These studies were supported by grants from the Natural Sciences and Engineering Research Council (HM: NSERC Discovery Grant # RGPIN-2018-06040), and Children Hospital Research Institute of Manitoba (HM: Grant # 320035), and the ALS Canada-Brain Canada Arthur J. Hudson Translational Team Grant (JK, HM).

All animal procedures were performed in accordance with institutional regulations and the Guide to the Care and Use of Experimental Animals from the Canadian Council for Animal Care and has been approved by local authorities (&ldquo;the Bannatyne Campus Animal Care Committee&rdquo;). All efforts were made to minimize the number and suffering of animals used. Adequate depth of anesthesia was confirmed by observing that there was no change in respiratory rate associated with manipulation and toe pinch or corneal reflex.
1. Preparation
NOTE: Schedule the provision of timed pregnant mice, based on the research plan, for post-conception E12&ndash;E18. Choice of the timing depends on the desired cell characteristics (see below). Prepare the cover slips and plates at least 2 days before the experiment. Poly-L-ornithine is used as a coating material to enhance cell attachment to the cover slips.
<ol>
<li>Coat the cover slips 2 days before cell isolation.
<ol>
<li>Place the round cover slips in a 24 well plate, under sterile conditions in a biosafety cabinet. Leave a gap between each cover slip to avoid any contamination.</li>
<li>Add 90 &micro;L of poly-L-ornithine (PLO, 500 &micro;g/mL) to the center of each cover slip. Close the cap and gently place the plate in a 37 &deg;C/5% CO2 incubator for 2 days. NOTE: A greater volume may spill off the cover slips during incubation. The cover slip does not need to be completely covered with PLO after placing a drop. During overnight incubation, PLO distributes equally to the edge of the cover slips.</li>
</ol>
</li>
<li>One day before cell isolation, prepare the culture medium I (DMEM/F-12 containing putrescine 100 &micro;M, sodium selenite 30 nM, L-glutamine 3.9 mM, gentamicin 3.5 &micro;g/mL, tri-iodothyronine (T3) 0.5 ng/mL, and N3 supplements [progesterone 4 &micro;M, insulin 20 &mu;g/mL, transferrin 20 mg/mL]) and seeding medium (culture medium I [without N3 and T3] containing 10% fetal bovine serum [FBS]) and store them in 4 &deg;C. Prepare the trypsin working solution (0.25%) in DMEM/F12 and keep it at 4 &deg;C. NOTE: The medium compositions are provided in Table 1.</li>
<li>On the day of cell isolation, place culture medium I and seeding medium in the incubator at 37 &deg;C. NOTE: Before starting the cerebellum isolation, make sure that all the tools and working surfaces are sterile.</li>
<li>Wash cover slips.
<ol>
<li>Take the 24 well plate out of the incubator.</li>
<li>Wash the cover slips in the 24 well plate 3x with double distilled water (DDW) in a biosafety cabinet under sterile conditions. Each time let the cover slips soak in DDW for 5 min before starting aspiration.</li>
<li>Leave the cover slips in a biosafety cabinet for at least 2 h to completely dry.</li>
</ol>
</li>
</ol>
2. Cerebellum collection
<ol>
<li>Prepare three 10 cm sterile plastic Petri dishes filled with ice-cold 1x phosphate-buffered saline (PBS), 3 Petri dishes filled with ice-cold 1x Hank&rsquo;s balanced salt solution (HBSS), and approximately 5 Petri dishes filled with ice-cold dissection medium (1x HBSS containing gentamicin 10 &mu;g/mL). Keep them on ice.</li>
<li>Anesthetize the E18 CD1 pregnant mouse with 40% isoflurane. Perform cervical dislocation on the mouse. Sterilize the abdomen with 70% ethanol solution.</li>
<li>Use a pair of scissors to make a skin incision from the pubic symphysis to the xiphoid process. Then hold the skin with forceps and open the abdominal cavity.</li>
<li>Excise the uterine horns with forceps and wash them 3x in the ice-cold 1x PBS on ice. NOTE: The following steps should all be conducted on ice to minimize the metabolic rate and prevent tissue and cell damage.</li>
</ol>
3. Dissecting the cerebellum
<ol>
<li>After the last washing step, using a pair of fine forceps separate the embryos from the uterus in 1x HBSS and transfer them to the ice-cold dissection medium. In the dissection medium, decapitate the embryos with scissors. Place the tissue in a clean dissection medium. NOTE: Using a stereomicroscope for microdissection is optional.</li>
<li>Hold the skull with fine forceps and cut through the calvarium with a pair of small scissors from the lateral aspect of the skull in a line from the foramen magnum to the external acoustic meatus and inferior border of the orbital cavity. NOTE: Taking this step exposes the cranial cavity at the level of the skull base and makes it easier to remove the brain.</li>
<li>Using a pair of fine forceps, remove the skull base and peel the skull away from the brain.</li>
<li>Carefully remove the meninges on the cerebellum, starting from the lateral surface of the middle cerebellar peduncle and pons.</li>
<li>Cut both cerebellar peduncles and separate the cerebellum from the rest of the brain (Figure 1).</li>
<li>Immediately place the collected cerebella in a sterile 15 mL conical tube filled with 14 mL of DMEM/F12 on ice.</li>
<li>Centrifuge the tube at 1,000 x g, 4 &deg;C for 1 min, 3x. Each time gently remove the supernatant with a pipette and resuspend the pellet in fresh, ice-cold DMEM/F12. NOTE: To avoid losing the samples, use the cabinet suction as little as possible.</li>
</ol>
4. Cerebellum dissociation
<ol>
<li>Add 2 mL of prewarmed (37 &deg;C) trypsin to the pellet from step 3.7 and gently pipet for adequate mixing.</li>
<li>Place the tube in a 37 &deg;C water bath for 12 min.</li>
<li>After incubation, bring the tube to a biosafety cabinet and add 10 mL of DMEM/F12 to inactivate the trypsin.</li>
<li>Centrifuge the mixture at 1,200 x g for 5 min. Discard the supernatant and resuspend the pellet in fresh DMEM/F12. Repeat 3x.</li>
<li>Prewet a sterile plastic transfer pipet with DMEM/F12.</li>
<li>After washing and the final centrifugation, add 3.5 mL of DNase working solution (1 mL of DNase I stock solution [0.05% DNase + 12 mM MgSO4 + 1x HBSS] in 500 &micro;L of heat-inactivated FBS and 2 mL of DMEM/F12) to the pellet in the same tube.</li>
<li>Triturate the tissue with a pipet at least 30x until the mixture attains a homogenous milky color.</li>
</ol>
5. Cell collection
<ol>
<li>Add 10 mL of ice-cold DMEM/F12 to the mixture.</li>
<li>Centrifuge the sample at 1,200 x g, 4 &deg;C for 5 min.</li>
<li>Carefully remove the supernatant without disturbing the pellet.</li>
<li>Remove the seeding medium from the incubator.</li>
<li>Add 500 &micro;L of prewarmed seeding medium to the pellet and mix it very well using a pipet to resuspend the cells.</li>
<li>Count the cells using a hemocytometer.</li>
<li>Dilute the cell suspension with the seeding medium to a density of 5 x 105 cells/mL.</li>
<li>Under a biosafety cabinet, add 90 &micro;L of the diluted mixture to each well at the center of the cover slip. NOTE: Do not load the particles that did not suspend in the DNase.</li>
<li>Place the plate in the incubator at 37 &deg;C for 3&minus;4 h.</li>
<li>After incubation, add 500 &micro;L of prewarmed culture medium I to each well and place the plate back into the incubator (37 &deg;C).</li>
</ol>
6. Treatment of the recovered cells
<ol>
<li>After 7 days, replace the old medium with fresh culture medium II (culture medium I supplemented with cytosine arabinoside [Ara-C, 4 &mu;M] and 100 &mu;g/mL bovine serum albumin [BSA]; see Table 1). NOTE: This step is critical to avoid growth of nonneuronal cells.</li>
<li>Monitor the culture medium once a day. If the pH changes (indicated by a marked change in color, usually yellower), remove half (almost 250 &micro;L) of the old medium from all wells and add 300 &micro;L of prewarmed culture medium I to each of them to avoid nutrient loss. NOTE: Phenol red in culture medium is a good indicator of the pH of the medium and activity of the cells. Try to minimize the exposure time of the cells to out of incubator conditions. This prevents any stress that might affect their viability.</li>
</ol>
7. Cell collection and fixation
NOTE: Depending on the experimental design, the cells can be collected at any day, at any timepoint.
<ol>
<li>Prepare a separate 24 well plate with the corresponding numeric organization and add 100 &micro;L of 4% paraformaldehyde (PFA) to each well.</li>
<li>To harvest cells on the desired days (depending on the experimental protocol), gently remove the cover slips from the wells of the original culture plate and place them in the corresponding wells of the PFA-filled plate.</li>
<li>Add PFA to the wells to completely immerse the cover slips. NOTE: To avoid cell detachment from the cover slip, do not add the PFA directly on the cover slip.</li>
<li>Keep the PFA plate at 4 &deg;C for 30&minus;120 min.</li>
<li>After incubation, restore the plate to room temperature.</li>
<li>Gently wash the cover slips 3x for 5 min with 1x PBS.</li>
<li>Proceed to the immunostaining process. NOTE: In this study, a fluorescence microscope equipped with a camera was used to capture the images and assembled into montages using an image editing software application.</li>
</ol>

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The authors have nothing to disclose.

The use of primary cultures is a well-known method applicable for all types of neurons17,18,19. In the presented protocol, we explain how to isolate cerebellar neurons and maintain their viability with optimum survival in vitro for a maximum of 3 weeks. Primary culture of cerebellar cells, which were isolated at E15&minus;E18, confirms the collection of three classes of large neurons: PCs, Golgi cells, and CNs. The cell bodies a...

For several decades, cell lines have been widely used as a high throughput tool in preclinical studies and biological research. Cost-effectiveness, fast growth, and reduction of live animal use are some benefits of using these cells. However, genetic alterations and phenotypical changes accumulate after several passages in vitro1. Misidentification of cell lines and genetic dissimilarity from primary cells can lead to irreproducible experiments and false conclusions2,3,4,5. Therefore, in spite of some similarities to differentiated cells such as neurons (e.g., neurotransmitters, ion channels, receptors, and other neuron-specific proteins), neuronal cell lines cannot replicate the full phenotype of neurons. Using mature neurons is another option; however, these cells are non-dividing postmitotic cells that are difficult to propagate in culture. Moreover, re-entry into the cell cycle may precipitate apoptosis6.
Three-dimensional (3D) cell culture, organotypic slice cultures, and organoid cultures have been developed to provide an environment in which cells can arrange into a 3D form that mimics the in vivo setting. Thus, cell-to-cell communication, migration, invasion of tumor cells into surrounding tissues, and angiogenesis can be studied7. However, additional costs of using extra cellular matrix (ECM) proteins or synthetic hydrogels as a bedding, difficulty in imaging, and compatibility with high-throughput screening instruments are considerable drawbacks of 3D cell culturing. A major disadvantage of organotypic tissue slice culture is the use of a large number of animals and the adverse effects of axotomy, which leads to inaccessibility of targets and growth factors for axons, and consequently neuronal death8.
Therefore, an alternate approach, which avoids the problems with cell lines, the difficulty of growing mature cells, and complexity of tissues, is in vitro maturation of immature primary cells. Primary cells are derived directly from human or animal tissue and dissociated using enzymatic and/or mechanical methods9. The main principles of isolation, seeding, and maintenance in culture medium are similar regardless of the tissue source. However, the trophic factors necessary to promote proliferation and maturation are highly cell specific6.
Knowing the &lsquo;birthdate&rsquo; of each cerebellar cell type is a prerequisite for designing a primary culture experiment. In general, Purkinje cells (PCs) and the neurons of the cerebellar nuclei (CN), are born before the smaller cells, including interneurons (e.g., basket, stellate cells) and granule cells. In mice, PCs emerge between embryonic day (E)10&ndash;E13, whereas CN neurons at approximately E9&ndash;E1210.
Other cerebellar neurons are born much later. For example, in mice, the Golgi subpopulation of interneurons are generated from VZ at (~E14&minus;E18) and the remaining interneurons (basket cell and stellate cells) located in the molecular layer emerge from dividing progenitor cells in the white matter between early postnatal (P)0&ndash;P711. Granule cells are generated from the external germinal zone (EGZ), a secondary germinal zone that is derived from the rostral rhombic lip and goes through terminal division after birth. But before their precursors arise from the rhombic lip from E13&ndash;E16, the cells have already migrated rostrally along the pia surface to make a thin layer of cells on the dorsal surface of the cerebellum anlage. Nonneuronal macroglial cells such as astrocytes and oligodendrocytes, which originate from the ventricular neuroepithelium, are born at E13.5&minus;P0 and P0&minus;P7 respectively11,12,13,14,15. Microglia are derived from yolk-sac primitive myeloid progenitor cells between E8&ndash;E10 and after invasion into the central nervous system can be detected in the mouse brain by E916.
The method presented in this article is based on the one originally developed by Furuya et al. and Tabata et al.17,18, which was optimized for primary culturing of Purkinje cells derived from Wistar rat cerebella. We have now adapted this method and carefully modified it to study the growth of mouse cerebellar neurons19. Unlike in our new protocol, cold dissection medium is the main washing buffer used during dissection and dissociation steps before adding seeding medium in Furuya&rsquo;s protocol17. This buffer lacks the nutrition, growth factors, and hormones (all in Dulbecco&rsquo;s modified Eagle medium:nutrient mixture F-12 [DMEM/F12]) that are necessary to support cell growth and survival during the aforementioned steps. In addition, based on our extensive experience with murine primary cerebellar cultures, we have used 500 &mu;L of culture medium in each well (instead of 1 mL) and increased the tri-iodothyronine concentration to 0.5 ng/mL, which improves growth of neuronal cells, in particular those with a Purkinje cell phenotype, and promotes the outgrowth of dendritic branches in culture. The principal method featured in this article can be broadly applied to other small rodents (e.g., squirrels and hamsters) during embryonic development and can be used to study cerebellar neurogenesis and differentiation in the various embryonic stages, which differ between species.

<table>
	<tbody>
		<tr>
			<td>Adobe Photoshop CS5 Version 12</td>
			<td>Adobe Inc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Sodium Channel (SCN)PN4 (Nav1.6)</td>
			<td>Sigma</td>
			<td>S0438</td>
			<td>3 &#956;g/mL</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Millipore Sigma</td>
			<td>A3608</td>
			<td></td>
		</tr>
		<tr>
			<td>CALB1</td>
			<td>Swant Swiss Antibodies (Polyclonal)</td>
			<td>CB38</td>
			<td>1/5000 dilution</td>
		</tr>
		<tr>
			<td>CALB1</td>
			<td>Swant Swiss Antibodies (Monoclonal)</td>
			<td>300</td>
			<td>1/1000 dilution</td>
		</tr>
		<tr>
			<td>Cytosine &#946;-D-arabinofuranoside or Cytosine Arabinoside (Ara-C)</td>
			<td>Millipore Sigma</td>
			<td>C1768</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase I from bovine pancreas</td>
			<td>Roche</td>
			<td>11284932001</td>
			<td></td>
		</tr>
		<tr>
			<td>Dressing Forceps</td>
			<td>Delasco</td>
			<td>DF-45</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM-F12)</td>
			<td>Lonza</td>
			<td>12-719F</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Cover Glasses: Circles</td>
			<td>fisher scientific</td>
			<td>12-545-81</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Gibco</td>
			<td>15710-064</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks&#8217; Balanced Salt Solution (HBSS)</td>
			<td>Gibco</td>
			<td>14185-052</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin from bovine pancreas</td>
			<td>Millipore sigma</td>
			<td>I5500, I6634, I1882, and I4011</td>
			<td></td>
		</tr>
		<tr>
			<td>Large Scissor</td>
			<td>Stoelting</td>
			<td>52134-38</td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>Gibco</td>
			<td>25030-081</td>
			<td></td>
		</tr>
		<tr>
			<td>Metallized Hemacytometer</td>
			<td>Hausser Bright-Line</td>
			<td>3100</td>
			<td></td>
		</tr>
		<tr>
			<td>Microdissection Forceps</td>
			<td>Merlan</td>
			<td>624734</td>
			<td></td>
		</tr>
		<tr>
			<td>Pattern 5 Tweezer</td>
			<td>Dixon</td>
			<td>291-9454</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Fisher BioReagents</td>
			<td>BP399-26</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-Ornithine</td>
			<td>Millipore Sigma</td>
			<td>P4638</td>
			<td></td>
		</tr>
		<tr>
			<td>Progesteron (P4)</td>
			<td>Millipore sigma</td>
			<td>P8783</td>
			<td></td>
		</tr>
		<tr>
			<td>PVALB</td>
			<td>Swant Swiss Antibodies</td>
			<td>235</td>
			<td>1/1500 dilution</td>
		</tr>
		<tr>
			<td>Samll Scissor</td>
			<td>WPI Swiss Scissors, 9cm</td>
			<td>504519</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Selenite</td>
			<td>Millipore Sigma</td>
			<td>S9133</td>
			<td></td>
		</tr>
		<tr>
			<td>Transferrin</td>
			<td>Millipore Sigma</td>
			<td>T8158</td>
			<td></td>
		</tr>
		<tr>
			<td>Tri-iodothyronine (T3)</td>
			<td>Millipore Sigma</td>
			<td>T2877</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Gibco</td>
			<td>15090-046</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss Fluorescence microscope</td>
			<td>Zeiss</td>
			<td>Z2 Imager</td>
			<td></td>
		</tr>
	</tbody>
</table>

primary culture of neurons isolated from embryonic mouse cerebellum

The use of primary cell cultures has become one of the major tools to study the nervous system in vitro. The ultimate goal of using this simplified model system is to provide a controlled microenvironment and maintain the high survival rate and the natural features of dissociated neuronal and nonneuronal cells as much as possible under in vitro conditions. In this article, we demonstrate a method of isolating primary neurons from the developing mouse cerebellum, placing them in an in vitro environment, establishing their growth, and monitoring their viability and differentiation for several weeks. This method is applicable to embryonic neurons dissociated from cerebellum between embryonic days 12&ndash;18.

Based on the different birthdates of neuronal subtypes in the cerebellum, cultures from E12&#8722;E18 mouse embryos yielded different cell types. Large projection neurons, such as CN neurons (E9&#8722;E12) and PCs (E10&#8722;E13), emerged early during cerebellar development. In mice, granule and Golgi cells arose between ~E13&#8211;E18 and underwent terminal divisions up to postnatal week 4.
Replacing old medium I with fresh culture medium II at days in vitro (DIV) 7 will eventually prevent gl...

Watch this Scientific Journal Video about Primary Culture of Neurons Isolated from Embryonic Mouse Cerebellum at JoVE.com

Primary Culture of Neurons Isolated from Embryonic Mouse Cerebellum

Conducting in vitro experiments to reflect in vivo conditions as adequately as possible is not an easy task. The use of primary cell cultures is an important step toward understanding cell biology in a whole organism. The provided protocol outlines how to successfully grow and culture embryonic mouse cerebellar neurons.

The use of primary cell cultures has become one of the major tools to study the nervous system in vitro. The ultimate goal of using this simplified model ...

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Anatomy of the Central and Peripheral Nervous System

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19.4K Views.  University of Manitoba. Conducting in vitro experiments to reflect in vivo conditions as adequately as possible is not an easy task. The use of primary cell cultures is an important step toward understanding cell biology in a whole organism. The provided protocol outlines how to successfully grow and culture embryonic mouse cerebellar neurons.

Video: Primary Culture of Neurons Isolated from Embryonic Mouse Cerebellum

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 ...

Lectin-based Isolation and Culture of Mouse Embryonic Motoneurons

Spinal motoneurons develop towards postmitotic stages through early embryonic nervous system development and subsequently grow out dendrites and axons. Neuroepithelial cells of the neural tube that express Nkx6.1 are the unique precursor cells for spinal motoneurons1. Though postmitotic motoneurons move towards their final position and organize themselves into columns along the spinal tract2,3. More than 90% of all these differentiated and positioned motoneurons express the transcription factors Islet 1/2. They innervate the muscles of the limbs as well as those of the body and the inner organs. Among others, motoneurons typically express the high affinity receptors for brain derived neurotrophic factor (BDNF) and Neurotrophin-3 (NT-3), the tropomyosin-related kinase B and C (TrkB, TrkC). They do not express the tropomyosin-related kinase A (TrkA)4. Beside the two high affinity receptors, motoneurons do express the low affinity neurotrophin receptor p75NTR. The p75NTR can bind all neurotrophins with similar but lower affinity to all neurotrophins than the high affinity receptors would bind the mature neurotrophins. Within the embryonic spinal cord, the p75NTR is exclusively expressed by the spinal motoneurons5. This has been used to develop motoneuron isolation techniques to purify the cells from the vast majority of surrounding cells6. Isolating motoneurons with the help of specific antibodies (panning) against the extracellular domains of p75NTR has turned out to be an expensive method as the amount of antibody used for a single experiment is high due to the size of the plate used for panning. A much more economical alternative is the use of lectin. Lectin has been shown to specifically bind to p75NTR as well7. The following method describes an alternative technique using wheat germ agglutinin for a preplating procedure instead of the p75NTR antibody. The lectin is an extremely inexpensive alternative to the p75NTR antibody and the purification grades using lectin are comparable to that of the p75NTR antibody. Motoneurons from the embryonic spinal cord can be isolated by this method, survive and grow out neurites.

An alternative way of isolating mouse embryonic motoneurons from the spinal cord is described. The method takes into account the fact that lectin can bind to the low affinity nerve growth factor receptor p75NTR. This lectin-based preplating allows a purification similar to that with a specific antibody against the p75NTR.

Spinal motoneurons develop towards postmitotic stages through early embryonic nervous system development and subsequently grow out dendrites and axons. ...

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 ...

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls gastrointestinal motility. A procedure for the isolation and culture of a mixed population of neurons and glia from the myenteric plexus is described. The primary cultures can be maintained for over 7 days, with connections developing among the neurons and glia. The longitudinal muscle strip with the attached myenteric plexus is stripped from the underlying circular muscle of the mouse ileum or colon and subjected to enzymatic digestion. In sterile conditions, the isolated neuronal and glia population are preserved within the pellet following centrifugation and plated on coverslips. Within 24-48 hr, neurite outgrowth occurs and neurons can be identified by pan-neuronal markers. After two days in culture, isolated neurons fire action potentials as observed by patch clamp studies. Furthermore, enteric glia can also be identified by GFAP staining. A network of neurons and glia in close apposition forms within 5 - 7 days. Enteric neurons can be individually and directly studied using methods such as immunohistochemistry, electrophysiology, calcium imaging, and single-cell PCR. Furthermore, this procedure can be performed in genetically modified animals. This methodology is simple to perform and inexpensive. Overall, this protocol exposes the components of the enteric nervous system in an easily manipulated manner so that we may better discover the functionality of the ENS in normal and disease states.

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

We demonstrate a cell culture protocol for the direct study of neuronal and glial components of the enteric nervous system. A neuron/glia mixed culture on coverslips is prepared from the myenteric plexus of adult mouse providing the ability to examine individual neuron and glia function by electrophysiology, immunohistochemical, etc.

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls ...

Primary Culture of Mouse Dopaminergic Neurons

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

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

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

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

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

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

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

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...

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 ...

Purification of Prominin-1+ Stem Cells from Postnatal Mouse Cerebellum

Most cerebellar neurons arise from two embryonic stem niches: a rhombic lip niche, which generates all the cerebellar excitatory glutamatergic neurons, and a ventricular zone niche, which generates the inhibitory GABAergic Purkinje cells, which are neurons that constitute the deep cerebellar nuclei and Bergman glia. Recently, a third stem cell niche has been described that arises as a secondary germinal zone from the ventricular zone niche. The cells of this niche are defined by the cell surface marker prominin-1 and are localized to the developing white matter of the postnatal cerebellum. This niche accounts for the late born molecular layer GABAergic interneurons along with postnatally generated cerebellar astrocytes. In addition to their developmental role, this niche is gaining translational importance in regards to its involvement in neurodegeneration and tumorigenesis. The biology of these cells has been difficult to decipher because of a lack of efficient techniques for their purification. Demonstrated here are efficient methods to purify, culture, and differentiate these postnatal cerebellar stem cells.

Purification of Prominin-1+ Stem Cells from Postnatal Mouse Cerebellum

Demonstrated here is an efficient and cost-effective method to purify, culture, and differentiate white matter stem cells from postnatal mouse cerebellum.

Most cerebellar neurons arise from two embryonic stem niches: a rhombic lip niche, which generates all the cerebellar excitatory glutamatergic neurons, ...