This work was funded by NIH grants DA013137, DA031604, and HD043680. Partial support was provided by a NIH T32 training grant in Biomedical-Behavioral science.

All animal protocols were reviewed and approved by the Animal Care and Use Committee at the University of South Carolina (assurance number: A3049-01).
1. Low-density Embryonic Neuronal Culture
<ol>
	<li>Preparation for primary cortical neuron culture
	<ol>
		<li>Solutions:
		<ol>
			<li>Prepare Poly-L-Lysine stock solution by dissolving 5 mg of Poly-L-Lysine in 10 ml Borate buffer.</li>
			<li>Prepare working solution by diluting 1 ml Poly-L-Lysine Stock in 49 ml Borate Buffer.</li>
			<li>Prepare borate Buffer, pH 8.4 by starting with 395 ml dH2O and then adding all ingredients (1.24 g boric acid + 1.9 g of Borax). Adjust pH to 8.4 by 1 M NaOH. Adjust to 400 ml and filter with 0.2 &#181;m nylon membrane filter under hood.</li>
			<li>Prepare HBSS solution, pH 7.2 by starting with 445 ml dH2O and adding all other ingredients (50 ml 10X HBSS Stock + 1.2 g HEPES). Adjust pH to 7.2 by 1 N HCl, bring to 500 ml and filter with 0.2 &#181;m nylon membrane filter under hood.</li>
			<li>Prepare plating medium: DMEM/F12 with 10% fetal bovine serum. Filter with 0.2 &#181;m nylon membrane filter under hood.</li>
			<li>Prepare complete growth medium: To 50 ml Neurobasal medium, add the supplements (500 &#181;l GlutaMax (100X) + 500 &#181;l Glucose + 1 ml B-27 (50X) + 500 &#181;l of Antibiotic-Antimycotic solution (100X) + 100 &#181;l 7.5% Sodium Bicarbonate. Filter with 0.2 &#181;m nylon membrane filter under hood.</li>
		</ol>
		</li>
		<li>Two days before culture:
		<ol>
			<li>Coat twelve 35 mm glass-bottom dishes with 2 ml of Poly-L-Lysine working solution, and keep under the hood O/N.</li>
		</ol>
		</li>
		<li>One day before culture:
		<ol>
			<li>Empty coating agent from the dishes and rinse with ddH2O. Allow dishes to dry for an hour under hood.</li>
			<li>Prepare plating medium (DMEM/F12 with 10% fetal bovine serum (10% of total solution). Pipette 2 ml medium into each dish, and incubate O/N at 37 oC, 5% CO2. 
			Note: It is important to avoid plastic culture dishes when culturing. Use of glass-bottom dishes (for inverted microscopes) or glass coverslips (for upright microscopes) yields sharp and clear images. Compared to a plastic-bottom dish, the glass-bottom dish avoids unwanted hues or glare during fluorescent microscopy.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Cell culture protocol:
	<ol>
		<li>Put refrigerated HBSS Buffer (100-150 ml), 100 mm petri dishes, forceps, scissors, 50 ml tubes, and 15 ml tubes under UV light in hood.</li>
		<li>Euthanize pregnant rat with lethal inhalation of 5% sevoflurane under the hood outside of culture room.</li>
		<li>Sterilize with 70% EtOH, tent the skin of the lower abdomen using forceps while cutting from the tail up in the middle of abdominal cavity.Open the uterus using scissors to expose the placenta.</li>
		<li>Remove 8-10 E18 fetuses. Place 8-10 fetuses in petri dishes containing HBSS solution.</li>
		<li>Hold the back of the head of the fetus with forceps, then use scissors to sever the head from body. Place it in another petri dish filled with 5-7 ml of HBSS. Transfer dish to hood.</li>
		<li>Fill two additional 100 mm petri dish with cold HBSS.</li>
		<li>Peel aside skull, and scoop brain into a new petri dish filled with HBSS.</li>
		<li>Use sharp curved forceps, separate cerebellum and brain stem from brain, then transfer brain to a new petri dish with cold HBSS in it.</li>
		<li>Use tweezers to secure, separate the hemispheres with curved forceps. Remove meninges; isolate frontal cortex and transfer pieces into marked 15 ml centrifuge tubes.</li>
		<li>Fill the 15 ml tube with fresh 2 ml HBSS and add 20 &#181;l Trypsin EDTA (10x concentrated mixture with 0.5% Trypsin and 5.3 mM EDTA.) to each 15 ml tube.</li>
		<li>Incubate 10-15 min at RT. Gently swirl tube every few minutes, do not allow settling.</li>
		<li>After 10-15 min, use glass pipette to remove old HBSS and rinse twice with new HBSS.</li>
		<li>Immerse in trypsin inhibitor (1 mg/ml, 2 mg Trypsin inhibitor in 2 ml HBSS), mix well and let it sit 5 min. Wash twice with fresh HBSS.</li>
		<li>Using glass pipette with rubber bulb, triturate brain pieces 10-15 times very slowly so as to avoid bubbles.Triturate again using a calibrated pipette and a sterile tip with a reduced tip diameter very slowly until homogeneous.</li>
		<li>Transfer dissociated cells at desired density to pre-prepared coated culture dishes (50 cells/mm2). Incubate O/N at 37 oC, 5% CO2.</li>
		<li>After 24 hr, replace DMEM/F12 medium with freshly made complete growth serum-free Neurobasal medium.</li>
		<li>Maintain the cells at all times in a 5% CO2/95% room air humidified incubator for at least 12-14 days before experimentation. Supplement the cells every 5-6 days with fresh Neurobasal medium replacing approximately 50% of the old medium.</li>
	</ol>
	</li>
</ol>
2. Fluorescent Labeling and Immunocytochemistry
Note: The immunofluorescent labeling of primary cortical cell cultures was carried out in glass bottom 35 mm cell culture dishes with a working volume of 1 ml.
<ol>
	<li>Wash the glass-bottom dish twice with phosphate-buffered saline pH 7.4 (PBS).</li>
	<li>Fix cells with 4% paraformaldehyde for 15 min at RT.</li>
	<li>Wash cells twice with PBS and permeabilize with 0.1% Triton X-100 in PBS for 5 min.</li>
	<li>Treat cells for 20 min at RT with an F-actin specific stain, AlexaFluor 488 Phalloidin (1:40, 25 &#181;l of Phalloidin in 1 ml PBS).</li>
	<li>Rinse cells twice with PBS and block with 10% normal goat serum at RT for 1-2 hr.</li>
	<li>Dilute the chicken polyclonal anti-MAP2 antibody (1:2,500) in PBS with 2% normal goat serum and incubate O/N at 4 oC.</li>
	<li>Following incubation, rinse in PBS twice.</li>
	<li>Dilute the secondary antibody (Alexa Red 594-conjugated goat anti-chicken IgG (1:500) with 2% normal goat serum and incubate for 2 hr at RT.</li>
	<li>Rinse with PBS and add 10 &#181;l/dish of Hoescht dye.</li>
	<li>Incubate 3 min at RT and wash with PBS twice. 
	Note: The labeled cells now ready for step 3 (F-actin puncta counting).</li>
	<li>Preserve cell sample with 100 &#181;l of antifade reagent as a coverslipping agent and keep at 4 oC in the dark for long term storage. 
	Note: Phalloidin is relatively stable in PBS at +4 &#176;C in the dark for up to 1 week and fluorescence can be preserved with antifade coverslipping reagents. However, optimal images are obtained from 1-3 days after staining since Phalloidin may start to diffuse out with extended storage.</li>
</ol>
3. F-actin Puncta Counting
<ol>
	<li>Turn on fluorescent microscope and switch to 20&#215; objective. Open microscope software, and set up program at 1280&#215;960 pixel image size, and 0.17 &#181;m/pixel image resolution at 1&#215; zoom.</li>
	<li>Acquire images of co-labeled F-actin/MAP2 neurons under green (495 nm)/red (613 nm) fluorescent channels.</li>
	<li>Choose 5 Green (F-actin)/Red (MAP2) immunolabeled/Blue (Hoescht) fluorescent images of individual neuron with clearly defined dendritic arbors.</li>
	<li>Identify the F-actin rich structures in second order dendritic segment (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence.</li>
	<li>Rotate the selected region of images to a horizontal level. Copy and paste as a new image.</li>
	<li>Subtract the background of images, using a consistent adjustments for each dish.</li>
	<li>Count the bright green F-actin puncta and trace the length of selected dendritic segment manually by trained independent observers. Export the data to a spreadsheet file.</li>
	<li>Calculate the density by dividing total F-actin labeled puncta (N) by the length (L) of the MAP2 labeled dendrites. Express data as number of F-actin puncta per 10 &#181;m of dendrite 
	Note: Puncta (size &#8804;1.5 &#181;m) of F-actin fluorescence with a peak intensity of at least 50% above the average intensity of staining in the dendritic shaft were included in each selected dendritic segment.</li>
</ol>

<ol>
	<li>Heller, E. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+biochemical+anatomy+of+cortical+inhibitory+synapses.">The biochemical anatomy of cortical inhibitory synapses.</a> PLoS One. 7, e39572 (2012).</li><li>Craig, A. M., Blackstone, C. D., Huganir, R. L., 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=Selective+clustering+of+glutamate+and+gamma-aminobutyric+acid+receptors+opposite+terminals+releasing+the+corresponding+neurotransmitters.">Selective clustering of glutamate and gamma-aminobutyric acid receptors opposite terminals releasing the corresponding neurotransmitters.</a> Proc Natl Acad Sci USA. 91, 12373-12377 (1994).</li><li>Johnson, O. L., Ouimet, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+regulatory+role+for+actin+in+dendritic+spine+proliferation.">A regulatory role for actin in dendritic spine proliferation.</a> Brain Res. 1113, 1-9 (2006).</li><li>Rao, A., Craig, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signaling+between+the+actin+cytoskeleton+and+the+postsynaptic+density+of+dendritic+spines.">Signaling between the actin cytoskeleton and the postsynaptic density of dendritic spines.</a> Hippocampus. 10 (5), 527-541 (2000).</li><li>Matus, A., Ackermann, M., Pehling, G., Byers, H. R., Fujiwara, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+actin+concentrations+in+brain+dendritic+spines+and+postsynaptic+densities.">High actin concentrations in brain dendritic spines and postsynaptic densities.</a> Proc Natl Acad Sci USA. 79, 7590-7594 (1982).</li><li>Allison, D. W., Gelfand, V. I., Spector, I., Craig, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+actin+in+anchoring+postsynaptic+receptors+in+cultured+hippocampal+neurons:+differential+attachment+of+NMDA+versus+AMPA+receptors.">Role of actin in anchoring postsynaptic receptors in cultured hippocampal neurons: differential attachment of NMDA versus AMPA receptors.</a> J Neurosci. 18, 2423-2436 (1998).</li><li>Sekino, Y., Kojima, N., Shirao, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+actin+cytoskeleton+in+dendritic+spine+morphogenesis.">Role of actin cytoskeleton in dendritic spine morphogenesis.</a> Neurochem Int. 51, 92-104 (2007).</li><li>Zhang, W., Benson, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+synapse+development+defined+by+dependence+on+F-actin.">Stages of synapse development defined by dependence on F-actin.</a> J Neurosci. 21 (14), 5169-5181 (2001).</li><li>Bertrand, S. J., Aksenova, M. V., Mactutus, C. F., Booze, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=HIV-1+Tat+protein+variants:+Critical+role+for+the+cysteine+region+in+synaptodendritic+injury.">HIV-1 Tat protein variants: Critical role for the cysteine region in synaptodendritic injury.</a> Exp Neurol. 248, 228-235 (2013).</li><li>Bertrand, S. J., Mactutus, C. F., Aksenova, M. V., Espensen-Sturges, T. D., Booze, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptodendritic+recovery+following+HIV+Tat+exposure:+neurorestoration+by+phytoestrogens.">Synaptodendritic recovery following HIV Tat exposure: neurorestoration by phytoestrogens.</a> J Neurochem. 128 (1), 140-151 (2014).</li><li>Lau, P. M., Zucker, R. S., Bentley, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+filopodia+by+direct+local+elevation+of+intracellular+calcium+ion+concentration.">Induction of filopodia by direct local elevation of intracellular calcium ion concentration.</a> J Cell Biol. 145, 1265-1275 (1999).</li></ol>

None of the authors have conflicts of interest to declare.

In this protocol, we describe culturing rat cortical neurons at low density in 35 mm glass-bottom dishes which allows us to identify dendrites of individual neurons. Next, we use Phalloidin and MAP2 staining to detect dendritic changes. Then, we used specialized software to quantify changes in F-actin puncta.
To determine changes in F-actin puncta the entire neuronal network of an individual neuron must be clearly visible, this allows selection of appropriate second order dendritic segments from a single neuron. Low-density plating is critical in order to both visualize individual neurons as well as to minimize the presence of glial cells. Astrocytes also contain F-actin and without a neuronal marker, could confound the analysis of F-actin positive puncta. Low-density cultures and the use of Neurobasal medium are helpful in decreasing astrocytic proliferation in the cell cultures. However, given the caveat that the F-actin protein is not specific to neurons, specific neuronal protein markers, such as MAP2, should be used together with Phalloidin. Other protein antibodies may also be used in conjunction with Phalloidin, such as TH (tyrosine hydroxylase) for identification of specific neuronal populations in culture.
One general technique for studying synapses and synaptic morphology is immunocytochemistry (ICC). ICC is popular since many antibodies for synaptic proteins are readily available including PSD 95, NMDAR (post-synaptic) as well as synaptophysin, bassoon and synapsin I (pre-synaptic)11. However, some structures cannot be detected by ICC (such as thin filopodia), but a combination of F-actin Phalloidin labeling with antibody detection (or double labeling with two different antibodies) may allow for specific and complex investigation of synaptic subpopulations and differential responses to experimental treatments.
F-actin puncta may be particularly sensitive to experimental treatments. We recently reported that treatment with HIV-1 Tat protein produced a decrease in F-actin puncta (24 hr) prior to any evidence for overt neuronal death (48 hr)10. It is of note that experimental treatments may either increase (memantine) or decrease (methamphetamine+HIV-1 Tat) F-actin puncta. We also found that F-actin puncta loss is a reversible response following specific experimental treatments10. As such, F-actin puncta quantification is a valuable tool to monitor not only acute synaptopathy, but also for studying synaptic recovery and experimental neurorestoration processes.

The primary goal of this study is to develop a reliable method of measurement (estimation) of synaptic integrity of the neuronal dendritic network. Here we describe quantification of F-actin puncta in primary rat cultured neurons using a combination of Phalloidin staining and immunocytochemical (ICC) detection of dendrites with subsequent analysis using specialized (NIS-Elements) software.
Labeled phallotoxins have similar affinity for both large and small filaments (F-actin) but do not bind to monomeric globular actin (G-actin), unlike some actin antibodies 1. Nonspecific binding of Phalloidin is negligible, thus providing minimal background during cellular imaging. Phalloidin is much smaller than antibodies that would typically be used to label cellular proteins for fluorescent microscopy, which allows for much more intense labeling of F-actin by Phalloidin. Thus, detailed images of F-actin localization in neurons can be obtained through the use of labeled Phalloidin.
Phalloidin (F-actin) staining of neuronal dendrites generates discrete &#34;hot spots&#34; or bright &#34;puncta&#34;, which represent a variety of dendritic structures, including mature spines, non-spiny synapses 2 and immature spines. Immature spines include thin filopodia and some forms of patch morphology, and may represent the initiation of spinogenesis 3. Immature spines and non-spiny patches lack PSD95 4. Changes in production of F-actin lead to subsequent changes in not only spines but also additional dendritic structures, thus making Phalloidin an important tool for investigating synaptodendritic integrity 5-7. In general, numbers of Phalloidin-positive (F-actin) puncta reflect a balance among active synapses (excitatory and inhibitory), actin dynamics and synapse stability 8.
Although it is important to study specific types of synapses (i.e., excitatory spines), when the target of a treatment is unknown it is necessary to first estimate the general integrity of a variety of dendritic structures. Since F-actin is a major component of dendritic spines and other structures, including inhibitory synapses, an altered number of F-actin puncta may indicate a synaptopathy. This synaptopathy may then be investigated further for more specific alterations. Our quantification method for detecting multiple synaptic types/structures yields an overall estimate of dendritic synaptic alterations (increases and decreases) following various experimental treatments.

<table border="0" cellpadding="0" cellspacing="0" width="626">
	<tbody>
		<tr height="50">
			<td height="50" style="height:50px;width:211px;">35 mm Glass Bottom Dishes No. 1.5 coverglass</td>
			<td style="width:155px;">MatTek Corporation</td>
			<td style="width:121px;">P35G-1.5-20-C</td>
			<td style="width:139px;"></td>
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		<tr height="37">
			<td height="37" style="height:37px;width:211px;">DMEM/F12 medium</td>
			<td style="width:155px;">Life Technologies</td>
			<td style="width:121px;">10565-018</td>
			<td style="width:139px;"></td>
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			<td style="width:121px;">15400-054</td>
			<td style="width:139px;"></td>
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		<tr height="32">
			<td height="32" style="height:32px;width:211px;">Poly-L-Lysine</td>
			<td style="width:155px;">Sigma</td>
			<td style="width:121px;">P9155</td>
			<td style="width:139px;"></td>
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		<tr height="33">
			<td height="33" style="height:33px;width:211px;">Boric acid</td>
			<td style="width:155px;">Sigma</td>
			<td style="width:121px;">B0252</td>
			<td style="width:139px;"></td>
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			<td height="25" style="height:25px;width:211px;">Borax</td>
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			<td style="width:121px;">B9876</td>
			<td style="width:139px;"></td>
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			<td height="32" style="height:32px;width:211px;">GlutaMax</td>
			<td style="width:155px;">Life Technologies</td>
			<td style="width:121px;">35050-061</td>
			<td style="width:139px;">100X</td>
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			<td height="32" style="height:32px;width:211px;">Glucose</td>
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			<td style="width:139px;"></td>
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			<td height="26" style="height:26px;width:211px;">HBSS</td>
			<td style="width:155px;">Sigma</td>
			<td style="width:121px;">H4641</td>
			<td style="width:139px;">10X</td>
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			<td height="39" style="height:39px;width:211px;">Iris Forceps</td>
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			<td style="width:121px;">15914</td>
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			<td height="46" style="height:46px;width:211px;">Dumont #7 Forceps</td>
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			<td style="width:121px;">14097</td>
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			<td height="44" style="height:44px;width:211px;">Dumont #5 Forceps</td>
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			<td style="width:121px;">14095</td>
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			<td height="39" style="height:39px;width:211px;">Paraformaldehyde&#160;</td>
			<td style="width:155px;">Sigma</td>
			<td style="width:121px;">P6148</td>
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			<td height="37" style="height:37px;width:211px;">Cover glass</td>
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			<td height="35" style="height:35px;width:211px;">AlexaFluor 488 Phalloidin</td>
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			<td height="50" style="height:50px;width:211px;">Chicken polyclonal anti-MAP2</td>
			<td style="width:155px;">abcam</td>
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			<td style="width:139px;"></td>
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			<td height="69" style="height:69px;width:211px;">Alexa Red 594-conjugated goat anti-chicken IgG</td>
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			<td height="60" style="height:60px;width:211px;">NIS-Elements software package</td>
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			<td height="81" style="height:81px;width:211px;">Nikon Eclipse TE2000-E inverted fluorescent computer-controlled microscope&#160;</td>
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			<td style="width:121px;">151-4020</td>
			<td style="width:139px;"></td>
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</table>

quantification of filamentous actin (f-actin) puncta in rat cortical neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

In the present methods, we first culture rat cortical neurons at low density in 35 mm glass-bottom dishes, which allows us to identify the dendrites of individual neurons. In Figure 1, the differential interference contrast (DIC) images show the morphological changes in developing fetal rat cortical neurons at days 4, 6, 10, 14, 21 and 27 in vitro. Note that the length and number of dendrites increase with maturation of cultured rat primary neurons. Neurons are used in experiments only after 14 days maturation.
To detect dendritic and synaptic changes, we combine the Phalloidin F-actin labeling with MAP2 antibody detection of dendrites. Since Phalloidin labeling of F-actin is very rapid (20-30 min), it is possible to visually estimate the integrity of synaptodendritic network before proceeding with ICC antibody labeling (Figure 2). MAP2 indicates the intact dendrites and overlies the Phalloidin staining. This allows confirmation that the F-actin puncta are located on neuronal dendrites and not localized to other cells, such as astrocytes.
Next, we acquire high resolution images of co-labeled Phalloidin (F-actin)/MAP2 neurons and analyzed randomly selected neurons. Fine filopodia, spine protrusions, and F-actin patches were considered F-actin rich structures and were included in our studies (Figure 3). Segments of the second order dendrites (MAP2 positive) were selected for the analysis of F-actin puncta densities. MAP2 positive staining is used to confirm the neuronal localization of the F-actin puncta. Computer-assisted detection and counting of Phalloidin (F-actin) labeled (green fluorescence channel) synaptic puncta was performed via use of specialized software. There is a step by step detailed protocol in Figure 4 describing the use of the software package. We have reported that computer-assisted F-actin counting correlates very well with manual counting of F-actin and that the inter-observer correlation in F-actin puncta counts between two trained observers is very high (r2=0.97).
Previously, we used quantification of F-actin puncta to assess synaptodendritic injury induced by HIV-1 Tat9 and recovery from HIV-1 Tat-induced synaptopathy10. In Figure 5, rat cortical neurons were co-labeled with Phalloidin and MAP2 antibody after 50 nM of HIV-1 Tat treatment. MAP2 staining revealed fewer dendritic branches and diminished F-actin following HIV-1 Tat-treatment.
We found that F-actin puncta may either increase or decrease in response to experimental treatments. In Figure 6, cultured neurons were treated with the uncompetitive NMDA receptor antagonist memantine, significantly increasing F-actin positive puncta. In contrast, treatment with a combination of Methamphetamine+HIV-1 Tat resulted in significant loss of F-actin puncta.
<img alt="Figure 1" src="/files/ftp_upload/53697/53697fig1.jpg" /> 
Figure 1. Fetal rat cortical neurons in cell culture. Differential interference contrast (DIC) images of fetal rat cortical neurons between 4 - 27 days in vitro (20X). Neurons appear mature at 14 days in vitro. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53697/53697fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53697/53697fig2.jpg" /> 
Figure 2. Phalloidin/MAP2 co-labeling in rat cortical neurons. Cultured neurons labeled with MAP2 antibody (red) and Phalloidin (green). Merged images allow determination that the Phalloidin staining is localized to neuronal dendrites (20X). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53697/53697fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53697/53697fig3.jpg" /> 
Figure 3. F-actin synaptic structures of rat cortical neurons. (A) Left - F-actin positive structures labeled with Phalloidin (60X). The arrowheads indicate F-actin labeled structures include mushroom spines (purple), patch-like morphology (blue) and long filopodia (orange). Middle - Merged image demonstrating these F-actin structures are localized to dendrites (20X). Right - Box in the lower right indicates the second order dendritic branch selected for analysis (20X). (B) Dendritic segments of Phalloidin (F-actin) (Green)/MAP2(Red) co-labeled cortical neurons (20X) are used to verify neuronal origin of F-actin puncta. The green only image (Phalloidin/F-actin) is further processed. (C) Phalloidin/F-actin (green) images of three dendritic segments selected for puncta counting. Densities are determined by dividing N/L. N=number of puncta, L=length of dendritic segment. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53697/53697fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53697/53697fig4.jpg" /> 
Figure 4. Demonstration of software package: step by step instructions. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53697/53697fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53697/53697fig5.jpg" /> 
Figure 5. HIV-1 Tat mediated synaptodendritic injury in rat cortical neurons. Cell cultures were co-stained for F-actin and MAP2 after HIV-1 Tat protein treatment (50nM). Upper panels- untreated neurons showing robust F-actin, complex branching patterns, and extensive fine neuronal processes. Lower panels - HIV-1 Tat protein treated neurons with diminished F-actin and decreased dendritic branching. (20X) <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53697/53697fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/53697/53697fig6.jpg" /> 
Figure 6. F-actin puncta in rat cortical neurons: different effects produced by Memantine vs. Methamphetamine+Tat treatment. Images (20X) of untreated cultured neurons, neurons treated with Memantine (10 &#181;M), and neurons treated with Methamphetamine (20 &#181;M) +10 nM of Tat (10nM). Memantine treatment increased the F-actin staining, whereas Methamphetamine+HIV-1 Tat treatment decreased F-actin staining and decreased dendritic branching. Memantine treatment significantly increased F-actin puncta density, relative to untreated control cultures. In contrast, Methamphetamine+HIV-1 Tat treatments significantly decreased F-actin puncta density, relative to untreated control cultures. Mean + SEM, * p&#60;0.05. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53697/53697fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons at JoVE.com

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

quantification-filamentous-actin-f-actin-puncta-rat-cortical

Nanyang Technological University

Research

JoVE Journal

Neuroscience

11.9K Views.  University of South Carolina. Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Video: Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

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 ...

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.

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 ...

Labeling F-actin Barbed Ends with Rhodamine-actin in Permeabilized Neuronal Growth Cones

The motile tips of growing axons are called growth cones. Growth cones lead navigating axons through developing tissues by interacting with locally expressed molecular guidance cues that bind growth cone receptors and regulate the dynamics and organization of the growth cone cytoskeleton3-6. The main target of these navigational signals is the actin filament meshwork that fills the growth cone periphery and that drives growth cone motility through continual actin polymerization and dynamic remodeling7. Positive or attractive guidance cues induce growth cone turning by stimulating actin filament (F-actin) polymerization in the region of the growth cone periphery that is nearer the source of the attractant cue. This actin polymerization drives local growth cone protrusion, adhesion of the leading margin and axonal elongation toward the attractant.
Actin filament polymerization depends on the availability of sufficient actin monomer and on polymerization nuclei or actin filament barbed ends for the addition of monomer. Actin monomer is abundantly available in chick retinal and dorsal root ganglion (DRG) growth cones. Consequently, polymerization increases rapidly when free F-actin barbed ends become available for monomer addition. This occurs in chick DRG and retinal growth cones via the local activation of the F-actin severing protein actin depolymerizing factor (ADF/cofilin) in the growth cone region closer to an attractant8-10. This heightened ADF/cofilin activity severs actin filaments to create new F-actin barbed ends for polymerization. The following method demonstrates this mechanism. Total content of F-actin is visualized by staining with fluorescent phalloidin. F-actin barbed ends are visualized by the incorporation of rhodamine-actin within growth cones that are permeabilized with the procedure described in the following, which is adapted from previous studies of other motile cells11, 12. When rhodamine-actin is added at a concentration above the critical concentration for actin monomer addition to barbed ends, rhodamine-actin assembles onto free barbed ends. If the attractive cue is presented in a gradient, such as being released from a micropipette positioned to one side of a growth cone, the incorporation of rhodamine-actin onto F-actin barbed ends will be greater in the growth cone side toward the micropipette10.
Growth cones are small and delicate cell structures. The procedures of permeabilization, rhodamine-actin incorporation, fixation and fluorescence visualization are all carefully done and can be conducted on the stage of an inverted microscope. These methods can be applied to studying local actin polymerization in migrating neurons, other primary tissue cells or cell lines.

A method to visualize and quantify F-actin barbed ends in neuronal growth cones is described. After culturing neurons on glass coverslips, cells are permeabilized with a saponin-containing solution. Then, a short incubation with the saponin buffer containing rhodamine-actin incorporates fluorescent actin onto free actin barbed ends.

The motile tips of growing axons are called growth cones. Growth cones lead navigating axons through developing tissues by interacting with locally ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

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 ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons

The cerebral cortex is characterized by multiple layers and many distinct cell-types that together as a network are responsible for many higher cognitive functions including decision making, sensory-guided behavior or memory. To understand how such intricate neuronal networks perform such tasks, a crucial step is to determine the function (or electrical activity) of individual cell types within the network, preferentially when the animal is performing a relevant cognitive task. Additionally, it is equally important to determine the anatomical structure of the network and the morphological architecture of the individual neurons to allow reverse engineering the cortical network. Technical breakthroughs available today allow recording cellular activity in awake, behaving animals with the valuable option of post&#160;hoc identifying the recorded neurons. Here, we demonstrate the juxtasomal biocytin labeling technique, which involves recording action potential spiking in the extracellular (or loose-patch) configuration using conventional patch pipettes. The juxtasomal recording configuration is relatively stable and applicable across behavioral conditions, including anesthetized, sedated, awake head-fixed, and even in the freely moving animal. Thus, this method allows linking cell-type specific action potential spiking during animal behavior to reconstruction of the individual neurons and ultimately, the entire cortical microcircuit. In this video manuscript, we show how individual neurons in the juxtasomal configuration can be labeled with biocytin in the urethane-anaesthetized rat for post&#160;hoc identification and morphological reconstruction.

To understand the structure of neuronal networks, functional and morphological characterization of individual neurons is a necessity. Here, we demonstrate juxtasomal biocytin labeling, which allows electrophysiological recordings in the extracellular configuration, yet maintaining the ability to intracellularly label the neuron for post&#160;hoc reconstruction of dendritic and axonal architecture.

The cerebral cortex is characterized by multiple layers and many distinct cell-types that together as a network are responsible for many higher cognitive ...

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, ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

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 ...