tube in a 37&deg;C incubator for 20 minutes.</li>
<li>Remove the supernatant and wash cortices or hippocampi with 1.0 mL PM (section 2.2) by gently inverting the Eppendorf tube. Repeat the wash twice, leaving 1 mL of PM in the tube. </li>
<li>Triturate the chunks 15 times with a P1000 pipette, and remove supernatant/cells to a new 15 mL conical tube containing 4 mL of PM, leaving any chunks that remain in Eppendorf tube.</li>
<li>Spin the 15 mL tube at 20xg (350rpm) for 7 minutes with the brake off. Discard the supernatant and add 100 &mu;L of premixed, room temperature nucleofection solution (Lonza) for each transfection. Triturate 5 times with a gentle up and down motion of the P1000 pipette.</li>
<li>Remove 100 &mu;L of the nucleofection solution/cell mixture to each new Eppendorf tube and add the appropriate amount of DNA. For long term cultures we generally use 1-2&mu;g of DNA per transfection. However, this only labels a small &lt;10% proportion of neurons in the culture. We generally use 5-10&mu;g of DNA per transfection if want higher transfection efficiency for short term culture. We have used up to a total of 40&mu;g of DNA when transfecting with two different plasmids. Plasmids are stored in TE buffer at 1&mu;g/ &mu;L. </li>
<li>Add cell suspension/DNA to the cuvette (Lonza) and electroporate the cells in the Nucleofector (Lonza), using program O-005 (Mouse CNS neurons).</li>
<li>Working quickly, add 500 &mu;L of pre-warmed and equilibrated PM to the cuvette and remove solution/cells to a new 1.5 mL Eppendorf tube. Add enough PM to bring the volume of each transfection to 1.0 mL. Count the cells with a hemocytometer and plate at 3-5x103 cells/cm2 for young cultures, or 5-10x103 cells/cm2 for long-term cultures.</li>
<li>For short-term cultures, flood the 35mm culture dishes with 2.0 mL of warmed, CO2-equilibrated SFM after one hour of plating. If using coverslips, we remove one-half of the PM and replace it with SFM, then repeat two more times. Either flooding the imaging chambers or washing the coverslips results in a very low serum content (&lt;0.5%). Short-term cultures do not need to be cultured with a glial feeder layer and do not need to be re-fed. </li>
<li>For long-term cultures, we remove a glia covered coverslip containing three dots of paraffin/petroleum jelly and invert it over the 15mm hole in the 35mm dish, one hour after initial plating. Two milliliters of the conditioned SFM from the glial dish is then added to the imaging chamber. To feed the long-term cultures, we remove one-third of the SFM every 2-3 days and replace it with fresh, pre-warmed and CO2-equilibrated SFM. </li>
</ol> 
5. Representative Results:
<img src="/files/ftp_upload/2373/2373fig1.jpg" alt="Figure 1" /> 
Figure 1. Living hippocampal neurons in successive stages of development. Paired images of representative living hippocampal neurons are shown as both a differential interference contrast image and a corresponding fluorescent micrograph. Each of these cells has been transfected with EGFP-Tubulin and DsRed2 in pCAX vectors. The neurons were imaged at the following days in vitro (DIV): Stage 1(1DIV), Stage 2 (1DIV), Stage 3 (2DIV), Stage 4 (11DIV) and Stage 5 (32DIV). Scale bar is 20&mu;m.

<ol>
	<li>Goslin, K., Asmussen, H., Banker, G., Goslin, K., Banker, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+13.">Chapter 13.</a> Culturing Nerve Cells. , 339-370 (1998).</li><li>Kaech, S., Banker, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+hippocampal+neurons.">Culturing hippocampal neurons.</a> Nat Protoc. 1, 2406-2415 (2006).</li><li>Dent, E. W., Callaway, J. L., Szebenyi, G., Baas, P. W., Kalil, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reorganization+and+movement+of+microtubules+in+axonal+growth+cones+and+developing+interstitial+branches.">Reorganization and movement of microtubules in axonal growth cones and developing interstitial branches.</a> J Neurosci. 19, 8894-8908 (1999).</li><li>Dent, E. W., Kalil, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+imaging+of+neuronal+cytoskeleton.">Dynamic imaging of neuronal cytoskeleton.</a> Methods Enzymol. 361, 390-407 (2003).</li><li>Dent, E. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Filopodia+are+required+for+cortical+neurite+initiation.">Filopodia are required for cortical neurite initiation.</a> Nat Cell Biol. 9, 1347-1359 (2007).</li><li>Hu, X., Viesselmann, C., Nam, S., Merriam, E., Dent, E. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activity-dependent+dynamic+microtubule+invasion+of+dendritic+spines.">Activity-dependent dynamic microtubule invasion of dendritic spines.</a> J Neurosci. 28, 13094-13105 (2008).</li><li>Lebrand, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Critical+role+of+Ena/VASP+proteins+for+filopodia+formation+in+neurons+and+in+function+downstream+of+netrin-1.">Critical role of Ena/VASP proteins for filopodia formation in neurons and in function downstream of netrin-1.</a> Neuron. 42, 37-49 (2004).</li><li>Meberg, P. J., Miller, M. W., Hollenbeck, P. J., Bamburg, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+7.">Chapter 7.</a> Neurons: Methods and Applications for the Cell Biologist. , 112-129 (2003).</li><li>Osumi, N., Inoue, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+transfer+into+cultured+mammalian+embryos+by+electroporation.">Gene transfer into cultured mammalian embryos by electroporation.</a> Methods. 24, 35-42 (2001).</li><li>Zeitelhofer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-efficiency+transfection+of+mammalian+neurons+via+nucleofection.">High-efficiency transfection of mammalian neurons via nucleofection.</a> Nat Protoc. 2, 1692-1704 (2007).</li></ol>

No conflicts of interest declared.

This protocol for culturing embryonic hippocampal and cortical mouse neurons was developed as a modification of the Banker protocol, which uses rat neurons1,2. We have used this protocol for culturing mouse and hamster neurons as well3,4,5,6,7. This protocol works equally well for both hippocampal and neocortical neurons and is similar to a protocol published by Meberg and Miller8. Generally, we use hippocampal neurons for long-term culture because they are well characterized and a more established model system. Furthermore, they are likely to contain a more homogeneous population of neurons than neocortex. However, neocortical neurons cultured using this protocol also survive and differentiate similarly (unpublished data). We routinely use hippocampal and neocortical neurons for short term culture. Dissection of neocortex also results in substantially more neurons (1.5x106 neurons per pair of cortices) than hippocampal dissection (2.5x105 neurons per pair of hippocampi), which makes it a better choice of material for Western blotting, for example.
As with any primary culture, it is essential to minimize the time that it takes from the death of the animal to the plating of the cells. It will generally take 10-20 dissections to become consistently fast at dissection and plating. Also, when working with the Lonza Nucleofector, it is critical to work quickly during the electroporation procedure, as the viability of the neurons decreases rapidly if they are left in the nucleofection buffer.
Much of our imaging is conducted with total internal reflectance fluorescence microscopy (TIRFM). This type of microscopy is only capable of imaging several hundred nanometers beyond the coverslip. Therefore, the areas of the neurons that we frequently image, the axonal growth cone and dendritic spines, need to be adhered directly to the coverslip. Thus, we use low density cultures that require glial feeding for long-term culture. We have used higher density cultures (>2x104 cells/cm2), without glial feeder layers, for long-term cultures and found that they survive very well with little feeding. However, the dendritic spines of these neurons are oftentimes too far away from the substrate to image in TIRFM, although they can be readily detected with wide-field microscopy or confocal microscopy.
In most of our studies we transfect neurons before plating, and have imaged fluorescently-labeled proteins for up to three months in culture. This long-term expression of fluorescently-labeled proteins gives us confidence that by using low concentrations of DNA (1-2&mu;g) we are not producing overexpression artifacts in the neurons. However, this procedure can also be used to study overexpression of proteins if high amounts of DNA are used (10-20&mu;g). The plasmids that we use to transfect neurons usually contain EGFP or mCherry fusion proteins, although we also label the neuronal cytoplasm with DsRed2 or EGFP alone. This electroporation technique works well with a number of vectors. We prefer plasmids that contain a &beta;-actin promoter with a CMV enhancer and &beta;-globin poly-A tail (pCAGGs or pCAX plasmids)9, due to the relatively high levels of expression, and the fact that they are well tolerated by the neurons in both short- and long-term culture. Generally, proteins begin to express within about 4 hours of plating and reach levels sufficient for imaging within 10-24hrs10. We have successfully used CMV-promoter-driven plasmids in short term cultures, but have found that they can cause high levels of overexpression that kill neurons in long-term culture. Nevertheless, we have found that the glial conditioning of low density cultures helps with the survival of neurons transfected with CMV-promoter driven plasmids, compared to higher density (non-glial fed) cultures.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Sodium Borate</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>S93356</td>
<td>Store borate buffer at room temperature (RT).</td>
</tr>
<tr>
<td>Poly-d-Lysine</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P7886-100mg</td>
<td>Dissolve in 0.1M borate buffer. Sterile filter and store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>Trypsin (10x)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15090-046</td>
<td>Store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>DNase (10x)</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>DN25-1G</td>
<td>Store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>MEM</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11095-080</td>
<td>Store at 4&deg;C.</td>
</tr>
<tr>
<td>HBSS (10x)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>14185-052</td>
<td>Store at RT.</td>
</tr>
<tr>
<td>HEPES (100x)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15630-080</td>
<td>Store at 4&deg;C.</td>
</tr>
<tr>
<td>Penicillin/Streptomycin (100x)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15140-122</td>
<td>Store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>Horse Serum</td>
<td>&nbsp;</td>
<td>Hyclone</td>
<td>SH3007403</td>
<td>Store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>Trypsin/EDTA</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>25200-056</td>
<td>Store at 4&deg;C.</td>
</tr>
<tr>
<td>Neurobasal E (NB)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>21103-049</td>
<td>Store at 4&deg;C.</td>
</tr>
<tr>
<td>B27 Supplement (50x)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>17504-044</td>
<td>Store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>Glutamine (100x)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>25030-081</td>
<td>Store aliquots at -80&deg;C</td>
</tr>
<tr>
<td>30% Glucose in NB (100X)</td>
<td>&nbsp;</td>
<td>Fisher/Invitrogen</td>
<td>BP350-500</td>
<td>Dissolve glucose in NB and sterile filter. Store at 4&deg;C.</td>
</tr>
<tr>
<td>3.75M NaCl in NB (100X)</td>
<td>&nbsp;</td>
<td>Fisher/Invitrogen</td>
<td>BP358-1</td>
<td>Dissolve NaCl in NB and sterile filter. Store at 4&deg;C.</td>
</tr>
<tr>
<td>Fetal Bovine Serum (FBS)</td>
<td>&nbsp;</td>
<td>Hyclone</td>
<td>SH3007003</td>
<td>Store aliquots at -80&deg;C.</td>
</tr>
<tr>
<td>Paraffin/Petroleum Jelly (3:1 w/w)</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>Paraffin &#x2013; P31-500 Petroleum &#x2013; P66-1</td>
<td>Combine in conical tube and melt in boiling water.</td>
</tr>
<tr>
<td>Concentrated Nitric Acid (HNO3)</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>A509-212</td>
<td>Store at room temperature. Can be reused several times. Discard if it yellows.</td>
</tr>
<tr>
<td>5mM cytosine &#x2013;B-D-arabinofuranoside hydrochoride (AraC) in NB (1000X)</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>C6645</td>
<td>Dissolve AraC in NB and sterile filter. Store at -80&deg;C</td>
</tr>		</tbody>
		</table>
		</div>
			<div>
			*Most reagents that we store at -80&deg;C can be stored at -20&deg;C as well. Storing them at -80&deg;C lengthens their shelf life and results in slightly more consistent cultures.		</div>

nucleofection and primary culture of embryonic mouse hippocampal and cortical neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

Watch this Scientific Journal Video about Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons at JoVE.com

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

nucleofection-primary-culture-embryonic-mouse-hippocampal-cortical

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JoVE Journal

Neuroscience

33.3K Views.  University of Wisconsin-Madison. Questo protocollo delinea i passaggi necessari per analizzare, attraverso trasfezione elettroporazione e neuroni dell'ippocampo e del mouse cultura corticale. Colture a breve termine possono essere utilizzati per gli studi della crescita degli assoni e la guida, mentre a lungo termine le culture possono essere utilizzati per studi di sinaptogenesi e analisi spine dendritiche.

Video: Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

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 ...

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

In vitro study of primary neuronal cultures allows for quantitative analyses of neurite outgrowth. In order to study how genetic alterations affect neuronal process outgrowth, shRNA or cDNA constructs can be introduced into primary neurons via chemical transfection or viral transduction. However, with primary cortical cells, a heterogeneous pool of cell types (glutamatergic neurons from different layers, inhibitory neurons, glial cells) are transfected using these methods. The use of in utero electroporation to introduce DNA constructs in the embryonic rodent cortex allows for certain subsets of cells to be targeted: while electroporation of early embryonic cortex targets deep layers of the cortex, electroporation at late embryonic timepoints targets more superficial layers. Further, differential placement of electrodes across the heads of individual embryos results in the targeting of dorsal-medial versus ventral-lateral regions of the cortex. Following electroporation, transfected cells can be dissected out, dissociated, and plated in vitro for quantitative analysis of neurite outgrowth. Here, we provide a step-by-step method to quantitatively measure neuronal process outgrowth in subsets of cortical cells.
The basic protocol for in utero electroporation has been described in detail in two other JoVE articles from the Kriegstein lab 1, 2. We will provide an overview of our protocol for in utero electroporation, focusing on the most important details, followed by a description of our protocol that applies in utero electroporation to the study of gene function in neuronal process outgrowth.

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

In utero electroporation is a valuable method for transfecting neuronal progenitor cells in vivo. Depending upon the placement of the electrodes and the developmental timepoint of electroporation, certain subsets of cortical cells can be targeted. Targeted cells can then be analyzed in vivo or in vitro for effects of genetic alteration.

In vitro study of primary neuronal cultures allows for quantitative analyses of neurite outgrowth. In order to study how genetic alterations affect ...

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

Isolation and Culture of Hippocampal Neurons from Prenatal Mice

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual neurons, researchers are able to analyze properties related to cellular trafficking, cellular structure and individual protein localization using a variety of biochemical techniques. Results from such experiments are critical for testing theories addressing the neural basis of memory and learning. However, unambiguous results from these forms of experiments are predicated on the ability to grow neuronal cultures with minimum contamination by other brain cell types. In this protocol, we use specific media designed for neuron growth and careful dissection of embryonic hippocampal tissue to optimize growth of healthy neurons while minimizing contaminating cell types (i.e. astrocytes). Embryonic mouse hippocampal tissue can be more difficult to isolate than similar rodent tissue due to the size of the sample for dissection. We show detailed dissection techniques of hippocampus from embryonic day 19 (E19) mouse pups. Once hippocampal tissue is isolated, gentle dissociation of neuronal cells is achieved with a dilute concentration of trypsin and mechanical disruption designed to separate cells from connective tissue while providing minimum damage to individual cells. A detailed description of how to prepare pipettes to be used in the disruption is included. Optimal plating densities are provided for immuno-fluorescence protocols to maximize successful cell culture. The protocol provides a fast (approximately 2 hr) and efficient technique for the culture of neuronal cells from mouse hippocampal tissue.

We provide a protocol for the culture of highly purified hippocampal neurons from prenatal mouse brains without the use of a feeder glial cell layer.

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual ...

Isolation and Culture of Mouse Cortical Astrocytes

Astrocytes are an abundant cell type in the mammalian brain, yet much remains to be learned about their molecular and functional characteristics. In vitro astrocyte cell culture systems can be used to study the biological functions of these glial cells in detail. This video protocol shows how to obtain pure astrocytes by isolation and culture of mixed cortical cells of mouse pups. The method is based on the absence of viable neurons and the separation of astrocytes, oligodendrocytes and microglia, the three main glial cell populations of the central nervous system, in culture. Representative images during the first days of culture demonstrate the presence of a mixed cell population and indicate the timepoint, when astrocytes become confluent and should be separated from microglia and oligodendrocytes. Moreover, we demonstrate purity and astrocytic morphology of cultured astrocytes using immunocytochemical stainings for well established and newly described astrocyte markers. This culture system can be easily used to obtain pure mouse astrocytes and astrocyte-conditioned medium for studying various aspects of astrocyte biology.

Astrocytes have been recognized to be versatile cells participating in fundamental biological processes that are essential for normal brain development and function, and central nervous system repair. Here we present a rapid procedure to obtain pure mouse astrocyte cultures to study the biology of this major class of central nervous system cells.

Astrocytes  are an abundant cell type in the mammalian brain, yet much remains to be  learned about their molecular and functional characteristics. In ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Primary Culture of Mouse Dopaminergic Neurons

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions such as motor control, motivation, and working memory. Nigrostriatal dopaminergic neurons selectively degenerate in Parkinson&#39;s disease (PD). This progressive neuronal loss is unequivocally associated with the motors symptoms of the pathology (bradykinesia, resting tremor, and muscular rigidity). The main agent responsible of dopaminergic neuron degeneration is still unknown. However, these neurons appear to be extremely vulnerable in diverse conditions. Primary cultures constitute one of the most relevant models to investigate properties and characteristics of dopaminergic neurons. These cultures can be submitted to various stress agents that mimic PD pathology and to neuroprotective compounds in order to stop or slow down neuronal degeneration. The numerous transgenic mouse models of PD that have been generated during the last decade further increased the interest of researchers for dopaminergic neuron cultures. Here, the video protocol focuses on the delicate dissection of embryonic mouse brains. Precise excision of ventral mesencephalon is crucial to obtain neuronal cultures sufficiently rich in dopaminergic cells to allow subsequent studies. This protocol can be realized with embryonic transgenic mice and is suitable for immunofluorescence staining, quantitative PCR, second messenger quantification, or neuronal death/survival assessment.

Dopaminergic neurons play a vital regulatory role in the brain. Their loss is associated with Parkinson's disease. In this video, we show how to generate primary cultures of central dopaminergic neurons from embryonic mouse mesencephalon. Such cultures are useful to study the extreme vulnerability of these neurons to various stresses.

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions ...

Isolation, Culture and Long-Term Maintenance of Primary Mesencephalic Dopaminergic Neurons From Embryonic Rodent Brains

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes involved in physiological functions and vulnerability and death of these neurons is imparative to understanding the underlying causes and unraveling the cure for this common neurodegenerative disorder. Primary cultures of mesDA neurons provide a tool for investigation of the molecular, biochemical and electrophysiological properties, in order to understand the development, long-term survival and degeneration of these neurons during the course of disease. Here we present a detailed method for the isolation, culturing and maintenance of midbrain dopaminergic neurons from E12.5 mouse (or E14.5 rat) embryos. Optimized cell culture conditions in this protocol result in presence of axonal and dendritic projections, synaptic connections and other neuronal morphological properties, which make the cultures suitable for study of the physiological, cell biological and molecular characteristics of this neuronal population.

The causes of degeneration of midbrain dopaminergic neurons during Parkinson&#8217;s disease are not fully understood. Cellular culture systems provide an essential tool for study of the neurophysiological properties of these neurons. Here we present an optimized protocol, which can be utilized for in vitro modeling of neurodegeneration.

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes ...

Low-Density Primary Hippocampal Neuron Culture

The ability to probe the structure and physiology of individual nerve cells in culture is crucial to the study of neurobiology, and allows for flexibility in genetic and chemical manipulation of individual cells or defined networks. Such ease of manipulation is simpler in the reduced culture system when compared to the intact brain tissue. While many methods for the isolation and growth of these primary neurons exist, each has its own limitations. This protocol describes a method for culturing low-density and high-purity rodent embryonic hippocampal neurons on glass coverslips, which are then suspended over a monolayer of glial cells. This &#39;sandwich culture&#39; allows for exclusive long-term growth of a population of neurons while allowing for trophic support from the underlying glial monolayer. When neurons are of sufficient age or maturity level, the neuron coverslips can be flipped-out of the glial dish and used in imaging or functional assays. Neurons grown by this method typically survive for several weeks and develop extensive arbors, synaptic connections, and network properties.

This article describes the protocol for culturing low-density primary hippocampal neurons growing on glass coverslips inverted over a glial monolayer. The neuron and glial layers are separated by paraffin wax beads. The neurons grown by this method are suitable for high-resolution optical imaging and functional assays.

The ability to probe the structure and physiology of individual nerve cells in culture is crucial to the study of neurobiology, and allows for flexibility ...

Generation of Neurospheres from Mixed Primary Hippocampal and Cortical Neurons Isolated from E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have a robust in vitro model that can be exploited for various neurobiology studies. Though primary neuron cultures (i.e., long-term hippocampal cultures) have provided scientists with models, it does not yet represent the complexity of brain network completely. In the wake of these limitations, a new model has emerged using neurospheres, which bears a closer resemblance to the brain tissue. The present protocol describes the plating of high and low densities of mixed cortical and hippocampal neurons isolated from the embryo of embryonic day 14-16 Sprague Dawley rats. This allows for the generation of neurospheres and long-term primary neuron culture as two independent platforms to conduct further studies. This process is extremely simple and cost-effective, as it minimizes several steps and reagents previously deemed essential for neuron culture. This is a robust protocol with minimal requirements that can be performed with achievable results and further used for a diversity of studies related to neuroscience.

Presented here is a protocol for the spontaneous generation of neurospheres enriched in neural progenitor cells from high density plated neurons. During the same experiment, when neurons are plated at a lower density, the protocol also results in prolonged primary rat neuron cultures.

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have ...