We thank Dr. Michael Wooten for his help in preparing the manuscript. This work was supported by NIH 2RO1NS033661 (MWW).

1. Set-up Prior to Harvest
<ol>
 <li>To generate prenatal pups for neuron harvest, schedule breeding between adult mice 19 days prior to the day of neuron isolation. (C57BL/6 mice ages 2-8 months were used in matings for the purposes of developing this protocol). Successful mating can be confirmed by detection of a vaginal plug in the female, palpitation or visual confirmation of pregnancy.</li>
 <li>The day prior to neuron isolation:
 <ol class="ualpha">
 <li>For immunofluorescence applications, coat glass coverslips in a 24-well plate with a light coating of 3:1 Collagen 1, Rat Tail:poly-D-Lysine solution.</li>
 <li>For cell culture applications, coat appropriate size tissue culture plastic ware with a light coating of 3:1 Collagen 1, Rat Tail:poly-D-Lysine solution.</li>
 </ol>
 </li>
 <li>Rest the plates uncovered in a tissue culture hood under a UV light overnight.</li>
 <li>Wash the plates with sterile Hank's Balance Salt Solution (HBSS) prior to use. Coated plates can be filled with HBSS and stored up to one week at 4 &deg;C in the dark.</li>
</ol>
2. Tissue Harvest
<ol>
 <li>Sterility is always a factor when growing primary cell cultures and as such, the greatest caution should be exercised to ensure the most sterile environment possible. With careful attention to sterile technique, initial dissection and harvest of neural tissue for this protocol can be completed outside of a laminar flow hood with minimal risk of contamination. After initial harvest, all subsequent steps should be conducted under maximum sterile conditions within a hood rated for cell culture.</li>
 <li>Euthanize a pregnant mouse at approximately 19 days post-fertilization by decapitation. Use of anesthesia to euthanize the pregnant female is not recommended as anesthesia is known to cause brain cell death (Stratmann, et al., 2010). Using sterile dissecting scissors and forceps, create an opening in the mid-ventral side of the mouse to completely reveal body cavity. Instruments can be sterilized using alcohol and an open flame.</li>
 <li>Prenatal pups will be located towards the posterior of the mouse's body cavity and should be easily visible in the uterus. With autoclaved sterile forceps, open the uterus and remove pups. Decapitate pups with fresh sterile scissors and place removed head on sterile gauze under a dissecting microscope. Sterile, autoclaved instruments can be flame cleaned using alcohol and an open flame prior to and during use. </li>
 <li>Using sterile scissors or scalpel, open cranium of pup from back of the neck to the nose. This procedure can normally be completed by inserting one tip of the scissors into the vertebral foramen and then proceeding anteriorly. Carefully remove the entire brain with forceps. Place the brain on sterile gauze. Using a sterile scalpel, remove the cerebellum and incise down the midline of the brain to separate it into two hemispheres (Figure 1).</li>
 <li>Grasp a small section of meninges surrounding the hippocampus with sterile forceps and pull it gently away. Although it is not strictly necessary to remove the meninges before isolating the hippocampus, the presence of the meninges can make the dissection of the hippocampus more difficult due to the toughness of the membrane. In either case, the hippocampus will be more clearly visible after the meninges have been removed. The hippocampus is a curved structure that starts in the distal part of the hemisphere and bends ventrally (Figure 2). As the inner, concave, side (caudal) is facing a ventricle, it is already free. Therefore, to isolate the hippocampus, one needs to cut along the convex outer side. After the dissection, gently lift each hippocampi with sterile tissue forceps and transfer into a small tissue culture dish with warmed (37 &deg;C) HBSS under a cell culture hood. Brain tissue can be combined from multiple pups.</li>
</ol>
3. Tissue Dissociation
<ol>
 <li>Using a sterile scalpel, gently mince brain tissue in 3 ml of sterile HBSS in a 100 mm Tissue Culture dish.</li>
 <li>Transfer the minced tissue and HBSS to a 15 ml conical tube. Add 1.5 ml of HBSS and 0.5 ml of 0.25% Trypsin solution to a total volume of 5 ml.</li>
 <li>Cap and gently invert the tube 4-5 times to mix. Try to avoid producing bubbles as DNA released from the digested tissue will adhere to the bubbles and cause the minced tissue to float instead of settling to the bottom of the tube (Figure 3).</li>
 <li>Incubate hippocampal tissue at 37 &deg;C for 15 minutes, inverting tube as above every 5 minutes.</li>
 <li>Allow the tissue to settle to the bottom of the tube.</li>
 <li>Carefully remove excess solution using a sterile pipette, leaving tissue undisturbed at the bottom of the tube.</li>
 <li>Wash tissue pellet with 5 ml of HBSS at 37 &deg;C for 5 minutes. Repeat a total of 3 times. Allow tissue to completely settle to the bottom of the tube each time before proceeding to the next wash step.</li>
 <li>Remove the final wash from the tissue pellet and replace with 2 ml of fresh HBSS.</li>
</ol>
4. Neuron Trituration
<ol>
 <li>Before beginning trituration steps, you will need to prepare fire polished Pasteur pipettes. Using a Bunson burner, hold a sterile 9 inch Pasteur pipette tip (Figure 4a) in the flame until diameter of the pipette opening is approximately 0.5 mm in size and edges of the pipette opening have been rounded slightly (Figure 4b). Allow pipette to completely cool before beginning the trituration process.</li>
 <li>Using a normal sterile 9 inch Pasteur pipette, gently triturate the tissue a total of 7 times. Larger tissue pieces are normal at this point and should be allowed to settle to the bottom of the tube prior to moving to the next step.</li>
 <li>Transfer the supernatant to a fresh sterile 50 ml conical tube.</li>
 <li>To the remaining tissue, add 2 ml of sterile HBSS and triturate a total of 5 times using the fire polished Pasteur pipette.</li>
 <li>Allow all remaining larger tissue pieces to settle to the bottom of the tube and combine supernatant with the previous supernatant for a total of 4 ml dissociated neuronal cells.</li>
</ol>
5. Cell Plating
<ol>
 <li>Count the dissociated cells using a hemocytometer.</li>
 <li>As a general rule, once cell numbers have been determined, subtract 20% from that final number to account for any cell death that may occur after plating.</li>
 <li>Cells can be plated using the recommendations below: 
 For coverslips in a 24 well plate - 6 x 104 cells/well in 0.5 ml 
 For 60 mm Tissue Culture plates - 4 x 105 cells/plate in 3 ml 
 For 100 mm Tissue Culture plates - 6 x 106 cells/plate in 6 ml</li>
 <li>Mix appropriate cell numbers with indicated volume of Neurobasal Plating Media (Neurobasal Media containing B27 Supplement [1 ml / 50 ml], 0.5 mM Glutamine Solution, 25 &mu;M Glutamate (Mr 147.13 g / mol), Penicillin (10,000 units / ml)/Streptomycin (10,000 &mu;g / ml) [250 &mu;l / 50 ml], 1mM HEPES (Mr 238.3 g / mol), 10% Heat Inactivated Donor Horse Serum) and add cells to plates. Swirl plates gently to distribute cells evenly. HI-Donor Horse Serum is added to the Plating Media to enrich the cells during the first 24 hours of growth. The cells are subsequently weaned from the serum and returned to a Serum-Free environment by serial reduction of the serum at each media replacement. It should also be noted that while glutamate at higher concentrations is toxic to neuronal cell cultures, at the lower concentrations added here, it will inhibit the attachment of non-neuronal cells11. However, it should only be added to the plating media for the first 24 hours in culture and subsequently left out of any Feeding Media to prevent neurotoxicity of the cells.</li>
 <li>Place neurons in a 37 &deg;C, 5% CO2 incubator overnight.</li>
 <li>Remove half the volume of media from the cells and replace with same volume of Neurobasal Feeding Media (Neurobasal Media containing B27 Supplement [1 ml / 50 ml], 0.5 mM Glutamine Solution, Penicillin (10,000 units / ml) / Streptomycin (10,000 &mu;g / ml) [250 &mu;l / 50 ml], 1 mM HEPES (Mr 238.3 g / mol).</li>
 <li>Neurons should be fed every 4 days by removing half of the old media and replacing it with the same volume of fresh Neurobasal Feeding media. Neuronal processes should begin to be visible on Day 1 (Figure 5a) and become prevalent by Day 10 (Figure 5b).</li>
</ol>
6. Representative Results
The ability to grow and culture primary neuronal cells has become an indispensible part of neuroscience. Primary cultures allow the researcher to analyze specific cellular pathways, chemical modification and treatment, target localization and growth patterns in a controlled environment. Many of these procedures utilize sophisticated methodology to visualize specific changes in cell responses. In this case, hippocampal neurons are used to study specific neuronal pathways that would prove difficult, if not impossible to analyze in the intact brain. Preparation of near homogeneous populations of neurons from specific areas of the brain is critical for studying brain function. Molecular effects in individual neurons can be instrumental in delineating higher order pathways such as memory or learning. As this protocol yields relatively pure cultures of hippocampal neurons, without the need of a feeder layer of glial cells, these neurons are easily utilized for immunofluorescence studies. However, as with all primary culture from organs containing multiple cell types, some contamination by less desired cells can occur. In isolation of neuronal cells, contamination by glial cells can be a common problem. Glial cells can be easily detected upon microscopic visualization of the culture as their morphology differs significantly from the target neurons (Figure 6). The impact of glial cell contamination will depend on the planned use of the cultures. If cells are being used for immuno-fluorescence examination, glial contamination can be nothing more than an inconvenience when trying to photograph individual neurons. However, if the neuronal cultures are to be used for biochemical analysis, any significant contamination by glial cells could cause major changes in the results. Ways to address glial cell contamination are outlined further in the Discussion.
Once neurons have been successfully isolated and grown in culture, one typical application is to examine cellular processes immuno-fluorescence techniques. As illustrated in Figure 7, organelles, such as the mitochondria, can be stained using vital dyes added to the culture media prior to fixation. Endogenous cellular proteins can be visualized from fixed cells using standard immuno-fluorescence techniques (Figure 8). Once neuronal cells are fixed, specific antibodies for proteins of interest can be introduced to the cell and these proteins can be visualized using a fluorescence microscope. Cultured neurons also provide the researcher with the means to examine individual protein effects on neuronal functions. Using a variety of techniques including DNA transfections, electroporation or viral transduction, proteins can be overexpressed in neuronal cells (Figure 9). How neural cells respond to the effects of over-expressed proteins can have direct inferences on how the brain may respond and offers the possibility of identifying cellular targets for drug treatments. The details of these types of experiments go beyond the scope of this paper but they do illustrate that cultures prepared by this technique are suitable for a wide array of down-stream applications. However, the overall simplicity of this protocol, as well as, the short time period required to prepare these neuronal cultures make this an ideal method for use in today's neuroscience laboratory.
<img alt="Figure 1" src="/files/ftp_upload/3634/3634fig1.jpg" /> 
Figure 1. Dissection of the prenatal mouse brain. The first incision is down the midline of the brain separating it into two hemispheres.
<img alt="Figure 2" src="/files/ftp_upload/3634/3634fig2.jpg" /> 
Figure 2. Location of the hippocampus in the prenatal mouse brain. The striatum is moved aside to visualize the hippocampus and is noted by the curved "kidney bean" type structure in the distal region of each hemisphere.
<img alt="Figure 3" src="/files/ftp_upload/3634/3634fig3.jpg" /> 
Figure 3. Dissociation of hippocampal tissue in trypsin solution.
<img alt="Figure 4" src="/files/ftp_upload/3634/3634fig4.jpg" /> 
Figure 4. Pasteur pipette tips used in trituration of hippocampal tissue. (a) Normal Pasteur pipette tip. (b) Fire-polished Pasteur pipette tip. Take note of the rounded edges and the approximate 50% decrease in pipette opening size.
<img alt="Figure 5" src="/files/ftp_upload/3634/3634fig5.jpg" /> 
Figure 5. Hippocampal neurons isolated using this procedure and plated in NB Media. (a) Cell growth 1 day post-plating. Neuronal processes begin to be visible during Day 1. (b) Cell growth 10 day post-plating, neurites are branched and overlapping.
<img alt="Figure 6" src="/files/ftp_upload/3634/3634fig6.jpg" /> 
Figure 6. Hippocampal neurons contaminated with glial cells grown for 7 days and stained with the organelle marker MitoTracker Red CM-H2XRos (Invitrogen #M7515) and transfected with GFP-LC3 using Lipofectamine 2000 (Invitrogen #11668019). Mitochondria are visible in all cells however only a single neuron was successfully transfected with the fluorescent construct. Contamination with glial cells makes analysis of GFP-LC3 expression in the neuronal processes difficult to visualize.
<img alt="Figure 7" src="/files/ftp_upload/3634/3634fig7.jpg" /> 
Figure 7. Hippocampal neurons grown for 7 days and stained with the organelle marker MitoTracker Red CM-H2 XRos (Invitrogen #M7513). This vital dye is used to stain active mitochondria in tissue culture cells. The cells were fixed in 4% paraformaldehyde/PBS and visualized by fluorescent microscopy. The dye itself is non-fluorescent until oxidized in the mitochondria. Active mitochondria can be seen throughout the neuronal processes.
<img alt="Figure 8" src="/files/ftp_upload/3634/3634fig8.jpg" /> 
Figure 8. Hippocampal neurons grown for 7 days, fixed with 4% paraformaldehyde/PBS and immuno-stained with monoclonal anti-tubulin &beta; antibody (Sigma #T0198). Following primary antibody, Oregon Green labeled goat-anti-mouse secondary antibody (Invitrogen #O11033) was added and fluorescence visualized by microscopy.
<img alt="Figure 9" src="/files/ftp_upload/3634/3634fig9.jpg" /> 
Figure 9. Hippocampal neuronal cultures were grown for 5 days and transfected with GFP-LC3&beta; DNA construct using Lipofectamine 2000 (Invitrogen #11668019). At Day 7, cells were fixed using 4% paraformaldehyde/PBS and aggresomes with GFP tagged LC3&beta; incorporated into their outer membrane were visualized using fluorescent microscopy. Aggresomes are located throughout the cell body and neurites and are denoted with arrows.

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No conflicts of interest declared.

Hippocampal cultures have been used in molecular biology for more than 20 years. While in principle, neuronal cultures can be made from any part of the brain, hippocampal cultures have proven to be the most popular due to the relatively simple architecture of the nerve cell population in the hippocampus7. Hippocampal cultures are typically made from late-stage embryonic tissue. This tissue is easier to dissociate and contains fewer glial cells than does mature brain tissue1. Isolation of hippocampal...

<table border="1">
<tbody>
 <tr>
 <td>Name of Reagent</td>
 <td>Vendor</td>
 <td>Catalog Number</td>
 </tr>
 <tr>
 <td>Rat Tail Collagen 1</td>
 <td>BD Biosciences</td>
 <td>354236</td>
 </tr>
 <tr>
 <td>Poly-D-lysine Solution</td>
 <td>Chemicon</td>
 <td>A-003-E</td>
 </tr>
 <tr>
 <td>Hanks Balanced Salt Solution</td>
 <td>Invitrogen</td>
 <td>14175-095</td>
 </tr>
 <tr>
 <td>Trypsin Solution (1X) 0.25%, liquid</td>
 <td>Invitrogen</td>
 <td>15050-065</td>
 </tr>
 <tr>
 <td>NeuroBasal Medium (1X) liquid</td>
 <td>Invitrogen</td>
 <td>21103-049</td>
 </tr>
 <tr>
 <td>B27 Supplement (50X) liquid</td>
 <td>Invitrogen</td>
 <td>17504-044</td>
 </tr>
 <tr>
 <td>L-Glutamine 200 mM (100X) liquid</td>
 <td>Invitrogen</td>
 <td>25030-149</td>
 </tr>
 <tr>
 <td>Penicillin (10,000 units/ml) / Streptomycin (10,000 &mu;g/ml)</td>
 <td>Invitrogen</td>
 <td>15140-148</td>
 </tr>
 <tr>
 <td>HI-Donor Horse Serum</td>
 <td>Atlanta Biologicals</td>
 <td>S12150H</td>
 </tr>
</tbody>
</table>

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.

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Isolation and Culture of Hippocampal Neurons from Prenatal Mice

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 ...

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Video: Isolation and Culture of Hippocampal Neurons from Prenatal Mice

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

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

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

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

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 ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

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

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

Isolation and Culture of Dissociated Sensory Neurons From Chick Embryos

Neurons are multifaceted cells that carry information essential for a variety of functions including sensation, motor movement, learning, and memory. Studying neurons in vivo can be challenging due to their complexity, their varied and dynamic environments, and technical limitations. For these reasons, studying neurons in vitro can prove beneficial to unravel the complex mysteries of neurons. The well-defined nature of cell culture models provides detailed control over environmental conditions and variables. Here we describe how to isolate, dissociate, and culture primary neurons from chick embryos. This technique is rapid, inexpensive, and generates robustly growing sensory neurons. The procedure consistently produces cultures that are highly enriched for neurons and has very few non-neuronal cells (less than 5%). Primary neurons do not adhere well to untreated glass or tissue culture plastic, therefore detailed procedures to create two distinct, well-defined laminin-containing substrata for neuronal plating are described. Cultured neurons are highly amenable to multiple cellular and molecular techniques, including co-immunoprecipitation, live cell imagining, RNAi, and immunocytochemistry. Procedures for double immunocytochemistry on these cultured neurons have been optimized and described here.

Cell culture models provide detailed control over environmental conditions and thus provide a powerful platform to elucidate numerous aspects of neuronal cell biology. We describe a rapid, inexpensive, and reliable method to isolate, dissociate, and culture sensory neurons from chick embryos. Details of substrata preparation and immunocytochemistry are also provided.

Neurons are multifaceted cells that carry information essential for a variety of functions including sensation, motor movement, learning, and memory. ...

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

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

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

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

Spinal Cord Neurons Isolation and Culture from Neonatal Mice

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on postnatal day 1-3. A mouse litter, usually 4-10 pups born from one breeding pair, is gathered for one experiment, and spinal cords are collected individually from each mouse after euthanasia with isoflurane. The spinal column is dissected out and then the spinal cord is released from the column. The spinal cords are then minced to increase the surface area of delivery for an enzymatic protease that allows for the neurons and other cells to be released from the tissue. Trituration is then used to release the cells into solution. This solution is subsequently fractionated in a density gradient to separate the various cells in solution, allowing for neurons to be isolated. Approximately 1-2.5 x 106 neurons can be isolated from one litter group. The neurons are then seeded onto wells coated with adhesive factors that allow for proper growth and maturation. The neurons take approximately 7 days to reach maturity in the growth and culture medium and can be used thereafter for treatment and analysis.

This study presents a technique for the isolation of neurons from WT neonatal mice. It requires the careful dissection of the spinal cord from the neonatal mouse, followed by the separation of neurons from the spinal cord tissue through mechanical and enzymatic cleavage.

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on ...

Isolation and Culture of Oculomotor, Trochlear, and Spinal Motor Neurons from Prenatal Islmn:GFP Transgenic Mice

Oculomotor neurons (CN3s) and trochlear neurons (CN4s) exhibit remarkable resistance to degenerative motor neuron diseases such as amyotrophic lateral sclerosis (ALS) when compared to spinal motor neurons (SMNs). The ability to isolate and culture primary mouse CN3s, CN4s, and SMNs would provide an approach to study mechanisms underlying this selective vulnerability. To date, most protocols use heterogeneous cell cultures, which can confound the interpretation of experimental outcomes. To minimize the problems associated with mixed-cell populations, pure cultures are indispensable. Here, the first protocol describes in detail how to efficiently purify and cultivate CN3s/CN4s alongside SMNs counterparts from the same embryos using embryonic day 11.5 (E11.5) IslMN:GFP transgenic mouse embryos. The protocol provides details on the tissue dissection and dissociation, FACS-based cell isolation, and in vitro cultivation of cells from CN3/CN4 and SMN nuclei. This protocol adds a novel in vitro CN3/CN4 culture system to existing protocols and simultaneously provides a pure species- and age-matched SMN culture for comparison. Analyses focusing on the morphological, cellular, molecular, and electrophysiological characteristics of motor neurons are feasible in this culture system. This protocol will enable research into the mechanisms that define motor neuron development, selective vulnerability, and disease.

Isolation and Culture of Oculomotor, Trochlear, and Spinal Motor Neurons from Prenatal Islmn:GFP Transgenic Mice

This work presents a protocol to yield homogeneous cell cultures of primary oculomotor, trochlear, and spinal motor neurons. These cultures can be used for comparative analyses of the morphological, cellular, molecular, and electrophysiological characteristics of ocular and spinal motor neurons.

Oculomotor neurons (CN3s) and trochlear neurons (CN4s) exhibit remarkable resistance to degenerative motor neuron diseases such as amyotrophic lateral ...

Isolation and Culture of Primary Neurons and Glia from Adult Rat Urinary Bladder

The lower urinary tract has two main functions, namely, periodic urine storage and micturition; these functions are mediated through central and peripheral neuroregulation. Although extensive research on the lower urinary tract nervous system has been conducted, most studies have focused on primary culture. This protocol introduces a method for the isolation and culture of bladder neurons and glia from Sprague&#8211;Dawley rats. In this method, the neurons and glia were incubated in a 37 &#176;C, 5% CO2 incubator for 5&#8211;7 days. As a result, they grew into mature shapes suitable for related subsequent immunofluorescence experiments. Cells were morphologically observed using an optical microscope. Neurons, synaptic vesicles, and glia were identified by &#946;-III-tubulin and MAP-2, Synapsin-1, and GFAP staining, respectively. Meanwhile, immunocytochemistry was performed on several neurotransmitter-related proteins, such as choline acetyltransferase, DYNLL2, and SLC17A9.

This protocol attempts to establish a repeatable protocol for primary neurons and glia isolation from rat bladder for further cellular experiments.

The lower urinary tract has two main functions, namely, periodic urine storage and micturition; these functions are mediated through central and ...

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high fidelity. Exploration of LTP and LTD mechanisms, single neuron dendritic computation, and experience-dependent changes in circuitry, would not have been possible without this classical preparation. However, with a few exceptions, most basic research using acute hippocampal slices has been performed using slices from rodents of relatively young ages, ~P20-P40, even though synaptic and intrinsic excitability mechanisms have a long developmental tail that reaches past P60. The main appeal of using young hippocampal slices is preservation of neuronal health aided by higher tolerance to hypoxic damage. However, there is a need to understand neuronal function at more mature stages of development, further accentuated by the development of various animal models of neurodegenerative diseases that require an aging brain preparation. Here we describe a modification to an acute hippocampal slice preparation that reliably delivers healthy slices from adult and aging mouse hippocampi. The protocol&#8217;s critical steps are transcardial perfusion and cutting with ice-cold sodium-free NMDG-aSCF. Together, these steps attenuate the hypoxia-induced drop in ATP upon decapitation, as well as cytotoxic edema caused by passive sodium fluxes. We demonstrate how to cut transversal slices of hippocampus plus cortex using a vibrating microtome. Acute hippocampal slices obtained in this way are reliably healthy over many hours of recording, and are appropriate for both field-recordings and targeted patch-clamp recordings, including targeting of fluorescently labeled neurons.

This is a protocol for acute slice preparation from adult and aging mouse hippocampi that takes advantage of transcardial perfusion and slice cutting with ice-cold NMDG-aCSF to reduce hypoxic damage to the tissue. The resulting slices stay healthy over many hours, and are suitable for long-term patch-clamp and field-recordings.

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high ...

Isolation and Culture of Chick Ciliary Ganglion Neurons

Chick ciliary ganglia (CG) are part of the parasympathetic nervous system and are responsible for the innervation of the muscle tissues present in the eye. This ganglion is constituted by a homogenous population of ciliary and choroidal neurons that innervate striated and smooth muscle fibers, respectively. Each of these neuronal types regulate specific eye structures and functions. Over the years, neuronal cultures of the chick ciliary ganglia were shown to be effective cell models in the study of muscle-nervous system interactions, which communicate through cholinergic synapses. Ciliary ganglion neurons are, in its majority, cholinergic. This cell model has been shown to be useful comparatively to previously used heterogeneous cell models that comprise several neuronal types, besides cholinergic. Anatomically, the ciliary ganglion is localized between the optic nerve (ON) and the choroid fissure (CF). Here, we describe a detailed procedure for the dissection, dissociation and in vitro culture of ciliary ganglia neurons from chick embryos. We provide a step-by-step protocol in order to obtain highly pure and stable cellular cultures of CG neurons, highlighting key steps of the process. These cultures can be maintained in vitro for 15 days and, hereby, we show the normal development of CG cultures. The results also show that these neurons can interact with muscle fibers through neuro-muscular cholinergic synapses.

Chick ciliary ganglia (CG) are part of the parasympathetic nervous system. Neuronal cultures of chick CG neurons were shown to be effective cell models in the study of nerve muscle interactions. We describe a detailed protocol for the dissection, dissociation and in vitro culture of CG neurons from chick embryos.

Chick ciliary ganglia (CG) are part of the parasympathetic nervous system and are responsible for the innervation of the muscle tissues present in the ...