We thank Dr. Sean Freeman, Dr. Nathalie Sol-Foulon and Dr. Thomas Roux for valuable comments on the manuscript. This work was funded by INSERM, ICM, ARSEP Grants R13123DD, ANR R17127DD (to A.D.) and FRM fellowship, SPF20110421435 (to A.D.), FDT20170437332 (to M.T.).&#160;We thank the CELIS cell culture facility and the icm.Quant imaging platform.

All work involving animals complied with institutional policies and guidelines established by the UPMC, INSERM and the French and European Community Council Directive 86/609/EEC.
1. Preparation of Culture Medium and Culture Inserts (Hands-on Time &#8776; 10&#8211;15 min)
NOTE: Perform this step in a flow culture hood under sterile conditions
<ol>
	<li>Prepare 40 mL of culture medium, which consists of 50% BME, 25% Hank&#8217;s Balanced Salt Solution (1x), 25% heat-inactivated horse serum, supplemented with 2 mM glutamine derivative, 100 IU/mL penicillin-streptomycin and 4.5 mg/mL D-Glucose. Adjust the pH to 7.4.</li>
	<li>Filter-sterilize the medium through a 0.22 &#956;m filter adapted on a 50 mL plastic syringe. 
	NOTE: Culture medium can be stored at 4 &#176;C for at least a week.</li>
	<li>For a standard litter of 6 pups, use two sterile 6-well plates. For each plate, remove the cover and add 1 mL of culture medium per well.</li>
	<li>Place one culture insert into each individual well by holding the plastic edge with sterile forceps. Put the cover back on the plate. 
	NOTE: The cerebellar slices collected from one pup (6-8) can be dispatched on two membranes (3-4 cerebellar slices per membrane insert).</li>
</ol>
2. Preparation of Dissection Medium (Hands-on Time &#8776; 5 min)
NOTE: Perform this step in a flow culture hood under sterile conditions
<ol>
	<li>Prepare 100 mL of dissection medium, which consists of Gey&#8217;s Balanced Salt Solution supplemented with 4.5 mg/mL D-Glucose and 1x penicillin&#8211;streptomycin (100 IU/mL each).</li>
	<li>Filter-sterilize the medium through a 0.22 &#956;m filter unit. 
	NOTE: The dissection medium can be stored at 4 &#176;C for at least a month.</li>
	<li>Add 5 mL of cold dissection medium into 60 mm cell culture dishes. Prepare one 60 mm cell culture dish for each pup to be dissected.</li>
	<li>Keep the dissection dishes on ice until use.</li>
</ol>
3. Preparation of dissection material (Hands-on Time &#8776; 15 min)
<ol>
	<li>Clean the bench with 100% ethanol.</li>
	<li>Sterilize all dissection tools, a razor blade and the tissue chopper&#8217;s plastic platform with 100% ethanol.</li>
	<li>Fix the razor blade and the plastic platform on the tissue chopper and adjust the cutting thickness to 300 &#181;m.</li>
	<li>Make sure all the ethanol has evaporated before starting the dissection.</li>
</ol>
4. CerebellumDissection and Sections Preparation (Hands-on Time &#8776; 15&#8211;20 min per animal)
<ol>
	<li>Place one of the 60 mm cell culture dishes containing the ice-cold dissection medium under the binocular microscope and remove the top plate.</li>
	<li>Swab the head fur with 70% ethanol (carefully avoiding touching the eyes of the animal) and decapitate the animal using large sharp scissors, according to the approved procedure. 
	NOTE: The cerebellar slices are generated from P9-P10 mice without any specific sex or strain bias.</li>
	<li>Use small scissors to cut the skin along the midline of the head starting from the neck and going up to the mouse muzzle.</li>
	<li>To hold the head and expose the skull, fold back the skin ventrally under the head, and pinch the two folds of skin between one&#8217;s fingers.</li>
	<li>Insert the small scissors gently into the foramen magnum and cut the skull by making one lateral incision towards the side and then cut all around the head skull. Keep the tip of the scissors as parallel and close to the skull as possible while cutting, to avoid damaging the brain tissue.</li>
	<li>Retrieve the dorsal part of the skull using fine-straight forceps. While carefully lifting the dorsal part of the skull, cut the adhering meninges (translucent irrigated membrane surrounding the brain tissue) if needed with small scissors to avoid damaging the brain tissue.</li>
	<li>Carefully introduce fine-straight forceps (or alternatively use a spatula) between the ventral skull and the brain, gently flip out the brain and cut the optic and trigeminal nerves with small scissors.</li>
	<li>Help the brain to drop by gravity into the 60 mm cell culture dish containing ice-cold dissection medium by turning the head upside down just above the dish and release the last adhesions with small scissors if needed.</li>
	<li>Using fine forceps, orientate the brain with the dorsal side facing the researcher, and the ventral side lying on the bottom of the dish.</li>
	<li>Under the binocular microscope, use the fine-straight forceps to immobilize the brain on the forebrain side and avoid touching the hindbrain, as it could get damaged. Separate the hindbrain from the rest of the brain (fore- and midbrain, see schematic on Figure 1Aa), using fine-straight forceps. To separate the cerebellum from the rest of the hindbrain, use the fine forceps to cut the cerebellar peduncles underneath the cerebellum (see schematic on Figure 1Aa&#8217;).</li>
	<li>Once the cerebellum is isolated, carefully tear away the meninges using fine-straight forceps (see schematic on Figure 1Aa&#8217;).</li>
	<li>Hold the cerebellum gently with the fine curved forceps and place it with the dorsal side up onto the plastic platform perpendicularly to the chopper razor blade (see schematic on Figure 1Bb).</li>
	<li>Aspirate any excess of dissection medium around the cerebellum using a sterile thin-end pipette tip adapted on a 1 mL pipette (see schematic on Figure 1Bb).</li>
	<li>Ensure that the cerebellum lays flat, but is not stretched, once placed onto the platform to get optimal sagittal slices during sectioning (see schematic on Figure 1Bb).</li>
	<li>Slice 300 &#956;m-thick sagittal sections of the cerebellum with the tissue chopper (see schematic on Figure 1Bb&#8217;). The force and speed of the blade have to be optimized on site.</li>
	<li>Gently add a drop of dissection medium onto the sliced cerebellum. Using a wide-bore pipette tip placed onto a 1 mL pipette, slowly aspirate the sliced cerebellum and transfer it back into the 60 mm cell culture dish containing ice-cold dissection medium (see schematic on Figure 1Bb&#8217;&#8217;).</li>
	<li>Clean the tissue chopper&#8217;s plastic platform with 100% ethanol between each cerebellum slicing. Let the ethanol evaporate before placing the next cerebellum onto the platform.</li>
	<li>Use two fine-straight forceps (or alternatively titanium needles) to separate individual slices and select 6-8 slices from the vermis (see schematic on Figure 1Cc and 1Cc&#8217;).</li>
	<li>Take a 6-well plate with culture inserts and transfer up to 4 selected slices onto one culture insert along with some dissection medium, using a wide-bore pipette tip adapted on a 1 mL pipette (see schematic on Figure 1Cc&#8217;).</li>
	<li>Place the slices in the middle of the culture insert using the dissection medium or forceps to push and pull them gently into the right location, avoiding them to be in contact with each other (see schematic on Figure 1Cc&#8217;).</li>
	<li>Remove any excess of dissection medium around the slices using a thin-end pipette tip. Ensure that the slices lay flat on the membrane (see schematic on Figure 1Cc&#8217;).</li>
	<li>Place the 6-well plate into the incubator immediately after having added the slices on the membrane.</li>
</ol>
5. Slices Culture and Demyelination (Hands-on Time &#8776; 15&#8211;20 min)
NOTE: Perform this step in a flow culture hood under sterile conditions
<ol>
	<li>Replace the culture medium with 1 mL per well of pre-warmed and buffered fresh culture medium every 3 days after slice preparation.</li>
	<li>For demyelination, remove all culture medium below the culture inserts after 6 days in vitro (DIV) and replace it with 1 mL per well of pre-warmed fresh culture medium containing 0.5 mg/mL LPC. Incubate for 15-17 h at 37 &#176;C, 5% CO2.</li>
	<li>After incubation, wash the inserts by placing them in a 25 mm petri dish containing 1 mL of pre-warmed culture medium. 
	NOTE: The inserts should be in contact with the medium, but not be covered by it.</li>
	<li>Immediately transfer the culture inserts in a new 6-well plate containing fresh pre-warmed culture medium.</li>
	<li>Replace the culture medium with 1 mL of pre-warmed fresh culture medium every 2-3 days until slices fixation.</li>
</ol>
6. Immunohistochemistry (Hands-on Time &#8776; 1:30&#8211;2 h)
NOTE: Carry out the steps 6.1. to 6.3. under a fume hood. Avoid exposition to paraformaldehyde (PFA) and refer to the product safety datasheet for adequate manipulation and protection.
<ol>
	<li>Use forceps to lift each culture inserts by holding the plastic edge and remove the culture medium beneath the membrane inserts from each well.</li>
	<li>Fix the cerebellar slices for 30 min by adding 2 mL of 4% PFA in 1x PBS pH 7.4 onto the membrane inserts. 
	NOTE: The time point of fixation depends on the stage of myelination studied: 4 DIV corresponds to the onset of myelination, 7 DIV to fully myelinated slices. In case of LPC treatment, 9 DIV corresponds to the peak of demyelination, 11 DIV to the onset of remyelination and 14 DIV to fully remyelinated slices.</li>
	<li>Wash the inserts with the slices three times with 2 mL 1x PBS for 10 min.</li>
	<li>Separate the slices from the membrane inserts under the binocular microscope using a 25x to 30x magnification, by gently pushing them using a scalpel or a brush. Alternatively, to avoid the risk of damaging the slices while detaching them from the membrane, use a scalpel to cut the membrane with the slices still attached.</li>
	<li>Transfer the floating slices into the wells of a 4-well plate containing 1x PBS, using a brush.</li>
	<li>Aspirate the PBS from each well and incubate the slices in pre-cooled 100% ethanol at -20 &#176;C for 15 to 20 min. This step facilitates antibody penetration in myelinated fibers.</li>
	<li>Aspirate the 100% ethanol and wash once briefly with 1x PBS, then twice in 1x PBS for 10 min at room temperature.</li>
	<li>Aspirate the PBS and incubate the slices for 30 to 45 min with a solution containing 1x PBS, 5% NGS, 0.3% non-ionic detergent at room temperature to block non-specific antibody fixation sites.</li>
	<li>Aspirate the blocking solution. No washing is necessary.</li>
	<li>Incubate the slices overnight with the primary antibodies diluted in blocking solution at 4 &#176;C. To visualize the myelin on cerebellar organotypic slices, use MBP (Chicken, 1/150 or Mouse IGg2b, Smi99, 1/200) or PLP (Rat, hybridoma, 1/5 to 1/10) antibodies. 
	NOTE: To reveal the expression of more than one antigen in the same slice, perform a double- or triple staining procedure (see Figure 1 as an example). To perform the incubation simultaneously, ensure primary antibodies are produced in different species. Then, use their corresponding secondary antibodies coupled to different fluorophores (see Table of Materials for nodal and paranodal marker18 antibodies).</li>
	<li>On the second day, wash the slices three times in 1x PBS for 10 min.</li>
	<li>Incubate for 3h with the secondary antibodies diluted in blocking solution (1/500 dilution) in the dark at room temperature. 
	NOTE: Optimization of antibody dilution and incubation time might be required when using other antibodies than the ones described here.</li>
	<li>Wash the slices three times in 1x PBS for 10 min in the dark.</li>
	<li>Under a binocular microscope, mount the slices onto a glass slide. Place 100 &#181;L of 1x PBS on the slide and transfer the slices into the PBS using a brush. Unfold the slices if needed and flatten them on the slide. Remove any excess of PBS. In case the immunolabeling is performed with the slices still attached to membranes, mount with the slices facing the coverslip, with the membrane on the glass slide.</li>
	<li>Place a drop of mounting medium directly onto the glass coverslip and gently cover the slices. Mounted slides can be stored for several months at 4 &#176;C in the dark.</li>
</ol>

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None of the authors have competing interests or conflicting interests.

Here, we detail a protocol to generate an ex vivo model corresponding to the mouse cerebellar organotypic slice cultures, adapted from previously published methods15,16,19 and the subsequent myelin immunostaining of these preparations. This strategy offers the possibility to visualize myelin components with a high-resolution in both healthy and pathological states.
Cerebellar organotypic slice cultures taken from 10-day-old mice are a well-established experimental model to investigate molecular and cellular mechanisms underlying myelination and remyelination processes, as it allows to reproduce most anatomical and functional features of the corresponding tissue in vivo including not only the preservation of a well-defined cellular architecture but also the maturation timeline17. Furthermore, cerebellar slices have a preserved architectural organization after two weeks in vitro&#160;(Figure 1C). The transgenic PLP-GFP strain is an alternative, allowing, in particular, to observe the myelination status under a fluorescent binocular microscope on live slices, which is particularly convenient for demyelination-remyelination studies (Figure 1C).
These preparations are of particular interest as a fast approach to assess myelination rate with the developed automated quantification, including western-blot analysis20, CNPase assay16 or quantification of PLP/Calbindin (or MBP/Calbindin) staining17,21, thus representing a high-throughput system allowing for the testing of pharmacological drugs in demyelinating disorders22,23.
One of the main critical issues when performing these experiments might be related to the quality of the cerebellar slices. Firstly, the age of mice is of importance for cerebellar organotypic slice cultures. The survival of Purkinje cells, the only myelinated neuron in cerebellar cultures, is compromised if cultures are prepared from P2 to P7 rodents or with donors older than P1317. More generally, organotypic brain slice cultures are difficult to obtain from adults animals. Secondly, the duration of tissue processing during dissection is critical, as a dissection step of more than 15-20 minutes per animal leads to a decline in the survival rate of the slices. Furthermore, when selecting the slices to be cultured, it is best to avoid using the ones from either end of the cerebellum due to the poor survival of neurons and resultant scarcity of myelinated fibers.
Other parameters such as the stability of temperature and CO2 concentration while in culture are determining factors to ensure slice health. Lastly, the quality of the slices is influenced by the composition of the horse serum, which cannot be controlled by the users. Therefore, testing several batches of serum is highly recommended.
To obtain an adequate labeling, tissue fixation is a key step. In order to facilitate the penetration of the antibodies to reach myelin antigens, as myelin is a very compact structure, slices pre-treatment with absolute ethanol is advised following fixation. A similar protocol of staining can further be applied to in vivo fixed tissues, though Triton concentration may need to be adapted to the thickness of vibratome or cryostat generated in vivo sections. The availability of non-commercial antibodies should also be considered. Preparing slices from transgenic mice such as PLP-GFP24 or CNP-GFP25 thus represents an alternative to visualize myelin and oligodendrocytes processes.
In addition to the immunochemical methods described in this protocol, other approaches are commonly used to study myelin architecture. Complementary techniques also exist to investigate myelin ultrastructure, with electron microscopy being the gold standard26. Other techniques exist, such as label-free approaches including optical coherence tomography27,28, Raman scattering28,29, third-harmonic generation30 or spectral confocal reflectance microscopy31; these approaches have the additional advantage of enabling the observation of dynamic cellular processes. However, some of these techniques are complex and further require state-of-the-art microscopy set-ups. For these reasons, myelin study by immunofluorescence labeling remains a standard approach to investigate myelination and its defects in various developmental or acquired diseases and their models, as well as to assess the potential of future therapeutic treatments. It can further be used to study myelination in multiple contexts, such as learning and plasticity, as well as aging, depression, or autism in which myelin is known to be altered32.

Neurons are highly polarized cells, which comprise a somato-dendritic compartment that receives inputs from its environment and an axon that ensures the generation and propagation of electrical impulses to other cells. Rapid propagation and timely delivery of information is essential for the proper functioning of the nervous system. In vertebrates, it is facilitated by myelination, which allows increasing axonal conduction velocity1. Myelin is a specialized structure formed by compacted layers of plasma membrane generated by the myelinating glia, namely oligodendrocytes in the central nervous system (CNS) and Schwann cells in the peripheral nervous system (PNS). Both in the CNS and the PNS, axoglial interactions drive the formation of specialized axonal domains: the nodes of Ranvier and their surrounding domains, the paranodes, and juxtaparanodes2. The axonal segments insulated by myelin, or internodes, alternate with the nodes of Ranvier, which correspond to small unmyelinated domains enriched in voltage-gated sodium channels (Nav). The high concentration and rapid activation of Nav channels at the nodes of Ranvier allow the regeneration of action potentials, and together with the insulating properties of the myelin sheaths, ensure the efficient and fast saltatory conduction of the nerve impulse along the axon3.
In addition to its role in accelerating the conduction velocity of the nerve impulse, myelinating glia provides metabolic support to the axon, preserving its long-term integrity and participating in its survival4,5. Furthermore, it has become clear in recent years that myelin is dynamically modulated throughout life, thus presumably participating in the regulation and plasticity of various nervous system functions. Adjustments of the distribution, number, length, and thickness of myelin sheaths along axons might thus represent a novel way to finely tune various networks6,7,8. Therefore, the evolutionary acquisition of myelin is a key process for sensory, motor and cognitive functions and the perturbation of the interaction between axons and glia is increasingly considered as contributing to the developmental or acquired neurological diseases9.
Myelin composition has been characterized, with the specific feature of a high proportion of lipids (70%) compared to proteins (30%) in contrast to other cellular membranes10. However, unlike myelin lipids, most of myelin proteins are specific to myelin, including myelin basic protein (MBP), proteolipid protein (PLP), 2&#39;,3&#39;-cyclic nucleotide 3&#39;-phosphodiesterase (CNP), myelin-associated glycoprotein (MAG), myelin-oligodendrocyte glycoprotein (MOG), PMP-22 and P010. Various histological methods to stain myelin exist based on its lipid composition, such as Luxol fast blue11, Sudan Black B12, Baker&#39;s acid hematin method13, as well as silver staining14. Nevertheless, these approaches do not always allow for an adequate contrast and resolution to visualize individual fibers. An alternative approach to detect myelin is through immunohistochemistry directed against myelin proteins. Various antibodies target myelin-specific antigens with a high specificity and can be used routinely to detect myelinated structures. The antibody-antigen interaction can be further revealed using a secondary antibody coupled to a fluorophore directed against the primary antibody and visualized with adequate fluorescence microscopy. Here, we describe an immunochemical protocol to stain myelin on ex vivo cerebellar slices, a model which allows for a good preservation of the nervous tissue architecture. In addition, the organization and size of the Purkinje cells (the sole myelinated neuron of the cerebellum) make them a classical model for electrophysiological studies and they are similarly ideal to perform fixed or live-imaging studies.
The cerebellar slices are generated from P9-P10 mice, a time corresponding to the early onset of Purkinje cell myelination, a process that is mostly achieved by one week ex vivo (6-7 days in vitro, DIV)15. Furthermore, this model is adapted to investigate demyelinating disorders such as multiple sclerosis (MS), as an extensive demyelination can be induced in cerebellar slices using the myelinotoxic compound lysophosphatidylcholine (or lysolecithin, LPC), which is followed by a spontaneous remyelination16,17. Endogenous remyelination takes place from two days after LPC removal from the culture medium and is almost complete a week post treatment.
The completion of this protocol takes approximatively 3 weeks, including half a day for cerebellar slice cultures preparation, a week to obtain fully myelinated slices, followed by 2 days to reach the peak of demyelination and another week for their full remyelination. In addition, immunohistochemistry can be completed in 2 days. The protocol described here is adapted to a standard litter of 6 mice pups and needs to be adapted regarding the number of animals used for the planned experiment.

<table>
	<tbody>
		<tr>
			<td>BME medium</td>
			<td>ThermoFisher Scientific</td>
			<td>41010026</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#8217;s Balanced Salt Solution (10X HBSS)</td>
			<td>ThermoFisher Scientific</td>
			<td>14180046</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutaMAX (100X)</td>
			<td>ThermoFisher Scientific</td>
			<td>35050038</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat-inactivated Horse Serum</td>
			<td>ThermoFisher Scientific</td>
			<td>26050088</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin&#8211;Streptomycin (10.000 IU/mL)</td>
			<td>ThermoFisher Scientific</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Gey&#8217;s Balanced Salt Solution</td>
			<td>Sigma Aldrich</td>
			<td>G9779-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Glucose Solution (45%)</td>
			<td>Sigma Aldrich</td>
			<td>G8769-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Lysophosphatidylcholine (LPC)</td>
			<td>Sigma Aldrich</td>
			<td>L4129-100MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde (PFA)</td>
			<td>Electron Microscopy Sciences</td>
			<td>15714</td>
			<td></td>
		</tr>
		<tr>
			<td>Absolute ethanol (100% ethanol)</td>
			<td>VWR Chemicals</td>
			<td>20821.330</td>
			<td>Cooled at -20&#176;C</td>
		</tr>
		<tr>
			<td>Triton&#174; X-100</td>
			<td>Sigma Aldrich</td>
			<td>X100-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>10% Normal Goat Serum (NGS)</td>
			<td>ThermoFisher Scientific</td>
			<td>500622</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffer Solution</td>
			<td>EuroMEDEX</td>
			<td>ET330-A</td>
			<td>pH 7.4</td>
		</tr>
		<tr>
			<td>Anti-GFP Antibody (Polyclonal, Chicken)</td>
			<td>Merck Millipore</td>
			<td>06-896</td>
			<td>Dilution 1/300</td>
		</tr>
		<tr>
			<td>Anti-Myelin Basic Protein (MBP) Antibody (Polyclonal, Chicken)</td>
			<td>Merck Millipore</td>
			<td>AB9348</td>
			<td>Dilution 1/150</td>
		</tr>
		<tr>
			<td>Anti-Myelin Basic Protein (MBP) Antibody (Monoclonal, Mouse IgG2b)</td>
			<td>Merck Millipore</td>
			<td>NE1019</td>
			<td>Dilution 1/200</td>
		</tr>
		<tr>
			<td>Anti-PLP Antibody (Rat, Hybridoma)</td>
			<td>Gift from Dr. K. Ikenaka; Okasaki, Japan</td>
			<td></td>
			<td>Dilution 1/5 to 1/10</td>
		</tr>
		<tr>
			<td>Anti-Sodium Channel, Pan Antibody (Monoclonal, Mouse IgG1, clone K58/35)</td>
			<td>Sigma Aldrich</td>
			<td>S8809</td>
			<td>Dilution 1/150</td>
		</tr>
		<tr>
			<td>Anti-Caspr Antibody (Polyclonal, Rabbit)</td>
			<td>Abcam</td>
			<td>ab34151</td>
			<td>Dilution 1/500</td>
		</tr>
		<tr>
			<td>Goat Secondary Antibodies conjugated to Alexa Fluor 488, 594, 647 or 405</td>
			<td>ThermoFisher Scientific</td>
			<td></td>
			<td>Dilution 1/500</td>
		</tr>
		<tr>
			<td>Fluoromount</td>
			<td>SouthernBiotech</td>
			<td>0100-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue chopper</td>
			<td>McIlwain</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Razor blades</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Large scissors</td>
			<td>F.S.T</td>
			<td>14101-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Small scissors</td>
			<td>F.S.T</td>
			<td>91500-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine-straight forceps</td>
			<td>F.S.T</td>
			<td>91150-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved-fine forceps</td>
			<td>F.S.T</td>
			<td>11297-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture dishes (60-mm and 100-mm)</td>
			<td>TPP</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4-, 6-well culture plates</td>
			<td>TPP</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Millicell culture inserts (0,4 &#181;m, 30-mm diameter)</td>
			<td>Merck Millipore</td>
			<td>PICM0RG50</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture incubator</td>
			<td></td>
			<td></td>
			<td>37&#176;C, 5% CO2</td>
		</tr>
		<tr>
			<td>Fine-end pipette tips</td>
			<td>Dutscher</td>
			<td>134000CL</td>
			<td></td>
		</tr>
		<tr>
			<td>Wide-bore pipette tips</td>
			<td>ThermoFisher Scientific</td>
			<td>2079G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile syringe</td>
			<td>Terumo Europe</td>
			<td></td>
			<td>20 or 50 mL</td>
		</tr>
		<tr>
			<td>Sterile syringe filters</td>
			<td>Terumo Europe</td>
			<td></td>
			<td>0.22 &#181;m</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Swann-Morton</td>
			<td>0510</td>
			<td></td>
		</tr>
		<tr>
			<td>Brush</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>RS France</td>
			<td></td>
			<td>76 x 26 x 1.1 mm</td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td>RS France</td>
			<td></td>
			<td>22 x 22 mm</td>
		</tr>
		<tr>
			<td>Kimtech Sciences Tissue Wipers</td>
			<td>Kimberly-Clark Professional</td>
			<td>5511</td>
			<td></td>
		</tr>
		<tr>
			<td>Binocular microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

preparation and immunostaining of myelinating organotypic cerebellar slice cultures

In the nervous system, myelin is a complex membrane structure generated by myelinating glial cells, which ensheathes axons and facilitates fast electrical conduction. Myelin alteration has been shown to occur in various neurological diseases, where it is associated with functional deficits. Here, we provide a detailed description of an ex vivo model consisting of mouse organotypic cerebellar slices, which can be maintained in culture for several weeks and further be labeled to visualize myelin.

Examples of representative myelin immunostainings in organotypic cerebellar slices obtained from P9-P10 C57black6 wild-type (WT) (Figure 2A), as well as PLP-GFP transgenic mice (Figure 2B), together with Purkinje cells staining. Cerebellar slices myelinate from the white matter tracks region of the slices towards the periphery of the folia and myelination of the Purkinje cells is mostly achieved after 6 to 7 DIV. At 7 DIV, the induction of a full demyelination is possible through LPC treatment (Figure 2Ci-ii). Following demyelination, the slices spontaneously remyelinate and are fully remyelinated 6-7 days after the peak of demyelination (Figure 2Ciii).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59163/59163fig1.jpg" /> 
Figure 1: Illustration of cerebellum slices generation. The dissection is divided in three main steps. (A) The cerebellum is isolated and the meninges removed (steps 4.10 and 4.11). (B) The cerebellum is then transferred to the chopper platform and sliced (steps 4.12 to 4.16). (C) Lastly, the slices are isolated from the vermis and placed on a membrane insert. The dissection medium transferred with the slices is removed and the 6-well plate is placed into the incubator (steps 4.18 and 4.22). <a href="https://www.jove.com/files/ftp_upload/59163/59163fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59163/59163fig2.jpg" /> 
Figure 2: Example of myelinated cerebellar slices cultured ex vivo. Image stacks (with orthogonal projection) were obtained using an upright confocal microscope following free floating immunolabeling and flat mounting as described in the protocol. The slices are robustly myelinated at 11 DIV (A, PLP; B, GFP in green)&#160;and the axonal domains are assembled as observed in vivo with nodes of Ranvier enriched in voltage-gated sodium channels (Nav, in red) flanked by the paranodal axoglial junction domain (Caspr, in white) in C57black6 WT (A) as well as PLP-GFP (B) transgenic mice slices. (C) The cerebellar cortex architecture is preserved in the cultured slices, as observed on cerebellar slices from PLP-GFP mice. (i) Purkinje cells axon (Calbindin, Calb, in blue) are robustly myelinated (GFP, in green) after one week in culture. (ii) LPC treatment fully demyelinates the slices, which remyelinate spontaneously and are fully remyelinated 6 days post demyelination (iii, 14 DIV). Scale bars = 20 &#181;m (A, B); 100 &#181;m (C). <a href="https://www.jove.com/files/ftp_upload/59163/59163fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures at JoVE.com

Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

Here we present a method to prepare organotypic slice cultures from mouse cerebellum and myelin sheath staining by immunohistochemistry suitable for investigating mechanisms of myelination and remyelination in the central nervous system.

In the nervous system, myelin is a complex membrane structure generated by myelinating glial cells, which ensheathes axons and facilitates fast electrical ...

preparation-immunostaining-myelinating-organotypic-cerebellar-slice

Nanyang Technological University

Research

JoVE Journal

Neuroscience

10.9K Views.  Sorbonne Université, Inserm, CNRS, Institut du Cerveau et de la Moelle épinière, ICM-GH Pitié-Salpétrière. Here we present a method to prepare organotypic slice cultures from mouse cerebellum and myelin sheath staining by immunohistochemistry suitable for investigating mechanisms of myelination and remyelination in the central nervous system.

Video: Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

The Organotypic Hippocampal Slice Culture Model for Examining Neuronal Injury

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of the hippocampus is relatively preserved 1. The organotypic culture system allows for the examination of neuronal, astrocytic and microglial effects, but as an ex vivo preparation, does not address effects of blood flow, or recruitment of peripheral inflammatory cells. To that end, this culture method is frequently used to examine excitotoxic and hypoxic injury to pyramidal neurons of the hippocampus, but has also been used to examine the inflammatory response. Herein we describe the methods for generating hippocampal slice cultures from postnatal rodent brain, administering toxic stimuli to induce neuronal injury, and assaying and quantifying hippocampal neuronal death.

The organoptypic hippocampal slice culture model is an in vitro model used to examine neuronal injury in a variety of paradigms. In this article, we describe the methods for generating slice cultures and quantifying neuronal injury.

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of ...

Slice Preparation, Organotypic Tissue Culturing and Luciferase Recording of Clock Gene Activity in the Suprachiasmatic Nucleus

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the anterior hypothalamus. The clock is directly synchronized by light via the retina and optic nerve. Circadian oscillations are generated by interacting negative feedback loops of a number of so called "clock genes" and their protein products, including the Period (Per) genes. The core clock is also dependent on membrane depolarization, calcium and cAMP 1. The SCN shows daily oscillations in clock gene expression, metabolic activity and spontaneous electrical activity. Remarkably, this endogenous cyclic activity persists in adult tissue slices of the SCN 2-4. In this way, the biological clock can easily be studied in vitro, allowing molecular, electrophysiological and metabolic investigations of the pacemaker function.
The SCN is a small, well-defined bilateral structure located right above the optic chiasm 5. In the rat it contains ~8.000 neurons in each nucleus and has dimensions of approximately 947 &mu;m (length, rostrocaudal axis) x 424 &mu;m (width) x 390 &mu;m (height) 6. To dissect out the SCN it is necessary to cut a brain slice at the specific level of the brain where the SCN can be identified. Here, we describe the dissecting and slicing procedure of the SCN, which is similar for mouse and rat brains. Further, we show how to culture the dissected tissue organotypically on a membrane 7, a technique developed for SCN tissue culture by Yamazaki et al. 8. Finally, we demonstrate how transgenic tissue can be used for measuring expression of clock genes/proteins using dynamic luciferase reporter technology, a method that originally was used for circadian measurements by Geusz et al. 9. We here use SCN tissues from the transgenic knock-in PERIOD2::LUCIFERASE mice produced by Yoo et al. 10. The mice contain a fusion protein of PERIOD (PER) 2 and the firefly enzyme LUCIFERASE. When PER2 is translated in the presence of the substrate for luciferase, i.e. luciferin, the PER2 expression can be monitored as bioluminescence when luciferase catalyzes the oxidation of luciferin. The number of emitted photons positively correlates to the amount of produced PER2 protein, and the bioluminescence rhythms match the PER2 protein rhythm in vivo 10. In this way the cyclic variation in PER2 expression can be continuously monitored real time during many days. The protocol we follow for tissue culturing and real-time bioluminescence recording has been thoroughly described by Yamazaki and Takahashi 11.

The procedure of preparing slices containing the adult mouse hypothalamic suprachiasmatic nucleus (SCN), and a rapid way to culture the SCN tissue in organotypic culture condition, are reported. Further, the measurement of oscillatory clock gene protein expression using dynamic luciferase reporter technology is described.

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the ...

Organotypic Hippocampal Slice Cultures

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the strength of their connections after brief periods of strong activation. This phenomenon, known as long-term potentiation (LTP) can last for hours or days and has become the best candidate mechanism for learning and memory 2. In addition, the well defined anatomy and connectivity of the hippocampus 3 has made it a classical model system to study synaptic transmission and synaptic plasticity4.
As our understanding of the physiology of hippocampal synapses grew and molecular players became identified, a need to manipulate synaptic proteins became imperative. Organotypic hippocampal cultures offer the possibility for easy gene manipulation and precise pharmacological intervention but maintain synaptic organization that is critical to understanding synapse function in a more naturalistic context than routine culture dissociated neurons methods.
Here we present a method to prepare and culture hippocampal slices that can be easily adapted to other brain regions. This method allows easy access to the slices for genetic manipulation using different approaches like viral infection 5,6 or biolistics 7. In addition, slices can be easily recovered for biochemical assays 8, or transferred to microscopes for imaging 9 or electrophysiological experiments 10.

We describe a method to prepare organotypic hippocampal slices that can be easily adapted to other brain regions. Brain slices are laid on porous membranes and culture media is allowed to form an interface. This method preserves the gross architecture of the hippocampus for up to 2 weeks in culture.

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the ...

Organotypic Cerebellar Cultures: Apoptotic Challenges and Detection

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. Since then, the technique was developed further and currently there are many different ways to prepare organotypic cultures. The method presented here was adapted from the one described by Stoppini et al. for the preparation of the slices and from Gogolla et al. for the staining procedure 3,4.
A unique feature of this technique is that it allows you to study different parts of the brain such as hippocampus or cerebellum in their original structure, providing a big advantage over dissociated cultures in which all the cellular organization and neuronal networks are disrupted. In the case of the cerebellum it is even more advantageous because it allows the study of Purkinje cells, extremely difficult to obtain as dissociated primary culture. This method can be used to study certain developmental features of the cerebellum in vitro, as well as for electrophysiological and pharmacological experiments in both wild type and mutant mice.
The method described here was designed to study the effect of apoptotic stimuli such as Fas ligand in the developing cerebellum, using TUNEL staining to measure apoptotic cell death. If TUNEL staining is combined with cell type specific markers, such as Calbindin for Purkinje cells, it is possible to evaluate cell death in a cell population specific manner. The Calbindin staining also serves the purpose of evaluating the quality of the cerebellar cultures.

This method describes the generation of organotypic cerebellar cultures and the effect of certain apoptotic stimuli on the viability of different cerebellar cell types.

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. ...

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

Derivation of Enriched Oligodendrocyte Cultures and Oligodendrocyte/Neuron Myelinating Co-cultures from Post-natal Murine Tissues

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications for understanding the pathogenesis of demyelinating diseases such as Multiple Sclerosis (MS). Cellular development is commonly studied with primary cell culture models. Primary cell culture facilitates the evaluation of a given cell type by providing a controlled environment, free of the extraneous variables that are present in vivo. While OL cultures derived from rats have provided a vast amount of insight into OL biology, similar efforts at establishing OL cultures from mice has been met with major obstacles. Developing methods to culture murine primary OLs is imperative in order to take advantage of the available transgenic mouse lines. 
Multiple methods for extraction of OPCs from rodent tissue have been described, ranging from neurosphere derivation, differential adhesion purification and immunopurification 1-3. While many methods offer success, most require extensive culture times and/or costly equipment/reagents. To circumvent this, purifying OPCs from murine tissue with an adaptation of the method originally described by McCarthy &amp; 
de Vellis 2 is preferred. This method involves physically separating OPCs from a mixed glial culture derived from neonatal rodent cortices. The result is a purified OPC population that can be differentiated into an OL-enriched culture. This approach is appealing due to its relatively short culture time and the unnecessary requirement for growth factors or immunopanning antibodies. 
While exploring the mechanisms of OL development in a purified culture is informative, it does not provide the most physiologically relevant environment for assessing myelin sheath formation. Co-culturing OLs with neurons would lend insight into the molecular underpinnings regulating OL-mediated myelination of axons. For many OL/neuron co-culture studies, dorsal root ganglion neurons (DRGNs) have proven to be the neuron type of choice. They are ideal for co-culture with OLs due to their ease of extraction, minimal amount of contaminating cells, and formation of dense neurite beds. While studies using rat/mouse myelinating xenocultures have been published 4-6, a method for the derivation of such OL/DRGN myelinating co-cultures from post-natal murine tissue has not been described. Here we present detailed methods on how to effectively produce such cultures, along with examples of expected results. These methods are useful for addressing questions relevant to OL development/myelinating function, and are useful tools in the field of neuroscience.

This article describes methods to derive enriched populations of murine oligodendrocyte precursor cells (OPCs) in primary culture, which differentiate to produce mature oligodendrocytes (OLs). In addition, this report describes techniques to produce murine myelinating co-cultures by seeding mouse OPCs onto a neurite bed of mouse dorsal root ganglion neurons (DRGNs).

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications ...

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

The mouse is an excellent model organism to study mammalian brain development due to the abundance of molecular and genetic data. However, the developing mouse brain is not suitable for easy manipulation and imaging in vivo since the mouse embryo is inaccessible and opaque. Organotypic slice cultures of embryonic brains are therefore widely used to study murine brain development in vitro. Ex-vivo manipulation or the use of transgenic mice allows the modification of gene expression so that subpopulations of neuronal or glial cells can be labeled with fluorescent proteins. The behavior of labeled cells can then be observed using time-lapse imaging. Time-lapse imaging has been particularly successful for studying cell behaviors that underlie the development of the cerebral cortex at late embryonic stages 1-2. Embryonic organotypic slice culture systems in brain regions outside of the forebrain are less well established. Therefore, the wealth of time-lapse imaging data describing neuronal cell migration is restricted to the forebrain 3,4. It is still not known, whether the principles discovered for the dorsal brain hold true for ventral brain areas. In the ventral brain, neurons are organized in neuronal clusters rather than layers and they often have to undergo complicated migratory trajectories to reach their final position. The ventral midbrain is not only a good model system for ventral brain development, but also contains neuronal populations such as dopaminergic neurons that are relevant in disease processes. While the function and degeneration of dopaminergic neurons has been investigated in great detail in the adult and ageing brain, little is known about the behavior of these neurons during their differentiation and migration phase 5. We describe here the generation of slice cultures from the embryonic day (E) 12.5 mouse ventral midbrain. These slice cultures are potentially suitable for monitoring dopaminergic neuron development over several days in vitro. We highlight the critical steps in generating brain slices at these early stages of embryonic development and discuss the conditions necessary for maintaining normal development of dopaminergic neurons in vitro. We also present results from time lapse imaging experiments. In these experiments, ventral midbrain precursors (including dopaminergic precursors) and their descendants were labeled in a mosaic manner using a Cre/loxP based inducible fate mapping system 6.

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

A method to generate organotypic slices from the E12.5 murine embryonic midbrain is described. The organotypic slice cultures can be used to observe the behavior of dopaminergic neurons or other ventral midbrain neurons.

The  mouse is an excellent model organism to study mammalian brain development due  to the abundance of molecular and genetic data. However, the ...

The Analysis of Purkinje Cell Dendritic Morphology in Organotypic Slice Cultures

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly oriented in the sagittal plane and develops mostly in the postnatal period in small rodents 3. Furthermore, several antibodies are available which selectively and intensively label Purkinje cells including all processes, with anti-Calbindin D28K being the most widely used. For viewing of dendrites in living cells, mice expressing EGFP selectively in Purkinje cells 11 are available through Jackson labs. Organotypic cerebellar slice cultures cells allow easy experimental manipulation of Purkinje cell dendritic development because most of the dendritic expansion of the Purkinje cell dendritic tree is actually taking place during the culture period 4. We present here a short, reliable and easy protocol for viewing and analyzing the dendritic morphology of Purkinje cells grown in organotypic cerebellar slice cultures. For many purposes, a quantitative evaluation of the Purkinje cell dendritic tree is desirable. We focus here on two parameters, dendritic tree size and branch point numbers, which can be rapidly and easily determined from anti-calbindin stained cerebellar slice cultures. These two parameters yield a reliable and sensitive measure of changes of the Purkinje cell dendritic tree. Using the example of treatments with the protein kinase C (PKC) activator PMA and the metabotropic glutamate receptor 1 (mGluR1) we demonstrate how differences in the dendritic development are visualized and quantitatively assessed. The combination of the presence of an extensive dendritic tree, selective and intense immunostaining methods, organotypic slice cultures which cover the period of dendritic growth and a mouse model with Purkinje cell specific EGFP expression make Purkinje cells a powerful model system for revealing the mechanisms of dendritic development.

We present a protocol that permits to view and to quantitatively asses the morphology of the dendritic tree of individual Purkinje cells grown in organotypic cerebellar slice cultures. This protocol is intended to promote studies on the mechanisms of Purkinje cell dendritic development.

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly ...

Organotypic Slice Cultures to Study Oligodendrocyte Dynamics and Myelination

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During embryonic and postnatal development they actively proliferate and generate myelinating oligodendrocytes. These cells have commonly been studied in primary dissociated cultures, neuron cocultures, and in fixed tissue. Using newly available transgenic mouse lines slice culture systems can be used to investigate proliferation and differentiation of oligodendrocyte lineage cells in both gray and white matter regions of the forebrain and cerebellum. Slice cultures are prepared from early postnatal mice and are kept in culture for up to 1 month. These slices can be imaged multiple times over the culture period to investigate cellular behavior and interactions. This method allows visualization of NG2 cell division and the steps leading to oligodendrocyte differentiation while enabling detailed analysis of region-dependent NG2 cell and oligodendrocyte functional heterogeneity. This is a powerful technique that can be used to investigate the intrinsic and extrinsic signals influencing these cells over time in a cellular environment that closely resembles that found in vivo.

A technique to study NG2 cells and oligodendrocytes using a slice culture system of the forebrain and cerebellum is described. This method allows examination of the dynamics of proliferation and differentiation of cells within the oligodendrocyte lineage where the extracellular environment can be easily manipulated while maintaining tissue cytoarchitecture.

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During ...

The Analysis of Neurovascular Remodeling in Entorhino-hippocampal Organotypic Slice Cultures

Ischemic brain injury is among the most common and devastating conditions compromising proper brain function and often leads to persisting functional deficits in the affected patients. Despite intensive research efforts, there is still no effective treatment option available that reduces neuronal injury and protects neurons in the ischemic areas from delayed secondary death. Research in this area typically involves the use of elaborate and problematic animal models. Entorhino-hippocampal organotypic slice cultures challenged with oxygen and glucose deprivation (OGD) are established in&#160;vitro models which mimic cerebral ischemia. The novel aspect of this study is that changes of the brain blood vessels are studied in addition to neuronal changes and the reaction of both the neuronal compartment and the vascular compartment can be compared and correlated. The methods presented in this protocol substantially broaden the potential applications of the organotypic slice culture approach. The induction of OGD or hypoxia alone can be applied by rather simple means in organotypic slice cultures and leads to reliable and reproducible damage in the neural tissue. This is in stark contrast to the complicated and problematic animal experiments inducing stroke and ischemia in vivo. By broadening the analysis to include the study of the reaction of the vasculature could provide new ways on how to preserve and restore brain functions. The slice culture approach presented here might develop into an attractive and important tool for the study of ischemic brain injury and might be useful for testing potential therapeutic measures aimed at neuroprotection.

A protocol for entorhino-hippocampal organotypic slice cultures, which allows reproducing many aspects of ischemic brain injury, is presented. By studying changes of the neurovasculature in addition to changes in the neurons, this protocol is a versatile tool to study plastic changes in neural tissue after injury.

Ischemic brain injury is among the most common and devastating conditions compromising proper brain function and often leads to persisting functional ...