This study was supported by an NIH/NIMH grant to S.Q. (R00MH087628).

All experimental procedures involving mice were approved by the Institutional Animal Care and Use Committee of the University of Arizona, and conformed to NIH guidelines.
1. Tissue Source for Hippocampal Neuron Culture
<ol>
	<li>To generate prenatal mouse pups for hippocampal neuron culture, use time-pregnant mice (C57Bl6/J) that are bred in house17. The day with vaginal plug detection is designated as E0.5. The planned harvest time for culture is E16.5-E17.5. 
	Note: This protocol describes cultures of two 24-well plates of two high density neurons co-culturing with low-density neurons (48 coverslips total). For more plates, materials and reagents can be scaled correspondingly.</li>
</ol>
2. 24-well Plates Preparation for High Density Neuron Cultures
Note: Perform the following steps (2-3) the day before the planned harvest of embryos.
<ol>
	<li>Bring two 24 well plates into the cell culture hood, open the individual packaging.</li>
	<li>Connect an 18 G syringe needle to a 1 ml plastic single-use syringe.</li>
	<li>Hold the syringe, and aim the needle tip at ~1 mm left to the center of the bottom of the well. Apply moderate force (~250 g) to etch the bottom of the well (~1 mm long). A groove will be formed and the plastic materials displaced will form an elevated support above the flat bottom well surface. Etch another ~1 mm-long parallel groove at ~1 mm to the right of the center of the bottom in a similar way.</li>
	<li>Repeat this step for all the wells of the two 24-well plates. The two 24-well plates used for high density culture are now ready for coating with poly-D-lysine (see step 3.8 below).</li>
</ol>
3. Coverslips Preparation for Low Density Neuron Cultures
<ol>
	<li>Use 12-mm diameter #1 round glass.</li>
	<li>Place 50 coverslips inside a 100 mm diameter sterile plastic petri dish, and submerge coverslips in 20 ml 70% ethanol solution. Place this petri dish on a rotating platform with moderate (70 rpm) speed rotation for an hour. Remove 70% ethanol and then replace with a fresh batch of 70% ethanol. Rotate and wash for another hour.</li>
	<li>Discard 70% ethanol cleaning solution. Add sterile water (filtered through 0.2 &#181;m filter unit). Rinse the coverslips rigorously by rotating the petri dish by hand. Rinse three times, each time replacing with fresh sterile water.</li>
	<li>Place the petri dish with coverslips inside a sterile cell culture hood with laminar flow. Aspirate the rinsing water through the vacuum line, and add 10 ml filtered sterile water to immerse the coverslips. The coverslips should be sterile and the surface should be clean enough for poly-D-lysine coating.</li>
	<li>Open two 24-well plates from their individual packing, and place them in the hood.</li>
	<li>Turn on the vacuum line in the hood. Connect a 200 &#181;l pipette tip to the vacuum line, and use a scissor to cut the tip to create a larger opening.</li>
	<li>Use a fine-tip #5 tweezer to pick up coverslips from the petri dish, one at a time, and use the vacuum line to completely dry the coverslip. Then place one coverslip into each of the wells of the 24-well plate. The 50 cleaned coverslips should be enough to fill each of the wells of the two 24-well plates.</li>
	<li>Dilute the poly-D-lysine stock solution (1 mg/ml) into working solution (0.1 mg/ml) with borate buffer (pH 8.5). For the four 24-well plates (including two high density plates with etched bottom), prepare 30 ml of total working solution.</li>
	<li>Add 300 &#181;l 0.1 mg/ml poly-D-lysine work solution to each well with glass coverslips. Make sure the coverslips are completely immersed in solution. If not, use the #5 tweezer tip to press down coverslips against the bottom of the well, and use the tip to guide poly-D-lysine solution to cover all the surface of the coverslips. Visually inspect all the coverslips to make sure no air was trapped between the plate and coverslips.</li>
	<li>Add 300 &#181;l 0.1 mg/ml poly-D-lysine work solution to each well of the high density culture plate with etched bottom. Rotate by hand to spread the solution to cover the entire bottom of the well.</li>
	<li>Return all the four plates with poly-D-lysine to the culture incubator, and incubate O/N.</li>
</ol>
4. Washing and Pre-conditioning Plates for Culture
Note: The following steps (4-9) are carried out on the day of tissue harvest.
<ol>
	<li>After incubation/coating of coverslips and 24-well plates O/N, bring all four plates (two with donor coverslips, two acceptor high density plates with etched plastic bottom) into the culture hood. Use the vacuum line to remove the poly-D-lysine solution.</li>
	<li>Immediately following the removal of the poly-D-lysine solution, add ~1 ml sterile water to each well using a plastic transfer pipette. After all the wells are filled with water, let stand for 10 min. Remove water by vacuum, then add 300 &#181;l Wash medium in each well to cover the bottom of the well (with or without glass coverslips). Do not let the poly-D-lysine coating dry out.</li>
	<li>Return all four plates to the cell incubator. The plates are ready to use once the dispersed hippocampal neurons are prepared.</li>
</ol>
5. Preparation of Complete Culture Medium, and Trypsin Solution for Enzymatic Digestion
<ol>
	<li>Prepare 60 ml Complete Culture medium by mixing the ingredients (see Materials). Use a 50 ml syringe connected with a 0.2 &#181;m-pore size syringe filter, filter the Complete Culture medium into two 50&#160;ml conical tubes. Each tube contains 30 ml Complete Culture medium.</li>
	<li>In a separate 50 ml conical tube, add ~30 ml Wash medium (see Materials) and warm it up to 37 &#176;C. This will be used to wash the tissue after enzymatic digestion.</li>
	<li>Prepare the enzyme digestion medium. In a 15 ml conical tube, add 4.5 ml Wash medium and 0.5 ml 2.5% trypsin solution, and 50 &#181;l 100X DNase I.</li>
	<li>Place the Complete Culture medium, Wash medium, and enzyme digestion medium in a 37 &#176;C water bath.</li>
</ol>
6. Preparation of Surgical Tools
<ol>
	<li>Sterilize all the surgical tools in a sterilization pouch: two small scissors, a pair of forceps and two #5 tweezers.</li>
</ol>
7. Removal of Brains from E16.5-E17.5 Mouse Embryos, and Dissection of Hippocampi
<ol>
	<li>Prepare two 10-cm petri dishes filled with ice-cold, sterile Hank&#39;s balanced salt solution (HBSS).</li>
	<li>Euthanize the mouse dam with an intraperitoneal injection of sodium phenobarbital (100 mg/kg in a volume of ~ 0.5 ml). After injection, once the dam loses response to toe pinch, sacrifice with cervical dislocation.</li>
	<li>Spray the dam&#39;s abdomen area with 70% ethanol, and make a 2-cm long incision along the midline of abdomen to expose the uterus.</li>
	<li>Remove the embryos and place individual ones in a 10-cm diameter petri dish with cold dissecting HBSS buffer.</li>
	<li>Hold the embryo by the neck using a forceps, and use a small scissor to make the first cut right in the midline below the cerebellum level and section across the brain tissue. Do not decapitate the embryo.
	<ol>
		<li>Insert the right side tip of the small scissor from caudal region that is cut open, all the way through the rostral part of the embryonic brain (do not cross midline), and make a cut on the left side at the level of skull base.</li>
		<li>Insert the left side scissor tip again to make a cut on the right side. Leave a narrow band of uncut skull bone and skin tissue at the rostral end, so that the whole brain can be flipped aside for easy extraction.</li>
		<li>Use the tip of scissor to scoop out the embryonic brain into the HBSS solution. Inspect the brain under a dissecting microscope. The brain should be intact with no gross cut-off regions.</li>
		<li>Repeat these steps to collect all the embryo brains.</li>
	</ol>
	</li>
	<li>Collect all the intact brains in a new petri dish containing 20 ml cold HBSS buffer. Place the petri dish under a dissection microscope.</li>
	<li>View the brain from ventral side. While holding down the brain with forceps at the caudal end, insert a sharp #5 tweezer diagonally to make a cut between the middle rostral point and the caudal-temporal region. Repeat this for the other hemisphere. After cutting, separate two hemispheres with intact hippocampus from the rest of brain stem tissues.
	<ol>
		<li>Repeat this for each of the brains.</li>
	</ol>
	</li>
	<li>Pool all the dissected hemispheres, and remove the meninges using two pairs of #5 tweezers.</li>
	<li>Identify the developing hippocampus as a curved structure that is folded inside the temporal lobe. Alternate the two pairs of tweezers to separate the hippocampus from adjoining cortical tissues. Collect and pool all the hippocampi tissue (see Figure 1).</li>
</ol>
8. Enzymatic Digestion, Separate into Single Neurons
<ol>
	<li>Using a plastic transfer pipette, transfer all the dissected hippocampi into the enzymatic digestion solution containing 5 ml Wash medium with 0.25% trypsin + 1X DNase I in a 15 ml conical tube.</li>
	<li>Place the tube inside a water bath and incubate at 37 &#176;C for 25 min, with intermittent gentle inverting of the tube once in every 5 min.</li>
	<li>After 25 min, carefully remove the enzyme solution using a hand held plastic pipette by aspiration. Add warm Wash medium to 10 ml and gently wash the tissue by slowly inverting the tube 5 times. Remove the Wash medium, and add 10 ml Wash medium again for an additional wash.</li>
	<li>Remove the Wash medium, and add 3 ml of fresh warm Wash medium again. Shake the tube by hand rigorously for 15 times to dislodge the digested neurons from the tissue. The medium should become turbid once the cells are separated from the tissue into the Wash medium. Collect the cell suspension in a new 15 ml conical tube while leaving the remaining tissue in the tube.</li>
	<li>Add another 2 ml Wash medium to the tissue, and gently separate the remaining tissue by triturating through a plastic pipette for ~15 times. Let sit for 5 min. Pool the cell suspension into the new 15 ml conical tube that contains collected &#39;shake-off&#39; cells.</li>
	<li>Count the density of neurons with a hemocytometer. Typically, 3,000-5,000 cells per microliter can be obtained depending on the number of embryos dissected. An average number of 2.5 x 107 neurons are estimated if assuming six embryos are dissected.</li>
	<li>Calculate the volume needed of this cell suspension to yield a density of 250,000 neurons/ml when added to the 30 ml Complete Culture medium. Add this volume to one of the two conical tubes containing 30 ml Complete Culture medium to yield the high density neuron plating medium.</li>
	<li>Transfer 1.2 ml of this high density neuron solution to another 30 ml Complete Culture medium to yield a concentration of ~10,000 neurons/ml. Use this dilution to seed the low-density coverslips.</li>
</ol>
9. Plating of Neurons and Long Term Co-culture
<ol>
	<li>Bring the two 24-well plates with etched bottoms for high density neurons into the cell culture hood. Remove the 300 &#181;l precondition medium by vacuum. Plate 0.6 ml high density cell suspensions at 250,000 neurons/ml.</li>
	<li>Bring the other two 24-well plates with coverslips into the culture hood, and remove the 300 &#181;l precondition medium by vacuum. Plate 0.6 ml low density cell suspensions at 10,000 neurons/ml on the coverslips inside the two 24-well plates.</li>
	<li>Return all four 24-well plates to the incubator. Let sit for 2 hr.</li>
	<li>Flip the low density coverslips with adhering neurons to the high density culture. Make sure the neurons are facing each other (i.e., not separated by the glass coverslip). Then return the two plates with co-culture to the incubator.</li>
</ol>
10. Co-culture Sustaining
<ol>
	<li>Feed co-cultures by adding 300 &#181;l fresh Feed medium (see Materials) at day in vitro (DIV) 5. There is no need to remove 50% of original Complete Culture medium, as reported by many other studies. The feeding medium also contains 15 &#181;M cytosine arabinoside (Ara-C, to yield a final concentration of 5 &#181;M) to minimize the chance of glia contamination and proliferation.</li>
	<li>Following this initial feeding, add 300 &#181;l fresh Feed medium without Ara-C to the well every week. Should the culture be kept for more than one month, replace half of the medium with fresh Feed medium every week beyond one month time point (with the first medium half-replacement at DIV33). Morphologically, both high density and low density neurons look healthy up to three months (the longest time observed by the experimenters).</li>
</ol>
11. Illustration of an Experimental Manipulation-low Density Neuron Transfection
Note: When transfection in low density neurons is desired, a simple calcium phosphate protocol can be adopted. We have found that low density cultures have better transfection efficiency with calcium phosphate protocol before DIV12, but they can be transfected with much lower efficiency at DIV21 or older, during which time dendritic spines are prominent. The feasibility of transfection of the cultured low density neuron is illustrated by transfecting neurons with a pEGFP-C3 plasmid (see below).
<ol>
	<li>At DIV21, remove 600 &#181;l conditioned medium from each well of the co-culture, and transfer it to a new 24-well plate. Then add 600 &#181;l fresh Feed medium to the co-culture to bring it back to the original volume.
	<ol>
		<li>Transfer the coverslips into the new 24-well plate with 600 &#181;l conditioned medium. Make sure the low density neurons are facing upwards. Place the high density plates and the new plates with coverslips back in the incubator.</li>
	</ol>
	</li>
	<li>Prepare two 1.5 ml sterile tubes. In one tube, add 600 &#181;l 2X HEPES buffered saline (HBS) solution. Calculate the volume of 12 &#181;g pEGFP-C3 DNA based on plasmid concentration. In the other tube, add 74 &#181;l CaCl2 (2 M stock) and the volume of water that after adding plasmid will make 600 &#181;l. Mix well by pipetting and then add plasmid DNA. Slowly mix well. Let both tubes sit for 5 min at RT.</li>
	<li>While hand-holding the 2X HBS tube and moderately agitating the solution on a mixer, add all of the 600 &#181;l the DNA-CaCl2 solution drop by drop to the 600 &#181;l 2X HBS tube. It is critical that the precipitate formed is in the shape of &#39;fine sand&#39; granules. Mixing too strong and too fast is not desirable, since it will more likely produce large &#39;snow flake&#39;-like precipitates, which has dramatically lower transfection efficiency based on our observations.</li>
	<li>Let the precipitate continue to form in the dark at RT for 30 min.</li>
	<li>Add 100 &#181;l precipitate to each well that contains low density neuron on the coverslips, drop by drop and evenly. Return the plate to incubator, and incubate for 2 hr. Assuming most of the CaCl2 is in free form, this results in a concentration of ~17.6 mM Ca2+, which may be toxic to neurons following prolonged incubation.</li>
	<li>Prepare a 50 mL conical tube with ~50 ml Wash medium, warm it up to 37 &#176;C.</li>
	<li>At the end of 2 h, bring the low density neuron with DNA-calcium phosphate precipitates into the hood. Pick up each individual coverslip with a sharp #5 tweezer, and dip it three times in the 50 ml Wash medium to wash it briefly. Then return it to the high density neurons in its original co-culture plates, with low density neurons facing downwards.</li>
	<li>Continue the culture until the low density culture neurons on coverslips are harvested for studies.</li>
</ol>

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The authors declare that they have no competing financial interests.

We present a detailed protocol for long-term culture of ultra-low density hippocampal glutamatergic neurons under serum free conditions. At ~2000 neurons/cm2, the density is at least two fold lower than most &#39;low density&#39; culture preparations with or without glia support reported by the existing literature2,3,11,13,14. In addition to being ultra-low density, this protocol is novel and significant in two more ways. First, no glia feeder layer is needed as the low density neurons obtain trophi...

Growing hippocampal neurons under in vitro conditions enables observation and experimental manipulation of these neurons that are otherwise not possible in vivo. This experimental approach is widely used to reveal neuronal mechanisms of growth, polarity, neurite specification, trafficking and subcellular localization of proteins, synapse formation and functional maturation1. These in vitro cultured hippocampal neurons, when harvested from late embryonic stages, are relatively pure (&#62;90%) glutamatergic cells of pyramidal morphology2. Because neurons were grown in a 2-D surface under in vitro conditions, this method allows easy observation, such as live imaging or immunocytochemistry (ICC) staining through a single focal plane3; or manipulations, such as drug treatment and transfections3-6. When grown at high density, neurons tend to have high rates of survival because of higher concentrations of secreted growth factors in addition to alimentary support from the growth media, and also because of neurite contact-dependent mechanisms7. However, low density hippocampal neurons are desirable for morphological studies, where an individual neuron can be imaged in its entirety or stained for ICC analysis. Low density neurons are hard to maintain in culture due to lack of paracrine support and thus often require trophic support from a glial (typically cortical astrocyte) feeder layer, which has to be prepared prior to neuron culture2. When co-cultured with a glial cell feeder layer, low density neurons are grown on coverslips, and then flipped on top of the glia layer so that the low density neurons and glia are facing each other. A small confined space between glia and neurons is created by placing paraffin wax dots on the corners of the coverslips, therefore creating a &#39;sandwich&#39; layout2,8,9. The low density neurons will grow within the confined space between glia and the coverslip, which creates a permissive microenvironment with concentrated factors secreted by neurons and glia. This approach yields low-density, fully developed neurons that are spaced reasonably apart, therefore facilitating ICC labeling or live imaging studies.
An apparent drawback of neuron-glia co-culture, aside from being time-consuming and laborious, is that it prevents study of neuron-specific, or cell-autonomous, mechanisms. Although this system is far less complex than in vivo neural tissue, glia impact on neuron development through secreted, not-yet-fully-defined factors can confound the experiments10. Therefore, in experiments that require the investigation of neuron specific mechanisms, defined culture conditions that remove serum and glial support layer are necessary. A previous study has succeeded in culturing low concentrations of neurons (~9,000 cells/cm2) using a three dimensional hydrogel matrix11. Since a relatively pure neuronal population can be cultured at high density under serum free conditions without glial support, we hypothesize that ultra-low density of hippocampal neurons can be grown in serum free defined culture medium by co-culturing them with high density neurons, in a way that is analogous to the conventionally adopted neuron-glia co-culture. Indeed, high density hippocampal neuron cultures in a &#39;sandwich&#39; configuration have been recently used to support a small number of specialized magnocellular endocrine neurons12.
Therefore, co-culture with high density neurons may allow low density neurons to receive trophic factors support that is sufficient to enable long term survival. This protocol of culturing ultra-low density neurons was thus formulated and validated. The protocol can be implemented within one single experiment, by preparing high density (~250,000 cells/ml) dissociated hippocampal neurons first, and then making a dilution to yield a density ~10,000 neurons/mL (~3,000 neurons/coverslip, or ~2,000 neurons/cm2), which is much lower than most reported low density cultures2,3,9,11,13. This culture condition is loosely referred to as &#39;ultra-low density&#39; culture and used with &#39;low-density&#39; inter-changeably. The high density neurons are plated on the poly-D-lysine coated 24-well plates; while the low density neurons are seeded on poly-D-lysine coated 12-mm glass coverslips that are placed inside another 24-well plate. The coverslips with adhering low density neurons are flipped on top of the high density neurons 2 hr later after the neurons are descended and attached to the coverslips. In addition, instead of using paraffin wax dots to elevate the coverslips above the high density neuron layer, an 18 G syringe needle was used to etch the bottom of the 24 well plates with two parallel stripes. The resulting displaced, adjoining plastics provide an elevated support for the glass coverslips. This space is consistently measured at 150-200 &#956;m, which allows sufficient oxygen and culture medium exchange while providing a microenvironment with concentrated trophic factors. Under this condition, the low density neurons grow extensively, and can survive beyond three months in culture. When these neurons are transfected with GFP plasmid after three weeks in culture, the dendrites are profusely studded with dendritic spines. As a proof of principle, data are presented to show that this co-culture system supports ultra-low density cultures of hippocampal neurons seeded on poly-D-Lysine &#39;micro-islands&#39;, where neurons form autaptic connections that may facilitate investigation of cell-autonomous, network-independent mechanisms.

<table><tbody><tr><td>Neurobasal medium</td><td>Life Technologies</td><td>21103-049</td><td>Protect from light</td></tr><tr><td>B27 supplement</td><td>Life Technologies</td><td>17504-044</td><td>aliquot, store in 0.6ml size</td></tr><tr><td>GlutaMAX-I</td><td>Life Technologies</td><td>35050-061</td><td>dilute 100X&amp;nbsp;</td></tr><tr><td>antibiotic-antimycotic (AA)</td><td>Life Technologies</td><td>15240-096</td><td>dilute 100X&amp;nbsp;</td></tr><tr><td>Complete Culture medium</td><td></td><td></td><td>Neurobasal medium with 1X B27, 1X AA, 1X GlutaMAX-I</td></tr><tr><td>Wash medium</td><td></td><td></td><td>same as &#039;Neurobasal medium&#039;</td></tr><tr><td>Feed medium</td><td></td><td></td><td>Neurobasal with 1X B27 supplement</td></tr><tr><td>DNAse I</td><td>Sigma-Aldrich</td><td>D5025</td><td>prepare 100X stock at 0.6mg/ml</td></tr><tr><td>poly-D-lysine</td><td>Sigma-Aldrich</td><td>P6407</td><td>M.W. 70000-150000</td></tr><tr><td>borate buffer</td><td>Sigma-Aldrich</td><td>B6768 (boric acid); 71997(borax)</td><td>1.24g boric acid &amp;amp; 1.9g borax in 400ml H2O, pH to 8.5 use HCl</td></tr><tr><td>12-mm round glass coverslips</td><td>Glasswarenfabrik Karl Hecht GmbH</td><td>1001/12</td><td>No. 1 glass, purchase from Carolina Biological Supply</td></tr><tr><td>proFection transfection kit</td><td>Promega</td><td>E1200</td><td>see&amp;nbsp; protocol for details</td></tr><tr><td>2X HEPES buffered saline (HBS)</td><td>Promega</td><td>E1200</td><td>see&amp;nbsp; protocol for details</td></tr><tr><td>Syringe filter</td><td>Pall Corporation</td><td>4192</td><td>0.2um pore size</td></tr><tr><td>Endofree plasmid prep kit</td><td>Qiagen</td><td>12362</td><td>for preparation of transfection grade plasmid DNA</td></tr><tr><td>anti-MAP2 antibody</td><td>Millipore</td><td>MAB3418</td><td>mouse antibody, clone AP20</td></tr><tr><td>anti-p-Tau antibody</td><td>Millipore</td><td>AB10417</td><td>rabbit polyclonal antibody</td></tr><tr><td>anti-NR1 antibody</td><td>Millipore</td><td>MAB1586</td><td>mouse antibody, clone R1JHL</td></tr><tr><td>anti-GluR1 antibody</td><td>Millipore</td><td>AB1504</td><td>rabbit polyclonal antibody</td></tr><tr><td>Hank&#039;s balanced salt solution</td><td>ThermoFisher</td><td>14025092</td><td>500ml size</td></tr></tbody></table>

a simplified method for ultra-low density, long-term primary hippocampal neuron culture

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic hippocampal neurons can often grow successfully on glass coverslips at high density under serum-free conditions, but low density cultures typically require a supply of trophic factors by co-culturing them with a glia feeder layer, preparation of which can be time-consuming and laborious. In addition, the presence of glia may confound interpretation of results and preclude studies on neuron-specific mechanisms. Here, a simplified method is presented for ultra-low density (~2,000 neurons/cm2), long-term (&#62;3 months) primary hippocampal neuron culture that is under serum free conditions and without glia cell support. Low density neurons are grown on poly-D-lysine coated coverslips, and flipped on high density neurons grown in a 24-well plate. Instead of using paraffin dots to create a space between the two neuronal layers, the experimenters can simply etch the plastic bottom of the well, on which the high density neurons reside, to create a microspace conducive to low density neuron growth. The co-culture can be easily maintained for &#62;3 months without significant loss of low density neurons, thus facilitating the morphological and physiological study of these neurons. To illustrate this successful culture condition, data are provided to show profuse synapse formation in low density cells after prolonged culture. This co-culture system also facilitates the survival of sparse individual neurons grown in islands of poly-D-lysine substrates and thus the formation of autaptic connections.

The protocol described here enables successful ultra-low density, long-term culture of pure glutamatergic neurons without the need of glia cells serving as a feeder layer. The protocol is diagramed in Figure 1, which involves preparation of high density (on poly-D-lysine coated 24 wells) and low-density neurons (on poly-D-lysine coated glass coverslips) separately, and subsequent co-culture that can be maintained up to three months.
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A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture

Low density cultures of primary hippocampal neurons usually require glia feeder layer to supply neurotrophic factors and sustain longevity. We describe here a simplified method to culture ultra-low density neurons on glass coverslips in the presence of a high density neuronal feeder layer, which facilitates investigation of specific neuronal-autonomous mechanisms.

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic ...

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15.7K Views.  University of Arizona College of Medicine-Phoenix. Low density cultures of primary hippocampal neurons usually require glia feeder layer to supply neurotrophic factors and sustain longevity. We describe here a simplified method to culture ultra-low density neurons on glass coverslips in the presence of a high density neuronal feeder layer, which facilitates investigation of specific neuronal-autonomous mechanisms. 

Video: A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture

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 ...

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

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 ...

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 ...

High-resolution Volume Imaging of Neurons by the Use of Fluorescence eXclusion Method and Dedicated Microfluidic Devices

Volume is an important parameter regarding physiological and pathological characteristics of neurons at different time scales. Neurons are quite unique cells regarding their extended ramified morphologies and consequently raise several methodological challenges for volume measurement. In the particular case of in vitro neuronal growth, the chosen methodology should include sub-micrometric axial resolution combined with full-field observation on time scales from minutes to hours or days. Unlike other methods like cell shape reconstruction using confocal imaging, electrically-based measurements or Atomic Force Microscopy, the recently developed Fluorescence eXclusion method (FXm) has the potential to fulfill these challenges. However, although being simple in its principle, implementation of a high-resolution FXm for neurons requires multiple adjustments and a dedicated methodology. We present here a method based on the combination of fluorescence exclusion, low-roughness multi-compartments microfluidic devices, and finally micropatterning to achieve in vitro measurements of local neuronal volume. The high resolution provided by the device allowed us to measure the local volume of neuronal processes (neurites) and the volume of some specific structures involved in neuronal growth, such as growth cones (GCs).

Volume is an important parameter regarding physiological and pathological characteristics of cells. We describe a fluorescent exclusion method allowing full-field measurement of in vitro neuronal volume with sub-micrometric axial resolution required for the analysis of neurites and dynamic structures implied in neuronal growth.

Volume is an important parameter regarding physiological and pathological characteristics of neurons at different time scales. Neurons are quite unique ...

Super-Resolution Imaging to Study Co-Localization of Proteins and Synaptic Markers in Primary Neurons

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. Imaging studies are widely used to investigate their function and plasticity in physiological and pathological conditions. Because of their size and structure, localization studies of proteins require high-resolution imaging techniques. In this protocol, we describe a procedure to study in primary neurons the co-localization of target proteins with synaptic markers at a super-resolution level using structured illumination microscopy (SIM). SIM is a patterned-light illumination technique that doubles the spatial resolution of wide-field microscopy, reaching a detail of around 100 nm. The protocol indicates the required controls and settings for robust co-localization studies and an overview of the statistical methods to analyze the imaging data properly.

This protocol shows how to employ super-resolution microscopy to study protein co-localization in primary neuronal cultures.

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. ...

Easy and Reproducible Low-Density Primary Culture using Frozen Stock of Embryonic Hippocampal Neurons

Neuronal culture is a valuable system for evaluating synaptic functions and drug screenings. In particular, a low-density culture of primary hippocampal neurons allows the study of individual neurons or subcellular components. We have shown subcellular protein localization within a neuron by immunocytochemistry, neuronal polarity, synaptic morphology, and its developmental change using a low-density primary hippocampal culture. Recently, ready-to-use frozen stocks of neurons have become commercially available. These frozen stocks of neurons reduce the time needed to prepare animal experiments and also contribute to the reduction of the number of animals used. Here, we introduce a reproducible low-density primary culture method using a 96-well plate. We used a commercially available frozen stock of neurons from the rat embryonic hippocampus. The neurons can be stably cultured long-term without media changes by reducing the growth of glial cells at particular timepoints. This high-throughput assay using low-density culture allows reproducible imaging-based evaluations of synaptic plasticity.

A ready-to-use frozen stock of neurons is a powerful tool for evaluating synaptic functions. Here, we introduce an easy low-density primary culture from frozen stock using a 96-well plate.

Neuronal culture is a valuable system for evaluating synaptic functions and drug screenings. In particular, a low-density culture of primary hippocampal ...

A Technique to Culture Mouse Hippocampal Neurons

Source: Yang, R., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/61411">Use of Primary Cultured Hippocampal Neurons to Study the Assembly of Axon Initial Segments.</a> J. Vis. Exp. (2021)
The video demonstrates a method for culturing neurons from murine hippocampi. Initially, the hippocampi undergo enzymatic digestion to break down the extracellular tissue matrix. Subsequently, the digested tissue is mechanically disrupted to isolate individual neurons. Finally, these neurons are cultured on glial feeder cells.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Preparing glia cell feeder dishes ...

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Neuronal Culture Techniques

Primary Neuronal Culture

Generating a Low-Density Primary Neuron Culture From Frozen Embryonic Neurons

Source: Koganezawa, N. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/64872">Easy and Reproducible Low-Density Primary Culture using Frozen Stock of Embryonic Hippocampal Neurons.</a>&#160;J. Vis. Exp. (2023)
This video demonstrates the method of culturing cryopreserved embryonic neurons, involving controlled thawing, dilution with a neurobasal medium, and seeding onto a poly-L-lysine-coated plate. Subsequent incubation supports the growth and maintenance of neuronal connections.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Plate coating

	Coat a 96-well ...

An Indirect Neuron-Astrocyte Co-Culture Technique for Studying Neuron-Glia Interactions

Source: Gottschling, C. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/54757">The Indirect Neuron-astrocyte Coculture Assay: An In Vitro Set-up for the Detailed Investigation of Neuron-glia Interactions.</a>&#160;J. Vis. Exp.&#160;(2016)
This video illustrates a method for establishing an indirect neuron-astrocyte co-culture using a compartmentalized approach. In this setup, neurons and astrocytes are spatially separated but share a conditioned medium. Neurotrophic factors released by astrocytes promote neuron differentiation and the formation of synaptic connections, suggesting neuron-glia interactions.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...